Genetics, Vol. 175, 993-1010, March 2007, Copyright © 2007
doi:10.1534/genetics.106.065987

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Contribution of Trf4/5 and the Nuclear Exosome to Genome Stability Through Regulation of Histone mRNA Levels in Saccharomyces cerevisiae

Clara C. Reis^*, and Judith L. Campbell^*,1

^* Braun Laboratories, California Institute of Technology, Pasadena, California 91125 and Gulbenkian Ph.D. Program in Biomedicine, Instituto Gulbenkian de Ciência, 2780-156 Oeiras, Portugal

¹ Corresponding author: Braun Laboratories, 147-75, California Institute of Technology, Pasadena, CA 91125.
E-mail: jcampbel{at}caltech.edu

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS LITERATURE CITED

 
Balanced levels of histones are crucial for chromosome stability, and one major component of this control regulates histone mRNA amounts. The Saccharomyces cerevisiae poly(A) polymerases Trf4 and Trf5 are involved in a quality control mechanism that mediates polyadenylation and consequent degradation of various RNA species by the nuclear exosome. None of the known RNA targets, however, explains the fact that trf mutants have specific cell cycle defects consistent with a role in maintaining genome stability. Here, we investigate the role of Trf4/5 in regulation of histone mRNA levels. We show that loss of Trf4 and Trf5, or of Rrp6, a component of the nuclear exosome, results in elevated levels of transcripts encoding DNA replication-dependent histones. Suggesting that increased histone levels account for the phenotypes of trf mutants, we find that TRF4 shows synthetic genetic interactions with genes that negatively regulate histone levels, including RAD53. Moreover, synthetic lethality of trf4 rad53 is rescued by reducing histone levels whereas overproduction of histones is deleterious to trf's and rrp6 mutants. These results identify TRF4, TRF5, and RRP6 as new players in the regulation of histone mRNA levels in yeast. To our knowledge, the histone transcripts are the first mRNAs that are upregulated in Trf mutants.

Despite being initially identified biochemically as DNA polymerases (WANG et al. 2000), it is widely accepted today that TRF4 and TRF5 encode for nuclear poly(A) polymerases in budding yeast, as predicted from their primary sequence (ARAVIND and KOONIN 1999; SAITOH et al. 2002; HARACSKA et al. 2005). In fact, TRF4 has also been designated PAP2 [poly(A) polymerase 2]. The structure and biochemical functions of Trf4, and of Trf5, which is 58% identical to Trf4, are conserved throughout evolution, as orthologs in Schizosaccharomyces pombe (READ et al. 2002; SAITOH et al. 2002; WIN et al. 2006a,b), Caenorhabditis elegans (WANG et al. 2002a), Xenopus (BARNARD et al. 2004), human, and mouse (KWAK et al. 2004) have been associated with poly(A) polymerase activity. There is also functional conservation at the physiological level, since S. pombe cid14 can complement the lethality of S. cerevisiae trf4-ts top1 at the restrictive temperature (WIN et al. 2006a).

Although polyadenylation had generally been thought to increase the stability of eukaryotic mRNAs, current studies challenge this view. In fact, Trf4 was recently shown to be involved in polyadenylation of hypomodified forms of tRNA^met, targeting them for degradation by the nuclear exosome, and as such to participate in an unanticipated RNA quality control mechanism in yeast (KADABA et al. 2004). RNAs encoded by intergenic regions transcribed by RNA polymerase II, as well as a number of Pol I and Pol III rRNA transcripts, pre-snRNAs, and pre-snoRNAs, are also targets of this RNA surveillance system (LACAVA et al. 2005; VANACOVA et al. 2005; WYERS et al. 2005; EGECIOGLU et al. 2006; KADABA et al. 2006). Trf4 protein has little activity on its own but is the catalytic subunit of a poly(A) polymerase complex, termed TRAMP, that contains Air1 and/or Air2, putative RNA-binding subunits, and Mtr4, a putative helicase (LACAVA et al. 2005; VANACOVA et al. 2005; WYERS et al. 2005). Trf5 also has poly(A) polymerase activity and is a component of the TRAMP5 complex, which contains Trf5, Air1, Mtr4, and targets such as rRNA (HARACSKA et al. 2005; EGECIOGLU et al. 2006; HOUSELEY and TOLLERVEY 2006). Degradation of the Trf targets is mediated at least in part by Rrp6, a component of the nuclear exosome, whose nucleolytic activity is promoted by association with the TRAMP4/5 complexes, which, in turn, are stimulated to make their associated RNAs better substrates for the exosome (HOUSELEY et al. 2006). However, none of the RNAs thus far identified as targets of these new poly(A) polymerases accounts for their protein/protein interactions, the genetic interactions trf mutants display, or the defects in genome stability that are the hallmark of trf4 and trf5 mutants. Both Trf4 and Trf5 are nuclear proteins and are chromatin bound (WANG et al. 2002b; HUH et al. 2003). trf4 is synthetic lethal with mutants lacking topoisomerase I, which retains superhelical tension within chromosomes in a viable range, with the DNA replication helicase/nuclease, dna2 (M. E. BUDD, C. C. REIS and J. L. CAMPBELL, unpublished data), and with many genes involved in chromatin dynamics and histone modification (SADOFF et al. 1995; CASTANO et al. 1996a; PAN et al. 2006). trf5 is synthetic lethal with genes involved in DNA replication, such as cdc45 and cdc8 (TONG et al. 2004). In addition, Trf5 interacts with pol , a replicative DNA polymerase, by two-hybrid assays, and both Trf4 and Trf5 interact with pol in in vitro pull-down assays (EDWARDS et al. 2003). trf4 mutants are viable at 30° but are inviable at 16° (SADOFF et al. 1995). Deletion of TRF5 alone produces no phenotype but is lethal in a trf4 background, and overexpression of TRF5 suppresses trf4 phenotypes, suggesting that the two genes perform redundant functions (CASTANO et al. 1996b; WALOWSKY et al. 1999). trf4-ts trf5 mutants are temperature sensitive for growth and show an altered cell cycle (CASTANO et al. 1996b; WANG et al. 2000). trf4 mutants show specific defects in DNA metabolism: hyperrecombination in the rDNA and sensitivity to DNA damaging agents, DNA replication inhibitors, and microtubule poisons (SADOFF et al. 1995; CASTANO et al. 1996a,b; WALOWSKY et al. 1999; WANG et al. 2002b; EDWARDS et al. 2003). The Trf mutants have been reported to have chromosome condensation and cohesion defects (WANG et al. 2000; CARSON and CHRISTMAN 2001; EDWARDS et al. 2003), although subsequent studies did not find a defect in trf4 cells in sister chromatid cohesion (PETRONCZKI et al. 2004). All of these observations suggest that the Trf's may have targets that regulate stable chromosome transmission in addition to the targets identified to date, which consist of stable RNA species, such as rRNA, snRNA, snoRNA, and tRNA.

In this study, we show that inactivation of Trf4 and Trf5 poly(A) polymerases or deletion of Rrp6 nuclease of the exosome leads to abnormally high levels of mRNAs encoding the core histones of the nucleosome. Furthermore, we observe strong synthetic interactions between trf4 mutants and genes required for histone homeostasis both at the protein and at the mRNA levels. Since we also find that the trf and rrp6 mutants are hypersensitive to ectopic histone overexpression, we propose that Trf4, Trf5, and Rrp6 contribute to a previously undocumented level of histone mRNA regulation in yeast.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS LITERATURE CITED

 
Yeast strains and plasmids:
Strains and plasmids used in this study are listed in Tables 1 and 2, respectively. All strains constructed in this study are derived from W303 RAD5+ except HA-tagged versions of TRF4 and TRF5, which were derived from OAy470 (W303 MATa, bar1::hisG rad5-535). W303 strains received from other labs were crossed at least once to W303 RAD5+ and used in this study (the RAD5 allele was determined by DNA sequencing where indicated in Table 1). Construction of Trf4 and Trf5 C-terminal HA-tagged strains and deletion mutants was performed as described (LONGTINE et al. 1998). Correct chromosomal integrations were confirmed by colony PCR using appropriate flanking primers. Strains carrying combinations of multiple mutant alleles were generated by genetic crosses. Manipulations and growth of S. cerevisiae were performed by standard procedures. Tetrad analysis was performed following standard techniques.

For site-directed mutagenesis of TRF4 and TRF5, plasmids pDLT4 (Table 2) and pRS316GALTRF5 were amplified with Pfu Ultra (Stratagene, La Jolla, CA) using primers Trf4DXDF/Trf4DXDR and Trf5DXDF/Trf5DXDR, respectively. The PCR reactions were digested with DpnI for 3 hr. DpnI was heat inactivated, and 5 µl of the reactions were used to transform Escherichia coli. Correct mutagenesis was confirmed by DNA sequencing.

Cell synchrony studies:
Trf4 and Trf5 HA-tagged strains were arrested in G1, S phase, and G2/M by supplementing the media of exponential cultures with 10 µg/ml -factor, 200 mM hydroxyurea, or 10 µg/ml nocodazole, respectively, for 2 hr. For the trf4-ts trf5 strain, arrest in G1 was performed by adding 10 µg/ml -factor to exponentially growing cells at the permissive temperature of 30° for 1 hr 30 min, after which cultures were shifted to 37° and -factor was readded to a final total concentration of 15 µg/ml. Cells were held at the restrictive temperature for 1 hr before the release from -factor to reduce the effects of heat shock.

Flow cytometry:
Approximately 5 x 10⁶ cells were fixed in 70% ethanol and kept at –20° until further processing. Cells were washed in 1 ml of 50 mM sodium citrate buffer, pH 7.4 and resuspended in 500 µl of the same buffer. RNase A was added to a final concentration of 1 mg/ml and the reactions were incubated at 50° overnight. Subsequently, proteinase K was added to the samples to a final concentration of 1 mg/ml and incubated for 1 hr at 50°. After the incubation, 500 µl of the sodium citrate buffer and propidium iodide (to a final concentration of 16 µg/ml) were added to each sample. After this point cells were protected from light. The samples were sonicated, filtered with mesh (3-64/32 NITEX; Tetko), and analyzed by flow cytometry.

Viability assays:
Cells were grown to saturation in medium lacking leucine (–LEU) overnight. Equivalent numbers of cells were serially diluted and plated on –LEU plates, or on freshly prepared –LEU plates containing indicated concentrations of hydroxyurea (HU) or methyl methanesulfonate (MMS), and grown for 4 days at the indicated temperatures.

Protein analysis:
Rad53 phosphorylation status was analyzed as described (BUDD et al. 2006). For quantification of the cellular levels of HA-Trf4 and HA-Trf5, whole-cell extracts were prepared by alkaline lysis and separated in SDS–8% polyacrylamide gels. After transfer to nitrocellulose membrane, protein blots were probed by anti-PSTAIRE (1:10,000) and anti-HA (1:2000) antibodies. Intensity of bands was quantified by densitometry and normalized against Cdc28 (anti-PSTAIRE).

RNA analysis:
For real-time PCR total RNA was isolated and cDNA was synthesized as described (LESUR and CAMPBELL 2004). Cultures of wild-type and trf4-ts trf5 cells (strain AC1968) were grown to early exponential phase at 30° and treated with 10 µg/ml -factor for 1.5 hr. Cultures were shifted to 37°, -factor was readded to a final total concentration of 15 µg/ml, and cells were allowed to grow at the restrictive temperature for 1 hr. Cells were then collected by centrifugation, washed, and resuspended in YPD prewarmed at 37°. Cell cycle progression was monitored by flow cytometry, and RNA extraction was carried out in cells collected 45 min and 65 min after G1 release for wild type and AC1968, respectively. Real-time PCR was performed exactly as described (LESUR and CAMPBELL 2004) except that 12.5 pmol primers were used in each reaction and that normalization was to ACT1 mRNA. For Northern blot analysis, total RNA was extracted as described (COLLART and OLIVIERO 1993). Poly(A)+ purification from total RNA was performed by using the MicroPoly(A)Purist kit (Ambion, Austin, TX). For agarose–formaldehyde Northern blots, 8 µg total RNA or 1 µg poly(A) RNA were dissolved in RNA sample loading buffer (Sigma, St. Louis), heated at 65° for 7 min, and separated in a 1.2% denaturing agarose–formaldehyde gel in 1x MOPS buffer pH 7.0. RNA Marker (no. G3191; Promega, Madison, WI) was used as a ruler. RNA was blotted onto a positively charged nylon membrane (no. 1417240; Roche, Indianapolis) by overnight capillary transfer in 20x SSC. The membranes were washed in 2x SSC after transfer. For acrylamide–urea Northern blots, 8 µg of total RNA were dissolved in loading buffer II (Ambion), heated at 80° for 8 min, and separated in a 6.5% polyacrylamide-bis-acrylamide (19:1) gel containing 6.5 M urea and 0.5x TBE. RNA ladder, low range (Fermentas) was run in adjacent lanes. After the run, RNA was transferred to a nylon membrane by electrotransfer in 0.5x TBE using a Trans-Blot Cell apparatus (Bio-Rad, Hercules, CA). Blots were UV crosslinked using a UV Stratalinker 2400 from Stratagene (autocrosslink at 120 mJ/cm²). Transfer efficiency and equivalence of loadings were assessed by staining the filters with 0.03% methylene blue in 0.3 M NaAC, pH 5.2. The oligonucleotide probes (Table 3) were radiolabeled by incubating [-³²P]ATP (Amersham Pharmacia, Piscataway, NJ) and T4 polynucleotide kinase (New England Biolabs, Beverly, MA) with 25 pmol of oligonucleotide probe at 37° for 1 hr in 50-µl reactions and purified with Micro Bio-Spin 30 columns (Bio-Rad). The blots were probed using radiolabeled oligonucleotides in ULTRAhyb-Oligo hybridization buffer (Ambion) according to the manufacturer's instructions. Following hybridizations, filters were exposed to Phosphor-Imager screens and analyzed using Storm Scanner (Molecular Dynamics, Sunnyvale, CA).

RNase H reactions:
Total RNA (20 µg) was annealed with 300 ng H4hyb2 oligo or simultaneously to 300 ng H4hyb2 and 400 ng oligo(dT) (Table 3) in 25 mM Tris pH 7.5, 1 mM EDTA, 50 mM NaCl in 100 µl volume. Samples were heated at 65° for 10 min and allowed cool to room temperature. Ten microliters of 10x RNase H buffer (200 mM Tris pH 7.5, 100 mM MgCl₂, 500 mM NaCl, 10 mM DTT, 300 µg/ml BSA) were added to the hybridization reaction together with 0.75 unit of RibonucleaseH (Promega). The reactions were incubated at 30° for 90 min. Five hundred microliters of 5:5:1 phenol:chloroform:LET buffer (100 mM LiCl, 20 mM EDTA, 25 mM Tris pH 8.0) were added to the reaction. After vortexing for 3 min and centrifugation for 3 min, the supernatant was recovered and precipitated.

	RESULTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS LITERATURE CITED

 
Trf mutants display a slow S phase but are not defective in the DNA damage or replication stress checkpoints:
We wished to further understand the mechanism underlying the phenotypes of TRF4/PAP2 mutants with respect to genome stability. One clue was the fact that Trf4/Pap2 and Trf5 interact with Pol2, the catalytic subunit of pol (EDWARDS et al. 2003). Specifically, these interactions occur via the C-terminal domain of the Pol2 protein, which is required not only for DNA replication but also for the DNA replication stress checkpoint and for the replication of specific chromatin domains (NAVAS et al. 1995, 1996; EDWARDS et al. 2003; IIDA and ARAKI 2004; TACKETT et al. 2005). Here, we have carefully analyzed two of these functions, S-phase progression and replication checkpoint status in trf4, trf5, and trf4-ts trf5 mutants. Each trf single mutant was arrested in G1 phase with -factor, the cells were released from the block, and the ensuing synchronous cell cycle was monitored by flow cytometry. Deletion of TRF4 leads to a delay in entry into and progression through S phase, while trf5 cells display a flow cytometry profile similar to wild-type cells (Figure 1A). Although the trf4 cells show an S-phase delay, they complete S phase and enter new cell cycles. Since TRF4 and TRF5 are thought to have at least partially redundant functions (CASTANO et al. 1996b), we also studied trf4-ts trf5, which grows at 30° but not at 37° (WANG et al. 2000). The double-mutant cells, synchronized with -factor, also present a significant delay in entry into S phase when released from G1 arrest at the restrictive temperature and show a clear delay in progression through S phase (Figure 1B). The delayed entry into and completion of S phase observed in trf mutants are consistent with a role for these genes in S phase. To test this idea further, we compared the steady-state levels of both Trf4 and Trf5 proteins in cells arrested in G1, S, or G2/M phases of the cell cycle. Using HA-tagged versions of Trf4 and Trf5, we found that both Trf4 and Trf5 protein expression levels peak in S-phase-arrested cells (Figure 1C). We also compared the steady-state levels of Trf4 and Trf5 protein in asynchronous cells, and, consistent with other reports (GHAEMMAGHAMI et al. 2003), Trf4 is expressed at about four times the level of Trf5 (Figure 1D). The absence of a growth defect or slow S phase in a single trf5 mutant is perhaps explained by the lower levels of Trf5 compared to Trf4. To investigate the state of the S-phase checkpoint in the trf mutants, we monitored the level of phosphorylation of the checkpoint effector kinase Rad53. We found that trf4 cells proficiently phosphorylate Rad53 when exposed to MMS (Figure 2A). Thus, the trf4 mutant is proficient in activating the checkpoint response to exogenous DNA damage. We also asked whether the slow S-phase progression (Figure 1A) resulted from endogenous DNA damage that activated the S-phase checkpoint by monitoring Rad53 phosphorylation in the absence of exogenous DNA damage. We observed no phosphorylation of Rad53 in the trf4 mutant in the absence of MMS, at 30° (Figure 2A) or at the restrictive temperature of 16° (not shown), indicating that there is insufficient spontaneous damage to activate the checkpoint and therefore probably insufficient DNA damage to account for the slowing of S phase in trf4 mutants. Rad53 was also phosphorylated in a trf4-ts trf5 mutant after addition of MMS but not in the absence of MMS, even after prolonged incubation at the restrictive temperature (Figure 2B). S-phase checkpoint activation is known to slow the progression of S phase, and we also found that the trf4-ts trf5 mutant is competent for slowing S phase in response to HU (Figure 1B, column 4). Thus, the DNA damage checkpoint, while intact, also fails to account for the slow S-phase progression of cells with both Trf4 and Trf5 functions compromised.

View larger version (52K): [in this window] [in a new window] [Download PPT slide]   FIGURE 1.— Characterization of the S-phase progression in trf4/5 mutants and determination of endogenous levels of expression of Trf4 and Trf5. (A) Cells from wild type, trf4 (AC1946), and trf5 (AC1947) were released from -factor arrest at 30°. Samples were collected each 15 min and analyzed by flow cytometry. (B) Cells from wild type and trf4-ts trf5 (AC1968) were arrested with -factor at 37° as described in MATERIALS AND METHODS. G1-arrested cells were released into the cell cycle at 37° into YPD media (columns 1 and 2) or into YPD containing 200 mM hydroxyurea (HU). Samples were collected at the indicated time points and processed for flow cytometry. Note the slow S-phase progression of trf4-ts trf5 (compare columns 1 and 2). (C) Cells from HA-tagged versions of Trf4 and Trf5 were arrested in G1 with -factor, in S phase with hydroxyurea (HU), and at G2/M with nocodazole (Noc). Equivalent numbers of cells were lysed and whole-cell extracts (WCE) were analyzed on 8% SDS–PAGE gels as described in MATERIALS AND METHODS. The levels of HA-Trf4 and HA-Trf5 were detected by anti-HA Western blot. Anti-PSTAIRE was used as loading control. The indicated ratio is calculated as intensity of anti-HA/anti-PSTAIRE signals as determined by densitometry and normalized to 1 for G1-arrested cells. (D) The steady-state levels of Trf4 and Trf5 were determined by analysis of WCE of asynchronous exponential growing cells as in C. The indicated ratio is calculated as intensity of anti-HA/anti-PSTAIRE and normalized to 1 for HA-Trf4.

View larger version (39K): [in this window] [in a new window] [Download PPT slide]   FIGURE 2.— Mutations in TRF4 and TRF5 do not lead to defects in Rad53 phosphorylation. (A) Cultures from the indicated genotypes were released from -factor arrest into YPD containing 0.02% methyl methanesulfonate (MMS) at 30°. Samples were collected before release into MMS or 30 min later and analyzed by anti-Rad53 Western blot. The asterisk indicates a cross-reacting band that was used as loading control. mec1 (U953-61A) and rad53 (U960-5C) strains were used as control for Rad53 antibody. (B) Wild-type and trf4-ts trf5 (AC1968) cells were released from -factor arrest at the restrictive temperature into YPD media containing 0.02% MMS where indicated and analyzed by anti-Rad53 Western blot at 30 min and 90 min after release from G1 into MMS.

View larger version (77K): [in this window] [in a new window] [Download PPT slide]   FIGURE 3.— Genetic interactions between TRF4 and genes involved in the DNA damage checkpoint and replication stress pathways and genes required for transcriptional repression of histones. (A–E) trf4 is synthetically lethal with rad53 and synthetically sick with a rad53 kinase-dead mutant (rad53K227A) but not with other genes involved in the checkpoint activation pathway. Strains of the indicated genotypes were crossed and tetrads dissected. In each case, circles indicate expected and recovered double mutants. Squares indicate trf4 spores. All dissection plates were incubated at 30° for at least 3 days. Strains crossed were (A) AC2061 and AC2123, (B) AC2055 and AC2163, (C) U953-61A and AC2115, (D) SPY40 and AC1959, and (E) AC1957 and AC1959. (F) trf4 asf1 shows reduced fitness and is inviable at 37°. Strains AC1946 and AC2122 were used for this cross. (G) trf4 hir1 shows reduced fitness. Strains AC1946 and AC2225 were crossed as described for A–F.

View larger version (96K): [in this window] [in a new window] [Download PPT slide]   FIGURE 5.— Deletion of HHT2 and HHF2 suppresses the inviability of trf4 rad53 and histone overexpression is toxic to trf mutants. (A) Crosses were done in sml1 background between strains AC2192 and AC2062. trf4 rad53 sml1 expected spores are indicated by circles. trf4 rad53 sml1 hht2-hhf2 spores are indicated by squares. (B) Overexpression of the four histones exacerbates the ts phenotype of trf4-ts trf5 cells. Wild-type and AC2193 trf4-ts trf5 strains were transformed with control vector pRS425 or with pPK128, a 2µ plasmid expressing the four core histones (HHT1-HHF1-HTA1-HTB1, Table 2). Serial dilutions were spotted on plates lacking leucine and allowed to grow for 4 days at the indicated temperatures. (C) Overexpression of the four core histones increases the MMS and HU sensitivities of trf4 (AC1946). Dilution assays were performed as in B on the indicated media and at indicated temperatures. (D) Overexpression of histones is toxic to mutants in subunits of NuA3 and NuA4 histone acetyltransferase complexes. Strains from the indicated genotypes were transformed with control empty vector or pPK128 carrying the four histone genes. Transformants were streaked in –LEU plates at 37° and allowed to grow for 4 days.

Rad53 activation/phosphorylation in response to DNA damage or to replication stress relies on the action of the two upstream kinases, Mec1 and Tel1. Rad53 phosphorylation in turn leads to the activation of the downstream protein kinase, Dun1. If the synthetic lethality of trf4 rad53 were due to failure of the DNA damage or replication stress checkpoint, one would expect the trf4 mutant also to require MEC1, TEL1, and DUN1 for viability and, hence, that trf4 mec1, trf4 tel, and/or trf4 dun1 would be synthetically lethal. However, mec1 trf4 mutants are viable and do not present a negative synergistic genetic interaction at 30° (Figure 3C). Even at 37° or 16°, temperatures at which the slow growth of trf4 is even more evident than at 30° (not shown), double mutants grow like the trf4 single mutant. (sml1 is present in the trf4 mec1 crosses, as the lethality of mec1 is suppressed by inactivation of SML1.) Moreover, there is no synthetic interaction between trf4 and tel1 (Figure 3D) or between trf4 and dun1 (Figure 3E). Although some of the double trf4 dun1 mutant spores appear smaller than either single mutant, when restreaked the colonies from single and double mutants are similar. Therefore, the essential role of RAD53 in the absence of TRF4 is independent of Mec1 and Tel1 and also independent of the downstream checkpoint effector Dun1 and, thus, does not appear to be related to disruption of the DNA damage or replication stress-induced checkpoint.

Synthetic genetic interactions between TRF4 and genes required for histone mRNA regulation:
In addition to and independent of its role in responding to DNA damage, Rad53 was recently reported to monitor histone protein levels and target excess soluble histones for degradation (GUNJAN and VERREAULT 2003). Interestingly, the role of Rad53 in histone regulation requires its kinase activity and is Mec1 and Tel1 independent, but excess histone levels do not lead to phosphorylation of Rad53 (GUNJAN and VERREAULT 2003). The results shown above are consistent with a role for Trf's, like Rad53, in contributing to the regulation of histone levels. If so, we reasoned that trf4 might be synthetically lethal with asf1, a histone H3/H4 chaperone important for nucleosome assembly in vivo during DNA replication, repair, and transcription (TYLER et al. 1999; SCHWABISH and STRUHL 2006). In addition, asf1 cells are defective in repression of histone mRNA transcription upon S-phase arrest with HU (SUTTON et al. 2001) and are sensitive to histone overexpression (SHARP et al. 2005). We find that deletion of TRF4 in combination with ASF1 deletion results in significantly reduced fitness at 30° and inviability at 37° (Figure 3F). This synthetic lethal interaction agrees with the prediction of a role for Trf4 in regulation of histone levels.

The HIR genes are transcriptional corepressors required for regulation of histone expression (SHERWOOD et al. 1993; SPECTOR et al. 1997). For that reason, we looked at the effect of disruption of HIR1 on trf4 cells. By genetic crosses we find that trf4 hir1 cells are synthetically sick. Double mutants in this cross were recovered; however, the double mutant spores form extremely slow-growing colonies (Figure 3G).

In conclusion, TRF4 exhibits strong synthetic interactions with ASF1 and HIR1, two genes that have been previously implicated in repression of histone transcription in yeast.

Synthetic genetic interactions between TRF4 and genes involved in histone acetylation:
The synthetic lethality between TRF4 and TOP1 that led to the initial discovery of Trf4 is intriguing, but remains unexplained. The synthetic lethal interaction between TOP1 and YNG2, a nonessential subunit of the NuA4 histone acetyltransferase (HAT) complex (CHOY and KRON 2002), raised the possibility that the lethality of trf4 top1 could be related to chromatin dynamics. To test this idea, we investigated genetic interactions between trf4 and genes involved in histone acetylation. Table 4 is a summary of the genetic interactions performed in this study. We found that trf4 is synthetically lethal with yng2 and yaf9, subunits of the NuA4 HAT complex, and with gcn5, the catalytic subunit if the ADA/SAGA HAT complexes. In addition, trf4 is synthetically lethal with a deletion of HTZ1, which codes for a replacement variant of histone H2A (see DISCUSSION). Interestingly rad53, but not mec1, is also synthetically lethal with htz1, asf1, and yaf9 (PAN et al. 2006). Shared synthetic lethal interactions have been interpreted to imply function in a common regulatory module, and therefore our results taken together with those on rad53 suggest parallel roles of Trf4 and Rad53 in histone homeostasis.

View this table: [in this window] [in a new window]   TABLE 4 Summary of genetic interactions between trf4 and several deletion mutants

trf4-ts trf5 cells display abnormally high levels of mRNA coding for the core histones:

View larger version (37K): [in this window] [in a new window] [Download PPT slide]   FIGURE 4.— Quantitative analysis of histone and other transcripts in the trf4-ts trf5 mutant. (A) Real-time PCR analysis of the levels of several transcripts in the trf4-ts trf5 mutant at the restrictive temperature. Cells were released from -factor arrest. RNA extraction was performed on cells collected at the time points after G1 release indicated in B (45 min for wild type and 65 min for the trf4-ts trf5 mutant, circled sections) at 37°. RNA levels were determined as described in MATERIALS AND METHODS. Reactions were run in triplicate, and results are plotted as fold increase in expression of the mutant over wild type after normalizing to ACT1 mRNA levels. (B) Flow cytometry used to monitor the stage of S-phase progression in trf4-ts trf5 (AC1968) in comparison to wild type. (C) Total RNA from exponentially growing trf4-ts trf5 cells after a 3-hr shift to 37° was separated on a 1.2% agarose–formaldehyde gel. Northern blot analysis was performed using the HHF2 probe to detect HHF2 transcripts, as described in MATERIALS AND METHODS. TSA1 mRNA was detected in the same blot and used as loading control because it did not vary under these conditions. The numbers beneath the blots were derived through quantification of RNA levels and indicate the ratio of mutant to wild type. (D) The same total RNA samples used for cDNA preparation for real-time PCR (A and B) were used to prepare the poly(A)+-enriched fraction and analyzed by Northern blot as in C. The numbers beneath the blots were derived through quantification of RNA levels and indicate the ratio of mutant to wild type. (E) HHF2 is polyadenylated by Pap1. Exponentially growing cells of wild type or pap1-1 (AC2207) were shifted to 37° for 1 hr to inactivate Pap1. Total RNA was analyzed by PAGE–urea Northern blots. As a loading control, ADH1 mRNA was used because it is not affected by Pap1 inactivation. (F) Introduction of TRF4 in trf4-ts trf5 cells lowers the levels of HHF2 transcript. Total RNA from exponentially growing cells after shift to 37° for 3 hr was separated in 6.5% PAGE–urea gels. Wild-type cells carried the pRS316 vector; trf4-ts trf5 carried pRS316 or pRS316TRF4 plasmids, as indicated. TSA1 was detected in the same blot and used as loading control. Note the lower loading in the middle lane. (G) Real-time PCR analysis of the levels of several transcripts in the trf4 mutant in asynchronous cells. Reactions were run in triplicate, and results are plotted as fold of expression of trf4 over wild type after normalizing to ACT1 mRNA levels. (H) Analysis of HHF2 transcript in trf4 and trf5 single mutants. Total RNA from exponentially growing cells of wild type, AC1946, and AC1947 was analyzed by PAGE–urea Northern blot.

Since the replication-dependent histones are polyadenylated in yeast, we also asked if the excess RNA was represented in the poly(A)+ RNA fraction. As shown in Figure 4D, analysis of the same RNA samples used in Figure 4, A and B, revealed significant accumulation of poly(A)+ RNA in the mutant compared to wild type. Since Trf4 and Trf5 are either defective or missing, this suggests that Pap1, the conventional poly(A) polymerase, may polyadenylate histone mRNAs. Indeed, we find that thermal inactivation of Pap1 in a pap1-1 mutant leads to a reduction in the amount of HHF2 transcript, with a fraction of HHF2 mRNA migrating faster (Figure 4E, arrows). Thus, the histone mRNAs are targets of Pap1. There is clearly residual polyadenylated HHF2 in the pap1-1 strain (Figure 4E), as verified by enrichment on oligo(dT) cellulose, but deleting TRF4 or TRF5 in the pap1-1 mutant did not reduce the amount of residual polyadenylated HHF2 (data not shown).

Finally, we show that upregulation of histone mRNA levels is specifically attributable to the mutations in TRF4/5, since introduction of TRF4 in a single-copy plasmid into trf4-ts trf5 cells reduces the HHF2 mRNA levels back to wild-type amounts (Figure 4F).

We also examined the level of HHF2 RNA in the trf4 and trf5 single mutants, respectively. Quantitative PCR, performed on exponentially growing trf4 cells, shows no significant alteration in the levels of any of the transcripts coding for the four core histones when compared to wild type (Figure 4G). Lack of histone mRNA overexpression was confirmed for trf4 and extended to trf5 by Northern blotting (Figure 4H). Note that in our hands, quantification of acrylamide–urea Northern blots tends to overestimate the difference in the levels of the RNA species analyzed when compared to agarose–formaldehyde gels. For instance, compare Figure 4, C and D, with Figure 4F in which the extent of HHF2 mRNA overexpression in the trf4-ts trf5 strain is 2.4-fold in agarose Northern blots—an estimation in the range of the one obtained by real-time PCR (Figure 4A)—whereas the value rises to 6.8-fold in acrylamide Northern analysis.

Our observations suggest that Trf4 and Trf5 have a redundant role in histone regulation and that each can compensate for the loss of the other.

Deletion of HHT2-HHF2 suppresses the synthetic lethality between TRF4 and RAD53 and histone overexpression is toxic to trf mutants:
Given the multiple functions of Rad53 in yeast cells and the lack of previously reported protein-encoding mRNA targets for Trf4, we sought further evidence that the synthetic lethality of trf4 rad53 was related to the observed histone mRNA expression and presumed histone imbalance. We asked if the synthetic lethality between rad53 and trf4 could be suppressed by lowering the histone dosage in the cell. We crossed a double mutant rad53 hht2-hhf2 with trf4. As observed above, rad53 trf4 spores either were missing or formed extremely sick spores. On the contrary, all the expected rad53 trf4 hht2-hhf2 spores were recovered and formed viable colonies (Figure 5A), supporting the proposal that the inviability of rad53 trf4 is due to the combined failure of two pathways to control the cellular levels of histone mRNA and protein, one governed by Trf4 and the other by Rad53.

Conversely, we investigated the effect of histone overexpression in trf mutants. Delivery of the four core histones in a multicopy vector under the control of a constitutive promoter results in balanced overexpression of the four core histones and does not affect wild-type growth (see below). However, as shown in Figure 5B, constitutive histone overexpression exacerbates the ts phenotype of the tr4-ts trf5 mutant. In addition, trf4 cells become hypersensitive to MMS and HU when histones are overexpressed (Figure 5C). Thus, excess of histones may explain, at least in part, the sensitivity of trf4 to exogenous DNA damage and replication stress. Ectopic histone overexpression has no effect on the growth of mutants deficient in the classical poly(A) polymerase, pap1-1 (not shown).

Because the trf4 mutant is synthetically lethal with yng2, gcn5 (this study), and yaf9 (PAN et al. 2006), we decided to evaluate the effect of histone overexpression in these mutants. As shown in Figure 5D, histone overexpression has a strong deleterious effect in each strain. Conversely, we do not detect a reduction in the amount of acetylated bulk histone H4 in the trf4 or the trf4-ts trf5 strains (data not shown). Altogether, these results suggest that deregulation of histones underlies the synthetic lethality between TRF4 and YNG2/YAF9/GCN5.

rrp6 cells, but not air1/2, have increased histone mRNA levels and are sensitive to histone overexpression:
We reasoned that if Trf's are required to prevent excess transcripts coding for the core histones, then mutants in other members of the TRAMP complex or in the Rrp6 exoribonuclease subunit of the exosome should also display unusually high levels of histone mRNAs and hypersensitivity to histone overexpression. When we analyzed the effect of deleting two putative RNA-binding subunits of TRAMP and TRAMP5 complexes, AIR2 and AIR1, respectively, we found that air1 and air2 single mutants, as well as air1 air2 double mutants, show wild-type levels of transcript encoding HHF2 (Figure 6). Consistent with this, overexpression of histones has no deleterious effect on the growth of air1, air2, or air1 air2 strains even when these mutants are challenged with exogenous DNA damage (not shown). Therefore, Trf4 and Trf5 can maintain appropriate histone mRNA levels in the absence of Air1 and Air2, suggesting that there may be other RNA-binding proteins besides Air1 and Air2 and that mediate Trf interaction with specific substrates that lead to histone mRNA upregulation. Evidence for this is presented in supplemental Figure 1 at http://www.genetics.org/supplemental/.

View larger version (55K): [in this window] [in a new window] [Download PPT slide]   FIGURE 6.— The RNA-binding proteins Air1 and Air2 do not play a role in histone mRNA level regulation. HHF2 mRNA levels were analyzed in total RNA samples from asynchronous cells of indicated genotypes (strains AC2164, AC2208, AC2231, and AC2232) by PAGE–urea Northern blots. HHF2 RNA levels were normalized with respect to loading controls and the ratio of mutant to wild-type levels is shown.

View larger version (46K): [in this window] [in a new window] [Download PPT slide]   FIGURE 7.— rrp6 cells are sensitive to histone overexpression and display abnormally high levels of HHF2 mRNA. (A) Histone overexpression exacerbates the temperature-sensitive phenotype of rrp6. Serial dilutions were performed as in Figure 5B, using strain AC2161 and the pRS425/pPK128 plasmids. Plates were incubated at the indicated temperatures for 3 days. (B) Levels of transcripts encoding for HHF2 were analyzed in exponentially growing cells of rrp6 and isogenic wild-type cells at 30°. Total RNA samples were analyzed in PAGE–urea gels and Northern blotting was performed as in Figure 4, F and H. Numbers are RNA levels normalized to wild type. (C) Cell cycle profile of HHF2 mRNA levels in rrp6 or wild-type cells following G1 release at 30°. RNA was collected from samples at the indicated times after -factor release and analyzed by PAGE–urea Northern blots. PhosphorImager analysis of the Northern blot is shown in E. (D) Cell cycle progression of the cells analyzed in C was monitored by flow cytometry. (E) HHF2 mRNA levels from C were quantified using a PhosphorImager and normalized to the ADH1 control. The maximum intensity of the HHF2 band in wild type was defined as 1.0. ,wild type; , rrp6. (F) Cell cycle profile of HHF2 levels in trf4-ts trf5 or wild-type cells following G1 release at 37°. Analysis was carried out as in C. (G) Cell cycle progression of the samples analyzed in F was monitored by flow cytometry. (H) HHF2 mRNA levels from F were quantified using a PhosphorImager and normalized to ADH1 mRNA levels. The maximum intensity of the HHF2 band in wild type was defined as 1.0. , wild type; , trf4-ts trf5.

We next investigated the temporal pattern of HHF2 mRNA expression in trf4-ts trf5, as shown in Figure 7, F–H. We observed that the increase in accumulation of HHF2 mRNA, as in the rrp6 strain, is pronounced in S phase and that negative regulation is mostly intact in G1, G2, and M phases. Higher levels of HHF2 mRNA are observed outside of S phase in the trf4-ts trf5 mutant when compared to wild type or rrp6. However, the slightly elevated levels of expression seen outside of S phase in the trf4-ts trf5 may be related to inability to fully synchronize this mutant with -factor and resulting asynchrony, or there may be some derepression in trf4-ts trf5 cells. Thus, it appears that it is the DNA replication-related histone mRNA accumulation that is deregulated in the trf and rrp6 mutants. This phenotype is very similar to that observed with asf1 or with hir1 mutants (SUTTON et al. 2001).

Analysis of poly(A) tail length of HHF2 mRNA in trf's and rrp6 mutants:
One possible substrate for polyadenylation in the Trf/Rrp6-mediated regulatory system is the histone mRNAs themselves, although, to date, no mRNA transcripts have been identified as Trf substrates. We therefore inquired about the length of the HHF2 poly(A) tail in different mutant backgrounds to determine if histone mRNAs are directly polyadenylated by Trf4 and Trf5. To analyze poly(A) lengths at high resolution, RNase H-directed cleavage was used. A DNA oligonucleotide complementary to the terminal portion of the HHF2 transcript 214 nucleotides upstream of the first major transcription termination site was annealed to total RNA extracted from wild-type and tr4-ts trf5 cells. Following RNase H digestion we estimated the lengths of the poly(A) tail by Northern blot by comparison to samples hybridized to oligo(dT) before RNase H treatment to remove poly(A) tails. As shown in Figure 8A, HHF2 transcripts carry heterogeneous poly(A) tails that extend up to 70 bp. We found no significant increase in the length of the polyadenylated HHF2 mRNA population derived from the wild-type or rrp6 strains upon TRF4 (Figure 8A) or TRF5 overproduction (not shown) or decrease in HHF2 mRNA length in the tr4-ts trf5 strain (Figure 8B). Perhaps the presence of Pap1 in the strains, instability of the RNAs polyadenylated by Trf polymerases, or the multiple transcription termination sites render any changes in poly(A) tail length undetectable (see DISCUSSION).

View larger version (58K): [in this window] [in a new window] [Download PPT slide]   FIGURE 8.— Analysis of poly(A) tail length in rrp6 and trf4-ts trf5 mutants. (A) Overexpression of TRF4 does not lead to a detectable increase in the poly(A) tail length of HHF2 in wild-type or rrp6 backgrounds. Strains of indicated genotypes were grown in –URA raffinose medium to exponential phase. Overexpression of TRF4 and trf4DADA alleles was induced by adding 2% galactose to the medium for 3 hr. Total RNA was digested with RNase H and indicated oligonucleotides. Digested RNAs were analyzed by PAGE–urea Northern blots. (B) Analysis of poly(A) tail length of HHF2 mRNA in wild-type and trf4-ts trf5 cells. RNase H digestion and Northern blot are as in A.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION ACKNOWLEDGEMENTS LITERATURE CITED

This study provides genetic and biochemical evidence that Trf4 and Trf5 make a redundant contribution to genome stability in yeast through control of histone mRNA levels during S phase. We show that the mRNAs coding for the four core histones, but not other cell cycle-regulated transcripts tested, accumulate to abnormally high levels in S phase in a trf4-ts trf5 mutant whereas deletion of each TRF gene per se does not result in a significant increase in histone mRNAs. So far, no other true mRNA products of RNA Pol II transcription have been found to be upregulated in Trf mutants. That this overexpression has a physiological impact is supported by the fact that additional ectopic overproduction of histones decreases the viability and increases the DNA damage sensitivity of trf mutants with no effect in the wild-type isogenic strain. Moreover, trf4 cells require RAD53 for viability. This supports the idea that Trf4 and Trf5, like Rad53, are involved in regulation of histone amounts and that histone abundance reaches lethal levels in the absence of both pathways. Although trf4 and trf4-ts trf5 mutants exhibit a slow S phase, the synthetic lethal interaction between TRF4 and RAD53 is unlikely to result from DNA damage due to slow or aberrant DNA synthesis, since induction of the DNA damage response is not observed in the trf mutants. This is true even in the trf4-ts trf5 double mutant at the restrictive temperature, even though the checkpoint is intact and can be activated in the presence of exogenous DNA-damaging agents. Supporting this, no synthetic interactions are observed between TRF4 and the master checkpoint kinases, MEC1 and TEL1, indicating that a function specifically carried out by Rad53 and independent of the general DNA damage checkpoint activation pathway is required for trf4 viability. Finally, trf4 rad53 lethality is suppressed by lowering histone dosage in the cell, demonstrating that excess histones underlie trf4 rad53 inviability. We therefore propose that the slow S phase and related defects in chromosome maintenance in trf mutants are related, at least partly, to defects in regulation of the S-phase histone mRNAs.

The effects on histone mRNA accumulation in trf4-ts trf5 are furthermore likely attributable to the established role of the interplay between Trf4/Trf5 and the nuclear exosome in RNA degradation. An attractive feature of this model is that it provides an explanation to why loss of Rrp6 leads to the same histone mRNA phenotypes as trf4-ts trf5. In as much as Trf4 and Trf5 are thought to be exclusively nuclear and Rrp6 is found only in the nuclear exosome and not in the cytoplasmic, histone mRNA regulation may constitute an essential function of the nuclear exosome. The rrp6 null mutant is temperature sensitive for growth at 37°, has slow growth at 30°, and, like the trf mutants, shows benomyl and MMS sensitivity (BEGLEY et al. 2002; DANIEL et al. 2006). A recent study found that a considerable number of mRNAs are upregulated in rrp6 cells (HOUALLA et al. 2006). However, an increase in histone mRNA levels upon inactivation of Rrp6 was not detected. Interestingly, the effect of a deletion of RRP6 on other transcripts analyzed in this study was reduced at 37° (HOUALLA et al. 2006), in complete agreement with our observations for HHF2 mRNA levels, whose increase in rrp6 cells is less elevated at higher temperatures (data not shown). Air1 and Air2, RNA-binding components of TRAMP4 and TRAMP5 complexes, are not required for the maintenance of normal histone mRNA levels (Figure 6 and supplemental Figure 1 at http://www.genetics.org/supplemental/), suggesting that other RNA-binding subunit/s might be involved in the regulation of histone transcripts, even if indirectly, by Trf's and the exosome. Interestingly, the RNA-binding protein Nrd1p was recently reported to associate with and stimulate the nuclear exosome for 3'-end processing of RNA polymerase II transcripts. Nrd1, which also interacts with the TRAMP complex, has been proposed to assist the exosome in identifying certain mRNAs as targets for degradation (GAVIN et al. 2006; VASILJEVA and BURATOWSKI 2006), indicating that other RNA-binding proteins, besides Air1 and Air2, might modulate Trf4 and Trf5 action. Our genetic studies on the toxic effect of TRF5 overexpression on air1 air2 double mutants (supplemental Figure S1 at http://www.genetics.org/supplemental/) also support this hypothesis.

While the simplest model to explain our results is that histone mRNAs are substrates of Trf4- and Trf5-mediated degradation, to establish this has been more difficult than it has been for other Trf4/5 targets studied to date. There may be several reasons for this. First, the generally rapid turnover of mRNAs relative to the stable rRNA, tRNA, snRNA, and snoRNA species may pose an obstacle to the detection of mRNA degradation intermediates. Second, there are at least two major transcription termination sites for HHF2 [Figure 8, A and B, oligo(dT) lanes], as reported by others (CROSS and SMITH 1988). Third, HHF2 RNA is a substrate of the classical poly(A) polymerase, Pap1, which adds heterogeneous length poly(A) tails (Figure 4E). In addition to the heterogeneity of histone mRNAs making it difficult to determine whether Trf4 and/or Trf5 polyadenylate histone mRNAs, it is also possible that the instability or low abundance of histone species polyadenylated by Trf's precludes their detection. At this point we still cannot discard the possibility that the inability to see effects of inactivation or overproduction of TRF's on histone mRNA length, even in rrp6 strains, is due to the heterogeneity, instability, or low abundance of the mRNAs, putatively polyadenylated by Trf4/5. It is possible that the histones and perhaps other mRNA levels are regulated through a competition between stabilization and degradation involving multiple Paps.

Genetic analysis reported here (Table 4) and additional findings reported elsewhere (PAN et al. 2006) support the existence of a module of interconnected pathways that regulate histone homeostasis and thereby ensure effective chromatin assembly (see Figure 9 for schematic). We show that the trf mutants exhibit synthetic fitness defects with asf1, hir1, and rad53, each of which causes elevated histone levels. Given the wide spectrum of functions assigned to Asf1 and the Hir1 complex (OSLEY and HEREFORD 1982; OSLEY and LYCAN 1987; XU et al. 1992; SHARP et al. 2001, 2005; SUTTON et al. 2001; GUNJAN and VERREAULT 2003; ADKINS and TYLER 2004; GROTH et al. 2005; RECHT et al. 2006; SCHWABISH and STRUHL 2006), it is hard to stipulate as to the exact reason why trf4 asf1 or trf4 hir1 are synthetically lethal, but the synthetic effects may be due to the shared roles of Asf1 and Hir1 in regulating transcription of the histone mRNAs. Regulation of histone mRNAs by Trf's could operate either at the level of histone transcription or at the level of the stability of histone mRNAs. Negative controls operate on the histone genes during inhibition of DNA replication and in G1, G2, and M phases of the cell cycle, and positive controls activate transcription in S phase. Although the precise mechanism of histone transcriptional regulation in yeast is unknown, the histone promoters (with the exception of HTA2–HTB2) contain well-characterized cis-acting positive and negative regulatory promoter elements (OSLEY et al. 1986; OSLEY 1991; FREEMAN et al. 1992). The Hir1 complex and Asf1 contribute to negative regulation through these sequences (OSLEY and LYCAN 1987; CROSS and SMITH 1988; MORAN et al. 1990; XU et al. 1992; SHERWOOD et al. 1993; DOLLARD et al. 1994; COMPAGNONE-POST and OSLEY 1996; SPECTOR et al. 1997; SUTTON et al. 2001). Since we demonstrate that HTA2 and HTB2 transcripts are upregulated in trf4-ts trf5 cells, we do not favor the idea that Trf's are involved in the Hir1 or Asf1 transcriptional repression pathway because the promoter of the HTA2–HTB2 locus lacks the negative element required for this regulation (OSLEY and LYCAN 1987; SUTTON et al. 2001). More likely, they participate in a parallel pathway, such as post-transcriptional regulation (LYCAN et al. 1987; XU et al. 1990). The importance of post-transcriptional regulation of histone mRNAs is underscored by the fact that even when expressed from a constitutive promoter, full-length HTB1 mRNA is periodically regulated, indicating that regulation at the post-transcriptional level is sufficient to confer cyclic oscillation of histone mRNA (LYCAN et al. 1987; XU et al. 1990). Genes involved in this mechanism in yeast have not been identified, although 3'-end sequences of HTB1 required for this level of regulation have been mapped (CAMPBELL et al. 2002).

View larger version (8K): [in this window] [in a new window] [Download PPT slide]   FIGURE 9.— Summary of pathways regulating histone levels in yeast. See text for details.

An additional aspect of our genetic analysis worth comment is the synthetic lethality between the trf genes and components of the histone acetyltransferase complexes (Table 4) a