Article |
Address correspondence to Vladimir I. Titorenko, Department of Biology, Concordia University, 7141 Sherbrooke Street, West, Montreal, Quebec H4B 1R6, Canada. Tel.: (514) 848-2424-3424. Fax: (514) 848-2424-2881. email: VTITOR{at}vax2.concordia.ca
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Abstract |
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Key Words: organelle division; membrane fission; peroxisome assembly; peroxin; liposomes
Abbreviations used in this paper: Aox, acyl-CoA oxidase; ICL, isocitrate lyase; MLS, malate synthase; PIC, protease inhibitor cocktail; PMP, peroxisomal membrane protein; THI, thiolase.
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Introduction |
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Data on purification, protein profiling, and electron microscopic analysis of mammalian and yeast peroxisomes have provided important information regarding the process of peroxisomal development. It was established that the population of peroxisomes in a cell consists of several peroxisomal subforms that differ in their size, morphology, buoyant density, and protein composition (Lüers et al., 1993; van Roermund et al., 1995; Wilcke et al., 1995; Titorenko et al., 1996, 2000; Faber et al., 1998). Furthermore, data on the in vivo dynamics of peroxisomal protein localization to several peroxisomal subforms demonstrated that these subforms also differ in their import competency for various proteins. In fact, newly synthesized peroxisomal proteins in mammalian (Heinemann and Just, 1992) and yeast (Titorenko et al., 2000) cells are imported primarily into small peroxisomal vesicles of intermediate buoyant density that subsequently convert to mature peroxisomes of high density. Recent findings in human (South and Gould, 1999; Gould and Valle, 2000) and yeast (Snyder et al., 1999; Subramani et al., 2000; Titorenko et al., 2000) cells have suggested that several peroxisomal subforms are organized into a multistep peroxisome assembly pathway. The pathway operates by the conversion of subforms in a temporally ordered manner, involves the stepwise import of distinct subsets of matrix and membrane proteins into different intermediates along the pathway, and leads to the assembly of mature peroxisomes (Gould and Valle, 2000; Subramani et al., 2000; Titorenko and Rachubinski, 2001a,b).
The peroxisome assembly pathway operating in the yeast Yarrowia lipolytica leads to the formation of mature peroxisomes, P6 (Titorenko et al., 2000). In this yeast, five immature peroxisomal subforms, termed P1P5, differ in their import competency for various proteins and are related through a time-ordered conversion of one subform to another. The current study utilizes several approaches to investigate whether growth and division of immature peroxisomal vesicles and mature peroxisomes are coordinated in Y. lipolytica cells. Furthermore, we have previously demonstrated that peroxisome division in this yeast is regulated by the intraperoxisomal peripheral membrane protein Pex16p (Eitzen et al., 1997), a member of the peroxin family of proteins required for peroxisome assembly, division, and inheritance (Sacksteder and Gould, 2000; Subramani et al., 2000; Purdue and Lazarow, 2001; Titorenko and Rachubinski, 2001b). Here, we further investigate the role of Pex16p in peroxisome division. We describe an unusual mechanism that controls peroxisome division from inside the peroxisome. A temporally and spatially regulated interaction between Pex16p and a heteropentameric complex of acyl-CoA oxidase (Aox), one of the proteins imported into the early peroxisomal precursor P2, plays a pivotal role in this control mechanism.
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Results |
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The membrane-bound Aox complex interacts with the peripheral membrane peroxin Pex16p inside mature peroxisomes
The membrane-associated Aox complex of mature peroxisomes of wild-type cells coimmunoprecipitated under native conditions with the peroxin Pex16p (Fig. 1 F), which is attached to the matrix face of the peroxisomal membrane (Eitzen et al., 1997). Neither Aox nor Pex16p was recovered in the flowthrough when native immunoprecipitation was done with anti-Aox1p or anti-Pex16p antibodies (Fig. 1 F). Thus, the membrane-bound pools of both Aox and Pex16p in mature peroxisomes of wild-type cells were present only as components of a complex, and none of these proteins could be found in its free form. No other peroxisomal membrane peroxin tested, including Pex2p (Eitzen et al., 1996), Pex5p (Szilard et al., 1995), and Pex8p (Smith et al., 1997), interacted with the membrane-bound Aox or Pex16p (Fig. 1 F).
The different Aox subunits and Pex16p are present in equimolar amounts in their membrane-associated complex, as judged by quantitation of their stoichiometry in L-[35S]methioninelabeled complex immunoprecipitated from mature peroxisomes of wild-type cells (Fig. S2, A and C, available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1). No other radiolabeled membrane protein coimmunoprecipitated with the components of the AoxPex16p complex under native conditions (Fig. S2 B). Whereas the molecular mass of the Aox complex recovered from the matrix of mature peroxisomes was 443 kD (Fig. S3 A, available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1) (Titorenko et al., 2002), the molecular mass of the AoxPex16p complex attached to the matrix face of the peroxisomal membrane in wild-type cells was
900 kD (Fig. S3 B). From these observations and a consideration of the molecular masses of each Aox subunit (
80 kD) and Pex16p (
45 kD), we conclude that relocation of a significant portion of the Aox complex from the matrix to the membrane at the last step of the assembly of mature peroxisomes leads to the formation of a supramolecular complex containing two molecules of Aox complex and two molecules of Pex16p. Relocation of Aox complex to the matrix face of the peroxisomal membrane requires two Aox subunits, Aox4p and Aox5p, and Pex16p. In fact, no membrane-bound form of the Aox complex was detected in mature peroxisomes recovered from mutant cells lacking any of these three proteins (Fig. 1, D, E, and G).
The inability of the Aox complex to titrate all membrane-bound Pex16p causes a defect in the division of mature peroxisomes
Overexpression of the PEX16 gene by the highly active THI promoter in the strain pex16-TH results in a reduced number of greatly enlarged mature peroxisomes (Fig. 3) (Eitzen et al., 1997). Similar to Aox in mature peroxisomes of wild-type cells, a significant portion of all five Aox subunits inside the mature peroxisomes of pex16-TH cells is relocated from the matrix to the membrane (Fig. 1 H), where they form a complex with each other and with Pex16p. In fact, all five Aox subunits and Pex16p recovered from the membranes of mature peroxisomes of pex16-TH cells coimmunoprecipitated under native conditions with anti-Aox3p or anti-Pex16p antibodies (Fig. 1 I). However, unlike the membrane-attached Pex16p in mature peroxisomes of wild-type cells, which is present only as a component of the AoxPex16p complex (Fig. 1 F), most of the Pex16p in mature peroxisomes of pex16-TH cells cannot be immunoprecipitated under native conditions with anti-Aox3p antibodies (Fig. 1 I) and, therefore, does not interact with the membrane-bound Aox complex.
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Importantly, the accumulation of greatly enlarged peroxisomes in aox4KO and aox5KO cells was not due to a deficiency in peroxisomal fatty acid ß-oxidation. In fact, no mutation knocking out a single Y. lipolytica AOX gene affected the enzymatic activity of Aox, one of the key enzymes of peroxisomal ß-oxidation, or impaired the utilization of oleic acid as a carbon source (Wang et al., 1999). Thus, the observed changes in peroxisome size and number in aox4KO and aox5KO cells (Fig. 3) cannot be attributed to a defect in the so-called metabolic control of peroxisome abundance (Chang et al., 1999), which operates in yeast, mammalian, and human cells (Fan et al., 1998; Poll-Thé et al., 1988; Chang et al., 1999; Smith et al., 2000; van Roermund et al., 2000).
Morphometric analysis of random electron microscopy sections was used to evaluate the dynamics of change in the size and number of peroxisomes in wild-type and aox mutant cells transferred from glucose- to oleic acidcontaining medium. In wild-type cells of Y. lipolytica, such a transfer greatly increases peroxisome size and number (Fig. S4, available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1) (Smith et al., 2000). Data from morphometric analysis further confirmed that the inability of the Aox complex to relocate from the matrix to the membrane at the last step of the assembly of mature peroxisomes impairs their ability to divide. During the first 3 h of incubation in oleic acidcontaining medium, the size of peroxisomes in wild-type, aox4KO, and aox5KO cells significantly increased (Fig. 4 A; Figs. S4S6, available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1), while their number did not change (Fig. 4 B and Figs. S4S6). Similar dynamics of change in peroxisome size and number by 3 h after the shift from glucose- to oleic acidcontaining medium was observed in aox1KO, aox2KO, and aox3KO cells (unpublished data). By 6 and 9 h after the shift to oleic acidcontaining medium, the number of peroxisomes in wild type (Fig. 4 B and Fig. S4), and in aox1KO, aox2KO, and aox3KO cells (unpublished data), dramatically increased, attaining 14.6 ± 2.0 peroxisomes per µm3 of cell section volume. Concomitantly, the proportion of small peroxisomes in these cells gradually increased, leading to significant variability in peroxisome size by 9 h after the shift (Fig. 4 A and Fig. S4). In contrast, the size of peroxisomes in aox4KO and aox5KO mutant cells continued to increase by 6 and 9 h after the shift to oleic acidcontaining medium (Fig. 4 A and Figs. S5 and S6), with only greatly enlarged peroxisomes visible by 9 h after the shift. During the entire period of incubation after the shift from glucose- to oleic acidcontaining medium, the number of peroxisomes in aox4KO and aox5KO cells did not change significantly, attaining only 1.7 ± 0.3 and 2.3 ± 0.4 peroxisomes per µm3 of cell section volume, respectively, by the end of the incubation (Fig. 4 B and Figs. S5 and S6). Thus, the inability of the Aox complex lacking either the Aox4p or the Aox5p subunit to relocate from the matrix to the membrane and, therefore, to titrate all membrane-bound Pex16p results in the inability of Aox to prevent the negative effect of Pex16p on the division of mature peroxisomes.
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Relocation of Aox from the matrix to the membrane of mature peroxisomes is due to an increase in the total mass of matrix proteins above a critical level
Comparison of the spectra and relative distributions of peroxisomal matrix and membrane proteins demonstrated that even the earliest intermediates in the multistep peroxisome assembly pathway, the immature peroxisomal vesicles P1 and P2, contain most of the peroxisomal membrane proteins (PMPs) associated with mature peroxisomes, P6 (Fig. 6 A). P1 and P2 undergo fusion to generate larger and more dense immature peroxisomal vesicles, P3 (Titorenko et al., 2000), containing PMPs derived from both fusion partners. The quantities of PMPs in P4, P5, and P6 peroxisomes were significantly lower than in P1, P2, and P3 peroxisomes, and gradually decreased from P4 to P6 (Fig. 6 B). In contrast, only a few matrix proteins found in mature peroxisomes were seen in the immature peroxisomal vesicles P1, P2, and P3 (Fig. 6 A). Most matrix proteins were associated with P4, P5, and P6, and the complexity of their spectra increased from P4 to P6 (Fig. 6 A). The quantities of matrix proteins in P4, P5, and P6 peroxisomes were significantly higher than in P1, P2, and P3 peroxisomes, and gradually increased from P4 to P6 (Fig. 6 B). Taken together, these results strongly suggest that the stepwise import of distinct subsets of matrix proteins into different immature intermediates along the peroxisome assembly pathway provides them with an increasing fraction of the matrix proteins present in mature peroxisomes, P6 (see Fig. 8). P6 peroxisomes contain the highest levels of matrix proteins (Fig. 6, A and B).
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A comparative analysis of PLB and PLC, which differ in their amounts of Aox (Fig. 7 C) but contain similar low amounts of other matrix proteins (Fig. 7 B), showed that a significant increase in the amount of matrix-associated Aox did not result in its relocation to the membrane (Fig. 7 C). Taken together, these findings suggest that overloading mature peroxisomes with matrix proteins other than Aox is a major factor in the relocation of Aox complex from the matrix to the membrane.
Finally, although PLA and PLD were loaded with very similar high amounts of matrix proteins (Fig. 7 B), including Aox (Fig. 7 C), the Aox complex was attached to the membrane only inside PLA (Fig. 7 C). PLA contain Pex16p, whereas PLD lack this membrane-bound peroxin (Fig. 7 C). Therefore, Pex16p is the only attachment factor for the Aox complex in the PLA liposomes containing high amounts of matrix proteins and, perhaps, also in mature peroxisomes P6 (Fig. 1 G) containing the greatest percentage of matrix proteins as compared with immature peroxisomal vesicles P1P5 (Fig. 6, A and B).
It should be noted that the above data cannot rule out the possibility that a distinct, yet unknown, matrix protein or a limited set of such proteins rather than protein mass in the peroxisomal matrix initiates the relocation of Aox complex from the matrix to the membrane, thereby terminating the negative action of Pex16p on peroxisome division. Although Aox in the matrix of mature peroxisomes does not form a stable complex with any protein (Titorenko et al., 2002), even its transient interaction with a specific, yet unknown, soluble factor may promote the redistribution of Aox from the matrix to the membrane. Alternatively, overloading mature peroxisomes with matrix proteins may ultimately lead to the relocation of an unknown specific factor from the matrix to the membrane. Once bound to the peroxisomal membrane, this specific factor may cause perturbations in its physical properties, thereby promoting the assembly of the AoxPex16p complex at the matrix face of the membrane. The development of a reliable in vitro assay for reconstructing the relocation of Aox from the matrix to the membrane and its interaction with membrane-bound Pex16p creates the opportunity to test individual peroxisomal matrix proteins for their ability to initiate these processes.
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Discussion |
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The stepwise import of distinct subsets of matrix proteins into P1P5 intermediates provides them with an increasing fraction of the matrix proteins present in mature peroxisomes (Fig. 6, A and B). This increase in the total mass of matrix proteins above a critical level causes the redistribution of the heteropentameric complex of Aox, which is imported into the early intermediate P2 (Titorenko et al., 2000), from the matrix to the matrix surface of the membrane (Fig. 1, A and F). A significant redistribution of Aox complex from the matrix to the membrane occurs only in mature peroxisomes (Fig. 1 A), which contain the greatest percentage of matrix proteins (Fig. 6, A and B). Overloading mature peroxisomes with matrix proteins other than Aox can be a major factor in the relocation of Aox complex to the membrane. The available data cannot rule out the possibility that a distinct, yet unknown, matrix protein, rather than protein mass, in the peroxisomal matrix initiates the relocation of Aox complex from the matrix to the membrane.
Inside mature peroxisomes, the membrane-bound pool of Aox complex interacts, via its Aox4p and Aox5p subunits, with Pex16p (Fig. 1 F). This interaction leads to the formation of a supramolecular complex containing two molecules of Aox complex and two molecules of Pex16p (Figs. S2 and S3) and terminates the negative action of Pex16p on scission of the peroxisomal membrane, thereby allowing mature peroxisomes to divide. The temporally and spatially regulated interaction between membrane-attached Aox and Pex16p ensures the temporal and spatial separation of the processes of peroxisome assembly and division in Y. lipolytica. Such a separation may provide an important advantage for the efficient, stepwise assembly of mature, metabolically active peroxisomes.
Different organisms exhibit different temporal patterns of peroxisome growth and division
A combination of morphometric electron microscopic analysis, pulse-chase analysis of the trafficking of peroxisomal proteins in vivo, and the isolation and protein profiling of structurally distinct peroxisomal subforms has convincingly demonstrated that yeast peroxisomes do not grow and divide at the same time (Veenhuis and Goodman, 1990; Tan et al., 1995; Titorenko et al., 2000; this study). It seems that evolution has generated at least two different temporal patterns of peroxisome growth and division. In the yeast C. boidinii (Veenhuis and Goodman, 1990), the massive proliferation of immature peroxisomal vesicles containing only minor amounts of matrix proteins is a primary event in peroxisomal development. This significant increase in the number of immature peroxisomes by their division precedes the growth of these early peroxisomal precursors by membrane and matrix protein import and their conversion to mature organelles containing the complete set of peroxisomal proteins (Veenhuis and Goodman, 1990). We demonstrated that the timing of events of peroxisome growth and division is different in the yeast Y. lipolytica. In this organism, the growth of immature peroxisomal vesicles, which is accomplished by the import of matrix proteins, and their development into mature peroxisomes occur before completely assembled mature peroxisomes undergo division (this study; Titorenko et al., 2000). Similar temporal patterns of peroxisome growth and division have been observed for the yeast Hansenula polymorpha (Tan et al., 1995).
In human cells, both immature peroxisomal vesicles and mature peroxisomes are proposed to be able to divide (Gould and Valle, 2000). However, the division of immature peroxisomes before their growth and maturation by peroxisomal protein import can only be seen in some peroxin-deficient human fibroblasts after reactivation or reexpression of an originally defective peroxin-encoding gene (Matsuzono et al., 1999; South and Gould, 1999; Sacksteder and Gould, 2000). On the other hand, in normal human cells, growth of immature peroxisomal vesicles by membrane and matrix protein import, resulting in their conversion to mature peroxisomes, may occur before peroxisomes undergo division (Gould and Valle, 2000).
Two mechanisms regulate peroxisome division in Y. lipolytica in response to a signal from inside the peroxisome
This study and our published data (Eitzen et al., 1997; Smith et al., 2000) provide evidence that the membrane scission event required for peroxisome division in Y. lipolytica is regulated by two mechanisms. Both mechanisms control peroxisome division in response to a specific signal transmitted from inside the peroxisome. One mechanism acts through the Pex16p- and Aox-dependent intraperoxisomal signaling cascade (Fig. 8). In this cascade, the ability of Pex16p to inhibit membrane scission is inversely proportional to the level of membrane-bound Aox, which, in turn, depends on the intraperoxisomal level of matrix proteins other than Aox. Only inside mature peroxisomes, which are assembled by the stepwise import of distinct subsets of matrix proteins into different immature peroxisomal vesicles, does the total mass of matrix proteins other than Aox reach its critical level. This triggers the relocation of a significant portion of Aox from the matrix to the matrix face of the peroxisomal membrane, ultimately terminating the negative action of Pex16p on membrane scission. Importantly, whereas the pex16-TH, aox4KO, and aox5KO mutations impair the Pex16p- and Aox-dependent control of peroxisome division and result in a reduced number of greatly enlarged mature peroxisomes (Fig. 3) (Eitzen et al., 1997), they do not affect metabolic flux through the peroxisomal fatty acid ß-oxidation pathway (Eitzen et al., 1997; Wang et al., 1999). On the other hand, loss of the activity of multifunctional enzyme type 2, one of the key enzymes of peroxisomal ß-oxidation, but not the absence of this protein, causes pronounced changes in peroxisome size and number in Y. lipolytica (Smith et al., 2000). Therefore, the Pex16p- and Aox-dependent control of peroxisome division in Y. lipolytica coexists with another mechanism regulating scission of the peroxisomal membrane from inside the peroxisome, the so-called metabolic control of peroxisome abundance (Chang et al., 1999). This second mechanism depends on metabolic flux through the peroxisomal fatty acid ß-oxidation pathway and regulates peroxisome division in yeast (Smith et al., 2000; van Roermund et al., 2000), mammalian (Fan et al., 1998; Poll-Thé et al., 1988), and human (Chang et al., 1999) cells. The metabolic control of peroxisome abundance may be due to the ability of peroxisomes to generate a signaling molecule, perhaps an intermediate of peroxisomal fatty acid ß-oxidation, that initiates a cascade of events ultimately promoting peroxisome division (van Roermund et al., 2000; Li and Gould, 2002).
Controlling the size and number of different organelles is essential to the normal physiology and the viability of cells. Here we have described an unusual mechanism that controls the division of peroxisomes from within the peroxisome itself. We have shown that the temporally and spatially regulated interaction between a peroxin, Pex16p, required for peroxisome biogenesis and the heteropentameric complex of the peroxisomal ß-oxidation enzyme Aox plays a pivotal role in the control of peroxisome division in the yeast Y. lipolytica. A challenge for the future will be to understand how perturbations in the physical properties of the peroxisomal membrane promote the membrane scission event required for peroxisome division and how the Pex16p- and Aox-dependent intraperoxisomal signaling cascade triggers this process.
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Materials and methods |
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Subcellular fractionation and peroxisome isolation
Subcellular fractionation of Y. lipolytica cells grown in oleic acidcontaining medium (Szilard et al., 1995), isolation of highly purified mature peroxisomes P6 (Titorenko et al., 1998), and purification of immature peroxisomes P1P5 (Titorenko et al., 2000) were performed as described previously.
Immunoaffinity chromatography
Covalent coupling of affinity-purified antibodies to protein ASepharose was performed as described previously (Xu et al., 1998). For immunoaffinity chromatography under native conditions, peroxisomal matrix proteins recovered in the supernatant fraction after centrifugation of osmotically lysed peroxisomes and peroxisomal liposomes were diluted with an equal volume of 50 mM Tris-HCl, pH 7.5, buffer containing 300 mM NaCl, 1% (vol/vol) Triton X-100, and 2x protease inhibitor cocktail (PIC) (Szilard et al., 1995). The pellets of PMPs recovered after centrifugation of osmotically lysed peroxisomes and peroxisomal liposomes were resuspended in 25 mM Tris-HCl, pH 7.5, buffer containing 150 mM NaCl, 0.5% (vol/vol) Triton X-100, and 1x PIC. Samples were cleared of any nonspecifically binding proteins by incubation for 20 min at 4°C with protein ASepharose washed five times with 10 mM Tris-HCl, pH 7.5. The cleared samples were then subjected to immunoaffinity chromatography. Bound proteins were washed five times with 25 mM Tris-HCl, pH 7.5, 150 mM NaCl, and 0.5% (vol/vol) Triton X-100 and eluted with 100 mM glycine, pH 2.8. Proteins were precipitated by addition of trichloroacetic acid to 10%, washed in ice cold 80% (vol/vol) acetone, and then subjected to SDS-PAGE followed by immunoblotting or by fluorography (Titorenko et al., 1998).
For immunoaffinity chromatography under denaturing conditions, PMPs purified by immunoaffinity chromatography under native conditions and proteins recovered in the 200KgS (cytosolic), 200KgP, and 20KgP subcellular fractions were diluted with an equal volume of 4% SDS, and samples were warmed at 65°C for 10 min. Samples were then allowed to cool to room temperature, and four volumes of 62.5 mM Tris-HCl, pH 7.5, buffer containing 190 mM NaCl, 1.25% (vol/vol) Triton X-100, and 6 mM EDTA were added. Samples were cleared of any nonspecifically binding proteins by incubation for 20 min at 4°C with protein ASepharose washed five times with 10 mM Tris-HCl, pH 7.5. The cleared samples were then subjected to immunoaffinity chromatography. Bound proteins were washed five times with 50 mM Tris-HCl, pH 7.5, 150 mM NaCl, 1% (vol/vol) Triton X-100 and eluted with 2% SDS at 95°C for 5 min. Eluted proteins were subjected to a second immunoprecipitation (recapture) step (Bonifacino and Dell'Angelica, 1998), resolved by SDS-PAGE, and analyzed by immunoblotting or visualized by fluorography (Titorenko et al., 1998).
Flotation gradient analysis
The pellet of PMPs recovered after centrifugation of osmotically lysed P6 peroxisomes was resuspended in 100 µl of buffer M (10 mM MES-KOH, pH 5.5, 1 mM KCl, 0.5 mM EDTA, 0.1% [vol/vol] ethanol, 1x PIC), transferred to the bottom of a 5-ml ultraclear centrifuge tube (Beckman Coulter), and supplemented with five volumes of 65% (wt/wt) sucrose in buffer M in order to adjust the sucrose concentration of the sample to 54% (wt/wt). The sample was then overlaid with 1.1 ml of 45% sucrose, 1.1 ml of 30% sucrose, 1.1 ml of 10% sucrose (all wt/wt in buffer M), and lastly with 1.1 ml of buffer M alone. After centrifugation at 200,000 g for 18 h at 4°C in a SW50.1 rotor (Beckman Coulter), 18 fractions of 275 µl each were collected.
Peroxisomal matrix proteins recovered in the supernatant fraction after centrifugation of osmotically lysed P6 peroxisomes were incubated for 2 h at 75°C. Under these conditions, all matrix proteins formed insoluble aggregates, as judged by light scattering at 320 nm and as confirmed by SDS-PAGE followed by Coomassie staining. Aggregates of peroxisomal matrix proteins were pelleted by centrifugation at 20,000 g for 30 min at 4°C and resuspended in 100 µl of buffer M. This material was subjected to flotation on a multistep sucrose gradient as described above.
Electron microscopy
Electron microscopy (Goodman et al., 1990), morphometric analysis of random electron microscopic sections of cells (Titorenko et al., 1998), and electron microscopic analysis of purified peroxisomal liposomes (Titorenko et al., 2000) were performed as previously described.
Other methods
Preparation of peroxisomal liposomes (supplemental Materials and methods, available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1), SDS-PAGE and immunoblotting (Titorenko et al., 1998), pulse-chase analysis (Titorenko et al., 1998), and fractionation of peroxisomal proteins by centrifugation on a linear 535% glycerol gradient (Titorenko et al., 2002) were performed as previously described. Osmotic lysis of peroxisomes, protein extraction, and protease protection analysis of purified peroxisomes were performed according to established procedures (Szilard et al., 1995).
Online supplemental material
The supplemental material (available at http://www.jcb.org/cgi/content/full/jcb.200305055/DC1) includes additional Materials and methods and figures (Figs. S1S7). The supplemental Materials and methods describe preparation of peroxisomal liposomes. Fig. S1 demonstrates that all remaining Aox subunits form a membrane-attached complex that interacts with Pex16p inside mature peroxisomes of mutant cells lacking Aox1p, Aox2p, or Aox3p. Fig. S2 shows that all five Aox subunits and Pex16p are present in equimolar amounts in their membrane-associated complex inside mature peroxisomes of wild-type cells. Fig. S3 provides evidence that relocation of a 443-kD Aox complex from the matrix to the membrane at the last step of the assembly of mature peroxisomes results in the formation of a 900-kD complex containing Aox and Pex16p. Figs. S4S6 provide data on electron microscopic analysis of the dynamics of change in the size and number of peroxisomes in wild-type, aox4KO, and aox5KO cells transferred from glucose- to oleic acidcontaining medium. Fig. S7 shows the amounts of individual components of mature peroxisomes, P6, used for the reconstitution of peroxisomal liposomes PLA to PLD and provides transmission electron micrographs of these liposomes purified by flotation on a multistep sucrose gradient.
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Acknowledgments |
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This work was supported by grants from the Canadian Institutes of Health Research to V.I. Titorenko (MOP-57662) and R.A. Rachubinski (MOP-9208) and by a grant from the Canada Foundation for Innovation to V.I. Titorenko. J.M. Nicaud and M.T. Le Dall were supported by the Institut National de la Recherche Agronomique and by the Centre National de la Recherche Scientifique. R.A. Rachubinski is a Canada Research Chair in Cell Biology and an International Research Scholar of the Howard Hughes Medical Institute.
Submitted: 12 May 2003
Accepted: 20 August 2003
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