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Vol. 20, Issue 2, 708-720, January 15, 2009
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*Department of Biology, Harvey Mudd College, Claremont, CA 91711; Department of Zoology, Miami University, Oxford, OH 45056; and Howard Hughes Medical Institute, Chevy Chase, MD 20815
Submitted July 21, 2008;
Revised October 28, 2008;
Accepted November 7, 2008
Monitoring Editor: Paul Forscher
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Zariwala et al., 2007). For example, cilia are essential for Tetrahymena swimming, feeding, and cell division. The cytoskeleton of the cilium/flagellum is the axoneme comprising more than 600 proteins that assemble into the "9 + 2" arrangement of microtubules, outer and inner rows of dynein arms, and the radial spokes (Piperno et al., 1977; Pazour et al., 2005; Smith et al., 2005).
Because of the essential roles of cilia and flagella, mechanisms have evolved to ensure the proper assembly and disassembly of the axoneme. Ciliary components are synthesized in the cell body but are incorporated at the tip of the axoneme; this requires a transport mechanism to carry components from base to tip. Intraflagellar transport (IFT; reviewed in Rosenbaum and Witman, 2002; Scholey, 2003; Blacque et al., 2008) is this transport mechanism producing bidirectional movement of material along the axoneme (Kozminski et al., 1993). IFT can be divided into two distinct stages. Anterograde IFT, mediated by complex B components, carries axonemal components from the cell body to the growing tip. Retrograde IFT, mediated by complex A components, carries the anterograde IFT machinery back from the tip to the cell body in order to recycle the IFT components. Both stages of IFT are powered by microtubule motors: kinesin-II is the anterograde motor and dynein-2 is the retrograde motor.
Dynein-2 is one of the several dynein complexes present in cells with cilia and flagella. The dynein-2 heavy chain, originally called DHC1b, was first discovered in a search of the dynein heavy chain genes expressed in sea urchin embryos (Gibbons et al., 1994). The dynein-2 motor complex is composed of four subunits: the heavy chain, Dyh2, contains the microtubule motor activity; the dynein-2 light intermediate chain, D2LIC, is thought to stabilize the dynein-2 complex; and the dynein-2 IC, D2IC, and light chain 8, LC8, are hypothesized to mediate the connection between the motor and the retrograde complex A components (Rompolas et al., 2007). In Tetrahymena, the dynein-2 components are encoded by the DYH2 (heavy chain), D2LIC (light intermediate chain), D2IC (intermediate chain), and LC8 (light chain) genes (Lee et al., 1999; Wilkes et al., 2007).
In several organisms, dynein-2 is required for the formation of normal cilia/flagella. In Chlamydomonas, the individual disruption of the heavy chain, D2LIC or light chain results in two striking effects: 1) the flagella are stumpy and swollen and 2) the flagella are filled with electron-dense material due to the inability of the system to recycle the IFT components (Pazour et al., 1998, 1999; Porter et al., 1999; Hou et al., 2004). In Caenorhabditis, the individual disruption of the heavy chain or the D2LIC results in shortened and swollen sensory cilia (Wicks et al., 2000; Schafer et al., 2003). Both dynein-2 heavy chain and D2LIC mutations produce bulbous primary cilia in mice, consequently affecting body patterning during development (Rana et al., 2004; Huangfu and Anderson, 2005; May et al., 2005). In Leishmania, deletion of the nonessential isoform of the two dynein-2 heavy chains, LmxDHC2.2, results in rounded cells with no apparent flagella (Adhiambo et al., 2005). Taken together, these observations demonstrate a well-conserved requirement of dynein-2 in normal cilia formation, and that loss of any of the dynein-2 components results in shortened and bulbous flagella or cilia.
The ciliated protozoan Tetrahymena thermophila is an important system in which to study IFT because its cilia are essential and are frequently generated. The >1000 surface cilia propel the cell through the medium; cilia draw nutrients into the oral apparatus; and cilia power rotokinesis, which is the final stage of cell division (Brown et al., 1999a). In the laboratory, Tetrahymena divide rapidly, thus forming new arrays of cilia every 3 h (Frankel, 2000). Tetrahymena anterograde IFT appears to be the same as what has been found in other systems. Disruption of the kinesin-II genes KIN-1 and KIN-2 results in the complete absence of cilia (Brown et al., 1999b). As expected, individual disruptions of the Tetrahymena complex B genes, IFT52, IFT80, and IFT172, result in extremely short or no cilia (Brown et al., 2003; Beales et al., 2007; Tsao and Gorovsky, 2008a).
Although Tetrahymena anterograde IFT is the same as that observed in other species, the role of Tetrahymena retrograde IFT appears to be different. In contrast to what has been found in other model organisms, the knockdown of DYH2 in Tetrahymena did not result in the absence of functional cilia (Lee et al., 1999). These DYH2 knockdown cells continued to produce cilia and were motile, but the transformants were often mis-sized and mis-shaped. This earlier study used a macronuclear knockdown of DYH2, and, because of the high copy number of structural genes in the Tetrahymena somatic macronucleus, one interpretation of our previous results is that there were sufficient copies of the wild-type DYH2 gene remaining to permit ciliogenesis to proceed.
To understand more clearly the role of dynein-2 in Tetrahymena ciliogenesis, we produced separate knockout heterokaryons for both the DYH2 and D2LIC genes. Knockout heterokaryons have both copies of the targeted gene disrupted in their transcriptionally silent diploid germline micronuclei and are wild-type in their somatic macronuclei (Hai et al., 2000). When two heterokaryons are mated, the gametic micronuclei undergo syngamy and produce new macronuclei; thus, the macronuclei of the progeny of such a mating completely lack the targeted gene. In this way, we have created knockout cell lines of DYH2, called KO-DYH2, and D2LIC, called KO-D2LIC. As expected, the macronuclei of the knockout cell lines are completely devoid of the targeted genes. Both KO-DYH2 and KO-D2LIC cells produce motile cilia, though the cilia are fewer in number and of more variable lengths than wild-type cilia. The cilia do not have bulbous tips nor do they appear to collect aggregated IFT particles. Thus, dynein-2 is not essential for ciliogenesis in Tetrahymena.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Gene and Protein Sequence Analyses
The full length T. thermophila D2LIC, D2IC, and D1LIC gene sequences and 4986 base pairs (bp) of the DYH2 gene encoding the region past the AAA-4 domain (including the microtubule-binding domain) were determined initially by a tBLASTn search of the T. thermophila genome (Eisen et al., 2006; http://tigrblast.tigr.org/er-blast/index.cgi?project=ttg). Introns were confirmed by RT-PCR and sequencing at the Rancho Santa Ana Botanical Garden (Claremont, CA) sequencing facility. Total RNA was isolated from B2086.1 cells by the method of Chirgwin et al. (1979). The RNA was treated with amplification grade DNaseI to remove any contaminating genomic DNA (Invitrogen, Carlsbad, CA). Reverse transcription was performed with Superscript III RNaseH– reverse transcriptase according to the manufacturer's protocol (Invitrogen, Carlsbad, CA).
Sequence analyses were done by DNASTAR (DNAStar Lasergene, Madison, WI). Domain analyses of D2LIC and D2IC sequences were done with the Protean program (DNAStar Lasergene) and SMART (http://dylan.embl-heidelberg.de/) programs, respectively. Multiple sequences were aligned by ClustalW (ebi.ac.uk/clustalw/index.html). Phylogenetic and molecular evolutionary analyses were conducted using MEGA version 2.1 (www.megasoftware.net) (Kumar et al., 2001). UPGMA, neighbor-joining, and maximum parsimony trees were constructed with known D2LIC and D1LIC sequences or known dynein-2 IC, dynein-1 IC, and outer arm dynein IC3/IC69 sequences. Only the neighbor-joining trees are shown in Figure 1. Similar results were obtained with all three methods. Accession numbers for the sequences compared are shown in Tables 1 and 2.
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18 h at a density of 3 x 105 cells/ml. For deciliation the method of Calzone and Gorovsky (1982) was followed. At 2 h after the first deciliation the cells were deciliated for a second time. Total RNA was isolated 30 min after the second deciliation (Chirgwin et al., 1979). As a baseline, total RNA was isolated from B2086.1 cells that were not deciliated (time 0). RNA was DNaseI-treated and reverse-transcribed as described above. Quantitative real-time PCR was performed using Platinum SYBR Green qPCR SuperMix UDG according to the manufacturer's directions (Invitrogen). Real-time reactions were done in the GeneAmp 5700 Sequence Detector (Applied Biosystems, Foster City, CA). The expression levels of DYH2, D2LIC, D2IC, and LC8 genes in nondeciliated and deciliated cells were determined and compared with DYH1 (noninduced control) and DYH4 (induced control). Gene-specific primers that spanned introns were used for all genes examined in this study. Real-time reactions were done with two different RNA preparations and in duplicate for each. The cDNAs were used over a 100-fold range of dilutions. Regulation relative to DYH1 was determined by the method of Pfaffl (2001) as described in Wilkes et al. (2007).
Knockout Constructs
The DYH2 and D2LIC knockout constructs were built using the NEO3 cassette kindly provided by Martin Gorovsky (University of Rochester, Rochester, NY). The NEO3 cassette has a metallothionein promoter that drives the expression of the neomycin resistance gene when induced with cadmium (Shang et al., 2002). In the DYH2 knockout construct, the upstream flanking region is 1623 bp and corresponds to +6938 to +8561 in the DYH2 gene sequence, where the A of the start codon ATG is +1. The downstream flanking region is 516 bp and corresponds to +8694 to +9210 in the DYH2 gene sequence. The upstream and the downstream flanking regions were ligated into either side of the NEO3 gene using SacI/SpeI and XhoI/ApaI restriction sites, respectively. For the D2LIC knockout construct, the upstream flanking region is 1070 bp and corresponds to –1094 to –24 of the D2LIC gene sequence (the A of the start codon ATG is at +1). The downstream flanking region is 2026 bp and corresponds to +574 and +2600 in the D2LIC gene sequence. The upstream and the downstream flanking regions of the D2LIC knockout construct were ligated into either side of NEO3 using NotI/BamHI and XhoI/ApaI restriction sites, respectively. Before biolistic transformation, both plasmids were purified using QIAGEN Plasmid Maxi Kit (QIAGEN, Valencia, CA). The DYH2 and D2LIC knockout plasmids were linearized by digestion with SacI/ApaI and NotI/ApaI enzymes, respectively. The digested plasmids were purified by organic extraction. For each bombardment, 10 µg of DNA was used.
Germline Knockouts Using Biolistic Bombardment
Tetrahymena strains B2086.1 (mating type II) and CU428.1 (mating type VII) were grown to midlogarithmic phase ( 4 x 105 cells/ml), washed once, and starved in 10 mM Tris (pH 7.5) at 30°C. After 18–20 h, the starved cells were mixed and left unshaken at 30°C. Biolistic bombardment was performed using Model PDS 1000/He Biolistic Particle Delivery System (Bio-Rad Laboratories, Hercules, CA) at 3.0, 3.5, 4.0, and 4.5 h after initiating mating (Bruns and Cassidy-Hanley, 2000). Bombarded cells were left at 30°C unshaken overnight to complete mating. About 18 h later, they were refed and 1 µg/ml final concentration of CdCl2 was added to induce the MTT promoter. Five hours after MTT induction, transformants were selected by adding paromomycin (neomycin analog) to a final concentration of 120 µg/ml. Homozygous heterokaryons were produced and screened by the method of Hai et al. (2000).
Homozygous heterokaryons of different mating types carrying the targeted deletion in their germline micronuclei were grown to 4 x 105 cells/ml, washed, and starved overnight at 30°C in 10 mM Tris (pH 7.5). Equal volumes of parental cell lines were mixed and allowed to mate at 30°C. Single-pair isolation (SPI) was done as described by Hamilton and Orias (2000). Each pair was isolated into separate wells of modified Neffs medium, and each pair was allowed to grow at 30°C. As a control, wild-type B2086.1 and CU428.1 cells were mated with one another, and SPI was used to establish clones. After the individual pairs grew up, one clone each of wild-type, KO-DYH2, and KO-D2LIC cell lines was used for genotypic and phenotypic analyses.
Verification of Targeted Gene Disruptions
Genomic DNA (gDNA) was isolated from wild-type, KO-DYH2, and KO-D2LIC cell lines (Gaertig et al., 1994). About 200 ng of gDNA of each cell line was used for each of the PCR reactions. For each knockout cell line, two sets of primers were designed to test the deletion of the targeted gene sequence and the integration of NEO3 in the targeted locus. The presence of wild-type copies of DYH2 and D2LIC genes in the knockout cell lines was tested with gene-specific primers. The DYH2 gene-specific primers were as follows: A: 5' CTGAGCATAATTCTAATGGCAG 3', B: 5' TACTCTTGAAATCCAATCCCTC 3', C: 5' CAGAGTCAGTCAAGGCAC 3', and D: 5' GAGGTTTATCTCCAGACAAAG 3'. The D2LIC gene-specific primers were as follows: F: 5' GACAGCATTCACTACACCTG 3', G: 5' CGAGTGAGCCATTTACG 3', H: 5' CCACTCAAAAGTATCTTCTTCAG 3', and I: 5' GCTCTCTTTCTTTTACTGCCTC 3'. The integration of the NEO3 cassette into the targeted gene loci was tested with the following NEO3-specific primers: N1: 5' CGCCTTCTATCGCCTTC 3' and N2: 5' CTACAAAGAATCAAGAGCGTTGC 3'. The PCR products were separated on a 1.0% (wt/vol) agarose gel and visualized with ethidium bromide using the Bio-Rad Gel Doc EQ gel documentation system (Bio-Rad Laboratories).
RNA was isolated from wild-type, KO-DYH2, and KO-D2LIC cell lines grown to low log-phase densities using Tri-reagent according to the manufacturer's directions (Sigma-Aldrich, St. Louis, MO). About 5 µg of each RNA was used for reverse-transcription as described earlier. For each knockout cell line, a pair of gene-specific primers, one within the deleted region, was used to test for the lack of expression of the gene in the deleted region. The DYH2 gene-specific primers were as follows: C: 5' CAGAGTCAGTCAAGGCAC 3' and E: 5' CAGAGTAGAGAAGAGTTTCAG 3'. The D2LIC gene-specific primers were as follows: H: 5' CCACTCAAAAGTATCTTCTTCAG 3' and J: 5' AATGCCTCTAAGTAGGTC 3'. Equal volumes (1 µl) of each cDNA were used as template for PCR reactions. The PCR products were separated on a 2.0–2.5% (wt/vol) agarose gel and visualized as described above.
For DYH2 tail expression tests, total RNA was isolated from B2086.1 (control), KO-DYH2, and KO-D2LIC cells by the method of Chirgwin et al. (1979). A total of 10 µg of RNA from each cell line were DNaseI-treated and reverse- transcribed as described before. Equal volumes (1 µl) of each cDNA were used as template for PCR reactions. The primers a: 5' CAAGAACACACTGCCCCTTC 3' and b: 5' CGGTTGAGCAAGCCATTCCAGC 3' in the tail region of the DYH2 gene were used for PCR reactions. The PCR products were separated on a 1.0% (wt/vol) agarose gel and visualized as described before.
Swimming Characteristics
For determination of swimming speeds, wild-type, KO-DYH2, and KO-D2LIC cultures grown to 5 x 104 cells/ml were used. Each cell type was further diluted in modified Neffs medium so that the field of view contained only a few cells. The tracks of moving cells were viewed by dark-field microscopy using a 2.5x objective and 6-s time-lapse images were captured using a CCD camera. A micrometer scale was also photographed and the lengths of the tracks were measured. The swimming path linearity coefficient was defined as the ratio of the linear distance from the starting point to the end point divided by the total path length (perfect linearity = 1.00). p values were determined by unpaired one-tailed t tests.
Scanning Electron Microscopy
Wild-type and knockout cells for scanning electron microscopy (SEM) were grown in modified Neffs medium to 4 x 105 cells/ml. They were fixed with a final concentration of 1.25% glutaraldehyde, 2% paraformaldehyde in 0.05 M cacodylate buffer (pH 7.0) for 25 min at room temperature. A drop of cells was placed on precoated poly-L-lysine (0.01%) coverslips and incubated for 20 min at room temperature, washed with 0.05 M cacodylate buffer (pH 7.0) and ddH2O, and dehydrated using a series of ethanol concentrations. Coverslips were critical-point dried using Tousimis SAMDRI-780A critical-point dryer and sputter-coated with 90-nm gold particles using Anatec Hummer VI Sputter Coater. Samples were observed using JEOL 840A (Peabody, MA) or Zeiss Supra 35 scanning electron microscope (Thornwood, NY) using an accelerating voltage of 12 KeV.
Transmission Electron Microscopy
Tetrahymena cells were fixed and prepared for transmission electron microscopy (TEM) similar to the methods of Allen (1967) and Jerka-Dziadosz (1981). Cultures were grown to midlog phase, centrifuged at 750 x g, and the supernatant was aspirated. The remaining pellets were fixed with 3% glutaraldehyde in 0.1 M cacodylate buffer (pH 7.4) for 30 min at room temperature and washed with 0.1 M cacodylate buffer. Cells were then transferred to microfuge tubes, pelleted at 325 x g and postfixed in 1% osmium tetroxide (OsO4) for 1 h at 4°C. After the excess OsO4 was removed, 1% liquid agar was added and pelleted at 325 x g for 1 min at room temperature. The microfuge tubes were placed in the freezer for 10 min to harden the agar. The agar was then diced into 1-mm3 pieces, washed with 0.1 M cacodylate buffer, and enbloc-stained with 0.1% tannic acid for 30 min at room temperature. Samples were dehydrated in an acetone series and embedded in Firm Spurr's resin. Ultrathin sections of 70-nm thickness were cut and stained with 2% uranyl acetate and 2% lead citrate. Sections were examined with a JEOL 100S TEM using a high voltage of 80 KeV.
Cilia Density and Length Analyses
Wild-type, KO-DYH2, and KO-D2LIC cells were grown to 5 x 104 cells/ml, washed once in 1x PHEM buffer (60 mM PIPES, 25 mM HEPES, 10 mM EGTA, and 2 mM MgCl2, pH 6.9), and fixed with 2% paraformaldehyde, 0.4% Triton-X 100 in 1x PHEM buffer. Cells were stained with mAb 1–6.1, an antibody specific for acetylated 
-tubulin (Asai et al., 1982), at 1:50 dilution in 0.1% BSA in PBS. Goat anti-mouse IgG antibody was used as the secondary antibody (Kirkegaard & Perry Laboratories, Gaithersburg, MD). The stained cells were examined by confocal fluorescence microscopy, using a Zeiss LSM510 system. An image of the widest confocal section of each cell was captured, and the ciliary densities and lengths were determined using the LSM510 software. A total of 12 cells for the control and 36 cells for each of the knockout cell lines were evaluated. The density of cilia was defined as the number of measurable cilia per micrometer of the cell circumference. p values were determined by unpaired, one-tailed t tests.
Sequential Cell Division Analyses
Growth curves were determined by isolating a single cell to begin the culture. For each cell type, triplicate cultures were analyzed by measuring the cell density every day. Cells were fixed with formaldehyde (1% final concentration) and counted using a hemacytometer. After 6 d, a single cell from each of the initial cultures was picked and placed in fresh medium, and the second round of cell growth measurements was determined. After 6 d, single cells were again picked, and a third round of cell densities was determined. The generation times of the different cell types were calculated from the initial slopes of their growth curves.
Phagocytosis
For phagocytosis experiments, freshly seeded cultures of cells at 1.2 x 105 cells/ml were used. Red fluorescent latex beads of 2-µm diameter (Sigma-Aldrich) were added to cultures, and the cells were incubated at 30°C with shaking at 100 rpm. At 1.5 and 24 h, cells were washed three times in 10 mM Tris (pH 7.5) and then fixed with 2% paraformaldehyde, 0.4% Triton X-100 in 1x PHEM buffer. The percentage of cells that contained beads was determined (n = 100). Cells that contained 
5 beads were considered as having no beads because it is not unusual to have small numbers of beads adhere to the outside of cells.
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RESULTS |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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; Rompolas et al., 2007). The Tetrahymena DYH2 gene encodes a protein of 4236 amino acids with a molecular weight of 487,769 Da (GENBANK accession no. AAV37189). The single Tetrahymena D2IC gene (accession no. ABX82716) encodes a protein of 594 amino acids with a molecular weight of 68,231 Da. Tetrahymena D2IC includes four WD40 repeats, which are characteristic of dynein ICs. A comparison of several dynein intermediate chains is shown in Figure 1A. It is clear that the Tetrahymena D2IC belongs to the cytoplasmic dynein-2 IC group and not the cytoplasmic dynein-1 IC group or an axonemal dynein IC group. Tetrahymena expresses a conventional LC8 gene and five LC8-like genes (accession nos. ABF38950, ABF38951, ABF38952, ABF38953, ABF38954, and ABF38955; Wilkes et al., 2007). The conventional Tetrahymena LC8 shares 76–90% sequence identity with the LC8 proteins from other organisms; the Tetrahymena LC8-like proteins are 33–56% identical to conventional LC8s across species.
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; Perrone et al., 2003; Schafer et al., 2003). However, the Tetrahymena D2LIC sequence does not contain the same coiled-coil domain.
Expression of Tetrahymena Dynein-2 Genes Increases after Deciliation
To determine if the dynein-2 gene products may be involved in IFT in Tetrahymena, the expression of each gene after deciliation was measured by quantitative real-time PCR. An increase in the expression of a gene after deciliation is an indication that the gene product is either a component of the cilia or is involved in the process of ciliogenesis (Lefebvre et al., 1980; Schloss et al., 1984; Soares et al., 1993). For example, the expression levels of Chlamydomonas dynein-2 heavy chain, D2LIC, and IC genes are up-regulated after deflagellation (Pazour et al., 1999; Porter et al., 1999; Perrone et al., 2003; Rompolas et al., 2007). Tetrahymena were twice deciliated with a calcium shock, and the cells were allowed to regenerate their cilia. Total RNA was isolated before deciliation and 30 min after the second deciliation and reverse-transcribed into cDNA. The levels of expression of all four of the Tetrahymena dynein-2 component genes significantly increased 30 min after deciliation (Figure 1C). In this experiment, the change in expression of each gene was normalized to that of DYH1, which should not be affected by deciliation. The positive control was the ciliary outer arm β dynein gene DYH4.
Complete Disruption of Tetrahymena DYH2 and D2LIC
To examine the function of dynein-2 in Tetrahymena, germline knockouts of the DYH2 and D2LIC genes were produced. The DYH2 gene was disrupted by a deletion of 132 bp that includes the region encoding the essential catalytic P1 loop (Lee-Eiford et al., 1986; Figure 2A). The D2LIC gene was disrupted by a deletion of 598 bp that begins 24 bp upstream of the ATG initiation codon and extends 574 bp into the coding region (Figure 2D). For both genes the deletions were replaced with the 3.6 kb NEO3 gene placed in opposite orientation to the direction of transcription of the dynein gene. Homozygous heterokaryons were produced in which both copies of the targeted gene were disrupted in the diploid micronuclei (Hai and Gorovsky, 1997). Mating of the homozygous heterokaryons produced progeny that completely lacked wild-type copies of the targeted gene in both the transcriptionally silent germline micronucleus and the somatic macronucleus. These knockout progeny are called KO-DYH2 and KO-D2LIC.
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KO-DYH2 and KO-D2LIC Cells Are Motile But Swim More Slowly and Less Straight
In other organisms, dynein-2 disruptions result in short, bulbous cilia that do not support motility. However, both the KO-DYH2 and KO-D2LIC cells were motile indicating that they had functioning cilia. Time-lapse dark-field microscopy of 27 wild-type, 36 KO-DYH2, and 41 KO-D2LIC cells was used for determining swimming velocities and the relative linearity of the swimming path. The swimming speed of wild-type cells was 181.8 ± 34.5 µm/s. The two knockout cell lines swam 3.5-fold slower than control cells; the average velocities of the KO-DYH2 and KO-D2LIC cell lines were 52.8 ± 21.2 and 51.3 ± 19.9 µm/s, respectively (Figure 3D, Table 3). Both knockout cell lines also showed a reduction in straight swimming; the difference in straightness was statistically significant between KO-DYH2 and wild-type cells (Figure 3E, Table 3). Many of the KO-DYH2 and KO-D2LIC cells exhibited a corkscrew swimming pattern or frequent turns (Figure 3, A–C). Thus, Dyh2 and D2LIC contribute in similar ways to cell motility.
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, 1999; Porter et al., 1999; Hou et al., 2004). Both SEM and TEM examinations of the Tetrahymena cilia were conducted using a blinded protocol. One of us (A.S.) was provided coded cultures that she prepared for EM analyses and evaluated without knowing the coding. SEM showed that the cilia on Tetrahymena KO-DYH2 and KO-D2LIC cells were of varying lengths, including some very short cilia (Figure 4, A–C). However, none of the cilia had bulbous tips (Figure 4, D–F). Examination of the thin-section TEM images found no differences among the three samples and no evidence of the accumulation of material in the cilia of the Tetrahymena knockouts (Figure 4, G–L). The axonemal organization was normal in the knockouts with the 9 + 2 pattern of microtubules, radial spokes, and inner and outer arm dyneins (Figure 4, G–I).
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,b, 2003; Williams et al., 2006). To examine the ability of the dynein-2 knockouts to divide, single cells were isolated and grown to high density; then a single cell from the first culture was isolated to start a second clone of cells, and then the process was repeated a third time. Results of such experiments are shown in Figure 6. Over this long protocol, the KO-DYH2 and KO-D2LIC cells continued to divide, albeit more slowly. At 30°C without shaking, wild-type Tetrahymena cells divided every 2.9 h and grew to a density of 1–1.5 x 106 cells/ml. The KO-DYH2 and KO-D2LIC grew with longer generation times of 4.6 and 4.2 h, respectively, and achieved densities of 0.4–0.5 x 106 cells/ml.
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, 2003; Williams et al., 2006). Examples of knockout cells are shown in Figure 7. We conclude that the impaired cell motility of the dynein-2 knockout cells resulted in inefficient cell division that underlies the longer generation times and lower cell densities.
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). A simple assay for phagocytosis is to incubate cells with latex beads and then to evaluate ingestion of the beads by light microscopy. Both KO-DYH2 and KO-D2LIC cells had oral apparatuses as visualized by immunostaining with anti-tubulin antibody and SEM, but neither phagocytosed beads normally. After 1.5 h, 71% of wild-type cells contained beads, whereas none of the KO-DYH2 and only 4% of the KO-D2LIC cells contained beads. After 24 h, neither KO-DYH2 nor KO-D2LIC cells contained significant numbers of beads (Figure 8).
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Dynein-2 Is Not Required for Ciliogenesis in Tetrahymena
In other organisms, the loss of dynein-2 results in the failure to produce normal and motile cilia or flagella. In Chlamydomonas, disruptions of the dynein-2 genes result in very short and swollen flagella containing massive accumulations of electron-dense IFT material in the bulbous distal tips (Pazour et al., 1998, 1999; Porter et al., 1999; Hou et al., 2004). In Leishmania, disruption of one of the DYH2 isoforms results in the complete loss of flagella (Adhiambo et al., 2005). Dynein-2 mutants in C. elegans have short, stumpy sensory cilia (Signor et al., 1999; Wicks et al., 2000; Schafer et al., 2003), and in mouse, dynein-2 mutations result in bloated and bulbous embryonic nodal cilia that are shorter than normal or completely absent (Rana et al., 2004; Huangfu and Anderson, 2005; May et al., 2005). In striking contrast to what has been observed in these other model systems, the disruption of dynein-2 components in Tetrahymena did not prevent the formation of motile, normal-appearing cilia.
Because the parental heterokaryon cell lines expressed wild-type levels of the Dyh2 and D2LIC proteins, some dynein-2 may remain in the progeny soon after mating even though the knockout cells do not have the capacity to express the genes. Therefore, it is important to consider the possibility that this residual dynein-2 is sufficient to drive ciliogenesis. The growth rates experiment shown in Figure 6 provides the basis for evaluating this possibility. In this experiment, each culture was begun with a single cell, the culture was allowed to grow up for 6 d, and then a single cell from the 6-d culture was used to begin a new culture. This protocol was repeated a third time. From the cell densities measured at the end of every expansion, we estimate that the contents of the single dynein-2 mutant cell used to start the experiment were effectively diluted more than 1016-fold. Of course, this is a gross underestimation because it does not consider protein degradation. We conclude that the fact that the dynein-2 knockout cells continue to produce motile cilia is not due to residual carry over of dynein-2 proteins.
In light of our result with the retrograde motor, it is interesting to ask whether loss of function of the retrograde complex A results in a more severe phenotype. The disruption of the Tetrahymena complex A IFT122A gene resulted in cells that continued to assemble cilia similar to our KO-DYH2 and KO-D2LIC cells (Tsao and Gorovsky, 2008b). Our lab also has produced a knockdown of another Tetrahymena complex A gene, IFT140, again resulting in only a mild effect on ciliogenesis (J. DuMond, V. Rajagopalan, D. Wilkes, unpublished results). Together, these results demonstrate that retrograde IFT—at least as it is powered by dynein-2—is not required for ciliogenesis in Tetrahymena.
Dynein-2 Mutants Have Lost the Ability to Regulate the Lengths of Their Cilia
The dynein-2 knockout cells have impaired motility, as measured by several parameters. The mutants swim more slowly and less straight, fail to consume fluorescent beads, and exhibit inefficient cell division. A close examination of the cilia on individual cells reveals that the principal effect of the gene knockouts was on ciliary length. Although average lengths of cilia on knockout cells were less than that of wild-type cells, there was an abnormally wide range of ciliary lengths on the knockout cells, including some unusually long cilia (Figure 5). This loss of ciliary length homogeneity is reminiscent of what was observed when the IFT complex A component 122A was knocked out (Tsao and Gorovsky, 2008b).
The metachronous coordination of ciliary beating is expected to be sensitive to variations in lengths among individual cilia (Nelsen and DeBault, 1978; Frankel, 2000). Thus, the impaired motility exhibited by the knockout cells is interpreted to be a secondary effect caused by the loss of ciliary length regulation. These results suggest that in Tetrahymena retrograde IFT is responsible for the close control of cilia length.
Retrograde IFT and Ciliogenesis
The intraflagellar transport model is an important framework in which to understand ciliogenesis. Anterograde transport, which depends on kinesin-II and complex B components, carries materials to the tip, which is the site of incorporation of new ciliary proteins. Retrograde transport, which depends on dynein-2 and complex A components, is responsible for returning the anterograde machinery to the base of the cilium so that it can be recycled. The IFT model predicts that loss of function of the anterograde motor or complex B will result in no ciliary growth and that loss of function of the retrograde apparatus will result in accumulation of IFT material at the ciliary tip, producing short, stumpy, and bulbous cilia. Despite some variations in different organisms—e.g., two different anterograde motors in C. elegans versus one in Chlamydomonas (Kozminski et al., 1995; Snow et al., 2004; Mukhopadhyay et al., 2007) —the general model holds well for anterograde IFT. The disruption of the kinesin or individual complex B components results in no cilia or flagella in all of the organisms examined, including Tetrahymena (Kozminski et al., 1995; Collet et al., 1998; Nonaka et al., 1998; Brown et al., 1999b; Snow et al., 2004; Pedersen et al., 2005; Beales et al., 2007; Absalon et al., 2008; Tsao and Gorovsky, 2008a).
In contrast to the highly conserved requirement for anterograde IFT in ciliogenesis in all organisms, the requirement for retrograde IFT is not universal. As this study shows, the disruption of dynein-2 components does not prevent ciliogenesis in Tetrahymena, and the cilia on the knockout cells do not have accumulations of IFT material as seen in other systems. Other exceptions to the retrograde IFT model include Toxoplasma gondii and the marine diatom Thalassiosira pseudonana. Both organisms lack the genes for dynein-2 heavy chain and D2LIC even though they form flagella (Wickstead and Gull, 2007).
Why Is Tetrahymena Different?
Clearly, dynein-2 and IFT complex A (Tsao and Gorovsky, 2008b) do not make the same contributions to Tetrahymena ciliogenesis as they do in other model organisms, notably Chlamydomonas. Why is Tetrahymena different? Here we consider two possibilities.
One possibility is that retrograde IFT is required for Tetrahymena ciliogenesis, but that motors other than dynein-2 can power retrograde IFT. The KO-DYH2 cells were produced by the deletion of the region of the DYH2 gene encoding the catalytic P-loop, which is essential for dynein motor activity (Lee-Eiford et al., 1986). Because the deleted portion of the DYH2 gene is several kilobasepairs away from the initiation codon, a large portion of the 5' end of the DYH2 gene remains in the KO-DYH2 cells. As expected, the deleted portion of the DYH2 gene was not expressed in the KO-DYH2 cell line (Figure 2C). However, RNA-directed PCR revealed that the portion of the DYH2 gene located upstream of the catalytic P-loop was transcribed (Figure 9). This is reminiscent of earlier work in which transcripts corresponding to truncated versions of DYH genes were found to persist in Tetrahymena dynein transformants (Liu et al., 2004).
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). This possible redundancy of IFT motors may extend to the Tetrahymena anterograde machinery (see Awan et al., 2004). Future experiments—currently not feasible because of the lack of specific antibody reagents—aimed at localizing dynein-2 subunits in the knockout cells may provide evidence for the formation of complexes of dynein-2 subunits with different motor proteins.
A second possibility is that retrograde IFT is simply not important in Tetrahymena. The cilia on Tetrahymena are shorter and less dynamic than Chlamydomonas flagella which undergo cycles of lengthening and resorption (Johnson and Porter, 1968; Marshall and Rosenbaum, 2001). The static nature of Tetrahymena cilia is underscored by our preliminary experiments in which we have measured the rate of ciliary growth in nondividing cells. Even after 22 h of reciliation, the dynein-2 mutants continue to exhibit motile cilia of varying lengths (V. Rajagopalan, D. Wilkes, and D. Asai, unpublished results). Thus, whereas anterograde IFT is essential for Tetrahymena cilia formation, as is the case in all organisms examined, Tetrahymena retrograde IFT may only contribute to the regulation of ciliary length. If this is the case, then we would expect to find that dynein-2 mutants will be unable to resorb their cilia, for example, by the overexpression of NIMA-related kinases (Wloga et al., 2006). An earlier study reported that partial deciliation of cells causes the remaining cilia to stiffen and then to undergo resorption (Rannestad, 1974). Although we have not purposely attempted to obtain partial deciliation, we have not observed unresorbed cilia on dynein-2 knockout cells after their deciliation.
In conclusion, the elimination of dynein-2 subunits in Tetrahymena resulted in cells that continue to produce motile cilia. The cilia are normally shaped and show no signs of accumulation of unrecycled IFT material. Unlike what is observed on wild-type cells, there is a significant variation of the lengths of cilia on individual dynein-2 mutant cells. We conclude that in Tetrahymena dynein-2 is not required for ciliogenesis, but that retrograde IFT plays an important role in regulating cilia length.
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ACKNOWLEDGMENTS |
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Footnotes |
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Address correspondence to: David J. Asai (david_asai{at}hmc.edu).
Abbreviations used: IFT, intraflagellar transport; DYH2, cytoplasmic dynein-2 heavy chain; D2LIC, cytoplasmic dynein-2 light intermediate chain.
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REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
|---|
Adhiambo, C., Forney, J. D., Asai, D. J., and LeBowitz, J. H. (2005). The two cytoplasmic dynein-2 isoforms in Leishmania mexicana perform separate functions. Mol. Biochem. Parasitol 143, 216–225.[CrossRef][Medline]
Allen, R. D. (1967). Fine structure, reconstruction and possible functions of components of the cortex of Tetrahymena pyriformis. J. Protozool 14, 553–565.[Medline]
Asai, D. J., Brokaw, C. J., Thompson, W. C., and Wilson, L. (1982). Two different monoclonal antibodies to alpha-tubulin inhibit the bending of reactivated sea urchin spermatozoa. Cell Motil 2, 599–614.[CrossRef][Medline]
Awan, A., Bernstein, M., Hamasaki, T., and Satir, P. (2004). Cloning and characterization of Kin5, a novel Tetrahymena ciliary kinesin II. Cell Motil. Cytoskelet 58, 1–9.[CrossRef][Medline]
Beales, P. L. et al. (2007). IFT80, which encodes a conserved intraflagellar transport protein, is mutated in Jeune asphyxiating thoracic dystrophy. Nat. Genet 39, 727–729.[CrossRef][Medline]
Blacque, O. E., Cevik, S., and Kaplan, O. I. (2008). Intraflagellar transport: from molecular characterisation to mechanism. Front. Biosci 13, 2633–2652.[CrossRef][Medline]
Brown, J. M., Hardin, C., and Gaertig, J. (1999a). Rotokinesis, a novel phenomenon of cell locomotion-assisted cytokinesis in the ciliate Tetrahymena thermophila. Cell Biol. Int 23, 841–848.[CrossRef][Medline]
Brown, J. M., Marsala, C., Kosoy, R., and Gaertig, J. (1999b). Kinesin-II is preferentially targeted to assembling cilia and is required for ciliogenesis and normal cytokinesis in Tetrahymena. Mol. Biol. Cell 10, 3081–3096.
Brown, J. M., Fin, N. A., Pandiyan, G., Thazhath, R., and Gaertig, J. (2003). Hypoxia regulates assembly of cilia in suppressors of Tetrahymena lacking an intraflagellar transport subunit gene. Mol. Biol. Cell 14, 3192–3207.
Bruns, P. J., and Cassidy-Hanley, D. (2000). Biolistic transformation of macro- and micronuclei. Methods Cell Biol 62, 501–512.[Medline]
Calzone, F. J., and Gorovsky, M. A. (1982). Cilia regeneration in Tetrahymena. A simple reproducible method for producing large numbers of regenerating cells. Exp. Cell Res 140, 471–476.[CrossRef][Medline]
Chirgwin, J. M., Przybyla, A. E., MacDonald, R. J., and Rutter, W. J. (1979). Isolation of biologically active ribonucleic acid from sources enriched in ribonuclease. Biochemistry 18, 5294–5299.[CrossRef][Medline]
Collet, J., Spike, C. A., Lundquist, E. A., Shaw, J. E., and Herman, R. K. (1998). Analysis of osm-6, a gene that affects sensory cilium structure and sensory neuron function in Caenorhabditis elegans. Genetics 148, 187–200.
Eisen, J. A. et al. (2006). Macronuclear genome sequence of the ciliate Tetrahymena thermophila, a model eukaryote. PLoS Biol 4, e286.[CrossRef][Medline]
Frankel, J. (2000). Cell Biology of Tetrahymena thermophila. Methods Cell Biol 62, 27–125.[Medline]
Gaertig, J., Thatcher, T. H., Gu, L., and Gorovsky, M. A. (1994). Electroporation-mediated replacement of a positively and negatively selectable beta-tubulin gene in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 91, 4549–4553.
Gibbons, I. R. (1981). Cilia and flagella of eukaryotes. J. Cell. Biol 91, 107–124.[CrossRef]
Gibbons, B. H., Asai, D. J., Tang, W. J., Hays, T. S., and Gibbons, I. R. (1994). Phylogeny and expression of axonemal and cytoplasmic dynein genes in sea urchins. Mol. Biol. Cell 5, 57–70.[Abstract]
Grissom, P. A., Vaisberg, E. A., and McIntosh, J. R. (2002). Identification of a novel light intermediate chain (D2LIC) for mammalian cytoplasmic dynein 2. Mol. Biol. Cell 13, 817–829.
Hai, B., Gaertig, J., and Gorovsky, M. A. (2000). Knockout heterokaryons enable facile mutagenic analysis of essential genes in Tetrahymena. Methods Cell Biol 62, 513–531.[Medline]
Hai, B., and Gorovsky, M. A. (1997). Germ-line knockout heterokaryons of an essential alpha-tubulin gene enable high-frequency gene replacement and a test of gene transfer from somatic to germ-line nuclei in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 94, 1310–1315.
Hamilton, E. P., and Orias, E. (2000). Genetic crosses: setting up crosses, testing progeny, and isolating phenotypic assortants. Methods Cell Biol 62, 219–228.[Medline]
Hou, Y., Pazour, G. J., and Witman, G. B. (2004). A dynein light intermediate chain, D1bLIC, is required for retrograde intraflagellar transport. Mol. Biol. Cell 15, 4382–4394.
Huangfu, D., and Anderson, K. V. (2005). Cilia and Hedgehog responsiveness in the mouse. Proc. Natl. Acad. Sci. USA 102, 11325–11330.
Jerka-Dziadosz, M. (1981). Cytoskeleton-related structures in Tetrahymena thermophila: microfilaments at the apical and division-furrow rings. J. Cell Sci 51, 241–253.[Abstract]
Johnson, U. G., and Porter, K. R. (1968). Fine structure of cell division in Chlamydomonas reinhardi. Basal bodies and microtubules. J. Cell Biol 38, 403–425.
Kozminski, K. G., Beech, P. L., and Rosenbaum, J. L. (1995). The Chlamydomonas kinesin-like protein FLA10 is involved in motility associated with the flagellar membrane. J. Cell Biol 131, 1517–1527.
Kozminski, K. G., Johnson, K. A., Forscher, P., and Rosenbaum, J. L. (1993). A motility in the eukaryotic flagellum unrelated to flagellar beating. Proc. Natl. Acad. Sci. USA 90, 5519–5523.
Kumar, S., Tamura, K., Jakobsen, I. B., and Nei, M. (2001). MEGA 2, Molecular Evolutionary Genetics Analysis software, Tempe, AZ: Arizona State University.
Lee-Eiford, A., Ow, R. A., and Gibbons, I. R. (1986). Specific cleavage of dynein heavy chains by ultraviolet irradiation in the presence of ATP and vanadate. J. Biol. Chem 261, 2337–2342.
Lee, S., Wisniewski, J. C., Dentler, W. L., and Asai, D. J. (1999). Gene knockouts reveal separate functions for two cytoplasmic dyneins in Tetrahymena thermophila. Mol. Biol. Cell 10, 771–784.
Lefebvre, P. A., Silflow, C. D., Wieben, E. D., and Rosenbaum, J. (1980). Increased levels of mRNAs for tubulin and other flagellar proteins after amputation or shortening of Chlamydomonas flagella. Cell 20, 469–477.[CrossRef][Medline]
Liu, S., Hard, R., Rankin, S., Hennessey, T., and Pennock, D. G. (2004). Disruption of genes encoding predicted inner arm dynein heavy chains causes motility phenotypes in Tetrahymena. Cell Motil. Cytoskelet 59, 201–214.[CrossRef][Medline]
Marshall, W. F., and Rosenbaum, J. L. (2001). Intraflagellar transport balances continuous turnover of outer doublet microtubules: implications for flagellar length control. J. Cell Biol 155, 405–414.
May, S. R., Ashique, A. M., Karlen, M., Wang, B., Shen, Y., Zarbalis, K., Reiter, J., Ericson, J., and Peterson, A. S. (2005). Loss of the retrograde motor for IFT disrupts localization of Smo to cilia and prevents the expression of both activator and repressor functions of Gli. Dev. Biol 287, 378–389.[Medline]
Mukhopadhyay, S., Lu, Y., Qin, H., Lanjuin, A., Shaham, S., and Sengupta, P. (2007). Distinct IFT mechanisms contribute to the generation of ciliary structural diversity in C. elegans. EMBO J 26, 2966–2980.[CrossRef][Medline]
Nelsen, E. M., and DeBault, L. E. (1978). Transformation in Tetrahymena pyriformis: description of an inducible phenotype. J. Protozool 25, 113–119.[Medline]
Nonaka, S., Tanaka, Y., Okada, Y., Takeda, S., Harada, A., Kanai, Y., Kido, M., and Hirokawa, N. (1998). Randomization of left-right asymmetry due to loss of nodal cilia generating leftward flow of extraembryonic fluid in mice lacking KIF3B motor protein. Cell 95, 829–837.[CrossRef][Medline]
Pazour, G. J., Agrin, N., Leszyk, J., and Witman, G. B. (2005). Proteomic analysis of a eukaryotic cilium. J. Cell Biol 170, 103–113.
Pazour, G. J., Dickert, B. L., and Witman, G. B. (1999). The DHC1b (DHC2) isoform of cytoplasmic dynein is required for flagellar assembly. J. Cell Biol 144, 473–481.
Pazour, G. J., Wilkerson, C. G., and Witman, G. B. (1998). A dynein light chain is essential for the retrograde particle movement of intraflagellar transport (IFT). J. Cell Biol 141, 979–992.
Pedersen, L. B., Miller, M. S., Geimer, S., Leitch, J. M., Rosenbaum, J. L., and Cole, D. G. (2005). Chlamydomonas IFT172 is encoded by FLA11, interacts with CrEB1, and regulates IFT at the flagellar tip. Curr. Biol 15, 262–266.[CrossRef][Medline]
Perrone, C. A., Tritschler, D., Taulman, P., Bower, R., Yoder, B., and Porter, M. E. (2003). A novel dynein light intermediate chain colocalizes with the retrograde motor for intraflagellar transport at sites of axoneme assembly in Chlamydomonas and mammalian cells. Mol. Biol. Cell 14, 2041–2056.
Pfaffl, M. W. (2001). A new mathematical model for relative quantification in real-time RT-PCR. Nucleic Acids Res 29, e45.
Piperno, G., Huang, B., and Luck, D. J. (1977). Two-dimensional analysis of flagellar proteins from wild-type and paralyzed mutants of Chlamydomonas reinhardtii. Proc. Natl. Acad. Sci. USA 74, 1600–1604.
Porter, M. E., Bower, R., Knott, J. A., Byrd, P., and Dentler, W. (1999). Cytoplasmic dynein heavy chain 1b is required for flagellar assembly in Chlamydomonas. Mol. Biol. Cell 10, 693–712.
Rana, A. A., Barbera, J. P., Rodriguez, T. A., Lynch, D., Hirst, E., Smith, J. C., and Beddington, R. S. (2004). Targeted deletion of the novel cytoplasmic dynein mD2LIC disrupts the embryonic organiser, formation of the body axes and specification of ventral cell fates. Development 131, 4999–5007.
Rannestad, J. (1974). The regeneration of cilia in partially deciliated Tetrahymena. J. Cell Biol 63, 1009–1017.
Rompolas, P., Pedersen, L. B., Patel-King, R. S., and King, S. M. (2007). Chlamydomonas FAP133 is a dynein intermediate chain associated with the retrograde intraflagellar transport motor. J. Cell Sci 120, 3653–3665.
Rosenbaum, J. L., and Witman, G. B. (2002). Intraflagellar transport. Nat. Rev. Mol. Cell Biol 3, 813–825.[CrossRef][Medline]
Schafer, J. C., Haycraft, C. J., Thomas, J. H., Yoder, B. K., and Swoboda, P. (2003). XBX-1 encodes a dynein light intermediate chain required for retrograde intraflagellar transport and cilia assembly in Caenorhabditis elegans. Mol. Biol. Cell 14, 2057–2070.
Schloss, J. A., Silflow, C. D., and Rosenbaum, J. L. (1984). mRNA abundance changes during flagellar regeneration in Chlamydomonas reinhardtii. Mol. Biol. Cell 4, 424–434.
Scholey, J. M. (2003). Intraflagellar transport. Annu. Rev. Cell Dev. Biol 19, 423–443.[CrossRef][Medline]
Shang, Y., Song, X., Bowen, J., Corstanje, R., Gao, Y., Gaertig, J., and Gorovsky, M. A. (2002). A robust inducible-repressible promoter greatly facilitates gene knockouts, conditional expression, and overexpression of homologous and heterologous genes in Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA 99, 3734–3739.
Signor, D., Wedaman, K. P., Orozco, J. T., Dwyer, N. D., Bargmann, C. I., Rose, L. S., and Scholey, J. M. (1999). Role of a class DHC1b dynein in retrograde transport of IFT motors and IFT raft particles along cilia, but not dendrites, in chemosensory neurons of living Caenorhabditis elegans. J. Cell Biol 147, 519–530.
Smith, J. C., Northey, J. G., Garg, J., Pearlman, R. E., and Siu, K. W. (2005). Robust method for proteome analysis by MS/MS using an entire translated genome: demonstration on the ciliome of Tetrahymena thermophila. J. Proteome Res 4, 909–919.[CrossRef][Medline]
Snow, J. J., Ou, G., Gunnarson, A. L., Walker, M. R., Zhou, H. M., Brust-Mascher, I., and Scholey, J. M. (2004). Two anterograde intraflagellar transport motors cooperate to build sensory cilia on C. elegans neurons. Nat. Cell Biol 6, 1109–1113.[CrossRef][Medline]
Soares, H., Galego, L., Cóias, R., and Rodrigues-Pousada, C. (1993). The mechanisms of tubulin messenger regulation during Tetrahymena pyriformis reciliation. J. Biol. Chem 268, 16623–16630.
Tsao, C. C., and Gorovsky, M. A. (2008a). Different effects of Tetrahymena IFT172 domains on anterograde and retrograde intraflagellar transport. Mol. Biol. Cell 19, 1450–1461.
Tsao, C. C., and Gorovsky, M. A. (2008b). Tetrahymena IFT122A is not essential for cilia assembly but plays a role in returning IFT proteins from the ciliary tip to the cell body. J. Cell Sci 121, 428–436.
Wicks, S. R., de Vries, C. J., van Luenen, H.G.A.M., and Plasterk, R.H.A. (2000). CHE-3, a cytosolic dynein heavy chain, is required for sensory cilia structure and function in Caenorhabditis elegans. Dev. Biol 221, 295–307.[CrossRef][Medline]
Wickstead, B., and Gull, K. (2007). Dyneins across eukaryotes: a comparative genomic analysis. Traffic 8, 1708–1721.[CrossRef][Medline]
Wilkes, D. E., Rajagopalan, V., Chan, C. W., Kniazeva, E., Wiedeman, A. E., and Asai, D. J. (2007). Dynein light chain family in Tetrahymena thermophila. Cell Motil. Cytoskelet 64, 82–96.[CrossRef][Medline]
Wilkes, D. E., Watson, H., Mitchell, D. R., and Asai, D. J. (2008). 25 dyneins in Tetrahymena: a re-examination of the multi-dynein hypothesis. Cell Motil. Cytoskelet 65, 342–351.[CrossRef][Medline]
Williams, N. E., Tsao, C. C., Bowen, J., Hehman, G. L., Williams, R. J., and Frankel, J. (2006). The actin gene ACT1 is required for phagocytosis, motility, and cell separation of Tetrahymena thermophila. Eukaryot. Cell 5, 555–567.
Wloga, D., Camba, A., Rogowski, K., Manning, G., Jerka-Dziadosz, M., and Gaertig, J. (2006). Members of the NIMA-related kinase family promote disassembly of cilia by multiple mechanisms. Mol. Biol. Cell 17, 2799–2810.
Zariwala, M. A., Knowles, M. R., and Omran, H. (2007). Genetic defects in ciliary structure and function. Annu. Rev. Physiol 69, 423–450.[CrossRef][Medline]
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, wild-type; ·, KO-DYH2; 
, KO-D2LIC.
