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KTI11 and KTI13, Saccharomyces cerevisiae genes controlling sensitivity to G1 arrest induced by Kluyveromyces lactis zymocin
Article first published online: 7 MAY 2002
Volume 44, Issue 3, pages 865–875, May 2002
How to Cite
Fichtner, L. and Schaffrath, R. (2002), KTI11 and KTI13, Saccharomyces cerevisiae genes controlling sensitivity to G1 arrest induced by Kluyveromyces lactis zymocin. Molecular Microbiology, 44: 865–875. doi: 10.1046/j.1365-2958.2002.02928.x
- Issue published online: 7 MAY 2002
- Article first published online: 7 MAY 2002
The Kluyveromyces lactis zymocin and its γ-toxin subunit inhibit cell cycle progression of Saccharomyces cerevisiae. To identify S. cerevisiae genes conferring zymocin sensitivity, we complemented the unclassified zymocin-resistant kti11 and kti13 mutations using a single-copy yeast library. Thus, we identified yeast open reading frames (ORFs) YBL071w-A and YAL020c/ATS1 as KTI11 and KTI13 respectively. Disruption of KTI11 and KTI13 results in the complex tot phenotype observed for the γ-toxin target site mutants, tot1–7, and includes zymocin resistance, thermosensitivity, hypersensitivity to drugs and slow growth. Both loci, KTI11 and KTI13, are actively transcribed protein-encoding genes as determined by reverse transcriptase–polymerase chain reaction (RT–PCR) and in vivo HA epitope tagging. Kti11p is highly conserved from yeast to man, and Kti13p/Ats1p is related to yeast Prp20p and mammalian RCC1, components of the Ran–GTP/GDP cycle. Combining disruptions in KTI11 or KTI13 with a deletion in TOT3/ELP3 coding for the RNA polymerase II (RNAPII) Elongator histone acetyltransferase (HAT) yielded synthetic effects on slow growth phenotype expression. This suggests genetic interaction and possibly links KTI11 and KTI13 to Elongator function.
The Kluyveromyces lactis zymocin, a plasmid-encoded three-subunit (αβγ) toxin complex, inhibits proliferation of Saccharomyces cerevisiae by a G1 cell cycle arrest (Stark et al., 1990; Butler et al., 1991a; Schaffrath and Breunig, 2000; Meinhardt and Schaffrath, 2001). Conditional expression of its γ-subunit (or γ-toxin) is sufficient to mimic this arrest (Butler et al., 1991b). The α- and β-subunits are likely to be required for zymocin docking, a process involving interaction with cell wall chitin (Butler et al., 1991c; Jablonowski et al., 2001a).
Zymocin-resistant S. cerevisiae mutants have been isolated (skt: Kawamoto et al., 1990; kti: Butler et al., 1994; iki: Kishida et al., 1996; tot: Frohloff et al., 2001). Ten distinct kti class II complementation groups suggest that a number of proteins participate in the process blocked by zymocin by either acting in a pathway or forming a multiprotein target (Butler et al., 1994). In favour of the latter, analysis of the tot mutants identified the six-subunit RNA polymerase II (RNAPII) Elongator as TOT, the putative γ-toxin target complex (Frohloff et al., 2001; Jablonowski et al., 2001b; Winkler et al., 2001). Mutations in the TOT genes, six of which are identical to Elongator genes ELP1–6, lead to zymocin resistance (Yajima et al., 1997; Otero et al., 1999; Wittschieben et al., 1999; Fellows et al., 2000; Frohloff et al., 2001; Jablonowski et al., 2001b; Krogan and Greenblatt, 2001; Li et al., 2001; Winkler et al., 2001). The histone acetyltransferase (HAT) activity encoded by ELP3 (TOT3) is essential for both Elongator function in vivo and K. lactisγ-toxin action (Wittschieben et al., 2000; Frohloff et al., 2001; Winkler et al., 2001). TOT3 mutagenesis revealed that zymocin sensitivity can be uncoupled from Elongator function, indicating that TOT interacts genetically with zymocin in a manner dependent on Elongator’s HAT (Jablonowski et al., 2001b). Thus, TOT/Elongator appears to be the key player in K. lactis zymocicity. Cells lacking the Sit4p phosphatase (Sutton et al., 1991) display zymocin resistance and all other tot phenotypes, suggesting that TOT function may be a Sit4p-dependent process (Jablonowski et al., 2001c). As Elongator is dispensable for life and the γ-toxin arrest is TOT dependent, the K. lactis zymocin cannot simply block TOT/Elongator function. Instead, zymocin may rather modify Elongator function, so that RNAPII activity becomes poisoned (Frohloff et al., 2001; Jablonowski et al., 2001b).
To study the relationship between uncharacterized KTI genes and TOT/Elongator function, we identified KTI11 (YBL071w-A) and KTI13 (YAL020c/ATS1). KTI11 and KTI13 deletions confer a tot phenotype in common with tot1–7Δ cells including zymocin resistance, slow growth, thermosensitivity and hypersensitivity to drugs. Reverse transcriptase–polymerase chain reaction (RT–PCR) and in vivo epitope tagging identify both KTI11 and KTI13 as transcribed protein-encoding genes. Kti11p is conserved from yeast to man, and Kti13p is related to RCC1, a component of the Ran–GTP/GDP cycle. Combining tot3Δ with either kti11Δ or kti13Δ affects slow growth phenotype expression, implying interaction between the KTI and the TOT genes.
Genetic complementation of the kti11 and kti13 mutations
Inspired by our findings that tot mutants are hypersensitive towards caffeine, we checked the kti11 and kti13 mutants and observed that their growth was significantly inhibited by caffeine (Fig. 1). We next set out a genetic complementation approach involving the yeast library YPH1 (CEN4/LEU2) and selecting for Leu+kti11 and kti13 transformants able to tolerate up to 7.5 mM caffeine. Candidates obtained this way were subjected to killer eclipse assays to check for complementation of zymocin resist-ance; one cafR clone for each of the kti11 and kti13 mutants was found to be zymocin sensitive. To verify that complementation was based on the input YPH1 library, the kti11- and kti13-derived cafR zymS Leu+ transformants were subjected to plasmid DNA rescue. After retrans-formation of starting kti11 and kti13 mutant cells, we observed full complementation of the kti-associated phenotypes by the YPH1-based plasmid DNAs, termed pLF11.1 and pLF13.1 respectively (Fig. 1).
KTI11 is allelic to orphan YBL071w-A
Sequencing pLF11.1 revealed that kti11 was complemented by a 9.5 kb chromosome II fragment (co-ordinates 84 564–94 818) carrying almost nine complete open read-ing frames (ORFs), seven of which are non-essential (Fig. 2A). To check whether any of these may represent KTI11, we subjected defined null alleles (ybl075cΔ, ybl072cΔ, ybl071cΔ, ybl070cΔ, ybl069wΔ and ybl068wΔ) from the Euroscarf disruption collection to K. lactis killer eclipse assays. All but one null allele displayed zymocin sensitivity, with ybl071cΔ conferring zymocin resistance (Fig. 2B). To test whether YBL071c is identical to KTI11, we subcloned a pLF11.1 fragment containing YBL071w-A, YBL071c and YBL070c (pLF90) and YBL071c on its own (pLF91) and asked for gene-specific complementation. Surprisingly, although pLF90 was able to complement kti11 cells, pLF91 failed to do so (Figs 2A, C and D; data not shown). On testing YBL071w-A on its own (pLF93), however, we observed that complementation of kti11 cells was accomplished by single-copy YBL071w-A (Figs 2A, C and D). Thus, although a Euroscarf deletion in YBL071c conferred zymocin resistance similar to the kti11 mutant, it is not YBL071c but its neighbouring ORF YBL071w-A that complements kti11 cells. This apparent inconsistency can be explained when analysing YBL071c and YBL071w-A organization on chromosome II; both genes partially overlap on opposite DNA strands. So, deletion of YBL071c will affect the integrity of YBL071w-A, eventually leading to a non-functional latter gene. Indeed, by disrupting YBL071w-A individually without affecting YBL071c, we observed zymocin resistance within the genetic background of LS20 (see below: kti11Δ). In conclusion, these analyses indicate that KTI11 is allelic with yeast orphan YBL071w-A.
KTI13 is allelic to YAL020c/ATS1
DNA sequence analysis of the pLF13.1 insert revealed that kti13 was complemented by a 9.9 kb chromosome I fragment (co-ordinates 108 758–118 649). It carries four non-essential ORFs (Fig. 3A), systematic Euroscarf deletants of which (yal022cΔ, yal021cΔ, yal020cΔ and yal019wΔ) were subjected to K. lactis killer eclipse assays. All but one null allele displayed zymocin sensitivity, with yal020cΔ conferring zymocin resistance (Fig. 3B). To check whether YAL020c is allelic to KTI13, we next crossed kti13 cells with the yal020cΔ knock-out strain. The resulting diploid was found to be as zymocin resist-ant as the haploid parents, whereas crosses between kti13 and wild-type YAL020c led to zymocin sensitivity (Fig. 3C). Next, we subcloned YAL020c (pLF88), retransformed into starting kti13 mutant cells and found full complementation of caffeine sensitivity and zymocin resistance with YAL020c in single copy (Figs 3A and D). Taken together, this indicates that KTI13 is allelic to YAL020c. YAL020c has previously been isolated as an α-tubulin multicopy suppressor, ATS1 (Kirkpatrick and Solomon, 1994).
Disruption of KTI11 and KTI13 results in the complex tot phenotype
Based on our findings that, besides zymocin resistance, tot mutants phenocopy thermosensitivity, slow growth and hypersensitivity towards drugs (Frohloff et al., 2001; Jablonowski et al., 2001b), we created disruptions of KTI11 and KTI13 in LS20, our standard zymocin tester strain. kti11Δ and kti13Δ null alleles obtained by PCR gene targeting were subjected to phenotypic analysis and shown to display the complex tot phenotype indistinguishable from tot1–7Δ cells. The kti11Δ and kti13Δ strains each showed resistance towards exozymocin (Fig. 4A) and endogenously expressed γ-toxin (Fig. 4B). Moreover, deletion of KTI11 or KTI13 conferred a slow growth phenotype comparable to tot1–7Δ cells (see Fig. 6) and resulted in thermosensitivity above 38°C (Fig. 4C), a phenotype that could be partially rescued by the addition of 1 M sorbitol (data not shown). To check whether kti11Δ and kti13Δ cells might be affected in cell wall integrity, we tested their growth behaviour in the presence of caffeine and the cell wall poison Calcofluor white (CFW). Caffeine sensitivity was shown by kti11Δ and kti13Δ cells, although to a lesser extent for kti13Δ than for kti11Δ and tot3Δ cells (Fig. 5A). Compared with wild-type strain LS20 and the CFW-resistant control chs3Δ, both kti11Δ and kti13Δ cells were found to be hypersensitive towards CFW (Fig. 5B) to a similar extent to the tot3Δ control (Fig. 5B). This suggests that, as for tot1–7Δ strains, cell wall integrity may be affected in kti11Δ and kti13Δ cells. We next checked the effect of KTI11 and KTI13 deletions on transcriptional processes using 6-azauracil (6-AU) as an indicator. As shown in Fig. 5C, kti11Δ cells showed sensitivity against 6-AU, whereas kti13Δ cells behaved almost indistinguishably from wild-type strain LS20, a situation well in line with a generally weaker tot phenotype expressed by kti13Δ cells. Sensitivity of kti11Δ against 6-AU, however, was less pronounced than in a ppr2Δ strain lacking the elongation factor TFIIS. In conclusion, this drug-induced phenotype suggests a role for KTI11 in transcription elongation in vivo and is consistent with previous observations on 6-AU sensitivity of tot1–7Δ strains (Frohloff et al., 2001; Jablonowski et al., 2001b). In summary, loss of KTI11 and KTI13 gene function has a pleiotropic effect on a yeast cell’s performance, leading to the complex tot phenotype. As deletion of KTI11 and KTI13 phenocopies the effects of disrupting TOT1–7, it is likely that KTI11 and KTI13 may be involved in TOT/Elongator function too. To study this, we combined a disruption of the TOT3/ELP3 Elongator HAT gene (tot3Δ) with a null allele of either KTI11 or KTI13 and asked whether the double mutants displayed synthetic phenotypes. This analysis revealed that tot3Δkti11Δ and tot3Δkti13Δ double mutants were significantly affected with regard to slow growth phenotype expression compared with tot3Δ single deletants. Thus, tot3Δkti11Δ cells (Fig. 6A) and the double tot3Δkti13Δ mutant (Fig. 6B) grew more slowly than tot3Δ cells on their own. Intriguingly, compared with kti11Δ, the double tot3Δkti11Δ mutant performed better (Fig. 6A), indicating that kti11Δ-associated slow growth is partially suppressed by concomitant deletion of TOT3. This shows that combining KTI11 or KTI13 null alleles with tot3Δ affects growth behaviour and suggests that a cell’s dependence on TOT3 function for normal growth is determined in part by KTI11 and KTI13. Thus, KTI11 and KTI13 interact genetically with TOT3 and are possibly linked to TOT/Elongator function.
TOT gene expression analysis in kti11 and kti13 backgrounds
We next studied whether KTI11 and KTI13 constitute actively transcribed genes. As judged from RT–PCR analysis, both ORFs are transcribed in wild-type cells, although to a lesser extent than histone H3 gene HHT1 (Fig. 7A). Consistently, total RNA preparations from kti11Δ and kti13Δ knock-outs did not produce amplifiable RT–PCR KTI11 or KTI13 fragments (Fig. 7A). Intriguingly, KTI11 and KTI13 transcription itself was found to be mutually affected: kti13Δ cells were significantly reduced in KTI11 mRNA, whereas the KTI13 message fell in the kti11Δ deletant (Fig. 7A). As for the putative roles of KTI11 and KTI13, it has been formulated that both genes might act upstream of TOT4/KTI12, a gene whose deletion and overexpression results in zymocin resistance (Butler et al., 1994; Frohloff et al., 2001; Fichtner et al., 2002). As multicopy maintenance of KTI12/TOT4 in either the kti11 or the kti13 mutant partially suppressed the zymocin resistance phenotype, Butler et al. (1994) speculated that little or no Tot4p (which in turn confers zymocin resistance) was being produced in these kti backgrounds and that elevated TOT4 copy number would restore basal expression levels enough to account for the observed partial suppression. To address this hypothesis, we next examined TOT4 gene expression in either kti11Δ or kti13Δ cells by RT–PCR and one-step in vivo epitope tagging. TOT4 gene expression at the transcriptional level was hardly affected in the absence of KTI11 or KTI13 (Fig. 7A). As for the analysis of TOT4 gene expression at the translational level, we epitope tagged the TOT4 gene product in the kti backgrounds and found that, although expression of tagged Tot4p in the kti13 mutant was indistinguishable from that of the wild-type strain LL20 (Fig. 7B), kti11 cells produced slightly elevated Tot4p levels (Fig. 7B). The kti12 mutant itself, despite being tagged, did not produce any Tot4p at all (Fig. 7B). Although the latter result indicates that the kti12 mutant might have acquired a nonsense or promoter mutation, the former result shows that KTI13 cannot be considered to code for a component obligatory for TOT4 expression under normal conditions. As for the role of KTI11 in TOT4 function, slightly elevated Tot4p levels suggest that either TOT4 gene expression is upregulated or the stability/half-lifetime of the Tot4 gene product is aggravated when KTI11 is mutated. As the former can be excluded from the above RT–PCR study on TOT4 transcription in the absence of KTI11 (Fig. 7A), we favour the latter conclusion. Consistently, TOT1–3 and TOT5–7 expression was not affected in the absence of KTI11 or KTI13, suggesting that both genes are dispensable for TOT gene transcription (data not shown).
Identification of the Kti11 and Kti13 polypeptides
KTI13 has been isolated previously as a high-copy suppressor of α-tubulin mutants, ATS1. In addition, its gene product, Ats1p/Kti13p, is related to Prp20p, the yeast homologue of RCC1, which serves as GEF in the Ran–GTP/GDP cycle (Kirkpatrick and Solomon, 1994; Nigg, 1997). Database searches using the amino acid sequence predicted for the KTI11 gene product as a query revealed significant homology with as yet functionally unassigned proteins from human, Drosophila, fission yeast and Caenorhabditis elegans (Fig. 8A). Baker’s yeast Kti11p is 22/36% identical and 57/55% similar to two human Kti11 homologues over the entire sequence (Fig. 8A). Otherwise, homology between the proteins ranges from 50% identical and 70% similar (S. cerevisiae and Schizosaccharomyces pombe) to 41/63% (S. cerevisiae and C. elegans) and 41/57% (S. cerevisiae and Drosophila melanogaster). Using PCR-mediated one-step in vivo tagging, the Kti11 and Kti13 proteins were C-terminally marked with the HA epitope tag to analyse gene expression (Fig. 8B and C). Epitope tagging identified KTI11 and KTI13 as protein-encoding structural genes. Thus, total-protein extracts from exponentially grown KTI11- and KTI13-(HA)6 cells contained single Kti11 and Kti13 polypeptides that cross-reacted with the anti-HA antibody 3F10 (Fig. 8B and C) and were consistent with the predicted molecular weights (Kti11-(HA)6p: 18.3 kDa; Kti13-(HA)6p: 45.5 kDa). We next performed cell fractionation studies with yeast strains co-expressing HA-tagged Kti11p with c-Myc-tagged Tot4p and HA-tagged Kti13p with c-Myc-tagged Tot3p. Protein fractions were analysed by SDS–PAGE and immunoblotting to follow their migration patterns in comparison with the largest RNAPII subunit, Rpb1p, and three Elongator proteins, Elp2p/Tot2p, Elp3p/Tot3p and Elp5p/Tot5p. Studying Kti11p, this analysis revealed that some of the Kti11p fractions co-maintained Tot4p (Fig. 9, lanes 3–4) and, to a lesser extent, Elp2p/Tot2p (Fig. 9, lanes 3–4). Peak Kti11p fractions (Fig. 9, lanes 1–3), however, were devoid of Rpb1p and the Elongator HAT Elp3p/Tot3p and contained little Elp5p. Also, peak Rpb1p, Elp2p/Tot2p and Elp3p/Tot3p fractions, which overlapped (Fig. 9, lane 4), contained minimal Kti11p levels. Intriguingly, Kti11p was found to migrate as two distinct protein bands, suggesting HA-specific degradation or protein modification. Similarly, Kti13p fractionation paralleled that of Kti11p, although Kti13p itself was found to be much less abundant than the latter. The Kti13p peak fraction (Fig. 9, lane 3) overlapped with Tot4p (Fig. 9, lane 3) and did not contain Rpb1p or the Elongator HAT Elp3p/Tot3p. In summary, Kti11p and Kti13p partially co-migrated with Tot4p, an Elongator-associated protein (Fichtner et al., 2002), and peak fractions contained little or no RNAPII and Elongator subunits. As for the role that KTI11 and KTI13 may play in TOT function, this behaviour suggests that Kti11p and Kti13p are not Elongator subunits. Consistently, immunoprecipitation studies did not detect physical interaction between Elongator and either Kti11p or Kti13p (data not shown).
Zymocin-resistant S. cerevisiae tot mutants have recently been shown to be affected in seven TOT genes, six of which are identical to ELP1–6 coding for RNAPII Elongator (Frohloff et al., 2001; Jablonowski et al., 2001b; Li et al., 2001; Winkler et al., 2001). Together with our findings that six mutations defined through the kti screen (Butler et al., 1994) map to genes encoding Elongator subunits and/or Elongator-associated protein(s) too (Fichtner et al., 2002; M. J. R. Stark, L. Fichtner and R. Schaffrath, unpublished data), these results indicate that Elongator plays a crucial role in signalling toxicity of the K. lactis zymocin. This is reinforced by the data presented here showing that deletion of either KTI11 or KTI13 results in the complex tot phenotype common to tot1–7Δ cells and strongly suggests that the majority of mutations identified by the kti screen affect TOT/Elongator function. As for the putative roles played by KTI11 and KTI13, they are not expected to encode additional structural subunits of Elongator, as cell fractionation studies showed that Kti11p and Kti13p migrated in patterns distinct from the Elongator subunits. Also, because combining tot3Δ with kti13Δ led to a synthetic phenotype, and concomitant deletion of TOT3 in kti11Δ cells partially suppressed slow growth, these data suggest that the proteins in question are not part of the same complex. Consistently, immunoprecipitation techniques did not detect interaction between Elongator and Kti11p or Kti13p (data not shown). Therefore, other possible roles may include functions involved in TOT/Elongator regulation such that, in the absence (or by mutation) of the appropriate Kti proteins, individual TOT/Elongator subunits are not allowed to be assembled or are prevented from being associated with RNAPII. The former speculation that KTI11 and KTI13 might act upstream of TOT4/KTI12 by positively influencing its expression (Butler et al., 1994) could not be confirmed by our RT–PCR and in vivo epitope-tagging studies. Thus, TOT4 expression was not significantly downregulated at the transcriptional or translational level in kti11Δ or kti13Δ cells. So, the assumption of Butler et al. (1994) that multiple copies of TOT4 would restore basal expression levels enough to account for the observed partial suppression of kti11- and kti13-associated zymocin resistance can be ruled out.
Concerning KTI11 function, the more severe tot phe-notype seen as a result of KTI11 deletion (in comparison with loss of KTI13 function) points towards a regulatory role of Kti11p in TOT function. Our finding that Tot4p levels appear to be elevated in kti11Δ cells may indicate that, under normal wild-type conditions, TOT4 is under negative control of KTI11. However, this assumption contrasts with the observation that zymocin resistance associated with kti11 cells can be counteracted by elevated TOT4 copy number (Butler et al., 1994) and our finding that TOT4 transcription is largely unaffected by KTI11 gene function. Whatever its precise role, KTI11 has been shown in this report to encode a protein, Kti11p, that is highly conserved among eukaryotes. Together with the observation that a kti11Δ null mutation combined with a tot3Δ knock-out results in a synthetic effect on slow growth phenotype expression (in comparison with the single tot3Δ deletant) and the finding that a KTI11 deletion is partially suppressed by loss of TOT3 function, these data imply that KTI11 interacts genetically with TOT3, linking KTI11 to TOT/Elongator function.
KTI13/ATS1 was identified as a high-copy suppressor of conditional α-tubulin mutants that arrest in G1 with excess microtubules (Kirkpatrick and Solomon, 1994). ATS1 is conserved to PRP20, a yeast homologue of the mammalian mitotic checkpoint gene RCC1 (Kirkpatrick and Salomon, 1994). Prp20p and RCC1 serve as gua-nine nucleotide exchange factor (GEF) during the Ran–GTP/GDP cycle involved in nucleocytoplasmic transport (Nigg, 1997). As Tot4p/Kti12p has been shown to carry a potential GTP-binding P-loop motif that is essential for TOT4 function in vivo (Fichtner et al., 2002), it is possible that Kti13p serves as a GEF for Tot4p. Consistent with this assumption is the genetic interaction seen between TOT4 and KTI13 that has been shown to result in partial suppression of zymocin resistance (Butler et al., 1994). Moreover, both deletion of KTI13 and elevated TOT4 gene copy number result in mild tot phenotypes (Fichtner et al., 2002) that are considerably less pronounced than loss of TOT1–7 or KTI11 functions (Frohloff et al., 2001; Jablonowski et al., 2001b). If KTI13 acted upstream of TOT4 by activating Tot4p post-translationally, one would assume that total Tot4p levels were not affected in the absence of KTI13 function. Consistently, we found that, regardless of whether cells encode KTI13 or not, total Tot4p levels remained unchanged. Together with the finding that multicopy TOT4 partially suppresses kti13-associated zymocin resistance (Butler et al., 1994), this may suggest that Tot4p overproduction can bypass the potential requirement of KTI13 for TOT4 function.
Strains, media and general methods
All yeast strains used or generated in this study are described in Table 1. For cultivation, standard rich and minimal growth media, YPD and SD were used. For phenotypic analysis, these media were supplemented with 6-AU (0–100 μg ml−1), caffeine (0–7.5 mM) or CFW (0–25 μg ml−1). For 6-AU assays, Ura+ transformants of the ktiΔ strains were used. Yeast transformations involved the lithium acetate method (Gietz et al., 1992). Bacterial transformations used Escherichia coli strain DH5α and TOP10 grown in LB supplemented with one or several of the following: ampicillin (100 μg ml−1), Xgal (80 μg ml−1) and IPTG (50 μg ml−1).
|AWJ137||αleu2 trp1 [k1+ k2+]||Frohloff et al. (2001)|
|NK40||αade1 ade2 leu2 [k1− k2+]||Frohloff et al. (2001)|
|FY1679-08A||MATa ura3-52 leu2Δ1 trp1Δ63 his3Δ200 GAL||Euroscarf, Frankfurt|
|BY4741||MATa his3Δ1 leu2Δmet15Δura3Δ||Euroscarf, Frankfurt|
|Y03094||As BY4741, but ybl068wΔ::kanMX4||Euroscarf, Frankfurt|
|Y03095||As BY4741, but ybl069wΔ::kanMX4||Euroscarf, Frankfurt|
|Y03096||As BY4741, but ybl070cΔ::kanMX4||Euroscarf, Frankfurt|
|Y03097||As BY4741, but ybl071cΔ::kanMX4||Euroscarf, Frankfurt|
|Y03098||As BY4741, but ybl072cΔ::kanMX4||Euroscarf, Frankfurt|
|Y03101||As BY4741, but ybl075cΔ::kanMX4||Euroscarf, Frankfurt|
|Y00386||As BY4741, but yal022cΔ::kanMX4||Euroscarf, Frankfurt|
|Y00387||As BY4741, but yal021cΔ::kanMX4||Euroscarf, Frankfurt|
|Y00388||As BY4741, but yal020cΔ::kanMX4||Euroscarf, Frankfurt|
|Y00389||As BY4741, but yal019wΔ::kanMX4||Euroscarf, Frankfurt|
|LL20||MATαleu2-3,112 his3-11,15||Butler et al. (1994)|
|ARB69||As LL20, but kti11||Butler et al. (1994)|
|ARB90||As LL20, but kti13||Butler et al. (1994)|
|ARB18||As LL20, but kti12||Butler et al. (1994)|
|LFYA||ARB90 × BY4741||This study|
|LFYB||ARB90 × Y00388||This study|
|LS20||As LL20, but ura3||Schaffrath et al. (1997)|
|FFY6||As LS20, but dst1Δ/ppr2Δ::KlLEU2||Frohloff et al. (2001)|
|LFY11||As LS20, but kti11Δ::KlLEU2||This study|
|LFY13||As LS20, but kti13Δ::KlLEU2||This study|
|DJY3||As LS20, but chs3Δ::KlLEU2||Jablonowski et al. (2001a)|
|FFY3||As LS20, but tot3Δ::KlLEU2||Frohloff et al. (2001)|
|LFY13-3||As FFY3, but kti13Δ::SpHIS5||This study|
|LFY11-3||As FFY3, but kti11Δ::SpHIS5||This study|
|LFY11-4dt||As FY1679-08A, but KTI11-(HA)6::KlTRP1, TOT4-(c-myc)3::SpHIS5||This study|
|LFY13-3dt||As FY1679-08A, but KTI13-(HA)6::KlTRP1, TOT3-(c-myc)3::SpHIS5||This study|
|ARB69-4t||As ARB69, but TOT4/KTI12-(c-myc)3::SpHIS5||This study|
|ARB90-4t||As ARB90, but TOT4/KTI12-(c-myc)3::SpHIS5||This study|
|ARB18-4t||As ARB18, but tot4/kti12-(c-myc)3::SpHIS5||This study|
|LL20-4t||As LL20, but TOT4/KTI12-(c-myc)3::SpHIS5||This study|
|FFY2/5-dt||As FY1679-08A, but TOT2-(c-myc)3::SpHIS5, TOT5-(HA)6::KlTRP1||Fichtner et al. (2002)|
Genetic complementation of the kti11 and kti13 mutants
Complementation of the kti11 and kti13 mutants involved transformation with the single-copy library YPH1 (ATCC 77162: CEN4/LEU2). Leu+ transformants (5000 for each of the kti11 and kti13 backgrounds representing roughly three or four genome equivalents) were selected on SD medium supplemented with histidine and replica plated onto YPD medium containing 7.5 mM caffeine. Using killer eclipse assays (see below), 26 and 50 cafR Leu+ transformants raised in the kti11 and kti13 backgrounds, respectively, were checked for complementation of zymocin resistance. One cafR clone for each of the kti11 and kti13 backgrounds was found to be zymocin sensitive. After DNA rescue and retransformation, the kti-associated phenotypes were complemented by the rescued plasmid DNAs termed pLF11.1 and pLF13.1 respectively. DNA sequence analysis of the pLF11.1 and pLF13.1 inserts used primers LF50.1 (5′-GCT TCG CTA CTT GGA GCC-3′) and LF50.2 (5′-TAT AGG CGC CAG CAA CCG-3′). Complementation was tested for kti13 with pLF88, a YCplac111-based plasmid carrying a 1.8 kb NsiI–BglII fragment of pLF13.1, which contains YAL020c/ATS1. For kti11, a 1.7 kb PstI–HindIII genomic fragment from pLF11.1 containing YBL071w-A, YBL071c and YBL070c (pLF90), a 0.6 kb PCR fragment carrying just YBL071c (pLF91) and a genomic 0.8 kb HindIII–SacI fragment coding for YBL071w-A (pLF93) were tested for complementation.
Kluyveromyces lactis zymocin methods
Zymocin assays used the killer eclipse system (Kishida et al., 1996) and K. lactis strains AWJ137 (zymocin producer) and NK40 (non-producer) (Table 1). Growth was monitored after incubation for 1 day at 30°C. Galactose-inducible expression of the γ-toxin was done with pHMS14, a vector carrying the UASGAL1–γ-toxin fusion (Frohloff et al., 2001). Plasmid pHMS22 served as a UASGAL1 vector control (Frohloff et al., 2001). Assays included testing growth on S agar containing 2% glucose or 2% galactose by replica spotting. Diploid yeast strains were assessed for zymocin sensitivity/resistance by mating LL20-derived cells (MATα) with BY4741 derivatives (MATa) and subjecting them to killer eclipse assays as above.
Construction, isolation and analyses of kti11Δ and kti13Δ mutants
For PCR-mediated construction of defined kti11Δ and kti13Δ null alleles, the YDp-KlL plasmid carrying the K. lactis LEU2 marker was used (Frohloff et al., 2001) together with the following knock-out primer pairs: FW-ko-KTI11: 5′-ACA TAC CAC GAC TGT AAG CAC ATC ATT TGT ACA ATA CAT TAC CAG CTG AAC GAC GGC CAG TGA ATT CCC GG-3′; RV-ko-KTI11: 5′-CTT TAT TTC TAT TTG TAT TCT CGA TCT AGC CTC TCA TCT TTA GGC AGC AGA GCT TGG CTG CAG GTC GAC GG-3′; FW-ko-KTI13: 5′-GCT ATA ACA GGC TTG TAT CGA TGA GTT GTG TGT ATG CGT TTG GGT CTA ATC GAC GGC CAG TGA ATT CCC GG-3′; RV-ko-KTI13: 5′-ATA GTG GGT ATA TAG TTA CTT ATC AGT GCT AGA GCA CGA TCC ACG TGG TGA GCT TGG CTG CAG GTC GAC GG-3′. Leu+ transformants were verified by PCR using the knock-out primer pairs to amplify the foreign marker and ORF-specific promoter and terminator primer pairs (FW-KTI11: 5′-ATG AAC ACT GCG TAA GAG AAA GCC C-3′; RV-KTI11: 5′-TAG TAA TTC CAA CCG GAG TCC AGC G-3′; FW-KTI13: 5′-CCA CAC TTG CAT CCG GAA CTG TTG GC-3′; RV-KTI13: 5′-CGC AAG TAC GGG CGA TAA CAA AAG GC-3′) to check for proper integration. Phenotypic analyses of the kti11Δ and kti13Δ mutants were performed as described previously (Frohloff et al., 2001; Jablonowski et al., 2001b). kti11Δtot3Δ and kti13Δtot3Δ double mutants were created by PCR-mediated disruption of either KTI11 or KTI13 in FFY3 (Table 1; Frohloff et al., 2001) using the S. pombe HIS5 marker gene on plasmid YDp-SpH (Jablonowski et al., 2001a) and the above knock-out primer pairs.
Epitope tagging, cell fractionation and immunological techniques
Epitopes were fused to KTI11 and KTI13 using PCR-based one-step in vivo tagging as described for TOT1–5 (Frohloff et al., 2001). For KTI11 tagging, the following primers were used (S3-KTI11: 5′-ACT ACG AAG AGG CAG GCA TCC ACC CCC CTG AGC CTA TTG CCG CTG CTG CCC GTA CGC TGC AGG TCG AC-3′ and S2-KTI11: 5′-TAT AGC TCT TTC TTT ATT TCT ATT TGT ATT CTC GAT CTA GCC TCT CAT CTA TCG ATG AAT TCG AGC TCG-3′). KTI13 tagging involved primer S3-KTI13 (5′-CTG GAA AAC CTC GCG TGT TTG GCG GAT GTG CCA CCA CGT GGA TCG TGC TCC GTA CGC TGC AGG TCG AC-3′) and S2-KTI13 (5′-TGA ATG GAC ATC TAT GTA TAT GAT AGT GGG TAT ATA GTT ACT TAT CAG TGA TCG ATG AAT TCG AGC TCG-3′). TOT4/KTI12 tagging was carried out according to the method of Frohloff et al. (2001). Detection of tagged Tot/Kti proteins involved 9E10 mouse antibody recognizing the c-Myc epitope (Roche) and 3F10 rat antibody recognizing the HA epitope (Roche) as described previously (Frohloff et al., 2001). Monoclonal mouse antibody 8WG16 (Covance) was used to detect RNAPII subunit Rpb1p. Cell fractionation used yeast growth conditions and sucrose gradient ultracentrifugation techniques essentially as described previously (Kölling and Hollenberg, 1994).
Total RNA isolation and RT–PCR were performed as described previously (Frohloff et al., 2001). HHT1 and TOT1–4 primers are described by Jablonowski et al. (2001c). RT–PCR involved these and primers for TOT5 (5′-TATT GACGCTACGCAGATGG-3′ and 5′-CTCCTCTTCTTGCTT AGTGG-3′), TOT6 (5′-GATGCTACCTTCGTCAACTC-3′ and 5′-TACGTCCTTTGCAAAACCGG-3′), TOT7 (5′-TTTGCAAA GGAGCTACCTGG-3′ and 5′-GGAAGCAACAGTACAACCC-3′), KTI11 (5′-CGAAGATATGACGTTTGAGCC-3′ and 5′-CCT CTTCGTAGTACTCAGCC-3′) and KTI13 (5′-TAGTCAGGAA GATAGCGTGC-3′ and 5′-GTATCGTACACCAATACGGG-3′) to amplify fragments specific for histone H3 (HHT1: 0.32 kb), TOT1/ELP1 (0.49 kb), TOT2/ELP2 (0.57 kb), TOT3/ELP3 (0.53 kb), TOT4/KTI12 (0.47 kb), TOT5/ELP5 (0.5 kb), TOT6/ELP6 (0.5 kb), TOT7/ELP4 (0.5 kb), KTI11 (0.18 kb) and KTI13/ATS1 (0. 42 kb) genes.
Thanks are due to M. Stark, M. Larsen, K. Hager and C. Mehlgarten for providing the kti mutants, help with homology searches, cell fractionation studies and phenotypic analysis of ktiΔ mutants. L.F. acknowledges receipt of a grant sponsored by the Martin-Luther-Universität. The work was supported by a grant from the DFG (Scha 750/2) to R.S.
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