• cell cycle;
  • DsRed–SKL;
  • mitochondria;
  • oleic acid;
  • peroxisomal-targeting sequence;
  • Schizosaccharomyces pombe


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

Peroxisomes were visualized for the first time in living fission yeast cells. In small, newly divided cells, the number of peroxisomes was low but increased in parallel with the increase in cell length/volume that accompanies cell cycle progression. In cells grown in oleic acid, both the size and the number of peroxisomes increased. The peroxisomal inventory of cells lacking the dynamin-related proteins Dnm1 or Vps1 was similar to that in wild type. By contrast, cells of the double mutant dnm1Δ vps1Δ contained either no peroxisomes at all or a small number of morphologically aberrant organelles. Peroxisomes exhibited either local Brownian movement or longer-range linear displacements, which continued in the absence of either microtubules or actin filaments. On the contrary, directed peroxisome motility appeared to occur in association with mitochondria and may be an indirect function of intrinsic mitochondrial dynamics. We conclude that peroxisomes are present in fission yeast and that Dnm1 and Vps1 act redundantly in peroxisome biogenesis, which is under cell cycle control. Peroxisome movement is independent of the cytoskeleton but is coupled to mitochondrial dynamics.

Despite their distinct structure, protein import mechanisms and evolutionary origins (1,2), the activities of peroxisomes and mitochondria are inextricably intertwined. The most obvious expression of this is their co-operativity in a number of biochemical pathways (3). More recently, it has become clear that the biogenesis of peroxisomes and mitochondria also has elements in common (4). Peroxisomes have their origins at the endoplasmic reticulum (ER) (5) and undergo a stepwise maturation and division process, the details of which differ in fungi, plants and animals (6,7). A unifying theme is the involvement at a late stage of the maturation process of one or more dynamin-related proteins [DRPs; (8–10)]. DRPs are large guanosine triphosphatases (GTPases) that orchestrate membrane fission and fusion events associated with membrane traffic and organelle biogenesis (11). In the budding yeast Saccharomyces cerevisiae, arguably the organism in which the function of the dynamin superfamily has been most comprehensively studied, two DRPs, Dnm1 and Vps1, are involved in peroxisome biogenesis (12–14), and peroxisomes become greatly enlarged in cells lacking either protein (12–14). Homologues of these proteins exist in both mammalian cells [Dlp1; (15)] and plants [Drp3A; (16)]. Vps1 participates in a variety of other cellular activities, while Dnm1 has a single additional role in mitochondrial fission (17,18). Dnm1 is anchored to the mitochondrial surface by the outer membrane protein Fis1 (19–24), and Fis1 appears to play the same role at the peroxisome membrane (25,26). Peroxisomes not only vary in their maturation pathway but also in their association with the cytoskeleton. In budding yeast and plants, peroxisomes are actin associated (12,27,28), whereas in mammalian cells, they interact with microtubules (29,30). Nothing is known about the biogenesis or cytoskeletal association of peroxisomes in the fission yeast Schizosaccharomyces pombe, although, like mammalian cells, mitochondrial segregation in fission yeast is microtubule based (31). S. pombe possesses homologues of the budding yeast DRPs (32) plus most of the so-called pex genes involved in peroxisome biogenesis in other fungi (33). However, peroxisomes have never been visualized in fission yeast and, hence, nothing is known of their cellular role. Here, we show that peroxisomes do indeed exist in S. pombe. We further show that the number of peroxisomes per cell is under cell cycle control and that Dnm1 and Vps1 act redundantly in peroxisome biogenesis. Finally, we demonstrate that peroxisomes are intimately associated with, and move along, mitochondria.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

Identification of fission yeast peroxisomes

To identify peroxisomes in S. pombe, we took advantage of the ability of the tripeptide peroxisomal-targeting sequence SKL to import green fluorescent protein (GFP) and its derivatives into the peroxisome compartment (34). A construct in which SKL was fused to the C-terminus of DsRed was expressed in wild-type fission yeast cells under the control of an attenuated version of the thiamine repressible nmt1 promoter. Individual cells expressing DsRed–SKL revealed an average of 11 small spots (11.25 ± 0.45, n = 175), each having a diameter of approximately 0.5 μm (0.49 ± 0.02, n = 191) (Figure 1A,C). These appeared to be randomly distributed throughout the cytosol, with no obvious accumulation at sites of cell growth and division, the cell poles in interphase cells or the equatorial division plane during mitosis and cytokinesis. To confirm that these structures were indeed peroxisomes, we expressed DsRed–SKL in wild-type cells in the presence of oleic acid, a known inducer of peroxisome biogenesis (35). Within 4 h of oleate addition (roughly one cell cycle), an increase in peroxisome number and diameter was evident (data not shown). Cells maintained in oleate for 24 h showed a 43% increase in peroxisome number (15.93 μm ± 1.19, n = 135) and a 17% increase in peroxisome size (0.62 μm ± 0.02, n = 112) (Figure 1C,D). To eliminate potential variation in the appearance of peroxisomes in cells grown under different conditions on different days, we grew parallel cultures in the presence (EMM-OA) and absence (EMM-TE, see Methods) of oleic acid and mixed them prior to photography, the EMM-TE cells being first stained with Hoechst for later recognition. As can be seen in Figure 1B, oleic acid-grown cells displayed a larger diameter than the cells grown in glucose-based medium, and both the number and the fluorescence intensity of peroxisomes were substantially enhanced. This effect was not because of an increase in the level of expression of DsRed–SKL, as verified by Western blotting (data not shown). Cells expressing yellow fluorescent protein (YFP)-tagged (36) Pex5, a peroxisomal import receptor (37,38), and the peroxisomal-docking protein Pex14 (39) were examined by fluorescence microscopy. Both proteins were primarily cytoplasmic but were also localized to spots whose number, diameter and dynamics were comparable to the DsRed–SKL-labelled structures (Pex14, Figure 1E; Pex5, data not shown). Both the number and the diameter of these YFP-labelled spots increased in the presence of oleate. Although we failed to covisualize the Pex–YFP products and DsRed–SKL in the same cells, we are confident that, in both cases, the structures we observed are indeed peroxisomes (see further evidence below) and that, as in other organisms (34), DsRed–SKL is a good tool for studying peroxisomes in S. pombe.


Figure 1. Expression of DsRed–SKL identifies peroxisomes in Schizosaccharomyces pombe. A) Wild-type cells expressing DsRed–SKL were grown in minimal medium (EMM) and switched to either EMM or EMM containing oleic acid (EMM-OA) for 20 h. Cells grown in EMM-OA show an enlarged diameter and an increase in the number and intensity of peroxisomes. B) Parallel cultures in either EMM-TE or EMM-OA for 20 h were mixed and mounted together on the same slide. The EMM-TE-grown cells were stained with Hoechst for subsequent identification. C and D) The average number of peroxisomes per cell and the mean peroxisome diameter in EMM (white), EMM-TE (red) and EMM-OA (orange). Note that counts and measurements were made on single focal plane pictures, and values are probably underestimated. At least 112 cells were surveyed for each growth condition, and the results shown are a mean of three independent experiments. E) Wild-type cells expressing Pex14–YFP were grown in EMM + thiamine and switched to either EMM or EMM containing oleic acid (EMM-OA) with no thiamine for 20 h. Bars = 5 μm. DIC, differential interference contrast.

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Differential interference contrast images revealed the presence of small refractile bodies, which increased in size and number in the presence of oleate (Figure 1A,E). These structures were originally described as ‘oscillating lipid droplets’ in fission yeast (40) and were found in budding yeast to be partially associated with peroxisomes (41). Figure S1 shows that indeed some ‘lipid droplets’ may be associated with peroxisomes.

In the course of the above observations, we noticed that the number of peroxisomes per cell was heterogeneous, ranging from as few as four in newly divided cells to up to 25 in cells prior to division. To determine whether peroxisome number was dependent on cell length, i.e. was coupled to cell cycle progression, we counted peroxisomes over one cell cycle. As can be seen in Figure 2, peroxisome number increased in parallel to cell elongation. We observed frequent peroxisomal docking, fusion and fragmentation events, and it is probable that the organelles use these mechanisms to accommodate the increase in cell volume. No significant modification in the organization of peroxisomes was observed at mitosis, suggesting that there is a stochastic partitioning of the peroxisome pool to each of the two daughter cells.


Figure 2. The number of peroxisomes increases during the cell cycle. A) Wild-type cells expressing DsRed–SKL grown in EMM and observed for one division cycle. Bar = 5 μm. B) Average peroxisomes number per wild-type cell as a function of cell length. Cells were grown for 20 h in either EMM-TE or EMM-OA. DIC, differential interference contrast.

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Dynamin-dependent peroxisome biogenesis

In S. cerevisiae, the DRPs Vps1 and Dnm1 were shown to control peroxisome abundance (13). We therefore expressed DsRed–SKL in cells lacking either dnm1(32) or vps1 (this study) and in the double mutant dnm1Δ vps1Δ. vps1 was identified in S. pombe by homology to the VPS1 gene of S. cerevisiae(42) and was deleted by gene replacement. vps1Δ was viable, as was the double mutant dnm1Δ vps1Δ. Vps1 contains a putative Pex19-binding site (Figure S2), suggesting that it is involved in peroxisome biogenesis in fission yeast. Strains lacking each dynamin individually revealed only minor differences in the number and size of peroxisomes compared with the wild type (Figure 3). By contrast, the double deletion dnm1Δ vps1Δ strain contained either no peroxisomes at all (61.73% ± 7.09, n = 300) or aberrant organelles (Figure 3). It is not clear whether these structures were giant peroxisomes or aggregates of smaller ones, but the fact that their shape was irregular and changed over time suggests that the latter is most likely. Further analysis of these peroxisomes over one cell cycle revealed a failure to distribute properly in the mother cell, resulting in one daughter cell inheriting the entire peroxisome pool and the other none at all (Figure 4). Cells with no peroxisomes were able to divide. This was a surprising finding given that the dnm1Δ vps1Δ strain grows normally at all temperatures and shows no obvious morphology defects. We conclude that Dnm1 and Vps1 are dispensable individually for normal peroxisome biogenesis, but at least one DRP is essential. We next examined the effect of oleate on peroxisome size and number in each dynamin deletion strain. As can be seen in Figure 3B and C, both dnm1Δ and vps1Δ showed an increase in peroxisome number, indicating that these organelles remain competent to import enzymes for fatty acid oxidation. In the case of the double mutant, however, oleate had no effect on the morphology of peroxisomes, at least at the light microscope level.


Figure 3. Peroxisomes in dynamin deletion mutants. A) Wild-type (WT), vps1Δ, dnm1Δ and vps1Δ dnm1Δ cells expressing DsRed–SKL were grown in EMM and transferred to EMM-TE or EMM-OA for 20 h. Note the absence of peroxisomes in some cells in the double mutant and the enlargement of peroxisomes in others. Bar = 5 μm. B and C) Average number of peroxisomes per cell and mean peroxisome diameter in EMM-TE and EMM-OA. Pink: WT, white: vps1Δ, brown: dnm1Δ and black: vps1Δ dnm1Δ. A minimum of 85 cells were measured or counted for each growth condition, and the results shown are a mean of at least two independent experiments.

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Figure 4. Distribution of peroxisomes during the cell cycle of the vps1Δ dnm1Δ double mutant. Bar = 3 μm.

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Peroxisome motility

Peroxisomes exhibited very obvious motility, moving over a broad range of speeds from 0.01 to 0.29 μm/second (n = 30; DsRed–SKL peroxisomes, Movie S1; Pex14–YFP peroxisomes, Movie S2). Tracking individual peroxisomes over time revealed their motility to be remarkably complex. Nevertheless, we were able to identify three types of motile behaviour: a small proportion of organelles showed no movement at all (Figure 5A,B, orange), a larger number showed apparently Brownian movement (Figure 5A,B, purple) and another, extremely variable, population showed longer-range, linear displacements (Figure 5A,B, light blue). The number of peroxisomes showing each type of movement was impossible to quantify because percentages varied from cell to cell and even within individual cells, populations frequently interconverting from one form of motility to another. Figure 5C shows an individual peroxisome with two main phases of local motions [t = 5–11 seconds and t = 32–47 seconds (dashed boxes); speed = 0.17 μm/second] interrupted by saltatory displacements [e.g. t = 13–20 seconds (dotted box); speed = 0.29 μm/second]. Such behaviour suggests that there is no global regulation of peroxisome movement; rather, individual peroxisomes are free to interconvert between different motile states.


Figure 5. Peroxisome movement. A) Movement of individual peroxisomes in a wild-type cell observed by time lapse microscopy at intervals of 1 second for 1 min in a single focal plane. Every fifth image is shown; the outline of the cell is shown by the dashed line in panel ‘0’. Orange arrowhead: static peroxisome, light blue arrowhead: long-distance translocation and purple arrowhead: local movement. Note that some peroxisomes align along linear tracks. See also Movies S1 and S2. Bar = 3 μm. B) Patterns of movement of individual peroxisomes highlighted in (A). The colour of the track in (B) corresponds to the colour of the arrow in (A). Only peroxisomes that could be followed for at least 20 seconds are shown. C) Schematic representation of the distance covered by a single peroxisome from its first position in the track over time. The peroxisome is the one indicated by the purple arrowhead in (A) and (B). Dashed boxes illustrate phases of oscillatory movement. The dotted box shows an example of a long-range, linear displacement.

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A clue to the basis of linear movement was suggested by the occasional observation that peroxisomes formed files reminiscent of the organization of cytoplasmic microtubules (Figure 5A and Movies S1 and S2). To investigate this further, we expressed DsRed–SKL into atb2–GFP (43), a strain in which peroxisome excursions and microtubule dynamics could be imaged in the same cell. As shown in Figure 6A, some peroxisomes were found in close proximity to microtubule bundles (Figure 6A, open arrowhead), while others appeared to be associated with microtubule ends (Figure 6A, plain arrowhead). Peroxisomes made both anterograde and retrograde movements, but these were often non-linear, an individual peroxisome moving circumferentially around a microtubule bundle (Figure 6A, left panel, open arrowhead). It was clear that in some cases at least, peroxisomes aligned in the absence of microtubules (Movie S3). To investigate this point further, we examined the situation in cells entering mitosis where interphase microtubules are absent and the only microtubule structure is an intranuclear mitotic spindle. In such cells, no major change was observed in terms of either peroxisome distribution or movement [Figure 6A, dimethyl sulphoxide (DMSO), right panel, arrows]. To exclude the possibility that peroxisomes behaved differently during mitosis, we eliminated cytoplasmic microtubules in interphase cells by treatment with thiabendazole (TBZ). atb2–GFP cells incubated in 100 μg/mL TBZ for up to 2 h showed no modification in peroxisome distribution, motion (Figure 6, TBZ and Movie S4) or mean velocity (DMSO = 0.140 μm/second, n = 38; TBZ = 0.144 μm/second, n = 38). Hence, despite their close association, microtubules and peroxisomes might not interact directly, but, rather, peroxisomes are associated with a cellular component that follows the microtubule network. An obvious candidate was mitochondria (31,44).


Figure 6. Peroxisomes align with microtubules but move in their absence. atb2–GFP cells expressing DsRed–SKL were grown in EMM and transferred to EMM containing either 0.2% DMSO, 100 μg/mL TBZ or 1 mm LAT-A in 0.2% DMSO. A) The GFP and DsRed signals were captured alternatively every 1 second for 1 min in a single focal plane. Every sixth image is shown. DMSO: interphase cell (left panel) and early mitotic cell (right panel). Open arrowhead: peroxisome moving alongside and around a microtubule bundle and filled arrowheads: peroxisomes at the end of a microtubule bundle. See also Movie S3. Interphase cell treated with TBZ or LAT-A: peroxisome movement continues in the absence of cytoplasmic microtubules or the actin cytoskeleton (arrows). Bottom panels confirm that microtubule and actin depolymerization were complete. See also Movie S4. B) Patterns of movement of individual peroxisomes in the presence of TBZ. The colour of the track corresponds to the colour of the arrow in (A). Orange arrowhead: static peroxisome and purple arrowhead: long-distance translocation. Bars = 3 μm.

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As can be seen in Figure 7A, peroxisomes did indeed colocalize with mitochondria (Figure 7A and Movie S5) and appeared to move along them (Figure 7A and Movie S6). Such movements were not always linear but were also rotational, tracking around the mitochondrial circumference (Figure 7A, DMSO, arrowhead). Pex5 and Pex14–YFP-labelled peroxisomes displayed a similar localization and movement along mitochondria (Pex14, Figure 7B; Pex5, data not shown). Following microtubule depolymerization with TBZ, mitochondria fragment (45). In many cases, mitochondrial fragments aggregated and peroxisomes showed a corresponding accumulation (Figure 7A, TBZ, left panel). Non-aggregated fragments also showed associations with peroxisomes (Figure 7A, TBZ, right panel), which seemed to be able to move from fragment to fragment (Figure 7C). Confocal images revealed that mitochondrial fragments were in fact interconnected and that consequently, peroxisomes were able to move between these structures (Figure 7D and Movie S7).


Figure 7. Peroxisomes interact with mitochondria. A) Wild-type cells expressing DsRed–SKL were grown in EMM and stained with Mitotracker Green. The green and red signals were captured alternatively every 1 second for 1 min in a single focal plane. In control (DMSO) cells, peroxisomes colocalize with mitochondria and translocate along them (arrowhead). See also Movies S5 and S6. In the presence of 100 μg/mL TBZ, mitochondria fragment (right panel) and eventually further aggregate (left panel). Filled arrowheads: local motion of peroxisomes associated with mitochondrial fragments and open arrowheads: moving peroxisomes along larger mitochondrial fragments (on the left panel, note the circular pattern around the aggregated mitochondria). B) The peroxisomal-docking protein Pex14 (green) is found in close proximity to mitochondria (red). C) Cells treated with TBZ as in (A) show peroxisomes (red) apparently moving by bridging two mitochondrial fragments (green). See also Movie S7. D) Confocal images of cells treated as in (A) and (C). Depending on the z-section observed, mitochondria appear either conventionally fragmented (e.g. z = 1.2 μm) or connected (e.g. z = 3 μm). Peroxisomes are associated with branched mitochondrial fragments. E) Wild type (WT) and dynamin mutants expressing DsRed–SKL stained with Mitotracker Green and visualized by confocal microscopy. Whole cell projections are shown. Mitochondria are fused in dnm1Δ, to a lesser extent also fused and branched in vps1Δ and coiled in vps1Δ dnm1Δ. In all three cases, peroxisomes are associated with mitochondria. See also Movies S8, S9 and S10. Bars = 3 μm.

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Mitochondria in dnm1Δ were shown previously to form nets as a result of their inability to undergo fission [(32); Figure 7E, dnm1Δ and Movie S8]. In vps1Δ, mitochondria were often branched and formed small, unconnected nets (Figure 7E, vps1Δ and Movie S9). The vps1Δ dnm1Δ double mutant was different from each of the single mutants, mitochondria appearing twisted and forming nodes made of tight loops (Figure 7E, vps1Δ dnm1Δ and Movie S10). In each of these three aberrant mitochondrial phenotypes, peroxisomes were unambiguously associated with mitochondria (Figure 7E and Movies S8, S9 and S10). The motility of peroxisomes in vps1Δ dnm1Δ was dependent upon their size, the larger and most abundant ones being static, but nevertheless these also coaggregated with mitochondria. Although showing no overt motility, the enlarged peroxisomes in vps1Δ dnm1Δ were dynamic, changing shape perhaps as the result of vesicle docking or frustrated fission. Because mitochondria in this mutant segregate properly (data not shown), the mis-segregation of peroxisomes is probably a consequence of their inability to distribute along the surface of mitochondria.

In budding yeast, peroxisomes are associated with the actin cytoskeleton (12,46). To examine this possibility in fission yeast, we expressed DsRed–SKL in crn1–GFP (47), a strain in which actin patches can be visualized. We noticed no obvious colocalization between the two structures (data not shown). Further, peroxisome distribution and motility were unaffected by the presence of 1 mm latrunculin A (LAT-A), which depolymerizes all actin structures in fission yeast (48) (Figure 6, LAT-A). We also monitored peroxisome motility in myo52Δ(49), lacking one of two fission yeast type V myosins, and for3Δ, in which actin cables are absent (50), and found no difference to the wild type (data not shown). We conclude that peroxisomes are dynamic organelles whose distribution and displacement are independent of the cytoskeleton but are mitochondria dependent.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

Peroxisomes were the last eukaryotic organelle to be discovered (51) and, appropriately, are the final membrane-bound compartment to be described in fission yeast. Although extensively studied in budding yeasts and filamentous fungi (52), peroxisomes have not previously been described in a non-budding yeast such as S. pombe. Few fission yeast peroxisomal proteins contain the canonical SKL type I peroxisomal-targeting signal per se (our unpublished data), but this sequence fused to the C-terminus of a fluorescent marker such as GFP has proved to be imported into peroxisomes in a wide range of organisms (34). Unlike S. cerevisiae, which contains few to no peroxisomes in glucose-based medium (53), similarly grown fission yeasts contain an average of 11 peroxisomes per cell. Peroxisome number was related to cell volume, being low in newly divided (small) cells and roughly doubling as cells increased in length prior to cell division. No specific mechanism for peroxisome segregation was observed in wild-type mitotic cells, and we conclude that, as in mammalian cells (30), peroxisomes are inherited through a simple partition process [the ‘stochastic’ mechanism of Warren and Wickner (54)]. The unequal inheritance of large peroxisomes in the dnm1Δ vps1Δ mutant was a consequence of their asymmetric distribution and low motility, although we cannot exclude the possibility that undetected peroxisome membranes are still present or that the absence of peroxisomes in some cells is because of their removal by pexophagy (55,56). Whether peroxisomes arise de novo from the ER or by division of pre-existing organelles is a matter of debate. A recent study in budding yeast showed that peroxisomes can form de novo, but only in cells temporarily having no peroxisomes (57). In wild-type S. cerevisiae, peroxisomes multiply by division (57). The fact that we observed regular fusion and fission events suggests that the latter is also possible in S. pombe. In dnm1Δ vps1Δ cells with no peroxisomes, we saw no reacquisition of these organelles in the two following cell cycles. In a growing population of this mutant, the percentage of cells containing no peroxisomes increased by approximately 8% each successive generation (data not shown), and this net loss is consistent with an absence of peroxisome biogenesis in those cells. However, the situation might be different in wild-type cells where both DRPs, Vps1 and Dnm1, may also have a role in the remodelling of the ER (58). In S. cerevisiae, however, the formation of peroxisomes from the ER appears to be independent of both Dnm1 and Vps1 (57). Whether de novo peroxisome formation does not occur in S. pombe or whether dnm1Δ vps1Δ fission yeast cells are defective in peroxisome biogenesis, it is clear that the two yeasts do not deal with peroxisome production in the same way.

The presence of peroxisomes in glucose-grown fission yeast suggests that these organelles play a role in cell metabolism, albeit a non-essential one as dnm1Δ vps1Δ cells containing no peroxisomes continue to divide. Fission yeast cells grew in the presence of fatty acid, although they showed an enlarged cell diameter. Peroxisome number increased by about 0.5-fold in medium supplemented with oleate, and peroxisome diameter also increased. The level of peroxisome proliferation was considerably less than that seen in oleate-grown S. cerevisiae (∼2.5-fold), but so little is known about fatty acid metabolism in fission yeast that it is premature to speculate as to the significance of these differences.

Peroxisomes exhibited complex motile behaviour. A small subpopulation showed no overt motility, whereas the majority showed local Brownian movement, interspersed with occasional longer-range, saltatory movements. A formal analysis of peroxisome motility was complicated by variability from cell to cell and even within an individual cell over longer periods of time. Mammalian cells also contain populations of peroxisomes with distinct motile properties, although a function for such movements remains elusive (59). Most, as in fission yeast, show Brownian movement, but a minor population of peroxisomes exhibit long-range linear movements that are sensitive to microtubule depolymerization (29,30,60). In budding yeast, peroxisomes are carried from mother cell to bud along actin cables by the type V myosin Myo2p (12,46). Peroxisome motility in plants is also actin based (27,28). In fission yeast, by contrast, we observe motility in the absence of both microtubules and actin. Peroxisome movements continued in mitotic cells that lack cytoplasmic microtubules and in interphase cells following depolymerization with TBZ. Such movements were also insensitive to the actin-depolymerizing drug LAT A and were unaffected by the absence of actin cables in the mutant for3Δ(50) and in cells lacking the S. pombe homologue of Myo2p, Myo52 (49). Further, fission yeast has no homologues of the Inp1 and Inp2 proteins that mediate the interaction of peroxisomes to the actomyosin cytoskeleton in S. cerevisiae(46). Although we cannot definitively exclude the possibility that peroxisomes interact with the dynamic ends of cytoplasmic microtubules (61), the rate of microtubule growth and shrinkage was at least an order of magnitude slower than the fastest rate of peroxisome movement (62). Based on a careful analysis of the disposition of microtubules and organelles in fission yeast by electron tomography, Höög et al. (44) found no evidence to suggest that vesicles of the size of peroxisomes were transported along microtubules, and our results are consistent with this view. Rather, peroxisome movements appeared to occur in close association with the surface of mitochondria. Mitochondria themselves align closely with microtubules (44) and are disrupted when microtubules are depolymerized (45). However, the peroxisome–mitochondria interaction survives this modification. The functional significance of this observation is unknown, although mitochondria and peroxisomes have an intimate metabolic relationship (63). Whether this interaction is direct or not, and what drives the movement of peroxisomes at the surface of mitochondria, is unclear. Mitochondria are themselves dynamic, undergoing repeated dynamin-driven fusion and fission events (64,65). Mitochondrial motility is unaffected by the depolymerization of the cytoskeleton (our unpublished data), and it is possible that the movements of peroxisomes we observe are a consequence of this. It is unlikely that the peroxisomal movements we observe are associated with the ER. In S. pombe, relatively little is known about the ER, except that it extends from the nuclear envelope to the plasma membrane (66), distinct from the location of microtubules. Moreover, no interaction between the ER and microtubules was found in the most careful study of fission yeast ultrastructure to date (44). Static peroxisomes however are often observed close to the plasma membrane. Further studies may determine whether they are nascent organelles arising from the ER.

Because peroxisome biogenesis and mitochondrial fission have elements in common (6,13), we investigated the contribution of the DRPs Dnm1 or Vps1 to peroxisome formation in fission yeast. Schizosaccharomyces pombe cells lacking Dnm1 contain fused mitochondria (32). vps1Δ cells also revealed a fused mitochondria phenotype although less severe. Single dnm1Δ or vps1Δ mutant cells showed no major change in peroxisome morphology or motility. In the double mutant, however, many cells contained no peroxisomes but, in those that did, these were hugely enlarged. In budding yeast, peroxisomes in dnm1Δ vps1Δ cells are a single, fused structure that undergoes amoeboid-like movements (13). In fission yeast, the large peroxisomes appear to be aggregates of numerous smaller structures that dock but are unable to fuse. Hence, the DRPs in S. pombe may play a role in both peroxisome fusion and fission. Vps1 is known to play such a dual role in vacuole biogenesis in budding yeast (67). Strikingly, those peroxisomes, still associated with the surface of mitochondria, were non-motile, reinforcing our view that there is an intimate relationship between Dnm1, Vps1, mitochondrial fission and fusion and peroxisome biogenesis and motility. That peroxisomes formed in the absence of either DRP individually are functional, at least in terms of importing new protein, is suggested by the fact that in both dnm1Δ and vps1Δ strains, peroxisome size and number increased in response to oleate. Hence, Dnm1 and Vps1 appear to serve redundant functions, each being able to support peroxisome biogenesis in the absence of the other. Attempts to localize either Dnm1 or Vps1 to peroxisomes have met with limited success (12,13). In fission yeast, Vps1–GFP localizes to spots of the same size and with the same motile properties as peroxisomes, but we have also been unable to confirm that these are the same structures (data not shown). In budding yeast, Vps1 interacts with the peroxisomal membrane through the peroxin Pex19 (14,68). The fission yeast genome contains a Pex19 homologue, and Vps1 contains a recognizable Pex19-binding motif (69) that includes the conserved valine at position 491 (V516 in S. cerevisiae) that is essential for Pex19 interaction (14). Hence, the interaction of DRPs with peroxisomes in fission yeast would appear to be conventional.

The relationship between DRPs and peroxisome biogenesis in fission yeast is quite different to that in S. cerevisiae where both vps1Δ and dnm1Δ contain one or two giant peroxisomes in oleate-grown, but not glucose-grown, cells (12,13). That peroxisomes are associated with the actin cytoskeleton in budding yeast (12,46) and not in S. pombe also encourages us to believe that peroxisome biology is different in the two yeasts. This first paper on peroxisomes in fission yeast poses as many questions as it answers, but it encourages us to believe that S. pombe will prove to be a useful model system in which to explore many aspects of peroxisome biology.

Materials and Methods

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

Plasmid constructs

The DNA coding for the DsRed tag fused to the N-terminus of the SKL tripeptide was amplified by polymerase chain reaction (PCR) from plasmid pRS31DsRed-SKL kindly provided by Dr J. Aitchison. The 5′-TATATGGATCCGGTATGAGGTCTTCCAAGAATG and the 3′-ATATACCATGGCTAAAgCTTTgAAAGGAACAGA (Sigma-Proligo) primers contained BamHI and NcoI restriction sites, respectively. The pREP41-CGFP plasmid was opened by digestion with BamHI and NcoI, which removed the gfp cassette and allowed the insertion of the digested PCR product. The final vector was named pREP41DsRed-SKL. The construct was verified by sequencing (Allan Wilson Centre, Palmerston North, New Zealand). The pDUAL-YFH1-Pex5 and pDUAL-YFH1-Pex14 plasmids were obtained from the RIKEN Institute, Japan (36).

Yeast strains and cultures

The S. pombe strains used in this study are listed in Table S1. Media, growth, genetics and maintenance of strains were as described previously (70). The fission yeast gene coding for Vps1 (SPAC767.01c) was identified on the basis of its homology with S. cerevisiae Vps1 (62.9% amino acid identity; Figure S2). The GTPase, middle and GTP effector domains that are features of dynamins were conserved, and two putative Pex19-binding domains, characteristic of Vps1 proteins, were found (14,69). The vps1::ura4 strain was constructed by homologous recombination. The high-performance liquid chromatography-purified primers 5′-ATGGATCCCTCATTGATTAAAGTTGTCAATCAACTTCAGGAGGCTTTCTCCACAGTTGGCGTTCAAAACTTGATTGACTTCTATATGTATGCATTTGTGT and 3′-TTAAACGTTAGACACAATCTCACTTGCCTGTAGGAGAGATTCTACCATTTGTTCGCATTCTTTACGCGTTGCACTGTCAACCAATGTTTATAACCAAGT were used to amplify the ura4 gene from the pREP41 plasmid. The deletion was verified by PCR. pREP41DsRed-SKL was expressed in the continuous absence of thiamine. Cells expressing pDUAL-YFH1-Pex5 and pDUAL-YFH1-Pex14 plasmids were pre-cultured in the presence of 60 μm thiamine, washed and grown for approximately 20 h in the absence of thiamine.

A 10× stock solution of oleic acid [2% oleic acid (Sigma-Aldrich), 0.2% Tween-40 and 10% ethanol] was made fresh and diluted to 1× in EMM plus the appropriate supplements (EMM-OA). EMM containing 0.02% Tween-40 and 1% ethanol was used as a control (EMM-TE). Cells were pre-cultured in 5 mL EMM, diluted into 50 mL EMM-OA or EMM-TE and grown at 25°C for 20–24 h to a density of 2–4 × 106 cells/mL.

Western blot

Total protein extracts were isolated as previously described (71). Concentrations were determined by Bradford assay, and 10 μg of each sample was separated on an 11% sodium dodecyl sulphate–polyacrylamide gel. The membrane was incubated with anti-DsRed (Clontech; 1:1000) and anti-tubulin TAT-1 primary antibodies (provided by K. Gull; 1:750) and then with anti-rabbit and anti-mouse horseradish peroxidase-conjugated secondary antibodies (Promega; 1:2500). Signals were detected using the Super Signal kit (Pierce).


Live cell imaging was performed in an imaging chamber (CoverWell, 20-mm diameter, 0.5-mm deep) (Molecular Probes) filled with 800 μL of 2% agarose in EMM and sealed with a 22 × 22-mm glass coverslip. Mitotracker (Molecular Probes) was dissolved into DMSO at a concentration of 1 mm and diluted in EMM to 1 μm. Nine hundred microlitres of mid-log phase cells expressing DsRed–SKL was incubated with 100 μL MitoTracker (final concentration 100 nm) for 30 min. The cells were washed three times in EMM before being applied to an imaging chamber. For microtubule depolymerization experiments, the chamber contained 100 μg/mL TBZ in 0.2% (final concentration) DMSO or DMSO alone. For LAT-A experiments, 1 μL of a 1 mm LAT-A stock solution or the equivalent concentration of DMSO was pipetted on the top of cells pre-applied to a chamber containing no drug. Cells were left to equilibrate for between 15 min and 2 h in the presence of the drugs before observation. Hoechst-stained cells were prepared as described previously (72). For peroxisome movement on mitochondria, a picture of Mitotracker fluorescence was taken 1 second after each DsRed–SKL image and the two stacks were aligned to make a montage. In the experiment aimed at colocalizing peroxisomes and lipid droplets, cells were fixed in 3.3% paraformaldehyde. The DsRed signal survived this procedure. Schizosaccharomyces pombe cells were observed at room temperature using an Olympus IX71 microscope with a ×100 oil immersion lens. Pictures were captured with a Hamamatsu ORCA-ER C4742-80 digital charge-coupled device camera (Hamamatsu Corporation). For confocal microscopy, a Leica TCS SP5 microscope and a ×63 oil immersion lens NA = 1.4, were used. Counts and measurements were made using metamorph software (Molecular Devices Corporation) and downloaded to Microsoft Excel for analysis.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

We thank Suresh Subramani and Chris Danpure for their help and advice, John Aitchison, Pascale Belenguer, Fred Chang and Yasushi Hiraoka for strains and plasmids and Keith Gull for the TAT-1 antibody. This study was supported by a grant from the Palmerston North Medical Research Foundation.


  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information
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Supporting Information

  1. Top of page
  2. Abstract
  3. Results
  4. Discussion
  5. Materials and Methods
  6. Acknowledgments
  7. References
  8. Supporting Information

Table S1: List of strains used in this study

Figure S1:Partial association of peroxisomes with lipid droplets. Wild-type cells expressing DsRed–SKL were grown in EMM-OA for 24 h and fixed. Lipid droplets were observed by DIC and DsRed–SKL peroxisomes by fluorescence. Framed: lipid droplet associated with a peroxisome; open arrowhead: free lipid droplet and filled arrowhead: free peroxisome.

Figure S2:Sequence alignment of Saccharomyces cerevisiae Vps1 (Sc) and predicted Schizosaccharomyces pombe Vps1 (Sp). The two proteins share 62.9% identity. Red bar: GTPase domain, blue bar: GTP effector domain (GED), yellow boxes: Pex19-binding sites and star: conserved valine essential for interaction with Pex19.

Movie S1: Movement of DsRed–SKL peroxisomes in a wild-type cell. Note on the top left corner of the cell, a static peroxisome and along the left side, a peroxisome moving down rapidly. Other peroxisomes display mostly local motions. The time is indicated in min:seconds. Bar = 3 μm.

Movie S2: Movement of Pex14–YFP peroxisomes in a wild-type cell. Arrowheads point at two peroxisomes displaying long-range linear movements. The time is indicated in min:seconds. Bar = 3 μm.

Movie S3: Confocal images reconstituted in three dimensions of peroxisomes in three atb2–GFP cells. Peroxisomes are close to microtubules but not bound to them. Bar = 3 μm.

Movie S4: Movement of peroxisomes in an atb2–GFP cell treated with TBZ. The time is indicated in min:seconds. Two arrows at times 5and 19 seconds, respectively, show peroxisome displacement over long, linear distances. The last image confirms that microtubule depolymerization was complete. Bar = 3 μm.

Movie S5: Confocal images reconstituted in three dimensions of peroxisomes in a wild-type cell stained with Mitotracker Green. Most peroxisomes bind to mitochondria. Note that the free ends of some mitochondria are capped with a peroxisome. Bar = 3 μm.

Movie S6: Peroxisomes moving along or around a Mitotracker Green-labelled mitochondria in a wild-type cell. The green and red pictures were taken every 1 second alternatively, and the stacks were aligned. The time is indicated in min:seconds. The arrow at time 28 seconds points at a peroxisome about to display a long-range movement on the mitochondria. Bar = 3 μm.

Movie S7: Peroxisomes in a wild-type cell stained with Mitotracker Green and treated with TBZ. Arrow at time 0 seconds shows a peroxisome displacement along a long mitochondrial fragment. Arrowhead at time 1 second shows a peroxisome apparently bridging two mitochondrial fragments. The time is indicated in min:seconds. Bar = 3 μm.

Movie S8: Peroxisomes in a dnm1Δ cells stained with Mitotracker Green. Confocal images were reconstituted in three dimensions. Mitochondria are fused (top of the cell), and peroxisomes are equally associated with mitochondrial nets and arms. Bar = 3 μm.

Movie S9: Confocal images reconstituted in three dimensions of peroxisomes in a vps1Δ cell stained with Mitotracker Green. This mitochondria organizes as three distinct nets, which also branch. Peroxisomes colocalize with mitochondria regardless of their architecture. Bar = 3 μm.

Movie S10: Confocal images reconstituted in three dimensions of peroxisomes in a vps1Δ dnm1Δ cell stained with Mitotracker Green. This cell shows a long and twisted mitochondria terminated by node-like structures. The few and large peroxisomes characteristic of this double mutant colocalize with mitochondria and tend to coaggregate with the nodes. Bar = 3 μm.

Supplemental materials are available as part of the online article at

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TRA_685_sm_FigureS1.pdf253KSupporting info item
TRA_685_sm_FigureS2.pdf229KSupporting info item
TRA_685_sm_MovieS1.mov73KSupporting info item
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TRA_685_sm_MovieS7.mov116KSupporting info item
TRA_685_sm_MovieS8.mov120KSupporting info item
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