Präsident der Humboldt-Universität zu Berlin
Prof. Dr. Jürgen Mlynek
The essential histone H3 variant Cse4 plays a crucial role at the centromere in S. cerevisiae, where it replaces histone H3 in that it assembles centromere specific (Cse4-H4)2 tetrameres. We found in our study that the histone H3 variant was able to interact over its unique N-Terminus with two subunits of the histone acetyltransferase complex SAS-I: Sas2 and Sas4. Mutations within the acetyl-CoA binding site (HAT domain) or the zink-finger of Sas2 disrupted the binding to Cse4, although an indirect interaction was found with co-immunoprecipitation experiments.
Additionally, the N-terminus of Cse4 interacted with Cac1, the largest subunit of the chromatin assembly factor CAF-I and Asf1 – two histone chaperones that assemble histones H3 and H4 into nucleosomes. Our findings further suggest a role of Cac1 independent of Cac2 and Cac3 as no binding to Cse4 could be detected. A role for Sas2 at the centromere was further confirmed in that a sas2 deletion (sas2 delta) disrupted the binding of Cse4 to Ctf19. Additionally, sas2 delta partially rescued the temperature sensitivity of a cse4-103 mutated strain at elevated temperatures, suggesting a role for Sas2 in improving centromere stability. An important question resulted from our studies: is Sas2 able to acetylate the histone H3 variant Cse4 ? We have circumstantial evidence that Cse4 was indeed acetylated in the cell, but whether Sas2 accounts for the acetylation remains to be determined.
Die essentielle Histon H3 Variante Cse4 ersetzt am Centromer das Standard Histon H3 und bildet zusammen mit Histon H4 funktionelle Cse4-H4 Tetramere aus. In dieser Studie konnte gezeigt werden, das Cse4 über seinen einzigartigen N-Terminus mit zwei Komponenten des Histon-Acetyltransferase-Komplexes SAS-I interagiert: der enzymatischen Untereinheit Sas2 und Sas4. Mutationen innerhalb des atypischen C2HC Zink-Fingers oder der HAT-Aktivierungsdomäne von Sas2 verhindern eine Bindung an Cse4, obwohl mit Hilfe von Co-Immunopräzipitationsexperimenten eine indirekte Interaktion nachgewiesen werden konnte.
Weiterhin wurde gezeigt, dass Cse4 mit Cac1, der größten Untereinheit des Chromatin-Assemblierungsfaktors CAF-I und Asf1 interagiert – zwei Histon Chaperonen, die Histon H3 und H4 in Chromatin assemblieren. Unsere Ergebnisse lassen weiterhin auf eine separate Rolle von Cac1, unabhängig von den beiden anderen Untereinheiten schließen. Die Interaktion von Cse4 und Ctf19 wird durch eine Deletion von Sas2 verhindert. Ebenfalls kann die Temperatur-Sensitivität eines cse4-103 mutierten Hefestamms durch eine Sas2-Deletion partiell supprimiert. Somit kann man darauf schließen, dass Sas2 eine Funktion bei der Stabilisierung des Centromers aufweist.
Die bisherigen Ergebnisse lassen die Frage aufkommen, ob Cse4 in der Zelle acetyliert ist und ob es möglicherweise als Histon H3 Variante ebenfalls ein Substrat von SAS-I darstellt. Wir konnten zeigen, dass Cse4 tatsächlich in einem acetylierten Status vorliegt, ob SAS-I jedoch für die Acetylierung verantwortlich ist bleibt nachzuweisen.
In eukaryotic cells, genomic DNA is packaged into chromatin in the nucleus. Chromatin is able to undergo dynamic changes during replication, recombination, transcription and DNA repair, but also to control temporal gene expression (Wolffe and Kurumizaka, 1998). The fundamental core structure of chromatin are nucleosomes, which are repetitive units of approximately 147 bp DNA wrapped around a histone core in a left-handed superhelix (Fig. 1) (Luger, et al., 1997). The histone proteins H2A, H2B, H3 and H4 are evolutionarily conserved. They form a tripartite protein helix with a H3-H4 tetramere in the middle flanked by an H2A-H2B dimer within the nucleosome.
In mammals, the linker histone H1 binds between the single nucleosome core particles. Although an H1-homologue (HHO1) has been found in S. cerevisiae, it is still unknown whether it has the same function in stabilizing the nucleosome (Freidkin and Katcoff, 2001);(Landsman, 1996). Six to eight nucleosomes form a solenoid structure by further coiling with a diameter of 30 nm.
Chromatin can be divided into heterochromatin and euchromatin. Heterochromatin is more compact than euchromatin and contains genes that are not actively transcribed. One of the best known examples for heterochromatin is the X-chromosome in female mammals, which is inactivated in a process called dosage compensation and forms the so-called Barr body. In general, heterochromatin replicates late in the S-phase of the cell cycle and can be found in regions containing no or only few genes, such as the telomeres and the centromere. In contrast, euchromatin is “active” chromatin, which contains DNA sequences that are transcribed into RNA.
After DNA replication, recombination or repair, DNA is re-assembled into nucleosomes. This mechanism involves several protein complexes, which function as chaperones and help to integrate histones and DNA into a highly organized chromatin structure. During S-phase of the cell cycle, the parental nucleosomes become temporarily separated. After they pass the replication fork, nucleosomes are newly assembled onto the two DNA daughter strands by integrating pre-existing histones as well as newly synthesized histones.
Chromatin assembly factor I (CAF-I)
In S. cerevisiae, CAF-I is a chromatin assembly factor that delivers histone H3 and H4 to DNA during DNA replication or DNA repair (Gaillard, et al., 1996);(Kamakaka, et al., 1996);(Kaufman, et al., 1997). In order to bind to the replication fork, CAF-I interacts with the proliferating cell nuclear antigen (PCNA) (Verreault, et al., 1996).
CAF-I is an evolutionary conserved heterotrimeric complex with the subunits Cac1, Cac2 and Cac3. Deletion of any of the three CAC genes leads to an increase
Anti-silencing factor 1 (Asf1)
Asf1 is thought to act in concert with CAF-I as a chromatin assembly factor. It also promotes assembly of nucleosomes in vitro (Tyler, et al., 1999) and the D. melanogaster homologue of Asf1 was shown to interact with histones H3 and H4 that carry the acetylation pattern of newly synthesized histones (Tyler, et al., 2001).
ASF1 was originally identified in a screen for high-dosage disrupters of silencing at the mating-type loci in S. cerevisiae (Le, et al., 1997). When overexpressed, it also leads to reduced silencing at the telomeres (Le, et al., 1997);(Singer, et al., 1998) and at the rDNA locus (Singer, et al., 1998). Asf1 is not essential for the cell, but a deletion causes defects in heterochromatic gene silencing, slow growth due to a lengthened S-phase, sensitivity to DNA damaging and replication blocking agents (Le, et al., 1997);(Tyler, et al., 1999) and an increase in chromosome loss (Le, et al., 1997).
Additionally, Asf1 interacts with the CAF-I subunit Cac2 and increases CAF-I activity in nucleosome assembly in Drosophila as well as in yeast (Mello, et al., 2002);(Tyler, et al., 2001). Inactivation of both CAF-I and Asf1 leads to a synergistic reduction in heterochromatic gene silencing, since double mutants display more severe phenotypes than strains with either single mutant (Tyler, et al., 1999). These results imply that CAF-I and Asf1 are both chromatin assembly
Histone regulation genes (Hir)
Gene products from the HIR (histone regulatory) genes HIR1 and HIR2 have been shown to interact with Asf1 in vitro and in vivo (Sharp, et al., 2001);(Sutton, et al., 2001), implying that they function together in a silencing pathway that is also PCNA dependent and partially overlaps with the CAF-I silencing pathway (Krawitz, et al., 2002).
The HIR genes HIR1, HIR2, HIR3 and HPC2 encode proteins that tightly regulate histone gene transcription (Osley and Lycan, 1987);(Xu, et al., 1992). They code for repressors proteins that bind to the histone promotors in early G1-, late S- and in G2/M-phase and therefore prevent the histone genes from being transcribed (Osley, et al., 1986). Additionally, Hir proteins contribute to Asf1-mediated nucleosome assembly (Sharp, et al., 2001). The HIR-genes are exclusively expressed during G1/S transition in yeast, whereas in other phases of the cell cycle they become repressed by a mechanism that is thought to involve a specialized chromatin structure (Dimova, et al., 1999).
Mutations in HIR genes have only minor effects on silencing at the telomeres and the HM loci (Kaufman, et al., 1998), but when combined with mutations in CAF-I subunits, yeast cells display synergistic reduction of position-dependent gene silencing both at the HM loci and the telomeres, increased sensitivity to DNA damaging agents and slow growth (Kaufman, et al., 1998);(Qian, et al., 1998);(Sharp, et al., 2001). A new role for histone interacting and –deposition proteins has been described at the centromere, where CAF-I and Hir proteins function in
Nucleosome assembly protein 1 (Nap1)
After deposition of histone H3 and H4, histone H2A and H2B are integrated into chromatin to form the core nucleosome. The nucleocytoplasmatic shuttle protein Nap1 (nucleosome assembly protein 1) binds to histone H2A and H2B and helps to assemble chromatin from newly synthesized DNA (Ito, et al., 1996). Since Nap1 has been localized in several cell compartments, the current model posits that Nap1 functions as a histone chaperone that binds H2A and H2B already in the cytoplasm. Due to interaction with Kar114p, the primary karyopherin/importin responsible for the nuclear import of H2A and H2B, it helps to import them into the nucleus, where it incorporates the histones into newly replicated DNA (Mosammaparast, et al., 2002).
Histones consist of a highly conserved globular histone-fold domain and an N-terminal domain that remains outside of the nucleosome (Luger, et al., 1997). The N-termini as well as the core region are targets for posttranslational modifications, e.g. acetylation, phosphorylation, methylation and ubiquitination, which result in changes of gene activity (Spencer and Davie, 1999).
Methylation or acetylation occurs on the ε-amino group of positively charged lysines (Fig. 2). In the case of acetylation, the charge of the amino group is then
Several histone methyltransferases (HMTs) have been characterized. One example are the SUV39 family proteins that exist in D. melanogaster, humans and S. pombe, and selectively methylate histone H3 lysine 9 (Rea, et al., 2000). This methylation has been shown to be a prerequisite for binding of the heterochromatin binding protein 1 (HP1), which connects H3 K9 methylation to silencing and heterochromatin formation (Bannister, et al., 2001);(Lachner, et al., 2001). Additionally, methylation on H3 K9 prevents phosphorylation of H3 S10, which is known as a modification involved in gene activation (Lo, et al., 2001);(Lo, et al., 2000). Methylation can also lead to transcriptional activation, e.g. methylation of H3 K4 in S. cerevisiae (Noma, et al., 2001). It is becoming clear that histone modifications are able to influence each other and work together in the form of a “histone code”, thereby creating a specific chromatin structure that is connected with gene activity.
Histone acetylation is carried out by several known histone acetyltransferase (HATs), and as acetylation is a reversible process, there also exist histone deacetylases (HDACs) as antagonists in the cell. In general, histones tend to be hyperacetylated in actively transcribed regions, whereas histones are hypoacetylated in transcriptionally repressed regions. The simplest model of gene
Histone deacetylases (HDACs)
Histone deacetylation has been linked to the establishment of transcriptional inactive heterochromatin. At least ten HDACs exist in S. cerevisiae: Hda1, Hos1-3, Rpd3, Hst1-4 and Sir2. Hos 1/Hos3 and Hos 2 mainly affect ribosomal DNA and ribosomal protein genes, whereas Rpd3 and Hda1, which preferentially deacetylate distinct promotors, function in histone deacetylase complexes and their absence results in hyperacetylation of histone H3 and H4 (Kuo and Allis, 1998);(Robyr, et al., 2002).
The first hint that Sir2 functions as a histone deacetylase was the observation that SIR2 overexpression leads to hypoacetylation of histones in yeast (Braunstein, et al., 1993). Sir2 has now been characterized as a phylogenetically conserved NAD+-dependent HDAC that efficiently deacetylates histone H4 lysine 16 and histone H3 lysine 9 (Imai, et al., 2000);(Landry, et al., 2000);(Smith, et al., 2000). One role of Sir2 is the establishment of silencing at the rDNA locus (Bryk, et al., 1997);(Smith and Boeke, 1997), as well as at silent mating-type loci HML and HMR and at the telomeres in a complex with other Sir proteins (Sir3 and Sir4, at the HM loci additionally Sir1) (Moretti, et al., 1994);(Rine and Herskowitz, 1987). Recently, a role in ageing has been described for Sir2 in C. elegans and in other organisms, including yeast (Guarente, 2000);(Guarente, 2001);(Imai, et al., 2000). Overexpression of Sir2 leads to an increase in life span (Kaeberlein, et al., 1999), which is determined in yeast by how many times a mother yeast cell is able to divide, whereas SIR2 deletion or NAD+-deprivation leads to a increased recombination rate and prevents longevity (Lin, et al., 2000).
In contrast to HDACs, histone acetyltransferases have the opposite effect on silencing, since the acetylation of lysine residues generally leads to the formation of euchromatin and transcriptional activation. The various existing HATs are grouped into five families based on sequence similarities. These families also include the MYST family with its representatives in yeast (Sas2, Sas3, Esa1), D. melanogaster (MOF, Chameau) and human (MOZ, MORF, Tip60, HBO1). Members of the evolutionary conserved MYST family share the MYST homology domain with a binding site for acetyl-CoA (HAT domain) and an atypical C2HC zink-finger motif.
SAS2 was originally identified in a screen for suppressors of silencing defects at HMR (Ehrenhofer-Murray, et al., 1997). In contrast to its role at the HMR locus, a deletion of SAS2 (sas2Δ) decreases silencing at HML in sir1Δ strains and derepresses silencing at the telomeres (Ehrenhofer-Murray, et al., 1997);(Reifsnyder, et al., 1996). The conserved acetyl-CoA binding domain as well as the HAT domain is essential for HML and telomeric silencing, because mutations in these domains lead to the same silencing phenotypes as a SAS2 deletion (Meijsing and Ehrenhofer-Murray, 2001);(Osada, et al., 2001). Additionally, (Meijsing and Ehrenhofer-Murray, 2001) have shown that a mutation in histone H4 lysine 16 to arginine phenocopies the effect of a sas2Δ, implying that H4 K16 is one target for Sas2 acetylation.
Sas2 exists in a complex called SAS-I together with two other proteins involved in silencing, Sas4 and Sas5 (Meijsing and Ehrenhofer-Murray, 2001);(Osada, et al., 2001) and acetylates next to histone H4 lysine 16 also H3 lysine 14 in vitro (Sutton, et al., 2003). The association with Sas4 is essential for Sas2’s acetyltransferase activity, whereas binding of Sas5 is required for improving the activity of the complex (Sutton, et al., 2003). Furthermore, Sas2 has been shown to interact with the chromatin assembly factors CAF-I and Asf1 (Meijsing and Ehrenhofer-Murray, 2001);(Osada, et al., 2001),
Three silenced regions are known in the yeast S. cerevisiae: the silent mating-type loci HML and HMR on chromosome III, the rDNA locus on chromosome XII and the telomeres. Both silencing at the HM loci and at the telomeres requires a common set of trans-acting factors, which include the Sir (silent information regulator)-proteins, the N-termini of histone H3 and H4, Abf1 and the repressor-activator protein Rap1. The structure of the different silenced regions and their common and distinct features are discussed below.
Silencing at the mating-type loci HML and HMR
Haploid yeast cells can display either an a- or a α-mating-type. The cell type is defined by the allele present in the MAT locus on chromosome III. Whereas cells with a MATa allele are of a-mating-type, and cells with the MATα allele are of α-mating-type. Under certain circumstances, a cell can switch its mating-type by cleaving the mating-type locus with HO endonuclease and replacing the MAT with information from HML or HMR.
Additionally to MAT, the cell contains unexpressed copies of the mating-type information genes near the telomeres on the same chromosome: HMR generally codes for a-information and HML for α-information. The repression of the HM loci is maintained by flanking silencing elements (E and I silencer) as well as by silencer binding proteins. Whereas the E-silencer is absolutely essential for
To exemplify silencing at the HM loci, silencing at the well-studied HMR silencer is explained subsequently and in Fig. 3. The a-information at HMR is flanked by the HMR-E and the HMR-I silencers. The ~140 bp HMR-E contains binding sites for the origin recognition complex ORC, Rap1 and Abf1, whereas the HMR-I silencer lacks the Rap1 binding site. Rap1 and Abf1 are regulatory proteins that function as transcriptional activators in other positions in the genome beside their role in silencing (Shore, 1997).
ORC, Rap1 and Abf1 serve as a platform for binding of the Sir-proteins to HMR-E. Deletion of Sir1 only causes a weak silencing defect, whereas deletion of SIR2-4 leads to complete derepression. Through the interaction with ORC, Sir1 facilitates the binding of Sir2, Sir3 and Sir4 to the silencer. Hereby, Sir3 and Sir4 interact directly with Rap 1 and the N-termini of histone H3 and H4 (Moretti, et al., 1994);(Triolo and Sternglanz, 1996). Sir2 further improves efficient spreading
Silencing at the rDNA locus
Another silencing locus in S. cerevisiae is the highly repetitive rDNA locus on chromosome XII. The rDNA locus encodes the ribosomal RNAs, which are not translated into proteins, but establish the ribosome together with ribosomal proteins. One rDNA unit consists of ~9.1 kb, which is tandemly repeated in 100-200 copies per cell. Silencing at this locus is thought to regulate the access of RNA polymerase I, but prevents RNA polymerase II from transcription (Shou, et al., 2001).
At the rDNA locus, Sir2 assembles together with Net1, Cdc14, Nan1 and PolI into the so-called RENT (regulator of nucleolar silencing and telophase exit) complex. In sir2Δ cells, the rDNA silencing is decreased and the chromatin structure is less compact (Smith and Boeke, 1997), whereas the recombination rate is increased (Gottlieb and Esposito, 1989). Thus, Sir2 improves rDNA silencing via its deacetylation activity.
The structural core component of the RENT complex is Net1, which recruits the other RENT-proteins to the rDNA locus. Net1 binds within the RENT complex to PolI, the RNA polymerase I, and stimulates its enzymatic activity (Shou, et al., 2001). Whether Net1 directly binds to DNA is still unknown, but it is possible that other still unknown proteins contribute to rDNA silencing. Interestingly, when NET1 is overexpressed, it is also associated with the HMR-E silencer, whereas net1-1 acts indirectly in HMR silencing by releasing Sir2 from the nucleolus (Kasulke, et al., 2002).
Telomeres can be found at the ends of all 16 chromosomes in yeast. They consist of a 300-350 bp region with irregular TG1-3 repeats, which are disrupted by X- and Y-elements of variable size (Palladino and Gasser, 1994). The Rap1 binds on average every 18 bp in the telomeric repeats and is a structural core component of the telomeres. The Rap1 binding region is also referred to as the telosome and is characterized by its nuclease resistance.
Rap1 in turn recruits Sir2, Sir3 and Sir4 to the telomeres, where Sir3 and Sir4 bind to the N-termini of histone H3 and H4. The Sir proteins spread in a histone-dependent manner up to 2.8 kb into the core telomeric heterochromatin. (Strahl-Bolsinger, et al., 1997) have proposed a model in which telosomal Rap1 folds back onto subtelomeric regions. This allows a condensation at telomeric heterochromatin due to the interaction between Rap1 and the Sir’s, as well as among the Sir proteins themselves.
Silencing at the telomeres and subsequent condensation prevents the chromosome ends from being degraded and improves their replication (Palladino, et al., 1993). Additionally, the positioning of the telomeres to the nuclear periphery is maintained.
Centromeres are the site of kinetochore formation and mictotubuli attachment on the chromosome. They vary in size and form in eukaryotes, but all have a key role in chromosome inheritance and are therefore essential for the transfer of genetic information from one cell to its daughter cell. Most eukaryotes contain monocentric centromeres, where the centromere-kinetochore complex is formed at a single point on the chromosome. One exception are holocentric organisms like the nematode Caenorhabditis elegans, which have holocentric kinetochores formed along the entire chromosome (Albertson and Thomson, 1982).
Centromeres consist of centromeric DNA (CEN DNA) and an assembled specialized nucleoprotein complex termed kinetochore (Clarke, 1998). After chromosomes have replicated during S-phase, sister chromatids are still attached to each other at the kinetochore. During mitosis, several spindle microtubuli bind to the kinetochore and mediate chromosome movement to the nuclei of the daughter cells. In order to assure accurate segregation, microtubuli attachment is closely monitored, so that the cell cycle can be arrested in case of incomplete or inappropriate attachment (Nicklas, et al., 1995);(Rudner and Murray, 1996).
Although the centromere was first identified by Walther Flemming more than 120 years ago, the complex structure and function is still not completely elucidated. In multicellular organisms, the centromere is embedded within constitutive centric heterochromatin (Fig. 4). The outer kinetochore is responsible for attachment of spindle microtubuli, the transition from metaphase to anaphase and poleward movement. Additionally, it has a key role in binding proteins that keep the attachment of microtubuli to the kinetochore under a surveillance mechanism called the spindle assembly checkpoint (SAC) or the mitotic checkpoint (Shah and Cleveland, 2000). The inner kinetochore/centromere serves as an attachment site for kinetochore proteins and is important for centromere identity.
The mammalian centromere is composed of highly repetitive α-satellite DNA that consists of 171 bp monomer sequences (Willard, 1991). The proteins binding to the inner kinetochore can be divided into constitutive proteins, e.g. CENP-A, -B, -C and –G, and facultative proteins like tubulin, CENP-E and CENP-F (Van Hooser, et al., 1999). Whereas CENP-B binds specifically to a 17 bp α-satellite motif, CENP-A, -C and –G are located near/at the inner kinetochore plate. CENP-A is one of the best-studied centromeric proteins. It was originally identified as an essential 17 kD antigen recognized by anti-centromere autoantibodies from calcinosis/Raynaud’s phenomenon/esophageal dysmotility/ sclerodactyly/ telangiectasia (CREST) serum (Earnshaw and Rothfield, 1985). CENP-A is 62%
The organization of the centromere of S. cerevisiae is relatively simple compared to higher eukaryotes. It consists of 125 bp centromeric DNA sequences, which can be subdivided into three conserved regions named CDEI, CDEII and CDEIII (Hyman and Sorger, 1995). CDEI (8 bp) and CDEIII (25bp) are palindromic sequences flanking the A/T-rich 78-86 bp CDEII. Whereas CDEIII is absolutely necessary for centromeric function, mutations in CDEI and CDEII can be tolerated.
CDEIII contains the binding site for the essential four-protein complex CBF3 (Cbf2-Cep3-Ctf13-Skp1) (Connelly and Hieter, 1996); to CDEI binds a Cbf1 homodimer that increases the accuracy of chromosome segregation to the factor
Recently, (Sharp, et al., 2003) discovered, that the silencing protein Sir1 is a functional component of centromeric chromatin. This was the first time that a protein that functions in heterochromatic gene silencing at the HM loci was found to play a role at the centromere. Here, Sir1 functions in a partially overlapping pathway with CAF-I and Hir1 to maintain chromosome stability. It interacts with Cac1, the largest subunit of CAF-I and helps to maintain the chromatin assembly factor at the centromere.
Figure 4 illustrates the current model of the yeast centromere. Many protein-protein and DNA-protein interactions have been demonstrated by two-hybrid analysis, co-immunoprecipitation and chromatin-immunoprecipitation (ChIP) experiments, but it is expected that other still unknown factors might also play a role at the centromere.
Histone H3 variants are a common element found at functional centromeres in eukaryotes. Centromere specific histone H3 variants have first been described with CENP-A in mammals (Palmer, et al., 1987);(Sullivan, et al., 1994), but CENP-A homologues have also been found in D. melanogaster (Cid) (Henikoff, et al., 2000), S. pombe (Cnp1) (Takahashi, et al., 2000), C. albicans (CaCse4) (Sanyal and Carbon, 2002), C. elegans (HCP-3) (Buchwitz, et al., 1999) and in S. cerevisiae (Cse4) (Stoler, et al., 1995). They all share highly conserved C-terminal
The histone H3 variant CSE4 from S. cerevisiae was first identified in a screen for mutants with defects for c hromosome se gregation. Here, the mutant cse4-1 increases the frequency of chromosome non-disjunction and leads to a G2-M cell cycle arrest at elevated temperatures (Stoler, et al., 1995).
CSE4 encodes an essential 27 kD protein with a unique 135 aa N-terminus containing two putative nuclear localization signals (NLS), and a C-terminus with >64% homology to the histone-fold-domain of histone H3. The N-terminus is localized outside the centromeric nucleosome core and has at least one essential function, because deletion of the first 50 amino acids leads to cell death (Keith, et al., 1999). This region was further characterized by (Chen, et al., 2000), who delineated a 33 amino acid region (aa 28-60) termed END domain (essential N-terminal domain) by deletion mutagenesis, which is essential and sufficient for wildtype Cse4 function.
One function of the END domain is the interaction with the kinetochore protein Ctf19 (Chen, et al., 2000);(Ortiz, et al., 1999), which exists in a complex together with Okp1 and Mcm21. MCM21 is a suppressor of END domain mutations,
The C-terminus of Cse4 is highly homologous to the histone-fold-domain of histone H3. The histone-fold-domain consists of three helices (helix I-III) that are separated by two β-loop stands. In front of helix I an N-loop can be found and in case of histone H3 an additional N-terminal helix. Substitutions of helix II or helix III of Cse4 with analogous histone H3 amino acids are critical and often lead to increased chromosome loss and lethality. Replacing the histone-fold-domain of Cse4 with that of CENP-A is unable to rescue the cse4 null phenotype, so leading to the conclusion that specific domains within the histone-fold-domain are essential for the function of the yeast centromere (Keith, et al., 1999).
Both histone-histone interactions as well as histone-DNA interactions occur via the globular histone-fold domain, which was demonstrated in a genetic screen for Cse4 histone-fold-mutants. The current model suggests that Cse4 replaces both copies of histone H3 at the centromere and forms together with histone H4 centromere specific (Cse4-H4)2 tetrameres (Glowczewski, et al., 2000). Supporting this model, overexpression of Cse4 can suppress a temperature sensitive phenotype in a mutant histone H4-allel (hhf1-20), which leads to a G2-M cell cycle arrest and increased chromosome missegregation (Meluh, et al., 1998);(Smith, et al., 1996). Reciprocally, overexpressing histone H4 can repress centromere defects in cse4 mutants.
Histone H2A has also been described to have a function at the centromere (Pinto and Winston, 2000). hta1-mutants lead to a cell cycle delay in G2-M, display increased chromosome loss and show genetic interaction with mutations in genes
Cse4 competes against histone H3 for binding to histone H4. Normal core histones are transcribed exclusively during S-phase of the cell cycle from histone genes that have a special 3’ untranscribed region instead of poly(A) tails (Hereford, et al., 1981);(Osley, 1991). In contrast, Cse4 mRNA is polyadenylated and transcribed at low constant levels during the whole cell cycle. How the competition between Cse4 and H3 is regulated is still unknown, but overexpression of histone H3 in cse4 mutant cells is lethal (Glowczewski, et al., 2000).
So far no modifications have been found at the histone H3 variant Cse4, although acetylation and phosphorylation have been investigated within the END domain of Cse4 (Chen, et al., 2000). We found with two-hybrid analysis that Cse4 interacted with the histone acetyltransferase Sas2 and further analyzed if Cse4 is able to interact with other components of the SAS-I complex. Additionally, we investigated the role of Sas2 at the centromere and if the acetyltransferase is able to acetylate Cse4.
Strains between horizontal lines are isogenic
The Cse4 peptide used in this study was synthesized by Sigma Genosys. It consists of the N-terminal amino acids 112-134 with the following sequence (putative acetylation sites are marked in red):
1 mg/ml peptide was solubilized in dH2O and stored at -80°C.
The primers used in this study were synthesized by Metabion and applied in a dilution of 10 pmol/μl. Sequences derived from the Saccharomyces Genome Database (
The cultivation of bacteria with plasmids was carried out in LB-media or on LB-plates at 37°C unless indicated otherwise. For the bacterial expression of 3xHA-Cse4, nutrient rich TY-median was used. For the maintenance of plasmids the appropriate antibiotics were added to the media (ampicillin, kanamycin, chloramphenicol).
Chemically competent bacterial strains were obtained from Gibco BRL (DH5α) and Invitrogen (TOP10, BL21 (DE3) Star). The transformation was carried out as suggested by the manufacturer.
For the preparation of competent yeast cells and the transformation of DNA into yeast we used a method described from (Klebe, et al., 1983). Competent yeast cells were used immediately or were stored after the addition of DMSO to 5.5% at
-80°C.
The isolation of plasmids from bacteria was performed with the Mini- or Midi-Kit from QIAGEN according to the manufacturers instruction.
Genomic yeast DNA was isolated as described from (Hoffman and Winston, 1987). For PCR (polymerase chain reaction) mediated analysis of gene knock-outs in yeast, a single yeast colony was heated for one minute in a microwave. The PCR mix was added subsequently and the DNA amplified in a thermocycler.
The epitope tagged HA-CSE4 plasmid was constructed by inserting three HA-tags into the XbaI-site of CSE4 analogous to (Stoler, et al., 1995). For the insertion of the N- and C-terminal CSE4 fragments into the pACT2 and pBTM117c vector the amino acids 11-139 and the amino acids 137-229 were PCR- amplified with CSE4-primers containing specific restriction sites and subcloned into pCR®BluntII-TOPO (Invitrogen). For insertion into pACT2, the N-terminus was excised with PstI/SpeI and the C-terminus with SacI/PstI. The overlapping ends were filled in with T4 DNA-polymerase and the fragments were cloned into SmaI-linearized pACT2. To obtain the N-/C-terminal CSE4 in the LexA-vector, the fragments were excised with AccI/NotI and inserted into AccI/NotI-linearized pBTM117c.
To generate yeast strains with a specific genotype, the appropriate strains were crossed, sporulated and the resulting tetrads were dissected. For this purpose, small amounts of the yeast strains to be crossed were mixed in dH2O and were allowed to mate for ≥ 8 h. To select for diploid cells, the cells were then transferred onto supplemented YM and grown at 30°C. After two days, diploids were restreaked onto YPD before they were transferred to sporulation plates and incubated at 30°C for 3-4 days. The cell wall of the sporulated strain was digested with zymolyase solution for 10 minutes at room temperature; the reaction was stopped by the addition of 160 μl dH2O. The spores from the asci were separated under the microscope (Zeiss Axioskop FS) with a micromanipulator (Narishige) and analyzed for their genotype.
For specific gene knock-outs, we took advantage of homologous recombination. In most cases the kanMX-system was used, which lead after successful homologous recombination to geneticin (G418) resistance in S. cerevisiae.
Standard conditions:
PCR for subsequent cloning: repeat step 2-4 for 20-25 cycles
Control PCR: repeat step 2-4 for 25-30 cycles
Sequencing of DNA templates was performed with the ABI PRISMTM Dye Terminator Cycle Sequencing Ready Reaction Kit. For this purpose, 0.5-1 μl DNA was mixed with 4 μl Terminator Ready Reaction Mix, 1 μl Primer and dH2O and amplified in the thermocycler.
After amplification, the DNA was precipitated with Na-acetate/EtOH, dried and analyzed in the sequencing department of the MPIMG.
The Two-hybrid system can be used to detect protein-protein interactions in yeast. The method is based on the GAL4 gene from Saccharomyces cerevisiae, whose gene product activates several genes from the galactose synthesis pathway. For their activation, Gal4p binds with its DNA-binding domain (DNA-BD) to sequences upstream of the genes (UAS).
Additionally, Gal4p consists of an activation domain (AD), which binds via proteins to the DNA-binding domain and leads to transcription of the reporter genes (lacZ, HIS3). If the two domains remain separated from each other, the transcription from the reporter genes is blocked.
For our experiments we used the pACT2-vector with the activation domain and pBTM117c with the DNA-BD. pBTM117c contains instead of the Gal4 DNA-BD LexA, a bacterial protein that normally binds to lexA promotors. The advantage of LexA based two-hybrid is that less false positive interactions occur and that weak interactions are effectively amplified due to multiple LexA operators. The proteins to be tested were cloned into the vectors and both plasmids were
Upon activation of the reporter gene lacZ, the enzyme β-galactosidase is expressed in the cells. In this assay, the enzyme uses the colourless X-Gal (5-Brom-4-chlor-3-indolyl-β-D-galactopyranoside) as a substrate instead of galactose, and cleaves it into the dark blue 5-brom-4-chlor-indigo. For this purpose the yeast cells were transferred onto a nitrocellulose filter and broken in liquid nitrogen. The filter was incubated with 2,5 ml buffer Z and 50 μl X-gal for up to 2 h at 30°C. During this time course, a positive interaction between the proteins should lead to a blue coloration.
The reporter gene expression was also tested with the HIS3 reporter assay. The two-hybrid strain with the appropriate plasmids was streaked onto media lacking histidine. L40c itself is auxotroph for histidine, but due to the interaction of the proteins to be tested, HIS3 gene expression is activated, and the strain can grow in the absence of histidine.
To perform FACS analysis, yeast cells were cultured in an appropriate volume (~3 ml) until they reached OD600=0.05-0.1. After brief centrifugation and washing, cells were fixed in 70% EtOH at 4°C over night. Cells were then incubated for 4 h in 20x TE + 1 μg/ml RNase A, washed twice in PBS and subsequently stained over night in PBS + 100 μg/ml propidium-iodide (Sigma). After the suspension was 10x diluted, cell were separated from each other by sonication (3x 5 sec, 60%) and maintained in the dark. FACS analysis was performed with a flow cytometer (FACSCalibur) at the Deutsche Rheumaforschungszentrum in Berlin.
Protein extract preparation from bacteria
After reaching the desired optical density (OD), the bacterial culture was centrifuged for 10 minutes at 5000 rpm and the pellet resuspended in column buffer. The cells were broken with ultrasound (4 x 1 minute, 60 %) or with the french press. To obtain the protein extract, the broken cells were centrifuged for 30 minutes at 20.000 rpm and the supernatant was frozen at -80°C.
Protein extract preparation from yeast
Yeast cell cultures were grown to an OD600= 0,8-1 and harvested by centrifugation for 20 minutes at 5000 rpm at 4°C. After washing the cells with dH2O, the cell pellet was resuspended in the appropriate buffer.
Native whole cell extracts from yeast cultures <200 ml were prepared in 2x buffer L by glass bead lysis as described by (Moazed and Johnson, 1996), except that the concentration of NaCl was adjusted to 200 mM. For denatured protein extracts, the cells were lysed in Bead Buffer, boiled for 10 minutes at 95°C and diluted in IP dilution buffer for subsequent immunoprecipitation.
Larger cell cultures were resuspended in 2x buffer L or bead buffer, respectively, and the cells were broken using a french press. After 1 h centrifugation at 40.000 rpm, protein extracts were analyzed or frozen at -80°C for further use.
Proteins were analyzed according to their molecular weight with 8/10/12% SDS-PAGE gels. The transfer from the gel to a nitrocellulose membrane was performed either at 0,8 A/cm2 for 1 h with a Semi-Dry Blot from BIO-RAD, or at 110 V for 45 minutes with a wet blot from BIO-RAD. The efficiency of the transfer was visualized with Poinceau S dye. Subsequently, the membrane was blocked for 1 h with 5% fat free milk/TBST and incubated with the primary antibody in 5% fat free milk/TBST at 4°C overnight. After washing the membrane four times with TBST, the secondary HRP-conjugated antibody was added for 30 minutes. The membrane was washed with TBST for six times and proteins were then detected with ECL-solution from Amersham.
To detect proteins directly in SDS-PAGE gels, the gel was stained for 1 h with Coomassie Brilliant Blue R250. Subsequently, the gel was destained with 25% methanol/10% acetic acid, such that the staining of the proteins was maintained.
If small protein concentrations (~0,1 ng/mm2) were to be detected, we used the Silver Stain Kit from Bio-Rad. The staining of the proteins in an SDS-PAGE gel was performed according to the manufacturer’s instructions. Protein concentrations in solutions were determined with the Bradford Assay (Bio-Rad Protein Assay Kit).
To concentrate proteins in a smaller volume, Centricon-columns (amicon) with an exclusion matrix of 10 kD were used. After one hour centrifugation at 5000 g, protein solutions were concentrated from 2 ml to approximately 50-200 μl. If
Immunoprecipitation experiments were performed to isolate proteins and protein complexes. If the immunoprecipitation was carried out under denaturing conditions, the protein extract was boiled before use for 10 minutes at 95°C. The appropriate antibody was added to the whole cell protein extract and incubated for one hour with shaking at 4°C. Subsequently, protein G sepharose beads (Pharmacia) were added and the lysate-antibody-protein G mix was incubated over night.
Immunoprecipitates from native extracts were washed four times with 1x buffer L and resuspended in SDS sample buffer; precipitates from denatured extracts were washed with Urea wash buffer, IP buffer and detergent free wash buffer before resuspension in SDS sample buffer. After boiling for 10 min at 70°C and centrifugation, the immunoprecipitates were analyzed by immunoblotting.
For bacterial expression of large amounts of Cse4, N-terminal His-tagged Cse4 was generated by inserting CSE4 into the XhoI/BamHI-site of pET15b and transforming the resulting plasmid (pAE994) into BL21(DE3) Star cells. The expression of His-Cse4 in 2 l LB-culture was induced by adding 1 mM IPTG to the medium at OD600=0.8 and subsequent growth for additional 2h at 37°C. The cells were harvested and proteins extracted in Lysis Buffer by sonication (3x 20 sec, 40-50%). After centrifugation, the supernatant was added to 2 ml 50% Ni-NTA matrix and incubated with rotation for 1 h at 4°C. Proteins that did not bind the to matrix were washed off with 15 ml Wash Buffer.
To investigate the acetylation of proteins, we took advantage of an in vitro acetylation assay. In a total volume of 25 μl we mixed 2 μg recombinant histone H4 and/or 2 μg His-Cse4 together with the enzyme (200 μg rSAS-I or 500 μg recombinant PCAF), 5xHAT-Buffer and 0,5 μl [14C] acetyl-CoA (50-62 mCi/mmol Amersham). After one hour incubation at 30°C, the mix was run on a 15 % SDS-PAGE gel. The gel was dried in a gel dryer (BioRad) for 1 hour at 80°C, and the acetylated proteins were detected after over night exposure with a phosphoimager.
Sas2 belongs to the MYST family of HATs and acetylates lysine 16 on histone H4 (Meijsing and Ehrenhofer-Murray, 2001);(Suka, et al., 2002) and lysine 14 on histone H3 (Sutton, et al., 2003). To identify proteins that interact with Sas2, we performed a two-hybrid screen and identified a fragment of the histone H3 variant Cse4 (Table 1).
In order to test whether Cse4 and Sas2 also interact in vivo, we performed co-immunoprecipitation experiments (Fig. 7A). We constructed a 3x-HA-tagged version of CSE4 that was fully functional in that it rescued the lethality of a CSE4 deletion (data not shown). In an immunoprecipitation with a Sas2-antibody, we were able to detect HA-Cse4, whereas the antibody did not precipitate HA-Cse4 in the absence of Sas2.
Since Sas2 exists in the SAS-I complex together with Sas4 and Sas5 (Meijsing and Ehrenhofer-Murray, 2001), we asked whether Cse4 was able to interact with Sas4 and Sas5. Whereas Cse4 interacted via its N-terminus also with Sas4, we were unable to find two-hybrid interactions between Cse4 and Sas5. The binding of Cse4 to Sas4 in vivo was confirmed by co-immunoprecipitation. Interestingly, Sas5 could be precipitated in this assay (Fig. 7A). Thus, we suggest that Cse4 and Sas5 did not interact directly with each other, but that they can be found in the same complex in vivo. Possible mediators between these two proteins are Sas2 or Sas4.
As a member of the MYST family of HATs, Sas2 contains an acetyl-CoA-binding site (HAT) and an atypical zink-finger (C2HC motif), which is necessary for substrate recognition in the Sas2 homologue MOF in D. melanogaster (Clarke, et al., 1999). We tested the relevance of these motifs for the interaction with Cse4 (Table 1, Figure 7B). Both a mutation in the atypical zink-finger (C106L) and in the acetyl-CoA-binding site (P213A/P214V) resulted in a loss of the Cse4-Sas2 two-hybrid interaction, showing that both motifs were essential for Sas2 binding to Cse4.
However, the Sas2 mutants still co-immunoprecipitated with Cse4 (Fig. 7B), suggesting that the Cse4-Sas2 interaction was stabilized in vivo by other proteins. Alternatively, the folding of Sas2 in the two-hybrid may be partially compromised.
As Cac1 exists in the CAF-I complex together with Cac2 and Cac3 (Kaufman, et al., 1997), we asked whether Cse4 interacted with the whole CAF-I complex or
We found that the chromatin assembly factor Asf1 was also able to interact with Cse4. This interaction might be important at earlier steps in the assembly pathway, e.g. by delivering Cse4 to CAF-I, which in turn incorporates Cse4 at the centromere. As Cse4 is able to interact with subunits of the Asf1 and CAF-I histone deposition factors, there may exist different steps or mechanisms in assembling Cse4 into chromatin. In contrast to cacΔhirΔ cells, cacΔasf1Δ cells showed a G2/M delay independent of spindle assembly checkpoint and segregated a reporter minichromosome at wildtype frequencies. This result indicates, that the centromere needs Hir1, but not Asf1 in the absence of CAF-I function, so that the HIR/ASF1 chromatin assembly pathway might be bifurcated at some point.
Since Cse4 interacted with Cac1 as well as with SAS-I, we next sought to determine the dependence of these interactions on SAS-I components and Cac1, respectively. As can be seen in Figure 9A, a deletion of SAS2 did not disturb the interaction between Cse4 and Sas4 or Cac1, respectively. This suggested that the interaction between Cse4 and Sas4 or Cac1 was stable and that the presence of Sas2 was not necessary for maintaining the association between Cse4, Sas4 and Cac1.
In contrast, the chromatin assembly factor Cac1 was not essential to maintain the interaction between Cse4 and the SAS-I complex (Fig. 8C). Sas2 as well as Sas4 were still associated with Cse4 in a cac1Δ strain, showing that the interaction between Cse4 and SAS-I was not affected by the presence or absence of Cac1.
The results from the two-hybrid assay and the co-immunoprecipitation experiments indicated, that Sas2 might indeed have a role at the centromere. As Sas2 is a histone acetyltransferase that acetylates histone H3 K14 and histone H4 K16, we hypothesized that the histone acetylation might be important for the centromere as well. One possibility is that Sas2 interacts with the histone H3 variant Cse4 because it is an additional acetylation target. We next asked, if a sas2 deletion, and therefore a missing acetylation, had any effect in combination with cse4 mutations at the centromere. As a CSE4 deletion is lethal, we used a temperature sensitive mutant for the subsequent experiment.
The cse4-103 strain, which has two amino acid exchanges in the histone-fold-domain (I156V, L193Q), was generated by random mutagenesis. This temperature sensitivity is due to the weakened interaction between cse4-103 and histone H4, because the L193Q substitution is predicted to disrupt the Cse4-H4 interface (Glowczewski, et al., 2000). We inserted both cac1 and sas2 single deletions as well as cac1 sas2 double deletions into the cse4-103-strain and investigated the temperature sensitivity at 23°C, 34°C and 37°C. At the permissive temperature all strains grew equally well, whereas at 37°C all cse4-103 mutants were unable to survive. At 34°C, the cse4-103 mutant strain grew poorly compared to wildtype (Fig. 10).
When SAS2 was deleted, the cse4-103 strain grew better at 34°C as compared to the single cse4-103 mutation. Perhaps a missing acetylation on histones H3, H4 or Cse4 bySas2 stabilizes the structure and function of the centromere. As Cse4 together with histone H4 forms stable (Cse4-H4)2 tetrameres, one possible explanation is that the target of acetylation was histone H4, because the acetylation of free histone H4 at lysine 16 has previously been described (Sutton, et al., 2003). One hypothesis is that the missing acetylation on histone H4 lysine
Thus, the cse4-103 sas2Δ strain is still temperature sensitive, but not to the same extent as the single cse4-103 mutation. Another possibility with a similar result would be if Cse4 also was a target for acetylation by Sas2.
Drugs like benomyl or nocodazole affect the correct formation of microtubuli. When a yeast strain has an additional mutation in a gene that is important for the correct formation of the centromere-kinetochore complex, it becomes sensitive and is unable to grow on plates with benomyl or nocodazole. We investigated if a deletion of ASF1 or components of the SAS-I and CAF-I complex were benomyl or nocodazole sensitive, which might point to a role during centromere-kinetochore formation. None of the investigated deletions resulted in an increased or decreased benomyl sensitivity. We deduced from our data that all examined genes were dispensable for the correct formation of the centromere-kinetochore complex.
(Sharp, et al., 2003) found that a deletion of SIR1 lead to a greater resistance to the microtubule-depolymerizing drug benomyl in a point mutated cse4-107 background. cse4-107 contains a single point mutation at L175F, and is predicted to be in close association with the histone fold domain of histone H4 (Glowczewski, et al., 2000). We were now interested whether a SAS2 deletion also had an effect on benomyl sensitivity in combination with a cse4-103 mutation, which is also thought to disrupt binding to histone H4. However, benomyl sensitivity was neither increased nor decreased. In light of our results
Next to components of the SAS-I complex, Cse4 additionally interacted in co-immunoprecipitation experiments with chromatin assembly factors. Chromatin assembly factors like CAF-I and Asf1, but also associated proteins like Hir1 have distinct roles in the cell cycle. Apart from their functions in heterochromatic gene silencing, they also display various cell cycle dependent phenotypes.
A single deletion of cac1 or hir1 does not have any effects on cell cycle progression as can be seen in their FACS profiles or in assays monitoring chromosome loss (Sharp, et al., 2002). However, if yeast cells have an asf1 deletion or a cac1 hir1 double deletion, respectively, are delayed in G2-M phase in the cell cycle (Sharp, et al., 2002);(Tyler, et al., 1999) and have an increased rate of chromosome loss compared to wildtype cells (Le, et al., 1997);(Sharp, et al., 2002).
We were interested in examining the effect of a sas2Δ on cell cycle progression. In order to investigate if a deletion causes a delay in the cell cycle, we performed FACS analysis with wildtype, cac1Δ, hir1Δ, sas2Δ, asf1Δ, cac1Δhir1Δ, hir1Δsas2Δ, and asf1Δsas2Δ cells. As expected, cac1Δhir1Δ as well as asf1Δ cells displayed a G2-M arrest in the cell cycle. In contrast, neither a single sas2 deletion nor a sas2 hir1 double deletion caused any effect. When Sas2 and Asf1 were both missing, the cells displayed the same phenotypes as an asf1 deletion alone (data not shown).
Another method to investigate defects within the cell cycle is to monitor loss of plasmids or chromosomes. (Stoler, et al., 1995) constructed a strain where chromosome loss could be monitored. This strain is disomic for chromosome III and contains several markers on the additional cen 130-3 mutated chromosome. Loss of the original or the additional chromosome III could be followed by
Another method to investigate correct segregation is to monitor plasmid loss. In order to do so, we transformed sas2Δ, hir1Δ, cac1Δ single, double and triple mutated strains with a plasmid carrying SUP11 as a marker (pUN90). We were able to verify the elevated chromosomal instability in cac1Δhir1Δ cells that has already been reported by (Sharp, et al., 2002). A single SAS2 deletion or sas2Δ in combination with cac1Δ and hir1Δ did not cause a further increase in plasmid loss. In summary we concluded that a single or an additional SAS2 deletion had no detectable influence on cell cycle progression or chromosome and plasmid stability under the conditions tested here.
Ctf19 is a Cse4-interacting protein at the centromere that mediates the connection between CDEI and CDEIII in a complex with Okp1 and Mcm21 (Ortiz, et al., 1999). Ctf19 is nonessential, and mutations result in chromosome missegregation, increased benomyl sensitivity and accumulation in the G2/M phase of the cell cycle (Grienenberger, et al., 2002). Since Ctf19 and Cse4 interact by two-hybrid analysis, we asked whether a SAS2-deletion had an effect on their association.
In contrast to the wildtype situation, where Ctf19 and Cse4 interacted with each other, no association was found when SAS2 was deleted (Fig.11). This suggested that Sas2 had a role at the centromere, e.g. by stabilizing the cohesion between the two centromeric proteins Ctf19 and Cse4 via putative acetylation of Cse4.
As Sas2 is a histone acetyltransferase for histone H4 K16 and histone H3 K14 (Meijsing and Ehrenhofer-Murray, 2001);(Osada, et al., 2001);(Sutton, et al., 2003) and additionally interacts with the histone H3 variant Cse4, we asked whether Sas2 was also able to acetylate Cse4 in vivo. To determine the putative acetylation, we analyzed denatured protein extracts from wildtype and sas2Δ strains with α-acetyl-lysine-antibodies. If Sas2 acetylated Cse4, we expected to be able to precipitate Cse4 in the wildtype protein extract but less or none in the sas2Δ protein extract. It was important to denature the proteins, because otherwise Cse4 might be precipitated in a complex together with histone H4, which would lead to a signal due to histone H4’s acetylation pattern.
Furthermore, to reduce the complexity of the α-acetyl-lysine precipitation, we isolated the protein extracts and performed a first immunoprecipitation with α-HA-antibody to isolate Cse4 from the extract. This additional immunoprecipitation step was necessary to partially purify Cse4 for the second immunoprecipitation with α-acetyl-lysine-antibody. After washing off the other proteins, we removed the bound Cse4 with Bead Buffer containing 1 % SDS that was then diluted to 0.1 % SDS for the subsequent second immunoprecipitation
As shown in Figure 13, the amount of HA-Cse4 applied was comparable in both wildtype and sas2Δ protein extracts (Input). After immunoprecipitation with α-acetyl-lysine-antibody, HA-Cse4 could be detected in wildtype and sas2Δ extracts, whereas the control without antibody (-Ab) did not show a HA-Cse4 specific signal. We concluded that an acetylated Cse4 form existed in the cell, although differences in the intensity of the signal from wildtype and sas2Δ extracts could not be detected. As the signal varied in independent experiments, we cannot rule out the possibility that Cse4 might only be temporarily acetylated during the cell cycle, so that the acetyl group is removed at some point. Furthermore, an associated histone deacetylase might be co-purified in the assay. In this case, the acetyl-group from acetylated Cse4 could be removed during the purification, so that in consequence we were unable to detect Cse4. Additionally, it might still be possible that after removing the bound HA-Cse4 with 1% SDS from the α-HA-antibody, protein complexes containing Cse4 and histone H4 are restored upon dilution of SDS to 0,1%. In this scenario, the precipitation of HA-Cse4 with the α-acetyl-lysine-antibody may be via acetylated histone H4.
Because it was difficult to determine if Cse4 is acetylated in the cell with immunoprecipitation experiments, we also took advantage of another independent method to determine a putative Cse4 acetylation via Sas2: an in vitro acetylation assay. For this purpose, we used the bacterially expressed and purified SAS-I-complex (pAS134, plasmid and purification as described by (Sutton, et al., 2003) and Cse4. The advantage of this method was that large quantities of purified proteins could be purified from bacteria. On the other hand, co-factors or other modifications that are important for the reaction might be missing.
As can be seen in Figure 13, PCAF as well as the SAS-I complex were functional and able to acetylate recombinant histone H4 (lane 1+2). Additionally, self-acetylation of components from the SAS-I complex could be detected in lane 2. When His-Cse4 was added to PCAF in the assay (lane3), no Cse4-specific acetylation signal could be detected. The signals obtained in lanes 3 and 4 could
The main question of this assay was, whether the SAS-I complex was able to acetylate the histone H3 variant Cse4. Thus, we incubated SAS-I together with purified His-Cse4, but we were unable to detect acetylated His-Cse4 with the applied concentrations, even when self-acetylation of the SAS-I complex could be seen (lane5). The addition of histone H4 to the sample lead to its acetylation, but acetylated His-Cse4 was still not detectable (lane 6).
Additionally to the purified His-Cse4 from bacteria, we used a Cse4 peptide for our in vitro acetylation assays. As Cse4 contains several putative acetylation sites (9 lysines in the N-terminus, 7 lysines in the C-terminal histone fold domain), we chose a lysine rich region (aa 112-134 with K115, 119, 122, 126, 130, 131) within the N-terminus for this purpose. As Cse4 interacted via its N-terminus with the acetyltransferase Sas2, we hypothesized to find a putative acetylation site in this region, but we were unable to find a Cse4 specific acetylation signal with SAS-I in the in vitro assay (data not shown).
Cse4 is an essential, evolutionarily conserved core component of the centromere. Homologues have been found in S. pombe, C. elegans, D. melanogaster and mammals. In all organisms, Cse4 and its homologues replace the histone H3 at the centromere by building specific (Cse4-H4)2 tetrameres.
In this work we were able to demonstrate that the histone H3 variant Cse4 was able to interact with subunits of the SAS-I complex as well as with chromatin assembly factors. All interactions could be shown by two-hybrid analysis and co-immunoprecipitation experiments and occurred mainly over the unique 135 aa N-terminus of Cse4. The absence of single proteins did not result in destabilizing the complex, except for the Cac1-Cse4 interaction in a sas4Δ-strain. The biochemical interaction between Cse4 and Sas2 could further be confirmed with genetic interactions. We observed that the temperature sensitivity of the cse4-103 mutant at 34°C was partially suppressed by an additional sas2 deletion. Sas2 was also shown to have a direct role at the centromere in that its deletion disrupted the interaction between Cse4 and Ctf19. Additionally, we were able to show that Cse4 existed in an acetylated state in the cell, but if Cse4 was an acetylation target for the acetyltransferase Sas2 could not be verified.
In this study we were able to show with two-hybrid experiments that the histone H3 homologue Cse4 interacted with the acetyltransferase Sas2. We further narrowed the Sas2-interacting region of Cse4 down a region within the N-terminus (amino acids 11-139). Therefore, we propose that Sas2 interacted with a
As Sas2 is a member of the MYST-family of HATs, it contains an acetyl-CoA-binding site (HAT) and an atypical zink finger (C2HC motif). Both motifs are essential for the acetylation activity of Sas2. We tested if the interaction between Cse4 and Sas2 was still intact when the HAT-domain or the atypical zink finger, respectively, was mutated. Clearly, the interaction between Cse4 and the point-mutated Sas2 was disturbed, so that we concluded that both motifs were important for the binding of Sas2 to Cse4. As the zink-finger is absolutely required for substrate recognition in the Drosophila Sas2-homologue MOF (Akhtar and Becker, 2001), one way of explanation could be that Cse4 was an acetylation target for Sas2.
Since Sas2 functions in a complex together with Sas4 and Sas5 as a histone acetyltransferase, we hypothesized that Cse4 also binds to Sas4 and Sas5. Indeed we could verify an interaction over the N-terminal tail of Cse4 and Sas4, whereas no binding to Sas5 was detected. From our two-hybrid data we concluded that Cse4 binds via its N-terminus to two subunits of the SAS-I complex, Sas2 and Sas4.
All two-hybrid interactions were further confirmed with co-immunoprecipitation experiments in vivo. Interestingly, we were also able to precipitate Sas2-HAT-, Sas2-Zn- and Sas5 with Cse4, although no direct interaction was detected with the two-hybrid assay. However, as Cse4 interacted with two subunits from the SAS-I complex, Sas2 and Sas4, the interaction with the point mutated Sas2 as well as with Sas5 could be bridged by them. In this case a positive precipitation could be found even if the proteins did not bind directly to each other.
Two other proteins could also be able to bridge these interactions – Cac1, the largest subunit of the chromatin assembly factor CAF-I, and Asf1. CAF-I and Asf1 are histone chaperones that deliver histone H3 and histone H4 to DNA after
Both proteins, Cac1 and Asf1, have been shown to interact with the SAS-I complex (Meijsing and Ehrenhofer-Murray, 2001);(Osada, et al., 2001), and we found with two-hybrid and co-immunoprecipitation experiments that they also bind to Cse4. The interaction with Cac1 could further be narrowed down to the amino acids 94-137. Although Cac1 exists in a complex, no interaction could be detected with Cac2 or Cac3, which argues for a function of Cac1 separate of CAF-I. Recently, (Sharp, et al., 2002) reported that CAF-I and Hir1 have a role in maintaining the centromeric chromatin structure. They are not absolutely required for Cse4 deposition, but their absence leads to additional deposition outside of the centromere.
Since Cac1 as well as Asf1 are both chromatin assembly factors that bind histones H3 and H4, our findings together with the results from (Sharp, et al., 2002) support a model where Cac1 and Asf1 were also responsible for delivering the histone H3 variant Cse4 to the centromere. Since CAF-I and Hir1 are not essential for recruiting Cse4 to the centromere (Sharp, et al., 2002), our results could point out an important role for the nucleosome assembly factor Asf1. In Figure 14 all interactions between Cse4, SAS-I and the chromatin assembly factors are summarized.
In our experiments we further found that neither deletion of sas2, sas4 or cac1 affected the association of Cse4 with components of the SAS-I complex or Cac1. One exception was found in a sas4Δ strain, where Cac1 and Cse4 were unable to bind to each other. If sas4 is deleted, the whole SAS-I complex is disturbed, because Sas5 binds only to Sas4 and not to Sas2. In light of our results, one
This model would include that Cac1 is only able to bind to the complex in the presence of the active SAS-I complex. Sas4 has been shown to be essential for histone acetyltransferase activity for Sas2 (Sutton, et al., 2003). If SAS4 is deleted, the SAS-I complex is both incomplete and inactive. In this case, Sas2 would be unable to acetylate its target, probably the histone H3 homologue Cse4 or histone H4, which would then prevent binding of Cac1 to the complex. This hypothesis would further include that Cac1 is also unable to bind to Sas2, which remains to be tested.
(Meijsing and Ehrenhofer-Murray, 2001) have reported that Sas2, Sas4 and Sas5 coeluted in gel filtration experiments in a peak of ~220 kD, even if the calculated molecular masses of myc-Sas2, myc-Sas4 and Sas5 add up to ~140 kD. Additionally, (Osada, et al., 2001) detected by Superose 6-size exclusion chromatography and a subsequent western blot a 450 kD Sas2 containing
The histone acetyltransferase Sas2 functions in the SAS-I complex together with Sas4 and Sas5 and acetylates histone H3 K14 and histone H4 K16 (Meijsing and Ehrenhofer-Murray, 2001);(Sutton, et al., 2003). Additionally, Sas2 has a role in heterochromatic gene silencing at the HM loci and at the telomeres (Ehrenhofer-Murray, et al., 1997);(Reifsnyder, et al., 1996).
Does Sas2 also have a role at the centromere ? A direct function for Sas2 at the centromere has not been described so far, but (Sharp, et al., 2003) discovered that the silencing protein Sir1 is a functional component of centromeric chromatin and helps to maintain CAF-I at the centromere. As Sas2 genetically interacts with Sir1 in silencing at HML (Ehrenhofer-Murray, et al., 1997), an additional function at the centromere might be possible. Furthermore, our two-
In addition to our findings that the SAS-I complex was associated with Cse4, we found that a SAS2 deletion leads to better growth in the temperature-sensitive cse4-103 strain at 34°C. The cse4-103 strain has a point mutation in two amino acids within the histone-fold-domain (I156V, L193Q). This mutation is thought to affect the interaction with the globular domain of histone H4. In consequence, the (cse4-103/H4)2 tetrameres are destabilized at elevated temperatures and the cells are unable to survive.
How can a SAS2 deletion have a positive influence on the stability of (cse4-103/H4)2 tetrameres ? It is known that Sas2 acetylates histone H4 at lysine 16, which is also present at the centromere in contrast to histone H3. Histone acetylation has mainly been found in the neighborhood of transcriptionally active promotors and enhancers (Kuo and Allis, 1998), where they are thought to restrict the folding of nucleosomes into the condensed 30 nm fiber (Garcia-Ramirez, et al., 1995);(Tse, et al., 1998). As the yeast centromere is not packed into heterochromatin like in other organisms, one might hypothezise that the histone tails exist in an acetylated state. In SAS2 wildtype cells, this acetylation may further destabilize the already weak interaction between cse4-103 and histone H4, so that in consequence the cells are unable to grow at elevated temperatures. If the histone acetyltransferase Sas2 is absent, the acetylation of histone H4 K16 is missing and the nucleosomes would be able to form a more compact structure that in turn helps to tighten the (cse4-103/H4)2 tetrameres. Another possibility could be that Sas2 also acetylates Cse4 next to histone H4. A missing acetylation on Cse4 would therefore have the same consequences on the stability as on histone H4 in a sas2Δ strain.
How can Sas2 influence the interaction between Cse4 and Ctf19 ? Post-translational modifications of histones have already been shown to be critical for protein-protein interactions. One example are the silent information regulator (SIR) proteins Sir3 and Sir4 that interact with deacetylated tails of histone H3 and histone H4 in order to build up a repressive chromatin state. An explanation for our observation could be that Sas2 acetylates one or more lysine residues on the N-terminus of Cse4. Thereby, the acetylation may provoke a conformational change in the (Cse4-H4)2 tetrameres that are now able to interact with other proteins, e.g. Ctf19. In a sas2Δ cell, the N-terminus of Cse4 may remain more unflexible due to the missing acetylation so that the accessibility for other interacting proteins is decreased. In that case, the interaction between Cse4 and Ctf19 would be disrupted as can be seen in a two-hybrid assay.
Further analysis with co-immunoprecipitation experiments need to be carried out to confirm this hypothesis. Additionally, chromatin immunoprecipitation or indirect immunofluorescence experiments with Sas2 could be used to test whether it can be found at some point of the cell cycle at the centromere.
We were interested in exploring the possibility that Cse4 exists in an acetylated state in the cell and if the histone acetyltransferase Sas2 is a Cse4-modifying
The N-termini of the core histones H2A, H2B, H3 and H4 are post-translationally modified by methylation, acetylation and phosphorylation. These modifications are essential for modulating nucleosome structure and therefore changing gene activity. Little is known about modification of histones and histone variants at the centromere, but it has been reported that histone H3 phosphorylation in mammals precedes the phosphorylation of the Cse4-homologue CENP-A in prophase (Zeitlin, et al., 2001). This phosphorylation is necessary for directing the subcellular localization of enzymes required for completion of cytokinesis.
So far no modifications have been shown for the histone H3 variant Cse4, although they may occur because its homologue in mammals is phosphorylated. We were now able to show that Cse4 is acetylated at least at one stage in the cell cycle. This acetylation could be a specific mark for chromatin assembly factors like CAF-I and Asf1. In consequence, Cse4 is exclusively delivered to the centromere, where it replaces histone H3. One putative acetyltransferase involved in Cse4 modification could be Sas2, although no direct activity was detected. An explanation for the missing acetylation activity in our in vitro assay could be that the Cse4 concentration was too low. In that case, the radioactive signal of [14C] may be too weak to be detected. Another possibility could be that additional cofactors or pre-existing modifications at the N-terminus are needed for Cse4 acetylation. It has already been described for the histone H3 N-terminus that modifications are able to influence each other. One example is that histone H3 K4 methylation by Set7 inhibits methylation of lysine 9 by Su(var)3-9, but promotes acetylation of histone H3 by p300 (Wang, et al., 2001).
First, Cse4 and histone H3 have a highly homologous C-terminal histone-fold domain, but their N-terminus is distinctly different. The 135 aa N-terminus of Cse4 shows no homology to known proteins and contains at least one essential function, since the deletion of the END domain (amino acids 28-60) is lethal to the cell (Chen, et al., 2000). The N-terminus of Cse4 may extend from the core and is able to interact with other proteins, so that in consequence chromatin assembly factors could distinguish between the standard core (H3-H4)2 tetramere and centromere specific (Cse4-H4)2 tetrameres. Another mechanism for helping the chromatin assembly factors to separate H3-H4 assembly from Cse4-H4 assembly is to divide their assembly in time. Whereas the standard core histones are exclusively expressed in early S-phase, Cse4 mRNA can be found at low levels throughout the whole cell cycle. Thus, the equilibrium is shifted towards Cse4-H4 after early S-phase, which may help to deposit them after H3-H4 deposition took place.
In this model, the histone acetyltransferase complex SAS-I may first bind to free Cse4 and specifically modifies one or more lysine residues on its N-terminus. The nucleosome assembly factor Asf1 is a histone chaperone that delivers histones H3 and H4 to the replication fork. Asf1 binds to the SAS-I complex via Sas4 (Meijsing and Ehrenhofer-Murray, 2001);(Sutton, et al., 2001) and we were able to show that Asf1 also interacted with the histone H3 variant Cse4. Thus, Asf1 might bind to the (Cse4-H4)2 tetrameres after identification via the N-terminus of Cse4. Here, the specific Sas2-dependent acetylation may already have an important role for recognizing the specificity for centromere deposition. (Sharp, et al., 2002) reported that Asf1 didn’t play a role at the centromere, because cacΔasf1Δ cells segregated a CFIII(D8B.d) reporter minichromosome at wildtype frequencies and displayed a G2-M delay independent of spindle assembly checkpoint activation. In contrast to their observation, we found that a single asf1Δ already resulted in a significant increase of loss of an additional chromosome III (data not shown). Thus, the conclusion that the centromeres did not require Asf1 requires re-investigation.
In a subsequent step, the whole complex may be delivered to the centromere. Here, the chromatin assembly factor CAF-I may still be bound to the DNA after replication took place. Asf1 interacts with Cac2 (Tyler, et al., 2001) and may deliver the (Cse4-H4)2 tetrameres to CAF-I that binds the histone H3 variant Cse4 via the Cac1 subunit, and histone H4 via Cac3 (Verreault, et al., 1996).
In this model, CAF-I releases Asf1 as well as SAS-I and now deposits Cse4 and histone H4 into centromere-specific nucleosomes and therefore replaces pre-assembled (H3-H4)2 tetrameres. The acetylation of Cse4 may further stabilize the centromere by mobilizing the N-terminus that can now interact with other centromeric protein, e.g. Ctf19. Of course, further studies are necessary to prove this model and also to elucidate the role of the nucleosome assembly factor Asf1 as well as the dependence of the Cse4-Ctf19 interaction on the histone acetyltransferase Sas2.
Education
October 2000 Diploma (Masters degree) summa cum laude in Biology, Humboldt University of Berlin
1995-2000 Undergraduate Education in Biology at Justus-Liebig University of Gießen, Technical University of Dresden and Humboldt University of Berlin
19994-1996 Study of Pedagogic at the Justus-Liebig University of Gießen
1994 High School Altkoenigschule Kronberg
Work Experience
November 2000 to present: Ph.D. student at the Max-Planck Institute of Molecular Genetics, OWL, AG Ehrenhofer-Murray. Project: “Connecting the histone acetyltransferase complex SAS-I to the centromere in S. cerevisiae”
May-October 2000: Diploma thesis at the Max-Planck Institute of Molecular Genetics, OWL, AG Ehrenhofer-Murray. Project: “Interaction of the centromeric histone H3-homologue Cse4 with the acetyltransferase Sas2 and the chromatin assembly factor CAF-I in S. cerevisiae”
July-December 1999: Practical course at the Max-Planck Institute of Molecular Genetics, OWL, AG Ehrenhofer-Murray. Project: “A screen for high-copy suppressors of silencing defects at the silent mating-type locus HML of S. cerevisiae”
1998 Supervising and teaching undergraduates in a practical course in zoology, Technical University of Dresden
Courses
July/August 2001: C. elegans course at Cold Spring Harbor Laboratory, USA
2003: Scientific Writing
Publications
Kasulke D, Seitz S, Ehrenhofer-Murray AE., Genetics. 2002 Aug;161(4):1411-23, A role for the Saccharomyces cerevisiae RENT complex protein Net1 in HMR silencing.
An erster Stelle möchte ich Ann für die Betreuung und Unterstützung während der Doktorarbeit, sowie generell der letzten 4,5 Jahre danken. Ich hatte die Möglichkeit selbständig zu arbeiten und dadurch auch viel zu lernen, was sicherlich nicht selbstverständlich ist.
Prof. Saumweber danke ich für seine Bereitschaft mich von der HU Berlin aus zu betreuen, sowie für seine erneute Gutachter-Tätigkeit. Prof. Francis Stewart möchte ich ebenfalls für seine spontane Zusage als Gutachter danken.
Meinen lieben (Ex-) Kollegen aus Labor 005 möchte ich gerne für eine sehr lustige und schöne Zeit danken. Vor allem Jacqueline, Horst und Anne, mit denen ich sehr viel Spaß hatte und auf deren Hilfsbereitschaft ich immer zählen konnte. Ich hoffe, dass wir einige Traditionen weiterführen können.
Für die hervorragende Unterstützung bei den Experimenten meiner Arbeit danke ich ganz besonders Uta, Jacqueline und Anne. Ich danke natürlich auch allen anderen (Ex-) Kollegen, es wäre eine sehr lange Liste, wenn ich Euch alle aufführen würde. Ich sag es einfach mal mit einem Satz: hoffentlich habe ich bei meiner nächsten Stelle ähnlich viel Glück. Wenn ich könnte, würde ich Euch alle mitnehmen! :)
Meinen Eltern und Caroline danke ich für Ihre Unterstützung in allen Bereichen.
Von ganzem Herzen danke ich Daniel – einfach dafür, dass er immer für mich da ist. Und natürlich auch Marlin, der durch seine langen Schlafphasen das Schreiben der Arbeit unglaublich erleichtert hat. In den Schreibpausen hingegen konnte ich mit ihm die Welt neu entdecken...
Ich erkläre hiermit, dass ich die vorliegende Arbeit selbst und nur unter Verwendung der angegebenen Quellen und Hilfsmitteln verfasst habe. Der Inhalt der Promotionsordnung der Mathematisch-Naturwissenschaftlichen Fakultät I ist mir bekannt.
Stefanie Seitz