the actin gene act1 is required for phagocytosis, motility, and cell ... · cell separation of...

EUKARYOTIC CELL, Mar. 2006, p. 555–567 Vol. 5, No. 31535-9778/06/$08.00�0 doi:10.1128/EC.5.3.555–567.2006Copyright © 2006, American Society for Microbiology. All Rights Reserved.

The Actin Gene ACT1 Is Required for Phagocytosis, Motility, andCell Separation of Tetrahymena thermophila†

Norman E. Williams,1 Che-Chia Tsao,2 Josephine Bowen,2 Gery L. Hehman,1Ruth J. Williams,1 and Joseph Frankel1*

Department of Biological Sciences, University of Iowa, Iowa City, Iowa,1 and Department of Biology,University of Rochester, Rochester, New York2

Received 20 September 2005/Accepted 2 December 2005

A previously identified Tetrahymena thermophila actin gene (C. G. Cupples and R. E. Pearlman, Proc. Natl.Acad. Sci. USA 83:5160–5164, 1986), here called ACT1, was disrupted by insertion of a neo3 cassette. Cells inwhich all expressed copies of this gene were disrupted exhibited intermittent and extremely slow motility andseverely curtailed phagocytic uptake. Transformation of these cells with inducible genetic constructs thatcontained a normal ACT1 gene restored motility. Use of an epitope-tagged construct permitted visualization ofAct1p in the isolated axonemes of these rescued cells. In ACT1� mutant cells, ultrastructural abnormalities ofouter doublet microtubules were present in some of the axonemes. Nonetheless, these cells were still able toassemble cilia after deciliation. The nearly paralyzed ACT1� cells completed cleavage furrowing normally, butthe presumptive daughter cells often failed to separate from one another and later became reintegrated. Clonalanalysis revealed that the cell cycle length of the ACT1� cells was approximately double that of wild-typecontrols. Clones could nonetheless be maintained for up to 15 successive fissions, suggesting that the ACT1gene is not essential for cell viability or growth. Examination of the cell cortex with monoclonal antibodiesrevealed that whereas elongation of ciliary rows and formation of oral structures were normal, the ciliary rowsof reintegrated daughter cells became laterally displaced and sometimes rejoined indiscriminately across theformer division furrow. We conclude that Act1p is required in Tetrahymena thermophila primarily for normalciliary motility and for phagocytosis and secondarily for the final separation of daughter cells.

Actin, although not abundant (26), has been detected atnumerous intracellular sites within ciliates. These sites includethe oral apparatus (8, 21, 29, 41, 45), phagosomes (food vacu-oles) (8, 31), the cytoproct (8, 28), basal bodies (29, 40, 41, 64),and cilia (41, 46, 64) of both Tetrahymena (reviewed in refer-ence 15) and Paramecium, and the division furrow (22, 28) ofTetrahymena. However, despite the numerous places whereactin has been found, its role in the life of these cells is incom-pletely understood.

Perhaps the best-established function of ciliate actin is as-sociated with phagocytosis, as cytochalasins and other drugsthat affect actin polymerization (9) inhibit formation of foodvacuoles in Tetrahymena pyriformis (49, 50), T. vorax (23), T.thermophila (72), and Paramecium tetraurelia (8, 14). Cytocha-lasin B also binds to isolated oral apparatuses of T. thermophila(21), and inhibits the formation of this organelle, but does so ata much higher concentration than is sufficient to inhibit feeding(20). A recent study also has shown that latrunculin, an inhib-itor of actin polymerization, interferes with the posteriorlydirected movement of phagosomes within this cell (31).

From past work it is unclear whether perturbing actin func-tion interferes with cell division. Cytochalasin B did not pre-vent cell division of synchronized Tetrahymena pyriformis cellsat concentrations that inhibit food vacuole formation (50) and

also failed to prevent synchronized division of T. thermophila(strain WH-6) when added at a high concentration to cellsafter (predivision) stage 4 of oral development (20). However,two novel methods of interference with actin function didsucceed in bringing about division arrest. First, microinjectionof skeletal muscle actin into T. thermophila cells just prior tothe onset of constriction prevented cytokinesis but had noeffect on macronuclear division (53). Second, green fluorescentprotein (GFP)-tagged actin expressed at a high level from anrRNA gene-based replicative vector blocked both macro-nuclear elongation and cytokinesis in some T. thermophilacells (32).

None of the reports on effects of cytochalasins on ciliatesmention changes in cell motility, suggesting that it was notimpaired. Likewise, a study of the effects of cytochalasin D onflagellar microtubule dynamics in the alga Chlamydomonasreinhardtii stated that “the cells were able to swim normally” inthis drug, and also were capable of nearly normal flagellarregeneration after amputation (11). However, a null mutationof the IDA5 gene, later shown to encode the sole conventionalactin of this species (39), reduced swimming speed to less thanhalf that of wild-type Chlamydomonas cells (38) without affect-ing cell growth or division.

The actin gene, ACT1, was cloned from Tetrahymena ther-mophila by Cupples and Pearlman (10), and a close orthologwas cloned from T. pyriformis (26). The predicted amino acidsequence of the protein encoded by the T. thermophila ACT1gene was found to be highly divergent relative to actins in otherorganisms, showing, for example, only 77% identity with themost similar human actins, compared to 87 to 89% identity

* Corresponding author. Mailing address: Department of BiologicalSciences, The University of Iowa, 143 Biology Bldg., Iowa City, IA52242. Phone: (319) 335-1110. Fax: (319) 335-1069. E-mail: [email protected].

† Supplemental material for this article may be found at http://ec.asm.org/.

555

on June 16, 2020 by guesthttp://ec.asm

.org/D

ownloaded from

https://crossmark.crossref.org/dialog/?doi=10.1128/EC.5.3.555-567.2006&domain=pdf&date_stamp=2006-03-01

http://ec.asm.org/

between Saccharomyces cerevisiae actin (GenBank accessionnumber CAA24598) and human actins.

A search of the latest assembly of the T. thermophila macro-nuclear genome from The Institute for Genomic Research(http://www.tigr.org/tdb/e2k1/ttg/) and the Tetrahymena GenomeDatabase (http://www.ciliate.org) revealed that ACT1 is one offour genes for actins in this organism. The predicted aminoacid sequences of the proteins encoded by the other three actingenes were found to be even more divergent than those en-coded by ACT1, and these proteins also are less highly ex-pressed (C.-C. Tsao and M. A. Gorovsky, unpublished obser-vations). Therefore, despite its divergent amino acid sequenceand its atypical properties (27), we consider the protein en-coded by the ACT1 gene to be the “classical” Tetrahymenaactin, and have named it Act1p.

As a vital step in elucidating the role of actins in T. thermophila,we disrupted the ACT1 gene of T. thermophila with the neo3inactivation cassette. In spite of the multiplicity of actin genes, weobtained a clear mutant phenotype from this disruption. Themost obvious direct effect was the presence of almost com-pletely nonmotile cilia. Food vacuole formation was also in-hibited. However, macronuclear division and cytoplasmiccleavage furrowing were unimpaired; also, the ACT1 gene ap-parently was not essential for viability. As reported earlier (3,5), arrest of ciliary motility brought about failure of the finalstep of separation of presumptive daughter cells. This failureimpeded clonal growth and also generated a unique cellularreintegration phenotype.

MATERIALS AND METHODS

Cells and culture media. The Tetrahymena thermophila strains used wereCU428 {Mpr/Mpr, Chx�/Chx� [6-methylpurine sensitive (mp-s), cycloheximidesensitive (cy-s), paromomycin sensitive (pm-s), VII]} and B2086 {Mpr�/Mpr�,Chx�/Chx�, (mp-s, cy-s, pm-s, II)}, originally constructed by P. J. Bruns, CornellUniversity. The media employed in this investigation were PPY (1% proteosepeptone plus 0.5% yeast extract); PPYGFe (2% proteose peptone, 0.5% glucose,0.2% yeast extract, plus 9 � 10�5 M Fe-EDTA); SPP (1% proteose peptone,0.2% glucose, 0.1% yeast extract, plus 9 � 10�5 M Fe-EDTA); and MEPP (2%proteose peptone, 0.06% Na3 citrate · 2H2O, 0.027% FeCl3 · 6H2O, 0.0003%CuSO4 · 5H2O, and 0.0001% folinic acid, Ca salt, prepared as described inreference 54). The Fe-EDTA used in PPYGFe and SPP in some cases came froma stock made up as described in footnote 1 of reference 48. In all experimentsfollowing transformation, 100 units/ml penicillin G, 100 �g/ml streptomycinsulfate, and 0. 25 �g/ml amphotericin B were added to the culture media toprevent bacterial or fungal contamination.

Cloning of ACT1. The complete coding sequence of T. thermophila ACT1,along with 250 bp of 5� untranslated region (UTR) (10), was amplified by PCRusing both a cDNA library (courtesy of Aaron Turkewitz, University of Chicago)and genomic DNA as templates. No introns were found. The forward primer was5�-AGATCTCATCAAACAATTA-3� and the reverse primer was 5�-TCAGAAGCACTTTCTGTGGA-3�. The 1,382-bp PCR product was cloned using theInvitrogen Topo TA cloning kit; the final construct is pAct-35-TOPO.

Somatic disruption of ACT1. The targeting construct (Fig. 1A) was created byinsertion of the neo3 disruption cassette (58), which confers resistance to paro-momycin, into pAct-35-TOPO at a unique SfuI site found at position 625 of theACT1 gene. Plasmid pMNBL containing the neo3 disruption cassette was a giftfrom Martin Gorovsky, University of Rochester. The neo3 cassette consists of thecadmium-inducible MTT1 promoter, the neomycin coding region, and the BTU23�-flanking region.

The pAct-35-TOPO-neo3 targeting construct (Fig. 1A) digested with BstXIwas transformed into CU428 cells using the biolistic PDS-1000/He particle de-livery system (Bio-Rad) (7). Cells were transferred to SPP medium containing 1�g/ml CdCl2, and 4 hours later 80 �g/ml paromomycin was added. Transformedcells were subsequently plated in wells of microtiter plates containing increasingconcentrations of paromomycin and were found to tolerate up to 2 mg/ml. Stockswere propagated by serial transfer in 160-�l batches of SPP medium in wells of

96-well flat-bottomed microtiter plates maintained at 30°C with transfer everysecond day. Additional cultures derived from single-cell isolates were established inwells of microtiter plates containing SPP medium with 1.8 mg/ml paromomycin.

Four somatically transformed clones were analyzed 200 generations aftertransformation by whole-cell PCR to detect the presence of ACT1 genes with andwithout the neo3 cassette (43). For each sample, cells in 150 �l of medium wereconcentrated by centrifugation, incubated for 1 h in 100 �l buffer K (0.03 mMproteinase K in 1� PCR buffer from Promega) at 55°C, and then boiled for 20min. Two aliquots from each lysate were subjected to PCR as described above.Using the two ACT1 primers described above, a PCR product of 1.38 kb is obtainedif the wild-type ACT1 gene is present. PCR using the ACT1 forward primer plus areverse primer that anneals to the neo3 internal sequence, 5�-AGAGTCCTTGGTCTTAACACT-3�, gives a 1.04-kb product when the transgene is present.

Germ line disruption of ACT1. The pAct-35-TOPO-neo3 construct (Fig. 1A)digested with BstXI was introduced into conjugating CU428 and B2086 cells by

FIG. 1. Disruption of the ACT1 gene. (A) A diagram showingpreparation of the ACT1 disruption construct and the targeting strat-egy. The neo3 disruption cassette was inserted into the unique SfuI sitein the ACT1 coding region of pAct-35-TOPO, thereby creating thepAct-35-TOPO-neo3 targeting construct. The BstXI sites were used tofree that construct prior to transformation. The targeting constructreplaced the endogenous ACT1 gene by homologous recombination,generating a “knockout allele” that contained a 3,070-bp insertion ofthe neo3 cassette. (B) Genotypic analysis of the wild-type cells andhomozygous ACT1� homokaryons. Total genomic DNA isolated fromCU428 wild-type cells and ACT1� cells was analyzed by PCR usingACT1-specific primers (arrowheads) whose locations are indicated inA. A 1.2-kb band was amplified from the wild-type (WT) cells (centerlane). In the homozygous ACT1� “knockout” (KO) homokaryons, the1.2-kb band was absent, and a 4.2-kb major band containing the neo3insert was obtained (right lane). The left lane shows the molecular sizemarkers. (C) RT-PCR analysis to assay ACT1 expression. Total RNAextracted from CU428 and ACT1� cells was analyzed by RT-PCRusing the same ACT1-specific primers described in B. A 1.2-kb ACT1cDNA band (arrowhead) was amplified from the wild-type (WT) cellsbut not from the ACT1 knockout (KO) cells. For the control, primersspecific to ribosomal protein gene RPL21 were included in the samereaction. The 0.36-kb RPL21 cDNA band (asterisk), but not the0.84-kb band that would be expected from the amplification of con-taminating genomic DNA, was obtained from both wild-type andknockout cells. Illustrations were prepared for publication on a Macin-tosh G4 computer using Photoshop CS software. All adjustments wereapplied solely to improve the clarity of the images.

556 WILLIAMS ET AL. EUKARYOT. CELL


.org/D

ownloaded from

http://ec.asm.org/

biolistic particle bombardment 3 h after mixing (7). The germ line transformantswere selected, assorted, and made homozygous for the knockout allele (Fig. 1A)in the micronucleus as described (25). ACT1�F10ns and ACT1�C3ns, twoheterokaryon lines with different mating types, were obtained. These cells eachhave the ACT1 gene with the drug resistance disruption cassette (the knockoutallele in Fig. 1A) in their micronuclear genome and have phenotypically assortedtheir macronuclear genome to wild-type ACT1 alleles.

ACT1�F10ns and ACT1�C3ns heterokaryon cells were mated and individualconjugating pairs were isolated between 6 and 10 h postmixing and transferredinto SPP medium drops to finish conjugation. To determine if progeny exhibitedthe same phenotypes as the somatic knockout cells, these progeny were exam-ined with an Olympus SZH-ILLD dissecting microscope every 6 to 8 h. After 2days, progeny cells were transferred to microtiter plates and further selected with120 �g/ml paromomycin and MEPP medium with 1 �g/ml CdCl2 to eliminate theparomomycin-sensitive cells derived from aborted conjugants. The true progenycells (homozygous ACT1� homokaryons) were then maintained in rich medium(PPYGFe or MEPP) in Erlenmeyer flasks with vigorous shaking.

For analysis of the genotype of the homozygous ACT1� homokaryons,genomic DNA extracted from CU428 and ACT1� cells was analyzed by PCR(43) using primers (B1-ACT1-F10, 5�-ATAGGATCCATGGCTGAAAGTGAATCCCCCGCT-3�, and A1-ACT1-R10, 5�-TAAGGCGCGCCGAAGCACTTTCTGTGGACGATG-3�) designed to amplify the whole ACT1 open readingframe (ORF) (Fig. 1A). PCR was performed using TripleMaster mix (Eppen-dorf).

For reverse transcription (RT)-PCR analysis to assess ACT1 expression, totalRNA from CU428 and ACT1� cells in vegetative growth was extracted usingTrizol reagent (Invitrogen) following the manufacturer’s instructions; 2 �g oftotal RNA was reverse-transcribed using SuperScript III (Invitrogen) with oli-go(dT) primers at 50°C for 60 min following the manufacturer’s instructions.

Multiplex PCR (94°C, 30 seconds; 50°C, 30 seconds; 72°C, 1 min for 23 cycles)was performed using the B1-ACT1-F10 and A1-ACT1-R10 primers to detect theACT1 message. A pair of specific primers (L21Fw: 5�-AAGTTGGTTATCAACTGTTGCGTT-3�, and L21Rv: 5�-GGGTCTTTCAAGGACGACGTA-3�), whichflank the first two introns of the ribosomal protein RPL21 gene and amplify a 0.36-kbregion from the cDNA or a 0.84-kb region from the genomic DNA template,respectively, were used in the same reaction as an internal control.

Rescue experiment. The whole ACT1 ORF was obtained by PCR using B1-ACT1-F10 and A1-ACT1-R10 primers and CU428 genomic DNA template. ThePCR product was directionally ligated between a 1.6-kb 5�-flanking sequence anda 0.5-kb 3�-flanking sequence of the MTT1 gene (58) and cloned into pBlueScriptvector (Stratagene).

For the MTT-ACT1-3�HA construct, a triple hemagglutinin (3�HA) oligo-nucleotide was inserted in frame downstream of the initiator ATG of MTT1 andupstream of the 5� end of the ACT1 ORF to add a 3�HA tag at the aminoterminus of Act1p. For the MTT-ACT1-FLAG construct, an oligonucleotidecarrying a FLAG epitope was incorporated in frame at the 3� end of ACT1 ORFto add a single FLAG tag at the carboxyl terminus of Act1p. To retransform theACT1� cells, the MTT-ACT1-3�HA and MTT-ACT1-FLAG rescue constructswere digested with SacII and XhoI and introduced into vegetative cells bybiolistic bombardment. For a negative control, cells were bombarded with theempty vector. Cells were incubated in SPP medium with 1 �g/ml of CdCl2 for 4 hat 30°C and then aliquoted to microtiter plates. Rescued swimming cells wereobserved after about 2 to 3 days.

Phenotypic analyses of living cells. Cells were examined using an OlympusCK2 inverted microscope, and photographed using a Leica inverted modelDMIRBE bright-field microscope equipped with a Hamamatsu Orca digitalcamera, model C4742-5. Living homozygous ACT1� homokaryons were ob-served in drops on a petri dish kept in a moist chamber. More extensive micro-scopic observations on individual cells and their vegetative progeny were made atintervals after isolation of single cells into 2- to 5-�l drops of culture mediumplated on plastic petri plates (Fisher) and then covered with a thin layer of lightmineral oil (Fisher, 021-1) (17, 44). These microdrop cultures were maintainedat room temperature (22 to 24°C) and periodically examined using a ZeissStereomicroscope-II and an Olympus CK2 inverted microscope.

Preliminary low-power observations of food vacuole uptake by live cells werecarried out using carmine (Alum Lake, Fisher) suspended in culture medium ata concentration of 0.1 �g/ml. In a subsequent more detailed analysis, live cellswere washed in 10 mM Tris (pH 7.5) and incubated in Higgin’s ink (concentra-tion of about 0.5 �l ink per 1 ml of cells) at 30°C for 1 h. Cells were pelleted at365 � g for 2 min and then fixed for 30 min with 2% paraformaldehyde in PHEMbuffer (60). After fixation, cells were washed and resuspended in phosphate-buffered saline (60), and wet-mount samples were observed directly by interfer-ence microscopy.

Cilium regeneration and measurement of cilium length. Cells were deciliatedand then allowed to regenerate cilia for 2 h, as previously described (6, 55).Measurements were made either on cilia still attached to cells or on isolatedaxonemes taken from the supernatant from cells that had regenerated cilia; bothwere fixed with 2% paraformaldehyde in PHEM buffer with 0.05% Triton X-100.Cells or axonemes were stained with rabbit polyclonal antibody raised againstTetrahymena tubulin (1:500) (66) and images were taken with the Leica TCS SPconfocal microscope. The length of axonemes was measured by ImageJ softwareobtained from http://rsb.info.nih.gov/ij.

Immunofluorescent studies. The axonemes from MTT-ACT1-3�HA rescuedcells were prepared as described (24). Cells were fixed in 2% paraformaldehydein PHEM or in Triton-ethanol (68) followed by staining with anticentrin mono-clonal antibody 20H5 (1:500), kindly provided by Jeffrey Salisbury of the MayoClinic. In some preparations, cells were doubly stained with anticentrin (1:500)and with the monoclonal antibody 9A9 (1:5) that stains contractile vacuole pores(33). Cilia and cortical microtubule bands were visualized with polyclonal anti-bodies raised against T. pyriformis tubulins (66, 67) and secondary goat anti-rabbit immunoglobulin G fluorescein isothiocyanate conjugate (1:500, Sigma-Aldrich). 3�HA-tagged Act1p was stained with mouse anti-HA antibody(16B12, 1:500, Sigma-Aldrich) and secondary Alexa Fluor 568 goat anti-mouseimmunoglobulin G (1:500, Invitrogen). Nuclei were stained with 4�,6�-diamidino-2-phenylindole (DAPI) as described (60), and stained cells were examined in theDMIRBE inverted microscope equipped with a Chroma fluorescent cube. Fixedand stained cells were observed with a Leica TCS SP confocal microscope (PLApo 100� lens, N.A. 1.4) (Fig. 3A and B and 4) or with an Olympus BH-2fluorescent microscope (Fig. 3C and 9).

Electron microscopy. Cells were fixed in 1.2% glutaraldehyde in 0.1 M phos-phate buffer (pH 7.0), postfixed in 1% osmium tetroxide in the same buffer,dehydrated in an ethanol series, and embedded in Eponate 12. Sections were cutand stained with uranyl acetate and lead citrate, and cilia were examined in aHitachi H7000 transmission electron microscope.

RESULTS

Targeted disruption of ACT1. Somatic ACT1� cells wereobtained by transformation of macronuclei with the ACT1disruption plasmid construct (pAct-35-TOPO-neo3) followedby selection in paromomycin with cadmium chloride present.Genes in the ciliate macronucleus are borne by DNA mole-cules that lack centromeres and segregate randomly duringmacronuclear division, bringing about the phenomenon knownas phenotypic assortment (47, 70). In time, genes being selectedagainst will no longer be represented among the progeny. In-creasing concentrations of paromomycin impose a strong se-lection favoring macronuclear chromosomes containing theneo3 disruption cassette such that all cells should eventuallyhave macronuclei that are fully assorted for the knockout al-lele, unless there is countervailing selection for maintenance ofcopies of the wild-type allele.

The persistence of phenotypic heterogeneity as revealed byclonal analysis (see below) as well as the strong signals ob-tained in PCR-based assays with both nondisruption and so-matic disruption primer sets (see Materials and Methods) to-gether strongly suggest that the somatic ACT1� transformantsmaintained some wild-type alleles in the macronucleus. Weconclude that the ACT1� knockout allele probably could notbe driven to fixation in growing populations following somaticdisruption of the wild-type ACT1 allele, although such fixationmay have occurred in some individual cells within these pop-ulations (see below).

Using the pAct-35-TOPO-neo3 disruption construct, homo-zygous ACT1� homokaryon cells were created by germ line trans-formation (25). To examine the genotype of these homozygousACT1� homokaryon cells, genomic DNA was extracted and sub-jected to PCR analysis. As shown in Fig. 1B, the ACT1 primers

VOL. 5, 2006 DISRUPTION OF THE ACT1 GENE OF T. THERMOPHILA 557


.org/D

ownloaded from

http://ec.asm.org/

used specifically amplified the 1.2-kb ACT1 coding region us-ing DNA from wild-type cells. When the same primers wereused with ACT1� DNA, however, a 4.2-kb band correspondingto the disrupted ACT1 allele was obtained and no 1.2-kb bandwas visible, confirming that all of the ACT1 genes had beendisrupted in both the macronucleus and micronucleus.

In wild-type cells, a 1.2-kb band (Fig. 1C, arrowhead) wasamplified by RT-PCR using ACT1-specific primers. However,RT-PCR failed to amplify an ACT1 transcript from ACT1�cells under the same conditions, while the control primersspecific to ribosomal protein RPL21 amplified the 0.36-kbcDNA band (Fig. 1C, asterisk) in the same multiplex PCR.This result showed that ACT1 message was not detected in theACT1� cells.

Phenotypes of living cells with disrupted ACT1 genes. Themost obvious effect of the somatic ACT1 disruption was theappearance of many cells that seemed to be blocked in fissionafter at least 50 generations in progressively increasing con-centrations of paromomycin (Fig. 2A). Examination by lightmicroscopy of living cells (e.g., Fig. 2F) and by DAPI stainingindicated that the macronuclei of these fission-blocked cellswere completely divided (see also Fig. 9B).

The homozygous ACT1� homokaryons exhibited the samephenotype, but much more uniformly. Individual conjugatingpairs of ACT1� heterokaryons were isolated into drops of SPPmedium on a petri dish and examined with a dissecting micro-scope. Initially the exconjugants could swim and divide afterconjugation was complete. After three to four divisions, how-ever, cells started to lose their motility and cells that had failedto complete cytokinesis could easily be identified. After five orsix divisions, cells became severely paralyzed and lost theirnormal motility; the motility that remained consisted of a slowand spasmodic oscillatory motion (see films A and B in thesupplemental material to compare locomotion in wild-typecells and in ACT1� cells). Tandem chains of two to four cellsthat had failed to separate (similar to those shown in Fig. 2)also became more common. These observations indicated thatafter five to six rounds of divisions from a single pair, theremaining wild-type Act1p from the parental pool was nolonger sufficient to support motility in the progeny cells. Suchnearly paralyzed cells were used for a more detailed clonalanalysis (see below).

Close observation of dividing ACT1� cells indicated that allsuch cells developed an advanced cleavage furrow. Cell sepa-ration was delayed or arrested at a terminal stage of cytokine-sis, when the connection between the two daughters was ex-tremely narrow. There was no evidence for blockage at earlierstages of cytokinesis, strongly suggesting that the various ap-parent fission-arrested phenotypes seen in Fig. 2A were con-sequences of progressive reintegration of cells that had suc-ceeded in furrowing but not in separating.

Figures 2B to D, albeit taken of different cells, illustrate themost likely course of division in the affected cells (see also filmB in the supplemental material). Since all living cells observedin division completed constriction but failed to separate priorto arrest, the configurations seen in Fig. 2E and F as well as inFig. 2A can most easily be interpreted as stages of regression ofpreviously completed furrows. Cells may then persist with re-gressed furrows, form new furrows while retaining a linearorder (Fig. 2G), undergo an anterior telescoping of the poste-

rior presumptive daughter cell (cell 5 in Fig. 2A), or carry outother spatial readjustments (Fig. 2 H and I) ultimately leadingto the formation of irregular monsters (Fig. 7D and 9I).

Rescue of motility. To further confirm that the loss of mo-tility was caused by the inactivation of ACT1, a rescue exper-iment was performed by reintroducing the wild-type ACT1DNA with a triple HA tag into the nonmotile ACT1� cells.The ACT1� cells were transformed with the MTT-ACT1-3�HA construct or empty vector and maintained in microtiter

FIG. 2. Living, nearly immobile cells from cultures of somaticACT1� transformants photographed on the bottom of the microtiterculture wells. (A) A small region at the bottom of a well, showing fivecells in various (numbered) stages of furrow regression following arrestat the final stage of cleavage, as well as four nondividing cells. The cellnumbered 5 has undergone an anteriorward telescoping of the poste-rior daughter cell. (B to F) Different cells photographed in successivestages of cleavage and reintegration. (B) Early fission. (C) Completionof furrowing. (D) Re-elongation of presumptive daughter cells thathave failed to separate. The posterior daughter is shown pivoted rel-ative to the anterior daughter, probably in an attempt to separate fromit. (E) A dividing cell beginning to reintegrate; the connection betweenthe presumptive daughter cells is broadening. (F) A more fully inte-grated divider, with a broad cytoplasmic union and separated macro-nuclei (arrowheads). (G to I) Examples of alternative developmentalconsequences of the permanent reintegration of presumptive daughtercells. (G) An attempted second division of one of the two daughtercells. (H) Displacement of the axes of the two daughter cells. (I) Prob-able twisting of the two presumptive daughter cells into a heteropolarconfiguration. All of the photographs are printed at the same magni-fication. Bar, 50 �m.



.org/D

ownloaded from

http://ec.asm.org/

plates in SPP with 1 �g/ml of CdCl2 to induce expression of thetagged Act1p. After 3 days, swimming rescued cells could befound in more than two-thirds of total microtiter wells, while inthe negative control plates transformed with vector DNA nocell gained normal motility. The rescued cells looked indistin-guishable from wild-type cells. Thus, restoration of normalmotility in the mutant cells demonstrated that knockout ofACT1 indeed caused the ciliary defect. A similar rescue wasachieved with the MTT-ACT1-FLAG construct.

Analysis of the ciliary defect. Examination of nearly para-lyzed cells both among the somatic ACT1� transformants and

in the homozygous ACT1� homokaryons indicated that full-length cilia were present (Fig. 3 and 6), suggesting that in thiscase, unlike the previously described T. thermophila mutantswith paralysis and subsequent failure of cell separation (3, 5),there was no loss of cilia. Cilia could also regenerate nearlynormally following deciliation. First, both wild-type cells andhomozygous ACT1� homokaryons regenerated cilia with sim-ilar kinetics in the recovery buffer (data not shown). Second,regenerated cilia measured on intact ACT1� cells averaged 4.8 �0.3 �m (n � 36) in length compared to 5.4 � 0.5 �m (n � 37)in parallel CU428 cells; when isolated, regenerated cilia weremeasured, the corresponding lengths were 4.8 � 0.6 �m (n �83) for the ACT1� cells, 4.9 � 0.8 �m (n � 84) for CU428cells, and 5.6 � 0.8 �m (n � 77) for the MTT-ACT1-3�HArescued cells.

While the average ciliary measurements for the wild-typeand rescued cells were always slightly greater than those for theACT1� cells (significantly so for cilia measured on intact cells),it was nonetheless clear that the disruption of the ACT1 geneimposed no major impairment on the formation of new cilia.Close observations of ACT1� and wild-type cells under phase-contrast microscopy revealed that the cilia in mutant cells beatat a much lower frequency than those in wild-type cells, andthe beating was not metachronal (see film B in the supplemen-tal material). Unlike the constant and fast ciliary beating inwild-type cells, the mutant cilia made one or several strokes,paused, and then made another one or several strokes. All ofthese observations argue that ACT1 plays a role in ciliarymotility but is not required for the assembly of an axoneme.

Our evidence for a functional deficit in cilia of ACT1� cellsmade it desirable to confirm an earlier report (46) that Act1pis present in ciliary axonemes of T. thermophila. This was dem-onstrated in homozygous ACT1� homokaryons rescued by theMTT-ACT1-3�HA construct (Fig. 4). Comparison of theslightly offset antitubulin and anti-HA images (Fig. 4B) indi-cated that Act1p and tubulin were largely colocalized in theisolated cilia. There were very few regions where Act1p wasvisualized in the absence of coincident tubulin, but more places

FIG. 3. Nonmotile cilia in homozygous ACT1� homokaryons (Aand B) and in a division-arrested somatic ACT1� transformant (C) im-munostained with antitubulin antibodies. A and P indicate the anteriorand posterior ends of cells, respectively. Bar, 10 �m. (See also film Bin the supplemental material).

FIG. 4. Immunolocalization of ciliary actin. Three frames from the same confocal image of isolated axonemes from homozygous ACT1�homokaryons rescued with an MTT-ACT1-3�HA construct and stained with antitubulin (A, fluorescein isothiocyanate) and with anti-HA (C,Alexa Fluor 568). The central panel (B) is an overlay of A and C with a slight displacement to allow comparison of the structures labeled by thetwo fluorochromes. The arrows in A and B point to a single ciliary axoneme labeled by antitubulin, whereas the arrowheads in B and C indicatethe same axoneme labeled by anti-HA. Bar, 10 �m.



.org/D

ownloaded from

http://ec.asm.org/

where tubulin immunofluorescence was seen without corre-sponding anti-HA fluorescence. This was most likely due to thepresence of an additional ATG in the MTT-ACT1-3�HA con-struct, located between the 3�HA epitope tag and the ACT1gene. The existence of a second translational start signal down-stream from the sequence encoding the 3�HA tag implies thatsome of the Act1p present in the cilia might not possess theepitope tag. Intracellular immunostaining of cells transformedwith this construct was variable; however, in no case did wedetect localization of the 3�HA-Act1p in the division furrow.

Cilia of cells with disrupted ACT1 genes, both somaticACT1� transformants and homozygous ACT1� homokaryons,were studied by transmission electron microscopy. The major-ity of ciliary cross sections were similar to those of wild-typecells (as shown, for example, in Fig. 8 of reference 69), but 1 to5% had relatively subtle but distinct abnormalities, notablyincomplete doublet microtubules, most commonly the B tubule(Fig. 5A, B, and D), but occasionally the A tubule (Fig. 5C).Very few cilia were grossly abnormal (not shown). Thisstrongly suggests that the ciliary defect is related to an inabilityto assemble fully normal axonemes.

Impairment of food vacuole formation. Preliminary feedingexperiments using carmine particles, with observations at lowpower, revealed that whereas all nondividing wild-type cells

took up numerous large particles within 5 min after addition,most of the homozygous ACT1� homokaryons failed to takeup carmine, and only a minority of about 10% took up a few ofthe smallest particles over a period of hours. A more detailedexamination using black ink indicated that while the wild-typecells took up numerous large ink particles that became con-centrated near the rear end of the cells (Fig. 6A and B), thegreat majority of the ACT1� cells took up only a small amountof ink, and that was almost invariably wedged in the oralapparatus (Fig. 6C and D). These results are reminiscent ofthose obtained earlier (49) for cells of T. pyriformis exposed tocytochalasin B, except that the amount of ink trapped in theoral region was far less in the ACT1� cells of T. thermophilathan in the cytochalasin-inhibited cells, which were not re-ported to have impaired motility. These results strongly sug-gest that Act1p is directly as well as indirectly required forsuccessful phagocytosis (see Discussion).

Clonal analysis. Clones of homozygous ACT1� homokary-ons were followed for extended periods in microdrops of cul-ture medium to determine the effects of absence of the ACT1gene on cell division, the cell cycle, and long-term survival.

About 2 weeks after the creation of the ACT1� homokary-ons, 50 normal-appearing cells were isolated into microdrops,25 into SPP and 25 into PPYGFe, and 24 control cells of theACT1 heterokaryon parental strain were isolated into SPP.Control isolates all generated vigorously growing populationsof cells swimming rapidly throughout the microdrop (Fig. 7A).In contrast, cells isolated from ACT1� cultures all crept ex-tremely slowly on the plastic substratum of the drop, and eachgenerated variable numbers of nearly paralyzed progeny (Fig.7B to D). No ACT1� cell ever recovered normal or near-normal motility.

Among ACT1� cells that were followed closely through theirfirst two division cycles, all progressed to the late constrictionstage. The proportion of success of separation in the first twoattempts at division was 43% in SPP (n � 40) and 69% inPPYGFe (n � 48), a statistically significant difference (2 �

FIG. 5. Electron micrographs of sections through cilia having ab-normal outer doublet microtubules in somatic ACT1� transformants(A to C) and in a homozygous ACT1� homokaryon (D). (A) A mod-erately abnormal cilium with abnormal detachment of B tubules fromthe A tubules (arrowheads). (B) A more severely abnormal ciliumincluding another example of a detached B tubule (arrowhead). (C) Amoderately abnormal cilium with a detached A tubule (arrowhead).(D) A moderately abnormal cilium with a detached B tubule (arrow-head) and an extra incomplete microtubule (arrow). Magnification,120,000�.

FIG. 6. Differential interference contrast images of uptake of inkparticles in fixed CU428 cells (A and B) and in homozygous ACT1�homokaryons (C and D). Arrows indicate ink particles in the region ofthe oral opening (cytostome), most likely in transit (A) or trapped (C).Cilia fixed in various phases of the normal beat cycle are also faintlyvisible in A and B, whereas outstretched nonmotile cilia are moreeasily seen in C and D. Bar, 10 �m. (See also films A and B in thesupplemental material).



.org/D

ownloaded from

http://ec.asm.org/

6.12, 0.02 P 0.01). We have also observed that this diffi-culty in separation is associated with a defect in the finaltwisting apart of the nearly separated daughter cells, a processthat was named rotokinesis by its discoverers (4).

The average duration of the first cell cycle, measured as thetime between successive onsets of furrowing, was 6.6 h (range, 5.0to 9.5 h, n � 14) in SPP, and 5.9 h (range, 4.25 to 7.5 h, n � 15) inPPYGFe, both at 23 � 1°C. The average interfission interval in thecontrol heterokaryon clones was 3.1 h (range, 2.5 to 3.75 h, n � 44).

While the control heterokaryon clones reached stationaryphase with 200 or more cells per microdrop within 2 days,ACT1� clones grew slowly and variably, probably as conse-quences both of frequent failure to complete constriction andof lengthened cell cycles. The largest clones (Fig. 7B) included

many nearly normal cells and went through as many as 15 cellcycles after inception. Average clones, consisting of 20 to 30cells, generally included a variety of abnormal reintegratedcells, irregular monsters, and tiny cells that may have buddedoff from abnormal larger cells (Fig. 7C). The smallest cloneswere generally those that persistently failed in cell separation;they often included one or a few giant irregular monsters,derived from repeated unsuccessful attempts at cell separation(Fig. 7D).

Figure 8 gives a summary of the quantitative subclonal data.A comparison of histograms A and B indicates that the max-imum clone sizes of ACT1� cells were greater in the richerPPYGFe medium than in SPP, but there was no statisticallysignificant difference in the average clone sizes (Mann-WhitneyU test). Histogram C indicates that differences in clone size andproliferation rate are not heritable. Two subclonal sets, initi-ated with normal appearing cells from clones (a and c in Fig.8B) of very different sizes, both reiterated the range of popu-lation sizes seen in the original clones, with no significantdifferences between them and no consistent loss of prolifera-tive vigor. The subclonal set derived from clone b was strikinglydifferent; there were few normal-appearing cells left in clone bat the time of subcloning and the subclones were uniformlysmall and mostly died before the end of the experiment.

A clonal analysis was also carried out (in SPP) for cellsselected from mass cultures of somatic ACT1� transformants.In the majority (50 out of 60) of such clones, cells swam anddivided normally, and produced vigorous populations muchlike those of the heterokaryon controls described above. How-ever, 10 of the 60 selected cells gave rise to clones very similarto those described above for the homozygous ACT1� homo-karyons. In these clones, speed of swimming and frequency ofdivision slowed dramatically within a day after the selection,and many cells underwent a late-fission arrest. These resultsled us to suspect that the culture was heterogeneous; whereasmost somatic ACT1� transformants had not fully assorted,those cells that produced the “slow” clones probably had as-sorted to fixation or near-fixation of macronuclear chromo-somes with disrupted ACT1 alleles. This interpretation wassupported by the uniform results (described above) that werelater obtained with the homozygous ACT1� homokaryons.

These clonal analyses thus indicate that (i) the division ar-rest phenotype of the ACT1� cells is due solely to failure offully furrowed cells to complete their separation, (ii) this fail-ure of separation is invariably associated with loss of cell mo-tility, (iii) cell cycles of the ACT1� cells are substantiallylengthened but not halted, (iv) an abnormal cellular geometryresulting from a failure of cell separation impedes but does nottotally prevent subsequent cell division, and (v) T. thermophila.cells have the potential to live, grow and divide for a long timewithout the ACT1 gene, although that potential is not alwaysrealized.

Observations of immunostained cells with disrupted ACT1genes. Cortical configurations were examined in somaticACT1� transformants, primarily using the 20H5 anti-centrinmonoclonal antibody, which detects basal bodies, oral cres-cents, and the filament ring underlying the anterior crown ofbasal body couplets (33, 59). Oral crescents are absent at theonset of cell division (Fig. 9A) and develop in both sets of oralstructures during cell division (12, 18, 34). Therefore, cells that

FIG. 7. Photographs of clones of ACT1�C3ns heterokaryons(A) and homozygous ACT1� homokaryons (B, C, and D) cultivated inmicrodrops containing SPP (A, C, and D) or PPYGFe (B) medium.(A) A clone of phenotypically normal (�) heterokaryons, photo-graphed 24 h after clone initiation with a single cell. Approximately100 swimming cells are uniformly dispersed and therefore mostly outof focus. At least eight cells in various stages of division are in or nearthe focal plane at the bottom of the drop. (B) A portion of a microdropcontaining about one-half of the cells of the largest ACT1� clone (a inFig. 8B) photographed 14 days after clone initiation. These cells arenearly stationary in a single focal plane just above the plastic surface.Some cells are in various stages of division or integration. Othernondividing cells have single macronuclei and appear nearly normal.(C) An average ACT1� clone, photographed 11 days after initiation.All of the 27 nearly stationary cells are in focus just above the plasticsubstratum. One cell is in late division (arrowhead), another abnormalcell may be budding off a small daughter cell (long thin arrow), andthere is one large monster (short thick arrow). Most cells have theirregular rotund shape characteristic of cells that have reintegratedafter an aborted cytokinesis, and a few other cells are smaller and morespherical than normal. (D) An ACT1� clone, photographed 11 daysafter clone initiation. All of the cells in the clone are visible in theportion of the drop shown here. A large monster with many cellularsubunits is in the process of attempting to bud off a smaller cell(arrow). One small daughter cell had successfully separated from themonster on the eighth day after isolation. A third small cell appearedon the eleventh day. One of these two small cells is attempting todivide again (arrowhead). Bar, 100 �m.



.org/D

ownloaded from

http://ec.asm.org/

have well-developed oral crescents (OC in Fig. 9C, D, and F)are likely to have undergone cell division.

Examination of at least 200 somatic ACT1� transformantsfixed at all stages of cortical development (staging from refer-ence 1) up to the onset of division furrowing revealed noabnormalities in cortical patterns visible with anticentrin andantitubulin immunostaining. Most relevant, the oral primor-dium was always seen at its normal equatorial position alongthe same cell longitude as the anterior oral apparatus. Thisheld true even at the final stage before the onset of cytokinesis(Fig. 9A), when the fission zone has already formed as a com-plete equatorial ring of gaps in the longitudinal ciliary rows.

Cells that had undergone previous failure of cell separationalmost invariably had oral structures with normal compoundciliary structures (membranelles and undulating membrane)and normal oral crescents; however, they were abnormallypositioned, along different cell longitudes in the two presump-tive daughter cells (Fig. 9B). In cells that were at advanced

stages of regression of the division furrow, the posterior oralapparatus was virtually always on a different cell longitude thanthe anterior oral apparatus (Fig. 9C, D, and F). In one tally, 46out of 50 reintegrated cells, diagnosed as such by their matureoral apparatuses, had their two sets of oral structures on dif-ferent cell longitudes. Since this condition was never seen inpredividing cells, it was presumably generated by a lateraldisplacement of presumptive daughter cells during the abortedcell separation.

A consequence of this lateral displacement was that differentciliary rows abutted one another across the division furrow.Often, these rows did not rejoin, and polar structures weresometimes formed (Fig. 9E, AC2). Disorder and displacementof ciliary rows in the furrow region were commonly observed(Fig. 9F, arrow, and 9G, rows a and b).

Surprisingly, some ciliary rows did rejoin across the furrowplane. This is seen for a single row (arrowhead) in Fig. 9D, andfor three rows (a, b, and c) in Fig. 9F. This clearly was not a

FIG. 8. Cumulative number of cells produced in clones of homozygous ACT1� homokaryons cultivated in microdrops in SPP (A) or PPYGFe(B) medium for 15 days, and in subclones cultivated in microdrops of PPYGFe medium for 14 days (C). Each symbol represents a clone or subclone(circles, SPP; squares, PPYGFe), and the position of the symbol on the abscissa indicates the number of cells counted in that clone on day 15 (or14 for the subclones), or on days 10 to 14 in the few cases when there were late declines in cell number or losses for other reasons. Some but notall of the clones and subclones with 1 to 5 cells became moribund at various times before days 14 to 15, though few cells actually lysed. The threeclones used as founders of the subclones are labeled a, b, and c in the histogram (B); they each have their own distinctive symbol, which is usedto identify the source of each of the subclones in histogram (C).

FIG. 9. Images of somatic ACT1� transformants, immunostained with anticentrin monoclonal antibody 20H5 (A to F and H to I) or polyclonalantitubulin antibody (G). “Left” and “right” refer to the cell’s left and right, which is opposite to the viewer’s left and right, except in E. (A) Apredividing cell with compound ciliary structures of the anterior oral apparatus (OA) and the posterior oral primordium (OP) located along thesame longitudinal meridian. Ciliary rows (CR) with basal bodies (BB) are equatorially subdivided to make up a fission zone (FZ), the site of thefuture division furrow. (B) An optical cross section of a divider in an early stage of reintegration. The division furrow (DF), oral apparatuses (OA1and OA2), and divided macronuclei (Mac1 and Mac2) are visible. (C) A more fully reintegrated ex-divider (DF, regressed division furrow) witha fully formed oral crescent (OC) of the posterior oral apparatus (OA2) and lateral displacement of OA2 relative to its anterior counterpart (OA1).The presumptive anterior daughter cell possesses a normal anterior crown of basal body couplets (AC1), whereas the anterior end of the posteriordaughter cell has only a few brightly stained developing anterior basal body couplets (dAC2). (D) One surface of a partially reintegrated formerdivider. One ciliary row of the posterior presumptive daughter cell (marked with an arrowhead at its posterior end) appears to be continuous witha ciliary row of the anterior daughter cell. (E) The opposite surface of the same cell as in D. About eight ciliary rows of the posterior daughtercell have formed a partial anterior crown of basal body couplets (AC2). (F) A more completely reintegrated former divider with three ciliary rows(marked a, b, and c) secondarily rejoined across the former fission zone. Other ciliary rows (arrow) have become disrupted and fragmented nearthe fission zone. (G) Antitubulin staining of a reintegrated cell. OA2 is on the viewer’s left side of the cell, and OA1 is on the opposite cell surfaceand therefore is not visible in this section. Ciliary basal bodies (BB) are indistinctly visible, with prominent longitudinal microtubule bands (LM)immediately to their right, and transverse microtubule bands (TM) extending to their left. Cilia (C), some detached from their cellular moorings,are brightly stained. The LMs on two ciliary rows (marked c and d) appear to be continuous across the former fission zone, and those on a third(marked e) have a small equatorial gap (arrow). Two other ciliary rows, marked a and b, are laterally offset at the fission zone. (H) A failed dividerthat has attempted to divide again. The less distorted posterior fission zone (arrow) is likely to be the more recent one. (I) An irregular monsterrepresenting the end state of successive aborted divisions. An oral apparatus (arrow) has normal membranelles, undulating membrane, and an oralcrescent. Bar, 10 �m.



.org/D

ownloaded from

http://ec.asm.org/



.org/D

ownloaded from

http://ec.asm.org/

rejoining of the same rows that had been together prior todivision. Note, in Fig. 9F, the major lateral displacement of theposterior relative to the anterior oral structures, as well as thecircumstance that three ciliary rows are interposed betweenthe rejoined ciliary rows labeled b and c posterior to OA2, withonly one such row interposed anterior to OA2. The microtu-bular assemblies (longitudinal microtubule bands) associatedwith the basal bodies sometimes also rejoined (Fig. 9G, rows cand d).

When the presumptive daughter cells retained their linear or-der, they could attempt to divide again, probably along newlyformed fission zones (Fig. 9H). Eventually, many such cellsformed irregular monsters in which the oral apparatuses werechaotically distributed albeit still internally normal (Fig. 9I, arrow).

DISCUSSION

Phenotypic effects of ACT1 disruption. Our analysis of thecellular role of Act1p took advantage of homologous genereplacement by biolistic transformation in Tetrahymena ther-mophila (7) to disrupt ACT1. The effects of this disruption,achieved in two different ways, are summarized in Table 1.First of all, this gene disruption led to a nearly complete loss ofcellular motility, despite the continued presence of full-lengthcilia that retained their normal capacity to regenerate afteramputation. We found that this near-paralysis is due to a greatdecrease of the ciliary beating frequency and the loss of syn-chronization, indicating that, as in Chlamydomonas reinhardtii(39), a conventional actin is required for ciliary motility inTetrahymena thermophila. Consistent with this conclusion, weconfirmed the presence of Act1p in Tetrahymena thermophilacilia (Fig. 4). As the tagged actin that was used to visualize thislocalization probably coexisted in the cell with an untaggedrescuing actin (see Results), the images in Fig. 4 might notreveal the full extent of localization of actin within cilia.

The contrast between a dramatic effect of gene disruptionand an absence of any reported effect of cytochalasin on cellularmotility is consistent with the same contrast reported earlier inChlamydomonas reinhardtii (see the introduction). Among sev-eral possible explanations for this discrepancy, cytochalasinscould have difficulty in entering the intraciliary space, or theconformation or molecular associations of actin in cilia mightrender them inaccessible to these drugs; there is also a remotepossibility that Act1p might be insensitive to cytochalasins, sothat the reported effects of these drugs would be due to inac-tivation of one or more of the other three Tetrahymena actins.

An association of actin with inner-arm dynein chains wasdemonstrated in C. reinhardtii. (39, 71) and in T. thermophila(46) by fractionation and Western blotting. Because Tetrahy-mena actin differs significantly from other actins, the associa-tion of actin with inner-arm dynein may be a conserved prop-erty of actin (46). The association was also visualizedultrastructurally in Chlamydomonas reinhardtii by showing asubtle but definite alteration of the inner arms in the actin-minus ida5 mutant (52). For this reason, we examined theultrastructure of cilia in T. thermophila cells with disruptedACT1 genes. Although we were unable to assess whether therewas a subtle change in the dynein arms, we did find a lowproportion of ciliary cross sections with incomplete micro-tubule doublets (Fig. 5).

The images encountered most frequently suggest that theACT1� cells experience difficulty in the proper attachment ofthe B tubule to the A tubule. This suggests that Act1p may beinvolved, either directly or indirectly, in the assembly of axo-nemal microtubules in Tetrahymena cells. Interestingly, anidentical defect of incomplete B tubules was recently observedin a T. thermophila mutant lacking sites of �-tubulin polyglycy-lation (57), a posttranslational modification found mainly incilia (2). It has been suggested that polyglycylation regulatesbinding of axonemal microtubule-associated proteins (63). Wecan speculate that �-tubulin polyglycylation is required forproper binding or activity of Act1p on axonemal microtubules.Alternatively, both �-tubulin polyglycylation and Act1p couldbe cooperatively required for another component(s) which sta-bilizes the B tubule.

A second effect of ACT1 gene disruption (Table 1) was anearly complete absence of food vacuole formation in cellslacking normal ACT1 genes. The simultaneous inhibition ofcell motility complicates our interpretation of this result, sincethe absence of food vacuoles might be caused indirectly (byfailure to sweep particles into the buccal cavity) or directly (byfailure to form normal food vacuoles). A comparison of ourresults with an earlier observation (49) leads us to concludethat both causes operate simultaneously. Exposure of Tetrahy-mena pyriformis to cytochalasin B caused a large amount of amarker (carmine) to accumulate in the food vacuole-formingregion, while very little got into the cell proper (49). We ob-served a much smaller accumulation of our marker (ink) in thesame region (Fig. 6). This reduced uptake is most likely due toweaker ciliary currents in the oral region, but the identical siteof accumulation suggests a defect in internalization of foodvacuoles, in this case brought about genetically rather thanpharmacologically. This interpretation is consistent with manyreports that actin is required for phagocytosis in ciliates.

Our initial observations suggested that Act1p might also beinvolved in division furrow constriction, since the most obviousphenotype emerging after disruption of the normal ACT1 genewas the appearance of “chains” of connected cells. However,this phenotype was found to be a consequence of cellularreintegration following a failure of separation of cells in whichcleavage furrowing had been completed, and it invariably oc-curred in cells that were virtually immobile and therefore ham-pered in the final twisting apart (rotokinesis) that is necessaryfor separation of most cells (4). The observed course of fissionand reintegration seen in living ACT1-disrupted cells was iden-tical to that recorded after disruption of two genes (KIN1 and

TABLE 1. Summary of effects of loss of Act1p in T. thermophila

Condition Impairment inACT1� cells

Direct orindirect

Ciliary and cellular motility Severe DirectPhagocytosis Severe BothCell division furrowing NoneMacronuclear division NoneCell separation Frequent IndirectBasal body formation NoneAxoneme assembly Slight Direct?Cell cycle Moderate IndirectLong-term viability Inconsistent Uncertain



.org/D

ownloaded from

http://ec.asm.org/

KIN2) encoding the Tetrahymena homologs of kinesin-2 (seeFig. 9 in reference 5). More recently, disruption of anothergene required for assembly of cilia, IFT52, was shown to alsolead to arrest of cytokinesis (3). Thus, cellular immobility,whether caused by loss of cilia (3, 5) or by impairment of ciliaryfunction as a consequence of ACT1 disruption (this study),prevents or retards rotokinesis (4) and thereby impairs cellseparation.

We also observed that even when cell separation failed andvarious bizarre cortical configurations arose as a consequence,new oral apparatuses remained normal, as did the generalarchitecture of the ciliary rows. Actin has been reported inthese structures in T. thermophila. (29) and in P. tetraurelia(41), but there is no evidence available yet that it is essentialfor their formation. It is possible that one of the other actinsmight be necessary and sufficient for formation of basal bodiesand of the oral apparatus in the absence of Act1p.

We conclude, therefore, that Act1p is required directly forcellular locomotion and for food vacuole formation in Tetra-hymena spp. and is required indirectly for normal cell separa-tion (Table 1). Act1p is not essential for the constriction of thedivision furrow or for the formation and arrangement of cor-tical structures made of basal bodies and associated cytoskel-etal components. The presence of Act1p in the division furrowhad been strongly suggested by its immunostaining with anantibody against an N-terminal peptide specific to Act1p (22).We failed to confirm this observation, but since we also failedfor technical reasons to get consistent internal immunostainingwith our MTT-ACT1-3�HA construct, our observations donot conclusively indicate that Act1p is absent from the divisionfurrow, only that if it is present there, it is not essential forfurrowing.

The contribution of the other genes in the actin gene familyto these various activities remains to be determined. In ourpreliminary investigation of the mRNA levels of other actingenes in the homozygous ACT1� homokaryons, expression ofat least one of the other actin genes was greatly induced whenACT1 was deleted (C.-C. Tsao and M. A. Gorovsky, unpub-lished data). We suggest that Act1p and other actin isotypesmight share partially, but not completely, redundant functions.

The multiplicity of Tetrahymena actins might also help toaccount for the differences between the results of ACT1 dis-ruption reported here and the effects of injection of skeletalmuscle actin (53) and of amplification of GFP-actin (32). Inour study, we specifically examined the physiological role ofAct1p; in both of the previous studies, the addition of a per-turbing agent may have had dominant-negative effects thatimpinge upon several different actins. Injected skeletal muscleactin might form complexes with any (or all) of the severalTetrahymena actins. For GFP-actin (32), this possibility is lessobvious, since the GFP coding sequence was fused to the sameACT1 gene that was disrupted in our study. However, this genewas carried on an rRNA gene-based processing vector (37)that was highly amplified; hence, it is likely to be even morehighly overexpressed than the ACT1 gene under control of theMTT1 promoter. If different actins compete for common li-gands, an abundant GFP-Act1p could interfere not only withthe function of native Act1p but also with the functions ofother actins by titrating shared factors; it might also generate

abnormalities on its own, as is known, for example, for GFP-centrin constructs (59).

The inability of ACT1� cells to take up food vacuoles limitsa major route of entry of nutrients and therefore might beexpected to retard cell growth and impair long-term viability.However, the availability of alternative routes of nutrient entrysuch as clathrin-mediated endocytosis (13, 51) and carrier-mediated uptake (30) allows survival and growth in the ab-sence of food vacuole formation in certain media (54). Wepropose that the doubling of generation time that we observedin ACT1� cells was not a direct consequence of loss of theACT1 gene but rather was an indirect consequence of forcingthe cells to use alternative, less efficient systems of nutriententry. A similar increase in generation time was seen in cellsthat were prevented from feeding normally by disruption of theciliary assembly gene IFT52 (D. Dave and J. Gaertig, personalcommunication). Likewise, our cloning experiments suggestedthat cells lacking a normal ACT1 gene have the potential tosurvive indefinitely since some (but not all) of the subclones ofACT1� cells maintained undiminished vigor when the experi-ment was terminated 6 weeks after the loss of the normalACT1 gene.

Laterally displaced cortical reintegration: a serendipitousexperiment. The cortical reintegration phenotype describedhere, in which fusion of different ciliary rows can take placeafter a lateral displacement of the presumptive daughter cellsrelative to each other, has no precedent in the great variety offission-block mutants collected over the course of two decades(17, 19, 35, 36, 42; Frankel, unpublished observations). Thisnovel phenotype arises because, unlike any of the conditionalfission arrest mutants, every reintegrated divider bearing anACT1 disruption cassette had at some previous time normallycompleted cytokinesis, with the two daughters connected onlyby a thin strand of cytoplasm, which in normal cells gets brokenby rotary twisting (rotokinesis). If that twisting should be in-sufficient to separate the daughters, they will then reintegrateat whatever alignment they happen to be in when the effort atseparation by rotation finally ends (Fig. 9B; notice that OA1and OA2 are not aligned). This seemingly random reintegra-tion puts different ciliary rows of the two daughter cells intoclose juxtaposition, after which they can either ectopically healtogether or remain separate. Cells lacking KIN1 and KIN2appear to undergo a similar lateral displacement during roto-kinesis (Fig. 3E in reference 5), followed by rejoining of ciliaryrows (Fig. 1C in reference 4) or anterior shifting of the pre-sumptive posterior daughter cell after reintegration (Fig. 3F inreference 5).

A reintegration of the longitudinal microtubule bands (Fig.9G) does not necessitate a rejoining of severed microtubules,because these bands consist of short microtubules that aretilted relative to the longitudinal axis of the cell, so that eachindividual microtubule of the longitudinal band is much shorterthan the cell (56).

If our interpretation of the origin of the lateral displace-ments observed in the reintegrated cells is correct, then the lossof motility brought about by disruption of the ACT1 genecaused Tetrahymena thermophila to carry out a type of exper-iment that was performed microsurgically on the large ciliateStentor coeruleus (61, 65). These experiments involved first ahorizontal severing of anterior and posterior portions of the



.org/D

ownloaded from

http://ec.asm.org/

same cell, then a rotation of the two subcells relative to oneanother, and finally a rejoining with a circumferential misalign-ment. The cell’s response could be assessed by observing thesevered longitudinal pigment stripes, which in Stentor coeruleusare located between the less visible ciliary rows. In the controlexperiment, in which the anterior and posterior subcells wererejoined without any lateral displacement, the pigment stripes,and presumably the ciliary rows between them, healed. How-ever, if there was a misalignment, the stripes did not heal, andthe cortex of either the basal or apical subcell subsequentlygrew anteriorly or posteriorly, replacing the other subcell;which of these two misaligned subcells prevailed depended onthe level at which the severing and rejoining took place, with atendency for the posterior subcell to dominate (65). Theseresults can be interpreted to reflect a structural “incompatibil-ity gradient” (see reference 16, p. 126–127) as a correlate of thevisible circumferential gradient in pigment stripe widths thatcharacterizes this organism (62); the different longitudes in thisorganism appear to be different in some way that makes mu-tual adjustment impossible.

A similar rotation and rejoining, accomplished in Tetrahy-mena thermophila by aborted rotokinesis, appeared to generateno such radical readjustments; at least some ciliary rows re-joined while the posterior oral apparatus remained at its nor-mal equatorial position (Fig. 9F and G). The absence of acircumferential incompatibility gradient is in accord with theabsence in Tetrahymena spp. of the visible structural gradationthat is obvious around the circumference of Stentor coeruleus.This also indicates that in some respects Tetrahymena mayexceed even Stentor’s legendary capacity for regulative reorga-nization.

ACKNOWLEDGMENTS

We express our gratitude to the following contributors: MartinGorovsky and Jacek Gaertig (valuable consultation), Martin Gorovsky(pMNBL plasmid), Michael Dailey and Leah Fuller (biolistic transfor-mation), Shelley Plattner (light microscopy) Chris Stipp (movie con-version), David Hoskinson (help with images), and Jeff Salisbury (anti-centrin antibody 20H5). We also thank Martin Gorovsky, Jacek Gaertig,Anne Frankel, and Nels Elde for critical reading of the manuscript.

This research was supported in part by a grant from the U.S. Na-tional Institutes of Health (GM-26973) to Martin A. Gorovsky of theUniversity of Rochester.

REFERENCES

1. Bakowska, J., E. M. Nelsen, and J. Frankel. 1982. Development of the ciliarypattern of the oral apparatus of Tetrahymena thermophila. J. Protozool.29:366–382.

2. Bre, M. H., V. Redeker, M. Quibell, J. Darmanaden-Delorme, C. Bressac, J.Cosson, P. Huitorel, J. M. Schmitter, J. Rossler, T. Johnson, A. Adoutte, andN. Levilliers. 1996. Axonemal tubulin polyglycylation probed with two mono-clonal antibodies: widespread evolutionary distribution, appearance duringspermatozoan maturation and possible function in motility. J. Cell Sci. 109:727–738.

3. Brown, J. M., N. A. Fine, G. Pandiyan, R. Thazhath, and J. Gaertig. 2003.Hypoxia regulates assembly of cilia in suppressors of Tetrahymena lacking anintraflagellar transport subunit gene. Mol. Biol. Cell 14:3192–3207.

4. Brown, J. M., C. Hardin, and J. Gaertig. 1999. Rotokinesis, a novel phe-nomenon of cell locomotion-assisted cytokinesis in the ciliate Tetrahymenathermophila. Cell Biol. Int. 23:841–848.

5. Brown, J. M., C. Marsala, R. Kosoy, and J. Gaertig. 1999. Kinesin II ispreferentially targeted to assembling cilia and is required for ciliogenesis andnormal cytokinesis in Tetrahymena. Mol. Biol. Cell 10:3081–3096.

6. Calzone, F. J., and M. A. Gorovsky. 1982. Cilia regeneration in Tetrahymena:a simple reproducible method for producing large numbers of regeneratingcells. Exp. Cell Res. 140:471–476.

7. Cassidy-Hanley, D., J. Bowen, J. H. Lee, E. Cole, L. A. VerPlank, J. Gaertig,

M. A. Gorovsky, and P. J. Bruns. 1997. Germline and somatic transforma-tion of mating Tetrahymena thermophila by particle bombardment. Genetics146:135–147.

8. Cohen, J., N. Garreau de Loubresse, and J. Beisson. 1984. Actin microfila-ments in Paramecium: localization and role in intracellular movements. CellMotil. 4:443–468.

9. Cooper, J. A. 1987. Effects of cytochalasin and phalloidin on actin. J. CellBiol. 105:1473–1478.

10. Cupples, C. G., and R. E. Pearlman. 1986. Isolation and characterization ofthe actin gene from Tetrahymena thermophila. Proc. Natl. Acad. Sci. USA83:5160–5164.

11. Dentler, W. L., and C. Adams. 1992. Flagellar microtubule dynamics inChlamydomonas: cytochalasin D induces periods of microtubule shorteningand elongation; and colchicine induces disassembly of the distal, but notproximal, half of the flagellum. J. Cell Biol. 117:1289–1298.

12. Diogon, M., C. Henou, V. Ravet, P. Bouchard, and B. Vigues. 2001. Evidencefor regional differences in the dynamics of centrin: cytoskeletal structures inthe polymorphic hymenostome ciliate Tetrahymena paravorax. Eur. J. Pro-tistol. 37:223–231.

13. Elde, N. C., G. Morgan, M. Winey, L. Sperling, and A. P. Turkewitz. 4November 2005. Elucidation of clathrin-mediated endocytosis in Tetrahy-mena reveals an evolutionarily convergent recruitment of dynamin. PLOSGenet. 1:e52. [Epub ahead of print.]

14. Fok, A. K., S. S. Leung, D. P. Chun, and R. D. Allen. 1985. Modulation of thedigestive lysosomal system in Paramecium caudatum. II. Physiological effects ofcytochalasin B, colchicine, and trifluoroperazine. Eur. J. Cell Biol. 37:27–34.

15. Frankel, J. 2000. Cell biology of Tetrahymena thermophila. Methods CellBiol. 62:27–125.

16. Frankel, J. 1989. Pattern formation: ciliate studies and models. OxfordUniversity Press, New York, N.Y.

17. Frankel, J., J. Mohler, and A. W. K. Frankel. 1980. Temperature-sensitiveperiods of mutations affecting cell division in Tetrahymena thermophila.J. Cell Sci. 43:59–74.

18. Frankel, J., and E. M. Nelsen. 2001. The effects of supraoptimal tempera-tures on population growth and cortical patterning in Tetrahymena pyriformisand Tetrahymena thermophila. J. Eukaryot. Microbiol. 48:135–146.

19. Frankel, J., E. M. Nelsen, and L. M. Jenkins. 1977. Mutations affecting celldivision in Tetrahymena pyriformis, syngen 1. II. Phenotypes of single anddouble homozygotes. Dev. Biol. 58:255–275.

20. Gavin, R. H. 1976. The oral apparatus of Tetrahymena pyriformis strainWH-6. II. Cytochalasin B inhibition of oral apparatus morphogenesis. J. Exp.Zool. 197:59–64.

21. Gavin, R. H. 1976. The oral apparatus of Tetrahymena pyriformis strainWH-6. III. The binding of 3H-cytochalasin B by the isolated oral apparatus.J. Exp. Zool. 197:65–70.

22. Gonda, K., M. Katoh, K. Hanyu, Y. Watanabe, and O. Numata. 1999. Ca2�

calmodulin and p85 cooperatively regulate an initiation of cytokinesis inTetrahymena. J. Cell Sci. 112:3619–3626.

23. Gronlien, H. K., T. Berg, and A. M. Løvlie. 2002. In the polymorphic ciliateTetrahymena vorax, the non-selective phagocytosis in microstomes changes toa highly selective process in macrostomes. J. Exp. Biol. 205:2089–2097.

24. Guerra, C., Y. Wada, V. Leick, A. Bell, and P. Satir. 2003. Cloning, local-ization, and axonemal function of Tetrahymena centrin. Mol. Biol. Cell 14:251–261.

25. Hai, B., J. Gaertig, and M. A. Gorovsky. 2000. Knockout heterokaryonsenable facile mutagenic analysis of essential genes in Tetrahymena. MethodsCell Biol. 62:513–531.

26. Hirono, M., H. Endoh, N. Okada, O. Numata, and Y. Watanabe. 1987.Tetrahymena actin: cloning and sequencing of the Tetrahymena actin geneand identification of its gene product. J. Mol. Biol. 194:181–192.

27. Hirono, M., Y. Kumagai, O. Numata, and Y. Watanabe. 1989. Purification ofTetrahymena actin reveals some unusual properties. Proc. Natl. Acad. Sci.USA 86:75–79.

28. Hirono, M., M. Nakamura, M. Tsunemoto, T. Yasuda, H. Ohba, O. Numata,and Y. Watanabe. 1987. Tetrahymena actin: localization and possible biolog-ical roles of actin in Tetrahymena cells. J. Biochem. (Tokyo) 102:537–545.

29. Hoey, J. G., and R. H. Gavin. 1992. Localization of actin in the Tetrahymenabasal-body cage complex. J. Cell Sci. 103:629–641.

30. Hoffmann, E. K., and L. Rasmussen. 1972. Phenylalanine and methioninetransport in Tetrahymena pyriformis. Characteristics of a concentrating, in-ducible transport system. Biochim. Biophys. Acta 266:206–216.

31. Hosein, R. E., S. A. Williams, and R. H. Gavin. 2005. Directed motility ofphagosomes in Tetrahymena thermophila requires actin and Myo1p, a novelunconventional myosin. Cell Motil. Cytoskel. 61:49–60.

32. Hosein, R. E., S. A. Williams, K. Haye, and R. H. Gavin. 2003. Expression ofGFP-actin leads to a failure of nuclear elongation and cytokinesis in Tetra-hymena thermophila. J. Eukaryot. Microbiol. 50:403–408.

33. Jerka-Dziadosz, M., L. M. Jenkins, E. M. Nelsen, N. E. Williams, R. Jaeckel-Williams, and J. Frankel. 1995. Cellular polarity in ciliates: persistence ofglobal polarity in a disorganized mutant of Tetrahymena thermophila thatdisrupts cytoskeletal organization. Dev. Biol. 169:644–661.

34. Jerka-Dziadosz, M., I. Strzyzewska-Jowko, U. Woysa-Lugowska, W. Krawc-



.org/D

ownloaded from

http://ec.asm.org/

zynska, and A. Krzywicka. 2001. The dynamics of filamentous structures inthe apical band, oral crescent, fission line and the postoral meridional fila-ment in Tetrahymena thermophila revealed by monoclonal antibody 12G9.Protist 152:53–67.

35. Kaczanowska, J., L. Buzanska, and M. Frontczak. 1992. The influence offission line expression on the number and positioning of oral primordia inthe cdaA1 mutant of Tetrahymena thermophila. Dev. Genet. 13:216–222.

36. Kaczanowska, J., L. Buzanska, and M. Ostrowski. 1993. Relationship be-tween spatial pattern of basal bodies and membrane skeleton (epiplasm)during the cell cycle of Tetrahymena: cdaA mutant and anti-membrane im-munostaining. J. Eukaryot. Microbiol. 40:747–754.

37. Karrer, K. M. 2000. Tetrahymena genetics: two nuclei are better than one.Methods Cell Biol. 62:127–186.

38. Kato, T., O. Kagami, T. Yagi, and R. Kamiya. 1993. Isolation of two speciesof Chlamydomonas reinhardtii flagellar mutants, ida5 and ida6, that lack anewly identified heavy chain of the inner dynein arm. Cell Struct. Funct.18:371–377.

39. Kato-Minoura, T., M. Hirono, and R. Kamiya. 1997. Chlamydomonas inner-arm dynein mutant, ida5, has a mutation in an actin-encoding gene. J. CellBiol. 137:649–656.

40. Kersken, H., J. Vilmart-Suewen, M. Momayezi, and H. Plattner. 1986. Fil-amentous actin in Paramecium cells: mapping by phalloidin affinity labelingin vivo and in vitro. J. Histochem. Cytochem. 34:443–454.

41. Kissmehl, R., I. M. Sehring, E. Wagner, and H. Plattner. 2004. Immuno-localization of actin in Paramecium cells. J. Histochem. Cytochem. 52:1543–1559.

42. Krzywicka, A., M. Kiersnowska, D. Wloga, and J. Kaczanowska. 1999. Anal-ysis of the effects of the cdaK1 mutation of Tetrahymena thermophila on themorphogenesis of the fission line. Eur. J. Protistol. 35:342–352.

43. Liu Y., K. Mochizuki, and M. A. Gorovsky. 2004. Histone H3 lysine 9methylation is required for DNA elimination in developing macronuclei inTetrahymena. Proc. Natl. Acad. Sci. USA 101:1679–1684.

44. Løvlie, A. 1963. Growth in mass and respiration rate during the cell cycle ofTetrahymena pyriformis. Compt. Rend. Trav. Lab. Carlsberg 33:377–411.

45. Metenier, G. 1984. Actin in Tetrahymena paravorax: ultrastructural localiza-tion of HMM-binding filaments in glycerinated cells. J. Protozool. 31:205–215.

46. Muto, E., M. Edamatsu, M. Hirono, and R. Kamiya. 1994. Immunologicaldetection of actin in the 14S ciliary dynein of Tetrahymena. FEBS Lett.343:173–177.

47. Nanney, D. L. 1964. Macronuclear differentiation and subnuclear assortmentin ciliates, p. 253–273. In M. Locke (ed.), The role of chromosomes indevelopment (Society for Developmental Biology Symposium, vol. 23). Ac-ademic Press, New York, N.Y.

48. Nelsen, E. M., J. Frankel, and E. Martel. 1981. Development of the ciliatureof Tetrahymena thermophila. I. Temporal coordination with oral develop-ment. Dev. Biol. 88:27–38.

49. Nilsson, J. R. 1977. Fine structure and RNA synthesis of Tetrahymena duringcytochalasin B inhibition of phagocytosis. J. Cell Sci. 27:115–126.

50. Nilsson, J. R., T. R. Ricketts, and E. Zeuthen. 1973. Effects of cytochalasinB on cell division and vacuole formation in Tetrahymena pyriformis GL. Exp.Cell Res. 79:456–459.

51. Nilsson, J. R., and B. Van Deurs. 1983. Coated pits and pinocytosis inTetrahymena. J. Cell Sci. 63:209–222.

52. Ohara, A., T. Kato-Minoura, R. Kamiya, and M. Hirono. 1998. Recovery offlagellar inner-arm dynein and the fertilization tubule in Chlamydomonasida5 mutant by transformation with actin genes. Cell Struct. Funct. 23:273–281.

53. Ohba, H., M. Hirono, M. Edamatsu, and Y. Watanabe. 1992. Timing offormation of Tetrahymena contractile ring microfilaments investigated byinhibition with skeletal muscle actin. Dev. Genet. 13:210–215.

54. Orias, E., and L. Rasmussen. 1976. Dual capacity for nutrient uptake inTetrahymena. IV. Growth without food vacuoles and its implications. Exp.Cell Res. 102:127–137.

55. Pennock, D. G. 2000. Selection of motility mutants. Methods Cell Biol.62:281–290.

56. Pitelka, D. R. 1961. Fine structure of the silverline and fibrillar systems ofthree tetrahyminid ciliates. J. Protozool. 8:75–89.

57. Redeker, V., N. Levilliers, E. Vinolo, J. Rossier, D. Jaillard, D. Burnette, J.Gaertig, and M.-H. Bre. 2005. Mutations of tubulin glycylation sites revealcross-talk between the C-termini of �- and �-tubulin and affect the ciliarymatrix of Tetrahymena. J. Biol. Chem. 280:596–606.

58. Shang, Y., X. Song, J. Bowen, R. Constanje, Y. Gao, J. Gaertig, and M. A.Gorovsky. 2002. A robust inducible-repressible promoter greatly facilitatesgene knockouts, conditional expression, and overexpression of homologousand heterologous genes in Tetrahymena thermophila. Proc. Natl. Acad. Sci.USA 99:3734–3739.

59. Stemm-Wolf, A., G. Morgan, T. H. Giddings, Jr., E. A. White, R. Marchione,H. B. McDonald, and M. Winey. 2005. Basal body duplication and mainte-nance requires one member of the Tetrahymena thermophila centrin genefamily. Mol. Biol. Cell 16:3606–3619.

60. Stuart, K. R., and E. S. Cole. 2000. Nuclear and cytoskeletal fluorescencemicroscopy techniques. Methods Cell Biol. 62:291–311.

61. Tartar, V. 1956. Grafting experiments concerning primordium formation inStentor coeruleus. J. Exp. Zool. 131:75–122.

62. Tartar, V. 1960. The biology of Stentor. Pergamon Press, Oxford, UnitedKingdom.

63. Thazhath, R., C. Liu, and J. Gaertig. 2002. Polyglycylation domain of �-tu-bulin maintains axonemal architecture and affects cytokinesis in Tetrahy-mena. Nat. Cell Biol. 4:256–259.

64. Tiggemann, R., and H. Plattner. 1981. Localization of actin in the cortex ofParamecium tetraurelia cells by immuno- and affinity-fluorescence micros-copy. Eur. J. Cell Biol. 24:184–190.

65. Uhlig, G. 1960. Entwicklungsphysiologische Untersuchungen zur Morpho-genese von Stentor coeruleus. Ehrbg. Arch. Protistenk. 105:1–109.

66. Van Dewater, L., III, S. D. Guttman, M. A. Gorovsky, and J. B. Olmsted.1982. Production of antisera and radioimmunoassays for tubulin. MethodsCell Biol. 24:79–86.

67. Williams, N. E., and J. E. Honts. 1987. The assembly and positioning ofcytoskeletal elements in Tetrahymena. Development 100:23–30.

68. Williams, N. E., J. E. Honts, V. M. Dress, E. M. Nelsen, and J. Frankel. 1995.Monoclonal antibodies reveal complex structures in the membrane skeletonof Tetrahymena. J. Eukaryot. Microbiol. 42:422–427.

69. Williams, N. E., and J. H. Luft. 1968. Use of a nitrogen mustard derivativein fixation for electron microscopy and observations on the ultrastructure onTetrahymena. J. Ultrastruct. Res. 25:271–292.

70. Wong, L., L. Klionsky, S. Wickert, V. Merriam, E. Orias, and E. P. Hamilton.2000. Autonomously replicating macronuclear DNA pieces are the physicalbasis of genetic co-assortment groups in Tetrahymena thermophila. Genetics155:1119–1125.

71. Yanagisawa, H. A., and R. Kamiya. 2001. Association between actin and lightchains in Chlamydomonas flagellar inner-arm dyneins. Biochem. Biophys.Res. Commun. 288:443–447.

72. Zackroff, R. V., and L. A. Hufnagel. 2002. Induction of anti-actin drugresistance in Tetrahymena. J. Eukaryot. Microbiol. 49:475–477.



.org/D

ownloaded from

http://ec.asm.org/

the actin gene act1 is required for phagocytosis, motility, and cell ... · cell separation of...

Documents