Aurora A (also known as STK15/BTAK in humans), a putative oncoprotein naturally overexpressed in many human cancers, is a member of the conserved Aurora protein serine/threonine kinase family that is implicated in the regulation of G2–M phases of the cell cycle. In vitro studies utilizing antibody microinjection, siRNA silencing and small molecule inhibitors have indicated that Aurora A functions in early as well as late stages of mitosis. However, due to limitations in specificity of the techniques, exact functional roles of the kinase remain to be clearly elucidated. In order to identify the physiological functions in vivo, we have generated Aurora A null mouse embryos, which show severe defects at 3.5 d.p.c. (days post-coitus) morula/blastocyst stage and lethality before 8.5 d.p.c. Null embryos at 3.5 d.p.c. reveal growth retardation with cells in mitotic disarray manifesting disorganized spindle, misaligned and lagging chromosomes as well as micronucleated cells. These findings provide the first unequivocal genetic evidence for an essential physiological role of Aurora A in normal mitotic spindle assembly, chromosome alignment segregation and maintenance of viability in mammalian embryos.
Aurora A, a member of the conserved serine/threonine protein kinase family represented by the prototypic Ipl1 kinase in yeast (Carmena and Earnshaw, 2003), has been identified as a mitosis regulatory protein and a putative oncoprotein that is naturally overexpressed in many human cancers (Katayama et al., 2003). Aurora A expression peaks during G2–M phase of the cell cycle localizing predominantly on the centrosomes and proximal mitotic spindle.
In vitro studies have implicated Aurora A in the regulation of centrosome maturation separation, mitotic entry, bipolar spindle assembly, chromosome alignment and mitotic exit (Berdnik and Knoblich, 2002; Giet et al., 2002; Conte et al., 2003). Identification of microtubule-associated proteins as substrates of Aurora A (Yu et al., 2005; Koffa et al., 2006) as well as their localization on k fibers in prometaphase human cells (Sillje et al., 2006) indicates that Aurora A forms complex with mitotic spindle assembly factors in the organization of the spindle microtubules. Additionally, while silencing of Aurora A has been reported to cause delay or block in mitotic entry (Marumoto et al., 2002; Hirota et al., 2003; Du and Hannon, 2004; Satinover et al., 2006), accelerated initiation of mitosis in presence of active Aurora A has also been observed (Ma et al., 2003; Liu and Ruderman, 2006). These effects may, in part, be explained by the reported role of Aurora A in facilitating expression and activation of critical mitosis regulators cyclin B and Cdc25B (Mendez et al., 2000; Tay et al., 2000; Dutertre et al., 2004). However, involvement of Aurora A in regulating specific cellular phenotypes still remain uncertain in view of the inherent limitations in assessing the contributions of the endogenous proteins in ex vivo experiments and also due to occasional contradictory results published on the consequence of ablating Aurora A function in cells grown in vitro (Girdler et al., 2006; Hoar et al., 2007).
In order to elucidate the physiological functions of Aurora A in vivo, we have carried out targeted disruption of the mouse Aurora kinase A (Aurka) gene through Cre-loxP-mediated recombination in mice. The results reveal that Aurora A is essential for proper mitotic spindle assembly, chromosome alignment segregation and viability in mammalian embryos.
Results and discussion
Targeted disruption of the Aurka gene was achieved as outlined in Figure 1a. Ten percent of the recombinant clones, selected for neomycin resistance in G418 and against TK in gancyclovir to eliminate random insertions, were properly targeted giving rise to the conditional neomycin (CN) allele based on the Southern blot hybridization results with the 5′ and 3′ probes. KpnI and XbaI digestion of DNA from these clones revealed the correct sized genomic fragments of 2 kb with the 5′ probe and 5.4 kb with the 3′ probe in addition to the 8 kb fragments from the wild-type allele (+). Microinjection of two targeted cell lines resulted in germ-line chimeras. The heterozygotes with the CN allele, derived from the chimeras, were used to generate the conditional (C) allele through Flp-mediated recombination. The CN and the C mice were crossed with ZP3-Cre mice to remove the floxed exon 2, generating the null allele (−). Homozygous and heterozygous mice with the wild-type, the CN, the C and the null alleles were verified by Southern blotting of the respective genomic DNA with the 5′ and 3′ probes (Figure 1b) and also by PCR analyses (Figure 1c) with five primers, ‘a’ through ‘e’, utilized in three different combinatorial pairs (referred to as ‘CN PCR’, ‘+/C PCR’ and the ‘Exc PCR’) as mentioned in the Supplementary Table 1. Functional status of the mutant alleles was determined by western blot analyses performed with testis tissues lysate from mice with different genotypes since Aurora A protein is clearly detectable in this tissue. The band intensities suggested that the CN and the C alleles were expressing comparable amounts of the protein like the wild-type allele and the null allele was silenced (Figure 1d).
Aurka+/− intercrosses did not result in any homozygous nulls among live born offspring and 8.5 d.p.c. (days post-coitus) embryos indicating early embryonic lethality of this genotype. Among 61 live born mice and 40 embryos at 8.5 d.p.c., wild-type genotype was detected in 20 and 14, while the numbers of heterozygote were 41 and 26, respectively. Thus in contrast to the expected ratio of 1:2:1 for the wild-type homozygote: heterozygote and homozygote nulls, the observed ratio of 1:2:0 suggested embryonic lethality of the null embryos before 8.5 d.p.c. To determine the timing of embryonic lethality, 40 embryos at 3.5 d.p.c. (morula/blastocyst) were genotyped. This analysis revealed the presence of 8 Aurka−/− in addition to 12 wild-type and 20 heterozygotes (Figures 2a and b). The results, therefore, demonstrated that Aurka−/− embryos were viable at 3.5 d.p.c. in utero but failed to grow further and implant.
The 3.5 d.p.c. embryos from heterozygous crosses were then analyzed for ex vivo growth potential by culturing in vitro. The embryos were photographed immediately (3.5 d.p.c.) and up to 3 days of in vitro culture (E 4.5–E 6.5) followed by genotyping. Total of four wild-type homozygous, nine heterozygous and five null homozygous embryos were analyzed. The 3.5 d.p.c. null mutants were invariably smaller with fewer cells and cells of irregular size in the inner cell mass compared with the wild-type and the heterozygous embryos. After 1 day culture (E 4.5), the wild-type and the heterozygous embryos showed similar growth but the null embryos remained noticeably smaller with some revealing degeneration of the inner cell mass (Figure 2c). The embryos genotyped at the end of the second (E 5.5) and third (E 6.5) days of culture did not reveal any null mutants with only the wild-type homozygote or heterozygote detected (data not shown). The results indicated that the null mutants degenerated sometime between days 4.5 and 6.5 of in vitro culture.
The 3.5 d.p.c. embryos derived from Aurka+/− intercrosses were also analyzed for total cellular content, nuclear morphology, M phase distribution of cells and mitotic structures. Due to the small size and fragility of these embryos, both immunofluorescence staining and PCR genotyping was difficult and in most instances only one of the two methods could be successfully performed. About a quarter of the embryos that were Aurora A-negative and distinctly smaller in size consisted of fewer cells with signs of degeneration, as in the case of the null embryos utilized for ex vivo growth study. Examination of seven 3.5 d.p.c. Aurora A-negative and ten positive embryos revealed clear evidence of growth impairment in the cells of the null embryos. The positive embryos consisted of an average of 40 cells but the negative embryos had about 18 cells, with some appearing more like morulae rather than blastocysts. The average numbers of mitotic cells and micronuclei were about 5- and 10-fold higher, respectively, in the negative embryos compared with the positive embryos (Figure 3a). The mitotic cells of the negative embryos displayed disorganized spindle assembly and majority (∼69%) of these cells were in prometaphase, with condensed chromosomes not congressed or aligned on the metaphase plate (Figure 3b), while about 27 and 4% were in metaphase and anaphase, respectively (Figure 3c). The null cells also manifested misaligned and lagging chromosomes at metaphase and anaphase stages (Figure 3d). The Aurora A-positive embryos revealed a more even distribution of cells at different phases of mitosis with about 33% in prometaphase, 44% in metaphase and 22% in anaphase displaying predominantly normal alignment and segregation of chromosomes. Presence of an average of 18 cells in the null 3.5 d.p.c. morulae/blastocysts indicates that these embryos are able to undergo at least four mitotic cell divisions despite lacking a functional Aurora A allele. It is plausible that initial cell divisions in the null embryos are facilitated by the presence of a low amount of maternal protein since the Unigene cluster database reveals existence of murine Aurka transcript (Mm 249363) in oocyte and zygote. Additionally, the presence of a relatively elevated number of cells in mitosis in the null embryos indicate that mitosis may be initiated with a delay rather than inhibited in absence of Aurora A as has been reported in case of Xenopus early embryonic cell cycles investigated with egg extracts depleted of endogenous Aurora A (Liu and Ruderman, 2006; Satinover et al., 2006).
Aberrant spindle assembly is expected due to impaired formation of k fibers requiring active Aurora A complex with spindle assembly factors, such as TPX2 and HURP, among others (Gruss et al., 2001; Koffa et al., 2006; Sillje et al., 2006; Tulu et al., 2006). It has been hypothesized that microtubules can be organized at multiple, often transient, structures that can catalyze γ-tubulin-dependent microtubule nucleation from their minus ends, plus ends or sides (Luders and Stearns, 2007) with the polar spindle microtubules ‘collecting’ preassembled chromosome-generated minispindles into a single bipolar spindle within the cells (Rieder, 2005). These proposed mechanisms imply that microtubules nucleated on chromosomes and k fibers mediate capture and stabilization of the sister kinetochores on the bipolar spindle and raise the interesting scenario that absence of Aurora A in the null embryos may be interfering with the kinetochore capture process giving rise to the misaligned and lagging chromosomes. The possibility appears conceivable in view of our recent observation that Aurora A is a key regulator of kinetochore-associated microtubules formation process (Katayama et al., submitted).
Severe mitotic defects and early embryonic lethality in Aurora A null mutant are similar to those seen in case of null mutants for other known proteins regulating chromosome segregation and cytokinesis, such as CenpC (Kalitsis et al., 1998), Incenp (Cutts et al., 1999), CenpA (Howman et al., 2000) and Survivin (Uren et al., 2000), as well as those involved in the activation of the spindle assembly checkpoint network like, Mad2 (Dobles et al., 2000), BubR1 (Wang et al., 2004), Mad1 (Iwanaga et al., 2007) and Bub1 (Perera et al., 2007). While defect in kinetochore-microtubule attachments is expected to disrupt normal mitotic progression, apparent delay or arrest of mitosis and also chromosome segregation anomalies in the Aurora A null cells indicate that loss of Aurora A function influences the spindle assembly checkpoint network. Induction of similar chromosome segregation and cytokinesis abnormalities in Aurora A overexpressing cells also tend to favor this notion. Although elevated expression of Aurora A has been reported to override the spindle assembly checkpoint (Anand et al., 2003), specific molecular targets of Aurora A in the checkpoint network remain unknown. Aurora A null allele containing cells will provide an ideal model system for these investigations.
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This study was supported by grants from the National Institutes of Health (RO1 CA89716) and Institutional Research Grant award from the University of Texas MD Anderson Cancer Center to SS. We thank Dr Richard Behringer for critical reading of the paper and Dr Jan Parker-Thornburg for help in this study. Institutional Biospecimen Extraction Facility, DNA Analysis Facility are supported by the Cancer Center Support Grant CA16672.
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