* Department of Biology, University of Utah, Salt Lake City, Utah 84112; Department of Developmental Biology and
Department of Genetics, Stanford University School of Medicine, Stanford, California 94305; and § Department of Molecular and
Cellular Biology, University of California, Davis, California 95616
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Abstract |
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Membrane fusion is required to establish the morphology and cellular distribution of the mitochondrial compartment. In Drosophila, mutations in the fuzzy onions (fzo) GTPase block a developmentally regulated mitochondrial fusion event during spermatogenesis. Here we report that the yeast orthologue of fuzzy onions, Fzo1p, plays a direct and conserved role in mitochondrial fusion. A conditional fzo1 mutation causes the mitochondrial reticulum to fragment and blocks mitochondrial fusion during yeast mating. Fzo1p is a mitochondrial integral membrane protein with its GTPase domain exposed to the cytoplasm. Point mutations that alter conserved residues in the GTPase domain do not affect Fzo1p localization but disrupt mitochondrial fusion. Suborganellar fractionation suggests that Fzo1p spans the outer and is tightly associated with the inner mitochondrial membrane. This topology may be required to coordinate the behavior of the two mitochondrial membranes during the fusion reaction. We propose that the fuzzy onions family of transmembrane GTPases act as molecular switches to regulate a key step in mitochondrial membrane docking and/or fusion.
Key words: GTPase; membrane fusion; mitochondria; mtDNA; organelle morphology ![]() |
Introduction |
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THE homotypic, or self-fusion, of mitochondrial membranes plays an important role in controlling the
copy number and cellular distribution of these essential organelles (Bereiter-Hahn and Voth, 1994). In
budding yeast and some mammalian cells, frequent mitochondrial fusion events are balanced by mitochondrial fission, maintaining the compartment as a highly branched,
tubular network (Hoffman and Avers, 1973
; Stevens, 1981
;
Koning et al., 1993
; Nunnari et al., 1997
; Hermann, 1998
).
Mitochondrial fusion also functions to remodel mitochondrial morphology during differentiation. Individual
mitochondria in rat skeletal muscle cells undergo postembryonic fusion to generate a branched mitochondrial reticulum (Bakeeva et al., 1981
). During spermatogenesis in insects, fusion generates giant mitochondria that associate
with the axoneme of the sperm flagella (Lindsley and
Tokuyasu, 1980
; Fuller, 1993
). The generation of such
large, interconnected mitochondrial compartments is
thought to facilitate the distribution of energy and metabolites as well as chemical and electrical signals throughout
these cells (Bakeeva et al., 1978
; Ichas et al., 1997
). The genetic and molecular mechanisms that control mitochondrial fusion have not been defined.
Molecules that regulate the heterotypic and homotypic
fusion of membranes in the secretory and endocytic pathways have been studied in some detail. Both types of
fusion require the cytosolic factors NSF and SNAPs (or
homologues) as well as compartment-specific integral
membrane proteins termed v- and t-SNAREs (Denesvre
and Malhotra, 1996; Pfeffer, 1996
; Rothman, 1996
; Hay
and Scheller, 1997
; Edwardson, 1998
; Götte and Fischer
von Mollard, 1998
; Patel et al., 1998
; Rabouille et al.,
1998
). During heterotypic fusion, v/t-SNARE pairing promotes the stable association of the vesicle with the target
membrane and may be sufficient to catalyze bilayer mixing (Weber et al., 1998
). In contrast, the homotypic fusion of ER and Golgi membranes depends on self-interactions
among resident t-SNAREs (Patel et al., 1998
; Rabouille et
al., 1998
). Yeast vacuole fusion requires a v/t-SNARE
pairing for optimal fusion but can occur (with lower efficiency) between vacuoles containing only the t-SNARE
partner (Nichols et al., 1997
). Thus, in some cases, the recognition of "like" membranes in homotypic fusion reactions may be mediated by homomeric interactions between compartment-specific t-SNAREs. NSF and SNAPs
appear to act generally during fusion to disrupt unproductive SNARE associations within a membrane before docking (Mayer et al., 1996
; Otto et al., 1997
) and may also regulate changes in SNARE conformation after docking but
before membrane fusion (Söllner et al., 1993
).
Although an isoform of the VAMP-1 SNARE has been
reported to localize to mitochondria in transfected epithelial cells (Isenmann et al., 1998), molecules in the NSF,
SNAP, and SNARE families have not been implicated in
mitochondrial fusion. This is not entirely unexpected since
mitochondrial fusion, unlike most other membrane fusion
reactions in the cell, requires the sequential mixing of two
distinct lipid bilayers. Moreover, the mitochondrial compartment is evolutionarily distinct from compartments of
the secretory and endocytic pathways. Thus, the molecular
machinery that mediates mitochondrial fusion is likely to
be unique, even though the general mechanism of membrane fusion may be conserved. A novel gene required for
mitochondrial fusion, fuzzy onions (fzo),1 was recently
identified through the analysis of Drosophila mutants defective in spermatogenesis (Hales and Fuller, 1997
). During spermatid differentiation in flies, individual mitochondria coalesce, fuse into two giant organelles, and coil
tightly around one another forming the Nebenkern, a
spherical structure that resembles an onion in cross section
(Lindsley and Tokuyasu, 1980
; Fuller, 1993
). Mutations in
the fzo gene disrupt mitochondrial fusion and result in
disorganized aggregates of individual mitochondria that
do not associate properly with the elongating axoneme
(Hales and Fuller, 1997
). fzo encodes a predicted transmembrane GTPase that is detected on the mitochondrial
compartment just as fusion begins and disappears soon after fusion is complete. Together, the fzo mutant phenotype and the expression pattern of the Fzo protein suggest
that this molecule either regulates, or is a direct mediator of, mitochondrial fusion.
The discovery of fzo homologues in yeasts, nematodes,
and mammals defined a new family of multiple domain,
high molecular weight GTPases, and raised the possibility
that these molecules act generally to control mitochondrial fusion events in different organisms and cell types
(Hales and Fuller, 1997). Fzo family members contain an
amino-terminal GTPase domain, two adjacent carboxy-terminal transmembrane domains, and multiple heptad repeats (see Fig. 1 A) (Hales and Fuller, 1997
). Here we
show that the Saccharomyces cerevisiae orthologue of
Drosophila Fzo, Fzo1p, is a mitochondrial integral membrane protein required to maintain a tubular mitochondrial reticulum during mitotic growth and demonstrate a
direct role for this protein in mitochondrial fusion during yeast mating. Mutations in two conserved FZO1 GTPase
motifs disrupt mitochondrial fusion but do not affect
Fzo1p localization. Subcellular fractionation and protease
protection experiments reveal that the amino terminus of
the protein (containing the essential GTPase domain and
multiple heptad repeats) extends into the cytoplasm where
it could participate in organelle docking/fusion and interact with molecules that regulate GTP binding or hydrolysis.
In addition, the distribution of Fzo1p in submitochondrial
fractionation studies suggests that the carboxy-terminus of
this protein may interact with both mitochondrial membranes. This topology is similar to that of SNARE molecules and viral fusion proteins and suggests that Fzo1p
could play a direct role in the docking and fusion of mitochondrial compartments.
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Materials and Methods |
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Strains and Genetic Techniques
All yeast strains (Table I) are isogenic to FY10 (Winston et al., 1995). All
media including YPDextrose, YPGlycerol, SDextrose, SGalactose, and
SRaffinose were prepared as described by Sherman et al. (1986)
. Strains
were grown at 30°C unless otherwise noted. Genetic and molecular cloning manipulations followed standard techniques (Maniatis et al., 1982
;
Sherman et al., 1986
; Baudin et al., 1993
). The Escherichia coli host strain
JM109 (Promega Corp., Madison, WI) was used for all bacterial manipulation of plasmids.
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Plasmid Construction
For pRS416-FZO1, a genomic fragment containing the FZO1 coding region
plus 500 bp of 5'and 3' flanking sequence was PCR amplified from JSY999
using primers containing engineered EcoRI sites: P297 (5'-GGGGGAATTCCCAGGTGACAGAATGTCTGGGTTGAAAG-3') and P298
(5'-GGGGGAATTCCTTGCTCCTTGTTGTCTTTTAAATGGAG-3'), and cloned into the unique EcoRI site in pRS416 (verified by sequencing; Stratagene, La Jolla, CA). An EcoRI fragment from pRS416-FZO1 was
subcloned into pRS414 (Stratagene) to generate pRS414-FZO1. The
pRS414-FZO1 and pRS416-FZO1 plasmids complemented both the mitochondrial morphology defects and the loss of mtDNA in fzo1. To generate a GAL1 regulated form of FZO1, a PCR fragment containing a
3XMYC tag at the amino terminus of FZO1 (N-3XMYC-FZO1) was amplified from JSY2028 (see below) using primers with engineered SalI,
P379 (5'-GGGGGTCGACTATCTAATCGATGTCTAAATTTATTTCTTC-3'), and XbaI, P380 (5'-GGGGTCTAGATTAACGATGTCTAGGGAACAAAAGCTGGAG-3'), sites. The PCR fragment was cloned
into pRS415-GAL1 (Mumberg et al., 1994
) (American Type Culture Collection, Rockville, MD) to generate pGAL1-N-3XMYC-FZO1 (pRS415-
GAL1-N-3XMYC-FZO1, verified by sequencing). Growth in SRaffinose
provided sufficient expression of N-3XMYC-Fzo1p from the GAL1 promoter to complement the mitochondrial morphology and growth defects
of fzo1
cells. To generate mutations in the conserved FZO1 GTPase domain, a 3.5-kb EcoRI fragment from pRS416-FZO1 was cloned into the
EcoRI site of pALTER-1 (Promega Corp.) to create pALTER-1-FZO1.
Site-directed mutagenesis (Altered Sites II; Promega Corp.) was performed with the following mutagenic oligonucleotides: K200A (5'-GATGTAAATACTGGCGCCTCAGCTCTTTGCAAC-3', introduces an EheI
site); S201N (5'-CAGGTGATGTAAATACTGGTAAAAATGCATTATGCAACTCTCTATTAAAGCAGCG-3', introduces an NsiI site);
T221A (5'-GGATCAGCTACCATGCGCAAATGTATTTTCCGAA-3', introduces an Fsp I site); K371A (5'-GTTTTTTGTTGTGAAAGCTTTTGACAAAATCAGGG-3', introduces a HindIII site). The mutants were identified by restriction digest and confirmed by sequence analysis. The mutagenized fzo1 genes were subcloned into the EcoRI sites of
pRS414 and pRS424.
Generation and Characterization of fzo1::HIS3 Cells
The fzo1::HIS3 mutation was generated by transforming the diploid
strain JSY1373 with a PCR fragment containing 50 bp of FZO1 flanking
sequence interrupted by the HIS3 gene (Baudin et al., 1993
). The fzo1
::
HIS3 disruption precisely removed the entire FZO1 coding sequence and
was verified by PCR analysis. In 39 tetrads from the sporulated heterozygous diploid (JSY1808), the HIS3 marker segregated 2:2 with a slow
growth defect on YPDextrose and an inability to grow on YPGlycerol.
Mitochondrial morphology was visualized in four tetrads using a primary
anti-porin antiserum (1:200 dilution; Molecular Probes, Eugene OR) and
a secondary goat anti-mouse FITC antibody (1:100 dilution) (Jackson ImmunoResearch Laboratories, West Grove, PA) (Pringle et al., 1991
).
DAPI (4',6-diamidino-2-phenylindole; 25 ng/ml) was included in the
mounting medium to visualize mtDNA. The loss of mtDNA was confirmed by mating JSY1810 (fzo1
) to a rhoo tester strain JSY2555 (FZO1,
rhoo). pRS416-FZO1 was transformed into a fzo1
strain (JSY1810) to generate JSY2579 (wild-type mitochondrial morphology, no detectable mtDNA). To generate the rhoo strain JSY2555 (wild-type mitochondrial
morphology, no detectable mtDNA), a rho+ strain (JSY999) was grown
twice to saturation in synthetic minimal medium containing 25 µg/ml
ethidium bromide (Fox et al., 1991
).
Electron microscopy of wild-type (JSY1812) and fzo1 (JSY1810) cells
was performed essentially as described with the following modifications
(Yaffe, 1995
). The strains were grown in YPDextrose before fixation, and
two additional changes of anhydrous Spurr resin (Polysciences, Inc., Warrington, PA), followed by overnight incubation, were used to achieve
maximum infiltration of the samples.
Depletion of N-3XMYC-Fzo1p
To deplete the N-3XMYC-Fzo1p, the pGAL1-N-3XMYC-FZO1 plasmid
was transformed into JSY2038 (fzo1 rho+ + pRS416-FZO1) and cells
that had lost the pRS416-FZO1 plasmid were selected on SRaffinose medium containing 5-FOA (5-fluoro-orotic acid) to yield JSY2273 (fzo1
rho+ + pGAL1-N-3XMYC-FZO1). JSY2273 was grown for 24 h in
SRaffinose medium lacking leucine to select for pGAL1-N-3XMYC-
FZO1. To block expression of N-3XMYC-Fzo1p, the cells were collected,
rinsed, and grown in SDextrose minus leucine to a density of 2.5 × 106
cells/ml. Mitochondrial morphology and mtDNA nucleoid distribution were evaluated at the indicated time points by staining with DiOC6 (3,3'
dihexyloxacarbocyanine) (Molecular Probes Inc.) (Hermann et al., 1997
)
or DAPI (Pringle et al., 1991
), respectively. JSY2270 (FZO1 rho+ + pGAL1-N-3XMYC-FZO1), containing the wild-type FZO1 gene, did not
exhibit any changes in mitochondrial morphology or loss of mtDNA during the N-3XMYC-Fzo1p depletion (data not shown). N-3XMYC-Fzo1 protein levels in total cell extracts were analyzed by Western blotting
(anti-MYC antibody) at the indicated time points (Harlow and Lane,
1988
). Blots were stripped and reprobed with anti-3-PGK (3-phosphoglycerate kinase) (1:1,000 dilution) (Molecular Probes Inc.) to control for
differences in protein loading. Mitochondrial morphologies were scored
by GFP (green fluorescent protein) staining in JSY2273 cells containing
the pRS416-ADH-COXIVpre-GFP plasmid (JSY2634; mito-GFP).
Generation and Characterization of the fzo1-1 Mutation
The fzo1-1 temperature-sensitive allele was generated by low-fidelity
PCR (Muhlrad et al., 1992). Mutagenized pRS414-FZO1 plasmids were
transformed into a strain in which fzo1
::HIS3 was covered with pRS416-
FZO1 (JSY2287). The loss of pRS416-FZO1 from these cells was selected
by growth on medium containing 5-FOA. Cells containing the pRS414-
FZO1 mutagenized plasmids were tested for growth at 25° and 37°C on
SGlycerol medium. One strain that was inviable at 37°C was identified
(fzo1-1). The plasmid containing the mutant version of FZO1 was recovered, and when retransformed into a cell lacking FZO1 caused temperature-sensitive growth on SGlycerol medium. Transformation of the
pRS414-fzo1-1 plasmid into a wild-type strain revealed that the fzo1-1
temperature-sensitive growth defect was recessive. Mitochondrial morphology was examined in fzo1-1 cells by staining with DiOC6 (not shown)
or mito-GFP.
To examine mitochondrial fusion during mating, fzo1 + pRS414-
fzo1-1 cells labeled with either mito-GFP (JSY2802) or Mitotracker red
(JSY2804) (Molecular Probes Inc.), were mated at 25° or 37°C and analyzed as described by Nunnari et al. (1997)
.
Construction and Analysis of MYC-tagged Fzo1p
A 3XMYC epitope was introduced immediately downstream of the initiating Met in FZO1 (JSY2028) as described by Schneider et al. (1995). To
add additional MYC epitope tags, the N-3XMYC-FZO1 coding region
plus 500 bp of 5' and 3' flanking sequence was amplified from JSY2028 using P297 and P298 and cloned into the EcoRI site of pRS426 generating
pRS426-N-3XMYC-FZO1 (verified by sequence analysis). N-3XMYC-
FZO1 was subcloned into the EcoRI site of pALTER-1 (lacking a NotI
site) and digested with NotI to release the 3XMYC tag. NotI fragments
containing 3XMYC epitopes from the plasmid pMPY-3XMYC (Schneider et al., 1995
) were inserted into the amino-terminal NotI site of FZO1.
PCR screening and sequence analysis was used to identify a clone containing
a 9XMYC insert (pALTER-1-N-9XMYC-FZO1). The N-9X-MYC-FZO1
was cloned into the EcoRI site of pRS414 to generate pRS414-N-9XMYC-
FZO1. fzo1
cells expressing only the 3XMYC-tagged (pRS426-N-3XMYC-
FZO1; JSY2028) or the 9XMYC-tagged (pRS414-N-9XMYC-FZO1; JSY2394)
Fzo1 protein grew normally on nonfermentable carbon sources, retained
mtDNA, and had wild-type mitochondrial morphology, indicating that the fusion proteins were functional. All MYC-tagged forms of Fzo1p were detected by Western blotting of total protein extracts using an anti-MYC
mouse monoclonal antibody (9E10; 1:1,000) (Berkeley Antibody Co.,
Richmond, CA) (Hermann et al., 1997
). The N-9XMYC-Fzo1p was localized in JSY2392 (fzo1
+ pRS414-FZO1) and JSY2394 by indirect immunofluorescence with a primary anti-MYC antibody (1:100 dilution) and a
goat anti-mouse FITC secondary antibody (1:100 dilution) (Jackson ImmunoResearch) (Pringle et al., 1991
).
Biochemical Analysis of N-9XMYC-Fzo1p
A strain expressing N-9XMYC-Fzo1p (JSY2394) was grown in SGalactose medium lacking tryptophan to select for the pRS414-N-9XMYC- FZO1 plasmid. Cell lysates (cytosol) were fractionated by differential sedimentation to generated a mitochondrial pellet and a postmitochondrial supernatant (Daum et al., 1982; Zinser and Daum, 1995
). Samples (cell
equivalents) from each fraction were analyzed by Western blotting. To determine the membrane association of N-9XMYC-Fzo1p, 100 µg of mitochondria purified from JSY2394 were pelleted at 12,000 g for 10 min, resuspended in 75µl of 100 mM Na2CO3, pH 11.5, or 1% Triton X-100
containing 1 mM PMSF, 1 µg/ml aprotinin, and 1 µg/ml leupeptin, incubated on ice for 30 min, and then sedimented at 100,000 g for 30 min.
Equal volumes of pellet and supernatant fractions were analyzed by Western blotting. Mitochondrial inner and outer membranes (Daum et al.,
1982
) were loaded onto 5 ml of 0.85-1.6 M sucrose step gradients (0.8, 1.1, 1.35, and 1.6 M) and then centrifuged for 16 h at 30,000 rpm in a Beckman
SW50.1 rotor at 2°C (Fullerton, CA). The gradient was partitioned into 13 individual 400-µl fractions that were analyzed by Western blotting. Comparison of protein levels was performed using NIH Image 1.60 (National
Institutes of Health, Bethesda, MD). The orientation of N-9XMYC-Fzo1p
on mitochondria was determined by incubating 100 µg of purified mitochondria (from JSY2394) in breaking buffer (0.6 M mannitol, 20 mM
Hepes-KOH, pH 7.4) containing 100 µg/ml trypsin (Sigma Chemical Co.,
St. Louis, MO) on ice. To disrupt the outer membrane, mitochondria were
diluted with nine volumes of OS buffer (20 mM Hepes-KOH, pH 7.4) and
trypsin was added to a final concentration of 100 µg/ml. After 20 min, the
reaction was stopped by the addition of soybean trypsin inhibitor (2.5 mg/
ml, Sigma Chemical Co.) and 1 mM PMSF. Samples were analyzed by
Western blotting. Western blots were performed with the indicated antibodies at the following dilutions: anti-MYC (1:1,000), anti-porin (1:1,000),
anti-3-PGK (1:1,000), anti-CoxIV (cytochrome oxidase subunit IV;
1:20,000; Molecular Probes Inc.), and anti-cytochrome b2 (1:10,000).
Analysis of FZO1 GTPase Point Mutants
pRS414, pRS414-FZO1, and pRS414 containing the FZO1 GTPase point
mutations were shuffled into JSY2287 (fzo1 rho+ + pRS416-FZO1), to
generate the following strains: (a) JSY2354 (fzo1
+ pRS414); (b)
JSY2392 (fzo1
+ pRS414-FZO1); (c) JSY2355 (fzo1
+ pRS414-
fzo1[T221A]); (d) JSY2356 (fzo1
+ pRS414-fzo1[K371A]); (e) JSY2357
(fzo1
+ pRS414-fzo1[K200A]); and (f) JSY2358 (fzo1
+ pRS414-
fzo1[S201N]). None of the mutant Fzo1 proteins caused defects in mitochondrial morphology (anti-porin staining), mtDNA nucleoid retention
(DAPI staining) or function (growth on glycerol) when introduced on low
or high copy plasmids into a wild-type FZO1 strain (JSY2288) (data not
shown). Subcellular fractionation (see above) and Western blotting with
an anti-Fzo1p antibody confirmed that the mutant proteins were expressed and localized to mitochondrial membranes in fzo1
cells (data
not shown).
Microscopic Techniques
Cells were viewed on a Zeiss Axioplan microscope (1.25× optivar setting;
Carl Zeiss Inc., Thornwood, NY) as described in Roeder et al. (1998). Images were captured using a Hamamatsu C5810 color-chilled 3CCD camera (Hamamatsu Photonics, Hamamatsu City, Japan) interfaced to a Macintosh Quadra 840AV computer. For three-dimensional fluorescence microscopy, data collection was carried out using a Leica confocal microscope and a 100× 1.4 N.A. objective (Leica Inc., St. Gallen, Switzerland).
All shutters, stage motion, and image acquisition were under computer
control. Images were acquired by moving the stage in ~0.2-µm intervals.
Thin sections were viewed with a Hitachi H-700 electron microscope (Tokyo, Japan) and images were captured using the Kodak 161 digital camera
system v1.55b (Eastman Kodak Co., Rochester, NY). Digital images were
assembled into figures and printed as described in Roeder and Shaw
(1996)
.
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Results |
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S. cerevisiae FZO1 Is Required for Maintenance of Mitochondrial Morphology and Retention of Mitochondrial DNA
To determine whether Fzo family members act generally
to control mitochondrial fusion, we disrupted one copy of
S. cerevisiae FZO1 in a diploid strain. After sporulation
and dissection, fzo1 haploid cells exhibited a significant
growth defect relative to wild type on medium containing
the fermentable carbon source dextrose and were unable
to grow on medium containing the nonfermentable carbon
source glycerol (Fig. 1 B). This growth pattern, also referred to as a "petite" phenotype (Dujon, 1981
), is characteristic of strains with defective mitochondrial respiration
and indicates that the loss of FZO1 disrupts normal mitochondrial function.
Indirect immunofluorescence revealed that Fzo1p is required for normal organization of the mitochondrial network in vegetatively growing cells (Fig. 2). In a wild-type
strain, porin-stained mitochondrial membranes appeared
as branched, tubular networks distributed at the cell surface (Fig. 2, A and B). In contrast, less than 1% of fzo1
cells contained a wild-type mitochondrial network. Instead, fzo1
mutants contained between one and five
spherical or slightly elongated mitochondrial structures
that were localized to the cell cortex but were not distributed evenly around the cell periphery (Fig. 2, G and H). In
addition, numerous small mitochondria were occasionally
observed scattered throughout the cytoplasm in the mutant (data not shown). The mitochondrial morphology in
fzo1
cells resembles that in mutants with abnormal actin
cytoskeletons (Drubin et al., 1993
; Lazzarino et al., 1994
).
However, we did not observe defects in the organization
of the actin and microtubule cytoskeletons in the fzo1
strain (data not shown). Although many of the yeast mitochondrial morphology mutants identified to date also have
an associated mitochondrial inheritance defect (Burgess et
al., 1994
; Sogo and Yaffe, 1994
; Berger et al., 1997
; Hermann and Shaw, 1998
), our analysis did not reveal a mitochondrial inheritance defect in fzo1
cells relative to wild
type (data not shown). Thus, Fzo1p controls mitochondrial network organization in yeast but is not required for
mitochondrial transport into daughter cells during division.
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Transmission electron microscopy indicated that mitochondrial morphology and distribution were also abnormal at the ultrastructural level in fzo1. In wild-type cells,
mitochondrial profiles were distributed throughout the peripheral cytoplasm and contained elaborate cristae (invaginations of the inner mitochondrial membrane) (Fig. 2 Y).
In contrast, mitochondrial profiles in fzo1
cells appeared
to cluster together in one or two regions near the plasma membrane (Fig. 2, Z and A'), similar to the mitochondrial
distribution observed by indirect immunofluorescence
(Fig. 2, G and H). The clustered mitochondrial profiles in
fzo1
could represent closely opposed tubules of a collapsed, but still interconnected, mitochondrial reticulum.
Alternatively, the clusters could be composed of individual unfused mitochondrial fragments (see below). Cross
sections also revealed that fzo1
mitochondria often contained fewer cristae or lacked cristae altogether (Fig. 2, Z
and A').
The wild-type function of Fzo1p is also required for the
maintenance of mtDNA (mitochondrial DNA) nucleoids.
Wild-type cells labeled with the DNA-specific dye DAPI
always contained brightly stained nuclei as well as 25-50
punctate mtDNA nucleoids localized at the cell periphery
(Fig. 2, C and D). In contrast, mitochondrial nucleoids
were never detected in fzo1 cells (Fig. 2, I and J). Crosses
of fzo1
with a known rhoo strain (lacking mtDNA) confirmed that mitochondrial genomes were absent in fzo1
cells (data not shown). Since mtDNA encodes RNAs and
proteins essential for mitochondrial respiratory function (Pon and Schatz, 1991
), these results raised the possibility
that fzo1
mitochondrial morphology defects were an indirect consequence of mtDNA loss. However, reintroduction of the wild-type FZO1 gene was sufficient to restore
elongated and branched mitochondrial networks in fzo1
cells (Fig. 2, M and N) in the absence of mtDNA (Fig. 2, O
and P). The morphology of the restored mitochondrial networks was identical to that observed in an isogenic rhoo
strain containing a wild-type FZO1 gene (Fig. 2, S-V).
These results demonstrate that the loss of mtDNA, and
the resulting defects in mitochondrial respiration, do not
cause mitochondrial morphology defects in wild-type cells
(Guan et al., 1993
) or in fzo1
mutants and suggest that
Fzo1p is directly required for the maintenance of normal
mitochondrial morphology.
Fzo1p depletion studies revealed that changes in mitochondrial morphology preceded the loss of mtDNA nucleoids. A GAL1 regulated, N-3XMYC-tagged form of Fzo1p
on a plasmid was introduced into the fzo1 mutant strain
using a plasmid shuffling technique to prevent mtDNA
loss (refer to Materials and Methods). Growth in raffinose-containing medium (which neither induces nor represses transcription from the GAL1 promoter) resulted
in sufficient expression of N-3XMYC-Fzo1p to fully support wild-type growth and restore wild-type mitochondrial
morphology in fzo1
cells (Fig. 3, A, solid squares, t = 0, and D, Wildtype). After transfer to glucose-containing medium to block expression of N-3XMYC-Fzo1p from the
GAL1 promoter, aliquots were harvested at the indicated
times and mitochondrial morphology (DiOC6 staining)
and mtDNA distribution (DAPI staining) were examined.
During the 8 h after transfer, wild-type mitochondrial networks broke down into smaller segments which eventually
collapsed, forming between one and five spherical or
slightly elongated membrane clusters per cell (Fig. 3, A
and D). The number, shape, and size of these mitochondrial clusters at the 8-h time point were identical to those
visualized in the fzo1
mutant by anti-porin staining (Fig.
2, G and H). The defects in mitochondrial morphology observed 8 h after the shift did not severely affect mitochondrial function, since these misshapen organelles were still
able to accumulated the membrane potential-sensitive dye
DiOC6 (Chen, 1988
). Western blotting with an anti-MYC
antiserum showed a 10-fold reduction in N-3XMYC-Fzo1
protein levels during the course of the depletion experiment (Fig. 3 C). Cells lacking DAPI-stained mtDNA nucleoids were not detected, even after 8 h when the majority of cells exhibited defective mitochondrial morphology
(Fig. 3 B). Moreover, 24 h after the initial N-3XMYC-Fzo1p expression block (11 doublings), only 25% of the
cells lacked nucleoids, making it unlikely that Fzo1p is required for mtDNA partitioning into buds (data not
shown). Together, these results indicate that the loss of
Fzo1p leads first to defects in mitochondrial membrane
morphology and only later to defects in mtDNA maintenance.
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Mitochondrial Networks Fragment at 37°C in a Conditional fzo1-1 Mutant
Mitochondrial morphology is likely to be regulated, in
part, by opposing membrane fusion and membrane fission
reactions (Nunnari et al., 1997). If the primary effect of an
fzo1 mutation is to block membrane fusion, we predicted
that ongoing mitochondrial fission would initially cause
the mitochondrial network to fragment. To test this, a conditional FZO1 allele (fzo1-1) was identified by replacing a
wild-type FZO1 plasmid with mutagenized FZO1 plasmids in an fzo1
strain (refer to Materials and Methods).
fzo1-1 allowed growth of the fzo1
strain at 25° but not
37°C on SGlycerol medium (Fig. 3 E). The fzo1-1 sequence contained three different nucleotide changes, resulting in amino acid substitutions K538I, N543I, and P553Q in the predicted polypeptide (refer to Materials
and Methods). Staining with DiOC6 (data not shown) or
GFP (Fig. 3, F and G) demonstrated that mitochondrial
network morphology was wild type in the majority (76%)
of fzo1-1 cells grown at the permissive temperature (Fig. 3,
F, fzo1-1, 25°C, and G, t = 0 min). 10 min after shifting to
37°C, however, the networks in 99% of the fzo1-1 cells had
fragmented into many small, uniformly distributed, mitochondrial compartments (Fig. 3, F, fzo1-1, 37°C, and G,
t = 10 min). Upon extended incubation at 37°C, these mitochondrial fragments clustered together forming large
membrane aggregates similar to those observed in the
fzo1
strain (data not shown). If after 20 min at 37°C the
temperature was reduced to 25°C, the mitochondrial network regained its wild-type morphology within 60 min
(Fig. 3 G). In control experiments, mitochondrial fragmentation was never observed in wild-type strains grown at 25°
or 37°C (Fig. 3 F, FZO1, 25°C and 37°C). The rapid fragmentation of the mitochondrial network observed in the
conditional fzo1-1 strain provides a direct demonstration that Fzo1p controls mitochondrial morphology in yeast
and is consistent with a role for this protein in mitochondrial membrane fusion.
fzo1-1 Blocks Mitochondrial Fusion during Mating
Mitochondrial fusion and content mixing has been shown
to occur in yeast zygotes soon after cell fusion (Azpiroz
and Butow, 1993; Nunnari et al., 1997
). To test directly the
requirement of Fzo1p in mitochondrial fusion, we examined the effect of the fzo1-1 mutation on mitochondrial
content mixing during mating. Mitochondrial networks
were visualized by labeling one haploid parent with the
fluorescent vital dye Mitotracker red and the other haploid parent with a matrix targeted form of GFP (mito-GFP) (Nunnari et al., 1997
). After mating and zygote formation, the distribution of the fluorophores was examined
by fluorescence confocal microscopy. In matings between
wild-type cells performed at 25° or 37°C, the two fluorescent markers rapidly and completely colocalized in zygotes, indicating that the parental mitochondrial membranes had fused and mitochondrial contents had mixed
(Nunnari et al., 1997
) (Table II). Mitochondrial fusion also
occurred efficiently in fzo1-1 × fzo1-1 zygotes formed at
25°C (Fig. 4, A-D; Table II). In contrast, mitochondrial
networks fragmented and failed to fuse in fzo1-1 × fzo1-1
zygotes formed at 37°C (Fig. 4, E-L; Table II). When fzo1-1
zygotes were optically sectioned using the confocal
microscope, mitochondrial membranes containing the haploid-derived green and red fluorescent markers never
colocalized (Fig. 4, E-L). Moreover, fzo1-1 × fzo1-1 zygotes formed at the restrictive temperature contained unfused mitochondria even after they had completed karyogamy and formed a new bud (Fig. 4, I-L). Thus, fzo1-1
causes a block, and not simply a delay, in mitochondrial docking and/or fusion. Finally, mitochondrial fusion was
also reduced in matings between fzo1-1 and wild-type parents at 37°C (Table II), suggesting that the function of
Fzo1p is required in both haploid parents for efficient mitochondrial fusion.
|
|
We think it is unlikely that the fzo1-1 mutation prevents mitochondrial fusion indirectly by interfering with mitochondrial motility and/or distribution. First, although mitochondrial networks fragment at 37°C in fzo1-1 × fzo1-1 zygotes, these membrane fragments were segregated normally into daughter cells, indicating that mitochondrial motility was not severely impaired (Fig. 4, I-L, arrow). Second, the fragmentation of mitochondria in fzo1-1 × fzo1-1 zygotes does not appear to block their association. We often observed red and green mitochondrial compartments in close apposition near the zygote neck or in the diploid daughter cell (Fig. 4, H and L, arrows). Together, these results provide compelling evidence that Fzo1p plays an essential and direct role in mitochondrial fusion.
Fzo1p Is a Mitochondrial Outer Membrane Protein with Its GTPase Domain Facing the Cytoplasm
The Fzo1 protein is localized on the mitochondrial network in vegetatively growing yeast cells (Fig. 5). A plasmid encoding Fzo1p tagged near the amino terminus with
nine MYC epitopes (N-9XMYC-Fzo1p) was constructed
and shown to rescue the mitochondrial morphology defect, the mtDNA loss phenotype, and the glycerol growth
defect in the fzo1 strain (data not shown). Anti-MYC antibodies recognized a 116-kD protein in extracts prepared
from wild-type cells expressing the N-9XMYC-Fzo1p (Fig.
6 A, lane 2) but not the native Fzo1 protein (Fig. 6 A, lane
1). Indirect immunofluorescence with anti-MYC antibodies revealed that N-9XMYC-Fzo1p localized to the mitochondrial network in wild-type cells and was uniformly
distributed on this compartment (Fig. 5, A-D). The pattern of DAPI-stained mtDNA nucleoids in these cells
overlapped with the fluorescent signal confirming that
N-9XMYC-Fzo1p was located on the mitochondrial network (Fig. 5, B and D; compare white arrows in A and C
with B and C). Similar results were obtained when cells
were stained with polyclonal antibodies generated against
the native Fzo1 protein (data not shown). In control experiments, no signal was detected in cells expressing only
the wild-type Fzo1p (Fig. 5 E).
|
|
Subcellular fractionation confirmed the mitochondrial localization of Fzo1p (Fig. 6 B). N-9XMYC-Fzo1p cofractionated with the mitochondrial pellet, along with the outer mitochondrial membrane protein porin, during differential centrifugation of a postnuclear cell extract (Fig. 6 B, Mito). No N-9XMYC-Fzo1p was detected in the postmitochondrial supernatant fraction that contained the cytoplasmic protein 3-PGK (Fig. 6 B, PMS). Similar results were obtained from wild-type cells using anti-Fzo1p antiserum to follow fractionation of the native Fzo1 protein (data not shown).
Fzo1p behaved like an integral membrane protein, as
predicted based on the two closely spaced and conserved
hydrophobic domains near its carboxy terminus (refer to
Fig. 1 A) (Hales and Fuller, 1997). When mitochondria
containing N-9XMYC-Fzo1p were extracted with 100 mM
Na2CO3, pH 11.5, to release peripheral membrane proteins, both the N-9XMYC-Fzo1p and the integral outer-membrane protein porin were resistant to sodium carbonate extraction and remained associated with the membrane
pellet (Fig. 6 C). In contrast, Fzo1p and porin were released into the supernatant when mitochondrial membranes were solubilized with 1% Triton X-100 (Fig. 6 C).
Fzo1p also remained associated with mitochondrial membranes after the organelles were disrupted by osmotic and
mechanical methods to release soluble intermembrane
space and matrix proteins (Fig. 6 D and data not shown).
Analysis of submitochondrial membrane fractions separated on sucrose density gradients indicated that Fzo1p associates with both the inner and outer mitochondrial
membranes. N-9XMYC-Fzo1p fractionated at an intermediate density that overlapped with, but was distinct from,
the distribution of both inner membrane vesicles containing the integral membrane protein CoxIV and outer membrane vesicles containing porin (Fig. 6, D and E). Although this result does not prove that Fzo1p crosses both
mitochondrial membranes, this fractionation pattern is
characteristic of "trapped" translocation intermediates
that span both mitochondrial membranes at contact sites
formed by the translocation pore (Pon et al., 1989). Proteins that fractionate with both inner and outer mitochondrial membranes and are enriched at contact sites have
been observed previously (Pon et al., 1989
).
Fzo1p is oriented with its amino-terminal GTPase domain and adjacent heptad repeats exposed on the cytoplasmic face of the mitochondrial compartment. Treatment of isolated, intact mitochondria with trypsin resulted in the complete digestion of the MYC tag on the amino terminus of Fzo1p (Fig. 6 F, lane 3). In contrast, cytochrome b2, a protein in the intermembrane space, was resistant to proteolysis, indicating that the outer mitochondrial membrane remained intact (Fig. 6 F, lane 3). Cytochrome b2 could be digested, however, if the mitochondrial outer membrane was disrupted by osmotic shock (Fig. 6 F, lane 4). Since Fzo1p behaves like an integral membrane protein, these data also indicate that at least one of the hydrophobic domains at the carboxy-terminus of Fzo1p is embedded in the mitochondrial outer membrane.
The signals and machinery that target Fzo proteins to
mitochondria may be conserved. When a cDNA encoding
the Drosophila melanogaster Fzo protein was introduced
into wild-type cells on a GAL1 low copy plasmid, the
Drosophila protein cofractionated with mitochondrial membranes as assayed by Western blotting with antibodies specific for the D. melanogaster homologue (data not
shown). However, the Drosophila Fzo protein did not rescue mitochondrial morphology or mtDNA loss phenotypes in the S. cerevisiae fzo1 strain.
Fzo1p Function Requires the Conserved GTPase Domain
The Fzo1p GTPase domain contains four conserved motifs designated G1-G4 (Fig. 7 A). In most GTPases, these
domains are required for GTP binding and hydrolysis as
well as conformational changes elicited by nucleotide
binding (Bourne et al., 1991). Conserved residues in three
of the four G motifs in Fzo1p (K200A and S201N in G1,
T221A in G2, and K371A in G4) were altered by site-directed mutagenesis (Fig. 7 A). All of these amino acid
substitutions are known to disrupt either nucleotide binding or interactions with effector proteins in other GTPases
(Sigal et al., 1986
; Adari et al., 1988
; Cales et al., 1988
; Feig
and Cooper, 1988
; Farnsworth and Feig, 1991
; Vojtek et
al., 1993
; Murphy et al., 1997
). When low copy plasmids
containing the mutated fzo1 genes were introduced into
fzo1
cells, the mutant Fzo1 proteins were expressed at
wild-type levels and were targeted to the mitochondrial
compartment as assayed by differential centrifugation and
Western blotting with anti-Fzo1p antiserum (data not
shown). However, the fzo1(K200A), fzo1(S201N), and
fzo1(T221A) mutant genes failed to rescue the glycerol
growth defect or the mitochondrial morphology defects in
the fzo1
strain (Fig. 7 B). Interestingly, a significant percentage (13 and 12%, respectively) of cells containing the
fzo1(K200A) and fzo1(T221A) mutant genes contained
detectable mtDNA nucleoids (Fig. 7 B), although the total
number of nucleoids was reduced relative to wild type (1-5
instead of 25-50) (data not shown). It is possible that these
mutant Fzo1 proteins retain some residual functions required for mtDNA maintenance. Alternatively, cells containing these mutant proteins may simply lose their mitochondrial genomes more slowly than fzo1-null cells.
Mutation of a conserved residue in the G4 domain
(K371A) did not disrupt the function of FZO1 (Fig. 7 B).
This result is somewhat surprising, since lysine 371 is conserved throughout the GTPase superfamily and is known
to be required for high-affinity GTP binding by Ras (Der
et al., 1988
; Bourne et al., 1991
). In addition, the same mutation in Drosophila fzo had a modest but significant effect on the function of the protein (Hales and Fuller,
1997
). Finally, wild-type cells overexpressing the mutant
Fzo1 proteins (10-20 fold from a 2-µ vector) did not exhibit dominant growth or mitochondrial morphology defects when compared with cells overexpressing the wild-type Fzo1p (10-20 fold from a 2-µ vector) (data not
shown). These observations indicate that GTP binding
and/or hydrolysis is essential for Fzo1p function.
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Discussion |
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Fzo1p Regulates Mitochondrial Fusion
We have shown that Fzo1p is a transmembrane GTPase
required for mitochondrial fusion in yeast. Mitochondrial
membranes rapidly fragment at the nonpermissive temperature in a conditional fzo1-1 strain. Since the opposing
processes of mitochondrial fusion and fission are responsible for the reticular structure of the mitochondrial network in mitotically dividing cells (Nunnari et al., 1997), the
simplest explanation for this result is that fzo1-1 causes a
selective block in fusion and that fragmentation occurs as
a result of continuing mitochondrial fission. fzo1-1 is the
only yeast mitochondrial morphology mutant with a fragmentation phenotype, consistent with its novel role in mitochondrial fusion. Mitochondrial fusion and content mixing were also blocked in homozygous fzo1-1 × fzo1-1
zygotes and reduced in heterozygous fzo1-1 × FZO1+
matings. Functional Fzo1 protein could be required on opposing mitochondrial membranes for efficient fusion. If
this is the case, then the residual mitochondrial fusion observed in fzo1-1 × FZO1+ matings could result from new
protein synthesis and complementation in the heterozygous zygote. Alternatively, functional Fzo1 protein on
only one of the fusion partners may allow fusion between
Fzo1p+ and Fzo1p
mitochondrial membranes at lower efficiency.
Disruption of the FZO1 gene, depletion of the Fzo1 protein, or prolonged incubation of the fzo1-1 strain at 37°C
lead to the severe clumping and aggregation of mitochondrial membranes. It seems likely that a constitutive block
in fusion is responsible for the dramatic defects in mitochondrial morphology we observed under all of these conditions. Although mitochondrial inheritance is often defective in yeast mutants with abnormal mitochondrial morphology (Burgess et al., 1994; Sogo and Yaffe, 1994
;
Berger et al., 1997
; Hermann, 1998
), the loss of Fzo1p
function did not affect the motility or transport of mitochondrial membranes during mitotic division and the abnormal mitochondrial compartments in fzo1-1 and fzo1
cells were efficiently transmitted to daughter buds (refer
to Fig. 2 G). Defects in Fzo1p function also lead to the loss
of mitochondrial genomes. Although we cannot rule out
that Fzo1p participates directly in mtDNA replication
and/or segregation, several lines of evidence suggest that
the loss of mtDNA in fzo1
cells is a secondary consequence of changes in mitochondrial morphology. First,
when Fzo1p is depleted from wild-type cells, defects in mitochondrial membrane structure are observed many hours
before DAPI-stained mtDNA nucleoids are lost. Second, a number of studies indicate that mutations affecting mitochondrial morphology result in decreased mtDNA stability (Guan et al., 1993
; Burgess et al., 1994
; Sogo and Yaffe,
1994
; Berger et al., 1997
; Hermann, 1998
). Further experiments are required to determine the mechanism by which
mtDNA is lost from fzo1 mutant strains.
Fzo1 Mitochondrial Membrane Association and Topology
Immunolocalization, fractionation, and protease digestion
studies indicated that Fzo1p is an integral membrane protein with its amino terminus displayed on the mitochondrial surface. This topology positions Fzo1p's GTPase domain and adjacent heptad repeats in the cytoplasm where
they could interact with binding partners and/or regulatory molecules that might regulate Fzo1p function. Given
the conservation of the domain structure and overall
charge distribution of the different Fzo family members
(Fig. 1 A; Hales and Fuller, 1997), we predict that Fzo homologues from other organisms will display a similar mitochondrial distribution and topology.
Mitochondrial fusion has been reported to initiate at
stable contact sites between the inner and outer mitochondrial membranes (Bereiter-Hahn and Voth, 1994). The
carboxy terminus of Fzo proteins could play an important
role in coordinating the behavior of the two lipid bilayers
at these sites during the fusion reaction. Protease digestion
and protein solubilization studies suggested that at least
one of Fzo1p's carboxy-terminal hydrophobic domains is
embedded in the outer mitochondrial membrane. In addition, Fzo1p migrated in sucrose gradients with an intermediate density fraction that partially overlapped both inner
and outer mitochondrial membrane markers. This fractionation pattern has been observed for proteins that are
physically associated with mitochondrial contact sites (Pon
et al., 1989
), suggesting that Fzo1p is located in these structures. There are a number of topologies that could account
for the ability of Fzo1p to fractionate with both mitochondrial membranes. It is possible that Fzo1p spans both
membranes with its carboxy terminus in the matrix as originally proposed by Hales and Fuller (1997)
. Alternatively,
the carboxy-terminal tail of Fzo1p could extend into the
intermembrane space and interact with proteins in the inner membrane. Finally, Fzo1p might be exclusively associated with the outer membrane at contact sites formed by other proteins. Localization of the carboxy terminus of
Fzo1p will help to distinguish between these models.
Although the Drosophila and yeast Fzo proteins are
both required for mitochondrial fusion, several observations suggest that they may be regulated differently. First,
our studies indicate that the Drosophila Fzo protein is efficiently targeted to mitochondrial membranes in wild-type
yeast but cannot rescue mitochondrial phenotypes in the
fzo1 strain (data not shown). Second, the Drosophila Fzo and yeast Fzo1 proteins exhibit distinct expression patterns in the two organisms. Drosophila Fzo was only detected on sperm mitochondria during a short period of
time when mitochondrial fusion was occurring (Hales and
Fuller, 1997
), suggesting that the timing of mitochondrial fusion could be developmentally regulated by controlling
Fzo expression, localization, and/or degradation. In contrast, yeast Fzo1p protein levels and mitochondrial localization did not change during mitotic growth, mating, or
meiosis (data not shown), consistent with the observation
that mitochondrial fusion occurs during all stages of the
yeast life cycle (Pon and Schatz, 1991
; Nunnari et al., 1997
;
Hermann and Shaw, 1998
). In time-lapse studies, yeast mitochondrial fusion is observed when the tip of a mitochondrial tubule encounters the tip or side of another mitochondrial tubule (Nunnari et al., 1997
). This has led to the
suggestion that key fusion components are localized or
specifically activated at the tips of mitochondrial tubules
(Nunnari et al., 1997
). Our observation that Fzo1p is uniformly distributed on the mitochondrial compartment suggests that the fusion machinery is not localized at tips.
Instead, Fzo1p could be activated locally at sites of membrane contact.
Fzo1 Function Requires the Amino-terminal GTPase Domain
The GTPase domain of Fzo1p is essential for its function and could act as a molecular switch to regulate mitochondrial docking and/or fusion. The fzo1(K200A) and
fzo1(S201N) mutations in the G1 motif are equivalent to
mutations in the Ras GTPase that reduce guanine nucleotide binding (Sigal et al., 1986; Feig and Cooper, 1988
;
Farnsworth and Feig, 1991
). The fzo1(T221A) substitution in the G2 motif is based on a Ras mutation which eliminates interactions with effector molecules (Adari et al.,
1988
; Cales et al., 1988
; Vojtek et al., 1993
; Murphy et al.,
1997
). None of these mutations disrupted Fzo1p localization suggesting that GTP binding and/or hydrolysis is not
required to target Fzo1p to mitochondria.
In contrast, the fzo1(K371A) mutation in the G4 motif
retained wild-type function and was able to rescue defects
in mitochondrial fusion, mtDNA maintenance, and glycerol growth in fzo1 mutant cells. This lysine is highly conserved among the superfamily of GTPases (Bourne et al.,
1991) and is required for efficient nucleotide binding and
stablilization of the GTP binding pocket in Ras (Lys117) (Der et al., 1988
; Pai et al., 1990
). Although mutating
Lys117 in Ras significantly reduces its affinity for GTP, it
does not disrupt its oncogenic potential (Clanton et al.,
1986
; Der et al., 1988
). In addition, the analogous mutation
in the G4 motif of Drosophila Fzo did not completely disrupt mitochondrial fusion (Hales and Fuller, 1997
). These
results suggest that the yeast and fly Fzo proteins may
have a high intrinsic affinity for GTP that is not completely compromised by alterations in the G4 lysine. Alternatively, this lysine may be completely (yeast Fzo1p) or partially (fly Fzo) dispensable with respect to nucleotide
binding.
Mutant forms of the yeast dynamin-like proteins Vps1p
and Dnm1p (Vater et al., 1992; Otsuga et al., 1998
), mammalian dynamin (Herskovits et al., 1993
; van der Bliek et
al., 1993
), and Ras (Sigal et al., 1986
; Feig and Cooper,
1988
; Farnsworth and Feig, 1991
) induce dominant-interfering phenotypes when overexpressed in wild-type cells.
These dominant phenotypes are thought to result because
the mutant forms of the proteins either titrate out or block
the activities of binding partners required for their function. In contrast, none of the disabled forms of Fzo1p we
tested induced dominant mitochondrial phenotypes in
wild-type cells (data not shown). The simplest interpretation of these results is that Fzo1p acts alone or that mutations in the GTPase domain of Fzo1p completely disrupt
its ability to interact with itself or other proteins.
Models for the Mechanism of Fzo1p Action
Given what is known regarding the role of other integral
membrane proteins in fusion, we propose that Fzo GTPases act as molecular switches to directly mediate mitochondrial docking and/or membrane fusion. Both the domain structure of the Fzo family members and the
topology we have determined for Fzo1p are consistent
with this model. Fzo molecules share structural features
with two classes of integral membrane proteins that mediate membrane fusion events. The SNAREs regulate membrane docking and fusion during vesicle transport and during the homotypic fusion of organelle membranes (Denesvre
and Malhotra, 1996; Hay and Scheller, 1997
; Edwardson, 1998
; Götte and Fischer von Mollard, 1998
; Weber et al.,
1998
). The viral-type fusion proteins regulate extracellular
membrane docking and fusion events (Hernandez et al.,
1996
; Huovila et al., 1996
). Both SNAREs and viral fusion
proteins contain multiple heptad repeats in their amino-terminal domains. These repeats form parallel coiled coils
that are proposed to mediate the direct association of v-
and t-SNAREs on opposing membranes and the oligomerization of viral fusion proteins within a membrane (Kee et
al., 1995
; Hernandez et al., 1996
; Hanson et al., 1997
; Hay
and Scheller, 1997
; Hughson, 1997
). Our studies indicate
that the two amino-terminal heptad repeats in Fzo1p extend into the cytoplasm. If Fzo1p serves a SNARE-like
function, these repeats could mediate the direct or indirect
association of Fzo1p with itself or with another protein binding partner on an opposing mitochondrial membrane.
The cytoplasmic GTPase domain of Fzo1p might function
to control the rate or fidelity of such Fzo1p interactions,
similar to the manner in which Rab GTPases regulate
SNARE-SNARE associations (Lupashin and Waters, 1997
).
The idea that Fzo1p functions as a novel type of mitochondrial SNARE is attractive, given that none of the v- or
t-SNAREs encoded by the S. cerevisiae genome localize to
the mitochondrial compartment (H.R.B. Pelham, personal
communication).
Alternatively, Fzo1p may operate more like viral fusion
proteins (White, 1990, 1992
; Hernandez et al., 1996
; Hughson, 1997
; Qiao et al., 1998
). These integral membrane
proteins contain additional hydrophobic fusion peptides
that normally remain masked. When activated, the fusion
peptide inserts into the target membrane generating a
docked state. Conformational changes in the protein pull
the membranes together and promote fusion. Fzo1p contains two hydrophobic sequences in its cytoplasmic domain that resemble fusion peptides (Hermann, 1998
). It is
possible that Fzo1p uses its GTPase activity as a switch to
stimulate the insertion of these putative fusion peptides
into a neighboring mitochondrial membrane. The GTPase
domain might also regulate conformational changes in
Fzo1p that pull the two docked mitochondrial membranes together after peptide insertion. The observation that
Fzo1p is tightly associated with both mitochondrial membranes suggests that it might regulate the fusion of the inner membrane as well. Whether the predicted transmembrane domain and heptad repeat closest to the carboxy
terminus of Fzo1p are in a position to influence inner
membrane behavior remains to be seen.
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Footnotes |
---|
Received for publication 4 August 1998 and in revised form 25 August 1998.
Address all correspondence to Janet M. Shaw, Department of Biology,
University of Utah, Salt Lake City, UT 84112. Tel.: (801) 585-6205. Fax:
(801) 581-4668. E-mail: shaw{at}bioscience.utah.edu, or Jodi Nunnari, Department of Molecular and Cellular Biology, University of California,
Davis, CA 95616. Tel.: (530) 754-9774. Fax: (530) 752-7522. E-mail:
fzmito{at}peseta.ucdavis.edu
We thank R. Jensen (Johns Hopkins University, Baltimore, MD) for the mito-GFP plasmid and cytochrome b2 antibody, the staff of the Research Microscopy Facility (K.H. Albertine and N.B. Chandler) at the University of Utah Health Sciences Center (Salt Lake City, UT) for assistance with the ultrastructural studies, and Q. Tieu (University of California, Davis, CA) for technical assistance isolating the fzo1-1 allele.
This work was supported by grants from the American Cancer Society (CB-97) and the National Institutes of Health (NIH) (GM-53466) to J.M. Shaw, the National Science Foundation (MCB-9724143) to J. Nunnari, and the NIH (HD-29194) to M.T. Fuller. G.J. Hermann, K.G. Hales, and J.P. Mills were supported by NIH training grants (GM-07464, HG-00044, and GM-07790, respectively). The University of Utah Research Microscopy Facility is supported by a grant from the NIH (S10-RR-10489). The Utah Health Sciences Sequencing Facility is supported by a National Cancer Institute grant (5-P30CA42014).
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Abbreviations used in this paper |
---|
3-PGK, 3-phosphoglycerate kinase; 5-FOA, 5-fluoro-orotic acid; CoxIV, cytochrome oxidase subunit IV; DAPI, 4',6-diamidino-2-phenylindole; DiOC6, 3,3' dihexyloxacarbocyanine; fzo, fuzzy onions gene; GFP, green fluorescent protein; mtDNA, mitochondrial DNA.
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References |
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---|
1. | Adari, H., D.R. Lowy, B.M. Willumsen, C.J. Der, and F. McCormick. 1988. Guanosine triphosphatase activating protein (GAP) interacts with the p21 ras effector binding domain. Science 240: 518-521 |
2. | Azpiroz, R., and R.A. Butow. 1993. Patterns of mitochondrial sorting in yeast zygotes. Mol. Biol. Cell 4: 21-36 [Abstract]. |
3. | Bakeeva, L.E., Y.S. Chentsov, and V.P. Skulachev. 1978. Mitochondrial framework (reticulum mitochondriale) in rat diaphragm muscle. Biochim. Biophys. Acta. 501: 349-369 |
4. | Bakeeva, L.E., Y.S. Chentsov, and V.P. Skulachev. 1981. Ontogenesis of mitochondrial reticulum in rat diaphragm muscle. Eur. J. Cell Biol. 25: 175-181 |
5. | Baudin, A., O. Ozier-Kalogeropoulos, A. Denouel, F. Lacroute, and C. Cullin. 1993. A simple and efficient method for direct gene deletion in Saccharomyces cerevisiae. Nucleic Acids Res. 21: 3329-3330 |
6. | Bereiter-Hahn, J., and M. Voth. 1994. Dynamics of mitochondria in living cells: shape changes, dislocations, fusion, and fission of mitochondria. Microsc. Res. Tech. 27: 198-219 |
7. |
Berger, K.H.,
L.F. Sogo, and
M.P. Yaffe.
1997.
Mdm12p, a component required
for mitochondrial inheritance that is conserved between budding and fission
yeast.
J. Cell Biol.
136:
545-553
|
8. | Bourne, H.R., D.A. Sanders, and F. McCormick. 1991. The GTPase superfamily: conserved structure and molecular mechanism. Nature 349: 117-127 |
9. | Burgess, S.M., M. Delannoy, and R.E. Jensen. 1994. MMM1 encodes a mitochondrial outer membrane protein essential for establishing and maintaining the structure of yeast mitochondria. J. Cell Biol. 126: 1375-1391 [Abstract]. |
10. | Cales, C., J.F. Hancock, C.J. Marshall, and A. Hall. 1988. The cytoplasmic protein GAP is implicated as the target for regulation by the ras gene product. Nature 332: 548-551 |
11. | Chen, L.B.. 1988. Mitochondrial membrane potential in living cells. Annu. Rev. Cell Biol. 4: 155-181 . |
12. | Clanton, D.J., S. Hattori, and T.Y. Shih. 1986. Mutations of the ras gene product p21 that abolish guanine nucleotide binding. Proc. Natl. Acad. Sci. USA. 83: 5076-5080 [Abstract]. |
13. |
Daum, G.,
P.C. Bohni, and
G. Schatz.
1982.
Import of proteins into mitochondria.
J. Biol. Chem.
257:
13028-13033
|
14. | Denesvre, C., and V. Malhotra. 1996. Membrane fusion in organelle biogenesis. Curr. Opin. Cell Biol. 8: 519-523 |
15. | Der, C.J., B. Weissman, and M.J. MacDonald. 1988. Altered guanine nucleotide binding and H-ras transforming and differentiating activities. Oncogene 3: 105-112 . |
16. | Drubin, D.G., H.D. Jones, and K.F. Wertman. 1993. Actin structure and function: roles in mitochondrial organization and morphogenesis in budding yeast and identification of the phalloidin-binding site. Mol. Biol. Cell 4: 1277-1294 [Abstract]. |
17. | Dujon, B. 1981. Mitochondrial genetics and functions. In Molecular Biology of the Yeast Saccharomyces. J.M. Strathern, E.W. Jones, and J.R. Broach, editors. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 505-635. |
18. | Edwardson, J.M.. 1998. Membrane fusion: All done with SNAREpins? Curr. Biol. 8: R390-R393 |
19. | Farnsworth, C.L., and L.A. Feig. 1991. Dominant inhibitory mutations in the Mg2+-binding site of RasH prevent its activation by GTP. Mol. Cell Biol. 11: 4822-4829 |
20. | Feig, L.A., and G.M. Cooper. 1988. Inhibition of NIH 3T3 cell proliferation by a mutant ras protein with preferential affinity for GDP. Mol. Cell Biol. 8: 3235-3243 |
21. | Fox, T.D., L.S. Folloy, J.J. Mulero, T.W. McMullin, P.E. Thorsness, L.O. Hedin, and M.C. Costanzo. 1991. Analysis and manipulation of yeast mitochondrial genes. Methods Enzymol. 194: 149-165 |
22. | Fuller, M.T. 1993. Spermatogenesis. In The Development of Drosophila melanogaster. M. Bate and A. Martinez-Arias, editors. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 71-147. |
23. | Götte, M., and G. Fischer von Mollard. 1998. A new beat for the SNARE drum. Trends Cell Biol. 8: 215-218 . |
24. | Guan, K., L. Farh, T.K. Marshall, and R.J. Deschenes. 1993. Normal mitochondrial structure and genome maintenance in yeast requires the dynamin-like product of the MGM1 gene. Curr. Genet. 24: 141-148 |
25. | Hales, K.G., and M.T. Fuller. 1997. Developmentally regulated mitochondrial fusion mediated by a conserved, novel, predicted GTPase. Cell 90: 121-129 |
26. | Hanson, P.I., R. Roth, H. Morisaki, R. Jahn, and J.E. Heuser. 1997. Structure and conformational changes in NSF and its membrane receptor complexes visualized by quick-freeze/deep-etch electron microscopy. Cell 90: 523-535 |
27. | Harlow, E., and D. Lane, editors. 1988. Antibodies: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, New York. 726 pp. |
28. | Hay, J.C., and R.H. Scheller. 1997. SNAREs and NSF in targeted membrane fusion. Curr. Opin. Cell Biol. 9: 505-512 |
29. |
Hermann, G.J.,
E.J. King, and
J.M. Shaw.
1997.
The yeast gene, MDM20, is
necessary for mitochondrial inheritance and organization of the actin cytoskeleton.
J. Cell Biol.
137:
141-153
|
30. | Hermann, G.J. 1998. Mitochondrial inheritance and morphology in yeast. Ph.D. thesis. University of Utah, Salt Lake City, UT. 242 pp. |
31. | Hermann, G.J., and J.M. Shaw. 1998. Mitochondrial dynamics in yeast. Annu. Rev. Cell Dev. Biol. 14: 265-303 . |
32. | Hernandez, L.D., L.R. Hoffman, T.G. Wolfsberg, and J.M. White. 1996. Virus-cell and cell-cell fusion. Annu. Rev. Cell Dev. Biol. 12: 565-578 . |
33. | Herskovits, J.S., C.C. Burgess, R.A. Obar, and R.B. Vallee. 1993. Effects of mutant rat dynamin on endocytosis. J. Cell Biol. 122: 565-578 [Abstract]. |
34. | Hoffman, H., and C.J. Avers. 1973. Mitochondrion of yeast: ultrastructural evidence for one giant, branched organelle per cell. Science 181: 749-751 |
35. | Hughson, F.M.. 1997. Enveloped viruses: a common mode of membrane fusion? Curr. Biol. 7: R565-R569 |
36. | Huovila, A.J., E.A.C. Almeida, and J.M. White. 1996. ADAMs and cell fusion. Curr. Opin. Cell Biol. 8: 692-699 |
37. | Ichas, F., L.S. Jouaville, and J.P. Mazat. 1997. Mitochondria are excitable organelles capable of generating and conveying electrical and calcium signals. Cell 89: 1145-1153 |
38. |
Isenmann, S.,
Y. Khew-Goodall,
J. Gamble,
M. Vadas, and
B. Wattenberg.
1998.
A splice-isoform of vesicle-associated membrane protein-1 (VAMP-1)
contains a mitochondrial targeting signal.
Mol. Biol. Cell.
9:
1649-1660
|
39. | Kee, Y., R.C. Lin, S. Hsu, and R. Scheller. 1995. Distinct domains of syntaxin are required for synaptic vesicle fusion complex formation and dissociation. Cell 14: 991-998 . |
40. | Koning, A.J., P.Y. Lum, J.M. Williams, and R. Wright. 1993. DiOC6 staining reveals organelle structure and dynamics in living yeast cells. Cell Motil. Cytoskeleton 25: 111-128 |
41. | Lazzarino, D.A., I. Boldogh, M.G. Smith, J. Rosand, and L.A. Pon. 1994. Yeast mitochondria contain ATP-sensitive, reversible actin-binding activity. Mol. Biol. Cell 5: 807-818 [Abstract]. |
42. | Lindsley, D., and K.T. Tokuyasu. 1980. Spermatogenesis. In Genetics and biology of Drosophila. Vol. 2. M. Ashburner and T.R. Wright, editors. Academic Press, New York. 225-294. |
43. |
Lupashin, V.V., and
M.G. Waters.
1997.
t-SNARE activation through transient
interaction with a rab-like guanosine triphosphatase.
Science
276:
1255-1258
|
44. | Maniatis, T., E.F. Fritsch, and J. Sambrook. 1982. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 545 pp. |
45. | Mayer, A., W. Wickner, and A. Haas. 1996. Sec18p (NSF)-driven release of Sec17p (alpha-SNAP) can precede docking and fusion of yeast vacuoles. Cell 85: 83-94 |
46. | Muhlrad, D., R. Hunter, and R. Parker. 1992. A rapid method for localized mutagenesis of yeast genes. Yeast 8: 79-82 |
47. | Mumberg, D., R. Muller, and M. Funk. 1994. Regulatable promoters of Saccharomyces cerevisiae: comparison of transcriptional activity and their use for heterologous expression. Nucleic Acids Res. 22: 5767-5768 |
48. |
Murphy, G.A.,
M.S. Moore,
G. Drivas,
P.P. de la Ossa,
A. Villamarin,
P. D'Eustachio, and
M.G. Rush.
1997.
A T42A Ran mutation: differential interactions with effectors and regulators, and defect in nuclear protein import.
Mol. Biol. Cell
8:
2591-2604
|
49. | Nichols, B.J., C. Ungermann, H.R.B. Pelham, W. Wickner, and A. Haas. 1997. Homotypic vacuolar fusion mediated by v- and t-SNAREs. Nature 387: 199-202 |
50. | Nunnari, J., W.F. Marshall, A. Straight, A. Murray, J.W. Sedat, and P. Walter. 1997. Mitochondrial transmission during mating in Saccharomyces cerevisiae is determined by mitochondrial fusion and fission and the intramitochondrial segregation of mitochondrial DNA. Mol. Biol. Cell 8: 1233-1242 [Abstract]. |
51. |
Otsuga, D.,
B.R. Keegan,
E. Brisch,
J.W. Thatcher,
G.J. Hermann,
W. Bleazard, and
J.M. Shaw.
1998.
The dynamin-related GTPase, Dnm1p, controls mitochondrial morphology in yeast.
J. Cell Biol.
143:
333-349
|
52. |
Otto, H.,
P.I. Hanson, and
R. Jahn.
1997.
Assembly and disassembly of a ternary complex of synaptobrevin, syntaxin, and SNAP-25 in the membrane of
synaptic vesicles.
Proc. Natl. Acad. Sci. USA
94:
6197-6201
|
53. | Pai, E.F., U. Krengel, G.A. Petsko, R.S. Goody, W. Kabsch, and A. Wittinghofer. 1990. Refined crystal structure of the triphosphate conformation of H-ras p21 at 1.35 angstrom resolution: implications for the mechanism of GTP hydrolysis. EMBO (Eur. Mol. Biol. Organ.) J. 9: 2351-2359 [Abstract]. |
54. | Patel, S.K., F.E. Indig, N. Oliviera, N.D. Levine, and M. Latterich. 1998. Organelle membrane fusion: a novel function for the syntaxin homolog Ufe1p in ER membrane fusion. Cell 92: 611-620 |
55. | Pfeffer, S.R.. 1996. Transport vesicle docking: SNAREs and associates. Annu. Rev. Cell Biol. 12: 441-461 |
56. | Pon, L., T. Moll, D. Vestweber, B. Marshallsay, and G. Schatz. 1989. Protein import into mitochondria: ATP-dependent protein translocation activity in a submitochondrial fraction enriched in membrane contact sites and specific proteins. J. Cell Biol. 109: 2603-2616 [Abstract]. |
57. | Pon, L., and G. Schatz. 1991. Biogenesis of Yeast Mitochondria. In The Molecular Biology of the Yeast Saccharomyces. J.R. Broach, J.R. Pringle, and E.W. Jones, editors. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 334-406. |
58. | Pringle, J.R., A.E.M. Adams, D.G. Drubin, and B.K. Haarer. 1991. Immunofluorescence methods for yeast. Methods Enzymol. 194: 565-602 |
59. |
Qiao, H.,
S.L. Pelletier,
L. Hoffman,
J. Hacker,
R.T. Armstrong, and
J.M. White.
1998.
Specific single or double proline substitutions in the "spring-loaded" coiled-coil region of the influenza hemagglutinin impair or abolish
membrane fusion activity.
J. Cell Biol.
141:
1335-1347
|
60. | Rabouille, C., H. Kondo, R. Newman, N. Hui, P. Fremont, and G. Warren. 1998. Syntaxin 5 is a common component of the NSF- and p97-mediated reassembly pathways of Golgi cisternae from mitotic Golgi fragments in vitro. Cell 92: 603-610 |
61. |
Roeder, A.D., and
J.M. Shaw.
1996.
Vacuole partitioning during meiotic division in yeast.
Genetics
144:
445-458
|
62. |
Roeder, A.D.,
G.J. Hermann,
B.R. Keegan,
S.A. Thatcher, and
J.M. Shaw.
1998.
Mitochondrial inheritance is delayed in Saccharomyces cerevisiae cells
lacking the serine/threonine phosphatase, PTC1.
Mol. Biol. Cell
9:
917-930
|
63. |
Rothman, J.E..
1996.
The protein machinery of vesicle budding and fusion.
Prot.
Sci.
5:
185-194
|
64. | Schneider, B.L., W. Seufert, B. Steiner, Q.H. Yang, and A.B. Futcher. 1995. Use of polymerase chain reaction epitope tagging for protein tagging in Saccharomyces cerevisiae. Yeast 11: 1265-1274 |
65. | Sherman, F., G.R. Fink, and J.B. Hicks. 1986. Methods in Yeast Genetics. Cold Spring Harbor Press, Cold Spring Harbor, NY. 186 pp. |
66. | Sigal, I.S., J.B. Gibbs, J.S. D'Alonzo, G.L. Temeles, B.S. Wolanski, S.H. Socher, and E.M. Scolnick. 1986. Mutant ras-encoded proteins with altered nucleotide binding exert dominant biological effects. Proc. Natl. Acad. Sci. USA. 83: 952-956 [Abstract]. |
67. | Sogo, L.F., and M.P. Yaffe. 1994. Regulation of mitochondrial morphology and inheritance by Mdm10p, a protein of the mitochondrial outer membrane. J. Cell Biol. 126: 1361-1373 [Abstract]. |
68. | Söllner, T., S.W. Whitehart, M. Brunner, H. Erdjumentbronage, S. Geromanos, P. Tempst, and J.E. Rothman. 1993. SNAP receptors implicated in vesicle targeting and fusion. Nature 362: 318-324 |
69. | Stevens, B. 1981. Mitochondrial Structure. In The Molecular Biology of the Yeast Saccharomyces. J.M. Strathern, E.W. Jones, and J.R. Broach, editors. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 471-504. |
70. | van der Bliek, A.M., T.E. Redelmeier, H. Damke, E.J. Tisdale, E.M. Meyerowitz, and S.L. Schmid. 1993. Mutations in human dynamin block an intermediate stage in coated vesicle formation. J. Cell Biol. 122: 553-563 [Abstract]. |
71. | Vater, C.A., C.K. Raymond, K. Ekena, I. Howald-Stevenson, and T.H. Stevens. 1992. The VPS1 protein, a homologue of dynamin required for vacuolar protein sorting in Saccharomyces cerevisiae. J. Cell Biol. 119: 773-786 [Abstract]. |
72. | Vojtek, A.B., S.M. Hollenberg, and J.A. Cooper. 1993. Mammalian Ras interacts directly with the serine/threonine kinase Raf. Cell 74: 205-214 |
73. | Weber, T., B.V. Zemelman, J.A. McNew, B. Westermann, M. Gmachi, F. Parlati, T.H. Sollner, and J.E. Rothman. 1998. SNAREpins: minimal machinery for membrane fusion. Cell 92: 759-772 |
74. | White, J.M.. 1990. Viral and cellular membrane fusion proteins. Annu. Rev. Physiol. 52: 675-697 |
75. | White, J.M.. 1992. Membrane fusion. Science 258: 917-924 |
76. | Winston, F., C. Dollard, and S.L. Ricupero-Hovasse. 1995. Construction of a set of convenient Saccharomyces cerevisiae strains that are isogenic to S228C. Yeast 11: 53-55 |
77. | Yaffe, M.P.. 1995. Isolation and analysis of mitochondrial inheritance mutants from Saccharomyces cerevisiae. Methods Enzymol. 260: 447-453 |
78. | Zinser, E., and G. Daum. 1995. Isolation and biochemical characterization of organelles from the yeast, Saccharomyces cerevisiae. Yeast 11: 493-536 |