A Class II Histone Deacetylase Acts on Newly SynthesizedHistones in Tetrahymena�
Joshua J. Smith, Sharon E. Torigoe,† Julia Maxson,‡ Lisa C. Fish,§ and Emily A. Wiley*Joint Science Department, Claremont McKenna, Scripps, and Pitzer Colleges, Claremont, California 91711
Received 7 November 2007/Accepted 19 December 2007
Newly synthesized histones are acetylated prior to their deposition into nucleosomes. Following nucleosomeformation and positioning, they are rapidly deacetylated, an event that coincides with further maturation of thechromatin fiber. The histone deacetylases (HDACs) used for histone deposition and de novo chromatinformation are poorly understood. In the ciliate Tetrahymena thermophila, transcription-related deacetylation inthe macronucleus is physically separated from deposition-related deacetylation in the micronucleus. Thisfeature was utilized to identify an HDAC named Thd2, a class II HDAC that acts on newly synthesized histonesto remove deposition-related acetyl moieties. The THD2 transcript is alternatively spliced, and the major formcontains a putative inositol polyphosphate kinase (IPK) domain similar to Ipk2, an enzyme that promoteschromatin remodeling by SWI/SNF remodeling complexes. Cells lacking Thd2, which retain deposition-relatedacetyl moieties on new histones, exhibit chromatin and cytological phenotypes indicative of a role for Thd2 inchromatin maturation, including the proteolytic processing of histone H3.
In eukaryotes, DNA complexes with an octamer of histoneproteins (two each of histones H2A, H2B, H3, and H4) to forma repeating unit of chromatin called a nucleosome (23, 29, 50).The precise positioning of nucleosomes can regulate DNA-templated processes, such as replication, transcription, andrecombination. Nucleosome assembly and initial positioningoccur during the S phase of the cell cycle and are closelycoupled to passage of replication forks (28). During replica-tion, the existing nucleosomes are randomly segregated ontothe newly synthesized DNA (47), while new nucleosomes areformed de novo through the assembly of free histones in aspecific order. First, two H3/H4 heterodimers are depositedonto the DNA, followed by two H2A/H2B dimers (14). Prior totheir deposition, free histones H3 and H4 are acetylated onlysine (Lys) residues by cytoplasmic histone acetyltransferases(HATs) in patterns distinct from transcription acetylationpatterns (2, 21, 42, 53). The deposition acetylation promoteshistone assembly into nucleosomes through interactions withhistone chaperones, such as chromatin assembly factor 1(CAF-1), Hif1, and Asf1, that incorporate new H3 and H4specifically onto newly replicated DNA (1, 14, 45, 52). Thedeposition-related pattern on histone H4 is highly conservedfrom yeasts to humans and consists of diacetylated Lys5 andLys12, which correspond to Lys4 and Lys11 in the ciliate Tet-rahymena thermophila (2, 8, 46). Recently, acetylation of Lys91in the globular domain of H4 was identified as another mod-ification important for nucleosome assembly (60). The acety-
lation pattern on newly synthesized histone H3 is more vari-able between organisms. In Tetrahymena, Lys9 and Lys14 areacetylated, whereas in Drosophila, Lys14 and Lys23 are thepreferred sites (46), and in budding yeast, most new H3 mol-ecules are monoacetylated on Lys9, Lys14, Lys23, Lys27, orLys56 (25, 32). In contrast to H3 and H4, nascent H2A andH2B histones do not display any acetylation patterns that aredistinct from parental forms.
Once assembled into nucleosomes, new histones H3 and H4are rapidly deacetylated (2, 21), an event important for chro-matin maturation. When chromatin is replicated in vivo in thepresence of sodium butyrate to inhibit deacetylation, the re-sulting nucleosome structure and distribution on the chroma-tin fiber appear normal, but the “mature” fiber is abnormallyhypersensitive to DNase I (7, 44). Chromatin assembled invitro with hyperacetylated histones is also more nuclease sen-sitive (24). Aside from a recent demonstration that deacetyla-tion of Lys91 on histone H4 facilitates the formation of a saltbridge to histone H2B, little is known about the role of histonedeacetylation in chromatin maturation. Further studies in thisarea have in part awaited identification of the histone deacety-lases (HDACs) involved.
Most HDACs identified to date fall into three phylogeneticclasses, depending on their homology to the yeast deacetylasesRpd3 (class I), Hda1 (class II), or NAD-dependent Sir2 (classIII). Enzymes in these classes can differ in localization andtissue-specific expression (11). Generally, class I HDACs re-side in the nucleus, while many class II enzymes shuttle be-tween the nucleus and cytoplasm in response to cellular sig-nals. Some class I yeast HDACs that regulate transcriptiondeacetylate Lys5/Lys12, in addition to other residues on H4,and thus may also play a role in chromatin maturation (41).However, their transcription function complicates investigationsinto possible maturation functions, since the two processes occursimultaneously in the same organelle. Most systems share thedifficulty of separating chromatin maturation-related from tran-scription-related deacetylation events. To circumvent this bar-
* Corresponding author. Mailing address: W. M. Keck Science Cen-ter, 925 N. Mills Ave., Claremont, CA 91711. Phone: (909) 607-9698.Fax: (909) 621-8555. E-mail: [email protected].
† Present address: Division of Biological Sciences, University ofCalifornia at San Diego, La Jolla, CA 92093.
‡ Present address: Department of Cell and Developmental Biology,Oregon Health and Science University, Portland, OR 97219.
§ Present address: Fred Hutchinson Cancer Research Center, Seat-tle, WA 98109.
rier to identifying a maturation-related HDAC, we utilized theciliated protozoan T. thermophila for its two distinct nuclei,called the macronucleus and the micronucleus. Global chro-matin in the macronucleus is highly acetylated, while that inthe micronucleus is entirely unacetylated. Both nuclei divideand assemble new chromatin, but transcription-related acety-lation occurs only in the macronucleus during vegetativegrowth; the micronucleus is transcriptionally silent throughoutthe cell cycle (2, 51). The only acetylation observed in micro-nuclei is highly transient and occurs in deposition-related pat-terns (8). Thus, HDACs that act on micronuclear chromatinduring vegetative growth must function in the removal of dep-osition acetylation. Compelling evidence for such a micro-nuclear HDAC activity was observed during a period in thesexual-conjugation pathway when the macronucleus remains ina nonreplicative state (G1) while the micronucleus undergoessuccessive rounds of mitosis and DNA replication (12).Throughout this period, newly synthesized histones weredeacetylated by an unknown micronuclear HDAC (2, 8). Fol-lowing deacetylation, micronuclear chromatin is thought tomature through evolutionarily conserved pathways, but oneunique feature is the proteolytic processing of a fraction of newhistone H3 molecules into a truncated form (3, 5, 6). Produc-tion of this form is related to mitotic chromosome condensa-tion and segregation in the micronucleus (54).
Here, we report the identification of a class II HDAC calledThd2 (Tetrahymena histone deacetylase 2) that localizes to themicronucleus, where it removes deposition-related acetylationfrom histones H3 and H4. Cells lacking Thd2 display micro-nuclear chromatin phenotypes, including reduced H3 proteol-ysis. THD2 is an alternatively spliced gene, the first reportedfor a coding sequence in Tetrahymena. Of the two resultingsplice variants, the predominant form encodes an HDAC en-zyme containing a putative inositol polyphosphate kinase(IPK) domain that in other systems has been implicated inthe regulation of chromatin remodeling. From this study, pos-sible roles for HDACs in the maturation of new chromatin arerevealed.
MATERIALS AND METHODS
Bioinformatics. Eighteen HDACs (Thds) were identified as containing puta-tive HDAC domains from The Institute for Genomic Research-annotated genesin the Tetrahymena macronuclear genome database (http://www.ciliate.org).Their respective gene sequence identification numbers are listed in Table 1.Since experimental confirmation of the coding sequences was lacking, only theHDAC domains were used for phylogenetic analysis. The sequences of the 18Tetrahymena HDAC domains were aligned with the S. cerevisiae Rpd3, Hda1,Sir2, Hst2, and Hst4 HDAC domains using Multiple Sequence Alignment-CLUSTALW (Kyoto University Bioinformatics Center [http://align.genome.jp/]). The CLUSTAL protein alignment was performed using a gap open penaltyof 10, a gap extension penalty of 0.05, a hydrophobic gap, no weight transition,and a BLOSUM weight matrix. Distances were computed using the PoissonCorrection Distance method in Molecular Evolutionary Genetic Analysis(MEGA) software version 4.0 (MEGA4) (49). The unweighted-pair groupmethod using average linkages tree was constructed from the matrix of distancesaccording to the model using MEGA4, and the robustness of the tree topologywas tested with 1,000 bootstrap replicates.
Strains and cell culture conditions. T. thermophila strains B2086 (II), CU428(chx1/chx1 [cy-s] VII), and SB1969 (mpr1/mpr1 [6-mp-s] II) were used as thewild-type strains (Tetrahymena Stock Center, Cornell University). Unless oth-erwise indicated, cells were grown at 30°C with shaking in either 1% (wt/vol)enriched proteose peptone (17) or 2% proteose peptone, 0.2% yeast extract, and0.1% sequestrene (61) liquid medium with 1� penicillin, streptomycin, andfungizone (Gibco-BRL) to a density of 1 � 105 to 3 � 105 cells/ml for all
experiments. The cells were starved in 10 mM Tris-HCl (pH 7.4) for 14 to 20 hat a density of 3 � 105 cells/ml at 30°C with no shaking.
RT-PCR. Genomic DNA was isolated as described previously (61). Total RNAwas isolated from vegetatively dividing, starved, and conjugating cells using theRNeasy Total RNA kit (Qiagen, Valencia, CA). The cDNA was made as pre-viously described (31) using 2 �g of total RNA for each reaction. PCR wasperformed on dilutions of the cDNA to determine the linear range of conditionsfor the sample set (data not shown). Experiments were performed using condi-tions that were within the linear amplification range to obtain semiquantitativeresults. PCR and reverse transcriptase (RT)-PCR were performed using 2�GoTaq master mixture (Promega, Madison, WI) following the manufacturer’sdirections and using the following primer sets: THD2a (�) (5�-GTTTATTTTGATATCTGCTG-3�) and THD2b (�) (5�-TTAGACTTCAAATGAATTTAC-3�); THD2 (�) (5�-CTAAATGCGATCCTTTAATTC-3�) and HHP1 (�) (5�-TTAGCAATGATAAACCTTAGAC-3�); HHP1 (�) (5�-TGTGTAAAGAGATTTTCCATC-3�) and ACT1 (�) (5�-GAACAGAGAAAAGATGACCAAG-3�);ACT1 (�) (5�-GGTAAGTTCGTGGATACCAGG-3�) and CYP1 (�) (5�-AGAGTAACCCTAATAACACC-3�); and CYP1 (�) (5�-CCGTTGAAAATTCCAGACG-3�).
THD2 gene and splice variant analysis. The 5� terminus of the THD2 mRNAwas deduced by RT-PCR with the THD2 (�) primer listed above and thefollowing primers upstream of the predicted translation start site: THD2-4 (�)(5�-CCAAAACAGACAAACTATGCAAC-3�), THD2-132 (�) (5�-ATTTCATATGATAAGATTGAATTCATTTAC-3�), THD2-219 (�) (5�-CATTTTTTCTATCTATTATGAGTCATAATC-3�), THD2-313 (�) (5�-ATGTTAAATTTTTACGTAATTTTAAATTGCG-3�), and THD2-360 (�) (5�-ATTAACTAATGCTTACATGATATTATTC-3�).
The 3� poly(A) addition site for the THD2a and THD2b mRNA sequence wasdetermined by RT-PCR using THD2E2 (5�-CATGACGATGGATCATTCTATCC-3�), a plus-sense primer in exon 2 (E2), and a poly(dT)20 primer as thereverse primer. The PCR product was subcloned using the Invitrogen TA cloningkit (Invitrogen, California) and sequenced on an Applied Biosystems 3130xlGenetic Analyzer (Rancho Santa Ana Botanic Garden Molecular Laboratory,Claremont, CA) to determine the correct intron junctions and the poly(A)addition sites for the splice variants.
Cell elutriation and conjugation. Logarithmically growing cells (1 � 105 to3 � 105 cells/ml) were synchronized using a centrifugal elutriator rotor as pre-viously described (31), and RNA was collected every 20 min after the elutriatedcells were resuspended in fresh 2% proteose peptone, 0.2% yeast extract, and0.1% sequestrene. For conjugation experiments, starved CU428 (VII) andSB1969 (II) were mixed in equal concentrations (3 � 105 cells/ml), and 10-mlaliquots were placed in 100-mm by 15-mm prewarmed petri dishes and incubatedin a humidity chamber at 30°C (no shaking). RNA was collected from 10 ml ofconjugating cells every hour initially after the mixing (0 h) through 14 h and againat 24 h after the mixing.
TABLE 1. Putative Tetrahymena HDACs with similarity to humanand S. cerevisiae proteins
Plasmid construction. The NEO construct used to replace THD2 was madefollowing a previously described method (56), except that the THD2 3� and 5�flanking regions were amplified by PCR using the following primers: 5�FlankBamHI(�) (5�-GAGGGATCCAAAGTGTGATGTGTTTGAAG-3�), 5�FlankXmaI (�)(5�-GTGCCCGGGCTAAAAAGAAATCAAGAAAG-3�), 3�FlankClaI (�) (5�-CGCATCGATAGAAACAACTCCTGTCTGTC-3�), and 3�FlankXhoI (�) (5�-AGACTCGAGTCAAATACCTAGCCCTCTCC-3�).
The PCR products were digested with appropriate restriction enzymes andligated into p4T2-1 containing the Neor drug resistance cassette to makepTHD2-NEO (56). For transformation, pTHD2-NEO was digested to produce alinear fragment containing the 5� and 3� THD2 flanking sequence interrupted bythe neomycin drug resistance cassette. Gold beads were coated with this, aspreviously described (31).
Construction of a strain expressing green fluorescent protein (GFP) fused tothe THD2 genomic sequence was done as previously described (59), using prim-ers THD2GFP (�) (5�-CAACAGGATAATATAGAAGGTATAG-3�) andTHD2GFP (�) (5�-TCATTTTGCCTTTTTGTTAAGAG-3�), and transformedvia electroporation as previously described (15).
Macronuclear THD2 replacement. Tetrahymena cells were transformedthrough biolistic bombardment to delete THD2 as previously described (30).Transformants were selected by growth in the presence of 100 �g/ml paromo-mycin (Sigma Chemicals) after 5 h of incubation at 30°C without paromomycin(no shaking). The thd2� allele was assorted to 100% by growing the transfor-mants in media with increasing concentrations of paromomycin to a final con-centration of 10 mg/ml. Complete macronuclear replacement of THD2 wasconfirmed by PCR as previously described (56) using the following primers:THD2Flank (�) (5�-TGGAAGTCAAGCATTTCTTCC-3�), THD2-3 (�) (5�-GTCATCATTCAGGTGAATC-3�), NEOF2 (�) (5�-CTGGGCACAACAFACAATCG-3�), NEOR2 (�) (5�-GTTTCGCTTGGTGGTCGAAT-3�), and NEOS(�) (5�-CCTTAAATTAAAAATTCAATATATATTTTAC-3�).
The resulting strain lacking macronuclear copies of THD2 was named JS01.
Lack of THD2 expression in strain JS01 was confirmed by RT-PCR using theTHD2a (�) and THD2 (�) primers listed above.
Immunoblot analysis. Nuclei from B2086 (wild type) and JS01 (thd2�) wereisolated as previously described (17). The nuclei (1 � 106) were lysed by incu-bation in 30 �l of sodium dodecyl sulfate (SDS) gel loading buffer (50 mMTris-HCl [pH 6.8], 100 mM dithiothreitol, 2% [wt/vol] SDS, 0.1% bromophenolblue, 10% glycerol) and boiled for 5 min. Micronuclei (5 � 105) and macronuclei(1.6 � 105) were resolved by SDS-polyacrylamide gel electrophoresis (PAGE) ona 15% (for histone H4) or 22% (for histone H3) polyacrylamide gel, transferredto a nitrocellulose membrane, and probed with the following antibodies: �-his-tone H4 (1:5,000; a gift from C. D. Allis), �-histone H3 (1:5,000; a gift from C. D.Allis), �-histone H4K16ac (1:2,000; Upstate Biotechnology catalog no. 06-762),and �-histone H3S10ph (1:5,000; a gift from C. D. Allis).
Indirect immunofluorescence and DAPI staining of cells. B2086 (wild-type)and JS01 (thd2�) cells in logarithmic growth (1 � 105 to 3 � 105 cells/ml) werefixed in paraformaldehyde and processed for immunofluorescence as previouslydescribed (9). For detection of acetylated histones, cells were incubated with�-acetylated histone H4 antiserum (1:200; Upstate Biotechnology catalog no.06-598), �-acetylated histone H3 antiserum (1:200; a gift from C. D. Allis), or�-histone H3K9ac antiserum (1:200; Upstate Biotechnology catalog no. 06-942),which also detects H3K14ac in Tetrahymena. For detection of only micronuclei,cells were incubated with anti-micronuclear linker histone 1 (�-Mlh1) (micro-nuclear-specific linker histone) antiserum (1:200; a gift from C. D. Allis). Primaryantibodies were detected with rhodamine-conjugated goat anti-rabbit immuno-globulin G (1:100; Jackson ImmunoResearch catalog no. 111-025-003). The cellswere counterstained with 0.1 �g/ml 4�,6 diamino-2-phenylindole dihydrochloride(DAPI; Sigma Chemicals) in 0.1% bovine serum albumin–phosphate-bufferedsaline for 10 min according to the common protocol (37).
Nucleotide sequence accession numbers. The cDNA sequences for the splicevariants were submitted to GenBank under accession numbers EU254713(Thd2a) and EU254714 (Thd2b).
RESULTS
Thd2 localizes to the micronucleus. Thds were first identi-fied by searching the Tetrahymena Genome Database to findpredicted open reading frames that contained putative HDACdomains. Eighteen putative HDAC proteins were identified(Table 1) and classified by their similarity to the yeast HDACsRpd3 (class I), Hda1 (class II), and Sir2 (class III) (Fig. 1). TheHDAC domains fell into all three classes—3 in class I (Thd1,Thd3, and Thd6), 2 in class II (Thd2 and Thd4), and 11 in classIII (Thd8 through Thd18)—and 2 were HDAC-like (Thd5 andThd7). Most of the putative Tetrahymena class III HDACs
FIG. 1. Phylogenetic tree of the putative Thds (unweighted-pairgroup method using average linkages). Shown is an alignment of theputative HDAC domains of the 18 Thd protein sequences. S. cerevisiaeRpd3 (class I), Hda1 (class II), Sir2, Hst2, and Hst4 (class III) wereused as references to sort the putative Thd proteins into their respec-tive classes (boldface). The positions of human homologs are shown inlight-gray capital letters. (Note: HDAC6 and SIRT5 appear twice dueto construction of the tree with yeast HDAC domains that are lesssimilar to Thd proteins than those of humans.)
FIG. 2. GFP-Thd2 is localized to the micronucleus. Live-cell im-ages of Tetrahymena transformed with a GFP-Thd2 fusion constructand viewed using the fluorescein isothiocyanate channel to detect GFPsignal. The positions of nuclei in each cell were visualized by stainingthem with DAPI. GFP expressed alone, without fusion to anotherprotein, remained in the cytoplasm; the regions devoid of fluorescencecorrespond to the macronucleus (M) and the micronucleus (m). Cellsexpressing GFP-Thd2 showed fluorescence in both the macronucleusand micronucleus, which persisted throughout the cell cycle (see themitotically dividing cell in the right panel).
(Thd8 through Thd18) were more closely related to humansirtuin homologs than to the yeast sirtuins Sir2, Hst2, and Hst4(Fig. 1).
An HDAC that resides in the micronucleus would likelyremove deposition-related acetyl marks from newly synthe-sized histones, since only deposition acetylation occurs on his-tones in this nucleus. To identify a micronuclear HDAC, sev-eral of the putative HDAC genes (Fig. 1) were cloned in framewith GFP coding sequence at their 5� ends and expressed fromthe metallothionein promoter. The resulting amino-terminalGFP fusion proteins were induced, and their localization in livecells was assessed by fluorescence microscopy. Of four tested,only GFP-Thd2 localized to the micronucleus in addition tothe macronucleus (Fig. 2); the other putative HDACs tested
localized exclusively to the macronucleus or to the cytoplasm(data not shown) (57).
THD2 is alternatively spliced. The THD2 coding sequencewas determined by cloning and sequencing its cDNA (as de-scribed in Materials and Methods). Amplifying the cDNA withgene-specific primers revealed two cDNAs of different lengths,indicating that the THD2 gene was alternatively spliced. Themost abundant splice variant, called THD2a, contained onlyE1, E2, and E4), while the minor form, called THD2b, con-tained E1 through E4 (Fig. 3A). Sequence analysis showed thatthe inclusion of E3 in THD2b caused a frameshift that pro-duced a premature stop codon within E4 and loss of a putativeIPK domain (Fig. 3B). The relative transcription level ofTHD2a was 15- to 20-fold higher than that of THD2b during
FIG. 3. Thd2a is a splice variant of THD2 that contains both an HDAC and an IPK domain. (A) Diagram of the THD2 locus with the flankingsequence (thin black), coding sequence (thick gray) with exons labeled below (E1 to E4), and introns (white) labeled below (i1 to i3). The arrowsrepresent primers used to amplify splice variants of THD2. Displayed below the THD2 sequence is a schematic of the mRNA splice variants forThd2a (only exons E1, E2, and E4) and Thd2b. The locations of putative HDAC (light gray) and putative IPK (dark gray) domains are indicated.(B) Protein sequence alignment of Thd2a and Thd2b. HDAC domain, light gray; IPK domain, dark gray; change in coding sequence due to splicingframeshift, #; and stop codon, *. (C) RT-PCR of wild-type vegetatively growing (V) and starved (S) cells. PCR on a cDNA template using primersTa(�) and T2(�) was used to detect THD2a transcripts, and primers Tb(�) and T2(�) were used to detect the THD2b variant transcripts. ThecDNA was made with (�) and without (�) RT as a control for genomic DNA contamination. Genomic DNA (G) was amplified as a control.RT-PCR yielded two bands for THD2b; the fastest-migrating band corresponds to THD2b (arrow). (Note: THD2b amplification was detected onlywith an additional seven cycles of PCR amplification over that used for THD2a detection.)
vegetative growth when normalized to the amplification ofgenomic DNA for each PCR (Fig. 3C and data not shown).When cells were starved and no longer progressing through thecell cycle, both THD2a and THD2b transcription levels de-creased two- to threefold, suggesting that both enzyme formsmay have more significant functions during periods of activegrowth and cell division.
The HDAC domain of Thd2 was aligned with the yeastHDACs Hda1 (class II) and Rpd3 (class I) and found to havegreater similarity to Hda1 (37% identical and 56% similaramino acid residues) (Fig. 4A). Thd2 had a high degree ofsimilarity to Hda1 in regions of the HDAC domain that areconserved in most class II enzymes (Fig. 4A) (26, 63). Tofurther classify the putative IPK domain in Thd2, the sequencewas compared to two well-characterized yeast IPKs, Ipk1 andIpk2, the primary enzymes in the pathway for conversion ofinositol triphosphate (IP3) to inositol hexaphosphate (IP6)(Fig. 4B). The putative IPK domain of Thd2a was more closelyrelated to Ipk2 (27% identical and 36% similar amino acidresidues) than to Ipk1 (10% identical and 22% similar aminoacid residues). Notably, the cofactor binding regions and ino-sitol polyphosphate binding region for Ipk2 were similar inThd2a. Thd2a also contains a conserved aspartate residue thatis essential for Ipk2 kinase activity (Fig. 4B) (20).
Expression of THD2 coincides with DNA replication. Tobegin characterizing THD2 function, we first examined its tran-scription over a normal cell cycle during vegetative growth,since cells produced two- to threefold more THD2 mRNAwhen growing than in the noncycling, starved state. In Tetra-hymena cells, the micronucleus progresses through the cellcycle without resting in G1 phase. Following mitotic division, a
cell proceeds directly into S phase and then rests in an ex-tended G2 phase while the macronucleus undergoes amitoticdivision followed by DNA synthesis (22, 58). Logarithmicallygrowing cells were synchronized by isolating new daughter cellsthrough centrifugal elutriation and then allowing them to pro-ceed through the cell cycle. The synchrony of cells in theculture was monitored at frequent intervals throughout severalcell cycles by counting the cells at different stages (Fig. 5A).RT-PCR analysis of THD2 expression at regular intervalsthroughout two cell cycles (from 20 min to 280 min afterelutriation) revealed that THD2 was expressed in a cyclicalpattern (Fig. 5B). Maximum expression of THD2 occurred at80 and 200 min, which were the times when the greatest num-bers of cells were in micronuclear mitosis. The expression ofACT1, which does not change throughout the cell cycle, wasmonitored as a control (34, 62). Since micronuclear DNAsynthesis occurs immediately following anaphase, the observedpeak of THD2 expression coincident with anaphase was con-sistent with the possibility that Thd2a might deacetylate newlydeposited histones.
To further examine whether THD2a expression coincideswith DNA replication and division, transcript levels were mon-itored throughout the sexual-conjugation pathway. Early inthis process, the transcriptionally silent micronucleus under-goes three successive rounds of DNA replication and two mi-toses while the transcriptionally active macronucleus remainsin G1 (Fig. 6A). Exploiting this feature, we tested whetherTHD2a expression coincided with DNA synthesis and histonedeposition in the micronucleus at the time (5 to 7 h) when amicronuclear HDAC activity that removes deposition acetyla-tion was previously detected (2). Cells of two different mating
FIG. 4. Thd2 is a class II HDAC with a putative IPK domain similar to Ipk2. (A) Alignment of the Thd2a HDAC domain with the HDACdomains of S. cerevisiae Hda1 (class II) and Rpd3 (class I). Residues conserved between Thd2 and either Hda1, Rpd3, or both are shaded in gray.Regions that are highly conserved between most class II HDACs are marked with a black line below the sequence. A conserved histidine residuecritical for HDAC activity is marked by an asterisk. (B) Alignment of the Thd2a putative IPK domain with S. cerevisiae Ipk1 and Ipk2. Residuesconserved between Thd2 and either Ipk2, Ipk1, or both are shaded in gray. The inositol polyphosphate binding domain is marked with a gray linebelow the sequence, and regions involved in cofactor binding are marked by a black line below the sequence. A conserved aspartate residue criticalfor Ipk2 activity in yeast is designated by an asterisk.
types were starved, mixed together, and allowed to conjugateover a 24-hour period. To monitor population synchrony, thepercentage of cells in each stage was determined at regularintervals throughout the conjugation time course. By 2 hoursafter they were mixed, 66% of the cells were paired (Fig. 6A).By 3 hours after they were mixed, 66% were in meioticprophase and 85% were paired. Throughout the entire conju-gation time course, 60 to 70% of the cells were tightly synchro-nized and an additional 20 to 25% of the culture was within 60min of this primary synchronized population. RT-PCR oncDNA made from conjugating cells harvested at 1-hour inter-vals revealed that THD2a was most highly expressed between5 and 7 h after the mixing (Fig. 6B). At the peak of expression(6 h postmixing), 70% of the cells were in one of the mitoticdivisions (30% in prezygotic mitosis and 40% in postzygoticmitosis) (Fig. 6A). This result demonstrated a strong correla-tion between THD2a expression and periods of DNA replica-tion, evidence that THD2a encodes an HDAC that acts duringthis time.
Similar analysis revealed that Thd2b expression remainedlow during conjugation compared to that of Thd2a. This minorvariant also appeared to be more ubiquitously expressedthroughout conjugation, with only a slight peak in expression at6 h (Fig. 6B). The band corresponding to genomic DNA con-tamination was amplified to a greater extent at some conjuga-
tion points when THD2a levels were relatively low. Curiously,when reactions lacking RT were performed on these mRNAsamples and they were used as templates for PCR, no ampli-fication of Thd2 cDNA was observed (Fig. 3C and data notshown). This result raises the possibility that there may be yetanother variant, one that retains the intron between E3 andE4. Such a variant would produce an HDAC similar to Thd2bthat lacks the putative IPK domain but that has an earlier stopcodon than Thd2b.
Thd2 deacetylates micronuclear histones. To test whetherThd2 deacetylates micronuclear chromatin, a mutant cell linelacking THD2 was first engineered. A genetic construct for thesomatic replacement of THD2 with a paromomycin resistancegene (NEO) was transformed into wild-type cells (Fig. 7A).Complete replacement of the THD2 allele with the NEO con-struct was confirmed by PCR (Fig. 7B, THD2 and NEO), aswas the correct integration of the construct into the THD2locus (Fig. 7B, THD2-NEO). The resulting cell line lacking allmacronuclear copies of THD2 was called thd2�. RT-PCR wasconducted on the thd2� cells to confirm the absence of THD2expression (Fig. 7C). Obtaining a complete knockout strainindicated that THD2 was a nonessential gene in vegetativelygrowing cells.
The micronucleus is transcriptionally inactive and lacksacetylated chromatin in growing cells. Acetylated histone H4 in
FIG. 5. Thd2 is expressed during DNA replication and cell division in growing cells. (A) Graph of the cell cycle stages represented in samplesof a culture synchronized by centrifugal elutriation. Samples were taken every 20 min, and the percentages of cells in each stage of cell divisionwere scored. The concentration of the culture is provided above the graph and a key describing the four different stages of division scored is tothe right of the graph. (B) Samples from the synchronized culture were subjected to RT-PCR to detect THD2 transcripts at regular intervalsthroughout the cell cycle. Primers Ta(�) and T2(�) were used in RT-PCR to detect the predominant form, THD2a. ACT1 cDNA, which remainsconstant throughout the cell cycle, was used as a control. Relative levels of THD2 (Rel. THD2) were determined by quantifying band intensitiesof THD2 and ACT1 at each time point, dividing the values for THD2 by the values for ACT1 and then normalizing the resulting value to that atthe 20-min time point.
micronuclei was previously detected only when cells weretreated with general HDAC inhibitors, such as sodium bu-tyrate and trichostatin A (2, 8, 13). To test the acetylation stateof micronuclear chromatin in thd2� cells, immunofluorescencewas performed using antiserum specific for only acetylatedspecies of histones H3 and H4 (�-H3ac and �-H4ac). Consis-tent with previous studies of wild-type cells (51), acetylatedhistones H3 and H4 were detected only within the transcrip-tionally active macronucleus and not in the micronucleus (Fig.8, WT). In thd2� cells, however, acetylated histones H3 and H4were detected in the micronucleus, as well (Fig. 8, thd2�).Micronuclear histones were acetylated in every cell and in eachstage of the cell cycle observed, in contrast to wild-type cells, inwhich micronuclear acetylation was never detected (Fig. 8).This analysis demonstrated that Thd2 is an HDAC that nor-mally deacetylates histones in the micronucleus. Whether italso acts on macronuclear histones was not determined.
It was deduced that the observed micronuclear acetylation inthd2� cells resulted from the retention of deposition-relatedacetyl moieties. To help confirm this, we performed an immu-nofluorescence assay using an antiserum against acetylatedlysine 9 and/or 14 on histone H3 (�-H3K9/14ac), the acetylmarks found on newly synthesized Tetrahymena histones (2,25). Whereas this antiserum normally does not hybridize withmicronuclear chromatin in wild-type cells, it did hybridize withchromatin in thd2� micronuclei, indicating that H3Lys9 and/or-14 was acetylated in the mutant cells. Other deposition site-specific acetylation antisera (anti-acetylated Lys5 and anti-acetylated Lys12) were not included in this analysis due to theirlack of specificity; Tetrahymena histone H4 is shorter by 1
FIG. 7. THD2 is a nonessential gene. (A) Diagram of the THD2 deletionconstruct used to replace THD2 with NEO in the somatic macronucleus.Depicted in the diagram are the flanking regions (thin black lines), the codingsequence (thick dark gray lines), introns (white boxes), the histone H4 pro-moter (thick black lines), the neomycin resistance gene (NEO), and the BTU2polyadenylation region (light gray line). The arrows represent the primersused to confirm correct integration of the replacement allele. (B) PCR am-plification of genomic DNA from wild-type (WT) and thd2� (�) cells con-firmed that all THD2 alleles were replaced with the NEO cassette. THD2PCR was performed using Ta(�) and T2-3 primers (WT allele), NEO PCRwith NF and NR primers (NEO cassette), and THD2-NEO PCR with F1 andNS primers (incorporation of the NEO cassette in the THD2 locus). HHP1PCR was performed as a positive control for the genomic DNA. (C) TotalcDNA derived from WT and thd2� cells was used in PCRs to test for thepresence of THD2 mRNA in these cells. HHP1 was used as a control forcDNA synthesis and PCR amplification. Genomic DNA (G) was used as atemplate to control for genomic-DNA contamination in cDNA.
FIG. 6. Thd2 is expressed during conjugation coincident with micronuclear DNA synthesis and mitosis. (A) Diagram of Tetrahymena conju-gation stages. Two different mating types were mixed to initiate conjugation. Samples were taken every hour and stained with DAPI, and thepercentage of cells in each stage of conjugation was determined by fluorescence microscopy, as indicated above the diagram. The black barsindicate periods of DNA synthesis and mitosis (the short bar is the prezygotic mitosis just prior to the pronuclear exchange; the long bar is thepostzygotic mitoses I and II following zygotic fusion). (B) Total RNA was harvested from vegetatively growing (V) cells, starved (S) cells, and cellsduring conjugation (0 through 14 and 24 h after mixing) and used as a template in RT-PCRs with primers specific for THD2a or THD2b variants.Primers for CYP1 and HHP1, two genes showing consistent expression throughout conjugation, were used as controls in this analysis. GenomicDNA (G) was used to control for genomic-DNA contamination in the RNA samples. (Note: THD2b RT-PCR contained two bands; thefaster-migrating band corresponds to the spliced form of Thd2b.)
amino acid (it is missing Arg3) at the N terminus. To help ruleout possible aberrant transcription-related acetylation in themicronuclei of thd2� cells, an immunoblot analysis was per-formed on proteins from purified micronuclei or macronucleiwith antiserum detecting acetylated Lys16 (�-H4K16ac), aknown transcription-specific mark found exclusively in the ma-cronuclei of wild-type cells (8, 25). This antiserum was shownto have correct specificity for a Tetrahymena H4 peptide acety-lated at this position (E. Wiley, unpublished data). As shown inFig. 8D, neither wild-type nor mutant (thd2�) micronuclear hi-stone H4 molecules were acetylated on Lys16. Consistent withprevious studies, Lys16 acetylation was detected only in macro-nuclei. Due to high background, results from immunofluores-cence experiments with this antiserum are not shown. Combined,these results suggest that Thd2 is normally required for mainte-nance of the deacetylated state of micronuclear histones H3 andH4 through removal of deposition-related acetyl modifications.
Chromatin defects in thd2� cells. Immunofluorescencestudies revealed a number of elongated micronuclei in thd2�
cells. Although this is the normal appearance of micronuclei inanaphase, many of these elongated micronuclei in mutant cellswere closely associated with the macronucleus, an aberrantposition for an anaphase nucleus (Fig. 8C, bottom right). Insome cases it was difficult to define the micronuclei thatstretched across portions of the macronucleus by DAPI stain-ing. To improve the visualization of micronuclei, immunoflu-orescence was performed with antiserum specific for Mlh1,which is present only in the micronucleus (Fig. 9A). As ex-pected, Mlh1 localized specifically to the micronucleus for bothwild-type and thd2� cells. Micronuclei were scored for elon-gated shape when in close proximity to the macronucleus (inthe “pocket”) for both wild-type and thd2� cells (Fig. 9A). Thepercentage of cells with abnormal micronuclei in the pocketwas approximately 10-fold higher in thd2� cells than that inwild-type cells (50.5% � 3.3% and 4.7% � 3.2%, respectively).Micronuclei were more variable in size, as well. Although celldivision appeared to progress normally, the doubling time forthd2� cells was three times that of wild-type cells (9 h and 3 h,
FIG. 8. Thd2 removes deposition-related acetylation from micronuclear histones. (A) Immunofluorescence using antiserum against acetylatedhistone H3 (�-H3ac). Cells were counterstained with DAPI to visualize both the macronucleus (M) and the micronucleus (m). Acetylated histoneH3 was detected exclusively in the macronuclei of wild-type (WT) cells and additionally in the micronuclei of thd2� cells. (B) Immunofluorescenceusing antiserum against acetylated histone H4 (�-H4ac). DAPI stain was used to detect both the macronucleus and the micronucleus. Acetylatedhistone H4 was detected exclusively in the macronucleus of WT cells and additionally in the micronuclei of thd2� cells throughout every stage ofthe cell cycle. A high proportion of cells contained elongated micronuclei in close proximity to the macronucleus (the phenotype is depicted in thelast panel). (C) Immunofluorescence using antiserum against acetylated Lys9 on histone H3 that additionally detects acetylated Lys14 inTetrahymena (�-H3K9/14ac). Cells were counterstained with DAPI to visualize both the macronucleus and the micronucleus. Histone H3Lys9/Lys14 acetylation was detected only in the macronuclei of wild-type cells, but also in the micronuclei of thd2� cells. (D) Total nuclear proteinsfrom purified macronuclei and micronuclei were resolved by SDS-PAGE, transferred to a nitrocellulose membrane, and subjected to immunoblotanalysis using antiserum against acetylated Lys16 on histone H4 (�-H4K16ac) or against general histone H4 (�-H4). In both wild-type and thd2�cells, H4 Lys16 was acetylated only in macronuclei.
respectively) (data not shown). Together, these aberrant mor-phologies and weak DAPI staining are consistent with chro-matin condensation abnormalities in thd2� micronuclei.
In micronuclei, histone H3 (called H3s) is proteolyticallycleaved to form a faster-migrating form, called H3f, which ismissing 6 amino acids from the extreme amino terminus. H3f isthe only form of histone H3 in Tetrahymena that is phosphor-ylated on Ser10, a mitotic modification necessary for chromo-some condensation and segregation (4, 6, 54). In thd2� cells,the elongated micronuclear phenotype, reminiscent of eitherdecondensed chromatin or mitotic micronuclei in anaphase,prompted us to examine H3 processing and related phosphor-ylation on Ser10. Immunoblot analysis revealed that while theabundance of micronuclear H3s was similar to that in wild-typecells, cleavage of histone H3s to H3f was greatly reduced inthd2� cells (Fig. 9B). As expected, the reduced amount of H3f
correlated with reduced Ser10 phosphorylation, likely the re-sult of the failed H3 processing in these cells (Fig. 9B). Theseresults suggest that Thd2 acts upstream of the proteolyticcleavage and subsequent phosphorylation of Ser10 on his-tone H3.
DISCUSSION
HDACs in Tetrahymena. Completion of the Tetrahymena ge-nome-sequencing project facilitated the identification and clas-sification of putative HDACs according to the three classical
yeast deacetylases, Rpd3, Hda1, and Sir2 (11). All three classesof HDACs (classes I, II, and III, respectively) are representedin Tetrahymena (Fig. 1). Interestingly, their deacetylase do-mains are more similar to those in human enzymes than tothose in budding yeast. Tetrahymena Thd1, Thd3, and Thd6enzymes are class I HDACs due to their similarity to yeastRpd3, and their HDAC domains are most similar to humanHDAC1. Thd2 and Thd4 are class II homologs with similarityto yeast Hda1 but that have deacetylase domains most similarto human HDAC6. There are two HDAC-like proteins thatdid not group well with any of the yeast HDACs. One of these,Thd5, is highly similar to HDAC11, a class IV HDAC thoughtto be a hybrid of class I and II enzymes present in primates,rodents, Drosophila, and plants (16, 18). Tetrahymena also con-tains 11 sirtuins (class III HDACs), more than have beenreported for any other organism to date. The putative Tetra-hymena sirtuins are similar to yeast Sir2 but are more closelyrelated to protein sequences for the human SIRT2 (Thd16),SIRT3 (Thd13, Thd14, and Thd15), SIRT5 (Thd10, Thd17,and Thd18), and SIRT6 (Thd8, Thd9, Thd11, and Thd12)proteins. Tetrahymena will likely prove a useful model systemfor further studies addressing distinct roles for differentHDACs, especially the sirtuins, within a single cell.
Identification of an HDAC that deacetylates new histones.The nuclear dimorphism of Tetrahymena was exploited to iden-tify an HDAC that removes deposition-related acetylation onnewly synthesized histones. Previous work demonstrated thatthe transcriptionally active macronucleus contains highlyacetylated histones H2A, H2B, H3, and H4 (51). In contrast,the micronucleus is transcriptionally inactive in growing cells,and little to no acetylation of the core histones is observed (2).The only histone acetylation ever detected in the micronucleiof actively dividing cells was related to histone deposition, andthese acetyl moieties were rapidly removed by a micronuclearHDAC activity (2, 40). Surprisingly, the micronucleus wasshown to contain HAT activity. However, this activity is dis-tinct from the transcription-related HAT activities of the mac-ronucleus (40). Whereas macronuclear HATs produce tran-scription-related acetylation patterns (Lys8 and Lys16 on H4[25, 36]), micronuclear and cytoplasmic HAT activities (40)produce only deposition-related patterns (Lys4/Lys11 on H4[8, 27, 46], analogous to Lys5/Lys12 in other organisms). Thus,an HDAC(s) in the micronucleus must maintain the unacety-lated state of micronuclear chromatin by removing deposition-related acetylation, the only acetylation observed in micronu-clei. Previously, GFP tagging was used to show that Thd1, aclass I Tetrahymena HDAC, localized exclusively to the macro-nucleus and not to the micronucleus (57). The same strategywas used in the present study to identify Thd2 as an enzyme(class II) that localized to the micronucleus, in addition to themacronucleus (Fig. 2). As yet, there is no evidence that Thd2,like class II enzymes in other systems, shuttles between thenucleus and cytoplasm.
Our results indicate that in the micronucleus, Thd2 deacety-lates histones H3 and H4. Immunofluorescence experimentson deletion mutants of THD2 (thd2�) revealed the presence ofacetylated micronuclear histones H3 and H4 at different stagesthroughout the cell cycle, suggesting that deposition acetyla-tion was retained in the mutant cells (Fig. 8A and B). Anti-serum that detected deposition-related acetylation on histone
FIG. 9. Cells lacking Thd2 exhibit chromatin phenotypes. (A) Im-munofluorescence using antiserum against micronuclear linker histoneH1 (�-Mlh1) was performed on wild-type (WT) and thd2� cells. DAPIstaining was used to visualize both the macronucleus (M) and themicronucleus (m). A higher incidence of elongated micronuclei inclose association with macronuclei was observed in the mutant cells.(B) Total proteins from purified macronuclei and micronuclei wereresolved by SDS-PAGE, transferred to a nitrocellulose membrane, andhybridized with antiserum against general histone H3 (�-H3) or withantiserum against phosphorylated serine 10 on histone H3 (�-H3S10ph). The full-length form of histone H3 (H3s) was detected inboth macronuclei and micronuclei of all cells, but only wild-type mi-cronuclei contained the faster-migrating proteolytically cleaved form(H3f). Likewise, phosphorylation of Ser10, which is specific for H3f,occurred only in wild-type micronuclei. (Note: the blot was first hy-bridized with �-H3S10ph and then stripped and hybridized with �-H3.)
H3 (Lys9 and Lys14 in Tetrahymena) showed that acetylationof these residues was retained in thd2� micronuclear chroma-tin (Fig. 8C). Although it is unlikely, these experiments did notrule out the possibility that micronuclear acetylation in thd2�cells resulted from aberrant localization of a transcription-related HAT to micronuclei. However, antiserum against thetranscription-related acetyl-Lys16 on H4 (�-H4K16ac) con-firmed that Lys16 remained unacetylated in thd2� micronuclei(Fig. 8D). Combined with the fact that only deposition acety-lation has ever been observed in micronuclei, these resultswere taken as strong evidence that micronuclear Thd2 deacety-lates newly deposited histones. Further support was providedby the finding that THD2a is maximally transcribed duringperiods of DNA replication (late anaphase) (Fig. 5). The clear-est example of this was observed early in the sexual-conjuga-tion cycle (hours 5 to 7), when micronuclei undergo tworounds of DNA replication and mitotic division in rapid suc-cession while macronuclei remain in G1. Over this period,there was a pronounced peak of THD2 transcription (Fig. 6B),the same window in which rapid deacetylation of newly syn-thesized histones occurs (2).
Histone deacetylation and chromatin maturation. Histonedeacetylation is an important event in chromatin maturation(7). When chromatin is replicated and assembled in vivo in thepresence of sodium butyrate to inhibit deacetylation, it retainsthe DNase I sensitivity typical of immature chromatin, a fea-ture that is rapidly lost when chromatin is replicated undernormal conditions (7, 44). Similarly, chromatin assembled invitro with hyperacetylated histones displays increased sensitiv-ity to DNase I (24). The nuclease sensitivity can be attributedin part to reduced H1-mediated internucleosomal interactionsthat require core histone amino termini (38, 39). Recently, itwas found that deacetylation of Lys91 in the globular domainof histone H4 is important for interaction of H3-H4 tetramerswith H2A-H2B dimers during nucleosome formation. Lys91 isinitially acetylated prior to deposition, and Lys91 deacetylationfacilitates the formation of a salt bridge between Lys91 and aglutamic acid residue on histone H2B (10, 60). Outside ofthese observations, little is known about the role of deacetyla-tion in the chromatin maturation process.
Following nucleosome segregation and de novo formationon replicated DNA strands, some nucleosome remodeling islikely required to position nucleosomes for further chromatinmaturation. Interestingly, the predominant splice variant ofTHD2 (THD2a) contains a putative IPK domain related toIpk2 in yeast. This putative IPK domain is the only one of itstype in the predicted Tetrahymena coding sequences (J. Smith,unpublished data). Recent studies demonstrated a functionallink between the IPK signaling pathway and chromatin remod-eling; Ipk2 was shown to be involved in the recruitment ofchromatin-remodeling complexes for transcription (43, 48). Inyeast, mutations in Ipk2, the enzyme that normally convertsIP3 to IP4 and IP5, prevented specific gene induction due tofailed recruitment of the chromatin-remodeling complexesSWI/SNF and INO80 (48). Thd2 contains most of the resi-dues within the conserved cofactor binding and IP bindingdomains found in polyphosphate kinases (Fig. 4B). In Ipk2,a D131A mutation (underlined) within the IP kinase motif(PXXXDXKXG . . . SSLL) impaired chromatin remodelingand gene transcription in yeast (43, 48). Thd2a contains this
highly conserved region and the critical aspartate residuewithin the IPK motif (Fig. 4B). Other than regulating chroma-tin remodeling, other possible nuclear roles for an Ipk2-likedomain include promoting mRNA export (33, 35) and regu-lating nonhomologous end joining (19). Future studies willaddress these possibilities, especially in relation to the HDACactivity residing on the same enzyme.
Although this is the first report of an IPK motif on anHDAC enzyme, broad gene database searches revealed otherThd2 homologs with putative IPK domains nearly identical insequence and structure across the putative polyphosphatebinding and kinase motifs. Interestingly, they were found onlyin other protozoans (Paramecium tetraurelia, Plasmodium fal-ciparum, Plasmodium yoelii, Theileria annulata, and Theileriaparva) (data not shown). In each case, like Thd2, the putativeIPK domain was at the C terminus of the HDAC enzyme. It ispossible that in other organisms, instead of residing on thesame polypeptide, IPKs exist as separate polypeptides thatcomplex with HDACs to mediate chromatin maturation.
The alternative splicing of THD2 transcripts yields a variantwithout the putative IPK domain, a process that could be usedto regulate the IPK activity on this HDAC. Splice variantTHD2a contains the putative IPK domain, whereas splice vari-ant THD2b does not. Moreover, the two variants are differen-tially expressed throughout sexual conjugation. Whereas theyshare a common peak of transcription at 6 hours coincidentwith micronuclear mitosis, THD2b is expressed to almost thesame degree at later times, as well (Fig. 6). In this study, GFPwas cloned in frame with the THD2 genomic sequence. It is notclear which form, Thd2a or Thd2b (or both), localizes to themacronucleus; future studies will address whether these formsare differentially localized. Our experiments revealed a possi-ble third splice variant that retains intron 3 (Fig. 3C and 6B).This form, like THD2b, would lack the putative IPK domain.We were unable to rule out the possibility that this variant wassimply an unprocessed intermediate of THD2b in which intron3 had not yet been removed (Fig. 3A). Regardless of whetherthere are two or three splice variants of THD2, this is the firstreported example of an alternative spliced coding sequencefrom Tetrahymena.
The cytological phenotypes of thd2� cells were suggestive ofchromatin defects. Although the doubling time of thd2� cellswas 3 times that of wild-type cells (data not shown), nucleardivision and cytokinesis appeared normal. However, 50% ofelongated micronuclei (a shape typical of anaphase) in thd2�cells were in close proximity to or associated with the macro-nucleus, an aberrant position for an anaphase micronucleus.Determination of whether these micronuclei are arrested inanaphase awaits further studies. A prolonged anaphase wouldaccount for the longer population doubling time observed inmutant cells. It is noteworthy that micronuclei also elongatenormally when their chromatin decondenses over a short in-terval early in sexual conjugation. It is thus tempting to spec-ulate that chromatin is less condensed in growing thd2� cells,something that could result from defects in chromatin matu-ration. Although the sizes and shapes of micronuclei in mutantcells were more variable than those in wild-type cells (manywere quite small), cells completely lacking a micronucleus werenot observed, perhaps due to activation of a mitotic checkpointupon loss of micronuclear DNA.
Although a more extensive analysis of cytological pheno-types in relation to chromatin maturation awaits future studies,one chromatin abnormality detected in thd2� cells was defec-tive proteolytic processing of new histone H3 (H3s). It is stillunclear what purpose this processing normally serves; bothforms (H3s and H3f) are present in mononucleosomes (3).Specific to histone H3f is the phosphorylation of Ser10, a mod-ification present only in micronuclei during the early stages ofmitosis (54). Phosphorylation is not required for formation ofthe H3f species, as demonstrated with an unphosphorylated H3mutant (H3S10A), but micronuclei in these cells displayedchromosome segregation defects (55). In contrast, proteolyticprocessing of H3 was defective in thd2� cells; H3f was greatlyreduced (Fig. 9B). This result was confirmed by immunoblot-ting with antiserum against phosphorylated Ser10 on histoneH3 (�-H3S10ph). As expected, phosphorylation was greatlyreduced compared to that in wild-type cells, consistent withthere being less H3f in the thd2� cells (Fig. 9B). These findingssuggest that Thd2 is required for H3 processing and subse-quent phosphorylation. As Ser10 phosphorylation is linked tochromatin condensation and progression of mitosis in manyorganisms, including Tetrahymena, the reduction in phosphor-ylation may account for some of the observed growth andcytological phenotypes observed in the mutant cells. Futurestudies will explore these possible relationships in the contextof roles for HDAC enzymes in chromatin maturation andfunction.
ACKNOWLEDGMENTS
We thank Dan Romero and Eric Cole for their assistance with cellelutriation and Meng-Chao Yao for his assistance with biolistic trans-formation. We are grateful to Doug Chalker for providing the pIGFplasmid and protocols for making the GFP-THD2 construct and to C.David Allis for providing various antisera. We give special acknowl-edgment to Alison Plumley, Paige Chung, Carey Wickham, and Chris-tine Wong for their supporting experimental and intellectual contri-butions to this study.
This work was supported by NSF Career Award no. 0545560 toE.A.W.
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