©1995 by The American Society for Biochemistry and Molecular Biology, Inc.
Novel Peroxisomal Populations in Subcellular Fractions from Rat Liver
IMPLICATIONS FOR PEROXISOME STRUCTURE AND BIOGENESIS (*)

(Received for publication, April 4, 1994; and in revised form, September 13, 1994)

Mona Wilcke (1) (3) Kjell Hultenby (2) Stefan E. H. Alexson (3)(§)

From the  (1)Department of Metabolic Research, The Wenner-Gren Institute, Arrhenius Laboratories F3, Stockholm University, Stockholm and the (2)Clinical Research Center, Huddinge Hospital, Huddinge and (3)Department of Clinical Chemistry, Karolinska Institutet, Huddinge University Hospital, Huddinge, Sweden

ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
FOOTNOTES
ACKNOWLEDGEMENTS
REFERENCES

ABSTRACT

According to current concepts, new peroxisomes are formed by division of pre-existing peroxisomes or by budding from a peroxisomal reticulum. Recent cytochemical and biochemical data indicate that protein content in peroxisomes are heterogenous and that import of newly synthesized proteins may be restricted to certain protein import-competent peroxisomal subcompartments (Yamamoto, K., and Fahimi, H. D.(1987) J. Cell Biol. 105, 713-722; Heinemann, P., and Just, W. W.(1992) FEBS Lett. 300, 179-182; Lüers, G., Hashimoto, T., Fahimi, H. D., and Völkl, A.(1993) J. Cell Biol. 121, 1271-1280).

We have observed that substantial amounts of peroxisomal proteins are found together with ``microsomes'' (100,000 times g pellet) after subcellular fractionation of rat liver homogenates. In this study we have investigated the origin of these peroxisomal proteins by modified gradient centrifugation procedures in Nycodenz and by analysis of enzyme activity distributions, Western blotting, and immunoelectron microscopy. It is concluded that much of this material is confined to novel populations of ``peroxisomes.'' Immunocytochemistry on gradient fractions showed that some vesicles were enriched in acyl-CoA oxidase and peroxisomal multifunctional enzyme (``catalase-negative'') whereas others were enriched in catalase and thiolase (``acyl-CoA oxidase-negative''). Double immunolabeling experiments verified the strong heterogeneity in the protein contents of these vesicles and also identified peroxisomes varying in size from about 0.5 µm (``normal peroxisomes'') to extremely small vesicles of less than 100 nm in diameter. The possibility that these vesicles may be related to different subcompartments of a larger peroxisomal structure involved in protein import and biogenesis will be discussed.


INTRODUCTION

Peroxisomes are nearly ubiquitous organelles, generally spherical in shape with a finely granular matrix surrounded by a single membrane. The abundance, size, and appearance in electron microscopy of peroxisomes vary considerably. Peroxisomes contain enzymes which produce and degrade hydrogen peroxide, and it is clear that fatty acid degradation is a common function of peroxisomes in eukaryotic cells (for reviews, see (1) and (2) ). Earlier studies on the incorporation and turnover of peroxisomal proteins suggested that peroxisomes form a homogenous population without any signs of maturation and with an apparently random degradation of the organelle(3, 4) . Subsequent studies showed that peroxisomal proteins are synthesized on unbound ribosomes and appear in the cytosol before import into pre-existing organelles(4, 5, 6, 7) . These data, together with cytochemical studies(8) , showed that the biogenesis of peroxisomes is independent of ER. (^1)The current view implies that peroxisomes are formed by division of preexisting organelles or by budding from a peroxisomal reticulum(9, 10) . The finding that proliferation of peroxisomes results in a heterogenous induction of peroxisomal proteins (11) may offer a useful tool in the exploration of the mechanisms of peroxisome biogenesis. Several reports have now demonstrated profound heterogeneity under proliferative conditions. The first biochemical indications were obtained by analytical differential centrifugation demonstrating that clofibrate treatment induced a polydispersity of peroxisomes in rat liver(12) . Similar polydispersities in rat liver occur after thyroxine treatment(13) , ischemia reperfusion(14) , and cold exposure(15) . A catalase-negative subpopulation of peroxisomes, which was induced by clofibrate treatment, was identified in mouse liver by subcellular fractionation (16) . In addition, cytochemical studies have indicated heterogenous labeling of peroxisomal proteins(17, 18, 19) .

Recent studies on inherited peroxisomal disorders and peroxisome assembly mutants have added much new information on the structure, function, and biogenesis of peroxisomes. The first ultrastructural observations on Zellweger syndrome indicated that the defect may be due to a total lack of peroxisomes(20) . However, subsequent studies utilizing immunocytochemistry demonstrated that peroxisomal membranes are present in cells of Zellweger patients(21, 22) , and subcellular fractionations of fibroblasts from Zellweger patients demonstrated that peroxisomal proteins are particulate to varying extents(23, 24) . Some fibroblast cell lines from Zellweger patients were able to import thiolase into peroxisomes (25) but failed to import proteins containing the carboxyl-terminal-SKL targeting signal(26) . Thus, studies on Zellweger patients have provided evidence of at least two, possibly three, distinct pathways for import of peroxisomal matrix proteins. Immunocytochemical studies on the heterogeneity of peroxisomes, and the characterization of peroxisome assembly mutants from various yeasts have shed light on both functional and developmental aspects of peroxisomes(27, 28, 29, 30, 31) . Taken together these observations now call for a revision of present peroxisome biogenesis models. A review presenting current concepts on the ultrastructural basis of the biogenesis of peroxisomes was recently published(32) .

In our earlier unpublished experiments on subcellular fractionations of liver homogenates from di(2-ethylhexyl)phthalate (DEHP)-treated rats, we found substantial amounts of peroxisomal proteins recovered in the microsomal fraction. In the present study we have further fractionated microsomal fractions by modified gradient centrifugations in Nycodenz. The enzyme activity distributions of the gradients were analyzed by Western blotting and fractions enriched in peroxisomal proteins were analyzed by immunoelectron microscopy. We conclude that with the present protocol we can separate several classes of ``peroxisomes'' from livers of DEHP-treated rats (that can be distinguished by different protein contents and sedimentation properties which are interpreted to represent peroxisomal subcompartments, probably formed during homogenization of the tissue), rather than populations of peroxisomes. These subcompartments are discussed in relation to peroxisome structure and biogenesis.


EXPERIMENTAL PROCEDURES

Materials

DEHP was obtained from Aldrich (Steinheim, Germany). Fatty acyl-CoA esters, o-nitrophenyl acetate, cytochrome c (type II from horse heart), and horseradish peroxidase were all obtained from Sigma. Nycodenz was purchased from Nycomed AS (Oslo, Norway). ECL chemiluminescence detection kit and nitrocellulose membranes were from Amersham (Buckinghamshire, United Kingdom), and Percoll was obtained from Pharmacia (Uppsala, Sweden).

Methods

Animals

Male Sprague-Dawley rats were obtained from Eklunds (Stockholm, Sweden). Rats were fed ordinary laboratory chow (R3, Ewos, Södertälje, Sweden) or chow supplemented with 2% (w/w) of DEHP for at least 10 days in order to induce peroxisomes. After fasting overnight, the animals were sacrificed by CO(2) anesthesia followed by decapitation.

Subcellular Fractionations

The livers were homogenized with four up and down strokes in a Potter-Elvehjem type homogenizer in ice-cold 0.25 M sucrose, containing 10 mM Tris-HCl (pH 7.4), 1 mM EDTA (pH 7.4), and 0.1% ethanol. The homogenates were diluted to 20% (w/v) and centrifuged twice for 10 min at 750 times g to obtain the nuclear fraction (N-fraction). A heavy mitochondrial fraction (HM fraction) was prepared by centrifugation of the combined supernatants at 3,670 times g for 10 min, followed by re-centrifugation of the resuspended pellet. The peroxisome-enriched light-mitochondrial fraction (LM fraction) was prepared by centrifugation at 24,500 times g for 20 min. The pellet was resuspended and centrifuged again. The microsomal fraction (P-fraction) was prepared by centrifugation of the combined supernatants at 48,400 times g for 2 h.

Gradient Centrifugations

Linear Nycodenz gradients were prepared with density ranges of 1.09-1.19 g/ml for fractionations of P-fractions and 1.15-1.25 g/ml for fractionation of LM fractions. Nycodenz solutions were prepared by solving Nycodenz in 250 mM sucrose containing 10 mM Tris, pH 7.4, 1 mM EDTA, and 0.1% ethanol. The total volume of the gradients was 23 ml resting on 8-ml cushions consisting of 60% Nycodenz. The total volume of the applied sample and overlay (consisting of homogenization buffer) was 8 ml, making up to a total volume of 39 ml. Gradient centrifugations of microsomes were performed at 60,000 times g for 60 min, and LM fractions were centrifuged for 35 min at the same g-force in a Beckman VTi 50 rotor with slow acceleration and deceleration. The density of the gradient fractions was calculated from the measured refractive index of gradient fractions.

In some experiments linear Nycodenz gradients with density range of 1.15-1.27 g/ml were prepared and centrifuged for 1, 20, and 29-32 h at 60 000 times g to compare the enzyme distributions after rate sedimentation and equilibrium density centrifugations.

Enzyme Assays

Acyl-CoA oxidase(33) , catalase(34) , esterase(35) , protein(36) , cytochrome c oxidase(37) , 3-hydroxyacyl-CoA dehydrogenase(33) , and acid phosphatase (38) were determined as described.

Gel Electrophoresis and Western Blotting

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) was carried out with 10% acrylamide gels and stained with Coomassie Brilliant Blue or silver. For Western blotting, proteins were separated by SDS-PAGE and transferred to nitrocellulose membranes in a semi-dry blotter for 1 h at 150 mA. The membranes were blocked overnight in TST buffer (50 mM Tris base, 150 mM NaCl, and 0.5% Tween 20) containing 1% bovine serum albumin, followed by incubation of primary antibodies diluted in TST buffer containing 0.1% bovine serum albumin. After washing of the membranes in TST, the membranes were incubated with peroxidase-conjugated goat anti-rabbit IgG and visualized by a chemiluminescence system.

Preparation of Antibodies

Antibodies against PMP70 were prepared by immunization of rabbits with a synthetic peptide corresponding to a predicted cytosolic domain (amino acids 403-417) of the sequence reported by Kamijo et al.(1990). The peptide was synthesized with an extra NH(2)-terminal cysteine which was used for the coupling of the peptide to activated keyhole limpet hemocyanin. The preparation and characterization of this antibody is described elsewhere. (^2)

Antibodies against peroxisomal acyl-CoA oxidase (anti-Aox) and thiolase (anti-thiolase) were kindly provided by Dr. W. W. Just. Affinity-purified antibodies against peroxisomal MFE (anti-MFE) were kindly provided by Dr. J. K. Hiltunen.

Catalase was purified from livers of DEHP-treated rats. Briefly, after solubilization and centrifugation (at 200 000 times g for 45 min in a Beckman TL-100 table top ultracentrifuge) of isolated peroxisomes, the matrix fraction was applied to chromatography on a DEAE column. The bound activity was eluted with a linear NaCl gradient, and the fractions containing activity were pooled and precipitated by 35% (NH(4))(2)SO(4). The pellet was dissolved in 50 mM Tris buffer containing 200 mM NaCl and subjected to size exclusion chromatography in Sephacryl S-300 (Pharmacia) at a flow rate of 0.8 ml/min. The fractions containing activity were precipitated with 65% (NH(4))(2)SO(4) and applied to a MEMSEP 1010 (DEAE cartridge, Millipore). Catalase activity was eluted with a linear NaCl gradient (0-0.5 M) in 50 mM Tris-HCl (pH 7.4). One rabbit (of the Loop strain) was immunized intramuscularly with 330 µg of purified catalase emulsified with Freund's complete adjuvant, followed by three booster injections of 165 µg of catalase protein emulsified with Freund's incomplete adjuvant. The rabbit was bled from the ear vein at 2-week intervals after the third booster injection, and sera were prepared. This antiserum was monospecific for catalase at dilutions up to 1:500 000 on Western blot. (^3)

Immunocytochemistry

Aliquots of gradient fractions were immediately embedded in an equal volume of 8% luke warm gelatin (Merck, Darmstadt, Germany) and allowed to congeal. The samples were fixed in 1% paraformaldehyde in 0.1 M phosphate buffer (pH 7.4), overnight at 4 °C. Small pieces were infiltrated with 2.0 M sucrose containing 15% polyvinylpyrrolidone and frozen in liquid nitrogen. Sectioning was performed according to (39) at -110 °C. Sections were placed on carbon-reinforced formvar-coated 50-mesh grids and placed directly on drops of 2% bovine serum albumin (Fraction V, Sigma) in 0.1 M phosphate buffer (pH 7.4), containing 20 mM glycine for 30 min. Subsequently, the sections were incubated with the primary antibody for 2 h in a humidified chamber at room temperature. All antibodies were diluted in 0.1 M phosphate buffer (pH 7.4), containing 0.1% bovine serum albumin (PBB). The sections were thoroughly washed in PBB, and bound antibodies were detected with protein A coated with 10-nm gold (Janssens, Olen, Belgium, diluted 1:100). In the double labeling procedure, the protein A was added to a final concentration of 20 µg/ml for at least 5 min, after which the sections were rinsed in PBB and incubated with the second antibodies. Sections were then washed, contrasted by uranyl acetate-oxalate for 5 min, and embedded in 2% methyl cellulose containing 0.3% uranyl acetate. The grids were then examined in a Philips 420 electron microscope at 80 kV. Specificity of labeling was checked with normal peroxisomes prepared by centrifugation of LM fractions (from DEHP-treated rats) in Nycodenz gradient fractions and with pre-immune rabbit antiserum.


RESULTS

Subcellular Fractionation

After fractionation of rat liver homogenates into nuclear, HM, LM, microsomal, and supernatant fractions, it was noted that substantial amounts of the peroxisomal enzymes catalase and Aox were present both in the LM fractions and microsomal fractions. After DEHP treatment, almost 10% of the recovered Aox activity was found in the microsomal fractions, and about 17% of the activity was found in the LM fractions. Since DEHP treatment causes a 10-20-fold induction of peroxisomal beta-oxidation enzymes(40) , the amount of peroxisomal Aox and catalase sedimenting with microsomes must be considered to be substantial. In order to study eventual heterogeneity of peroxisomes, LM fractions and microsomal fractions were further fractionated by gradient centrifugation, and enzyme distributions were compared by means of enzyme activity determinations, Western blot analysis and immunoelectron microscopy.

Fractionation of LM Fractions by Nycodenz Gradient Centrifugation

LM fractions prepared from livers of DEHP-treated rats were centrifuged in Nycodenz gradients according to standard procedures (see ``Methods''). Electron microscopy on the peak peroxisomal fraction showed a mixture of peroxisomes, some containing crystalloid cores of urate oxidase (not shown). The distributions of catalase and Aox in the gradients were as expected with most of the activities found at high density (fractions 3 and 4) with small additional peaks near the top of the gradient (Fig. 1, upper panel). However, it was apparent that the small peaks of catalase and Aox in the low density region were displaced. Catalase activity peaked in fractions 19 and 20 and Aox activity peaked in fractions 17 and 18. These data were confirmed by Western blot analysis of the same fractions (Fig. 1, lower panel) which also showed that the distribution of peroxisomal thiolase closely resembled the distribution of catalase. Western blot analysis with anti-PMP70 showed signal in both the high and low density fractions, indicating that peroxisomal membranes are present also in the low density fractions of the gradient.


Figure 1: Subcellular fractionation of a LM fraction in Nycodenz. A LM fraction was prepared from livers of DEHP-treated rats and subsequently layered on top of a linear Nycodenz density gradient ranging from 25 to 50% Nycodenz. The gradient was centrifuged at 60,000 times g for 35 min, and the gradient was fractionated from the bottom (left to right). The gradient was analyzed for catalase and Aox activities, and aliquots of the fractions were analyzed by Western blotting after SDS-PAGE. The blots were probed with primary antibodies against catalase, Aox, PMP70, and peroxisomal thiolase.



Immunoelectron Microscopy of Peroxisomal Peak Fractions after Nycodenz Gradient Centrifugation

Immunocytochemistry on the high density gradient fractions showed intense labeling with all antibodies tested (anti-MFE, anti-thiolase, anti-catalase, and anti-PMP70) (data not shown). Additionally, the low density fraction containing highest Aox activity was analyzed by immunoelectron microscopy (Fig. 2). The immunogold labeling was highly specific for vesicular structures. The antibody labeling was intense in the labeled vesicles, whereas other vesicles were devoid of labeling, as could be expected from the presence of microsomes in this region of the gradient. Double labeling experiments showed profound heterogeneity in the ratios of the different peroxisomal proteins. Sections incubated with anti-MFE and anti-catalase showed varying ratios of the number of gold particles between different structures. Some were strongly positive for catalase with relatively weak labeling for MFE (Fig. 2a). Other vesicles were strongly labeled for MFE and weakly labeled for catalase (Fig. 2b). Similar data were obtained in double labeling experiments with anti-Aox and anti-catalase (Fig. 2. c and d) in which Aox labeling resembled labeling for MFE. Double labeling experiments with anti-PMP70, compared with anti-catalase and anti-Aox, verified that the immunolabelings were confined to vesicles which were labeled for the peroxisome-specific membrane protein PMP70 (Fig. 2, e-g). In some cases we observed structures which were very strongly labeled for PMP70, although the morphology of these structures more resembled aggregates (Fig. 2g). It is possible that these structures are related to the ``double-membraned loops'' that were recently described by Baumgart et al.(18) .


Figure 2: Immunoelectron microscopy on low density peroxisomes isolated by centrifugation of a LM fraction in Nycodenz. The low density fraction containing highest Aox activity was prepared for immunoelectron microscopy as described under ``Methods.'' Double labeling experiments were carried out with anti-catalase (5-nm gold particles) and anti-MFE (10-nm gold particles, a and b), anti-Aox (10-nm gold particles, c and d), and anti-PMP70 (10-nm gold particles, e-g). Bar = 0.2 µm.



Centrifugation of Microsomal Fractions in Nycodenz Gradients

Microsomal fractions were fractionated by centrifugation for 60 min into Nycodenz gradients (density range 1.09-1.19 g/ml). The distributions of peroxisomal and marker enzyme activities are shown in Fig. 3(upper panel). Most of the gradient protein was found near the top, almost completely co-sedimenting with esterase activity. Catalase separated into two main peaks, the first was a high density peak in fractions 1-4 corresponding to a density of about 1.20 g/ml, with a shoulder of catalase at densities of 1.15-1.20 g/ml. The second peak banded at about 1.06-1.10 g/ml. In these gradients it was obvious that catalase did not band at distinct densities but rather distributed in broad peaks, indicating a profound heterogeneity. Aox activity showed a bimodal distribution with some of the activity found in fractions 1-4 and most of the activity found at a density of about 1.12 g/ml (fractions 18 and 19). In contrast to catalase, most of the activities of Aox (about 75%) and 3-hydroxyacyl-CoA dehydrogenase were associated with the low density peaks near the top. The mean distribution of catalase between the high and low density peaks was 35% (range: 25-52%) and 27% (range: 21-30%), respectively; and the mean distribution of Aox was 18% (range: 10-28%) and 53% (range: 41-75%), respectively (means of four different fractionations). Marker enzymes for mitochondria and lysosomes (cytochrome oxidase and acid phosphatase, respectively) distributed mainly in fractions between the two peaks of Aox and 3-hydroxyacyl-CoA dehydrogenase, respectively. Esterase, the marker enzyme for microsomes, banded in fractions 18 and 19 similar to Aox and MFE. In control experiments (not shown) microsomal fractions pretreated with different concentrations of Triton X-100 showed that released catalase sedimented to the same fractions as the main low density peak of catalase. Centrifugation of 100,000 times g supernatants showed that ``free'' catalase banded in these fractions. Thus the low density peak of catalase is mainly due to protein which is apparently not associated with vesicles. Fractions corresponding to the high and low density peaks of Aox and MFE activities were analyzed by transmission and immunoelectron microscopy. The morphology of peroxisomal structures found at high density was similar to normal peroxisomes, although the average sizes appeared somewhat smaller with most peroxisomes having diameters less than 0.5 µm. Urate oxidase cores appeared less frequently in these peroxisomes (data not shown). The structure of the vesicles observed in the low density peak enriched in Aox and MFE resembled smooth ER and the sizes of these vesicles were usually below 100 nm.


Figure 3: Subcellular fractionation of a microsomal fraction in Nycodenz. Microsomal fractions were prepared from livers of rats treated with DEHP, layered on top of linear Nycodenz gradients, and centrifuged at 60,000 times g for 60 min. Fractions were collected from the bottom of the tubes (from left to right) and analyzed for enzyme activities (upper panels). It should be noted that the amount of mitochondria found in the microsomal fractions is very low and does not contribute significantly to the distributions of protein and 3-hydroxyacyl-CoA dehydrogenase activity in the gradients. Gradient fractions (10 µl of each) were electrophoresed by SDS-PAGE in 10% acrylamide gels. The separated proteins were transferred to nitrocellulose membranes and probed with the indicated antibodies. The blots were visualized by a chemiluminescence method (lower panels).



Western blot analysis of the same gradient fractions (Fig. 3, lower panel) verified the enzymatic data for catalase and 3-hydroxyacyl-CoA dehydrogenase (MFE). In addition, Western blotting showed that peroxisomal thiolase distributed similarly to catalase and that substantial amounts of PMP70 were found in fractions 15-21, indicating the presence of peroxisomal membranes in these fractions. The very close association of most of the Aox and MFE found in the microsomal fraction with microsomes after gradient centrifugation demonstrates that these proteins are not associated with peroxisomes of normal size and/or density. From these experiments it was not possible to rule out the possibility that Aox and MFE were sticking to microsomes. However, in parallel experiments where microsomal fractions were centrifuged in Percoll gradients, a clear separation of Aox activity from microsomes was obtained (data not shown).

The results shown are typical of more than 15 fractionation experiments on enzyme activity distributions and at least 5 experiments by Western blot analysis.

Immunoelectron Microscopy on High Density Nycodenz Fractions

Immunocytochemistry on high density fractions from Nycodenz gradients (corresponding to fractions 1-3 in Fig. 3a) showed that the labeling with the different antibodies were highly specific (Fig. 4). Antibodies to catalase specifically labeled membrane surrounded organelles evenly (Fig. 4a) as did antibodies to MFE (Fig. 4b). Labeling for PMP70 was confined to the membranous structures of the particles (Fig. 4c). The immunocytochemistry on these gradient fractions showed highly specific labeling of vesicles with average diameters of about 0.2-0.3 µm. It thus appears to be an enrichment of smaller peroxisomes in the microsomal fraction as compared with the LM fraction.


Figure 4: Immunoelectron microscopy on high density peroxisomes isolated by centrifugation of microsomal fractions in Nycodenz. Gradient fractions corresponding to the high density peak of peroxisomes obtained after centrifugation of microsomal fractions in Nycodenz were prepared and incubated with antibodies and colloidal gold as described under ``Methods.'' a, incubated with anti-catalase; b, incubated with anti-peroxisomal MFE; c, incubated with anti-PMP70. The peroxisomes found at high densities were generally 0.1-0.5 µm in diameter. Bar = 0.2 µm.



Immunoelectron Microscopy of Low Density Fractions from Nycodenz Gradients

The low density fractions from Nycodenz gradients contained mainly microsomes, and it was not possible to identify peroxisomes by means of morphology. However, immunolabeling experiments showed that few particles, with a ghost-like appearance, and some particles, with a more granular matrix, contained peroxisomal proteins. The structures of these particles resembled peroxisomes in that they were surrounded by a single membrane, relatively round in shape with some of them also having irregular protrusions (Fig. 5b) as reported by Lüers et al.(41) . In double immunolabeling experiments, some particles were positively labeled for catalase but were apparently negative for MFE (Fig. 5a), whereas other particles were labeled by both antibodies (Fig. 5b). Heterogeneous staining was also observed in double labeling experiments for MFE and thiolase, demonstrating MFE-positive particles (apparently negative for thiolase) and particles containing both enzymes (Fig. 5, c and d, respectively). We also observed heterogeneities in the labeling for catalase and PMP70 (Fig. 5, e-f), and catalase and thiolase (Fig. 5, g and h). Fig. 5g shows a catalase-positive particle, lacking thiolase, and Fig. 5h shows a particle labeled for both proteins. The ratio of labeling for thiolase and catalase was very different in various particles (data not shown). Although the frequency of labeled vesicles were low in these fractions, the antibody labeling was highly specific for vesicular structures. The low frequency is due to the small amount of peroxisomal vesicles present in these fractions which mainly contain microsomes and to the method of choice for preparing the fractions for immunoelectron microscopy. We considered it important to avoid fixation of dilute fractions, followed by pelleting by high speed centrifugation. Instead, the milder treatment of embedding peroxisomes in gelatin and fixing in formaldehyde probably reduced the release of proteins from the vesicles, although this procedure resulted in a final 2-fold dilution of the fraction. It should also be emphasized that the immunolabeling experiments were carried out on consecutive cryosections from the same block within each experiment.


Figure 5: Immunoelectron microscopy on low density gradient fractions after centrifugation of microsomal fractions in Nycodenz. Microsomal fractions were centrifuged in Nycodenz gradients, and fractions corresponding to the low density peak of Aox (also containing catalase) were prepared and incubated with antibodies and colloidal gold as described under ``Methods.'' Labeled for: a and b, catalase (10 nm gold particles) and MFE (5 nm gold particles), showing a MFE-negative vesicle labeled for catalase (a) and a double-labeled vesicle positive for MFE (arrows, b); c and d, MFE (10-nm gold particles) and thiolase (5-nm gold particles). MFE-positive vesicles lacking thiolase (c) and vesicles labeled for both proteins (thiolase at arrows, d); e and f, catalase (10-nm gold particles) and PMP70 (5-nm gold particles), catalase-positive particles also labeled for PMP70 (arrows, e) and membrane material labeled for PMP70 apparently lacking catalase (f); g and h, catalase (10-nm gold particles) and thiolase (5-nm gold particles), catalase-positive particles lacking thiolase (g) and vesicles strongly labeled for thiolase (arrows, h) containing catalase. The sizes of the labeled vesicles are generally below 200 nm. Bar = 0.1 µm.



The immunolabeling experiments on the low density fractions clearly demonstrated that the peroxisomal proteins found at low density during gradient centrifugation are at least partially confined to membrane surrounded vesicles, rather than becoming released proteins in soluble form or sticking to other membranes such as microsomes. The double labeling experiments also indicated the existence of profound heterogeneities in the contents of peroxisomal enzymes in these vesicles.

Effect of Prolonged Density Gradient Centrifugation of Microsomal Fractions

In order to investigate the effect of centrifugation of the various peroxisomal subcompartments to their equilibrium in Nycodenz, microsomal fractions were divided into three aliquots and layered on top of three identical gradients that were centrifuged for 1, 20, and 29 h, respectively. The gradients were fractionated and catalase and Aox activities were measured. Fig. 6shows the enzyme distributions after 1 and 20 h of centrifugation from one of these experiments. The enzyme distributions after 20- and 29-h centrifugation were identical, indicating that these enzymes had reached equilibrium at 20 h. After 1-h centrifugation, catalase was distributed into two peaks of high (about 1.22 g/ml) and low (about 1.08 g/ml) densities. After 20 h, catalase activity was found in only one, although rather broad, peak at a density of about 1.16-1.22 g/ml. Essentially no catalase activity was found at densities higher than 1.25 g/ml.


Figure 6: Comparison of the sedimentation of catalase and Aox after centrifugation of microsomal fractions for 1 and 20 h in Nycodenz gradients. Liver microsomal fractions were prepared from DEHP-treated rats and centrifuged in linear Nycodenz gradients as described under ``Methods.'' The microsomal fraction was divided into three aliquots which were layered on top of Nycodenz gradients and centrifuged for 1, 20, and 29 h (not shown), respectively. The gradients were fractionated from the bottom (left to right) and analyzed for catalase and Aox activities, protein, and density. a, centrifugation for 1 h; b, centrifugation for 20 h.



Aox was also distributed into two peaks after 1-h centrifugation, one peak at about 1.22 g/ml and the other peak at about 1.15 g/ml, clearly different from catalase. After 20 and 29 h of centrifugation, Aox still tended to band in two peaks, one at about 1.22 g/ml and the other still at about 1.15 g/ml. However, it is evident that a larger part of the Aox activity also sedimented to a density of 1.25 g/ml or more, clearly higher than catalase.

These experiments imply that there exists differences also in the equilibrium densities among the vesicles containing catalase and Aox.


DISCUSSION

Isolation and characterization of peroxisomes have been hampered by the facts that peroxisomes have densities in gradient media similar to, or only slightly different from, the densities of other organelles and because peroxisomal protein normally constitutes only 2% (in rat liver) or less of the cellular protein. It is also well established that peroxisomes are leaky and that the leakiness is selective. Catalase and thiolase are very easily released, whereas other proteins, such as Aox and the MFE, are much less prone to be released(42) . Other peroxisomal proteins may be expected to cover the range. This leakiness appears to be time-dependent and dependent on the treatment of peroxisomes. It has also been implied that peroxisomes are very sensitive to hydrostatic pressure, since enzyme distributions in gradients often show activity of peroxisomal proteins throughout the gradients. However, in the light of the results presented here, the peroxisomal proteins found in the low density fractions are at least partially present in vesicles.

Identification and Separation of Different Peroxisomal Subcompartments

In this study we report on the separation and identification of peroxisomal subcompartments (``populations'') of different sizes, equilibrium densities, and heterogenous contents of peroxisomal proteins after centrifugation of LM and microsomal fractions in Nycodenz gradients. Besides the normal peroxisomes sedimenting to a density of >1.20 g/ml (after fractionation of LM fractions), catalase, Aox, and MFE each separated into two peaks after fractionation of LM and microsomal fractions in Nycodenz gradients. One peak was found at a density around 1.19 g/ml, possibly representing ``immature'' peroxisomes. The second peak sedimented to a lower density. Equilibrium density centrifugations for 20 and 29 h in Nycodenz-sedimented catalase into one broad peak at 1.15-1.22 g/ml, whereas Aox still showed a somewhat bimodal distribution. By electron microscopy it was evident that the structures of the low density organelles were quite different from normal peroxisomes which typically contained crystalline cores and granular matrix, indicating a substantial content of protein. The organelles found at low density, that were labeled with antibodies to peroxisomal proteins, often consisted of a membrane surrounded vesicle in which most of the matrix area was apparently empty and sometimes had only a rim of protein associated with the inside of the membrane. Although it is well established that peroxisomes are leaky, which may result in the formation of ``ghost-like'' peroxisomes, we conclude that the peroxisomal proteins present in the low density fractions are at least in part confined to peroxisomal vesicles. These are not likely to be the result of selective protein release based on the following arguments: (i) Immunogold labeling showed that the peroxisomal proteins were mainly confined to vesicles surrounded by membranes that contained PMP70. (ii) Our previous findings showed that thiolase and catalase are the most leaky enzymes of those studied so far(42) , which was supported by subsequent studies of Thompson and Krisans(43) . It is unlikely that extraction of peroxisomal proteins should result in the formation of catalase- and thiolase-positive particles which are nearly devoid of MFE and Aox, proteins known to be retained in peroxisomes. However, independent of the mechanisms for release of proteins, any heterogeneity in the protein contents between various vesicles indicates hetrogeneity of peroxisomal structures. (iii) The peroxisomal ``ghosts'' isolated after extraction with pyrophosphate (at pH 9) still contained the crystalline cores of urate oxidase(42) . The low density particles found in the present study lack such crystalline cores, but cores were observed in peroxisomes banding at high density in these gradients. The very small size of the vesicles lacking cores also argues against the presence of cores in these vesicles. (iv) Although we have looked only at a limited number of peroxisomal proteins so far, there appears to be a rather strong correlation between the selective protein content of these vesicles and proposed targeting mechanisms for protein import.

Formation of Peroxisomal Subcompartments by Fragmentation of a Peroxisome Reticulum

So how do these membranous structures originate? The classical model for peroxisome biogenesis is based on the assumption that peroxisomes are formed by budding from ER. This assumption was derived from ultrastructural studies describing apparent membrane connections between peroxisomes and ER (44, 45, 46) and similarities in membrane composition(47) . Later biochemical data (on import of peroxisomal proteins) as well as cytochemistry on catalase and glucose 6-phosphatase, support a concept where peroxisomal proteins are synthesized on free ribosomes and posttranslationally imported into peroxisomes(5, 6, 7, 8) . Recent ultrastructural work showed that peroxisomes are, at least temporarily, interconnected to form a reticulum(10, 48, 49, 50, 51, 52) . Staining of peroxisomes for various peroxisomal enzymes have demonstrated heterogeneity both in the intensity between different peroxisomes and in the staining intensity in interconnections(18, 51, 52, 53) . The weak labeling of interconnections suggest that a barrier may exist that prevents diffusion of matrix proteins in these interconnections or that matrix proteins are transported through the interconnections and accumulating in the buds.

The most likely interpretation of our results is that the different peroxisomal subcompartments described here are formed during homogenization of a peroxisome reticulum. Fig. 7illustrates how the different peroxisomal populations, found during subcellular fractionation of rat liver homogenates, may form by vesiculation. Besides normal peroxisomes (1), the peroxisomes found at a density of about 1.19 g/ml (corresponding to the high density peaks after centrifugation of microsomes) may be formed by the pinching off of buds that are not yet complete (2). The very small vesicles with heterogenous protein contents may be formed by vesiculation of the ``stalks'' connecting the buds to the ``body'' of the reticulum (3, 4, 5). The heterogeneity of these vesicles could be explained by a nonrandom distribution of the receptors involved in binding and import of peroxisomal proteins. Such a heterogeneity could also explain why we are able to isolate different populations of peroxisomes enriched in proteins containing different PTS.


Figure 7: Hypothetical model for the formation of various populations of peroxisomes during subcellular fractionation of rat liver. If peroxisomes are formed by budding from a peroxisomal reticulum, fragmentation of such a reticulum by homogenization could explain the observed heterogeneity of vesicles containing peroxisomal proteins. Normal peroxisomes (1) are formed by budding and correspond to the bulk of peroxisomes found at densities around 1.24 g/ml. Fragmentation of ``buds'' (at the stalks) may thus correspond to ``immature'' peroxisomes with a complete set of peroxisomal proteins sedimenting to about 1.19 g/ml (2). Fragmentation of the stalks can be expected to result in the formation of ``microperoxisomes'' (3, Aox- and MFE-enriched vesicles; 4, catalase-enriched vesicles; and 5 thiolase-enriched vesicles). The heterogeneity could be explained by uneven distribution of receptors for import of peroxisomal proteins. It is possible that the ``body'' of the reticulum forms another population (?) of vesicles that is not yet identified due to lack of an appropriate marker.



Recent information concerning peroxisome structure and protein content in Zellweger syndrome has demonstrated that peroxisomal particles may have a selective set of proteins. The presence of peroxisomal ghosts containing peroxisomal membrane proteins(21, 22) , and the finding of a low density particle (w-particle) in fibroblasts of Zellweger patients which contained catalase(24) , also demonstrated a heterogeneity in peroxisomal protein content. Suzuki et al.(23) showed by indirect immunofluorescence staining and subcellular fractionation that some catalase and Aox was associated with organelles in some fibroblasts from Zellweger patients. Interestingly, substantial amounts of 3-ketoacyl-CoA thiolase and PMP70 were detected in organelles in all cells. These data suggest that the transport and processing of peroxisomal proteins carrying different PTS's are different, which is compatible with the present knowledge on sequence requirements for import of these proteins into peroxisomes. Several peroxisomal proteins contain a conserved COOH-terminal -Ser-Lys-Leu sequence, or acceptable variations of this sequence, that directs proteins to microbodies in mammalian cells, yeasts, plants, insects, and trypanosomes(54, 55, 56, 57, 58, 59, 60, 61) . Import of proteins containing this conserved tripeptide is probably mediated by a common receptor. However, import of peroxisomal thiolase appears to be dependent on a cleavable NH(2)-terminal presequence(59, 62, 63) . The targeting sequence for catalase is not clear at present. It appears that the internal -Ser-His-Leu-sequence may not target catalase to peroxisomes, as it was demonstrated that the addition of one or two amino acids COOH-terminal of the -SKL PTS abolishes import into peroxisomes(64) . This implies that catalase may follow another, still unknown, mechanism for targeting and import.

Fahimi et al.(32) suggested that synthesis and incorporation of membranous material, containing PMP70, is an early event, resulting in the growth of the membrane followed by import of newly synthesized proteins. Our results are compatible with such a model favoring synthesis of membrane material as an early event which is followed by incorporation of highly expressed beta-oxidation enzymes (Aox, MFE, and thiolase). This is reflected by the high content of Aox and MFE in the low density compartment after DEHP treatment. However, neither this model nor our present results can fully explain the detailed mechanism for the heterogenous distribution of peroxisomal proteins in different peroxisomes.

Proposed Model for Peroxisome Biogenesis

Based on previous data in the literature and our present data, we favor a model where peroxisomes are formed by budding from a peroxisome reticulum (Fig. 7). Import of proteins may be restricted to specific locations in the reticulum with Aox and the multifunctional protein, both carrying a COOH-terminal -SKL PTS, and being imported at a common site, possibly in a ``body'' of the reticulum. A zonation of protein import could be due to nonrandom distributions of receptors for different PTS or differences in lipid composition affecting protein targeting. Catalase and thiolase, probably carrying different PTS's, may be imported at different sites by specific receptors. However, the nature of this hypothetical peroxisomal reticulum remains to be characterized. In a recent study, Ohno and Fujii (10) showed that in cultured hepatocytes, the peroxisomal reticulum consisted of a smooth ER-like structure which they termed ``peroxisome-forming sheet.'' By DAB staining for catalase they identified three types of peroxisomal segments: one type strongly DAB-positive, a second type containing weak DAB reaction, and the third type of sheet-like extensions containing no DAB reaction. They concluded that the smooth membranous structures lacking DAB reaction may represent the peroxisome forming sheet. Assuming that import of peroxisomal proteins is restricted to specific locations in such a reticulum, this model can explain the observations of small particles (formed by homogenization of the narrow interconnections) containing mainly Aox, MFE, and PMP70. It may also explain the formation of catalase/thiolase-rich vesicles formed by homogenization, assuming that the import of these proteins takes place, e.g. closer to the ``peroxisome bud.'' Urate oxidase import is also mediated by the -SKL PTS(65) , but crystalloid cores may only form in the growing buds after substantial accumulation of the protein. Withdrawal of peroxisome proliferators quickly reverses the effects of most parameters. It should be noted that heterogenous staining for at least catalase appears after withdrawal of clofibrate treatment in mice (66) . Such a heterogeneity could be explained by deterioration of the reticulum containing immature buds. If there is only a limited import of protein into released peroxisomes, the heterogeneity may persist for some time. One could speculate that the limited success so far in achieving a high efficiency of in vitro import of peroxisomal proteins into normal peroxisomes may not be due to leakiness of these organelles, but may in fact be due to the presence of few adequate PTS receptors.

Strong support for this model is also obtained from studies in yeasts where a functional heterogeneity among microbodies has been suggested to exist(27, 28) . These studies demonstrated that addition of methanol to Candida boidinii cells grown on oleic acid resulted in incorporation of newly synthesized proteins into smaller, presumably ``immature'' peroxisomes. This heterogeneity was only transient, and continuous cultures grown on a mixture of oleic acid and methanol at steady-state conditions showed that both the enzymes of the beta-oxidation pathway and methanol metabolism were found in one and the same compartment(28) . This transient heterogeneity is likely to reflect an incompetence of mature peroxisomes to incorporate newly synthesized proteins. This idea is further supported by the recent pulse-chase experiments on in vitro import which showed that Aox is imported into a compartment of intermediate density (pulse) and subsequently chased into a high density compartment (normal peroxisomes)(67) . During our present investigation, Lüers et al.(41) reported on a modified scheme of subcellular fractionation of regenerating livers which resulted in the separation of two apparently different populations of peroxisomes. They found that [S]methionine injected into rats was first incorporated into a ``light peroxisomal'' fraction (5-30 min after injection) and that the label appeared in the ``heavy peroxisomal'' fraction at 90 min.

The formation of a peroxisomal reticulum distinguishes peroxisomes from the other intracellular membrane systems of mitochondria, chloroplasts, and lysosomes. Mitochondria are generally believed to represent an ancient endosymbiont, probably after ``invasion'' of a bacteria. ER and lysosomal proteins are synthesized by co-translational insertion of the proteins into ER followed by glycosylation. ER resident proteins are apparently retrieved in the ER by a receptor-mediated process involving a COOH-terminal tetrapeptide (KDEL or HXEL) (see (68) for review). Lysosomal proteins are transported to the Golgi apparatus where the lysosomes are formed by budding of vesicles containing lysosomal proteins which are then transported to the acidic compartment. Peroxisomal proteins may be transported from the site of import (at the peroxisome-forming sheet) by ``bulk flow'' (similar to protein transport in the rough ER) to the buds. Experiments aimed at functionally characterizing the different peroxisomal subcompartments, which are under way in our laboratory, may clarify the origin and functions of these vesicles. It still remains to biochemically identify the peroxisome-forming sheet during subcellular fractionation of rat liver homogenates.


FOOTNOTES

*
This work was supported by grants from the Swedish Natural Science Research Council, The Bank of Sweden Tercentenary Foundation, ``Magnus Bergvalls Stiftelse,'' ``Lars Hiertas Minne,'' ``Tore Nilsons Fond för Medicinsk Forskning,'' ``Hierta-Retzius' fond för vetenskaplig forskning,'' and the ``Ax:son Johnsons Stiftelse.'' The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore by hereby marked ``advertisement'' in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§
To whom correspondence should be addressed: Dept. of Clinical Chemistry, Karolinska Institutet, Huddinge University Hospital, S-141 86 Huddinge, Sweden. Tel.: 46-8-746-1601; Fax: 46-8-746-1698.

(^1)
The abbreviations used are: ER, endoplasmic reticulum; Aox, acyl-CoA oxidase; DEHP, di(2-ethylhexyl)phthalate; MFE, peroxisomal multifunctional enzyme; PBB, phosphate-buffered bovine serum albumin; PMP70, the 70-kDa peroxisomal integral membrane protein; HM, heavy mitochondrial fraction; LM, light mitochondrial fraction; SKL, Ser-Lys-Leu; PTS, peroxisomal targeting signal; DAB, 3,3`-diaminobenzidine. Note: from the distributions of marker enzymes we conclude that nearly all of the 3-hydroxyacyl-CoA dehydrogenase activity found in the microsomal fraction is due to the peroxisomal multifunctional enzyme (MFE), expressing hydratase/dehydrogenase/isomerase activity(69) . The abbreviation MFE is used to denote this protein when detected with a monospecific antibody in Western blotting and immunoelectron microscopy, whereas 3-hydroxyacyl-CoA dehydrogenase is used when the activity is measured in the gradient fractions.

(^2)
T. Svensson, M. Wilcke, S. Alexson, H. Häyrinen, R. Sormunen, and K. Hiltunen, submitted for publication.

(^3)
A. Messing-Eriksson and S. Alexson, unpublished results.


ACKNOWLEDGEMENTS

We thank Dr. Björn Afzelius for helpful discussions and Dr. Henrik Garoff and Dr. J. Kalervo Hiltunen for critical reading of this manuscript.


REFERENCES

  1. Lazarow, P. B., and Fujiki, Y. (1985) Annu. Rev. Cell Biol. 1, 489-530 [CrossRef]
  2. Lazarow, P. B., and Moser, H. W. (1989) in The Metabolic Basis of Inherited Disease (Scriver, C. R., Beaudet, A. L., Sly, W. S., and Valle, D., eds) pp. 1479-1509, McGraw-Hill, New York
  3. Poole, B., Leighton, F., and de Duve, C. (1969) J. Cell Biol. 41, 536-546 [Abstract/Free Full Text]
  4. Lazarow, P. B., and de Duve, C. (1973) J. Cell Biol. 59, 507-524 [Abstract/Free Full Text]
  5. Goldman, B. M., and Blobel, G. (1978) Proc. Natl. Acad. Sci. U. S. A. 75, 5066-5070 [Abstract]
  6. Fujiki, Y., Rachubinski, R. A., and Lazarow, P. B. (1984) Proc. Natl. Acad. Sci. U. S. A. 81, 7127-7131 [Abstract]
  7. Rachubinski, R. A., Fujiki, Y., Mortensen, R. M., and Lazarow, P. B. (1984) J. Cell Biol. 99, 2241-2246 [Abstract]
  8. Shio, H., and Lazarow, P. B. (1981) J. Histochem. Cytochem. 29, 1263-1272 [Abstract]
  9. Lazarow, P. B., Shio, H., and Robbi, M. (1980) in 31st Mosbach Colloquium. Biological Chemistry of Organelle Formation (Weiss, H., Bucher, T., and Sebald, W., eds) pp. 187-206, Springer-Verlag, New York
  10. Ohno, S., and Fujii, Y. (1990) Histochem. J. 22, 143-154 [Medline] [Order article via Infotrieve]
  11. Lazarow, P. B., and de Duve, C. (1976) Proc. Natl. Acad. Sci. U. S. A. 73, 2043-2046 [Abstract]
  12. Flatmark, T., Christiansen, E. N., and Kryvi, H. (1981) Eur. J. Cell Biol. 24, 62-69 [Medline] [Order article via Infotrieve]
  13. Just, W. W., Hartl, F. U., and Schimassek, H. (1982) Eur. J. Cell Biol. 26, 249-254 [Medline] [Order article via Infotrieve]
  14. Gulati, S., Singh, A. K., Irazu, C., Orak, J., Rajagopalan, P. R., Fitts, C. T., and Singh, I. (1992) Arch. Biochem. Biophys. 295, 90-100 [Medline] [Order article via Infotrieve]
  15. Goglia, F., Liverini, G., Lanni, A., Iossa, S., and Barletta, A. (1989) Exp. Biol. 48, 127-133 [Medline] [Order article via Infotrieve]
  16. Klucis, E., Crane, D. I., Hughes, J. L., Poulos, A., and Masters, C. J. (1991) Biochim. Biophys. Acta 1074, 294-301 [Medline] [Order article via Infotrieve]
  17. Baumgart, E., Stegmeier, K., Schmidt, F. H., and Fahimi, H. D. (1987) Lab. Invest. 56, 554-564 [Medline] [Order article via Infotrieve]
  18. Baumgart, E., Völkl, A., Hashimoto, T., and Fahimi, H. D. (1989) J. Cell Biol. 108, 2221-2231 [Abstract]
  19. Lüers, G., Beier, K., Hashimoto, T., Fahimi, H. D., and Völkl, A. (1990) Eur. J. Cell Biol. 52, 175-184 [Medline] [Order article via Infotrieve]
  20. Goldfischer, S., Moore, C. L., Johnson, A. B., Spiro, A. J., Valsamis, M. P., Wisniewski, H. K., Ritch, R. H., Norton, W. T., Rapin, I., and Gartner, L. M. (1973) Science 182, 62-64 [Medline] [Order article via Infotrieve]
  21. Santos, M. J., Imanaka, T., Shio, H., and Lazarow, P. B. (1988) J. Biol. Chem. 263, 10502-10509 [Abstract/Free Full Text]
  22. Santos, M. J., Imanaka, T., Shio, H., Small, G. M., and Lazarow, P. B. (1988) Science 239, 1536-1538 [Medline] [Order article via Infotrieve]
  23. Suzuki, Y., Shimozawa, N., Yajima, S., Orii, T., Yokota, S., Tashiro, Y., Osumi, T., and Hashimoto, T. (1992) Cell Struct. Funct. 17, 1-8 [Medline] [Order article via Infotrieve]
  24. Aikawa, J., Chen, W., Kelley, R. J., Tada, K., Moser, H. W., and Chen, G. L. (1991) Proc. Natl. Acad. Sci. U. S. A. 88, 10084-10088 [Abstract]
  25. Balfe, A., Hoefler, G., Chen, W. W., and Watkins, P. A. (1990) Pediatr. Res. 27, 304-310 [Abstract]
  26. Walton, P. A., Gould, S. J., Feramisco, J. R., and Subramani, S. (1992) Mol. Cell. Biol. 12, 531-541 [Abstract]
  27. Veenhuis, M., Sulter, G., van der Klei, I., and Harder, W. (1989) Arch. Microbiol. 151, 105-110 [Medline] [Order article via Infotrieve]
  28. Waterman, H. R., Keizer-Gunnink, I., Goodman, J., Harder, W., and Veenhuis, M. (1992) J. Bacteriol. 174, 4057-4063 [Abstract]
  29. Wiebel, F. F., and Kunau, W. H. (1992) Nature 359, 73-76 [CrossRef][Medline] [Order article via Infotrieve]
  30. Waterman, H. R., Titorenko, V. I., Swaving, G. J., Harder, W., and Veenhuis, M. (1993) EMBO J. 12, 4785-4794 [Abstract]
  31. Zhang, J. W., Luckey, C., and Lazarow, P. B. (1993) Mol. Biol. Cell 4, 1351-1359 [Abstract]
  32. Fahimi, H. D., Baumgart, E., and Völkl, A. (1993) Biochimie (Paris) 75, 201-208 [Medline] [Order article via Infotrieve]
  33. Osumi, T., and Hashimoto, H. (1978) Biochem. Biophys. Res. Commun. 83, 479-483 [Medline] [Order article via Infotrieve]
  34. Baudhuin, P., Beaufay, H., Rahman-Li, Y., Zellinger, O. Z., Wattiaux, R., Jacques, P., and de Duve, C. (1964) Biochem. J. 92, 179-184 [Medline] [Order article via Infotrieve]
  35. Beaufay, H. A., Amar-Costesec, A., Feytmans, E., Thines-Sempoux, D., Wibo, M., Robbi, M., and Berthet, J. (1974) J. Cell Biol. 61, 188-200 [Abstract/Free Full Text]
  36. Bradford, M. M. (1976) Anal. Biochem. 72, 248-254 [CrossRef][Medline] [Order article via Infotrieve]
  37. Leighton, F., Poole, B., Beaufay, H., Baudhuin, P., Coffey, J. W., Fowler, S., and de Duve, C. (1968) J. Cell Biol. 37, 482-513 [Abstract/Free Full Text]
  38. Bergmayer, H. V., Gawehn, K., and Grassel, M. (1974) in Methods in Enzymatic Analysis (eds) Vol. 1, pp. 425-522, Academic Press, New York
  39. Tokuyasu, K. T. (1973) J. Cell Biol. 57, 551-565 [Abstract/Free Full Text]
  40. Hashimoto, T. (1982) Ann. N. Y. Acad. Sci. 386, 5-12 [Medline] [Order article via Infotrieve]
  41. Lüers, G., Hashimoto, T., Fahimi, H. D., and Völkl, A. (1993) J. Cell Biol. 121, 1271-1280 [Abstract]
  42. Alexson, S. E. H., Fujiki, Y., Shio, H., and Lazarow, P. B. (1985) J. Cell Biol. 101, 294-304 [Abstract]
  43. Thompson, S. L., and Krisans, S. K. (1990) J. Biol. Chem. 265, 5731-5735 [Abstract/Free Full Text]
  44. Essner, E. (1967) Lab. Invest. 17, 71-87 [Medline] [Order article via Infotrieve]
  45. Novikoff, A. B., and Shin, W. Y. (1964) J. Microsc. (Paris) 3, 187-206
  46. Novikoff, P. M., Novikoff, A. B., Quintana, N., and Davis, C. (1973) J. Histochem. Cytochem. 21, 540-558 [Medline] [Order article via Infotrieve]
  47. Donaldson, R. P., Tolbert, N. E., and Schnarrenberger, C. (1972) Arch. Biochem. Biophys. 152, 199-215 [Medline] [Order article via Infotrieve]
  48. Gorgas, K. (1982) Ann. N. Y. Acad. Sci. 386, 519-522
  49. Gorgas, K. (1984) Anat. Embryol. 169, 261-270 [Medline] [Order article via Infotrieve]
  50. Gorgas, K. (1985) Anat. Embryol. 172, 21-32 [CrossRef][Medline] [Order article via Infotrieve]
  51. Yamamoto, K., and Fahimi, H. D. (1987) J. Cell Biol. 105, 713-722 [Abstract]
  52. Yamamoto, K., and Fahimi, H. D. (1987) Eur. J. Cell Biol. 43, 293-300 [Medline] [Order article via Infotrieve]
  53. Stäubli, W., Schweizer, W., Suter, J., and Weibel, E. R. (1977) J. Cell Biol. 74, 665-689 [Abstract]
  54. Keller, G. A., Gould, S., Deluca, M., and Subramani, S. (1987) Proc. Natl. Acad. Sci. U. S. A. 84, 3264-3268 [Abstract]
  55. Keller, G. A., Krisans, S., Gould, S. J., Sommer, J. M., Wang, C. C., Schliebs, W., Kunau, W., Brody, S., and Subramani, S. (1991) J. Cell Biol. 114, 893-904 [Abstract]
  56. Gould, S. J., Keller, G. A., Hosken, N., Wilkinson, J., and Subramani, S. (1989) J. Cell Biol. 108, 1657-1664 [Abstract]
  57. Gould, S. J., Keller, G. A., Schneider, M., Howell, S. H., Garrard, L. J., Goodman, J. M., Distel, B., Tabak, H., and Subramani, S. (1990) EMBO J. 9, 85-90 [Abstract]
  58. Aitchison, J. D., Murray, W. W., and Rachubinski, R. A. (1991) J. Biol. Chem. 266, 23197-23203 [Abstract/Free Full Text]
  59. Swinkels, B. W., Gould, S. J., and Subramani, S. (1992) FEBS Lett. 305, 133-136 [CrossRef][Medline] [Order article via Infotrieve]
  60. Blattner, J., Swinkels, B., Dörsam, H., Prospero, T., Subramani, S., and Clayton, C. (1992) J. Cell Biol. 119, 1129-1136 [Abstract]
  61. Miura, S., Kasuya-Arai, I., Mori, H., Miyazawa, S., Osumi, T., Hashimoto, T., and Fujiki, Y. (1992) J. Biol. Chem. 267, 14405-14411 [Abstract/Free Full Text]
  62. Tsukamoto, T., Hata, S., Yokota, S., Miura, S., Fujiki, Y., Hijikata, M., Miyazawa, S., Hashimoto, T., and Osumi, T. (1994) J. Biol. Chem. 269, 6001-6010 [Abstract/Free Full Text]
  63. Miura, S., Miyazawa, S., Osumi, T., Hashimoto, T., and Fujiki, Y. (1994) J. Biochem. (Tokyo) 115, 1064-1068 [Abstract]
  64. Gould, S. J., Keller, G.-A., Hosken, N., Wilkinson, J., and Subramani, S. (1989) J. Cell Biol. 108, 1657-1664 [Abstract]
  65. Miura, S., Oda, T., Funai, T., Ito, M., Okada, Y., and Ichiyama, A. (1994) Eur. J. Biochem. 223, 141-146 [Abstract]
  66. Meijer, J., Starkerud, C., and Afzelius, B. A. (1993) Eur. J. Cell Biol. 60, 291-299 [Medline] [Order article via Infotrieve]
  67. Heinemann, P., and Just, W. W. (1992) FEBS Lett. 300, 179-182 [CrossRef][Medline] [Order article via Infotrieve]
  68. Pelham, H. R. B. (1990) Trends Biochem. Sci. 15, 483-486 [CrossRef][Medline] [Order article via Infotrieve]
  69. Palosaari, P. M., and Hiltunen, J. K. (1990) J. Biol. Chem. 265, 10750-10753

©1995 by The American Society for Biochemistry and Molecular Biology, Inc.