Systematic identification of proteins regulated by the TOR Complex 2-dependent kinase Ypk1 in
Saccharomyces cerevisiae
By
Alexander Muir
A dissertation submitted in partial satisfaction of the
requirements for the degree of
Doctor of Philosophy
in
Molecular and Cell Biology
in the
Graduation Division
of the
University of California, Berkeley
Committee in charge:
Professor Jeremy W. Thorner, Chair
Professor David G. Drubin
Professor John Kuriyan
Professor Kevan M. Shokat
Spring 2015
Abstract
Systematic identification of proteins regulated by the TOR Complex 2-dependent kinase Ypk1 in
Saccharomyces cerevisiae
by
Alexander Muir
Doctor of Philosophy in Molecular and Cell Biology
University of California, Berkeley
Professor Jeremy W. Thorner, Chair
Abstract:
Eukaryotic plasma membranes are highly complex structures, both with respect to molecular
composition and spatial organization. This composition and organization is essential for cellular
function and must be maintained during dramatic shifts in extracellular environment that
challenge membrane homeostasis. In the model eukaryote Saccharomyces cerevisiae, signaling
by TOR Complex 2 (TORC2) and its effector kinases Ypk1 and Ypk2 is modulated in response
to various membrane stresses and cells deficient in TORC2-Ypk signaling have membrane
defects, suggesting that this signaling pathway is important for maintaining membrane
homeostasis. To understand the molecular mechanisms by which TORC2-Ypk1 signaling
regulates membrane homeostasis, I devised a three-tiered genome-wide screen to identify
substrates of Ypk kinases. This screen identified 12 novel Ypk substrates in a variety of
processes linking TORC2-Ypk signaling to membrane homeostasis including: lipid metabolism,
glycerol metabolism, peroxisome function and autophagy. I then performed detailed studies of
the regulation of identified substrates Lac1 and Lag1, components of the ceramide synthase
complex, a lipid metabolism enzyme. In response to various membrane stresses, TORC2-Ypk1
activates ceramide synthase. This activation is essential in allowing cells to survive these
stresses. Interestingly, it appears that activation of ceramide synthase promotes stress survival
not by increasing ceramide levels, but rather by preventing the accumulation of ceramide
precursors. Lastly, I characterized the TORC2-Ypk regulation of identified substrates Fps1, Gpt2
and Smp1, proteins involved in glycerol metabolism and the cellular response to hyperosmotic
shock. I show that TORC2-Ypk is a novel hyperosmotic shock responsive pathway and that
TORC2-Ypk coordinately regulates glycerol metabolism substrates to promote accumulation of
glycerol, balance osmolarity and promote cellular survival. Thus, this work has identified at least
some of the molecular mechanisms by which TORC2-Ypk signaling restores membrane
homeostasis in the face of environmental stress.
1
Table of Contents
Acknowledgements
iii
List of tables
iv
List of figures
v
List of abbreviations
vii
Chapter 1 - Introduction
Molecular composition of the eukaryotic plasma membrane
Molecular organization of the plasma membrane
Role of plasma membrane organization and composition
in cellular physiology
TORC2-Ypk signaling and maintenance of membrane homeostasis
Outstanding questions and outline of this thesis
Figures
1
1
2
Chapter 2 - Materials and methods
Construction of yeast strains
Plasmids and recombinant DNA methods
Bioinformatic prediction of Ypk1 substrates
Yeast growth assays and synthetic dosage lethality screening
Purification of Ypk1-as purification of putative Ypk substrates
Ypk1-as in vitro kinase assays
Preparation of cell extracts and immunoblotting
Analysis of sphingolipid species
In vitro ceramide synthase assay
Fluorescence microscopy of Smp1-GFP and Fps1-GFP
Measurement of STL1 promoter transcription by fluorescence
activated cell sorting (FACS)
Measurement of intracellular glycerol accumulation
Co-immunoprecipitation of Fps1 and Rgc2
Tables
12
12
12
12
13
13
14
14
15
16
17
Chapter 3 - A three-tiered screen identifies novel Ypk substrates
Introduction
Results
A three-tiered screen to identify new Ypk1 substrates
Bioinfomatic prediction of Ypk substrates
A synthetic dosage lethality screen to identify Ypk
interacting proteins
Biochemical screening of bioinformatically and genetically
predicted Ypk substrates
Discussion
29
29
29
29
29
i
3
4
7
9
17
17
17
19
30
31
32
Figures
Tables
36
39
Chapter 4 - Ypk1 phosphorylates ceramide synthase to stimulate synthesis
of comples sphingolipids
Introduction
Results
Global screen indicates Lac1 and Lag1are Ypk1 substrates
Lac1 and Lag1 are phosphorylated by Ypk1 in vivo
Lac1 and Lag1 phosphorylation increases under stress and is
required for cell survival
Calcineurin down-regulates Lac1 and Lag1 phosphorylation
Ypk1-mediated phosphorylation of Lac and Lag1 stimulate
ceramide synthase activity
Ypk1-mediated stimulation of Lac1 and Lag1 prevents
autophagy induction during TORC2-driven up-regulation
of sphingolipid biosynthesis
Is Atg21 a physiologically-relevant Ypk1 substrate?
Discussion
Figures
Chapter 5 - TOR Complex 2-Ypk is a MAPK-independent osmolarity
sensing pathway that promotes glycerol accumulation and stress survival
Introduction
Results
Smp1 is phosphorylated by Ypk kinases and this modification
influences Smp1 function and localization
Fps1 is phosphorylated by Ypk kinases and this modification
is blocked when cells are subjected to hyperosmotic stress
TORC2-Ypk regulation of Fps1 is independent of the known
Ypk-mediated phosphorylation of Fps1 promotes channel
opening and glycerol efflux
TORC2-Ypk regulation of Fps1 is independent of the
known Fps1 regulators Hog1 and Rgc1/2 and allows
cells to survive hyperosmotic stress in the absence of Hog1
Discussion
Figures
Tables
45
45
46
46
46
47
48
49
50
51
52
56
69
69
71
71
72
74
75
76
78
90
Chapter 6 - Concluding remarks
Conclusions
Future directions
Final remarks
91
91
92
94
Literature cited
96
Appendix – A study of places to walk around Berkeley and the Bay Area
118
ii
Acknowledgements
I grew up watching Total Request Live on MTV. No, no, don’t worry. I am not going to thank
Carson Daly for raising me or anything like that. I just liked the part where he’d ask audience
members to give a “shout out” to someone special. I am very happy to shout out the many people
to whom I am indebted for making my life in Berkeley happy, fulfilling and productive.
First of all, I have to thank the members of the Thorner lab for creating such an exciting place to
work. I owe much of my scientific training to Françoise Roelants, an amazing scientist whom I
am tremendously lucky to count as a friend. As a researcher, I would be pretty hopeless without
her training, and her goodwill helped me tremendously through my difficult early years in the
lab. Additionally, I have to thank the Thorner lab post-doctorates, in particular, Subu
Ramachandran, for their input into my work over the years. My fellow graduate students also
contributed tremendously to my scientific growth. In particular, Dr. Jesse Patterson’s
doggedness, technical skill, and willingness to help out were a source of inspiration. I would also
like to thank Chris Alvaro for being thoughtful, being razor sharp, and singing beautifully in the
lab late at night. What more could you ask of a colleague? I am also incredibly fortunate to have
worked with two undergraduate students, Jeff Liu and Garrett Timmons. Jeff, Garrett and I
collaborated on much of the work described here and we made an excellent team. I am also
profoundly indebted to Garrett for helping me cajole the lab into a skydiving trip.
I am grateful to the scientific mentors I have had throughout my training. My advisor Jeremy
Thorner taught me much, but two lessons stand out. First, science is not merely a body of facts,
but a community of people who collect the facts and interpret them. Jeremy taught by example
that teaching and being a good member of this community is of utmost importance. I remember
once a student came well over an hour late to an appointment with him. He was in the middle of
something else, but he stopped and made time to answer her questions. I aspire to be so caring.
Jeremy also taught me, however, that the facts do matter. The usefulness of his encyclopedic
knowledge continues to inspire me to be as well read as possible. I am also grateful to my thesis
committee members, David Drubin, John Kuriyan and Kevan Shokat. They always had
interesting ideas for other directions I could take my projects. I regret that I didn’t always have
the time to try all their suggestions out. Lastly, I am very thankful to my undergraduate advisor
Ilaria Rebay, who took a chance, asked me to work in her lab, and introduced me to research.
I have to say thank you to my amazing friends. Jeremy Amon, Alex Wu and I spent many hours
bowling, eating salads at the Sizzler, and discussing everything from contemporary music videos
to new frontiers in salads. I would not have had 1/10th the fun or be 1/10th the person I am today
(for better or worse) without them. I certainly would have never experienced children’s day at
the Chabot Gun Club. I am also extremely grateful to have found kindred adventuring spirits in
Eric Estrin, Crystal Pierce, Erik Lokensgard, Debbie Lehmann and Sarah Price. I’ll never forget
camping in the snow and rain, road trips and wanderings with all of you. If I could still request a
music video for y’all, it would be Kelly Clarkson’s “My Life Would Suck Without You”.
Finally, I thank Mom, Dad, Faith, Alyssa, Caelin, Jeremy, Grandma, Ben, Christine, Emerson
and my favorite aunt Dixie. It is impossible to thank you enough for all that you have done for
me. I won’t try to put anything in words, other than to offer the following work as thanks to you.
iii
List of tables
Table 2.1
Yeast strains used in this study
19
Table 2.2
Plasmids used in this study
24
Table 3.1
Known Ypk1 substrates and potential substrates predicted
by MOTIPS listed under GO Slim terms
39
Synthetic dosage lethality of various Fps1 mutants in
ypk1-as ypk2Δ cells
90
Table 5.1
iv
List of figures
Figure 1.1
Sequence alignment of Ypk1 and Ypk2
9
Figure 1.2
TORC2-Ypk signaling pathway
10
Figure 1.3
Initially identified Ypk substrates give insight into TORC2-Ypk
regulation of membrane homeostasis
11
Figure 3.1
A three-part screen to identify likely Ypk1 substrates
36
Figure 3.2
Identified Ypk substrates
38
Figure 4.1
Lac1 and Lag1 are components of the ceramide synthase
enzyme complex
56
Genetic and biochemical screening indicates Lac1 and Lag1
are Ypk1 substrates
57
Figure 4.3
Ypk1 phosphorylates Lac1 and Lag1 at S23 and S24 in vivo
58
Figure 4.4
Enhanced Ypk1 phosphorylation of Lac1 and Lag1 under
sphingolipid and heat
59
Activation of calcineurin leads to rapid dephosphorylation
of Lac1 and Lag1 without affecting Ypk1 function
60
Ypk1 phosphorylation of Lac1 and Lag1 stimulates ceramide
synthase activity
61
Failure of Ypk1 to upregulate ceramide synthase causes LCBP
accumulation that triggers autophagy
63
Figure 4.2
Figure 4.5
Figure 4.6
Figure 4.7
Figure 4.8
TORC2-Ypk1 signaling globally activates sphingolipid synthesis,
selectively directs flux toward ceramide metabolites, and
preventsLCBP cross-talk to the autophagy pathway
65
Figure 4.9
Loss of Ypk activity or putative Ypk phosphorylation of
Atg21 does not appear to prevent CVT or amino acid starvation
induced autophagy
67
Figure 5.1
Gpt2 is not phosphorylated by Ypk or other basophilic kinases
78
Figure 5.2
Smp1 is phosphorylated at predicted Ypk sites and
phosphorylation of these residues is important for Smp1 function 79
v
Figure 5.3
Figure 5.4
Figure 5.5
Figure 5.6
Figure 5.7
Figure 5.8
Changes in Fps1 dosage does not affect TORC2 of Pkh
phosphorylation of Ypk1
Ypk in response to TORC2 signaling phosphorylates
Fps1 at three sites in vivo
81
82
TORC2 phosphorylation of Ypk1 decreases rapidly in response
to hyperosmotic shock resulting in Fps1 dephosphorylation
84
Phosphorylation of Fps1 by TORC2-Ypk increases channel
transport activity
85
TORC2-Ypk regulates Fps1 independently of known
regulators Hog1 and Rgc1/2
87
Saccharomyces cerevisiae utilizes two independent
osmosensing systems to rapidly increase intracellular
glycerol upon hyperosmolarity
89
vi
List of abbreviations
1-NM-PP1
3-MB-PP1
ATP
BSA
CoA
CVT
DAPI
DHAP
DHS
DHS-1-P
DMSO
DNA
DTT
EDTA
EGTA
GFP
GST
IPC
IPTG
LC
LCB
LCB-1-P
MAPK
MIPC
MOTIPS
MS/MS
OD600
PCR
PHS
PHS-1-P
PQD
SC
SEM
SDL
SDS-PAGE
SPOTS
TAP
TCA
TLC
TOR
TORC1
TORC2
YP
1-(1,1-dimethylethyl)-3-(1-naphthalenylmethyl)-1H-pyrazolo[3,4-d]pyrimidin-4amine
1-(tert-Butyl)-3-(3-methylbenzyl)-1H-pyrazolo[3,4-d]pyrimidin-4-amine
Adenosine triphosphate
Bovine serum albumin
Co-enzyme A
Cytoplasmic to vacuole pathway
4',6-diamidino-2-phenylindole
Dihydroyacetone phosphate
Dihydrosphingosine
Dihydrosphingosine-1-phosphate
Dimethylsulfoxide
Deoxyribonucleic acid
Dithiothreitol
Ethylenediaminetetraacetic acid
Ethylene glycol tetraacetic acid
Green fluorescent protein
Glutathione S-transferase
Inositol phosphorylceramide
Isopropyl β-D-1-thiogalactopyranoside
Liquid chromatography
Long chain base
Long chain base-1-phosphate
Mitogen activated protein kinase
Mannosylinositol phosphorylceramide
Motif analysis pipeline
Tandem mass spectrometry
Optical density at 600 nm
Polymerase chain reaction
Phytosphingosone
Phytosphingosone -1-phosphate
Pulsed-Q dissociation
Synthetic complete medium
Standard error of the mean
Synthetic dosage lethality
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis
Serine palmitoyl-CoA transferase complex
Tandem affinity purification
Trichloroacetic acid
Thin layer chromatography
Target of rapamycin
Target of rapamycin complex 1
Target of rapamycin complex 2
Yeas extract peptone medium
vii
Chapter 1: Introduction
Molecular composition of the eukaryotic plasma membrane
All cells are bound by a ~30 Å thick hydrophobic membrane that defines the cell’s boundary,
ensures cellular integrity and mediates interactions between the individual cell and its
environment. In eukaryotic cells, this plasma membrane is a complex supramolecular assembly
composed of thousands of different chemically diverse lipid and protein species (Guan and
Wenk, 2006; Shevchenko and Simons, 2010). It is clear that the plasma membrane and its
compositional complexity is important to eukaryotic organisms, as ~5% of the protein coding
capacity of the human genome is dedicated to lipid metabolism (van Meer et al., 2008) and
~26% of the genome codes for membrane proteins (Fagerberg et al., 2010). Below I will outline
the major molecular classes that compose the plasma membrane, before moving to an overview
of how these species are spatially organized in biological membranes.
Lipids are the major structural component of biological membranes and they represent ~50% of
eukaryotic membrane mass (Cooper and Hausman, 2013). Membrane lipids are of three major
classes: glycerophospholipids, sphingolipids and sterols (van Meer et al., 2008). The main lipid
class of eukaryotic membranes is glycerophospholipids; ~70-80% of membrane phospholipids
are of this type. These lipids consist of a hydrophobic diacylglycerol moiety, containing fatty
acyl chains of various lengths and degrees of unsaturation. Various hydrophilic head groups can
be attached to diacylglycerol giving rise to the large variety of glycerophospholipids observed in
eukaryotic membranes. The most common glycerophospholipids observed in cellular membranes
are phosphatidylcholine (~40% membrane phospholipid by mole), phosphatidylethanolamine
(~20%), phosphatidylserine (~10%) and various phosphatidylinositols (~2-5%) (van Meer et al.,
2008).
Sphingolipids are the second major membrane lipid class and are structurally distinct from
glycerophospholipids. The hydrophobic sphingolipid backbone is ceramide, a molecule
composed of a long chain (sphingoid) base that is amide linked to a very long chain saturated
fatty acid. As with glycerophospholipids, various polar head groups (typically composed of
carbohydrates) can be appended to ceramide to form the full complement of cellular
sphingolipids (Futerman and Hannun, 2004). Sphingolipids typically represent ~25% of all
plasma membrane lipids (van Meer et al., 2008).
Lastly, sterols are a third structurally different class of lipid. Sterols are planar molecules
composed of multiple carbon rings resulting from cyclization of squalene derived from the
mevalonate biosynthetic pathway (Buhaescu and Izzedine, 2007). They are, with the exception
of a hydroxyl group, non-polar molecules and they are the major non-polar lipid species in
eukaryotic membranes (van Meer et al., 2008).
In addition to lipid species, thousands of various membrane proteins also reside in the membrane
(Fagerberg et al., 2010). These proteins play diverse biological roles, primarily acting as
receptors, transporters and enzymes (Sachs and Engelman, 2006; Almén et al., 2009). Thus,
proteins are also a major component and source of compositional complexity in membranes. It is
also becoming clear that these molecules are not merely passengers in the lipid membrane where
1
they perform functions, but many of these proteins also play important roles in generating and
maintaining membrane organization, as discussed below.
Molecular organization of the eukaryotic plasma membrane
In recent decades, it has become increasingly clear that the thousands of lipid and protein species
that compose the plasma membrane are not homogenously mixed, as often illustrated in textbook
models of the membrane, for example, as when depicting the fluid mosaic model (Singer and
Nicolson, 1972). Rather, the plasma membrane is a highly structured organelle. Localization and
organization of specific lipid species has been observed along two axes: laterally, with respect to
the plane of the bilayer; and, trans-bilayer, with respect to each of the two leaflets.
In the lateral axis, the physical properties of sphingolipid and sterol species allows them to
cluster away from glycerophospholipids and form local phase separations termed “lipid rafts”
(Simons and van Meer, 1988). Lipid rafts were initially defined as small (10-200 nm diameter)
domains of the membrane enriched in sphingolipids and sterols. The very long saturated acyl
chains of membrane sphingolipids was presumed to lead to thickening of membrane in these
domains (Gandhavadi et al., 2002), which could in turn lead to recruitment of certain membraneembedded and membrane-associated proteins, especially GPI-anchored protein or Spalmitoylated proteins (Brown and Rose, 1992; Zacharias et al., 2002; Foster et al., 2003). The
function and even the existence of lipid-mediated membrane domain organization in live cells
has been controversial (Simons and Sampaio, 2011). However recent advances in resolution and
dynamics of live cell microscopy has revealed the presence of such structures in cells (Eggeling
et al., 2009).
In addition to lipid-mediated lateral membrane organization, cells use other protein-driven
processes to generate membrane organization. Local restriction of lipids and membrane proteins
by cortical cytoskeleton “fences” has been observed to form membrane domains (Chichili and
Rodgers, 2009). Membrane-associated proteins also generate lateral membrane organization. For
example, cytokinetic septin proteins alter lipid diffusion, generating regions of distinct lipid
composition (Beise and Trimble, 2011; Clay et al., 2014) and BAR domain-containing proteins
bend membranes forming stable domains (Ziółkowska et al., 2011; Zhao et al., 2013). Integral
membrane proteins themselves generate lateral membrane organization. Caveolin proteins form
sphingolipid- and sterol-enriched domains, termed caveolae (Monier et al., 1996). In the context
of integral membrane proteins, both protein-protein and protein-lipid interactions have been
observed to generate domains (Douglass and Vale, 2005). Lastly, cells also use directed vesicular
transport to form stable lateral membrane domains (Dustin and Groves, 2012).
In addition to lateral membrane organization, bilayer asymmetry is also a well established
property of all biological membranes and has been demonstrated in many cell types (Devaux,
1991). Sphingolipids and phosphatidylcholine predominate in the extracellular leaflet of the
membrane bilayer, whereas other glycerophospholipids are enriched in the inner leaflet (van
Meer et al., 2008). Lipids are able to spontaneously “flip” from one leaflet to the other, although
kinetically this is a slow process (Kornberg and McConnell, 1971). Therefore, to maintain
bilayer asymmetry, cells use a variety of membrane-embedded enzymatic machines (e.g.
"flippases, floppases and scramblases") to translocate lipids from one bilayer leaflet to the other
2
(Sharom, 2011).
Lastly, beyond the structuring and ordering of lipid species in the plasma membrane, there is also
dramatic ordering of the protein content of the plasma membrane. Oligomeric membrane-binding
proteins have been observed to form microdomains (Monier et al., 1995; Ziółkowska et al.,
2011) and represent one such class of membrane protein organization. Additionally, signaling
proteins have been observed to cluster at “signaling synapses” and play important role in cellular
signaling events (Dustin and Groves, 2012). Recently, high-throughput microscopy experiments
have found that membrane proteins in general are not diffusely localized in the membrane.
Rather they are found in multiple non-overlapping domains (Spira et al., 2012). Thus, distinct
clustering of lipids and proteins leads to a patchwork organization of the membrane - a "mosaic
tile" model (Engelman, 2005) rather than the rapid diffusion and homogeneity of the previously
proposed "fluid mosaic" model (Singer & Nicolson, 1972).
Role of plasma membrane organization and composition in cellular physiology
It is increasingly being appreciated that the above described membrane organization is crucial for
determining the physical properties of biological membranes and thus their function. Studies
have found that loss of bilayer asymmetry reduces the mechanical stability of cellular
membranes (Manno et al., 2002) and leads to dramatic defects on the ability of membranes to
bend and bud off vesicles (Devaux, 1991; Williamson and Schlegel, 1994; Devaux et al., 2008).
Accordingly, flippase function and regulation has been found to be important for aspects of
cellular physiology including intracellular membrane trafficking (Hua et al., 2002; Fairn et al.,
2011), polarized cell growth (Saito et al., 2007; Das et al., 2012), cell signaling (Sartorel et al.,
2015), and cytokinesis (Roelants et al., 2015). Recent work demonstrates that absence of a
flippase causes mislocalization of a Drosophila olfactory receptor (Ha et al., 2014). At the
organismal level, loss of flippases and membrane asymmetry results in a number of pathologies
(Ikeda et al., 2006), including profound defects in lymphocyte development (Clark, 2011). Thus,
asymmetric lipid organization in the bilayer axis is of functional importance.
In addition to the functional roles ascribed to bilayer asymmetry, lateral membrane asymmetry is
increasingly understood to have functional roles in cellular physiology as well. This lateral
compartmentation of the plasma membrane has been well documented in fungi and plants
(Malinsky et al., 2010; Malinsky et al., 2013). In animal cells, lateral membrane microdomains
are important for propagation of extracellular signals across membranes (Dustin and Groves,
2012), examined particularly thoroughly in the case of Ras (Schmick et al. 2015; Zhou &
Hancock, 2015), and membrane trafficking processes (Simons and Sampaio, 2011). Recent
studies have found that just displacing membrane proteins from their appropriate microdomains,
even while leaving them intact in the plasma membrane can drastically affect membrane protein
function (Berchtold et al., 2012; Spira et al., 2012). Thus, lateral membrane organization does
indeed seem to play important roles in various cellular processes.
Lastly, membrane composition is also incredibly important for determining function. Classically,
in order to understand the functional role of certain lipids in cellular function, cells defective in
biosynthesis of certain lipids (either by genetic manipulation or pharmacological inhibition) were
phenotypically compared to appropriate control cells. For example, by studying mutants
3
defective in sphingolipid metabolism, it was found that sphingolipids are essential for
endocytosis (Zanolari et al., 2000). This experimental modality has successfully identified
functional roles for certain lipid classes and by this approach composition has been shown to
affect membrane fluidity, the function of membrane proteins and membrane trafficking (Spector
and Yorek, 1985). Recent advances in mass spectrometry based analytical chemistries have made
it possible to quantitatively examine all membrane lipids, termed the lipidome (Ejsing et al.,
2009; Brügger, 2014). These techniques have been used to identify and quantify dynamic
changes to the cellular lipidome in response to various stimuli, during transit through the cell
division cycle, and in various mutant backgrounds (Atilla-Gokcumen et al., 2014; da Silveira
Dos Santos et al., 2014; Casanovas et al., 2015). Excitingly, these unbiased lipidomics
approaches have identified dynamic changes in lipid composition that are required for various
cellular processes, such as a requirement for glycosphingolipid biosynthesis in cytokinesis
(Atilla-Gokcumen et al., 2011; Atilla-Gokcumen et al., 2014) and a requirement for increased
cardiolipin synthesis during the diauxic shift to mitochondrial respiration in the yeast
Saccharomyces cerevisiae (Casanovas et al., 2015). These recent findings from lipidomic
analysis suggest that dynamic changes in membrane composition are an important component of
the cellular response to many stimuli and during developmental processes, and further that many
new functional roles for lipids in these processes remain to be discovered.
TORC2-Ypk signaling and maintenance of membrane homeostasis
Given that membrane composition and organization (collectively, I will refer to these properties
as 'membrane status') is of vital importance to cellular function, and that cells appear to
dynamically alter their membranes in response to environmental stimuli, it is important to
understand what mechanisms cells use to sense membrane status and take action to restore
functional homeostasis.
The model eukaryote S. cerevisiae has proven useful in identifying a number of signaling
pathways that respond to various types of membrane stress. Using this organism, mitogenactivated protein kinase (MAPK) pathways have been shown to be important in eliciting the
responses required by the cell to survive a number of different stressful conditions. Signaling
pathways that activate the MAPK Hog1 are important for viability under hyperosmotic
conditions, a stress that leads to cell shrinkage and membrane compression (Hohmann, 2015;
Saito and Posas, 2012). The MAPK Slt2/Mpk1 downstream of Pkc1 signaling is important for
survival under hypotonic conditions and after physical wounding of the cell wall (Levin, 2011;
Kono et al., 2012). These signaling pathways are conserved across eukaryotes [mammalian
ortholog of Hog1 is MAPK p38 and the mammalian ortholog of Slt2 is MAPK Erk5], and
mammalian cells similarly use MAPKs (Zarubin and Han, 2005) and PKC (Togo et al., 2000;
Liu et al., 2003; Hermoso et al., 2004; Bullard et al., 2007) to respond to osmolarity change or
membrane stress.
Another less well characterized, yet important, membrane stress-responsive signaling pathway
controls two highly related effectors: protein kinases Ypk1 and Ypk2 (formerly Ykr2). As
assessed by genetic analysis, these two enzymes appear to carry out vital functions in membrane
homeostasis. Cells lacking both Ypk1 and Ypk2/Ykr2 are inviable (Chen et al., 1993; Roelants et
al., 2002). Phenotypic characterization of cells deficient in Ypk activity revealed that they have
4
dramatic defects in endocytosis, fail to couple cell wall growth to membrane growth, and are
dramatically sensitive to inhibitors of sphingolipid biosynthesis (Sun et al., 2000; deHart et al.,
2002; Roelants et al., 2002; Schmelzle et al., 2002; Roelants et al., 2004; Roelants et al., 2010).
All of these phenotypes suggested that Ypk1 and Ypk2 play an important role in maintaining
membrane homeostasis.
Ypk1 and Ypk2 are very similar proteins (especially in their kinase domain) (see Fig. 1.1) and
are functionally semi-redundant (Chen et al., 1993; Roelants et al., 2002; Roelants et al., 2004).
These paralogs likely arose from a wholesale genome duplication that occurred during the
evolutionary path that led to modern-day S. cerevisiae (Byrne and Wolfe, 2005). Ypk1 and Ypk2
are members of the AGC family of protein kinases, one of the conserved groups in this enzyme
super-family (Pearce et al., 2010). Mammalian SGK1/SGK (serum-and glucocorticoid-induced
protein kinase) is the sequence and functional ortholog of yeast Ypk1 (Casamayor et al., 1999).
Activation of the catalytic activity of AGC kinases requires phosphorylation of a conserved Thr
in their activation loop ("T-loop"). This modification causes conformational changes that make
the enzyme competent to catalyze phosphoryl transfer from ATP to a target protein (Pearce et al.,
2010). The upstream activating protein kinases that mediate T-loop phosphorylation of Ypk1 and
Ypk2 are the protein kinases Pkh1 and Pkh2. Mammalian PDK1 (2-phosphoinositide-dependent
protein kinase) is the sequence and functional ortholog of Pkh1 and Pkh2 (Casamayor et al.,
1999; Roelants et al., 2002; Roelants et al, 2004). Pkh1 and Pkh2 localize in cortical puncta at
the cell periphery (Roelants et al., 2002) in tight association with two PtdIns4,5P2-binding
proteins (Pil1 and Lsp1) that are major components of plasma membrane-bound complexes,
referred to as eisosomes (Walther et al., 2006; Walther et al., 2007). Thus, Pkh1 and Pkh2 could
potentially serve as sensors of some aspect of membrane status.
In addition to T-loop phosphorylation, AGC kinases can be further activated by phosphorylation
of conserved sites - the so-called turn and hydrophobic motifs sites - in the carboxy-terminal
extension downstream of their catalytic domain (Pearce et al., 2010). Complexes containing the
atypical protein kinase TOR (target of rapamycin) are responsible for phosphorylation of AGC
kinases at these C-terminal sites (Pearce et al., 2010). In yeast, there are two TOR paralogs, Tor1
and Tor2 (Heitman et al. 1991), whereas in mammalian cells there is a single gene product
(mTOR/RAFT/RAPT/FRAP) (Sabatini et al., 1994). One of the TOR-containing complexes in
yeast, TORC1, contains either Tor1 or Tor2, as well as other subunits diagnostic of this protein
ensemble that ensure its localization to the cytosolic face of the lysosome and dictate its substrate
specificity (Loewith et al., 2002; Wedaman et al., 2003; Reinke et al., 2004; Loewith & Hall,
2011). The other TOR-containing complex in yeast, TORC2, contains only Tor2, as well as other
subunits diagnostic for this assembly that direct its localization to the plasma membrane and
impose its substrate specificity (Loewith et al., 2002; Wedaman et al., 2003; Reinke et al., 2004;
Loewith & Hall, 2011). Mammalian cells have highly homologous TORC1 and TORC2
complexes that contain mTOR (Kim et al., 2002; Sarbassov et al., 2004; Huang & Manning,
2009; Betz & Hall, 2013).
TORC1 and TORC2 regulate different aspects of cell physiology. TORC1, but not TORC2, is
specifically sensitive to inhibition by the macrolide rapamycin and its structural relatives
("rapalogs") (Benjamin et al., 2011; Chiarini et al., 2015), which has allowed for extensive
characterization of the cellular functions of TORC1 (Loewith et al., 2002; Loewith & Hall, 2011;
5
Albert & Hall, 2015). TORC1 is most active when nutrients, especially amino acids, and in
particular Leu and Gln (Jewell et al. 2015), are available in both yeast (Panchaud et al., 2013;
Chantranupong et al., 2015) and animal cells (Bar-Peled & Sabatini, 2014; Efeyan et al., 2015)
and, in animal cells, when cells have been concomitantly stimulated with a growth factor (e.g.
EGF, PDGF, insulin, etc.) that activates PtdIns 3-kinase (Huang & Manning, 2009; Chiarini et
al., 2015). Active TORC1 phosphorylates targets that promote anabolic processes, such as
ribosome biogenesis, protein synthesis, pyrimidine biosynthesis and lipogenesis, while inhibiting
catabolic processes, such as autophagy and lysosome biogenesis (Laplante and Sabatini, 2012;
Howell et al. 2013; Albert & Hall, 2015). In contrast, there has not been (until recently, see
Kliegman et al., 2013; Rispal et al., 2015), a specific inhibitor of TORC2; hence, our
understanding of the physiological functions of this complex is more rudimentary. What is
known is that TORC2 mediates phosphorylation of AGC kinases distinct from those
phosphorylated by TORC1 (Pearce et al., 2010).
In yeast, phosphorylation of Ypk1 and Ypk2 at their C-terminal turn and hydrophobic motifs is
mediated by TORC2 and phosphorylation at these sites does considerably increase their catalytic
activity (Kamada et al., 2005; Roelants et al., 2011). In yeast, tor1∆ mutants are viable, whereas
tor2∆ are inviable, indicating that TORC2 function is essential. Strikingly, the absence of
TORC2 activity can be rescued by alleles of Ypk1 or Ypk2 that make these enzymes
constitutively hyperactive, suggesting that the sole function of TORC2 that is essential for cell
viability is activation of Ypk1 and Ypk2 (Kamada et al., 2005; Roelants et al., 2011. Because
Ypk1 and Ypk2 are the effectors that execute the essential function(s) of TORC2, its stands to
reason, therefore, that characterization of the substrates of Ypk1 and Ypk2 should shed
considerable light on those aspects of cellular physiology that are under TORC2 control. In this
regard, and given the phenotypes of Ypk-deficient cells, initial research on the roles of Ypk1 and
Ypk2 centered on attempts to understand how membrane status is sensed and relayed to the Ypk
kinases. Based on in vitro biochemical assays, two groups (Friant et al., 2001; Liu et al., 2005)
reported that the activity of Pkh1 and Pkh2 could be stimulated by addition of long chain
sphingoid base (mainly, phytosphingosine in yeast), suggesting that the Pkh kinases might serve
as sensors of plasma membrane composition that could, in turn, modulate Ypk kinase activity
appropriately to adjust the rates of the reactions necessary to maintain homeostasis. However,
more recent results demonstrated that, in vivo, T-loop phosphorylation of the Ypk kinases by the
Pkh kinases is not subject to control by sphingolipids or their precursors (Roelants et al., 2010).
Therefore, it appears that other Ypk kinase regulators must be the sensors of membrane status,
and TORC2 clearly represents one of the most likely candidates for that role.
Indeed, more recent work has found that TORC2 activity is sensitive to a variety of membrane
stresses. Three types of membrane stress - inhibition of sphingolipid synthesis (Roelants et al.,
2011), heat shock (Sun et al., 2012), and hypotonic conditions (or mechanical stretch) (Berchtold
et al., 2012) - clearly elevate TORC2 function. In these cases, change in TORC2 activity was
measured indirectly by monitored the status of Ypk phosphorylation at its hydrophobic motif, by
assessing the specific activity of immunoprecipitated Ypk1, and/or by evaluating the
phosphorylation state of downstream Ypk1 substrates (e.g., Orm2). In the case of hypotonic
stress, it appears that the increase in TORC2 activity is due to dissociation of two TORC2
components (Slm1 and Slm2) from the rest of the TORC2 complex and their sequestration into a
separate plasma membrane microdomain (Berchtold et al., 2012; Niles et al., 2012). Conversely,
6
upon exposure of cells to hyperosmotic conditions, our laboratory has found, in collaborative
studies, that TORC2 activity dramatically decreases (Lee et al., 2012b), although the precise
mechanism by which this rapid and marked reduction in apparent activity occurs has not yet
been determined.
In summary, although T-loop phosphorylation by the Pkh kinases is necessary to license Ypk
activity, it appears that TORC2 senses and responds to membrane stress and conveys this
information to Ypk1. Furthermore, the downstream outputs under the control of TORC2-Ypk1
are important for maintaining membrane homeostasis, as judged by the fact that cells defective in
TORC2-Ypk signaling exhibit dramatic membrane-related phenotypes. Pkh- and TORC2depndent Ypk signaling and its membrane phenotypes are schematized in Figure 1.2.
Outstanding questions and outline of this thesis
These findings raised an obvious and important question - what processes does TORC2-Ypk
signaling modulate in order to maintain membrane homeostasis in response to stressful stimuli?
In contrast to the hyperosmotic stress responsive Hog1 MAPK pathway and the hyposmotic
stress responsive Rho1-dependent Pkc1 pathway (sequence ortholog of mammalian RHOactivated PKN2/PRK2), where the downstream functional outputs are largely known (Levin,
2011; Saito and Posas, 2012; Hohmann, 2015), relatively little was understood about the
downstream responses of TORC2-Ypk to membrane stress. However, two approaches provided
initial insight about the downstream effectors of TORC2-Ypk stress-induced signaling. The first
tactic was a genetic approach. To find factors whose full activity requires Ypk-mediated
phosphorylation, a selection for dosage suppressors of the temperature sensitivity of a ypk1-ts
ypk2∆ strain was conducted (Roelants et al., 2002), which yielded plasmids containing the SMP1
gene, which encodes a MADS box-family transcription factor that I have now demonstrated (see
Chapter 5) is an authentic in vivo substrate of Ypk1. Conversely, to find factors whose activity is
inhibited by Ypk-mediated phosphorylation, a selection for transposon insertion mutations that
suppressed the temperature sensitivity of ypk1-ts ypk2∆ cells was conducted (Roelants et al.,
2002). This screen recovered Orm2, which is an inhibitor of the first enzyme unique to
sphingolipid biosynthesis, L-serine:palmitoyl-CoA acyltransferase comples (SPOTS) (Breslow et
al., 2010). Subsequently, it was found that Ypk1 phosphorylates and inhibits the Orm proteins,
and thus TORC2-Ypk1 signaling increases the rate of formation of the long-chain base precursor
(phytoshingosine) to yeast sphingolipids (Berchtold et al., 2012; Roelants et al., 2011). Thus,
increased TORC2-Ypk1 signaling in response to sphingolipid depletion or membrane stretch
allows cells to increase sphingolipid production to manage these stresses (Berchtold et al., 2012;
Roelants et al., 2011). A more recent chemical genetic screen in which a small molecule
inhibitable tor2-as allele was systematically introduced to a collection of yeast mutants deleted
for single genes similarly found genetic interactions between TORC2 and sphingolipid
biosynthetic genes (Kliegman et al., 2013). This screen also found intriguingly that TORC2
function is required for the pentose phosphate pathway that generates high energy metabolites
7
needed for nucleotide and lipid biosynthesis. Thus, genetic screening has proven useful in
identifying functionally important Ypk substrates and TORC2 regulated processes.
Another approach for identifying downstream effectors of TORC2-Ypk1 signaling was
biochemical. Use of synthetic peptides indicated that Ypk1 has a rather stringent phosphoacceptor site sequence preference (Casamayor et al., 1999), which was fully confirmed wben
much more extensive combinatorial peptide libraries were employed (Mok et al., 2010). By
scanning the yeast proteome for proteins with occurrences of this motif that also had some
connection to plasma membrane physiology, a candidate substrate emerged, namely Fpk1 (and
its paralog Fpk2), a protein kinase responsible for activation of plasma membrane-localized
aminophospholipid flippases (Nakano et al. 2008). Indeed, it was then demonstrated that Ypk1
phosphorylates and negatively regulates Fpk1 both in vitro and in vivo (Roelants et al., 2010).
Thus, in addition to stimulating production of sphingolipid precursor, TORC2-Ypk fine-tunes the
glycerophospholipid composition of the inner leaflet of the membrane bilayer. Similarly, based
in its phosphoacceptor motif, it was found that Ypk1 also phosphorylates and inhibits one of the
two isoforms (Gpd1) of glycerol-3-phosphate dehydrogenase, a primary source of glycerol-3-P
for production of glycerophospholipids (Lee et al., 2012b), but also, when dephosphorylated, a
source of the glycerol that yeast cells produce as a innocuous osmo-protectant when subjected to
hyperosmotic conditions (Hohmann, 2009). As discussed above, we have found that TORC2
activity is markedly decreased during hyperosmotic shock, which prevents Ypk1-mediated
inhibition of Gpd1, thereby increasing synthesis of glycerol-3-P, which can be dephosphorylated
to produce the glycerol osmolyte that yeast cells accumulate as a means to counteract the
increase in external osmotic pressure (Lee et al., 2012b).
Thus, by identifying only a handful of Ypk substrates (Fpk1, Fpk2, Orm1, Orm2 and Gpd1), a
great deal was learned at the mechanistic level about how Ypk function maintains membrane
homeostasis downstream of stimuli that both increase and decrease TORC2 activity. This degree
of progress is outlined in Figure 1.3. Nonetheless, an important outstanding question remains what other substrates in the cell are regulated by Ypk action? I felt that identification of
previously uncharacterized substrates would undoubtedly give further insight not only about how
TORC2-Ypk signaling is connected to membrane biology, but about those essential aspects of
cell physiology that are under the control of TORC2. Thus, the main aim of my doctoral
dissertation research was to develop a genome-wide strategy to systematically identify and
characterize new Ypk kinase substrates.
As described in this thesis, I devised a novel approach towards matching substrates with cognate
kinases and applied it to identify Ypk1 substrates (Chapter 3). I then proceeded to characterize in
depth a number of the candidate Ypk1 substrates that I pinpointed by applying my methodology.
Chapter 4 documents TORC2-Ypk-mediated regulation of Lac1 and Lag1, conserved catalytic
subunits of the ceramide synthase complex, an enzyme central for the production of
sphingolipids. Finally, in Chapter 5, I present studies showing that TORC2-Ypk regulates Fps1
(an aquaglyeroporin) and Smp1 (a transcription factor), two substrates involved in the cellular
response to hyperosmotic shock, which led me to the discovery that TORC2-Ypk signaling is a
novel MAPK-independent pathway that cells use to survive the challenge of hyperosmotic stress.
8
Figures
Fig. 1.1: Sequence alignment of Ypk1 and Ypk2. Primary sequence diagrams of Ypk1 and
Ypk2 are shown with regions of significant sequence similarity shown. Darker coloring indicates
higher sequence similarity. The percent identity between each region is indicated.
9
Fig. 1.2: TORC2-Ypk signaling pathway. Diagram outlines known relationships between
TORC2 [complex members and interactions adopted from (Loewith et al., 2002; Wullschleger et
al., 2005)], Pkh and Ypk kinases. Downstream of Ypk, membrane related phenotypes that are
observed in Ypk mutants are shown.
10
Fig. 1.3: Initially identified Ypk substrates give insight into TORC2-Ypk regulation of
membrane homeostasis. Diagram of the TORC2-Ypk signaling pathway from Fig. 1.1 is
shown. However, the known substrates of Ypk kinases are now shown. Arrows indicate
interactions that increase activity, while blunted lines indicate interactions that decrease activity
of the indicated protein. Identification of these 5 substrates gives mechanistic insight into the
membrane phenotypes associated with loss of TORC2-Ypk signaling. Identification of the
flippase activating kinases Fpk1/2 as Ypk1 substrates mechanistically links TORC2-Ypk
signaling to membrane asymmetry. Identification of TORC2-Ypk regulation of the Orm1/2
proteins , negative regulation of the SPOTS complex, implicates TORC2-Ypk as an activator of
sphingolipid metabolism. This explains why Ypk deficient strains are sensitive to sphingolipid
biosynthetic inhibitors. Lastly, identification of Gpd1 as a Ypk substrate links TORC2-Ypk
signaling to osmotic induced glycerol metabolism, an essential feature of the hyperosmotic shock
response in S. cerevisiae. This provides mechanistic insight to the osmotic shock sensitivity
observed in Ypk deficient strains.
11
Chapter 2: Materials and Methods
Construction of yeast strains
All S. cerevisiae strains used in this study are listed in Table 2.1. Strains were constructed using
standard yeast genetic manipulations (Burke et al., 2005). For all strains constructed, integration
of the desired DNA fragment into the correct genomic loci was confirmed by PCR using an
oligonucleotide complementary to the integrated DNA fragment and an oligonucleotide
complementary to genomic sequence at least 150 bases away from the integration site. After
confirmation that the DNA fragment was integrated at the correct locus, additional PCRs were
performed to generate fragments covering the integrated DNA fragment. These reaction products
were sequenced to confirm the DNA fragment introduced into the genome had the correct
sequence.
Plasmids and recombinant DNA methods
All plasmids used in this study (except the library of PGAL1-based overexpression plasmids for
synthetic dosage lethality (SDL) screening) are listed in Table 2.2. All plasmids were constructed
and progated in E. coli using standard laboratory methods (Green and Sambrook, 2012) or by
Gibson assembly (Gibson et al., 2009) using the Gibson Assembly Master Mix Kit according to
manufacturers specifications (New England Biolabs). For SDL screening, the entire open reading
frame of each predicted and known Ypk1 substrate was amplified by PCR from BY4741
genomic DNA and ligated into the multiple cloning site of YCpLG (CEN, PGAL1, LEU2),
generating a vector allowing galactose-inducible overexpression of each substrate. All constructs
generated in this study were confirmed by sequence analysis covering all promoter and coding
regions in the construct.
Bioinformatic prediction of Ypk1 substrates
A Nx20 position weight matrix defining Ypk1 phosphoacceptor site specificity was made
merging previously published data sets defining Ypk1 peptide substrate selectivity (Casamayor
et al., 1999; Mok et al., 2010). This position weight matrix was then used with MOTIPS (Lam et
al., 2010) to identify proteins with likely phosphorylated occurrences of this motif. Yeastmine
(Balakrishnan et al., 2012) was used to identify all S. cerevisiae genes that showed genetic
interactions with Ypk1, Ypk2, any component of the sphingolipid biosynthetic pathway or any
component of known Ypk1 regulators (TORC2 and protein phosphatase 2 A). Genes that showed
significant growth phenotypes on myriocin, aureobasidin A and caspifungin (all compounds that
cause severe growth phenotypes in ypk1Δ strains) were identified from the literature
(Hillenmeyer et al., 2008). Lastly, MOTIPS predicted Ypk1 phosphorylation sites were
compared to Phosphogrid (Sadowski et al., 2013) to identify those sites known to be
phosphorylated in vivo. To be considered a potential Ypk1 substrate in this study, a protein had
to have a myriocin, aureobasidin or caspofungin phenotype or a Yeastmine identified genetic
interaction and: (a) 4 or more MOTIPS predicted sites, (b) 3 sites with at least one above a
MOTIPS likelihood score of 0.7 or a Phosphogrid identified site or (c) 1-2 sites with a MOTIPS
likelihood score of 0.7 and a Phosphogrid identified site. I also considered for further analysis a
limited number of proteins that did not meet these criteria, but which still contained Ypk1 motifs.
12
These are indicated in Table 3.1.
Yeast growth assays and synthetic dosage lethality screening
For SDL screening, yAM135-A and BY4741 strains were both transformed with each SDL
plasmid. Transformants were then cultured overnight in SC medium containing 2% raffinose and
0.2% sucrose. Ten-fold serial dilutions of overnight cultures starting from A600 nm = 1.0 were then
made in sterile water and spotted onto SC solid medium with 2% galactose (to induce protein
expression) or 2% dextrose (no protein expression). These plates also contained 1:1000 DMSO,
1 μM 3-MB-PP1 or 2 μM 3-MB-PP1 to inhibit to varying degrees Ypk1-as kinase activity in
yAM135-A. Serially spotted cultures were allowed to grow in the dark at 30˚C for 3 days. Plates
were then scanned on a flatbed scanner and growth phenotypes were assessed and scored.
For all SDL overexpression constructs that caused toxicity upon overexpression in WT
BY4741 strains, I also performed Ypk1 dosage rescue growth assays. PMET25-Ypk1 or PMET25Ypk1(K367A) (kinase-dead) plasmids were co-transformed into BY4741 with the toxic SDL
plasmid. Transformants were then cultured overnight in SC medium containing 2% raffinose and
0.2% sucrose. Ten-fold serial dilutions of overnight cultures starting from A600 nm = 1.0 were
then made in sterile water and spotted onto SC solid medium with 2% galactose (to induce
protein expression) or 2% dextrose (no protein expression), in the presence of methionine to
repress Ypk1 expression or in the absence of methionine to derepress Ypk1 expression. Serially
spotted cultures were allowed to grow in the dark at 30˚C for 3 days. Plates were then scanned
on a flatbed scanner and growth phenotypes were assessed to determine if Ypk1 overexpression
could rescue toxicity of overexpression of the SDL protein.
For broth growth assays, cultures in exponential phase in rich (YP) medium (Burke et al.,
2005) with 2% dextrose were diluted to A600 nm = 0.1. A portion (100 μL) of each culture was
placed in a well in a 96-well plate with vehicle or drug at the indicated concentration. Cultures
were grown with orbital shaking at 30˚C in a Tecan Infinite M-1000 PRO plate reader (Tecan
Systems Inc.) for 24 h. Absorbance measurements were taken every 15 min Absorbance values
were converted to A600 nm values using a standard curve of absorbance values of cultures at
known A600 nm taken on the same plate reader.
Purification of Ypk1-as and purification of putative Ypk substrates
To purify Ypk1-as kinase, pAX50 (2μ, PGAL1-Ypk1(L424A)-TAP, URA3) transformed yAM135A yeast were diluted to A600 nm = 0.125 in 3L of SC with 2% raffinose+0.2% sucrose and grown
with shaking at 30˚C to mid-exponential phase. Expression was induced for ~18 h by the
addition of 2% galactose. Cells were harvested by centrifugation and frozen in liquid nitrogen.
The cells were then lysed cryogenically using Mixer Mill MM301 (Retsch). The lysate was
resuspended at 2 mL/g in TAP-B [50 mM Tris-Cl pH 7.5, 200 mM NaCl, 1.5 mM MgOAc, 1
mM DTT, 2 mM NaVO4, 10 mM NaF, 10 mM Na-PPi, 10 mM β-glycerol phosphate, 1x
cOmplete protease inhibitors™ (Roche)]. The lysate was clarified by centrifugation at 15,000xg
for 20 min Clarified lysate was then further centrifuged at 100,000xg for 1 h and then brought to
0.15% NP-40 using 10% NP-40 detergent stock. Ypk1-as-TAP fusion was then affinity purified
from the lysate using IgG-agarose resin (GE Healthcare). The resin was extensively washed with
Protease 3C Buffer (50 mM Tris-Cl pH 7.5, 200 mM NaCl, 1.5 mM MgOAc, 1 mM DTT, 0.01%
NP-40, 10% Glycerol, 2 mM NaVO4, 10 mM NaF, 10 mM Na-PPi, 10 mM β-glycerol
13
phosphate) and then resuspended in 1 mL Protease 3C Buffer. Ypk1-as was eluted by the
addition of 80 U Prescission Protease™ (human rhinovirus 3C protease) (GE Healthcare) for 5h
at 4˚C. Protease 3C was removed by the addition of glutathione-agarose (GE Healthcare).
Each putative Ypk1 substrate (or a large fragment thereof) was fused to the C-terminus of GST
in the vector pGEX-6P-1 and the resulting GST-substrate fusion proteins expressed in and
purified from E. coli strain BL21. Cultures (1L) were grown at 37˚C to mid-exponential phase
and then induced at room temperature for 4 h with 0.5 mM IPTG. Cells were harvested by
centrifugation and the fusion proteins were purified by affinity chromatography using
glutathione-agarose and standard procedures.
Ypk1-as in vitro kinase assays
For kinase assays, 0.25 μg of purified Ypk1-as was incubated with a purified GST-substrate in
Kinase Assay Buffer (50 mM Tris-Cl, pH 7.5, 200 mM NaCl, 10 mM MgCl2, 0.1 mM EDTA)
with 2 μCi [γ-32P]ATP at 30˚C in the presence of absence of 10 μM 3-MB-PP1 for 30 min
Reactions were terminated by the addition of SDS-PAGE sample buffer containing 6% SDS
followed by boiling for 5 min Labeled proteins were resolved by SDS-PAGE and analyzed by
Coomassie blue staining and autoradiography with Phosphorimager plates (Molecular
Dynamics) on a Typhoon imaging system (GE Healthcare).
Preparation of cell extracts and immunoblotting
To extract proteins from yeast cells, an alkaline lysis and trichloroacetic acid (TCA) precipitation
method were used, as previously described (Westfall et al., 2008). Cell extracts were then
resolved by various SDS-PAGE methods described below prior to immunobotting.
To resolve Lac1, Lag1, Fps1, Smp1(1-274) and Ypk1 phosphorylated species, 15 μL of TCA
extract was resolved by SDS-PAGE [8% acrylamide, 35 µM Phos-tag™ (Wako Chemicals USA,
Inc.), 35 µM MnCl2 at 160 V]. Inclusion of the Phos-tag™ reagent retards the mobility of and
improves the resolution of phosphorylated species (Kinoshita et al., 2009). To resolve Gpt2
phosphorylated species, 15 μL of the TCA extract was resolved by SDS-PAGE [8% 75:1
acrylamide:bisacrylamide), at 120 V]. Reducing the degree of gel crosslinking by increasing the
ratio of acrylamide monomer to bisacrylamide is a strategy also known to improve the separation
between phosphorylated and unphosphorylated species (Huang et al., 1997). To resolve full
length Smp1 phosphorylated species 15 μL of TCA extract was resolved by SDS-PAGE [8%
acrylamide, 45 µM Phos-tag (Wako Chemicals USA, Inc.), 45 µM MnCl2 at 160 V].
After separation by SDS-PAGE, the resolved proteins were transferred to nitrocellulose and
incubated with primary antibody in Odyssey buffer (Licor Biosciences), washed, and incubated
with IRDye680LT-conjugated anti-mouse IgG (Licor Biosciences) in Odyssey buffer with 0.1%
Tween-20 and 0.02% SDS. Blots were imaged using an Odyssey infrared scanner (Licor
Biosciences). Primary antibodies and dilutions used in this study were: 1:1000 rabbit anti-HA
(Covance Inc.), 1:5000 mouse anti-FLAG (Sigma-Aldrich), 1:5000 rabbit anti-FLAG (SigmaAldrich), 1:100 tissue culture medium containing mouse anti-c-myc mAb 9E10 (Monoclonal
Antibody Facility, Cancer Research Laboratory, Univ. of California, Berkeley; 1:100), 1:500
14
mouse anti-GFP (Roche), 1:500 rabbit anti-pSGK (T256) (to recognize Ypk1 phosphorylated at
T504) (Santa Cruz Biotechnology), 1:20000 rabbit anti-Ypk1(pT662) (a generous gift from Ted
Powers), 1:10000 rabbit anti-Ape1 (a generous gift from Daniel Klionsky), and 1:10000 rabbit
anti-PGK1 (our laboratory).
Analysis of sphingolipid species
Complex sphingolipids were analyzed by thin layer chromatography. Logarithmically growing
cultures of strains were adjusted to A60o nm = 1.0 and 2 ml cultures were labeled with 100 µCi of
[32P] H3PO4 and cells allowed to grow for 3 h. Lipids were extracted and resolved as previously
described (Hanson and Lester, 1980; Momoi et al., 2004) with minor modifications. The cell
pellet was washed twice with 2 ml water and treated with 5% trichloroacetic acid for 20 min on
ice. Pellets were extracted twice with 0.75 ml of ethanol/water/diethyl-ether/pyridine/NH4OH
(15:15:5:1:0.018) at 60°C for 1 h. Glycerophospholipids in the extract were hydrolyzed by
treating with 0.1 M monomethylamine at 50°C for 1 h after which the base was neutralized by
addition of 12 µl of glacial acetic acid. Lipids were extracted with 1 ml chloroform, 0.5 ml
methanol and phases separated with addition of 1 ml water. For some samples 1 ml of 4 N NaCl
was used, in order to facilitate the separation of the aqueous and organic phases. The organic
layer was dried under vacuum and resuspended in 50 µl of chloroform/methanol/water (16:16:5)
and resolved on a silica gel TLC plate with chloform/methanol/4.2 N NH4OH (9:7:2).
Radioactivity on the TLC plate was visualized with a Phosphorimager screen and Typhoon
imaging system.
LCBs and LCB-1-Ps levels were monitored by liquid chromatography-mass spectrometryOvernight cultures were adjusted to A60o nm = 0.2 and allowed to grow to an A60o nm = 1.0. Cell
pellets from 10 ml of this culture equivalent to 10 A600 nm units was used for analysis. C17sphingosine (Avanti Polar Lipids), (5 nmol) was added to all samples as an internal standard.
Lipids were extracted as described above and the final dried lipid extract was dissolved in
methanol/water/formic acid (79:20:1), centrifuged to remove insoluble material and an aliquot of
this material was injected into the HPLC for analysis.
Lipid extracts were analyzed using an Agilent 1200 liquid chromatograph (LC; Santa Clara, CA)
that was connected in-line with an LTQ Orbitrap XL mass spectrometer equipped with an
electrospray ionization source (ESI; Thermo Fisher Scientific, Waltham, MA).
The LC was equipped with a C4 analytical column (Viva C4, 150 mm length × 1.0 mm inner
diameter, 5 µm particles, 300 Å pores, Restek, Bellefonte, PA) and a 100 µL sample loop.
Solvent A was 99.8% water/0.2% formic acid and solvent B was 99.8% methanol/0.2% formic
acid (v/v). Solvents A and B both contained 5 mM ammonium formate. The sample injection
volume was 85 µL (partial loop). The elution program consisted of isocratic flow at 30% B for 2
min, a linear gradient to 65% B over 0.1 min, a linear gradient to 100% B over 4.9 min, isocratic
flow at 100% B for 4 min, a linear gradient to 30% B over 0.1 min, and isocratic flow at 30% B
for 18.9 min, at a flow rate of 170 µL/min The column and sample compartments were
maintained at 40°C and 4°C, respectively. The injection needle was rinsed with a 1:1
methanol/water (v/v) solution after each injection.
The column exit was connected to the ESI probe of the mass spectrometer using PEEK tubing
15
(0.005” inner diameter × 1/16” outer diameter, Agilent). Mass spectra were acquired in the
positive ion mode over the range m/z = 250 to 1200 using the Orbitrap mass analyzer, in profile
format, with a mass resolution setting of 100,000 (at m/z = 400, measured at full width at halfmaximum peak height). In the data-dependent mode, the five most intense ions exceeding an
intensity threshold of 30,000 counts were selected from each full-scan mass spectrum for tandem
mass spectrometry (MS/MS) analysis using pulsed-Q dissociation (PQD). MS/MS spectra were
acquired in the positive ion mode using the linear ion trap, in centroid format, with the following
parameters: isolation width 3 m/z units, normalized collision energy 40%, and default charge
state 1+. A parent mass list was used to preferentially select ions of interest for targeted MS/MS
analysis. To avoid the occurrence of redundant MS/MS measurements, real-time dynamic
exclusion was enabled to preclude re-selection of previously analyzed precursor ions, using the
following parameters: repeat count 2, repeat duration 30 s, exclusion list size 500, exclusion
duration 180 s, and exclusion width 0.1 m/z unit. Data were analyzed using Xcalibur software
(version 2.0.7 SP1, Thermo) and LIPID MAPS Online Tools (Fahy et al., 2007).
Exact masses of precursor ions in positive ion (M+H+) state obtained from LIPID MAPS Online
Tools were as follows- phytosphingosine- 318.3003, dihydrosphingosine-302.3053,
phytosphingosine-1 phosphate-398.2666 and dihydrosphingosine-1 phosphate- 382.2717. The
ions were further confirmed by tandem MS and identification of the fragments; product ions for
the phytosphingosine headgroup was 282.3 and dihydrosphigosine headgroup was 266.4.
In vitro ceramide synthase assay
Yeast expressing 3xFLAG-Lag1 (wild-type and phospho-site mutations) were grown at 30°C to
mid-exponential phase and microsomes were prepared as described previously (Schorling et al.,
2001) and resuspended in B88 buffer (20 mM HEPES-KOH pH 6.8, 150 mM KAc, 5 mM
MgOAc, and 250 mM sorbitol). Ceramide synthase was then immunopurified from these
microsomes using anti-FLAG agarose (Sigma) as described previously (Vallée and Riezman,
2005). Prior to assembling ceramide synthase reactions, a small alioquot of each (15 μL)
immunoprecipitate was taken and proteins were resolved by SDS-PAGE and immunoblotted
with anti-FLAG to determine the relative amount of immunopurified ceramide synthase in each
reaction. Recovered ceramide levels for each reaction were normalized to the amount of
ceramide synthase determined by this procedure.
Ceramide synthesis reaction was assembled in B88 buffer with 5 mg/ml BSA, in a total reaction
volume of 0.1 ml containing 47.5 µL anti-FLAG agarose immunoprecipitates, 50 µM
phytosphingosine and 0.1 mM of steroyl (C18)- CoA. Reactions were incubated at 30 °C for 1 h.
Reactions were terminated by addition of 0.4 ml methanol/CHCl3 (2:1) and further extracted
with 0.4 ml CHCl3 and 0.4 ml water. The CHCl3 phase was separated by centrifugation and
removed and dried under vacuum. The dried lipids resuspended in methanol/water/formic acid
(79/20/1) and an aliquot analyzed by LC-MS as described above. The product from the reaction,
C18-phytoceramide was identified by the exact mass of its precursor ion at 584.5612 and its
product ion upon fragmentation at 282.3. Additionally, authentic C18-phytoceramide (Matreya,
LLC.) standard was used to obtain a standard curve in order to establish that values from
ceramide synthase reaction were in the linear range of estimation.
16
Fluorescence microscopy of Smp1-GFP and Fps1-GFP
Cells transformed with plasmids that express GFP-tagged proteins were grown in selective
medium to mid-exponential phase (A60o nm ~0.6) or stationary phase. For cells stained with DAPI
to visualize nuclear DNA, 2.5 μg/mL of DAPI was added to medium for 30 min Stained cells
were then washed in fresh medium. Cells were viewed directly with an epifluorescence
microscope (model BH-2; Olympus America, Inc.) using a 100X objective fitted with
appropriate band-pass filters (Chroma Technology Corp.). Images were collected using a
CoolSNAP MYO charge-coupled device camera (Photometrics).
Measurement of STL1 promoter transcription by fluorescence activated cell sorting
(FACS)
Cells with PSTL1-GFP integrated at the HIS3 locus were grown to early exponential phase in SCD
medium lacking tryptophan. Cells with the ypk1-as were continuously grown in the presence of
10 μM 3-MB-PP1 to inhibit Ypk1 activity. Cells were then switched into fresh medium with or
without 0.4 M NaCl to trigger hyperosmotic stress. Cells were grown under these conditions for
90 min Cells were then pelleted and 100 μL of paraformaldehyde solution (4% w/v
paraformaldehyde, 3.4% sucrose) was added to cells. Cells were incubated for 15 min at room
temperature. Cells were then pelleted again and washed in wash solution (1.2 M sorbitol, 100
mM potassium phosphate, pH 7.5). The cell pellet was then resuspended in the wash solution.
Cells were sonicated in a sonicator bath and then cellular GFP fluorescence was then measured
by FACS using a Guava easyCyte (Merck Millipore). For each sample, fluorescence in 10 5 cells
was measured. The average cellular fluorescence was then plotted.
Measurement of intracellular glycerol accumulation
Measurement of intracellular glycerol was conducted as described in Albertyn et al. (1994).
Briefly, cells from ~40 mL of exponentially-growing culture were harvested by centrifugation
and washed with 1 mL of medium. The resulting cell pellets were frozen in liquid N2 and stored
at -80˚C prior to analysis. Each cell pellet was then boiled at 100˚C for 10 min in 1 mL of 50
mM Tris-Cl, pH 7.0. The lysate was then clarified by centrifugation at 16,000xg for 15 min.
Glycerol concentration in the resulting supernatant solution was measured using an enzymatic
assay kit (Sigma Aldrich) and was normalized to extract protein concentration, as measured by
Bradford assay (Bradford, 1976).
Co-immunoprecipitation of Fps1 and Rgc2
Co-immunoprecipitation experiments were performed as previously described (Lee et al., 2013),
with only minor modifications. Cells expressing Fps1-3xFLAG (yAM271 – A), Fps13A-3xFLAG
(yAM272 – A) or untagged Fps1 (BY4742) were transformed with a plasmid expressing Rgc23xHA under the control of the MET25 promoter (Lee et al., 2013). Cultures of each were grown
to mid-exponential phase in SCD -uracil medium. Cultures were then diluted to A60o nm = 0.2 in 1
L of SCD-Met-Ura medium to induce expression of Rgc2-3xHA. Cultures were grown at 30˚C
for 5 h. Cells were harvested by centrifugation and resuspended in 5 mL of TNE+Triton+NP-40
[50 mM Tris-Cl, pH 7.5, 150 mM NaCl, 4 mM NaVO4, 50 mM NaF, 20 mM Na-PPi, 5 mM
17
EDTA, 5 mM EGTA, 0.5% Triton-X100, 1.0% NP-40 1x cOmplete protease inhibitor™
(Roche)]. The cells were then lysed cryogenically using Mixer Mill MM301 (Retsch). The lysate
was thawed on ice and then clarified twice by centrifugation at 13,000xg for 20 min. Protein
concentration of the clarified lysate was measured by BCA assay (Thermo Scientific) and
normalized between the lysates. Fps1-3xFLAG was then isolated from ~10 mg of lysate using 50
uL of mouse anti-FLAG antibody coupled agarose resin (Sigma Aldrich) equilibriated in
TNE+Triton. Binding was allowed to occur for 2 h at 4˚C. The resin was then extensively
washed with TNE+Triton and precipitates were then processed for SDS-PAGE resolution and
immunoblot detection of Fps1-3xFLAG and Rgc2-3xHA.
18
Tables
Table 2.1 Yeast strains used in this study
Strain
Relevant Genotype
Source/reference
BY4741
MATa his3∆1 leu2∆0 met15∆0 ura3∆0
Research Genetics, Inc.
BY4742
MATα his3∆1 leu2∆0 lys2∆0 ura3∆0
Research Genetics, Inc.
CGA65
BY4741 snf1∆::KanMX
Christopher Alvaro, this
lab
DL3188
BY4742 rgc1∆::KanMX rgc2∆::KanMX
(Beese et al., 2009)
JTY5173
W303-1B MATα ade2-1 can1-100 his3-11,15 leu2-
(Zaman et al., 2009)
3,112 ura3∆0 trp1-1 tpk1-as tpk2-as tpk3-as
JTY5468
BY4741 tor2-29::KanMX
(Li et al., 2011)
JTY5537
BY4742 pbs2∆::KanMX
Research Genetics, Inc.
JTY5538
BY4742 ssk2∆::KanMX
Research Genetics, Inc.
JTY5539
BY4742 ssk22∆::KanMX
Research Genetics, Inc.
JTY5540
BY4742 sho1∆::KanMX
Research Genetics, Inc.
JTY5541
BY4743 ssk1∆::KanMX/ssk1∆::KanMX
Research Genetics, Inc.
JTY5574
BY4741 cna1∆::KanMX4 cna2∆::KanMX4
M.S. Cyert, Stanford
Univ.
JTY6142
BY4741 ypk1∆::KanMX4
Research Genetics, Inc.
JTY6322
BY4743 fps1∆::KanMX/fps1∆::KanMX
Research Genetics, Inc.
JTY6328
BY4741 gpd1∆::KanMX4 gpd2∆::HIS3
(Lee et al., 2012)
yAM120-A
BY4741 ypk2∆::KanMX4
This study
yAM134-A
BY4741 Ypk1(L424A)::URA3-
This study
19
yAM135-A
BY4741 Ypk1(L424A)::URA3- ypk2∆:: KanMX4
This study
yAM159-A
BY4741 3xFLAG-Lag1::LEU2
This study
yAM163-A
BY4741 3xFLAG-Lag1(S23A S24A)::LEU2
This study
yAM165-A
BY4742 3xHA-Lac1::HIS3
This study
yAM166-A
BY4742 3xHA-Lac1 (S23A S24A)::HIS3
This study
yAM168
BY4741 3xHA-Lac1::HIS3 3xFLAG-Lag1::LEU2
This study
yAM181 - A
BY4742 fps1∆::natNT2
This study
yAM184
BY4741 3xHA-Lac1(S23A S24A)::HIS3 3xFLAG-
This study
Lag1(S23A S24A)::LEU2
yAM192-A
BY4741 MET15+ 3xHA-Lac1(S23E S24E)::HIS3
This study
3xFLAG-Lag1(S23E S24E)::LEU2
yAM197
BY4742 smp1∆::LEU2
This study
yAM205-A
BY4742 Lac1::LEU2 Lag1::LEU2
This study
yAM207-B
BY4742 Lac1(S23A S24A)::LEU2 Lag1(S23A
This study
S24A)::LEU2
yAM210
BY4742 Lac1(S23E S24E)::LEU2 Lag1(S23E
This study
S24E)::LEU2
yAM219 – A
BY4741 gpt2∆:: Hygr
This study
yAM221 - A
BY4741 atg21∆::Hygr
This study
yAM231
BY4741 gpt2∆:: Hygr sct1∆::natNT2 [pRS315
This study
PGPT2-Gpt2-3xFLAG; pAX240]
yAM232
BY4741 gpt2∆:: Hygr sct1∆::natNT2 [pRS315
PGPT2-Gpt23A-3xFLAG; pAX248]
20
This study
yAM233
BY4741 gpt2∆:: Hygr sct1∆::natNT2 [pRS315
This study
PGPT2-Gpt23E-3xFLAG; pAX245]
yAM234 - B
BY4741 PTPI1-GFP-Atg8::HIS3
This study
yAM235 - A
BY4741 PTPI1-GFP-Atg8::HIS3 atg21∆::Hygr
This study
yAM236 - B
BY4741 PTPI1-GFP-Atg8::HIS3
This study
Ypk1(L424A)::URA3- ypk2∆:: KanMX4
yAM241 - A
BY4741 PSTL1-GFP::HIS3
This study
yAM242 - A
BY4741 PSTL1-GFP::HIS3 ypk1-as::URA3-
This study
yAM243
BY4741 PSTL1-GFP::HIS3 smp1∆::LEU2
This study
yAM244 – A
BY4741 PSTL1-GFP::HIS3 ypk1-as::URA3-
This study
smp1∆::LEU2
yAM246 - A
BY4741 Smp1-3xFLAG::URA3
This study
yAM247 - A
BY4741 Smp1(T20A T107A T221A)-
This study
3xFLAG::URA3
yAM248 - A
BY4741 Smp1(T20E T107E T221E)-
This study
3xFLAG::URA3
yAM275
BY4742 LYS2+ Fps1-3xFLAG::URA3
This study
hog1∆::KanMX
yAM278
BY4742 Fps1(S181A S185A S570A)-
This study
3xFLAG::URA3 hog1∆::KanMX
yAM281
BY4742 Ypk1(L424A)::URA3- ypk2∆:: KanMX4
This study
Fps1-3xFLAG::URA3
yAM284 - A
BY4742 Ypk1(L424A)::URA3- ypk2∆:: KanMX4
21
This study
Fps1(S181A S185A S570A)-3xFLAG::URA3
yAM291 - A
BY4742 Fps1(S570A)-3xFLAG::URA3
This study
hog1∆::KanMX
yAM301 - A
BY4742 Fps1(S181A S185A)-3xFLAG::URA3
This study
yAM307 - A
BY4742 Fps1(Δ544-581)-3xFLAG::URA3
This study
yAM308 - A
BY4742 Fps1(I218A V220A)-3xFLAG::URA3
This study
yAM309 - A
BY4742 Fps1(S181A S185A I218A V220A
This study
S570A)-3xFLAG::URA3
yAM310 - A
BY4742 Fps1(T147A)-3xFLAG::URA3
This study
yAM315
BY4741 Rgc2(S344A T808A S948A S75A S827A
This study
S1021A S1035A)-3xHA:: Hygr
yAM318
BY4741 Fps1(S181A S185A S570A)-
This study
3xFLAG::URA3 Rgc2(S344A T808A S948A S75A
S827A S1021A S1035A)-3xHA:: Hygr
YDB379
BY4741 Ypk1-3xFLAG::natNT2
J.S. Weissman, Univ. of
California, San Francisco
yGT12
BY4742 LYS2+ Lac1::LEU2 Lag1::LEU2
This study
lcb3∆::natNT2
yGT13
BY4742 LYS2+ Lac1(S23A S24A)::LEU2
This study
Lag1(S23A S24A)::LEU2 lcb3∆::natNT2
yGT14
BY4742 LYS2+ Lac1(S23E S24E)::LEU2
This study
Lag1(S23E S24E)::LEU2 lcb3∆::natNT2
yGT21
BY4742 Fps1-3xFLAG::URA3
This study
22
yGT22
BY4742 Fps1(S181A S185A S570A)-
This study
3xFLAG::URA3
yGT24
BY4742 Fps1(S570A)-3xFLAG::URA3
This study
YJP544
BY4741 hog1∆::KanMX
Jesse Patterson, this lab
yKL4
BY4741 TOR2+::Hygr
Kristin Leskoske, this lab
yKL5
BY4741 Tor2(L2178A)::Hygr
Kristin Leskoske, this lab
23
Table 2.2 Plasmids used in this study
Plasmid
Description
Source/reference
pGEX6P-1
GST tag, bacterial expression vector
GE Healthcare, Inc.
pGEX4T-1
GST tag, bacterial expression vector
GE Healthcare, Inc.
YCpLG
CEN, LEU2, PGAL1 vector
(Bardwell et al., 1998)
BG1805
2µm, URA3, PGAL1, C-terminal tandem affinity
Open Biosystems, Inc.
(TAP) tag vector
pRS313
CEN, HIS3, vector
(Sikorski and Hieter, 1989)
pRS316
CEN, URA3, vector
(Sikorski and Hieter, 1989)
pRS416
CEN, URA3, vector
(Sikorski and Hieter, 1989)
pRS426
2µm, URA3, vector
(Sikorski and Hieter, 1989)
pBC111
CEN, LEU2, vector
(Iida et al., 2007)
CHp282
pRS416 PMET25-GFP
Chris Huynh, this laboratory
p3151
pRS316 PMET25-Rgc2-3xHA
(Lee et al., 2013)
pAX131
pGEX4T-1 Lac1(1-76)
This study
pAX132
pGEX4T-1 Lac1(1-76)(S23A S24A)
This study
pAX133
pGEX4T-1 Lag1(1-80)(S23A S24A)
This study
pAX134
pGEX6P-1 Muk1(1-305)
This study
pAX135
pGEX6P-1 Fps1(531-669)(S570A)
This study
pAX136
pRS313 PLAC1-3xHA-Lac1
This study
pAX215
pGEX6P-1 Cyk3
This study
pAX223
pGEX6P-1 Gpt2(1-35)
This study
pAX224
pGEX6P-1 Gpt2(570-743)
This study
24
pAX225
pGEX6P-1 Bre5
This study
pAX226
pGEX6P-1 Npr1(1-437)
This study
pAX227
pGEX6P-1 Pal1
This study
pAX228
pGEX6P-1 Ysp2(97-665)
This study
pAX229
pGEX6P-1 Ysp2(1072-1282)
This study
pAX230
pGEX6P-1 Atg21
This study
pAX231
pGEX6P-1 Pex31(250-462)
This study
pAX238
pRS316 PGPT2-Gpt2-3xFLAG
This study
pAX239
CEN, LEU2, PTPI1-GFP-Atg8
This study
pAX240
pRS315 PGPT2-Gpt2-3xFLAG
This study
pAX241
pRS316 PATG21-Atg21-3xFLAG
This study
pAX244
pRS316 PGPT2-Gpt2(S649A S650A S651A)-
This study
3xFLAG
pAX245
pRS315 PGPT2-Gpt2(S649E S650E S651E)-
This study
3xFLAG
pAX246
pRS316 PATG21-Atg21(S237E)-3xFLAG
This study
pAX247
pRS316 PATG21-Atg21(S237A)-3xFLAG
This study
pAX248
pRS315 PGPT2-Gpt2(S649A S650A S651A)-
This study
3xFLAG
pAX250
pRS313 PTPI1-GFP-Atg8
This study
pAX253
pRS316 PSMP1-Smp1-3xFLAG
This study
pAX260
pRS316 PSMP1-Smp1(T20A T107A T221A)-
This study
3xFLAG
25
pAX263
pRS316 PSMP1-Smp1(T20E T107E T221E)-
This study
3xFLAG
pAX270
pRS316 PSMP1-Smp1-GFP
This study
pAX274
pRS316 PFPS1-Fps1-3xFLAG
This study
pAX275
pRS316 PFPS1-Fps1(S181A S185A S570A)-
This study
3xFLAG
pAX286
pRS316 PSMP1-Smp1(1-274)-3xFLAG
This study
pAX287
pRS316 PSMP1-Smp1(1-274)(T20A T107A
This study
T221A)-3xFLAG
pAX290
pRS316 PFPS1-Fps1-GFP
This study
pAX293
pRS316 PFPS1-Fps1(S570A)-GFP
This study
pAX294
pRS316 PFPS1-Fps1(S181A S185A)-GFP
This study
pAX295
pRS316 PFPS1-Fps1(S181A S185A S570A)-GFP
This study
pAX50
BG1805 Ypk1(L424A)
This study
pAX53
pRS416 PMET25-Ypk1(K376A)
This study
pAX55
pGEX6P-1 Lcb3(1-79)
This study
pAX56
pGEX6P-1 Cdc1(1-41)
This study
pAX58
pGEX6P-1 Her1(1-224)
This study
pAX59
pGEX6P-1 Rts3
This study
pAX62
pGEX6P-1 Fkh1
This study
pAX63
pGEX6P-1 Yhp1
This study
pAX66
pGEX6P-1 YNR014W
This study
pAX67
pGEX6P-1 YHR097C
This study
26
pAX94
pGEX6P-1 Mds3(545-1016)
This study
pAX96
YCpLG Fps1
This study
pBCT-
pBC111 PTDH3-Cch1
(Iida et al., 2007)
pBT12
pGEX6P-1 Smp1
This study
pBT6
pGEX6P-1 Fps1(1-255)
This study
pBT7
pGEX6P-1 Fps1(531-669)
This study
pFR203
pGEX4T-1 Orm1(1-85)
(Roelants et al., 2011)
pFR252
pRS315 PYPK1-Ypk1(S51A S57A S71A T504A
This study
CCH1H
S644A S653A T662A)-myc
pFR273
pRS316 PYPK1-Ypk1(D242A)
(Roelants et al., 2011)
pFR291
pGEX4T-1 Lag1(1-80)
This study
pGT02
pRS426 PFPS1-Fps1-3xFLAG
This study
pGT05
pRS426 PFPS1-Fps1(Δ12-231) -3xFLAG
This study
pGT06
pRS426 PFPS1-Fps1(Δ534-650)-3xFLAG
This study
pGT11
YCpLG Fps1(Δ12-231)
This study
pGT12
YCpLG Fps1(Δ534-650)
This study
pGT13
YCpLG Fps1(Δ544-581)
This study
pGT14
YCpLG Fps1(S181A S185A S570A)
This study
pGT15
YCpLG Fps1(S181E S185E S570E)
This study
pGT17
pRS426 PFPS1- Fps1(S181A S185A S570A)-
This study
3xFLAG
pGT18
pRS426 PFPS1- Fps1(S181A S185A S570A)-
27
This study
3xFLAG
pLB215
pRS416 PMET25-Ypk1
(Niles et al., 2012)
28
Chapter 3: A three-tiered screen identifies novel Ypk substrates
This chapter is based on work published in Muir A et al., eLife, 2014. I devised the threetiered screen for Ypk substrates with Prof. Jeremy Thorner. I was responsible for all
bioinformatic and biochemical screening. I constructed the substrate overexpression
library and performed all synthetic dosage lethality genetic screening with the assistance of
undergraduate Garrett Timmons.
Introduction
To gain further insight into how the TORC2-Ypk1 signaling axis contributes to plasma
membrane homeostasis and potentially other cellular processes, I aimed to identify novel Ypk
substrates. A variety of mass spectrometry-based approaches for identifying kinase substrates are
available (Koch and Hauf, 2010). In general, these approaches involve inhibiting the kinase of
interest in vivo, extracting phosphopeptides from this sample and comparing levels of
phosphopeptides to an appropriate control. Those peptides that change upon inhibition are
candidate substrates for the kinase of interest. However, these mass spectrometry-based
approaches have the drawback that they are not sensitive enough to detect low-copy number
signaling proteins and can also fail to detect membrane proteins (Koch and Hauf, 2010). As
nearly all of the previously identified Ypk substrates fall into these categories, use of mass
spectrometry based approaches seemed ill advised.
Instead, I devised a three-step procedure that would avoid the drawbacks of mass spectrometrybased approaches to screen in an unbiased and genome-wide manner for additional candidate
Ypk1 substrates whose physiological relevance I could then evaluate. I first used bioinformatics
to search the yeast proteome for presumptive targets, then applied a genetic method to narrow
down the hits to likely, functionally important substrates, and then used biochemical analysis
both in vitro and in vivo to validate the best prospects. In this chapter, I will describe in detail
each step of the screen and report here the general outcomes of this screen. Chapters 4 and 5
describe characterization of the functional role of Ypk phosphorylation of substrates involved in
sphingolipid metabolism and substrates involved in glycerol metabolism, respectively.
Results
A three-tiered screen to identify new Ypk1 substrates
I devised a three-step strategy (Fig. 3.1A) to pinpoint bona fide cellular targets of Ypk1, utilizing
bioinformatics to predict potential Ypk1 substrates, then an in vivo genetic test involving a novel
variation of the synthetic dosage lethality method to winnow the list to likely candidates, and
finally biochemical analysis in vitro to confirm whether the identified gene product serves as a
direct substrate of Ypk1. The physiological relevance of Ypk1-dependent modification of each
protein on the resulting final list could then been evaluated.
Bioinfomatic prediction of Ypk substrates
For initial bioinformatic search of the yeast proteome, I developed a position-weighted
29
consensus sequence logo (Fig. 3.1B) for the preferred Ypk1 phospho-acceptor site based on two
primary criteria: (a) the known Ypk1 sites in five, validated in vivo targets (Fpk1, Fpk2, Orm1,
Orm2, and Gpd1) (Roelants et al., 2010; Roelants et al., 2011; Lee et al., 2012b; Niles et al.,
2012; Sun et al., 2012); and, (b) the sequence preference displayed by Ypk1 for phosphorylation
of synthetic peptides in vitro (Casamayor et al., 1999; Mok et al., 2010). All demonstrated
substrates either in vivo or in vitro contain Arg at positions -5 and -3 with respect to the
phosphorylated Ser (or Thr); thus, these positions were invariant in the search motif. Given that
nearly all the verified sites within known in vivo targets possess a hydrophobic residue (V, I, F)
at position +1, the search motif gave preference to sites with a hydrophobic residue at the +1
position. I then took advantage of the existing MOTIPS motif analysis package (Lam et al.,
2010) to identify those S. cerevisiae gene products that contain occurrences of the search logo.
Several authentic Ypk1 substrates (e.g., Fpk1, Orm1, and Orm2) contain multiple Ypk1
phosphorylation sites. Thus, I filtered our search further by prioritizing candidates containing
multiple matches to the search logo, as predicted by MOTIPS. However, to avoid disregarding
potential Ypk1 substrates with a single predicted match to the sequence logo, I also considered
MOTIPS hits wherein there was existing evidence in the PhosphoGRID database (Sadowski et
al., 2013) indicating that a predicted site is phosphorylated in vivo. To narrow down the list of
potential substrates further, I chose to pursue those gene products containing matches to the
sequence logo for which there was existing information in the literature suggesting a phenotypic
relationship to known Ypk1-dependent processes: (a) a loss-of-function mutation in the
candidate gene exhibits elevated sensitivity to agents (aureobasidin A, caspofungin and/or
myriocin) toward which a ypk1∆ mutant is also sensitive (Hillenmeyer et al., 2008); (b) the
candidate gene product is reported to be involved in a genetic or biochemical interaction with
Ypk1, as curated in YeastMine (Balakrishnan et al., 2012); (c) the candidate gene product is
connected in some way to known Ypk1 regulators (e.g., TORC2, PP2A); and/or, (d) the
candidate gene product is involved in a known Ypk1-regulated process (e.g., sphingolipid
metabolism) (for further details, see Chapter 2). Reassuringly, our approach identified three
known Ypk1 substrates (Fpk1, Orm1 and Orm2); absence of Fpk2 and Gpd1 from the list
generated solely by the MOTIPS search criteria arose from the fact that these substrates contains
only a single predicted site that is not presently recorded in PhosphoGRID. For this reason, I also
restored for consideration additional gene products that contain a single match to the consensus
sequence logo that YeastMine indicated are involved in processes in which Ypk1 is implicated.
The resulting candidates, grouped via cellular process on the basis of current GO Slim
terminology (http://www.geneontology.org/GO.slims.shtml), are cataloged in Table 3.1, and
represent fewer than 100 gene products out of the approximately 6,000 protein-coding genes in
the yeast genome (Lin et al., 2013) [although the number of authentic open-reading-frames
undergoes constant revision (http://www.yeastgenome.org/cache/genomeSnapshot.html)].
A synthetic dosage lethality screen to identify Ypk interacting proteins
As the secondary filter for the candidates recognized bioinformatically, I developed an in vivo
approach to identify those gene products that manifested an expected hallmark of proteinsubstrate interaction. I reasoned that under conditions where the level of activity of an essential
kinase, like Ypk1, is near-limiting for normal growth, high-level over-expression of an authentic
substrate might tie up the available pool of active enzyme and prevent efficient phosphorylation
of other cellular substrates necessary for growth and/or viability (Fig. 3.1C). This scheme is a
30
novel variation on a genetic approach referred to as synthetic dosage lethality (SDL) (Sopko et
al., 2006; Sharifpoor et al., 2012). To limit Ypk1 activity, I used ypk1-as ypk2∆ cells, which
express from the YPK1 locus an analog-sensitive allele, Ypk1(L424A), and titrated down its
activity by addition of a low concentration of an efficacious inhibitor,1-(tert-butyl)-3-(3methylbenzyl)-1H-pyrazolo[3,4-d]pyrimidin-4-amine (3-MB-PP1) (Burkard et al., 2007) that has
no effect on wild-type cells. To achieve high-level over-expression, each bioinformatic hit was
expressed from the galactose-inducible GAL1 promoter on a CEN plasmid. As proof of concept,
I used two known Ypk1 substrates, Orm1 and Orm2, as positive controls and GFP, which is not a
Ypk1 substrate (data not shown), as a negative control. In the absence of limiting the activity of
Ypk1(L424A) with inhibitor, overexpression of neither Orm proteins nor GFP was deleterious to
cell growth (Fig, 3.1D, left panels, compare lower to upper). However, in the presence of a low
dose of 3-MB-PP1, overexpression of Orm1 and Orm2 on galactose medium prevented cell
growth, whereas overexpression of GFP did not (Fig. 3.1D, middle and right panels, compare
lower to upper). I was able to test the majority of the candidates (90/96) that arose in the
bioinformatic search in this same fashion [however, 10/90 caused toxicity upon over-expression
even in wild-type cells and, hence, could not be scored].
Those candidates that, like Orm1 and Orm2, exhibited toxicity only on galactose medium and
only when Ypk1(L424A) activity was limited in the presence of 3-MB-PP1, but not when
inhibitor was absent, were designated SDL hits (Table 3.1, column 4). Moreover, use of a series
of 3-MB-PP1 concentrations allowed for quantification of the strength of the SDL effect (from +
to ++++). In one case (Fps1), a marked SDL effect was observed upon overexpression in the
ypk1-as ypk2Δ cells in the absence of chemical inhibition; I considered this a valid SDL hit
because GAL promoter-driven over-expression of Fps1 was not growth inhibitory in wild-type
(YPK1+ YPK2+) cells. Thus, as summarized in Table 3.1 (column 4), a significant fraction
(20/90) of the candidates identified bioinformatically that were tested in this fashion, but far
from all, displayed an SDL phenotype. In this regard, it is important to note that all known Ypk1
substrates tested (Gpd1, Fpk1, Orm1 and Orm2) displayed an SDL phenotype, whereas nearly
80% of the bioinformatic hits, like GFP, did not. Consistent with the view that the SDL
phenotype could arise from the over-expressed target serving as a decoy substrate that titers a
limited pool of active Ypk1 away from acting on its essential substrates, I observed that overexpressed catalytically-inactive Fpk1 caused an SDL phenotype equivalent to or stronger than
wild-type Fpk1 (Table 3.1). If such SDL phenotypes reflect occlusion of a limited pool of
enzyme by over-expressed substrate, then, conversely, co-overexpression of Ypk1 or even of a
kinase-dead allele Ypk1(K376A) (driven from the MET25 promoter) might rescue the toxicity.
Indeed, the deleterious effect of Smp1 over-expression was rescued by co-over-expression of
either Ypk1 or Ypk1(K376A) (Table 3.1), suggesting that the SDL phenotype of over-expressed
Smp1 also arises from titration of a limited amount of Ypk1 away from essential substrates.
Biochemical screening of bioinformatically and genetically predicted Ypk substrates
Lastly, to determine whether the gene products that displayed an SDL phenotype are indeed
substrates for Ypk1, I incubated those (17/20) that I was able to successfully express and purify
as recombinant proteins or protein fragments (as GST fusions) from E. coli with [-32P]ATP and
Ypk1(L424A), which was highly purified from yeast cells as described in Chapter 2. I chose to
use the analog-sensitive allele, even though it is only about 50% as active as wild-type Ypk1
31
(Roelants et al., 2011), because ablation of activity in the presence of 3-MB-PP1 allowed us to
confirm that any 32P incorporation into substrate observed was due to phosphorylation by
Ypk1(L424A) itself (and not due to some other protein kinase that might be present in the
preparation). All known in vivo substrates of Ypk1 (Fpk1, Fpk2, Gpd1, Orm1 and Orm2) display
robust incorporation (as judged by autoradiography) in this in vitro assay (Lee et al., 2012b;
Roelants et al., 2010; Roelants et al., 2011). Therefore, in testing each candidate, an appropriate
positive control, Orm1(1-85) was included (Fig. 3.1E), which also allowed comparison between
independent assays. Gratifyingly, 12/17 (70%) of the SDL hits tested in this manner displayed
readily detectable and Ypk1-specific phosphorylation (Table 3.1, right column). Moreover, the
results of such assays clearly demonstrated that Ypk1 is not a "promiscuous" enzyme. For
example, Fps1 has three possible N-terminal Ypk1 sites (RPRGQT147T, RRRSRS181R and
RSRATS185N) and one C-terminal site. The C-terminal site is a Ypk1 target (see Chapter 5);
however, none of the N-terminal motifs serves as an efficient Ypk1 phospho-acceptor site (Fig.
3.1E), perhaps because each lacks a hydrophobic residue at +1.
Thus, I considered it very likely that the dozen candidates (shown schematically in Fig. 3.2 and
highlighted in bold in Table 3.1) identified bioinformatically that also displayed an SDL
phenotype and served as Ypk1 substrates in vitro would be functionally important Ypk1
substrates in vivo. To validate this conclusion and confirm that these candidates are indeed
physiologically relevant Ypk1 targets, I chose to characterize Lac1 and Lag1, two of the dozen
candidates (Table 3.1), because they are the catalytic subunits of the ceramide synthase complex
and might further our understanding about how sphingolipid production is regulated by the
TORC2-Ypk1 signaling axis (see Chapter 4).
Discussion
Identification of new TORC2-Ypk1 substrates
Our approach identified 12 new Ypk substrates. I was able to generated reagents to examine the
physiological relevance of five of these twelve candidates. In Chapters 4 and 5, I report that four
of the five tested substrates are indeed bona fide targets of Ypk in vivo. These findings validate
the ability of our methods for discovery of physiologically relevant protein kinase substrates.
Phospho-acceptor site motif pattern-matching alone, although useful in identifying kinase
substrates in some cases (Gwinn et al., 2008; Holt et al., 2007; Hutti et al., 2009; Linding et al.,
2007; Mah et al., 2005; Manning et al., 2002; Rennefahrt et al., 2007; Yaffe et al., 2001), can
yield a large number of false positives. This possibility was a significant concern for us because
certain features of the basophilic Ypk1 motif are shared with other protein kinases (Mok et al.,
2010). As a means to avoid this problem, I devised a novel SDL-based genetic approach to apply
as a secondary filter to parse the bioinformatically selected candidates further.
Although not the only possible explanation for detecting an SDL hit, one mechanism our genetic
method should be able to assess is whether the candidate gene product displays a primary
characteristic of a true substrate, namely the ability to compete with other substrates for
association with Ypk1. A genuine Ypk1 substrate should, when highly over-expressed, sequester
a large fraction of the available pool of active kinase, and thus impede its actions on its essential
cellular targets. In the absence of Ypk2, over-expression of an authentic Ypk1 substrate should
32
therefore be deleterious for growth when the activity of an analog-sensitive Ypk1 allele is
reduced with a selective inhibitor. The fact that 70% of the SDL hits were indeed substrates for
Ypk1-mediated phosphorylation in vitro verified that our use of this genetic method as a
secondary filter to pick out true substrates from the list of bioinformatically identified candidates
was well justified. In this regard, overexpression of Fpk1(KD), a catalytically-inactive nonessential Ypk1 substrate, yielded a readily detectable SDL phenotype. Thus, using the protocol I
devised, SDL may be a generally useful way to identify substrates and perhaps other binding
partners of protein kinases, not just those directly connected to any output phenotype being
measured.
Other genetic approaches have also been useful in identifying physiologically relevant targets of
Ypk1. For example, a transposon insertion that suppressed the growth defect of a ypk1-ts ypk2Δ
strain at an otherwise non-permissive temperature initially identifed Orm2 as a potential Ypk1
substrate (Roelants et al., 2002), a finding later corroborated by us (Roelants et al., 2011) and
others (Berchtold et al., 2012; Liu et al., 2012; Niles et al., 2012; Sun et al., 2012). Similarly,
Smp1, which I confirmed here is a likely Ypk1 substrate (see also Chapter 5), was first identified
as a potential Ypk1 target because it was isolated as a dosage suppressor of the temperaturesensitive phenotype of ypk1-ts ypk2Δ cells (Roelants et al., 2002). Smp1 is a transcription factor
(Dodou & Treisman, 1997; de Nadal et al., 2003) that mediates iron toxicity (Lee et al., 2012a)
and, similarly, Ypk1 is required for iron toxicity (Lee et al., 2012a). Thus, TORC2-Ypk1
signaling might be mechanistically coupled to iron metabolism by modulation of Smp1
transcriptional output (see Chapter 5).
Likewise, other situations where chemical genetics can be applied have proven useful in gleaning
what aspects of cell function are controlled by a protein kinase. For example, mutagenesis of
yeast Tor2 to confer susceptibility to a chemical inhibitor and thereby selectively inhibit TORC2
action has been achieved (Kliegman et al., 2013). This tool was combined with a collection of
deletion mutants to identify what processes, when eliminated, are especially deleterious to cell
growth and survival when TORC2 action (and presumably Ypk1 activity) is limiting. This
analysis suggested some connection between TORC2 action and the pentose-phosphate pathway
(Kliegman et al., 2013), in keeping with the growth-promoting roles of both TORC1 and TORC2
and the demand for NADPH in many cellular anabolic reactions. Similarly, use of TOR
inhibitors has implicated TORC2-Ypk1 signaling in regulation of actin filament formation that is
somehow required for yeast cell survival in response to low levels of DNA damage (Shimada et
al., 2013).
By contrast, the genetic approach in our SDL method is quite different, in that it scores the
deleterious effect arising from overexpression of a gene product (increased protein dosage) rather
than from the total absence of a gene product, when the activity of the kinase of interest is
limited by the presence of a moderate concentration of a specific inhibitor. Theoretically, under
our conditions, I should also have been able to observe synthetic dosage rescue ("SDR");
however, I found no such examples. In any event, our method independently identified nearly all
of the previously known in vivo substrates of Ypk1, as well as nearly a dozen genuine Ypk1
targets, including Lac1 and Lag1 (Chapter 4), that have only been pinpointed by our three-tiered
method. Thus, our SDL screening technique provides a complementary approach for identifying
substrates of a protein kinase above and beyond those accessible through genetic interactions
33
between the kinase and single-gene deletions or other genetic schemes.
Many of the gene products identified by our screen are involved in processes that the TORC2Ypk1 signaling axis is already known to regulate. For example, Ypk1 regulates glycerol-3phosphate production via phosphorylation and inhibition of glycerol-3-phosphate dehydrogenase
Gpd1 (Lee et al., 2012b). Interesting, a potential Ypk1 target that met all of the criteria in our
screen is Gpt2, sn-glycerol-3-phosphate 1-acyltransferase (Zheng and Zou, 2001), an enzyme
that esterifies glycerol-3-phosphate as the first step in glycerolipid formation. In this same
regard, as another very likely Ypk1 target I also identified Fps1, a membrane channel
(aquaglyceroporin) that regulates efflux of glycerol (a glycerol-3-phosphate-derived metabolite)
(Luyten et al., 1995). These results strengthen the conclusion that TORC2-Ypk1 signaling is
intimately involved in modulating the level of the precursor to both glycerophospholipids and the
osmolyte glycerol (Lee et al., 2012b) (see Chapter 5).
Ypk1 action has been implicated in regulation of both fluid phase and receptor-mediated
endocytosis (deHart et al., 2002). In this regard, I identified as a very likely Ypk1 target an
endocytic adaptor, the α-arrestin Rod1, which is necessary for ubiquitinylation-triggered
internalization of nutrient permeases (Becuwe et al., 2012; Lin et al., 2008) and the pheromone
receptor Ste2 (Alvaro et al., 2014). Thus, as it does for plasma membrane lipids, TORC2-Ypk1
signaling may modulate plasma membrane protein composition via this α-arrestin, a possibility
being pursued by my fellow graduate student Christopher G. Alvaro and Ann Aindow, an
undergraduate working with him.
Ypk1 function has also been implicated in regulating production of reactive oxygen species
(ROS) (Niles et al., 2014), but an as yet undefined mechanism. In our screen, I found Ysp2, a
protein that regulates mitochondrial morphology and ROS levels (Sokolov et al., 2006), as a very
likely Ypk1 substrate, possibly providing insight into the molecular basis of the connection
between TORC2-Ypk1 signaling and ROS levels.
Similarly, several other prospects that were identified by our screen as very likely Ypk1
substrates remain to be validated. Such candidates include Muk1, a guanine nucleotide exchange
factor (GEF) for yeast Rab 5-type small GTPases (Vps21, Ypt52, and Ypt53) involved in
vesicle-mediated Golgi body-to-endosome trafficking (Paulsel et al., 2013), suggesting that
Ypk1 may also control switches that direct the flow of lipids. Muk1 is also intriguing for another
potential reason. In S. pombe, a Rab 5-like GTPase (Ryh1) and its Muk1-like GEF were
identified in a screen for TORC2 activators (Tatebe et al., 2010). Thus, if Ypk1-mediated
phosphorylation inhibits Muk1 function, it could represent a negative feedback mechanism
exerted on TORC2; conversely, if Ypk1-mediated phosphorylation stimulates Muk1 function, it
could represent a mechanism for self-reinforcing maintenance of TORC2 activity and, thus, a
high level of activated Ypk1. Clearly, by further investigating the physiological relevance of
these and other remaining candidates much new biology may be learned. Indeed, several gene
products of totally unknown function, such as Yhr097c and Ynr014w, as well as gene products
(e.g., Atg21 and Pex31) not previous linked to either TORC2 or Ypk1, if validated, may provide
new mechanistic insight into additional cellular processes regulated by TORC2-Ypk1 signaling.
In conclusion, our screening approach has uncovered a significant number of candidate
34
substrates. These appear to be high-quality candidates because the majority that have been tested,
to date, are functionally relevant in vivo substrates of Ypk1. Characterization of the remaining
substrates will very likely shed further light on aspects of cellular function that are regulated by
TORC2-Ypk1 signaling.
35
Figures
36
Fig. 3.1: A three-part screen to identify likely Ypk1 substrates. (A) The three-part screening
strategy to identify Ypk1 substrates is shown schematically as a flow chart. Numbers indicate the
number of hits/considered genes at each step in the screen. (B) The bioinformatic approach
towards identifying Ypk1 substrates is schematized as a flowchart with each filter as a box.
Genes were first filtered by MOTIPS on the basis of having likely phosphorylatable Ypk1
motifs. Subsequently, substrates were filtered by having many Ypk1 motifs or having a Ypk1
site known to be phosphorylated in published data sets. Lastly, genes were filtered by requiring
the gene to have a published chemical sensitivity like Ypk1 does, or a published interaction with
Ypk1, Ypk1 regulators (TORC2 or PP2A) or sphingolipid biosynthetic machinery.(C) A possible
explanation for Ypk1 synthetic dosage lethality interactions is shown. Normally, the cell has
enough kinase activity to buffer overexpression of a substrate (Substrate 2), so that essential
substrates are regulated and normal growth is unperturbed. However, concurrent decrease in
kinase activity coupled with substrate overexpression causes loss of regulation of essential
substrate(s) (Substrate 1) leading to observable growth defects. (D) ypk1-as ypk2Δ (yAM135 –
A) cells were transformed with PGAL1-GFP (negative control), PGAL1-Orm1 or PGAL1-Orm2
(known Ypk1 substrates, positive SDL controls) plasmids. Overnight cultures were then serially
diluted onto either dextrose (to repress substrate overexpression) or galatose (to induce substrate
overexpression) containing media with increasing concentrations of the Ypk1-as inhibitor 3-MBPP1. (E) GST-Orm1(1-85) (pFR203) and GST-Fps1(1-255) (pBT6) were purified from E. coli
and incubated with [γ-32P]ATP and Ypk1-as, purified from S. cerevisiae, in the absence or
presence of 3-MB-PP1. The products were then resolved by SDS/PAGE and analyzed as
described in chapter 2.
37
Fig. 3.2: Identified Ypk kinases substrates. Diagram of the TORC2-Ypk pathway from chapter
1. Known Ypk substrates and newly predicted substrates are grouped by the biological processes
(if known) to which the substrates contribute. Known substrates are shown in black whereas
newly predicted substrates are shown in red.
38
Tables
Table 3.1 Known Ypk1 substrates and potential substrates predicted by MOTIPS listed
under GO Slim terms.a
Gene
GPD1/YDL022W*
FPK1/YNR047W
FPK1(D621A)
[Kinase-dead
mutant]
ORM1/YGR038W
ORM2/YLR350W
AVO1/YOL078W
AVO2/YMR068W
BEM2/YER155C
BNI1/YNL271C
CDH1/YGL003C
ENT1/YDL161W
GIC2/YDR309C
LSB3/YFR024C-A
PAL1/YDR348C
SLA1/YBL007C
TSC11/YER093C
YHR097C
YSC84/YHR016C
MOTIPS
Sites
24P
37, 200,
244, 436,
481
37, 200,
244, 436,
481
52, 53
47, 48
552P,597,
1078
273P, 305,
407
83, 168,
1810,
1813
119,
1138P,
1533
51, 195,
213P
160P
90, 312,
345P
262P
49P, 391,
436
445, 447P,
449P, 477
19P, 97,
188
58, 288P,
294P
274, 374P
Chemical
Sensitivity/YeastMine
Interaction(s)
Known Ypk1 Substrates
YeastMine
SDL
Score
Ypk1
Dosage
Rescue
In vitro
substrate
+++
N/A
+
YeastMine
+
N/A
+
YeastMine
++
N/A
+
YeastMine
YeastMine
Cytoskeleton Organization
+++
++++
N/A
N/A
+
+
YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
Myr, YeastMine
-
N/A
N/A
AbA, Casp, YeastMine
-
N/A
N/A
AbA, YeastMine
TOXIC
-
N/A
YeastMine
-
N/A
N/A
Myr, AbA
-
N/A
N/A
YeastMine
-
N/A
N/A
AbA, Casp
++++
N/A
-
YeastMine
-
N/A
N/A
YeastMine
N/A
N/A
N/A
Myr
+++
N/A
+/-
Myr, YeastMine
-
N/A
N/A
39
Biological Process
Unknown
COM2/YER130C
ECM3/YOR092W*
ICS2/YBR157C
JIP4/YDR475C
KKQ8/YKL168C
RTS3/YGR161C*
SEG1/YMR086W
YDR186C
YHR080C
YNR014W
YPK3/YBR028C
EPL1/YFL024C
FKH1/YIL131C
GAL11/YOL051W
HCM1/YCR065W*
HOT1/YMR172W*
RLM1/YPL089C*
SMP1/YBR182C*
SSN2/YDR443C
YHP1/YDR451C*
BCK2/YER167W
ESC2/YDR363W
251, 370,
380
312, 350
14, 95,
136, 172P
348, 352,
592, 649
83, 113,
144, 146,
212, 293
30, 238P
56, 118P,
217, 634,
752P
334P,
540P,
542P, 620,
715P
401, 513,
667
54, 115,
156, 197
72, 73,
90P
24, 28, 61
404
1003P
80
387, 520,
586
20
20, 107
608P
180, 182
12, 38,
373, 430
114, 143,
145
Myr, YeastMine
-
N/A
N/A
Myr, AbA, YeastMine
-
N/A
N/A
Myr
-
N/A
N/A
Myr, AbA
N/A
N/A
N/A
YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
Myr
+
N/A
N/A
YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
YeastMine
+++
N/A
+
Myr, Casp
-
N/A
N/A
Transcription from RNA
Polymerase II Promoter
YeastMine
Myr, YeastMine
Myr, YeastMine
AbA, YeastMine
TOXIC
-
N/A
N/A
N/A
N/A
N/A
N/A
Myr
-
N/A
N/A
Myr
YeastMine
Myr
Myr
Mitotic Cell Cycle
TOXIC
TOXIC
-
+
N/A
N/A
N/A
+
N/A
N/A
YeastMine
TOXIC
-
N/A
YeastMine
-
N/A
N/A
40
PTK2/YJR059W
SET3/YKR029C
SWI4/YER111C
VHS2/YIL135C
ZDS1/YMR273C*
ZDS2/YML109W
59P, 82,
91, 171,
275
236, 405,
428
816P
316, 318,
325P
78, 370
183, 267,
345
Myr, YeastMine
+
N/A
N/A
YeastMine
-
N/A
N/A
Myr, YeastMine
-
N/A
N/A
Myr, Casp
-
N/A
N/A
AbA, YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
YeastMine
+
N/A
N/A
Myr
-
N/A
N/A
Myr, AbA, YeastMine
TOXIC
-
N/A
YeastMine
++++
N/A
-
Myr, YeastMine
-
N/A
N/A
Myr, Casp, YeastMine
-
N/A
N/A
YeastMine
-
N/A
N/A
Myr, AbA
-
N/A
N/A
N/A
-
N/A
+
Myr, AbA, YeastMine
-
N/A
N/A
Myr
Myr, YeastMine
+++
+++
N/A
N/A
+
+
24P
Myr, YeastMine
+++
N/A
+
16P
Myr, YeastMine
Cellular Ion Homeostasis
and Transport
-
N/A
+
Myr, AbA, YeastMine
-
N/A
N/A
YeastMine
-
N/A
+
Protein Phosphorylation
P
HAL5/YJL165C
KIN1/YDR122W
KIN2/YLR096W
KSP1/YHR082C
NPR1/YNL183C
SKY1/YMR216C
YAK1/YJL141C
YPL150W
P
17 , 217 ,
233
652, 791P,
879, 986P
665P, 818,
1020
594, 827P,
884P
125P,
255P,
257P, 317P
383P
128P, 206,
240
371P, 452,
890
Lipid Metabolic Process
ADR1/YDR216W
CDC1/YDR182W*
CKI1/YLR133W
GPT2/YKR067W
LAC1/YKL008C*
LAG1/YHL003C
LCB3/YJL134W
AVT3/YKL146W
b
CCH1/YGR217W
180, 230P,
756
9
P
14 , 25P,
30P
27, 652
23, 24
55, 59P,
172, 174
146, 148,
347
41
FPS1/YLL043W
MNR2/YKL064W
NHA1/YLR138W*
PPZ1/YML016C
DED1/YOR204W
HCR1/YLR192C*
HEF3/YNL014W*
RPL3/YOR063W
SUI2/YJR007W*
TEF1/YPR080W*
BPH1/YCR032W
CSR2/YPR030W
ROM2/YLR371W
SSD1/YDR293C
BRE5/YNR051C
EXO84/YBR102C
MUK1/YPL070W
RGP1/YDR137W
147, 181,
185, 570P
165, 620,
621, 826
544, 830
122, 203,
250P
P
84, 576
223
898
24P, 337
58
72P
1334,
1336,
1963
61, 103,
525, 987
76P, 193p,
396
164P,
482P, 503
398P
76, 313,
494, 554
173, 184P,
185P
220, 364P,
450, 452
Myr, YeastMine
+++++
N/A
+
AbA
-
N/A
N/A
Myr, YeastMine
-
N/A
N/A
Myr, YeastMine
TOXIC
-
N/A
-
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
N/A
Casp
-
N/A
N/A
Myr
-
N/A
N/A
YeastMine
-
N/A
N/A
Myr, AbA, YeastMine
TOXIC
-
N/A
Golgi Vesicle Transport
Myr, YeastMine
+++
N/A
-
YeastMine
-
N/A
N/A
Myr
+++
N/A
+/-
YeastMine
-
N/A
N/A
YeastMine
N/A
N/A
N/A
Myr, AbA
-
N/A
N/A
Myr, AbA, YeastMine
+++
N/A
-
YeastMine
N/A
N/A
N/A
Translation
YeastMine
Myr, AbA, YeastMine
Myr, AbA, YeastMine
Myr, AbA, YeastMine
Myr
Myr
Cell Wall Organization or
Biogenesis
Signaling
IRA2/YOL081W
GIS3/YLR094C*
MDS3/YGL197W
SYT1/YPR095C
882, 884,
1578,
1745,
3069
249, 333
757, 824,
842, 851,
1204
277P, 410,
728
DNA Replication
42
CDC13/YDL220C
CTI6/YPL181W
RIM4/YHL024W
314, 333P
155, 216P
93, 429,
525, 607
YeastMine
Myr, YeastMine
TOXIC
-
N/A
N/A
N/A
YeastMine
-
N/A
N/A
Myr
-
N/A
N/A
Myr
+++
N/A
+/-
YeastMine
-
N/A
N/A
Myr, YeastMine
-
N/A
N/A
Myr, AbA
TOXIC
-
+
Myr, YeastMine
+++
N/A
+
YeastMine
-
N/A
N/A
Myr
N/A
N/A
N/A
+++
N/A
-
-
N/A
N/A
++
N/A
+
-
N/A
N/A
+++
N/A
+
Endocytosis
ALY2/YJL084C
ROD1/YOR018W
ROG3/YFR022W
166P, 201,
225, 803
563, 617,
807, 823
425, 584,
718
Other
FRT1/YOR324C
HER1/YOR227W
YSP2/YDR326C
167, 201,
203, 228P,
385
28P, 102p,
157P
326, 518,
1237
RNA Catabolic Process
P
JSN1/YJR091C
PUF2/YPR042C
174, 275 ,
600
55, 143,
246, 902
Cytokinesis
CYK3/YDL117W
159, 207P,
746
DSN1/YIR010W
240, 250P
PEX31/YGR004W
432P
PAM1/YDR251W
471, 553P,
625
ATG21/YPL100W
237P
AbA, YeastMine
Chromosome Segregation
YeastMine
Peroxisome Organization
YeastMine
Pseudohyphal Growth
Myr, AbA, Casp,
YeastMine
Response to Starvation
Myr
a
Genes that are not bioinformatically predicted Ypk1 substrates, but contain Ypk1 motifs and
were included in this study are marked with an asterisk. SDL assay results are listed for each
bioinformatically predicted Ypk1 substrate. The scoring system reports growth phenotypes of the
ypk1-as ypk2Δ strain transformed with the indicated PGAL1-SUBSTRATE plasmid upon
overexpression on galactose with varying levels of 3-MB-PP1-imposed Ypk1-as inhibition. A
growth phenotype is defined as at least 1 serial dilution spot less growth than YCpLG-GFP
control at the given 3-MB-PP1 concentration. A strong growth phenotype is defined as no
growth at the given 3-MB-PP1 concentration. (+++++) indicates a growth phenotype with no 3MB-PP1. (++++) is a strong growth phenotype on 1 μM 3-MB-PP1. (+++) indicates a growth
43
phenotype on 1 μM 3-MB-PP1. (++) is defined as no phenotype on 1 μM 3-MB-PP1, but a
strong growth phenotype on 2 μM 3-MB-PP1. (+) indicates no phenotype on 1 μM 3-MB-PP1,
but a detectable growth phenotype on 2 μM 3-MB-PP1. (-) indicates no growth phenotype at any
concentration of 3-MB-PP1 tested. TOXIC indicates overexpression of the putative substrate on
galactose-containing media was deleterious to growth even in the wild-type control strain
(BY4741). These toxic genes were then tested for Ypk1 dosage rescue (for details, see Materials
and Methods); here, (+) indicates that Ypk1 overexpression could at least partially rescue the
overexpression toxicity of the indicated gene and (-) indicates that Ypk1 overexpression could
not rescue the overexpressiion toxicity. Lastly, the results of testing the indicated purified
predicted Ypk1 target as a substrtate in the in vitro protein kinase assay with purified Ypk1-as;
here, (+) indicates that Ypk1-as- dependent (3-MB-PP1 inhibitable) incorporation was detectable
for the substrate at a level comparable seen for incorporation into the positive control (the known
Ypk1 substrate, GST-Orm1(1-85); (+/-) indicates that readily detectable Ypk1-as-dependent
incorporation was found, but at a level lower than that seen for an equivalent amount of GSTOrm1(1-85) protein. (N/A) indicates that the indicated gene product was not tested in the
indicated assay. Genes listed in bold are those that are both SDL interactors and in vitro Ypk1
substrates and are considered hits in our screen. bThe Cch1 SDL assay was performed with a
plasmid constitutively overexpressing Cch1 under the TDH3 promoter (pBCT-CCH1H, (Iida et
al., 2007)), as efforts to generate a PGAL1-Cch1 vector failed.
44
Chapter 4: Ypk1 phosphorylates ceramide synthase to stimulate synthesis of
complex sphingolipids
This chapter is based on work published in Muir et al., 2014, eLife. I was responsible for all
in vivo experiments examining Ypk1 regulation of ceramide synthase and with Research
Specialist Françoise Roelants, Ph.D., I performed experiments examining Ypk1
phosphorylation of ceramide synthase in vitro. Postdoctoral researcher Subramaniam
Ramachandran, Ph.D., and I performed all the lipid analyses and in vitro ceramide
synthase experiments. Unpublished results on Ypk1 regulation of Atg21, in which I
performed all of the studies, are also reported here.
Introduction
The proteins Lac1 and Lag1 were identified as likely Ypk substrates from the screen described in
Chapter 3. Lac1 (418 residues) and Lag1 (411 residues) are apparent paralogs at the primary
sequence level (69% identity, 77% similarity) (Byrne and Wolfe, 2005) and are polytopic
integral proteins in the ER membrane with the predicted Ypk1 site in each protein residing near
its N-terminus (Fig. 4.1A). It has been shown that the N-termini of these proteins are exposed to
the cytosol (Kageyama-Yahara and Riezman, 2006). Along with a small accessory subunit Lip1
(150 residues), Lac1 and Lag1 are demonstrated constituents of the ceramide synthase complex
(Schorling et al., 2001; Vallée and Riezman, 2005), which catalyzes N-acylation of the free
amino group on the long-chain base (LCB), mainly phytosphingosine in yeast, with a preference
for very long chain fatty acyl-CoAs as the acyl donor, thereby forming phytoceramide (Fig.
4.1B). Genetically, Lac1 and Lag1 appear to play overlapping functional roles; lac1Δ or lag1Δ
single mutants are viable, whereas a lac1Δ lag1Δ double mutant is reportedly either inviable
(Jiang et al., 1998) or extremely slow growing (Barz and Walter, 1999; Schorling et al., 2001;
Vallée and Riezman, 2005). The ceramide synthase reaction lies at an important branch point in
the sphingolipid metabolic network (Fig. 4.1B) because de novo synthesis of ceramides both
consumes LCBs and prevents conversion of LCBs to their 1-phosphorylated derivatives
(LCBPs). Thus, the rate of ceramide synthesis is tightly coupled to the levels of both LCBs and
LCBPs (Kobayashi and Nagiec, 2003); and, moreover, the balance between ceramides and total
LCBs and LCBPs affects growth rate in both fungi (Kobayashi and Nagiec, 2003) and
mammalian cells (Spiegel and Milstien, 2003). By virtue of their position in the pathway, Lac1
and Lag1 are situated to be important regulators of this balance.
TORC2-Ypk signaling is already known to regulate the first step of sphingolipid biosynthesis via
Ypk1-mediated phosphorylation of Orm1 and Om2, alleviating their inhibition of the LCBproducing SPOTS complex (Roelants et al., 2011), and this regulation is important for surviving
sphingolipid depletion (Roelants et al., 2011) and membrane stretch under hypotonic stress
(Berchtold et al., 2012). Therefore, I sought to understand what type of control TORC2-Ypk1mediated phosphorylation exerts on Lac1 and Lag1 and what role their regulation of ceramide
synthase plays in sphingolipid homeostasis and metabolism.
45
Results
Global screen indicates Lac1 and Lag1are Ypk1 substrates
Among the bioinformatically predicted substrates, both Lac1 and Lag1 displayed a readily
detectable SDL phenotype (Fig. 4.2A) and both GST-Lac1(1-76) and GST-Lag1(1-80) served as
in vitro substrates for Ypk1, albeit with the phosphorylation of Lac1 being reproducibly much
more robust than Lag1 (Fig. 4.2B). Site-directed mutagenesis confirmed that the Ypk1-mediated
phosphorylation of both substrates occurred exclusively at the predicted phospho-acceptor
site(s), specifically Ser23 and Ser24 in both proteins (Fig. 4.2B).
Lac1 and Lag1 are phosphorylated by Ypk1 in vivo
To determine whether both Lac1 and Lag1 are phosphorylated in vivo in a Ypk1-dependent
manner and at their Ypk1 consensus sites, an integrated 3xHA-tagged version of each protein
and its corresponding S23A S24A mutant was expressed in yeast and extracts of the cells were
analyzed by phosphate-affinity SDS-PAGE (Phos-tag gels) (Kinoshita et al., 2009). In this
separation technique, the more highly phosphorylated the protein, the more its migration is
retarded. Both Lac1 (Fig. 4.3A, left) and Lag1 (Fig. 4.3A, right) migrated as two species, and the
slower of the two could be attributed to phosphorylation because it was eliminated if the sample
was pre-treated with calf intestinal phosphatase. This slower mobility species represented
phosphorylation at S23 and S24 because the band was also eliminated in Lac1(S23A S24A) and
Lag1(S23A S24A) mutants (Fig. 4.3A). I noted that phosphatase treatment, even of the
Lac1(S23A S24A) and Lag1(S23A S24A) mutants, resulted in appearance of a third, even faster
migrating species, presumably due to removal of a phosphorylation(s) elsewhere in these
proteins, consistent with indirect evidence that Lac1 and Lag1 might be subject to casein kinase
II (yeast Cka2)-dependent modification (Kobayashi and Nagiec, 2003).
In agreement with their relative efficacies as in vitro substrates (Fig. 4.2B), I found that,
reproducibly, the majority of Lac1 was present in the cell as the slower mobility isoform,
whereas the opposite was true for Lag1 (Fig. 4.3A). Because the behavior of Lac1 gave us
greater sensitivity of detection, and for the sake of conciseness, some of our subsequent findings
are illustrated with data for Lac1 only. However, all experiments were repeated with both
proteins with virtually identical results and conclusions.
To further confirm that Ypk1 is the protein kinase responsible for phosphorylation at Ser23
Ser24 in vivo, plasmids encoding 3xHA-tagged tagged Lac1 and Lag1 were introduced by DNAmediated transformation into wild-type, ypk1Δ and ypk2Δ strains. Cultures of the resulting cells
were grown in the absence or presence of a sub-lethal dose of myriocin, a condition which
several previous studies have shown activates TORC2- and Ypk1-mediated signaling (Roelants
et al., 2011; Berchtold et al., 2012; Sun et al., 2012), and the resulting extracts were analyzed on
Phos-tag gels. As observed previously for two other bona fide substrates, Orm1 and Orm2
(Roelants et al., 2011), absence of Ypk1 totally abrogated the appearance of phosphorylated
Lac1 (Fig. 4.3B) and phosphorylated Lag1 (data not shown), whereas elimination of Ypk2 had
no effect. Thus, Ypk1 is the paralog solely responsible for the observed in vivo phosphorylation
of Lac1 and Lag1 at Ser23 and Ser24. Furthermore, under conditions that stimulate TORC246
Ypk1 signaling, nearly all of the Lac1 (Fig. 4.3B) and much more of the Lag1 (data not shown)
were converted to the phosphorylated isoform indicating that TORC2 activation is relayed to
ceramide synthase via Ypk1. To further confirm that TORC2 signaling is essential for Ypk1
mediated phosphorylation of ceramide synthase, Lac1 phosphorylation was monitored in TOR2
and tor2-as strains after the addition of the tor2-as specific inhibitor BEZ-235 (Kliegman et al.,
2013). TORC2 inhibition led to rapid and sustained dephosphorylation of Lac1 (Fig. 4.3C),
confirming that TORC2 activity is necessary for Ypk1-mediated ceramide synthase
phosphorylation. Thus, our screening approach was successful in revealing two, previously
uncharacterized, cellular targets of the TORC2-Ypk1 signaling axis.
Lac1 and Lag1 phosphorylation increases under stress and is required for cell survival
The fact that impeding LCB production with a sub-lethal dose of the SPOTS inhibitor myriocin
stimulated Ypk1-mediated Lac1 and Lag1 phosphorylation suggested that this modification is
important for their physiological function. As one means to confirm that reduction in
sphingolipid biosynthesis capacity results in up-regulation of Lac1 and Lag1 phosphorylation,
we subjected the pathway to blockade near its end by treating the cells expressing integrated
3xHA-tagged Lac1 or Lag1 with aureobasidin A (Heidler and Radding, 1995), an antibiotic that
prevents formation of complex sphingolipids in yeast by inhibiting Aur1 (phosphatidylinositol:
ceramide phosphoinositol transferase) (Nagiec et al., 1997). As observed for treatment with
myriocin, the amount of phosphorylated Lac1 (Fig. 4.4A) and Lag1 (data not shown) was
markedly increased in response to treatment with aureobasidin A.
Another perturbation that has been shown to transiently up-regulate TORC2-Ypk1-mediated
signaling is heat shock (Sun et al., 2012). Consistent with that response, it has been shown
previously that heat shock leads to a transient increase in sphingolipid production and that
sphingolipid production is important for heat shock survival (Jenkins et al., 1997; Cowart et al.,
2003). Moreover, measurement of pathway intermediates and mathematical modeling also
suggested that a sharp spike of increased ceramide synthase activity may occur after heat shock
(Chen et al., 2013). Therefore, I subjected the same cells to heat shock and monitored Lac1 and
Lag1 phosphorylation at various times thereafter. Within 5 min, the amount of phosphorylated
Lac1 (Fig. 4.4B) and Lag1 (data not shown) increased markedly, but was back to the resting
level by 30 min
If these changes in phosphorylation state at Ser23 and Ser24 in Lac1 and Lag1 are important for
the metabolic adjustments that the cell needs to adapt appropriately, then preventing
phosphorylation at these sites should impair cell survival. To test this prediction, I integrated as
the sole source of ceramide synthase, mutant versions of Lac1 and Lag1 in which Ser23 and
Ser24 were mutated to Ala and, hence, cannot be phosphorylated under any circumstances. As a
control, I also generated integrated versions of Lac1 and Lag1 in which Ser23 and Ser24 were
mutated to Glu, to mimic conversion of the entire population to the phosphorylated state. Indeed,
I found that the cells co-expressing Lac1(S23A S24A) and Lag1(23A S24A) grew detectably
less well when challenged with either myriocin or aureobasidin A than either otherwise isogenic
wild-type cells or cells co-expressing Lac1(S23E S24E) and Lag1(S23E S24E) (Fig. 4.4C). This
growth difference could not be attributed to differences in protein levels between the ceramide
synthase alleles, as immunoblot analysis indicates similar steady state levels of ceramide
47
synthase components under these various stresses (Fig. 4.4D). Thus, Ypk1-dependent
phosphorylation of these sites in Lac1 and Lag1 is functionally important for cell survival in
response to the stress of limiting sphingolipid biosynthesis. Furthermore, the fact that the
Lac1(S23E S24E) Lag1(S23E S24E) strain phenocopied a LAC1+ LAG1+ strain under conditions
that promote phosphorylation of Lac1 and Lag1 indicates that these mutations generated
effective phosphomimetic alleles.
Calcineurin down-regulates Lac1 and Lag1 phosphorylation
As a means to delineate what cellular phosphatase is responsible for counteracting the Ypk1mediated phospho-regulation of Lac1 and Lag1, plasmid-borne 3xHA-tagged Lac1 was
expressed in a collection of deletion strains lacking each of the non-essential protein phosphatase
genes or their associated factors, and the phosphorylation state of Lac1 was assessed using Phostag gels. By this approach, I was unable to find any phosphatase-deficient mutant that exhibited a
significant increase in the amount of phosphorylated Lac1 compared to control cells (data not
shown). However, considerable genetic evidence indicates a strong connection between Ca2+
signaling and sphingolipid biosynthesis (Beeler et al., 1998). Moreover, it has been reported
previously that TORC2-Ypk-dependent regulation of sphinglipid biosynthesis is antagonized by
the action of the Ca2+/calmodulin-dependent protein phosphatase calcineurin (also known as
phosphoprotein phosphatase 2B), although the level at which the phosphatase acted was
unknown (Aronova et al., 2008). Hence, I conducted the converse experiment by stimulating the
cells expressing 3xHA-tagged Lac1 acutely with 0.2 M CaCl2, a condition known to robustly
activate calcineurin in yeast (Stathopoulos-Gerontides et al., 1999). Within <10 min, I found
total abrogation of phospho-Lac1 (Fig. 4.5A, left) and total abrogation of phospho-Lag1 (data not
shown) in wild-type cells, whereas in otherwise isogenic cna1Δ cna2Δ mutants (which lack both
calcineurin catalytic subunit paralogs) a substantial portion of the phosphorylated species
remained (Fig. 4.5A, right).
These results suggested that calcineurin may directly reverse the phoshorylations introduced into
Lac1 and Lag1 by Ypk1. However, it is also the case that two of the ancillary subunits associated
with TORC2, Slm1 and Slm2, are demonstrated calcineurin-binding proteins (Bultynck et al.,
2006; Tabuchi et al., 2006) and that calcineurin action appears to oppose TORC2-dependent
signaling (Mulet et al., 2006; Daquinag et al., 2007; Berchtold et al., 2012). Hence, it was
possible, therefore, that the observed loss of phospho-Lac1 and -Lag1 might be due to a Ca2+stimulated and calcineurin-mediated reduction in TORC2 and/or Ypk1 activity, rather than to
direct action of calcineurin on phospho-Lac1 and -Lag1. To rule out the former possibility, I
examined Lac1 and Lag1 phosphorylation in cells expressing constitutively-active
Ypk1(D242A), which I have demonstrated bypasses the need for its TORC2-dependent
activation (Roelants et al., 2011). I found that Ca2+ addition still led to nearly complete Lac1 and
Lag1 dephosphorylation in these cells (Fig. 4.5B). Furthermore, as judged by immunoblotting
with a phospho-site specific antibody, Pkh1- and Pkh2-dependent phosphorylation of the
activation loop in Ypk1 (Fig. 4.5C) [or in Ypk1(D242A) (data not shown)] was not diminished
in cells stimulated with Ca2+, confirming that there was no calcineurin-mediated decrease in the
amount of active Ypk1 present. Therefore, it seems clear that calcineurin dephosphorylates the
sites in Lac1 and Lag1 phosphorylated by Ypk1 directly, rather than through down-regulation of
Ypk1 function.
48
Ypk1-mediated phosphorylation of Lac and Lag1 stimulates ceramide synthase activity
Given the phenotypic evidence that TORC2-Ypk1-dependent modulation of Lac1 and Lag1 is
physiologically important (Fig. 4.4C), I next conducted biochemical analysis to determine how
Ypk1-mediated phosphorylation affects the function of Lac1 and Lag1 in ceramide synthesis.
Toward that end, I first used LC-MS to monitor the levels of LCBs and LCBPs extracted from
equivalent numbers of cells from exponentially-growing cultures of wild-type cells or otherwise
isogenic cells expressing as the sole source of Lac1 and Lag1 either the non-phosphorylatable
Lac1(S23A S24A) and Lag1(S23A S24A) mutants or the phosphomimetic Lac1(S23E S24E)
and Lag1(S23E S24E) mutants. Relative to wild-type cells, the Lac1(S23A S24A) Lag1(S23A
S24A) cells accumulated significantly more PHS (Fig. 4.6A, top left), as well as more
dihydrosphingosine (DHS) (Fig. 6A, top right), which is a more minor LCB in yeast (note the
difference in scale of the ordinate), whereas the Lac1(S23E S24E) Lag1(S23E S24E) cells
displayed a level of both PHS and DHS that was somewhat lower, and a similar trend was
observed even for PHS-1-P, an even less abundant metabolite (Fig. 4.6A, bottom left). The level
of DHS-1-P (Fig. 4.6A, bottom right) was so low as to make reliable measurement difficult, but
no differences between strains could be detected. Given that the ceramide synthase complex is
responsible for the conversion of LCBs into ceramides, these findings indicate that, in the
absence of Ypk1-mediated phosphorylation, the rate of LCB utilization is significantly reduced,
consistent with the conclusion that Ypk1-dependent modification of Lac1 and Lag1 promotes
ceramide synthase function.
As an independent means to measure flux through the sphingolipid pathway, and because all
complex sphingolipids in yeast contain inositol-phosphate, an equivalent number of cells of the
same three strains were pulse-labeled, in triplicate, with [32P]PO4-3, and the acidic sphingolipids
extracted and analyzed by thin-layer chromatography. Strikingly, the amount of complex
sphingolipids generated during the pulse was higher in the Lac1(S23E S24E) Lag1(S23E S24E)
cells than in the wild-type controls, while Lac1(S23A S24A) Lag1(S23A S24A) cells generated
lower levels of complex sphingolipids (Fig. 4.6B). This finding is again consistent with the
conclusion that Ypk1-mediated phosphorylation of Lac1 and Lag1 stimulates the production of
the ceramide precursors to complex sphingolipids.
Ypk1-mediated phosphorylation could stimulate the ceramide synthase reaction in vivo by
stabilizing Lac1 and Lag1 thereby increasing their steady-state level, by enhancing their
association with the small ancillary subunit Lip1, and/or by direct activation. Immunoblotting of
exponentially-growing cultures expressing the 3xHA-tagged versions of wild-type Lac1 and
Lag1 and the Lac1(S23A S24A) Lag1(S23A S24A) and Lac1(S23E S24E) Lag1(S23E S24E)
indicated no discernible difference in their steady-state level (see Fig. 4.4D). Likewise, in cells
co-expressing the same proteins and FLAG-tagged Lip1 (gift of Howard Riezman, Univ. of
Geneva), I observed no difference in the efficiency of Lip1 co-immunoprecipitation between
wild-type Lac1 and Lag1 and either the Lac1(S23A S24A) Lag1(S23A S24A) or Lac1(S23E
S24E) Lag1(S23E S24E) mutants (data not shown). These results suggested that Ypk1-mediated
phosphorylation may directly enhance the catalytic efficiency of Lac1 and Lag1. To test this
possibility directly, 3xFLAG-tagged versions of Lac1 and Lag1 were immuno-purified from
detergent-solubilized microsomes isolated from exponentially-growing cells and the resulting
protein assayed in vitro, monitoring the formation of ceramide from PHS and steroyl-CoA by
49
LC-MS. No product was observed in the absence of added steroyl-CoA (data not shown), and
product formation was reduced by 85% in the presence of PHS and steroyl-CoA if 1 μM
australifungin, a demonstrated and specific ceramide synthase inhibitor (Mandala and Harris,
2000), was added (data not shown), confirming that the reaction measured was catalyzed by
ceramide synthase. I found that the specific activity of the Lac1(S23E S24E) Lag1(S23E S24E)
complex was reproducibly ~2 higher than either the Lac1(S23A S24A) Lag1(S23A S24A)
mutant or the wild-type complex (Fig. 4.6C upper panel), similar to the degree of difference in
sphingolipid metabolites between these same strains that I measured by other means (Fig. 4.6A
and 4.6B). In two independent trials (each performed in triplicate), the identical trend was found
when microsomes from these same cells were assayed directly (i.e. without detergent
solubilization and enzyme enrichment by immunoprecipitation (data not shown). Additionally, I
found that immunopurified ceramide synthase from wild-type cultures treated with myriocin had
higher activity than ceramide synthase prepared from untreated cells (Fig. 4.6C, lower panel).
Furthermore, this increase in ceramide synthase activity in response to myriocin treatment was
not observed in Lac1 (S23A S24A) Lag1 (S23A S24A) cells, consistent with TORC2-Ypk1
signaling increasing ceramide synthase activity by phosphorylation at these residues. These
findings are also in agreement with a reported ~2 fold decrease in the rate of ceramide
production by ceramide synthase complex isolated from TORC2-deficient yeast (Aronova et al.,
2008). Hence, I conclude that Ypk1 phosphorylation directly increases the catalytic activity of
ceramide synthase.
Ypk1-mediated stimulation of Lac1 and Lag1 prevents autophagy induction during
TORC2-driven up-regulation of sphingolipid biosynthesis
The observed increase in ceramide synthase activity in response to Ypk1-mediated
phosphorylation could serve two roles that are not mutually exclusive: (i) to produce more
ceramide and the derived complex sphingolipids; and, (ii) to prevent inadvertent accumulation of
LCBs and the derived LCBPs when TORC2-driven Ypk1 activation stimulates metabolic flow
into the sphingolipid pathway by alleviating Orm1- and Orm2-imposed inhibition of SPOTS (see
Fig. 4.8). I reasoned that if the latter were one of the important physiological functions of Ypk1dependent stimulation of ceramide synthase, then TORC2 activation would be detrimental to
cells in which the Lac1(S23A S24A) Lag1(S23A S24A) mutant is the sole source of this
enzyme. To mimic TORC2-stimulated elevation of Ypk1 activity, the constitutively-active
Ypk1(D242A) allele (hereafter referred to as Ypk1*) was expressed from the YPK1 promoter on
a CEN plasmid in either wild-type cells or the Lac1(S23A S24A) Lag1(S23A S24A) mutant.
Indeed, compared to the empty vector control, expression of Ypk1* was well tolerated by cells
containing wild-type Lac1 and Lag1, but deleterious to the growth of the Lac1(S23A S24A)
Lag1(S23A S24A) mutant cells, whether measured on agar plates (Fig. 4.7A, top) or in liquid
culture (Fig. 4.7A, bottom).
The most likely metabolic perturbation responsible for the observed decrease in growth in the
Lac1(S23A S24A) Lag1(S23A S24A) expressing Ypk1* is the accumulation of either LCBs or
LCBPs. I asked if levels of these metabolites were altered in Ypk1* expressing cells. As
previously noted (see Fig. 4.6A) Lac1(S23A S24A) Lag1(S23A S24A) cells had higher PHS
levels than wild-type controls, but the addition of the Ypk1* plasmid caused no further increase
(Fig. 4.7B upper panel). This indicates even in the absence of TORC2-Ypk1 mediated ceramide
50
synthase regulation, cells are able to buffer the levels of LCBs even with active TORC2
signaling. However, the addition of Ypk1* did cause an increase in PHS-1-P levels in
Lac1(S23A S24A) Lag1(S23A S24A) but not wild-type cells (Fig. 4.7B lower panel). Thus, I
reasoned that abnormally high LCBP levels might cause the growth defect observed in Ypk1*
expressing Lac1(S23A S24A) Lag1(S23A S24A) cells. If accumulation of LCBPs in Lac1(S23A
S24A) Lag1(S23A S24A) co-expressing Ypk1* is indeed responsible, then reduction in LCBP
synthesis by elimination of the gene LCB4, which encodes the major LCB kinase (Nagiec et al.,
1998), should alleviate the growth inhibition. As expected, introduction of an lcb4∆ null
mutation into the Lac1(S23A S24A) Lag1(S23A S24A) mutant suppressed the growth inhibitory
effect of Ypk1* (Fig. 4.7C). Conversely, and consistent with toxicity arising from accumulation
of LCBPs when Lac1 and Lag1 cannot be stimulated by Ypk1, I found that the poor growth
phenotype of cells lacking the gene (LCB3) encoding the major LCBP phosphatase (Mandala et
al., 1998), was markedly exacerbated by introduction of the Lac1(S23A S24A) Lag1(S23A
S24A) alleles, but not by introduction of the Lac1(S23E S24E) Lag1(S23E S24E) alleles (Fig.
4.7D). In fact, and strikingly, presence of the Lac1(S23E S24E) Lag1(S23E S24E) allele
afforded nearly complete rescue of the slow-growth phenotype of the lcb3∆ mutation (Fig.
4.7D). Consistent with the toxicity of LCBPs when LCB utilization by ceramide synthase is
inefficient, an elo3 mutation, which prevents efficient formation of C26-CoA (an acyl chain
found in yeast ceramides), was synthetically lethal with lcb3∆ (Kobayashi and Nagiec, 2003).
It has been reported that aberrant increases in LCBP level impede growth by triggering
autophagy even under nutrient-rich conditions (Zimmermann et al., 2013). In agreement with the
poor growth arising from LCBP-evoked induction of autophagy, I found that preventing
autophagy by deletion of ATG1, which encodes a protein kinase necessary for induction of
autophagophore formation and its elongation (Papinski and Kraft, 2014), provided substantial
rescue of the growth debilitating effect of Ypk1* expression in Lac1(S23A S24A) Lag1(S23A
S24A) mutant cells (Fig. 4.7F). This rescue was not a result of ATG1 mutation indirectly
affecting LCBP levels as LCBPs did not significantly change in atg1Δ strains and still
accumulated in Ypk1* expressing atg1Δ Lac1(S23A S24A) Lag1(S23A S24A) cells (Fig. 4.7G).
Collectively, these results indicate that, aside from stimulating ceramide synthesis per se, a
physiologically important role of Ypk1-dependent ceramide synthase activation is also to prevent
accumulation of LCBPs and thereby avoid inadvertent induction of autophagy under nutrientsufficient conditions (Fig. 4.8).
Is Atg21 a physiologically-relevant Ypk1 substrate?
Given that TORC2-Ypk signaling appears to play an important role in preventing autophagy
induction due to LCBP accumulation, I was also interested in the fact that Atg21, a PtdIns3Pbinding -propeller (WD-40 repeat) protein required for vesicle formation in the cytoplasm-tovacuole targeting (Cvt) pathway (Krick et al., 2008; Nair et al., 2010), was another potential
Ypk1 substrate that I identified in my screen (see Fig. 3.2). CVT is a form of selective autophagy
that occurs under basal conditions and is used to deliver certain enzymes into the vacuole
(Lynch-Day & Klionsky, 2010; Umekawa & Klionsky, 2012). Therefore, I asked whether I
could detect any alteration of CVT pathway function either in cells with diminished Ypk activity
or in cells expressing mutant alleles of Atg21 where the predicted Ypk sites were mutated either
to Ala to mimic the absence of Ypk1-mediated phosphorylation or to Glu to mimic permanent
51
Ypk1-mediated phosphorylation. To examine CVT pathway activity, I monitored the status of
GFP-Atg8 and the processing of Ape1 under nutrient rich (basal autophagy) conditions. Under
these circumstances, proteolytic maturation of both these proteins requires CVT pathway activity
and Atg21 function (Umekawa and Klionsky, 2012). However, I did not observe any significant
differences in these two biochemical read-outs of CVT pathway activity, compared to control
cells, when Ypk activity was inhibited (Fig. 4.9A) or for cells expressing either type of Atg21
phospho-site mutant (Fig. 4.9B). Recently, it has been reported that Ypk1 activity is important
for autophagy initiated in response to amino acid starvation, but not in response to general
nitrogen source starvation (Vlahakis et al., 2014). Processing of GFP-Atg8 provides a robust
read-out for amino acid starvation evoked autophagy (Guimaraes et al., 2015). Hence, I
monitored the status of GFP-Atg8 in response to leucine limitation in strains expressing my
Atg21 phospho-site mutants. Again, however, I observed no difference between control cells and
cells expressing either class of Atg21 alleles in the predicted Ypk1 sites (Fig. 4.9C).
Thus, I have not yet uncovered any evidence that Ypk1-mediated phosphorylation of Atg21
plays any role in controlling its function. It is possible that pulse-chase experiments might
provide a better way to observe whether the kinetics of the CVT and autophagy pathway are
affected than the steady-state analysis I conducted. Furthermore, I have not yet conclusively
shown that Atg21 is phosphorylated at its predicted Ypk sites in vivo. Thus, future experiments
examining Ypk1 phosphorylation of Atg21 in vivo, and more careful phenotypic analysis of the
consequences of this phosphorylation, will be necessary to determine if Atg21 is an authentic
Ypk1 substrate or a false positive from my screen.
Discussion
Mechanism of phosphoregulation of ceramide synthase
As I have demonstrated here, TORC2-dependent Ypk1-mediated phosphorylation of Lac1 and
Lag1 stimulates the function of the ceramide synthase complex. Consistent with our findings, a
previous study found a consistent decrease in ceramide synthase activity in microsomal fractions
isolated from cells in which TORC2 had been inactivated (and thus Ypk1 activity was
presumably reduced) (Aronova et al., 2008), although indirect effects of the loss of TORC2
function on ceramide synthase activity could not be ruled out. Our findings make it clear that the
role of TORC2 is to promote the Ypk1-dependent phosphorylation of Lac1 and Lag1 subunits of
this enzyme. However, the precise molecular mechanism by which this post-translational
modification stimulates this enzyme is still not completely clear. In this regard, it has been
shown that mammalian ceramide synthase activity increases upon heterodimerization of different
catalytic subunit isoforms (Laviad et al., 2012). However, as judged by co-immunoprecipitation,
I found no difference in the state of Lac1-Lag1 association between the wild-type proteins and
either our Lac1(S23A S24A) Lag1(S23A S24A) or Lac1(S23E S24E) Lag1(S23E S24E)
mutants (data not shown). As mentioned in Results, I found no difference in the steady-state
level of these same complexes or in their content of Lip1, a non-catalytic component of the
complexes also essential for ceramide synthase activity (Vallée and Riezman, 2005). Thus,
understanding of how phosphorylation of Ser23 and Ser24 in Lac1 and Lag1 stimulate ceramide
synthase activity may require detail structural information, which will be challenging to obtain
for these polytopic integral membrane proteins.
52
TORC2-Ypk1 control of ceramide synthesis is conserved
Although the enzymic steps that carry out sphingolipid biosynthesis have been largely
elucidated, much less was known, until recently, about regulation of these enzymes (Breslow &
Weissman, 2010; Breslow, 2013). The first insight came when it was demonstrated (Roelants et
al., 2011; Berchtold et al., 2012; Sun et al., 2012) that, in response to membrane stresses,
including treatment with myriocin and aureobasidin A, TORC2-Ypk1 signaling is activated and
alleviates inhibition of the SPOTS complex by phosphorylating the negative regulatory proteins
Orm1 and Orm2 (Fig. 8). As a direct consequence, the rate of de novo production of the LCB
precursor to sphingolipids is increased. Although TORC2 signaling had been implicated in
promoting synthesis of ceramide, the product of LCB N-acylation (Aronova et al., 2008), it was
unknown whether that role was simply the result of TORC2-Ypk1-dependent stimulation of
SPOTS function and the resulting increase in LCB supply. As I demonstrated here, Ypk1mediated phosphorylation of Lac1 and Lag1 also increases in response to both myriocin and
aureobasidin A, suggesting that ceramide synthesis per se, and not simply general elevation of
LCB levels, is important for allowing the cells to cope with the effects of these antibiotics.
Indeed, collectively, the findings I describe here demonstrate unequivocally that, in addition to
up-regulation of SPOTS, TORC2-Ypk1 exerts direct control on the ceramide synthase step of the
sphingolipid biosynthetic pathway by phosphorylating and stimulating the function of the Lac1
and Lag1 subunits of the ceramide synthase complex. Moreover, as I also demonstrated, the
ceramide synthase reaction represents an important branch point in sphingolipid biosynthesis
(Figs. 2B and 8). LCBs produced by the SPOTS reaction can either be converted to ceramides or
become phosphorylated by LCB kinase Lcb4 (and its paralog Lcb5) to form LCBPs (Nagiec et
al., 1998). Accumulation of LCBPs has been shown to be toxic to yeast cell growth (Kim et al.,
2000), as I have also confirmed here, at least in large part because, as is now known, these
metabolites trigger inappropriate induction of autophagy (Zimmermann et al., 2013). Thus, the
rate of ceramide production must be properly adjusted to maintain the pool of LCBs and derived
LCBPs at a non-deleterious level, in agreement with evidence in yeast and other organisms that
ceramides and LCBPs generally play antagonist roles and must be maintained in the proper
dynamic balance (Kobayashi and Nagiec, 2003; Spiegel and Milstien, 2003; Kihara et al., 2007;
Dickson, 2008; Breslow and Weissman, 2010; Bikman and Summers, 2011). Hence, the function
I have discovered and described here for TORC2-Ypk1 in stimulating ceramide synthase
promotes utilization of the increased LCB generated upon TORC2-Ypk1-mediated up-regulation
of SPOTS. This metabolic control has multiple clear-cut physiological benefits to the cell: (a)
directing flow in the sphingolipid pathway toward complex sphingolipids to populate the plasma
membrane barrier; (b) reduction of the level of potentially toxic LCBPs; and, (c) avoidance of
inappropriate induction of autophagy under nutrient-sufficient conditions (Fig. 8). Indeed, our
results indicate that a significant role for the coordination exerted by TORC2-Ypk1 between the
level of SPOTS activity and the level of ceramide synthase activity is to prevent metabolic
"cross-talk" to the autophagy pathway.
If this function of TORC2-Ypk1 signaling is important, then it should be conserved. Indeed,
alignments of the primary structures of Lac1 and Lag1 homologs predicted from sequenced
fungal genomes, from Saccharomyces sensu stricto species to the very distantly related Ustilago
maydis, nearly all contain a basophilic sequence that matches the Ypk1 phospho-acceptor site
consensus and is located, as in S. cerevisiae Lac1 and Lag1, in their N-terminal cytosolic
53
extensions, suggests that Ypk1-related protein kinases may regulate ceramide synthase across
essentially all fungal species. Consistent with this suggestion, a genetic interaction between a
Ypk1 homolog (YpkA) and a Lac1/Lag1-like ceramide synthase component (BarA1) has been
reported in Aspergillus nidulans (Colabardini et al., 2013). However, this mode of regulation
may not extend to multicellular eukaryotes; of the six distinct ceramide synthase catalytic
subunts encoded in the human genome, only CerS6 has a sequence that resembles a Ypk1 site
(43-R-x-R-x-x-x-S-Hpo-50), and it serves as only a weak phospho-acceptor for SGK1, the
human ortholog of Ypk1 (S. Ramachandran and J. Thorner, unpublished results).
Orm1 and Orm2, when phosphorylated at unique sites by protein kinase Npr1, appear to
promotes steps in the sphingolipid pathway that lead to more complex sphingolipids
(Shimobayashi et al., 2013). In contrast to Ypk1, which is activated by TORC2, Npr1 is inhibited
by TORC1 (MacGurn et al., 2011). Thus, this control mechanism will only be exerted under
conditions that inactivate TORC1, such as amino acid starvation (Loewith and Hall, 2011), a
condition that presumably requires adjustment of both plasma membrane lipid and protein
composition to maximize the cell's ability to scavenge and assimilate nutrients. Under the same
condition, autophagy is induced because, like Npr1, TORC1 also negatively regulates the
autophagy-inducing protein kinase Atg1-Atg13 complex (Alers et al., 2014). Conversely, under
nutrient sufficient conditions, TORC1 is active, and phosphorylates and stimulates protein kinase
Sch9. Interestingly, Sch9 action should act in concert with TORC2-Ypk1 signaling to help keep
the levels of LCBs and LCBPs low and ceramides high. This is likely because Sch9 promotes
transcriptional repression of genes (YDC1 and YPC1) that encode ceramidases and inhibits a
phosphosphingolipid phospholipase C (Isc1) that hydrolyzes complex sphingolipids (Swinnen et
al., 2014). Such complex multi-component controls may be a general feature of signaling
modalities that interface with biosynthetic pathways that have intermediates, like LCBPs, that are
not innocuous, but are themselves bioactive metabolites.
Calcineurin negatively regulates ceramide synthase
Prior studies had suggested that calcineurin negatively regulates sphingolipid production via
effects on the function of the ancillary TORC2 subunits, Slm1 and Slm2 (Bultynck et al., 2006;
Mulet et al., 2006; Tabuchi et al., 2006; Daquinag et al., 2007), although the molecular
connection between Slm1 and Slm2 and sphingolipid biosynthesis was unclear. Subsequently, it
was observed that, in cells lacking the regulatory subunit (Cnb1), there was an increase in C26containing ceramides, suggesting that calcineurin somehow antagonizes TORC2-dependent
signaling (Aronova et al., 2008). I found that calcineurin negatively regulates ceramide synthesis
by directing the dephosphorylation of Lac1 and Lag1. This Ca2+-activated calcineurin-dependent
dephosphorylation occurred even in cells expressing a TORC2-independent constitutively-active
Ypk1 allele. Moreover, I showed here that calcineurin does not affect Pkh1- (and Pkh2-)
mediated phosphorylation of the activation loop of Ypk1, and I demonstrated previously that
presence or absence of calcineurin does not alter TORC2-mediated phosphorylation of Ypk1 or
cause any substantial change in Ypk1 specific activity (Roelants et al., 2011). Thus, direct downmodulation of either TORC2 or Ypk1 by calcineurin cannot account for the negative regulation it
exerts on sphingolipid biosynthesis. What our findings now make clear is that calcineurin
negatively regulates the sphingolipid pathway at the level of ceramide synthesis, at least in large
part, by direct dephosphorylation of the stimulatory phosphorylations in the ceramide synthase
54
subunits Lac1 and Lag1 that are installed by TORC2-Ypk1 signaling. Calcineurin recognizes
substrates via a docking motif (PxIxIT or variants thereof), typically also accompanied quite a
distance upstream by a secondary docking site (LxVP) (Roy and Cyert, 2009). However, these
sites can be quite degenerate; for example, the more hydrophobic variant PVIVIT is much more
potent in recruiting calcineurin when it is used to replace the native "PxIxIT" sequences in either
the transcription factor Crz1 (PIISIQ) (Roy et al., 2007) or the endocytic adaptor Aly1 (PILKIN)
(O'Donnell et al., 2013). In both Lac1 and Lag1, there is a similar hydrophobic sequence located
at the identical position in both proteins (355PIVFVL360). Whether this or any other degenerate
match represents a calcineurin-binding site remains to be determined.
55
Figures
Fig. 4.1: Lac1 and Lag1 are components of the ceramide synthase enzyme complex. (A) A
diagram of the membrane topology of Lac1 and Lag1 derived from (Kageyama-Yahara and
Riezman, 2006). Lac1 and Lag1 are experimentally determined to have 6 transmembrane
domains. The N terminus of these proteins is cytosolic and therefore accessible for Ypk1
phosphorylation. (B) Diagram of yeast de novo sphingolipid biosynthesis derived from (Dickson,
2008). Skeleton structures of metabolites are shown and enzymes are indicated as ovals.
Metabolites shaded in green are those directly produced or derived from ceramide synthase while
those in red are alternative products at the ceramide synthase branch point.
56
Fig. 4.2: Genetic and biochemical screening indicates Lac1 and Lag1 are Ypk1 substrates.
(A) SDL results for Lac1 and Lag1. ypk1-as ypk2Δ (yAM135 – A) or wild-type (BY4741) cells
were transformed with PGAL1-GFP (negative control), PGAL1-Lac1 or PGAL1-Lag1 plasmids. The
SDL assay was then performed as described in Materials and Methods. (B) GST-Lac1(1-76)
(pAX131), GST-Lag1(1-80)(pFR29), GST-Lac1(1-76)(S23A S24A)(pAX132) and GST-Lag1(180)(S23A S24A)(pAX133) were purified from E. coli and Ypk1-as kinase assays were
performed as described in chapter 2.
57
Fig. 4.3: Ypk1 phosphorylates Lac1 and Lag1 at S23 and S24 in vivo. (A) 3xHA-Lac1
(yAM165-A) 3xHA-Lac1(S23A S24A) (yAM166-A), 3xFLAG-Lag1 (yAM159-A) and
3xFLAG-Lag1(S23A S24A) (yAM163-A) strains were grown exponentially in YPD. Cells were
then harvested and whole-cell extracts were prepared. Extracts were split and one fraction was
then treated with calf intestinal phosphatase. The extracts resolved by Phos-tag SDS-PAGE and
immunoblotted with HA or FLAG and Pgk1 (loading control) antibodies. P-Lac1 and P-Lag1
indicate the band corresponding to S23 S24 phosphorylation. * indicates a non-specific band that
appears in HA blots of yeast whole cell extracts. (B) Wild-type (BY4741), ypk1Δ (JTY6142) and
ypk2Δ (yAM120 – A) strains were transformed with a plasmid centromeric plasmid encoding
3xHA-Lac1 expressed under control of its endogenous promoter (pAX136). Cells were grown to
mid-exponential phase and then treated with a sublethal dose of myriocin (0.625 μM) or
methanol (vehicle) for 2 h. Cell extracts were then prepared, resolved by Phos-tag SDS-PAGE
and blotted as above in (A). (C) TOR2 (yKL04) or tor2-as (yKL05) strains were transformed
with a 3xHA-Lac1 expressing plasmid (pAX136). Cells were grown to mid-exponential phase
and treated with 1 uM BEZ-235 for the indicated times. Cell extracts were then prepared,
resolved by Phos-tag SDS-PAGE and blotted as above in (A).
58
Fig. 4.4: Enhanced Ypk1 phosphorylation of Lac1 and Lag1 under sphingolipid and heat
stress. (A) 3xHA-Lac1 (yAM165-A) cells were grown to early exponential phase in YPD.
Cultures were then treated with sublethal doses of myriocin (0.625 μM) or methanol (vehicle) or
aureobasidin A (1.8 μM) of ethanol (vehicle) for 2 h. (B) 3xHA-Lac1 (yAM165-A) cells were
grown to exponential phase in YPD at 30˚C. A sample of this culture was then harvested. The
remaining culture was then moved to 42˚C to initiate heat shock and samples were harvested at
the indicated time points. Whole cell extracts were prepared from each sample, resolved by
Phos-tag SDS-PAGE, and immunoblotted as in Figure 4.3. P-Lac1 indicates the band
corresponding to S23 S24 phosphorylation. * indicates a non-specific band that appears in HA
blots of yeast whole cell extracts. (C) LAC1 LAG1 (yAM205 - A), Lac1SSAA Lag1SSAA (yAM207
- B) and Lac1SSEE Lag1SSEE (yAM210) were grown to exponential phase in YPD. Serial dilutions
of each culture were made and spotted on YPD plates containing vehicle or the indicated
concentration of myriocin or aureobasidin A. Cells were allowed to grow for 3 days at 30˚C prior
to imaging. (D) 3xHA-Lac1::HIS3 3xFLAG-Lag1::LEU2 (yAM168) 3xHA-Lac1(S23A
S24A)::HIS3 3xFLAG-Lag1(S23A S24A)::LEU2 (Lac1SSAA Lag1SSAA) (yAM184) and 3xHALac1(S23E S24E)::HIS3 3xFLAG-Lag(S23E S24E)1::LEU2 (Lac1SSEE Lag1SSEE) (yAM192 –
A) cells were treated with 1.0 uM myriocin or 18.2 nM aureobasidin A for 2h. Extracts were
prepared from each sample, resolved by SDS-PAGE, and immunoblotted as indicated.
59
Fig. 4.5: Activation of calcineurin leads to rapid dephosphorylation of Lac1 and Lag1
without affecting Ypk1 function. (A) Wild-type (BY4741) or cna1Δ cna2Δ (JTY5574) strains
were transformed with a centromeric plasmid encoding 3xHA-Lac1 expressed under control of
its endogenous promoter (pAX136). Cultures were grown to mid-exponential phase in selective
media and then treated with 200 mM CaCl2 for 10 minutes to activate calcineurin. Cultures were
then harvested and whole cell extracts were prepared from each sample, resolved by Phos-tag
SDS-PAGE, and immunoblotted as in Figure 4.3. (B) 3xHA-Lac1 (yAM165-A) cells were
transformed with a centromeric plasmid encoding the hyperactive TORC2 independent
Ypk1D242A allele expressed under its own promoter (pFR273) or vector (pRS316). Cultures were
grown in selective media to mid-exponential phase before treatment with 200 mM CaCl2 for 10
min Cultures were then harvested and whole cell extracts were prepared from each sample,
resolved by Phos-tag SDS-PAGE, and immunoblotted as above. (C) 3xFLAG-Ypk1 (YDB379)
cells were grown to midexponential phase in YPD. Cultures were treated with or without 200
mM CaCl2 for 10 min Cells were then harvested and whole extracts prepared. Extracts were
resolved by SDS-PAGE and blotted with anti-pSGK(T256), which recognizes Ypk1 T504
activation loop phosphorylation and anti-FLAG antibody (Ypk1 loading control). The blot is
representative of triplicate samples and the quantitation of the ratio of pT504/FLAG from these
replicates is shown below the blot.
60
61
Fig. 4.6: Ypk1 phosphorylation of Lac1 and Lag1 stimulates ceramide synthase activity.
(A) Cultures of LAC1 LAG1 (yAM205 - A), Lac1SSAA Lag1SSAA (yAM207 - B) and Lac1SSEE
Lag1SSEE (yAM210) strains were grown to mid-exponential phase and then harvested.
Sphingolipids were extracted and analyzed as described in chapter 2. Values represent the mean
of three independent experiments (each performed in triplicate) and error bars represent SEM.
(B) Triplicate exponentially growing cultures of LAC1 LAG1 (yAM205 - A), Lac1SSAA Lag1SSAA
(yAM207 - B) and Lac1SSEE Lag1SSEE (yAM210) were adjusted to OD600 = 1.0. Complex
sphingolipids were labeled and analyzed by thin layer chromatography (TLC) as in chapter 2. A
representative TLC plate is shown with the origin at the bottom of the image. The assigned
identity of species as IPCs and MIPCs was confirmed by pharmacological or genetic inhibition
of the production of these species in control cultures (data not shown). Quantification of total
complex sphingolipids was performed in ImageJ by integrating the phosphorimager screen
intensity across the lane for each sample and normalized to 1 for wild-type ceramide synthase
samples. (C) (upper) Ceramide synthase was purified by FLAG immunopurification from 3xHALac1::HIS3 3xFLAG-Lag1::LEU2 (yAM168) 3xHA-Lac1(S23A S24A)::HIS3 3xFLAGLag1(S23A S24A)::LEU2 (Lac1SSAA Lag1SSAA) (yAM184) and 3xHA-Lac1(S23E S24E)::HIS3
3xFLAG-Lag(S23E S24E)1::LEU2 (Lac1SSEE Lag1SSEE) (yAM192 – A) cells.
Immunoprecipitates were then split into 3 fractions and in vitro ceramide synthase assays (60
min reactions) were performed in triplicate. A small sample of each ceramide synthase assay was
resolved by SDS-PAGE and immunoblotted. The signal intensity quantified from the
immunoblot was then used to normalize ceramide synthase activity in each sample. (lower)
Ceramide synthase was immunopurified from 3xHA-Lac1::HIS3 3xFLAG-Lag1::LEU2
(yAM168) 3xHA-Lac1(S23A S24A)::HIS3 or 3xFLAG-Lag1(S23A S24A)::LEU2 (Lac1SSAA
Lag1SSAA) (yAM184) cells as above except cultures were treated with 1.0 uM myriocin or
methanol vehicle prior to harvesting. Values represent the mean of three independent
experiments (each performed in triplicate) and error bars represent SEM. Statistical significance
of values (Student's t-test): *, p = <0.05, **, p = <0.009; and, ***, p <0.0009.
62
63
Fig. 4.7: Failure of Ypk1 to upregulate ceramide synthase causes LCBP accumulation that
triggers autophagy. (A) LAC1 LAG1 (yAM205 - A) or Lac1SSAA Lag1SSAA (yAM207 - B) were
transformed with PYPK1-Ypk1D242 (shown as Ypk1*) (pFR273) or pRS316 (E.V.). Transformants
were grown strains were grown to exponential phase in synthetic complete media and then
diluted to OD600=0.1 and grown in microtiter plates (lower) or on agar plates (upper). For liquid
cultures, each was grown in at least quadruplicate and the error bars indicate the SEM of
replicates at each time point. (B) cells from (A) were grown to mid-exponential phase in
selective synthetic complete media and then harvested. Sphingolipids were extracted and
analyzed as described in chapter 2. Values represent the mean of three independent experiments
(each performed in triplicate) and error bars represent SEM. (C) LAC1 LAG1 lcb4Δ (yAM237) or
Lac1SSAA Lag1SSAA lcb4Δ (yAM238 – A) were transformed with PYPK1-Ypk1D242 (shown as
Ypk1*) (pFR273) and growth experiments performed as in (A). (D) Liquid growth assays were
performed as in (A) for LAC1 LAG1 (yAM205 - A), Lac1SSAA Lag1SSAA (yAM207 - B), Lac1SSEE
Lag1SSEE (yAM210), LAC1 LAG1 lcb3Δ (yGT12), Lac1SSAA Lag1SSAA lcb3Δ (yGT13) and
Lac1SSEE Lag1SSEE lcb3Δ (yGT14) strains. (E) LAC1 LAG1 (yAM205 - A) or Lac1SSAA Lag1SSAA
(yAM207 - B) or LAC1 LAG1 lcb3Δ (yGT12) strains were transformed with PYPK1-Ypk1D242
(shown as Ypk1*) (pFR273) or pRS316 (E.V.) and additionally PTPI1-GFP-Atg8. Growing
cultures treated with vehicle or 2 μg/mL rapamycin for 2h and then harvested and whole extracts
prepared. Extracts were resolved by SDS-PAGE and blotted with anti-GFP to detect GFP-Atg8
and free GFP arising from GFP-Atg8 autophagic processing and anti-Pgk1 antibody. The blot is
representative of triplicate samples and the quantitation of the ratio of free GFP/Pgk1 from these
replicates is shown below the blot. (F) LAC1 LAG1 atg1Δ (yAM239 - A) or Lac1SSAA Lag1SSAA
atg1Δ (yAM240 – A) sensitivity to PYPK1-Ypk1D242 was measured as in (A). (G) Cells from (F)
were grown to mid-exponential phase in selective synthetic complete media and then harvested.
Sphingolipids were extracted and analyzed as described in (B).
64
65
Fig. 4.8: TORC2-Ypk1 signaling globally activates sphingolipid synthesis, selectively
directs flux toward ceramide metabolites, and prevents LCBP cross-talk to the autophagy
pathway. (A) Diagram of yeast de novo sphingolipid biosynthesis shown is derived from
(Dickson, 2008). Enzymes are in ovals. Skeletal structures of metabolites are shown. Increased
color intensity indicates level of metabolite increase in response TORC2-Ypk1 activation.
TORC2-Ypk1 signaling globally activates de novo sphingolipid biosynthesis via derepression of
the SPOTS complex (Berchtold et al., 2012; Roelants et al., 2011; Sun et al., 2012), potentially
increasing levels of all metabolites. However, Ypk1 also upregulates ceramide synthesis via
phosphorylation of Lac1 and Lag1, thus primarily directing this increased flux towards
ceramides and away from LCBs and LCBPs. (B) In the absence of Ypk1 mediated ceramide
biosynthesis regulation, increased sphingolipid flux raises LCB and LCBP levels. This slows cell
growth by activating autophagy. Thus, TORC2-Ypk1 signaling not only activates sphingolipid
biosynthesis in response to stress, but also insulates this flux towards ceramides to prevent
metabolite mediated crosstalk to the autophagy machinery.
66
67
Fig. 4.9: Loss of Ypk activity or putative Ypk phosphorylation of Atg21 does not appear to
prevent CVT or amino acid starvation induced autophagy. (A) Wild-type (yAM234 - B),
atg21Δ (yAM235-A), or ypk1-as ypk2Δ (yAM236-B) strains expressing GFP-Atg8 from a
constitutive promoter (TPI1) integrated at the HIS3 locus were grown with the indicated
concentrations of 3-MB-PP1 into mid-exponential phase in rich (YPD) medium. Based on
doubling time of ypk1-as ypk2Δ (yAM135–A) cells in YPD, 0.5 μM 3-MB-PP1 leads to ~30%
reduction in Ypk1 activity, 1.0 μM 3-MB-PP1 leads to ~50% reduction in Ypk1 activity, and 2.0
μM 3-MB-PP1 leads to ~90% reduction in Ypk1 activity (data not shown). Cells were harvested,
and then whole extracts were prepared and blotted as in Fig. 4.7E. (B) atg21Δ (yAM221-A) cells
were co-transformed with a centromeric PTPI1-GFP-Atg8 plasmid (pAX239) and empty vector
(pRS316), Atg21-3xFLAG (pAX241), Atg21 (S237A)-3xFLAG (pAX247) or Atg21(S237E)3xFLAG (pAX246) plasmids. Cells were grown in selective SCD medium into mid-exponential
or stationary phase. Cells were harvested and blotted for GFP-Atg8 processing as in (A).
Additionally, blotting was performed for Ape1. (C) atg21Δ (yAM221-A) cells were cotransformed with a centromeric PTPI1-GFP-Atg8 plasmids (pAX250) and empty vector (pRS316),
Atg21-3xFLAG (pAX241), Atg21 (S237A)-3xFLAG (pAX247) or Atg21(S237E)-3xFLAG
(pAX246) plasmids. Cells were grown to mid-exponential phase in selective SCD medium. Cells
were then washed and put into SCD medium without leucine (yAM221–A is a leucine
auxotroph) to trigger amino acid starvation. Cells were harvested and blotted for GFP-Atg8
processing as in (A).
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Chapter 5: TORC2-Ypk1 signaling is a MAPK-independent mechanism for
promoting glycerol accumulation and survival in response to hyperosmotic
stress
Introduction
Cells routinely encounter many diverse environmental and developmental stresses that physically
challenge the plasma membrane. For example, mechanical stretching of skin keratinocytes or
environmental osmotic shocks in microbes generate forces that threaten membrane integrity and
ultimately, if unresolved, cell viability. Understanding how cells cope with commonly occurring
membrane stresses is essential for understanding how membrane homeostasis is maintained in
changing environments.
How S. cerevisiae cells respond to hyperosmotic shock has served as a model system for
understanding how the cell copes with reduced turgor pressure and relaxed membrane tension. In
yeast, two pathways have been identified, to date, that are triggered when cells are subjected to
hypertonic stress. In response to such a challenge, plasma membrane-embedded osmosensors
(Sln1 and Sho1) associated with heavily O-glycosylated cell wall-embedded mucins (Msb2 and
Hkr1) (Cullen, 2007; Tanaka et al., 2014; Tatebayashi et al., 2015) initiate pathways that activate
a MAPK family member, Hog1 (Brewster, 1993; Brewster & Gustin, 2014). The sequence and
functional ortholog of Hog1 in mammals is p38/SAPK/MAPK14 (Han et al., 1994; Cuenda &
Rousseau, 2007; Cuadrado & Nebreda, 2010). Activated Hog1 drives a transcriptional and posttranscriptional program that increases glycerol production and retention (Hohmann, 2009; Saito
and Posas, 2012; Hohmann, 2015). Intercellular glycerol acts as a counter-balancing osmolyte
critical for optimal survival when cells are placed in hyperosmotic medium. In addition,
hyperosmotic shock increases cytosolic Ca2+ concentration (Denis and Cyert, 2002) to a level
sufficient to activate phosphoprotein phosphatase 2B/calcineurin (CN), which then specifically
associates with and dephosphorylates the PtdIns4,5P2 5-phosphatase Inp53/Sjl3 to stimulate
retrieval of excess plasma membrane via clathrin-mediated endocytosis (Guiney et al., 2015).
Conversely, when yeast cells are placed in hypotonic conditions, different mechanosensors
(Wsc1, Wsc2, Wsc3, Mid2 and Mtl1) that span the plasma membrane and the cell wall (Kock et
al. 2015) activate GEFs for the small GTPase Rho1, which in its GTP-bound state stimulate a
protein kinase, Pkc1, that initiates a signaling cascade that activates another MAPK family
member, Mpk1/Slt2 (de Nobel et al., 2000; Levin, 2011). Activated Slt2/Mpk1 drives the
expression of factors that strengthen the cell wall layers (Orlean, 2012; Free, 2013) to combat the
increased turgor pressure and, hence, is a response referred to as the cell wall integrity pathway
(Levin, 2011; Engelberg et al., 2014). The sequence and functional ortholog of yeast Pkc1 in
mammals is the RHO- (and RAC-) activated PKC-related enzyme PRK2/PKN2 (Mukai, 2003;
Thumkeo et al., 2013) and the sequence and functional ortholog of yeast Slt2/Mpk1 in mammals
is ERK5 (Piper et al., 2006; Truman et al., 2006). Collectively, therefore, these studies of the
response to stress in yeast have helped to illuminate how mammalian cells respond to osmolarity
changes and/or the degree of membrane stretch.
Despite this progress in understanding the cellular response to extracellular osmolarity and
membrane stress, regulation of even well studied cellular survival mechanisms still remains
incomplete. For example, an important part of the yeast response to hyperosmolarity is
69
prevention of glycerol efflux from cells that is mediated by the aquaglyceroporin Fps1 (Luyten et
al., 1995; Tamás et al., 1999; Ahmadpour, et al., 2014). Although Hog1 can negatively regulate
Fps1 by displacing the Fps1-activating protein Rgc2 in response to hyperosmotic shock (Lee et
al., 2013), Fps1 still closes in response to hyperosmotic shock in the absence of Hog1 (Tamás et
al., 1999; Babazadeh et al., 2014). Thus, there must be at least one additional osmosensing
mechanism that exists that is able to transduce the hyperosmolarity signal to Fps1 and prevent
efflux of glycerol and, thus, promote adaptation to hypertonic stress.
Recently, we and others have found that the TORC2 complex in S. cerevisiae, which was known
to respond to membrane stresses such as depletion of sphingolipids and heat shock (Roelants et
al., 2011; Berchtold et al., 2012; Sun et al., 2012) also responds to changes in osmotic conditions
and/or membrane stretch. As judged by phosphorylation of the TORC2 effector Ypk1, TORC2
activity apparently increases under hypo-osmotic stress (or by mechanical stretch) and the
increase in TORC2-Ypk1 signaling elevates the level of membrane sphingolipids to manage this
stress (Berchtold et al., 2012). Conversely, as judged by the same criterion, TORC2 activity
seems to decrease dramatically during hyperosmotic shock and the loss of TORC2-Ypk1
signaling helps attenuate the hyperosmotic shock, at least in part, through alleviating Ypk1dependent inhibition of glycerol-3-phosphate dehydrogenase Gpd1, leading to the production of
more glycerol-3-P, which, after dephosphorylation, increases intracellular glycerol (Lee et al.,
2012b). As with the other osmosensing machinery in yeast, mammalian TORC2 activity is also
responsive to changes in osmotic conditions (Sengupta et al., 2013) and membrane stretch
(Kippenberger et al., 2005). Thus, TORC2 is a conserved, membrane stress-responsive signaling
module whose output modulates both membrane composition and intracellular osmolyte level.
To understand how TORC2-Ypk1 signaling contributes to cellular osmotic and membrane stress
response, I recently performed a genome-wide screen to identify novel Ypk1 substrates. In this
way, I demonstrated that activated Ypk1 stimulates the ceramide synthase complex to to promote
the formation of sphingolipids (Muir et al., 2014). In this screen, I also identified two enzymes
involved in glycerol metabolism. The aquaglyceroporin Fps1 and Gpt2/Gat1, a glycerol-3phosphate/dihydroxyacetone phosphate acyltransferase that consumes glycerol-3-phosphate to
generate phosphatidic acid, a precursor to phospholipids or triacylglycerol (Zheng and Zou,
2001; Bratschi et al., 2009; Henry et al., 2012). Additionally, the screen uncovered a
transcription factor Smp1, that has been previously implicated in the response to hyperosmotic
shock (de Nadal et al., 2003) and also in Ypk1 physiology (Roelants et al., 2002).
In this chapter, I document that, although Gpt2 does not appear to be under TORC2-Ypk1
regulation, Fps1 and Smp1 are bona fide substrates of Ypk1 in vivo and their Ypk1-dependent
phosphorylation is physiologically important. I found that, under normal growth conditions,
Ypk1 phosphorylates Fps1 and that this phosphorylation is essential for Fps1 channel activity.
Thus, in the absence of stress, TORC2-Ypk1 signaling licenses glycerol efflux. However, upon
hyperosmotic shock, TORC2 activity and, in turn, Ypk1 activity are rapidly lost, resulting in
closure of the Fps1 channel. Therefore, the reduction in TORC2-Ypk1 signaling that occurs
under hyperosmotic conditions allows cells to cope with this stress by promoting accumulation
of intracellular glycerol not only by increasing glycerol production (Lee et al., 2012b), but also
by ensuring glycerol retention. Lastly, I show that abrogation of TORC2-Ypk1 phosphorylation
of Fps1 occurs independently of the MAPK Hog1 and other components of known hyperosmotic
70
sensing pathways. Thus, my work has uncovered a previously uncharacterized mechanism that
contributes to cellular adaptation to high osmolarity by driving a coordinated program of
glycerol production and retention.
Results
Smp1 is phosphorylated by Ypk kinases and this modification influences Smp1 function
and localization
To examine the role of TORC2-Ypk1 signaling in glycerol metabolism, I examined, first,
phosphorylation of a candidate Ypk1 substrate that emerged from my screen (Muir et al. 2014)
that is connected to utilization of glycerol-3-P, namely the glycerol-3-P/dihydroxyacetone-P sn-1
O-acyltransferase Gpt2/Gat1. There is one perfect match to the Ypk1 consensus phosphoacceptor
sequence (646RSRSSSI652) in this 743-residue enzyme. Based on past experience, we have found
that at such sites containing multiple Ser residues more than just the canonical one (in red) can
be phosphorylated in a Ypk1-dependent manner (Roelants et al., 2011). For that reason, I
prepared a Gpt2(S649A S650A S651A) mutant as well as a phosphomimetic Gpt2(S649E S650E
S651E) mutant. One or more of these three Ser residues is phosphorylated in vivo because,
compared to wild-type Gpt2, the Gpt23A mutant exhibited a distinctly faster mobility upon SDSPAGE, a hallmark of decreased phosphorylation (Fig. 5.1A). However, this phosphorylation did
not appear to be dependent on Ypk1 activity because there was no change in Gpt2 mobility when
a ypk1-as ypk2Δ strain was treated with inhibitor 3-MB-PP1 (Fig. 5.1A). Other AGC kinase
family members with prominent roles in metabolic regulation have, like Ypk1, basophilic
consensus phosphoacceptor motifs, including yeast AMPK (Snf1) and yeast PKA (Tpk1, Tpk2
and Tpk3). However, like loss of Ypk1 function, absence of these kinases did not prevent Gpt2
phosphorylation (Fig. 5.1A). Furthermore, based on a phenotypic criterion, it appears that
phosphorylation at the predicted Ypk1 consensus site(s) has no physiological role. In the absence
of Gpt2, the growth of yeast cells is inhibited by high concentrations of the fatty acid oleate,
whereas wild-type cells grow well under this condition (Zaremberg & McMaster, 2002).
Moreover, in the absence of both Gpt2 and its paralog Sct1/Gat2, cells are inviable (Zaremberg
& McMaster, 2002). I found that gpt2∆ sct1∆ cells expressing either the Gpt23A or Gpt23E alleles
grew just as well as cells expressing wild-type Gpt2 (Fig. 5.1B). Thus, in the absence of any
evidence for Ypk1-mediated phosphorylation of Gpt2 or any evidence for an effect of
phosphorylation at the putative Ypk1 site, I turned my attention to another candidate Ypk1
substrate that emerged from our screen (Muir et al., 2014), namely the MADS-box transcription
factor Smp1 (Shore & Sharrocks, 1995; Dodou & Treisman, 1997), which had been connected
previously to both Ypk1 function (Roelants et al, 2002) and in the response of yeast cells to
hyperosmotic stress (de Nadel et al., 2003).
The mammalian orthologs of Smp1 (and its paralog Rlm1) are the MEF2 family of transcription
factors (Potthoff & Olson, 2007). There are three predicted Ypk1 phosphorylation sites
(15RNRTVTF21, 102RRRRATV108 and 216RIRPDTH222) in the 452-residue Smp1 protein. To
enhance the ability to detect phosphorylation-dependent mobility shifts, I generated a C-terminal
truncation, Smp1(∆275-452), because the resulting Smp1(1-274) fragment contains all three
predicted Ypk1 sites, but lacks sequences that contain predicted Hog1 (MAPK) phosphorylation
sites (de Nadel et al., 2003). Using Smp1(1-274) produced a readily interpretable banding pattern
upon SDS-PAGE containing Phos-tag™, a separation method designed to resolve phospho71
isoforms (2009). I found that a slower mobility species of Smp1(1-274) was detectable in wildtype cells that became markedly more prominent when cells were treated with a stimulus
(addition of the antibiotic myriocin) known to lead to Ypk1 activation (Roelants et al. 2011)
(Fig. 5.2A). Moreover, this species was completely eliminated when a Smp1(1-273; T20A
T107A T221A) mutant was examined (Fig. 5.2A). Resolution of full-length Smp1 by Phos-tag
SDS-PAGE yielded a complicated banding pattern; yet, absence of the three predicted Ypk1 sites
caused a readily detectable decrease in the mobility of many of these species (Fig. 5.2B).
Collectively, these observations indicate that Smp1 is phosphorylated in cells in a Ypk1dependent manner and at its predicted Ypk1 sites. Furthermore, based on a phenotypic criterion,
it appears that phosphorylation at the predicted Ypk1 consensus site(s) has an important
physiological role. Unlike wild-type cells, cells lacking Smp1 are resistant to high concentrations
of ferric (Fe3+) ions (Lee et al., 2012a). I found that both an Smp1(T20A T107A T221A) allele
and a Smp1(T20E T107E T221E) allele were just as resistant to the toxic effects of Fe3+ as a
complete null mutant (smp1∆) (Fig. 5.2C), even though wild-type Smp1 and the two mutants
were expressed at comparable levels (Fig. 5.2D). Thus, authentic phosphorylation at its Ypk1
sites is necessary for Smp1 function (and, in the case of this protein, Glu residues at the same
positions cannot replace the presence of actual phosphate groups). Ypk1 activity also appears to
affect the subcellular localization of Smp1. Using an Smp1-GFP fusion, which has been shown
previously to be functional (de Nadal et al., 2003), I found that Smp1 localizes to the cytoplasm
during exponential growth, but translocates to the nucleus during stationary phase, in agreement
with what others have observed before (de Nadal et al., 2003). However, in cells expressing the
ypk1-as allele, when 10 μM 3-MB-PP1 was added to completely inhibit Ypk1 activity (while
leaving Ypk2 activity, and thus cell viability unaffected), I observed that Smp1 was present in
the nucleus even during exponential growth (Fig. 5.2E). Thus, one role for Ypk1-mediated
phosphorylation of Smp1 is to prevent its nuclear import. Lastly, Smp1 mediates part of the
transcriptional response to hyperosmotic shock (de Nadal et al., 2011), including contributing to
full transcriptional upregulation of the STL1 gene (de Nadal et al., 2003), which encodes a
glycerol/H+ symporter responsible for glycerol import (Duskova et al., 2015). Using a
STL1promoter-GFP gene fusion as a transcriptional reporter, I measured STL1promoter-driven
expression in smp1Δ, ypk1-as or double mutant cells both basally and after exposure to
hyperosmotic shock (0.4 M NaCl). I found that cells lacking Smp1 had only a modest diminution
in its salt stress-induced expression, similar to what has been described previously (de Nadal et
al., 2003) (Fig. 5.2F). Strikingly, however, in cells in which Ypk1-as had been inhibited with 3MB-PP1, salt stress induction of STL1 expression was almost completely prevented. The
requirement for Ypk1 activity for salt stress-evoked STL1 expression was observed even in cells
lacking Smp1. Hence, Ypk1 function clearly modulates Smp1 localization and function, but also
influences gene expression independently of its modification of Smp1. It would be worthwhile to
construct and examine additional reporter plasmids containing other predicted Smp1-responsive
and hyperosmotic shock-regulated promoters to try to characterize how Ypk1 modulates
transcription. Whether TORC2-Ypk1 signaling has a role in regulating other types of membrane
stress-induced transcription is also of significant interest.
Fps1 is phosphorylated by Ypk kinases and this modification is blocked when cells are
subjected to hyperosmotic stress
A third candidate Ypk1 substrate that emerged from my screen (Muir et al. 2014) that is directly
72
connected to control of intracellular glycerol concentration is the aquaglyceroporin Fps1, which
regulates glycerol efflux (Karlgren et al., 2004; Lee et al., 2013; Duskova et al. 2015). I observed
before that GAL promoter-driven overexpression of Fps1 in ypk1-as ypk2Δ cells was growth
inhibitory, whereas Fps1 overexpression had no effect on wild-type (YPK1+ YPK2+) cells (Muir
et al. 2014). Moreover, in a global screen for interactors with all yeast protein kinases and
phosphatases, it has been reported that the paralog of Ypk1, Ypk2, is able to physically associate
with Fps1 (Breitkreutz et al., 2010). For these reasons, I initially entertained the possibility that
the reason for the observed toxicity of Fps1 overexpression in cells with only limiting Ypk1
activity might be that the elevated level of Fps1 sequestered the available pool of Ypk1 away
from the membrane microdomains that harbor the essential Ypk1 activators Pkh1 (and Pkh2) and
TORC2, resulting in a lack of activation of Ypk1, a kinase that is essential for cell viability
(Chen et al. 1993; Roelants et al., 2002). Therefore, I examined Ypk1 phosphorylation at T504
(the T-loop/activation loop residue that is the target of Pkh1 and Pkh2) and T662 (a residue in
the so-called hydrophobic motif that is a target of TORC2) in strains lacking or overexpressing
Fps1. I found that the level of Fps1 expression had no significant effect on Ypk1 phosphorylation
at either of these sites (Fig. 5.3). Therefore, it did not appear that Fps1 was identified in our
screen because it interferes with Ypk1 activation. Hence, it seemed more likely that Fps1 was
recovered in our screen because it is, in fact, a bona fide Ypk1 substrate. Despite the technical
difficulties in examining an integral membrane protein, I confirmed this conclusion, as follows.
In Fps1, there are four, predicted, Ypk1 consensus phosphorylation sites (142RPRGQTT148,
176RRRSRSR182 , 180RSRATSN186 and 565RARRTSD148) (Fig. 5.4A). A C-terminal fragment of
Fps1 (and its proteolysis products) containing the single Ypk1 site Ser570 was phosphorylated in
a Ypk1-specific manner in vitro and mutation of Ser570 to Ala demonstrated that the observed
phosphorylation was specific to the predicted Ypk site (Fig. 5.4B). Using Phos-tag SDS-PAGE
analysis, I confirmed that Ser570 is phosphorylated in vivo because a corresponding S570A
mutation caused a dramatic shift to a faster mobility species (Fig. 5.4C, rightmost lanes). An Nterminal fragment of Fps1 containing the three other predicted Ypk1 sites was not detectably
phosphorylated by Ypk1 in vitro (Fig. 5.4B); however, when expressed in vivo, and based on the
effect on mobility in Phos-tag SDS-PAGE of mutating Ser181 and Ser185 to Ala, these two sites
are clearly phosphorylated in the cell (Fig. 5.4C). Likewise, the mobility of Fps1 underwent the
same dramatic shift to faster mobility in ypk1-as ypk2Δ cells treated with 3-MB-PP1, but not in
wild-type cells (Fig. 5.4D). Moreover, the mobility of the resulting species was not different
from that of an Fps1(S181A S185A S570A) mutant, Fps13A hereafter, and inhibition of Ypk1
activity did not further increase its mobility. One likely possibility to explain the difference
between our in vitro results (performed with N- and C-terminal fragments of Fps1) and our in
vivo results (examining intact Fps1) is that Ypk1 docks at a binding site in the C-terminal domain
in order to phosphorylate sites in the N-terminal domain. Furthermore, as expected for a Ypk1dependent modification, phosphorylation of these same sites in Fps1 in vivo was also TORC2dependent, as judged by the fact that inhibition of TORC2 activity with a pharmacological agent
(NVP-BEZ235) (Maira et al., 2008; Kliegman et al., 2013) also drastically reduced Fps1
phosphorylation (Fig. 5.4E). I conclude that Fps1 is an authentic in vivo substrate of Ypk1.
Recently, we documented that TORC2-mediated phosphorylation of Ypk1 is greatly decreased
when cells are subjected to hyperosmotic shock (Lee et al., 2012b). Using a Ypk17A mutant,
which allows for facile detection of mobility shifts arising from its TORC2-specific
73
phosphorylation, I found that loss of TORC2-mediated phosphorylation of Ypk1 after
hyperosmotic shock occurs very rapidly (within 1 min) (Fig. 5.5A). This loss of TORC2mediated phosphorylation of Ypk1 is transient, however. By 20 min after hyperosmotic shock,
TORC2-mediated phosphorylation of Ypk1 is again readily detectable and is back to its prestress level by 75 min after application of the stress (Fig. 5.5A). The reduction in TORC2mediated phosphorylation of Ypk1 in response to hypertonic stress was still observed in mutants
that abrograte signal propagation in either the Sho1- or Sln1-dependent branches of the HOG
pathway, or in the absence of their common terminal effector, Hog1 MAPK (Fig. 5.5B). Given
that TORC2-Ypk1 action results in direct Fps1 phosphorylation and given that TORC2-mediated
phosphorylation of Ypk1 is greatly decreased when cells are subjected to hyperosmotic shock, I
examined whether phosphorylation of Fps1 was affected when cells were subjected to
hyperosmotic shock. Indeed, as expected, hyperosmotic shock led to loss of phosphorylation of
Fps1 that mirrored the kinetics of the loss of TORC2-mediated phosphorylation of Ypk1 in
response to hypertonic stress and yielded a species with a mobility indistinguishable from that of
the Fps13A mutant lacking Ypk1 phosphorylation sites (Fig. 5.5C). Taken together, these results
demonstrate that loss of TORC2-mediated phosphorylation of Ypk1 in response to hyperosmotic
stress dramatically down-modulates Fps1 phosphorylation in a HOG pathway-independent
manner.
Ypk-mediated phosphorylation of Fps1 promotes channel opening and glycerol efflux
To determine whether this newly revealed mode of Fps1 regulation serves any function in
promoting cell survival in the face of hypertonic stress, I next investigated how Ypk1-mediated
phosphorylation may affect Fps1 function. First, I repeated the SDL assay (Muir et al., 2014) to
assess the phenotype caused by overexpression of wild-type Fps1, Fps13A, or previously
characterized mutants of Fps1 that are known to cause the channel to be constitutively open
(Tamás et al., 2003; Hedfalk et al., 2004) or constitutively closed (Lee et al., 2013). I found that
overexpression of wild-type, Fps13A, and a known closed channel mutant gave an SDL
phenotype, whereas known open channel mutants did not (Table 5.1). Based on this criterion, the
behavior of Fps13A resembles that of a closed channel, suggesting that Ypk1-mediated
phosphorylation promotes channel opening.
Fps1 can mediate the influx of xylitol (not a physiological carbon source for Saccharomyces
cerevisiae; Fraenkel, 2011) and expression of constitutively open channel mutants allows cells
defective in glycerol production (gpd1Δ gpd2Δ mutants) to survive when challenged with a high
concentration of exogenous xylitol by permitting the xylitol to equilibrate across the membrane,
thereby restoring osmotic balance (Hedfalk et al., 2004). Expression of an N-terminal truncation
mutant, Fps1(∆12-231), that is constitutively open allowed cells to survive on 1M xylitol,
whereas Fps13A did not (Fig. 5.6A). Thus, by this second criterion, the behavior of Fps13A
resembles that of a closed channel.
Fps1 can also mediate the entry of a toxic metalloid, arsenite, and thus arsenite sensitivity is a
read-out of Fps1 function (Thorsen et al., 2006). Unlike wild-type Fps1, all Fps1 mutants
predicted to lack transport activity, such as a null mutant (fps1∆), a cell lacking known activators
that promote Fps1 channel opening (rgc1Δ rgc2Δ double mutant) (Beese et al., 2009), and a
mutant (fps1ΔPHD) unable to bind Rgc1 or Rgc2 (Lee et al., 2013), exhibited resistant to arsenite
74
toxicity (Fig. 5.6B). Strikingly, cells expressing Fps13A as the sole source of this channel were
equally if not more arsenite resistant (Fig. 5.6B). Hence, by this third criterion, the behavior of
Fps13A resembles that of a closed channel.
If Ypk1-mediated phosphorylation at multiple sites promotes Fps1 channel opening in a graded
manner, then one might expect that phosphorylation at individual Ypk1 sites would contribute
additively to channel opening and, conversely, that absence of phosphorylation at individual
Ypk1 sites would contribute additively to arsenite resistance, which is exactly what I observed
(Fig. 5.6C).
Collectively, these results indicate that TORC2-stimulated Ypk1-mediated phosphorylation of
Fps1 promotes opening of this channel and its transport activity. It has been shown that loss of
Fps1 function leads to retention of intracellular glycerol (Tamás et al., 1999). Consistent with
Ypk1-dependent phosphorylation favoring the open state of Fps1, I found that cells expressing
Fps13A accumulated ~2 fold as much glycerol as wild-type cells (Fig. 5.6D). In agreement with
this finding, we previously documented that cells deficient in Ypk activity have a higher
intracellular glycerol level (Lee et al., 2012b). For this reason, I also examined the arsenite
resistance of cells in which I attempted to reduce Ypk activity because loss of the kinase should
mimic Fps13A and, perhaps, allow enhanced resistant to the toxic effect of arsenite. However, it
is not possible to completely inhibit Ypk function and maintain viability. However, even at a
sub-lethal dose of 3-MB-PP1, I did not detect any arsenite resistance in either ypk1-as or ypk1-as
ypk2Δ cells (Fig. 5.6E). Presumably, as with maintenance of viability, the residual Ypk activity
present is still sufficient to phosphorylate Fps1 and keep the channel open. I also cannot rule out
the possibility that reduction of Ypk activity may have additional consequences that might
increase the arsenite sensitivity of cells, even if there is some down-modulation of Fps1 function.
Nevertheless, taken together, all of our other results show conclusively that Ypk1 phosphorylates
and thereby promotes opening of Fps1.
I considered that TORC2-dependent Ypk1-mediated phosphorylation of Fps1 might also
promote its channel function in other ways, such as stabilizing the protein or increasing its dwell
time and, hence, its concentration, in the plasma membrane. However, immunoblotting (Fig.
5.6F) and fluorescence microscopy (Fig. 5.6G) indicate that steady-state level and localization of
wild-type Fps1 and Fps13A are indistinguishable. Therefore, the primary effect of Ypk1dependent phosphorylation of Fps1 is to favor the open state of this channel.
TORC2-Ypk regulation of Fps1 is independent of the known Fps1 regulators Hog1 and
Rgc1/2 and allows cells to survive hyperosmotic stress in the absence of Hog1
Fps1 is negatively regulated by Hog1 MAPK in two ways: (i) in response to challenge with
acetic acid or arsenite (but, curiously, not hyperosmotic stress), direct phosphorylation of Fps1
by Hog1 stimulates channel internalization and degradation (Thorsen et al., 2006; Mollapour and
Piper, 2007; and (ii) in response to hyperosmotic stress, direct phosphorylation of Fps1 by Hog1
causes dissociation of its positive activators Rgc1 and Rgc2 (Beese et al., 2009; Lee et al., 2013).
I found that Fps13A was still arsenite resistant even in a hog1Δ mutant (Fig. 5.7A), quite similar
to the behavior of a strain that expresses an Fps1 mutant unable to bind Hog1 (Lee et al., 2013)
75
(Fig. 5.7B). Thus, mechanistically, TORC2-Ypk1 regulation of Fps1 occurs in a manner
independent of any role for Hog1. In addition, as judged by their co-immunoprecipitation, I
found that the lack of Ypk1-mediated phosphorylation does not cause dissociation of the positive
Fps1 activator Rgc2 because as much or more Rgc2 is associated with Fps13A as with wild-type
Fps1 (Fig. 5.7C). Thus, although the exact mechanism by which TORC2-Ypk1 phosphorylation
favors Fps1 channel opening is not yet known, it is clear that it is not achieved through
modulation of the Hog1-Rgc1/2 system.
Our results suggest that loss of TORC2-dependent phosphorylation of Ypk1 that occurs when
cells are subjected to hypertonic conditions and the ensuing reduction of Ypk1-mediated
phosphorylation of Fps1 provides a novel means, independent from the HOG pathway, for both
sensing high osmolarity and responding to it because decreased Fps1 channel function should
permit intracellular glycerol accumulation. Indeed, tellingly, I found that a Fps13A hog1Δ double
mutant cells is more resistant to hyperosmotic stress than hog1Δ cells, exhibiting growth at a
near wild-type level at 0.5 and 0.75 M sorbitol concentrations (Fig. 5.7D). This striking rescue
confirms that loss of Ypk1-mediated Fps1 channel opening is sufficient for cells to accumulate
an adequate amount of glycerol to physiologically cope with hyperosmotic stress, even in the
absence of Hog1.
Discussion
Our screen for new substrates (Muir et al., 2014) reinforces the conclusion that glycerol
metabolism as an important process regulated by the TORC2-Ypk1 axis. Here I have
documented that the loss of TORC2 stimulation of Ypk1 that occurs in response to hyperosmotic
shock reduces Ypk1-mediated phosphorylation of the glycerol efflux channel Fps1. All evidence
supports the conclusion that Ypk1 phosphorylation of Fps1 favors its open state and, thus,
absence of Ypk1 phosphorylation closes the channel. By this means, in response to hyperosmotic
shock, Fps1-mediated glycerol efflux is prevented allowing intracellular glycerol to accumulate.
We demonstrated previously that Gpd1, the glycerol-3-P dehydrogenase that is the rate-limiting
enzyme in glycerol biosynthesis (Remize et al., 2001), is as bona fide in vivo substrate for Ypk1
and that Ypk1-mediated phosphorylation inhibits this enzyme (Lee et al., 2012b). This step is
rate-limiting for glycerol production because glycerol is formed directly by dephosphorylation of
the glycerol-3-P generated from DHAP by Gpd1 action (Fraenkel, 2011). Thus, the inactivation
of TORC2-Ypk1 signaling that occurs upon hyperosmotic shock has at least two coordinated
consequences that work synergistically to cause glycerol accumulation and promote cell survival
(Fig. 5.8). First, loss of TORC2-Ypk1 signaling alleviates inhibition of Gpd1, thereby increasing
glycerol production. Second, loss of TORC2-Ypk1 signaling results in closing of the Fps1
channel allowing for retention of the glycerol produced. In addition, my preliminary evidence
suggests that loss of TORC2-Ypk1 signaling will permit the nuclear entry of the Smp1
transcription factor whose function appears to contribute to survival under hypertonic conditions
(de Nadal et al., 2003). Thus, like the HOG response, the TORC2-Ypk1 signaling axis seems to
modulate both transcriptional and post-transcriptional processes that help the cell cope with
hyperosmotic stress. Future studies will be required to understand how Ypk1 (and Ypk2)
regulates Smp1 function and what roles Smp1 has in rewiring of cellular physiology in response
to hyperosmotic shock and other stresses that perturb plasma membrane structure and function.
76
Despite the similarities in stimuli sensed and pathway outputs, I have shown here that the
TORC2-Ypk1 response is independent of any connection to Hog1. Why S. cerevisiae has two
independent mechanisms for regulating glycerol levels in response to hyperosmotic stress is an
interesting question. The presence of two systems might allow cells to adjust their response
optimally to stresses that occur with different intensity, duration, or frequency. It has been shown
that Hog1 primarily responds to stresses occurring no more frequently than every 200 seconds
(Hersen et al., 2008; McClean et al., 2009). Perhaps TORC2-Ypk1 signaling allows response to
occur in environments that fluctuate even more rapidly. Another possibility is that TORC2-Ypk1
signaling drives adaptive glycerol accumulation in response to stimuli not sensed by the Sln1- or
Sho1-branches of the HOG pathway, or vice-versa.
How does TORC2 sense high osmolarity? A recent report provided some evidence that Sho1, an
integral plasma membrane protein that is thought to be a component of one of the osmosensors in
the Hog1 pathway, can interact directly with Tor2, the catalytic subunit of TORC2 (Lam et al.,
2015). This observation suggested the possibility that TORC2 activity could be subject to direct
negative regulation via the osmolarity sensors of the Hog1 pathway itself. However, as I have
shown here, in sho1Δ cells and mutants in other components of both the Sho1 and Sln1 branches
of the Hog1 pathway, TORC2 inactivation in response to hyperosmolarity is unaffected. My
results demonstrate that TORC2-Ypk1 is a novel osmosensing pathway independent of the
known MAPK response. Identification of mechanism by which TORC2-mediated
phosphorylation of Ypk1 is rapidly inactivated in response to high osmolarity is an important
goal for future work.
Osmosensing by TORC2 does not appear to be a phenomenon confined to S. cerevisiae.
mTORC2 activity has been found to be altered in response to mechanical stretch (Kippenberger
et al., 2005) and to osmotic shock (Sengupta et al., 2013) in human cell lines. Additionally, the
downstream mTORC2 effector kinase SGK1, which is the mammalian ortholog of Ypk1
(Casamayor et al., 1999), has been implicated in the response to increased salt (osmolyte) levels
in murine T cells (Wu et al., 2013). Furthermore, mTORC2 signaling is important for regulating
levels of ion transporters (Sengupta et al., 2013; Gleason et al., 2015). Collectively, these results
suggest that mTORC2 might respond to osmotic pressure by regulating ion transporters to
balance osmolarity, much like TORC2 in yeast responds to osmolarity and regulates Fps1 to
balance osmolarity. Thus, our screen has given us insight into a novel potentially conserved
signaling system by which eukaryotes respond to changes in osmotic pressure.
77
Figures
Fig. 5.1: Gpt2 is not phosphorylated by Ypk or other basophilic kinases. (A) Wild-type
(BY4741), ypk1-as ypk2Δ (yAM135 - A), snf1Δ (CGA65) or tpk1-as tpk2-as tpk3-as (JTY5173)
cells were transformed with Gpt2-3xFLAG (pAX238) or Gpt23A-3xFLAG (pAX244) plasmids.
Cells were grown to mid-exponential phase and then treated with 10 μM 3-MB-PP1 (ypk1-as
ypk2Δ), 5 μM 1NMPP1 (tpk1-as tpk2-as tpk3-as) (Zaman et al., 2009) or vehicle for 90 minutes.
Cells were harvested, extracts prepared, resolved by SDS-PAGE, and blotted as described in
chapter 2. (B) Wild-type (BY4741), gpt2Δ (yAM219 - A) or gpt2Δ sct1Δ strains covered with
Gpt2-3xFLAG (yAM231), Gpt23A-3xFLAG (yAM232) or Gpt23E-3xFLAG (yAM233) plasmids
were spotted and grown on YPD or YPD supplemented with 3.5 mM oleate for 3 days at 30C.
gpt2Δ sct1Δ mutants are not viable (Zaremberg and McMaster, 2002). Therefore, Gpt23A or
Gpt23E mutants do not completely abrogate Gpt2 activity as these plasmids would not be able to
sustain cell growth on YPD. Furthermore, these mutations do not appear to decrease the ability
of Gpt2 to allow cells to survive oleate stress. Cells with Gpt23A or Gpt23E as the sole source of
glycerol-3-phosphate/dihydroxyacetone phosphate acyltransferase do not exhibit sensitivity to
oleate as gpt2Δ cells do and are no more sensitive than wild-type cells.
78
79
Fig. 5.2: Smp1 is phosphorylated at predicted Ypk sites and phosphorylation of these
residues is important for Smp1 function. (A) Wild-type (BY4741) cells were transformed with
Smp1(1-274)-3xFLAG (pAX286) or Smp1(1-274)3A-3xFLAG (pAX287) plasmids. Cells were
grown to mid-exponential phase. Cells were harvested, extracts prepared, resolved by SDSPAGE, and blotted as described in chapter 2. (B) Wild-type (BY4741) cells were transformed
with Smp1-3xFLAG (pAX253) or Smp13A-3xFLAG (pAX260) plasmids. Cells were grown to
mid-exponential phase. Cells were harvested, extracts prepared, resolved by SDS-PAGE, and
blotted as described in chapter 2. (C) Upper panel Wild-type (BY4741), smp1Δ (yAM197),
Smp1-3xFLAG (yAM246 - A), Smp13A-3xFLAG (yAM247 - A) and Smp13E-3xFLAG
(yAM248 - A) cells were serially diluted, spotted and grown on SCD or SCD supplemented with
indicated ferric iron chloride concentrations [prepared as described in (Lee et al., 2012a)] for 3
days at 30C. Lower panel smp1Δ (yAM197) cells were transformed with Smp1-3xFLAG
(pAX253), Smp13A-3xFLAG (pAX260) and Smp13E-3xFLAG (pAX263) plasmids.
Transformants were serially diluted, spotted and grown on SCD medium lacking uracil (to select
for the plasmid) or SCD without uracil supplemented with indicated ferric iron chloride
concentrations for 3 days at 30C. (D) smp1Δ (yAM197), Smp1-3xFLAG (yAM246 - A),
Smp13A-3xFLAG (yAM247 - A) and Smp13E-3xFLAG (yAM248 - A) strains from (C Upper)
were grown to mid-exponential phase, harvested, extracts prepared, resolved by SDS-PAGE, and
blotted as described in chapter 2. (E) Wild-type (BY4741) or ypk1-as (yAM134 - A) cells were
transformed with a centromeric plasmids Smp1-GFP from its own promoter (pAX270). Cells
were then grown to mid-exponential phase or stationary phase as indicated and microscopy was
performed as described in chapter 2. ypk1-as cells were treated with 10 μM 3-MB-PP1 to inhibit
Ypk1 activity. Arrows indicate cell nuclei as determined by DAPI staining. (F) Wild-type
(yAM241 - A), smp1Δ (yAM243), ypk1-as (yAM242 - A) or smp1Δ ypk1-as (yAM244 - A) cells
with PSTL1-GFP integrated at the HIS3 locus were grown to mid-log phase in SCD without
tryptophan with 10 μM 3-MB-PP1 in ypk1-as strains to inhibit Ypk1 activity. Cells were then
transferred to SCD without tryptophan with or without 0.4 M NaCl to cause hyperosmotic shock.
Cells were then harvested, fixed and intracellular GFP was measured by FACS as described in
chapter 2.
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Fig. 5.3: Changes in Fps1 dosage does not affect either TORC2- or Pkh1-mediated
phosphorylation of Ypk1. Haploid wild-type (BY4742) and fps1Δ (yAM181 - A) cells or
diploid wild-type (BY4743) and fps1Δ/ fps1Δ (JTY6322) cells were co-transformed with
plasmids expressing Fps1 under the control of the GAL1/10 (pAX96) promoter or empty vector
control (YCpLG) and a plasmid expressing Ypk1-HA under the control of the MET25 promoter
(pLB215). Cells were grown to mid-exponential phase SC media without methionine to
stimulate Ypk1-HA expression with 2% raffinose before addition of 2% galactose to induce Fps1
expression for 3.5 hours. Cells were harvested, extracts prepared, resolved by SDS-PAGE, and
blotted as described in chapter 2. The upper hand corresponds to Ypk1-HA as judged by comigration between bands staining for HA and pT504. The faster migrating species seen in pT662
and pT504 blots result from these antibodies recognizing phosphorylation of endogenous Ypk1
and Ypk2.
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82
Fig. 5.4: Ypk in response to TORC2 signaling phosphorylates Fps1 at three sites in vivo.
(A) Diagram of Fps1 [adapted from (Lee et al., 2013)] with predicted Ypk sites indicated as red
circles. Primary sequence context of predicted sites are shown at left. (B) Upper GST-Fps1(1255) (pBT6), GST-Fps1(531-669)(pBT7) were purified from E. coli and Ypk1-as kinase assays
were performed as described in chapter 2. Lower GST-Fps1(531-669)(pBT7) and GSTFps1(531-669)(S570A) (pAX135) were purified from E. coli and Ypk1-as kinase assays were
performed as above. (C) Extracts from Fps1-3xFLAG (yGT21), Fps1(T147A)-3xFLAG
(yAM310 - A), Fps1(S181A S185A)-3xFLAG (yAM301 - A), Fps1(S570A)-3xFLAG (yGT24)
or Fps13A-3xFLAG (yGT22) expressing strains were prepared, resolved by Phos-tag SDS-PAGE
and immunoblotted as per chapter 2. Red asterisks indicate the appearance of higher mobility
non-phosphorylated Fps1 species. (D) Fps1-3xFLAG (yGT21), Fps13A-3xFLAG (yGT22), ypk1as ypk2Δ Fps1-3xFLAG (yAM281) and ypk1-as ypk2Δ Fps13A-3xFLAG (yAM284 - A) strains
were grown to mid-exponential phase and treated as indicated with 10 μM 3-MB-PP1 or vehicle
for 60 minutes. Fps1-3xFLAG was resolved by Phos-tag SDS-PAGE as in (C), and blotted as in
(C). Red asterisks indicate the appearance of higher mobility non-phosphorylated Fps1 species.
(E) tor2-as (yKL5) cells expressing Fps1-3xFLAG (pAX274) or Fps13A-3xFLAG (pAX275)
were grown to mid-exponential phase and then treated with 2μM BEZ-235 or vehicle for 30 min
Extracts were resolved and immunoblotted as in (C).
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Fig. 5.5: TORC2 phosphorylation of Ypk1 decreases rapidly in response to hyperosmotic
shock resulting in Fps1 dephosphorylation. (A) Wild-type (BY4741) or tor2-29 (JTY5468)
cells expressing Ypk17A-myc (pFR252) were grown at 26˚C to midexponential phase before
switching their media to include 1 M sorbitol for the indicated time periods. Extracts were
prepared, resolved and blotted as described in chapter 2. (B) As in (A) except extracts came from
hog1Δ (YJP544), sho1Δ (JTY5540), ssk1Δ (JTY5541), ssk22Δ (JTY5539), ssk2Δ (JTY5538) or
pbs2Δ (JTY5537) strains expressing Ypk17A-myc treated with 1 M sorbitol for 1 min (C) Fps13xFLAG (yGT21) or Fps13A-3xFLAG (yGT22) expressing cells were treated with 1 M sorbitol
for indicated time points and Fps1-3xFLAG was resolved and blotted as in (Fig. 5.4).
84
85
Fig. 5.6: Phosphorylation of Fps1 by TORC2-Ypk increases channel transport activity. (A)
gpd1Δ gpd2Δ cells (JT76328) that cannot balance osmolarity by glycerol production were
transformed with high copy number plasmids expressing Fps1-3xFLAG (pGT02), Fps1(Δ12231) -3xFLAG (pGT05), Fps1(Δ534-650)-3xFLAG (pGT06), Fps13A-3xFLAG (pGT17),
Fps13E-3xFLAG (pGT18) or empty vector (pRS426). Cells were then serially diluted and plated
on SCD lacking uracil to select for plasmid or SCD without uracil supplemented with 1 M
xylitol. Growth was allowed for 3 days at 30 ˚C prior to imaging. Constitutively open Fps1(Δ12231) allows cells to live. Interestingly, previous reports suggest that Fps1(Δ534-650) is
constitutively open as well (Hedfalk et al., 2004). However, this mutant does not allow cells to
survive this stress. (B) Cultures of Fps1-3xFLAG (yGT21), Fps13A-3xFLAG (yGT22),
Fps1ΔPHD-3xFLAG (yAM307 - A), rgc1Δ rgc2Δ (DL3188) and fps1Δ (yAM181 - A) were
adjusted to OD600=1.0 and serial dilutions were then spotted onto YPD plates containing the
indicated concentration of arsenite. Cells were allowed to grow for 4 days at 30˚C prior to
imaging. (C) As in (A) except Fps1-3xFLAG (yGT21), Fps1(T147A)-3xFLAG (yAM310 - A),
Fps1(S181A S185A)-3xFLAG (yAM301 - A), Fps1(S570A)-3xFLAG (yGT24) or Fps13A3xFLAG (yGT22) cultures were used and cells were grown for 2 days at 30˚C prior to imaging.
(D) Extracts were made from triplicate exponentially growing cultures of wild-type (BY4742),
fps1Δ (yAM181 - A), Fps1-3xFLAG (yGT21) and Fps13A-3xFLAG (yGT22) strains.
Concentrations of glycerol and protein in these extracts were then determined (see Materials and
Methods). These measurements were used to calculate the ratio of glycerol to protein mass in
each extract. (E) Wild-type (BY4741), fps1Δ (yAM181 - A), ypk1-as (yAM134 - A) and ypk1-as
ypk2Δ (yAM135 - A) cells were serially diluted and plated on YPD media containing 3-MB-PP1
to inhibit Ypk1as and/or arsenite as indicated. Growth was allowed for 3 days at 30 ˚C prior to
imaging. (F) Extracts were made from strains in (C) and resolved by standard SDS-PAGE using
8% acrylamide gels. (G) fps1Δ (yAM181 - A) cells expressing Fps1-GFP (pAX290),
Fps1(S181A S185A)-GFP, (pAX294), Fps1(S570A)-GFP (pAX293) or Fps13A-GFP (pAX295)
were imaged as described in chapter 2. Representative images are shown.
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87
Fig. 5.7: TORC2-Ypk regulates Fps1 independently of known regulators Hog1 and Rgc1/2.
(A) Cultures of Fps1-3xFLAG (yGT21), Fps1570A-3xFLAG (yGT24), Fps13A-3xFLAG (yGT22),
Fps1-3xFLAG hog1Δ (yAM275), Fps1570A-3xFLAG hog1Δ (yAM291 - A) and Fps13A-3xFLAG
hog1Δ (yAM278) strains were adjusted to OD600=1.0 and serial dilutions were then spotted onto
YPD plates containing the indicated concentration of arsenite. Cells were allowed to grow for 2
days at 30˚C prior to imaging. (B) As in (A) except Fps1IVAA-3xFLAG (yAM308 - A),
Fps1(3A)IVAA-3xFLAG (yAM309 - A), Rgc27A (yAM315) and Rgc27A Fps13A-3xFLAG
(yAM318) strains were tested. Fps1IVAA mutation prevents Hog1 binding to and regulation of
Fps1 while the Rgc27A cannot be phosphorylated and displaced from Fps1 by Hog1. These
mutations render the channel constitutively open and arsenite sensitive (Lee et al., 2013). (C)
Fps1-3xFLAG (yGT21) or Fps13A-3xFLAG (yGT22) strains were transformed with a PMET25Rgc2-HA plasmid (p3151). After Rgc2-HA expression, Fps1 was immunopurified (see chapter
2). Immunoprecipitates were resolved by SDS-PAGE and Fps1-Rgc2 interaction was determined
by blotting for Rgc2-HA in the precipitates. (D) Wild-type (BY4741), hog1Δ (YJP544) or
Fps13A-3xFLAG hog1Δ (yAM278) strains were plated onto synthetic complete medium lacking
tryptophan with 2% dextrose and the indicated concentration of sorbitol. Cells were grown for 3
days prior to imaging. Serial dilutions are shown from the same plate.
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Fig. 5.8: Saccharomyces cerevisiae utilizes two independent osmosensing systems to rapidly
increase intracellular glycerol upon hyperosmolarity. (A) Diagram of the Hog1 MAPK
mediated response to acute hyperosmotic stress [adapted from (Hohmann, 2015)]. Left shows the
unstressed condition while right denotes cells under hyperosmotic stress. Hog1 is activated
downstream of multiple osmosensing pathways. Hog1 increases glycolytic flux through
phosphorylation of Pkf26 allowing for more glycerol production and also prevents glycerol
efflux by phosphorylation and displacement of the Fps1 activators Rgc1/2 (Lee et al., 2013). On
longer time scales, Hog1 also translocates to the nucleus where it drives transcription of glycerol
production and uptake genes (de Nadal et al., 2011) (not depicted). (B) The TORC2-Ypk
response to hyperosmolarity similarly increases glycerol production and accumulation. Basal
TORC2-Ypk activity normally represses Gpd1 (Lee et al., 2012b) and is important for Fps1
channel activity. Hyperosmotic inactivation of TORC2 thus modulates the two rate limiting steps
of glycerol production (Remize et al., 2001) to promote glycerol accumulation. Diagram of
TORC2 adapted from (Liao and Chen, 2012; Wullschleger et al., 2005).
89
Tables
Table 5.1 Synthetic dosage lethality of various Fps1 mutants in ypk1-as ypk2Δ cellsa
Fps1 Mutant
Wild-type
Fps1(Δ12-231)
Fps1(Δ534-650)
Fps1(Δ544-581)
Fps13A
Fps13E
SDL Phenotype
+++++
+++++
+++++
+++++
Fps1 channel activity
Unknown
Constitutively open (Tamás et al., 2003)
Constitutively open (Hedfalk et al., 2004)
Constitutively closed (Lee et al., 2013)
Unknown (but phenotype indicates closed)
Unknown
a
SDL assays were performed as described in Chapter 2 and Table 3.1, except that the following
plasmids were used for overexpressing Fps1 (pAX96), Fps1(Δ12-231) (pGT11), Fps1(Δ534650) (pGT12), Fps1(Δ544-581) (pGT13), Fps13A (pGT14) and Fps13E (pGT15).
90
Chapter 6: Concluding Remarks
Conclusions
If a cell is to survive, it must adjust its plasma membrane lipid and protein composition properly
during growth and in response to environmental insults. Understanding the mechanisms that a
cell uses to maintain its plasma membrane homeostasis is a fundamental question in cell biology.
Many years of research using S. cerevisiae and its plasma membrane (van der Rest et al., 1995)
as a model has given tremendous insight into conserved molecular mechanisms and pathways
that a eukaryotic cell uses to cope with various membrane stresses (Ding et al., 2009; Shima &
Takagi, 2009; Bravim et al., 2010; Jendretzki et al., 2011; Levin, 2011; Saito and Posas, 2012;
Hohmann, 2015). Despite this progress, the pathways already described do not represent the
complete set of molecular mechanisms that have evolved to cope with such stresses. Recently,
our lab (Roelants et al., 2011) and others (Berchtold et al., 2012; Sun et al., 2012) demonstrated
that signaling from TORC2 and the downstream protein kinases that it stimulates, Ypk1 (and its
paralog Ypk2), is important for cell survival in response to stresses that perturb membrane lipid
composition and organization. Surprisingly, despite intense interest in TORC1, especially in
animal cells, because of its central role in growth control (Betz & Hall, 2013; Lemming &
Sabatini, 2013; Bar-Peled & Sabatini, 2014; Shimobayashi & Hall, 2014; Albert & Hall, 2015;
Efetan et al., 2015), relatively little was known about how TORC2 activity is regulated or how
its effectors influence cell physiology (Huang & Manning, 2009; Masui et el., 2014). Thus,
mechanistically how TORC2 action allows cells to cope with stress was unknown. The studies I
conducted for my doctoral research and that are presented in this dissertation describe my
discoveries about the mechanisms by which TORC2-Ypk1 signaling in yeast influences cellular
processes to allow cells to maintain plasma membrane homeostasis in the face of stresses that
otherwise would compromise cell viability.
In Chapter 3, I described a productive approach - combining bioinformatics, a novel genetic
screen that exploited synthetic dosage lethality, and in vitro biochemistry - to generate a high
quality list of previously uncharacterized Ypk1 substrates. The dozen substrates identified in this
manner include targets in processes in which TORC2 signaling was already implicated, such as
in control of sphingolipid metabolism, suggesting more complex and nuanced roles for TORC2
signaling in these processes. Other identified substrates indicate that TORC2 has roles in
modulating other areas of cell physiology in which it was not previously implicated. Thus, my
search for new Ypk1 substrates increased the breadth and depth of our understanding of the
regulation of cellular processes by TORC2. Of the five candidates that I explored in detail (Lac1,
Lag1, Gpt2, Smp1, Fps1), all except Gpt2 were demonstrably Ypk1 substrates in vivo and their
Ypk1-mediated phosphorylation had readily detectable physiological consequences. Thus, my
approach for identifying protein kinase substrates was very successful and might be very useful
in identifying substrates for protein kinases situated in other pathways.
Mass spectrometry-based methods for identifying protein kinase substrates have been very
successful (McLachlin & Chait, 2001; Graves & Haystead, 2003; Corthals et al., 2005; Cohen &
Knebel, 2006; Nita-Lazar et al., 2008; Macek et al., 2009; Yates et al., 2009; Dephoure et al.,
2013; Jünger & Aebersold, 2013) and have been applied usefully to yeast (Ficarro et al., 2002;
Chi et al., 2007; Bodenmiller & Aebersold, 2011). However, it is acknowledged by its
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practioners that, for technical reasons, mass spectrometry typically does not query easily either
membrane proteins or proteins present in low abundance, which is the case for many proteins
involved in signaling (Koch and Hauf, 2010). In this regard, it is striking that my approach
yielded three authentic Ypk1 targets that are polytopic integral membrane proteins (Fps1 in the
plasma membrane, and Lac1 and Lag1 in the ER membrane), as well as a low-abundance
transcription factor (Smp1) that shuttles between the cytosol and the nucleus. Thus, the approach
I developed for protein kinase substrate identification may be especially useful for matching
protein kinases to their cognate substrates in those circumstances where the targets are
anticipated to include membrane proteins and/or low-abundance proteins.
In Chapter 4, I presented detailed characterization of the role that Ypk1 phosphorylation plays in
regulation of Lac1 and Lag1, the two catalytic subunits of the ceramide synthase complex (Muir
et al., 2014). These proteins and their function have been conserved throughout eukaryotic
evolution. I found that Ypk1-mediated phosphorylation of Lac1 and Lag1 stimulated the activity
of the ceramide synthase complex and that this upregulation was essential for survival in
response to certain types of membrane stress. We had shown previously that TORC2-Ypk1
signaling increased the activity of the first step of sphingolipid biosynthesis (Roelants et al.,
2011). My new findings documented the further fact that TORC2-Ypk1 not only increases flux
into the sphingolipid pathway, but concomitantly channels the intermediates into end-product
(complex sphingolipids). In addition, my results also indicate that upregulation of ceramide
synthase also helps prevent accumulation of long chain base precursors and their phosphorylated
derivatives, which, when elevated, can inadvertently trigger autophagy under non-starvation
conditions (Zimmermann et al., 2013). This coordinate regulation may be a general feature of
signaling mechanisms that interface with biosynthetic pathways in order to prevent ‘metabolic
crosstalk’ by pathway intermediates that have multiple bioactive functions.
Lastly, in Chapter 5, I described characterization of TORC2-Ypk1 regulation of the
aquaglyceroporin Fps1 and a stress-responsive transcription factor, Smp1. I found that Fps1 is
phosphorylated in a TORC2- and Ypk1-dependent manner and that phosphorylation is required
for glycerol transport across the plasma membrane. Similarly, I found that Smp1 is also
phosphorylated by TORC2-Ypk1 and these phosphorylations are required both to impede the
nuclear entry of Smp1, but also for its function. Both of these substrates are involved in the
cellular response to hyperosmotic shock. This led me to examine the role of TORC2 signaling
during this stress. I confirmed that TORC2-mediated phosphorylation of Ypk1 is rapidly, but
transiently, down-regulated in response to hyperosmotic shock. This apparent inhibition does not
require any of the known high osmolarity sensing pathways in yeast. I found that the drop in
TORC2-Ypk1 signaling simultaneously increases glycerol production by alleviating Ypk1mediated inhibiton of the glycerol-3-P dehydrogenase Gpd1 (Lee et al., 2012b) and promote
glycerol retention by preventing Ypk1-dependent opening of the Fps1 channel. As a result,
intracellular glycerol accumulates to add the cell in coping with hypertonic conditions. Thus, my
screen for novel substrates revealed that TORC2-Ypk1 signaling provides a novel mechanism for
rapid response to hyperosmotic stress.
Future directions
The work presented here provides some insight into the mechanisms by which TORC2-Ypk1
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signaling promotes cellular survival during membrane stress. I believe future characterization of
additional substrates identified in Chapter 3 will continue to provide important new insights into
TORC2 signaling. For example, another candidate substrate I identified is Rod1, a member of
the family of endocytic adaptors known as -arrestins (O'Donnell et al., 2013; Alvaro et al.,
2014). Characterizing how TORC2-Ypk1 signaling regulates the α-arrestin Rod1 might explain
whether and how plasma membrane protein composition is regulated in concert with plasma
membrane lipid composition in response to stress. Additionally, after bioinformatic
identification, only the top ~100 hits were subjected to the secondary, more laborious SDL
screen described in Chapter 3. It should be possible to increase the throughput of the SDL screen
to permit more presumptive candidate substrates to be tested in an unbiased manner. If
additional candidate Ypk1 substrates are found, it would seem likely that their detailed
characterization would yield further insights into the mechanisms by which TORC2 signaling
regulates cellular physiology.
Identification of the substrates Fps1 and Smp1 has revealed that TORC2-Ypk1 signaling is a
novel hyperosmotic shock responsive pathway. TORC2-Ypk1 ouput is rapidly diminished upon
hyperosmotic shock; and, although in the converse direction from HOG pathway signaling with
respect to activation state, TORC2-Ypk1 function responds to the same environmental stress and
with the same kinetics and with similar output (accumulation of intracellular glycerol) as the
HOG pathway. This situation prompts an obvious question: why does the cell have two pathways
that trigger the same response to the same stress? I look forward to future studies that may shed
additional light on the nuances between these two stress-responsive pathways and that may
explain, hopefully, or at least in part, why yeast have evolved such seemingly redundant systems.
Additionally, identification of Smp1 prompted us to examine the role of TORC2-Ypk1 signaling
in hyperosmotic stress-evoked transcriptional changes. Interestingly, cells lacking Ypk1 activity
failed to activate transcription from the STL1 promoter, a classical hyperosmotic stress-induced
promoter, regardless of whether Smp1 was present or not. This observation suggests that the
nuclear entry and/or function of other transcription factors, or alleviation of repression by certain
transcriptional repressors, or the function of other components of the RNA polymerase II
transcription machinery, requires Ypk1-dependent phosphorylation, either directly or indirectly.
Indeed, among the candidate Ypk1 substrates identified by my approach (see Chapter 3) are
Gal11 and Ssn2, two subunits of the mediator complex that acts as a bridge between site-specific
DNA-binding transcriptional activators and the RNA polymerase II holoenzyme (Kornberg,
2005) and Epl1, a subunit of the essential NuA4 histone acetyl-transferase complex that also
promotes gene expression (Doyon & Cóté, 2004). Moreover, in addition to Smp1 itself, a
number of other DNA-binding transactivators were identified in my screen, including Rlm1
(another MADS-box family transcription factor and paralog of Smp1), Fkh1 and Hcm1 (two
Forkhead family transcription factors), and Hot1. Hot1 (for "high osmolarity-induced
transcription") is especially intriguing with regard to the necessity for Ypk1 action in STL1
induction in response to hyperosmotic shock because Hot1 has been shown to be required for the
induction of GPD1 (and its paralog GDP2) under hypertonic conditions (Alepuz et al., 2003;
Gomar-Alba et al., 2013). In contrast to all of the above factors that act positively on
transcription, among candidate Ypk1 targets identified by my methods only a single negative
regulator was identified, Yhp1, a transcriptional repressor of the Homeobox family. In any event,
my findings indicate that TORC2-Ypk1 signaling may play an important role in the induction of
93
the expression of gene products that aid the cell in coping with membrane stress. Therefore, I am
also looking forward to future studies that will address this aspect of TORC2-Ypk1 function.
Although the work described in this thesis addresses downstream aspects of TORC2-Ypk1
signaling, it is important to consider how TORC2 "senses" the status of the plasma membrane.
We know very little about how TORC2 function (state of assembly, localization, activity state,
stability) may be influenced by membrane lipid and protein composition during normal growth
and by stresses (organic solvent, heat shock, lipid synthesis inhibition, hypertonic and hypotonic
conditions, etc.) that perturb plasma membrane composition, structure and/or function. How
TORC2 activity is controlled is an important unanswered question in TOR signaling. By
comparison, significantly greater headway has been made in understanding how TORC1 activity
is regulated (Loewith & Hall, 2011; Laplante & Sabatini, 2012). Importantly, TORC2 activity
appears to be poised at an intermediate level. Some stresses, e.g., hypo-osmotic shock, appear to
upregulate TORC2 activity (Berchtold et al., 2012; Niles et al., 2012), whereas other stresses,
e.g., hyperosmotic shock, appear to inactivate TORC2 function (Lee et al., 2012b). Perhaps each
type of stress influences TORC2 by a unique and separate mechanism, or perhaps TORC activity
is coupled to a single property of the membrane, e.g., membrane tension, that can be
differentially affected by each type of stress. Either model can explain the effects of stress
observed to date. I am particularly excited for future studies that will hopefully define the
molecular mechanisms by which membrane status modulates TORC2 action.
TORC2 signaling is conserved in other eukaryotes, including humans (Cybulski & Hall, 2009;
Huang & Manning, 2009). TORC2 signaling in mammalian cells appears to be sensitive to
membrane stretch and osmolarity much like TORC2 is in yeast (Kippenberger et al., 2005;
Sengupta et al., 2013). Future studies examining regulation of TORC2 and how TORC2 then
regulates cell physiology in other organisms will be interesting as they identify common themes
and species-specific adaptive differences in signaling from this conserved pathway.
Lastly, I am looking forward to future experiments that will continue to give us new
understanding of the varied roles lipids play in cell physiology. My studies of TORC2 found that
strict regulation of LCB-1-P levels were necessary to prevent autophagy. However, due to the
relative difficulties of studying lipids compared to other biomolecules, it is still unknown how
LCB-1-P triggers autophagy. I look forward to studies that will address the role that intracellular
LCB-1-P plays in autophagy induction. In this regard, I am excited by the new lipidomics
techniques and in vivo chemistries to analyze, perturb and visualize lipids that are becoming
available (Atilla-Gokcumen et al., 2014; Muro et al., 2014; Casanovas et al., 2015). Recent
advances have made it possible to even determine the spatial distribution of lipid species on the
micron scale in situ (Soltwisch et al., 2015). It will be exciting to see these new tools being used
to study dynamic changes in the membrane composition of cells undergoing physiological
changes. These new techniques will hopefully give us new understanding of the role that
changing lipid composition plays in membrane function and cellular adaptation, and the
mechanisms that cells use to achieve these outcomes.
Final remarks
The cell membrane is the point of contact between the cell and the outside world. Through this
94
border the cell experiences and is assaulted by the environment. It is fascinating to learn the
mechanisms by which the cell keeps this essential organelle in working shape. TORC2 signaling
is one such conserved mechanism by which eukaryotes accomplish this task. I am very pleased
that my doctoral dissertation research has provided significant new insight and additional
understanding about the molecular mechanisms by which TORC2 action maintains plasma
membrane homeostasis.
95
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Appendix: A study of places to walk around Berkeley and the Bay Area
Introduction
As I was deciding where to go for graduate school, I was extremely fortunate to come across
“Berkeley’s Longest Paths or, why I took so long to graduate”, an appendix from Alexei Efros’
thesis (Efros, 2003). At the time I was reading (and quite enamored with) a whole bunch of stuff
on pedestrianism and its effect on thinking and mental processes. In particular, I was interested
in the links between walking and various philosophical schools from Aristotle to Rousseau,
especially as artfully and exuberantly detailed in Rebecca Solnit’s book Wanderlust. Thus, Dr.
Efros’ appendix and the promise of living in a place with some space to walk about and think
(and yeah, also, the excellent scientists) ultimately sent me to Berkeley.
Dr. Efros prescribed many interesting walks and I followed him around to many beautiful places
in town and in the near East Bay. However, my favorite recommendation of his stemmed from
the obvious observation that his list was by no means exhaustive. Thus, he recommended (in an
excellent pun on math textbook speak) that it would be “[…] an enjoyable exercise to the reader
to discover others”. So, I spent quite a bit of time (ssshhh, don’t tell my advisor!) finding some
other beautiful places, and decided to write my favorites down.
I admit I have been somewhat reticent to write this appendix - does Berkeley really need another
thesis appendix describing walks? Well, if Hollywood can make as many sequels as they want, I
think I can make one small one as well. I’ll focus on the two things I have the most experience
with and can offer the most insight into: (1) walking around Berkeley at night and (2) day hikes
accessible by public transportation. As with Dr. Efros, my experience is quite limited and this
guide is not complete in any sense. I hope it spurs you the reader onto your own investigations.
Lastly, I’d like to note that these recommendations will be loose descriptions, not detailed
instructions or guides. I highly recommend having a good map of Berkeley, such as the Berkeley
Path Wanderers Association map (http://berkeleypaths.org/maps/) or access to Google Maps to
sort out the details.
Night walks
“[…] fretted vault of heaven by night gives you the foil of the infinite, making your petty
exploit a brave adventure”
Stephen Graham, The Gentle Art of Tramping
“Out at night, you understand that wildness is not only a permanent property of land – it
is also a quality which can settle on a place with a snowfall, or with the close of day”
Robert Macfarlane, The Wild Places
Berkeley has many excellent places to walk at night. These make for great after work walks.
When I’ve felt stuck on projects and alienated with the normal routine, the darkness of these
night walks always transported me a thousand miles away from ordinary lab life with its cares
and worries. I’ve always felt like I was coming back into life after a much need long break or
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adventure, even though these walks are only an hour long give or take. I hope you the reader find
them similarly useful.
Panoramic Way tree tunnel
Panoramic Way is a beautiful road that runs up the ridge between Strawberry and Claremont
Canyons. Claremont Canyon drops off dramatically on the south side of the road, giving amazing
views of Berkeley blending into Oakland. As such, parking along the lower stretches of this road
and making out is a favorite past time of the younger set of Berkeley residents. However, an
even more amazing experience is to be had further up the road and away from the legions of
lovebirds. Walk up to Memorial Stadium, and keep walking south along Stadium Rim Way until
you get to Orchard Lane. Take these steps up to Arden Road and hang a left until you get to
Arden Path. Follow this path up to Panoramic Way. Walk along this road until you get to the
Upper Fire Trail. Take the very steep fire trail (commonly known as “the Connector”) up until it
rejoins Panoramic Way. Exit onto this road, and walk the road a bit until it forks. Take the lower
road on the right. As you walk along this narrow and potholed road, you should find yourself in a
tree tunnel. If you pick a moonless night (which I highly recommend), you’ll have the
impression of being in a perfectly black tunnel miraculously suspended above the city. It is quite
disorienting. The light at the end of the tunnel will be downtown Oakland blazing away. Traffic
along CA-24 moves like a snake in the foreground. As you continue, it feels quite a bit like
you’re walking out of the tunnel into the air right above town. Your own personal turn as
Harrison Ford in The Fugitive. Hopefully, Tommy Lee Jones isn’t chasing you with a gun
though. As you exit the tree tunnel, make sure to take a look down the slope to your right.
Someone has made two giant banana slug statues. You’ll see them down there, perpetually
sliming their way up. Just take Panoramic Way and Dwight Way past the lovers in their cars and
back into town. Welcome back to normal life.
Creepy redwood forest
Sometimes the best way to get out of a funk is to scare the hell out of yourself (or at least creep
yourself out a little bit). The Berkeley fire trails in Strawberry Canyon are an amazing place to
walk at night. During the day, they are packed with people and it is a fun park to go people
watching. During the night, the trails become deserted and walking here then takes you far away
from the normal routine and places you on adventure in a wild forest. It’s a great place and time
to see animals from deer to the occasional tree climbing fox! It’s also just spooky enough to pull
you out of everyday life. There is plenty to explore out there, but here is one route I recommend.
Walk up to the stadium, and take Centennial Drive up past Strawberry Canyon Pool. The fire
trail heads up from a small parking lot on the right. Take the trail up and keep your eyes open.
After ~10 minutes of walking up the trail, the trail bends and on the left there is a small plaque
marking the entrance to the redwood grove. Take one of the many trails heading up into the
grove on the left. The redwood grove is beautiful and uncrowded at night, except for the
occasional fellow walker or illegal Side-O mountain biker. The stars and moon disappear under
the canopy of the trees as you head further in, giving the grove a dark and spooky feel. I highly
recommend bringing a flashlight if you get easily spooked. The upper exit of the grove puts you
back at the fire trail near the top of the Connector. You can take the Connector down and head
back into town using the Orchard or Arden steps. Or you can head from here to the Panoramic
Way tree tunnel described above.
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Atlas and Scott Newhall Paths
The view from Atlas and Scott Newhall Paths cannot be beat! To get there, take Cedar Street to
La Loma Avenue and then head north on La Loma. Take a left onto Glendale Avenue and then
follow the Glendale Paths east up the hill. After the Glendale Paths end, take Arcade Avenue.
Then cross Grizzly Peak Boulevard. Atlas Path heads east from the left hand side of Grizzly
Peak. After about half an hour of non-stop uphill walking, there is a bench on Atlas Path that
provides a much needed rest, along with panoramic views from Richmond to the Golden Gate to
San Francisco. Keep going up Atlas Path and make a left hand turn on Hill Road. Follow this
road about 2 minutes until it dead ends. Scott Newhall Path (named after the famed San
Francisco Chronicle newspaperman) starts from the right hand side of the road and provides
again beautiful views as you walk along. From here take Hill Road down and then many routes
are available back to campus.
Northside Berkeley paths loop
This walk loops together several of my favorite north Berkeley paths. Take Spruce Street up
north. Take a right on Easter Path shortly before Marin Avenue and follow this path up. It ends
in Cragmont Park, which has good climbing (rock and tree) and views onto the Bay (and none of
the crowds of Indian Rock). From here, follow Regal Road east and take a left onto Pinnacle
Path. This path has a very interesting ceramic mosaic wall. It is especially beautiful at night,
where the tiles reflect and twinkle the light of your flashlight. From here, take a right onto Keeler
Avenue. This ends in Keeler Path, which winds through Remillard Park and leaves you feeling
on a trail in the forest far from town, if only for a second. When this path ends, take an
immediate right onto Sterling Path, which has dramatic views down into Berkeley and the Bay.
From here, Bret Hart Road and Bret Hart Way lead to Euclid Avenue and back to campus.
Garber Park
This walk leads through Garber Park and the local south Berkeley environs. It feels super secret
and leads through a lovely little park filled with many ferns. To get there, walk east on
Claremont Avenue past the Claremont Hotel. It’s the giant ostentatious white building, you can’t
miss it. It’ll feel like you’re walking past something out of The Great Gatsby. Keep on going
east, and a park will rise steeply on the right hand side of the road. There is a trail that heads up
the hillside and a little ~1 mile loop trail that goes through the park. Take the loop around and it
ends at Evergreen Lane. Take Evergreen Lane west where it ends in Evergreen Path. This path
goes to Alvarado Place which leads to the Shortcut Path. Shortcut Path leads back to Tunnel
Road, which you can then follow west back towards campus.
Public transit assisted hikes
“The end of automania would save open spaces, encourage wiser land use, and contribute
greatly to ending suburban sprawl. It would lead to the building of more compact,
sensitively planned communities in the future—and it would prompt many cities to build
quick, quiet, and convenient modes of transportation ranging from bicycle paths to mass
transit systems”
Stewart Udall, “The Last Traffic Jam”, The Atlantic
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One of the great things about the Bay Area is that it is easily possible to get to beautiful parks far
out of town by public transit alone. Many cities have great public transit systems, except when
you want to leave town to vacation. Then it becomes necessary to have a car. Berkeley is quite
the exception and it is possible to get far out of town, and go camping even, all without a car.
This unique aspect of public transit is an exceptionally good community resource, and I have
been glad to take advantage of it. I hope for its continued existence. Here are some of my
favorite day trips and the currently available transit options to get there.
Redwood Regional Park to Berkeley
A very nice ~6-7 hour trip is to follow the hills from Oakland’s Redwood Regional Park back to
Strawberry Canyon and Berkeley. To get to Redwood Park, take BART to Fruitvale Station. A
nice pre-walk treat can be found at Fruitvale. Powderface at Fruitvale BART serves excellent
coffee and beignets. From Fruitvale BART, ride AC Transit bus #39 east into the hills. Take the
stop at Skyline Boulevard and Crestmont Drive. Cross the street and you are in Redwood
Regional Park. There are many trail options now to head back north towards Berkeley. The most
direct is to take Dunn Trail to the Skyline National Recreational Trail and follow this north to
Berkeley. The Skyline Trail is a beautiful trail that follows the spine of the Oakland/Berkeley
hills. As you head north from Redwoods Park, you’ll cross through much varied scenery. From
the damp and forested Huckleberry Botanical Preserve, to the dry and windswept grasses as you
cross over the CA-24 Caldecott Tunnel into Tilden Park. The trail is well marked and easy to
follow. Many options for exploration and variation on the standard trail exist. An excellent
detailed description of the Skyline Trail and links to various resources can be found here:
http://gurmeet.net/hiking/hikes/East_Bay_Skyline_National_Recreation_Trail.html.
Once you have followed the Skyline Trail back to Tilden you might think you’re going crazy as
you hear the faint sounds of a stream train whistle. You have not. The park operates a small
model train near Vollmer Peak. It is a nice break to stop and ride the train. From here, you can
take any of the Tilden Park paths back towards Berkeley. A more direct route is to cross Grizzly
Peak Boulevard and where trailheads exist for the fire trails leading back into either Claremont
or Strawberry Canyons. All in all, it is quite the adventure, and a very satisfying feeling to bring
yourself home under your own power.
Another small note. AC Transit #39 runs right alongside Joaquin Miller Park before arriving at
Redwoods Park. This is another nice park to explore. It also hosts a permanent orienteering
course. The maps and directions can be downloaded from here:
http://baoc.org/wiki/Maps/Permanent_Courses. This makes for another nice afternoon walk and
adds some adventure while sharpening up your map and compass skills.
Lake Chabot to Redwood Park
It is possible to go even further afield in the Oakland hills using public transit. Rather than
starting in Redwood Park, it is possible to take public transit to Lake Chabot Regional Park. To
get to Lake Chabot, take BART to the Coliseum stop. Exit here and take AC Transit #46L to the
stop at Dunkirk Avenue and Golf Links Road. A short walk along Golf Links Road will take you
to a staging area with access to the many trails of Lake Chabot. A very enjoyable day long loop
is to take the Goldenrod Trail down to the end of Lake Chabot and the start of Grass Valley
Creek. If you are into fishing, and have the requisite permits, this trail takes you right into Bass
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Cove at Lake Chabot. I’ve never caught a bass here, but I can say for sure there are some catfish
there. After visiting Lake Chabot, cross over the creek and take the Columbine Trail back. This
trail follows the creek and eventually links with the Grass Valley Trail. This trail is quite
beautiful and follows a long grassy valley, as the name would imply. It is particularly beautiful in
the summer, where as far as you can see there is only yellow grass in the foreground, green
hillsides on the left and blue sky everywhere else. The entire world in three colors. This trail
connects to the MacDonald Trail further north, which can then be followed to the Golden Spike
Trail in Redwood Park. This trail can then be followed to an AC Transit #39 bus stop, which
takes you back to BART and Berkeley.
Briones and Layfayette Ridge
A favorite trip of mine is out to Briones Regional Park. This is a bucolic park with rolling green
hills, and it is a particularly nice place for full moon hikes (and dark and clear enough to see
meteor showers). The park also hosts another of the Bay Area Orienteering permanent courses as
well. One of the nicest things about Briones, is that BART can get you there in about an hour.
Take BART out to the Lafayette stop. Then walk east on Deer Hill Road before making a left
hand turn on Elizabeth Street. Follow this to the end, and you will end up at the park on the
Lafayette Ridge Trail. If you Google image search this trail, you’ll see a trail rolling endlessly
along a ridge of green grassy hills. Mount Diablo stands off in the distance. Being there is every
bit as awesome as it sounds. The whole ridge trail is quite long and makes an excellent morning
to afternoon trip. Unfortunately, as far as I’ve ever looked, there are no other transit options out
there beyond the BART station (I know, life is so hard sometimes), so this trip is an out and
back. Head back down to the Lafayette BART station when finished exploring.
Mount Tamalpais to the Marin Headlands
I found out about the Marin Transit coaches when I was a third year student, and I felt like I was
let in on a great secret. These buses will take you all along the west edge of Marin from Stinson
Beach to Point Reyes, all for only $2. This makes it possible to explore the huge park system and
beautiful beaches stretching from the Golden Gate all the way to Tomales Bay. Marin Transit
#61 takes you through Mount Tamalpais, to Muir Woods along the way, and down to Stinson
Beach. This bus makes it possible to do many trips in the Mount Tam area. Some examples.
Take the bus to Stinson beach to spend the morning relaxing, before walking up Mount Tam in
the afternoon and catching the bus home at Muir Woods. Circumambulate Mount Tam with Gary
Snyder’s poem in hands. The possibilities are many. I’ll describe a slightly longer tour (~12
miles) that has become my favorite. Take BART to San Francisco at catch Golden Gate Transit
#70 bus to the Donahue and Terners stop. From here take Marin Transit #61 to Pantoll Station.
From Pantoll, walk south on the Coast View Trail. At times I’ve started from Pantoll above the
fog line, and steadily descended below it. Quite an experience. Take the Heather Cutoff and
Redwood Creek Trail into Muir Beach. If you’re feeling snackish, the Pelican Inn offers nice
desserts and a bar. Great for a break. From Muir Beach, follow the Coast Trail into the Marin
Headlands. This trail hugs the coast with its dramatic cliffs falling into the Pacific. Quite
fetching. Keep following the Coast Trail (or one of many variations that are possible) into the
Rodeo Beach/Fort Cronkhite area. From here, take the MUNI #76X bus back into San Francisco.
Please note, as of now, the MUNI #76X only runs weekends.
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Grand adventure from Point Reyes to Mount Tamalpais
Lastly, Marin Transit #68 will take you to the visitor center at Point Reyes National Seashore.
I’ve still never gotten over how great that is. Point Reyes is one of the most beautiful places in
California. Using #68 bus, it is possible to do many day trips in this area. Point Reyes also has
many excellent camping sites and one excellent hostel, makeing overnight trips and further
exploration quite easy. I leave all of this for you to explore. Instead, I will describe one of my
favorite trips. This is a very long (~20 mile) trek from Point Reyes to Pantoll Station at Mount
Tam. It requires a bit of commitment and perhaps an overnight stay at Pantoll Station if you miss
the last bus home. For the challenge, you’ll be rewarded with a trip across much varied and
striking scenery. From the Point Reyes Visitor center, walk south along the Rift Zone and Olema
Valley Trails. You’ll pass many miles and sag ponds in the Olema Trough, where two tectonic
plates abut each other. Take a left and cross Highway 1 and go up to the Bolinas Ridge Trail on
either the Randall or McCurdle Trails. From here, continue south until you reach the FairfaxBolinas Road. Cross this road and head south along the Coastal Trail. This trail traverses steep
grassy hills that fall hundreds of feet to Bolinas below, a thin brown stripe edged into the huge
endless hills. Follow this trail to Pantoll. Along the way, you’ll pass a knocked over tree that
strikingly has sent all of its available branches skyward. If you ever need inspiration to keep
going when stuff isn’t going your way, there it is (I know this is gross anthropomorphization but
I don’t care). If you’re lucky enough, you might see some hang gliders launching from the hills
above you. If you’re really lucky, maybe even a fox. You’ll be sad when it’s over and you’re at
Pantoll. From here, you can catch the #61 coach back to town. Or if you’re lucky and you missed
the last bus, you’ll have to camp at Pantoll and wait for the next one in the morning. They have a
no-reservations, never filled camp site specifically for hikers. So, you needn’t worry about
having a place to stay at Pantoll. As of now, it is only $5 to stay in the hiker site.
References
Efros, A.A. (2003). Data-driven approaches for texture and motion (University of California,
Berkeley), pp. viii, 83 leaves.
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