YHR150w and YDR479c encode peroxisomal integral membrane proteins involved in the regulation of peroxisome number, size, and distribution in Saccharomyces cerevisiae

Peroxisomes, together with the glyoxysomes of plants and glycosomes of trypanosomes, make up the microbody family of organelles. Peroxisomes play diverse roles in the cell, compartmentalizing many activities related to lipid metabolism and functioning in the decomposition of toxic hydrogen peroxide (for reviews see Lazarow and Fujiki, 1985; van den Bosch et al., 1992; Subramani, 1993; Titorenko and Rachubinski, 2001).

Proteins destined for the peroxisomal matrix are synthesized on free polyribosomes and are imported posttranslationally (Lazarow and Fujiki, 1985; Subramani 1993, 1998; Subramani et al., 2000; Purdue and Lazarow, 2001). Most peroxisomal matrix proteins are sorted by tripeptide peroxisome-targeting signal 1 (PTS1)^* located at their carboxy termini (Gould et al., 1987, 1989, 1990; Aitchison et al., 1991; Swinkels et al., 1992), whereas a limited subset of proteins are sorted by a nonapeptide PTS2 at their amino termini (Osumi et al., 1991; Swinkels et al., 1991; Glover et al., 1994b; Waterham et al., 1994). A few exceptional matrix proteins are targeted by internal PTSs, which remain largely uncharacterized (Small et al., 1988; Purdue et al., 1990; Kragler et al., 1993; Elgersma et al., 1995).

Pex5p and Pex7p are receptors specific for PTS1- and PTS2-containing proteins, respectively. These receptors are mobile, interacting with their cargo in the cytosol and docking at the peroxisomal membrane through interaction with Pex13p and Pex14p (for reviews see Subramani, 1998; Hettema et al., 1999; Terlecky and Fransen, 2000; Purdue and Lazarow, 2001; Titorenko and Rachubinski, 2001). Strikingly, the PTS1 receptor has been shown to enter the peroxisomal matrix together with its cargo and to recycle back to the cytosol after dissociation from its cargo (Dammai and Subramani, 2001). The recycled PTS1 receptor is then available for another round of peroxisomal matrix protein import. The pathway of targeting of peroxisomal membrane proteins remains relatively uncharacterized, although it appears to be independent of the pathway of matrix protein targeting. Stretches of basic amino residues on the matrix side of membrane proteins, close to a transmembrane sequence, have been proposed to act in targeting proteins to the peroxisomal membrane (McCammon et al., 1994; Dyer et al., 1996; Elgersma et al., 1997; Pause et al., 2000).

A remarkable feature of peroxisomal protein import is that folded and assembled multimeric proteins can enter the organelle (Walton et al., 1992, 1995; Glover et al., 1994a; McNew and Goodman, 1994). The oligomerization of some of these multimeric proteins is facilitated by specific chaperone molecules, as in the case of the soluble peroxisomal matrix protein thiolase and its chaperone Pex20p (Titorenko et al., 1998), or is self-assisted, as in the case of the heteropentameric fatty acyl–CoA oxidase complex of the yeast Yarrowia lipolytica (Titorenko et al., 2002).

Failure to assemble functional peroxisomes leads to a class of genetic diseases known as the peroxisome biogenesis disorders (PBDs), the archetype of which is Zellweger syndrome (Lazarow and Moser, 1994; Brosius and Gärtner, 2002). Defining the molecular basis of the PBDs has been an area of intense research in recent years, particularly in regards to the identification of the genes controlling peroxisome assembly, the so called PEX genes. Of the 25 PEX genes identified so far, mutations in 11 of the 13 human orthologues have been shown to cause PBDs (for reviews see Fujiki, 2000; Gould and Valle, 2000; Subramani et al., 2000; Brosius and Gärtner, 2002). The identification of additional genes involved in peroxisome assembly and elucidation of the roles of the proteins they encode would provide greater understanding of the molecular basis of these lethal disorders. The completion of the Saccharomyces cerevisiae genome sequencing project has increased the utility of this model organism for the identification of novel genes involved in peroxisome assembly. Microarray transcriptional profiling of S. cerevisiae under conditions of peroxisome induction has already led to the identification of a novel PEX gene, PEX25 (Smith et al., 2002), and knowledge of the entire coding capacity of the S. cerevisiae genome has facilitated the identification of new proteins potentially involved in peroxisome assembly by their similarity with other proteins already shown to be involved in peroxisome assembly in other organisms.

We recently reported the isolation and characterization of YlPex24p, a 61-kD integral membrane protein of peroxisomes required for peroxisome assembly in Y. lipolytica (Tam and Rachubinski, 2002). A search of protein databases revealed that YlPex24p shares extensive sequence similarity with proteins encoded by the ORFs YHR150w and YDR479c of the S. cerevisiae genome. Here we report that both YHR150w and YDR479c code for peroxisomal integral membrane proteins, and we provide evidence for their role in peroxisomal dynamics in S. cerevisiae.

YlPex24p is an integral peroxisomal membrane protein that has been shown to be required for peroxisome assembly in the yeast Y. lipolytica (Tam and Rachubinski, 2002). A search of protein databases with the GENEINFO(R) BLAST Network Service of the National Center for Biotechnology Information revealed two proteins encoded by the ORFs YHR150w and YDR479c of the S. cerevisiae genome that exhibit extensive sequence similarity to YlPex24p (Fig. 1). YlPex24p and Yhr150p exhibit 21.1% amino acid identity and 44.4% amino acid similarity, YlPex24p and Ydr479p exhibit 20.8% amino acid identity and 42.1% amino acid similarity, while Yhr150p and Ydr479p exhibit 20.4% amino acid identity and 41.3% amino acid similarity. Yhr150p is predicted to be a protein of molecular weight 66,145 and to have two transmembrane helices at amino acids 246–268 and 393–415 (http://www.cbs.dtu.dk/services/TMHMM-2.0/) (Krogh et al., 2001). Ydr479p is predicted to be a protein of molecular weight 63,533 and to have four transmembrane helices at amino acids 146–165, 172–194, 265–284, and 291–308. Some potential functional redundancy between Yhr150p and Ydr479p may have prevented them from being identified as being involved in peroxisome assembly in S. cerevisiae by procedures involving random mutagenesis and negative selection for growth of yeast on oleic acid–containing medium.

The synthesis of many peroxisomal proteins is induced by the incubation of yeast cells in oleic acid-containing medium. Genomically encoded protein A chimeras of Yhr150p and Ydr479p were monitored to analyze the expression of YHR150w and YDR479c, respectively, under the control of their endogenous gene promoters. Yeast strains synthesizing Yhr150p-prA, Ydr479p-prA, thiolase-prA, and Pex17p-prA were grown in glucose-containing YPD medium and then shifted to oleic acid–containing YPBO medium. Aliquots of cells were removed at various times after the shift to YPBO medium, and their lysates were subjected to SDS-PAGE and immunoblotting (Fig. 2). Yhr150p-prA and Ydr479-prA, as well as Pex17p-prA (Huhse et al., 1998), were detected in glucose-containing YPD medium at the time of transfer, and their respective levels did not increase with time of incubation of cells in YPBO medium. In contrast, thiolase-prA was barely detectable in cells at the time of transfer to YPBO medium, and its levels were substantially increased with time of incubation in YPBO medium.

A carboxy-terminal PTS1 is sufficient to direct a reporter protein to peroxisomes (for review see Purdue and Lazarow, 2001). A fluorescent chimera between Discosoma sp. red fluorescent protein (DsRed) and the PTS1 Ser-Lys-Leu has been shown to target to peroxisomes of S. cerevisiae (Smith et al., 2002). Genomically encoded protein A chimeras of Yhr150p, Ydr479p, the peroxisomal peroxin Pex17p, (Huhse et al., 1998) and the mitochondrial translocon protein Tom20p (Lithgow et al., 1994) were localized in oleic acid–induced cells by indirect immunofluorescence microscopy combined with direct fluorescence from DsRed–PTS1 to identify peroxisomes (Fig. 3). Yhr150p-prA, Ydr479p-prA, and Pex17p-prA colocalized with DsRed–PTS1 to small punctate structures characteristic of peroxisomes by confocal microcopy. As expected, Tom20p-prA did not colocalize with DsRed–PTS1, as the respective individual green and red signals for these proteins remained separate in confocal microscopy.

Subcellular fractionation and organelle extraction were used to establish if Yhr150p and Ydr479p are associated with peroxisomes and to determine their suborganellar locations. Cells expressing Yhr150p-prA, Ydr479p-prA, and Pex17p-prA were incubated in oleic acid–containing medium and subjected to subcellular fractionation to yield postnuclear supernatant (PNS) fractions. The PNS fractions were subjected to further centrifugation to yield a supernatant fraction (20KgS) enriched for cytosol and a crude organellar pellet fraction (20KgP). Equal portions of the PNS, 20KgS, and the 20KgP were analyzed by immunoblotting. Yhr150p-prA, Ydr479p-prA, and Pex17p-prA all preferentially localized to the 20KgP fraction (Fig. 4 A). Peroxisomes were isolated from the 20KgP fractions of each of the strains expressing Yhr150p-prA, Ydr479p-prA, and Pex17p-prA. The gradients were fractionated, and equal portions of each fraction were analyzed by immunoblotting (Fig. 4 B). Yhr150p-prA and Ydr479p-prA coenriched with the peroxisomal peroxin Pex17p-prA and not with the mitochondrial protein, Sdh2p. Therefore, both microscopic analysis and subcellular fractionation showed Yhr150p and Ydr479p to be peroxisomal proteins. Some amount of Ydr479p-prA was always present in the 20KgS fraction and in the lighter fractions during the gradient isolation of peroxisomes. Whether this represents a selective liberation of a soluble form of Ydr479p-prA during the isolation of peroxisomes remains undetermined. It is unlikely that Ydr479p-prA is found in a cellular compartment in addition to peroxisomes, as immunostaining for Ydr479p-prA yielded exclusively a punctate pattern corresponding to the punctate pattern of peroxisomes defined by DsRed–PTS1 (Fig. 3). In addition, this punctate compartment does not correspond to mitochondria, as the fluorescence pattern generated with the mitochondria-specific dye MitoTracker did not overlap with the punctate pattern generated by detection of Ydr479p-prA by immunofluorescence (unpublished data).

Peroxisomes were hypotonically lysed by incubation in dilute alkali Tris buffer and subjected to centrifugation to yield a supernatant (Ti8S) enriched for matrix proteins and a pellet (Ti8P) enriched for membrane proteins (Fig. 4 C). The chimeras of Yhr150p and Ydr479p localized primarily to the Ti8P fraction, as did the chimeras of the peripheral peroxisomal membrane protein Pex17p (Huhse et al., 1998) and the integral peroxisomal membrane protein Pex3p (Höhfeld et al., 1991). The soluble peroxisomal matrix protein thiolase was found almost exclusively in the Ti8S fraction. The reproducible presence of some Ydr479p in the Ti8S fraction may again be representative of a soluble form of this protein that is selectively liberated during the isolation of peroxisomes (Fig. 4, A and B). The Ti8P fractions were then extracted with alkali sodium carbonate and subjected to centrifugation (Fig. 4 C). This treatment releases proteins associated with, but not integral to, membranes (Fujiki et al., 1982). Under these conditions, Yhr150p-prA and Ydr479p-prA fractionated with Pex3p-prA to the pellet fraction enriched for integral membrane proteins, whereas Pex17p-prA fractionated to the supernatant fraction enriched for soluble proteins, including peripheral membrane proteins. These data suggest that Yhr150p and Ydr479p are both primarily integral peroxisomal membrane proteins, as has been shown for YlPex24p (Tam and Rachubinski, 2002).

We next investigated the ultrastructure of cells incubated in oleic acid–containing YPBO medium by transmission EM. Wild-type cells (Fig. 5 A) consistently showed individual peroxisomes well separated from one another. In contrast, cells of the yhr150Δ (Fig. 5 B), ydr479Δ (Fig. 5 C), and, particularly, yhr150Δ/ydr479Δ (Fig. 5 D) strains contained peroxisomes that exhibited clustering. 28.0, 19.2, and 20.4% of peroxisomes of cells of the yhr150Δ/ydr479Δ, ydr479Δ, and yhr150Δ strains, respectively, showed clustering, in contrast to 4.0% of peroxisomes of wild-type cells (a cluster of peroxisomes was operationally defined as three or more adherent peroxisomes). The clustered peroxisomes often showed evidence of membrane thickening between adjacent peroxisomes in the cluster. Morphometric analysis showed that cells of the deletion strains contained a greater number of peroxisomes than wild-type cells and that, on average, these peroxisomes were smaller in size than those of wild-type cells (Table I). Cells of the deletion strains contained much greater numbers of peroxisomes with areas of 0.02 μm² or less than wild-type cells (Fig. 5 E). Nycodenz density gradient centrifugation analysis showed that peroxisomes purified from yhr150Δ/ydr479Δ cells have a greatly reduced density (peak fraction 8, 1.19 g/cm³) compared with peroxisomes from wild-type cells (peak fraction 1, 1.22 g/cm³) (Fig. 6 A). Peroxisomes isolated from cells deleted for YHR150w or YDR479c were also less dense than wild-type peroxisomes, although the differences in density were less than that observed between peroxisomes from yhr150Δ/ydr479Δ cells and wild-type peroxisomes (unpublished data). EM analysis showed that the peroxisomes purified from yhr150Δ/ydr479Δ cells still exhibited clustering and evidence of thickened peroxisomal membranes, whereas peroxisomes purified from wild-type cells were largely well separated from one another, with no evidence of membrane thickening (Fig. 6 B).

Because cells deleted for one or both of the YHR150w and YDR479c genes are compromised in their regulation of peroxisome number, size, and distribution, we investigated the effects of overexpression of three genes previously shown to be involved in these processes in S. cerevisiae. Cells of strains mutant for the genes encoding the peroxisomal peroxins Pex11p (Erdmann and Blobel, 1995; Marshall et al., 1995, 1996), Pex25p (Smith et al., 2002), and the dynamin-like protein Vps1p (Hoepfner et al., 2001) have reduced numbers of enlarged peroxisomes compared with wild-type cells (see Fig. S1). Overexpression of PEX25 (Fig. 7; Fig. 8 D) or VPS1 (Fig. 7; Fig. 8 E) in cells deleted for one or both of the YHR150w and YDR479c genes led to a partial restoration of the wild-type peroxisomal phenotype, with overexpressing cells containing increased numbers of separate, individual peroxisomes and reduced numbers of peroxisomal clusters (see Figs. S5 and S6). In contrast, overexpression of PEX11 did not appear to have any effect on the abnormal peroxisome morphology observed in cells mutant for the YHR150w and YDR479c genes (Fig. 7; Fig. 8 C; Fig. S4). It should be noted that overexpression of the YHR150w gene led to restoration of wild-type peroxisome morphology in yhr150Δ cells but not in ydr479Δ cells or in cells deleted for both the YHR150w and YDR479c genes (Fig. 7, Fig. 8 A; Fig. S2). In contrast, overexpression of YDR479c led to restoration of wild-type peroxisome morphology not only in ydr479Δ cells, but also in yhr150Δ cells and cells deleted for both genes (Fig. 7; Fig. 8 B; Fig. S3). Therefore, it appears that Yhr150p may be, for the most part, redundant in its functions in regulating peroxisome dynamics with respect to Ydr479p. Attempts at demonstrating physical interactions, either direct or indirect, between Yhr150p and Ydr479p or between these two proteins and Pex11p, Pex25p, or Vps1p were unsuccessful (unpublished data).

As reported previously (Erdmann and Blobel, 1995; Marshall et al., 1995, 1996; Hoepfner et al., 2001; Smith et al., 2002), cells deleted individually for the PEX11, PEX25, and VPS1 genes contained reduced numbers of enlarged peroxisomes (Fig. 7; Fig. S1). Overexpression of the PEX11 gene in wild-type cells or in cells deleted for PEX11, PEX25, or VPS1 led to a proliferation of smaller peroxisomes (Fig. 7; Fig. S4). However, overexpression of PEX25 or VPS1, although resulting in normal peroxisome morphology in their respective mutant backgrounds, did not affect the peroxisome morphology in the opposite mutant background (Fig. 7; Figs. S5 and S6). Overexpression of PEX25, but not VPS1, led to only a nominal increase in the number of normal peroxisomes in cells deleted for the PEX11 gene (Fig. 7; Figs. S5 and S6). Moreover, the pex11Δ cells retained their enlarged peroxisomal phenotype when either YHR150w or YDR479c was overexpressed (Fig. 7; Figs. S2 and S3). Our results suggest that Pex11p plays a central role in regulating peroxisome division. The results of gene overexpression studies are summarized in Table II.

Completion of the S. cerevisiae genome sequencing project proved invaluable for the identification of a novel PEX gene, PEX25, by transcriptome profiling of cells grown in oleic acid–containing medium versus cells grown in glucose-containing medium (Smith et al., 2002). The completed S. cerevisiae genome sequence has added value in that it provides the opportunity to identify novel proteins required for peroxisome biogenesis in S. cerevisiae through sequence similarity between proteins of unknown function encoded by S. cerevisiae and proteins already shown to be required for peroxisome biogenesis in other organisms. We recently demonstrated the requirement of the peroxisomal integral membrane protein Pex24p for peroxisome assembly in the yeast Y. lipolytica (Tam and Rachubinski, 2002). YlPex24p was shown to share extensive sequence similarity with two proteins of unknown function and unknown localization encoded by the ORFs YHR150w and YDR479c of the S. cerevisiae genome. In this manuscript, genomically encoded protein A chimeras of Yhr150p and Ydr479p were shown by a combination of microscopic and subcellular fractionation analyses to be peroxisomal proteins. In their response to extraction by different salts, Yhr150p and Ydr479p act primarily as integral membrane proteins. However, a fraction of Ydr479p consistently acts as a soluble protein. The exact nature of this soluble form of Ydr479p and its origin (is it in equilibrium with the peroxisomal membrane form of Ydr479p?) remain under investigation.

The proteins encoded by the YHR150w and YDR479c genes are not required for peroxisome assembly, as cells harboring deletions for one or both of these genes still contain peroxisomes. These peroxisomes were functional, at least to some degree, as the cells containing one or both of the gene deletions were able to grow in oleic acid–containing medium with essentially the same kinetics as the wild-type strain (unpublished data). However, the peroxisomes in the deleted strain are not normal and show phenotypic characteristics distinct from those of wild-type peroxisomes. The peroxisomes of cells deleted for one or both of the YHR150w and YDR479c genes are more abundant, smaller, and show extensive clustering, as compared with wild-type peroxisomes. In addition, the membranes of the clustered peroxisomes of the gene deletion strains are often thickened in appearance. These characteristics of peroxisomes of the deletion strains are consistent with a role for YHR150w and YDR479c in the control of peroxisome size, number, and distribution within cells. However, it does not appear that YHR150w and YDR479c are required for peroxisome inheritance per se, as all cells deleted for one or both of these genes still contained peroxisomes after numerous cell divisions. Also, if YHR150w or YDR479c had a direct role in the inheritance of peroxisomes, one might expect that a loss of peroxisomes from cells over time resulting from impaired segregation of peroxisomes into daughter cells would lead to decreased kinetics of growth in oleic acid–containing medium for the deletion strains when compared to the wild-type strain, which, as reported above, was not observed. It is interesting to note that Y. lipolytica cells deleted for the PEX24 gene also show evidence of abnormal peroxisomal divisional control. These cells lack mature peroxisomes but do accumulate membrane structures that contain both peroxisomal matrix and membrane proteins (Tam and Rachubinski, 2002). However, these membrane structures are not functional peroxisomes in Y. lipolytica, as pex24Δ cells cannot grow on medium containing oleic acid as the sole carbon source. Therefore, although YlPex24p, like Yhr150p and Ydr479p, most likely has a role in the regulation of peroxisome division, YlPex24p probably does not function identically to Yhr150p or Ydr479p, or is modulated in its actions differently than Yhr150p and Ydr479p.

The size, number, and distribution of peroxisomes are tightly controlled by the cell. Loss of the enzymatic activities of individual peroxisomal β-oxidation enzymes has been shown to result in pronounced changes in peroxisome size and/or number (Fan et al., 1998; Chang et al., 1999; Smith et al., 2000; van Roermund et al., 2000), due primarily to the increased levels of the remaining peroxisomal β-oxidation enzymes. The molecular mechanisms underlying this so-called metabolic control of peroxisome abundance (Chang et al., 1999) remain essentially unknown.

In contrast, members of the Pex11 family of peroxins have been implicated as effectors of peroxisome division in multiple species (Erdmann and Blobel, 1995; Marshall et al., 1995, 1996; Sakai et al., 1995; Abe and Fujiki, 1998; Lorenz et al., 1998; Passreiter et al., 1998; Schrader et al., 1998; Li and Gould, 2002). The recently reported PEX25 gene has also been implicated in the regulation of peroxisome size and number in S. cerevisiae (Smith et al., 2002), as has the dynamin-like protein Vps1p (Hoepfner et al., 2001). Like other dynamin-related proteins, Vps1p was proposed to be involved in a membrane fission event required for the regulation of peroxisome size and abundance. The thickened membranes between some peroxisomes of a peroxisome cluster seen in cells deleted for the YHR150w and/or YDR479c genes are also suggestive of a role for Yhr150p and Ydr479p in controlling fission of the peroxisomal membrane.

How might Yhr150p, Ydr479p, Pex11p, Pex25p, and Vps1p act and interact to control the abundance, size, and distribution of peroxisomes in the S. cerevisiae cell? We sought to get some insight into this question by determining the effects on peroxisome morphology of overexpressing the genes for these proteins in wild-type cells and cells deleted for the different genes.

Overexpression of the PEX11 gene in the pex11Δ genetic background has been reported to result in large numbers of small peroxisomes (Marshall et al., 1995). In contrast, overexpression of PEX25, VPS1, YHR150w, and YDR479c in their respective gene deletion backgrounds does not result in the production of large numbers of small peroxisomes but instead restores the wild-type peroxisomal phenotype. Considering the proliferation of peroxisomes as a two-step pathway, namely division of peroxisomes and separation of peroxisomes, overexpression of the PEX11 gene either in wild-type cells or in cells of the various deletion strains leads to a significant proliferation of peroxisomes, which remain, for the most part, adherent to one another. Thus, Pex11p plays a central and positive regulatory role in the division step of peroxisome proliferation but has little, or no readily apparent, role in the separation step of the process. The presence of reduced numbers of enlarged peroxisomes in pex25Δ (Smith et al., 2002; Fig. S1) and vps1Δ cells (Hoepfner et al., 2001; Fig. S1) suggests that Pex25p and Vps1p function, in addition to Pex11p, in the divisional step of peroxisome proliferation.

Upon completion of peroxisome division, peroxisomes must be separated from one another. Yhr150p and Ydr479p are two proteins required for this process, as their absence leads to an arrest or retardation of the peroxisome proliferation pathway, leading to the presence of clusters of peroxisomes with evidence of thickened membranes sometimes occurring between adjacent peroxisomes. Significant recovery of the wild-type peroxisomal phenotype by overexpression of PEX25 or VPS1 in cells deleted for one or both of the YHR150w or YDR479c genes implies that Pex25p and Vps1p have roles in the separation of peroxisomes in addition to their roles in peroxisome division discussed above. In contrast, overexpression of PEX11 in cells deleted for one or both of the YHR150w or YDR479c genes did not result in the reappearance of wild-type peroxisomes, and peroxisomes remained clustered and sometimes exhibited membrane thickening between adjacent peroxisomes, as in the original strains deleted for YHR150w and/or YDR479c. Therefore, Pex11p appears to function primarily or only at the divisional step of peroxisome proliferation but not at the separation step.

Organelles are highly dynamic structures that undergo fission and fusion processes to allow cells to respond to intracellular and extracellular cues and to allow for their correct segregation at cell division. The maintenance of compartmental integrity in the eukaryotic cell, therefore, requires tight control mechanisms for these events. In the control of peroxisome number, size, and distribution, our data suggest that Pex11p plays a preeminent role in controlling peroxisome division, whereas Pex25p, Vps1p, and the newly identified peroxisomal proteins Yhr150p and Ydr479p all play a prominent role in controlling the separation of peroxisomes from one another. Because of their role in peroxisome dynamics, we suggest that YHR150w and YDR479c be designated as PEX28 and PEX29, respectively, and their encoded peroxins as Pex28p and Pex29p. The challenge for the future lies in understanding further the interplay amongst these proteins and the signaling events they respond to and initiate in order to control peroxisomal dynamics in the cell.

The yeast strains used in this study are listed in Table III. All strains were cultured at 30°C. Media components were as follows: YPD, 1% yeast extract, 2% peptone, 2% glucose; YPBO, 0.3% yeast extract, 0.5% peptone, 0.5% K₂HPO₄, 0.5% KH₂PO₄, 1% Brij 35, 1% (vol/vol) oleic acid; synthetic minimal (SM) medium, 0.67% yeast nitrogen base without amino acids, 2% glucose, 1× Complete Supplement Mixture (Bio 101) without histidine and/or leucine; sporulation medium, 1% potassium acetate, 0.1% yeast extract, 0.05% glucose, 2% agar; and YNBD, 0.67% yeast nitrogen base without amino acids, 2% glucose, containing histidine, leucine, and uracil, each at 30 μg/ml.

The plasmids pDsRed–PTS1 (Smith et al., 2002) and pProtA/HIS5 (Rout et al., 2000) have been previously described. Genes to be overexpressed were amplified by PCR and cloned into the plasmid YEp13 (Broach et al., 1979). For overexpression, the YHR150w gene included 492 bp of upstream and 398 bp of downstream sequence, the YDR479c gene included 642 bp of upstream and 335 bp of downstream sequence, the PEX11 gene included 599 bp of upstream and 329 bp of downstream sequence, the PEX25 gene included 753 bp of upstream and 319 bp of downstream sequence, and the VPS1 gene included 555 bp of upstream and 324 bp of downstream sequence.

Genes were genomically tagged with the sequence encoding Staphylococcus aureus protein A by homologous recombination using PCR-based integrative transformation into parental BY4742 haploid cells (Aitchison et al., 1995; Dilworth et al., 2001).

Strains encoding protein A chimeras and transformed with the plasmid pDsRed–PTS1 were grown in SM medium for 12 h and then incubated in YPBO medium for 8 h. Cells were processed for immunofluorescence microscopy as previously described (Pringle et al., 1991; Tam and Rachubinski, 2002). Protein A chimeras were detected with rabbit antiserum to mouse IgG (ICN Biomedicals) and FITC-conjugated goat anti–rabbit IgG. Images were captured on an LSM510 META (Carl Zeiss MicroImaging, Inc.) laser scanning microscope or with a digital fluorescence camera (Spot Diagnostic Instruments). Whole cells and subcellular fractions were processed for EM as previously described (Eitzen et al., 1997).

For each strain analyzed, electron micrographs of 50 randomly selected cells at a magnification of 17, 000 were scanned, and the areas of individual cells and of individual peroxisomes were determined by counting the number of individual pixels in a particular cell or peroxisome with Image Tool for Windows, Version 2.00 (University of Texas Health Science Center). To determine the average area of a peroxisome, the total peroxisome area was calculated and divided by the total number of peroxisomes counted. To quantify peroxisome number, the numerical density of peroxisomes (number of peroxisomes per μm³ of cell volume) was calculated by the method of Weibel and Bolender (1973) for spherical organelles as follows. First, the total number of peroxisome profiles was counted and reported as the number of peroxisomes per cell area assayed (N_A). Next, the peroxisome volume density (V_V) was calculated for each strain (total peroxisome area/total cell area assayed). Using the values V_V and N_A, the numerical density of peroxisomes was determined (Weibel and Bolender, 1973).

Subcellular fractionation and peroxisome isolation were done essentially as previously described (Bonifacino et al., 2000; Smith et al., 2002). In brief, cells grown overnight in YPD medium were transferred to YPBO medium and incubated for 8 h. Cells were harvested, washed, and converted to spheroplasts by digestion with Zymolyase 100T (1 mg/g of cells) in 50 mM potassium phosphate, pH 7.5, 1.2 M sorbitol, 1 mM EDTA for 1 h at 30°C. Spheroplasts were lysed by homogenization in buffer H (0.6 M sorbitol, 2.5 mM MES, pH 5.5, 1 mM KCl) containing 1 mM EDTA and PINS (0.5 mM benzamidine, 2 μg leupeptin/ml, 2 μg aprotinin/ml, 1 μg pepstatin A/ml, 3 μg antipain/ml, 0.5 mg Pefabloc/ml). The homogenate was subjected to centrifugation for 10 min at 2,000 g to yield a PNS fraction. The PNS fraction was subjected to further differential centrifugation at 20,000 g for 30 min to yield supernatant (20KgS) and pellet (20KgP) fractions. The 20KgP fraction was resuspended in buffer H containing 11% Nycodenz and PINS, and a volume containing 5 mg of protein was overlaid onto either a 30-ml discontinuous gradient consisting of 17, 25, 35, and 50% (wt/vol) Nycodenz or a 30-ml continuous gradient of 30–45% (wt/vol) Nycodenz, both in buffer H containing PINS. Organelles were separated by centrifugation at 100,000 g for 90 min in a VTi50 rotor (Beckman Coulter). Fractions of 2 ml were collected from the bottom of the gradient.

Peroxisomes were extracted as previously described (Fujiki et al., 1982; Nuttley et al., 1990). Essentially, organelles in the 20KgP fraction (50 μg of protein) were lysed by incubation in 10 volumes of Ti8 buffer (10 mM Tris-HCl, pH 8.0) containing 3× PINS on ice for 1 h and separated into pellet (Ti8P) and supernatant (Ti8S) fractions by centrifugation at 245,000 g for 1 h at 4°C in a TLA120.2 rotor (Beckman Coulter). The Ti8P fraction was resuspended in Ti8 buffer to a final protein concentration of 0.5 mg/ml, and a portion of the resuspended fraction was extracted with 0.1 M Na₂CO₃, pH 11.3, for 1 h on ice and then separated into supernatant (CO₃S) and pellet (CO₃P) fractions by centrifugation at 245,000 g in a TLA120.2 rotor at 4°C for 1 h. Proteins in the Ti8S, Ti8P, CO₃S, and CO₃P fractions were precipitated by addition of TCA, and the precipitates were washed with acetone. Proteins in equal portions of each fraction were separated by SDS-PAGE and analyzed by immunoblotting.

The homozygous deletion diploid strain yhr150wΔ-HD (Giaever et al., 2002) was sporulated, and the tetrads were dissected to select for the haploid MATa strain. This strain was mated to the haploid MATα deletion strain ydr479cΔ by replica plating to obtain a heterozygous diploid strain harboring deletions for both YHR150w and YDR479c. The diploid strain was sporulated, and tetrads from 10 heterozygous diploids were dissected by micromanipulation. All spores were grown in YPD medium, and DNA was extracted. Haploid strains carrying deletions in both the YDR479c and YHR150w genes were selected by PCR analysis.

Antibodies to the carboxy-terminal SKL tripeptide (Aitchison et al., 1992), thiolase (Eitzen et al., 1996), and Sdh2p (Dibrov et al., 1998) have been previously described. FITC-conjugated anti–rabbit IgG and rhodamine-conjugated anti–guinea pig IgG (Jackson ImmunoResearch Laboratories) were used to detect primary antibodies in immunofluorescence microscopy. Rabbit antibodies to glucose-6-phosphate dehydrogenase (G6PDH) of S. cerevisiae were obtained from Sigma-Aldrich.

Extraction of nucleic acid from yeast lysates and manipulation of DNA were performed as previously described (Ausubel et al., 1994). Immunoblotting was performed using a wet transfer system (Ausubel et al., 1994), and antigen–antibody complexes in immunoblots were detected by ECL (Amersham Biosciences). Protein concentration was determined using a commercially available kit (Bio-Rad Laboratories) and bovine serum albumin as a standard.

The online version of this manuscript contains additional figures (Figs. S1–S6) showing the ultrastructure of cells of the gene deletion strains pex11Δ, pex15Δ, pex25Δ, and vps1Δ and of various strains overexpressing the YHR150w, YDR479c, PEX11, PEX25, and VPS1 genes. Cells were grown overnight in glucose-containing medium, transferred to oleic acid–containing medium, and incubated in oleic acid–containing medium for 8 h. Cells were then fixed and processed for EM.

We thank Honey Chan for her expert assistance with EM and Richard Poirier for help with confocal microscopy.

This work was supported by operating grant 39322 from the Canadian Institutes of Health Research to R.A. Rachubinski and J.D. Aitchison. R.A. Rachubinski holds the Canada Research Chair in Cell Biology and is an International Research Scholar of the Howard Hughes Medical Institute. Y.Y.C. Tam is the recipient of a studentship from the Alberta Heritage Foundation for Medical Research.