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Vol. 14, Issue 8, 3192-3207, August 2003
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* Department of Cellular Biology, University of Georgia, Athens, Georgia
30602;
Department of Biology, York University, Toronto, Ontario, Canada M3J 1P3
Submitted March 21, 2003;
Accepted April 2, 2003
Monitoring Editor: Paul Matsudaira
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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; Sloboda,
2002). During IFT, protein complexes are transported between the
cell body and the tips of axonemes
(Kozminski et al.,
1993). Anterograde IFT is believed to be driven by kinesin-II
motors moving along outer doublet microtubules
(Kozminski et al.,
1995; Signor et al.,
1999). In Chlamydomonas, Tetrahymena, and mouse,
loss-of-function mutations showed a requirement for kinesin-II in assembly of
cilia and flagella (Walther et
al., 1994; Nonaka et
al., 1998; Brown et
al., 1999b). Kinesin-II mutations also affected some
nonmotile cilia, including chemosensory neurons in Caenorhabditis
elegans and connecting cilia in mouse retinal rod cells
(Shakir et al., 1993;
Marszalek et al.,
2000). Cytoplasmic type dyneins are thought to be the motors that
return IFT complexes, kinesin-II, and perhaps other ciliary components to the
cell body. In Chlamydomonas, the dynein light chain mutant,
fla14, and the dynein heavy chain mutant, dhc1b, both have
extremely shortened flagella with tips filled with IFT materials, supporting
the role of dynein in retrograde IFT
(Pazour et al., 1998;
Pazour et al., 1999).
In the C. elegans che3 dynein mutant, IFT is specifically blocked in
the ciliary portion of chemosensory neurons
(Signor et al.,
1999).
In Chlamydomonas the IFT particles are composed of complex A,
containing four proteins, and complex B, containing 11 proteins
(Piperno and Mead, 1997;
Cole et al., 1998).
Many of the identified IFT polypeptides revealed similarity with potential
counterparts in other organisms. The complex B subunits p52, p88, and p172 of
Chlamydomonas are homologous to the C. elegans proteins
OSM-6, OSM-5, and OSM-1, respectively, all of which are implicated in the
function of the nematode chemosensory cilia
(Cole et al., 1998). A
mutation in the mouse homolog of IFT88, Tg737, caused murine
polycystic kidney disease associated with shortening of cilia in the kidney
cells (Pazour et al.,
2000), as well as affected the formation of mouse photoreceptor
outer segments (Pazour et al.,
2002).
Despite considerable progress in the identification of the molecular
components of IFT and growing evidence of their essential contribution to the
assembly of cilia and flagella, the specific biochemical function of IFT is
unclear. Currently, the best supported hypothesis is that IFT particles serve
as platforms that carry axonemal components from the cell body to their sites
of incorporation at the tips of the growing axoneme
(Rosenbaum et al.,
1999; Marszalek and Goldstein,
2000; Sloboda,
2002). However, so far it has not been possible to establish a
direct molecular connection between the IFT particles, motors, and axonemal
structural subunits. Thus, the molecular function of IFT beyond movement of
IFT particles remains undocumented.
We showed that knocking out two partly redundant Tetrahymena
kinesin-II genes, KIN1 and KIN2, led to loss of existing
cilia and inability to assemble new cilia. Unexpectedly, kinesin-II null cells
also underwent frequent arrests in cytokinesis
(Brown et al., 1999b).
Localization studies and observations of living cells led us to propose that
kinesin-II is not directly involved in cleavage furrow constriction
(Brown et al., 1999b).
Instead, we suggested that loss of motility in kinesin-II null mutants blocks
the final separation of daughter cells, which involves a series of rotational
movements facilitating cell fission (rotokinesis)
(Brown et al.,
1999a).
Herein, we report cloning of a Tetrahymena homolog of
IFT52. The IFT52 component of complex B is required for assembly of
flagella in Chlamydomonas
(Brazelton et al.,
2001), and its homolog OSM-6 is involved in assembly of sensory
cilia in C. elegans (Collet et
al., 1998). Elimination of IFT52 from
Tetrahymena cells led to a phenotype identical to the kinesin-II null
phenotype, including lack of cilia and inability to complete cytokinesis.
Unexpectedly, we isolated spontaneous IFT52 null suppressors that
have short cilia at low cell densities and grow longer cilia at higher cell
densities. We show that this effect of high cell density can be mimicked by
creating hypoxic conditions. Thus, suppressor cells at high cell density
influence each other mainly by creating pericellular hypoxia, which in turn
stimulates assembly of cilia. To our knowledge, this is the first report on
isolation of suppressors of a mutation in the IFT gene and on implicating
extracellular signaling in ciliary assembly in a way that involves a known IFT
component.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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) supplemented with 100 U/ml penicillin, 100
µg/ml streptomycin, and 0.25 µg/ml amphotericin B. Culture growth and
conjugation were done as described previously
(Brown et al.,
1999b).
IFT52 Cloning and Sequence Analysis
Polymerase chain reaction (PCR) was used to amplify a homologue of IFT52
from total Tetrahymena DNA. Degenerate primers were designed from
published peptide sequence alignments (Cole
et al., 1998). These primers amplified a 750-base pair
fragment with an open reading frame homologous to IFT52. A 3.4-kb
HindIII fragment was subsequently cloned that encompasses 
1.2 kb
of coding region along with two introns and the 3'-noncoding region. A
cDNA (#213) was located in the Tetrahymena expressed sequence tag
database (Fillingham et al.,
2002;
http://www.cbr.nrc.ca/reith/tetra/tetra.html)
and was found to be identical to the corresponding genomic sequence. Multiple
sequence alignments were prepared using the PILEUP program in the University
of Wisconsin (Madison, WI) GCG software (UWGCG). Shading was created with
Box-shade software
(http://www.ch.embnet.org/software/BOX_form.html).
Sequence similarities were calculated using BESTFIT (UWGCG).
IFT52 Knockout
The plasmid pIFT52-1, carrying the 3.4-kb genomic HindIII fragment
of IFT52, was digested at the unique Bsu36I site of the coding
region. Blunt ends were created, and the neo2 cassette
(Gaertig et al.,
1994) was inserted creating pIFT52–1bsuneo2. For germline
disruption of IFT52, 15 µg of pIFT52-1bsuneo2 digested with
HindIII to release the insert was used to coat 6 mg of 1-µm gold
particles (Bio-Rad, Hercules, CA). CU428.1 and B2086.1 cells conjugating for 3
to 4.5 h were bombarded every 30 min with coated particles by using the
biolistic gun (Cassidy-Hanley et
al., 1997). Bombarded cells were incubated in SPP medium at
30°C and selected with 120 µg/ml paromomycin 5.5 h after the last shot.
A single transformant heterozygous in the germline for IFT52::neo2
was brought to homozygosity and knockout heterokaryons were constructed
(Cassidy-Hanley et al.,
1997). To bring the phenotype to expression, two knockout
heterokaryon strains (UG7G5 and UG7G6) were induced to conjugate and
individual mating pairs were isolated into MEPP.
Rescue Transformation with Green Fluorescent Protein (GFP)-tagged
IFT52
In an attempt to construct a cadmium-inducible system for expression of
IFT52, the IFT52 cDNA was amplified with addition of
HindIII and BamHI sites to the 5' and 3' ends of
the coding region, respectively. The resulting fragment was used to replace
the coding region of pMTTG1 (Shang et
al., 2002), creating pMTTIFT52B2. In pMTTIFT52B2, the
cadmium-inducible MTT1 promoter
(Shang et al., 2002)
controls the IFT52 coding region and both are flanked by the 5'
and 3'-untranslated regions of the Tetrahymena BTU1 gene,
allowing insertion into the BTU1 locus as described previously
(Gaertig et al.,
1999). To create a GFP-tagged IFT52, PCR was used to add
a HindIII site to the 5' end and MluI, NcoI, and
BamHI sites to the 3' end of the IFT52 cDNA coding
region. The resulting fragment was digested with HindIII and
BamHI and inserted into pMTTG1 to create pMTTIFT52–3enz. A
MluI-NcoI-flanked GFP fragment
(Haddad and Turkewitz, 1997)
was inserted into digested pMTTIFT52–3enz to create pMTTIFT52GFP, which
encodes an IFT52 protein with a C-terminal GFP tag. IFT52
cells grown
in MEPP were biolistically bombarded with the 3.4-kb genomic IFT52 fragment
released from pIFT52-1, the 5'BTU1-IFT52coding-3'BTU1
(pMTTIFT52B2), or the
5'BTU1–5'MTT1-IFT52coding-GFP-3'BTU1 fragment released
from pMTTIFT52GFP. No drug selection was required because only transformants
recovered motility and normal cytokinesis and therefore could grow on the SPP
medium.
Immunocytochemistry and Electron Microscopy
Cells were prepared for confocal analysis as described previously
(Brown et al., 1999b)
with some modifications. Briefly, cells were isolated into drops of 10 mM Tris
pH 7.5 on poly-L-lysine–coated coverslips. For staining with
SG serum against total Tetrahymena tubulin
(Guttman and Gorovsky, 1979),
cells were simultaneously fixed and permeabilized and were then air dried at
30°C. For staining with anti-GFP rabbit polyclonal antibodies (BD
Biosciences Clontech, Palo Alto, CA), cells were permeabilized for a few
seconds in 0.5% Triton X-100 in the PHEM buffer with protease inhibitors (0.5
µg/ml leupeptin, 10 µg/ml E-64, 10 µg/ml chymostatin, 12.5 µg/ml
antipain) and 1 µM paclitaxel, followed by fixation with an equal volume of
2% paraformaldehyde in PHEM and air drying at 30°C. Nuclei were stained
with either propidium iodide or 4,6-diamidino-2-phenylindole (DAPI). An MRC
600 (Bio-Rad) confocal microscope was used for imaging. Ciliary lengths were
measured on individual confocal sections using Scion Image (Scion, Frederick,
MD). For consistency, cilia were measured using the widest section through the
nucleus for a given Z-series. Scion Image was also used to measure the total
length of the cell periphery on the section from which ciliary lengths were
measured by using the freehand tool to trace the cell periphery and the
Measure Accumulated Perimeter macro. The total number of measurable cilia was
divided by length of cell periphery to calculate measurable cilia/µm.
For electron microscopy (EM), cells were washed with 10 mM Tris, pH 7.5 and fixed in 4% glutaraldehyde in 10 mM Tris buffer at 4°C for 1 h, washed three times with 10 mM Tris, and postfixed in 4% osmium tetroxide for 1 h at 4°C. Cells were embedded in Epon after dehydration in graded steps from 30 to 100% ethanol. Sections were stained with uranyl acetate and lead citrate and were visualized on a 100CXII transmission electron microscope (JOEL, Tokyo, Japan).
Phenotypic Analysis
For dilution experiments, IFT52sm1 cells growing in MEPP with
shaking at 160 rpm at 30°C were washed and resuspended in fresh MEPP at a
concentration of 3 x 105/ml. Serial dilutions were prepared
and cells were incubated in 10 ml on Petri plates (10 cm diameter) with or
without gentle shaking (30 rpm) either at 30°C or at room temperature.
Cells were scored on an inverted microscope using a 10x objective
(100x total magnification) for motility and presence of the multiple
subcells indicative of cytokinesis failures. Cells were scored as motile if
there was clear cell body displacement. To account for growth of cells without
successful cytokinesis, average number of cortical subcells per milliliter was
calculated. First, average number of cells per 10x field of view on an
inverted microscope was converted to cells per milliliter by calibrating the
microscopic counts using a Beckman Coulter (Fullerton, CA) model ZF cell
counter. Average subcells per cell was calculated and multiplied by average
cells per milliliter to obtain subcells per milliliter. Stimulation of ciliary
assembly between strains was addressed by mixing IFT52sm1 cells with
IFT5210 or wild-type (OC21-12) cells in different proportions. For
stimulation by mutant cells, motility was quantified as described above. For
filter assays, 1.5 ml of potential stimulator cells were separated from 0.5 ml
of responder cells by a 3-µm pore filter (using Transwell inserts, 12 mm
diameter; Corning Costar Corp., Cambridge, MA) inserted in a 24-well tissue
culture plate well. After 24-h incubation, responder cells (from the inserts)
were scored for motility and subcells per chain. Statistical significance was
assessed with one- and two-tailed t tests.
For induction of partial hypoxia, IFT52sm7 cells were grown in MEPP,
washed, and suspended at 1.5 x 105 cells/ml in 3 ml of MEPP
supplemented with 0.2% normocin antibiotic (InvivoGen, San Diego, CA). Cells
were transferred to a 125-ml bottle with a ground glass closure covered with
vacuum grease, and exposed to a stream of nitrogen gas for about a minute. As
a control, another batch of cells was exposed to a stream of oxygen gas. The
bottles were closed tightly and incubated at 30°C with shaking at 130
rpm.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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). The IFT52 coding region encodes a
protein, Ift52p, of 434 amino acids with calculated molecular mass and pI of
49 kDa and 4.99, respectively. The predicted Ift52p protein sequence is 48%
identical to IFT52 of Chlamydomonas and 35% identical to OSM-6 of
C. elegans (Figure
1A).
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Cells Lacking Ift52p Do Not Assemble or Maintain Cilia and Are Unable
to Complete Cytokinesis
We created knockout heterokaryons (Hai
and Gorovsky, 1997) with disrupted copies of IFT52 in the
transcriptionally silent micronucleus (MIC) and wild-type (WT) copies in the
transcriptionally active macronucleus (MAC)
(Figure 1, B and C). These
heterokaryons maintain a WT phenotype during vegetative growth, because only
the genes in the MAC are expressed. When heterokaryons are mated to each
other, the progeny develop a new MAC from the MIC, thus expressing the
IFT52 null phenotype. Remarkably, the progeny (IFT52) lost
cilia with very similar timing to the loss of cilia in previously reported
kinesin-II knockouts (Brown et
al., 1999b). Anti-tubulin immunofluorescence revealed that
most cells nearly completely lost cilia by 30 h
(Figure 2). The IFT52
mutants died on the standard SPP medium but could grow in MEPP, which bypasses
the requirement for phagocytosis dependent on the oral cilia
(Rasmussen and Orias, 1975).
Many mutants consisted of multiple cortical "subcells" and sets of
nuclei (Figure 2, D–F),
showing that they fail to complete cytokinesis. This phenotype is essentially
identical in both the temporal progression and terminal morphology to the
phenotype of cells lacking kinesin-II motor subunits
(Brown et al., 1999b).
Thus, as kinesin-II, Ift52p is required for assembly and maintenance of cilia,
and both proteins likely function in the same pathway.
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GFP-tagged Ift52p Localizes to Cilia
We prepared a chimeric gene encoding an Ift52p-GFP fusion. We placed the
Ift52p-GFP coding region under the control of the cadmium inducible
MTT1 promoter and targeted the whole fragment into the nonessential
BTU1 locus (Shang et
al., 2002). Surprisingly, IFT52 cells could be rescued
with either the MTT1-IFT52 or MTT1-IFT52-GFP fragments without the addition of
cadmium. Attempts to remove possible traces of cadmium from the medium by
extensive washing and incubation in ultrapure water never resulted in a loss
of motility. It is known that in the absence of cadmium, the basal level of
expression controlled by the MTT1 promoter is extremely low
(Shang et al., 2002).
Thus, a very low level of production of Ift52p is sufficient for assembly of
cilia, suggesting that either the protein is needed in small amounts or that
it is very stable and can be recycled.
Staining with polyclonal anti-GFP antibodies revealed a predominant localization of Ift52p to cilia (Figure 3). In dividing cells, there was no colocalization of Ift52p-GFP with the cleavage furrow (Figure 3, D–O). During early stages of cell division, clusters of short immature oral cilia were intensely labeled, suggesting that as for kinesin-II, IFT particle proteins preferentially accumulate within cilia in the initial phase of assembly (Figure 3, D–F).
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We have previously described an elaborate type of cell motility called
rotokinesis, which seems to help WT cells complete cytokinesis by creating
mechanical strain within the cytoplasmic bridge connecting daughter cells
(Brown et al., 1999a).
As in kinesin-II mutants, IFT52 knockout cells are unable to undergo
rotokinesis, suggesting that for both mutations the cytokinesis defect is
caused by cell paralysis. To further test the importance of cell motility for
the completion of cytokinesis, we grew IFT52 knockouts with and
without vigorous shaking for 48 h. Cells grown without shaking were
polynucleated indicating cytokinesis failures
(Figure 4B). On the other hand,
although cells grown with shaking did not recover motility, they had
dramatically fewer cytokinesis failures
(Figure 4A). Similar results
were obtained for kinesin-II mutants lacking KIN1 and KIN2
genes (our unpublished data). These results further confirm that cleavage
furrow ingression is not substantially impeded in Tetrahymena IFT
mutants and that displacement of cells by an external mechanical force is
sufficient for final separation of daughter cells.
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Spontaneous Partial Suppressors of IFT52
The knockout strains of IFT52 used in this study were generated by
mating two parental strains with isogenic MIC genomes derived by genomic
exclusion (Orias and Bruns,
1976). Thus, we expected that all heterokaryon progeny cells would
show the same phenotype. To our surprise, 3% (n = 240) of the progeny of
knockout heterokaryons showed a partial suppression of the IFT52
phenotype. Within the synclones (mixed progenies derived from both
exconjugants of a single pair), the suppressed phenotype occurred within the
first few divisions after conjugation and not all cells showed the suppressed
phenotype. After subcloning, the phenotype of both suppressed and
nonsuppressed cells was stable. In the suppressor clones designated as
IFT52sm (semimotile), cells had short but partly functional oral cilia
(unlike nonsuppressed, IFT52sm produced some food vacuoles), and very
short, scattered locomotory cilia allowing for minimal movements. The density
of identifiable cilia on confocal sections of the nonsuppressed IFT52
cells was only 0.07 cilia/µm of cell periphery compared with 0.28
cilia/µm in WT cells. Compared with the nonsuppressed population, in
IFT52sm cells there was a dramatic increase to 0.21 cilia/µm.
Although the measurable cilia on IFT52sm cells were slightly longer
(mean length 1.2 µm, p = 0.000012) than on IFT52 (mean length 0.83
µm), these cilia were still much shorter than WT (mean length 4.2 µm)
(Figure 5, A–C, and E).
We did not find any suppressors among the 288 progenies of mating
heterokaryons lacking kinesin-II genes
(Brown et al., 1999b).
Thus, the observed suppression does not seem to be a general property of IFT
mutants in Tetrahymena.
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During an extended period of growth (130 generations) a more complete
suppression of the mutant phenotype occurred spontaneously in one of the eight
IFT52sm clones (IFT52sm1) studied. We established a subclone
resulting from this apparent second suppression event called IFT52mov1
(moving IFT52). These cells recovered normal cilia density (0.29/µm)
and the measurable cilia were of intermediate length (2.9 µm) between WT
and IFT52sm1 cells (p = 2.6 x 10–20;
Figure 5, D and E). Attempts at
obtaining such clones from the remaining seven IFT52sm strains failed,
despite growing some of them continuously for 20 mo.
Thin section EM revealed that most basal bodies in IFT52 cells
completely lacked axonemes (Figure
6A). Although some basal bodies did have associated extremely
short cilia, these cilia lacked a central pair
(Figure 6B). The semimotile
suppressor IFT52sm1 had fewer naked basal bodies, but cilia were short
with only 13% having a central pair (Figure
6C and Table 1).
These short 9 + 2 cilia may be responsible for the partial recovery of
motility (Figure 6, D and E,
and Table 1). The further
suppression in IFT52mov1 cells is correlated not only with an increase
in ciliary length but also with a dramatic increase in the proportion of 9 + 2
axonemes (Figures 5D and
6F and
Table 1). Thus, relatively
frequent, spontaneous suppression events allow IFT52 null cells to
assemble cilia without the Ift52p protein.
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IFT52 Suppression Occurs by a Novel Mechanism That Does Not
Involve a Heritable Change in the Germline Genome
To study the genetic basis of the suppression in IFT52 cells, we
performed a series of crosses and analyzed segregation of suppressed and
nonsuppressed phenotypes. We could not use IFT52sm clones for genetic
analyses because they did not recover motility in the starvation medium and
therefore would not mate. For this reason, we used the more complete
suppressor, IFT52mov1 and crossed it to a WT strain. All F1 progenies
analyzed had a WT phenotype. Six F1 clones after reaching maturity were
crossed to each other in three combinations. A total of 408 conjugation
progenies were analyzed (after exclusion of cells, which retained the parental
MACs or died). Among them, 304 had WT phenotype, 103 had nonsuppressed
IFT52 phenotype, and 1 had the IFT52sm suppression phenotype.
This result is inconsistent with the presence of a suppressor locus in the
germline MIC. Rather, heterozygotes show a standard 3:1 F2 segregation ratio,
and the suppression seems to occur among a small number of homozygous mutant
progenies as a secondary event, possibly mediated by a genetic alteration in
the newly developed somatic MAC or a nongenetic mechanism based on
physiological adaptation.
Maintenance of Cilia in IFT52sm Suppressor Cells Is Cell
Density and Temperature Dependent
The IFT52sm1 cells could grow with shaking at 30°C to a maximal
cell density of 5 x 105 cells/ml, which is considerably lower
than WT. In a shaken culture, the majority of cells complete cytokinesis, and
cells have very limited motility, even at the maximal density. Surprisingly,
we discovered that high-density cultures left unshaken gradually recovered
cell motility. The extent of motility in unshaken cultures was strongly
dependent on cell density (Figure 7,
A–D). The absence of motility at low cell density was
especially striking 9.5 h after dilution in cells diluted 20–100 x
(1.5 x 104–3 x 103 cells/ml). Such
cultures completely lost motility by 9.5 h postdilution and remained immobile
at 21.5 h (Figure 7, B and C).
As cell density increased in these diluted cultures over time (33.5 h), some
motility returned (Figure 7D).
Even after cells recovered some motility with increased cell density,
dilutions that initially led to the most severe loss of motility had many more
cells with evidence of cytokinesis defects
(Figure 7D). The remaining
seven IFT52sm clones also showed an identical cell density-dependent
effect on motility. Cell density did not affect motility in the more advanced
suppressor IFT52mov1 (our unpublished data).
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Immunofluorescence analysis of IFT52sm1 cells 21.5 h after dilution
revealed that the extent of motility was correlated with the density and
length of cilia (Figure 8 and
Table 2). Cells from a culture
with an initial density of 1.5 x 105 cells/ml were covered
with scattered short cilia (Figure
8A). Cells initially diluted to between 3 x 104
and 6 x 103 cells/ml maintained fewer short cilia that often
seemed to be concentrated near the anterior end and oral apparatus of the cell
(Figure 8, B and C). The nearly
complete absence of motility in cells that were diluted to 3 x
103 cells/ml was associated with a drastic reduction in ciliary
length (Figure 8, D and E). In
these cells, the density of measurable cilia was only 60% of the density in
the culture that was 50x more concentrated. The measurable cilia were
also short, their mean length being only 48% of the mean length of cilia from
cells diluted to 1.5 x 105
(Table 2).
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The above-mentioned experiments were all performed at the standard
temperature of 30°C. Unexpectedly, IFT52sm1 unshaken cells grown at
room temperature (22°C) could assemble and maintain cilia and had an
accompanying return to nearly normal cell motility and cytokinesis even at low
initial cell density (Figure 8F and Table 2). The remaining
seven IFT52sm clones showed the same temperature sensitivity (our
unpublished data). Thus, the partial suppression of IFT52 phenotype is
conditional and dependent on both cell density and temperature.
Finally, the effects of cell density and temperature on ciliogenesis in
IFT52sm cells were fully reversible. Cells that initially were highly
ciliated (due to growth at low temperature or high cell density without
shaking) lost cilia when diluted to low density and subsequently regained
motility when incubated unshaken at high cell density.
Imbalance between Rapid Growth Rate and Slow Ciliary Assembly Does
Not Account for the Cell Density-dependent IFT52sm Phenotype
Increased ciliogenesis in IFT52sm cells at lower temperatures, and
higher cell densities raised the possibility that suppressors are able to
maintain longer cilia only under conditions when their growth rate is reduced.
Thus, there could be an imbalance between the rate of ciliary assembly and the
rate of cell growth. To address this possibility, we diluted cells to 3
x 103 cells/ml and grew them at 30°C in several
modifications of the MEPP medium containing progressively lower concentrations
of proteose-peptone. Although reducing the proteose-peptone concentration
drastically slowed the growth of IFT52sm cells
(Figure 7E), it had no effect
on the motility. Cells in all media showed the same dilution-dependent loss of
cilia and cytokinesis defects. In fact, at later times, the cells grown at
higher concentration of peptone had more motility
(Figure 7F). This can be
explained by the faster growth rate, which leads to higher cell density. The
simplest explanation of these data is that the stimulation of ciliogenesis is
directly dependent on increased cell density, suggesting that cell-cell
signaling may be involved.
Ciliary Assembly in IFT52sm Can Be Stimulated by Wild-Type and
Nonsuppressed IFT52 Cells at High Cell Densities
To test whether cell-cell communication is involved in maintaining cilia in
IFT52sm cells, we performed mixed strain experiments. When we mixed
IFT52sm1 cells diluted to a concentration that inhibits ciliogenesis
among the suppressor cells alone with a high concentration of WT cells,
motility was clearly stimulated in the suppressor cells. To test whether the
effect is mediated by ciliogenesis, we measured ciliary length on tubulin
antibody-stained cells. To be sure we were measuring cilia only on mutant
cells, we chose only cells that showed clear arrests in cytokinesis
(Figure 9D).
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Clearly, this method prevents us from measuring cilia on cells that recover
near WT assembly and motility. Even so, we found that cilia on identifiable
IFT52sm1 cells were 39% longer when diluted and mixed with WT (p = 8.69
x 10–39) than when they were only diluted
(Figure 9D). On the other hand,
IFT52 (nonsuppressed) cells were unable to respond to the
assembly-promoting signal generated by WT cells
(Figure 9D). Similar mixed
strain experiments showed that concentrated IFT52 cells stimulate
ciliary assembly in diluted IFT52sm cells (our unpublished data). These
results suggest that IFT52sm cells posses a reception ability that
responds to the presence of other cells, whereas IFT52 cells lack such
a mechanism. Thus, the suppression mechanism could involved increased
sensitivity of IFT52sm cells to an unknown external signal.
The data presented above do not distinguish between direct cell-cell
contact and mediation by a diffusible factor. To distinguish between these two
possibilities, we performed coculture experiments where cells potentially able
to stimulate ciliary assembly were separated by a 3-µm pore filter from
cells potentially able to respond to the signal. Under these conditions,
direct cell contact is prevented. A stimulation of motility of diluted
IFT52sm1 cells was observed when they were exposed to concentrated
IFT52sm1 cells compared with incubation with diluted IFT52sm1 or
culture medium alone. Furthermore, high concentration nonsuppressed mutants
were also able to stimulate assembly on the suppressors in the filter assay
(our unpublished data). Thus, the ciliogenesis promoting effect of
concentrated cells in IFT52sm background does not require direct
contact between cells.
The Ciliation-stimulating Factor Is Equivalent to Pericellular
Hypoxia
We first attempted to detect a potential positive regulator of ciliogenesis
in medium containing high-density cells. Unexpectedly, the spent medium from
IFT52sm1 cells grown at high density did not stimulate assembly of
cilia in IFT52sm1 at low cell density. Also, spent medium from WT and
IFT52mov1 cells did not have an effect on diluted IFT52sm1. In
contrast, we observed that any kind of spent medium decreased motility
compared with fresh medium. Thus, the putative stimulatory factor could be
extremely unstable. This hypothesis was reinforced by the results of shaking
of suppressor cultures. We grew IFT52sm1 cells on a plate at low
density at room temperature, either without shaking or with very gentle
shaking (30 rotations/min). Slow shaking led to slower growth and less of
motility (more cytokinesis defects), even in high-density cells kept at room
temperature (Figure 9A). At
30°C we observed that shaken cells had fewer cytokinesis defects compared
with unshaken cells. Strikingly, more motile cells were still present in the
unshaken culture compared with the shaken culture
(Figure 9A). This result can be
explained if we assume that at 30°C, increased membrane fluidity allows
even gentle shaking to help cells complete cytokinesis by bypassing
rotokinesis. Thus, the results of shaking at 30°C are remarkable. Even
though the culture that was shaken grew to higher cell density, these cells
did not recover robust motility, as did cells grown at the same temperature
without shaking. Confocal imaging showed that the decreased motility of cells
in the shaking cultures was due to a dramatic inability to assemble cilia
(Figure 9, B and C). These data
suggested that the signal acts locally and may be extremely unstable.
If an unstable, autocrine stimulatory factor is involved, a single
IFT52sm cell should autostimulate itself for assembly of cilia when
grown in a extremely small volume. We therefore grew single isolated
IFT52sm1 cells in 1-µl drops of medium. Unexpectedly, after 24 h (at
30°C) cells in such extremely small drops did not recover motility and
invariably showed extensive cytokinesis defects (n = 21). Furthermore, even
initially highly motile IFT52sm cells (grown at room temperature), lost
motility within a few hours after isolation into 1-µl drops kept at
30°C. This surprising result was inconsistent with involvement of a
positive, autocrine-type regulator of ciliation. Intrigued by the effect of
culture volume on ciliation, we prepared IFT52sm1 cell suspension at
high cell density and grew cells in drops of varying size. After 24 h in
1-µl drops, we observed no motility despite the fact that cells were
suspended at the high cell density (3 x 105 cells/ml)
(Figure 10A). However, the
extent of motility increased with the increase of drop volume to 20 µl and
increased further when cells where incubated in 10-ml volume on a Petri plate
(Figure 10A). Thus, reducing
the culture volume overrides the stimulatory effect of high cell density. A
similar stimulatory effect of increased culture volume was observed when
IFT52sm cells were grown with shaking in Erlenmayer flasks
(Figure 10B). As the volume of
the culture medium increases, the surface/volume ratio decreases. This result
taken together with the inhibitory effect of culture shaking strongly
suggested that either a volatile stimulatory factor is released, or a volatile
inhibitory factor is acquired from the atmosphere. We therefore considered
that the atmospheric oxygen was responsible for the suppression of motility in
small volume and in shaken cultures, due to increased aeration. To reduce the
concentration of dissolved oxygen, we briefly exposed the suspended
IFT52sm cells to a stream of nitrogen, and incubate them in an airtight
culture vessel for 9 h with shaking. Remarkably, the cell population exposed
to a nitrogen stream showed a dramatic increase in cell motility, despite
shaking (Figure 10C). The
stimulatory effect of nitrogen stream was independent on cell density.
Furthermore, the nitrogen-flashed cultures showed a reduction in the growth
rate, indicating that hypoxic conditions were created (our unpublished data).
The stimulating effect of hypoxia on cell motility was correlated with the
increase in the length and density of cilia, as determined by
immunofluorescence (Figure
10D). Importantly, the hypoxia-induced newly assembled cilia were
concentrated mostly at the anterior ends of cells, the localization that also
was predominant in the cells exposed to high cell density without shaking
under normoxic conditions (Figure 8, A and
B). These observations indicate that the effect of high cell
density is equivalent to the effect of pericellular hypoxia. Thus, the
inhibition of cell density-mediated suppression by shaking and small culture
volume is caused by increased oxygen concentration in the medium. In
accordance, cells in the culture exposed to a stream of oxygen and grown for 9
h, showed lower motility, even compared with cells kept at normoxic conditions
(Figure 10C).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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)
and the requirement for IFT-52 in assembly of flagella in
Chlamydomonas (Brazelton et
al., 2001). Furthermore, the entire phenotype of IFT52 null
is essentially identical to the phenotype of cells lacking subunits of
kinesin-II (Brown et al.,
1999b). These results are consistent with Ift52p being a cargo of
kinesin-II. The Ift52p localizes mainly to cilia and, as in the case of
kinesin-II, does not occur in the cleavage furrow
(Brown et al., 1999b).
Furthermore, the cytokinesis defects of IFT52 and kinesin-II mutants
could be overcome by vigorous shaking, indicating that the cleavage furrow
progresses almost to completion so that cell fission can be accomplished by
mechanical force without damaging cells. It seems that the loss of a novel
type of whole cell motility, called rotokinesis, is sufficient to explain the
cytokinesis defects in IFT mutants. Rotokinesis involves the persistent,
unidirectional rotation of only the posterior daughter cell around an axis
passing through the cytoplasmic bridge connecting the daughter cells. We have
proposed that the torque generated by rotokinesis may help weaken the
cytoplasmic bridge between presumptive daughter cells, thereby increasing the
likelihood that cytokinesis will be successful
(Brown et al., 1999a).
Recently, a null mutant of FLA10, a Chlamydomonas kinesin-II
subunit, was found not to be affected in cell division
(Matsuura et al.,
2002). All these data together indicate the components of IFT are
not directly involved in cell division.
It was unexpected that spontaneous partial suppressor cells were identified
at a relatively high frequency among progenies of the IFT52 knockout
heterokaryons. The result was surprising because both the suppressed and
nonsuppressed cells were derived from crosses of two genetically identical
parental strains. Given that the partial suppression phenotype occurred with
such high frequency and after only a few generations, it is unlikely that it
was caused by spontaneous mutations. When a new MAC is produced during
conjugation, the genome undergoes extensive rearrangement, which involves
deleting 10% of MIC-specific DNA, as well as chromosome fragmentation.
Importantly, these genome-processing events are sometimes nonidentical. In
some cases, deletions occur at alternative sites separated by considerable
distance (Howard and Blackburn,
1985; Austerberry and Yao,
1988; Chau and Orias,
1996). It has never been reported that such alternative processing
could affect protein-coding genes. However, our results suggest that
alternative genome processing may have contributed to the suppression in the
original IFT52 strain. At the end of conjugation, each
exconjugant has two MACs, which undergo genome processing events independently
(Nanney and Caughey, 1953).
During the first cell division after conjugation, the two MACs do not divide
and are transmitted to the progeny cells, giving the so-called caryonidal
lines, which may therefore have genetically nonidentical MACs. The fact that
only a subset of cells derived from the same conjugation pair of
IFT52 knockout heterokaryons showed a suppressed phenotype is
consistent with a caryonidal inheritance and with the suppression mechanism
originating from alternative MAC genome processing. We found that a single
protein is more abundant in suppressor cilia compared with wild type (our
unpublished data). Thus, alternative processing may have affected the level of
expression of this or perhaps additional proteins which could compensate for
loss of Ift52p.
The most surprising result was the observation that the phenotype of
intermediate suppressor cells was dependent on cell density. We subsequently
found that the stimulating effect of high cell density can be opposed by
increasing medium aeration, either by shaking culture or by increasing the
surface/volume ratio of culture medium. Furthermore ciliogenesis could be
stimulated by decreasing the concentration of oxygen in the medium, even at
low cell density. It seems therefore that the cell density effect is based
mainly if not entirely on creation of local hypoxic conditions. The exact mode
of influence of hypoxia on ciliogenesis is unknown. First, it is possible that
the conformation of a molecule(s) involved in suppression is sensitive to the
oxygen presence, for example due to its direct oxidation. Alternatively, the
suppression could be based on induction of gene expression by hypoxia. It is
now well described that hypoxic conditions lead to a response at the level of
transcription of genes whose products mediate adaptation to low oxygen
conditions (Semenza, 2000).
Furthermore, high cell density was recently found to be responsible for
induction of signaling responses typical of hypoxia in prostate cancer cells
maintained under standard culture generally considered as normoxic
(Sheta et al., 2001).
The signaling hypothesis requires an existence of sensory and signal
transduction mechanisms that respond to the oxygen levels. In this regard, it
is unclear whether hypoxia has any effect on the assembly of cilia in
wild-type cells. We have not observed an increase in cell motility in unshaken
wild-type cultures. Furthermore, the more advanced suppressor,
IFT52mov1, did not show cell density effects. Thus, it seems that the
suppression occurred in two stages, and only the initial stage is dependent on
hypoxia. On the other side, there is evidence for existence of mechanisms that
sense oxygen concentration among ciliates. When Paramecium cells
where exposed to hypoxia, they swam along a provided temperature gradient
toward lower temperature, which seems to be a survival adaptation allowing for
lowering the rate of metabolism (Malvin
and Wood, 1992). Thus, hypoxia can modify the parameters of
ciliary beat and therefore hypoxia-responsive signaling components are likely
to exist inside the cilium. It is unlikely however, that oxygen is a general
regulator of ciliary assembly, because both aerobic and anaerobic species of
protists can assemble the conserved 9 + 2 axoneme
(van Hoek et al.,
2000). More likely, the suppression event might have coupled the
ciliary assembly machinery to hypoxia-mediated signaling and such a direct
connection may not be functioning in normal cells. Interestingly, we showed
that lower temperature also leads to increased ciliation in the IFT52sm
suppressors. In yeast, up-regulation of the gene encoding D9 fatty acid
desaturase also occurs both in hypoxia and at lower temperature
(Nakagawa et al.,
2002). Our observations provide additional evidence that low
temperature and hypoxia intersect in cellular signaling. Regardless of the
exact mechanism of suppression, our results indicate that Ift52p and its
homologues in other organisms, play some role in signaling pathways regulating
the assembly of cilia. It seems not to be a mere coincidence that NGD5, the
murine homolog of Ift52p, was originally identified as a gene whose expression
was down-regulated after exposure to an agonist of the 
-opioid receptor
(Wick et al., 1995).
It is therefore possible that Ift52p is located at the crossroads of some
unknown signaling pathways that regulate IFT.
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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Footnotes |
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Abbreviations used: IFT, intraflagellar transport; MAC, macronucleus; MIC, micronucleus; WT, wild-type.
Present address: Department of Biology, Young Harris College, Young Harris,
GA 30582. 
Corresponding author. E-mail address:
jgaertig{at}cb.uga.edu.
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONACKNOWLEDGMENTSREFERENCES |
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nonshaken (n = 62 for unshaken at 30
and 22°C, 50 for shaken at 30°C, and 70 for shaken at 22°C). (B)
Representative IFT52