Department of Cell Biology, University of Massachusetts Medical Center (UMMC), Worcester Foundation Campus, Shrewsbury, Massachusetts 01545
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Abstract |
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Several enzymes, including cytoplasmic and flagellar outer arm dynein, share an Mr 8,000 light chain termed LC8. The function of this chain is unknown, but it is highly conserved between a wide variety of organisms. We have identified deletion alleles of the gene (fla14) encoding this protein in Chlamydomonas reinhardtii. These mutants have short, immotile flagella with deficiencies in radial spokes, in the inner and outer arms, and in the beak-like projections in the B tubule of the outer doublet microtubules. Most dramatically, the space between the doublet microtubules and the flagellar membrane contains an unusually high number of rafts, the particles translocated by intraflagellar transport (IFT) (Kozminski, K.G., P.L. Beech, and J.L. Rosenbaum. 1995. J. Cell Biol. 131:1517-1527). IFT is a rapid bidirectional movement of rafts under the flagellar membrane along axonemal microtubules. Anterograde IFT is dependent on a kinesin whereas the motor for retrograde IFT is unknown. Anterograde IFT is normal in the LC8 mutants but retrograde IFT is absent; this undoubtedly accounts for the accumulation of rafts in the flagellum. This is the first mutation shown to specifically affect retrograde IFT; the fact that LC8 loss affects retrograde IFT strongly suggests that cytoplasmic dynein is the motor that drives this process. Concomitant with the accumulation of rafts, LC8 mutants accumulate proteins that are components of the 15-16S IFT complexes (Cole, D.G., D.R. Deiner, A.L. Himelblau, P.L. Beech, J.C. Fuster, and J.L. Rosenbaum. 1998. J. Cell Biol. 141:993-1008), confirming that these complexes are subunits of the rafts. Polystyrene microbeads are still translocated on the surface of the flagella of LC8 mutants, indicating that the motor for flagellar surface motility is different than the motor for retrograde IFT.
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Introduction |
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AN Mr 8,000 polypeptide, first identified as a light chain
LC8 of outer arm axonemal dynein in the green
alga Chlamydomonas reinhardtii (Piperno and Luck,
1979; Pfister et al., 1982
; King and Patel-King, 1995
), has
subsequently been shown to be associated with many different complexes, including cytoplasmic dynein (King et
al., 1996
), the Chlamydomonas inner arm dynein I1 (Harrison et al., 1998
), myosin V (Espindola, F.S., R.E. Cheney,
S.M. King, D.M. Suter, and M.S. Mooseker. 1996. Mol.
Biol. Cell. 7:372a), neuronal nitric oxide synthase (Jaffrey
and Snyder, 1996
), and a complex within the tegument of
the blood fluke Schistosoma (Hoffman and Strand, 1996).
The protein has been highly conserved throughout evolution, with homologues from humans, Drosophila melanogaster, Caenorhabditis elegans, and Chlamydomonas having
~90% sequence identity (King et al., 1996
). The function
of the polypeptide is not known, although it appears to
have a negative effect on nitric oxide synthase activity
(Jaffrey and Snyder, 1996
). The fact that the protein is associated with many different enzymes raises the possibility
that it has a common structural or functional role in diverse polypeptide complexes.
Because LC8 is associated with at least two complexes
in Chlamydomonas, Chlamydomonas provides a potentially unique and valuable system for investigating the cellular distribution and roles of LC8. Of particular importance is the ability to transform the nuclear genome of
Chlamydomonas (Kindle, 1990), which makes possible insertional mutagenesis (Tam and Lefebvre, 1993
; Gumpel
and Purton, 1994
; Pazour et al., 1995
). When Chlamydomonas is transformed, the transforming DNA inserts into
the genome at random, and either disrupts any gene at the
site of insertion or, more commonly, causes the deletion of
a block of DNA flanking the site of insertion. In either
case, the result is a mutation that is tagged by the exogenous DNA and can be identified as a restriction fragment length polymorphism. If a probe for a particular gene is
available, insertional mutants can be screened by Southern
blotting to identify those in which the gene is deleted or
disrupted. An advantage of insertional mutagenesis in
Chlamydomonas is that it usually results in a null mutant.
We have used this strategy to identify null mutants for
LC8 in Chlamydomonas. We used a cDNA encoding LC8
(King and Patel-King, 1995
) to screen a large collection of insertional mutants originally selected for defects in cell or flagellar morphology, flagellar motility, or phototaxis (Pazour et al., 1995
; Koutoulis et al., 1997
), and found two
strains that completely lack the LC8 gene, which we here
term FLA14. The mutants grow normally but have short,
paralyzed flagella that become progressively shorter during the light period. The mutants are rescued by transformation with the gene for LC8, confirming that the phenotype is due to loss of that polypeptide. Electron
microscopy of the mutant flagella revealed deficiencies in
the inner and outer arms, radial spokes, and beak-like projections that extend into the lumens of the B tubules of
outer doublet microtubules numbers 1, 5, and 6 (Witman et al., 1972
; Hoops and Witman, 1983
). In addition, there is
an abnormal accumulation of particles in the space between the flagellar membrane and doublet microtubules.
These particles appear to be identical to the "rafts" that
move rapidly to the tip of the flagellum and back again
during intraflagellar transport (IFT)1 (Kozminski et al.,
1993
, 1995
). Video-enhanced differential interference contrast (DIC) microscopy revealed that the mutants have
normal anterograde IFT but lack retrograde IFT, accounting for the accumulation of rafts in the flagellum. The
other structural abnormalities in the axoneme may reflect
a direct requirement for LC8 in the assembly of the affected components, or in the transport of the components
to the flagellum or within the flagellar shaft. In any case,
the results show that LC8 is essential for normal flagellar morphogenesis and stability.
Because LC8 is a subunit of cytoplasmic dynein, our
findings suggest that cytoplasmic dynein is the motor for
retrograde IFT. The outer arm dynein, which also contains
LC8 and is potentially in the correct location to generate
IFT, can be ruled out as the retrograde motor because mutants lacking the outer arms have normal IFT (Kozminski
et al., 1993). LC8 is not necessary for flagellar surface motililty (Bloodgood, 1989
), indicating that retrograde IFT
and retrograde surface motility are powered by different
motors.
Recently, novel 15-16S particles have been identified in
the cytoplasmic matrix fraction of Chlamydomonas flagella (Cole, D.G., and J.L. Rosenbaum. 1996. Mol. Biol.
Cell. 7:47a; Piperno and Mead, 1997; Cole et al., 1998
).
Concomitant with the accumulation of rafts in the flagella
of LC8 mutants, we find a massive accumulation of the
polypeptides that are postulated to be components of
these complexes. These results confirm that the 15-16S
particles are subunits of the rafts (Cole et al., 1998
).
Because Chlamydomonas is advantageous for both biochemical and molecular genetic studies, the null mutants described here should be very useful for definitive identification of the retrograde IFT motor, and for understanding the role of LC8 in the functioning of this motor. The mutants also should be useful for determining the full repertoire of complexes with which LC8 is associated in Chlamydomonas, and whether LC8 plays a direct role in the assembly of structures such as the inner and outer dynein arms.
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Materials and Methods |
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Strains
Chlamydomonas reinhardtii strains used in this work included: g1 (nit1,
NIT2, agg1, mt+) (Pazour et al., 1995), 137c (nit1, nit2, mt+), CC124 (nit1,
nit2, mt
), ida1 (ida1, mt+) (obtained from R. Kamiya, University of Tokyo, Tokyo, Japan), and pf18 (pf18, mt
). Strains 137c, CC124, and pf18
can be obtained from the Chlamydomonas Culture Collection (Duke University, Durham, NC). Strains produced in the course of this study included: V64 (fla14-1::NIT1, nit1, mt+) and V101 (fla14-2::NIT1, nit1,
mt+), both insertional mutants obtained by transforming g1 with cloned
NIT1 DNA; F5 (FLA14, fla14-1::NIT1, nit1, mt+) obtained by transforming V64 with the cloned FLA14 gene; and 2782.1 (FLA14, fla14-1::NIT1,
mt
) obtained by crossing F5 to 137c.
Growth Medium
Cells were grown in the following media: M (Sager and Granick [1953] medium I altered to have 0.0022 M KH2PO4 and 0.00171 M K2HPO4), M-N (M medium without nitrogen), R (M medium plus 0.0075 M sodium acetate), and SGII/NO3 (Sager and Granick [1953] medium II modified to have 0.003 M KNO3 as the nitrogen source).
Transformation
Transformation was performed using the glass bead method of Kindle
(1990) as described in Pazour et al. (1995)
. Insertional mutants were obtained by transforming nit1 cells (strain g1) with plasmid pGP505 (Pazour
et al., 1995
) which contains the Chlamydomonas nitrate reductase gene
(Fernandez et al., 1989
). After transformation, the cells were plated on
solid SGII/NO3 medium and allowed to grow into colonies. Individual colonies were picked into 5 ml of R medium and grown until the cultures
were light green in color. The cells were then examined for photoaccumulation and motility (Pazour et al., 1995
; Koutoulis et al., 1997
). From 2,978 transformants screened, 55 lines with defects in motility were identified.
DNA was isolated from the 55 lines and examined by Southern blotting using the LC8 cDNA clone as a probe.
For experiments in which fla14 cells were rescued by transformation with cloned FLA14 genomic DNA, transformants were selected using an enrichment technique based on the ability of transformants to swim. After vortexing in the presence of the exogenous DNA, cells were placed directly into flasks containing 125 ml of liquid R medium and then allowed to grow for ~1 wk. Untransformed cells were unable to swim and remained on the bottoms of the culture flasks, whereas transformants could swim and were distributed throughout the flasks. Transformants were observed in all flasks. After a few rounds of enrichment obtained by removing the inoculi from the tops of the cultures, most of the cells were swimming. At this point, cells were plated on solid medium; a single colony was kept from each flask.
Genetic Analysis
Mating and tetrad analysis were performed as described by Levine and
Ebersold (1960) and Harris (1989)
. Cells of each mating type were grown
on solid R medium, resuspended in M-N liquid medium, and then mixed
together. After pellicles became apparent in 1 or 2 d, the mixture was
plated on solid M medium, allowed to dry, and then placed in the dark for
6-10 d. Zygotes were hatched on solid R medium and dissected using a
glass needle. The meiotic progeny were allowed to grow for 3-5 d and
then transferred to 5 ml of liquid R medium. Cells were allowed to grow
for an additional 2-5 d and then scored for motility by microscopic observation of cells illuminated with dim red light.
Analysis of IFT and Flagellar Length
IFT was observed in living cells by video-enhanced DIC microscopy as described by Moss et al. (1992), except that a 20× projection lens was used,
and images were recorded on a Toshiba (Japan) PCM VHS recorder.
Cells were immobilized for observation by placing them between a slide
coated with a thin layer of 0.5% agarose in M medium and a coverslip.
Output was recorded as an AVI file using AV Master (Fast Multimedia,
Munich, Germany) to capture the data. IFT was quantitated by manually
counting the number of particles passing a point about halfway along the
flagellum while also measuring elapsed time.
For measurement of flagellar lengths, live cells were recorded at 2,000×. Subsequently, images of the flagella were displayed on a video monitor, traced onto transparent acetate overlays, and then measured with a flexible ruler. A slide micrometer was measured in the same way to establish the scale.
Isolation and Fractionation of Flagella
Flagella were isolated from the fla14 mutant strain V64 and from wild-type strain 137c by the dibucaine method (Witman, 1986). Isolated flagella
were demembranated with 1% NP-40 and the membrane plus matrix fraction was separated from the axonemes by centrifugation (Witman, 1986
).
The axonemes were then washed with a solution containing 10 mM ATP
in 30 mM Hepes pH 7.4, 10 mM MgSO4, 2 mM DTT, 0.5 mM EGTA, and
30 mM potassium acetate to remove any ATP-extractable components.
Finally, the washed axonemes were resuspended in the same solution but
without ATP. All three fractions from a particular preparation were equal
in volume. Samples from different preparations were normalized by comparing the amounts of axonemal tubulin on Coomassie blue-stained gels.
Western Blotting
Flagellar fractions were separated by SDS-PAGE (Pfister et al., 1982) and
electroblotted onto a polyvinylidene difluoride membrane (Immobilon-P;
Millipore Corp., Waters Chromatography, Milford, MA). Western blotting was carried out by standard procedures (Sambrook et al., 1987
) using
5% nonfat dry milk in 10 mM Tris, pH 7.5, 166 mM NaCl, and 0.05%
Tween to block the membrane. Primary antibodies were diluted in the
blocking solution as recommended by the supplier and then hybridized for
2 h at room temperature. Horseradish peroxidase-conjugated secondary
antibodies and enhanced chemiluminescence (Amersham Pharmacia Biotech. Inc., Piscataway, NJ) were used to detect the primary antibodies.
Primary antibodies used included: (a) R4058, specific for LC8 (King
and Patel-King, 1995) (gift of S. King, University of Connecticut Health
Center, Farmington, CT); (b) 1878
, specific for the outer arm dynein intermediate chain IC78 (King et al., 1985
); (c) EU51, specific for the inner
arm dynein intermediate chain IDA-IC140 (previously unpublished antibody gift of P. Yang and W. Sale, both from Emory University, Atlanta,
GA); (d) 2-10-
6, specific for
tubulin (gift of G. Piperno, Mount Sinai
School of Medicine, New York) (Piperno et al., 1987
; King et al., 1991
); (e) anti-FLA10N, specific for the anterograde IFT motor subunit FLA10
(Cole et al., 1998
) (gift of D. Cole and J. Rosenbaum, both from Yale University, New Haven, CT); and (f) a mixture of mAB172.1, mAB139.1,
mAB81.1, mAB81.2, mAB81.3, mAB81.4, and mAB57.1 specific for four
of the raft proteins (Cole et al., 1998
) (gift of D. Cole and J. Rosenbaum).
Additional antibodies used were specific for radial spoke proteins RSP1
(Williams et al., 1986
) and RSP3 (Williams et al., 1989
) (gifts of D. Cole
and J. Rosenbaum), the highly conserved kinesin sequences HIPYR and
LAGSE (gift of C. Walczak, University of California, San Francisco, CA),
and the molecular chaperone HSP70 (previously unpublished antibody
gift of E. Savino and J. Rosenbaum, both from Yale University).
Electron Microscopy
Cells were fixed in glutaraldehyde (Hoops and Witman, 1983) and processed as described in Wilkerson et al. (1995)
or Kozminski et al. (1995)
.
DNA Isolation and Analysis
DNA was isolated by digesting ~0.3 ml of packed cells with 0.5 ml of proteinase K (1 mg/ml) in 5% sodium lauryl sulfate, 20 mM EDTA, and 20 mM Tris, pH 7.5, at 50°C for 12-16 h. Ammonium acetate was added to
1.5 M, the mixture extracted once with 50% phenol/50% chloroform, once
with chloroform, and then precipitated with 0.7 vol of isopropyl alcohol.
DNA was resuspended in a solution containing 10 mM Tris, pH 8.0, and 1 mM EDTA and then digested with PstI. Gel electrophoresis and Southern
blotting were performed according to standard procedures (Sambrook et al.,
1987).
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Results |
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Identification of Mutants Deleted for the LC8 Gene
We previously used insertional mutagenesis to generate a
large number of mutants with defective flagella (Koutoulis
et al., 1997); these included slow-jerky swimmers, slow-smooth swimmers, uniflagellate cells, aflagellate cells, cells
with paralyzed flagella, and cells with long flagella. In an
effort to find mutations in the gene for LC8, we screened
the slow-jerky swimming cell lines for a restriction fragment length polymorphism detectable using a cDNA
clone encoding LC8. Slow-jerky swimming is indicative of outer dynein arm mutations (Kamiya, 1988
), and it seemed
reasonable that a defect in LC8 would affect the outer
arm. However, we found no LC8 mutations in this collection, suggesting that the phenotype of an LC8 mutation
could be quite different from that of other outer dynein
arm mutations. We therefore expanded our search by screening all of the insertional mutants in our collection.
This identified two cell lines, V64 and V101, that were
completely deleted for LC8 (Fig. 1).
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Both LC8 mutants have short, often unequal length flagella (Fig. 2 A) that sometimes have swellings on their sides or tip. The flagella are paralyzed and only rarely exhibit slight bending movements. They are longest at the beginning of the light cycle and shorten linearly at a rate of ~0.2 µm/h as the day progresses (Fig. 2 B). The mutations do not appear to affect growth or division of the cells, since doubling times and cell body morphology are normal (Fig. 2, A and C).
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We attempted to use tetrad analysis to investigate if the short paralyzed flagella phenotype was linked to the LC8 deletion. The V64 and V101 cell lines did not mate well with wild-type cells and only two partial tetrads were obtained after many attempts. The mutant offspring of these tetrads were missing the LC8 gene, whereas the normal offspring contained the LC8 gene, suggesting that the phenotype was indeed linked to the LC8 deletion.
More definitive evidence that loss of LC8 was responsible for the mutant phenotype was obtained by rescuing the mutants with the cloned LC8 gene. Genomic DNA encoding LC8 was isolated from a lambda phage library and used to transform mutant cells. The lambda phage clones and subclones as small as 3.1 kb complemented the mutation, restoring flagellar length (Fig. 2 A, Transformant) and the ability to swim. To rule out the possibility that the rescuing DNA contained genes in addition to LC8, the ends of the 3.1-kb complementing fragment were sequenced (data not shown). The LC8 gene occupied ~700 bp beginning at ~150 bp from one end. The sequence at the other end of the clone matched internal exons of the Arabidopsis propionyl-CoA carboxylase gene. The central ~1,200 bp of the fragment were not sequenced but presumably contained the remaining exon(s) and the 3' untranslated region of the propionyl-CoA carboxylase gene. This leaves very little room for any other gene, making it highly unlikely that this fragment contains any full-length genes except LC8.
To confirm that rescue was due to integration of the LC8 gene as opposed to suppression of the phenotype by spontaneous mutation of some other gene, we examined the segregation pattern of the exogenous DNA in crosses involving the rescued mutants. One of these transformants was mated to wild-type cells and then tetrads were dissected. A motile meiotic product, deleted for the endogenous copy of the LC8 gene but carrying a transformed copy of this gene, was identified and mated to wild-type cells. Seventeen complete tetrads were obtained. The motility of the offspring was observed by light microscopy and the genotype was determined by Southern blotting (Fig. 3). Because the transformed copy of the LC8 gene is integrated at a site unlinked to the original locus, the inserted gene segregated independently of the original gene, resulting in offspring with zero, one, or two functional copies of the gene. Cells with one or two copies of the gene exhibited a wild-type phenotype, whereas those with no copies had a mutant phenotype. Thus, the defects in V64 and V101 are caused by deletion of the gene encoding LC8, and rescue was due to restoration of the LC8 gene. This gene will be termed FLA14, and the mutations have been named fla14-1 and fla14-2, respectively.
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LC8 Deletion Mutants Have Severe Defects in Flagellar Ultrastructure
Electron microscopic analysis of V64 and V101 flagella revealed that the beak-like projections (Witman et al., 1972)
in the B tubule of the outer doublet microtubules are missing, most of the radial spokes are missing or very defective, and the outer and inner dynein arms are reduced in
number (Fig. 4). The central pair of microtubules and their
projections appeared normal, although the central pair
frequently is not centered in the axoneme, as previously
observed in the absence of radial spokes (Witman et al.,
1978
).
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The most striking feature of fla14 flagella is an unusual
abundance of electron-dense material between the flagellar membrane and the outer doublet microtubules. This
material looks very much like the rafts that have been correlated with the moving particles in IFT (Kozminski et al.
1993, 1995
). When viewed in flagellar cross-sections, rafts
appear as pairs of globular structures, each ~20 nm in diameter (Fig. 4 C). These paired structures occasionally were observed in wild-type flagella, but in the mutant they
were so abundant that they sometimes completely circumscribed the axoneme (Fig. 4, B and C). In longitudinal sections of flagella (Fig. 4 D) the rafts appeared as linear arrays of subunits, as reported by Kozminski et al. (1993)
.
Frequently, especially at the distal tips of the flagella, the
rafts were piled up, causing distension of the flagellar
membrane; this presumably gives rise to the bulges seen
by light microscopy. In such cases, some rafts were closely associated with the outer doublet microtubules, and others
with the flagellar membrane; these associations appeared
to be mediated by raft-microtubule or raft-membrane
crossbridges, respectively (Fig. 5).
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Retrograde IFT Is Defective in LC8 Mutants
It has been suggested that cytoplasmic dynein is the motor
responsible for retrograde IFT (Kozminski et al., 1995).
The fact that LC8 is a subunit of cytoplasmic dynein (King
et al., 1996
) prompted us to examine the flagella of the
fla14 mutants to determine if IFT was affected in any way.
IFT in flagella of control and mutant cells was assayed by
video-enhanced DIC microscopy. In control cells, particles
were observed moving in both the anterograde and retrograde directions (Fig. 6 A and Table I). Movement was smooth and continuous in each direction; the retrograde
rate was slightly greater than the anterograde rate, as previously reported (Kozminski et al., 1995
). In contrast, fla14
mutants almost completely lacked retrograde IFT, although anterograde IFT appeared to be normal (Fig. 6 B
and Table I). No discrete particles were observed to move
toward the cell body, although occasionally there appeared to be a peristaltic-like movement of material from
tip to base. The rate of anterograde transport in the mutants was indistinguishable from that in control cells (Fig.
6, A and B).
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fla14 Mutations Do Not Abolish Surface Motility
In addition to beating its flagella to swim through liquid
medium, Chlamydomonas also uses its flagella to glide
along solid surfaces. This occurs when the flagella bind to
the substrate and then glide over the substrate surface. If
the flagella are oriented in opposite directions, one of the
two becomes dominant and pulls the cell body and other
flagellum along after it. In a process that may be related to
gliding motility, Chlamydomonas flagella translocate exogenous microbeads that become bound to the outside of
their flagella (Bloodgood, 1989). However, in contrast to
gliding motility, where the apparent forces are unidirectional, bead movement occurs in both directions, indicating bidirectional forces.
The flagella of fla14 cells do not adhere to glass and so
we were unable to determine if they are capable of gliding
motility. However, polystyrene microspheres bind to fla14
flagella and are translocated (Fig. 7). In pf18 cells, which
have normal IFT (Kozminski et al., 1993) but have paralyzed flagella and thus facilitate observation of bead movement, the beads are seen moving along the flagella or
pausing near the base of the flagella (Fig. 7, A and C, open
bars). In contrast, the beads on fla14 flagella are much
more likely to be found at the tips of the flagella (Fig. 7, B
and C, closed bars). This is not due to a lack of retrograde movement of beads, since beads can be observed moving
in both directions on fla14 flagella (Fig. 7 D, closed circles). The rates of both anterograde and retrograde bead
movement (as indicated by the slopes of the lines in Fig. 7
D) are generally slower in the mutant than in wild type. It
is not clear if the slower rate is directly due to the lack of
LC8 or is an indirect effect related to the flagella being
filled with rafts.
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Many Flagellar Proteins Are Affected by Deletion of the LC8 Gene
The defects in axonemal structures and the accumulation
of raft-like material beneath the flagellar membrane suggested that there might be differences in the content of
proteins in the various flagellar fractions of fla14 mutants
as compared with wild-type cells. To investigate this, flagella were isolated from wild-type and fla14-1 cells and
separated into the following three fractions: (a) "membrane plus matrix," the detergent-soluble membrane proteins and the soluble proteins within the flagellum; (b)
"ATP wash," proteins that are bound to the axoneme only
by an ATP-sensitive bond, as might be expected for active
IFT motors and proteins associated with them; and (c) "axoneme," the insoluble proteins not released by detergent
or ATP. For each cell type (wild-type or mutant), the volumes of the fractions were adjusted so that equal aliquots
of each fraction contained protein from an equivalent number of flagella. The proteins in these fractions were
separated by SDS-PAGE and analyzed by silver staining
or immunoblotting; because of the difference in flagellar
length, loadings of fractions from wild-type versus mutant
cells were normalized based on tubulin content in the respective axonemal fractions (Fig. 8 B). In the membrane
plus matrix fraction, a number of polypeptides were elevated in fla14 flagella (Fig. 8 A, white dots), whereas only
two prominent proteins, migrating just behind tubulin in wild type, were missing from this fraction in the mutant
(Fig. 8 A, black dots). In contrast, in the axoneme fraction,
many proteins were decreased by the mutation.
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Western blotting with an antibody to LC8 showed that this protein was present in all fractions of the wild-type cells but in none of the fla14 fractions (Fig. 8 B, FLA14). This is expected as the fla14-1 allele is a complete deletion of the gene (refer to Fig. 1).
By EM, radial spokes appeared to be missing from fla14 axonemes. To confirm this result and to determine if the radial spokes had accumulated in an unassembled state in the fla14 membrane plus matrix fraction, blots were probed with antibodies to the radial spoke stalk protein RSP1 and the radial spoke head protein RSP3 (Fig. 8 B, RSP1 and RSP3). The blots revealed that these proteins are almost completely absent from the axonemes of the mutants, in agreement with the EM observations. The radial spoke proteins do not accumulate in an unassembled state in fla14 flagella, since these proteins also were reduced in the membrane plus matrix fraction of the mutant.
EM analysis indicated that the inner and outer dynein
arms are reduced in number in the mutant flagella. Inasmuch as LC8 is a subunit of both outer arm dynein (King
and Patel-King, 1995) and of the major inner arm dynein
species I1 (Harrison et al., 1998
), the absence of this subunit may affect the ability of the arms to assemble, be
transported to, or attach to their normal binding sites on
the doublet microtubules. Western blots using an antibody
to the outer arm dynein intermediate chain IC78 showed that this protein was absent from fla14 axonemes and
greatly reduced in the fla14 membrane plus matrix and
ATP wash fractions (Fig. 8 B, ODA-IC78). These results
are in agreement with the EM observations. It is likely that
the few outer arms observed in fla14 flagella by EM are
weakly attached to the axoneme and released by ATP. ATP-dependent release of at least some of the outer arms
from wild-type axonemes has been described previously
(Goodenough and Heuser, 1984
), but the mechanism of
this release is not understood. Western blots using an antibody to the inner arm I1 intermediate chain IC140 revealed a decrease in the amount of this chain in fla14 axonemes, nearly equivalent amounts of the chain in wild-type and fla14 membrane plus matrix fractions, but an increase in the amount of the chain in the ATP wash from
fla14 axonemes (Fig. 8 B, IDA-IC140). The former result
is consistent with our EM observations; the latter result
suggests that more of the I1 inner arms are attached to the
axoneme solely by ATP-dependent bonds in fla14 than in
wild-type cells. The results also indicate that the deficiency for inner arm I1 is not as severe as that for the outer arm.
The large amount of raft-like material seen in the mutant flagella by EM suggested that proteins of the rafts
should be elevated in the membrane plus matrix and/or
ATP wash fractions of fla14 cells. These rafts are thought
to be composed of two complexes (complex A and complex B) sedimenting at 15-16S and together containing at least 15 different polypeptides (Cole, D.G., and J.L.
Rosenbaum. 1996. Mol. Biol. Cell. 7:47a; Piperno and
Mead, 1997; Cole et al., 1998
). Western blotting using
monoclonal antibodies raised against one polypeptide
(p139) of complex A and three polypeptides (p172, p81,
and p57) of complex B (Cole et al., 1998
) showed that all
four proteins are greatly elevated in both the membrane
plus matrix and ATP wash fractions of fla14 cells (Fig. 8 C).
These findings provide further evidence that complexes A
and B make up at least part of the rafts. The fact that these
proteins were elevated in both fractions is consistent with
the EM observation that rafts are closely associated with
both flagellar outer doublet microtubules and membranes.
The molecular chaperone, HSP70, is found within the
flagellum and has been suggested to play a role in targeting tubulin and other axonemal components to the tip of
the flagellum where they are incorporated into the growing
structure (Bloch and Johnson, 1995). Within the flagellum,
HSP70 has been reported to be in both the membrane plus
matrix fraction and bound to the axoneme; furthermore,
part of the axonemal fraction is extractable by ATP (Bloch
and Johnson, 1995
). In our fractionation (Fig. 8 B), HSP70 had a distribution similar to that seen by Bloch and Johnson
(1995)
. HSP70 also has been suggested to be a component of
the 15-16S complexes believed to constitute the rafts (Piperno
and Mead, 1997
). We found that Hsp70 was slightly elevated
in fla14 flagella, but much less so than the p172, p139, p81,
and p57 components of the 15-16S complexes. Therefore,
if Hsp70 is a component of either complex A or complex
B, it is associated with only a subset of those particles.
Kinesin-related Proteins in fla14 Flagella
If retrograde transport is not functioning, anterograde motors may not be recycled back to the cell body and might
accumulate in the flagella. The anterograde motor is
thought to be a heterotrimeric kinesin, FLA10 kinesin-II
(Kozminski et al., 1995; Cole et al., 1998
). FLA10 kinesin-II contains two different kinesin-like motor subunits, one
of which is the ~90-kD FLA10 protein, and a slightly larger nonmotor subunit. We found that in wild-type flagella, FLA10 was most abundant in the ATP wash fraction,
although it was present in the other two fractions as well
(Fig. 9 A, FLA10). This protein was elevated in all fractions of fla14 flagella, consistent with the idea that the anterograde motors are not being returned to the cell body.
|
The pan-kinesin antisera, LAGSE and HIPYR, detect
several other proteins within the Chlamydomonas flagella
(Fox et al., 1994; Johnson et al., 1994
). To determine how
these are affected by fla14, Western blots were probed
with these sera. Both LAGSE and HIPYR detect a
FLA10-sized protein and a slightly smaller protein (Fig. 9,
B and C) that are greatly elevated in fla14 flagella. This
smaller protein may be the 85-kD motor subunit of the
heterotrimeric FLA10 kinesin-II (Cole et al., 1998
). However, the relative amounts of the two bands varied in the
wild-type fractions, suggesting that more than one species
is represented. There also is a >200-kD axonemal protein
(detected by LAGSE), a ~110-kD protein (detected by
both LAGSE and HIPYR in the ATP wash and the axoneme), and a ~40-kD protein (detected by HIPYR in the
ATP wash and axoneme fractions) that are elevated by the
mutation. Other proteins at ~150 and ~125 kD (detected
by HIPYR in the ATP wash) are not greatly affected by
the fla14 mutation. Interestingly, one protein at ~55 kD
(detected by LAGSE) in the membrane plus matrix fraction of wild type is completely missing in the mutant. This
may correspond to one of the two proteins indicated by
black dots in the membrane plus matrix fraction of Fig. 8 A.
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Discussion |
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Phenotype of LC8 Deletion Mutants
We have isolated mutant cell lines of Chlamydomonas
that are missing the gene encoding LC8, first identified as
a subunit of outer arm dynein (Piperno and Luck, 1979;
Pfister et al., 1982
; King and Patel-King, 1995
). Although
this chain also is a subunit of cytoplasmic dynein (King
and Patel-King, 1995
), and has been reported to be associated with mammalian myosin V (Espindola, F.S., R.E.
Cheney, S.M. King, D.M. Suter, and M.S. Mooseker. 1996. Mol. Biol. Cell. 7:37a) and neuronal nitric oxide synthase
(Jaffrey and Snyder, 1996
), deletion of the gene from
Chlamydomonas does not have a detectable effect on cell
growth or gross cell morphology. Therefore, the gene is
not required for any critical cellular process in Chlamydomonas. However, loss of the gene, which we have named
FLA14, does cause severe flagellar defects. Flagella of the
mutant cells are short, unstable, and exhibit only slight bending movements. They are defective in retrograde IFT
and are missing or have deficiencies in several axonemal
structures. In addition, the space between the axonemal
microtubules and the flagellar membrane of the mutants is
filled with an abnormal amount of electron-dense material. This material appears to be identical to the rafts, described by Kozminski et al. (1993
, 1995
), that have been
correlated with the moving particles of IFT. Consistent with the EM observations, there is an increase in many
proteins in the fla14 membrane and matrix fraction, and a
decrease in other proteins in the fla14 axonemal fraction,
relative to wild-type.
LC8 Is Required for Assembly of Multiple Axonemal Components
Both EM and Western blotting of flagellar fractions demonstrate that fla14 mutants have greatly reduced numbers
of outer dynein arms. This is most likely a direct result of
the loss of LC8, which is an outer arm subunit and, like the
outer arm dynein heavy and intermediate chains (Witman
et al., 1994), may be required for arm assembly or attachment to the doublet microtubule.
LC8 also has been reported to be a subunit of inner arm
I1 (Harrison et al., 1998). As with the outer arm, both EM
and Western blotting indicate that there is a reduced number of inner arms in fla14 axonemes, although the deficiency is not as severe as that for the outer arm. The 140-kD
intermediate chain of inner arm I1 is unique among the axonemal proteins that we examined in that it is elevated in
the ATP-wash fraction of fla14 mutants relative to that of
wild type. This could be because a substantial portion of I1
arms are more weakly attached than normal to their correct sites on the doublet microtubules of fla14 flagella, perhaps due to their deficiency in LC8. Alternatively, the I1
arms extracted by ATP could have been unassembled
arms which were in transit to the tip of the flagellum (see
below) and became bound directly or indirectly to the axonemal microtubules by rigor bonds after flagellar detachment.
Particularly intriguing is the nearly total loss of the radial spokes from fla14 axonemes. Similarly, fla14 axonemes are missing the beak-like projections present in
the lumens of the B tubules of doublet microtubules 1, 5, and 6 (Witman et al., 1972; Hoops and Witman, 1983
). A
possible explanation is that LC8 also is a subunit of these
components, and is directly required for their assembly
and/or attachment to the doublet microtubules. In this context, it may be relevant that axonemes contain a pool
of LC8 not associated with either the outer or inner dynein
arms (Benashski, S.E., A. Harrison, and S.M. King, unpublished data, cited in Benashski et al., 1997
).
The above explanations presume that LC8 is a subunit
of four different axonemal structures, and is necessary for
all four structures to assemble or bind to the doublet microtubules. An alternative explanation for the loss of inner
arms, radial spokes, and beak-like projections in fla14 mutants is that the absence of LC8 affects transport of these
structures in the flagellum. Both the radial spokes
(Johnson and Rosenbaum, 1992) and the inner arms (Piperno et al., 1996
) must be transported to the tip of the flagellum before binding to their normal sites on the doublet microtubules. In dikaryon rescue experiments (Segal et al.,
1984
), the beak-like projections are restored to fully
formed flagella of mutants specifically lacking the projections, raising the possibility that these structures also are
transported to the tip of the flagellum and then move back
down the lumen of the B tubule to their binding sites. At
least for the inner arms, movement to the tip of the flagellum is dependent upon FLA10 (Piperno et al., 1996
), one of the subunits of the anterograde IFT motor (Walther et al., 1994
; Kozminski et al., 1995
). If defects in the fla14 flagellum prevented this transport, deficiencies in assembly of
inner arms, radial spokes, and perhaps the beak-like projections would result. Although anterograde movement of
rafts appears to proceed unimpaired in fla14 flagella, the
movement of other components may be impeded.
It also is possible that loss of the structures from fla14
axonemes is due to a defect in the movement of axonemal
precursors into the flagellum, or to the base of the flagellum before their transport into the flagellum. In the accompanying paper, Cole et al. (1998) show that FLA10
kinesin-II and the IFT particle proteins are highly concentrated in the cell body near the basal bodies, and point
out that, inasmuch as the minus ends of the cytoplasmic
microtubules terminate near the basal bodies, this accumulation may involve a retrograde microtubule motor
such as cytoplasmic dynein. If the loss of LC8 impairs
transport of axonemal precursors to the base of the flagellum, the complexes may not enter the flagellum in sufficient quantities to build a complete axoneme. Indeed, neither outer arm nor radial spoke proteins accumulate in
fla14 flagella (refer to Fig. 8 B), consistent with there being a block to entry of these structures into the flagellum.
Retrograde IFT Is Dependent on LC8
A dramatic difference between wild-type and fla14 flagella is the presence of massive amounts of raft material at the tip of the fla14 flagella. This undoubtedly is due to the loss of retrograde IFT in the mutant; because anterograde IFT is present, the rafts are transported to the distal end of the flagellum where they accumulate.
Recently, two new complexes, sedimenting at 15-16S
and together containing ~15 different polypeptides, have
been found in the membrane and matrix fraction of Chlamydomonas flagella (Cole, D.G., and J.L. Rosenbaum. 1996. Mol. Biol. Cell. 7:47a; Piperno and Mead, 1997; Cole et al.,
1998
). These complexes are lost from the flagella concomitantly with the anterograde motor FLA10 kinesin-II when
the temperature-sensitive mutant fla10 is shifted to nonpermissive temperatures. As a result, it has been proposed
that the complexes are the cargo of the FLA10 kinesin-II motor. Because the rafts are moved by IFT (Kozminski et al.,
1995
), the 15-16S complexes may be subunits of the rafts.
The fact that the rafts are highly enriched in fla14 flagella provided an opportunity to test this hypothesis. Using antibodies specific for four different polypeptides of
complexes A and B, we found that all four of these proteins were greatly elevated in fla14 flagella. As expected,
the proteins were present primarily in the membrane plus
matrix fraction. These results strongly support the notion
that the 15-16S complexes are raft components (Cole et al.,
1998). That anterograde IFT continues unabated in the fla14 mutants despite the accumulation of large numbers
of rafts at the tip of the flagella suggests that the cell body
contains a large pool of raft components. This is consistent
with immunolocalization studies showing that proteins of
the 15-16S complexes are highly enriched in the basal
body region (Cole et al., 1998
).
Cytoplasmic Dynein Is Likely To Be the Motor for Retrograde IFT
Retrograde IFT moves particles toward the base of the flagellum, where the minus ends of the flagellar microtubules
are located. The motor driving retrograde IFT is unknown, but has been suggested to be either cytoplasmic
dynein or a kinesin that moves towards the minus ends of
microtubules (Kozminski et al., 1995). Inasmuch as cytoplasmic dynein contains LC8 (King et al., 1996
), our finding that deletion of the LC8 gene disrupts retrograde IFT
points to cytoplasmic dynein as the most likely motor. It
should be noted that loss of the outer arms and inner arms
in the LC8 mutants cannot be responsible for the loss of
retrograde IFT, since IFT is normal in mutants that specifically lack either or both of these structures (Kozminski,
1993, and Fig. 6 A of this article).
The involvement of cytoplasmic dynein in retrograde
IFT is further supported by studies of C. elegans mutants.
In the nematode, the ends of the sensory neurons are modified cilia (Perkins et al., 1986; White et al., 1986
). Defects
in the C. elegans CHE-3 gene, which encodes a dynein
heavy chain (DHC) (Grant, W., personal communication,
and see below), cause the tips of the ciliary neurons to become swollen and packed with electron-dense material
(Lewis and Hodgkin, 1977
; Albert et al., 1981
). The OSM-6
gene product, which is homologous to the p52 polypeptide
of the Chlamydomonas 15-16S complex B (Cole et al.,
1998
), is massively accumulated at the tips of the che-3
neurons (Collet et al., 1998
). Therefore, the electron-dense material in the tips of the C. elegans che-3 neurons is
likely to correspond to the rafts of Chlamydomonas flagella. The ciliated tips of the neurons also are much
shorter in che-3 than in wild-type worms (Lewis and
Hodgkin, 1977
). Thus, the che-3 mutation in C. elegans appears to cause a phenotype closely related to that reported
here for the Chlamydomonas fla14 mutant.
The occurrence of similar defects in organisms as evolutionarily distant as Chlamydomonas and Caenorhabditis
suggest that the underlying processpresumably retrograde IFT
is widely distributed among organisms. Likewise, the occurrence of similar defects in organelles as different as the motile flagellum and the sensory neuron
suggests that IFT is of importance in a wide range of cilia-based structures. This is likely to include the rods and
cones of the vertebrate retina, which are modified cilia and
contain a FLA10 kinesin-II homologue (Beech et al., 1996
).
Because C. elegans has no motile cilia and the doublet
microtubules of the ciliary neurons lack dynein arms, the
CHE-3 gene product must be a component of cytoplasmic
dynein. In a phylogenetic tree based on the predicted
amino acid sequences of members of the DHC family, the
C. elegans CHE-3 gene product clustered with the DHC1b
isoform from human, rat, mouse, sea urchin, and Tetrahymena (Pazour, G., and G. Witman, unpublished data).
DHC1b is more closely related to the conventional cytoplasmic DHC isoform (DHC1a) than to known axonemal
DHCs (Gibbons et al., 1994; Tanaka et al., 1995
; Porter et
al., 1996
), yet its expression is induced by deciliation (Gibbons et al., 1994
) and it accumulates at the apical ends of ciliated cells (Criswell et al., 1996
). A likely explanation
for these observations is that DHC1b is a cytoplasmic dynein that operates in cilia and flagella. Retrograde IFT
may be powered specifically by DHC1b.
Evidence that defects in LC8 cause defects in cytoplasmic dynein function is provided by a comparison of LC8
mutants and cytoplasmic dynein mutants in the fly Drosophila melanogaster and in the fungus Aspergillus nidulans. In Drosophila, strong LC8 alleles are embryonic lethals, whereas weak alleles cause sterility and defects in
the wings and bristles (Dick et al., 1996; Phillis et al.,
1996
). Similarly, DHC1a deletions are homozygous lethals,
whereas weaker alleles result in sterility and bristle and
eye defects (Gepner et al., 1996
). In Aspergillus, LC8
(nudG) and cytoplasmic DHC (nudA) mutations both affect nuclear migration (Xiang et al., 1994
; Beckwith, S.M.,
and N.R. Morris. 1995. Mol. Biol. Cell. 6:5a). Thus, in both
Drosophila and Aspergillus, the phenotypes of LC8 mutations closely resemble those of cytoplasmic DHC mutations. Assuming that LC8 is associated with both DHC1a
and DHC1b, defects in LC8 are likely to affect the functions of both isoforms.
The above arguments notwithstanding, one cannot yet rule out a role for a minus-end-directed kinesin in retrograde IFT. Chlamydomonas flagella contain several proteins that react with antisera against highly conserved kinesin sequences. One of these is an ~55-kD membrane plus matrix protein missing in fla14 flagella (Fig. 9 C), a pattern that might be expected for the IFT motor. Further study will be necessary to determine if this is a motor protein.
Retrograde IFT and Retrograde Surface Motility Are Powered by Different Motors
The discovery of IFT (Kozminski et al., 1993) raised the
possibility that the movement of beads in flagellar surface
motility (Bloodgood, 1989
) simply reflected the movement
of rafts to which the beads had become attached via a
transmembrane protein. Previous experiments to resolve
this have yielded ambiguous results. Both IFT and bead
movement ceased together within 60-90 min after elevation of fla10 cells to nonpermissive temperature, indicating
that both were dependent upon the anterograde IFT motor (Kozminski et al., 1995
); however, this dependency
might be due to a requirement for IFT to deliver the surface motor to the tip of the flagellum. Both IFT and bead
movement were inhibited by NaCl and sucrose (Kozminski et al., 1993
); this inhibition most likely was due to an
osmotic affect, which could have inhibited more than one process. Rafts and polystyrene beads are transported at
different rates (Kozminski et al., 1993
), but this may be
due to differences in the viscous drag acting on particles of
vastly different sizes. Low Ca2+ and EGTA blocked bead
movement without affecting IFT (Kozminski et al., 1993
),
although this might have been due to uncoupling of beads
from rafts. Our observation that retrograde bead movement continues in fla14 flagella even in the absence of retrograde IFT provides the strongest evidence to date that
these two types of movement are powered by different
motors. Possible candidates for the retrograde surface motility motor include the kinesin-like proteins that are not
affected by the loss of LC8 (Fig. 9).
The loss of LC8 does result in an abnormal accumulation of beads at the distal tip of the fla14 flagellum. Assuming that the motor responsible for bead movement is microtubule-based, this may occur because the accumulation of rafts at the tip of the flagellum pushes the membrane so far away from the axoneme that the surface motility motor can no longer span the gap between membrane and microtubules.
The Role of IFT
Observations on Chlamydomonas defective in the anterograde IFT motor FLA10 kinesin-II have shown that IFT is
essential for flagellar assembly. As discussed above, anterograde IFT may be necessary for transporting specific
axonemal components to the flagellar tip before their assembly into the axoneme. Whether retrograde IFT itself
has a role in flagellar assembly is less clear. Although the
flagella of fla14 mutants are short and paralyzed, this could
be a secondary consequence of loss of the radial spokes and inner and outer arms. Mutants with specific defects in
the radial spokes have paralyzed flagella (Witman et al.,
1978), and double mutants with defects in both inner and
outer arms have short flagella and are nonmotile (Brokaw
and Kamiya, 1987
; Kamiya et al., 1989
; Kurimoto and Kamiya, 1991
). It also is possible that axonemal assembly and
flagellar growth in fla14 strains are not affected by loss of
retrograde IFT per se, but by a defect in the transport of
axonemal precursors in the cytoplasm to the region of the
basal body, or from there into the flagellum (see above).
In related work, Morris and Scholey (1997) showed that
microinjection of kinesin-II antibodies into fertilized sea
urchin eggs blocked ciliogenesis. The injected embryos assembled only short, immotile flagella that lacked a central
pair. It is likely that this is due to an effect of the antibody
on IFT, and that IFT also is necessary for cilia formation in
sea urchin cells.
Although the importance of IFT in flagellar assembly is established, the purpose of this process in nongrowing flagella is less well understood. When fla10 cells are shifted to nonpermissive temperatures, their flagella slowly shorten. Similarly, the flagella of fla14 cells shorten throughout the day. Therefore, both anterograde and retrograde IFT are necessary for maintaining flagellar length. IFT may supply some component essential for axonemal stability or maintenance. Perhaps the continuous active movement of large numbers of rafts to and fro in the confined space between the doublet microtubules and the flagellar membrane stirs the flagellar matrix, facilitating mass transport of molecules along the flagellar shaft.
It is not clear why the flagella of fla14 cells form within the mother cell wall, but then shorten after hatching. The findings reported here indicate that retrograde IFT is necessary for returning the rafts to the base of the flagellum. By extension, retrograde IFT is likely to be involved in recycling of other components that are transported to the tip of the flagellum. The lack of recycling of certain components may result in a deficiency that accumulates with time, eventually affecting flagellar stability.
IFT also is likely to have an important role in the maintenance of the flagellar membrane. A 65-kD membrane
protein is one of the major species transported to the flagellum of nonregenerating Chlamydomonas cells (Remillard and Witman, 1982). Turnover of membrane proteins
probably represents replacement of proteins lost by blebbing of membrane vesicles from the flagellar tip (Bergman et al., 1975
), and failure to maintain the membrane could
have deleterious effects on axonemal stability.
Future Directions
Because LC8 is associated with multiple polypeptide complexes, it is not surprising that deletion of the LC8 gene has pleiotrophic effects. Importantly, loss of the chain is not lethal in Chlamydomonas. As a result, it should be quite feasible to learn more about the role of LC8 and the complexes of which it is a part in this organism. It also should now be possible to isolate mutants with defects in other polypeptides of the retrograde motor; these should permit the effects of loss of retrograde IFT per se to be distinguished from other effects due to loss of LC8.
![]() |
Footnotes |
---|
Received for publication 30 January 1998 and in revised form 6 April 1998.
Address all correspondence to George Witman, Department of Cell Biology, UMMC, Worcester Foundation Campus, 222 Maple Ave., Shrewsbury, MA 01545. Tel.: (508) 842-8921 Ext. 344. Fax: (508) 842-3915. E-mail: witman{at}sci.wfbr.eduWe thank J. Aghajanian (Worcester Foundation for Biomedical Research, Shrewsbury, MA) for electron microscopy, and D. Cole and J. Rosenbaum for discussion and for sharing unpublished results. Antibodies were graciously provided by D. Cole and J. Rosenbaum, P. Yang and W. Sale, S. King, G. Piperno, and C. Walczak.
These studies were supported by grants from the National Institutes of Health (GM30626) and the Campbell and Hall Charity Fund.
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Note Added in Proof |
---|
We recently have identified an insertional mutant of Chlamydomonas in which the gene for the dynein heavy chain isoform DHC1b is deleted. The mutant has short flagella, confirming that DHC1b is important for flagellar assembly.
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Abbreviations used in this paper |
---|
DHC, dynein heavy chain; DIC, differential interference contrast; IFT, intraflagellar transport.
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References |
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---|
1. | Albert, P.S., S. Brown, and D.L. Riddle. 1981. Sensory control of dauer larva formation in Caenorhabditis elegans. J. Comp. Neurol. 198: 435-451 |
2. |
Beech, P.S.,
K. Pagh-Roehl,
Y. Noda,
N. Hirokawa,
B. Burnside, and
J.L. Rosenbaum.
1996.
Localization of kinesin superfamily proteins to the connecting cilium of fish photoreceptors.
J. Cell Sci.
109:
889-897
|
3. |
Benashski, S.E.,
A. Harrison,
R.S. Patel-King, and
S.M. King.
1997.
Dimerization of the highly conserved light chain shared by dynein and myosin V.
J.
Biol. Chem.
272:
20929-20935
|
4. | Bergman, K., U.W. Goodenough, D.A. Goodenough, J. Jawitz, and H. Martin. 1975. Gametic differentiation in Chlamydomonas reinhardtii. II. Flagellar membranes and the agglutination reaction. J. Cell Biol. 67: 606-622 [Abstract]. |
5. |
Bloch, M., and
K.A. Johnson.
1995.
Identification of a molecular chaperone in
the eukaryotic flagellum and its localization to the site of microtubule assembly.
J. Cell Sci.
108:
3541-3545
|
6. | Bloodgood, R.A. 1989. Gliding Motility and Flagellar Glycoprotein Dynamics in Chlamydomonas. In Ciliary and Flagellar Membranes. R. Bloodgood, editor. Plenum Press, New York. 91-128. |
7. | Brokaw, C., and R. Kamiya. 1987. Bending patterns of Chlamydomonas flagella. IV. Mutants with defects in inner and outer dynein arms indicate differences in dynein arm function. Cell Motil. Cytoskel. 8: 68-75 |
8. |
Cole, D.G.,
D.R. Diener,
A.L. Himelblau,
P.L. Beech,
J.C. Fuster, and
J.L. Rosenbaum.
1998.
Chlamydomonas kinesin-II-dependent intraflagellar
transport (IFT): IFT particles contain proteins required for ciliary assembly
in Caenorhabditis elegans sensory neurons.
J. Cell Biol.
141:
993-1008
|
9. |
Collet, J.,
C.A. Spike,
E.A. Lundquist,
J.E. Shaw, and
R.K. Herman.
1998.
Analysis of osm-6, a gene that affects sensory cilium structure and sensory
neuron function in Caenorhabditis elegans.
Genetics
148:
187-200
|
10. |
Criswell, P.S.,
L.E. Ostrowski, and
D.J. Asai.
1996.
A novel cytoplasmic dynein
heavy chain: expression of DHC1b in mammalian ciliated epithelial cells.
J.
Cell Sci.
109:
1891-1898
|
11. | Dick, T., K. Ray, H.S. Salz, and W. Chia. 1996. Cytoplasmic dynein (ddlc1) mutations cause morphogenetic defects and apoptotic cell death in Drosophila melanogaster. Mol. Cell. Biol. 16: 1966-1977 [Abstract]. |
12. | Fernandez, E., R. Schnell, L.P.W. Ranum, S.C. Hussey, C.D. Silflow, and P.A. Lefebvre. 1989. Isolation and characterization of the nitrate reductase structural gene of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA. 86: 6449-6453 [Abstract]. |
13. |
Fox, L.A.,
K. Sawin, and
W.S. Sale.
1994.
Kinesin-related proteins in eukaryotic
flagella.
J. Cell Sci.
107:
1545-1550
|
14. |
Gepner, J.,
M. Li,
S. Ludmann,
C. Kortas,
K. Boylan,
S.J.P. Iyadurai,
M. McGrail, and
T.S. Hays.
1996.
Cytoplasmic dynein function is essential in
Drosophila melanogaster.
Genetics.
142:
865-878
|
15. | Gibbons, B.H., D.J. Asai, W.-J.Y. Tang, T.S. Hays, and I.R. Gibbons. 1994. Phylogeny and expression of axonemal and cytoplasmic dynein genes in sea urchins. Mol. Biol. Cell 5: 57-70 [Abstract]. |
16. | Goodenough, U., and J. Heuser. 1984. Structural comparison of purified dynein proteins with in situ dynein arms. J. Mol. Biol. 180: 1083-1118 |
17. | Gumpel, N.J., and S. Purton. 1994. Playing tag with Chlamydomonas. Trends Cell Biol. 4: 299-301 . |
18. | Harris, E.H. 1989. The Chlamydomonas Sourcebook. Academic Press, Inc., San Diego, CA. 780 pp. |
19. |
Harrison, A.,
P. Olds-Clarke, and
S.M. King.
1998.
Identification of the t complex-encoded cytoplasmic dynein light chain Tctex1 in inner arm I1 supports
the involvement of flagellar dyneins in meiotic drive.
J. Cell Biol.
140:
1137-1147
|
20. |
Hoffmann, K.F., and
M. Strand.
1996.
Molecular identification of a Schistosoma
mansoni tegumental protein with similarity to cytoplasmic dynein light
chains.
J. Biol. Chem.
271:
26117-26123
|
21. | Hoops, H.J., and G.B. Witman. 1983. Outer doublet heterogeneity reveals structural polarity related to beat direction in Chlamydomonas flagella. J. Cell Biol. 97: 902-908 [Abstract]. |
22. |
Jaffrey, S.R., and
S.H. Snyder.
1996.
PIN: An associated protein inhibitor of
neuronal nitric oxide synthase.
Science.
274:
774-777
|
23. | Johnson, K.A., and J.L. Rosenbaum. 1992. Polarity of flagellar assembly in Chlamydomonas. J. Cell Biol. 119: 1605-1611 [Abstract]. |
24. |
Johnson, K.A.,
M.A. Haas, and
J.L. Rosenbaum.
1994.
Localization of a kinesin-related protein to the central pair apparatus of the Chlamydomonas reinhardtii flagellum.
J. Cell Sci.
107:
1551-1556
|
25. | Kamiya, R.. 1988. Mutations at twelve independent loci result in absence of outer dynein arms in Chlamydomonas reinhardtii. J. Cell Biol. 107: 2253-2258 [Abstract]. |
26. | Kamiya, R., E. Kurimoto, H. Sakakibara, and T. Okagaki. 1989. A genetic approach to the function of inner and outer arm dynein. In Cell Movement. Vol. 1. The Dynein ATPases. F. Warner, P. Satir, and I. Gibbons, editors. Alan R. Liss, Inc., New York. 209-218. |
27. | Kindle, K.L.. 1990. High-frequency nuclear transformation of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA. 87: 1228-1232 [Abstract]. |
28. |
King, S.M., and
R.S. Patel-King.
1995.
The Mr = 8,000 and 11,000 outer arm dynein light chains from Chlamydomonas flagella have cytoplasmic homologues.
J. Biol. Chem.
270:
11445-11452
|
29. | King, S.M., T. Otter, and G.B. Witman. 1985. Characterization of monoclonal antibodies against Chlamydomonas flagellar dyneins by high-resolution protein blotting. Proc. Natl. Acad. Sci. USA. 82: 4717-4721 [Abstract]. |
30. |
King, S.M,
C.G. Wilkerson, and
G.B. Witman.
1991.
The Mr 78,000 intermediate chain of Chlamydomonas outer arm dynein interacts with ![]() |
31. |
King, S.M.,
E. Barbarese,
J.F. Dillman III,
R.S. Patel-King,
J.H. Carson, and
K.K. Pfister.
1996.
Brain cytoplasmic and flagellar outer arm dyneins share a
highly conserved Mr 8,000 light chain.
J. Biol. Chem.
271:
19358-19366
|
32. |
Koutoulis, A.,
G.J. Pazour,
C.G. Wilkerson,
K. Inaba,
H. Sheng,
S. Takada, and
G.B. Witman.
1997.
The Chlamydomonas reinhardtii ODA3 gene encodes a
protein of the outer dynein arm docking complex.
J. Cell Biol.
137:
1069-1080
|
33. | Kozminski, K.G., K.A. Johnson, P. Forscher, and J.L. Rosenbaum. 1993. A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA. 90: 5519-5523 [Abstract]. |
34. | Kozminski, K.G., P.L. Beech, and J.L. Rosenbaum. 1995. The Chlamydomonas kinesin-like protein FLA10 is involved in motility associated with the flagellar membrane. J. Cell Biol. 131: 1517-1527 [Abstract]. |
35. | Kurimoto, E., and R. Kamiya. 1991. Microtubule sliding in flagellar axonemes of Chlamydomonas mutants missing inner- or outer-arm dynein: Velocity measurements on new types of mutants by an improved method. Cell Motil. Cytoskel. 19: 275-281 |
36. | Levine, R.P., and W.T. Ebersold. 1960. The genetics and cytology of Chlamydomonas. Annu. Rev. Microbiol. 14: 197-216 . |
37. | Lewis, J.A., and J.A. Hodgkin. 1977. Specific neuroanatomical changes in chemosensory mutants of the nematode Caenorhabditis elegans. Comp. Neur. 172: 489-510 . |
38. |
Morris, R.L., and
J.M. Scholey.
1997.
Heterotrimeric kinesin-II is required for
the assembly of motile 9+2 ciliary axonemes on sea urchin embryos.
J. Cell
Biol.
138:
1009-1022
|
39. |
Moss, A.G.,
J.-L. Gatti, and
G.B. Witman.
1992.
The motile ![]() |
40. | Pazour, G.J., O.A. Sineshchekov, and G.B. Witman. 1995. Mutational analysis of the phototransduction pathway of Chlamydomonas reinhardtii. J. Cell Biol. 131: 427-440 [Abstract]. |
41. | Perkins, L.A., E.M. Hedgecock, J.N. Thomson, and J.G. Culotti. 1986. Mutant sensory cilia in the nematode Caenorhabditis elegans. Dev. Biol. 117: 456-487 |
42. | Pfister, K.K., R.B. Fay, and G.B. Witman. 1982. Purification and polypeptide composition of dynein ATPases from Chlamydomonas flagella. Cell Motil. 2: 525-547 |
43. |
Phillis, R.,
D. Statton,
P. Caruccio, and
R.K. Murphey.
1996.
Mutations in the 8 kDa dynein light chain gene disrupt sensory axon projection in the Drosophila imaginal CNS.
Development.
122:
2955-2963
|
44. | Piperno, G., and D.J.L. Luck. 1979. Axonemal adenosine triphosphatases from flagella of Chlamydomonas reinhardtii. Purification of two dyneins. J. Biol. Chem. 254: 3084-3090 |
45. |
Piperno, G., and
K. Mead.
1997.
Transport of a novel complex in the cytoplasmic matrix of Chlamydomonas flagella.
Proc. Natl. Acad. Sci. USA.
94:
4457-4462
|
46. |
Piperno, G.,
M. LeDizet, and
X.-j. Chang.
1987.
Microtubules containing acetylated ![]() |
47. | Piperno, G., K. Mead, and S. Henderson. 1996. Inner dynein arms but not outer dynein arms require the activity of kinesin homologue protein KHP1Fla10 to reach the distal part of the flagella in Chlamydomonas. J. Cell Biol. 133: 371-379 [Abstract]. |
48. |
Porter, M.,
J.A. Knott,
S.H. Myster, and
S.J. Farlow.
1996.
The dynein gene
family in Chlamydomonas reinhardtii.
Genetics
144:
569-585
|
49. | Remillard, S.P., and G.B. Witman. 1982. Synthesis, transport, and use of specific flagellar proteins during flagellar regeneration in Chlamydomonas. J. Cell Biol. 93: 615-631 [Abstract]. |
50. | Sager, R., and S. Granick. 1953. Nutritional studies with Chlamydomonas reinhardtii. Annu. NY Acad. Sci. 56: 831-838 . |
51. | Sambrook, J., E.F. Fritsch, and T. Maniatis. 1987. Molecular Cloning: A Laboratory Manual. Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY. 545 pp. |
52. | Segal, R.A., B. Huang, Z. Ramanis, and D.J.L. Luck. 1984. Mutant strains of Chlamydomonas reinhardtii that move backwards only. J. Cell Biol. 98: 2026-2034 [Abstract]. |
53. |
Tam, L.-W., and
P.A. Lefebvre.
1993.
Cloning of flagellar genes in Chlamydomonas reinhardtii by DNA insertional mutagenesis.
Genetics
135:
375-384
|
54. |
Tanaka, Y.,
Z. Zhang, and
N. Hirokawa.
1995.
Identification and molecular
evolution of new dynein-like protein sequences in rat brain.
J. Cell Sci.
108:
1883-1893
|
55. | Walther, Z., M. Vashishtha, and J.L. Hall. 1994. The Chlamydomonas FLA10 gene encodes a novel kinesin-homologous protein. J. Cell Biol. 126: 175-188 [Abstract]. |
56. | White, J.G., E. Southgate, J.N. Thomson, and S. Brenner. 1986. The nervous system of Caenorhabditis elegans. Phil. Trans. R. Soc. Lond. 314B:1-340. |
57. | Wilkerson, C.G., S.M. King, A. Koutoulis, G.J. Pazour, and G.B. Witman. 1995. The 78,000 Mr intermediate chain of Chlamydomonas outer arm dynein is a WD-repeat protein required for arm assembly. J. Cell Biol. 129: 169-178 [Abstract]. |
58. | Williams, B.D., D.R. Mitchell, and J.L. Rosenbaum. 1986. Molecular cloning and expression of flagellar radial spoke and dynein genes of Chlamydomonas. J. Cell Biol. 103: 1-11 [Abstract]. |
59. | Williams, B.D., M.A. Velleca, A.M. Curry, and J.L. Rosenbaum. 1989. Molecular cloning and sequence analysis of the Chlamydomonas gene coding for radial spoke protein 3: Flagellar mutation pf-14 is an ochre allele. J. Cell Biol. 109: 235-245 [Abstract]. |
60. | Witman, G.B.. 1986. Isolation of Chlamydomonas flagella and flagellar axonemes. Methods Enzymol. 134: 280-290 |
61. |
Witman, G.B.,
K. Carlson,
J. Berliner, and
J.L. Rosenbaum.
1972.
Chlamydomonas flagella.
J. Cell Biol.
54:
507-539
|
62. | Witman, G.B., J. Plummer, and G. Sander. 1978. Chlamydomonas flagellar mutants lacking radial spokes and central tubules. J. Cell Biol. 76: 729-747 [Abstract]. |
63. | Witman, G.B., C.G. Wilkerson, and S.M. King. 1994. The biochemistry, genetics, and molecular biology of flagellar dynein. In Microtubules. J. Hyams and C. Lloyd, editors. Wiley-Liss, Inc., New York. 229-249. |
64. | Xiang, X., S. Beckwith, and N.R. Morris. 1994. Cytoplasmic dynein is involved in nuclear migration in Aspergillus nidulans. Proc. Natl. Acad. Sci. USA. 91: 2100-2104 [Abstract]. |