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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.

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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

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segregation of peroxisomes and shaping of the organelle membrane.

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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

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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

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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

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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).

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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

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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.

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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.

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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).

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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

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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).

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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

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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’

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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.