EUKARYOTIC CELL, Aug. 2008, p. 1373–1386 1535-9778/08/$08.00⫹0 doi:10.1128/EC.00085-08 Copyright © 2008, American Society for Microbiology. All Rights Reserved.
Vol. 7, No. 8
Septins Stabilize Mitochondria in Tetrahymena thermophila䌤† D. Włoga,1 I. Strzyz˙ewska-Jo ´wko,2 J. Gaertig,1* and M. Jerka-Dziadosz2* Department of Cellular Biology, University of Georgia, Athens, Georgia 30602-2607,1 and Polish Academy of Sciences, M. Nencki Institute of Experimental Biology, Department of Cell Biology, 3 Pasteur Street, 02-093 Warsaw, Poland2 Received 8 March 2008/Accepted 18 June 2008
We describe phylogenetic and functional studies of three septins in the free-living ciliate Tetrahymena thermophila. Both deletion and overproduction of septins led to vacuolization of mitochondria, destabilization of the nuclear envelope, and increased autophagy. All three green fluorescent protein-tagged septins localized to mitochondria. Specific septins localized to the outer mitochondrial membrane, to septa formed during mitochondrial scission, or to the mitochondrion-associated endoplasmic reticulum. The only other septins known to localize to mitochondria are human ARTS and murine M-septin, both alternatively spliced forms of Sep4 (S. Larisch, Cell Cycle 3:1021–1023, 2004; S. Takahashi, R. Inatome, H. Yamamura, and S. Yanagi, Genes Cells 8:81–93, 2003). It therefore appears that septins have been recruited to mitochondrial functions independently in at least two eukaryotic lineages and in both cases are involved in apoptotic events. Septins are conserved GTP-binding and filament-forming proteins (51). Septins were discovered in Saccharomyces cerevisiae and later found to be ubiquitous in metazoans (39, 53). The cellular functions of septins are diverse and include participation in cytokinesis (22, 40), establishment of diffusion barriers for proteins and mRNAs during cell division (16, 75), vesicle trafficking, exocytosis (7, 28, 33, 37), and apoptosis (29, 49). Accordingly, cellular localizations of septins are diverse and include the bud neck (14), division furrow (1), presynaptic vesicles (86), dendritic spines (73, 85), and mitochondria (50, 74). Septins interact with microtubules (45, 71) and microfilaments (46, 56) as well as the endoplasmic reticulum (ER) and filaments emanating from the Golgi network (54, 71). Recently, the diverse patterns of septins were classified into types: projections, partitions, and dispersal over whole cells (52). The molecular functions of septins remain unclear (reviewed in reference 81). So far, septins have been studied only for fungi and metazoans. Since septins are known to be involved in cytoskeletal organization and membrane remodeling events, we anticipated that Tetrahymena thermophila septins are involved in partitions of distinct domains in the structurally elaborate cell cortex of ciliates, in particular during cytokinesis, when the cortex undergoes longitudinal segmentation. Surprisingly, we have found that septins of Tetrahymena thermophila localize to mitochondria and regulate mitochondrial dynamics.
sequences from NCBI (National Center for Biotechnology Information, http: //www.ncbi.nlm.nih.gov/); Drosophila melanogaster sequences from FlyBase (http: //flybase.bio.indiana.edu/); Caenorhabditis elegans sequences from the Sanger Institute (http://www.sanger.ac.uk/Projects/C_elegans/); and Chlamydomonas reinhardtii sequences from JGI (http://genome.jgi-psf.org/Chlre3/Chlre3.home .html). To search for septin sequences in the genomes of protists, we used S. cerevisiae and human septin sequences to perform tblastn or blastp searches for Giardia lamblia (http://www.giardiadb.org/giardiadb/), Toxoplasma gondii (http: //www.toxodb.org/toxo/home.jsp), Plasmodium falciparum (http://tigrblast.tigr.org /er-blast/index.cgi?project⫽pfa1), Trypanosoma brucei (http://www.sanger.ac.uk /cgi-bin/blast/submitblast/t_brucei/), Tetrahymena thermophila (http://seq.ciliate .org/cgi-bin/blast-tgd.pl), and Paramecium tetraurelia (http://paramecium.cgm .cnrs-gif.fr/tool/blast). The domain analysis of predicted septins was performed using SMART (http://smart.embl-heidelberg.de/). Coiled-coil domains were predicted using COILS (http://www.ch.embnet.org/software/COILS_form.html) and Coiled-Coil Prediction (http://www.russell.embl-heidelberg.de/cgi-bin/coils-svr .pl) (55). The transmembrane domains were predicted using THMMH (http: //www.cbs.dtu.dk/services/TMHMM/) and DAS (http://www.sbc.su.se/⬃miklos /DAS/). The molecular weight was calculated at http://ca.expasy.org/tools/pi_tool .html. The mitochondrial targeting signals were detected using PREDOTAR (http://urgi.versailles.inra.fr/predotar.html) and MITOPROT (http://ihg2 .helmholtz-muenchen.de/ihg/mitoprot.html). For phylogenetic analyses, septin sequences were aligned using ClustalX 1.82 and manually corrected using the SeaView program (26). The alignment of the central core of the septin proteins was used to calculate a neighbor-joining phylogenetic tree with the PHYLIP package (23), as described before (83). Cell culture. Cells were grown axenically as described previously (15). Strains were maintained in 7-ml aliquots of 1% proteose-peptone, 0.1% yeast extract. For most experiments, cells were grown in PPYGFe medium (2% proteosepeptone, 0.2% yeast extract, 0.5% glucose, 9 ⫻ 10⫺5 M iron chelate) (61) in 50-ml volumes in 250-ml Erlenmeyer flasks at 29°C with reciprocal shaking. Growth curves were calculated as described previously (84). Studies of expression of septin genes. Total RNA and cDNA were prepared from growing, conjugating, and cilium-regenerating cells as described previously (35). The following primers were used for reverse transcription PCR: forward primers Sep1-F (GAGAAAAAGGCAGTGAGAAAAGAAAATGGAATAAC TTTG), Sep2-F (GTAACTGGCATTCGTACTGA), and Sep3-F (GATTGAC ACTCTTGGTTATGG) and reverse primers Sep1-R (TTGGTAAACATCAG CCAATTTAATGAGTTTTGTGCT), Sep2-R (GTTAGTTTCTTAAGATCAG CG), and Sep3-R (CTTCAACAGATTTATACAAAT). Knockouts of septin genes. To prepare targeting plasmids, with the neo3 cassette under the control of the 0.9-kb MTT1 promoter, we amplified septin coding regions with 5⬘ and 3⬘ untranslated regions (UTR) from the genomic DNA. A 6.5-kb fragment of SEP1 was amplified with primers 5⬘-GGCAGTCT TTAGATAACTTTGCAT-3⬘ and 5⬘-CTTGAAATGAGTGGATAACACAAC3⬘. A 5.8-kb fragment of SEP2 was amplified with primers 5⬘-ACAATCAAAG AGTTCCATTTCC-3⬘ and 5⬘-AGAACGACTATAACTTAGCTTCAG-3⬘. A 4.9-kb fragment of SEP3 was amplified with primers 5⬘-GATTAAATGATACT
MATERIALS AND METHODS Bioinformatics. Protein sequences of the known septins were obtained from the following databases: Saccharomyces cerevisiae, Schizosaccharomyces pombe, Nannochloris bacillaris, Strongylocentrotus purpuratus, Danio rerio, and human
* Corresponding author. Mailing address for J. Gaertig: Department of Cellular Biology, University of Georgia, Athens, GA 306022607. Phone: (706) 542-3409. Fax: 48 22 822 53 42. E-mail: jgaertig @cb.uga.edu. Mailing address for M. Jerka-Dziadosz: Polish Academy of Sciences, M. Nencki Institute of Experimental Biology, Department of Cell Biology, 3 Pasteur Street, 02-093 Warsaw, Poland. Phone: 48 22 5892 233. Fax: (706) 542-4271. E-mail:
[email protected]. † Supplemental material for this article may be found at http://ec .asm.org/. 䌤 Published ahead of print on 27 June 2008. 1373
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FIG. 1. (A) Neighbor-joining phylogenetic tree based on conserved central domains of septins. Septins indicated in red are known to be associated with mitochondria. The following sequences were used: Tetrahymena thermophila (Tt) Sep1p TTHERM_00994080, Sep2p TTHERM_00316560, and Sep3p TTHERM_00989350; Paramecium tetraurelia (Pt) Sep1 GSPATP00027315001, Sep2 GSPATP00020551001, Sep3 GSPATP00037648001, and Sep4 GSPATP00017403001; Chlamydomonas reinhardtii (Cr) 148151; Nannochloris bacillaris (Nb) BAD42341.1; Saccharomyces cerevisiae (Sc) Cdc3 NP_013418.1, Cdc12 NP_011975.1, Cdc10 NP_009928.1, Cdc11 NP_012610.1, Spr28 NP_010504.1, Shs1 NP_010056.1, and Spr3 NP_011573.1; Schizosaccharomyces pombe (Sp) Spn1 NP_594754.1, Spn2 NP_593159.1, Spn3 P48008, Spn4 NP_593566.1, Spn5 NP_594040.1, Spn6 NP_588214.1, and Spn7 O60165; Caenorhabditis elegans (Ce) Unc59 NP_493388.1 and Unc61 NP_872156.1; Drosophila melanogaster (Dm) Pnut P40797, Sep1 NP_523430.1 (CG1403-PA), Sep2 NP_524417.1 (CG4173-PA), Sep4 NP_728003.1 (CG9699-PA), and Sep5 NP_651961.1 (CG2916PB); Strongylocentrotus purpuratus (Sp) Sep2 XP_788114.2, Sep3 XP_795119.2, Sep4 XP_001180939.1, Sep6 XP_790953.2, and Sep7 XP_001175696.1; Danio rerio (Dr) Sep2 AAH67625.1, Sep3 NP_001019589.1, Sep4a XP_001343014.1, Sep4b NP_001032456.1, Sep4c NP_001076284.1, Sep5 NP_956282.1, Sep6 NP_997791.1, Sep7 XP_001340758.1, Sep8 AAH55257.1, Sep9 NP_944593.1, Sep10 NP_001017557.1, and Sep12 XP_693022.2; and Homo sapiens (Hs) Sep1 NP_443070.1, Sep2 NP_004395.1, Sep3 NP_663786.1, Sep4 NP_004565.1, Sep5 NP_002679.2, Sep6 NP_665798.1, Sep7 NP_001779.3, Sep8 XP_034872.5, Sep9 AAF23374.1, Sep10 AAH20502.1, Sep11 NP_060713.1, Sep12 AAH35619.1, and Sep13 XP_001133108.1. The numbers at the branches represent bootstrap support values above 50%. (B) Domain analysis of septins. The following types of domains and motifs are marked: variable N-terminal domain (dark blue) followed by polybasic and hydrophobic residues implicated in membrane interactions (red), GTPase domain (light blue) followed by septin unique sequence (marine blue), transmembrane domain (green), coiled-coil domain (violet), and variable C-terminal domain (light green). aa, amino acids.
ATCAAGA-3⬘ and 5⬘-TCAGTTGAATCGTCTAGGTCA-3⬘. The fragments were ligated into pGEM-T Easy (Promega). The SEP1-T Easy plasmid was digested with SalI, and blunt ends were made with Klenow DNA polymerase and self-ligated to remove a HincII site of T Easy. The neo3 cassette was inserted in a reverse orientation between the HincII and NsiI sites. The SEP2-T Easy plasmid was digested with EcoRV and SnaBI, dephosphorylated, and ligated to a blunt-ended neo3 cassette inserted in a reverse transcriptional orientation. The SEP3-T Easy plasmid was digested with BglII, and blunt ends were made with Klenow DNA polymerase, digested with SpeI, and ligated with the neo3 cassette (digested with ClaI, polished with Klenow DNA polymerase, and digested with XbaI). The neo3 cassette was inserted in the reverse orientation. Before biolistic transformation, the targeting plasmids were digested with the following enzymes to separate the targeting fragment from the rest of the plasmid: SEP1-neo3 and SEP2-neo3 with EcoRI and SEP3-neo3 with ApaI and SalI. Conjugating CU428 and B2086 cells were transformed biolistically, as described previously (12), and transformants were selected with 100 g/ml paromomycin and 2.0 g/ml cadmium chloride. Strains lacking all three SEP genes were constructed by standard crosses and PCR-based typing of progeny, as described previously (69). GFP tagging. We used a total-cDNA (35) and a walking-primer PCR approach to verify the TIGR software-based prediction (Tetrahymena Genome Database, http://www.ciliate.org/) of 5⬘ and 3⬘ ends of the septin gene transcribed regions. We determined that the likely coding region for Sep3p is shorter than the prediction (TTHERM_00989350). To overexpress septins with an N-terminal green fluorescent protein (GFP) tag, we amplified the likely coding region with or without a fragment of the 3⬘ UTR from the genomic DNA, with the addition
of restriction sites, by using the following primers. The SEP1 coding region was amplified, with the addition of MluI and XhoI restriction sites (underlined), with primers 5⬘-ATAAACGCGTCGAGAACAATAGATGTGAT-3⬘ and 5⬘-TTTAT CTCGAGTTAGTTTTTTAGAATTTTGTATA-3⬘. The SEP2 coding region followed by 0.34 kb of the 3⬘ UTR was amplified, with the addition of MluI and XhoI restriction sites (underlined), with primers 5⬘-AATTACGCGTCATGGA CAATCAAAGAGTTC-3⬘ and 5⬘-TATTACTCGAGATTCATTTCCTTCAAT AAGTTG-3⬘. The SEP3 coding region followed by 0.6 kb of the 3⬘ UTR was amplified, with the addition of MluI and BamHI sites (underlined), with primers 5⬘-AATAAACGCGTCATGGACAGCTTTTATACTACT-3⬘ and 5⬘-AATAAT GGATCCTCATGAATGCTACTGCTAACA-3⬘. The SEP3 PCR fragment was digested with MluI and BamHI endonuclease restriction enzymes and ligated into pMTT1-GFP plasmid digested with the same enzymes. To clone SEP1 and SEP2 PCR fragments, we modified pMTT1-GFP by introducing a SalI site between the MluI and BamHI restriction sites. The SEP1 and SEP2 PCR fragments were digested with MluI and XhoI and ligated into the modified pMTT1GFP digested with MluI and SalI. The resulting pMTT1-GFP-Sep1, pMTT1GFP-Sep2, and pMTT1-GFP-Sep3 plasmids were digested with SacII and ApaI and used to transform CU522 cells (from Donna Cassidy-Hanley, Cornell University) as described previously (35). Transformed cells were selected with 20 M paclitaxel. To induce overexpression of GFP-tagged septins, transformants were grown overnight in PPYGFe medium without paclitaxel to the mid-log phase (about 2 ⫻ 105 cells/ml) and induced with cadmium chloride (2.5 g/ml). To express Sep1p as a C-terminal GFP fusion under the native promoter, the entire SEP1 coding region was amplified from the genomic DNA with the
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addition of SacII and MluI sites at the 5⬘ and 3⬘ ends, respectively, and cloned into pMTT1-Nrk2-GFP (83), digested with the same enzymes. The resulting pSEP1-GFP plasmid was digested with SacII and EcoRV, and the SEP1-GFP fragment followed by 0.6 kb of a 3⬘ BTU1 fragment was cloned into the neo3 plasmid with a 0.9-kb MTT1 promoter (pTvec-neo3) digested with SacII and SmaI to produce pSEP1-GFP-3⬘ BTU1-neo3. Next, 2.15 kb of the SEP1 3⬘ UTR was amplified from the genomic DNA with addition of ClaI and SacI sites at the 5⬘ and 3⬘ ends, respectively, and cloned into pSEP1-GFP-3⬘ BTU1-neo3 plasmid. Twenty micrograms of the resulting pSEP1-GFP-3⬘ BTU1-neo3-SEP1-3⬘ UTR plasmid was digested with SacI and SacII and used to transform CU428 cells. Transformants were selected in PPYGFe medium with 80 g/ml of paromomycin and 2 g/ml of cadmium chloride. Fluorescence microscopy. MitoTracker Red (Molecular Probes) and Mito Red (Fluka) were used at 20 to 200 nM. Cells were incubated for 30 min in the growth medium, washed with 10 mM Tris-HCl, pH 7.4, suspended in thick methylcellulose (76), and observed under a fluorescence microscope in the TRITC (tetramethyl rhodamine isothiocyanate) (excitation, 540 nm; emission, 590 nm) channel. Alternatively, the MitoTracker Red-labeled cells were fixed in 2% paraformaldehyde; washed in phosphate-buffered saline (PBS), 2% bovine serum albumin (BSA), 0.1% Tween 20; and mounted in Citifluor (Citifluor Ltd., London, United Kingdom). Cells were analyzed with a Leitz Wetzlar epifluorescence microscope or a Leica confocal microscope equipped with LAS AS software. Optical sections of 0.5 m were recorded, and images were processed with Adobe Photoshop 7.0. For detection of Sep1p-GFP expressed under the native promoter in live cells, we used an epifluorescence microscope equipped with a narrow-band G-excitation filter (excitation, 530 to 550 nm, and emission, 590 nm; Olympus America Inc., Melville, NY). Electron microscopy. Cells were washed with 10 mM Tris-HCl buffer, pH 7.4, and fixed for 1 h with a cold mixture of 2 parts of 0.05 M cacodylate buffer (pH 7.4), 1 part of 4% osmic acid, and 1 part of 6% glutaraldehyde on ice. After being washed three times for 20 min with cold 0.05 M cacodylate buffer, pH 7.4, cells were embedded in agar blocks, dehydrated in an ethanol series, and embedded in Durcupan (Sigma), as described previously (36). Ultrathin sections were contrasted with Reynolds lead citrate and uranyl acetate and observed under a JEOL 1200 EX electron microscope. Postembedding immunogold labeling of cells with septins tagged with GFP. Cells expressing GFP-tagged septins were washed with 10 mM Tris-HCl buffer, pH 7.4, and fixed with a mixture of 4% paraformaldehyde (Sigma) and 0.025% glutaraldehyde in 0.05 M cacodylate buffer, pH 7.4, for 1.5 h on ice. After four washes in cold 0.05 M cacodylate buffer, pH 7.4, cells were dehydrated in a series of ethanol and embedded in LR White resin (London Resin Company). After polymerization, embedded cells were cut into 80- to 100-nm sections and placed on nickel grids freshly covered with Formvar and coated with carbon. Sections were incubated in 3% BSA in PBS for 1.5 h and incubated with the anti-GFP antibodies (Santa Cruz) diluted 1:200 in 3% BSA in PBS for 1.5 h at room temperature. After two washes with 3% BSA in PBS and two washes with 1.5% BSA in PBS, grids were incubated overnight with anti-rabbit immunoglobulin G–10 nm gold-conjugated secondary antibodies (Sigma) at a 1:100 dilution in 3% BSA in PBS at 4°C. After several washes in PBS, grids were contrasted in aqueous saturated uranyl acetate mixed 1:1 with ethanol for 30 min. Sections were analyzed with a JEOL 1200 EX electron microscope. Western blot analysis. The GFP-tagged cells were grown in flasks to a density of 2 ⫻ 105 cells/ml. To extract total protein, 107 cells were collected by centrifugation, washed with 10 mM Tris-HCl, pH 7.5, and resuspended in cold 10 mM Tris-HCl, pH 7.5, with protease inhibitors (1 M leupeptin, 1 mM phenylmethylsulfonyl fluoride, 1 mM benzamidine, 1 mg/ml pepstatin A; Sigma Chemical Co.) to a 150-l final volume. Next, the cell suspensions were boiled in 5⫻ sodium dodecyl sulfate sample buffer for 5 min. Proteins were separated on 8% sodium dodecyl sulfate-polyacrylamide gels, transferred to a nitrocellulose membrane, blocked in 5% nonfat milk in TBST (150 mM NaCl, 20 mM Tris-HCl, pH 7.5, 0.05% Tween 20) for 1 h, and incubated with polyclonal anti-GFP antibodies (Santa Cruz) diluted 1:3,000 in 1% nonfat milk in TBST overnight at 4°C. After a wash in 5% nonfat milk in TBST, the membrane was incubated for 1 h in secondary horseradish peroxidase-conjugated antibody (Bio-Rad) diluted 1:10,000 in 1% nonfat milk in TBST. The blot was developed using an ECL kit (GE Healthcare).
RESULTS Septin genes of Tetrahymena thermophila. We used sequences of known septins from Saccharomyces cerevisiae and
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FIG. 2. GFP-tagged septins colocalize with mitochondria in live cells. (A to I) GFP images (A, D, and G), corresponding MitoTracker Red images (B, E, and H, respectively), and corresponding merged images (C, F, and I, respectively) are shown. (A to C) A GFP-Sep1poverproducing cell. The stars show bright foci of GFP fluorescence which do not stain with MitoTracker Red. The arrows show bright, vesicular structures. (D to F) A GFP-Sep2-overproducing cell. The arrows mark rods which do not colocalize with mitochondria. (G to I) A GFP-Sep3-overproducing cell. The arrows in panel I indicate the anterior region of the cell where a reduction in the density of mitochondria is seen. (J and K) GFP fluorescence in a WT cell (J) and a cell expressing Sep1-GFP under the native promoter (K). (L) A GFPSep1-overexpressing cell imaged at a higher magnification to show an intracytoplasmically located aggregate of mitochondria (g). Arrowheads point to cortically located mitochondria. Bar ⫽ 10 m.
mammals to search for related genes in the recently sequenced macronuclear genomes of free-living ciliates Tetrahymena thermophila (18) and Paramecium tetraurelia (5). Three septinencoding genes were identified in Tetrahymena (SEP1, SEP2, and SEP3), and four genes and a single pseudogene were found in Paramecium (L. Sperling, personal communication). The phylogenetic tree (Fig. 1A) calculated based on the conserved GTP-binding core domain of septins revealed several statistically supported clades. Most of these clades contain
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FIG. 3. Mitochondria are affected in septin-overproducing and septin-deficient cells. (A and B) Subcortical mitochondria in WT cells. Scission sites (s), intramitochondrial filamentous inclusions (f), and TMs are marked. (B) The scission sites are accompanied by flat ER cisternae. A Golgi stack (G) and a vesicular mitochondrion (vm) are indicated. (C) Abnormal morphotypes of mitochondrion in the Sep1-GFP strain. (D) Quantitative analysis of the ultrastructure of mitochondria in cells that either overproduce or are deficient in specific septins. The distributions of three morphotypes of mitochondria, swollen, vesicular, and normal, were determined on TEM sections. KO, knockout.
septins of invertebrates and vertebrates and some also contain fungal proteins, as earlier reported (39, 64). However, all ciliate septins belong to a clade that contains only protist sequences, including Chlamydomonas and Nannochloris. The sequences of the three predicted Tetrahymena septins are 29 to 31% identical (46 to 50% similar) to those of septins of metazoans or fungi. The Tetrahymena proteins have most of the conserved septin features (Fig. 1B). The core domains have the GDP/GTP exchange G1 (GxxxxGKS/T), G3 (DxxG), and G4 (xK/RxD) motifs, but the polybasic sequence is not apparent (see Fig. S1 in the supplemental material). A coiled-coil domain was predicted with high probability in Sep1p and with lower probability in Sep2p, but in contrast to that of fungal and metazoan septins, this domain was located near the N terminus. Surprisingly, a C-terminal transmembrane domain was predicted for all Tetrahymena septins as well as for Nannochloris and Chlamydomonas septins (Fig. 1B). None of the fungal or mammalian septins has a predicted transmembrane domain, but a putative transmembrane domain was predicted near the N-terminal end of the Unc61 septin of C. elegans (data not shown). Based on reverse transcription PCR, all three Tetrahymena septins are expressed in growing, conjugating, and cilium-regenerating cells (see Fig. S2A and B in the supplemental material). Tagged Tetrahymena septins colocalize with mitochondria. We expressed the predicted septin coding regions as GFP fusions under the cadmium-responsive promoter MTT1 (68). Based on Western blot analysis, fusion proteins were
detectable after 1.5 h of cadmium treatment (see Fig. S2C in the supplemental material) and their levels remained elevated for at least 24 h (data not shown). The rate of growth was reduced in all septin-overproducing strains in the presence of cadmium (see Fig. S2D in the supplemental material). Observations of live cells immobilized in methylcellulose revealed that all GFP-tagged septins localized primarily near the cell cortex (Fig. 2). Colabeling with MitoTracker Red showed that all GFP-tagged septins colocalized with mitochondria, although the patterns were not identical. In both wild-type (WT) cells (see Fig. 6A and B) and GFPtagged septin-overproducing cells (Fig. 2), the majority of mitochondria were localized in the proximity of the cell surface and were aligned along the ciliary rows (4, 48). Some septin-overproducing cells had mitochondria inside the cell body in the form of grapelike aggregates, and GFP-tagged septins also colocalized with these mislocalized internal mitochondria (Fig. 2A, C, and L). The cell body aggregates of Sep1-GFP produced an intense GFP signal, while the MitoTracker signal was relatively faint (Fig. 2B and C), indicating a reduced membrane potential in internal mitochondria (65). The GFP-Sep1p and GFP-Sep3p patterns closely matched the patterns of mitochondrial outlines revealed by MitoTracker Red (Fig. 2A, B, G, and H), while the GFP-Sep2p localization was distinct (Fig. 2D). In addition to mitochondrial outlines, the GFP-Sep2p signal was organized into rodlike structures and small patches in the proximity of mitochondria
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FIG. 4. GFP-tagged septins colocalize to and affect mitochondria and the ER. (A, C, E, and F) Standard TEM of strains overproducing septins. (B, D, G, and H) Postembedding immunogold electron microscopy of GFP-tagged septins. (A) In a GFP-Sep1p-overproducing cell, subcortical mitochondria (m) have an abnormally abundant associated ER. (B1 and B2) White arrows indicate gold grains corresponding to GFP-Sep1p,
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(Fig. 2D to F). The pattern of rods resembles the known pattern of transverse microtubule ribbons (TMs). The pattern of patches resembles the pattern of the Golgi apparatus (48). We also tagged Sep1p with GFP at the C terminus by targeting the endogenous locus. The pattern of distribution of Sep1p-GFP expressed under its own promoter closely resembled the pattern of ectopic GFP-Sep1 (Fig. 2J and K). Thus, the mitochondrial localization of Sep1p is not an artifact of overexpression and is not affected by the precise location of GFP within the tagged protein. Tetrahymena septins localize to endomembranes associated with mitochondria, and their overproduction destabilizes mitochondria. Transmission electron microscopy (TEM) showed that, in WT cells, mitochondria were located mainly near the cell surface along ciliary rows (Fig. 3A and B). As described previously (4, 44), the interior of the mitochondria was occupied by tubular cristae formed by the inner mitochondrial membrane (Fig. 3A and B). Rodlike inclusions that were previously identified as aggregates of the 14-nm filament protein (62) were seen in the matrix (Fig. 3A). Mitochondria were accompanied by the ER, with ribosomes located at the cytoplasmic side (Fig. 3B). Often, mitochondria were seen closely apposed to the alveolar membrane, especially those located under the longitudinal microtubules or TMs. Whenever two or more mitochondria were closely apposed, they were separated by flat cisternae of the ER (Fig. 3A and B). Dictyosomes of the Golgi apparatus consisting of one or two stacked cisternae accompanied by the transitional ER were found near cortical mitochondria (Fig. 3B), as described previously (24, 25, 48). Apparent dividing mitochondria with what appeared to be septa (44) were found in cells approaching cytokinesis (cells with an increased number of duplicating basal bodies). In such mitochondria, the invagination of the outer mitochondrial membrane was accompanied by ER membranes near the scission site (Fig. 3A and B). In addition to having normal mitochondria, WT cells occasionally had vesicles which resembled mitochondria based on their size but did not have internal tubular structures (Fig. 3B). These types of vesicles occasionally had rodlike filaments characteristic of mitochondria (data not shown). We suspect that these vesicles represent “old” mitochondria that undergo degradation. TEM analysis revealed that the addition of cadmium to WT cells did not change the ultrastructure (results not shown). However, GFP-tagged septin-overproducing cells showed severe alterations, primarily in the morphology of mitochondria and the associated ER. All three types of septin-overproducing cells had numerous vesicles smaller than mitochondria but larger than peroxisomes (Fig. 3C; see also Fig. 5 and Fig. S4 in the supplemental material). It appears that these structures correspond to vesicles with brightly labeled outlines seen in live cells (compare Fig. 2A and C and 3C) and represent remnants
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of degraded mitochondria prior to enclosure in autophagic vesicles. Depending on which GFP-tagged septin was overproduced, 30 to 40% of the mitochondria were swollen and/or in the process of degradation of tubules (Fig. 3D). Severe alterations in the morphology of the ER were found, especially in GFP-Sep1p- and GFP-Sep3p-overproducing cells (Fig. 4). Many Sep1p-GFP cells had a dilated ER (Fig. 4A). In GFPSep2p-overproducing cells, the ER appeared less abundant around cortical mitochondria. However, concentric stacks of the ER were present around abnormal mitochondria (see Fig. S4 in the supplemental material). In addition to having flat cisternae, the tubular ER was more abundant than that in control cells. These results are reminiscent of ER expansion in cells subjected to severe stress (8). Strikingly, GFP-Sep2p-overproducing cells had numerous mitochondria with an apparent septum which appeared as a fibrogranular material “squeezed in” between adjacent outer membranes (Fig. 4E). No such structures were found in WT cells, in cells expressing GFP-Sep1p or GFP-Sep3p, or in knockout cells lacking SEP2 (see below). In GFP-Sep3p-overexpressing cells, ectopic or abnormal mitochondrial fission events were observed (Fig. 4C). Quantitative immunogold labeling with anti-GFP antibodies confirmed that tagged septins colocalized with mitochondria (see Fig. S3 in the supplemental material). However, each septin was targeted to a distinct set of sites (Fig. 4; see also Fig. S3 in the supplemental material). GFP-Sep1p colocalized mostly with the outer mitochondrial membrane (Fig. 4B1 and B2). GFP-Sep2p localized to the outer membrane as well as to scission sites (that were induced by its overexpression) and TMs (Fig. 4F and H), while GFP-Sep3p localized primarily to the mitochondrial outer membrane (MOM) and the ER (Fig. 4D). A characteristic feature of cells overproducing each of the three septins was an increased number of autophagic vacuoles (Fig. 5) containing mitochondria in the course of degradation. Using the autophagic vacuole marker monodansylcadaverine (6, 60), we found that the number of cells containing more than five autophagosomes increased from 5% in WT cells (n ⫽ 100) to 30% in GFP-Sep1-overproducing cells (n ⫽ 100) (I. Sokolowska and M. Jerka-Dziadosz, unpublished data). Septins are required for normal stability of mitochondria and nuclei. Single knockouts of the SEP2 or the SEP3 gene did not affect the rate of cell multiplication, but cells lacking SEP1 grew more slowly (see Fig. S2E in the supplemental material). Furthermore, cells lacking SEP1 showed increased mortality when isolated. Among 53 clones, only 12 gave thriving cultures. The rates of cell motility of septin-deficient cells were similar to those for the WT (results not shown). Based on confocal imaging with MitoTracker, knockout cells (similarly to overexpressing cells) had an increased number of mitochondria located inside the cell body compared to that for the WT (compare Fig. 6C, E, G, I, K, and M to A). In some
associated with the outer mitochondrial membrane, ER, and cristae. (C and D) Subcortical mitochondria in GFP-Sep3p-overproducing cells. In panel C, arrows point to scission sites in the mitochondrion. In panel D, gold grains in immunolabeled cells (indicated by white arrows) are localized near the ER and the MOM. (E and F) TEM of subcortical mitochondria in Sep2-GFP-overproducing cells. (E) Abnormal scission events (s). Note the presence of electron-dense material in the septa. (F) TM overlying the mitochondrion adjacent to the scission figure (s). (G and H) Specific immunogold labeling at scission structures (s). bb, basal body; f, filament. Bar ⫽ 200 nm.
FIG. 5. Overexpression of GFP-Sep2 leads to an excessive number of mitochondria and autophagic vacuoles containing remnants of mitochondria (compare Fig. S6 in the supplemental material). Abundant mitochondria are visible both near the cell cortex and as subectoplasmic and endoplasmic groupings (arrowheads). Vesicular, apparently in situ-degraded mitochondria are also present, as is an excessive ER. AV, autophagic vacuole; bb, basal body; vm, vesicular mitochondrion. Bar ⫽ 200 nm. 1379
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FIG. 6. Cells lacking septins display abnormalities in the morphology and localization of mitochondria. Growing WT cells (A and B) and septin knockout (KO) cells (C to P) were fixed in paraformaldehyde and labeled with Mito Red (mitochondria) and Hoechst stain (nuclei) and observed with a confocal microscope. Twenty to 25 sections 0.5 m thick were recorded for each cell. Pairs of images represent sets of optical sections covering the cell interior (A, C, E, G, I, K, and M) and the ventral cortical layer (B, D, F, H, J, L, N, O, and the dorsal site [P]). In panels A, C, E, G, and M, arrows mark the Mic; in panels O and P, arrows point to abnormally long mitochondria. Cells lacking SEP1 (I to L) have a swollen Mac and lack a discernible Mic. Bar ⫽ 10 m.
cells lacking SEP1, the mitochondria appeared smaller and more numerous (Fig. 6K and L). Thus, in subtle ways, the absence of septins alters the organization and morphology of mitochondria. Cells lacking septin genes showed abnormalities in the structure of nuclei, based on Hoechst staining. In the WT (Fig. 7), 97% of cells had a normal nuclear composition, with one macronucleus (Mac) and one micronucleus (Mic). In strains lacking SEP1, 44% of cells lacked a detectable Mic (Fig. 6I and
K), which correlates with the high clonal mortality of this strain, as the presence of a Mic is needed for viability (31). In cells lacking SEP2, amicronucleate cells were not seen, but 15% of cells had more than one Mic. In cells lacking SEP3, about 50% of cells had various nuclear abnormalities (Fig. 7). TEM analysis of septin knockout strains revealed frequent defects in mitochondria. Cortical mitochondria with apparent fission sites were more frequent than in the WT (Fig. 8A to C). Many mitochondria in knockout strains were round and
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FIG. 7. Cells lacking septin genes (knockout, KO) have abnormalities in the composition of nuclei (either supernumerary or absent Mics). Samples of growing cultures were fixed and labeled with Hoechst stain. The numbers of cells with one Mac and one Mic (normal), one Mac and two or three Mics (abnormal), and one Mac but no Mic (abnormal) were scored on slides. Bars represent the percentages of different subpopulations in five studied strains. The numbers on top of the bars represent the sample sizes.
slightly swollen, and the tubular cristae were rarified, resulting in an “empty” interior (Fig. 8D; see also Fig. S5 and S6 in the supplemental material); these mitochondria were similar to swollen mitochondria in overexpressing cells (compare sm in Fig. S4 in the supplemental material). Vesicular mitochondria with an “empty” interior were frequent (Fig. 8E). The morphological features of damaged mitochondria were similar in all types of septin-deficient strains but were particularly severe in cells lacking Sep2p, where normal mitochondria comprised only 35% of the whole population (Fig. 3D; see also Fig. S6 in the supplemental material). Autophagic vacuoles with remnants of mitochondria, micronuclei (Fig. 8F and I), and occasionally cilia or basal bodies were encountered in all studied septin knockout strains. The morphology of the autophagic vacuoles resembled those described previously for ageing and starved Tetrahymena cells (21). TEM also revealed abnormal nuclei in septin knockout cells. Cases of decomposition of the nuclear envelope in Macs and Mics were seen (Fig. 8G, H, J, and K). The micronuclear envelope could be disrupted at multiple sites, and some Mics were swollen. Instead of the darkly contrasted chromatin bodies seen in the WT (see Fig. S7 in the supplemental material), the nuclear contents appeared as fine threads and no discernible chromatin bodies were present (Fig. 8H and J). Even in Mics engaged in mitosis, the nuclear envelope membrane was sometimes disrupted (Fig. 8K). These damaged Mics were sometimes enclosed in an autophagic vacuole or incorporated into a larger vacuole containing other organelles, such as mitochondria and cilia (Fig. 8F and I). On some sections, partially decomposed macronuclear envelopes were seen. In Macs with a missing envelope, the peripheral nucleoli were not present and mixing of the nucleoplasm with cytoplasmic organelles was observed (Fig. 8G). To test the possibility that Tetrahymena septins are functionally overlapping, we used crosses to create a strain that lacks SEP1, SEP2, and SEP3. Surprisingly, the growth and motility rate of this strain were similar to those of the WT (see Fig. S2E in the supplemental material). Labeling of mitochondria with MitoTracker and nuclei with Hoechst stain (Fig. 6M to P) did not reveal any obvious difference compared to the single knockouts, except for the occasional presence of unusually long and thin mitochondria (Fig. 6O and P). About 8% of cells (n ⫽ 109) had an abnormal composition of nuclei (Fig. 7).
TEM of triple knockout strains showed that about 20% of mitochondria were abnormal and swollen and lacked internal tubules and that 10% of mitochondria were in a process of in situ degradation (n ⫽ 293) (Fig. 3D and 9B). The ultrastructure of affected mitochondria resembled that of single knockouts, i.e., most were swollen, with rarified internal tubules. Thin, long mitochondria were located in the subcortical regions (Fig. 9A) and appeared as if the final splitting of dividing mitochondria was inhibited. Abnormal micronuclei were rare, and their morphology was similar to that of affected micronuclei in single knockouts but with less-severe membrane damage (compare Fig. 9B and 8H and J). Abundant autophagosomes similar to those found in single gene knockouts were present in all cells with affected mitochondria. It thus appears that the effects of individual knockouts are not synergistic. Moreover, in some respects, the single knockout strains showed more pronounced defects than the triple knockout strain. DISCUSSION Until recently, septin proteins were known to be present in fungi and metazoans but not in higher plants or parasitic protists, like Giardia (78) or Plasmodium fasciculatum (17). However, septin genes were recently found in the green algae Chlamydomonas reinhardtii and Nannochloris spp. (78). Here, we report the presence of septin genes in the free-living ciliates Tetrahymena thermophila and Paramecium tetraurelia. We complement earlier phylogenetic studies (51, 59, 64) by showing that septins of ciliates and green algae form a separate clade. Moreover, our functional studies support the proposed phylogeny by showing that ciliate septins have evolved unique functions. Septins of fungi and metazoans have a well-established role in cytokinesis. Thus, it was surprising that none of the tagged Tetrahymena septins colocalized with the fission furrow, known to contain a filamentous structure (36). Instead, the Tetrahymena septins were found to be associated with mitochondria. It will be important to investigate whether septins of other free-living protists, such as Chlamydomonas reinhardtii, also participate in mitochondrial functions. The Tetrahymena and Chlamydomonas (but not Paramecium) septins have a predicted transmembrane domain, and this feature could
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play a role in the mitochondrial functions of some protistan septins. This protein topology enables targeting of proteins to the ER and the MOM (9, 10). Tagged Tetrahymena septins localized to the MOM and the neighboring ER or formed septumlike structures. Among the known septins, mitochondrial localization is rare (39, 52). Two splice variants of Sep4, ARTS and M-septin, localize to mitochondria in mammalian cells (41, 49). ARTS has a proapoptotic activity (50). Deletion of ARTS in mice resulted in abnormal sperm development, with affected mitochondria (34). M-septin plays an undefined role in apoptosis of neural cells (74). A comparison of ARTS and M-septin with septinlike proteins of Tetrahymena revealed no significant sequence homology besides the central core domain that is conserved in all septins (Fig. 1B; see also Fig. S1 in the supplemental material). Moreover, it appears that ciliate and mammalian septins have nonidentical mitochondrial functions. First, unlike ARTS and M-septins, ciliate septins lack a mitochondrial targeting signal. Second, while ARTS has a proapoptotic activity, ciliate septins appear to protect against apoptosis-like damage of mitochondria and nuclear envelopes. We propose that the mitochondrial association of Tetrahymena and mammalian septins has resulted from the recruitment of septins to mitochondria, which has occurred independently in the lineages of protists and mammals. Studies of Tetrahymena have recently revealed a case of convergent evolution of another type of membrane-associated GTPases, dynamins. Elde and colleagues showed that the adaptation of some dynamins to endocytosis has occurred independently in the lineages of ciliates and metazoans (20). In contrast to mammalian and yeast cells, where the mitochondrial population is in constant flux, driven by organelle fusion and fission (30), genetic studies of Paramecium tetraurelia indicate that in this ciliate, mitochondria do not undergo fusion events (66). Several observations reported here indicate that, in Tetrahymena, septins play a role in fission of mitochondria. The disintegrating mitochondria in the single septin knockouts (Fig. 8D and E; see also Fig. S6 in the supplemental material) and abnormally long cortical mitochondria in triple knockouts (Fig. 9A and B) most likely are manifestations of defects in mitochondrial fission. In addition, the abnormal scission events in Sep3-overexpressing cells (Fig. 4C) and the induction of apparent mitochondrial septa (Fig. 4E and F), as well as the presence of clusters of small grapelike aggregates of mitochondria in septin-overproducing cells (Fig. 2A and L and 5), are consistent with the involvement of septins in mitochondrial fission. In Trypanosoma brucei, the parasitic protist that lacks septin sequences in the genome, the dynamics of mitochondria are regulated by another GTPase, dynamin. Deletion of the single dynamin gene in Trypanosoma inhibited both mitochondrial fission and endocytosis (13). Dynamins have been implicated in both fission and fusion of mitochondria in other organisms (30). Tetrahymena has several dynamin-related proteins. While some of these proteins participate in endocytosis at the parasomal sac, Drp7p dynamin localizes to subcortical mitochondria (20). This observation opens the possibility that, in Tetrahymena, septins and a subset of dynamins work together in the regulation of mitochondrial dynamics. This could explain why Tetrahymena cells lacking all septins remain viable.
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Mitochondria play a critical role in the induction of apoptosis in vertebrates by providing a source of apoptosis-inducing agents, such as cytochrome c (for a review, see references 47 and 77). The permeabilization of mitochondria and the resulting activation of cysteine proteases leading to cell death have been described for the unicellular eukaryote Leishmania major (2) and for Dictyostelium discoideum (3). While Tetrahymena does not appear to undergo cell apoptosis, the Mac undergoes elimination at the end of mating via an apoptosis-like mechanism (programmed nuclear death [PND]). It has been suggested that a mitochondrial endonuclease similar to human endonuclease G is involved in PND (42). Also, a caspaselike activity may be involved in PND in Tetrahymena (19, 42). We, however, were not able to find an obvious caspase or an endonuclease G-encoding sequence in the Tetrahymena genome (results not shown). This does not rule out the possibility that Tetrahymena has apoptotic cysteine proteases and endonucleases that are divergent. In addition, calpain proteases have been implicated in apoptosis (80) and the Tetrahymena genome encodes calpain homologs (18). On the other hand, AIF (apoptosis-inducing factor), when released from mitochondria, is sufficient to induce apoptosis of isolated nuclei independently of caspases (72). AIF is normally localized inside mitochondria and translocates to the nucleus when apoptosis is induced (71). The Tetrahymena genome encodes a protein with high homology to human AIF (TTHERM_00622710). We hypothesize that, in Tetrahymena, deficiencies in septin functions induce permeabilization of mitochondria and leakage of proapoptotic factors, causing damage to the nuclear envelope. The fact that both overexpression and depletion of septins lead to similar defects in mitochondria and nuclei suggests that a specific concentration of septins stabilizes mitochondria. An RNA interference knockdown of the Septin2 gene in Paramecium tetraurelia gave a nuclear phenotype similar to the gene knockout phenotypes described here for Tetrahymena (M. Jerka-Dziadosz and J. K. Nowak, unpublished observations). In starved Tetrahymena cells, a small fraction of mitochondria showed signs of autophagic degradation (43). This observation suggests that a turnover of mitochondria exists and that damaged or old mitochondria are moved away from the cortical region. In septin knockout cells (Fig. 6), the internal presence of mitochondria increases, suggesting that septins play a role in the turnover of mitochondria. The fact that septinlike proteins associate with the ER near mitochondria is, to our knowledge, a novel observation. It is known that mitochondrion-associated ER domains have clearly defined functions in the transfer of lipids between the ER and mitochondria. For example, the glycosylphosphatidylinositol biosynthetic pathway is confined to these ER domains (79) and a disruption of ER function also contributes to cellular apoptosis (11, 63, 82). Other studies revealed that defects in the ER affect the reorganization of the nuclear membrane during division (27, 32, 57). The ER stress is coupled to the mitochondrial intrinsic apoptotic pathway through BH-3 family proteins (67). We propose that depletion of septins in the ER contributes to the apoptosis-like events in mitochondria and nuclei by destabilizing the turnover of mitochondrial membrane. It was surprising that the Tetrahymena knockout strains lack-
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FIG. 9. Triple septin knockout (KO) cells have abnormal mitochondria. (A) Section of an abnormally long mitochondrion that spans five TMs of the ciliary row. (B) Section with many swollen and vacuolized mitochondria (sm and vm, respectively). The macronuclear and micronuclear envelopes are discontinuous (arrows). Some autophagic vacuoles are located near damaged mitochondria. AV, autophagic vacuole; bb, basal body; f, filament; m, mitochondrion.
ing single or multiple septins were viable. Moreover, some single knockout strains had more-severe defects than the triple knockout strain lacking septin functions entirely (Fig. 3D). However, septins form heteropolymers in vivo (70). It is possible that some phenotypic changes in the single knockout strains result from the accumulation of septins that cannot be
incorporated into a heteropolymer. Nevertheless, all septindeficient strains are viable. However, the detrimental effects of the lack of septins on mitochondria, possibly resulting in the release of nuclear-damage-inducing factors, could be compensated by massive induction of autophagic vacuoles. In other species, autophagy provides an important surveillance mecha-
FIG. 8. TEM reveals autophagic and apoptosis-like changes in septin gene knockout strains. (A to C) Mitochondrial scission sites (s) in cells lacking Sep1p (A), Sep2p (B), and Sep3p (C), which morphologically do not differ from WT cells (Fig. 3A and B). (D) A swollen mitochondrion (sm) in a SEP1 knockout (KO) cell. (E) A vesicular mitochondrion (vm) adjacent to a normal mitochondrion (m) closely apposed to the TM in a SEP2 knockout cell. (F) An autophagic vacuole (AV) in the SEP1 knockout cell, with remnants of nuclear material (nu) and compact mitochondria. (G) A Mac with a disrupted nuclear envelope (NE) (arrows). Remnants of the envelope are visible on the right side of the image. (H) A decomposing Mic in a SEP2 knockout cell. Arrows indicate regions missing the nuclear envelope. Arrowheads point to nuclear pores. (I) A decomposing Mic that appears to be in the process of incorporation into an autophagic vacuole. Arrows point to membranes of the forming autophagosome. (J) A decomposing Mic in a cell lacking SEP2. (K) A Mic in early mitotic prophase with a decomposed nuclear envelope (arrows) in a SEP3 knockout cell; the arrowhead indicates the autophagosome membrane. NPC, nuclear pore. Bar ⫽ 200 nm.
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nism which cells employ to degrade damaged or obsolete organelles and proteins (38, 85) and autophagy has been shown to play a central role in the degradation of mitochondria (58). Thus, we suggest that, in ciliates and possibly other free-living protists, septins play a role in the maintenance of mitochondria and that the deleterious consequences of their absence can be compensated by increased autophagy. ACKNOWLEDGMENTS We thank Linda Sperling (CGM, Gif-sur-Yvette) for annotations of septin genes in Paramecium. We also thank Linda Sperling, Janine Beisson (CGM, Gif-sur-Yvette), and Krzysztof Zablocki (The Nencki Institute) for comments on the manuscript. We thank the staff of the EM and Confocal Microscopy Laboratories of the Nencki Institute for skillful technical assistance. We thank Michelle Momany (University of Georgia, Athens, GA) for advice on imaging of GFP in live cells. This work was supported by statute grants from the Polish Ministry of Science and Higher Education to the Nencki Institute and by National Science Foundation grant MCB-033965 (to J.G.). REFERENCES 1. Adam, J. C., J. R. Pringle, and M. Peifer. 2000. Evidence for functional differentiation among Drosophila septins in cytokinesis and cellularization. Mol. Biol. Cell 11:3123–3135. 2. Arnoult, D., K. Akarid, A. Grodet, P. X. Petit, J. Estaquier, and J. C. Ameisen. 2002. On the evolution of programmed cell death: apoptosis of the unicellular eukaryote Leishmania major involves cysteine proteinase activation and mitochondrion permeabilization. Cell Death Differ. 9:65–81. 3. Arnoult, D., I. Tatischeff, J. Estaquier, M. Girard, F. Sureau, J. P. Tissier, A. Grodet, M. Dellinger, F. Traincard, A. Kahn, J. C. Ameisen, and P. X. Petit. 2001. On the evolutionary conservation of the cell death pathway: mitochondrial release of an apoptosis-inducing factor during Dictyostelium discoideum cell death. Mol. Biol. Cell 12:3016–3030. 4. Aufderheide, K. 1979. Mitochondrial associations with specific microtubular components of the cortex of Tetrahymena thermophila. I. Cortical patterning of mitochondria. J. Cell Sci. 39:299–312. 5. Aury, J. M., O. Jaillon, L. Duret, B. Noel, C. Jubin, B. M. Porcel, B. Segurens, V. Daubin, V. Anthouard, N. Aiach, O. Arnaiz, A. Billaut, J. Beisson, I. Blanc, K. Bouhouche, F. Camara, S. Duharcourt, R. Guigo, D. Gogendeau, M. Katinka, A. M. Keller, R. Kissmehl, C. Klotz, F. Koll, A. Le Mouel, G. Lepere, S. Malinsky, M. Nowacki, J. K. Nowak, H. Plattner, J. Poulain, F. Ruiz, V. Serrano, M. Zagulski, P. Dessen, M. Betermier, J. Weissenbach, C. Scarpelli, V. Schachter, L. Sperling, E. Meyer, J. Cohen, and P. Wincker. 2006. Global trends of whole-genome duplications revealed by the ciliate Paramecium tetraurelia. Nature 444:171–178. 6. Bampton, E. T., C. G. Goemans, D. Niranjan, N. Mizushima, and A. M. Tolkovsky. 2005. The dynamics of autophagy visualized in live cells: from autophagosome formation to fusion with endo/lysosomes. Autophagy 1:23–36. 7. Beites, C. L., H. Xie, R. Bowser, and W. S. Trimble. 1999. The septin CDCrel-1 binds syntaxin and inhibits exocytosis. Nat. Neurosci. 2:434–439. 8. Bernales, S., K. L. McDonald, and P. Walter. 2006. Autophagy counterbalances endoplasmic reticulum expansion during the unfolded protein response. PLoS Biol. 4:e423. 9. Borgese, N., S. Brambillasca, and S. Colombo. 2007. How tails guide tailanchored proteins to their destination. Curr. Opin. Cell Biol. 19:368–375. 10. Borgese, N., I. Gazzoni, M. Barberi, S. Colombo, and E. Pedrazzini. 2001. Targeting of a tail-anchored protein to endoplasmic reticulum and mitochondrial outer membrane by independent but competing pathways. Mol. Biol. Cell 12:2482–2496. 11. Breckenridge, D. G., M. Germain, J. P. Mathai, M. Nguyen, and G. C. Shore. 2003. Regulation of apoptosis by endoplasmic reticulum pathways. Oncogene 22:8608–8618. 12. Cassidy-Hanley, D., J. Bowen, J. H. Lee, E. Cole, L. A. VerPlank, J. Gaertig, M. A. Gorovsky, and P. J. Bruns. 1997. Germline and somatic transformation of mating Tetrahymena thermophila by particle bombardment. Genetics 146:135–147. 13. Chanez, A. L., A. B. Hehl, M. Engstler, and A. Schneider. 2006. Ablation of the single dynamin of T. brucei blocks mitochondrial fission and endocytosis and leads to a precise cytokinesis arrest. J. Cell Sci. 119:2968–2974. 14. Chant, J. 1996. Septin scaffolds and cleavage planes in Saccharomyces. Cell 84:187–190. 15. Cole, E., J. Frankel, and L. Jenkins. 1988. Interactions between janus and bcd cortical pattern mutants in Tetrahymena thermophila. An investigation of intracellular patterning mechanisms using double-mutant analysis. Rouxs Arch. Dev. Biol. 197:476–489. 16. Dobbelaere, J., and Y. Barral. 2004. Spatial coordination of cytokinetic events by compartmentalization of the cell cortex. Science 305:393–396.
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