University of Groningen
Yeast peroxisomesManivannan, Selvambigai
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1
Yeast peroxisomes: de novo formation and
maintenance
2
The studies presented in this thesis were performed in the research group Molecular Cell Biology
of the Groningen Biomolecular Sciences and Biotechnology Institute (GBB) of the University of
Groningen, The Netherlands.
3
Yeast peroxisomes
De novo formation and maintenance
PhD thesis
to obtain the degree of PhD at the University of Groningen on the authority of the
Rector Magnificus Prof. E. Sterken and in accordance with
the decision by the College of Deans.
This thesis will be defended in public on
Friday 2 May 2014 at 11.00 hours
by
Selvambigai Manivannan
born on 18 January 1986
in Velur, India
4
Supervisor
Prof. I. J. van der Klei
Assessment committee
Prof. M. Schrader
Prof. B. M. Bakker
Prof. D. J. Slotboom
5
Contents
Aim and Outline 13
Chapter 1: Introduction 17
Chapter 2: Lumenal peroxisomal protein aggregates are removed 57
by concerted fission and autophagy events
Chapter 3: Preperoxisomal vesicles can form in the absence 91
of Pex3
Chapter 4: Pre-peroxisome formation in Hansenula polymorpha 129
pex19 cells does not require the ER
Chapter 5: The Hansenula polymorpha WSC homolog is involved 157
in peroxisomal membrane shaping and organelle
partitioning
Summary 189
Samenvatting 197
6
Aim and Outline
Hallmark of eukaryotic cells is the presence of different membrane bound components
termed organelles, each comprised of a unique structure and function. Of these, peroxisomes are
important organelles that characteristically contain at least one hydrogen peroxide producing
oxidase together with catalase to remove the hydrogen peroxide by-product. Peroxisomes are
inducible in nature and dynamic which facilitates a rapid adaptation of their number and function
in relation to the metabolic needs of the cell. Indeed, the number of peroxisomes per cell is
carefully regulated. When required, they are induced but when redundant for growth, they may
be rapidly removed via selective autophagy (also termed pexophagy). Peroxisomes are the
primary sites of lipid metabolism of the cell. Presently, the major contentious issues in
peroxisome field are the role of the ER in peroxisome formation and sorting of peroxisomal
membrane proteins (PMPs). In yeast, peroxisomes are pre-dominantly formed by division of pre-
existing organelle. However, peroxisomes can also be formed from ER by de novo particularly in
PEX mutants which lacks peroxisomes such as Δpex3 and Δpex19 upon complementation with
their corresponding gene. Hitherto, the sorting and insertion pathway of PMPs is a conundrum.
Various models have been proposed ranging from the view that all PMPs travel via the ER to the
opposite that PMPs are directly post-translationally sorted. Also intermediate have been
proposed in that only few PMPs travel via the ER whereas others insert directly. This Thesis
project aims at further elucidating the molecular mechanisms behind peroxisome biosynthesis
and maintenance.
Chapter 1 contains a synopsis of the contemporary views on peroxisome formation, homeostasis
(biogenesis and degradation), and their implications in cellular aging, apoptosis and cell death.
7
Chapter 2 describes the relation that exists between peroxisome fission and degradation. Using
mutants of Saccharomyces cerevisiae and Hansenula polymorpha that are defective in
peroxisome fission (∆dnm1 and ∆pex11) we show that peroxisome degradation is inhibited, like
in the ∆atg1 control strain. Moreover, we demonstrated that, upon introduction of catalase
protein aggregates into peroxisomal lumen, these aberrant structures are rapidly removed from
the organelles by a concerted action of peroxisome fission and selective autophagy. Removal of
these aggregates was important to sustain cell health as their presence was associated with
distinct physiological disadvantages as it affected growth and caused enhanced levels of reactive
oxygen species. Removal required the normal peroxisome fission machinery as it was inhibited
in peroxisome fission mutants (∆dnm1 and ∆pex11). Taken together, these data demonstrate that
peroxisomal housekeeping mechanisms exist to remove unwanted or harmful components from
the organelles, thus keeping them vital and functional.
Chapter 3 deals with de novo peroxisome formation in a Hansenula polymorpha pex3 mutant
which occurs when the PEX3 gene is reintroduced in this mutant. The current view was that pex3
cell lack peroxisome membrane remnants and new peroxisomes were created using the ER as
membrane template. Our current data indicate that this view in not generally valid and requires
adaptation. This is based on our finding that pex3 cell do have peroxisomal membrane remnants
which are very unstable and subject to rapid degradation. However, subsequent deletion of
ATG1, a gene essential for autophagy, resulted in stabilization of these peroxisomal membrane
vesicles. Biochemical and microscopy studies revealed that these vesicles contained Pex8
together with the docking site proteins Pex13 and Pex14. Other PMPs (i.e. Pex10, Pex11, and
Pmp47) were unstable and localized to the cytosol. Finally, we showed that the Pex14-containing
structures were the target for peroxisome development after reintroduction of Pex3 in pex3 atg1
8
cells. Together, these findings suggest that i) peroxisomal membrane structures can be formed in
the absence of Pex3, and ii) alternative sorting mechanisms exist for Pex13 and Pex14, which
reach their target membrane independent of the Pex3/Pex19 docking machinery.
Chapter 4 is a continuation of chapter 3, as it aims to study de novo formation of peroxisomes in
a second mutant described to lack peroxisomal membranes, namely pex19. We show that also H.
polymorpha pex19 cells do contains cluster of peroxisomal vesicles, which are unstable but could
be stabilized by blocking autophagy (ATG1 deletion). Our data further showed that the
peroxisomal membrane proteins such as Pex3, Pex13, Pex14 and Pex8 are targeted to the
vesicular structures of pex19 atg1 cells, whereas the Pex10, Pex11, Pex26 and Pmp47 are not.
Finally our data revealed that upon complementation with PEX19, the vesicles present in pex19
cells acts as template for peroxisome formation and not the ER. Therefore, our findings suggest
that Pex19 is not essential for peroxisomal vesicles formation from ER and also for sorting of
Pex3.
The work in Chapter 5 is aimed to investigate the function of putative H. polymorpha orthologs
of the PMP22/MPV17 family of proteins (WSC, SYM1, and YOR292C). In silico analyses
revealed the presence of homologs in H. polymorpha. Fluorescence microscopy (FM), using
GFP fusions of these proteins revealed that under peroxisome inducing conditions, the woronin
sorting complex (Wsc) protein is localized to peroxisomal membrane whereas the Sym1 and
Yor292c levels were below the limit of detection by FM. WSC deletion cells (Δwsc) grew
normally on methanol and showed no alteration in peroxisome abundance. However on glucose,
a partial peroxisome segregation defect was observed during vegetative cell reproduction in
conjunction with aberrations in peroxisome morphology. Therefore, Wsc might be essential for
9
segregation of peroxisomes and shaping of the organelle membrane.
10
Chapter 1
Introduction
This chapter is an adapted and extended version of Manivannan S, Scheckhuber
CQ, Veenhuis M, van der Klei IJ (2012). The impact of peroxisomes on cellular
aging and death. Front Oncol. 2:50.
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Abstract
Peroxisomes are unique eukaryotic organelles, essential in man. Structurally, they are
characterized by a protein-rich matrix which is surrounded by a single membrane. In yeast,
peroxisomes primarily develop by fission of pre-existing ones but can also form de novo from
the endoplasmic reticulum (ER). They are highly dynamic; their number and functions varies
between organism and environmental conditions. Peroxisome homeostasis is regulated by a
delicate balance between peroxisome biogenesis and degradation. Various biochemical reactions
involving the production of reactive oxygen species (mostly hydrogen peroxide) are confined to
peroxisomes in order to prevent uncontrolled cellular damage by these potentially toxic
compounds. Additionally, peroxisomes may act as a crucial line of defense against ROS that are
produced elsewhere in the cell (e. g. release from mitochondria during oxidative
phosphorylation). In this contribution the current knowledge on the formation of peroxisomes,
their degradation via autophagy, maintenance of peroxisomal homeostasis and the role of these
fascinating organelles in ageing and cell death with a focus on studies performed in yeasts are
discussed.
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Introduction
Hallmark of eukaryotic cells is the existence of different membrane-bound
compartments, the organelles, which are specialized in various functions. Peroxisomes,
belonging to the organelle class of microbodies, are multifunctional compartments that were for
the first time biochemically characterized by Christian de Duve and Pierre Baudhuin (1966). The
morphology of peroxisomes is strikingly simple: a single membrane envelops a proteinaceous
matrix, which is devoid of DNA or protein synthesis machinery. Thus, all peroxisomal proteins
are nuclear encoded and synthesized in the cytosol. The size of the organelle ranges from 0.1-1
µm in diameter.
Peroxisomes are highly dynamic. Their morphology, abundance and function depends on
the species, the developmental stage and external stimuli (reviewed by Oku and Sakai, 2010;
Schrader et al., 2012). The predominant feature of peroxisomes in all eukaryotes is the presence
of H2O2-producing oxidases and the antioxidant enzyme catalase that detoxifies H2O2.
In yeast, peroxisomes are predominantly involved in the primary metabolism of unusual
carbon sources, such as oleic acid in Saccharomyces cerevisiae and methanol in methylotrophs.
In man, in addition to other vital functions, peroxisomes are involved in the α- and β-oxidation of
very long chain fatty acids, biosynthesis of ether phospholipids and bile acids (reviewed by
Wanders and Waterham, 2006). Moreover, peroxisomes also perform a number of species
specific functions as in the glyoxylate cycle in plants (Bernhard and Rouiller, 1967); Woronin
body biogenesis and inheritance in filamentous fungi N. crassa (Liu et al., 2008). Recently, also
novel, non-metabolic functions have been identified for mammalian peroxisomes, among which
are anti-viral innate immunity and anti-viral signaling (Dixit et al., 2010).
13
In this contribution we summarize the current knowledge on peroxisomes, with emphasis
on their formation and autophagic degradation. In addition, we describe the contribution of these
fascinating organelles to ageing and cell death processes.
Peroxisome biogenesis
Peroxisome formation involves a number of specific proteins called “peroxins”, which are
encoded by PEX genes. Most of the 34 PEX genes identified so far encode peroxins that are
involved in matrix protein import or membrane protein insertion. Others play a role in other
aspects of peroxisome biology such as the regulation of peroxisome number, peroxisome fission
and de novo peroxisome formation.
Matrix protein import
All peroxisomal matrix proteins are synthesized by free ribosomes in the cytosol and
post-translationally imported into peroxisomes. Sorting of matrix proteins depends on
peroxisomal targeting signals (PTSs). Most peroxisomal matrix proteins contain a PTS1
(typically containing a tripeptide with the consensus sequence S-A-C/K-R-H/I-L at the extreme
C-terminus), whereas a few have a PTS2 (consensus sequence R-K/L-V-I/H-Q/LA at the N-
terminus). These PTSs are recognized by the cytosolic receptors Pex5 and Pex7, respectively.
The peroxisomal matrix protein translocation machinery (translocon) involves docking
and RING-finger complexes, which needs to be assembled on the peroxisome membrane for
efficient import of matrix protein. Upon binding their cargo, the PTS receptors associate with a
docking complex at the peroxisomal membrane, which in yeast consists of Pex13, Pex14 and
14
Pex17 (Purdue and Lazarow, 2001; Gouveia et al., 2000; Gould and Collins, 2002). The release
of cargo in to the peroxisomal lumen is facilitated by further interaction with Pex8, a
peroxisomal protein located on the trans side of the membrane (Wang et al., 2003). Further, the
receptor disassociation and recycling is triggered by cascade of protein-protein interaction
events. The PTS1 receptor Pex5 is ubiquitinated by the components of RING finger complex
(Pex2, Pex10 and Pex12) together with Pex4 which possesses characteristics of E3 and E2
ubiquitin ligases, respectively (van der Klei et al., 1998; Williams et al., 2008; Platta et at.,
2009). Subsequently, the Pex5 is shuttled back to the cytosol by AAA (ATPase Associated with
various cellular Activities family) peroxins such as Pex1 and Pex6 which are associated with
peroxisome membrane via Pex15 in yeast (Platta et al., 2005). In comparison with PTS1
receptor, the studies concerning the recycling of PTS2 receptor is less, but evidences suggest that
possibly PTS2 receptor also shuttles between the cytosol and peroxisomal matrix by
ubiquitination (Leon et al., 2006; reviewed by Lazarow et al., 2006).
Sorting and insertion of PMPs
It is still debated whether peroxisomal membrane proteins (PMPs) are directly inserted
into the peroxisomal membrane upon their synthesis in the cytosol or traffick via the
endoplasmic reticulum (ER). The targeting signal of PMPs is called mPTS. However, no clear
consensus sequence of mPTSs is known yet. Two groups of PMPs have been classified, namely:
group I (Pex19-dependent), and group II PMPs (Pex19- independent). In group I PMPs, the
mPTS is recognized by the cytosolic mPTS receptor Pex19, which is recruited by Pex3 at the
peroxisomal membrane. The minor portion of PMPs belongs to group II whose mPTS are not
15
recognized by Pex19. Reports showed that peroxins such as Pex3 and Pex15 in yeast (Hoepfner
et al., 2005; Schuldiner et al., 2008) and Pex16 in mammals (Kim et al., 2006) are targeted to
peroxisomes by trafficking via ER.
Hence, Pex3 and Pex19 are pivotal for peroxisomal membrane protein import. However
the mechanism of PMPs insertion in the membrane is still unknown. Interestingly, recent data
suggest that Pex3 and Pex19 are not required for the insertion of PMPs into the peroxisomal
membrane, but instead for budding of Pex3-containing peroxisomal membrane vesicles from the
ER. According to this model, all PMPs first sort to the ER. Vesicles that bud off from the ER
fuse and subsequently mature into peroxisomes (van der Zand et al., 2010). Therefore, these
observations indicate that Pex19 seems to have distinct roles in assembling PMPs on
peroxisomal membrane.
Peroxisome proliferation by fission
The classical model for peroxisome proliferation is growth and division of pre-existing
organelles. According to this model, a nascent peroxisome grows by incorporating newly
synthesized peroxisomal membrane and matrix components. After attaining a matured size, it
asymmetrically divides, thereby forming a new small peroxisome, which subsequently follows
the same cycle (Fig. 1). According to this model, the peroxins Pex3p and Pex19p plays a major
role in membrane protein insertion. It is however unclear how lipids are transported to the
peroxisome in this model. The peroxisomal membrane predominantly comprises of
phospholipids, which are derived from the ER. Studies in Saccharomyces cerevisiae revealed as
16
major constituents ER derived phosphatidylcholine (48.2%), phosphatidylethanolamine (22.9%),
and phosphatidylinositol (15.8%). Notably, the organelles also contain a considerable amount of
cardiolipin (7%), which is synthesized in mitochondria (Zinser et al., 1991). For transport of
lipids from the ER and mitochondria to peroxisomes both vesicular and non-vesicular trafficking
pathways have been proposed (Raychaudhuri and Prinz, 2008). Others proposed a membrane
contact site that might exist between the ER and peroxisomes for non-vesicular lipid transfer.
Figure 1. Hypothetical model of peroxisome formation and degradation in yeast. De novo formation of
peroxisomes from the endoplasmic reticulum takes place in yeast cells which are devoid of pre-existing peroxisomes
(1). In wild-type yeast cells, the predominant mode of peroxisome proliferation occurs via fission of the pre-existing
organelles which involves Pex11p dependent elongation and DRP (Dynamin-related Protein) dependent scission of
the peroxisomal membrane. When peroxisomes are targeted for macropexophagy they are engulfed by
autophagosomes which ultimately fuse with the vacuolar membrane to deliver the organelle for degradation (2).
Peroxisomes can also be targeted for micropexophagy in which the protrusion of the vacuolar membrane engulfs the
organelle and they are subsequently degraded (3). MIPA: micropexophagy-specific membrane apparatus.
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Peroxisome fission occurs in three consecutive steps: organelle elongation, membrane
constriction and fission. The initial stage of peroxisome fission, organelle elongation, is mediated
by Pex11 (Koch et al., 2010). Studies in H. polymorpha showed that insertion of the N-terminal
amphipathic α-helix of Pex11 into the peroxisome membrane initiates membrane curvature that
causes organelle elongation (Opalinski et al., 2011). So far, it is unknown which proteins are
required for the constriction process. Dynamin-related proteins (DRPs), large GTPases, are
responsible for the final scission event. In S. cerevisiae Vps1 (at glucose-repressing condition)
and Dnm1 (at peroxisome-inducing conditions) are the DRPs involved in peroxisome fission
(Hoepfner et al., 2001; Koch et al., 2003; Kuravi et al., 2006), whereas in the yeast H.
polymorpha peroxisome fission entirely depends on Dnm1 (Nagotu et al., 2008). Recruitment of
Dnm1 to the peroxisomal membrane requires Fis1, a tail anchored protein. Interestingly, Fis1
and Dnm1 are also involved in mitochondrial fission. Similarly, mammalian Fis1 and Drp1 are
involved in both peroxisome and mitochondrial fission. Hence, peroxisomes and mitochondria
share key components of their fission machineries.
De novo biogenesis of peroxisomes from the ER
The observation that in mouse dendritic cells a membrane connection may exist between
the ER and peroxisomes, suggested that new peroxisomes may form from the ER (Geuze et al.,
2003). This mode of peroxisome formation was also proposed for yeast pex3 and pex19 mutants,
which are assumed to fully lack peroxisomal membranes. Upon functional complementation of
these mutants with the corresponding genes, new peroxisomes are formed de novo (Hohfeld et
al., 1991; Yan et al., 2005; Motley and Hettema, 2007). It has been proposed that in yeast pex3
mutants, re-introduced Pex3 is first sorted to the ER, from which new peroxisomes are formed.
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Based on this and other observations, it has been suggested that all PMPs first sort to the ER,
followed by exit from the ER, which depends on Pex3 and Pex19 (Hoepfner et al., 2005; Tabak
et al., 2008; Zipor et al., 2009). However, there is still a protracted dispute whether Pex3 and all
other PMPs invariably traffic via the ER to peroxisomes also in WT cells. In this regard it should
be noted that PMPs are never detected at the ER in WT cells under normal conditions, but solely
in pex3 or pex19 mutant strains (van der Zand et al., 2010).
A recent finding in S. cerevisae suggests that peroxisome formation from the ER involves
the budding of two biochemically distinct preperoxisomal vesicles each carrying half of the
matrix protein translocon machinery, from the ER. The fusion of these vesicles is mediated by
Pex1 and Pex6 and results in the formation of a functional translocon which enables the uptake
of peroxisomal matrix proteins from the cytosol (van der Zand et al., 2012). Hence, this data
suggests that peroxisomal contents are delivered from ER. But the factors involved in vesicles
formation at ER, nature of the vesicles and the mechanism of vesicles transport is still unclear.
In summary, de novo formation of peroxisomes so far observed only in the yeast mutants
which lacks peroxisomes (pex3 and pex19) and not in WT cells (Motley and Hettema et al.,
2007). However, the de novo formation may occur mostly in higher eukaryotes and it cannot be
excluded that in other species the formation of peroxisomes from the ER is the more prominent
process of peroxisome proliferation (Geuze et al., 2003; reviewed by Tabak et al., 2003; Kim et
al., 2006; Yonekawa et al., 2011).
Peroxisome inheritance in yeast
In yeast peroxisomes are carefully partitioned over mother and daughter cell during cell division.
Transport of peroxisomes from the mother cell to the bud is mediated in a Myo2 (an actin based
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motor protein)-dependent manner (Hoepfner et al., 2001). Myo2 binds to the peroxisome to be
transported to the bud, via the peroxisomal membrane protein Inp2. Surprisingly, recent studies
showed that Pex19 also plays a role in peroxisome inheritance by associating peroxisomes with
Myo2 (Otzen et al., 2012).
Inp1 is a peripheral membrane protein recruited to the peroxisomal membrane by Pex3
(Munck et al., 2009). The absence of Inp1 leads to complete depletion of peroxisomes in mother
cells, because no organelles are retained in the mother (Fagarasanu et al., 2005). Similarly, cells
lacking Inp2 fail to segregate peroxisomes to the bud, resulting in buds lacking peroxisomes. In
yeast, it has been proposed that peroxisomes are also formed de novo in mutant cells which lack
peroxisomes due to a segregation defect. Yeast cells lacking Inp2 develop new buds that are
initially devoid of peroxisomes, but at later stages peroxisomes are detected again (Motley and
Hettema, 2007). Similarly, in inp1 cells, new peroxisomes are formed de novo in mother cells
(Fagarasanu et al., 2005).
Peroxisome homeostasis
In order to establish a ‘healthy’ population of peroxisomes, cells continuously form new
peroxisomes whereas old or damaged/exhausted ones are degraded by autophagy. Peroxisomes
can be degraded by autophagy non-selectively or selectively. Selective degradation is termed as
“pexophagy” and mostly studied in the yeasts H. polymorpha, P. pastoris and S. cerevisiae.
In yeasts peroxisomes can be degraded by processes resembling micro- and macro-
autophagy. Micropexophagy involves the formation of finger-like protrusions by the vacuole to
20
engulf a cluster of peroxisomes that is subsequently degraded. At the same time, a double
membrane structure termed micropexophagy-specific membrane apparatus (MIPA) is assembled
at the peroxisome surface. This ultimately completes the sequestration process (reviewed by
Dunn et al., 2005). Macropexophagy involves the sequestration of individual peroxisomes from
the cytosol by the autophagosome and its subsequent fusion with the vacuole for degradation
(Veenhuis et al., 1978). In H. polymorpha micropexophagy is induced at nitrogen starvation
conditions and macropexophagy occurs when methanol-grown cells are shifted to glucose-
containing medium. Under the latter conditions, the key enzymes for methanol utilization are
repressed and the organelle becomes redundant for growth (Komduur et al., 2003; Monastyrska
et al., 2005a). In the yeast P. pastoris, the induction of micro- or macropexophagy depends on
the ATP levels in the cell (Ano et al., 2005).
Although autophagy has been studied for many years, it still remains unclear where
autophagosomes originate from. However, recent studies in mammalian cells showed that ER
(especially the ER-Golgi intermediate compartment) and mitochondria are likely the essential
components of autophagosome membrane formation at starvation-induced autophagy (Axe et al.,
2008; Hailey et al., 2010; Ge et al., 2013). Hence, ER and mitochondria may also significantly
contribute to pexophagosome formation.
In H. polymorpha it has been shown that peroxisomes are constitutively degraded at
normal growth conditions in order to prevent the accumulation of damaged components, which
can be detrimental to the cells (Aksam et al., 2007; van Zutphen et al., 2011). Studies in H.
polymorpha demonstrated that peroxisomes containing aberrant protein aggregates are removed
by asymmetric fission and selective autophagy in order to maintain a healthy organelle
21
population. However, the cells which are affected in fission (dnm1 and pex11) and selective
autophagy (atg11) failed to remove these aggregate containing peroxisomes, suggesting that
these mechanisms are essential for selectively removing the damaged organelles (Manivannan et
al., 2013). Timely rejuvenation of a cell’s organelle population is most likely essential for cell
viability. Currently, there are two different hypotheses for the factors that are capable to induce
degradation of dysfunctional peroxisomes. They may become degraded if i) the organelle loses
protein import competence or if ii) the intraperoxisomal redox balance is disturbed so that
excessive peroxisomal ROS production takes place (Monastyrska and Klionsky, 2006a; van
Zutphen et al., 2011; Manivannan et al., 2013).
Pexophagy-related genes in yeast
Both micro- and macro-pexophagy requires a number of specific proteins identified as autophagy
related (Atg) proteins (Xie and Klionsky, 2007). Atg1, a serine threonine kinase involved in
autophagy and Cytoplasm-to-vacuole targeting (Cvt) pathways (Straub et al., 1997), is also
essential for both micro- and macropexophagy. In the yeast Hansenula polymorpha, ATG1
deleted cells peroxisome degradation is blocked even at the conditions that induce peroxisome
degradation (Kommudar et al., 2003). In yeast, Atg11 is involved in selective pexophagy. It is
required for the cargo recognition and delivery to the PAS (phagopore assembly site; this is the
site where formation of the autophagosome membrane is initiated) via actin cables (Yorimitsu
and Klionsky, 2005; Monastyrska et al., 2006b). Studies in H. polymorpha showed that Atg8 is
involved in autophagosome formation during macropexophagy, because the absence of this
protein showed defects in the formation of pexophagosomes (Monastyrska et al., 2005b). Next to
22
Atg8, Atg25 has a role in the closure of the pexophagosome structure (Monastyrska et al.,
2005a).
Similar to the S. cerevisiae mitophagy receptor Atg32 (Kanki et al., 2009; Okamoto et al.,
2009), Atg36 was found to physically interact with both Atg8 and Atg11 (Motley et al., 2012).
Under pexophagy-inducing conditions, Atg8 and Atg11 are tightly bound to Atg36. This ternary
complex is supposed to mediate the irreversible uptake and subsequent breakdown of the
peroxisome into the vacuole lumen. The notion that Atg36 is a bona fide pexophagy receptor is
also supported by the experimental finding that overexpression of the gene encoding Atg36
strongly increases pexophagy (under conditions that induce pexophagy) (Motley et al., 2012).
The genome of methylotrophic yeasts encodes no sequence homolog of Atg36. However,
a protein with similar functions is present which is not found in baker’s yeast: Atg30 (Farre et
al., 2008). Atg30 and Atg36 share common features like their interaction with Pex3 and Atg11,
post-translational modification by phosphorylation under the conditions of nitrogen starvation
(Farre et al., 2013) and the induction of pexophagy when the Atg30/Atg36-encoding genes are
being overexpressed. It should be noted that there are also striking differences between
pexophagy in methylotrophic yeasts and baker’s yeast: (i) removal of Pex3 from peroxisomes is
a prerequisite for pexophagy in H. polymorpha (Veenhuis et al., 1996; Bellu et al., 2002; van
Zutphen et al., 2011) but not in baker’s yeast and (ii) Pex14 is required for directing Atg30 to
peroxisomes in P. pastoris (Farre et al., 2008) but not to those in S. cerevisiae (Motley et al.,
2012).
23
Peroxisomal damage prevention and repair mechanisms
The production and accumulation of ROS is a profound stress factor in living cells due to
the fact that ROS can oxidize and therefore damage vital macromolecules such as nucleic acids,
proteins and lipids. Intracellular accumulation of these damaged components leads to ageing - a
process defined as the deterioration of cells in time which is accompanied by gradual loss of cell
viability. Until recently, mitochondria were considered as the main players in ROS production
and hence in ageing of eukaryotic cells. However, recent reports proposed that peroxisomes also
produce significant amounts of ROS. Therefore, like mitochondrial dysfunction, peroxisomal
dysfunction also may contribute to ageing. The role of mitochondria in cell death pathways such
as apoptosis and necrosis is well established. However, the importance of peroxisomes in these
processes is much less understood.
Peroxisomal ROS and anti-oxidant enzymes
Peroxisomes counteract the compromising effects of ROS by anti-oxidant enzymes. Among
these are peroxisomal catalase, glutathione peroxidase, peroxiredoxin I and PMP20 to degrade
hydrogen peroxide and CuZnSOD and MnSOD to detoxify superoxide anions (Bonekamp et al.,
2009). S. cerevisiae contains two catalases, namely the peroxisomal Cta1p and the cytosolic
Ctt1p. Surprisingly, inactivation of both catalases in this yeast was shown to result in an increase
of the chronological lifespan (Mesquita et al., 2010). This unexpected finding could be explained
by the activation of superoxide dismutase by the elevated hydrogen peroxide levels. In this view
elevated levels of hydrogen peroxide in catalase-deficient S. cerevisiae cells prolong
chronological lifespan. However, earlier studies indicated that deletion of the gene encoding
peroxisomal catalase in S. cerevisiae resulted in a shortened lifespan (Petriv and Rachubinski,
2004). These seemingly contradictory results may be related to differences in the experimental
24
procedures to determine the lifespan. The same authors also reported that in Caenorhabditis
elegans the deletion of peroxisomal catalase (Ctl-2) in the genetic background of the long lived
Δclk-1 mutant shortens the maximum life-span as well. The shortened lifespan was accompanied
by altered peroxisome morphology that might point to compromised peroxisomal function with
increased production of reactive oxygen species (Petriv and Rachubinski, 2004).
In H. polymorpha, the role that peroxisomal catalase (Cat) exerts on ageing has been studied
recently (Kawalek et al., 2013). Deletion of the CAT gene has no detrimental effects on the
chronological lifespan (CLS) of cells grown in medium containing glucose (or glycerol) and
ammonium sulphate. Under these growth conditions, peroxisomal function is not obligatory. In
contrast, when cat cells were grown in medium containing glucose and methylamine (which is
oxidized by peroxisomal amine oxidase), their CLS was shortened significantly compared to the
wild type control. Methylamine was previously demonstrated to extend the CLS of wild type H.
polymorpha cells due to the generation of NADH from formaldehyde which is formed by its
oxidation (Kumar et al., 2012). The life span reduction in cat cells grown under
glucose/methylamine conditions is speculated to be based on consumption of NADH by
cytochrome-c peroxidase which (probably rather inefficiently) serves as a substitute for catalase
(Kawalek et al., 2013). Surprisingly, CLS of cat cells grown in glycerol/methanol/ammonium
sulphate-containing medium was found to be increased compared to wild type cells. It was
shown that the transcription factor Yap1 is up-regulated in the cat cells so that anti-oxidant
defense systems (i. e. cytochrome-c peroxidase and superoxide dismutase) are strongly induced.
In this regard, Δcat cells might benefit from a cellular hormesis response which ultimately results
in an increased CLS (Kawalek et al., 2013). Similar to catalase, the peroxisomal peroxiredoxin
Pmp20p is also involved in degradation of H2O2. Peroxiredoxins are thiol-specific evolutionary
25
conserved antioxidant enzymes. In addition to H2O2 breakdown they are also involved in
degradation of organic hydroperoxides (ROOH) and therefore important for maintaining the
integrity of lipid membranes. In the catalytic center of peroxiredoxins there is at least one
conserved cysteine residue that cycle between peroxide-dependent oxidation and thiol-dependent
reduction (Yamashita et al., 1999).
Studies in methylotrophic yeasts H. polymorpha and C. boidinii demonstrated that
Pmp20p is important for cell viability during growth on methanol due to its capability to repair
ROS generated damages (i. e., lipid peroxidation) at the peroxisomal membrane surface
(Horiguchi et al., 2001; Aksam et al., 2007). In the fission yeast Schizosaccharomyces pombe it
was shown that Pmp20p (in addition to thioredoxin peroxidase and glutathione peroxidase)
inhibited thermal aggregation of citrate synthase at a high temperature (43°C) in vitro. This
suggests that at least in S. pombe peroxisomal Pmp20p may have a second function as a
molecular chaperone which could be important for organelle quality control (Kim et al., 2010).
In human catalase gene mutations have been identified and linked to detrimental
conditions like diabetes, hypertension, macular degeneration, cataracts, cancer and skin
pigmentation disorders (Goth et al., 2004). A severe decrease of catalase activity is found in
clinical cases of hypo- or acatalasia and has been predominantly identified in Japan and certain
European countries (e. g., Hungary, Switzerland). Clinical symptoms are severe (among them
hemolytic anemia), illustrating the importance of functional catalase for health. Re-direction of
functional catalase to peroxisomes in catalase deficient cell lines led to increased detoxification
of H2O2 and also restoration of cellular plasmalogen and fatty acid levels (Sheikh et al., 1998).
ROS are usually associated with their potent damaging capabilities but they are also
involved in crucial cellular signaling processes. With regard to ageing, low levels of ROS might
26
induce a hormesis response which increases chances of cellular survival by up-regulating
pathways dedicated to high-stress adaptation like the retrograde response (Jazwinski, 2012) and
the TOR (target of rapamycin) and AMPK (adenosine mono phosphate kinase) pathways (Gems
and Partridge, 2008). Although it is clear that mitochondria are supposedly the main players in
this regard, it is likely that also peroxisomes integrate into the signaling network as important
mediators of aging processes.
Peroxisomal Lon protease
So far one conserved peroxisomal protease is known that plays a role in peroxisomal
protein quality control, namely the peroxisomal Lon protease, Pln (Kikuchi et al., 2004; Aksam
et al., 2007). Unexpectedly, S. cerevisiae and the closely related yeast Candida glabrata lack this
protein. Pln belongs to the AAA (ATPases Associated with diverse cellular Activities) protein
family which is supposed to be involved in degradation of unfolded and non-assembled
peroxisomal matrix proteins (Aksam et al., 2007; Ngo and Davies, 2007). Studies in H.
polymorpha showed that the deletion of the gene encoding Pln leads to a pronounced decrease in
cell viability accompanied by enhanced ROS production (Aksam et al., 2009). One possible
explanation could be that the accumulation of unfolded proteins leads to the formation of protein
aggregates, resulting in a disturbance of ROS homeostasis, which ultimately leads to cell death.
Indeed, electron dense aggregates are often observed in peroxisomes of H. polymorpha cells
lacking Pln (Aksam et al., 2007). Recently, it was show in P. chrysogenum deficiency of Pln
activity leads to formation of protein aggregates in the peroxisomal matrix and enhanced
oxidative stress (Bartoszewska et al., 2012). These findings suggest that Pln might also function
in quality control of peroxisomal matrix proteins. However, the precise molecular mechanism
27
and interplay between protein aggregation, ROS production and cell death still remains unclear
and needs to be studied in more detail.
Several studies have investigated the function of Pln in mammalian cells after the
identification of this enzyme in rat liver peroxisomes (Kikuchi et al., 2004). In mammals Pln also
seems to play a role in peroxisomal matrix protein import as catalase import is compromised
when Pln is overproduced (Omi et al., 2008). Recently it was also demonstrated that the protease
Tysnd1 (trypsin domain-containing 1, involved in the processing of several PTS1-containing
proteins and cleavage of N-terminal pre-sequences from PTS2-containing protein precursors)
and Pln cooperatively regulate essential peroxisomal functions like fatty acid β-oxidation in
mammals (Okumoto et al., 2011).
The role of peroxisomes in yeast ageing
Like mitochondria, peroxisomes can multiply by division of pre-existing ones (Fig. 1).
However, so far no evidence has been obtained that peroxisomes fuse. In two fungal model
systems for ageing, the filamentous ascomycete Podospora anserina and baker’s yeast it was
shown that down-regulation of mitochondrial fission by deletion of the DNM1 gene leads to a
robust increase in replicative life-span (Scheckhuber et al., 2007; Scheckhuber et al., 2008).
Moreover, deletion of DNM1 also has a positive effect on chronological ageing in baker’s yeast
(Palermo et al., 2007). These beneficial effects might be based on improved content mixing of
mitochondria so that molecular damage to proteins, lipids and mtDNA can be ameliorated more
efficiently. However, it has to be stressed that the effect of DNM1 deletion on peroxisome fission
was not investigated in these studies. Hence, the observed effects may also be partially due to
defects in peroxisomal fission. Several data suggest that peroxisomes divide asymmetrically,
28
resulting in larger mature organelles and small, nascent ones (Koch et al., 2003; Koch et al.,
2010; Cepinska et al., 2011; Huber et al., 2012). As a consequence cells contain a heterologous
population of peroxisomes, ranging from relatively young and vital nascence organelles to
relatively old, mature ones, in which dysfunctional components accumulate in time due to
damage caused by products of peroxisomal metabolism.
Mechanistic insights into the role of peroxisomes in aging have been gained mainly from
studies conducted in S. cerevisiae. When this yeast is grown in glucose rich media, neutral lipids
(e. g., triacylglycerols) are produced in the endoplasmic reticulum (ER) and incorporated in lipid
bodies (Olofsson et al., 2009).When these storage lipids need to be mobilized, peroxisomes and
lipid bodies come into close contact to allow uptake of the neutral lipids for subsequent oxidation
of the non-esterified fatty acids (free fatty acids) (Binns et al., 2006). An intriguing model has
been put forward linking life span determination and the synthesis and degradation of lipids in
the ER, lipid bodies and peroxisomes (Goldberg et al., 2009a; Goldberg et al., 2009b; Titorenko
and Terlecky, 2011). According to this model, repression of essential components of fatty acid β-
oxidation by ethanol (produced as a side-product of glucose fermentation) leads to the
accumulation of free fatty acids. This challenges the cell with detrimental effects, i.e. stimulation
of necrotic cell death (Jungwirth et al., 2008; Aksam et al., 2008) and lipoapoptosis (shown in
the fission yeast S. pombe) (Low et al., 2005). Moreover, as a result of impaired β-oxidation in
peroxisomes diacylglycerol accumulates in the ER and mediates the induction of protein kinase
C-dependent signaling which ultimately affects cellular pathways involved in various stress
responses (demonstrated in the nematode C. elegans) (Feng et al., 2007).
Peroxisomes and peroxisomal enzymes also play a vital role in the phenomenon of
retrograde response (RTG) in baker’s yeast (Butow and Avadhani, 2004; Jazwinski, 2012). RTG
29
is activated in yeast cells that are confronted with mitochondrial respiratory dysfunction.
Interestingly, induction of RTG leads to an increased replicative lifespan (Kirchman et al., 1999).
The more pronounced RTG induction is, the larger the beneficial effect on ageing. RTG leads to
the induction of transcription of several nuclear genes that help to promote the survival of the
cell despite mitochondrial dysfunction (Table 1). Genes encoding Cit1p, Aco1p, Idh1/2p (first
enzymes of the citric acid cycle), Ald4p and Acs1p (enzymes for cytosolic biosynthesis of
acetyl-CoA), Crc1p, Ctp1p, Dic1p and Odc2p (membrane transporters for shuttling metabolites
between mitochondria, peroxisomes and the cytosol) and Pex11p, Pxa1p, Cit2p, Fox1-3p
(peroxisomal proteins) belongs to this group. Collectively, these (and further) proteins enable
yeast cells to enhance oxidation of fatty acids and to synthesize essential metabolic intermediates
of the TCA cycle that otherwise would not be available.
The role of peroxisomes in ageing in mammals
Early studies on age-related changes of peroxisomal function were conducted on livers
isolated from mice (Perichon and Bourre, 1995; Perichon and Bourre, 1996). These studies
showed that the rate of β-oxidation in peroxisomes declines sharply by 40-70%, respectively.
However, similar work in rat did not show substantial changes in β-oxidation in young and old
animals (Cattley et al., 1991; Huber et al., 1991; Badr and Birnbaum, 2004). These virtually
contradicting results can be possibly explained by species differences or usage of different age
groups. Peroxisomal β-oxidation in hepatocytes is, of course, essential, because the example
subjection of rat to hyperinsulinemia lead to inhibition of β-oxidation and a concomitant
acceleration of ageing in these animals (Xu et al., 1995). Clearly, the activity of the peroxisomal
marker enzyme catalase decreases by 30 to 40% in liver samples isolated from old mice and rats
30
(Haining and Legan, 1973; Semsei et al., 1989; Perichon and Bourre, 1995; Xia et al., 1995;
Perichon and Bourre, 1996). Decreased levels/activity of this H2O2-decomposing protein is
contributing to the phenomenon of peroxisome senescence. This process leads to diminished
regulation of organelle maturation and division, protein import and overall dysfunction (Legakis
et al., 2002; Koepke et al., 2008). It has also been reported that the import efficiency of proteins
into peroxisomes via the Pex5p pathway decreases with age in fibroblasts. These cells also
produce more ROS due to an imbalance of peroxisomal pro- and antioxidants (Legakis et al.,
2002).
A comparative study with the goal to identify differences between liver subproteomes from
young and old mice revealed several upregulated peroxisomal proteins. One of the upregulated
enzymes was epoxide hydrolase 2, which detoxifies epoxides and converts these to excretable
dihydrodiols (Amelina et al., 2011). This may constitute a counteracting mechanism in old
animals. Another up-regulated enzyme, peroxisomal 3-ketoacyl-thiolase A, was correlated to
increased cholesterol levels in old animals (Amelina et al., 2011).
Recently, an inter-organelle crosstalk between mitochondria and peroxisomes regarding the
production of ROS was described (Ivashchenko et al., 2011). Enhanced formation of these
compounds in peroxisomes profoundly disturbs the redox balance within mitochondria, which
results in fragmentation of these organelles. This finding proves that peroxisome dysfunction can
have a pronounced effect on mitochondrial structure and function. It is thus conceivable that
certain scenarios of mitochondrial dysfunction involving elevated cellular ROS levels can also be
linked to peroxisomes.
31
Peroxisomes and cell death in yeast and mammals
Cellular death has many faces. Two of the most common ones, apoptosis and necrosis,
have been studied in much detail over the past few years (Golstein and Kroemer, 2007; Taylor et
al., 2008). Apoptosis is a ubiquitous mode of programmed cell death, strictly regulated and
conserved in eukaryotes (Madeo et al., 2004). This highly organized process is manifested by
condensation and cleavage of nuclear DNA, release of pro-apoptotic proteins from mitochondria
and, at later stages, “blebbing” of the plasma membrane. Mitochondria are key factors when it
comes to the execution of apoptosis. Mitochondrial fragmentation, depolarization and release of
various proteins that contribute to apoptosis are hallmarks of this process (Wang and Youle,
2009). Necrosis, on the other hand, is accompanied by rupture of organelles and the plasma
membrane. Recent evidence demonstrates that necrosis can be also tightly controlled, similar to
apoptosis (Golstein and Kroemer, 2007). Peroxisomes, seemingly not being important for
apoptosis induction and progression, are however involved in the regulation of necrosis. For
example, S. cerevisiae PEX6 deletion cells display hallmarks of necrosis and strongly elevated
formation of ROS (Jungwirth et al., 2008). Pex6p belongs to the family of AAA proteins and is
involved in import of proteins into the peroxisomal matrix (Ma et al., 2011). Moreover, in H.
polymorpha deletion of Pmp20 leads to pronounced induction of necrosis when the cells are
grown on methanol-containing media (Aksam et al., 2008). Interestingly, matrix proteins of the
peroxisomes were leaking into the cytosol of Pmp20 deletion cells during necrosis. This process
is reminiscent of peptide/protein release by mitochondria during apoptosis and constitutes an
interesting parallel between mitochondria and apoptosis on the one hand and peroxisomes and
necrosis on the other hand (Eisenberg et al., 2010).
32
Perspectives
The principles of peroxisome biogenesis and sorting/assembly of PMPs are still
controversial and intensely debated topics. At present, consensus has been reached only on the
peroxisome fission machinery. But not up to the extent that fission contributes to the total
organelle population per cell. This mechanism in fact may differ among species. However, in
case a rapid proliferation of the organelles is required, due to metabolic need, fission may be the
preferred machinery since the fission machinery is a much faster process relative to the de novo
formation pathway. De novo peroxisome formation requires extensive investigations; only two
proteins (Pex25 and Rho1) have yet been identified to be involved in this process. Recently it
has been clear with the snags in identifying proteins involved in de novo synthesis during earlier
genetic screens.
Saraya et al (2011) convincingly showed that H. polymorpha mutants affected in either
fission (pex11) or de novo synthesis (pex25) contain lowered numbers of peroxisomes and do not
exhibit a peroxisome-deficient phenotype. A complete lack of peroxisome was attained by the
combination of these two mutations, which simultaneously affects both fission and de novo
synthesis (as in a Pex25.Pex11 mutant). Consequently, an elegant approach is now available for
searching proteins involved in de novo synthesis by mutating a H. polymorpha pex11 strain and
select for peroxisome deficient mutants.
A second strongly debated topic is the routing of PMPs. Current views range from
routing of all PMPs via the ER to direct posttranslational sorting to the target organelle.
Considering the current literature, a plausible option is that both pathways exist. In PMPs
targeting information is observed for the ER (Thoms et al., 2012) but also signals for binding to
33
Pex19, the proposed PMP receptor those docks together with its cargo at the peroxisomal
membrane via Pex3. Hence, it can be envisaged that the two targeting signals have different
affinities for translocons in the ER and peroxisomes. Hence, if the affinity for the peroxisomal
one is stronger relative to the putative ER translocon, PMPs will travel to peroxisomes in WT
cells, but routed to the ER in cells lacking peroxisomes. This fits with the various observations
that Pex3-GFP can travel to the ER in pex3 mutants, but can directly sort to peroxisomes in WT
cells (Fujiki et al., 1984).
Research on the role of peroxisomes during cellular degeneration is supposed to unravel exciting
new mechanisms and will likely integrate these fascinating organelles into the network of
cellular pathways mediating ageing. Some of the interesting lines of research that will yield
promising insights might comprise (i) identification and characterization of a peroxisomal
unfolded protein response (UPR) as a mechanism to signal organelle dysfunction (and
subsequent degradation), (ii) studying selective inheritance of peroxisomes during replicative
ageing so that the daughter cells receive ‘healthy’ peroxisomes and (iii) investigating the role of
de novo formation of peroxisomes versus fission in maintaining a functional peroxisomal
population (see above) during chronological and replicative ageing. Recent reports in
mammalian system showed that the mitochondrial fission/fusion and mitophagy machineries are
dynamically coupled, thereby allowing a selective segregation of dysfunctional mitochondria
which are ultimately eliminated (Twig et al., 2008). Studies in yeast also revealed that mutants
deficient in autophagy accumulate dysfunctional mitochondria and are also characterized by
enhanced ROS production (Zhang et al., 2007). Hence autophagy plays a vital role in
maintaining cell viability and in the delay of cell aging. Possibly, similar mechanisms (except
peroxisomal fusion) might be involved acting on peroxisomes (Manivannan et al., 2013).
34
Peroxisomal quality control mechanisms and the impact of dysfunctional peroxisomes in cell
death and aging are relatively new and unexplored areas. Especially the signaling pathways
involved in the removal of dysfunctional peroxisomes are still unclear. Elucidating the role of
dysfunctional peroxisomes in cell viability and cell death may also provide new and important
insights into peroxisomal disorders. To achieve this goal it is relevant to reveal the mechanistic
details of recognition and removal of dysfunctional peroxisomes by auto(pexo)phagy.
35
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Name of gene S. cerevisiae H. polymorpha Mammals Description of protein Pathway
ACO1 + + + aconitase Krebs cycle/ RTG in S. cerevisiae
ACS1 + + + acetyl CoA synthetase acetate utilization/ RTG in S. cerevisiae
ALD4 + + + aldehyde dehydrogenase glucose fermentation/ RTG in S. cerevisiae
ATG30 - + - peroxisomal receptor pexophagy
CIT1 + + + citrate synthase 1 (mito.) Krebs cycle/ RTG in S. cerevisiae
CIT2 + - - citrate synthase 2 (peroxi.) RTG in S. cerevisiae
CRC1 + + - carnitine carrier fatty acid metabolism/ RTG in S. cerevisiae
CTA1 + + + peroxisomal catalase ROS detoxification
CTP1 + + + citrate transport protein mitochon. transporter/ RTG in S. cerevisiae
CTT1 + - - cytosolic catalase ROS detoxification
DIC1 + + + dicarboxylate carrier mitochon. transporter/ RTG in S. cerevisiae
DNM1/Drp1 + + + dynamin-related protein (DRP) 1 mitochondrial/ peroxisomal division
Ephx2 + + + epoxide hydrolase detoxification of epoxides
FIS1/hFis1 + + + binding partner for DRP 1 mitochondrial/ peroxisomal division
FOX1-2 + + + enzymes involved in β-oxidation of fatty acids
fatty acid oxidation/ RTG in S. cerevisiae
IDH1/2 + + + isocitrate dehydrogenase Krebs cycle/ RTG in S. cerevisiae
ODC2 + + + oxodicarboxylate carrier amino acid metabolism/ RTG in S. cerevisiae
PEX3 + + + peroxisomal membrane protein peroxisome biogenesis/ inheritance
PEX6 + + + AAA-peroxin recycling of peroxisomal signal receptor Pex5p
PEX11 + + + peroxisomal membrane protein peroxisome proliferation
PEX14 + + + peroxisomal membrane protein peroxisomal protein import
PLN - + + peroxisomal LON protease protein degradation
PMP20 - + + peroxisomal peroxiredoxin ROS detoxification
POT1/ Acaa1a + + + 3-ketoacyl-thiolase fatty acid oxidation
PXA1 + + + peroxisomal ABC transporter fatty acid transport/ RTG in S. cerevisiae
VPS1 + + - vacuolar protein sorting 1 vacuolar sorting/ peroxisomal division (S.c.)
Table 1: Overview of genes mentioned in this article. +: homolog present, -: homolog not present, RTG: retrograde response.
46
Chapter 2
Lumenal peroxisomal protein aggregates are removed by concerted
fission and autophagy events
Selvambigai Manivannan, Rinse de Boer, Marten Veenhuis and Ida J. van der Klei
Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB),
University of Groningen, Nijenborgh 7, 9747 AG Groningen, The Netherlands.
Published in Autophagy, 2013, 9(7):1044-56
47
Abstract
We demonstrate that in the yeast Hansenula polymorpha peroxisome fission and degradation are
coupled processes that are important to remove intra-organellar protein aggregates. Protein
aggregates were formed in peroxisomes upon synthesis of a mutant catalase variant. We show
that the introduction of these aggregates in the peroxisomal lumen had physiological
disadvantages as it affected growth and caused enhanced levels of reactive oxygen species.
Formation of the protein aggregates was followed by asymmetric peroxisome fission to separate
the aggregate from the mother organelle. Subsequently, these small, protein aggregate-containing
organelles were degraded by autophagy. In line with this observation we show that the
degradation of the protein aggregates was strongly reduced in dnm1 and pex11 cells in which
peroxisome fission is reduced. Moreover, this process was dependent on Atg1 and Atg11.
48
Introduction
Cellular homeostasis depends on a large array of quality control processes, which are important
for the recognition and removal or repair of cellular components. Detailed knowledge of such
processes is of major medical relevance as they are the molecular basis of devastating diseases
such as cancer,1 Alzheimer and Parkinson diseases.
2-4 Quality control processes include refolding
of unfolded polypeptides by molecular chaperones5 as well as proteolysis of individual proteins,
protein aggregates or cell organelles.6,7
A well-studied cytosolic proteolytic system is the ubiquitin-proteasome machinery for the
degradation of polypeptides. Endoplasmic-reticulum-associated protein degradation is
responsible for the export of aberrant proteins from the lumen of the endoplasmic reticulum
followed by cytosolic degradation by the proteasome.8 In both mitochondria and peroxisomes,
intra-organellar proteases have been identified that are crucial for the proteostasis of these
organelles.9
Autophagy is involved in the degradation of larger cellular compounds, such as protein
aggregates, ribosomes, pieces of cell organelles (e.g., piecemeal microautophagy of the
nucleus/micronucleophagy) or entire organelles.7 In yeast, autophagy is induced under starvation
conditions as a means to recycle unnecessary cellular components in order to allow the cell to
synthesize essential new ones.10
However, this process also serves as a cellular quality control
system to remove redundant, aberrant or toxic cell constituents.
Autophagic degradation of peroxisomes has been extensively studied in the yeast
Hansenula polymorpha. In this organism different peroxisome degradation processes can be
distinguished. The first one is glucose-induced selective macroautophagy (macropexophagy),
which serves to degrade peroxisomes containing enzymes that are redundant for growth. Second,
49
under nitrogen starvation conditions, peroxisomes are degraded by nonselective
microautophagy.11
Finally, constitutive autophagic degradation of peroxisomes has been
described and occurs during normal vegetative growth of the cells, likely as a mode to
continuously rejuvenate the organelle population.12
Consistent with this, an autophagic quality
control mechanism exists that most likely triggers the removal of dysfunctional or aberrant
peroxisomes.
To further analyze this phenomenon we stimulated the formation of aberrant peroxisomes
in H. polymorpha by introducing protein aggregates in the organelle matrix. This approach
recently became feasible as we observed that the production of a mutant variant of peroxisomal
catalase (containing a mutation in the heme binding site as well as the strong peroxisomal
targeting sequence –SKL) results in the formation of large intra-organellar protein aggregates.13
Protein aggregates are well known for being toxic in eukaryotic cells. Their accumulation often
causes the generation of reactive oxygen species (ROS), a major cause of ageing.14
Recently, a quality-control mechanism has been proposed in which mitochondrial fission,
fusion and selective autophagic degradation (mitophagy) cooperatively prevent the accumulation
of dysfunctional mitochondria.15,16
This poses the question as to whether comparable
mechanisms exist for peroxisomes to remove aberrant parts of the organelles. In turn, this
prompted us to investigate the putative role of peroxisome fission (these organelles do not
fuse17,18
and degradation in removing aberrant peroxisomes that contained lumenal protein
aggregates.
First, we show that the accumulation of these aggregates affects growth and results in enhanced
levels of ROS. Next, we demonstrate that peroxisome fission is important for both glucose-
induced pexophagy as well as for constitutive pexophagy. Finally, we show that peroxisomal
50
protein aggregates are removed from the organelles by a Dnm1- and Pex11-dependent
asymmetric peroxisome fission process, followed by degradation of the smaller aggregate-
containing organelles.
Results
Peroxisome fission is important for degradation.
To analyze whether peroxisome fission is important for glucose-induced selective peroxisome
degradation (macropexophagy), we analyzed this process in wild-type H. polymorpha as well as
in two mutant strains (dnm1 and pex11) that are strongly impaired in peroxisome fission.19,20
In
line with earlier observations, the levels of the peroxisomal marker protein alcohol oxidase (AO)
gradually decreased in the wild-type control upon induction of macropexophagy by glucose (Fig.
1A, B). However, in both dnm1 and pex11 cells, no significant reduction of AO protein levels
was observed. A similar result was obtained in the atg11 control strain, which is defective in
selective pexophagy.21
These results suggest that a reduction in peroxisome fission affects
glucose-induced macropexophagy.
To rule out that the observed block in pexophagy is not related to the fission defect in H.
polymorpha dnm1 or pex11 cells, but related to a direct function of fission proteins in
pexophagy, we performed a control study using Saccharomyces cerevisiae. Different from H.
polymorpha, in baker’s yeast Dnm1 and Vps1 have redundant functions in peroxisome fission.
Consequently, single dnm1 and vps1 mutants are only partially affected in peroxisome fission
and thus can be analyzed for a direct function of these proteins in pexophagy.22
As shown in Fig.
51
1C and D, single S. cerevisiae dnm1 and vps1 mutants were not blocked in glucose-induced
pexophagy, whereas cells of the dnm1 vps1 double mutant, which has a major peroxisome fission
Figure 1. Reduced peroxisome degradation in H. polymorpha and S. cerevisiae fission mutants. (A) Pexophagy
was induced by glucose in H. polymorpha cells grown for 20 h on methanol. Equal volumes of cultures were loaded
per lane. Western blots decorated with anti-alcohol oxidase (α-AO) antibodies show no significant reduction in AO
levels in the peroxisomal fission mutants dnm1 and pex11, similar to the atg11 control, which is blocked in
pexophagy, whereas AO levels gradually decreased in the wild-type control as expected. (B) Densitometry
quantification of the blots shown in (A). The amount of AO protein present at t = 0 h was set to 100%. The bar
represents the standard error of the mean (SEM). (** p<0.01). (C) Western blot analysis showing thiolase levels of
S. cerevisiae cells grown on oleate and subsequently diluted into SD(-N) medium to induce peroxisome degradation.
Samples were harvested at different time points, and equal amounts of protein were loaded per lane. There is no
significant decrease of thiolase in the dnm1∆ vps1∆ double mutant where the peroxisomal fission is completely
blocked. A similar result was obtained for the atg1 control strain. In the wild type and the single vps1 and dnm1
deletion strains degradation occurred. (D) Quantification of thiolase blots shown in (C). The level of thiolase protein
at t = 0 h was set to 100%. Levels were adjusted to the loading control glucose-6-phosphate dehydrogenase (not
shown). The bar represents the SEM (** p<0.01). (E) Constitutive peroxisome degradation is reduced in H.
polymorpha fission mutants. Cells were grown on methanol for 16 h. Western blots were prepared using crude
52
extracts and anti-GFP antibodies to detect Pmp47-GFP and GFP degradation products. The data show that the levels
of the cleaved fusion protein (lower band) are reduced in pex11 and dnm1 cells, relative to the wild-type control.
Equal concentrations of protein were loaded per lane. (F) Quantification of blots shown in (E). The levels of full-
length Pmp47-GFP proteins were arbitrarily set to 100%. The levels of cleaved GFP (lower band) are indicated as
percentage of the full-length fusion protein. The bar represents the SEM (* p<0.05; ** p<0.01).
defect, were impaired in peroxisome degradation. Using an H. polymorpha strain that produces
the peroxisomal membrane protein Pmp47 fused to green fluorescent protein (Pmp47-GFP), we
analyzed constitutive peroxisome degradation in methanol-grown cells of H. polymorpha wild
type and both dnm1 and pex11 mutant strains using western blot analysis and anti-GFP
antibodies (Fig. 1E). As expected, in extracts of wild-type cells in addition to the band
representing the full-length Pmp47-GFP fusion protein, a faster migrating band consisting of
cleaved GFP was also evident, indicative of constitutive pexophagy. The ratio of the amount of
cleaved GFP relative to the full-length fusion protein was reduced in dnm1 and pex11 cells
compared to wild-type controls (Fig. 1F), indicating that constitutive autophagy of peroxisomes
is also affected in H. polymorpha pex11 and dnm1 cells.23
Summarizing these data indicates that
in yeast peroxisome fission is required for pexophagy.
Intra-peroxisomal protein aggregates affect growth and cause oxidative stress.
Next, we addressed whether peroxisome fission acts in quality control of the organelles. For this
we took advantage of earlier observations that production of a mutant variant of catalase,
designated Catmut
, produced in wild-type H. polymorpha (i.e., also producing the endogenous
catalase protein) forms enzymatically inactive protein aggregates in the peroxisomal lumen.13
Peroxisomes containing these protein aggregates were used as a model for aberrant peroxisomes
in this study. First, we tested whether the presence of peroxisomal protein aggregates had
physiological consequences. As shown in Fig. 2A cultures of the Catmut
strain showed a reduced
53
yield. Remarkably, the specific catalase activities in cell extracts of cells of the Catmut
strain (185
U/mg) were enhanced relative to that of the wild-type control (130 U/mg). ROS measurements
indicated that at all time points examined the cells containing peroxisomal protein aggregates
had enhanced ROS levels relative to the wild-type control (Fig. 2B).
Figure 2. The effect of peroxisomal protein aggregates on growth and ROS levels. (A) Final optical densities of
wild-type, dnm1 and pex11 cells, producing or not producing Catmut
, upon growth on methanol as sole carbon source
for 40 h. Cell were extensively precultivated in glucose medium and subsequently shifted to medium containing
methanol. Final optical densities are expressed as adsorption at 660 nm. The bar represents the SEM (* p<0.05). (B)
ROS levels in cells at different time points after the shift of glucose-grown cells to methanol medium. The mean
intensity was measured by FACS. Catmut
cells show an enhanced ROS production relative to wild-type controls. The
bar represents the SEM (* p<0.05; ** p<0.01).
54
Intraperoxisomal protein aggregates are removed by fission and degradation.
To substantiate whether the protein aggregates induce peroxisome fission, we introduced Catmut
in the H. polymorpha atg1 mutant, in which autophagic degradation of peroxisomes is blocked.24
In order to visualize peroxisomes we introduced the fluorescent peroxisomal membrane marker
Pmp47-GFP. Fluorescence microscopy analyses of cells, cultivated for 16 h on methanol,
revealed that the Catmut
-producing atg1 strain contained, in addition to the normal organelles,
relatively small organelles that were not observed in the atg1 parental strain (Fig. 3A). This
result was confirmed by electron microscopy analysis of KMnO4-fixed cells (Fig. 3B) and also
showed the presence of aggregates in the small organelles (Fig. 3B). Subsequent analysis,
however, revealed that small organelles were not evident by fluorescence microscopy of wild-
type cells producing Catmut
(Fig. 3A), whereas electron microscopy revealed that these cells
harbored fewer small aggregate-containing peroxisomes relative to the atg1 strain producing
Catmut
. Apparently, the small separated organelles were rapidly removed in the wild-type
background. This process was further analyzed by electron microscopy. These analyses revealed
that once formed, aggregates migrated to the periphery of the organelle where they were
included in the buds that subsequently split off from the mother organelle (Fig. 3C).
To further address their degradation, we cultivated the H. polymorpha Catmut
strain on a mixture
of glycerol/methanol and subsequently administered 1 mM of the protease inhibitor
phenylmethanesulfonylfluoride (PMSF) to the culture to reduce the rate of degradation of
autophagic bodies in the vacuole. Electron microscopy analysis of these cells showed that at the
onset of the experiment vacuoles in Catmut
-producing cells contained very low numbers of
autophagic bodies (Fig. 4A, B). However, after 2 (Fig. 4C, D) and 4 h (Fig. 4E, F) of PMSF
administration the accumulation of aggregate containing organelles in the vacuole was evident.
55
Figure 3. Microscopy analysis of H. polymorpha atg1, atg1-Catmut
and wild-type- Catmut
cells. Cells were grown
on glycerol/methanol for 16 h. Peroxisomal membranes are marked by Pmp47-GFP. (A) Relative to the atg1
control, the atg1-Catmut
strain contains multiple small peroxisomes, which was not evident in wild-type-Catmut
cells.
The bar represents 1 µm. The peroxisomal phenotype was confirmed by electron microscopy (B). Note the presence
of protein aggregates in many of the organelles in atg1-Catmut
cells of which fewer are observed in wild-type-Catmut
cells. The bar represents 0.5 µm. (C) Ultrathin sections of KMnO4-fixed atg1-Catmut
cells showing different stages
of the formation of a small peroxisome containing a protein aggregate. The bar represents 0.2 µm. (D) Electron
micrographs, showing details of dnm1.atg1 cells (left panel) and pex11.atg1 cells (right panel) producing Catmut
,
demonstrating the presence of aggregates in the enlarged peroxisomes in these cells. The bar represents 0.2 µm. P,
peroxisomes; M, mitochondria; N, nucleus; V, vacuole.
56
These structures were not observed in the wild-type control. The presence of catalase protein was
confirmed by immunocytochemistry (Fig. 4G, H).
Figure 4. Catalase aggregates are degraded by autophagy. H. polymorpha wild-type and Catmut
cells were
cultivated on glycerol/methanol for 12 h and subsequently treated with 1 mM PMSF (t=0). Electron micrographs of
KMnO4-fixed Catmut
cells [at t=0 h (B)], [at t=2 h (D) and t=4 h (F)] revealed progressive accumulation of
autophagic bodies containing peroxisomes with protein aggregates that are not observed in wild-type cells (A, C, E).
Immunocytochemical localization of catalase shows that labeling is confined to peroxisomes of wild-type cells (G)
and is also abundant in the vacuole of Catmut
cells (H). M, mitochondria; N, nucleus; P, peroxisomes; V, vacuole.
The bar represents 0.5 µm.
57
These data suggest that peroxisomes with protein aggregates are delivered to the vacuole for
autophagic degradation. In conclusion, we showed that protein aggregates, which accumulated in
the peroxisomal lumen are removed by concerted fission and degradation events.
Degradation of aggregates is reduced in dnm1, pex11 and atg11 cells.
Clearly, the separation of small aggregate-containing peroxisomes is different from the fission
events that participate in normal peroxisome proliferation in wild-type cells. Therefore, we
analyzed if this fission process depends on Dnm1 and Pex11.19,20
To this end, the corresponding
deletion stains were constructed that produced Catmut
N-terminally fused to GFP (GFP-Catmut
) to
fluorescently mark the protein aggregates. Fluorescence microscopy analysis of cells also
producing the DsRed-SKL peroxisomal matrix marker protein revealed that small green
fluorescent spots were present in peroxisomes of both mutant strains, as well as in the wild-type
control (Fig. 5).
Figure 5. Visualization of catalase protein aggregates. H. polymorpha wild-type, dnm1 and pex11 cells,
producing DsRed-SKL and GFP-Catmut
, were grown on methanol for 16 h. The peroxisomes contained GFP spots in
the peroxisomal lumen, which represent the protein aggregates. The bar represents 1 µm.
58
To analyze the possible presence of GFP-Catmut
in the vacuole, the red vacuolar marker FM 4-64
was used instead of DsRed-SKL (Fig. 6). Fluorescence microscopy analysis confirmed that
green fluorescence was observed only infrequently in vacuoles of dnm1 and pex11 cells relative
to the wild-type control (Fig. 6).
Figure 6. Visualization of catalase protein aggregates. Identical experiment as shown in Fig. 5, using wild-type,
dnm1 and pex11 cells, producing GFP-Catmut
stained with the vacuolar marker dye FM 4-64. GFP fluorescence was
frequently observed in the vacuoles of wild-type cells. GFP fluorescence was also observed in the vacuoles of the
dnm1 and pex11 cells, albeit at reduced numbers compared to the wild-type control. The bar represents 1 µm.
Western blot analysis using crude extracts of these cells confirmed reduced cleavage of GFP-
Catmut
relative to that in the wild-type cells producing GFP-Catmut
(Fig. 7). The reduced
degradation of the aggregates also affected growth of the strains as lower growth yields on
methanol were observed in both dnm1 and pex11 cells for the strains producing Catmut
(Fig. 2A).
To strengthen the observation that Dnm1 and Pex11 are indeed required for the separation of the
small aggregate-containing peroxisomes, we also performed electron microscopy analysis of
59
Figure 7. Autophagic degradation of GFP-Catmut
is reduced in pex11 and dnm1 cells. (A) Western blot analysis
using crude extracts of cells described in Fig 5, decorated with anti-GFP antibodies. All strains contain both the full-
length GFP-Catmut
protein together with GFP (arrow), due to cleavage of the fusion protein. Pyruvate carboxylase
(Pyc1) was used as a loading control. (B) Quantification of the levels of GFP-Catmut
and GFP protein of the blots
shown in (A). The level of full-length GFP-Catmut
is set to 100%. The bar represents the SEM (** p<0.01). (C)
Quantification of GFP fluorescence in the vacuole. The percentage of cells containing GFP fluorescence in the
vacuole was calculated for wild-type, dnm1 and pex11 cells containing PAOXGFP-Catmut
. Per strain 2 samples of each
100 cells were counted.The bar represents the SEM (* p<0.05).
dnm1 atg1 and pex11 atg1 double mutant cells producing Catmut
. As evident from Fig. 3D, these
cells harbor large peroxisomes that contain protein aggregates. Degradation of the aggregates
was also reduced upon deletion of ATG11, a gene involved in selective peroxisome degradation.
In atg11 cells producing GFP-Catmut
numerous green spots were observed (Fig. 8A). However,
vacuoles did not accumulate GFP as observed in the wild-type background (compare Fig. 6 and
60
Fig. 8B). The block in autophagic degradation was furthermore evident from western blot
analysis of these cells showing that cleavage of GFP from the GFP-Catmut
fusion protein was
strongly reduced (Fig. 8C).
Figure 8. Autophagic degradation of GFP-Catmut
is reduced in atg11 cells. (A) In H. polymorpha atg11 cells
producing DsRed-SKL and GFP-Catmut
and grown on methanol for 16 h, most GFP-Catmut
spots colocalize with the
peroxisomal matrix marker DsRed-SKL. Some of the green spots do not colocalize with DsRed-SKL. Most likely
this is the result of the asymmetric peroxisome fission process, resulting in the formation of small aggregate-
containing peroxisomes that contain no or very little DsRed-SKL. (B) Vacuolar staining of atg11 GFP-Catmut
cells
with FM 4-64 dye. GFP fluorescence in the vacuoles is reduced in atg11 cells relative to the wild-type control (see
Fig. 6). The bar represents 1 µm. (C) Western blots decorated with anti-GFP antibodies fail to demonstrate the GFP
cleavage product (arrow) in atg11 cells producing GFP-Catmut
.
61
Discussion
Protein aggregates are harmful to cells and different machineries participate in the removal of
these structures from various cellular compartments. However, so far nothing was known on the
fate of protein aggregates in the peroxisomal matrix. In this study we present evidence that the
presence of protein aggregates in the peroxisomal matrix of H. polymorpha affects cell
performance and results in enhanced oxidative stress (Fig. 2). In order to remove these
aggregates from their lumen, peroxisomes undergo asymmetric fission, yielding small organelles
harboring these aggregates, which are degraded by pexophagy. The associated enhanced ROS
levels may be related to toxicity of the catalase protein aggregates, which could affect the
organelle’s redox homeostasis. The increased ROS levels may enhance expression of the
endogenous wild-type catalase gene, which is still present in the strains, explaining the higher
catalase activity observed in the cells producing the enzymatically inactive Catmut
.
Previous data show that pharmacological activation of macroautophagy contributes to a
decrease in aggregate-prone protein toxicity, such as that seen with mutant SNCA/α-synuclein
and HTT/huntingtin in cultured cells.25
Autophagy also plays a role in the degradation of
misfolded proteins, accumulated in the endoplasmic reticulum.26
Recent studies indicate that
aberrant organelles may also be subject to autophagic degradation.27
Hence, autophagy of protein
aggregates or cell organelles is very important to liberate eukaryotic cells of toxic and unwanted
components.
Similar to mitochondria, peroxisomes are major sites of ROS production. Oxidative
damage can result in protein unfolding and subsequent aggregate formation. Also, other
environmental conditions (e.g., heat shock) are likely to induce the formation of peroxisomal
protein aggregates. Although peroxisomes contain a Lon protease, which displays chaperone and
62
proteolytic activity, the capacity of this enzyme may be insufficient under certain conditions or
when high levels of aggregation-prone proteins are produced. Indeed, we recently showed that
the production of a mutant variant of peroxisomal catalase resulted in the formation of large
protein aggregates in peroxisomes.13
Also, we demonstrated that the absence of the peroxisomal
Lon protease in yeast and filamentous fungi9 results in the appearance of peroxisomal protein
aggregates.12
Hence, combining these data with our current findings, peroxisomal proteostasis
involves the concerted action of a chaperone/protease protein (Lon) for soluble proteins together
with autophagy for degradation of aggregates. This resembles the situation in the eukaryotic
cytosol, where molecular chaperones (e.g., Hsp70) together with the ubiquitin-proteasome
system and selective autophagy of aggregates (aggrephagy) combat protein aggregate
formation.28
Peroxisome fission is the major mode of peroxisome multiplication in yeast.17,20
So far,
this process was considered to be solely required to generate sufficient organelles to harbor
proper levels of peroxisomal enzymes to fulfill metabolic needs. We now show that peroxisomal
fission is also important for peroxisome degradation by pexophagy. First, we show that H.
polymorpha mutants (dnm1, pex11) defective in peroxisome fission show a major reduction in
glucose-induced pexophagy. Similarly, S. cerevisiae dnm1 vps1 cells, which are defective in
peroxisome fission, show reduced glucose-induced pexophagy. Second, we show that H.
polymorpha fission mutants display strongly reduced constitutive pexophagy. Hence, pexophagy
required peroxisome multiplication. Possibly, this is a mode to prevent the degradation of the
total peroxisome population within one cell, which would make the cell peroxisome deficient.
Indeed, in cells containing one peroxisome (in dnm1, pex11 cells and mpp1 cells)19,20,23
63
peroxisomes are not degraded. The residual pexophagy that is observed in dnm1 and pex11
mutants most likely is confined to the few cells containing more than one peroxisome.
An alternative explanation for the observation that the very large peroxisomes in dnm1 or
pex11 mutant cells are not degraded is that these organelles are too large to be enclosed within
autophagosomes. Interestingly, studies in Pichia pastoris indicate that indeed the components of
the autophagy machinery required for pexophagy differ for small and large organelles. Whereas
degradation of small peroxisomes only requires the core components of the autophagy machinery
(i.e., without pexophagy-specific Atg proteins), degradation of medium-sized organelles requires
Atg26, and large peroxisomes also need Atg11. Hence, the additional components required for
the larger organelles may be considered as an adaptation of the autophagy machinery.29
In this
explanation these adaptations, however, may not be sufficient to degrade the giant peroxisomes
in H. polymorpha dnm1 and pex11 cells.
For the removal of impaired regions of mitochondrial networks in mammalian cells,
organelle fragmentation caused by fission separates polarized and depolarized daughter
organelles, followed by selective autophagy of the depolarized ones.15,16
In this model,
mitochondrial fission is an important process in keeping the cellular mitochondrial population
healthy.30
However, whether this model is also true for yeast mitochondria is still debated. For S.
cerevisiae, data have been presented that mitophagy induced by nitrogen starvation31
or Mdm38
dysfunction32
depends on the function of Dnm1, a protein that is involved in mitochondrial
fission. Mendl and colleagues, however, suggest that there is no direct correlation between
mitochondrial fission and mitophagy,33
based on studies using mutant Dnm1 variants and
rapamycin induction. As is clear from the above, different modes of mitophagy induction have
been used in these studies, which may in part explain the virtually conflicting data.
64
Peroxisomes containing protein aggregates may also be considered as impaired cell
organelles. In line with the model proposed for mammalian mitochondria, we observed that upon
peroxisome fission aberrant, aggregate-containing daughter organelles are degraded. It is
tempting to speculate that these organelles are specifically degraded because they contain protein
aggregates. However, we cannot exclude the possibility that under the conditions studied the
autophagy machinery preferentially degrades smaller organelles, regardless of their aberrant
nature.29
However, previous studies in which glucose-induced pexophagy was studied in wild-
type H. polymorpha clearly indicate that under those conditions the larger peroxisomes are
preferentially degraded.34
This fission/degradation machinery is very effective judged from the observation that
aggregate-induced peroxisome fission is not paralleled by an increase in average peroxisome
numbers in wild-type cells, a phenomenon reinforced by the rapid accumulation of aggregate-
containing autophagic bodies in cells incubated in the presence of PMSF. As such, it stresses the
important role of autophagy in cellular housekeeping processes.
Acknowledgements.
We thank Dr. Tim van Zutphen, Dr. C.Q Scheckhuber, Dr. J.A.K.W. Kiel, Dr. Anita Kram and
A.M Krikken for skillful assistance in various parts of this study.
65
Materials and Methods
Organisms and growth.
The Hansenula polymorpha strains used in this study are listed in Table 1. Cells were grown at
37°C using either YPD (1% yeast extract, 1% peptone, 1% glucose) or mineral medium.35
For
analysis of peroxisome fission, cells were precultivated using mineral medium containing 0.25%
ammonium sulphate and 0.5% glucose as sole nitrogen and carbon sources, respectively.
Exponential cultures were subsequently shifted to media containing 0.5% methanol or a mixture
of 0.05% glycerol and 0.5% methanol as carbon sources. To induce pexophagy, cells were
precultivated in mineral medium containing 0.5% methanol and shifted to medium containing
0.5% glucose. When required, media were supplemented with 30 µg/ml leucine or appropriate
antibiotics. For cloning purposes, Escherichia coli DH5α was used as host for propagation of
plasmids using Luria broth supplemented with appropriate antibiotics (100 µg/ml) and grown at
37°C.
Saccharomyces cerevisiae strains used in this study are listed in Table 2. Cells were
grown at 30°C. Upon precultivation in mineral medium containing 0.25% ammonium sulphate
and 0.5% glucose, cells were shifted to mineral medium containing 0.1% oleate, 0.05% Tween
80, and 0.05% yeast extract. Whenever necessary, media were supplemented with leucine (30
mg/l), histidine (20 mg/l), uracil (30 mg/l) or lysine (30 mg/l). For peroxisome degradation
analysis, exponential oleate cultures were shifted to SD(-N) medium containing 0.17% yeast
nitrogen base without nitrogen and amino acids (YNB) (Boom, Y0626) supplemented with 2%
glucose.
66
Molecular and biochemical techniques.
Plasmids used in this study are listed in Table 3. Standard recombinant DNA techniques and
transformation of H. polymorpha was performed by electroporation as described previously.36
Cell extracts of trichloroacetic acid treated cells were prepared for sodium dodecyl sulfate
polyacrylamide gel electrophoresis as detailed previously.37
Equal volumes of the cultures were
loaded per lane, followed by western blot analysis.38
Blots were probed with rabbit polyclonal
antiserum against AO, thiolase or pyruvate carboxylase (Pyc1) or mouse monoclonal antiserum
against GFP (Santa Cruz Biotechnology, sc-9996). Blots were scanned by using a densitometer
(Biorad GS-710) and quantified using ImageJ (version 1.37); 3 measurements were performed
on each band of 3 individual blots per sample.
Preparation of crude cell extracts39
and catalase activity measurements40
were performed
as described previously. Protein concentrations were determined by the Bradford assay.
Construction of gateway plasmids.
For the construction of plasmid pSEM031, Gateway Technology (Invitrogen, 12537.023) was
used. A PCR fragment of 1587 bp of pHIPX9-Catmut
with Gateway sites was obtained by primers
CatFwB1 (GGGGACAAGTTTGTACAAAAAAGCAGGCTCGATGTCAAACCCCCCTGTTTTCAC)
& CatRvB2 (GGGGACCACTTTGTACAAGAAAGCTGGGTCGATTAAAGTTTGGATGGAGAAG)
on pHIPX9-CATmut
plasmid. The resulting PCR fragment was cloned into the standard Gateway
vector pDONOR-221 to obtain pENTR-221-Catmut
. Recombination of the entry vectors pENTR-
L4R1-PAOX GFP, pENTR-221- Catmut
and pENTR-23-TAMO and the destination vector pDEST-
R4-R3, resulted in plasmid pSEM031 (pDEST vector containing the PAOXGFP-Catmut
-TAMO
cassette). For stable integration of the plasmid into the H. polymorpha genome, the plasmid was
67
linearized with StuI in the PAOX region and transformed into H. polymorpha wild-type, atg11,
dnm1 and pex11 strains, which produce DsRed-SKL as a peroxisomal matrix marker. The
plasmid pHipZ4-DsRed-T1-SKL was linearized with KpnI and randomly integrated in the
genome of the above H. polymorpha strains.
The pDEST Pex11-HPH deletion cassette was obtained by the Gateway recombination of
plasmids pKVK106, pKVK107, pENTR-221-HPH and pDEST-R4-R3. From the resultant
plasmid, a 2.6 kb fragment of the pex11 deletion cassette was amplified by PCR with the primers
(PEX11-del3.1/PEX11-del3.2)19
and integrated in H. polymorpha wild-type and atg1 PMP47-
GFP cells. Similarly, the plasmid pSEM122 was made by the recombination of plasmids pENTR
DNM1 5′, pENTR DNM1 3′, pENTR-221-HPH and pDEST-R4-R3. A 2.4 kb fragment of the
DNM1 deletion cassette was amplified by PCR with the primers DNM1 del. fwd
(GCAGGACATTGTGACCAATACC) and DNM1 del. Rev (CCTCATATAGCAGCTCGTCGAA).
The amplified fragment was integrated in H. polymorpha wild-type and atg1 PMP47-GFP cells.
Fluorescence and electron microscopy.
Wide-field images were made using a Zeiss Axioscope fluorescence microscope (Carl Zeiss,
Sliedrecht, The Netherlands). Images were taken using a Princeton Instruments 1300Y digital
camera. The GFP signal was visualized by using a 470/40 nm bandpass excitation filter, a 495
nm dichromatic mirror and a 525/50 nm bandpass emission filter. DsRed fluorescence was
visualized with a 546/12 nm bandpass excitation filter, a 560 nm dichromatic mirror and a
575/640 nm bandpass emission filter. For vacuolar staining, 1 ml of cell culture was
supplemented with 1 µl FM 4-64 (Invitrogen, T-13320), incubated for 60 min at 37°C and
analyzed. For electron microscopy, whole cells were fixed in 1.5% KMnO4 for 20 min,
68
dehydrated in a graded ethanol series and embedded in Epon 812 (Serva, 21045). For
immunocytochemistry cells were fixed in 3% glutaraldehyde in 0.1 M cacodylate buffer pH 7.2
for 60 min and embedded in Unicryl (Aurion, 14660). Immunolabeling was performed on
ultrathin sections using specific polyclonal antisera and gold conjugated goat anti-rabbit
antiserum (Aurion, 815.011).41
ROS measurements.
Samples of methanol-grown H. polymorpha cells were harvested after 16 and 40 h of cultivation.
Approximately 5x106 cells were collected by centrifugation and subsequently resuspended in
250 µl of phosphate buffered saline (PBS) pH 7.4, containing 20 µM dihydrorhodamine
(Invitrogen, D-632). After 30 min incubation at room temperature in the dark, cells were washed
with PBS and resuspended in 500 µl of PBS for Fluorescence Activated Cell Sorting (FACS)
analysis using a BD FACS Aria TM
II.
69
Table 1. H. polymorpha strains used in this study.
Strains Characteristics Reference
Wild-type NCYC 495 leu1.1 42
Catmut
Strain containing pHIPX9-CATmut
This study
PAOX GFP-Catmut
Strain with pSEM031 This study
PMP47-GFP WT PPMP47 PMP47-GFP 43
PMP47-GFP Catmut
WT PPMP47 PMP47-GFP strain with pHIPX9-
CATmut
This study
DsRed-SKL GFP-Catmut
WT PAOX.DsRed-SKL integrated with pSEM031 This study
atg1 ATG1 deletion strain leu 1.1 24
atg1 Catmut
atg1 with pHIPX9-CATmut
This study
atg1 PMP47-GFP atg1 PPMP47-PMP47-GFP This study
atg1 PMP47-GFP Catmut
atg1 PPMP47-PMP47-GFP with pHIPX9-CATmut
This study
dnm1 DNM1 deletion strain, hygromycin This study
pex11 PEX11 deletion strain, hygromycin This study
dnm1 PMP47-GFP dnm1 PPMP47-PMP47-GFP This study
pex11 PMP47-GFP pex11 PPMP47-PMP47-GFP This study
dnm1 PMP47-GFP Catmut
dnm1 PPMP47-PMP47-GFP with pHIPX9-CATmut
This study
pex11 PMP47-GFP Catmut
pex11 PPMP47-PMP47-GFP with pHIPX9-CATmut
This study
dnm1 atg1 PPMP47-PMP47-GFP
Catmut
dnm1 atg1 PPMP47-PMP47-GFP with pHIPX9-
CATmut
This study
pex11 atg1 PPMP47-PMP47-
GFP Catmut
pex11 atg1 PPMP47-PMP47-GFP with pHIPX9-
CATmut
This study
atg11 NCYC 495 ATG11 deletion, URA3, leu1.1 21
atg11 GFP-catmut
atg11 strain with pSEM031
This study
atg11 DsRed-SKL GFP-Catmut
atg11 PAOX.DsRed-SKL strain with pSEM031 This study
dnm1 DNM1 deletion strain, ura3 20
dnm1 DsRed-SKL GFP-Catmut
dnm1 PAOX.DsRed-SKL strain with pSEM031 This study
dnm1 GFP-catmut
dnm1 strain with pSEM031 This study
pex11 PEX11 deletion leu1.1 19
pex11 DsRed-SKL GFP-Catmut
pex11 PAOX.DsRed-SKL strain with pSEM031 This study
pex11 GFP-Catmut
pex11 with pSEM031 This study
70
Table 2. S. cerevisiae strains used in this study.
Strains Characteristics Reference
BY4742 WT GFP-SKL BY4742 WT.URA3 PMET25 GFP.SKL 22
BY4742 dnm1 GFP-SKL BY4742 DNM1 deletion URA3 PMET25
GFP.SKL
22
BY4742 vps1 GFP-SKL BY4742 VPS1 deletion URA3 PMET25
GFP.SKL
22
BY4742 dnm1 vps1 GFP-SKL DNM1 VPS1 double deletion HIS3
URA3 PMET25 GFP.SKL
22
atg1 ATG1 deletion strain HIS3 URA3 LEU2 44
71
Table 3. Plasmids used in this study.
Plasmid Description Reference
pHIPX9–CAT mut
pHIPX9 containing PCAT CAT Y348G (point mutation in
the heme binding site of catalase) and the PTS1 –SKL
at the C terminus, kanR
13
pHipZ4-DsRed-T1-SKL Plasmid containing PAOX DsRed –SKL, ampR, zeo
R
20
pDONOR-221 Standard gateway vector Invitrogen
pDEST-R4-R3-NAT pDEST-R4-R3-containing nourseothricin marker, ampR
20
pENTR-23-TAMO pENTR-23 containing terminator of amine oxidase,
kanR
20
pHIPZ-PMP47-mGFP pHIPZ containing-PMP47-mGFP under the control of
PPMP47, zeoR, amp
R
43
pENTR-L4R1-PAOX-GFP pDONOR-P4-P1 containing GFP under the control of
PAOX, kanR
45
pENTR-221-Catmut
pDONOR-221 containing CAT mut
, kanR This study
pSEM031 pDEST-R4-R3-containing PAOX GFP- CAT mut
-TAMO,
nourseothricin marker, ampR
This study
pDONR-P4-P1R Standard Gateway vector Invitrogen
pDONR-P2-R3 Standard Gateway vector Invitrogen
pENTR-221-HPH pENTR-221 containing hygromycin marker, kanR
46
pENTR DNM1 5′ pDONR4 P4-1R containing 5′ region of DNM1, kanR
20
pENTR DNM1 3′ pDONR P2-P3 containing 3′ region of DNM1, kanR
20
pSEM122 pDESTR4-R3 containing DNM1 deletion cassette with
hygromycin marker, ampR
This study
pKVK106 pDONR4 P4-1R containing 5’-region of PEX11, kanR
19
pKVK107 pDONR P2-P3 containing 3’-region of PEX11, kanR
19
pDEST PEX11-HPH pDESTR4-R3 containing PEX11 deletion cassette with
hygromycin marker, ampR
This study
72
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76
Chapter 3
Preperoxisomal vesicles can form in the absence of Pex3
Kèvin Knoops1, Selvambigai Manivannan
1, Małgorzata N. Cepińska, Arjen M.
Krikken, Anita M. Kram, Marten Veenhuis and Ida J. van der Klei
Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB),
University of Groningen, Nijenborgh 7, 9747AG Groningen, The Netherlands
1These authors contributed equally to this work.
Article accepted in Journal of Cell Biology; December 2013.
77
Abstract
We demonstrate that Pex3 is not required for the formation of preperoxisomal vesicles, as yeast
pex3 cells already contain reticular and vesicular structures, which harbour key proteins of the
peroxisomal receptor docking complex, Pex13 and Pex14, as well as the matrix proteins Pex8
and alcohol oxidase. Other peroxisomal membrane proteins in these cells are unstable and
transiently localized to the cytosol (Pex10, Pmp47) or endoplasmic reticulum (Pex11). The
structures are more abundant in cells of a pex3 atg1 double deletion strain, as the absence of
Pex3 may render them susceptible for autophagic degradation, which is blocked in this double
mutant. Our data indicate, contrary to earlier suggestions, that peroxisomes are not formed de
novo from the ER when the PEX3 gene is re-introduced in pex3 cells. Instead, we find that re-
introduced Pex3 sorts to the preperoxisomal structures in pex3 cells, after which they mature into
normal peroxisomes.
78
Introduction
Peroxisomes are ubiquitous cell organelles that are involved in a large variety of metabolic
functions (Wanders and Waterham, 2006; Hu et al., 2012; Kohlwein et al., 2013). It is generally
accepted that peroxisomes proliferate by fission or form de novo from the endoplasmic reticulum
(ER). Although it is still debated which mechanism of organelle multiplication prevails in wild-
type (WT) cells, data obtained in yeast indicate that peroxisome fission is the most likely
mechanism of peroxisome proliferation in normal WT cells (Motley and Hettema, 2007; Nagotu
et al., 2008; Saraya et al., 2011).
In pex3 mutant cells, which are reported to lack peroxisomal membrane structures, new
organelles appear upon reintroduction of the PEX3 gene. A generally accepted view is that in
these cells the reintroduced Pex3 sorts to the endoplasmic reticulum (ER), followed by the
formation of pre-peroxisomal structures, which pinch off from the ER and develop into mature
peroxisomes. It has been suggested that all peroxisomal membrane proteins (PMPs) accumulate
at the ER in pex3 cells (van der Zand et al., 2010) and that upon reintroduction of Pex3, these
PMPs are incorporated in two types of pre-peroxisomal vesicles that fuse to form peroxisomes
(van der Zand et al., 2012). According to this model, Pex3 is important for the exit of PMPs from
the ER into pre-peroxisomal vesicles.
To date, relatively little is known about the molecular mechanisms involved in the reintroduction
of peroxisomes in pex3 cells. Here, we re-investigated this process focusing on the ultrastructure
of these cells and the subcellular localization of different PMPs prior to and after reintroduction
of Pex3 using a Hansenula polymorpha pex3 atg1 double deletion strain. The rationale for this
approach is that we have previously shown that removal of Pex3 from the peroxisomal
membrane is an essential early step in selective autophagic degradation of peroxisomes (Bellu et
79
al., 2002; Williams and van der Klei, 2013). This implies that the presence of Pex3 at the
peroxisomal membrane protects the organelles against autophagy. Hence, if peroxisomal
membrane structures develop in pex3 cells, they are likely to be rapidly degraded after their
formation. To prevent autophagy, we deleted ATG1, a gene essential for this process, in a H.
polymorpha pex3 strain. Our results show that pex3 atg1 cells contain preperoxisomal membrane
structures, which are the target for reintroduced Pex3 after which they mature into normal
peroxisomes.
Results and discussion
H. polymorpha pex3 atg1 cells contain vesicular structures that harbour PMPs
Careful electron microscopy (EM) analysis of H. polymorpha pex3 atg1 cells, grown at
peroxisome inducing conditions (mineral medium containing methanol and glycerol; MM-M/G),
revealed that these cells contain clusters of vesicular structures, which measure up to 70 nm in
diameter and have an electron dense content. These structures were not detected in WT control
cells (Fig. 1A). Immuno-EM (iEM) indicated that these structures contain Pex14, a PMP
involved in peroxisomal matrix protein import. The structures were generally observed in the
vicinity of the nuclear envelope, lateral ER and mitochondria (Fig. 1B). In support of our EM
results, mGFP- or mCherry-tagged Pex14 was observed as fluorescent spots adjacent to the
nuclear envelope, ER (Fig. 1C) or mitochondria (Fig. 1D). Electron tomography analysis
indicated that the clusters consist of reticular and vesicular structures (Fig. 1E). Distinct
connections with other cell organelles were not detected.
80
Figure 1. pex3 atg1 cells harbour Pex14-containing structures. (A) EM analysis of KMnO4-fixed pex3 atg1 and
WT cells grown for 16 h on MM-M/G. The inset shows a cluster of vesicles. (B) iEM analysis of pex3 atg1 cells
using α-Pex14 antibodies. (C,D) FM images of pex3 atg1 cells producing (C) Pex14-mCherry and the ER marker
BiPN30-eGFP-HDEL, or (D) Pex14-mGFP complemented with Mitotracker orange staining. (E) Electron
tomography analysis of (a) a serial-sectioned pex3 atg1 cell containing (b) a perinuclear membrane cluster (arrows).
(d-f) 10 nm thin digital slices through the tomogram reconstruction revealed vesicles (black arrows) and reticular
structures (red arrows). The surface-rendered reconstructions in (c) and (g) respectively show the viewing direction
of (d-g) and reticulo-vesicular structures in 3-D. Bars: (A) 500 nm, (Ainset,B) 100 nm, (C,D) 2.5 µm or (E) 250 nm.
CW – cell wall; M – mitochondrion; N – nucleus; P – peroxisome; V – vacuole.
81
The PMP containing structures in pex3 cells are susceptible to autophagic degradation
Although previous fluorescence microscopy (FM) studies suggested that, in H. polymorpha pex3
cells, Pex14-GFP is present in spots associated with mitochondria (Haan et al., 2006), iEM
revealed that these spots also represent clusters of vesicles, located adjacent to the nuclear
envelope, ER (not shown) or mitochondria at distances that cannot be resolved by FM (Fig. 2A).
The number of Pex14-mGFP spots is strongly reduced in pex3 cells, as was evident from
quantitative analysis of FM images (1.3 +/- 0.04 spots per cell in atg1 pex3 cells, relative to 0.6
+/- 0.04 in pex3 cells; Fig. 2B). In pex3 cells, but not in pex3 atg1 cells, mGFP fluorescence was
also observed in vacuoles (Fig. 2C), indicating autophagic degradation of the structures. This
was supported by western blot (WB) analysis, which revealed that the level of Pex14 was
strongly reduced in pex3 cells, compared to WT and pex3 atg1 cells (Fig. 2D).
Figure 2. pex3 atg1 cells contain enhanced numbers of Pex14-containing structures. (A) iEM analysis of pex3
cells using α-Pex14 antibodies, identifying structures (arrows) in the vicinity of mitochondria. CW – cell wall; M –
mitochondrion. (B) Quantification of Pex14-mGFP spots in pex3 and pex3 atg1 cells. (C) FM images of pex3 atg1
or pex3 cells, producing Pex14-mGFP complemented with FM4-64 vacuolar staining. The inset shows optimized
intensities for pex3 cells, highlighting the Pex14-mGFP spot and vacuolar mGFP. (D) WB analysis of cells grown
for 16 h on MM-M/G using α-Pex11 or α-Pex14 antibodies. Pyruvate carboxylase (Pyc1) was used as loading
control. Error bars: SEM. Scale bars: (A) 100 nm or (C) 1 µm.
82
Several peroxisomal proteins co-localize with Pex14 in pex3 atg1 cells
To examine whether other PMPs are also associated with the structures, we performed co-
localization studies using pex3 atg1 strains producing Pex14-mCherry together with different
PMP-mGFP fusion proteins, all under control of their endogenous promoter. We analysed Pex8,
Pex10 and Pex13, proteins of the importomer (Rucktaschel et al., 2011), as well as Pex11, a
PMP involved in peroxisome fission (Thoms and Erdmann, 2005), and Pmp47, a peroxisomal
carrier protein (Sakai et al., 1996). In WT cells grown for 16 h on MM-M/G, all mGFP fusion
proteins were readily detected at peroxisomes (“unpublished data”). In pex3 atg1 cells, Pex8-
mGFP and Pex13-mGFP co-localized with Pex14-mCherry, whereas the levels of Pex10-mGFP,
Pex11-mGFP and Pmp47-mGFP were below the limit of detection (Fig. 3A). To precisely
compare the levels of the above proteins, we monitored their induction after shifting cells from
peroxisome repressing (glucose; MM-Glu) to peroxisome inducing conditions (MM-M/G). In
pex3 atg1 cells, Pex8, Pex13 and Pex14 showed similar induction patterns (Fig. S1A-C) and
protein levels (Fig. 3B) as WT controls. Conversely, Pex10, Pex11 and Pmp47 were only
detected in pex3 atg1 cells at the initial stages after the shift, with the highest levels after 6 h of
induction (Fig. S1D-E) followed by a very strong reduction after prolonged cultivation.
However, at 6 h their levels were still strongly reduced compared to the WT controls (Fig. 3B).
FM revealed that in these cells Pex11-mGFP is predominantly localized to the nuclear envelope
and lateral ER, while Pmp47-mGFP was dispersed over the cytosol and Pex10-mGFP below the
limit of detection (Fig. 3C).
The strong reduction in Pex10, Pex11 and Pmp47 levels cannot be (fully) explained by a sudden
arrest in the synthesis of these PMPs, since growth was minimal between 6 and 8 h (Fig. S1G),
and hence must be caused by proteolytic degradation.
83
Figure 3. Pex10, Pex11 and Pmp47 do not localize with Pex14. (A) FM images of pex3 atg1 cells grown for 16 h on MM-M/G. Besides Pex14-mCherry, cells produced C-terminal
mGFP fusions of the indicated proteins. (B) WB analysis of (1) WT and (2) pex3 atg1 cells, grown for 6 or 16 h on
MM-M/G. (C) FM images showing mGFP-fluorescence in pex3 atg1 cells producing Pex14-mCherry (control) or
Pex14-mCherry together with the indicated mGFP fusion protein. Cells were grown for 6h on MM-M/G. (D,E) iEM
analysis of pex3 atg1 cells using (D) α-Pex5 or (E) α-alcohol oxidase antibodies. (F) Cell fractionation analysis of
the indicated strains. Post nuclear supernatants (PNS) were subjected to differential centrifugation resulting in a
30,000 x g organelle pellet (P) and supernatant fraction (S). (G) Flotation analysis of the organelle pellet showing
the distribution of the indicated proteins in the top (1) to bottom (10) fractions. (H) Co-localisation of Pex8-GFP and
Pex14-mCherry in pex10 cells. Bars: (A) 2.5 μm, (C) 5 μm, or (D, E) 100 nm. AOX – cytosolic alcohol oxidase
crystalloid; N – nucleus; V – vacuole.
84
Therefore, we conclude that in H. polymorpha pex3 atg1 cells two classes of PMPs can be
discriminated: i.e. those that sort independently of Pex3 to vesicular structures, where they are
relatively stable; and PMPs that require Pex3 for sorting and stability.
Pex13 and Pex14 are associated with membranes in pex3 atg1 cells
To study whether PMPs are membrane-bound in atg1 pex3 cells a flotation analysis of an
organelle pellet was performed. Pex8, Pex13 and Pex14, were detected in the organelle pellet
and migrated to fractions of low density upon flotation centrifugation (Fig. 3F, G). Pex10 and
Pmp47 could not be analysed because of strong degradation during the fractionation procedure
(“unpublished data”). Interestingly, the PTS1-receptor Pex5, as well as a minor portion of the
peroxisomal matrix protein alcohol oxidase, co-fractionated with Pex14 (Fig. 3F, G). Bulk of the
pelleted AO represents cytosolic crystalloids, which do not float. Localisation of Pex5 and AO at
the vesicles was confirmed by iEM (Fig. 3D, E). The accumulation of Pex5 at these structures
can be explained by the presence of a functional receptor docking complex, and the absence of
Pex10, which is essential for receptor recycling. Our observation that the structures contain
matrix protein is supported by the electron density of their lumen (Fig. 1A, B). Association of
Pex8 with the Pex14-containing structures is in line with observations obtained in Pichia
pastoris, which revealed that Pex8 import into peroxisomes only depends on PTS receptors and
Pex14 (Zhang et al., 2006; Ma et al., 2009). Also in H. polymorpha pex10 cells, Pex8 co-
localizes with Pex14 (Fig. 3H).
85
Pex14-containing structures in pex3 atg1 cells develop into peroxisomes upon
reintroduction of Pex3
To analyse whether the membrane structures can develop into peroxisomes upon re-introduction
of Pex3, we constructed a pex3 atg1 strain that contained PEX3-eGFP under control of the
inducible amine oxidase promoter (PAMO). Cells were extensively pre-cultivated on MM-Glu in
the presence of ammonium sulphate to fully repress PAMO. Subsequently, cells were shifted to
MM-M/G/methylamine to induce PAMO and peroxisome proliferation. Live cell imaging revealed
that the first eGFP fluorescence invariably co-localized with the Pex14-mCherry spots (Fig. 4B).
The Pex14-mCherry spots present in pex3 single deletion cells also appeared to be the sole
targets for reintroduced Pex3-eGFP (Fig. 4A). We then examined Pex10-mGFP and Pmp47-
mGFP upon reintroduction of Pex3 using strains that also produced Pex14-mCherry and PAMO-
driven PEX3. In cells pre-cultivated on MM-Glu with ammonium sulphate, these PMPs (with the
exception of Pex14-mCherry) were below the limit of detection (“unpublished data”). Upon
induction of PEX3 expression, the first Pex10-mGFP fluorescence signal appeared after 5 h, and
invariably co-localized with Pex14-mCherry (Fig. 4C). A similar result was observed for
Pmp47-mGFP, except that the first fluorescence was detected after 8 h (Fig. 4D).
Finally, we tested whether the Pex14-containing vesicles are capable of importing the matrix
marker GFP-SKL upon PEX3 induction. As shown in Figure 4E, GFP-SKL was cytosolic prior
to Pex3 reintroduction, but found to be concentrated at the Pex14-mCherry spots when Pex3
synthesis was induced.
These results indicate that in pex3 atg1 cells the Pex14-containing structures, rather than the ER,
are the target for re-introduced Pex3. Subsequently, Pex10 and Pmp47 also sort to these
structures, which mature into normal peroxisomes that import GFP-SKL.
86
Figure 4. The Pex14-containing vesicular structures mature into peroxisomes upon reintroduction of PEX3.
(A) FM images of pex3 cells with Pex14-mCherry upon Pex3-eGFP reintroduction after shifting cells from MM-Glu
with ammonium sulphate to MM-M/G with methylamine. (B-E) Live cell FM images of pex3 atg1 cells upon Pex3
reintroduction. pex3 atg1 cells producing Pex14-mCherry and (B) PAMOPEX3-eGFP, (C) PAMOPEX3.PEX10-GFP,
(D) PAMOPEX3.PMP47-GFP or (E) PAMOPEX3.PTEFGFP-SKL. Cells were grown similar as in (A). Bars: 2.5 µm. The
arrows in B-D indicate the first GFP signal that was detected.
87
Pex10, Pex11 and Pmp47 are stabilized upon Pex19 overproduction
One of the models of Pex19 function proposes that cytosolic Pex19 binds newly synthesized
PMPs, followed by recruitment of the complex by Pex3, and subsequent insertion of the PMPs
into the peroxisomal membrane (Schliebs and Kunau, 2004). This led us to speculate that in the
absence of Pex3, Pex19 may become saturated with PMPs that are dependent on the Pex3/Pex19
machinery, since cargo release is abolished. As a consequence, additionally synthesized PMPs
cannot bind to Pex19 and may become susceptible to degradation. To test this, we analysed the
effect of Pex19 overproduction, finding indeed an increase of the levels of Pex10, Pex11 and
Pmp47, but not of Pex14 (Fig. 5A, B). The enhanced protein levels allowed visualization of
Pex10 and Pmp47 by FM, which revealed that they both are cytosolic (Fig. 5C, D). These
findings are consistent with a model that newly synthesized Pex10, Pex11 and Pmp47 directly
insert into the peroxisomal membrane by a process that requires Pex3 and Pex19.
Pex19 and Pex25 are not required for vesicle formation in pex3 atg1 cells
Because in vitro assays suggested that Pex19, rather than Pex3, is essential to form peroxisomal
vesicles from the ER (Agrawal et al., 2011; Lam et al., 2010), we analyzed whether Pex14-
containing structures occur in H. polymorpha pex19 and pex19 atg1 cells. As shown in Fig. 5 E-
F, structures, similar to those observed in pex3 cells, are also present in these cells.
It is known that Pex25 is required for the reintroduction of peroxisomes in H. polymorpha pex3
cells (Saraya et al., 2011). However, as clusters of vesicular structures and Pex14-mGFP spots,
similar as those observed in pex3 atg1 cells, were also observed in cells of a pex3 atg1 pex25
triple deletion strain (Fig. 5G, H), this suggests that Pex25 is not required for the formation of
these vesicles.
88
Figure 5. Pex19 overproduction and PEX19 or PEX25 deletion. (A) WB analysis and (B) quantification of the
indicated proteins in WT (lane 1; blue), pex3 atg1 (lane 2; red), and pex3 atg1-PAOXPEX19 (lane 3; green) cells
grown for 6 h on MM-M/G. In B, the protein levels of WT cells were set to 100 %. (C) FM images and (D)
quantification of Pmp47-mGFP and Pex10-mGFP in pex3 atg1 (1; red) and pex3 atg1-PAOXPex19 (2; green) cells
grown for 6 h on MM-M/G. Control cells in D did not produce mGFP. Significance indications: NS=p<0.10,
*=0.10>p>0.05, **=0.05>p>0.01, ***=p<0.01. Error bars: SD. (E-F) iEM analysis of (E) pex19 and (F) pex19
atg1 cells using α-Pex14 antibodies. (G) EM analysis of KMnO4-fixed pex3 atg1 pex25 cells grown for 16 h on
MM-M/G showing clusters of membrane structures (arrows). (H) FM image of pex3 atg1 pex25 cells producing
Pex14-mGFP. Scale bars: (C) 2 µm, (E-F) 100 nm, (G) 250 nm, or (H) 2.5 µm. CW – cell wall; ER – endoplasmic
reticulum; M – mitochondrion; N – nucleus; V – vacuole.
89
Conclusions
Because peroxisomal membranes to which common marker PMPs co-localize were not detected
in yeast (Baerends et al., 1996; Hettema et al., 2000; Wiemer et al., 1996) or mammalian
(Shimozawa et al., 2000) cells lacking a functional PEX3 gene, it is generally accepted that cells
lacking Pex3 are unable to form peroxisomal membranes. Instead, FM analysis suggested that
PMPs were localized to the ER, mitochondria or were below the limit of detection, depending on
the marker PMP examined (South, 2000; Haan et al., 2006; Hettema et al., 2000; van der Zand et
al., 2010). Here, we show that in the absence of Pex3, the PMPs Pex13 and Pex14 co-localize at
membrane structures that are often located adjacent to other cell organelles at distances that
cannot be resolved by FM. Apparently, these PMPs can insert in membranes independent of
Pex3 (Fig. S2).
Based on FM, van der Zand et al. (2010) concluded that, in S. cerevisiae pex3 cells Pex13 and
Pex14 are present in foci at the ER. We consider it likely that these foci represented similar
structures. Indeed, our iEM analyses on S. cerevisiae pex3 atg1 cells revealed that these cells
also harbour Pex14-containing vesicles (“unpublished data”). Our observations are furthermore
supported by the presence of PMP containing membrane structures in P. pastoris pex3 cells
(Hazra et al., 2002).
In contrast to Pex13 and Pex14, the Pex10, Pex11 and Pmp47 apparently do require the
Pex3/Pex19 machinery for insertion into these membrane structures, given that in cells lacking
Pex3 they do not co-localize with Pex14 and are very instable. Instead, they are stabilized and
sorted to the structures upon Pex3 reintroduction. In addition, their levels increase in pex3 atg1
cells upon PEX19 overexpression, suggesting that Pex19 serves as cytosolic receptor for these
PMPs (Fig. S2). Pex10 and Pmp47 were invariably cytosolic in pex3 atg1 cells, which is
90
consistent with the cytosolic localization of the mammalian Ant1 (a homologue of Pmp47) in
PEX3 mutant cells (Fang et al., 2004).
Pex11 was the only PMP which we (transiently) observed at the ER, but only in minor amounts,
with the protein being very instable. The latter finding is consistent with pulse-chase experiments
using S. cerevisiae pex3 cells, which showed that Pex11 is normally synthesized, but – unlike in
the WT control – rapidly degraded (Hettema et al., 2000). This instability suggests that
localization at the ER may not be an intermediate stage of its normal sorting pathway. However,
at this stage it cannot be excluded that Pex11 traffics via the ER to peroxisomes and is degraded
in pex3 cells because of its inability to exit the ER. We note, however, that this pathway is not
consistent with our observation that Pex11 levels increase upon Pex19 overproduction in pex3
cells.
The most pressing question is the nature of the vesicles in pex3 cells. Our data indicate that they
have several properties in common with normal peroxisomal membranes as they appear to
contain a functional receptor docking site to which Pex5 associates, and are capable of importing
matrix proteins (Pex8, alcohol oxidase). This property is shared with peroxisomal membrane
ghosts that are present in H. polymorpha PEX deletion strains, which are defective in receptor
recycling, e.g. pex4 or pex10 (Koek et al., 2007). Thus, they may represent peroxisomal ghosts,
an assumption that is reinforced by the finding that they mature into normal peroxisomes upon
Pex3 reintroduction.
According to our model (Fig. S2), the vesicles may proliferate from a pre-existing peroxisomal
membrane structure. Alternatively, they may form from other membranes. If so, they are most
likely formed from the ER (Tabak et al., 2013; Fakieh et al., 2013), possibly by a similar
mechanism as the in vitro generated vesicles reported by Lam et al. (2010) and Agrawal et al.
91
(2011). Importantly, our current data demonstrate that, if these structures indeed derive from the
ER, their formation does not require Pex3.
Acknowledgements
We thank Ruchi Saraya, Geerke Maathuis and Chris Williams for their valuable contributions.
We thank Abraham Koster (LUMC) for making the electron tomography facilities available.
This work was supported by an EU Marie Curie IEF grant to K.K. (FP7-330150).
92
MATERIALS AND METHODS
Strains and growth conditions
The H. polymorpha strains used in this study are listed in Table S1. Yeast cultures were grown at
37°C, either on (1) YPD media containing 1% yeast extract, 1% peptone and 1% glucose, (2)
selective media containing 0.67% yeast nitrogen base without amino acids (YNB; Difco), or (3)
mineral media (MM) (van Dijken et al., 1976) supplemented with 0.5% glucose (MM-Glu),
0.5% methanol or a mixture of 0.5 % methanol and 0.05 % glycerol (MM-M/G) as carbon
sources and 0.25% ammonium sulphate or 0.25% methylamine as nitrogen sources. If required,
amino acids, uracil or leucine were added to a final concentration of 30 μg/ml. For growth on
agar plates, the medium was supplemented with 2% agar. For the selection of resistant
transformants, YPD plates containing 100 μg/ml zeocin (Invitrogen), 300 μg/ml hygromycine B
(Invitrogen) or 100 μg/ml nourseothricin (Werner Bioagents) were used.
For cloning purposes, Escherichia coli DH5α was used. Cells were grown at 37°C in LB media
supplemented with 100 μg/ml ampicillin or 50 μg/ml kanamycin, when required.
Molecular and biochemical techniques
Standard recombinant DNA techniques and transformation of H. polymorpha was performed by
electroporation as described previously (Faber et al., 1994). Cell extracts of TCA treated cells
were prepared for SDS-PAGE as detailed previously (Baerends et al., 2000). SDS-PAGE and
western blotting (WB) were performed by established methods. Equal amounts of protein were
loaded per lane and blots were probed with rabbit polyclonal antisera against Pex11, Pex14 or
pyruvate carboxylase-1 (Pyc1). mGFP-fusion proteins of Pex8, Pex10, Pex13 and Pmp47 were
detected using mouse monoclonal antiserum against green fluorescent protein (GFP; Santa Cruz
93
Biotechnology, sc-9996). Secondary antibodies conjugated to horseradish peroxidase were used
for detection. Pyc1 was used as a loading control. Blots were scanned by using a densitometer
(Biorad GS-710) and quantified using Image J (http://rsbweb.nih.gov/ij/). From two individual
blots per sample, the total intensity of the band of interest was measured and corrected for
background intensity and Pyc1 loading amount.
Construction of H. polymorpha strains
The plasmids and primers used in this study are listed in Table S2 and S3. All integrations were
confirmed by PCR. All deletions were confirmed by PCR and Southern blotting.
Construction of the pex3 atg1 and pex19 atg1 double deletion strain, and the pex3 atg1
pex25 triple deletion strain
The pex3 atg1 double deletion strain was obtained by crossing a H. polymorpha pex3 strain
(Baerends et al., 1996) with an atg1 strain (Komduur et al., 2003). Diploids were subjected to
random spore analysis and prototrophic segregants were subjected to complementation analysis
to determine their genotypes (Sudbery et al., 1988). The pex3 atg1 pex25 triple deletion strain
was made as follows. A PCR fragment of 2912 bp was obtained by PCR using primers Pex25-F
and Pex25-R and plasmid pRSA018 as a template (Saraya et al., 2011). This PCR fragment was
transformed to the pex3 atg1 double deletion strain. For the pex19 atg1 double deletion strain, a
PEX19-deletion cassette plasmid (pHOR30b) was digested with BglII and EcoRI to replace the
URA3 gene with the LEU2 gene, which was obtained after digestion of pBS-CaLeu2 with
BamHI and EcoRI. The final deletion PEX19-deletion plasmid (pSEM188) was digested with
94
BamHI and the resulting 4434 bp fragment was integrated in the genome of atg1 cells (Komduur
et al., 2003).
Construction of other strains
Plasmids pHIPZ-PEX8-mGFP (pMCE4), pHIPZ-PEX10-mGFP (pMCE5), pHIPZ-PEX25-
mGFP (pMCE1), pHIPZ-PMP47-mGFP (pMCE7) (Cepinska et al., 2011), pHIPZ5-PEX3-eGFP
and pHIPZ4-BiPN30-eGFP-HDEL (pRSA017) (Saraya et al., 2010) were linearized and integrated
in the endogenous promoter regions in the pex3 atg1 strain producing Pex14-mCherry essentially
as described previously (Cepinska et al., 2011; Saraya et al., 2010).
For the construction of plasmid pSEM01, a PCR fragment of 563 bp was obtained by using
primers Pex14-F and Pex14-R on genomic DNA. After digestion with HindIII and BglII, the
resulting fragment was inserted between the HindIII and BglII sites of pMCE02, resulting in
pSEM01 (5488 bp) containing pHIPN-PEX14-mCherry. For stable integration in the PEX14
promoter region, XhoI linearized plasmid was transformed to the pex3 atg1 double mutant,
resulting in a strain producing Pex14-mCherry under control of the endogenous promoter.
Plasmid pSEM02 (pHIPZ-PEX11-mGFP) was obtained as follows: Digestion of the pHIPZ-
mGFP fusinator plasmid with HindIII and BglII yields a fragment of 5077 bp. Similarly, the
pHIPN-PEX11-mCherry plasmid (pMCE3) was digested with HindIII and BglII to obtain a
fragment of 772 bp. Ligation of the 772 bp and 5077 bp fragments resulted in pSEM02 of
5849bp. The plasmid was linearized using Pst1 and integrated in the genome of pex3 atg1
producing Pex14-mCherry.
To construct plasmid pSEM03 (pHIPZ-PEX13-mGFP), PCR was performed on genomic DNA
using the primers Pex13-F and Pex13-R. The PCR product of 1146 bp was digested with HindIII
95
and BglII and the resulting fragment was inserted between the HindIII and BglII sites of pHIPZ-
mGFP fusinator plasmid. The resulting plasmid of 6223 bp, designed pSEM03, was linearized
with ApaI and transformed to H. polymorpha pex3 atg1 producing Pex14-mCherry.
For the construction of plasmid pSEM04, a PCR fragment of 2547 bp was obtained using
plasmid pHIPZ5-PEX3-eGFP plasmid (Table S2) as a template and primers H5-F and H5-R. The
PCR fragment was digested with NotI and PspXI and the resulting fragment was ligated in NotI
and SalI digested pHIPH4, resulting in plasmid pSEM04, which contains pHIPH5-PAMO-PEX3.
The plasmid was linearized with BsiWI and integrated in strain pex3 atg1.Pex14-mCherry
producing Pex10-mGFP or Pmp47-mGFP.
Similarly, plasmid pSEM05 was made by PCR amplification of an 885 bp fragment using
genomic DNA and primers Pex19-F and Pex19-R (Table S3). After digestion with HindIII and
XbaI, the resulting fragment was ligated in HindIII and XbaI digested pHIPH4, resulting in
plasmid pSEM05 containing pHIPH4-PAOX-PEX19. For stable integration, StuI linearized
plasmid was transformed to H. polymorpha pex3 atg1.Pex14-mCherry producing Pex10-mGFP
or Pmp47-mGFP.
For the construction of plasmid pAKW27, a vector of 5831 bp was obtained by BamHI and SalI
digestion of pHIPZ7, whereas the 736 bp eGFP-SKL insert was obtained by BamHI and SalI
digestion of pFEM35 followed by gel extraction. Ligation resulted in the plasmid pAKW27
containing pHIPZ7-PTEF1-GFPSKL. For stable integration, stuI linearized plasmid was
transformed to H. polymorpha pex3 atg1.Pex14-mCherry producing Pex3 under control of the
inducible PAMO.
96
Cell fractionation and membrane flotation
Crude extracts were prepared as described before (Baerends et al., 1997). Briefly, protoplasts
were prepared with Zymolyase (Brunschwig Chemie, Amsterdam, The Netherlands) and
homogenized using a Potter homogenizer. To remove cell debris, the homogenate was
centrifuged 2x at 3,000 x g (10 min, 4°C). The supernatant (PNS) was then subjected to
centrifugation at 30,000 x g (30 min, 4°C) to separate the soluble fraction (supernatant: S) from
the membrane pellet (P).
The 30,000 x g organelle pellet was used for flotation centrifugation as described before
(Baerends et al., 1997). Briefly the pellet was dissolved in 50% sucrose and over layered with
40%, 30% and 20% sucrose. Centrifugation was performed at 140,000 x g for 16 h at 4 °C. 10
fractions of 200 µl were collected from the top and analysed by SDS-PAGE and WB.
Fluorescence microscopy
All images were made using a 100x 1.30 NA Plan-Neofluar objective (Carl Zeiss). For wide-
field microscopy, the GFP signal was visualized with a 470⁄40 nm band pass excitation filter, a
495 nm dichromatic mirror, and a 525⁄50 nm band-pass emission filter. mCherry fluorescence
was visualized with a 587/25 nm band pass excitation filter, a 605 nm dichromatic mirror, and a
647/70 nm band-pass emission filter. DsRed, FM4-64 and Mitotracker Orange fluorescence were
visualized with a 546/12 band-pass excitation filter, a 560 nm dichromatic mirror, and a 575–640
nm band-pass emission filter. Images were captured using a Zeiss Axioskop50 fluorescence
microscope (Carl Zeiss, Sliedrecht, The Netherlands) using MetaVue software and a Princeton
Instruments 1300Y digital camera. The images were captured in the media in which the cells
were grown.
97
Mitochondria were stained by incubation of intact cells for 30 min at 37°C with 0.5 μg/ml
MitoTracker Orange (Invitrogen) followed by extensive washing with medium. For vacuolar
staining, 1 ml of cell culture was supplemented with 1 µl FM4-64 (Invitrogen), incubated for 60
min at 37°C and analysed.
Live cell imaging was performed on a Zeiss Observer Z1 using Axiovision software and a
Photometrics Coolsnap HQ2 digital camera. Cells were grown on 1% agar containing growth
medium and the temperature of the heating chamber XL was set at 37C. Three z –axis planes
were acquired for each time interval using 0.5 sec exposure times for both GFP and mCherry.
Confocal images were captured with a confocal microscope (LSM510; Carl Zeiss), equipped
with photomultiplier tubes (Hamamatsu Photonics) and Zen 2009 software. For live cell
imaging, the temperature of the objective and object slide was kept at 37C and the cells were
grown on 1% agar in medium. GFP fluorescence was analysed by excitation of the cell with a
488-nm argon ion laser (Lasos), and emission was detected using a 500–550 nm band-pass
emission filter. During simultaneous GFP and FM4-64 detection, both probes were excited with
a 488-nm argon ion laser and GFP was detected using 500–530 nm band-pass emission filter and
FM4-64 was detected using a 560 nm long-pass emission filter. Six z-axis planes were acquired
for each time interval.
Image analysis was carried out using ImageJ and figures were prepared using Adobe Photoshop
CS4. Unless indicated otherwise, the intensity minimum and maximum of the image were set
equal for all images represented within a single figure panel, thus facilitate direct fluorescence
intensity comparison between different strains.
For quantitative analysis of Pex14-mGFP fluorescent spots, Z-stacks were made of randomly
chosen fields. Quantification was done on 4 images per culture, containing at least 65 cells per
98
image. Cells were stained with FM4-64 to allow discrimination between vacuolar mGFP and
cytosolic mGFP spots. The average number of spots was calculated from 350 cells per culture.
The error bars indicate the standard error of the mean.
For the quantification of cytosolic mGFP intensity, single plane images were acquired on a Zeiss
Axioskop50, after which the total intensity of ~100 individual cells was measured and corrected
for the background intensity. The error bars indicate the standard deviation between individual
cells.
Electron microscopy
H. polymorpha pex3 atg1 were fixed in 1.5% potassium permanganate, stained en block with
0.5% uranylacetate and embedded in Epon 812 (Serva, 21045). For morphological studies,
ultrathin sections were viewed in a Philips CM12 TEM. For electron tomography, serial sections
of 150 nm thick were cut. The serial images of whole cells were stacked and aligned using
MIDAS (Kremer et al., 1996), after which individual cells could be scrutinized for peroxisomal
remnants. 10 nm gold beads were layered on top of the serial sections and acted as fiducial
markers for electron tomography. Two single-axis tilt series, each containing 141 images with 1°
tilt increments, were acquired with a pixel size of 0.7 nm on a FEI Tecnai12 at 120 kV using the
SerialEM acquisition software (Mastronarde, 2005) and a cooled slow-scan charge-coupled
device camera (4k Eagle; FEI Company) in 2x2 binned mode. The tilt series were aligned and
reconstructed using the IMOD software package and analysed using the AMIRA visualization
package (TGS Europe). To generate 3-dimensional surface rendered models in AMIRA, first,
masks of organelles were drawn manually and afterwards improved by nonlinear anisotropic
diffusion filtering followed by thresholding.
99
Cryosectioning and immuno-gold labelling
For immuno-gold labelling, cells were fixed in 3% glutaraldehyde in 0.1 M cacodylatebuffer pH
7.2 for 1 h on ice and treated afterwards with 0.4% Na-periodate (15 min) and 1% NH4Cl (15
min). Upon embedding in 12% gelatine in phosphate buffer pH 7.4, ~0.5 mm3 cubes were
infiltrated overnight in 2.3 M sucrose in the same buffer. Cryosections of 60 nm were cut using a
cryo diamond knife (Diatome) at -120 °C in a Reichert Ultracut. Sections were mounted on
carbon-coated formvar nickel grids. Gelatine was removed by incubating the grids for 30
minutes on 2% gelatine in phosphate buffer (pH 7.4) at 30°C. Pex14, Pex5 and alcohol oxidase
were localized using polyclonal antibodies raised against Pex14, Pex5 and alcohol oxidase,
respectively, and goat-anti-rabbit antibodies conjugated to 10 nm gold (Aurion). Sections were
stained with 2% uranyloxalate (pH 7.0) for 10 minutes, briefly washed on three drops of distilled
water and embedded in 0.5% methylcellulose and 0.5% uranylacetate on ice for 10 min before
viewing them in the CM12 electron microscope (Slot and Geuze, 2007).
100
Supplemental Figures
Figure S1. Induction of PMPs in WT and pex3 atg1 cells. (A-F) Cells, pre-cultivated on MM-Glu (0h), were
shifted to MM-M/G. Samples were taken at the indicated time points. (G) Growth curves of the WT and pex3 atg1
cultures. Error bars: SD of 5 experiments.
101
Figure S2. Model. In pex3 cells Pex13/Pex14-containing structures exist that import Pex8. These structures may
arise by proliferation of a pre-existing structure or form from the ER. They are constitutively degraded by
autophagy, unless ATG1 is deleted. Pex10, Pex11 and PMP47 are instable, because they are not inserted into these
structures. Re-introduced Pex3 is sorted to the structures, followed by insertion of the other PMPs by the
Pex3/Pex19 machinery and import of matrix proteins.
102
Table S1. H. polymorpha strains used in this study
Strains Characteristics Reference
wild-type (WT) NCYC 495 leu1.1 (Saraya et al., 2012)
atg1 ATG1 deletion strain, leu 1.1 (Komduur et al., 2003)
pex3 PEX3 deletion strain, ura3 (Baerends et al., 1996)
pex19 PEX19 deletion strain, ura3 (Otzen et al., 2004)
pex10 PEX10 deletion strain (Tan et al., 1995)
pex3 atg1 PEX3 ATG1 double deletion strain This study
pex19 atg1 PEX19 ATG1 double deletion strain This study
pex3 atg1.PEX14-mCherry pex3 atg1 with pSEM01 This study
pex3 atg1.PEX14-mCherry.PEX8-
mGFP
pex3 atg1 with pSEM01 and pMCE4 This study
pex3 atg1.PEX14-mCherry.PEX10-
mGFP
pex3 atg1 with pSEM01and pMCE5 This study
pex3 atg1.PEX14-mCherry.PEX11-
mGFP
pex3 atg1 with pSEM01 and pSEM02 This study
pex3 atg1.PEX14-mCherry.PEX13-
mGFP
pex3 atg1 with pSEM01 and pSEM03 This study
pex3 atg1.PEX14-mCherry.PMP47-
mGFP
pex3 atg1 with pSEM01 and pMCE7 This study
pex3 atg1.PEX13-mGFP pex3 atg1 with pSEM03 This study
pex3 atg1.PEX14-mCherry-BiPN30-
eGFP-HDEL
pex3 atg1 with pSEM01 and pRSA017 This study
pex3 atg1.PEX14-mCherry.PEX10-
mGFP – PAOX PEX19
pex3 atg1 with pSEM01, pMCE05 and
pSEM05
This study
pex3 atg1.PEX14-mCherry.PMP47-
mGFP – PAOX PEX19
pex3 atg1 with pSEM01, pMCE07 and
pSEM05
This study
pex3 atg1.PEX14-mGFP pex3 atg1 with pSNA12 This study
pex3 atg1 pex25 pex3 atg1 with pRSA018 This study
pex3 atg1 pex25.PEX14-mGFP pex3 atg1 pex25 with pSNA12 This study
pex3 atg1.PEX14-mCherry – PAMO
PEX3-eGFP
pex3 atg1 with pSEM01 and pHIPZ5-PAMO
PEX3-eGFP
This study
pex3 atg1.PEX14-mCherry.PEX10-
mGFP–PAMO PEX3
pex3 atg1 with pSEM01, pMCE5 and
pSEM04
This study
pex3 atg1.PEX14-mCherry.PMP47-
mGFP–PAMO PEX3
pex3 atg1 with pSEM01, pMCE7 and
pSEM04
This study
pex3 atg1.PEX14-mCherry – PAMO
PEX3-eGFP – PTEF1 eGFP-SKL
pex3 atg1 with pSEM01, pSEM04 and
pAKW27
This study
pex10. PEX8-mGFP.PEX14-mCherry pex10 with pSEM01 and pMCE4 This study
WT. PEX13-mGFP WT with pSEM03 This study
WT. DsRed-SKL.PEX8-mGFP WT. DsRed-SKL with pMCE4 (Cepinska et al., 2011)
WT. DsRed-SKL.PEX10-mGFP WT. DsRed-SKL with pMCE5 (Cepinska et al., 2011)
WT. DsRed-SKL.PMP47-mGFP WT. DsRed-SKL with pMCE7 (Cepinska et al., 2011)
WT. PEX10-mGFP.PEX14-mCherry WT with pSEM01 and pMCE7 This study
WT. PEX11-mGFP.PEX14-mCherry WT with pSEM01 and pMCE3 This study
WT. DsRed-SKL.PEX13-mGFP. WT. DsRed-SKL with pSEM03 This study
103
Table S2. Plasmids used in this study
Plasmid Characteristics Reference
pSEM01 pHIPN Plasmid containing C-terminal part of PEX14
fused to mCherry; natR; amp
R
This study
pSEM02 Plasmid containing C-terminal part of PEX11 fused to
mGFP; zeoR; amp
R
This study
pSEM03 Plasmid containing C-terminal part of PEX13 fused to
mGFP; zeoR; amp
R
This study
pSEM04 pHIPH5 containing PEX3 under control of PAMO, HphR,
;
ampR
This study
pSEM05 pHIPH4 containing PEX19 under control of PAOX, HphR,
;
ampR
This study
pSEM188 Plasmid containing PEX19 deletion cassette; LEU2;
ampR
This study
pHIPZ5-PEX3-eGFP pHIPZ5 containing PEX3-eGFP under control of PAMO,
zeoR; amp
R
(Nagotu et al., 2008)
pRSA017 pHIPZ4 containing BiPN30-eGFP-HDEL under control of
PAOX; zeoR; amp
R
(Saraya et al., 2011)
pHIPH4 pHIP containing hygromycine B marker, ampR (Saraya et al., 2011)
pHIPZ7 pHIP containing TEF1 promoter, zeo R
; ampR (Baerends et al., 1997)
pFEM35 pHIPX7 containing eGFP with C-terminal PTS1 under
control of PAOX; Leu2; kanR
(Krikken et al., 2009)
pHIPZ-mGFP fusinator pHIPZ containing mGFP and AMO terminator; zeoR;
ampR
(Saraya et al., 2010)
pMCE02
pHIPN plasmid containing mCherry; natR; amp
R (Cepinska et al., 2011)
pSNA12 Plasmid containing C-terminal part of PEX14 fused to
mGFP; zeoR; amp
R (Cepinska et al., 2011)
pMCE1 C-terminus of PEX25 fused to mGFP; zeoR; amp
R (Cepinska et al., 2011)
pMCE3 Plasmid containing C-terminal part of PEX11 fused to
mCherry; natR; amp
R
(Cepinska et al., 2011)
pMCE4 Plasmid containing C-terminal part of PEX8 fused to
mGFP; zeoR; amp
R
(Cepinska et al., 2011)
pRSA018 pDEST-R4-R3 containing PEX25 deletion cassette, natR;
ampR
(Saraya et al., 2011)
pMCE5 Plasmid containing C-terminal part of PEX10 fused to
GFP; zeoR; amp
R
(Cepinska et al., 2011)
pMCE7 Plasmid containing C-terminal part of PMP47 fused to
GFP; zeoR; amp
R
(Cepinska et al., 2011)
pAKW27 Plasmid containing eGFP-SKL under control of PTEF1;
zeoR; amp
R
This study
pHOR30b Plasmid containing PEX19 deletion cassette; URA3;
ampR
(Otzen et al., 2006)
pBS-CaLeu2 Plasmid containing C. albicans LEU2 gene; ampR (Otzen et al., 2006)
104
Table S3. Primers used in this study
Primer Sequence Reference
Pex25-F 5′-CTGGATGGAGGCTTCATCTC-3′ (Saraya et al., 2011)
Pex25-R 5′-GGAGCTGCTGTGCTTGTATG-3′ (Saraya et al., 2011)
H5-F 5′-GTGCTGCAAGGCGATTAAGT-3′ This study
H5-R 5′-AGAGCTCGAGGTTAAGCATCGAAATTAGAGTAGAC-3′ This study
Pex13-F 5′-AAAAGCTTATGACTACACCACGTCCAAAGCC-3′ This study
Pex13-R 5′-AAAGATCTGATCAATAGCTTTTGATCTTTCTTGAAC-3′ This study
Pex14-F 5′-CCCAAGCTTCGTTGCAGGAAGTCGACGAA-3′ This study
Pex14-R 5′-AGATCTTCCGGCATTCAGCTGCCACGCCG-3′ This study
Pex19-F 5′-AAAAGCTTATGAGCGAGAAAAAGTCC-3′ This study
Pex19-R 5′-ATCTAGACTATGTTTGTTTGCAAGTGTCTTCC-3′ This study
105
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Chapter 4
Pre-peroxisome formation in Hansenula polymorpha pex19 cells does
not require the ER.
Selvambigai Manivannan, Kèvin Knoops, Arjen M. Krikken, Anita Kram, Marten
Veenhuis and Ida J. van der Klei
Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute,
University of Groningen, Nijenborgh 7, 9747AG Groningen, The Netherlands
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Abstract
We demonstrate that H. polymorpha pex19 cells contain clusters of vesicles that contain the
peroxisomal membrane proteins (PMPs) Pex3, Pex13, Pex14 and Pex8 and thus are peroxisomal
in nature. In pex19 cells, these structures are unstable and subject to rapid degradation by
autophagy. The vesicles were stabilized in pex19 atg1 cells, in which autophagy is blocked
through deletion of the ATG1 gene. Their instability most likely is the major reason why these
structures have been overlooked so far. The vesicular structures lacked however other PMPs (i.e.
Pex10, Pex11, Pex26 and PMP47), which were unstable in pex19 atg1 cells and below the limit
of detection of fluorescence microscopy. The absence of Pex26 most likely explains the
instability of the organelles as Pex26 is essential to bind Pex1 and Pex6, proteins that recently
were shown to stimulate autophagy of peroxisome remnants. The vesicular structures, and not
the endoplasmic reticulum, served as the template for the formation of new peroxisomes upon
reintroduction of the PEX19 gene. This data challenge the current views on de novo peroxisome
formation and PMP sorting pathways.
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Introduction
Peroxisomes are essential eukaryotic organelles that perform a wide range of metabolic
functions dependent on species and environmental conditions. Generalized functions include β-
oxidation of fatty acids and the detoxification of hydrogen peroxide (reviewed by Van den Bosch
et al., 1992; Purdue and Lazarow, 2001).
Peroxisome formation and maintenance are controlled by PEX genes that encode
peroxins (Distel et al., 1996). Most peroxins identified so far are involved in organelle
development, i.e. membrane protein insertion and matrix protein import. Peroxisomal matrix
proteins are synthesized by free ribosomes in the cytosol and post-translationally imported into
peroxisomes (Fujiki et al., 1984). Correct sorting of matrix proteins requires the function of two
soluble receptors, namely Pex5 and Pex7, which recognize specific peroxisomal targeting signals
(PTS) located at the extreme C- (PTS1) or N-terminus (PTS2) (Gould et al., 1987; Brocard and
Hartg, 2006; Lazarow, 2006).
However, the sorting and insertion of peroxisomal membrane proteins (PMPs) is still
strongly debated (reviewed by Hettema and Motley, 2009; Ma et al., 2011; Nuttall et al., 2011;
Rucktäschel et al., 2011; Theodoulou et al., 2013; Dimitrov et al., 2013). Two pathways are
proposed namely i) PMPs are directly inserted in their target organelle mediated by the cytosolic
chaperone/receptor Pex19 together with Pex3, the peroxisomal docking protein for the Pex19-
cargo complex (Fang et al., 2004) or ii) they first insert into the ER from which vesicles derive in
a Pex3/Pex19-dependent way, which upon fusion form nascent peroxisomes. According to the
latter model peroxisomes can be defined as a branch of the endomembrane system. In both
scenarios Pex3 and Pex19 play a vital role in PMP insertion. The PMP sorting pathway via the
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endoplasmic reticulum (ER) is predominantly studied in pex3 or pex19 mutants that were
suggested to lack peroxisomal membrane remnants upon reintroduction of the corresponding
genes (Baerends et al., 1996; Hettema et al., 2000; Tam et al., 2005; Hoepfner et al., 2005).
Recently, Knoops et al. (2014) however demonstrated that pex3 cells do contain peroxisomal
membrane remnants that are the template for peroxisome re-introduction and not the ER upon
reintroduction of Pex3. These vesicles contained few PMPs, namely Pex13, Pex14 and Pex8,
which therefore did not require Pex3/Pex19 machinery for insertion in their membranes. Other
PMPs (i.e. Pex10, Pex11 and PMP47) did not insert in peroxisomal membranes in pex3 cells, but
did sort to nascent peroxisomes upon re-introduction of the PEX3 gene. This poses the question,
whether also in pex19 comparable structures are present which perform a similar function in
peroxisome re-introduction.
Extensive analysis of H. polymorpha pex19 cells indeed uncovered peroxisomal remnants
in these cells, which served as template for peroxisome re-introduction in these cells. The results
of the work are detailed in this paper.
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Results
Presence of peroxisomal vesicles in H. polymorpha pex19 cells
Fluorescence microscopy analyses of H. polymorpha pex19 cells producing Pex3-GFP, grown
for 16 h on glycerol/methanol, showed that in approximately 40% of the cells GFP fluorescence
was present in small spots, but GFP-fluorescence was predominantly observed in the vacuole
(Fig. 1B). Similar results were obtained for Pex14-mCherry (Fig. 1A).
Figure 1. Pex3 and Pex14 localization and protein levels in pex19 and pex19 atg1 cells. Glucose-grown cells of
the indicated strains producing Pex14-mCherry and Pex3-GFP, under control of their endogenous promoters, were
shifted to media containing glycerol/methanol and grown for 16 hours. Fluorescence microscopy analysis revealed
that pex19 cells infrequently contained small mCherry (A, arrows) or GFP (B, arrow) spots in conjunction with bulk
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of the fluorescence in the vacuole. In pex19 atg1 double mutant cells no vacuolar staining was observed, these cells
all contained distinct fluorescent spots (A, B). Pex3 and Pex14 co-localize in the same spots (Fig. 1C) in pex19 atg1
cells. The scale bar indicates 1µm (D). Western blots prepared from crude extracts of glycerol/methanol-grown cells
(16 h) decorated with α-Pex3 or α-Pex14 antibodies showing the Pex3 or Pex14 protein levels in pex19 (1), pex19
atg1(2) and WT (3) cells. Pyruvate carboxylase (Pyc1) was used as loading control.
Careful immunocytochemical analysis, using α-Pex14 antibodies, revealed specific labeling on
small clusters of vesicles, often located in the vicinity of the cell membrane or other cell
organelles (Fig. 2C). Since Pex3 appeared to be unstable in pex19 cells, these putative
peroxisomal structures likely are susceptible to degradation by autophagy as in pex3 cells (Bellu
et al, 2002; Knoops et al., 2014). To inhibit degradation, we constructed a pex19 atg1 double
mutant. Western blot analysis using crude extracts of glycerol/methanol-grown cells,
demonstrated that in these cells Pex3 and Pex14, produced under control of their endogenous
promoters, were stabilized to wild-type (WT) levels (Fig. 1D) and present in distinct spots (Fig.
1A, B, right row). Fluorescence microscopy analysis of pex19 atg1 cells producing both Pex14-
mCherry and Pex3-mGFP under control of their endogenous promoters demonstrated that these
proteins co-localized in the spots (Fig. 1C).
Subsequent electron microscopy analysis of KMnO4-fixed cells revealed the presence of
vesicular structures (Fig. 2A, B) that were enhanced in volume fraction relative to those in single
PEX19 deletion cells. These structures were not observed in WT cells that contained normal
peroxisomes (not shown). Immunocytochemistry revealed that these structures besides Pex14
(Fig. 2C) also contained the matrix protein alcohol oxidase (Fig. 2D). Moreover, also the PTS1
receptor Pex5 was associated to these structures (Fig. 2D, inset). Tomography, applied to resolve
the 3D structure of the vesicle complex, revealed that these structures consisted of separate
vesicles that were not connected with other cell components (Fig. 2E, F).
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Figure 2. pex19 atg1 cells harbor Pex14-containing vesicles. (A) Ultrathin section of KMnO4-fixed pex19 atg1
cell grown for 16 h on glycerol/ methanol showing the overall cell morphology and the presence of small vesicles
adjacent to the mitochondrion and nucleus (dotted box). (B) High magnification of the part indicated in A showing
the vesicular nature of the structure. (C) Immuno-electron microscopy of these pex19 atg1 cells, using antibodies
against Pex14, alcohol oxidase (D) and Pex5 (inset in D) demonstrating the association of these proteins with the
vesicular structures. (E) Electron tomography analysis of a serial-sectioned pex19 atg1 cell containing a membrane
cluster (arrow). (F) The surface-rendered model of the tomogram reconstructions in (E) revealed the vesicular
structures in 3D (arrow). Bars: (A) 500 nm, 50 nm, (B, C, D, E and F). M – mitochondrion; N – nucleus; V –
vacuole.
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Localization of peroxisomal membrane proteins in pex19 atg1
To obtain further insight in the peroxisomal nature of the vesicular structures in pex19 atg1 cells,
we analyzed the location of other PMPs. To this end, we constructed mGFP fusions with the
PMPs Pex8, Pex10, Pex11, Pex13, Pex26 and Pmp47, all produced under control of their
endogenous promoter in pex19 atg1 cells, also producing Pex14-mCherry. After 16 hours of
cultivation on glycerol/methanol medium, Pex8-GFP- and Pex13-GFP were observed in distinct
spots that co-localized with Pex14-mCherry suggesting that these proteins are localized to the
vesicular structures (Fig. 3A, B). However, Pex10-, Pex11-, Pex26- and Pmp47-GFP fusion
proteins were below the limit of detection in fluorescence microscopy (Fig. 3C-F). The low
levels of Pex10, Pex11, Pex26 and PMP47 after 16 h of cultivation on glycerol/methanol were
confirmed by Western blot analysis (Fig. 4A). Kinetic analyses performed on pex19 atg1 cells
shifted from glucose to glycerol/methanol revealed that the levels of these proteins increased
during the initial stage of adaptation to the new growth environment (until 3 - 6 h of cultivation),
but rapidly declined upon prolonged cultivation. This reduction was not observed in the WT
control (Fig. 4A, B). By contrast, the kinetics of Pex3, Pex8, Pex13 and Pex14 induction largely
followed the pattern observed in WT cells (Fig. 4B). Together, these data indicated that Pex8,
Pex13 and Pex14, but not Pex10, Pex11, Pex26 and PMP47, are stable in pex19 atg1 cells.
Moreover, Pex8, Pex13 and Pex14 apparently do not require Pex19 for routing/insertion in the
vesicular structures.
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Figure 3. Localization of various PMPs in pex19 atg1 cells. Fluorescence microscopy images of methanol-grown
pex19 atg1 cells producing Pex14-mCherry together with C-terminal mGFP fusions of the indicated proteins. GFP
was fused to the N-terminus of Pex16. Of these, Pex8 (A) and Pex13 (B) co-localize with Pex14, whereas Pex10
(C), Pex11 (D), Pmp47 (E) and Pex26 (F) are all below the limit of detection. The scale bar indicates 1µm.
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Figure 4. Induction patterns of various PMPs in WT and pex19 atg1 cells. (A, B) Glucose-grown inoculum cells
(0 h) producing mGFP fusions of the indicated proteins, were shifted to glycerol/methanol. Samples were taken at
the indicated time points and prepared for western blotting using antibodies against GFP. The data indicate that the
Pex8, Pex13 and Pex14 induction patterns were comparable to WT (B), whereas Pex10, Pex11, Pex26 and
PMP47were induced in the initial hours of growth but reduced again upon prolonged cultivation. Equal amounts of
protein were loaded per lane. Pyruvate carboxylase (Pyc1) was used as loading control.
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Pex14 containing vesicles in pex19 atg1 cells develop in peroxisomes upon re-introduction
of PEX19
Next, we investigated whether the Pex14 containing vesicles in pex19 atg1 cells are the target for
the formation of new peroxisomes upon re-introduction of the PEX19 gene under control of the
inducible alcohol oxidase promoter (PAOX). Glucose-grown pex19 atg1 PAOXPEX19 cells that
produce Pex14-mCherry and PADH1-GFP-SKL as a peroxisomal matrix marker were shifted to
agar slides containing glycerol/methanol medium and subjected to live cell imaging by confocal
laser scanning microscopy. As shown in selected still from the video (Fig. 5), GFP-SKL was
cytosolic before induction of PEX19 expression (T = 0 min). However, shortly after induction
the first GFP-SKL was observed co-localizing with the Pex14-mCherry spot (30 and 45 min).
After 120 min all GFP-SKL had accumulated in the Pex14 spot, representing a developing
peroxisome. These data are consistent with the view that the Pex14–containing vesicles are the
target for peroxisome re-introduction in these cells.
Discussion
We have re-investigated the phenotype of yeast PEX19 deletion (pex19) cells and demonstrated
that, in contrast to what is generally assumed, these cells contained peroxisome membrane
remnants. A similar phenomenon was recently described for pex3 cells (Knoops et al., 2014).
Remarkably, in both pex19 and pex3 cells these remnants were unstable and subject to
degradation by autophagy. In pex3 this instability was most likely related to the absence of Pex3,
a protein known to be essential to initiate degradation of intact peroxisomes (Bellu et al., 2002).
In pex19 cells, Pex3 was present on the peroxisomal vesicles and their instability possibly
relatesto the absence of Pex26 on these structures and hence also Pex1 and Pex6. Very recently,
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Figure 5. Kinetics of GFP-SKL import in newly formed peroxisomes in pex19 atg1 cells. Stills from a video,
taken of pex19 atg1 cells producing Pex14-mCherry and GFP-SKL cells showing the kinetics of GFP-SKL import
in Pex14 containing structures upon reintroduction of Pex19. Cells pregrown on glucose (0 min) were shifted to an
agar slide supplemented with glycerol/methanol (0.25 %) and followed in time by CLSM. The data indicated that
after 30 min of induction the first GFP-SKL fluorescence co-localized with the Pex14-mCherry spot. After 120 min
all GFP-SKL was incorporated in this spot. Scale bar indicates 2 µm.
Hettema and colleagues (personal communication) demonstrated that peroxisomal membrane
ghosts present in pex26, pex1 or pex6 cells are constitutively degraded by pexophagy. Obviously,
this instability is the major reason why these structures, except for one report, have been largely
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overlooked so far. Indeed, Snyder et al (1999) reported the presence of Pex3 containing
elongated membrane structures in P. pastoris pex19 cells, which most likely are the equivalent of
the vesicular structures observed in H. polymorpha. Surprisingly, the structures in H.
polymorpha pex19 cells contained, as in pex3 cells, only a subset of peroxisomal membrane
proteins namely Pex3, the components of the matrix protein receptor docking site Pex13 and
Pex14 as well as Pex8. All other PMPs tested were unstable and most likely mislocalized to the
cytosol and degraded by the proteasome. However, upon re-introduction of the PEX19 gene new
peroxisomes were formed to which these PMPs were sorted. This data challenge the current
views on de novo peroxisome formation and PMP sorting pathways.
At present, two models have emerged that explain the function of Pex3 and Pex19 in PMP
sorting. The first one prescribes that peroxisomes are autonomous organelles that are maintained
by growth and fission. In this model direct insertion of PMPs into peroxisomes depends on Pex3
and Pex19, in which Pex19 acts as chaperone/receptor for PMPs and Pex3 as the peroxisomal
docking protein for the Pex19-cargo complex at the peroxisomal membrane (Schliebs and
Kunau, 2004). After docking at the peroxisomal membrane the PMP inserts into the membrane
by a yet unknown mechanism, whereas Pex19 is recycled to the cytosol, where it can bind
another newly synthesized PMP. According to the second model PMPs are first inserted into the
ER from which vesicles derive in a Pex3/Pex19-dependent way (van der Zand et al., 2010). Data
obtained in S. cerevisiae (Van der Zand et al. 2012), suggest that two types of vesicles are
formed from the ER, each containing a different subset of PMPs. These vesicles subsequently
fuse to form a nascent peroxisome. Although both models differ fundamentally, they are both in
line with the generally accepted view that pex3 and pex19 cells lack peroxisomal membrane
structures. Our current data together with recent data on pex3 cells (Knoops et al., 2014) however
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demonstrated that both models require adaptation. The observation that pre-peroxisomal vesicles
exist in yeast pex19 and pex3 cells and contain Pex3, the PMPs of the matrix protein docking
site, Pex13 and Pex14, as well as Pex8 imply that these proteins insert into peroxisomal
membranes without the assistance of Pex3 and Pex19.
Previously, it has been suggested that all PMPs (including Pex3) first sort to the ER and exit
from the ER in vesicles, a process that depends on Pex3 and Pex19 (Hoepfner et al., 2005; Tabak
et al., 2008; van der Zand et al., 2012). However, our data showed that in H. polymorpha Pex19
is not required for vesicle formation and sorting of Pex3. Moreover, we did not observe ER
accumulation of PMPs at the ER of pex19 atg1 cells.
The origin of the peroxisomal vesicles is still unknown. A possible model is that Pex13 and
Pex14 first insert in the ER and is incorporated in the peroxisomal membrane vesicles after
budding from this membrane. A similar pathway can be envisaged for Pex3. However, fusion of
Pex13/Pex14 and Pex3-containing vesicles is less likely as the structures may lack Pex1 and
Pex6 due to the absence of Pex26. Pex1 and Pex6 were proposed to be essential for pre-
peroxisome vesicle fusion (van der Zand et al., 2012). Hence, Pex13/Pex14 and Pex3 could also
be included in the same vesicular structure via one vesicle formation event from the ER.
On the other hand, the presence of Pex13 in the initial stages of their formation may facilitate
association of Pex14 and subsequent import of Pex8. This model is in line with previous studies
in human cells which showed that Pex13 is capable to insert into peroxisomal membranes
independent of Pex19 (Fransen et al., 2004). Also, the presence of Pex13 is essential for the
association of Pex14 with the peroxisomal membrane (Girzalsky et al., 1999). Possibly,
membrane bound Pex13 is sufficient for Pex14 sorting. Because it has been demonstrated that P.
pastoris Pex8 only requires PTS receptors and Pex14 for import, a vesicle containing Pex14 may
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be capable of inserting this peroxin (Ma et al., 2009). Clearly, additional work has to be done to
unravel the mechanisms of this novel organelle assembly pathway.
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Materials and methods
Strains and growth conditions
H. polymorpha strains used in this study are listed in Table 1. Cells were grown at 37°C, either
on YPD media (containing 1% yeast extract, 1% peptone and 1% glucose) or mineral media
(MM) (van Dijken et al., 1976) supplemented with 0.5% glucose or a mixture of 0.5 % methanol
and 0.05 % glycerol as carbon sources and 0.25% ammonium sulphate as nitrogen source, unless
indicated otherwise. If required, amino acids or uracil were added to a final concentration of 30
μg/ml. For growth on agar plates, the media were supplemented with 2% agar. For the selection
of resistant transformants, YPD plates containing 100 μg/ml zeocin (Invitrogen), 300 μg/ml
hygromycine B (Invitrogen) or 100 μg/ml nourseothricin (Werner Bioagents) were used. For
cloning purposes, Escherichia coli DH5α was used. Cells were grown at 37°C in LB media
supplemented with 100 μg/ml ampicillin or 50 μg/ml kanamycin, when required.
Molecular and biochemical techniques
Standard recombinant DNA techniques and transformation of H. polymorpha was performed by
electroporation as described previously (Faber et al., 1994). Cell extracts of TCA treated cells
were prepared for SDS-PAGE as detailed previously (Baerends et al., 2000). SDS-PAGE and
western blotting were performed by established methods. Equal amounts of protein were loaded
per lane and blots were probed with rabbit polyclonal antisera against Pex3, Pex11, Pex14 or
pyruvate carboxylase-1 (Pyc1). The mouse monoclonal antiserum against green fluorescent
protein (GFP; Santa Cruz Biotechnology, sc-9996) was used to detect mGFP-fusion proteins of
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Pex8, Pex10, Pex13, Pex26 and Pmp47. Secondary antibodies conjugated to horseradish
peroxidase were used for detection. Pyc1 was used as a loading control.
Construction of H. polymorpha strains
The plasmids and primers used in this study are listed in Tables 2 and 3.
Plasmid pHIPN-PEX14-mCherry was linearized and integrated in the PEX14 promoter of pex19
atg1 cells as described previously (Knoops et al., 2014). Plasmids pHIPZ-PEX8-mGFP
(pMCE4), pHIPZ-PEX10-mGFP (pMCE5), pHIPZ–PEX11-mGFP (pSEM02), pHIPZ–PEX13-
mGFP (pSEM03) and pHIPZ-PMP47-mGFP (pMCE7) were linearized and integrated in the
endogenous promoter regions of the pex19 atg1 strain producing Pex14-mCherry essentially as
described previously (Cepinska et al., 2011; Saraya et al., 2010). All integrations were confirmed
by PCR.
Construction of plasmids
Plasmid pSEM61 (pHIPZ-PEX3-mGFP) was obtained as follows: PCR was performed on
the plasmid containing PAMO-PEX3 (pSEM04) using the primers Pex3-F and Pex3-R. The PCR
product of 1329 bp was digested with HindIII and BglII and the resulting fragment was inserted
between the HindIII and BglII sites of pHIPZ-mGFP fusinator plasmid. The resulting plasmid of
6406 bp, designed as pSEM61, was linearized with BstBI and transformed in H. polymorpha
pex19 atg1 cells producing Pex14-mCherry.
To construct the plasmid pHIPZ18-PADH1-GFP-SKL, PCR was performed on genomic
DNA using the primers Adh1-F and Adh1-R. The PCR product was further digested with
HindIII and NotI and the resulting fragment was inserted between the HindIII and NotI sites of
125
pHIPZ4-GFP.SKL plasmid. The resulting plasmid of 5878 bp, was linearized with EcoRI and
transformed in H. polymorpha pex19 atg1 producing Pex14-mCherry and PAOX PEX19.
For the production of an N-terminal mGFP fusion to Pex26, overlap PCR fragments of
the PEX26-promoter, mGFP- and PEX26 gene were made as follows. The promoter of PEX26
was amplified from wild-type H. polymorpha cells using the 131007GFPpex26F1 and
131007GFPpex26R1 primers. The amplified PEX26 promoter product containing a 24 bp GFP at
the 3’-end; whose 5’-end was further digested with HindIII. The mGFP gene was amplified from
pHIPZ-Pex25-mGFP using the 131007GFPpex26F2 and 131007GFPpex26R2 primers giving a
761 bp fragment that overlapped 25 bp with the PEX26 promoter at the 5’-end and overlapped 25
bp with the N-terminus of PEX26 at the 3’-end. The PEX26 gene was amplified from wild type
H. polymorpha cells using the 131007GFPpex26F3 and 131007GFPpex26R3 primers giving a
1081 bp fragment that overlapped 25 bp with the C-terminus of mGFP at the 5’-end and
contained a SalI digestion site at the 3’-end. The three PCR fragments were mixed in a molar
ratio of 1:1:1 and used for an overlap PCR-reaction with the 131007GFPpex26F1 and
131007GFPpex26R3 primers. A 3112 bp fragment was isolated by gel-extraction and digested
with HindIII and SalI. This fragment was inserted into the pHIPZ4-DsRed-SKL vector that was
linearized by HindIII and SalI digestion and of which the 5824 bp backbone was isolated by gel
extraction. The final resultant plasmid was named pHIPZ4-PPEX26.GFP-PEX26, checked by
sequencing and linearized by SphI for integration into H. polymorpha WT, DsRed-SKL and
pex19 atg1 cells producing Pex14-mCherry.
Fluorescence microscopy
Images were captured using a Zeiss Axioskop50 fluorescence microscope (Carl Zeiss,
Sliedrecht, The Netherlands) using MetaVue software and a Princeton Instruments 1300Y digital
126
camera. The GFP signal was visualized with a 470⁄40 nm band pass excitation filter, a 495 nm
dichromatic mirror, and a 525⁄50 nm band-pass emission filter; mCherry fluorescence was
visualized with a 587/25 nm band pass excitation filter, a 605 nm dichromatic mirror, and a
647/70 nm band-pass emission filter. Live cell imaging was performed on a Zeiss Observer Z1
using Axiovision software and a Photometrics Coolsnap HQ2 digital camera. Cells were grown
on 1% agar containing growth medium and the temperature of the heating chamber XL was set at
37C. Three z –axis planes were acquired for each time interval using 0.5 sec exposure times for
both GFP and mCherry. Image analysis was carried out using ImageJ and figures were prepared
using Adobe Photoshop CS4.
Electron microscopy
H. polymorpha cells were fixed in 1.5% potassium permanganate, post stained with 0.5%
uranylacetate and embedded in Epon 812 (Serva, 21045). For morphological studies, ultrathin
sections were viewed in a Philips CM12 TEM. For electron tomography, serial sections of 150
nm thick were cut. The serial images of whole cells were stacked and aligned using MIDAS
(Kremer et al., 1996), after which individual cells could be scrutinized for peroxisomal remnants.
10 nm gold beads were layered on top of the serial sections and acted as fiducial markers for
electron tomography. Two single-axis tilt series, each containing 141 images with 1° tilt
increments, were acquired with a pixel size of 0.7 nm on a FEI Tecnai12 at 120 kV using the
SerialEM acquisition software (Mastronarde, 2005) and a cooled slow-scan charge-coupled
device camera (4k Eagle; FEI Company) in 2x2 binned mode. The tilt series were aligned and
reconstructed using the IMOD software package and analyzed using the AMIRA visualization
package (TGS Europe). To generate 3-dimensional surface rendered models in AMIRA, first,
127
masks of organelles were drawn manually and afterwards improved by nonlinear anisotropic
diffusion filtering followed by thresholding.
Cryosectioning and immuno-gold labelling
For immuno-gold labelling, cells were fixed in 3% glutaraldehyde in 0.1 M Na-cacodylate buffer
pH 7.2 for 1 h on ice, incubated subsequently in 0.4% Na-periodate (15 min) and 1% NH4Cl (15
min). Upon embedding in 2% gelatine in phosphate buffer pH 7.4, ~0.5 mm3 cubes were
infiltrated overnight in 2.3 M sucrose in the same buffer. Cryosections of 60 nm were cut using a
cryo diamond knife (Diatome) at -120 °C in a Reichert Ultracut. Sections were mounted on
carbon-coated formvar nickel grids. Gelatine was removed by incubating the grids for 30
minutes on 2% gelatine in phosphate buffer (pH 7.4) at 30°C. Pex14, Pex5 and alcohol oxidase
(AOX) were localized using polyclonal antibodies raised against these proteins and goat-anti-
rabbit antibodies conjugated to 10 nm gold (Aurion). Sections were stained with 2%
uranyloxalate (pH 7.0) for 10 minutes, briefly washed with distilled water and embedded in 0.5%
methylcellulose on ice for 10 min before viewing in the CM12 electron microscope (Slot and
Geuze, 2007).
128
Table 1. The H. polymorpha strains used in this study.
Strains Characteristics Reference
wild-type (WT) NCYC 495 leu1.1 (Saraya et al., 2011)
atg1 ATG1 deletion strain, leu 1.1 (Komduur et al., 2003)
pex19 PEX19 deletion strain, ura3 (Otzen et al., 2004)
pex19 PEX14-mCherry pex19 with pSEM01 This study
pex19 PEX3-mGFP pex19 with pSEM61 This study
pex19 atg1 PEX19 ATG1 double deletion strain Knoops et al., 2014
pex19 atg1 PEX14-mCherry pex19 atg1 with pSEM01 This study
pex19 atg1 PEX14-mCherry.
PEX3-mGFP
pex19 atg1 with pSEM01 and pSEM61 This study
pex19 atg1 PEX14-mCherry PEX8-
mGFP
pex19 atg1 with pSEM01 and pMCE4 This study
pex19 atg1 PEX14-mCherry
PEX10-mGFP
pex19 atg1 with pSEM01and pMCE5 This study
pex19 atg1 PEX14-mCherry
PEX11-mGFP
pex19 atg1 with pSEM01 and pSEM02 This study
pex19 atg1 PEX14-mCherry.
PEX13-mGFP
pex19 atg1 with pSEM01 and pSEM03 This study
pex19 atg1 PEX14-mCherry GFP-
PEX26
pex19 atg1 with pSEM01 and pHIPZ4-GFP-
PEX26
This Study
pex19 atg1 PEX14-mCherry.
PMP47-mGFP
pex19 atg1 with pSEM01 and pMCE7 This study
pex19 atg1.PEX14-mCherry
PAOXPEX19
pex19 atg1 with pSEM01 and pSEM05 This study
pex19 atg1 PEX14-mCherry PADH1-
GFP.SKL–PAOX EX19
pex19 atg1 with pSEM01, pSEM05 and
pHIPZ18-GFP.SKL
This study
WT DsRed-SKL PEX13-mGFP WT with pSEM03 This study
WT DsRed-SKL PEX8-mGFP WT.DsRed-SKL with pMCE4 (Cepinska et al., 2011)
WT DsRed-SKL PEX10-mGFP WT.DsRed-SKL with pMCE5 (Cepinska et al., 2011)
WT PMP47-mGFP WT with pMCE7 (Manivannan et al., 2013)
WT DsRed –SKL GFP-PEX26 WT.DsRed-SKL and pHIPZ4-GFP-PEX26 This study
129
Table 2. Plasmids used in this study
Plasmid
Characteristics Reference
pSEM01 pHIPN Plasmid containing C-terminal part of PEX14 fused to
mCherry; natR; amp
R
Knoops et al., 2014
pSEM02 Plasmid containing C-terminal part of PEX11 fused to mGFP;
zeoR; amp
R
Knoops et al., 2014
pSEM03 Plasmid containing C-terminal part of PEX13 fused to mGFP;
zeoR; amp
R
Knoops et al., 2014
pSEM05 pHIPH4 containing PEX19 under control of PAOX, HphR, ;
ampR
Knoops et al., 2014
pSEM61 Plasmid containing C-terminal part of PEX3 fused to mGFP;
zeoR; amp
R
This study
pHIPZ18-GFP.SKL Plasmid containing GFP.SKL under the promoter of ADH1,
zeoR; amp
R
This study
pHIPZ4-PPEX26.GFP-
PEX26
Plasmid containing N-terminal part of PEX26 fused to GFP
under control of the PEX26 promoter, ZeoR; amp
R
This study
pHIPZ4-mGFP
fusinator
pHIPZ containing mGFP and AMO terminator; ZeoR; amp
R (Saraya et al., 2010)
pMCE4 Plasmid containing C-terminal part of PEX8 fused to mGFP;
ZeoR; amp
R
(Cepinska et al., 2011)
pMCE5 Plasmid containing C-terminal part of PEX10 fused to GFP;
ZeoR; amp
R
(Cepinska et al., 2011)
pMCE7 Plasmid containing C-terminal part of PMP47 fused to GFP;
ZeoR; amp
R
(Cepinska et al., 2011)
130
Table 3. Primers used in this study
Primer Sequence (5’- 3’)
Pex3-F AAGAAAAAGCTTCTTTTTGGCACGGGAGTGAT
Pex3-R AAAAGATCTAGCATCGAAATTAGAGTAGACAC
Adh1-F AAGGAAAAAAGCGGCCGCCCCCTGCATTATTAATCACC
Adh1-R AATCAATCAATCAATTTAAAAAGCTTGGG
131007GFPpex26F1 ATCTCCAAGCTTGTGGCTATACGAGAAGCTAGGATGTC
131007GFPpex26F2 TTCCGGAAGCTGCTTGAGTCAAATGAGCAAGGGCGAGGAGCTGTTCACC
131007GFPpex26F3 TCTCGGCATGGACGAGCTGTACAAGTCTCAGATTACAGTCAACGGTTCAA
131007GFPpex26R1 GGTGAACAGCTCCTCGCCCTTGCTCATTTGACTCAAGCAGCTTCCGGAA
131007GFPpex26R2 TTGAACCGTTGACTGTAATCTGAGACTTGTACAGCTCGTCCATGCCGAGA
131007GFPpex26R3 TCTAGAGTCGACCTATATATAAGTGATTTTCAGTGCCA
131
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134
Chapter 5
The Hansenula polymorpha Wsc homolog is involved in peroxisomal
membrane shaping and organelle partitioning
Selvambigai Manivannan, Jan A.K.W. Kiel, Arjen M. Krikken, Rinse de Boer,
Marten Veenhuis and Ida. J. van der Klei*
Molecular Cell Biology, Groningen Biomolecular Sciences and Biotechnology Institute (GBB),
University of Groningen, Nijenborgh 7, 9747AG Groningen, The Netherlands
135
Abstract
We have analyzed the Hansenula polymorpha homologue of Neurospora crassa Wsc, which is
involved in Woronin body biogenesis and partitioning. We show that Wsc locates to
peroxisomes in H. polymorpha. Deletion of the WSC gene does not appear to have a significant
effect on peroxisome biogenesis, proliferation and function, when cells are grown on methanol.
However, it affects peroxisome inheritance in conjunction with a major influence on the overall
morphology of the peroxisomes, which are highly irregular of shape in glucose-grown H.
polymorpha wsc cells and contain different long tubular extensions. Together, our data are
consistent with the view that Wsc is important for shaping the peroxisomal membrane and for
segregation of the organelles. We have not observed any indications for pore forming properties
of Wsc, as proposed for its human counterpart, Pxmp2.
Keywords: Peroxisome, WSC, Pxmp2, yeast, organelle morphology
136
Introduction
Peroxisomes are eukaryotic organelles, which consist of a protein-rich matrix surrounded
by a single membrane. Peroxisomes are well known for their role in a large variety of metabolic
pathways. Common functions include decomposition of hydrogen peroxide and β-oxidation of
fatty acids. Species specific functions include the biosynthesis of plasmalogens and bile acids in
mammals (Wanders and Waterham, 2006), the metabolism of methanol in methylotrophic yeasts
(van der Klei et al., 2006) and the biosynthesis of penicillin in filamentous fungi (Bartoszewska
et al., 2011). A highly specialized peroxisome, called Woronin body, only occurs in filamentous
ascomycetes (Jedd and Chua, 2000). This unique organelle has no metabolic function, but plays
a role in closing septal pores to prevent cytoplasmic leakage upon hyphal damage (Jedd and
Chua, 2000).
The wide range of peroxisomal metabolic pathways requires continuous metabolite transport
between the peroxisomal matrix and cytosol and vice-versa. So far only a few peroxisomal
transporters have been identified. For instance, peroxisomes harbor ATP binding cassette (ABC)
transporters, which use ATP to actively transport CoA esters of fatty acids across the membrane
(Higgins, 1992; Palmieri et al., 2001; Hettema et al., 1996). This provides support for the notion
that the peroxisomal membrane is not permeable to all solutes.
The permeability properties of the peroxisomal membrane are still a matter of debate. Until
recently, two major models have been proposed. One of these prescribes that the peroxisomal
membrane contains pore forming proteins and hence is permeable to almost all solutes.
According to the second model, the peroxisomal membrane contains specific transporter proteins
137
for all metabolites and co-factors that have to be transported across the peroxisomal membrane
(Antonenkov et al., 2007; Visser et al., 2007).
Pmp22 is an abundant integral peroxisomal membrane protein, which was first identified in rat
liver (Koster et al., 1986; Hartl and Just, 1987). Later, the gene encoding this protein was
designated PXMP2 and its translation product Pxmp2 (Luers et al., 2001). Based on in vitro
assays and biochemical studies with purified Pxmp2, it was proposed that this protein is a
channel forming protein that enables free diffusion across the membrane for molecules with
molecular masses up to 300 Da (Rokka et al., 2009). This observation indicates that probably the
peroxisomal membrane is permeable for small molecules, but requires specific transporters for
larger ones.
Pxmp2 is member of a small gene family, which includes the S. cerevisiae mitochondrial inner
membrane protein Sym1/Ylr251w (Trott and Morano, 2004) and its mammalian homologue
MPV17 (Spinazzola et al., 2006) as well as YOR292C, a protein of unknown function. In a high
throughput screen Scer-YOR292C was localized to the vacuolar membrane (Huh et al., 2003).
However, there are serious doubts concerning this localization (Visser et al., 2007). A Pxmp2
family member designated Woronin Sorting Complex (Wsc) was shown to be an integral
component of the peroxisomal and Woronin body membrane in the filamentous fungus
Neurospora crassa (Liu et al., 2008). Interestingly, this protein is implicated in Woronin body
biogenesis and the positioning and segregation of these specialized organelles (Liu et al., 2008).
This observation suggests that Pxmp2 proteins not only fulfill a function in solute transport.
Yeast are favorable organisms to study peroxisome function and biogenesis. As indicated above,
proteins of the Pxmp2 family may play an important role in peroxisome biology. In order to
138
obtain further insights into this protein family, we first identified all Pxmp2 family members in
yeast and filamentous fungi. Interestingly, a homologue of the Wsc protein was identified in
several yeast species including Hansenula polymorpha, despite the fact that these organisms do
not contain Woronin bodies.
Here we show that Wsc is localized to peroxisomes in H. polymorpha. Wsc however, is not
involved in peroxisome biogenesis, proliferation or function. As Wsc shows homology to
mammalian Pxmp2, we also analyzed the effect of deletion of WSC on the growth of cells on
various carbon and nitrogen sources. However, we have not observed any defect in peroxisome
function, independent of the growth conditions. In fact, we show that in glucose-grown cells Wsc
in involved in maintaining normal peroxisome morphology and normal partition of peroxisomes
over mother cell and bud during vegetative cell reproduction. The details of this work are
contained in this contribution.
139
Results
Identification of Pxmp2 homologues in Hansenula polymorpha
First, we set out to identify Pxmp2 family members in yeast species and filamentous
fungi by searching the NCBI databases. The methodology used was similar to that described
previously (Kiel et al., 2006). The results of this in silico analysis are listed in Table 4. The
results clearly indicate that there are significant differences in the number (2 to 7 members) of
Pxmp2 related proteins in various ascomycete and basidiomycete fungi. All organisms studied
contain Sym1 and YOR292C orthologs. Remarkably, these two proteins are the sole members of
the Pxmp2 family in the related yeast species S. cerevisiae and Candida glabrata, while other
species of budding yeast contain up to 5 family members. It is well established that both S.
cerevisiae and C. glabrata have evolved from an ancestor yeast species that underwent whole
genome duplication followed by massive gene loss (Ochman et al., 2005).
With the exception of S. cerevisiae and C. glabrata, the Wsc-ortholog is conserved in all yeast
and filamentous fungi analyzed (Table 4). Wsc has a highly specialized role in the formation of
Woronin bodies from peroxisomes (Liu et al., 2008). Woronin bodies are only present in
filamentous ascomycetous fungi. It is therefore rather surprising that orthologous proteins can be
readily identified in yeast as diverse as K. lactis (a rather close relative of S. cerevisiae),
methylotrophic yeast species and even basidiomycetes. This suggests that the WSC protein may
have additional functions beyond those described for N. crassa.
An alignment of all 12 fungal WSC proteins shows that WSC proteins contain 4 regions
of high similarity (Fig. 1) Hydropathy analysis of the alignment using the MEMGEN program
140
Figure 1. Sequence alignment of fungal WSC orthologs. Sequences were aligned using the Clustal_X
programme. Gaps were introduced to maximize the similarity. Residues that are similar in all 12 proteins are
represented by white letters that are shaded black. Similar residues in at least 10 of the proteins are shown as white
letters that are shaded dark grey, while those that are similar in at least 7 of the proteins are shown as black letters
that are shaded light grey. The lines above the sequences represent potential hydrophobic regions identified with the
MEMGEN program. (Filamentous ascomycetes: Anid, Aspergillus nidulans; Pchr, Penicillium chrysogenum; Ncra,
Neurospora crassa. Yeast species: Klac, Kluyveromyces lactis; Calb, Candida albicans; Ssti, Scheffersomyces
141
stipitis; Hpol, Hansenula polymorpha; Ppas, Pichia pastoris; Ylip, Yarrowia lipolytica; Pans, Podospora anserina.
Basidiomycetes: Umay, Ustilago maydis; Scom; Schizophyllum commune).
suggests that each of these conserved regions contains a hydrophobic motif that might constitute
a membrane spanning domain. A phylogenetic tree of these proteins including also Scer-Sym1
and S. cer-Yor292c is shown in Figure 2. Remarkably, the Wsc proteins have highly diverged
from S. cerevisiae-Sym1 and YOR292C and their orthologous.
Figure 2. Phylogenetic tree of Pxmp2 family proteins in S. cerevisiae, H. polymorpha and N. crassa. The tree
was constructed with TREECON for Windows using protein sequences aligned with Clustal-X. The distance scale
represents the number of differences between the sequences with 0.1 indicating a 10 % difference. (Scer,
Saccharomyces cerevisiea; Hpol, Hansenula polymorpha; Ncra, Neurospora crassa).
In order to further study these proteins, we analyzed the localization and function of the
putative mitochondrial protein Sym1, the putative vacuolar protein Yor292c as well as the WSC
homolog.
142
H. polymorpha Wsc is a peroxisomal protein
In order to localize three Pxmp2 family members in H. polymorpha, we constructed
hybrid genes encoding C-terminal mGFP fusions of Wsc, Sym1 and Yor292c. The fusion
proteins were produced under control of their endogenous promoter in strains producing the red
fluorescent peroxisomal matrix marker DsRed-SKL. Fluorescence microscopy of cells grown on
glucose medium for 4 hours revealed that Wsc-mGFP appeared as a spot, which co-localized
with DsRed-SKL (Fig 3A). Also, after 16 hours of cultivation in methanol-containing media
Wsc-mGFP co-localized with DsRed-SKL on peroxisomes (Fig 3B).
Figure 3. Localization of Wsc-mGFP in H. polymorpha. Fluorescence microscopy images of H. polymorpha cells
expressing Wsc-mGFP together with PTEF-driven DsRed-SKL (A) or PAOX-driven DsRed-SKL (B), grown for 4
hours on glucose (A) or 16 hours on methanol (B). The bar represents 1µm.
143
Analyses of glucose- or methanol-grown cells producing Sym1-mGFP or Yor292c-mGFP
revealed that the levels of both proteins were too low to determine their localization by
fluorescence microscopy (data not shown). The low levels of these proteins were confirmed by
western blot analysis using anti-GP antibodies (data not shown).
The levels of the fusion protein Wsc-mGFP were constant during exponential growth on glucose
(tested 2 and 4 hours after shifting glucose-grown cells to fresh glucose containing media; Fig
4A). When the glucose-grown cells were shifted to media containing methanol, the Wsc-mGFP
fusion protein levels remained similar during adaptation to methanol conditions until 16 hours;
Fig 4B), indicating that the WSC gene is constitutively expressed at peroxisome repressing and
inducing growth conditions.
Figure 4. Wsc is constitutively produced on glucose and methanol. Cells producing Wsc-mGFP were extensively
precultivated on glucose and subsequently shifted to glucose (A) or methanol (B) containing media. Samples were
taken at regular intervals after the shift. Blots were prepared from crude cell extracts and decorated using anti-GFP
antibodies. Equal amounts of protein were loaded per lane. Pyruvate carboxylase (Pyc1) was used as a loading
control.
Properties of the H. polymorpha WSC deletion strain.
We next addressed whether H. polymorpha Wsc plays a role in peroxisome biogenesis
and function using a WSC deletion strain (wsc). Defects in peroxisome biogenesis invariably
144
result in a deficiency of cells to grow on methanol (van der Klei et al., 2006). Growth
experiments (Fig. 5A) revealed that wsc cells grew normal on methanol as the wild-type control.
Also, similar final densities (OD660) were obtained.
Figure 5. (A) wsc cells show normal growth and peroxisome proliferation on methanol containing medium. A.
Growth curve of wsc and WT control cells in methanol medium. Cells were extensively pre-cultivated in media
containing 0.5% glucose and shifted to medium containing 0.5% methanol. The optical densities (OD) are expressed
as adsorption at 660 nm. (B) Fluorescence microscopy analysis of wsc cells producing the peroxisomal membrane
marker Pmp47-mGFP. Cells were grown for at 16 hours in methanol medium. The bar represents 1µm.
145
For growth on methanol, several metabolites have to be transported across the
peroxisomal membrane. These include formaldehyde, dihydroxyacetone, glyceraldehyde-3-
phosphate and xylulose-5-phosphate. Apparently, Wsc is not required for the transport/diffusion
of these molecules across the peroxisomal membrane. We also tested several other carbon
(ethanol) and nitrogen sources (methylamine, D-choline, D-alanine) that are (partially)
metabolized by peroxisome borne enzymes. However, invariably no significant differences in
growth properties were observed compared to the wild-type control (data not shown). Together,
this makes a role of Wsc in metabolite transport unlikely.
wsc cells show reduced peroxisome abundance and a partial segregation defect, when
grown on glucose, but not upon growth on methanol.
As in N. crassa WSC is involved in formation of a Woronin body from a peroxisome, H.
polymorpha Wsc may play a role in peroxisome proliferation. To investigate this, we constructed
a H. polymorpha wsc strain, which also produced the fluorescent peroxisomal membrane marker
PMP47-GFP. Cells were grown for 16 h on methanol and analyzed by FM (Fig. 5B). The data
revealed that the number of peroxisomes in wsc cells (average number of 3.2 peroxisomes per
cell) was comparable to those of the wild-type control (average 3.3 peroxisomes per cell). Hence,
Wsc does not appear to have a role in peroxisome abundance during growth on methanol. On
glucose, however, a significant difference in organelle numbers was observed, with an average
number of 0.9 in wsc cells relative to 1.3 in WT controls (Fig. 6). Subsequently, electron
microscopy analysis revealed that the peroxisomes in glucose-grown H .polymorpha wsc cells
were strongly irregular of shape relative to WT organelles and contained distinct extensions as
confirmed by the inspection of serial sections (Fig. 6C).
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Figure 6. Peroxisome distribution over mother cells and buds. Survey of wsc (A) and WT (B) cells grown on
glucose showing the differences of distribution of peroxisomes over mother cells and bud. Peroxisomes are marked
by GFP-SKL. (C) Serial sections of a KMnO4-fixed glucose-grown wsc cell showing tubular extensions
(arrowheads) at the peroxisome and an irregular organelle shape. Sclae bars: A and B - 5 µm; C – 500 nm. P –
peroxisome, M – mitochondrion, ER – endoplasmic reticulum.
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Budding WT cells generally contain two peroxisomes of which one is retained in the mother and
one transported to the bud, whereas budding glucose-grown wsc cells often contained a single
peroxisome. This organelle irregularly distributed over mother cell and bud in glucose-grown
cells (Fig. 7A-C: compare also surveys of Fig. 6A, B). This phenotype is different from that
observed in inp1 cells, which show a peroxisome retention defect, resulting in transport of bulk
of the organelles to the bud.
Discussion
We identified a homolog of N. crassa WSC in the yeast H. polymorpha that is localized to
peroxisomes. Before, WSC has been implicated in the formation and partitioning of Woronin
bodies that derive from peroxisomes. We could not find evidence in support of a role of H.
polymorpha WSC in peroxisome proliferation. In N. crassa WSC also is involved in Woronin
body inheritance via cortical association. During yeast fission, one or a few organelles segregate
to the developing bud, whereas most organelles are retained in the mother cell, via cortical
association. So far, Inp1 and Pex3 (Fagarasanu et al., 2005; Munck et al., 2009) have been
implicated in peroxisome retention in yeast mother cells, whereas Inp2 and Myo2 are required
for transport of the organelle to the bud (Motley and Hettema, 2007; Saraya et al., 2010). We
here show that Wsc does not play a role in this process. Mammalian Pxmp2 shows homology to
Wsc and has been implicated in solute transport. We showed that deletion of the WSC gene does
not influence growth on methanol or ethanol containing media. Also, the metabolism of D-amino
acids, D-choline or methylamine by peroxisomal oxidases was not defective in a WSC deletion
strain indicating that a role of Wsc in solute transport is very unlikely. Studies in mice also
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Figure 7. (A) Organelle quantification. Quantification (from Z-stack images) was carried out in budding cells for the
presence or absence of peroxisomes in the mother and daughter cell. Peroxisomes from 2 X 100 cells were counted
from two independent experiments. The bar represents standard error of mean (SEM). *, P < 0.05. (B, C) Selected
images were taken from time-lapse series of wsc budding cells revealed that peroxisomes either migrate into the
daughter cell at an early stage of bud formation, leaving the mother cell devoid of these structures (B) or stay in the
mother cell (C). Also, cells showing normal segregation like in WT cells can be observed (C). Peroxisomes are
marked by GFP–SKL. The bar represents 2 µm.
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showed no clear phenotype upon the deletion of PXMP2 and only a partial restriction of
membrane permeability to uric acid (Rokka et al., 2009).
Taken together, our data support a function of Wsc in maintaining the morphology of the
peroxisomal membrane. A similar function has been proposed for mitochondrial Sym1, i.e. in
maintaining the shape of the mitochondrial inner membrane in baker’s yeast (Dallabona et al.,
2010). We speculate that the aberrations in peroxisome partitioning are related to their unusual
morphology. However, the principles of this are unknown and require further investigations.
Clearly, the functions of Wsc and their homologues may largely differ, dependent on species and
subcellular location.
150
Materials and Methods
Organisms and growth
H. polymorpha strains used in this study are listed in Table I. Cells were grown at 37°C using
YPD (1% yeast extract, 1% peptone, 1% glucose) or mineral medium (van Dijken et al., 1976).
For microscopic and growth rate analysis, cells were extensively precultivated on mineral
medium containing 0.25% ammonium sulphate, methylamine, D-alanine or D-choline as sole
nitrogen sources and 0.5% glucose as carbon source. Exponential cultures were subsequently
shifted to media containing 0.5% methanol as carbon source and the nitrogen source that was
used in the preculture. When required, media were supplemented with 30 µg/ml leucine. For the
selection of transformants YPD plates contained 100 µg/ml nourseothricin or zeocin. For cloning
purposes, Escherichia coli DH5α was used as host for propagation of plasmids using Luria Broth
supplemented with appropriate antibiotics (100 µg/ml) and grown at 37°C.
Molecular and biochemical techniques
Plasmids used in this study are listed in Table II. Standard recombinant DNA techniques and
transformation of H. polymorpha was performed as described previously (Faber et al., 1994).
Cell extracts of trichloroacetic acid treated cells were prepared for sodium dodecyl sulfate
polyacrylamide gel electrophoresis as detailed previously (Baerends et al., 2000). Equal amounts
of protein were loaded per lane, followed by western blot analysis (Laemmli, 1970). Blots were
probed with rabbit polyclonal antiserum against pyruvate carboxylase (Pyc1) or mouse
monoclonal antiserum against GFP (Santa Cruz Biotechnology, sc-9996).
151
Plasmid constructions
For the construction of plasmid pSEM060, we performed PCR on the H. polymorpha WSC
(Hp32g403) genomic region using the primers P1 and P2. The obtained PCR fragment of 666 bp
encompassing the WSC ORF lacking a stop codon was digested with HindIII and BglII and
inserted between the HindIII and BglII sites of the pHIPZ-mGFP fusinator plasmid. The resulting
plasmid containing a WSC-mGFP fusion gene, designated as pSEM060, was linearized with
PflMI and integrated in the H. polymorpha genome resulting in expression of the fusion gene
under control of its endogenous promoter. H. polymorpha DsRed-SKL cells were used which
produce DsRed-SKL as a red-fluorescent peroxisomal matrix marker. The correct integration
was confirmed by PCR.
Construction of Gateway plasmids
A H. polymorpha WSC (Hp32g403) deletion strain was constructed by replacing the middle
portion of Hp32g403 genomic region comprising nucleotides +1659 to +2008 by the antibiotic
marker Hygromycine (Hph). To this end, pSEM027 [pDest-WSC (Hp32g403) deletion cassette]
was constructed using Gateway Technology (Invitrogen). By using H. polymorpha genomic
DNA as a template, two DNA fragments comprising the regions −1231 to +1658 and +2008 bp
to +2408 of the WSC genomic region were obtained by PCR using primers Fwd attB4/ Rev attb1
and Fwd attB2/ Rev attB3, respectively. The PCR fragments were recombined into the vectors
pDONR-P4-1R and pDONR-P2R-P3, respectively, resulting in the entry vectors pENTR-WSC
5′ and pENTR-WSC 3′. Recombination of the entry vectors pENTR-WSC 5′, pENTR-221-HPH,
and pENTR-WSC 3′, and the destination vector pDEST-R4-R3, resulted in pSEM027. A 2.6 kb
fragment of pSEM027 comprising the WSC deletion cassette was amplified by PCR with the
152
primers WSC del. fwd and WSC del. Rev. The amplified fragment was transformed into H.
polymorpha cells producing PMP47-GFP as a peroxisomal membrane marker. The integrations
and deletions were confirmed by PCR and Southern blot analysis.
Fluorescence microscopy.
Wide-field images were made using a Zeiss Axioscok50 fluorescence microscope (Carl Zeiss,
Sliedrecht, The Netherlands). Images were taken using a Princeton Instruments 1300Y digital
camera. The GFP signal was visualized by using a 470/40 nm bandpass excitation filter, a 495
nm dichromatic mirror and a 525/50 nm bandpass emission filter. DsRed fluorescence was
visualized with a 546/12 nm bandpass excitation filter, a 560 nm dichromatic mirror and a
575/640 nm bandpass emission filter.
Confocal imaging was performed on a Zeiss LSM510 confocal microscope, using Hamamatsu
photomultiplier tubes. GFP signal was visualized by excitation with a 488 nm argon laser
(Lasos), and emission was detected using a 500–550 nm bandpass emission filter. The DsRed
signal was visualized by excitation with a 543 nm helium neon laser and emission was detected
using a 565–615 nm bandpass emission filter. For live cell imaging, the temperature of the
objective and the object slide was kept at 37 °C. Five Z-axis planes were acquired for each time
interval to ensure that no fluorescent structures were missed. Image analysis was carried out
using ImageJ and Adobe Photoshop.
Quantification of peroxisomes
To quantify peroxisome inheritance, random pictures were taken as a stack in both bright field
and fluorescence mode. Z-stacks were made containing 12 optical slices of 0.9 μm thickness to
153
cover the entire cell. The Z-axis spacing was 0.5 μm, to ensure that no fluorescent signals were
missed. Using the Zeiss LSM Image Browser software, the cross-sectional area of the mother
and bud cell was determined. Assuming yeast cells to be spherical, the bud volume was
determined as percentage of that of the mother cell, tentatively set to 100%. Only cells for which
the bud volume was < 25% of the mother cell volume were counted. Quantification experiments
were performed using two independent cell cultures (100 cells per culture). To quantify the
average peroxisome number per cell, the same random pictures were used. The peroxisome
number per cell was quantified by counting the amount of fluorescent spot per cell only in single
non-budding cells. Statistical differences were determined by using student T-test.
Electron microscopy
H. polymorpha cells were fixed in 1.5% potassium permanganate, post stained with 0.5%
uranylacetate and embedded in Epon 812 (Serva, 21045). For morphological studies, ultrathin
sections were viewed in a Philips CM12 TEM.
In silico analysis
For the identification of homologs of the mammalian PMP22/MPV17 proteins in yeast
species and other fungi, we initially used the primary sequence of the Neurospora crassa WSC
protein (Genbank accession number EAA33867) as query in BlastP analyses (Altschul et al.,
1997) on the fungal dataset of the non-redundant (nr) protein database (Taxid: 4751) at the
National Center for Biotechnology Information (NCBI). During these screenings only proteins
from representative yeast and fungal species (listed in Table 4) were set apart for further
154
analyses. As expected, since WSC is a member of the Pxmp2 protein family, in addition to true
WSC orthologs many other homologous proteins were identified.
In the next step, reciprocal BlastP analyses were carried out using all the identified sequences as
queries. When these analyses resulted in highest similarity to the set of top scoring protein
sequences initially identified with the N. crassa WSC sequence, a protein sequence was judged
to be its ortholog. In a similar way orthologs to other Pxmp2 family proteins were identified.
These analyses also identified additional protein sequences that were absent in the initial screen
because of weak similarity to the N. crassa WSC sequence. These newly identified protein
sequences were also used as queries in reciprocal Blast analyses.
The BlastP screenings might not have resulted in identification of all putative homologues from a
specific fungal species present in the nr protein database, because the sequences were too
divergent to be identified. In such instances, the already identified WSC-related sequences were
used as queries in Position Specific Iterated (PSI)-Blast analyses (Altschul et al., 1997) on the
fungal dataset of the nr protein database. A statistical significance value of 0.001 was used as a
threshold for the inclusion of homologous sequences identified by the PSI-BLAST analysis on
iteration.
When these analyses had still not resulted in identification of all expected orthologous/
paralogous sequences in a specific organism, the identified protein sequences from a closely
related species were used as queries to search the fungal genome database using TBlastN
analyses (Altschul et al., 1997). In two cases we were able to identify a DNA sequence encoding
a MPV17/PMP22-family member in the fungal genome database, whilst its translation product
155
was absent in the protein database. This was caused by the presence of introns that had interfered
with the proper identification of coding sequences in the fungal genome.
Identification of MPV17/PMP22 members in the Hansenula polymorpha database
In addition to identifying fungal WSC-related sequences in the nr protein database at the NCBI,
we also searched the H. polymorpha genome sequence (Ramezani-Rad et al., 2003). TBlastN
searches were performed as described above using the N. crassa WSC sequence as well as all
identified MPV17/PMP22-related sequences from more closely related organisms (Pichia
pastoris, Yarrowia lipolytica, Candida albicans and Aspergillus nidulans) as queries. This
method enabled the identification of a putative H. polymorpha WSC ortholog as well as 3
paralogous proteins, which was again confirmed by reciprocal BlastP analyses.
Other methods
For DNA sequence analysis, the Clone Manager 5 program (Scientific and Educational Software,
Durham, USA) was used. The Clustal_X program (Thompson et al., 1997) was used to align
protein sequences. The GeneDoc program (available at http://www.psc.edu/biomed/genedoc)
was used to display the aligned sequences. TREECON for Windows (Van de Peer and De
Wachter, 1994) was used for the construction of phylogenetic trees. MEMGEN version 4.08 was
used to localize conserved hydrophobic regions in protein alignments (Lolkema and Slotboom,
1998).
156
Table I. Yeast strains used this study
Strains Characteristics Reference
Wild-type (WT) NCYC 495 leu1.1 (Sudbery et al., 1988)
WT. DsRed-SKL WT cells with integration of pSNA13 This study
WT. DsRed-SKL.WSC-GFP WT.DsRed-SKL with pSEM060 This Study
WT. PTEF. DsRed-SKL WT cells with plasmid containing PTEF DsRed-
SKL
This study
WT. PTEF. DsRed-SKL.WSC-
mGFP
WT. PTEF. DsRed-SKL with pSEM060 This Study
PMP47-GFP WT Ppmp47 PMP47-GFP (Manivannan et al., 2013)
wsc. PMP47-GFP WSC deletion with Ppmp47 PMP47-GFP This study
WT.GFP-SKL WT cells integrated with the plasmid containing
PTEF.GFP-SKL
(Krikken et al., 2009)
inp1.GFP-SKL INP1 deletion with PTEF.GFP-SKL (Krikken et al., 2009)
wsc.GFP-SKL WSC deletion with PTEF.GFP-SKL This study
157
Table II. Plasmids used in this study.
Plasmid Description Reference
pSNA13 Plasmid containing PAOX DsRed-SKL, ampR; nat
R (Cepinska et al., 2011)
pHIPZ-mGFP fusinator pHIPZ containing mGFP and AMO terminator; ZeoR;
ampR
(Saraya et al., 2011a)
pSEM060 Plasmid containing C-terminal part of WSC fused to
GFP; zeoR; amp
R
This study
pHIPX7. DsRed-SKL Plasmid containing PTEF DsRed-SKL, ampR; Leu
R (Krikken et al., 2009)
pDONOR-221 Standard gateway vector Invitrogen
pDONR-P4-P1R Standard Gateway vector Invitrogen
pDONR-P2-R3 Standard Gateway vector Invitrogen
pENTR-221-HPH pENTR-221 containing hygromycin marker, kanR (Saraya et al., 2011b)
pDEST-R4-R3-NAT pDEST-R4-R3-containing nourseothricin marker,
ampR; nat
R
(Nagotu et al., 2008)
pENTR-23-TAMO pENTR-23 containing terminator of amine oxidase,
kanR
(Nagotu et al., 2008)
pENTR-41-WSC 5’ pDONOR-P4-P1 containing 5’region of WSC, kanR This study
pENTR-23- WSC 3’ pDONOR-P2R-P3 containing 3’region of WSC, kanR This study
pSEM027 pDEST-R4-R3 containing WSC deletion cassette,
Hygromycine B marker, ampR
This study
PTEF. GFP-SKL Plasmid containing GFP-SKL under the control of
TEF promoter, kanR; Leu
R
(Krikken et al., 2009)
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Table III. Primers used in this study
Primer Name Sequence
P1 5’ AAAAAGCTTATGCTCGCCGATCTGAAC 3’
P2 5’ TTTAGATCTTTCATTCTTGTTCTGTTC 3’
Fwd attB4 5’GGGGACAACTTTGTATAGAAAAGTTGCCGCTCCGCCTCTTGGTGCTCCTCTAA3’
Rev attb1 5’GGGGACTGCTTTTTTGTACAAACTTGGCAAAGGGACGCGTTTTGTGACAGAG3’
Fwd attB2 5’GGGGACAGCTTTCTTGTACAAAGTGGCCACCAGTGGGCCGTGTTCTTC3’
Rev attB3 5’GGGGACAACTTTGTATAATAAAGTTGCGTGGACAAGGGCCGTCATAAACTGT3’
WSC del. Fwd 5’GCTCCGCCTCTTGGTGCTCCTCTAA3’
WSC del. Rev 5’GTGGACAAGGGCCGTCATAAACTGT3’
159
Table 4. Proteins of the MPV17/PMP22 family in ascomycete and basidiomycete fungi
Ascomycetes Basidiomycetes
Scer Cgla Klac Calb Ssti Hpol Ppas Ylip Anid Pchr Ncra Pans Umay Scom
WSC -- -- CAH00328 EAK98770 ABN64847 Hp32g403 CAY72077 CAG78284 CBF78752 CAP96827 EAA33867 CAP69185 EAK81496 EFJ02339
(WSC)
SYM1 NP_013352 CAG58022 CAG98810 EAK92584 ABN66872 Hp27g68 CAY69640 CAG82519 CBF82760 CAP96258 EAA34618 DNA1 EAK84312 EFJ03404
(SYM1)
Paralog1 -- -- -- EAK92585 ABN66871 -- -- -- -- -- -- -- -- --
Paralog2 -- -- -- -- -- -- -- -- -- -- -- -- -- EFJ00581
Unknown 1 -- -- -- -- -- -- -- -- CBF88890 CAP93899 EAA32569 CAP71852 -- --
Unknown 2 -- -- -- -- -- -- -- -- CBF71125 -- EAA36527 CAP67591 -- --
Paralog -- -- -- -- -- -- -- -- DNA2 CAP99639 -- -- -- --
Unknown 3 -- -- -- -- -- -- -- -- CBF87341 -- -- -- -- --
YOR292c NP_014935 CAG62813 CAH02288 EAK92072 EAZ64003 Hp24g381 CAY72196 CAG82751 CBF87860 CAP93144 EAA33195 CAP65570 EAK84020 EFJ02210
(YOR292c)
Unknown 4 -- -- -- EAK97649 ABN67061 Hp32g332 CAY70136 CAG79472 -- -- -- -- -- --
Key: The following organisms were used: Yeast species: Scer, Saccharomyces cerevisiea; Cgla, Candida glabrata; Klac, Kluyveromyces lactis; Calb, Candida
albicans; Ssti, Scheffersomyces stipitis; Hpol, Hansenula polymorpha; Ppas, Pichia pastoris; Ylip, Yarrowia lipolytica; Filamentous ascomycetes: Anid,
Aspergillus nidulans; Pchr, Penicillium chrysogenum; Ncra, Neurospora crassa; Pans, Podospora anserina; Basidiomycetes: Umay, Ustilago maydis; Scom;
Schizophyllum commune. DNA1, P. anserina accession number NW_001914859, nt 805177-804555 (with 1 intron); DNA2, A. nidulans accession number
NT_107003, nt 348404-349322 (with 3 introns).
160
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Summary
The term “cell” was coined by Robert Hooke and Antonie van Leeuwenhoek, which is the basic
unit of life. Organisms can be subdivided into three major domains, the bacteria, archaea and
eukaryotes. Generally, eukaryotes are characterized by the presence of a distinct double
membrane bound nucleus, which contains DNA, together with other internal membrane bound
compartments, called organelles. There are two categories of eukaryote organelles namely
autonomous ones (for instance mitochondria and chloroplasts) and the non-autonomous
organelles (e.g. the Golgi apparatus and vacuoles). It is generally accepted that autonomous
organelles multiply by growth and division and therefore cannot be generated de novo, whereas
the others all can derive from the endoplasmic reticulum. The peroxisome is one of the most
recently identified eukaryotic organelles. Peroxisomes contain a proteinaceous matrix, but lack
DNA, and are bound by a single membrane. These organelles are highly dynamic; their function,
morphology and abundance depend on the species in which they occur as well as on the
developmental stage and external stimuli. Peroxisomes are characterized by the presence of at
least one H2O2 producing oxidase in conjunction with catalase. Generalized functions include β-
oxidation of fatty acids and H2O2 metabolism.
Peroxisome biogenesis is governed by a number of specific proteins called “peroxins”, which are
encoded by PEX genes. Peroxisomal matrix proteins are synthesized by free ribosomes in the
cytosol and post-translationally imported into their target organelle. However, it is still debated
whether peroxisomal membrane proteins (PMPs) are directly inserted into the peroxisomal
membrane or traffic via the endoplasmic reticulum (ER).
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The formation of peroxisomes is yet an ardently debated topic. The classical view states that
peroxisomes are autonomous organelles that multiply by growth and fission. Recently, this is
defied by a second model which prescribes that peroxisomes are formed de novo from the ER.
This de novo formation was particularly observed in peroxisome-deficient mutants (pex3 and
pex19) that were thought to fully lack peroxisomal membrane remnants upon functional
complementation with their corresponding deleted genes. The most recent model describes that
de novo peroxisome formation initiates with the formation of two types of vesicles that derive
from the ER and carry different PMPs. These vesicles fuse, dependent of Pex1 and Pex6, to form
a pre-peroxisome that subsequently matures by the import of matrix proteins. It is very likely that
at least few PMPs travel via the ER. Together this would classify peroxisomes as semi-
autonomous organelles.
A proper balance between organelle biogenesis and degradation is important for peroxisome
homeostasis. Housekeeping processes exist that control repair or removal of redundant
peroxisomes via autophagy to maintain a vital peroxisome population.
Peroxisome biogenesis and degradation are highly conserved mechanisms in yeast. The different
stages of the peroxisome life cycle can be precisely manipulated by changing the growth
conditions. Hence, yeasts are attractive model organisms to study peroxisome homeostasis. The
work described in this thesis, mainly addresses the molecular mechanisms involved in de novo
peroxisome formation and organelle maintenance, using the methylotrophic yeast Hansenula
polymorpha, as a model organism.
Chapter 1 summarizes our current knowledge on peroxisome formation, the molecular
machineries involved in peroxisome fission and de novo synthesis from the ER, as well as
mechanisms of peroxisome quality control and their implications in cellular ageing.
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In Chapter 2 experiments are described that aim to elucidate whether peroxisome fission is
important for its degradation by autophagy. Our data showed that peroxisome degradation is
impaired in peroxisome fission mutants of Saccharomyces cerevisiae and H. polymorpha
(∆dnm1 and ∆pex11), like in the ∆atg1 (autophagy mutant) control strain. Subsequently, we
studied the significance of peroxisome fission and autophagy in removing aberrant peroxisomes.
To this purpose, we cloned a mutant form of catalase that contains a peroxisomal targeting signal
(Cat-Y348G.SKL) and at the same time easily forms aggregates in the peroxisomal lumen. We
produced these catalase protein aggregates in peroxisomes of H. polymorpha ∆atg1 cells and
tested whether this resulted in the induction of peroxisome fission. Using fluorescence and
electron microscopy, we observed the presence of protein aggregates in ∆atg1 cells, which
appeared to promote asymmetric peroxisome fission. This resulted in cells with numerous small
aggregate containing organelles, which did bud off from the mother organelle. Further, we
investigated whether these aggregates containing organelles are subjected to autophagic
degradation. Our electron microscopic data revealed that peroxisomes with protein aggregates
are often delivered to the vacuole for degradation in Cat-Y348G.SKL cells. However, the fission
mutants (∆dnm1 and ∆pex11) failed to remove these aggregate containing peroxisomes.
Additionally, we observed that the presence of protein aggregates affects the growth rate and
resulted in enhanced ROS levels. Hence, the timely removal of peroxisome with aberrant protein
aggregates is crucial. Consequently, our findings suggest that peroxisome fission and autophagic
degradation are coupled processes, which are essential for the removal of harmful components
from the organelles to prevent ROS induced cell death and maintaining organelle viability.
We demonstrated that Pex3 (Chapter 3) and Pex19 (Chapter 4) are not essential for the
formation of peroxisomal membranes. According to previous reports, deletion of PEX3 or
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PEX19 results in a complete lack of peroxisomal membrane structures and de novo formation of
peroxisomes from the ER occurs upon functional complementation with the corresponding
genes. In contrast, our data showed that H. polymorpha pex3 and pex19 cells do contain
peroxisomal remnants, which are however unstable and rapidly subjected to autophagic
degradation. This probably explains why they have been overlooked so long. When we blocked
autophagy in pex3 or pex19 cells via ATG1 deletion the peroxisomal membrane vesicles were
stabilized (i.e. in pex3 atg1 or pex19 atg1 cells). Additionally, our data showed that these
vesicles contained the PMPs Pex13 and, Pex14 as well as the matrix protein Pex8, whereas other
PMPs like Pex10, Pex11, Pmp47 and Pex26 (in pex19 atg1) are not targeted to the vesicles.
Moreover, the levels of these proteins were often near or below the limit of detection. We further
showed that Pex10, Pex11 and Pmp47 levels were enhanced and stabilized upon the
overexpression of Pex19 in pex3 atg1 cells. This suggests that these proteins depend on the
Pex3/Pex19 PMP insertion machinery, whereas Pex14 and Pex13 do not require these peroxins
for proper sorting.
Recent reports state that deletion of PEX19 results in the accumulation of Pex3 at the ER.
Conversely, we did not observe ER accumulation of Pex3 in pex19 atg1 cells; instead Pex3 was
present in the Pex14 containing vesicles in pex19 cells, or sorted to these structures upon
reintroduction of Pex3 in pex3 atg1 cells.
Subsequently, we investigated whether the Pex14-containing membrane vesicles are capable to
develop into peroxisomes. Using time-lapse videos, we showed that Pex3, Pex10 and Pmp47 are
targeted directly (independent of ER) to the Pex14 containing vesicles and mature in to
functional peroxisomes after PEX3 reintroduction in pex3 atg1 cells (Chapter 3). Hence, it is
clear from these results that the PMPs are not trafficking via ER to peroxisomes. Moreover, we
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showed that the Pex14 containing vesicles of pex3 atg1 and pex19 atg1 cells develop into
peroxisomes by importing matrix proteins (GFP-SKL) upon reintroduction of PEX3 or PEX19,
respectively (Chapter3 and 4).
Altogether, our findings challenge the most recent model which states that Pex3 and Pex19 are
required for the exit of PMPs containing pre-peroxisomal vesicles from ER. Conversely, our data
propose that i) Pex3 and Pex19 are not required for formation of vesicles from the ER; ii) the
existing vesicles in pex3/pex19 atg1 cells serves as template for peroxisome formation upon
reintroduction of the corresponding genes and not the ER; and iii) the PMPs Pex13 and Pex14
do not require the Pex3/Pex19 sorting machinery.
Finally, in Chapter 5 we investigated the localization and function of putative H. polymorpha
orthologs of the PMP22/MPV17 family of proteins (WSC, SYM1, and YOR292C). Using in
silico analysis, we identified 4 putative homologs in H. polymorpha. To localize these proteins,
we constructed C-terminal mGFP fusions of Wsc, Sym1 and Yor292c, and produced these in H.
polymorpha wild-type cells. Our fluorescence microscopy analysis showed that Wsc is localized
to the peroxisome. Reports state that Sym1 (yeast and mammals) is a mitochondrial inner
membrane protein and Yor292cp is a component of vacuolar membrane (mammals).
Nevertheless, we could not determine their exact localization since the protein levels were below
the limit of detection. Subsequently, we addressed the role of Wsc in H. polymorpha by making
WSC deletion strain. According to previous reports, Wsc is involved in Woronin body biogenesis
in the filamentous fungus Neurospora crassa and in transport of solutes in mammalian
peroxisomes. Our data showed that deletion of WSC does not influence growth on various carbon
and nitrogen sources. Also, this did not result in defects in peroxisome formation or proliferation
when cells were grown on methanol. However, glucose grown Δwsc cells showed reduced
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peroxisome abundance and irregular shaped organelles. Additionally, time-lapse video studies
demonstrated that single organelles present in glucose grown ∆wsc cells were frequently
distributed to the bud thereby leaving the mother cells devoid of peroxisomes. In other cells the
organelle remained in the mother cell during cell budding, resulting in a bud without a
peroxisome. This interesting finding indicates that glucose grown Δwsc cells have a partial
defect in peroxisome fission and segregation during vegetative cell reproduction. Apparently, the
function of Wsc and their homologous are highly species dependent.
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Samenvatting
Robert Hooke en Antonie van Leeuwenhoek gebruikten als eersten de term ‘cel’ voor de
basiseenheid van leven. Levende organismen kunnen onderverdeeld worden in drie domeinen, de
bacteriën, de archaea en eukaryoten. Karakteristiek voor de eukaryote cel is de aanwezigheid van
een kern en andere membraangebonden compartimenten, de cel organellen.
Eukaryote organellen worden onderverdeeld in twee categorieën: autonoom (zoals
mitochondriën en chloroplasten) en niet-autonoom (zoals het Golgi apparaat en vacuoles). Het is
algemeen geaccepteerd dat autonome organellen zich vermeerderen door groei en deling, dus
niet de novo, terwijl alle anderen vanaf het endoplasmatisch reticulum kunnen worden gevormd.
Het peroxisoom is één van de meest recent ontdekte organellen. Peroxisomen bevatten een
eiwitrijke matrix waarin zich geen DNA bevindt. De matrix wordt omsloten door een enkele
membraan. Deze organellen zijn zeer dynamisch; hun functie, aantal en morfologie zijn soort
specifiek en hangen tevens af van het ontwikkelingsstadium en het milieu waarin ze zich
bevinden. Karakteristiek voor peroxisomen is de aanwezigheid van tenminste één H2O2-
producerend oxidase in combinatie met katalase. Naast het metabolisme van H2O2 is de β-
oxidatie van vetzuren een andere algemene functie van peroxisomen.
De vorming van peroxisomen is afhankelijk van “peroxines”, specifieke eiwitten die worden
gecodeerd door PEX-genen. Peroxisomale matrixeiwitten worden gesynthetiseerd op vrije
ribosomen in het cytosol en vervolgens post-translationeel geïmporteerd in het organel. Er is
echter nog onduidelijkheid over de vraag of insertie van peroxisomale membraaneiwitten
(PMP’s) in de peroxisomale membraan direct vanuit het cytosol plaatsvindt of dat dit via het ER
gebeurt.
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De vorming van peroxisomen is ook nog steeds een onderwerp van hevige discussies. Het
klassieke model zegt dat peroxisomen autonome organellen zijn die zich vermenigvuldigen door
groei en deling. Recentelijk is dit ter discussie komen te staan door een tweede model dat
voorstelt dat peroxisomen de novo worden gevormd vanaf het ER. Dit laatste wordt vooral
waargenomen in peroxisoom-deficiënte mutanten (pex3 en pex19), na het terug brengen van de
corresponderende genen. Aanvankelijk dacht men dat in deze stammen restanten van
peroxisomale membranen (remnants) volledig ontbraken. Het meest recente model beschrijft hoe
de novo peroxisoom vorming begint met de vorming van twee soorten vesicles, elke met
verschillende PMPs, vanaf het ER. Deze vesicles fuseren vervolgens op een Pex1- en Pex6-
afhankelijke wijze en vormen een pre-peroxisoom dat verder uitgroeit door de import van
matrixeiwitten. Het is aannemelijk dat tenminste een aantal PMP’s via het ER worden
getransporteerd. Dit alles samengenomen kunnen peroxisomen wellicht worden geclassificeerd
als semi-autonome organellen.
Een goede balans tussen vorming en afbraak van organellen is van belang voor de homeostase
van peroxisomen. Huishoudprocessen in de cel zorgen voor herstel of afbraak van beschadigde
en overtollige peroxisomen, om zo een vitale populatie peroxisomen te behouden.
In gist zijn vorming en afbraak van peroxisomen sterk geconserveerde mechanismen. De
verschillende stadia van de levenscyclus van het peroxisoom kunnen precies worden
gemanipuleerd door de juiste groeicondities te gebruiken. Daardoor zijn gisten aantrekkelijke
modelorganismen voor het onderzoek naar homeostase van peroxisomen. Het werk beschreven
in dit proefschrift is vooral gericht op de moleculaire mechanismen die ten grondslag liggen aan
de novo formatie van peroxisomen en het behoud van vitale organellen. De methylotrofe gist
Hansenula polymorpha werd hierbij gebruikt als modelorganisme.
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Hoofstuk 1 vat onze huidige kennis samen over de vorming van peroxisomen, de moleculaire
mechanismen die betrokken zijn bij deling van peroxisomen en de novo synthese vanaf het ER,
en de mechanismen voor kwaliteitscontrole en impact hiervan op cel veroudering.
Hoofdstuk 2 beschrijft experimenten die tot doel hadden op te helderen of deling van een
peroxisoom nodig is alvorens autofagie kan plaatsvinden. De resultaten tonen aan dat het
afbraakproces van peroxisomen is verstoord in mutanten van Saccharomyces cerevisiae en H.
polymorpha waarin peroxisomen helemaal niet meer kunnen delen (∆dnm1 en ∆pex11), zoals we
dat ook zien in de controlestam ∆atg1 (autofagie mutant). Vervolgens keken we naar het belang
van peroxisoomdeling en -autofagie bij de afbraak van afwijkend gevormde peroxisomen. Voor
dit doel kloneerden we een gemuteerde vorm van het katalase gen, dat codeert voor een eiwit
met daaraan een peroxisomaal signaalpeptide (Cat-Y348G.SKL). Dit eiwit vormt aggregated in
de peroxisomale matrix. We produceerden dit eiwit in H. polymorpha ∆atg1 cellen, waardoor er
eiwit aggregaten in de peroxisomen ontstonden, en testten of peroxisoomdeling werd
geïnduceerd. Met behulp van fluorescentiemicroscopie zagen we eiwitaggregaten in
peroxisomen van ∆atg1 cellen, die asymmetrische peroxisoomdeling leken te bevorderen. Dit
resulteerde in cellen met daarin talloze organellen met kleine aggregaten die waren afgesplitst
van het moederorganel. Vervolgens onderzochten we of deze organellen met aggregaten werden
afgebroken via autofagie. Elektronenmicroscopie liet zien dat peroxisomen met eiwitaggregaten
vaak naar de vacuole werden getransporteerd om te worden afgebroken. De stammen gemuteerd
in peroxisoomdeling (∆dnm1 and ∆pex11) waren echter niet in staat de peroxisomen met
aggregaten af te breken. Daarbij zagen we dat de aanwezigheid van eiwitaggregaten van invloed
was op de groeisnelheid van de gistcellen en tot verhoogde ROS-niveaus leidde. Dit betekent dat
tijdige afbraak van peroxisomen met eiwitaggregaten cruciaal is. Uit deze resultaten leiden we af
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dat peroxisoomdeling en -afbraak via autofagie gekoppelde processen zijn, essentieel voor het
verwijderen van schadelijke componenten uit organellen. Op deze wijze wordt ROS-
geïnduceerde celdood voorkomen en daarmee de vitaliteit van organellen behouden.
In de volgende twee hoofdstukken lieten we zien dat Pex3 (Hoofdstuk 3) en Pex19 (Hoofdstuk
4) niet essentieel zijn voor de vorming van peroxisomale membranen. Volgens eerdere
publicaties resulteert de deletie van PEX3 of PEX19 in volledige afwezigheid van peroxisomale
structuren, waarbij peroxisomen de novo worden gevormd vanaf het ER, na functionele
complementatie met de corresponderende genen. Onze data laten zien dat H. polymorpha pex3
en pex19 cellen wel degelijk peroxisomale restanten bevatten die echter onstabiel zijn en snel
worden afgebroken door middel van autofagie. Dit kan verklaren waarom ze zolang
onopgemerkt zijn gebleven. Toen we autofagie blokkeerden in pex3 of pex19 cellen door deletie
van ATG1 werden de peroxisomale membraan structuren (vesicles) gestabiliseerd (in pex3 atg1
of pex19 atg1 cellen). Daarbij zagen we dat deze vesicles de PMP’s Pex13 en Pex14 bevatten en
het matrixeiwit Pex8, terwijl andere PMP’s zoals Pex10, Pex11, Pmp47 en Pex26 (in pex19
atg1) niet op de vesicles aanwezig waren. Bovendien lagen de niveaus van deze eiwitten vaak op
of net onder detectielimiet, terwijl ze in wild-type eenvoudig konden worden aangetoond. We
laten ook zien dat de niveaus van Pex10, Pex11 en Pmp47 worden verhoogd en gestabiliseerd na
overexpressie van PEX19 in pex3 atg1 cellen. Dit suggereert dat deze eiwitten afhankelijk zijn
van de Pex3/Pex19 PMP insertiemachinerie, terwijl Pex14 en Pex13 deze peroxines niet nodig
hebben voor de juiste sorting.
In recente publicaties staat dat deletie van PEX19 resulteert in de accumulatie van Pex3 op het
ER. Uit onze resultaten blijkt dat Pex3 in pex19 atg1 cellen niet ophoopt op het ER, maar
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aanwezig is de vesicles met Pex14 en in pex3 atg1 naar deze structuren gaat na re-introductie van
PEX3.
Vervolgens onderzochten we of de Pex14-bevattende vesicles uit konden groeien tot
peroxisomen. Met behulp van time-lapse video’s laten we zien dat na re-introductie van PEX3 in
pex3 atg1 cellen, Pex3, Pex10 en Pmp47 direct naar de Pex14-vesicles gaan (onafhankelijk van
het ER) die vervolgens uitgroeien tot functionele peroxisomen (Hoofdstuk 3). Op basis van deze
resultaten is het duidelijk dat de PMP’s niet via het ER naar de peroxisomen gaan. Daarbij laten
we zien dat de vesicles met Pex14 in pex3 atg1 en pex19 atg1 cellen zich ontwikkelen tot
peroxisomen door import van matrixeiwitten (GFP-SKL) na re-introductie van respectievelijk
PEX3 of PEX19 (Hoofdstuk 4).
Met onze resultaten vechten we de meeste recente modellen aan waarin wordt beweerd dat Pex3
en Pex19 nodig zijn voor het vormen van pre-peroxisomale vesicles met PMP’s vanaf het ER.
Uit onze resultaten blijkt dat i) Pex3 en Pex19 niet nodig zijn voor de vorming van vesicles vanaf
het ER; ii) in plaats van het ER, bestaande vesicles in pex3/pex19 atg1 cellen kunnen dienen als
sjabloon voor de vorming van peroxisomen na re-introductie van the corresponderende genen; en
iii) de PMP’s Pex13 en Pex14 niet afhankelijk zijn van de Pex3/Pex19 sorterings-machinerie.
Tot slot onderzochten we in Hoofstuk 5 de lokalisatie en functie van vermeende H. polymorpha
orthologen van de eiwitfamilie PMP22/MPV17 (Wsc, SYM1, en YOR292C). Door middel van
in silico analyses identificeerden we vier mogelijke homologen in H. polymorpha. Om deze
eiwitten te kunnen lokaliseren construeerden we C-terminale GFP-fusies van Wsc, Sym1 en
Yor292c en brachten deze tot expressie in H. polymorpha wild-type cellen. Analyse met behulp
van fluorescentiemicroscopie laat zien dat Wsc op het peroxisoom is gelokaliseerd. Eerdere
publicaties vermelden dat Sym1 (gisten en zoogdieren) een eiwit van de mitochondriële
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binnenmembraan is en Yor292cp een component van de vacuolemembraan (zoogdieren). We
konden de exacte locatie van deze eiwitten in H. polymorpha echter niet vaststellen omdat de
eiwitniveaus onder het detectieniveau lagen. Vervolgens onderzochten we de rol van Wsc in H.
polymorpha door een WSC deletiestam te maken. Volgens eerdere publicaties is Wsc betrokken
bij de biogenese van de zogenaamde ‘Woronin bodies’ in de filamenteuze schimmel Neurospora
crassa en is het betrokken bij het transport van kleine moleculen over de peroxisomale
membraan in zoogdieren. Onze resultaten tonen aan dat deletie van het WSC gen geen invloed
heeft op groei op verschillende koolstof- en stikstofbronnen. Ook had deletie geen invloed op de
vorming en proliferatie van peroxisomen wanneer de cellen werden gekweekt op methanol. In
wsc cellen gekweekt op glucose was het aantal peroxisomen echter gereduceerd. Tevens zijn de
organellen onregelmatig van vorm. Time-lapse video’s lieten zien dat het enkele organel in
cellen gekweekt op glucose vaak naar de knop gaat, daarbij de moedercel zonder peroxisoom
achterlatend. In andere cellen bleef het peroxisoom achter in de moedercel tijdens celdeling, wat
resulteerde in een knop zonder peroxisoom. Dit interessante resultaat duidt erop dat wsc cellen
gekweekt op glucose, een gedeeltelijk defect hebben in peroxisoom deling en segregatie tijdens
vegetatieve celdeling.