New Results

Unconventional kinetochore kinases KKT2 and KKT3 have a unique zinc finger that promotes their kinetochore localization

Gabriele Marcianò, Olga O. Nerusheva, Midori Ishii, Bungo Akiyoshi
doi: https://doi.org/10.1101/2019.12.13.875419

Abstract

Chromosome segregation in eukaryotes is driven by the kinetochore, a macromolecular protein complex that assembles onto centromeric DNA and binds spindle microtubules. Cells must tightly control the number and position of kinetochores so that all chromosomes assemble a single kinetochore. A central player in this process is the centromere-specific histone H3 variant CENP-A, which localizes specifically within centromeres and promotes kinetochore assembly. However, CENP-A is absent from several eukaryotic lineages including kinetoplastids, a group of evolutionarily divergent eukaryotes that have an unconventional set of kinetochore proteins. It remains unknown how kinetoplastids specify kinetochore positions or promote kinetochore assembly in the absence of CENP-A. Here we studied two homologous kinetoplastid kinases (KKT2 and KKT3) that localize constitutively at centromeres. KKT2 and KKT3 central domains were sufficient for centromere localization in Trypanosoma brucei. Crystal structures of the KKT2 central domain from two divergent kinetoplastids revealed a unique zinc finger domain, which promotes its kinetochore localization in T. brucei. Mutations in the equivalent zinc finger domain of KKT3 abolished its kinetochore localization and function. This study lays the foundation for understanding the mechanism of kinetochore specification and assembly in kinetoplastids.

Introduction

The kinetochore is the macromolecular protein complex that drives chromosome segregation during mitosis and meiosis in eukaryotes. Its fundamental functions are to bind DNA and spindle microtubules (Musacchio and Desai, 2017). In most eukaryotes, kinetochores assemble within a single chromosomal region, called the centromere. While components of spindle microtubules are highly conserved across eukaryotes (Wickstead and Gull, 2011; Findeisen et al., 2014), centromere DNA is known to evolve rapidly (Henikoff et al., 2001). It is critical that a single kinetochore is formed per chromosome and its position is maintained between successive cell divisions. A key player involved in this kinetochore specification process is the centromere-specific histone H3 variant CENP-A, which is found in most sequenced eukaryotic genomes (Talbert et al., 2009). CENP-A localizes specifically at centromeres throughout the cell cycle and recruits HJURP, a specific chaperone that incorporates CENP-A onto centromeres (Black and Cleveland, 2011; McKinley and Cheeseman, 2016; Stankovic and Jansen, 2017). Besides CENP-A, components of the constitutive centromere-associated network (CCAN) also localize at centromeres constitutively. CENP-A-containing nucleosomes are recognized by CCAN components, which in turn recruit the KNL1-Mis12-Ndc80 (KMN) network that has microtubule-binding activities. In addition to these structural kinetochore proteins, several protein kinases are known to localize at mitotic kinetochores, including Cdk1, Aurora B, Bub1, Mps1, and Plk1 (Cheeseman and Desai, 2008). These protein kinases regulate various aspects of mitosis, including kinetochore assembly, error correction, and the spindle checkpoint (Carmena et al., 2012; London and Biggins, 2014; Hara and Fukagawa, 2018).

Kinetoplastids are evolutionarily divergent eukaryotes that are defined by the presence of a unique organelle called the kinetoplast that contains a cluster of mitochondrial DNA (d’Avila-Levy et al., 2015). Centromere positions have been mapped in three kinetoplastids, Trypanosoma brucei (Obado et al., 2007), Trypanosoma cruzi (Obado et al., 2005), and Leishmania (Garcia-Silva et al., 2017), which all have regional centromeres. There is no specific DNA sequence that is common to all centromeres in each organism, suggesting that kinetoplastids also determine their kinetochore positions in a sequence-independent manner. However, none of CENP-A or any other canonical structural kinetochore proteins has been identified in kinetoplastids (Lowell and Cross, 2004; Berriman et al., 2005; Aslett et al., 2010). They instead have unique kinetochore proteins, such as KKT1–25 (Akiyoshi and Gull, 2014; Nerusheva and Akiyoshi, 2016; Nerusheva et al., 2019) and KKIP1–12 (D’Archivio and Wickstead, 2017; Brusini et al., 2019) in T. brucei. Among these proteins, six proteins localize at centromeres throughout the cell cycle (KKT2, KKT3, KKT4, KKT20, KKT22, and KKT23), implying close association with centromeric DNA. KKT2 and KKT3 are homologous proteins and have three domains conserved among kinetoplastids: a protein kinase domain that is classified as unique among known eukaryotic kinase subfamilies (Parsons et al., 2005), a central domain of unknown function, and divergent polo boxes (DPB) (Figure 1 and 2). Presence of an N-terminal kinase domain and a C-terminal DPB suggests that KKT2 and KKT3 likely share common ancestry with polo-like kinases (Nerusheva and Akiyoshi, 2016). Interestingly, a protein kinase domain is not present in any constitutively-localized kinetochore protein in other eukaryotes, making these protein kinases a unique feature of kinetoplastid kinetochores. In addition to the three domains that are highly conserved among kinetoplastids, AT-hook and SPKK DNA-binding motifs are found in some species, suggesting that these proteins are located close to DNA (Akiyoshi and Gull, 2014). Although RNAi-mediated knockdown of KKT2 or KKT3 leads to growth defects (Akiyoshi and Gull, 2014; Jones et al., 2014), little is known about their molecular function. In this report, we have revealed a unique zinc finger in the KKT2 and KKT3 central domain, which is important for their kinetochore localization and function in T. brucei.

Figure 1.
Figure 1. KKT3 central domain is able to localize at centromeres in T. brucei

(A) Schematic of the T. brucei KKT3 protein. Percentages of GFP-positive 2K1N cells that have kinetochore-like dots were quantified at 1 day post-induction (n > 24, each).

(B) Ectopically expressed TbKKT3 fragments that contain the central domain form kinetochore-like dots. Inducible GFP-NLS fusion proteins were expressed with 10 ng/mL doxycycline. Cell lines, BAP291, BAP292, BAP379, BAP296, BAP378, BAP377, BAP418. Scale bar, 5 μm.

Figure 2.
Figure 2. KKT2 has multiple domains that can promote centromere localization in T. brucei

(A) Schematic of the T. brucei KKT2 protein. Percentages of GFP-positive 2K1N cells that have kinetochore-like dots were quantified at 1 day post-induction (n > 22, each).

(B) Ectopically expressed TbKKT2 fragments that contain either the central domain (562–677), 672–1030, or the divergent polo boxes (1024–1260) form kinetochore-like dots. Inducible GFP-NLS fusion proteins were expressed with 10 ng/mL doxycycline. Cell lines, BAP327, BAP328, BAP381, BAP331, BAP457, BAP519, BAP517. Scale bar, 5 μm.

(C) TbKKT2 DPB WT, not W1048A, co-purifies with several kinetochore proteins. Inducible GFP-NLS fusion proteins were expressed with 10 ng/mL doxycycline, and immunoprecipitation was performed using anti-GFP antibodies. Cell lines, BAP517, BAP535.

(D) TbKKT2672–1030 co-purifies with KKIP1 and several kinetochore proteins. Cell line, BAP519.

Results

The central domain of KKT3 is able to localize at centromeres in T. brucei

To understand how KKT3 localizes at centromeres, we determined which domain was responsible for its centromere localization by expressing in T. brucei procyclic form (insect stage) cells, a series of truncated versions of TbKKT3, fused with a GFP-tagged nuclear localization signal peptide (GFP-NLS) (Figure 1A). We found that the N-terminal protein kinase domain or the C-terminal DPB of TbKKT3 did not localize at centromeres. In contrast, TbKKT3594–1058 and TbKKT3594–811 that contain the central region formed kinetochore-like dots (Figure 1B). TbKKT3594–728 also formed dots, suggesting that AT-hook and SPKK DNA-binding motifs (residues 771–780) are dispensable for centromere localization in this assay. These results revealed the importance of the central domain for the recruitment of TbKKT3 onto centromeres.

Multiple domains of KKT2 are able to localize at centromeres in T. brucei

We next performed a similar analysis for TbKKT2 and found that its central domain (residues 562– 677) was able to localize at centromeres (Figure 2A, B). Therefore, the central domains of both TbKKT2 and TbKKT3 are sufficient for centromere localization in this assay. In a previous study, we showed that ectopically-expressed TbKKT2 DPB (1024–1260) localized at centromeres (Nerusheva and Akiyoshi, 2016). The present study confirmed this result and also identified another region (residues 672–1030) that localized at centromeres (Figure 2A, B). These results show that TbKKT2 has multiple domains that are sufficient for centromere localization in this assay. Based on our previous finding that TbKKT2 co-immunoprecipitated with a number of kinetochore proteins (Akiyoshi and Gull, 2014), we reasoned that these fragments might localize at centromeres by interacting with other kinetochore proteins. To test this possibility, we immunoprecipitated TbKKT2 fragments and performed mass spectrometry to identify co-purifying proteins. We found that TbKKT2 DPB co-purified with several kinetochore proteins (KKT1, KKT6, KKT7, and KKT8), which was abolished in the TbKKT2 W1048A mutant that did not localize at kinetochores (Figure 2C and Table S1) (Nerusheva and Akiyoshi, 2016). Similarly, TbKKT2672–1030 co-purified with several kinetochore proteins, with KKIP1 being the top hit (Figure 2D and Table S1). These results suggest that, when ectopically expressed in trypanosomes, TbKKT2672–1030 and TbKKT2 DPB likely localize at centromeres by interacting with other kinetochore proteins. Because those co-purifying kinetochore proteins (e.g. KKT1, KKT6, KKT7, KKT8, KKIP1) localize at kinetochores only transiently, it is likely that the constitutively-localized TbKKT2 protein recruits them using TbKKT2672–1030 and DPB domains. Furtheremore, the fact that KKIP1 was detected in the immunoprecipitates of TbKKT2, but not other kinetochore proteins (Nerusheva et al., 2019), raises a possibility that KKIP1 is recruited to kinetochores by directly interacting with the 672–1030 region of TbKKT2. By contrast, the central domains of TbKKT2 and TbKKT3 did not robustly co-purify with other kinetochore proteins (Table S1).

The Bodo saltans KKT2 central domain adopts a unique zinc finger structure

To gain insights into how the central domains of KKT2 and KKT3 localize specifically at centromeres, we expressed and purified recombinant TbKKT2 and TbKKT3 proteins for their structure determination by X-ray crystallography. Our attempts to purify the TbKKT3 central domain were unsuccessful, but we managed to express and purify from E. coli the central domain of KKT2 from several kinetoplastids, including Bodo saltans (a free-living kinetoplastid (Jackson et al., 2016)) and Perkinsela (endosymbiotic kinetoplastids (Tanifuji et al., 2017)) (Figure S1). We successfully obtained crystals of the BsKKT2 central domain (residues 572–668), which allowed us to determine its structure to 1.8 Å resolution by zinc single-wavelength anomalous dispersion (Zn-SAD) phasing (Figure 3 and Table 1). Our analysis revealed the presence of two distinct zinc fingers: an N-terminal zinc finger (referred to as Znf1) consisting of 2 β-sheets (where β-strands 1, 4 and 5 comprise the first β-sheet, and β-strands 2 and 3 comprise the second β-sheet) and one α-helix, and a C-terminal zinc finger (referred to as Znf2) consisting of one β-sheet (comprising β-strands 6 and 7) and one α-helix (Figure 3A, B). BsKKT2 Znf1 coordinates two zinc ions and Znf2 coordinates one zinc ion.

Figure S1.
Figure S1. Purification of Bodo saltans KKT2 and Perkinsela KKT2a

Size exclusion chromatography of (A) Bodo saltans KKT2 central domain and (B) Perkinsela KKT2a central domain with respective SDS-PAGE gels showing pooled fractions.

Figure 3.
Figure 3. Crystal structure of Bodo saltans KKT2 central domain reveals the presence of two zinc fingers Znf1 and Znf2

(A) Topology diagram of the BsKKT2 central domain showing Znf1 in green and Znf2 in cyan.

(B) Cartoon representation of the BsKKT2 central domain in two orientations. Zinc ions are shown in grey spheres. The structure is colored as in (A).

(C) Surface representation of the BsKKT2 central domain colored according to sequence conservation using the ConSurf server (Landau et al., 2005; Ashkenazy et al., 2016). Structure orientation as in (B).

(D) Electrostatic surface potential of the BsKKT2 central domain generated by APBS (Jurrus et al., 2018). Structure orientation as in (B).

A structural homology search using the DALI server (Holm and Laakso, 2016) indicated that BsKKT2 Znf1 has weak structural similarity to proteins that have C1 domains (Table S2). C1 domains were originally discovered as lipid-binding modules in protein kinase Cs (PKCs) and are characterized by the HX12CX2CXnCX2CX4HX2CX7C motif (Colón-González and Kazanietz, 2006; Das and Rahman, 2014). C1 domains are classified into a typical C1 domain, which binds diacylglycerol or phorbol esters, and an atypical C1 domain, which is not known to bind ligands. The closest structural homolog of BsKKT2 Znf1 was the atypical C1 domain of the Vav1 protein (RMSD 2.7 Å across 52 Cα). Although BsKKT2 Znf1 and the Vav1 C1 domain share some structural similarity, their superposition revealed fundamental differences (Figure 4). Coordination of one zinc ion in BsKKT2 Znf1 occurs via the N-terminal residues Cys 580 and His 584, while that in the Vav1 C1 domain occurs via the N-terminal His 516 and C-terminal Cys 564. More importantly, KKT2 Znf1 does not have the HX12CX2CXnCX2CX4HX2CX7C motif that is present in all C1 domains. Therefore, KKT2 Znf1 and C1 domains are distinct domains, and their structural similarity is likely to be a product of convergent evolution.

Figure 4.
Figure 4. Bodo saltans KKT2 Znf1 is a unique domain

(A) Structure superposition of BsKKT2 Znf1 in green and its structural homolog Vav1 C1 domain in brown (PDB: 3KY9 (Yu et al., 2010)). Zinc ions are shown in grey spheres.

(B) Topology diagram of BsKKT2 Znf1 and Vav1 C1 domain.

(C) Close-up view showing a key difference in zinc coordination between BsKKT2 Znf1 and Vav1 C1 domain.

Structural analysis of BsKKT2 Znf2 revealed a classical C2H2 zinc finger domain (Table S3). C2H2 zinc fingers are known to bind DNA, RNA, or protein (Krishna et al., 2003; Brayer and Segal, 2008). In most known cases, two or more C2H2 zinc fingers are used to recognize specific DNA sequences (Wolfe et al., 2000). This recognition is typically achieved by specific interactions between the side chain of residues in positions -1, 2, 3, and 6 in the recognition α-helix (where -1 is the residue immediately preceding the α-helix) and DNA bases (Wolfe et al., 2000). Notably, some proteins with a single zinc finger can recognize specific DNA sequences (Omichinski et al., 1997; Dathan et al., 2002). BsKKT2 Znf2 structure consists of one C2H2 domain (−1: Ser 653, 2: Thr 655, 3: Lys 656, 6: Tyr 659). The sequence alignment of BsKKT2 Znf2 shows that residues at the position -1 and 3 are highly conserved in trypanosomatids, while those at position 2 and 6 are not (Figure 6A).

Figure 6.
Figure 6. KKT2 Znf1 is critical for the centromere localization in T. brucei

(A) Multiple sequence alignment of KKT2. Residues mutated in (B) as well as secondary structures of BsKKT2 are shown. Putative SPKK motifs are highlighted in boxes.

(B) Kinetochore localization of TbKKT2562–677 depends on Znf1, not Znf2. Percentages of GFP-positive 2K1N cells that have kinetochore-like dots were quantified at 1 day post-induction (n = 40, each). Inducible GFP-NLS fusion proteins were expressed with 10 ng/mL doxycycline. Cell lines, BAP457, BAP1700, BAP1702, BAP1710, BAP1712, BAP1715, BAP1717, BAP1649, BAP1719, BAP1837.

(C) TbKKT2 C576A and D622A mutants localize at kinetochores but fail to support normal cell growth. One allele of TbKKT2 was mutated and tagged with an N-terminal YFP tag, and the other allele was depleted using RNAi-mediated knockdown by targeting the 5’UTR of the TbKKT2 transcript. Top: cells were diluted every two days and cell growth was monitored for 8 days upon induction of RNAi. Controls are uninduced cell cultures. Similar results were obtained for at least three clones of TbKKT2 mutants. Bottom: Example of cells expressing the TbKKT2 mutants prior to RNAi induction, showing that they localize at kinetochores (n > 150, each). Maximum intensity projection are shown. RNAi was induced with 1 µg/mL doxycycline. Cell lines, BAP1789, BAP1779, BAP1786. Scale bars, 5 μm.

Znf1 is conserved in Perkinsela KKT2a

We next asked whether the central domain structure is conserved among kinetoplastids. Perkinsela is a highly divergent endosymbiotic kinetoplastid that lives inside Paramoeba (Tanifuji et al., 2017). Homology search identified three proteins that have similarity to KKT2 and KKT3. Interestingly, similarities among these Perkinsela proteins are higher than those between them and KKT2 or KKT3 in other kinetoplastids (Figure S2). Because these Perkinsela proteins overall have higher sequence similarity to KKT2 than KKT3, we call them PkKKT2a (XU18_4017), PkKKT2b (XU18_0308), and PkKKT2c (XU18_4564). PkKKT2c does not have a kinase domain, like KKT20 in other kinetoplastids (Nerusheva and Akiyoshi, 2016). Our sequence alignment suggests that PkKKT2a and PkKKT2b have a Znf1-like domain but do not have a Znf2-like C2H2 domain (Figure S2).

Figure S2.
Figure S2. Multiple sequence alignment of KKT2 and KKT3 from various kinetoplastids, as well as Perkinsela KKT2a and KKT2b

Position of the conserved aspartic acid is highlighted. A putative AT-hook motif in Perkinsela KKT2b is highlighted in a red box.

We determined the crystal structure of the PkKKT2a central domain (residues 551–679) at 2.9 Å resolution by Zn-SAD phasing (Figure S1 and Table 2), which confirmed the presence of a Znf1-like structure: 2 β-sheets (residues 551–647), followed by an extended C-terminal α-helix (residues 648–679) (Figure 5A, B). The Znf1-like domain of PkKKT2a overlaps closely with that of BsKKT2 (RMSD: 0.79 Å across 39 Cα), with the exception of some differences being localized to the loop insertion and the absence of a C2H2 domain in the PkKKT2a central domain (Figure 5C), consistent with our sequence analysis (Figure S2). Taken together, our structures have revealed that the KKT2 central domain contains a zinc finger domain that is conserved between Bodo saltans and Perkinsela. Given the sequence similarity of KKT2 between Bodo saltans and other trypanosomatids (Figure 6A), it is likely that the unique zinc finger structure is conserved among kinetoplaitids.

Figure 5.
Figure 5. Crystal structure of Perkinsela KKT2a central domain highlights the conservation of Znf1

(A) Cartoon representation of the PkKKT2a central domain in two orientations. Zinc ions are shown in grey spheres.

(B) Topology diagram of the PkKKT2a central domain structure.

(C) Structure superposition of the PkKKT2a central domain and BsKKT2 Znf1, showing that the core of the zinc finger structure is conserved. Variations between the two structures are due to sequence insertions within Znf1 and the absence of the Znf2 domain at the C-terminus in the PkKKT2a central domain.

Zinc finger Znf1 of KKT2 is important for long-term viability in T. brucei

To examine the functional relevance of the KKT2 Znf1 and Znf2 zinc fingers, we tested their mutants in T. brucei. Because KKT2 has multiple domains that can independently localize at kinetochores (Figure 2), we first expressed mutants in our ectopic expression of the central domain (TbKKT2562–677). We found that mutations in TbKKT2 Znf1 Zn-coordinating residues (C576A, H580A, C597A, C600A, C616A, C619A) all abolished kinetochore localization (Figure 6A, B). In contrast, similar mutations in TbKKT2 Znf2 (C640A C643A) did not affect the localization.

To gain insights into how KKT2 Znf1 may promote kinetochore localization, we analysed conservation and electrostatic potential of BsKKT2 Znf1 surface residues, to identify possible patches that may be involved in this process. Our analysis revealed a highly conserved acidic patch, centered around residue BsKKT2 D626 (Figure 3C, D, and Figure S3). Interestingly, this aspartic acid is strictly conserved in all KKT2 and KKT3 proteins (Figure S2). To test the importance of this residue, we mutated the corresponding residue and found that TbKKT2 D622A and TbKKT2 D622E mutants failed to localize at kinetochores (Figure 6B). Taken together, our results show that Znf1, but not Znf2, is important for the kinetochore localization of the TbKKT2 central domain.

Figure S3.
Figure S3. Bodo saltans KKT2 Znf1 surface sequence conservation and electrostatic potential reveal the presence of a conserved acidic patch centering on residue D626

(A) Surface sequence conservation of BsKKT2 Znf1 using the ConSurf server (Landau et al., 2005; Ashkenazy et al., 2016). Znf2 structure is shown as cartoon representation.

(B) Electrostatic surface potential of BsKKT2 Znf1 generated by APBS (Jurrus et al., 2018) reveals the presence of a conserved acidic surface in Znf1. The location of the residue D626 is marked by a black circle.

To test the importance of the TbKKT2 central domain for cell viability, we performed rescue experiments. We replaced one allele of TbKKT2 with an N-terminally YFP-tagged TbKKT2 construct that has either wild-type or mutant versions of the central domain, and performed RNAi against the 5’UTR of the TbKKT2 transcript to knock down the untagged allele of TbKKT2 (Figure S4) (Ishii and Akiyoshi, 2019). As expected, mutants in Znf1 (C576A and D622A) and Znf2 (C640A C643A) both localized at kinetochores (Figure 6C). Upon induction of RNAi, however, the Znf1 mutants failed to support normal growth after day 4, while the Znf2 mutants rescued the growth defects. These data confirm the importance of Znf1 for the function of TbKKT2 in vivo.

Figure S4.
Figure S4. RNAi of KKT2 and KKT3 causes growth defects in T. brucei

Growth curve of (A) KKT2 5’UTR RNAi, (B) KKT2-YFP, KKT2 5’UTR RNAi, (C) YFP-KKT2, KKT2 5’UTR RNAi, (D) KKT3 3’UTR RNAi, (E) YFP-KKT3, KKT3 3’UTR RNAi, and (F) KKT3-YFP, KKT3 3’UTR RNAi. 1 µg/mL doxycycline was added to induce RNAi. Controls are uninduced cell cultures. Similar results were obtained from at least two independent experiments. Cell lines, BAP1554, BAP1752, BAP1681, BAP1555, BAP1659, BAP1682.

Localization of KKT3 depends on the central domain in T. brucei

While TbKKT2 has multiple domains that can promote centromere localization, TbKKT3 has only the central domain that was able to localize at kinetochores (Figure 1). Consistent with this result, mutating TbKKT3 residues that align with Zn-coordinating histidine or cysteine residues in the Znf1 of KKT2 abolished the kinetochore localization of the ectopically-expressed full length TbKKT3 protein (Figure 7A, B). We also found that the conserved aspartic acid TbKKT3 D692 was essential for kinetochore localization. In contrast, mutations in TbKKT3 Znf2 (C707A C710A) did not affect the kinetochore localization.

Figure 7.
Figure 7. Kinetochore localization of KKT3 depends on the central domain in T. brucei

(A) Multiple sequence alignment of KKT3. Residues mutated in (B) or (C) are shown.

(B) Percentage of GFP-positive cells that have kinetochore-like dots were quantified at 1 day post-induction (n > 22, each). Inducible GFP-NLS fusion proteins were expressed with 10 ng/Ml doxycycline. Cell lines, BAP291, BAP359, BAP360, BAP446, BAP447, BAP1721, BAP1722, BAP362, BAP341.

(C) TbKKT3 C668A/C671A and D692A mutants do not localize at kinetochores and fail to support normal cell growth, while TbKKT3 C707A/C710A mutant is functional. One allele of TbKKT3 was mutated and tagged with a C-terminal YFP tag, and the other allele was depleted using RNAi-mediated knockdown by targeting the 3’UTR of the TbKKT3 transcript. Top: cells were diluted every two days and cell growth was monitored for 8 days upon induction of RNAi. Similar results were obtained for at least three clones of TbKKT3 mutants. Controls are uninduced cell cultures. Bottom: Example of cells expressing the TbKKT3 mutants prior to RNAi induction, showing that TbKKT3C668A C671A and TbKKT3D692A do not localize at kinetochores while TbKKT3C707A C710A localizes normally (n > 90, each). Maximum intensity projection are shown. RNAi was induced with 1 µg/mL doxycycline. Cell lines, BAP1791, BAP1793, BAP1783. Scale bars, 5 μm.

We next performed rescue experiments by replacing one allele of TbKKT3 with a C-terminally YFP-tagged construct that has either wild-type or mutant versions of the central domain, and performed RNAi against the 3’UTR of TbKKT3 to knock down the untagged allele of TbKKT3 (Figure S4). We first confirmed that TbKKT3 Znf1 mutants (C668A C671A and D692A) were unable to localize at kinetochores, while the TbKKT3 Znf2 mutant (C707A C710A) localized normally. Upon induction of RNAi, Znf1 mutants failed to rescue the growth defect, showing that kinetochore localization is essential for the TbKKT3 function (Figure 7C). In contrast, the TbKKT3 Znf2 mutant supported normal cell growth. These data show that the TbKKT3 Znf1 zinc finger is essential for the localization and function of TbKKT3, similarly to what was observed for TbKKT2 Znf1.

Discussion

A major open question concerning the unconventional kinetochores of kinetoplastids is how these organisms specify kinetochore positions without CENP-A. In other words, how do kinetochore proteins localize specifically at centromeres in a sequence-independent manner? Towards addressing this important question, we examined two constitutively localized kinetochore kinases KKT2 and KKT3, which have sequence similarity with polo-like kinases (Nerusheva and Akiyoshi, 2016). In addition to an N-terminal protein kinase domain and a C-terminal polo box domain present in polo-like kinases, KKT2 and KKT3 have a central domain that is highly conserved among kinetoplastids. By expressing fragments of KKT2 and KKT3 in trypanosomes, we first established that their central domains can localize specifically at centromeres. The crystal structure of the Bodo saltans KKT2 central domain then revealed a unique structure, which consists of two distinct zinc fingers (Znf1 and Znf2). It is likely that the central domain of T. brucei KKT2 has a similar structure based on high sequence similarity between BsKKT2 and TbKKT2 proteins. Importantly, mutational analyses of KKT2 in T. brucei revealed that Znf1 was important for the localization of the central domain, while Znf2 was not. Furthermore, although full-length TbKKT2 Znf1 mutants localized at kinetochores (likely due to interactions with other kinetochore proteins via other domains), they were not fully functional. Taken together, these data have established that Znf1 is essential for the function of TbKKT2, which is supported by the presence of Znf1, but not Znf2, in Perkinsela KKT2a. We speculate that Znf2 has affinity for DNA and stabilizes the localization of KKT2 at centromeres in T. brucei and Bodo saltans. In agreement with this hypothesis, putative DNA-binding SPKK motifs (Suzuki, 1989) are present right after Znf2 in many kinetoplastids (Figure 6A), while Perkinsela KKT2b has an AT-hook motif within the Znf1 domain (Figure S2). It remains unclear whether the structure of the central domain is conserved between KKT2 and KKT3. Nonetheless, our functional studies of TbKKT3 mutants showed that the Znf1 equivalent domain was also essential for the kinetochore localization and function, while the Znf2 equivalent domain was not, showing that the functional importance of Znf1 is conserved in KKT3. It will be important to obtain KKT3 central domain structures to reveal structural similarity or difference between KKT2 and KKT3.

The fact that KKT2/3 central domains manage to localize specifically at centromeres suggests that they are able to recognize something special at centromeres. What might be a unique feature at centromeres in kinetoplastids that lack CENP-A? Histone variants are one possibility. T. brucei has four histone variants, H2AZ, H2BV, H3V, and H4V. However, none of them is specifically enriched at centromeres (Lowell and Cross, 2004; Lowell et al., 2005; Siegel et al., 2009), and histone chaperones did not co-purify with any kinetochore protein (Akiyoshi and Gull, 2014). Alternatively, there might exist certain post-translational modifications on histones or DNA specifically at centromeres (e.g. phosphorylation, methylation, or acetylation). KKT2 Znf1 has a highly conserved acidic patch, which might act as a “reader” for such modifications. Although there is no known histone or DNA modification that occurs specifically at centromeres, KKT2/3 have a protein kinase domain and KKT23 has a Gcn5-related N-acetyltransferase (GNAT) domain (Nerusheva et al., 2019). It will be important to examine whether these enzymatic domains are important for recruiting the central domain and identify their substrates. Another unique feature at centromeres is the presence of kinetochore proteins, which could potentially recruit newly synthesized kinetochore components by direct protein-protein interactions. Finally, it is important to note that it remains unclear whether kinetoplastid kinetochores build upon nucleosomes. It is formally possible that the KKT2/3 central domains directly bind DNA and form a unique environment at centromeres. Understanding how the KKT2/3 central domains localize specifically at centromeres will be key to elucidating the mechanism of how kinetoplastids specify kinetochore positions in the absence of CENP-A.

Materials and Methods

Tryp cells and plasmids, microscopy, immunoprecipitation, and mass spectrometry

All trypanosome cell lines, plasmids, primers, and synthetic DNA used in this study are listed in Table S4. All trypanosome cell lines used in this study were derived from T. brucei SmOxP927 procyclic form cells (TREU 927/4 expressing T7 RNA polymerase and the tetracycline repressor to allow inducible expression) (Poon et al., 2012). Cells were grown at 28 °C in SDM-79 medium supplemented with 10% (v/v) heat-inactivated fetal calf serum (Brun and Schönenberger, 1979). Endogenous YFP tagging was performed using the pEnT5-Y vector (Kelly et al., 2007). Inducible expression of GFP-NLS fusion proteins or hairpin RNAi was performed using pBA310 (Nerusheva and Akiyoshi, 2016).

To make pBA1711 (KKT3 3’UTR hairpin RNAi), BAG95 was digested with HindIII/BamHI and subcloned into the pBA310 HindIII/BamHI sites. To make pBA1807, 4–3174bp of KKT3 coding sequence and 250bp of 3’UTR were amplified with primers BA2351/BA2352 and BA2353/BA2354, digested with HindIII/NotI and NotI/SpeI, respectively, and were cloned into the pEnT5-Y using HindIII/SpeI sites. Site-directed mutagenesis was performed using primers and template plasmids listed in Table S4. All constructs were sequence verified.

Plasmids linearized by NotI were transfected into trypanosomes by electroporation into an endogenous locus (pEnT5-Y derivatives) or 177bp repeats on minichromosomes (pBA310 derivatives). To obtain endogenously-tagged strains (pEnT5-Y derivatives), transfected cells were selected by the addition of 25 µg/ml hygromycin and cloned by dispensing dilutions into 96-well plates. Clones that express mutant versions of KKT2 or KKT3 were screened by Sanger sequencing of genomic DNA. pBA310 derivatives were selected by the addition of 5 µg/ml phleomycin. Expression of GFP-NLS fusion proteins (pBA310 derivatives) was induced by the addition of doxycycline (10 ng/ml). RNAi was induced by the addition of doxycycline (1 µg/ml).

Fluorescence microscopy, immunoprecipitation of GFP-fusion proteins, and mass spectrometry were performed essentially as described previously (Nerusheva and Akiyoshi, 2016; Ishii and Akiyoshi, 2019). Proteins identified with at least two peptides were considered as significant and shown in Table S1.

Multiple sequence alignment

Protein sequences and accession numbers for KKT2 and KKT3 homologs were retrieved from TriTryp database (Aslett et al., 2010), Wellcome Sanger Institute (https://www.sanger.ac.uk/), or UniProt (UniProt Consortium, 2019). Search for KKT2/3 homologs in Perkinsela was done using hmmsearch on its predicted proteome using manually prepared KKT2/3 hmm profiles (HMMER version 3.0 (Eddy, 1998)). Multiple sequence alignment was performed with MAFFT (L-INS-i method, version 7) (Katoh and Standley, 2013) and visualized with the Clustalx coloring scheme in Jalview (version 2.10) (Waterhouse et al., 2009).

Protein expression and purification

Multiple sequence alignment together with secondary structure predictions of the KKT2 central domain were used to design constructs in Bodo saltans and Perkinsela. To make pBA1660 (BsKKT2572–668 with an N-terminal TEV-cleavable hexahistidine (His6) tag), the central domain of Bodo saltans KKT2 (accession number BSAL_50690) was amplified from BAG50 (a synthetic DNA that encodes Bodo saltans KKT2, codon optimized for expression in Sf9 insect cells (Table S4)) with primers BA2117/BA2118 and cloned into the RSFDuet-1 vector (Novagen) using BamHI/EcoRI sites with an NEBuilder HiFi DNA Assembly Cloning Kit (NEB) according to the manufacturer’s instructions. To make pBA1139 (His6-PkKKT2551–679), the central domain of Perkinsela CCAP 1560/4 KKT2a (accession number XU18_4017) was amplified from BAG48 (a synthetic DNA that encodes Perkinsela KKT2a, codon optimized for expression in Sf9 insect cells (Table S4)) with primers BA1569/BA1570 and cloned into RSFDuet-1 using BamHI/EcoRI sites with an In-Fusion HD Cloning Plus kit (Takara Bio). Recombinant proteins were expressed in BL21(DE3) E. coli cells at 20 °C using auto induction media (Formedium) (Studier, 2005).

Briefly, 500 mL of cells were grown at 37 °C in 2.5 L flasks at 300 rpm until OD600 of 0.2–0.3 and then cooled down to 20 °C overnight (2 L for BsKKT2 and 6 L for PkKKT2a). Cells were harvested by centrifugation and resuspended in 50 ml per litre of culture of lysis buffer (25 mM Hepes pH 7.5, 150 mM NaCl, 1 mM TCEP, 10 mM imidazole, and 1.2 mM PMSF). Proteins were extracted by mechanical cell disruption using a French press (1 passage at 20,000 PSI) and the resulting lysate was centrifuged at 48,384 g for 30 min at 4 °C. Clarified lysate was incubated with 5 mL TALON beads (Takara Bio), washed with 150 mL lysis buffer and eluted in 22 mL of elution buffer (25 mM Hepes pH 7.5, 150 mM NaCl, 1 mM TCEP, and 250 mM imidazole) in a gravity column, followed by TEV treatment for the removal of the His6 tag. Salt concentration of the sample was subsequently reduced to 50 mM NaCl using buffer A (50 mM Hepes pH 7.5, and 1 mM TCEP) and the sample was loaded onto a 5 mL HiTrap Heparin HP affinity column (GE healthcare) pre-equilibrated with 5% buffer B (50 mM Hepes pH 7.5, 1 M NaCl, and 1 mM TCEP) on an ÄKTA pure 25 system. Protein was eluted by using a gradient from 0.05 to 1 M NaCl, and protein-containing fractions were combined, concentrated with an Amicon stirred cell using an ultrafiltration disc with 10 kDa cut-off (Merck), and then loaded onto a HiPrep Superdex 75 16/60 size exclusion chromatography column (GE healthcare) pre-equilibrated with 25 mM Hepes pH 7.5, 150 mM NaCl, and 1 mM TCEP. Fractions containing the KKT2 central domain were pooled together, concentrated with an Amicon stirred cell using an ultrafiltration disc with 10 kDa cut-off, and stored at -80 °C. Protein concentration was measured by Bradford assay.

Crystallization

Both BsKKT2 and PkKKT2a central domain crystals were optimized at 4 °C in sitting drop vapour diffusion experiments in 48-well plates, using drops of overall volume 400 nL, mixing protein and mother liquor in a 3:1 protein:mother liquor ratio. BsKKT2 central domain crystals grew from the protein at 26 mg/mL and mother liquor 40% PEG 400, 0.2 mM (NH4)2SO4, and 100 mM Tris-HCl pH 8. The 40% PEG400 in the mother liquor served as the cryoprotectant when flash-cooling the crystals by plunging into liquid nitrogen. PkKKT2a central domain crystals grew from the protein at 13 mg/mL and mother liquor 19% MPD, 50 mM Hepes pH 7.5, and 10 mM MgCl2. The crystals were briefly transferred into a cryoprotecting solution of 30% MPD, 50 mM Hepes pH 7.5, 10 mM MgCl2 prior to flash-cooling.

Data collection and structure determination

X-ray diffraction data from a BsKKT2 central domain crystal were collected at the I04 beamline at the Diamond Light Source (Harwell, UK) at the Zinc K-edge wavelength (λ=1.28297 Å). A set of 1441 images were processed in space group I222 using the Xia2 pipeline (Winter, 2010), with DIALS for indexing and integration (Winter et al., 2018) and AIMLESS for scaling (Evans and Murshudov, 2013) to 1.8 Å resolution. Initial 3 Zn atoms were localized by interpreting the anomalous difference Patterson, single anomalous dispersion (SAD) phases were estimated using Crank2 (Skubák and Pannu, 2013), and an initial model was built with BUCCANEER (Cowtan, 2006). The structure was completed by several cycles of alternating model building in Coot (Emsley et al., 2010) and refinement in autoBUSTER (Blanc et al., 2004; Bricogne et al., 2017).

PkKKT2a central domain X-ray diffraction data were collected at the I03 beamline at Diamond Light Source (Harwell, UK) also at the zinc K-edge (λ=1.28272 Å) and processed using the autoPROC pipeline (Vonrhein et al., 2011) using XDS (Kabsch, 2010) for indexing/integration and AIMLESS (Evans and Murshudov, 2013) for scaling, to a resolution of 3.8 Å. Two initial Zn positions were determined by interpreting the anomalous difference Patterson, and single anomalous dispersion (SAD) phases were estimated using Crank2 (Skubák and Pannu, 2013) and SHARP (Vonrhein et al., 2007) in space group P64. An initial model was manually built in Coot and refined once with RosettaMR (Terwilliger et al., 2012). The structure was completed by several cycles of alternating model building in Coot (Emsley et al., 2010) and refinement in autoBUSTER (Blanc et al., 2004; Bricogne et al., 2017).

A higher resolution dataset was collected from a PkKKT2a central domain crystal at the I24 beamline at Diamond Light Source (Harwell, UK), at a wavelength of λ=0.9686 Å. Data were processed using Xia2 pipeline (Winter, 2010), DIALS (Winter et al., 2018), and AIMLESS (Evans and Murshudov, 2013) in space group P64 to a resolution of 2.9 Å. The model obtained from the 3.8 Å dataset was used for further model building and refinement with autoBuster (Bricogne et al., 2017) and Coot (Emsley et al., 2010).

All the images were made with Pymol (Schrödinger LLC, Portland, OR) and CCP4mg (McNicholas et al., 2011). Topology diagrams were generated using TopDraw (Bond, 2003). Protein coordinates have been deposited in the RCSB Protein Data Bank (http://www.rcsb.org/) with accession codes 6TLY (Bodo saltans KKT2) and 6TLX (Perkinsela KKT2a).

Author contributions

Gabriele Marcianò purified recombinant proteins and solved crystal structures. Olga Nerusheva expressed KKT2 and KKT3 truncations and mutants in trypanosomes and performed immunoprecipitation and mass spectrometry. Midori Ishii performed rescue experiments. Bungo Akiyoshi expressed KKT2 and KKT3 truncations and mutants in trypanosomes. Gabriele Marcianò, Midori Ishii, and Bungo Akiyoshi wrote the manuscript.

Data availability

All data generated during this study are included in the manuscript and supplementary data. Protein coordinates have been deposited in the RCSB Protein Data Bank (http://www.rcsb.org/) with accession codes 6TLY (Bodo saltans KKT2) and 6TLX (Perkinsela KKT2a).

Acknowledgments

We thank Pietro Roversi and Matt Higgins for advice, crystallography facility manager Edward Lowe, as well as Pietro Roversi, Matt Higgins, Danny Huang, and Patryk Ludzia for comments on the manuscript. Midori Ishii was supported by a long-term fellowship from the TOYOBO Biotechnology Foundation. Bungo Akiyoshi was supported by a Wellcome Trust Senior Research Fellowship (grant no. 210622/Z/18/Z) and the European Molecular Biology Organization Young Investigator Program. The authors declare that no competing interests exist.

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Posted December 13, 2019.
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Supplementary Material

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Unconventional kinetochore kinases KKT2 and KKT3 have a unique zinc finger that promotes their kinetochore localization
Gabriele Marcianò, Olga O. Nerusheva, Midori Ishii, Bungo Akiyoshi
bioRxiv 2019.12.13.875419; doi: https://doi.org/10.1101/2019.12.13.875419
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