Membrane Stress and the Role of GYF Domain Proteins
Membrane Stress and the Role of
GYF Domain Proteins
Alexander Georgiev
Stockholm University
© Alexander Georgiev, Stockholm 2008
ISBN 978-91-7155-683-7
Printed in Sweden by Universitetsservice AB, Stockholm 2008
Distributor: Department of Biochemistry and Biophysics, Stockholm University
To my daughter Joanna
List of Publications
This thesis is based on the following papers, which are referred to in the text
by their Roman numerals.
I
II
III
IV
∗
Georgiev, A., Sjöström, M., Wieslander, Å. (2007). Binding specificities of the GYF domains from two Saccharomyces cerevisiae
paralogs. Protein Eng Des Sel, 20(9):443–52
Georgiev, A.∗ , Leipus, A.∗ , Olsson, I., Berrez, J-M., Mutvei, A.
(2008). Characterization of MYR1, a dosage suppressor of YPT6
and RIC1 deficient mutants. Current Genetics, 53(4):235–47
Georgiev, A., Ge, C., Wieslander, Å. (2008). Lipid-specific
interactions of Myr1 suggest a role in sensing membrane
stress. Manuscript
Wikström, M., Kelly, A.A., Georgiev, A., Eriksson, H.M., Rosén
Klement, M., Bogdanov, M., Dowhan, W., Wieslander, Å (2008).
Curvature-engineered Escherichia coli bilayers reveal critical
lipid head-group size for membrane protein function in vivo.
Manuscript
These authors have contributed equally to the paper.
Reprints were made with permission from the publishers.
Contents
Introduction . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Budding yeast as a model organism . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Classification, life cycle and biology . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Genetics . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Studying vesicular traffic in yeast . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Membrane traffic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Endomembrane systems in eukaryotic cells . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Protein translocation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
The life cycle of transport vesicles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Rab GTPases . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
ER to Golgi . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Intra-Golgi traffic and cisternal maturation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Exocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Vacuolar traffic and endocytosis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Retrograde traffic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lipids in Saccharomyces cerevisiae . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lipids in organelles . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Synthesis and distribution . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Regulation of synthesis . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Phosphoinositides . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lipids and membrane traffic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Protein-lipid interactions . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Membrane binding proteins . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Mechanisms of membrane binding . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Lipid binding and vesicular traffic . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Membrane stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Spontaneous curvature . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Physiological membrane stress . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Examples in eukaryotes and prokaryotes . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Importance of membrane stress for homeostasis in unicellular organisms . . . . . . . .
Translational control, RNA decay and stress . . . . . . . . . . . . . . . . . . . . . . .
Translation initiation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
RNA degradation . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Processing bodies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .
Nutrient deprivation and the TOR signaling pathway . . . . . . . . . . . . . . . . . . . . . . .
1
3
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4
4
5
5
6
6
8
9
10
10
11
12
13
13
14
15
16
17
19
19
19
22
23
23
24
25
26
27
27
28
29
30
The GYF domain . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33
Structural features . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 33
GYF domain protein families . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 34
Binding partners . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 35
Membrane traffic connections . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36
Relation to RNA metabolism . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 36
Integrating knowledge by computational biology . . . . . . . . . . . . . . . . . . . 37
Gene Ontologies . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38
Multivariate analysis of protein sequences . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 38
Summary of papers . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41
The GYF Domain of Myr1 and Smy2 (Paper I) . . . . . . . . . . . . . . . . . . . . . . . . . . . . 41
MYR1 – a suppressor of YPT6 and RIC1 mutants (Paper II) . . . . . . . . . . . . . . . . . . 42
Myr1 as a membrane sensor – interactions with lipids (Paper III) . . . . . . . . . . . . . . 43
Lipid-engineered bacteria (Paper IV) . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 45
Discussion and perspectives . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 47
Acknowledgments . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 51
References . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 53
Abbreviations
COPI
coat protein I
COPII
coat protein II
GAP
GTPase activating protein
GEF
guanine exchange factor
GFP
green fluorescent protein
GO
gene ontology
GYF
glycine-tyrosine-phenylalanine, signature amino acid motif of
the GYF domain
eIF
eukaryotic initiation factor
ER
endoplasmic reticulum
LUV
large unilamellar vesicle
MYR1
multi-copy suppressor of YPT6 and RIC1 (≡SYH1)
NSF
N-ethylmaleimide-sensitive factor
P-body
processing body, cytoplasmic site of mRNA degradation
PCA
principal components analysis
PLS
partial least squares
PA
phosphatidic acid
PC
phosphatidylcholine
PE
phosphatidylethanolamine
PI
phosphatidylinositol
PS
phosphatidylserine
PIPs
phosphoinositides, phosphorylated derivatives of PI
PITP
PI transfer protein
SMY2
suppressor of myo2 mutant
SNARE
soluble NSF attachment receptor
SRP
signal recognition particle
SYH1
Smy2 homolog, synonym for MYR1
Introduction
The divide between what we perceive as living or not is six nanometers thin,
and is known as the cellular membrane. It is the semi-permeable barrier
that topologically defines the cell and is responsible for maintaining the
gradients characteristic of living systems. In the following pages, I present a
study focused on the interplay between the two major constituents of biological membranes - lipids and proteins. The main objects of this study are
the two poly-proline binding GYF domain proteins Myr1 and Smy2 from
bakers’ yeast, both identified through genetic screens in trafficking mutants. Two independent lines of evidence connect these proteins to intracellular membrane traffic and to processing of messenger ribonucleic acid
(mRNA). These lines cross at the regulation of cellular membrane homeostasis through membrane-protein interactions, a research area characterized by a precipitous accumulation of data during the last few years.
Aiming for a deeper understanding of the nature of GYF domain proteins,
and of their role in the cellular response to membrane alterations, I outline the fundamental processes in which they take part. To establish the
biological context, I also briefly present the unicellular organisms Saccharomyces cerevisiae and Escherichia coli which, despite their idiosyncrasies,
have served as models to study membrane-protein interactions in general.
An emphasis is placed upon some evolutionary aspects of the different cellular processes and components. Thus, while most of the background is
heavily weighted towards budding yeast, examples from other domains of
life and especially those dear to us mammals are given when appropriate.
Separate chapters in the thesis are devoted to intracellular membrane
traffic, biogenesis and dynamics of lipids and membranes, and
lipid-protein interactions. Their connections to GYF domain proteins are
further evaluated in regard to transcript stability, regulation of translation
initiation, and membrane stress. Computational methods used to establish
some of these connections are also described. Finally, my contribution to
the advancement of knowledge in this field is summarized in a synopsis of
papers.
It is inevitable that once we succeed reaching the realms of the known
we are left facing the unknown, and therefore this work leaves many questions open. It is my conviction though that the years spent on it are merely
a preparation for the quest for answering those and many other questions.
1
Budding yeast as a model organism
The budding yeast Saccharomyces cerevisiae is one of the best studied eukaryotic organisms. It has been known to humans and used for millenia
for its ability to ferment sugars. In the past few decades, it was also established as an important research tool for genetics and molecular biology.
The advantages of yeast over other model systems combine rapid growth,
ease of cultivation, ease of genetic manipulation, low pathogenicity, and
the conservation of fundamental processes. Major metabolic and signaling pathways, such as cell cycle control and membrane trafficking, are very
similar in fungi and animals, and have often been initially studied in yeast
[131, 176].
Classification, life cycle and biology
The kingdom Fungi is widespread in all possible environments. Nearly 1000
species of yeasts have colonized the depths of oceans, the soils of earth, and
our own bodies. S. cerevisiae belongs to the Ascomycota phylum and was
named after the beer malt from which it was isolated in 1837 [217].
In nature yeasts are mostly diploid, and undergo meiosis under certain
nutrient limitations, only to mate again when conditions permit. Wild type
strains are normally homothallic, meaning that haploids are capable of
switching their mating type with a high frequency during budding, which
allows for rapid diploidization. To make genetic studies feasible, lab strains
are selected that have lost the mating type switching ability.
Yeast cells are oval and 5-10 µm in size. Both haploid and diploid cells
propagate by budding and optimally complete their cell cycle in 90 minutes. Several cellular structures may be observed by light microscopy and
with the aid of different staining techniques. Intracellular membranous
compartments in yeast include the endoplasmic reticulum (ER), the Golgi,
the vacuole, endosomes, multi-vesicular bodies, and transport vesicles
(fig. 1). Those are similar in function to the corresponding organelles in
metazoans, with the vacuole playing a comparable role to the lysosome
of animal cells. The nuclear envelope, a specialized extension of the ER
enclosing the genetic material, does not disassemble during cell division, a
phenomenon known as closed mitosis and also typical for some protozoans
[31].
3
Figure 1: Yeast cell - schematic representation. 1. ER, 2. Golgi cisternae at different levels of maturation, 3. Vacuole, 4. Nucleus, 5. Early endosome, 6. Pre-vacuolar
compartment, 7. Lipid droplets, 8. Bud, 9. Cell wall, 10. Mitochondrion. 11. Peroxisomes.
Genetics
Yeast is a widely used object of classical genetics, aided by the existence
of both haploid and diploid vegetative forms. Simple techniques for tetrad
dissection allow for the recovery of the products of meiosis, and the ability
to grow on synthetic media with completely defined composition makes
selection of diverse markers possible.
S. cerevisiae was the first eukaryote to have its entire genome sequenced,
in 1996 [73].The yeast genome is organized in 16 chromosomes, 12 million
nucleotides, and 5795 protein coding genes. While 1/4 of the yeast genes
are similar to human ones, a comparison between 13 Hemiascomicetous
species determined that another 1/3 are specific for Ascomycetes [144]. The
number of characterized yeast genes is steadily growing but despite earlier
predictions that the functions of all genes will be known already by 2007,
there are still more than 1000 uncharacterized genes, many of them yeastspecific [102, 187].
Studying vesicular traffic in yeast
Budding yeast has traditionally served as a model organism for the dissection of membrane traffic. One of the first studies dedicated to vesicular traffic and secretion is the work by Peter Novick and Randy Schekman
[173, 176]. The conservation of the mechanism of membrane traffic and the
lack of tissue specialization in yeast have allowed determining the function
of many protein regulators with homologs in metazoan species.
4
Membrane traffic
Living cells are highly organized structures in a fragile dynamic equilibrium. To maintain their organization during normal biosynthetic activity
and growth, they have developed pathways for regulated delivery of macromolecules. Membrane traffic is of special interest here and is presented in
this chapter. Other types of traffic, such as nucleo-cytoplasmic, import of
proteins into mitochondria and chloroplasts, or transport of molecules and
ions across membranes, are outside the scope of this work.
Endomembrane systems in eukaryotic cells
Nominally, the difference between prokaryotes and eukaryotes is the presence of a nucleus (karyon). It is in fact the existence of an evolved membrane system in eukaryotes that allows for the increased cell complexity by
compartmentalization; indeed, even the eponymous organelle organizing
the genetic material in eukaryotes may be regarded as a result of this compartmentalization [21].
The endomembrane compartments of animals and plants are similar
to those present in yeast as described above. Plant cells have vacuoles
while animal cells have lysosomes; due to the functional similarities and
homologous proteins in the two types of organelles we regard them as
analogous. Multicellular organisms with specialized tissues have evolved
types of membrane compartments specific for new functions, such as
melanosomes or secretory granules of neurons containing respectively
melanin or neurotransmitters.
Membrane compartments differ by the lipid and protein composition of
their membranes. Their identity is a result of their sequential biogenesis
and the directional routes of exchange of membrane material between
them. A typical biosynthetic pathway followed by a secretory protein
would begin with synthesis at membrane-bound polysomal complexes
at the ER, followed by translocation through the membrane leading
to either membrane insertion (for integral proteins) or to lumenal
localization. Membrane material travels further via vesicle and tubular
intermediates, to an intermediate compartment between ER and Golgi,
then to early, mid and late Golgi, and eventually by secretory vesicles to
the plasma membrane [181]. Outgoing traffic in this direction is referred
5
to as anterograde, in contrast to retrograde traffic. The latter includes
internalization of plasma membrane material by endocytosis, transport
of endocytic vesicles to a pre-endosomal compartment, with a final
destination either in the vacuole (for degradation) or backwards via Golgi
to the ER.
Protein translocation
Protein translocation at the ER, currently understood at the structural level
[234], is achieved through the translocon composed of Sec61, Sec62, Sec63,
Sec71 and Sec72 [191, 192]. The precursors of secretory proteins pass from
the cytosol into the lumen through a membrane channel formed by Sec61.
This process requires ATP and molecular chaperons of the hsp70 family,
exemplified by yeast Ydj1 and E. coli DnaJ.
Depending on whether the polypeptide synthesis is complete before
or after the onset of translocation, two major routes exist. For yeast,
post-translational translocation is typical. It requires that protein
precursors are completely synthesised in the cytosol and maintained there
in semi-folded state by chaperons homologous to E. coli DnaJ.
The co-translational translocation is used by other secretory precursors,
especially in mammalian cells. Upon such translocation the need for chaperons is relieved, however another set of cytosolic factors known as the signal recognition particle is needed [87, 123]. A ribosome–nascent-chain–SRP
complex binds to the translocon channel, after which the SRP is released
and the translating ribosome remains attached to the membrane via interactions with the channel [179, 192].
In the ER lumen another hsp70 chaperon exists known as BiP, encoded by
the KAR2 gene in S. cerevisiae. It binds to a peptide loop of Sec63 containing
a 70 residue region homologous to the ’J box’ of the DnaJ family of proteins.
The BiP protein serves as a molecular ratchet pulling the polypeptide chain
inside the lumen.
The life cycle of transport vesicles
For prokaryotes, the process of secretion is complete with protein translocation through the membrane. In contrast, this is the just the beginning for
eukaryotes. Once in the ER, proteins continue to traverse the endomembrane system in a regulated fashion, being carried inside transport vesicles.
Before going into more detail about the specific steps of the transport, one
needs to understand the general mechanism of formation of the carriers,
their molecular anatomy, and the way they are guided towards and fuse
with their targets.
6
Membrane vesicles emerge through invaginations of a donor membrane
that pinch off to form a spherical vesicle, typically 50-100 nm in diameter.
The initial bending of the membrane is caused by successive attachment
of protein molecules to its cytoplasmic side. These proteins include small
GTPases, coat proteins, and cargo adapters. Once the vesicle is formed the
membrane undergoes fission, thus releasing the completed carrier in the
cytosol. At this point, many vesicles “shed off” their protein coats to facilitate later fusion.
Figure 2: Vesicle cycle, adapted from Behnia & Munro [15].
Depending of the type of vesicle, it travels through the cell either by diffusion or through a vectorial transport along cytoskeletal elements. Upon
reaching its final destination, the target membrane, vesicles are tethered by
long filamentous proteins or multi-protein complexes. Specific attachment
to the correct membrane is ensured by the tethering mechanism, but also
enhanced by the next step, docking of the vesicle to the membrane with the
participation of both vesicle and target SNAREs. Finally, the membrane of
the vesicle fuses with the target membrane and the soluble cargo is released
inside the recipient compartment (fig. 2).
It is very important, from an evolutionary perspective, to realize that despite the strict specificity existing when it comes to the individual protein
regulators of the vesicle life cycle, the overall scheme of the process is the
same at each stage and indeed involves homologous proteins.
Sometimes an intermediate stage in vesicle formation is stabilized leading to formation of tubules instead. This is commonly observed in some
organelles, but can also be the result of aberrant vesicle formation due to
mutations in the protein regulator genes [51, 130].
7
Rab GTPases
The Rab family of small GTPases (as opposed to large GTPases, such as
various structural proteins or G proteins) is the largest subfamily of RASrelated GTPases. More than 60 homologs exist in humans, and 11 in yeast
[128, 188]. As their name suggests, they catalyse GTP hydrolysis to GDP and
phosphate, a reaction coupled to structural rearrangements within the GTPase molecule and hence to its activity.
The functional cycle of a Rab GTPase (fig. 3) begins with synthesis in the
cytoplasm, followed by post-translational prenylation necessary for membrane association. Initially Rabs bind to GDP which is more abundant than
GTP, and in complex with a Rab escort protein they travel to their site of action. The latter is marked by the specific guanine exchange factors (GEF)
which ensure that Rabs will only be activated at the correct site. Activation
itself is catalysed by the GEF and consists of ejecting the GDP molecule
and replacing it for a GTP [75]. Once activated, Rabs remain attached to
the membrane and may travel in a bound state with the transport vesicle.
When specific effectors become available, Rabs complete their cycle by interacting with the effectors and hydrolyzing GTP, a process stimulated by
GTPase activator proteins, GAPs. At this point, Rab-GDP is extracted from
the membrane to be reused for another cycle (see also Goody et al. [75] for
a recent review).
Figure 3: Functional cycle of Rabs.
Rab/Ypt proteins participate in four major steps of membrane traffic
(fig. 2), cargo sorting, myosin dependent transport, vesicle tethering, and
membrane fusion, recently reviewed by Grosshans et al. [78]. In contrast
to the evolutionary conservation of nucleoporins and SNAREs [117], the
Rab GTPase family has diversified in parallel with the specialization of
endomembrane systems in multicellular organisms [78, 80, 205]. Yeast
contains a minimal set of Rabs [128] which can be regarded as a reference
to the more diverse Rabs in animals and plants. The site of action and the
set of activators and effectors known for the yeast Rabs are listed in Table 1.
8
Table 1: Rab GTPases in S. cerevisiae and their known interaction partners.
GTPase
localization GEF
effectors
GAP
Sec4
PM
Sec2
Sec15 (exocyst),
Sro7
Msb3,
Msb4
Ypt1
Golgi
TRAPP
Sec34/35 (COG)
Gyp1
Ypt31/32
endosome,
Golgi
?
Sec2, Rcy1
?
Ypt51/52/53 endocytic
carriers
?
Vac1, CORVET
?
Ypt6
endosome,
Golgi
Ric1/Rgp1
complex
GARP/VFT
Gyp6
Ypt7
vacuole
Vps39
VPS/HOPS
Gyp7
ER to Golgi
Inside the ER, protein precursors are exposed to a changed environment
of higher oxidation and to a number of modification and maturation enzymes. First, the signal peptide is cleaved off by the Sec11/Spc1-3 complex
[23, 55, 160, 176]. Enzymes of the protein disulfide isomerase family are responsible for rearranging the disulfide bridges upon refolding of the protein
after translocation [57]. The ER is also site of initial post-translational modification by glycosylation [220].
From the ER, proteins may continue further by being exported in membrane vesicles. During the formation of vesicles, cargo selection is important since certain proteins, such as the processing enzymes, need to be retained in the ER. A typical retention signal for ER is KDEL (HDEL in yeast)
[186], while motifs such as FF or ExD serve as signals for packing into vesicles for transport.
Vesicles bud from the ER with the participation of the COPII complex,
consisting of the small GTPase Sar1 and the dimers Sec13-Sec31 and Sec23Sec24 [7, 130]. Sar1 is activated by Sec12 through binding to GTP [161, 162],
via a mechanism common to small GTPases. As a results, the N-terminal
amphipathic α-helix of Sar1 inserts into the membrane, causing a local deformation of the lipid bilayer and subsequently tubulation [130]. The presence of bulky hydrophobic residues in the α-helix is essential for its ability
to induce membrane curvature.
After attachment to the membrane, Sar1 recruits the Sec23/Sec24 dimer
[13]. Sec23 and Sec24 are structurally similar [129], however Sec23 is responsible for Sar1 binding, while Sec24 fulfills the task of recruiting cargo,
which additionally stabilizes the association with the membrane [7]. Fi9
nally, the tetramer composed of two Sec13 and two Sec31 subunits binds,
and subsequent polymerization forms the COPII cage [212].
Vesicles leaving the ER are exclusively targeted towards the Golgi. While
in mammals this proceeds through an intermediate compartment, ERGIC,
S. cerevisiae apparently delivers the vesicles directly to the cis-Golgi. Vesicles are tethered to the Golgi by the transport complex TRAPP [106] and by
Uso1 [32] and fuse under the regulation of Ypt1 [204]. This process is stimulated by the Ypt1 GEF, TRAPP [108], and inhibited by Gyp1, a GAP protein
for Ypt1 [52]. It also involves the v-SNAREs Sec22, Bet1, Bos1 and Ykt6, and
the t-SNARE Sed5 [43, 89, 168].
Intra-Golgi traffic and cisternal maturation
The Golgi is by far the most intriguing organelle in membrane traffic, due
both to its function as a central sorting station, and to its unique morphologies in different cell types and its diverse biosynthetic activities. Indeed,
until it reaches the Golgi, the vesicular pathway is practically unbranched,
since all vesicles generated at the ER target Golgi [24].
Yeast Golgi is functionally equivalent to its metazoan counterpart, however it is morphologically different. The Golgi apparatus cisternae are typically closely stacked in mammals, but in S. cerevisiae they are represented
by dispersed organelles. Those individual cisternae correspond to the functionally defined early (cis), medial, and late (trans) Golgi, and can be labeled by markers specific for these compartments. They are highly dynamic
structures, which are converted directionally from early to later forms, in
agreement with the cisternal maturation model [142, 148].
Golgi is the site of formation of COPI coated vesicles, the second coated
vesicle type to be described after clathrin [178]. Vesicle formation is initiated by Arf1, a “cousin” of Sar1 GTPase, by a similar mechanism. Membrane
insertion of the N-terminal helix of Arf1 follows activation by nucleotide exchange, catalysed by the Sec7 GEF. The COPI complex formed from seven
protein in the cytosol binds en block to the membrane [88]. COPI coated
vesicles mediate retrograde intra Golgi and Golgi to ER traffic.
Other routes leading from the Golgi to the plasma membrane or vacuole
are explained in more detail below.
Exocytosis
Secretion as the ultimate fate of most of the material participating in
membrane traffic has several functions, depending on the type of cell.
Non-specialized unicellular eukaryotes such as yeast use secretion in
order to expand their surface, therefore this is an essential process
10
needed for cell growth. Cells that possess any type of extracellular wall
or matrix produce and modify it by secreting building materials such as
polysaccharides and by exporting the modifying enzymes, which often
are retained in the periplasmic space. Finally, highly specialized cells
as the epithelial cells or neurons secrete unique enzymatic products or
neurotransmitters according to their function. All these processes occur
through the regulated formation of secretory vesicles at the Golgi and their
targeted transport and fusion with the plasma membrane.
The exocyst is a special landmark composed of eight proteins and responsible for the directing exocytosis [79, 224]. It interacts with the Sec4
GTPase, which is present on post-Golgi secretory vesicles after being activated by its GEF Sec2. Correct targeting of secretory vesicles involves the
cytoskeleton, and fusion with the plasma membrane requires a specific set
of SNAREs [174].
Vacuolar traffic and endocytosis
The vacuole is the important “back yard” of the yeast cell where various
compounds such as salts and amino acids are stored. The presence of hydrolytic enzymes is essential for the function of the vacuole and their activity is ensured by the environment characterized by lower pH. Aside from
the resident vacuolar enzymes, proteins are targeted there for degradation.
Membrane material reaches the vacuole via a pre-vacuolar compartment,
known as the late endosome, which on its part receives input from the late
Golgi or the early endosome. Through a special budding mechanism directed inwards, late endosome compartments are transformed into multivesicular bodies, thus allowing for the degradation of the enclosing membrane material as well as the cargo.
Cargo sorted for vacuolar delivery at the late Golgi includes several hydrolases such as carboxypeptidase Y, proteinase A and B [30, 77]. A peptide
signal is required for correct sorting to the vacuole, and in its absence the
proteins end up at the cell surface [30, 145]. Importantly, exit from the Golgi
towards a pre-vacuolar compartment is dependent of Vps34, a PI3 kinase.
Subsequent steps have been dissected through the analysis of vacuolar protein sorting (VPS) mutants [12, 193].
Another source of traffic to the vacuole is endocytosis. It is a means to internalize plasma membrane material in order to regulate signaling or permeability by retrieving receptors and transporters from the cellular exterior.
An evolutionary conserved process, endocytosis is largely associated with
clathrin in yeast as in mammals, even though it is possible in the absence
of clathrin in yeast [109].
Clathrin and Ede1 are the first two proteins to engage in endocytosis. The
presence of PI(4,5)P2 is necessary at endocytic cites. A specific structure as11
sociated with endocytosis is the eisosome, large complexes at the cell periphery containing the proteins Pil1, Lsp1, and Sur7 [110, 227].
Retrograde traffic
Following endocytosis, membrane material continues from the early
endosome either to the vacuole or potentially all the way to the ER. [24].
Endosome-to-Golgi recycling is of particular interest here, since we have
also addressed it by suppressor screening in a ypt6 null mutant (paper II).
The retrograde transport toward Golgi is of importance for retrieving
membrane material and regulators, to be reused in anterograde traffic, e.x.
α-factor maturation enzymes and the Vps10 receptor [39]. Retrieval from
the endosome requires as yet unknown components of a multimeric coat
complex [99, 171].
In a genetic screen for mutations that cause growth defects in combination with a mutant clathrin heavy chain, Bensen et al. [16] found an inactive
allele of Ric1. It was also shown that the SNAREs Ykt6 and Sed5, involved in
transport to cis-Golgi (see above), were able to alleviate the effects of both
Ric1 and Ypt6 deficiencies [16]. The protein complex Ric1/Rgp1 serves as
GEF for Ypt6 at the Golgi [210], while Gyp6 is a specific GAP [250]. It was
finally confirmed that Ypt6 is involved in retrograde Golgi transport using
also a temperature sensitive Ypt6 allele [143].
12
Lipids in Saccharomyces cerevisiae
Yeast lipidomics is a comparatively new research field, offering the advantages of yeast as a genetics model to explore the pathways and regulation
of lipid synthesis [68]. Thousands of lipid species exist in mammals [50]; in
contrast, the yeast lipidome is simpler and the existing collections of yeast
mutants allow addressing directly the effects of lipid metabolism perturbations.
Lipids in budding yeast include several derivatives of diacylglycerol, sphingolipids, and ergosterol, summarized in table 2. The major phospholipids
are represented by PC, PI, PE and PS. The acyl chains of diacylglycerol containing lipids are most commonly palmitoleic, palmitic, oleic and stearic
acid [200]. A few differences between yeast and animals should be noted:
yeast sphingolipids contain inositol instead of choline, they lack polyunsaturated fatty acids, and the major sterol in yeast cells is ergosterol [68, 260].
Lipids in organelles
Lipid composition of various membranous organelles is different, due to
both the site of biosynthesis of the lipid species as well as the their uneven
distribution with regard to their signaling qualities (fig. 4). Sub-cellular fractionation has been used in order to determine the amounts and distribution of different lipids [200, 260, 261]. The major lipid in yeast as well as
in most eukaryotes is PC, other abundant lipid species are PE and PS (Table 2). A special place is reserved for phosphatidylinositol (PI), one of the
most abundant lipid species, since inositol metabolism has a tremendous
impact on the entire cell [68]. Phosphorylated variants of PI occur in much
lower amounts than other lipids, however they play a disproportionately
large role as signaling molecules determining the identity of organelles [15].
Apart from building up cellular membranes, lipids serve a storage function
in yeast, as triglycerides accumulate in lipid droplets [190].
Lipid composition of yeast is characterized by stability upon constant
conditions and the possibility to change with the environment [200, 241,
259, 261]
13
Figure 4: Relative composition and sites of lipid synthesis in eukaryotic organelles.
Standard lipid abbreviations are used, in addition: Ch - cholesterol, SM - sphingomyelin, BMP - bis-monoacylglycerol-phosphate, Erg - ergosterol, ISL - yeast inositol sphingolipid. Flags indicate location of signaling PIPs. After van Meer et al.
[241].
Synthesis and distribution
The biosynthesis of lipids in yeast cells occurs in several specialized compartments [164, 200, 261]. Ergosterol and ceramide synthesis are carried out
mainly in the ER, while phospholipid production is more spread out, with
PS being decarboxylated either in the mitochondria by Psd1, or in the Golgi
by Psd2 [122, 229, 261]. Other organelles with a role in lipid biogenesis are
the peroxisomes and lipid droplets.
Phosphatidic acid (PA) is the precursor of three different phospholipid
types, PI, PC and CL. The pathway of direct generation of PC by PA is known
as the Kennedy pathway [114]. Phosphatidylserine is made directly from
CDP-DAG and serine by the Cho1 PS-synthase [135]. Upon decarboxylation
by Psd1 in the mitochondria or by Psd2 in the Golgi, it forms PE, which in
turn can be methylated in the Golgi to yield PC via an alternative route to
the Kennedy pathway.
The distribution of lipids from their site of synthesis to various
organelles in achieved by diffusion through the cytosol, through transport
14
Table 2: Phospholipid contents of budding yeast sub-cellular fractions, adapted
from [259, 261].
% of total phospholipid
Subcellular fraction
PC
PE
PI
PS
CL
PA
Others
Plasma membrane
16.8
20.3
17.7
33.6
0.2
3.9
6.9
Secretory vesicles
35.0
22.3
19.1
12.9
0.7
1.2
8.8
Golgi
25.5
24.5
26.5
23.5
–
–
–
Vacuoles
46.5
19.4
18.3
4.4
1.6
2.1
7.7
Nucleus
44.6
26.9
15.1
5.9
<1.0
2.2
4.3
Peroxisomes
48.2
22.9
15.8
4.5
7.0
1.6
–
Light microsomes
51.3
33.4
7.5
6.6
0.4
0.2
0.5
Mitochondria
40.2
26.5
14.6
3.0
13.3
2.4
–
of membrane material along the secretory pathways, or by specific
proteins [211]. Cholesterol is capable of flipping between the leaflets of
one membrane and of diffusing between membranes as a single molecule
[14]. PITPs such as Sec14 transfer PI and PC molecules inside an internal
cavity [158]. Ceramide travels with the cytoplasmic protein CERT [85].
Sphingolipids and sterols leave the Golgi after segregating from the
phospholipids in detergent-resistant microdomains [239].
Lipids segregate not only among organelles, but also between the two
leaflets of the same membrane. The most well known case is PS, greatly
enriched in the inner leaflet of the plasma membrane. This asymmetry is
energy requiring and is maintained by the plasma membrane aminophospholipid translocase at the expense of ATP hydrolysis [206]. Finally, lateral
segregation of lipids into lipid rafts is yet another means of increasing membrane complexity with major functional implications [9].
Regulation of synthesis
On the level of metabolites, lipid biogenesis is strongly influenced by
the presence of inositol and choline in the media. When supplied with
inositol, cells produce significantly more PI and less PS [113]. This
response is related to the increase in transcription level of INO1, the gene
for inositol-3-phosphate synthase [49]. INO1 is co regulated together
with a set of genes with a common UASI NO element in their promoter
regions. Loss-of-function ino1 mutants are auxotrophic for inositol [41]. In
contrast, cells overexpressing INO1 excrete inositol in the growth medium.
15
It was recently shown that Opi1, a PA binding protein, is a repressor of
INO1 [140].
Phosphoinositides
The hydroxyl groups of the inositol ring (fig. 5) can be modified by covalent
addition of a phosphate group, catalysed by various kinases [33]. The phosphorylation derivatives of PI are collectively known as phosphoinositides
and have emerged in the last decades as major cellular regulators in eukaryotes. Through specific interaction with a number of protein domains, recently review by Lemmon [133], phosphoinositides have been established
as the most important lipid species in membrane-related processes.
Figure 5: Phosphatidylinositol. The hydroxyl positions on the ring are numbered D1 to D6. Common modifications
are phosphorylation at positions D3, D4
and D5.
Phosphatidylinositol-3-phosphate, PI(3)P, is generated from PI
by the Vps34 kinase [201] and is necessary for protein sorting from
late Golgi to the vacuole and for various forms of autophagy [116].
Phosphatidylinositol-4-phosphate, PI(4)P, is synthesized at the Golgi and
plasma membrane by the PI 4-kinases Stt4 and Pik1, and to a lesser extent
by Lsb6 [61, 84, 255]. Diphosphorylated phosphoinositides are represented
by PI(4,5)P2 , generated by MSS4 at the plasma membrane [256], and
PI(3,5)P2 , synthesized by Fab1 at the vacuole [252].
PI phosphorylation is reversible and therefore the presence of
phosphoinositides is dependent also on the activity of phosphatases.
S. cerevisiae has seven distinct phosphoinositide phosphatases, recently
reviewed by Strahl & Thorner [216]. There are three types of activities
associated with phosphoinositide phosphatases, carried out by the
Sac1-like domain [175], by enzymes removing the D5 phosphate [247], or
by YMR1 [221]. The three synaptojanin-like phosphatases Inp51, Inp52
and Inp53 have both a Sac1-like domain and an inositol-5-phosphatase
16
domain, and in general most phosphatases have partially overlapping
functions in the cell [216].
Interestingly, all yeast synaptojanin-like phosphatases possess a
poly-proline motif known to be the target of the GYF domain, as explained
below. However, no potential interactions in this regard have been
investigated so far, except for an in vitro binding assay with synthetic
peptides [119]. Therefore, a putative link between synaptojanin-like
phosphatases and GYF domain proteins remains to be elucidated.
Lipids and membrane traffic
The first strong link of lipid biogenesis and vesicular traffic was provided by
the PITP protein Sec14, necessary for vesicle export from the Golgi [1, 11,
158, 215]. It was shown that the presence of PI(4)P is necessary for secretion, and that the defect associated with lack of Sec14 could be relieved by
either deleting the Sac1 phosphatase or overexpressing the Pik1 kinase [83],
essentially two alternative means of up-regulating the amount of PI(4)P at
the Golgi membrane (see above). Another example of the need for specific
lipids for traffic is the requirement for PI(4,5)P2 at the plasma membrane for
endocytic internalization of ion channels and clathrin dynamics [218, 243,
262]. Similarly, depletion of PI(3)P from endosomal compartments caused
block of endosomal trafficking [59]. The likeliest mechanism by which PIP
depletion affects traffic is by removing the “landmarks” specific for different
organelles, thus preventing effector protein binding [15, 240].
In addition to the signaling role played by the minor lipid species, the
more abundant lipids are also important for various steps in traffic. For instance, PA, produced by the action of phospholipase D upon PC, has been
shown to promote fusion of secretory vesicles with the plasma membrane
[101]. PE was recently found to be required for vacuolar protein targeting
[165]. Another recent study followed the intracellular distribution of PS with
a fluorescent biosensor, to find it enriched in endosomal and lysosomal
compartments, with the potential to attract there fusogenic effectors [253].
Ceramide, produced by hydrolysis of sphingomyelin, was required for formation of exosomes, likely for its conical shape, inducing spontaneous negative curvature at the endosome membrane [228]. Several lines of evidence
implicate sterols in targeting to the vacuole or the plasma membrane [9,
189, 233]. In a more indirect fashion, cholesterol was also suggested to affect
cellular polarization through the formation of PI(4,5)P2 enriched lipid domains [74]. In the majority of the above examples, it is the biophysical properties of the lipid species involved that cause the observed effects, rather
than effectors recognizing the lipids.
17
Protein-lipid interactions
Membrane binding proteins
The current view of the lipid bilayer membrane is formed after decades of
studies, from the discovery that the lipids are arranged in two monolayers
[76], to the detailed molecular models in modern simulations [156]. The
fluid mosaic model of Singer and Nickolson [209] is an important milestone,
since it grouped for the first time the membrane associated proteins into
integral and peripheral [157].
Unlike proteins that span the lipid bilayer and therefore are an integral
part of it, many others are capable of peripheral association with membranes (fig 6). This makes possible transient interactions subject to regulation by modification of both lipid bilayer and protein.
Figure 6: The lipid bilayer with associated proteins. Typical strategies for membrane
attachment are illustrated. A, lipid anchors; B, insertion of an amphiphillic helix; C,
electrostatic interactions with anionic lipids; D, specific lipid head group recognition.
Mechanisms of membrane binding
Covalent lipid modifications
Lipidation is common for the Ras-related small GTPases [86]. Members of
the Rab family are post-translationally prenylated at the conserved CxC or
CC motif at their C-terminus [115]. In complex with an escort protein, the
19
Rabs are modified by a type II geranylgeranyltransferase by addition of two
geranylgeranyl group to two C-terminal cysteines in the CC or CXC motif
[56], a modification that is essential for their function. Another trafficking
regulator modified by lipidation is the SNARE Ykt6 [150].
Amphipathic helices
Several groups of proteins employ amphipathic helices for membrane association. The crystal structure of MurG, member of the large family of glycosyltranferases, suggested an amphipathic helix as a likely membrane association site [82]. A combined structural/biophysical study of analogous
helix from the Acholeplasma laidlawii MGS verified its membrane-binding
capacity [139]. Another example for controlled membrane association by
similar structure are the GTPases of the ARF family (as described previously,
yeast Sar1 and Arf1 are involved in control of vesicle budding). The mechanism of membrane insertion of the helix upon activation by GTP binding
has been elucidated with the help of existing crystal structures of several
ARF proteins (fig. 7).
Figure 7: Interswitch toggle mechanism for Arf activation (after Pasqualato et al.
[184]). Upon binding, the γ−phosphate of GTP displaces a conserved Asp residue,
causing a shift in the interswitch and making the amphipathic N-terminal helix
available for membrane association. Arf6-GDP (left) and Arf6-GTPγS (right) are
shown in the same orientation.
Electrostatic interactions
Positive charge clusters in proteins cause commonly membrane
association via electrostatic attraction to the negatively charged anionic
lipid heads. Well studied examples are the BAR-domain proteins and
protein kinase C (PKC). BAR was named after the proteins where it
20
originally was found (bin/Amphiphysin/Rvs161-167), which are needed
for the final stage of vesicle formation, scission of the membrane [66]. They
are characterized by a concave shape stabilizing membrane curvature
upon binding; the latter is dominated by electrostatic interactions with
negatively charged lipids such as PS and PI.
The C2 domain of PKC-α, also found in phospholipases, binds to PS containing membranes in the presence of Ca2+ [20]. Interestingly, a negatively
charged amino acid residue (Asp55) and the bridging role of Ca2+ are required for this binding.
Specific lipid head recognition
The first mechanism of binding described above requires no special lipids
in the bilayer, while the next two generally depend on any negative charge
in the membrane. However, a large number of protein possess specialized
structural motifs targeting specific lipid head groups, as recently reviewed
by Lemmon [133].
The first example is the C1 domain from PKC (mentioned earlier regarding its C2 domain), identified as binding phorbol esters and DAG (reviewed
by Colon-Gonzalez & Kazanietz [38]). It is a zinc-finger domain of about
50 amino acids. For efficient binding, it requires not only the specific head
group but also an acidic membrane environment [112].
Another example are PH domains, named after a region of homology to
pleckstrin, a substrate of PKC in platelets [90]. They bind to the PI(4,5)P2
lipid, but also to its free head-group, inositol-1,4,5-triphosphate [67, 134].
Following the solution of a crystal structure of substrate-bound PH domain
[58], more than 150 articles in the past 20 years describe different aspects
and occurrences of PH domains, thus underlining their importance as effectors of lipid second-messengers. The phosphoinositide binding site of
PH domains is characterized by the KXn(K/R)XR motif in which the basic
side chains form the interactions with phosphate groups [104].
FYVE domains (an acronym for Fab1, YOTB/ZK632.12, Vac1, and EEA1)
recognize PI(3)P through another type of zinc-finger and the RR/KHHCR
basic motif [213]. Unlike PH domains, FYVE prefer membrane inserted
PI(3)P to the free I(1,3)P2 head-group for binding [124]. Importantly, FYVE
domain binding efficiency is increased by dimerization, allowing for
binding of multiple PI(3)P targets in the membrane. In the case of the EEA1
protein, the dimerization is achieved by coiled-coil domains [53].
Sorting nexins, important in membrane trafficking [99, 172], have a type
of PI(3)P selective domain known as PX domain, reviewed by Seet & Hong
[203]. PROPPINs are β−propeller proteins that bind PI(3,5)P2 [152]. A final
example are the ENTH (and related ANTH) domains, found in the epsin
N-terminal homology region of clathrin adapter proteins [63]. They bind
PI(4,5)P2 , however with low affinity.
21
Lipid binding and vesicular traffic
All protein regulators of membrane trafficking require some mechanism
of interaction, or association, with the membrane. A few are integral
membrane proteins, most notably the SNAREs, which usually have a
single C-terminal membrane spanning domain (19 out of 22 in yeast,
plus one lipidated). The majority of the trafficking regulators though
are peripherally attached to the membrane, often transiently and in
equilibrium with a cytosolic pool. Some examples given previously
include the two GTPase families, ARF and RAB; their GEFs; their effectors,
multimolecular tethering complexes such as TRAPP, HOPS and CORVET;
sorting nexins and amphiphysin.
It should already have become clear that the lipids in the organellar
membranes play a substantial role in the peripheral attachment of
the above listed proteins. Thus, a combination of protein-lipid and
protein-protein interactions drives the correct assembly of trafficking
regulators at specific membrane sites. While this idea may seem
transparent nowadays, it is in fact relatively young, with one of the first
hints been published only a few years ago by Levine & Munro [136].
22
Membrane stress
The biophysical notion of membrane stress is associated with the
accumulated curvature stress as the formation of a bilayer constrains
the non-bilayer prone lipids within a plane. This is separate from the
physiological membrane stress described later.
Spontaneous curvature
The planar arrangement of amphipathic lipid molecules minimizes the surface of the polar/non-polar interface, thus maximizing the internal contacts in the aqueous phase, and is therefore energetically profitable. This
does however come at a price, since the shape that the lipid molecules assume within a bilayer does not always guarantee a tight arrangement. The
most abundant phospholipid in eukaryotes, PC, has a nearly cylindrical
shape, however other common lipids such as PE, CL, and some glycolipids,
resemble a cone1 and are therefore non-bilayer prone (fig. 8). This results
in a spontaneous tendency of the monolayers to curve, suppressed inside
the bilayer formed of two monolayers with opposite topology [93]. Thus,
the presence of non-bilayer lipids in the membrane causes accumulation
of curvature stress.
Figure 8: Lipid shapes and the corresponding spontaneous monolayer curvature
Membrane curvature is also related to the lateral pressure along the normal of the bilayer [93, 146, 223]. The term lateral pressure is used to describe the forces within the plane of the bilayer. These forces are different at
1 The term “inverted” cone has been used to describe the shape of lysolipids. Since the math-
ematical definition of a cone does not imply an orientation in space, this term is regarded
here as confusing and is therefore avoided.
23
different depths, leading to the characteristic pressure profiles of bilayers
[146, 223].
The intrinsic curvature and lateral pressure are being actively modified
by membrane proteins (recently reviewed by Ces & Mulet [34], McMahon
& Gallop [149], Zimmerberg & Kozlov [258]). One line of research has elucidated the control of membrane homeostasis in Acholeplasma laidlawii by
maintaining constant spontaneous curvature [180]. This regulation can be
achieved by controlling the activity of lipid synthesizing enzymes dependent on curvature [42, 244]. Recently, effects on membrane curvature by
lipid modifications have been shown to be relevant to eukaryotes as well, in
a study on yeast depleted from PC [25].
A certain amount of stored curvature stress is essential for membrane
function. This stress becomes effective in events of membrane fusion or fission, such as those occurring during cell division or membrane traffic. One
example is the membrane intercalation of an amphipathic helix by the Sar1
GTPase during vesicle formation at the ER, described previously [184]. This
process introduces locally positive curvature, which is essential for membrane budding [18, 130]. Reciprocally, after GTP hydrolysis and dissociation of Sar1, the membrane maintains a certain amount of curvature stress,
which is relieved upon fusion with the target membrane [6, 130].
The energetics of membrane binding and bending by proteins determine
their function as either modulators of membrane curvature of merely sensors [258]. When the energy of attachment of a protein (such as one with a
BAR domain) is high, it is sufficient to modify the shape of the membrane to
follow that of the protein surface. On the other hand, when the free energy
released by the interaction is low, the protein will only bind if the membrane already has a suitable curvature, thereby the association event will
be equivalent to sensing the membrane curvature. Thus, the role of such
peripheral proteins is determined by their affinity to specific membranes
relative to the rigidity of the membranes.
Other mechanisms by which proteins modify membrane curvature include pulling by cytoskeletal elements or motor proteins and oligomerization of asymmetric transmembrane proteins [149].
As proteins can shape membranes, the opposite is also true – membrane
curvature and lateral pressure profiles affect the binding, conformation and
activity of some proteins, reviewed by Marsh [146], van den Brink-van der
Laan et al. [237].
Physiological membrane stress
In the context of this work, membrane stress is broadly defined as any nonphysiological change in the biophysical properties of biological membranes
affecting the function of membrane-associated proteins. Possible causes
24
may include altered bilayer composition, the effects of small, often amphipathic molecules, or deviation from a normal protein-to-lipid ratio.
Examples in eukaryotes and prokaryotes
Examples of membrane stress caused by changed composition are found in
bacterial strains in which the lipid biosynthetic pathways have been disrupted [45, 91, 155]. Escherichia coli defective in PG synthesis grows slowly
and requires sucrose and Mg++ supplemented medium [155]. Strains of E.
coli with inactive pss gene lack the major membrane phospholipid PE and
are characterized by divalent cation dependence and slow growth [45]. In
addition, electron transfer and transmembrane transport functions are affected in these strains [22, 153]. Interestingly, similar defects were not observed in Bacillus subtilis lacking PE due to disruption of PS decarboxylase,
however glucosyldiacylglycerol levels increased two- to fourfold in this organism [147].
Engineering of lipid composition in eukaryotes is more challenging,
since multiple biosynthesis pathways often exist (see above). PE was found
to be essential for growth of yeast [19]. Strains substantially depleted for
PE demonstrated strong negative effects on amino acid transporters [194].
These effects have been attributed to defective protein secretion in the
complete absence of PE [177].
Phospholipid composition has direct implication for membrane protein
structure and topology. In agreement with the "positive-inside" rule [245],
the anionic phospholipids PG and CL were shown as determinants of transmembrane protein orientation in E. coli [238]. A zwitterionic lipid, PE, was
also necessary for the correct organization of the γ−aminobutyric acid permease [257].
Some small molecules also have the capacity to cause membrane stress.
Alcohols and detergents have an effect on lipid packing and the gel – liquid
crystalline phase transition temperature [248]. Trifluoroethanol and other
small alcohols destabilize the KcsA tetramer by inserting in the headgroup
region of the lipid bilayer, resulting in a decreased lateral pressure in the
acyl chain region [235, 236].
The antipsychotic drug chlorpromazine has been used to cause effects
similar to blocking of membrane traffic after inserting in the membrane
interface and subsequent "stretching" of the lipid bilayer [46]. The mechanism of action of chlorpromazine relied on binding to negatively charged
phospholipids and disrupting association of traffic-regulator proteins with
the membrane [44].
25
Importance of membrane stress for homeostasis in
unicellular organisms
As illustrated by the above examples, physiological membrane stress is
common to unicellular organisms. They are more likely to be exposed
to effects of small molecules or to rapid changes in their environment.
Therefore, it can be expected that different mechanisms may have evolved
to deal with this stress, similar to the those controlling channel opening by
mechanosensation [5]. One such possible mechanism is described in more
detail in the next chapter.
26
Translational control, RNA decay and stress
Cells have evolved numerous coordinated reactions to changes in the environment. Some of these reactions are on the level of single molecules,
where the activity of an enzyme or a transporter changes together with alterations of metabolite concentrations or transmembrane potentials. Other
reactions involve signaling cascades of multiple molecules, and result in orchestrated responses on a large scale. Here, I will briefly outline the set of
events that connect sensing of stimuli including starvation and membrane
stress, to a rapid stall in protein translation and potentially degradation of
mRNAs.
Translation initiation
Upon export from the cell nucleus, mRNA has already undergone extensive
processing, most notably it has been spliced to remove non-coding regions
(introns), a polyadenine tail has been added to its 3´terminus, and a cap
structure has been added at its 5’ end [132]. Messenger RNAs are complexed
with proteins that maintain their transcriptional competence, such as the
poly(A) binding protein Pab1 [197].
Figure 9: Translation initiation in eukaryotes - 48S complex (after Lehninger et al.
[132]). See text for details.
Translation initiation in eukaryotes begins by association of the
cap-binding protein complex eIF4F with the cap structure of mRNA. eIF4F
consists of the cap-binding protein eIF4E, the RNA helicase eIF4A, and
the scaffolding protein eIF4G [98]. The latter mediates the binding of
mRNA to the 43S pre-initiation complex by interacting with eIF3 (fig. 9).
After binding, the 40S ribosomal subunit scans the mRNA to find the first
27
AUG codon, followed by dissociation of the initiation factors, binding of
the 60S ribosomal subunit, and the beginning of the elongation phase of
translation [71, 98, 132].
One regulatory step of translation initiation involves the inhibition of
eIF4E-eIF4F complex formation by the 4E binding proteins (4E-BPs) [40,
71]. These proteins compete with eIF4G for binding to eIF4E which necessarily precedes eIF4E-eIF4F binding. Phosphorylation of 4E-BPs in response to growth factors and as a result of the TOR PI-kinases (see below)
decreases their affinity for eIF4E and thus promotes translation initiation
[71]. Two 4E-BP proteins in yeast are Caf20 and Eap1 [2, 40]. Eap1 in particular was identified by a far-Western blotting assay for eIF4E interactors and
was shown to be a second 4E-BP in S. cerevisiae; its disruption leads to temperature sensitive growth and partial resistance to the macrolide antibiotic
rapamycin [40].
RNA degradation
The three types of ribonucleic acid (RNA) in cells have the same overall
primary structure, but generally different sizes and functions. Transfer
RNAs are small molecules with specific secondary structures that serve
as adapters during protein synthesis in the ribosome. Ribosomal RNAs
are larger, and together with the ribosomal proteins fulfill structural and
catalytic roles. The most variable type of RNA are the messenger RNAs,
transcripts of protein-coding genes with sizes roughly corresponding to
the size of the proteins they encode [132].
As all molecules in the cell, RNAs are turned over via constant production of new species and destruction of old ones. Messenger RNAs are subject to special regulation, since they are directly responsible for the protein
repertoire that the cell can synthesize. It has been of particular interest to
find out what determines mRNA stability, and different factors have been
considered in the past [96, 198, 199]. Currently, there is good knowledge
on several pathways of mRNA degradation [151, 208]. The degradation of
RNAs carrying premature termination codons by the nonsense-mediated
decay, as well as the miRNA mediated RNA decay (both recently reviewed
by Shyu et al. [208]) are more specialized mechanisms outside the scope of
this work. Instead, I will focus below on the more general aspects of RNA
degradation.
The main exonuclease in yeast acting in the 5’→3’ direction is Xrn1
(Kem1), a conserved enzyme homologous to pacman in Drosophila
[107, 159, 166]. In the opposite direction, 3’→5’, degradation is catalysed by
the exosome, a large multiprotein complex [154]. However, since none of
these enzymes has endonuclease activity, to digest RNA both Xrn1 and the
exosome need access to decapped or de-adenylated mRNA respectively.
28
Figure 10: Pathways of mRNA degradation, adapted from Newbury [167], Parker &
Song [183].
Depending on the order of events during mRNA degradation, three major pathways exist [151, 167, 183] (fig. 10). One starts with removal of the
cap structure from mRNA, catalysed by the decapping enzymes Dcp1 and
Dcp2 [54, 126]. Decapping is also controlled by Dhh1 [60] and the cytoplasmic heptameric complex of LSm1-7 proteins [225]. Decapped RNA can be
digested 5’→3’ by Xrn1. Another possibility is to initially de-adenylate the
mRNA by adenylase complexes such as those containing Ccr4 and Pop2
[231, 232], which would render it a substrate for the exosome in the 3’→5’
direction. Finally, cleavage of mRNA by endonucleases would generate unprotected 5’ and 3’ ends and result in sensitivity of the fragments to at least
one of the above degradation pathways [167].
Processing bodies
The site of mRNA degradation in the cytoplasm was shown to be organized
in foci containing the respective enzymes and named processing bodies,
or P-bodies [207]. In addition to mRNAs, they contain the proteins Dcp1,
Dcp2, LSm1-7, Edc3, Scd6, Pat1, Dhh1, Ccr4, Pop2, Not1-5, Ski7, Puf3 and
Xrn1 [182, 207]. Importantly, P-bodies are dynamic structures that rapidly
change their composition and appearance upon conditions promoting
mRNA degradation [28, 207, 222]. In yeast, P-bodies have been shown to
form upon stimuli such as glucose limitations and amino acid starvation,
osmotic stress and ultraviolet light.
It has been demonstrated that defective translation initiation leads to increased de-adenylation and decapping in S. cerevisiae strains with mutant
eIF4G, eIF4E and eIF4A [202]. On the other hand, activation of decapping
by Dhh1 and Pat1 is required for repression of translation during glucose
deprivation [37]. Thus, a functional connection exists between translation
and degradation of mRNA. This connection is manifested also in the re29
Figure 11: mRNA shuttles between polysomes and P-bodies, adapted from Newbury [167].
lation between the corresponding structures (fig. 11), since mRNA moves
from the polysomal complexes into P-bodies upon stimulation [28]. However, movement in the opposite direction was also observed, meaning that
mRNA can be stored rather than irreversibly degraded in the P-body [28].
Very recently, polyadenylated mRNAs and the translation initiation regulators Pab1, eIF4E, and eIF4G were also detected inside processing bodies
[27].
A general view emerging from the interactions between translation initiation and mRNA decay is that the two processes are in competition for
engaging with the mRNA [182] (fig. 11). Extrinsic factors, such as different types of stress or growth stimuli, can shift the equilibrium towards either process. Intrinsic factors such as mRNA defects will trigger irreversible
degradation and fulfill the quality control function of the mRNA decay machinery.
Nutrient deprivation and the TOR signaling pathway
The signals used routinely in the laboratory to trigger P-body assembly
are different types of nutrient deprivation [207]. Owing to the interplay
between mRNA translation and decay as described above, an interrupted
translation initiation step is a likely event preceding P-body formation.
Therefore, the question stands how the signal for nutrient limitation is
transferred to the protein synthesis machinery in the cell. One well studied
route for that signaling goes through the target of rapamycin (TOR) kinases
[151, 219].
TOR protein kinases were first identified in S. cerevisiae as the targets of
the antifungal agent rapamycin. While addition of rapamycin inhibits protein synthesis and promotes autophagy and glycogen accumulation, the
TOR kinases have the opposite effect, i.e. stimulating growth, translation
and ribosome biogenesis, as recently reviewed by [195].
30
In S. cerevisiae, two complexes exist (TORC1 and TORC2) [141]. They involve some common components and the highly similar Tor1 and Tor2 kinases. TOR complexes often localize to cellular membranes, which is important for their signaling function [195].
Signaling through the Tor pathway is branched, with several targets [195,
219]. One outcome is stimulation of the dephosphorylation of eIF2α by
the Sit4 phosphatase [36]. Another is through phosphorylation of the 4EBP proteins which competitively inhibit translation initiation (see above).
Adding onto the complexity of connections between TOR signaling and intracellular membranes, a recent study showed a requirement for functional
vacuolar protein sorting complexes for efficient TOR signaling [263].
It is becoming evident that signals coming from the cell membrane
through the TOR pathway could converge upon regulation of mRNA
translation and stability. One important example is the work by Deloche
et al. [46] on attenuation of translation initiation in membrane traffic
mutants and in cells treated with chlorpromazine (see above). The
authors observed a rapid response to shifting to non-permissive
temperature of several mutants at different steps of membrane transport.
Polysomal complexes were disassembled within 5-10 minutes and a
peak of 80S ribosomal subunits was observed [46]. The kinetics of the
response indicated it was too fast to be a result of the earlier reported
transcriptional repression of rRNA, tRNA and ribosomal protein genes
in secretion defective cells [137, 169]. Eventually, it was shown that the
TOR pathway acts during membrane stress, since (i) a gcn2∆ mutant was
resistant to chlorpromazine treatment; (ii) eIF2α was increasingly more
phosphorylated starting at 5 minutes after chlorpromazine treatment; and
(iii) the deletion of the 4E-BP Eap1 resulted in only minor redistribution of
polysomes during chlorpromazine treatment.
31
The GYF domain
The GYF domain, a key feature in the proteins subject of this study, was
discovered ten years ago in the CD2BP2 protein as responsible for binding
the repeated motif PPPGHR from the T cell adhesion molecule CD2 [170].
A conserved consensus sequence, W X Y X 6−11GP F X 4 M X 2W X 3GY F , was
found in various eukaryotic proteomes [64]. Two structural variants of the
GYF domain, the CD2BP2-type and the SMY2-type, differ by the length of
the first loop and by the residue found at position 8 of the CD2BP2-type
GYF domain [64, 120].
Figure 12: Peptide ligand bound to GYF domain from Arabidopsis thaliana GYN4
(PDB 1l2z).
Structural features
The GYF domain is a small fold with an α−helix packed against a β−sheet
[120]. The proline residues of the ligand form a PPII type helix. Conserved
aromatic residues from the domain are responsible for contacts with the
ligand. The structural difference between the two types of GYF domains,
W/D at position 8, determines the major difference in substrate specificity,
a preference for a tryptophan after the prolines/glycine in CD2BP2-type domains, as opposed to a preference for an aliphatic residue and against tryptophan in SMY2-type domains [120]. Another minor structural difference
33
determined a bias against negative charge and for longer proline repeats
for ligands of SMY2-GYF as opposed to the same type MYR1-GYF [70].
GYF domain protein families
Both structural types of GYF domains are present in numerous eukaryotic
proteins, although their expansion is not as wide as that of similar domains
such as WW or EVH1 domains [10, 120]. Examples of the CD2BP2-type are
CD2BP2, the first described instance of a GYF-domain protein, involved in
the immune response [170], GYN4 from A. thaliana (fig. 12), and Lin1 from
budding yeast. Importantly, representatives of this type of GYF domain are
spliceosome-associated proteins, as shown by mass spectroscopy, and are
readily detectable in the nucleus by fluorescence [17, 118, 125].
The SMY2-type GYF domain is found in Myr1 and Smy2, as well as in
homologous proteins from other yeasts. Another studied, albeit with an
yet unknown function, group of proteins with a SMY2-type GYF includes
the mouse GIGYF1/2 proteins and their human homologs PERQ1/2 [72].
GIGYF1 was described as an interactor of the Grb10 protein, involved
in insulin-receptor signaling [72]. Interestingly, the corresponding
proline-rich sequence in the human Grb10 does not conform to the
sequence requirement for recognition by the GYF domain, which casts a
shadow of uncertainty over the relevance of the GIGYF-Grb10 interaction
in mice [120]. Very recently, human GIGYF2 was found in a study of
familial Parkinson Disease (PD) on 249 patients [127]. Seven different
GIGYF2 missense mutations resulting in single amino acid substitutions
were present in 12 unrelated PD patients (4.8%) and not in controls; the
authors concluded that GIGYF2 is likely a gene with a causal role in familial
PD [127].
Figure 13: Examples of GYF domain proteins
While the region of high identity between yeast and mammalian SMY2type GYF domain proteins is limited to the GYF domain itself, additional
features such as a coiled-coil domain are present in both Myr1 and Smy2,
34
as well as in GIGYF/PERQ (fig. 13). However, when considering the possibility of a potential distant relation between these proteins, it is worthwhile
to mention that the couple MYR1/SMY2 was shortlisted among 17 “core
yeast genes” with homologs in other fungal genomes, but without known
homologs in other organisms [100].
Binding partners
Yeast Lin1 was identified as interacting with the yeast cohesin complex [17].
Thereafter, it was assigned a role linking chromosome segregation and premRNA splicing [17]. An earlier study using mass spectroscopy had already
detected Lin1 in a complex with the U4/U6.U5 small nuclear ribonucleoproteins from S. cerevisiae [214]. Subsequently, a splicing related function
was suggested also for the homologous CD2BP2 through binding to the
SmB/B’ core splicing component [92, 118]. Eventually CD2BP2 was also
found as a constituent of the evolutionary conserved U4/U6.U5 complex
[125]. It is currently accepted that Lin1 and CD2BP2 are homologs which
both serve auxiliary functions in the formation but not in the activity of the
U4/U6.U5 complex, through interactions either with the PRP8 protein or
with SmB/B’ [17, 92, 118].
A substantial body of data acquired by using the yeast two-hybrid
system (Y2H) has associated SMY2 and MYR1 to mRNA splicing as well
[65, 242, 246]. Nevertheless, no functional implications for splicing have
been related to these genes so far. While mass spectroscopy of purified
cellular components is a generally accepted method of dissecting protein
complexes, Y2H studies are notoriously prone to giving false positives. The
nuclear localization of most splicing factors is also inconsistent with the
demonstrated cytosolic localizations of Myr1 and Smy2 [69, 70, 97, 103].
It is likely that binding of SMY2-type GYF domains to various splicing
proteins is merely due to a coincidence of their liberal ligand requirements
and artificially created opportunities (in vitro, Y2H), rather than a
meaningful functional interaction.
A different line of evidence connected MYR1 and SMY2 to cytoplasmlocalized degradation of mRNA. GYF domains from Smy2 and Myr1 were
shown to bind strongly to Eap1 [119], an inhibitor of translation initiation
[40] (see above). In addition, the capacity to bind a number of factors associated with mRNA decay was shown [70, 119]. Finally, colocalization with
P-bodies supported possible interactions of Myr1 and Smy2 with the mRNA
decay machinery (Paper I). A very recent study [35] describing a connection
of Myo2 to processing bodies may turn out to be highly relevant in this context, see below.
Apart from interactions with Eap1, Myr1 and Smy2 have been shown to
interact with other translation initiation factors and with polysomal com35
plexes, by tandem affinity purification [62, 121] and Y2H [105]. Considering
the tight functional relations existing between control of translation initiation and mRNA decay, it is tempting to speculate that GYF domain proteins
may be the long-sought structural link between the two processes.
Membrane traffic connections
Functional data on SMY2-type GYF domain proteins have revealed another
aspect of their cellular roles. Smy2 was initially identified and named as a
suppressor of the myo2-66 mutant [138]. Myo2 and Myo4 are the two yeast
type V myosin motors, and Myo2 is required for the polarized delivery of secretory vesicles by actin-based transport [29]. Thus, the genetic interaction
with Smy2 suggested for the first time that Smy2 may be related to vesicular traffic as well. This relation was only recently confirmed by the group of
Akihiko Nakano at RIKEN, who described Smy2 as a suppressor of a mutant
of the Sec24 component of COPII coats [97].
Myr1 was also linked to membrane traffic by the finding that it is synthetic lethal with Ric1, part of Ypt6 GEF (see above) [226]. Indeed, Ric1 is
synthetic lethal with another 146 genes, which reduces the statistical importance of this finding for Ric1, however Myr1 has only been reported for
a synthetic phenotype with Ric1. Our own discovery of Myr1 among the
ypt6∆ suppressors along with the subsequent finding that Myr1 can rescue
ric1∆ confirmed the relevance of this interaction (Paper II). In conclusion,
the independent findings that both Smy2 and Myr1 are linked to membrane
traffic are corroborating evidence that these interactions are physiologically
meaningful.
Relation to RNA metabolism
A common theme in GYF domain protein binding partners is the relation
to RNA metabolism. Both structural types of GYF domains present in yeast,
the CD2BP2-type (Lin1), and the SMY2-type (Myr1 and Smy2) seem to mediate interactions with pre-mRNA splicing and mRNA decay respectively.
Thus, an ancient cellular role for all GYF domain containing proteins may
well be the regulation of mRNA processing. Indeed, one of the major points
of this thesis is that Myr1 and Smy2 affect the fate of mRNA in response to
membrane stress caused by trafficking defects.
36
Integrating knowledge by computational
biology
“It is unworthy of excellent men to lose hours like
slaves in the labor of calculation which could
safely be relegated to machines.”
– Gottfried Leibnitz, 1685
When the Saccharomyces genome was sequenced twelve years ago, the
functions of only about 1000 gene products were known. Today, this
number is closer to five thousand. This is indeed a giant leap for the
research community. Yet, how big a step is it for the individual scientist?
If one is to read about the function of one yeast gene product per day, it
would take about 15 years to go over all genes. It can easily be argued that
it is only human to begin to forget by the tenth year what one has been
reading during the first. Understandably enough, human minds are like
“running averages”, similar to 19 amino acid hydrophobicity windows, and
what one used to know 15 years ago is a poor predictor of one’s current
knowledge.
How then is it possible to take advantage of the wealth of information
that modern sequencing techniques and detection methods and computer
frameworks and databases provide? The answer could come from Nature
itself: what every biologist learns is that Life is repetitive – proteins have the
same structural fold over and over, different organisms share identical biochemical pathways, ontogeny recapitulates phylogeny. There is therefore a
huge redundancy in the biological information.
Thus, one way to deal with the overwhelming amount of data is to reduce
the redundancy. In doing so, one should take advantage of repetitive trends
in order to decrease the effects of randomness; afterwords, remaining large
differences would represent different aspects of the problem in sight; and
eventually, small differences between very similar objects could simply be
attributed to sampling errors and disregarded.
Translated into numbers, this is in fact the nature of the computational
technique principal component analysis (PCA), developed by Karl Pearson
in 1901 [185].
I will briefly describe here two computational approaches used to make
predictions for the possible structure and interactions of GYF domain proteins.
37
Gene Ontologies
Gene ontologies (GO) are controlled vocabularies describing gene and
gene product attributes in a given organism [8]. The Gene Ontology
project is a collaborative attempt to provide consistent descriptions of
gene products in many different databases. It can be accessed online at
http://www.geneontology.org/.
One important feature of gene ontologies is that they are controlled. This
means that a description of a gene product or function by a GO term is guaranteed to have a strictly defined meaning, rather than being subject of the
linguistic idiosyncrasy of the person using it. Another important fact is that
the terms are hierarchical. In this way, a more general term can describe all
the gene products that are also described by more specific terms lower in
the hierarchy.
Currently three independent ontologies are maintained: molecular function, biological process, and cellular component. A gene product or groups
of products of interest will have different annotations in the three different
ontologies, and each one will represent a different way to look at the data.
An example usage of the GO Term Finder [26] tool at Princeton University2 is shown on figure 14. We have used a similar approach in Paper I.
Multivariate analysis of protein sequences
GOs allow one to automatically search for text annotations, however protein sequence is another type of data that is problematic. Most analyses of
protein sequences have been based on their statistical nature, by using the
frequencies at which a certain pattern (or a single residue) will appear in a
given context or will be exchanged for another pattern/residue [3, 95, 111].
The assumption is that features within the sequences that are indispensable for function will be less prone to mutations. While generally correct,
this assumption fails to use explicitly the knowledge about protein structure acquired by other than statistical means.
In a different approach, the chemical and physical properties of amino
acids were analysed and used to develop numerical descriptors of each
amino acid [94]. A total of 29 properties were subjected to PCA and yielded
three numerical values for each amino acid naturally occurring in proteins.
Subsequently, we also used the above scale to extend the analysis of GYFdomain ligands. Employing PLS partial least squares, another multivariate
technique similar to PCA, we searched for specialization of two different
proteins based on their binding preferences (Paper I).
In conclusion, currently biological research seems unthinkable without
a secure grasp on the computational methods of the new millennium. Ade2 http://go.princeton.edu/
38
Figure 14: Results from GO Term Finder analysis of ypt6∆ suppressors. A list of 11
genes (Paper II, table 3) was submitted to the GO Term Finder service at Princeton
University. The ontology “process” was searched for terms to which several of the
11 genes are co-annotated. A strict threshold (2e10−3 ) was used in order to keep
the resulting graph small. The statistical significance of finding the displayed terms
is indicated on the legend.
39
quate approaches to deal with the steadily growing amounts of information
should make it possible to face even the challenge of an explosive expansion of the known organism diversity due to exploration of novel environments [254].
40
Summary of papers
The GYF Domain of Myr1 and Smy2 (Paper I)
In the first paper serving as basis to this thesis, we addressed the conserved
GYF domain as the most prominent feature in Myr1 (Syh1) and Smy2. Extensive studies by Kofler & Freund had provided in vitro binding data for the
GYF domains from the yeast proteins [119], and identified 153 potentially
interacting proteins. Clearly, not all possible interactions would prove to be
relevant in vivo. One obvious obstacle to binding of two proteins with mutual affinity would be spatial separation. Therefore, to gain more knowledge
about the potential role of Myr1 and Smy2, we tried to answer the following
questions:
1. Where in the cell are Smy2 and Myr1 localized (in regard to 153 interactors).
2. Can we group the putative interactors based on a common process?
3. Are there any differences in specificity supporting separate roles for
Myr1 and Smy2?
To answer these questions, a robust statistical procedure was necessary
in order to eliminate the inherent noise that we detected in the binding
affinity dataset. We used a PLS projection method (see above, also Paper I)
and modeled the binding strength against the amino acid structure of the
peptides used to assay binding. In this way, we could verify the conclusions
of Kofler et al. [119] for a consensus recognition sequence, while at the same
time detecting subtle differences in the preferences of Smy2-GYF and Myr1GYF. These differences were correlated with homology structural models of
the two domains (Paper I, suppl. data).
After constructing a reliable model relating ligand peptide structure
to GYF domain affinity, we re-assembled sets of good interactors, and
screened these sets for a common function using the GO Term Finder.
Degradation of mRNA appeared as a process with significantly enriched
associated proteins among GYF domain ligands. This was a strong
indication that Myr1 and Smy2 might take part in the cytoplasmic
processing of mRNA.
To verify the feasibility of the latter, we inspected GFP tagged GYF domains as well as complete GYF proteins for colocalization with the sites
of mRNA degradation. Using RFP-tagged DCP2 [207] to label processing
bodies, we were able to detect foci in the cytoplasm of nutrient-stressed
41
cells, corresponding to P-bodies and also containing GYF domain GFP fusions. This was a strong indication that the interaction of GYF proteins with
mRNA decay factors and translation initiation inhibitors detected in vitro is
also possible in vivo, and therefore likely to play a physiological role during
stress.
MYR1 – a suppressor of YPT6 and RIC1 mutants
(Paper II)
“... no time to wallow in the mire...”
∼ The Doors, 1967
In this paper, we had the following aims:
1. Search for additional gene products interacting with Ypt6-regulated
membrane trafficking.
2. Characterize novel suppressors of ypt6∆ (Myr1).
To this end, we transformed a Ypt6 deficient strain with a high-copy genomic library and selected survivors at non-stringent restrictive conditions
(as compared to previous screens in ypt6∆ [230], we chose a lower screening temperature of 35◦ C). The identified suppressors could be tentatively
categorized in two groups: genes with relation to vesicular membrane traffic, and genes with no obvious link to traffic (Paper II, table 3). A general
tendency was that traffic-related suppressors allowed for better growth of
the mutant strain, even at more than 35◦ C, while unrelated genes could
often rescue growth only at the original screening temperature (unpublished observation). Importantly, among the suppressors were Pab1, the
polyadenylyl-binding protein of yeast [197], and Lsm4 and Lsm8, involved
in mRNA processing [225]. Together with Myr1, these proteins form a functional group of mRNA-related suppressors of ypt6∆. This finding alone is
sufficient to suggest that membrane stress caused by lack of Ypt6 can be
relieved by effects on mRNA metabolism.
We further proceeded to characterize Myr1 by a broad range of techniques. Most importantly, genetic experiments demonstrated that suppression of ypt6∆-affected trafficking steps was specific, since a ypt1t s mutant
could not be rescued, but a ric1∆ strain was rescued by high dosage Myr1.
Biochemical experiments provided data on the nature of the Myr1 protein,
useful for determining its mechanism of cellular action.
It should be strongly emphasized that, in a parallel study, Higashio et al.
[97] collected very similar biochemical data on the Myr1 homolog, Smy2. In
particular, limited protein solubility resulting in sedimentation, membrane
binding, and punctate cytoplasmic localization were shown [97]. Moreover,
the genetic data strongly linked Smy2 to a membrane trafficking step, however distinct from the one we associated with Myr1. Thus, a likely possibility
42
emerges that Myr1 and Smy2 have been specialized to maintain separate
roles at successive steps of membrane trafficking.
An implied aim of the study was to find support for alternative functions
of Ypt6, however in retrospect, a synthetic lethal screen would have been
a more appropriate technique for that purpose. Our findings provided instead additional support for the already known functions of Ypt6 as regulator of Golgi-associated membrane traffic [16, 143, 210]. Importantly, we
collected a number of suppressors unrelated to vesicular traffic, which supports the view that the pleiotropic effects of Ypt6 deletion are indeed different manifestations of membrane stress.
One aspect of this study deserving special appreciation are the interactions between Myr1 and nuclear pore biogenesis. With pore complexes being anchored by transmembrane proteins, and the nuclear envelope being
an extension of the ER, clearly connections to membrane traffic could be
found [163, 196]. However, there is yet another important analogy emerging from recent discoveries about the similarities between nucleoporins
and vesicle coat proteins [47, 48, 80, 81], suggesting that an ancient role
of porins could have been maintenance of membrane curvature at the nuclear pore. Thus, the effects on nuclear pore distribution observed by us
might relate to common evolutionary origins of nucleoporins and protein
coats, and involve interactions of nature similar to the originally identified
suppression of ypt6∆ (Paper II) and sec24-20 [97].
Myr1 as a membrane sensor – interactions with lipids
(Paper III)
In agreement with the in vitro binding data of Kofler et al. [119], we could
also show that Myr1 and Smy2 are capable to associate with the mRNA decay machinery under physiologically relevant conditions (Paper I). On the
other hand, in Paper II we had established firmly that Myr1 interacts with
cellular membrane trafficking, an observation that was corroborated by the
data of Higashio et al. [97] about Smy2 participation in COPII formation.
Both our findings and those of Higashio et al. [97] suggested that a potential function in trafficking is mediated by the capacity of Myr1 and Smy2 to
associate with membranes. Therefore, in Paper III we addressed the following questions:
1. Does Myr1 restore Ypt6 functionality in membrane traffic or is its mechanism of suppression indirect?
2. What structural regions in the Myr1 molecule are relevant to membrane
binding?
3. How do changes in the lipid environment affect binding?
4. Do membrane stress-induced changes regulate Myr1 binding?
43
We detected partial restoration of vacuolar morphology in ypt6∆ and
ric1∆ mutants upon Myr1 overproduction. However, localization of Myr1
was not consistent with a direct role at late Golgi membranes. On the other
hand, stress effects on the membrane interface caused by chlorpromazine
induced P-body formation, showing that membrane stress and mRNA
decay are interconnected. Therefore, rescue of growth via indirect
effects on the stress response was a likely mechanism for suppression of
membrane trafficking defects by Myr1. We focused further on the ability of
Myr1 to detect membrane alterations.
Myr1 structural domains, described in Paper II, were re-evaluated
using BLAST [4] searches, and domain boundaries were predicted based
on sequence conservation. Thus, we considered three regions of the
Myr1 molecule for binding studies. The protein fragments were cloned
together with a GFP reporter, expressed in E. coli, and purified by affinity
chromatography. Thereafter, pure proteins were used in binding studies
with reconstituted liposomes (fig. 15).
Figure 15: Binding assay. GFP tagged pure protein is mixed with reconstituted sucrose-loaded liposomes of defined composition/origin. After ultracentrifugation, both protein and lipid content of supernatant and pellet are measured by fluorescence. GFP is used as a label for the protein, while a fluorescent
lipid is included in the liposomes to monitor lipid concentration.
The binding of Myr1-coiled coil and Myr1-C-terminus demonstrated
that:
1. Binding affinities are in the hundred µM range for total lipid and µM
range for proteins, concentrations consistent with those inside yeast
cells;
2. Liposome composition affects the binding strength;
3. Cooperative effects may exist upon simultaneous binding of different
domains.
We further focused on the role of positive charge clusters for binding. To
this end, peptides corresponding to specific regions of the Myr1 molecule
were subcloned in the GFP reporter constructs. Assays with these new constructs demonstrated that a limited region of the Myr1 sequence character44
ized by clusters of positive residues was sufficient for binding to negatively
charged phospholipids.
In conclusion, weak and selective binding of Myr1 sequence segments
carrying positive clusters to liposomes supported a potential role as a sensor of membrane stress.
Lipid-engineered bacteria (Paper IV)
This work reports important observations on the effects of glycolipids on
bacterial cells. I will focus mostly on aspects of this work relevant to membrane stress.
Normally, E. coli lipid bilayers contain a 3/4 molar fraction of PE, and
less PG and CL, and the propensity of PE to form nonbilayer aggregates and
induce curvature stress is of importance for membrane function. A mutant
strain (AD93) lacking PE was earlier shown to be adversely affected in terms
of growth and division capacity, response to osmotic stress, and function of
membrane transporters [45]. An envelope stress was also evident, with several upregulated proteins. However, when supplemented with the Acholeplasma laidlawii gene for a lipid glycosyltransferase, monoglucosyldiacylglycerol synthase (alMGS), AD93 synthesized substantial amounts of glucosyldiacylglycerol (GlcDAG), correcting some of the defects caused by the
absence of PE [249, 251].
An important question emerging from the above observation is whether
it is the zero net charge of GlcDAG that serves to dilute the membrane composed entirely of negatively charged lipids in AD93 (PG and CL), or whether
instead the small head group of GlcDAG is more important to introduce
curvature stress in the membrane. This question is addressed in the current paper by introducing a second A. laidlawii gene coding for diglucosyldiacylglycerol synthase (DGS). The product of this enzyme is GlcGlcDAG,
which like PE and GlcDAG has zero net charge. However, due to its larger
head group, consisting of two sugar moieties, it is bilayer-prone instead. A
parallel is drawn also to regulation in thylakoid membranes by using genes
from Arabidopsis thaliana for the corresponding galactosyl lipid species,
due to analogies between bacteria and organellar lipid membranes (fig 16).
Introducing the foreign genes to an E. coli mutant resulted in production of the corresponding lipid species. The differences in lipid composition between the various lipid clones caused differences in bilayer surface
charge and curvature stress, associated with effects on different functions
in the cell. It was shown that the type of lipid headgroup is important, in
regard to bilayer-forming tendencies. While surface charge can be deduced
by the lipid composition alone, measuring curvature stress required in vitro
liposome reconstitution with the addition of fluorescent probes. Since lipid
head group size and charge can affect the lateral area occupied by a single
45
Figure 16: Biosynthetic pathways in lipid-engineered Escherichia coli.
lipid molecule in a bilayer, chain ordering was estimated from the pyrenyllipid fluorescence spectra.
The two monosugar-glycolipid strains (GlcDAG and GalDAG) revealed
the highest chain ordering at larger bilayer depths, and higher than
PE-containing (wt) and PE-lacking (AD93) clones. Furthermore, the bilayer
of the GalGalDAG strain was somewhat more leaky than the GlcDAG,
GalDAG and GlcGlcDAG strains. Changed lipid composition in the
glycolipid-synthesising strains affected also protein function, as shown by
LacY studies. While a diglucosyl lipid maintained the correct topology of
LacY, it failed to provide the curvature stress profile needed for its uphill
transport function.
In conclusion, this study showed the importance of lipid identity for
membrane function in vivo. In regard to this thesis, it exemplified how
altering lipid composition in bacteria had membrane stress effects with
impact on the function of proteins and associated processes, and can be
studied by a range of biophysical and biochemical techniques.
46
Discussion and perspectives
Considering what is known about the mechanism of different stages of
membrane vesicle formation, targeting and fusion, what could possibly be
the role of GYF domain proteins therein? Higashio et al. [97] suggested a
role in vesicle formation for Smy2, however they have no direct support
that Smy2 indeed participates in the process [97]. Our data indicate
involvement of Myr1 at a different trafficking event and in vesicle fusion
rather than formation. Importantly, both Smy2 and Myr1 are dispensable
for vesicular traffic. Moreover, deleting both genes has no detectable
growth phenotype. Therefore, it seems unlikely that Smy2 and/or Myr1
have a direct role in vesicle formation/traffic.
Figure 17: Potential involvement of GYF domain proteins in sensing membrane
stress. Control of mRNA decay through interactions with P-body components (A),
control of translation initiation through binding to Eap1 (B) or to Tif34/Tif35 (C).
Yet another concern is the role that the GYF domain in particular might
have in membrane-related dynamics. Many available data point towards
involvement of GYF domain proteins in mRNA processing or translational
control [62, 70, 105, 119, 121]. Indeed, Higashio et al. [97] showed that suppression of sec24-20 required a functional GYF domain of Smy2, however
47
how that relates to GYF domain mediated binding is unclear, since neither
of the COPII subunits possess the signature ligand sequence.
From studies of GYF domain proteins in yeast [69, 70, 97, 138], combined with focused studies on membrane stress [44, 46] or mRNA degradation [35], a model emerges for the sequential interactions involving GYF
domain proteins (fig. 17). According to this model, membrane alterations
are sensed by bilayer-associating regions of Myr1 and Smy2. This causes a
signal to be transferred, via the GYF domain, to the cytoplasmic stress response elements. The latter include mRNA processing bodies (containing
several components with GYF domain target sequences), Eap1 (4E-BP, inhibitor of translation initiation), or translation initiation factors Tif34/Tif35
(physically interacting with Myr1 [105, 121]).
How is the signal transferred from the sensor to the effectors? It seems
most relevant that peripheral membrane association, typical for both Myr1
and Smy2, would play a key role. In Paper III, we established that positive
residues in the Myr1 membrane associating segments bind to liposomes
with negative phospholipids, therefore the extent of acidification of membrane surfaces can serve as a regulator of Myr1 attachment. Stress effects
are associated with PI phosphorylation, as suggested by the detection of
the Vps34 phosphatidylinositol 3-kinase in a synthetic lethal screen with
Tor pathway signaling [263].
One major question about the outcome of GYF protein signaling that
future studies should address is the sign of the effect it has on cellular
metabolism. One possibility is that Myr1/Smy2 regulate metabolism
negatively, by stimulating mRNA decay and inhibiting translation. This is
consistent with the lethal effects of Myr1 overexpression (Paper II). On
the contrary, a positive effect on metabolism can explain the rescuing of
various mutants in which a stress response was triggered.
The matter becomes further complicated if we consider simultaneously
membrane binding (fig. 18) and the possibility of effects on metabolism in
either direction. Overproducing Myr1 (and, in a sense, also Smy2, although
gene dosage was only doubled in the experiments by Higashio et al. [97])
will initially increase the cytoplasmic pool of the protein; subsequently, a
presumably constant fraction will associate with membranes. What is of interest here is whether it is the membrane-bound or the soluble form of the
protein that mediates signaling. One assumption could therefore be that a
certain soluble cytosolic concentration is needed to promote protein synthesis; thus, induced lipid binding upon stress would be counteracted by
increased production of soluble protein during overexpression. Unfortunately, this fairly simple model does not account for the complete dispensability of GYF proteins under normal conditions.
Alternatively, a lipid bilayer-associated fraction of the protein could mediate translation inhibition and transcript inactivation. Increasing this fraction by stress and PI phosphorylation would trigger inhibition; however,
48
Figure 18: Effects of stress and GYF protein expression on membrane-bound and
soluble levels.
in conditions of overproduction the soluble fraction may become substantially large to cause sequestration of translation inhibitors or mRNA decay
factors, preventing them from entering functional complexes. This would
allow cells to resume growth despite the presence of a stress condition,
consistent with the poor growth of suppressed mutants. Lack of the GYF
domain proteins would not necessarily result in a phenotype, according to
this model.
For a full understanding of the role of GYF domain proteins in yeast, conditions need to be found where the presence or absence of the proteins in
physiological concentrations will produce detectable cellular phenotypes.
However, this may not be achievable in standard lab settings, since some
genes are known to function only in very specific environments.
As a final note, my findings about the yeast GYF domain proteins Myr1
and Smy2 placed them in the context of intracellular membrane trafficking and mRNA degradation. Recent discoveries on the cellular responses
to membrane stress provided clues for the connection between these processes. Membrane-binding studies additionally confirmed the feasibility of
this connection with the participation of Myr1.
49
Acknowledgments
As the years are rolling by, I have become indebted to so many people for
helping me achieve my goals. I can hardly list all of you, or the occasions on
which you saved the day - my most sincere thanks go to you all!
Åke, for giving me a chance in your group in a difficult time, for your 24/7
enthusiasm for Science and for your openness to new ideas.
Arunas, we go a long way back, dude! Thank you for teaching me lots about
yeast and lab-work, and about good life and good beverages, for putting out
the fire in our kitchen and for being a generous host in California.
My old-time Södertörn roommates, Fergal, Marco, Ida, Stefan, Alia, for good
company.
Marco and Alessia, for the good old times, and for teaching me what mushrooms to pick.
Fergal, for hospitality and interesting discussions, for proofreading and encouragement. For the sentence “We work hard, we party hard”, the best
mental remedy to the anxiety the day after a party...
Ivo and Yann, my gym buddies. I miss our profound musings about Life,
and Universe, and... err... what was that other thing?
Per Kylsten, for generous hospitality in your lab and home, and trust in me.
SH groupleaders Swoboda, Nygård, Lönn, Wright, Johansson, for good advice.
The members of my adoptive group, Malin, Tuulia, Maria R→K, Amélie, and
Hanna E, for accepting me as a part of the group, for helping me into the
lipid field, for putting up with my deadly pessimism and even deadlier attempts at jokes a la Jack Handey.
Amélie, for excellent collaboration, for helping me get started in DBB, and
for understanding how I feel about the North (and for all the duty-free gifts
proving it!).
Current group members Hanna E and Changrong, for carrying on the torch.
Current and past office roommates at DBB, Helen, Pedro, Tiago, Kajsa,
Hanna G, Tomas, Anders - thanks for the good company.
Everyone at DBB, for always being nice and helpful, and for throwing great
Xmas parties.
Stefan, Joe and Buck, for an unforgettable “big exam”. Also all scientists who
took time to evaluate my work in committees in the past.
Everyone who ever fulfilled my request for a strain or a plasmid – my deepest gratitude and respect for your unselfishness.
51
Thanks to our Bulgarian expatriate community, for sharing holiday celebrations and perspectives. Especially my old friends Oggi, Assen, Ruslan, Ivo,
Volodya, Miro.
Respect to Drs Koumanov, Papazyan, Grantcharova, and Grancharov, for
showing me how to nail a PhD in Sweden with style.
Thanks to my parents, for lighting my interest in Science, and for being
there for me across the hundreds miles.
My sister Elena, for a roof above my head in San Fran, and for letting me
drive your car despite my hopelessness.
My Tanya, for your love and support during these years. Thank you for caring, and for putting up with my uncanny ability to be always right... You are
the only person who understands the true significance of my work ;)
Kurt Vonnegut and Douglas Adams, for depicting the world the way I see it.
AEW Smitts for financial support.
“When I was younger,
so much younger than today
I never needed anybody’s help in any way
But now these days are gone...”
∼ The Beatles, 1965
52
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