Introduction

During cell division, duplicated genetic material must be distributed equally into two daughter cells. The Aurora B kinase is a key mitotic regulator widely conserved among eukaryotes (Hochegger et al., 2012). It undergoes dynamic localization changes throughout mitosis to enable the spatially restricted phosphorylation of substrates involved in chromosome alignment, chromosome bi-orientation, spindle assembly checkpoint (SAC) signalling, and cytokinesis (Carmena et al., 2012). In early mitosis, Aurora B is first detected on chromosome arms and during prometaphase becomes enriched at centromeres, where it destabilizes incorrect kinetochore-microtubule attachments (Krenn and Musacchio, 2015). Upon anaphase onset, Aurora B translocates to the spindle midzone, and during cytokinesis associates with the equatorial cortex to regulate cell abscission (Adams et al., 2000; Cooke et al., 1987; Trivedi and Stukenberg, 2016).

The dynamic localization pattern of the Aurora B kinase is in part achieved through its association with a scaffold comprised of inner centromere protein (INCENP), Borealin and Survivin (Adams et al., 2001, 2000; Gassmann et al., 2004; Romano et al., 2003; Sampath et al., 2004; Vader et al., 2006; Wheatley et al., 2001). Together, these proteins form a tetrameric complex referred to as the chromosomal passenger complex (CPC). The CPC can be partitioned into two functional modules: The ‘catalytic module’ and the ‘localization module’. The catalytic module is composed of Aurora B in complex with the IN-box at the INCENP C-terminus, which is required for full activation of the Aurora B kinase (Bishop and Schumacher, 2002). The localization module comprises Borealin, Survivin and the N-terminus of INCENP, which are connected to one another via a three-helical bundle (Jeyaprakash et al., 2011, 2007; Klein et al., 2006). The two modules are linked by the central region of INCENP, composed of an intrinsically disordered domain and a single alpha helical (SAH) domain. INCENP harbors microtubule-binding domains within the N-terminus and the central SAH domain, which play key roles for CPC localization and function (Cormier et al., 2013; Fink et al., 2017; Kang et al., 2001; Mackay et al., 1993; Nakajima et al., 2011; Noujaim et al., 2014; Samejima et al., 2015; van der Horst et al., 2015; Wheatley et al., 2001; Wheelock et al., 2017).

In vertebrates, recruitment of the CPC to centromeric chromatin depends on two pathways, involving the Haspin and Bub1 kinases. Haspin phosphorylates histone H3 on Thr3 (H3T3ph), which is recognized by the baculovirus IAP repeat (BIR) domain of Survivin (Kelly et al., 2010; Wang et al., 2010; Yamagishi et al., 2010). H3T3ph is initially found along the entire length of chromosomes between sister chromatids but becomes enriched at the inner centromere (the space between sister kinetochores) during late prophase. In contrast, the kinetochore-associated Bub1 kinase phosphorylates histone H2A on Thr120 (H2AT120ph) (Kawashima et al., 2010). H2AT120ph recruits Shugoshin-like proteins (Sgo1 and Sgo2), which in turn are bound by Borealin (Tsukahara et al., 2010; Yamagishi et al., 2010). Recently, Sgo1 has also been demonstrated to interact with the BIR domain of Survivin through an N-terminal histone H3-like motif (Abad et al., 2022; Jeyaprakash et al., 2011). The interactions of Borealin and Survivin with Sgo1 form the basis for a kinetochore-proximal pool of the CPC which is distinct from the inner centromere pool (Broad et al., 2020; Hadders et al., 2020; Liang et al., 2020).

In most studied eukaryotes, ranging from yeast to humans, kinetochore assembly is scaffolded by a centromere-specific histone H3 variant, CENP-A (Allshire and Karpen, 2008; Black and Cleveland, 2011; Hori and Fukagawa, 2012; Maddox et al., 2012; Westhorpe and Straight, 2013). An assembly of inner kinetochore protein complexes, referred to as the constitutive centromere-associated network (CCAN), interacts with centromeric CENP-A chromatin throughout the cell cycle and provides a platform for recruitment of the outer kinetochore KNL1/Mis12 complex/Ndc80 complex (KMN) network that has microtubule-binding activity (Cheeseman et al., 2006; Foltz et al., 2006; Izuta et al., 2006; Okada et al., 2006). Some of these kinetochore proteins are present in nearly all sequenced eukaryotes, suggesting that key principles of chromosome segregation are widely shared among eukaryotes (Drinnenberg and Akiyoshi, 2017; Van Hooff et al., 2017; Meraldi et al., 2006; Tromer et al., 2019). However, a unique set of kinetochore proteins (KKT1–20, KKT22–25, KKIP1–12) are present in the evolutionarily divergent lineage called kinetoplastids (Akiyoshi, 2020; Akiyoshi and Gull, 2014; D’Archivio and Wickstead, 2017; Nerusheva et al., 2019; Nerusheva and Akiyoshi, 2016). Kinetoplastids are flagellated protists that are highly divergent from commonly studied eukaryotes (Cavalier-Smith, 2010). Trypanosoma brucei, Trypanosoma cruzi, and Leishmania spp. are causative agents of African trypanosomiasis, Chagas disease, and leishmaniasis, respectively, and as such pose a serious threat to public health and prosperity across the tropics and subtropics (Stuart et al., 2008; WHO, 2017).

Despite the absence of canonical kinetochore components (Berriman et al., 2005; Lowell and Cross, 2004), Aurora kinases are conserved in kinetoplastids. Early studies suggested that the Aurora B homolog (Aurora B^AUK1) in T. brucei forms a complex with chromosomal passenger complex proteins 1 and 2 (CPC1 and CPC2) and plays a crucial role in mitosis and cytokinesis (Li et al., 2008; Tu et al., 2006). CPC1 was later found to be a divergent INCENP homolog (hereafter referred to as INCENP^CPC1) based on the presence of a conserved C-terminal IN-box (Hu et al., 2014). However, INCENP^CPC1 lacks the central SAH domain and N-terminal residues, which in other eukaryotes interact with Survivin and Borealin. In addition, two orphan kinesins, KIN-A and KIN-B, have been proposed to transiently associate with Aurora B^AUK1 during mitosis (Li, 2012; Li et al., 2008). Although homologs of the ‘localization module’ proteins Survivin and Borealin have not been identified in kinetoplastids (Komaki et al., 2022), the trypanosome CPC displays a dynamic localization pattern similar to that of the metazoan CPC (Li et al., 2008): Aurora B^AUK1, INCENP^CPC1 and CPC2 localize to kinetochores in early mitosis and then translocate to the central spindle upon anaphase onset. From late anaphase onwards, an additional population of CPC proteins is detectable at the tip of the new flagellum attachment zone (FAZ), the point of cytokinesis initiation in T. brucei. It is presently not understood how the CPC assembles in these evolutionarily divergent eukaryotes nor how its localization dynamics are regulated during the cell cycle.

Here, by combining biochemical, structural and cell biological approaches in procyclic form T. brucei, we show that the trypanosome CPC is a pentameric complex comprising Aurora B^AUK1, INCENP^CPC1, CPC2, and the two orphan kinesins KIN-A and KIN-B. KIN-A and KIN-B interact via their coiled-coil domains to form a subcomplex within the CPC, which serves as a scaffold for the catalytic module (Aurora B^AUK1 + INCENP^CPC1). The C-terminal unstructured tail of KIN-A directs kinetochore localization of the CPC from S phase to metaphase, while the N-terminal motor domain promotes the central spindle enrichment in anaphase. Furthermore, we identify the KKT7 – KKT8 complex pathway as the main kinetochore recruitment arm of the trypanosome CPC.

Results

KIN-A and KIN-B are bona fide CPC proteins in trypanosomes

To identify additional interactors of the CPC in trypanosomes, we performed immunoprecipitation followed by liquid chromatography tandem mass spectrometry (IP-MS) of endogenously YFP-tagged Aurora B^AUK1 (Fig. S1A, Table S1 and Table S2). Besides Aurora B^AUK1, INCENP^CPC1 and CPC2, we observed notable enrichment of two orphan kinesins, KIN-A and KIN-B (Wickstead and Gull, 2006), as reported previously (Li et al., 2008). Both KIN-A and KIN-B were also highly enriched in immunoprecipitates of ectopically expressed GFP-INCENP^CPC1. Vice versa, IP-MS of GFP-tagged KIN-A and KIN-B identified Aurora B^AUK1, INCENP^CPC1, and CPC2 as top hits (Fig. 1A and Table S2).

Figure 1.

We next assessed the localization dynamics of fluorescently tagged KIN-A and KIN-B over the course of the cell cycle (Figs. 1, B-E). T. brucei possesses two DNA-containing organelles, the nucleus (‘N’) and the kinetoplast (‘K’). The kinetoplast is an organelle found uniquely in kinetoplastids, which contains the mitochondrial DNA and replicates and segregates prior to nuclear division. The ‘KN’ configuration serves as a good cell cycle marker (Siegel et al., 2008; Woodward and Gull, 1990). To our surprise, KIN-A-YFP and GFP-KIN-B exhibited a CPC-like localization pattern similar to that of Aurora B^AUK1: Both kinesins localized to kinetochores from S phase to metaphase, and then translocated to the central spindle in anaphase (Figs. 1, C-E). Moreover, like Aurora B^AUK1, a population of KIN-A and KIN-B localized at the new FAZ tip from late anaphase onwards (Figs. S1, B and C). This was unexpected, because KIN-A and KIN-B were previously reported to localize to the spindle but not to kinetochores or the new FAZ tip (Li et al., 2008). These data suggest that KIN-A and KIN-B are bona fide CPC proteins in trypanosomes, associating with Aurora B^AUK1, INCENP^CPC1, and CPC2 throughout the cell cycle.

A bioinformatic search for homologs of CPC proteins within Euglenozoa revealed that both KIN-A and KIN-B are present in trypanosomatids and bodonids, with KIN-A homologs detectable even in prokinetoplastids (Fig. 1F) (Materials and Methods). CPC2, on the other hand, was detectable only within trypanosomatids. Aurora B^AUK1 and INCENP^CPC1 are present in kinetoplastids as well as in diplonemids and euglenids (sister groups of kinetoplastids). Interestingly, homologs of Borealin, Survivin, KIN-A, or KIN-B were not detectable in diplonemids or euglenids, raising a possibility that these organisms may also possess ‘non-canonical’ CPC proteins. We conclude that the KIN-A and KIN-B kinesins are highly conserved within kinetoplastids, constituting integral components of the CPC in this evolutionary divergent group of eukaryotes.

KIN-A and KIN-B promote kinetochore localization of the CPC

To investigate which subunits of the CPC are responsible for its kinetochore targeting, we performed a series of RNAi experiments (Figs. 2, A-E). Because CPC2 is poorly conserved among kinetoplastids (Fig. 1F) and depletion of CPC2 using two different hairpin RNAi constructs (Table S1) was inefficient, we did not include CPC2 in these experiments. As previously reported (Li et al., 2008), depletion of Aurora B^AUK1, INCENP^CPC1, KIN-A or KIN-B resulted in a prominent growth defect (Fig. S2A) with cells arresting in G2/M (2K1N) (Fig. S2B). Knockdown of Aurora B^AUK1 did not affect the kinetochore localization of YFP-tagged INCENP^CPC1, KIN-A or KIN-B (Figs. 2, A and C-E). Knockdown of INCENP^CPC1 caused delocalization of Aurora B^AUK1 but not of KIN-A or KIN-B (Figs. 2, A and B, D and E). In contrast, both Aurora B^AUK1 and INCENP^CPC1 were delocalized upon depletion of KIN-A or KIN-B (Figs. 2, A-C). RNAi against KIN-A disrupted KIN-B localization and vice versa (Figs. 2, A, D and E). Moreover, total protein levels of KIN-B were affected by depletion of KIN-A (Fig. S2C), suggesting that the interaction with KIN-A is required to stabilize KIN-B.

Figure 2.

We next ectopically expressed GFP-tagged truncations of KIN-A and KIN-B to assess which domains promote their kinetochore targeting. Both KIN-A and KIN-B contain an N-terminal kinesin motor domain followed by several predicted coiled-coil motifs (Li et al., 2008), although KIN-B is predicted to be an inactive motor (Wickstead and Gull, 2006). In addition, KIN-A has a long C-terminal tail. Unlike full-length KIN-B, KIN-B^2-316 (inactive motor domain) failed to enrich at kinetochores and was instead found in the nucleolus (Fig. S2, D and E). KIN-A^2-309 (motor domain) was also primarily detected in the nucleolus, although we observed additional spindle and weak kinetochore-like signal in some metaphase cells (Fig. 2G). By contrast, both KIN-A^310-862 (coiled-coil domain + C-terminal disordered tail) and KIN-B^317-624 (coiled-coil domain) clearly localized to kinetochores from S phase to metaphase (Fig. 2H; Fig. S2F). Intriguingly, unlike endogenously YFP-tagged KIN-A, ectopically expressed GFP fusions of both full-length KIN-A and KIN-A^310-862 localized at kinetochores even in anaphase (Figs. 2, F and H). Weak anaphase kinetochore signal was also detectable for KIN-B^317-624 (Fig. S2F). GFP fusions of the central coiled-coil domain or the C-terminal disordered tail of KIN-A did not localize to kinetochores (data not shown). These results show that kinetochore localization of the CPC is mediated by KIN-A and KIN-B and requires both the central coiled-coil domain as well as the C-terminal disordered tail of KIN-A.

Structural model of the trypanosome CPC

To gain structural insights into the trypanosome CPC, we used AlphaFold2 (AF2) (Jumper et al., 2021; Mirdita et al., 2022) to predict the overall structure of the trypanosome CPC by testing combinations of full-length Aurora B^AUK1, INCENP^CPC1, CPC2, KIN-A, and KIN-B, and truncations thereof (Figs. S3, A-D). AF2 confidently predicted parallel coiled coils between KIN-A and KIN-B, with the main region of interaction contained within the central region of KIN-A (residues ∼310–550) and the C-terminal region of KIN-B (residues ∼320–580). The C-terminal tail (residues ∼550–862) of KIN-A is predicted to be intrinsically disordered (pLDDT scores <20, Figs. S3, A and B). The first ∼55 residues of INCENP^CPC1, predicted to form two α-helices (residues ∼7–20 and ∼36–55), interact with the coiled coils of KIN-A:KIN-B in close proximity to the kinesin motors. A flexible central linker in INCENP^CPC1 bridges the N-terminus of INCENP^CPC1 and the catalytic module of the CPC (Aurora B^AUK1 + INCENP^CPC1 IN-box). Similarly to INCENP^CPC1, an N-terminal α-helical region in CPC2 (residues ∼19–75) interacts with the KIN-A:KIN-B coiled coils immediately downstream of the INCENP^CPC1 binding site. Consistent with these predictions, GFP-tagged INCENP^{CPC1 2-147} or CPC2^2-120 displayed normal localization dynamics indistinguishable from the corresponding full-length constructs (Figs. S3, E-F and H-I). In contrast, deletion of the N-terminal domains of INCENP^CPC1 or CPC2 impaired kinetochore localization (Figs. S3, G and J). Together, these data suggest that Aurora B^AUK1 forms a subcomplex with the C-terminus of INCENP^CPC1, and that INCENP^CPC1 and CPC2 interact with the coiled-coil domain of KIN-A:KIN-B via their N-terminal domains.

To validate these findings biochemically, we performed bis(sulfosuccinimidyl) suberate (BS³) cross-linking of native CPC complexes isolated by immunoprecipitation of endogenously tagged YFP-Aurora B^AUK1 from cells arrested prior to anaphase, followed by mass spectrometry analysis (IP-CLMS) (Fig. 3A and Table S2). Indeed, our IP-CLMS data revealed high-score crosslinks between the predicted coiled-coil domains of KIN-A and KIN-B, suggesting that the two kinesins form a parallel heterodimer. As expected, Aurora B^AUK1 formed contacts with the C-terminal IN-box of INCENP^CPC1, consistent with these two proteins constituting the catalytic module of the CPC. The N-terminal region of INCENP^CPC1 interacted mainly with KIN-B and to a lesser degree with KIN-A, with most contacts confined to the N-terminal ends of the predicted coiled-coil domains of the kinesins close to their motor domains. The N-terminus of CPC2, on the other hand, formed crosslinks with the coiled-coil domain of KIN-A. We used PyXLinkViewer (Schiffrin et al., 2020) to map our IP-CLMS data onto the assembled AF2 model of the trypanosome CPC (Fig. 3B). Using a Euclidian distance cut-off of 30 Å, ∼85% of crosslinks were compatible with the model, providing confidence in the AF2 predictions. The few crosslinks that violated the distance constraints mainly represent intra-protein contacts between the INCENP^CPC1 N-and C-terminal domains or inter-protein contacts between the INCENP^CPC1 C-terminal domain and the kinesin motor domain of KIN-A. The central domain of INCENP^CPC1 is predicted to be unstructured (pLDDT scores <20, Figs. S3, C-D) and may act as a flexible linker, permitting multiple orientations of the catalytic module relative to the KIN-A:KIN-B scaffold. Taken together, these data indicate that KIN-A and KIN-B interact via their coiled-coil domains, which serve as a scaffold for the assembly of the remaining CPC subunits.

Figure 3.

The CPC is recruited to kinetochores through the KKT7 – KKT8 complex pathway

Core components of the Haspin-H3T3ph and Bub1-H2AT120ph-Sgo1 pathways that control CPC recruitment to the centromere in other model eukaryotes are not found in kinetoplastids (Berriman, 2005), and so far, no centromere-specific histone modifications and/or histone variants have been uncovered in T. brucei. We reasoned that the centromere receptor(s) of the trypanosome CPC may lie within the repertoire of unconventional kinetochore proteins present in kinetoplastids. Our IP-CLMS approach failed to detect crosslinks between CPC subunits and kinetochore components (Table S2), possibly due to the transient nature of these interactions. Nevertheless, several KKT proteins were commonly enriched in the immunoprecipitates of Aurora B^AUK1, CPC1, KIN-A and KIN-B, the most abundant ones being KKT6, KKT7, KKT8, KKT9, KKT10, KKT11 and KKT12 (Fig. 1A; Fig. S1A and Table S2). KKT7 is detected at kinetochores from S phase until the end of anaphase and recruits the KKT8 complex (comprising KKT8, KKT9, KKT11 and KKT12) (Akiyoshi and Gull, 2014; Ishii and Akiyoshi, 2020). The KKT8 complex localizes at kinetochores from S phase and dissociates at the metaphase-anaphase transition.

Using previously validated RNAi constructs (Akiyoshi and Gull, 2014; Llauró et al., 2018; Marcianò et al., 2021), we found that knockdown of KKT7 or KKT9 resulted in dispersal of YFP-Aurora B^AUK1 from kinetochores in ∼70% of cells (Figs. 4, A-D). In contrast, depletion of KKT1, KKT2, KKT3, KKT4, KKT6, KKT10/19, KKT14 and KKT16 had little or no effect on YFP-Aurora B^AUK1 localization (Figs. S4, A and B). KIN-A-YFP was also lost from kinetochores upon RNAi-mediated depletion of KKT8 complex subunits (Figs. S4, C and D). We next tested whether KKT7 or the KKT8 complex were able to recruit Aurora B^AUK1 to an ectopic locus using the LacI-LacO system (Landeira and Navarro, 2007). For these experiments, we expressed GFP tagged KKT7^2-261 or KKT8 fused to the Lac repressor (LacI) in trypanosomes containing an ectopic Lac operator (LacO) array stably integrated into rDNA repeats. We previously showed that KKT7 lies upstream of the KKT8 complex (Ishii and Akiyoshi, 2020). Indeed, GFP-KKT7^2-261-LacI recruited tdTomato-KKT8, -KKT9 and -KKT12 (Fig. S4E). Expression of GFP-KKT7^2-261-LacI or GFP-KKT8-LacI resulted in robust recruitment of tdTomato-Aurora B^AUK1 to LacO foci in S phase (Figs. 4, E and F). Intriguingly, we also noticed that, unlike endogenous KKT8 (which is not present in anaphase), ectopically expressed GFP-KKT8-LacI remained at kinetochores during anaphase (Fig. 4F). This resulted in a fraction of tdTomato-Aurora B^AUK1 being trapped at kinetochores during anaphase instead of migrating to the central spindle (Fig. 4F). We observed a comparable situation upon ectopic expression of GFP-KIN-A, which is retained on anaphase kinetochores together with tdTomato-KKT8 (Fig. S4F). In contrast, Aurora B^AUK1 was not recruited to LacO foci marked by GFP-KKT7^2-261-LacI in anaphase (Fig. 4E).

Figure 4.

KKT7 recruits the KKT8 complex via the KKT9:KKT11 subcomplex

To gain further insights into the structure and assembly hierarchy within the KKT7 – KKT8 complex pathway, we performed CLMS on native complexes isolated by immunoprecipitation of endogenously YFP-tagged kinetochore proteins and mapped the detected crosslinks onto AF2 structure prections (Figs. 4, G and H) (Materials & Methods). AF2 confidently predicted coiled coils between KKT8 and KKT12 and between KKT9 and KKT11 (Fig. 4G; Figs. S4, G and H), suggesting that KKT8:KKT12 and KKT9:KKT11 each form distinct subcomplexes. To validate these findings, we co-expressed combinations of 6HIS-KKT8, KKT9, KKT11, and KKT12 in E. coli and performed metal affinity chromatography (Fig. 4I). 6HIS-KKT8 efficiently pulled down KKT9, KKT11, and KKT12, as shown previously (Ishii and Akiyoshi, 2020). In the absence of KKT9, 6HIS-KKT8 still pulled down KKT11 and KKT12. Removal of either KKT9 or KKT11 did not impact formation of the KKT8:KKT12 subcomplex. In contrast, 6HIS-KKT8 could not be recovered without KKT12, indicating that KKT12 is required for formation of the full KKT8 complex. These results support the idea that the KKT8 complex consists of KKT8:KKT12 and KKT9:KKT11 subcomplexes. The two subcomplexes appear to be connected to each other through interactions between the C-terminal region of the KKT8:KKT12 coiled coils and the C-terminus of KKT11. Two alpha helices in KKT7^2-261 (residues ∼149–181) are predicted to interact with KKT9:KKT11. Using a distance cut-off of 30 Å, ∼70% of crosslinks were compatible with the model (Figs. 4, G and H). Of the crosslinks that failed to meet the distance criteria, ∼90% involved unstructured regions within KKT7 or the C-terminal tail of KKT8. Collectively, our results reveal that KKT7 recruits the KKT8 complex through interaction with the KKT9:KKT11 subcomplex.

We next examined the localization dependency of KKT8 complex components in cells. Using RNAi constructs against individual subunits of the KKT8 complex (Akiyoshi and Gull, 2014; Ishii and Akiyoshi, 2020; Marcianò et al., 2021), we assessed localization of endogenously YFP-tagged KKT8, KKT9 and KKT12 (Figs. S4, I-L). We found that knockdown of any subunit of the KKT8 complex affected protein levels and kinetochore localization of the other subunits, indicating that presence of all subunits is required to stabilize the full complex. YFP-KKT9 was least affected and was still detectable at kinetochores in ∼50% of cells depleted of KKT8, KKT11 or KKT12 (Fig. S4L). Thus, KKT9:KKT11 may lie upstream of KKT8:KKT12. Indeed our IP-CLMS data suggest that the KKT9:KKT11 subcomplex directly interacts with the N-terminus of KKT7 (Fig. 4H, Table S2). KKT7 also formed robust crosslinks with the KKT10/19 kinases (Table S2), supporting our previous finding that KKT7 and KKT10 form a stable complex (Ishii and Akiyoshi, 2020). Together, these data suggest that the KKT7 – KKT8 complex pathway serves as the main CPC recruitment arm in trypanosomes.

The KIN-A C-terminal tail interacts with the KKT8 complex through a conserved domain

By IP-CLMS we failed to detect reliable crosslinks between the CPC and the KKT7 – KKT8 complex or, in fact, any kinetochore proteins (Table S2). This suggests that the IP-CLMS approach, although well-suited for characterizing stable protein complexes, may not be sensitive enough to detect transient or lower affinity interactions. To overcome this, we used AF2 to probe for potential interactions between the KKT8 complex and CPC (sub)complex. AF2 did not predict interactions between the KKT8 complex and INCENP^CPC1, CPC2 or KIN-B (data not shown). Intriguingly, AF2 predicted with high confidence interactions between KKT9:KKT11 and a conserved region (residues ∼722–741) within the KIN-A C-terminal tail, which we termed conserved domain 1 (CD1) (Figs. 5, A and B; Figs. S5, A and B). This interaction involves a triple helix composed of KIN-A CD1, KKT9, and KKT11 in a region close to the KKT7-binding site. pLDDT scores improved significantly for KIN-A CD1 in complex with KKT9:KKT11 (>80) compared to KIN-A CD1 alone (∼20) (Fig. S3, A and B), suggesting that CD1 forms a helical structure upon binding to KKT9:KKT11. Sequence alignment revealed the presence of a second conserved domain (residues ∼816–862) within the C-terminal tail of KIN-A, hereafter referred to as CD2 (Fig. 5B; Fig. S5A). To assess the contributions of CD1 and CD2 for kinetochore recruitment of KIN-A in vivo, we ectopically expressed GFP fusions of the central coiled coils and C-terminal tail of KIN-A (residues ∼310–862) lacking either CD1, CD2, or both (Figs. S5, C-F). GFP-KIN-A^{310-862 ΔCD2} showed a moderate reduction in kinetochore localization in metaphase and was completely lost from kinetochores in anaphase (Figs. S5, D and G). By contrast, GFP-KIN-A^{310-862 ΔCD1} was largely dispersed in metaphase but reappeared at kinetochores in anaphase (Figs. S5, E and G). GFP-KIN-A^310-716 lacking both CD1 and CD2 failed to enrich at kinetochores both in metaphase and anaphase (Figs. S5, F and G). These data suggest that CD1 and CD2 synergistically promote kinetochore localization of KIN-A, with CD1 interacting with KKT9:KKT11 and CD2 possibly interacting with another receptor at the kinetochore.

Figure 5.

We next tested the relevance of KIN-A CD1 and CD2 for CPC localization and function by replacing one allele of KIN-A with C-terminally tagged wild-type or mutant constructs lacking either CD1 or CD2 and performed RNAi against the 3’UTR of KIN-A to deplete the untagged allele. We were unable to obtain a rescue cell line lacking both CD1 and CD2 as the double-mutant protein was not properly expressed. Wild-type KIN-A-YFP along with Aurora B^AUK1 robustly localized to kinetochores from S phase until anaphase onset (Figs. 1, C and D; Fig. 5C). KIN-A^ΔCD1-YFP was detectable at kinetochores in G2 but predominantly localized to the mitotic spindle from (pro)metaphase onwards (Figs. 5, D and F), indicating that removal of CD1 severely weakens the affinity of KIN-A for kinetochores and instead shifts the balance towards microtubule binding. Interestingly, expression of KIN-A^ΔCD1-YFP similarly affected the localization of Aurora B^AUK1 (Figs. 5, D and G; Fig. S5H). We also detected partial spindle localization of Aurora B^AUK1 in a small population (∼25%) of metaphase cells expressing KIN-A^ΔCD2-YFP (Figs. 5, E-G). Central spindle localization of KIN-A in anaphase was unaffected by deletion of either CD1 or CD2 (Figs. 5, D and E). Remarkably, despite a substantial loss of Aurora B^AUK1 from kinetochores in metaphase, ΔCD1 cells exhibited normal cell cycle profiles (Fig. S5I) and showed only a modest decrease in proliferation rates (Fig. 5H). This parallels the situation in budding yeast, in which error-free chromosome segregation can be sustained even when inner centromere localization of Aurora B is largely abolished (Campbell and Desai, 2013; García-Rodríguez et al., 2019).

CPC targeting to the central spindle in anaphase depends on KIN-A’s ATPase activity

Finally, we asked how translocation of the CPC to the spindle midzone in anaphase is achieved in trypanosomes. In mammalian cells, dephosphorylation of INCENP and the kinesin MKLP2 upon anaphase onset allows formation of a transient CPC-MKLP2 complex (Gruneberg et al., 2004; Hümmer and Mayer, 2009; Kitagawa et al., 2014; Serena et al., 2020). MKLP2 activity then drives plus-directed movement of this complex along microtubules of the anaphase spindle. We therefore speculated that anaphase translocation of the kinetoplastid CPC to the central spindle may involve the kinesin motor domain of KIN-A. KIN-B is unlikely to be a functional kinesin based on the absence of several well-conserved residues and motifs within the motor domain, which are fully present in KIN-A (Li et al., 2008). These include the P-loop, switch I and switch II motifs, which form the nucleotide binding cleft, and many conserved residues within the α4-L12 elements, which interact with tubulin (Fig. S6A) (Endow et al., 2010). Consistent with this, the motor domain of KIN-B, contrary to KIN-A, failed to localize to the mitotic spindle when expressed ectopically (Fig. S2E) and did not co-sediment with microtubules in our in vitro assay (Fig. S6B).

Ectopically expressed GFP-KIN-A and -KIN-A^2-309 partially localized to the mitotic spindle but failed to concentrate at the midzone during anaphase (Figs. 2, F and G), suggesting that N-terminal tagging of the KIN-A motor domain may interfere with its function. To address whether the ATPase activity of KIN-A is required for central spindle localization of the CPC, we replaced one allele of KIN-A with a C-terminally YFP-tagged G210A ATP hydrolysis-defective rigor mutant (Fig. 6A) (Rice et al., 1999) and used an RNAi construct directed against the 3’UTR of KIN-A to deplete the untagged allele. The rigor mutation did not affect recruitment of KIN-A to kinetochores (Figs. S6, C and D). However, KIN-A^G210A-YFP marked kinetochores were misaligned in ∼50% of cells arrested in metaphase, suggesting that ATPase activity of KIN-A promotes chromosome congression to the metaphase plate (Figs. S6, E-H). In anaphase, the KIN-A rigor mutant failed to concentrate at the central spindle and instead widely decorated the mitotic spindle, with increased signal observed at spindle poles possibly due to poleward flux (Figs. 6, B and C). Expression of the KIN-A^G210A rigor mutant prevented Aurora B^AUK1 from translocating to the central spindle and caused lagging chromosomes (Figs. 6, D-F). The KIN-A rigor mutation also slowed cell proliferation even in the presence of wild-type protein and caused accumulation of cells in anaphase (Figs. 6, G and H). We conclude that central spindle localization of the CPC depends on KIN-A’s ATPase activity and is required for proper chromosome segregation.

Figure 6.

Discussion

Whereas astonishing diversity in kinetochore composition is seen among eukaryotes (Hooff et al., 2017; Komaki et al., 2022; Tromer et al., 2019), the proteins of the regulatory circuitry underlying chromosome segregation, such as the APC/C, SAC and CPC, are more widely conserved. Homologs of the CPC proteins Aurora B kinase and its associated partner INCENP have been detected in almost all sequenced eukaryotes, including kinetoplastids. The dynamic localization pattern exhibited by the CPC (e.g. transferring from centromeres to the central spindle upon metaphase-anaphase transition) is likewise highly conserved across eukaryotes but appears to be achieved through a variety of mechanisms. For instance, while CPC recruitment to the centromeres in higher eukaryotes is governed by two histone phosphorylation marks (Haspin-mediated H3T3ph and Bub1-mediated H2AT120ph), budding yeasts employ a combination of histone modifications (Bub1-mediated H2AT121ph) and kinetochore proteins as CPC receptors. Borealin^Bir1 not only recognizes Sgo1 but also interacts with the CBF3 complex through Ndc10 (Cho and Harrison, 2011; Yoon and Carbon, 1999). Furthermore, INCENP^Sli15/Aurora B^Ipl1 interact with the Ctf19 subunit of the COMA complex at kinetochores (Fischböck-Halwachs et al., 2019). The proteins that form the localization module of the CPC in different species appear to mirror the diversity in centromeric CPC receptors. In fact, many phyla lack Borealin or Survivin homologs (Komaki et al., 2022). Komaki et al. recently identified two functionally redundant CPC proteins in Arabidopsis, Borealin Related Interactor 1 and 2 (BORI1 and 2), which engage in a triple helix bundle with INCENP and Borealin using a conserved helical domain, but employ an FHA domain instead of a BIR domain to read H3T3ph (Komaki et al., 2022).

In this study, we have identified KIN-A and KIN-B as components of the CPC in trypanosomes, and delineated a novel pathway for centromeric recruitment of the CPC in this evolutionary divergent group of eukaryotes. In agreement with our work, an early study on the CPC in T. brucei found that KIN-A and KIN-B co-purified with Aurora B^AUK1, INCENP^CPC1 and CPC2 based on a pull-down of an Aurora B^AUK1-PTP fusion protein followed by mass spectrometry analysis (Li et al., 2008). However, HA-tagged KIN-A and KIN-B were not detected at kinetochores (from late interphase until metaphase) nor at the new FAZ tip (from late anaphase), and hence were not interpreted as CPC proteins. Contrary to this report, our data clearly show that KIN-A and KIN-B are constitutive components of the CPC in T. brucei. First, YFP-tagged KIN-A and KIN-B co-localize with Aurora B^AUK1, INCENP^CPC1 and CPC2 throughout the cell cycle. Second, KIN-A and KIN-B are readily detected within native CPC complexes isolated by immunoprecipitation of CPC subunits from cells arrested prior to anaphase and form robust crosslinks with the other CPC subunits. Finally, YFP-Aurora B^AUK1 and INCENP^CPC1-YFP are crucially dependent on KIN-A and KIN-B for localizing both to metaphase kinetochores and to the anaphase central spindle. Thus, the KIN-A:KIN-B subcomplex constitutes the localization module of the trypanosome CPC.

Biochemical, cell biological and in silico modelling approaches indicate that the kinesins KIN-A and KIN-B form coiled coils between their central and C-terminal domains, respectively, which then serve as a scaffold onto which INCENP^CPC1 and CPC2 assemble via their N-terminal α-helical domains. The catalytic module of the trypanosomes CPC, consisting of Aurora B^AUK1 bound to the C-terminal IN-box of INCENP^CPC1, is connected to the KIN-A:KIN-B scaffold through a flexible linker in INCENP^CPC1. While our on-beads cross-linking of native CPCs suggests that the catalytic module is positioned in close proximity to the kinesin heads, this may not necessarily be true in vivo. For example, the catalytic module may exist in both ‘locked’ and ‘open’ conformations with regard to its association with the kinesin motor domains. We speculate that the interaction of the KIN-A motor domain with microtubules from prometaphase onwards may cause the catalytic module to disengage from its kinesin head-associated state (Fig. 7). In analogy to the SAH domain of INCENP in other model eukaryotes which has been proposed to function as a dog leash (Samejima et al., 2015; Santaguida and Musacchio, 2009), the INCENP^CPC1 flexible linker in trypanosomes (∼100 amino acids long) may then permit Aurora B^AUK1 to roam across a larger but nevertheless spatially constrained target area to phosphorylate its substrates while still being anchored to the kinetochore via KIN-A:KIN-B. Importantly, this mechanism would allow the CPC to act as an intrinsic ‘sensor’ of KT-MT attachments. Such models dealing with alternative conformations of the CPC in various cellular context will require further testing in the future.

Figure 7.

We propose that multiple weak interactions of the KIN-A C-terminal unstructured tail with kinetochore components act in synergy to stabilize the CPC at kinetochores. Critically, these interactions need to be of transient and reversible nature to permit the dynamic release of the CPC upon anaphase onset. Three mechanisms are likely to play a role in this context (Fig. 7). First, removal of the KKT8 complex (the ‘CD1 receptor’) at the metaphase-anaphase transition effectively eliminates one of the key CPC-kinetochore interfaces. Secondly, the affinity of the KIN-A C-terminal tail for its binding partners at the kinetochore may be further finetuned through reversible post-translational modifications. In support of this, the KIN-A C-terminal tail harbors many pututative CRK3 sites (10 sites matching the minimal S/T-P consensus motif for CDKs) and is also heavily phosphorylated by Aurora B^AUK1 in vitro (Ballmer et al., 2024). Finally, we speculate that the interaction of KIN-A motor domain with microtubules, coupled to the force generating ATP hydrolysis and possibly plus-end directed motion, eventually outcompetes the weakened interactions of the CPC with the kinetochore and facilitates the extraction of the CPC from chromosomes onto spindle microtubules during anaphase. Indeed, deletion of the KIN-A motor domain or impairment of its motor function through N-terminal GFP tagging causes the CPC to be trapped at kinetochores in anaphase. Central spindle localization is additionally dependent on the ATPase activity of the KIN-A motor domain as illustrated by the KIN-A rigor mutant.

It remains to be investigated whether KIN-A functions as a plus-end directed motor. The role of the KIN-B in this context is equally unclear. Because KIN-B does not possess a functional kinesin motor domain, we deem it unlikely that the KIN-A:KIN-B heterodimer moves hand-over-hand along microtubules as do conventional (kinesin-1 family) kinesins. Rather, the KIN-A motor domain may function as a single-headed unit and drive processive plus-end directed motion using a mechanism similar to the kinesin-3 family kinesin KIF1A (Okada and Hirokawa, 1999). Formation of transient complexes between the CPC and kinesins or other microtubule plus-end tracking proteins upon anaphase onset appears to be a common theme underlying the central spindle translocation of the CPC. For instance, the CPC interacts with MKLP2 or Bim1^EB1 in human and yeast cells, respectively (Gruneberg et al., 2004; Zimniak et al., 2012). Thus, deeper understanding of the CPC regulation in trypanosomes is bound to provide evolutionary insights into fundamental principles of chromosome segregation in eukaryotes and can lead to the discovery of druggable targets to combat kinetoplastid diseases (Saldivia et al., 2020).

Materials and Methods

Cloning

All primers, plasmids, bacmids, and synthetic DNA used in this study as well as their source or construction details are described in Supplemental Table S1. All constructs were sequence verified.

Trypanosome culture

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) or from PCF1339 procyclic form cells (TREU 927/4 expressing T7 RNA polymerase, tetracycline repressor and the Cas9 nuclease, Beneke et al., 2017) and are described in Supplemental Table S1. Cells were grown at 28°C in SDM-79 medium supplemented with 10% (vol/vol) heat-inactivated fetal calf serum, 7.5 μg/ml hemin (Brun and Schönenberger, 1979), and appropriate selection drugs. Cell growth was monitored using a CASY cell counter (Roche). PCR products or plasmids linearized by NotI were transfected into cells by electroporation (Biorad). Transfected cells were selected by the addition of 30 μg/ml G418 (Sigma), 25 μg/ml hygromycin (Sigma), 5 μg/ml phleomycin (Sigma), or 10 μg/ml blasticidin S (Insight biotechnology). To obtain endogenously tagged clonal strains, transfected cells were selected by the addition of appropriate drugs and cloned by dispensing dilutions into 96-well plates. Endogenous YFP tagging was performed using the pEnT5-Y vector (Kelly et al., 2007) or a PCR-based method (Dean et al., 2015). Endogenous tdTomato tagging was performed using pBA148 (Akiyoshi and Gull, 2014) and its derivatives. All constructs for ectopic expression of GFP fusion proteins include a short nuclear localization signal (NLS) (Marchetti et al., 2000). For doxycycline inducible expression of head-to-head (pBA3-based) and hairpin (pBA310-based) RNAi constructs, GFP-NLS (pBA310-based) and GFP-NLS-LacI fusion proteins (pBA795-based), the linearized plasmids were integrated into 177-bp repeats on minichromosomes. Expression of GFP-NLS or GFP-NLS-LacI fusions was induced by the addition of 10 ng/mL doxycycline for 24 h. RNAi was induced by the addition of 1 μg/ml doxycycline. LacO–LacI tethering experiments were carried out as described previously using the LacO array inserted at the rDNA locus (Ishii and Akiyoshi, 2020; Landeira and Navarro, 2007).

Immunoprecipitation followed by mass spectrometry (IP-MS)

For standard IPs, 400-ml cultures of asynchronously growing cells were grown to ∼5 – 10 million cells/ml. Cells were pelleted by centrifugation (800 g, 10 min), washed once with PBS, and extracted in PEME (100 mM Pipes-NaOH, pH 6.9, 2 mM EGTA, 1 mM MgSO₄, and 0.1 mM EDTA) with 1% NP-40, protease inhibitors (10 μg/ml leupeptin, 10 μg/ml pepstatin, 10 μg/ml E-64, and 0.2 mM PMSF) and phosphatase inhibitors (1 mM sodium pyrophosphate, 2 mM Na-β-glycerophosphate, 0.1 mM Na₃VO₄, 5 mM NaF, and 100 nM microcystin-LR) for 5 min at RT, followed by centrifugation at 1,800 g for 15 min. Samples were kept on ice from this point on. The pelleted fraction containing kinetochore proteins was resuspended in modified buffer H (BH0.15: 25 mM Hepes, pH 8.0, 2 mM MgCl₂, 0.1 mM EDTA, pH 8.0, 0.5 mM EGTA, pH 8.0, 1% NP-40, 150 mM KCl, and 15% glycerol) with protease and phosphatase inhibitors. Samples were sonicated to solubilize kinetochore proteins (12 s, three times with 1-min intervals on ice). 12 μg of mouse monoclonal anti-GFP antibodies (11814460001; Roche) preconjugated with 60 μl slurry of Protein-G magnetic beads (10004D; Thermo Fisher Scientific) with dimethyl pimelimidate (Unnikrishnan et al., 2012) were incubated with the extracts for 2.5 h with constant rotation, followed by four washes with modified BH0.15 containing protease inhibitors, phosphatase inhibitors and 2 mM DTT. Beads were further washed three times with pre-elution buffer (50 mM Tris-HCl, pH 8.3, 75 mM KCl, and 1 mM EGTA). Bound proteins were eluted from the beads by agitation in 60 μl of elution buffer (0.1% RapiGest [186001860; Waters] and 50 mM Tris-HCl, pH 8.3) for 25 min at RT. Eluates were then incubated at 100°C for 5 min. Proteins were reduced with 5 mM DTT at 37°C for 30 min and alkylated with 10 mM iodoacetamide at 37°C for 30 min. The reaction was quenched by adding 10 mM DTT at 37°C for 30 min, and 100 μl of 20 mM Tris-HCl (pH 8.3) was added. Proteins were digested overnight at 37°C with 0.2 μg trypsin (Promega). Formic acid was then added to 2% and the samples were incubated at 37°C for 30 min to cleave RapiGest, followed by centrifugation for 10 min. The supernatant was desalted over a C18 column and analyzed by electrospray tandem mass spectrometry over a 60-min gradient using Q-Exactive (Thermo Fisher Scientific) at the Advanced Proteomics Facility (University of Oxford). Peptides were identified by searching tandem mass spectrometry spectra against the T. brucei protein database with MaxQuant (version 2.0.1) with carbamidomethyl cysteine set as a fixed modification and oxidization (Met), phosphorylation (Ser, Thr, and Tyr), and acetylation (Lys) set as variable modifications. Up to two missed cleavages were allowed. The first peptide tolerance was set to 10 ppm. Results were filtered to remove contaminants and reverse hits. Proteins identified with at least two peptides were considered as significant and shown in Table S2 (protein FDR 1%).

Ex vivo cross-linking of the native CPC and kinetochore complexes (IP-CLMS)

For cross-linking IP-MS experiments, cell cultures were scaled up to 1,600 mL. Cell cultures were treated with 10 μM MG132 for 4 h to enrich for cells in metaphase. Cell lysis and immunoprecipitation steps were carried out as described above. After four washes with modified BH0.15 containing protease inhibitors, phosphatase inhibitors and 2 mM DTT, beads were washed three times with 25 mM HEPES pH7.5, 150 mM NaCl. Proteins were then cross-linked on beads with 0.4 mM BS³ (bis(sulfosuccinimidyl)suberate) (Thermo Fisher Scientific) for 30 min at RT with agitation, followed by three washes in 25 mM HEPES pH7.5, 150 mM NaCl and a further three washes in 0.1 M ammonium bicarbonate. Samples were then incubated in 8 M urea dissolved in 0.1 M ammonium bicarbonate for 10 min at RT with agitation. Proteins were reduced with 10 mM TCEP for 20 min and alkylated with 10 mM iodoacetamide for 40 min at RT. Proteins were then then pre-digested with 0.4 μg LysC for 2 h at 37°C. The urea concentration in the sample was brought down to < 1 M by addition of 0.1 M ammonium bicarbonate before adding CaCl₂ (to 2 mM) and 0.7 μg of trypsin for overnight digestion at 37°C. Formic acid was then added to 2% and the samples were frozen. The cross-linked samples were further processed and analyzed at the proteomics core facility at EMBL Heidelberg. Digested peptides were concentrated and desalted using an OASIS HLB µElution Plate (Waters) according to manufacturer instructions. Crosslinked peptides were enriched using size exclusion chromatography (Leitner et al., 2012). In brief, desalted peptides were reconstituted with SEC buffer (30% (v/v) ACN in 0.1% (v/v) TFA) and fractionated using a Superdex Peptide PC 3.2/30 column (GE) on a 1200 Infinity HPLC system (Agilent) at a flow rate of 0.05 ml/min. Fractions eluting between 50-70 μl were evaporated to dryness and reconstituted in 30 μl 4% (v/v) ACN in 1% (v/v) FA. Collected fractions were analyzed by liquid chromatography (LC)-coupled tandem mass spectrometry (MS/MS) using an UltiMate 3000 RSLC nano LC system (Dionex) fitted with a trapping cartridge (µ-Precolumn C18 PepMap 100, 5µm, 300 µm i.d. x 5 mm, 100 Å) and an analytical column (nanoEase M/Z HSS T3 column 75 µm x 250 mm C18, 1.8 µm, 100 Å, Waters). Trapping was carried out with a constant flow of trapping solvent (0.05% trifluoroacetic acid in water) at 30 µl/min onto the trapping column for 6 minutes. Subsequently, peptides were eluted and separated on the analytical column using a gradient composed of Solvent A (3% DMSO, 0.1% formic acid in water) and solvent B (3% DMSO, 0.1% formic acid in acetonitrile) with a constant flow of 0.3 µl/min. The outlet of the analytical column was coupled directly to an Orbitrap Fusion Lumos (Thermo Scientific, SanJose) mass spectrometer using the nanoFlex source. The peptides were introduced into the Orbitrap Fusion Lumos via a Pico-Tip Emitter 360 µm OD x 20 µm ID; 10 µm tip (CoAnn Technologies) and an applied spray voltage of 2.1 kV, instrument was operated in positive mode. The capillary temperature was set at 275°C. Only charge states of 4-8 were included. The dynamic exclusion was set to 30 sec. and the intensity threshold was 5e⁴. Full mass scans were acquired for a mass range 350-1700 m/z in profile mode in the orbitrap with resolution of 120000. The AGC target was set to Standard and the injection time mode was set to Auto. The instrument was operated in data dependent acquisition (DDA) mode with a cycle time of 3 sec between master scans and MSMS scans were acquired in the Orbitrap with a resolution of 30000, with a fill time of up to 100 ms and a limitation of 2e⁵ ions (AGC target). A normalized collision energy of 32 was applied. MS2 data was acquired in profile mode. RAW MS files were searched by the pLink2 software (Chen et al., 2019), with carbamidomethyl cysteine set as a fixed and oxidization (Met) set as variable modifications. Up to two missed cleavages were allowed. Precursor tolerance was set to 10 ppm. All the identified cross-links are shown in Table S2 (FDR 5%). Cross-links were plotted using xiView (Graham et al., 2019). All raw mass spectrometry files and custom database files used in this study have been deposited with the ProteomeXchange Consortium via the PRIDE partner repository (Deutsch et al., 2020; Perez-Riverol et al., 2019) with the dataset identifier PXD045987.

Fluorescence microscopy

Cells were washed once with PBS, settled onto glass slides, and fixed with 4% paraformaldehyde in PBS for 5 min as described previously (Nerusheva and Akiyoshi, 2016). Cells were then permeabilized with 0.1% NP-40 in PBS for 5 min and embedded in mounting media (1% wt/vol 1,4-diazabicyclo[2.2.2]octane, 90% glycerol, 50 mM sodium phosphate, pH 8.0) containing 100 ng/ml DAPI. Images were captured on a Zeiss Axioimager.Z2 microscope (Zeiss) installed with ZEN using a Hamamatsu ORCA-Flash4.0 camera with 63× objective lenses (1.40 NA). Typically, ∼20 optical slices spaced 0.2 or 0.24 μm apart were collected. Images were analysed in ImageJ/Fiji (Schneider et al., 2012). Kinetochore localization of endogenously tagged kinetochore proteins or ectopically expressed constructs were examined manually by quantifying the number of cells that clearly had detectable kinetochore-like dots at indicated cell cycle stages. Shown images are central slices.

In silico structure and interaction predictions

Structures and interactions were predicted with AlphaFold2-Multimer-v2 (Evans et al., 2022; Jumper et al., 2021) through ColabFold using MMseqs2 (UniRef+Environmental) and default settings (Mirdita et al., 2022). Structural predictions of KIN-A/KIN-B, KIN-A^310-862/KIN-B^317-624, CPC1/CPC2/KIN-A^300-599/KIN-B 317-624, and KIN-A^700-800/KKT9/KKT11 were performed using ColabFold version 1.3.0 (AlphaFold-Multimer version 2), while those of AUK1/CPC1/CPC2/KIN-A^1-599/KIN-B, KKT7^{1- 261}/KKT9/KKT11/KKT8/KKT12, KKT9/KKT11/KKT8/KKT12, and KKT7^1-261/KKT9/KKT11 were performed using ColabFold version 1.5.3 (AlphaFold-Multimer version 2.3.1) using default settings, accessed via https://colab.research.google.com/github/sokrypton/ColabFold/blob/v1.3.0/AlphaFold2.ipynb and https://colab.research.google.com/github/sokrypton/ColabFold/blob/v1.5.3/AlphaFold2.ipynb. All structure figures were made using PyMOL version 2.5.2 (Schrödinger, LLC). The following command was used to map pLDDT score onto the AF2 predicted structure models: spectrum b, rainbow_rev, maximum=100, minimum=50.

Protein purification from E. coli

Recombinant 6HIS-KKT8, KKT9, KKT11, KKT12 (pBA457) and derivatives were expressed in Rosetta 2(DE3)pLys E. coli cells (Novagen). 6HIS-KIN-A^2-309 (pBA2519) and 6HIS-KIN-B^2-316 (pBA2513) were expressed in BL21(DE3) cells. Proteins were purified and eluted from TALON beads as previously described (Llauró et al., 2018). Briefly, cells were grown in 2xTY media at 37°C to an OD₆₀₀ of ∼0.8, at which point protein expression was induced by 0.1 mM IPTG, and then incubated overnight at 20°C. Cells were pelleted at 3,400 g at 4°C and pellets were resuspended in P500 buffer (50 mM sodium phosphate, pH 7.5, 500 mM NaCl, 5 mM imidazole and 10% glycerol) supplemented with protease inhibitors (20 μg/ml leupeptin, 20 μg/ml pepstatin, 20 μg/ml E-64, 0.4 mM PMSF) and 1 mM TCEP, and were sonicated on ice. Lysates were treated with benzonase nuclease (500 U/1 l culture) and spun at 48,000 g at 4°C for 30 min. Supernatant was incubated with TALON beads (Takara Clontech) for 1 h at 4°C, rotating. The beads were washed three times with lysis buffer and proteins were then eluted with elution buffer (P500 buffer containing 250 mM imidazole with 1 mM TCEP). For microtubule co-sedimentation assays, 6HIS-KIN-A^2-309 and 6HIS-KIN-B^2-316 were buffer exchanged into BRB80 (80 mM Pipes-KOH, pH 6.9, 1 mM EGTA, and 1 mM MgCl₂) with 100 mM KCl using a Zeba columns (Thermo Fisher) and flash-frozen in liquid nitrogen for −80°C storage. Polyacrylamide gels were stained with SimplyBlue SafeStain (Invitrogen).

Microtubule co-sedimentation assay

Microtubule co-sedimentation assays were performed as described previously (Ludzia et al., 2021). Briefly, taxol-stabilised microtubules were prepared by mixing 2.5 ml of 100 μM porcine tubulin (Cytoskeleton) resuspended in BRB80 with 1 mM GTP (Cytoskeleton), 1.25 μl BRB80, 0.5 μl of 40 mM MgCl₂, 0.5 μl of 10 mM GTP, and 0.25 μl DMSO, and incubated for 20 min at 37C. 120 ml of pre-warmed BRB80 containing 12.5 μM Taxol (paclitaxel; Sigma) was added to the sample to bring the microtubule concentration to ∼2 μM. 20 μl of 6HIS-KIN-A^2-309 or 6HIS-KIN-B^2-316 (at final concentration of 4 μM) in BRB80 with 100 mM KCl were mixed with 20 μl of microtubules (final, 1 μM) and incubated for 45 min at room temperature. As a control, we incubated 6HIS-KIN-A^2-309 or 6HIS-KIN-B^2-316 with BRB80 (with 12.5 μM Taxol). The samples were spun at 20,000 g at room temperature for 10 min, and the supernatant was collected. To the tubes containing pelleted fractions, we added 40 μl of chilled BRB80 with 5 mM CaCl₂ and incubated on ice for 5 min to depolymerise microtubules. Following the incubation, samples were boiled for 5 min before SDS-PAGE. Gels were stained with SimplyBlue Safe Stain (Invitrogen). Co-sedimentation assays were performed at least twice with similar results.

Immunoblotting

Cells were harvested by centrifugation (800 g, 5 min) and washed with 1 ml PBS. The pellet was resuspended in 1× LDS sample buffer (Thermo Fisher) with 0.1 M DTT. Denaturation of proteins was performed for 5 min at 95°C. SDS-PAGE and immunoblots were performed by standard methods using the following antibodies: rabbit polyclonal anti-GFP (TP401, 1:5000) and mouse monoclonal TAT1 (anti-trypanosomal-alpha-tubulin, 1:5000, a kind gift from Keith Gull) (Woods et al., 1989). Secondary antibodies used were: IRDye 680RD goat anti-mouse (LI-COR, 926-68070) and IRDye 800CW goat anti-rabbit (LI-COR, 926-32211). Bands were visualized on an ODYSSEY Fc Imaging System (LI-COR).

Multiple sequence alignment

Protein sequences and accession numbers for Aurora B^AUK1, INCENP^CPC1, CPC2, KIN-A and KIN-B used in this study were retrieved from the TriTryp database (Aslett et al., 2010), UniProt (Bateman, 2019), or a published study (Butenko et al., 2020). Searches for homologous proteins were done using BLAST in the TriTryp database (Aslett et al., 2010) or using hmmsearch using manually prepared hmm profiles (HMMER version 3.0; Eddy, 1998). The top hit in each organism was considered as a true ortholog only if the reciprocal BLAST search returned the query protein as a top hit in T. brucei. Multiple sequence alignment was performed with MAFFT (L-INS-i method, version 7) (Katoh et al., 2019) and visualised with the Clustalx colouring scheme in Jalview (version 2.10) (Waterhouse et al., 2009).

Acknowledgements

We thank Miguel Navarro (Instituto de Parasitologıá y Biomedicina López-Neyra, Consejo Superior de Investigaciones Cientificas, Spain) for providing pMig75 and pMig96 plasmids and Midori Ishii Kanazawa and William Carter for helping trypanosome strain construction. We thank the Micron Advanced Bioimaging Unit and the Advanced Proteomics Facility at the University of Oxford, and the Proteomics Core Facility at the EMBL in Heidelberg, especially Mandy Rettel and Jennifer Schwarz, for their support. We also thank Patryk Ludzia and Midori Ishii Kanazawa for comments on our manuscript. D. Ballmer was supported by the Berrow Foundation. B. Akiyoshi was supported by a Wellcome Trust Senior Research Fellowship (grant 210622/Z/18/Z).

The authors declare no competing financial interest.

Rights retention: This research was funded in whole or in part by Wellcome Trust (grant 210622/Z/18/Z). For the purpose of Open Access, the author has applied a CC BY public copyright licence to any Author Accepted Manuscript (AAM) version arising from this submission.

Author contributions: D. Ballmer performed essentially all experiments except for some LacI/LacO tethering experiments that were performed by B. Akiyoshi. D. Ballmer wrote the manuscript. B. Akiyoshi edited the manuscript.