Aurora-A is a kinase involved in the formation and maturation of the mitotic spindle and chromosome segregation. This kinase is frequently overexpressed in human cancer, and its activity may confer resistance to antitumoral drugs such as Taxol. Inhibition of Aurora-A results in mitotic defects, and this kinase is considered as an attractive therapeutic target for cancer. Nevertheless, the specific requirements for this kinase in adult mammalian tissues remain unclear. Conditional genetic ablation of Aurora-A in adult tissues results in polyploid cells that display a DNA-damage–like response characterized by the upregulation of p53 and the cell-cycle inhibitor p21Cip1. This is accompanied by apoptotic, differentiation, or senescence markers in a tissue-specific manner. Therapeutic elimination of Aurora-A prevents the progression of skin and mammary gland tumors. However, this is not due to significant levels of apoptosis or senescence, but because Aurora-A–deficient tumors accumulate polyploid cells with limited proliferative potential. Thus, Aurora-A is required for tumor formation in vivo, and the differential response observed in various tissues might have relevant implications in current therapeutic strategies aimed at inhibiting this kinase in the treatment of human cancer. Cancer Res; 73(22); 6804–15. ©2013 AACR.

Aurora kinases play critical roles in centrosome biology and chromosome segregation (1). The founding member of this protein family, Aurora-A, is primarily implicated in centrosome maturation and spindle assembly (2). Indeed, Aurora-A localizes to the centrosomes during interphase, and to both spindle poles and spindle microtubules during early mitosis. The best-known Aurora-A regulator is TPX2. Binding with TPX2 induces a conformational change in Aurora-A in such a way that the phosphorylated activation domain of this kinase adopts a more compact position, providing a better substrate-binding platform and hiding the activating phosphoryl group from the activity of phosphatase PP1 (3). Aurora-A has been documented to be involved in p53 regulation because it phosphorylates p53 at Ser315, leading to its Mdm2-mediated ubiquitination and subsequent proteolysis (4). Other functions of Aurora-A include the regulation of the translation of RNAs that contain cytoplasmic polyadenylation elements (CPE) at their 3′ untranslated region, which has been shown in the regulation of meiosis (5), α-CamKII synthesis at synapses (6), and the regulation of the cell-cycle regulators cyclin B1 and Cdk1 (7).

Interfering with Aurora-A expression or activity via siRNA expression, immunodepletion, or by specific inhibitors induces mitotic alterations that impair cell-cycle progression. In fact, its disruption in Drosophila melanogaster, Caenorhabditis elegans, and Xenopus causes defects in centrosome maturation and spindle formation (8–13). In addition, RNA interference (RNAi)-mediated reduction of Aurora-A expression in mammalian cell lines has been associated with abnormalities in mitotic entry, bipolar spindle formation, and mitotic progression (14–18). Moreover, Aurora-A is necessary for proper mitotic progression during mouse embryonic development, and its genetic ablation results in lethality at the morula stage (19–21).

Aurora-A is overexpressed in multiple tumor types, including breast, pancreatic, ovarian, and gastric carcinomas (22). In addition, Aurora-A has been included in the top 70 list of genes of the cancer-associated chromosome instability signature (23). Several small-molecule inhibitors have been shown to block its function, and their in vivo antitumor activity is currently under evaluation in clinical trials for the treatment of human cancer (24). Treatment with Aurora-A inhibitors has resulted in some limited but promising responses in early-phase clinical trials (25). Improving future trials will require further understanding of the physiologic effects of inhibiting Aurora-A in adult tissues. Despite all the reported data on the role of Aurora-A in lower organisms and in cultured cells, the requirements for this kinase in adult mammals and the cellular consequences of its elimination remain unclear. We have generated a mouse model that allows the conditional elimination of Aurora-A upon the activation of the Cre recombinase by 4-hydroxytamoxifen (4-OHT), the active metabolite of tamoxifen. Here, we show that ubiquitous ablation of Aurora-A in vivo is associated with an age-related phenotype characterized by a significant alteration of the proliferative tissues. Aurora-A–depleted tissues are characterized by a significant increase in mitotic and DNA damage markers, and with the presence of tetraploid or aneuploid cells, eventually resulting in impaired proliferation and senescence.

Generation and characterization of Aurora-A–mutant mice

To generate Aurora-A conditional mouse models, we used mice carrying the AurAlox conditional allele (19), the RERTert allele (26), the K14-CreERT2T transgene (27), and the MMTV-PyVTT transgene (28). After the appropriate crosses, we obtained the experimental (Aurkalox/lox; RERTert/ert, Aurkalox/lox; TgK14-CreERT2+/T and Aurkalox/lox; RERTert/ert; MMTV-PyVT+/T), and control (Aurka+/+; RERTert/ert, Aurka+/lox; TgK14-CreERT2+/T and Aurka+/lox; RERTert/ert; MMTV-PyVT+/T) mice used in this work. These animals were maintained in a mixed background (129/Sv, CD1, C57BL/6J, and FVB/N), and were genotyped as reported previously (19, 26–28). Mice were housed in the pathogen-free animal facility of the Centro Nacional de Investigaciones Oncológicas (Madrid) in accordance with the animal-care standards of the institution. These animals were observed on a daily basis, and sick mice were killed humanely in accordance with the Guidelines for Humane End Points for Animals used in biomedical research. All animal protocols were approved by the Instituto de Salud Carlos III Committee for Animal Care and Research. For Cre activation, mice were either fed with tamoxifen-supplemented food (Harlan Laboratories Models) or topically painted with 5 mg/mL citrate tamoxifen (Cat. #T9262, Sigma-Aldrich). Conversion of Aurkalox allele to AurkaΔ allele was determined by Southern blot or PCR (complete protocols, primers, and probes are available from the authors upon request). For Western blotting, tissues were harvested and lysed in Laemmli buffer, and 50 μg of total protein was separated by SDS-PAGE and probed with antibodies against Aurora-A (ab13824, AbCam) and Vinculin (#V9131, Sigma-Aldrich).

For histologic observation, dissected organs were fixed in 10%-buffered formalin (Sigma) and embedded in paraffin wax. Sections of 3- or 5-μm thickness were stained with haematoxylin and eosin (H&E). Immunohistochemical examinations of the tissues were performed using specific antibodies against Aurora-A (610938, BD Transduction Laboratories), Ki67 (0003110QD, Master Diagnostica), p21 (sc-397-G, Santa Cruz Biotechnology), p53 (NCL-p53-CM5p, Leica), γ-H2AX (05-636; Millipore), or active caspase-3 (AF835, R&D Systems). Bone mineral density (BMD) and percentage of fat was determined from whole-body samples (excluding cranial region) using a Lunar PIXImus densitometer (GE Medical Systems). Hematopoietic parameters in blood samples were obtained using an Abacus Junior Vet Hematology Counter (Practice CVM).

Mouse depilation and regeneration assays were performed as previously described (29). Labeling retaining experiments were based on previously reported procedures (30). The main modification consisted in the use of paraformaldehyde-fixed skin samples embedded instead of whole-mount specimens.

Senescence assays

Senescence was examined in cells by a senescence-associated β-galactosidase assay (9860, Cell Signaling). Cells were fixed in a 2% formaldehyde and 0.2% glutaraldehyde solution for 15 minutes, washed in PBS, and stained overnight at 37°C in an X-gal staining solution. For tissues and tumor samples, whole-mount staining was performed using the same procedure. Stained samples were embedded and sectioned to visualize cell-specific expression of the reporter.

Tumorigenic experiments

Skin carcinogenesis induced by 7,12-dimethylbenz(a)-anthracene (DMBA) and 12-O-tetradecanoylphorbol-13-acetate (TPA) was performed as previously reported (31). Tumors were measured weekly in two bisecting diameters by using a caliper. Mice developed multiple lesions that were taken together to calculate total tumor mass per mouse. Mammary gland tumors were obtained by crossing the Aurkalox/lox model with MMTV-PyVT+/T transgenic mice (28). Females showed palpable tumors at 11 to 13 weeks of age. Since that moment, mice were fed with tamoxifen-supplemented food and tumor volumes scored weekly by microtomography using an eXplore Vista scanner (GE Healthcare). Tumor volume was calculated using the formula [sagittal dimension (mm) × cross dimension (mm)2]/2.

Generation, culture, and characterization of mouse embryonic fibroblasts

Mouse embryonic fibroblasts (MEF) were generated from E13.5 embryos and cultured using standard protocols (32). To eliminate Aurora-A from Aurkalox/lox; RERTert/ert MEFs, we followed the synchronization scheme depicted in Supplementary Fig. S1. We added 4-OHT (100 nmol/L final concentration; HT-904, Sigma-Aldrich) or infected the cells with adenoviruses expressing Cre (supplied by the Iowa University). The same MEFs incubated with vehicle or infected with adenoviruses expressing Flp were used as controls, respectively. For immmunfluorescence, cells were rinsed with PBS and fixed in 4% PFA-PBS for 7 minutes and, then, left in cold methanol overnight at 20°C. After being blocked with 1% BSA for 1 hour, phospho-histone H3 (P-H3) was detected using a specific antibody from Millipore (05-806). Images were acquired using a Leica D3000 microscope or confocal ultraspectral microscope Leica TCS-SP5. For time-lapse imaging experiments, asynchronous, H2B-GFP-expressing cells were recorded (5-minute frames during 13 hours) using a DeltaVision RT imaging system (Applied Precision, LLC; IX70/71; Olympus) equipped with a charge-coupled device camera (CoolSNAP HQ; Roper Scientific). Colony-formation assays were performed in immortal Aurkalox/lox; RERTert/ert MEFs that stably expressed the viral oncoproteins E1A or 16E6. A total of 5,000 cells were plated in triplicate and were nontreated or treated with 4-OHT. Three weeks later, colonies were stained with Giemsa and counted.

Karyotyping and scoring of aneuploidy

For metaphase spreads, cells were hypotonically swollen in 40% full-medium, 60% tap water for 5.5 minutes. Hypotonic treatment was stopped by adding an equal volume of Carnoy solution (75% pure methanol, 25% glacial acetic acid), cells were then spun down, and fixed with Carnoy solution for 10 minutes. After fixation, cells are dropped from a 5-cm height onto glass slides previously treated with 45% of acetic acid. Slides were mounted with ProLong Gold anti-fade reagent with 40,6-diamidino-2-phenylindole (DAPI; Invitrogen), and images were acquired with a Leica D3000 microscope and an ×60 PlanApo N 1.42 N.A. objective. Chromosomes from 30 cells per genotype were counted. In addition, aneuploidy was determined by measuring nuclear volumes in both cells growing in culture and tissues. Interphasic nucleus volumes were calculated using the following algorithm: 4/3*π*r3, where r = Feret diameter/2. Feret diameter was calculated using ImageJ on images of DAPI-stained fibroblasts or H&E-stained tissues.

Statistical and imaging analyses

Statistical analyses were performed using the Student t, χ2, or log-rank tests (GraphPad Prism 5). All data are shown as mean ± SD; probabilities of P < 0.05 were considered significant. Images were quantified using ImageJ (National Institutes of Health, Bethesda, Maryland).

Genetic ablation of Aurora-A induces mitotic defects, aneuploidy, and senescence in vitro

Because the lack of Aurora-A is not compatible with mouse development (19–21), a conditional mutant was generated by crossing the AurAlox conditional allele (19) with the RERTert allele expressing a 4-OHT-inducible Cre recombinase under the RNA polymerase II regulatory sequences (26). We first used Aurkalox/lox; RERTert/ert MEFs to eliminate Aurora-A upon the addition of 4-OHT. Aurka-null cultures were characterized by a decrease in Aurka protein levels and activity (Supplementary Fig. S1), and accumulated a high number of phospho-histone H3-positive cells, in correlation with a significant increase in the duration of mitosis (Fig. 1A and Supplementary Fig. S1). In agreement with previous reports (19–21), very few Aurka-null cells were able to normally segregate their DNA during mitosis (7% of the AurkaΔ/Δ vs. 97% of the control Aurkalox/lox cells, n = 88 and n = 143, respectively, P < 0.0001). These mutant cells mainly exited mitosis without chromosome segregation or with abnormal chromosome segregation in the presence of lagging chromosomes and chromosome bridges (41% and 50%, respectively, of Aurka-null cells entering mitosis, n = 88; Fig. 1A, Supplementary Fig. S2, and Supplementary Videos). This led to the accumulation of giant nuclei, characteristic of polyploid cells (Fig. 1B), as well as an increase in the aneuploid cell population (Fig. 1C). No differences were observed in the percentage of apoptotic cells in Aurkalox/lox; RERTert/ert MEFs treated or nontreated with 4-OHT throughout the 2-week observation period depicted in Supplementary Fig. S1 (data not shown). By contrast, the percentage of senescent cells was significantly increased among the Aurkalox/lox MEFs infected with Adeno-Cre when compared with the control ones infected with Adeno-EGFP (61.84 ± 5.83 vs. 14.59 ± 1.97, P < 0.0001; Fig. 1D). We next engineered Aurkalox/lox; RERTert/ert MEFs to stably express oncoproteins E1A and 16E6, which inactivate the retinoblastoma protein (pRb) or p53, respectively. These two different cell lines were equally sensitive to the lack of Aurora-A when subjected to a clonogenic assay (Fig. 1E), suggesting that the proliferative defects induced by Aurora-A loss are independent of p53 and pRb function.

Figure 1.

Cellular defects induced by Aurora-A depletion in mouse fibroblasts are p53 and pRb dependent. A, time-lapse imaging of wild-type and Aurora-A null cells stably expressing EGFP-H2B. Asynchronous, immortal Aurkalox/lox; RERTert/ert MEFs were treated with 4-OHT or vehicle for 48 hours and then analyzed by fluorescence microscopy. Pictures were collected every 5 minutes during 13 hours. Mitosis time zero was considered at the first frame showing condensed DNA. Mitosis was considered to have ended at the first frame in which the mitotic cell decondensed its DNA. Pictures are representative of the mitotic phenotypes observed in control and Aurora-A defective MEFs. Left, the bar graph shows a quantitation of mitotic length in Aurora-A null and control MEFs. Right, the bar graph represents the percentage of fibroblasts exiting mitosis without performing cytokinesis. The duration of mitosis was significantly longer in Aurora-A–depleted MEFs than in control cells (81.47 ± 10.33 vs. 159.6 ± 13.92 minutes, P < 0.0001). In addition, a significant high number of Aurora-A null cells were not able to segregate chromosomes and exited mitosis as a single tetraploid cell (63.00 ± 4.66 vs. 24.50 ± 8.37 in control cells, P = 0.0038; 34–38 cells were scored in each condition; ***, P < 0.0001). B, aneuploidy was induced upon Aurora deletion. Nuclear size of MEFs (four different clones incubated with 4-OHT or vehicle) was measured at the indicated times after serum re-addition. As shown in the bar graph, AurkaΔ/Δ nuclei were significantly bigge than the Aurkalox/lox control ones. Representative DAPI-stained images are shown. Scale bar, 40 μm. C, chromosomes were counted in individual cells of the previous experiment at day 5 after serum re-addition as indicated in Material and Methods. Bar graphs show the chromosome numbers (average of the four different clones) found in metaphases of Aurora-A–deficient and control MEFs. D, significant induction of senescence in AurkaΔ/Δ MEFs. Primary Aurkalox/lox; RERTert/erx MEFs were treated as in Supplementary Fig. S1A and assayed for SA-βGal activity 2 weeks after serum re-addition. Representative micrographs and the quantitation of SA-βGal–positive cells are shown. n = 6; ***, P < 0.0001. Scale bar, 40 μm. E, clonogenic experiment to test if the inactivation of either p53 or Rb pathways can rescue the lack of proliferation induced by Aurora-A abrogation. Aurkalox/lox; RERTert/erx (lox/lox) MEFs stably expressing E1A (for Rb pathway inactivation) or 16E6 (for p53 pathway inactivation) were seeded and treated with 4-OHT (+4-OHT) or with vehicle (−4-OHT). Two weeks later, colonies were stained and quantified. No colonies were observed in all the cases that were treated with 4-OHT.

Figure 1.

Cellular defects induced by Aurora-A depletion in mouse fibroblasts are p53 and pRb dependent. A, time-lapse imaging of wild-type and Aurora-A null cells stably expressing EGFP-H2B. Asynchronous, immortal Aurkalox/lox; RERTert/ert MEFs were treated with 4-OHT or vehicle for 48 hours and then analyzed by fluorescence microscopy. Pictures were collected every 5 minutes during 13 hours. Mitosis time zero was considered at the first frame showing condensed DNA. Mitosis was considered to have ended at the first frame in which the mitotic cell decondensed its DNA. Pictures are representative of the mitotic phenotypes observed in control and Aurora-A defective MEFs. Left, the bar graph shows a quantitation of mitotic length in Aurora-A null and control MEFs. Right, the bar graph represents the percentage of fibroblasts exiting mitosis without performing cytokinesis. The duration of mitosis was significantly longer in Aurora-A–depleted MEFs than in control cells (81.47 ± 10.33 vs. 159.6 ± 13.92 minutes, P < 0.0001). In addition, a significant high number of Aurora-A null cells were not able to segregate chromosomes and exited mitosis as a single tetraploid cell (63.00 ± 4.66 vs. 24.50 ± 8.37 in control cells, P = 0.0038; 34–38 cells were scored in each condition; ***, P < 0.0001). B, aneuploidy was induced upon Aurora deletion. Nuclear size of MEFs (four different clones incubated with 4-OHT or vehicle) was measured at the indicated times after serum re-addition. As shown in the bar graph, AurkaΔ/Δ nuclei were significantly bigge than the Aurkalox/lox control ones. Representative DAPI-stained images are shown. Scale bar, 40 μm. C, chromosomes were counted in individual cells of the previous experiment at day 5 after serum re-addition as indicated in Material and Methods. Bar graphs show the chromosome numbers (average of the four different clones) found in metaphases of Aurora-A–deficient and control MEFs. D, significant induction of senescence in AurkaΔ/Δ MEFs. Primary Aurkalox/lox; RERTert/erx MEFs were treated as in Supplementary Fig. S1A and assayed for SA-βGal activity 2 weeks after serum re-addition. Representative micrographs and the quantitation of SA-βGal–positive cells are shown. n = 6; ***, P < 0.0001. Scale bar, 40 μm. E, clonogenic experiment to test if the inactivation of either p53 or Rb pathways can rescue the lack of proliferation induced by Aurora-A abrogation. Aurkalox/lox; RERTert/erx (lox/lox) MEFs stably expressing E1A (for Rb pathway inactivation) or 16E6 (for p53 pathway inactivation) were seeded and treated with 4-OHT (+4-OHT) or with vehicle (−4-OHT). Two weeks later, colonies were stained and quantified. No colonies were observed in all the cases that were treated with 4-OHT.

Close modal

Conditional ablation of Aurora-A in adult mice

To analyze the consequences of Aurora-A ablation in an adult organism, Aurka+/+; RERTert/ert or Aurkalox/lox; RERTert/ert mice were fed with tamoxifen at 1 to 2 month of age for a minimum period of time of 4 weeks. As shown in Fig. 2A, a partial although significant reduction of the conditional alleles (Aurkalox) in favor of the deleted ones (AurkaΔ) was observed in Aurkalox/lox; RERTert/ert tissues after 4 weeks of treatment. This was associated with a significant reduction of the expression of Aurora-A protein in different tissues (Fig. 2B–D). Histopathologic studies of Aurora-A–deficient (hereafter, AurkaΔ/Δ) mice did not show major abnormalities in organs with low proliferative ratio such as the kidney, heart, pancreas, and brain (data not shown). However, tissues with high proliferation rates showed marked defects. We observed significant testicular atrophy, thinner and immature, epidermis as well as atrophy of the intestine, thymus, and spleen (Fig. 2C; Supplementary Fig. S3). Testis from AurkaΔ/Δ mice showed seminiferous tubules with reduced quantity of germinal cells and lack of mature spermatozoa. Clear-cell depletion with reduced organ size were found in both red and white pulp in the spleen of AurkaΔ/Δ mice, with the red pulp being the most affected by a significant reduction in the number of cells. In the skin, although control mice showed hair follicles reaching the paniculus carnosus muscle, AurkaΔ/Δ hair follicles were mainly at the catagen–telogen stage, therefore, located entirely within the dermis. The intestine showed decreased crypts and villi length with abnormal representations of normal cell populations.

Figure 2.

Conditional ablation of Aurora-A in adult mice affects proliferative tissues. A, Southern blot of genomic DNA isolated from different tissues of tamoxifen-treated wild-type (+/+; Aurka+/+; RERTert/ert) or Aurora-A null (Δ/Δ; Aurkalox/lox; RERTert/ert) mice. SP, spleen; KD, kidney; LG, lung. Wild-type (+), knock out (Δ), and conditional (lox) bands are indicated. B, immunoblot analysis in spleen, skin, and testis of wild-type (+/+) and Aurora-A null (Δ/Δ) mice shows a strong reduction in expression of Aurora-A protein. Detection of vinculin in the same samples was used for loading normalization. C, representative images for the abnormalities detected in spleen, skin, and testis of Aurora-null mice. Scale bars, 100 μm. Right, Aurora-A expression is significantly diminished upon tamoxifen treatment in Aurkalox/lox; RERTert/ert mice. Aurora-A immunohistochemical detection is shown for three representative proliferative tissues. Bar graphs indicate the quantification of the number of Aurora-A–positive cells (n = 3 mice per genotype; *, P < 0.05; **, P < 0.01). Scale bars, 100 μm.

Figure 2.

Conditional ablation of Aurora-A in adult mice affects proliferative tissues. A, Southern blot of genomic DNA isolated from different tissues of tamoxifen-treated wild-type (+/+; Aurka+/+; RERTert/ert) or Aurora-A null (Δ/Δ; Aurkalox/lox; RERTert/ert) mice. SP, spleen; KD, kidney; LG, lung. Wild-type (+), knock out (Δ), and conditional (lox) bands are indicated. B, immunoblot analysis in spleen, skin, and testis of wild-type (+/+) and Aurora-A null (Δ/Δ) mice shows a strong reduction in expression of Aurora-A protein. Detection of vinculin in the same samples was used for loading normalization. C, representative images for the abnormalities detected in spleen, skin, and testis of Aurora-null mice. Scale bars, 100 μm. Right, Aurora-A expression is significantly diminished upon tamoxifen treatment in Aurkalox/lox; RERTert/ert mice. Aurora-A immunohistochemical detection is shown for three representative proliferative tissues. Bar graphs indicate the quantification of the number of Aurora-A–positive cells (n = 3 mice per genotype; *, P < 0.05; **, P < 0.01). Scale bars, 100 μm.

Close modal

AurkaΔ/Δ mice showed external phenotypes, such as abnormal curvature of the upper spine and loss of hair—hallmarks typically associated with an age-related phenotype (Fig. 3A). Another common feature of the AurkaΔ/Δ mice was a reduction in their body weight. This reduction was observed as early as 1 week after the start of the tamoxifen treatment and, 3 weeks later, it reached a 30% of the body weight (Fig. 3B). These defects were not observed in control mice (Aurka+/+; RERTert/ert mice treated with tamoxifen; hereafter, Aurka+/+ mice), which showed a slight (8%) and transient reduction in body weight within the first week (Fig. 3B). Importantly, Aurora-A ablation compromised the survival of adult mice because 40% of treated animals died during the 40 weeks following the tamoxifen treatment whereas all control mice survived in the presence of tamoxifen (Fig. 3C). Histopathologic analysis of sick mice showed a pleiotropic phenotype preventing us from establishing a single common cause of death in all the cases. Several features found in Aurora-A null mice, such as thinner and immature epidermis, and atrophy of intestine, thymus, and spleen, could well be associated with death as a result of starvation, infection, dehydration, and/or multiorganic failure. Because the abnormal curvature of the upper spine, a condition called kyphosis, found in AurkaΔ/Δ mice, could appear as a result of osteoporosis, we decided to analyze the bone density of Aurkalox/lox; RERTert/ert mice treated with tamoxifen for a long period of time. As it is shown in Fig. 3D, the densitometer images from Aurkalox/lox; RERTert/ert mice treated with tamoxifen for 4 months displayed a 10% decrease in their bone density when compared with control mice (0.0524 ± 0.0004 vs. 0.058 ± 0.0012 g/cm3, P < 0.01). In addition, blood cells and a number of hematopoietic parameters were significantly reduced in the Aurora-A–depleted mice (Table 1). The reduction was significant in the case of lymphocytes, granulocytes, erythrocytes, platelets, hemoglobin, and the hematocrit.

Figure 3.

Significant reduction of Aurora-A protein levels in adult mice induces loss of weight, decrease bone density, low blood cell numbers, and severe abnormalities in proliferative tissues. A, external view of Aurka+/+; RERTert/ert and Aurkalox/lox; RERTert/ert mice treated with tamoxifen during the indicated times. B, Aurkalox/lox; RERTert/ert mice treated with tamoxifen (Δ/Δ) show a significant loss of weight when compared with control (+/+) mice (P < 0.0001, n = 3). C, Aurora-A deletion results also in lethality. Graph shows the survival curve of 1- to 2.5-month old mice fed with tamoxifen for a total of 38 weeks. Survival curves of Aurka+/+; RERTert/ert (n = 9) and Aurkalox/lox; RERTert/ert (n = 17) mice were compared using a log-rank (Mantel–Cox) test and showed differences close to the significance (P = 0.0814). D, bone densitometry of total body shows a significant reduction in the bone density of Aurora-A Δ/Δ mice when compared with control mice (0.0524 ± 0.0004, n = 4, vs. 0.058 ± 0.0012, n = 3, g/cm3, P < 0.01).

Figure 3.

Significant reduction of Aurora-A protein levels in adult mice induces loss of weight, decrease bone density, low blood cell numbers, and severe abnormalities in proliferative tissues. A, external view of Aurka+/+; RERTert/ert and Aurkalox/lox; RERTert/ert mice treated with tamoxifen during the indicated times. B, Aurkalox/lox; RERTert/ert mice treated with tamoxifen (Δ/Δ) show a significant loss of weight when compared with control (+/+) mice (P < 0.0001, n = 3). C, Aurora-A deletion results also in lethality. Graph shows the survival curve of 1- to 2.5-month old mice fed with tamoxifen for a total of 38 weeks. Survival curves of Aurka+/+; RERTert/ert (n = 9) and Aurkalox/lox; RERTert/ert (n = 17) mice were compared using a log-rank (Mantel–Cox) test and showed differences close to the significance (P = 0.0814). D, bone densitometry of total body shows a significant reduction in the bone density of Aurora-A Δ/Δ mice when compared with control mice (0.0524 ± 0.0004, n = 4, vs. 0.058 ± 0.0012, n = 3, g/cm3, P < 0.01).

Close modal
Table 1.

Aurora-A–deficient mice are defective in several hematologic parameters

Aurka+/+ (n = 8)AurkaΔ/Δ (n = 8)
Hematologic parameterMeanSEMMeanSEMP
WBC (×109/L) 9.82 1.34 4.86 0.98 ** 
LYM (×109/L) 7.23 1.07 3.75 0.82 
MID (×109/L) 0.41 0.08 0.27 0.82 0.11 
GRA (×109/L) 2.18 0.65 0.84 0.27 
RBC (×1012/L) 8.68 0.24 5.15 0.73 *** 
HGB (g/dL) 13.31 0.59 8.21 1.36 ** 
HCT (%) 40.20 2.21 24.14 2.95 *** 
MCV (fl) 46.00 2.09 48.88 2.84 0.21 
MCH (pg) 15.33 0.59 15.80 0.82 0.32 
MCHC (g/dL) 33.56 1.60 32.84 1.67 0.38 
RDWc (%) 18.21 0.75 21.79 1.88 
PLT (×109/L) 741.3 191.20 297.60 110.50 
PCT (%) 0.45 0.11 0.24 0.09 0.08 
MPV (fl) 5.87 0.15 7.40 0.69 
PDWc 32.15 0.53 37.26 1.57 ** 
Aurka+/+ (n = 8)AurkaΔ/Δ (n = 8)
Hematologic parameterMeanSEMMeanSEMP
WBC (×109/L) 9.82 1.34 4.86 0.98 ** 
LYM (×109/L) 7.23 1.07 3.75 0.82 
MID (×109/L) 0.41 0.08 0.27 0.82 0.11 
GRA (×109/L) 2.18 0.65 0.84 0.27 
RBC (×1012/L) 8.68 0.24 5.15 0.73 *** 
HGB (g/dL) 13.31 0.59 8.21 1.36 ** 
HCT (%) 40.20 2.21 24.14 2.95 *** 
MCV (fl) 46.00 2.09 48.88 2.84 0.21 
MCH (pg) 15.33 0.59 15.80 0.82 0.32 
MCHC (g/dL) 33.56 1.60 32.84 1.67 0.38 
RDWc (%) 18.21 0.75 21.79 1.88 
PLT (×109/L) 741.3 191.20 297.60 110.50 
PCT (%) 0.45 0.11 0.24 0.09 0.08 
MPV (fl) 5.87 0.15 7.40 0.69 
PDWc 32.15 0.53 37.26 1.57 ** 

NOTE: *, P < 0.05; **, P < 0.01; ***, P < 0.001.

Abbreviations: GRA, granulocytes count; HCT, hematocrit; HGB hemoglobin; LYM, lymphocyte count; MCH, mean corpuscular hemoglobin; MCHC, mean corpuscular hemoglobin concentration; MCV, mean corpuscular volume; MID, monocytes/eosinophils count; MPV mean platelet volume; PCT, platelet percentage; PDWc, platelet distribution width; PLT, platelet count; RBC, red blood cell count; RDWc red cell distribution width; WBC, total white blood cell count.

Cellular defects associated with Aurora-A loss in vivo

In the absence of Aurora-A, several proliferative tissues such as the skin, spleen, and thymus showed a significant reduction in the expression of the proliferation marker Ki67 (Fig. 4A, Supplementary Fig. S4). In addition, this pattern was observed in other tissues characterized by low proliferation rates, such as the lung, liver, kidney, heart, and pancreas (Supplementary Fig. S4). A detailed analysis of one of the proliferative tissues (spleen) allowed us to visualize among the AurkaΔ/Δ tissues an accumulation of mitotic figures that could correspond to cells arrested in prophase- or prometaphase–like stages (Fig. 4B). In fact, a significant increase of phospho-histone H3 signal, a typical mitotic marker, was found in the AurkaΔ/Δ spleen when compared with those of control mice (Fig. 4C). The AurkaΔ/Δ spleens are also characterized for the presence of cells with a significant increase in the nuclear size (Fig. 4D). In addition, this phenotype was also detected in other cell types, such as the skin, the erythroid lineage of the spleen as well as in specific lineages that undergo spontaneous polyploidization such as the hepatocytes (Supplementary Figs. S5 and S6).

Figure 4.

Depletion of Aurora-A in adult spleen is associated with lack of proliferation, mitotic arrest, aneuploidy, the accumulation of antiproliferative and DNA damage markers, and the induction of apoptosis and senescence. A, micrographs show, at different magnifications, the marked and prevalent signal of KI67 in control (+/+) spleen and the lack of, or weak, KI67 signal in Δ/Δ spleen. Bar graphs show the quantification of KI67-positive cells in +/+ and Δ/Δ mice. Three mice of each group were used in this analysis. **, P < 0.01. Scale bars are 200 μm in panels and 50μm in subpanels. B, a significant increased number of mitotic figures were detected in the cells from the red pulp of the spleen of Δ/Δ mice when compared with the +/+ control ones. Scale bar, 20 μm. Bar graphs show the quantification of mitotic arrested cells in +/+ and Δ/Δ spleens (n = 3; ***, P < 0.001). C, accumulation of P-H3 positive cells occurs in the red pulp of Aurora-A depleted spleen. Pictures show representative P-H3 detection in the spleen of Δ/Δ and +/+ mice. Scale bar, 50 μm. Bar graphs show the significant increase in the percentage of P-H3 positive cells found in Δ/Δ spleens when compared with control samples (n = 3; ***, P < 0.001). D, aneuploidy is also a common feature of Δ/Δ spleens. Nuclear size was used to compare DNA content of Aurora-A defective and control cells. Images show hematoxylin and eosin-stained spleen sections where nuclei have been highlighted. Scale bar, 20 μm. Bar graph shows the quantification of the nuclear volumes in Δ/Δ samples (1.95 ± 0.07, n = 279 nuclei) normalized to the control cases (0.99 ± 0.03, n = 280 nuclei). Three different mice per group (***, P < 0.001). E, representative p53, p21, and γH2AX staining in spleen sections from Aurka+/+; RERTert/erx (+/+) and Aurkalox/lox; RERTert/erx (Δ/Δ) mice treated with tamoxifen. Scale bar, 100 μm. Quantification of the number of positive cells in both groups (n = 3 in each one) is shown in bar graphs. *, P < 0.05; **, P < 0.01; ***, P < 0.001. F, apoptosis was analyzed in spleen sections using caspase-3 detection. Scale bar, 100 μm. Few caspase-3–positive cells were found in Δ/Δ and +/+ spleens (3.41 ± 0.19 and 1.88 ± 0.43, respectively). Importantly, the percentage of caspase-3–positive cells was significantly higher among the Aurora-A null samples than in the control samples (P < 0.05, n = 3). G, to study the possible induction of senescence upon Aurora-A deletion, senescence-associated β-Gal activity was assayed in whole-mount preparations of spleen specimens from +/+ and Δ/Δ mice. Low and high magnification micrographs of representative cases are shown. Scale bar, 50 μm. Bar graph shows the significant increase in senescent cells found among the Δ/Δ spleens when compared with the control spleens (61.56 ± 4.45% vs. 13.93 ± 0.33, P < 0.001, n = 3).

Figure 4.

Depletion of Aurora-A in adult spleen is associated with lack of proliferation, mitotic arrest, aneuploidy, the accumulation of antiproliferative and DNA damage markers, and the induction of apoptosis and senescence. A, micrographs show, at different magnifications, the marked and prevalent signal of KI67 in control (+/+) spleen and the lack of, or weak, KI67 signal in Δ/Δ spleen. Bar graphs show the quantification of KI67-positive cells in +/+ and Δ/Δ mice. Three mice of each group were used in this analysis. **, P < 0.01. Scale bars are 200 μm in panels and 50μm in subpanels. B, a significant increased number of mitotic figures were detected in the cells from the red pulp of the spleen of Δ/Δ mice when compared with the +/+ control ones. Scale bar, 20 μm. Bar graphs show the quantification of mitotic arrested cells in +/+ and Δ/Δ spleens (n = 3; ***, P < 0.001). C, accumulation of P-H3 positive cells occurs in the red pulp of Aurora-A depleted spleen. Pictures show representative P-H3 detection in the spleen of Δ/Δ and +/+ mice. Scale bar, 50 μm. Bar graphs show the significant increase in the percentage of P-H3 positive cells found in Δ/Δ spleens when compared with control samples (n = 3; ***, P < 0.001). D, aneuploidy is also a common feature of Δ/Δ spleens. Nuclear size was used to compare DNA content of Aurora-A defective and control cells. Images show hematoxylin and eosin-stained spleen sections where nuclei have been highlighted. Scale bar, 20 μm. Bar graph shows the quantification of the nuclear volumes in Δ/Δ samples (1.95 ± 0.07, n = 279 nuclei) normalized to the control cases (0.99 ± 0.03, n = 280 nuclei). Three different mice per group (***, P < 0.001). E, representative p53, p21, and γH2AX staining in spleen sections from Aurka+/+; RERTert/erx (+/+) and Aurkalox/lox; RERTert/erx (Δ/Δ) mice treated with tamoxifen. Scale bar, 100 μm. Quantification of the number of positive cells in both groups (n = 3 in each one) is shown in bar graphs. *, P < 0.05; **, P < 0.01; ***, P < 0.001. F, apoptosis was analyzed in spleen sections using caspase-3 detection. Scale bar, 100 μm. Few caspase-3–positive cells were found in Δ/Δ and +/+ spleens (3.41 ± 0.19 and 1.88 ± 0.43, respectively). Importantly, the percentage of caspase-3–positive cells was significantly higher among the Aurora-A null samples than in the control samples (P < 0.05, n = 3). G, to study the possible induction of senescence upon Aurora-A deletion, senescence-associated β-Gal activity was assayed in whole-mount preparations of spleen specimens from +/+ and Δ/Δ mice. Low and high magnification micrographs of representative cases are shown. Scale bar, 50 μm. Bar graph shows the significant increase in senescent cells found among the Δ/Δ spleens when compared with the control spleens (61.56 ± 4.45% vs. 13.93 ± 0.33, P < 0.001, n = 3).

Close modal

Furthermore, we tested if these abnormal phenotypes were accompanied by cell-cycle checkpoint activation. As represented in Fig. 4E, both p53 and its effector p21Cip1 were significantly upregulated in the spleen of Aurora-A–depleted mice. This was accompanied by phosphorylation of H2AX (γ-H2AX) (Fig. 4E), a marker of DNA damage, as well as a significant increase in apoptosis and senescence (Fig. 4F and G). Importantly, although the increase in apoptosis was moderate (3.41 ± 0.19% in AurkaΔ/Δ spleen cells vs. 1.88 ± 0.43 in Aurka+/+ mice, P = 0.032, n = 3 mice), a more clear induction of senescence was observed in the red pulp of the spleen (61.56 ± 4.45% in AurkaΔ/Δ mice vs. 13.93± 0.33 in Aurka+/+ mice, P = 0.0002, n = 3). Similar defects in mitosis, nuclear size, and p53 induction were found in the skin for p53 (Supplementary Fig. S5).

Aurora-A ablation in tissue regeneration and tumor growth

To investigate the consequences of Aurora-A ablation in tissue regeneration in vivo, we made use of the hair follicle cycle. Aurkalox/lox; RERTert/ert and Aurka+/+; RERTert/ert mice were depilated on 2-cm3 patches of their back-skin and treated with tamoxifen for 5 days. Seven days later, although Aurka+/+ control mice were able to repopulate the back-skin with new hair, this was not the case with the AurkaΔ/Δ mice (Fig. 5A). Follicular hairs in the Aurora-A null mice arrested at the last stage of the hair follicle cycle (telogene) and showed bigger nuclei (Fig. 5A), suggesting that Aurora-A–depleted cells were able to duplicate their genome but could not divide.

Figure 5.

Lack of Aurora-A affects hair regeneration. A, top scheme summarizes the protocol followed to test the regenerative capacity of Aurora-A–depleted hair follicles. Part of the back skin of Aurka+/+; RERTert/ert (n = 3) and Aurkalox/lox; RERTert/ert (n = 3) mice was depilated and topically treated with tamoxifen. Left, pictures show how, 1 week later, hair recovery was observed in +/+ control mice whereas no hair repopulation was detected in Δ/Δ mice. Histologic analysis (hematoxylin and eosin staining) shows defects in the skin of Δ/Δ samples. Specifically, hair follicles were in the last stage of their cycle and empty. Moreover, skin cells showed bigger nuclei. Bar graph shows the quantification of the nucleus volume in two wild-type and two Aurora-A null samples (n = 41–68 nuclei, ***, P < 0.001). Scale bar, 100 μm. B, labeling retaining experiment demonstrating that hair follicle stem cells do not proliferate upon Aurora-A depletion. BrdUrd was intraperitoneally injected during 4 days into Aurkalox/lox; RERTert/ert mice just before the first synchronized hair cycle ended. Thirty-one days later, before the second synchronized hair cycle was over, mice were depilated and topically treated on the depilated area and on the tail with either tamoxifen or vehicle (DMSO). Micrographs show, at different magnifications, BrdUrd detection in skin 14 days post tamoxifen and DMSO treatment. Scale bar, 100 μm. Bar graph shows nucleus volumes of BrdUrd-positive and -negative cells. Although no differences were detected between the nucleus size of the BrdUrd-negative and -positive cells of the lox/lox samples, Δ/Δ nuclei positive for BrdUrd were significantly bigger than those negative for BrdUrd (n = 7–12; **, P < 0.01).

Figure 5.

Lack of Aurora-A affects hair regeneration. A, top scheme summarizes the protocol followed to test the regenerative capacity of Aurora-A–depleted hair follicles. Part of the back skin of Aurka+/+; RERTert/ert (n = 3) and Aurkalox/lox; RERTert/ert (n = 3) mice was depilated and topically treated with tamoxifen. Left, pictures show how, 1 week later, hair recovery was observed in +/+ control mice whereas no hair repopulation was detected in Δ/Δ mice. Histologic analysis (hematoxylin and eosin staining) shows defects in the skin of Δ/Δ samples. Specifically, hair follicles were in the last stage of their cycle and empty. Moreover, skin cells showed bigger nuclei. Bar graph shows the quantification of the nucleus volume in two wild-type and two Aurora-A null samples (n = 41–68 nuclei, ***, P < 0.001). Scale bar, 100 μm. B, labeling retaining experiment demonstrating that hair follicle stem cells do not proliferate upon Aurora-A depletion. BrdUrd was intraperitoneally injected during 4 days into Aurkalox/lox; RERTert/ert mice just before the first synchronized hair cycle ended. Thirty-one days later, before the second synchronized hair cycle was over, mice were depilated and topically treated on the depilated area and on the tail with either tamoxifen or vehicle (DMSO). Micrographs show, at different magnifications, BrdUrd detection in skin 14 days post tamoxifen and DMSO treatment. Scale bar, 100 μm. Bar graph shows nucleus volumes of BrdUrd-positive and -negative cells. Although no differences were detected between the nucleus size of the BrdUrd-negative and -positive cells of the lox/lox samples, Δ/Δ nuclei positive for BrdUrd were significantly bigger than those negative for BrdUrd (n = 7–12; **, P < 0.01).

Close modal

To further understand whether the lack of Aurora-A affects activation and/or differentiation of hair follicle stem cells, we studied the presence of labeling-retaining cells at the hair follicles. Aurkalox/lox; RERTert/ert mice were injected with bromodeoxyuridine (BrdUrd) at P14 (when the first synchronized hair cycle ends) to label their stem cells. Just before the second hair cycle was completed, mice were depilated on their backs and treated topically (back skin and tail) with either dimethyl sulfoxide (DMSO) or tamoxifen. Two weeks later, hair growth had recovered in DMSO-treated mice (Fig. 5B). Hair follicles from the recovered area were, in their majority, negative for BrdUrd, indicating that the labeled stem cells were able to proliferate and differentiate. Similar results were observed for Aurka+/+ mice treated with either DMSO or tamoxifen (data not shown). However, Aurkalox/lox mice treated with tamoxifen were not able to repopulate the depilated area, and their hair follicles showed an accumulation of BrdUrd retaining cells from the back-skin and tail-skin confirming the loss of regenerative capacity (Fig. 5B). Furthermore, whereas BrdUrd-positive and -negative cells from Aurkalox/lox cells showed similar nuclear volume, AurkaΔ/Δ BrdUrd-positive cells were significantly bigger than control cells (Fig. 5B), suggesting that depletion of Aurora-A allows DNA replication but inhibits cell division, resulting in tetraploid cells.

The strong requirements for cell proliferation shown earlier for Aurora-A both in vitro and in vivo prompted us to analyze whether Aurora-A is also essential for tumor development. Skin tumors were induced using a two-stage carcinogenesis protocol in Aurkalox/lox and Aurka+/lox mice expressing an inducible Cre recombinase under the control of the Keratine 14 promoter. To test the effect of Aurora-A ablation in growing tumors, tamoxifen was topically applied on the tumors when they reached a volume of 50 mm3 (∼4 weeks after they were first observed). As shown in Fig. 6A, tumors continued to exponentially grow in the control mice (Aurka+/lox; TgK14-CreERT2+) whereas they completely arrested and never progressed in Aurkalox/lox; TgK14-CreERT2+/T mice. The tumor size was significantly different at the end of the experiment, 5 weeks after addition of tamoxifen (relative size of 647.9 ± 395.6 among the Aurka+/Δ tumors, n = 8, vs. 131.1 ± 60.67 among the AurkaΔ/Δ tumors, n = 21; P < 0.05). Immunohistochemical characterization of these tumors confirmed the loss of Aurora-A and lack of proliferation (low Ki67 levels; Fig. 6B). It is important to note that no differences were found in the expression of p53 (data not shown) nor in the levels of apoptotic (Fig. 6B) or senescent cells (data not shown). Therefore, the arrest observed in skin tumors treated with tamoxifen is not p53-dependent or due to an induction of apoptosis or senescence. Interestingly, Aurora-A–null tissues display increased levels of differentiation markers, such as loricrin or involucrin (Supplementary Fig. S6), in agreement with previous reports suggesting that the block of mitosis results in increased differentiation in the skin (33).

Figure 6.

Aurora-A ablation results in arrest of skin and mammary gland tumors in vivo. A, skin tumors were induced in Aurkalox/lox; TgK14CreERT2+/T mice using a DMBA + TPA treatment. The resulting papillomas were treated topically with tamoxifen or carrier (DMSO). Graph shows tumor volumes relative to the pretreatment size (mean ± SEM; *, P < 0.05). B, panels show representative pictures of skin tumor sections in which expression of Aurora-A, Ki67, and caspase-3 (Ca3) is detected. Scale bar, 100 μm. C, depletion of Aurora-A also inhibited the growth of polyomavirus middle T (PyVT) oncogene-induced mammary tumors. Tumor growth was compared in Aurka+/lox; RERTert/ertand Aurkalox/lox; RERTert/ert females that also expressed the MMTV-PyVT transgene. At 11 to 13 weeks of age, mice were treated with tamoxifen, and tumor size was measured every 2 weeks by computed tomography. Graph represents the longitudinal data obtained for seven mice of each group. At time 0, tamoxifen was administered to all the experimental mice. The experiment was completed 10 weeks later. Significant differences were detected in the tumor volumes at 8 and 10 weeks post tamoxifen treatment (*, P < 0.05; n = 4–12 per group in each time point). D, lung metastases were also reduced in Aurora-A null mice (Δ/Δ) when compared with controls (+/Δ). Bar graph shows the quantification of the metastases identified in 10 random fields from three lung sections (*, P < 0.05; n = 5 mice per group). Representative lungs are shown in pictures on the right. Scale bar, 2 mm. E, Δ/Δ tumors did not increase their metabolic activity with time. In 2 weeks (between week 1 and 3 of tamoxifen treatment), +/Δ, tumors significantly increased their metabolic activity, measured by PET (upper bar graph). However, Aurora-A null, Δ/Δ, tumors did not change their metabolic status (***, P < 0.001; n = 10). H, heart; B, bladder. F, nucleus volume in cells from mammary tumors is significantly larger in Δ/Δ samples than in controls. Tumor cells from mammary tumors were grown in culture for a week and, after DAPI staining, a minimum of 23 nuclei were quantified per sample. Bar graph shows nucleus volume of tumor cells from three mice of each type (*, P < 0.05 two-way ANOVA test). Images are representative DAPI-stained nuclei from Δ/Δ and +/Δ tumors. Scale bar, 10 μm.

Figure 6.

Aurora-A ablation results in arrest of skin and mammary gland tumors in vivo. A, skin tumors were induced in Aurkalox/lox; TgK14CreERT2+/T mice using a DMBA + TPA treatment. The resulting papillomas were treated topically with tamoxifen or carrier (DMSO). Graph shows tumor volumes relative to the pretreatment size (mean ± SEM; *, P < 0.05). B, panels show representative pictures of skin tumor sections in which expression of Aurora-A, Ki67, and caspase-3 (Ca3) is detected. Scale bar, 100 μm. C, depletion of Aurora-A also inhibited the growth of polyomavirus middle T (PyVT) oncogene-induced mammary tumors. Tumor growth was compared in Aurka+/lox; RERTert/ertand Aurkalox/lox; RERTert/ert females that also expressed the MMTV-PyVT transgene. At 11 to 13 weeks of age, mice were treated with tamoxifen, and tumor size was measured every 2 weeks by computed tomography. Graph represents the longitudinal data obtained for seven mice of each group. At time 0, tamoxifen was administered to all the experimental mice. The experiment was completed 10 weeks later. Significant differences were detected in the tumor volumes at 8 and 10 weeks post tamoxifen treatment (*, P < 0.05; n = 4–12 per group in each time point). D, lung metastases were also reduced in Aurora-A null mice (Δ/Δ) when compared with controls (+/Δ). Bar graph shows the quantification of the metastases identified in 10 random fields from three lung sections (*, P < 0.05; n = 5 mice per group). Representative lungs are shown in pictures on the right. Scale bar, 2 mm. E, Δ/Δ tumors did not increase their metabolic activity with time. In 2 weeks (between week 1 and 3 of tamoxifen treatment), +/Δ, tumors significantly increased their metabolic activity, measured by PET (upper bar graph). However, Aurora-A null, Δ/Δ, tumors did not change their metabolic status (***, P < 0.001; n = 10). H, heart; B, bladder. F, nucleus volume in cells from mammary tumors is significantly larger in Δ/Δ samples than in controls. Tumor cells from mammary tumors were grown in culture for a week and, after DAPI staining, a minimum of 23 nuclei were quantified per sample. Bar graph shows nucleus volume of tumor cells from three mice of each type (*, P < 0.05 two-way ANOVA test). Images are representative DAPI-stained nuclei from Δ/Δ and +/Δ tumors. Scale bar, 10 μm.

Close modal

Furthermore, we analyzed the effect of Aurora-A deletion in more aggressive tumors induced by the expression of the polyomavirus middle T (PyVT) oncogene in mammary glands (28). We generated Aurka+/lox; RERTert/ert and Aurkalox/lox; RERTert/ert females that also expressed the MMTV-PyVT transgene. As soon as they had palpable tumors (at ∼11–13 weeks of age), mice were treated with tamoxifen to induce the deletion of the Aurora-A gene. A clear increase in the size of multifocal adenocarcinomas was observed in control females (Aurka+/Δ) whereas AurkaΔ/Δ tumors grew at a significantly slower rate (Fig. 6C; tumor size 10 weeks after tamoxifen addition of 4,173 ± 1,380 among the Aurka+/Δ tumors, vs. 1,129 ± 599 among the AurkaΔ/Δ tumors; n = 4, P < 0.05). Furthermore, secondary metastatic tumors in the lung characteristic of this model were significantly reduced among AurkaΔ/Δ mice (P < 0.05; Fig. 6D). These Aurora-A–null tumors were characterized by a significant reduction of the proliferative markers (Ki67) without an increase in the apoptotic (C3A) or senescent (SA-βgal–associated activity) markers (Supplementary Fig. S7). However, AurkaΔ/Δ breast tumors were less active metabolically (PET data in Fig. 6E). Similar to AurkaΔ/Δ skin tumors, the AurkaΔ/Δ mammary gland tumors were characterized by increased levels of differentiation markers (p63 and cK8; Supplementary Fig. S6). In addition, these mutant tumors contained giant nuclei and mitotic abnormalities rarely observed in Aurka+/Δ tumors (Fig. 6F and Supplementary Fig. S2). A similar significant increase in the nuclear volume of AurkaΔ/Δ tumoral cells was observed in the skin adenocarcinomas induced by TPA plus DMBA treatment (Supplementary Fig. S7). All these data suggest that the lack of Aurora-A impairs cell division and induces a polyploid phenotype incompatible with cell proliferation, whereas apoptosis and senescence are rare events associated with Aurora-A elimination in these tumors.

Aurora-A is required for cellular proliferation in cultured cells, and its genetic inactivation results in early embryonic lethality in the mouse and severe defects in skin development (14, 15, 18–21, 34, 35). Due to its critical role in the cell cycle, several Aurora-A inhibitors are currently studied as antitumoral agents (24, 25). Because many of these inhibitors can act in a similar way against the three Aurora family members and also be active against other kinases, it is important to discriminate the specific effects of the inhibition of Aurora-A associated with the use of these inhibitors. Using a conditional knockout model, we show here that Aurora-A is critical for continued cell proliferation in adult animals. Importantly, this phenotype is not compensated by the other two Aurora family members, Aurora-B and Aurora-C, which are highly similar but quite different from Aurora-A (36). In fact, these two groups of Aurora kinases display different subcellular localization and bind to different partners (INCENP/survivin/borealin in the case of Aurora-B/C and TPX2/Bora in the case of Aurora-A). Interestingly, a single amino acid difference allows TPX2 to discriminate between Aurora-A and -B (37), and an Aurora-A mutant for that residue rescues Aurora-B loss of function (38, 39). Our results confirm that, despite the similarity and common origin of mammalian Aurora kinases, these proteins display specific characteristics that limit their possible compensatory roles.

Because Aurora-A inhibitors are not specific for tumor cells, our results provide critical genetic data to separate the relevance of this kinase in vivo and as a cancer target. We have added significant information on the effect of Aurora-A dosage in tumor development and progression. It had been previously shown that overexpression or heterozygous expression of Aurora-A induces malignant transformation (20, 40, 41). Here, we show that its complete loss prevents tumor formation and inhibits the progression of chemical or genetically induced tumorigenesis. As this apparent paradox is common to other mitotic regulators, one possibility is that both partial loss- or gain-of-expression induces chromosomal instability and favors tumorigenesis. On the other hand, as all these proteins are required for the cell cycle, their complete depletion impairs cell proliferation, a feature that has been critical for their consideration as putative cancer targets (25, 42).

Our results in vitro and in vivo suggest that lack of Aurora-A results in defective chromosome segregation and the generation of tetraploid or aneuploid cells. This is accompanied by a stress response characterized by DNA damage and the induction of p53 and p21Cip1. Aneuploid cells are frequently characterized by the induction of a DNA damage response as a consequence of a higher frequency of DNA damage and/or the induction of replicative stress (43). Moreover, abnormal and/or prolonged mitosis has been shown to induce accumulation of DNA breaks (44). More recently, Aurora-A has been identified in a genome-wide siRNA screen as one of the genes whose abrogation led to elevated levels of H2AX phosphorylation (45). This data is of special relevance in tumorigenesis, because it implies that the inhibition of Aurora-A could sensitize tumors to anticancer agents that work better against cancer cells with high levels of DNA damage.

Some Aurora-A inhibitors such as MLN8054 and MLN8237 (Milenium) induce senescence in myeloma and colon cell lines as well as in lung and melanoma xenografts (46–48). We have observed a significant induction of senescence in Aurora-A–depleted spleens. However, neither senescence nor apoptosis seem to be a frequent observation in other cases wherein Aurora-A is eliminated from normal or cancer cells. Nevertheless, our data suggest that Aurora-A inactivation seems to be sufficient to arrest tumor growth in the absence of both apoptosis and senescence (Fig. 6). Cell-cycle arrest caused by Aurora-A deficiency is also independent on p53 or Rb function (Fig. 1). However, Aurora-A–defective tissues show a clear induction of p53 (Fig. 4). The upregulation of p53 might be due to DNA damage induced upon Aurora-A depletion and/or could be a consequence of an increase in its stability, as it has been previously reported to occur when Aurora-A is silenced (4). However, depletion of p53 does not rescue the lack of proliferation of Aurora-A–null mouse fibroblasts (Fig. 1). Our data suggest that proliferative defects caused by Aurora-A deficiency are mostly a consequence of defective DNA synthesis in polyploid or aneuploid cells. Thus, although Aurora A-ablation allows DNA replication in cells with low ploidy, these cells finally arrest in a G0-like state with high ploidy. According to this hypothesis, Tsunematsu and colleagues have recently demonstrated that Aurora-A controls DNA replication by stabilizing Geminin in mitosis, which in turn inhibits SCFSkp2-mediated degradation of Cdt1 and ensures prereplicative complex formation in the subsequent cell cycle (49). Therefore, the lack of proliferation induced by Aurora-A depletion could be a consequence of a defective DNA replication rather than a p53-induced cell-cycle arrest. In addition, we have observed an intriguing induction of differentiation upon mitotic failure (Supplementary Fig. S6), an observation that may explain tumor arrest in the absence of apoptosis or senescence, at least in some specific tissues. Further understanding of these pathways will have important implications in future antitumoral therapies based on the use of inhibitors of Aurora-A or other mitotic targets (25). Given the lack of proper biomarkers for testing the effect of Aurora-A inhibition in clinical trials, a quantification of ploidy or nuclear size in treated tumors could be of help in future assays.

No potential conflicts of interest were disclosed.

Conception and design: I. Pérez de Castro, M. Malumbres

Development of methodology: M. Malumbres

Acquisition of data (provided animals, acquired and managed patients, provided facilities, etc.): C. Aguirre-Portoles, G. Fernández-Miranda, M. Canamero, D.O. Cowley, T. Van Dyke

Analysis and interpretation of data (e.g., statistical analysis, biostatistics, computational analysis): I. Pérez de Castro, M. Malumbres

Writing, review, and/or revision of the manuscript: I. Pérez de Castro, M. Malumbres

Administrative, technical, or material support (i.e., reporting or organizing data, constructing databases): C. Aguirre-Portoles, M. Malumbres

Study supervision: M. Malumbres

The authors are indebted to members of the Animal facility, as well as Histopathology, Molecular Imaging and Confocal Microscopy facilities of the CNIO for their excellent technical support. The authors thank Susana Temiño and Sara Rodrigo for their technical assistance, and Jorge Oller and Laura Bel for their help with cellular assays.

This work was financially supported by grants from the Ministerio de Economía y Competitividad (MINECO; SAF2010-19710 to I. Pérez de Castro and SAF2012-38215 to M. Malumbres), Fundación Ramón Areces, the OncoCycle Programme (S2010/BMD-2470) from the Comunidad de Madrid, the OncoBIO Consolider-Ingenio 2010 Programme (CSD2007-00017) from the MINECO, and the European Union Seventh Framework Programme (MitoSys project; HEALTH-F5-2010-241548).

The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked advertisement in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

1.
Carmena
M
,
Earnshaw
WC
. 
The cellular geography of aurora kinases
.
Nat Rev Mol Cell Biol
2003
;
4
:
842
54
.
2.
Dutertre
S
,
Descamps
S
,
Prigent
C
. 
On the role of aurora-A in centrosome function
.
Oncogene
2002
;
21
:
6175
83
.
3.
Bayliss
R
,
Sardon
T
,
Vernos
I
,
Conti
E
. 
Structural basis of Aurora-A activation by TPX2 at the mitotic spindle
.
Mol Cell
2003
;
12
:
851
62
.
4.
Katayama
H
,
Sasai
K
,
Kawai
H
,
Yuan
ZM
,
Bondaruk
J
,
Suzuki
F
, et al
Phosphorylation by aurora kinase A induces Mdm2-mediated destabilization and inhibition of p53
.
Nat Genet
2004
;
36
:
55
62
.
5.
Mendez
R
,
Hake
LE
,
Andresson
T
,
Littlepage
LE
,
Ruderman
JV
,
Richter
JD
. 
Phosphorylation of CPE binding factor by Eg2 regulates translation of c-mos mRNA
.
Nature
2000
;
404
:
302
7
.
6.
Huang
YS
,
Jung
MY
,
Sarkissian
M
,
Richter
JD
. 
N-methyl-D-aspartate receptor signaling results in Aurora kinase-catalyzed CPEB phosphorylation and alpha CaMKII mRNA polyadenylation at synapses
.
EMBO J
2002
;
21
:
2139
48
.
7.
Sasayama
T
,
Marumoto
T
,
Kunitoku
N
,
Zhang
D
,
Tamaki
N
,
Kohmura
E
, et al
Over-expression of Aurora-A targets cytoplasmic polyadenylation element binding protein and promotes mRNA polyadenylation of Cdk1 and cyclin B1
.
Genes Cells
2005
;
10
:
627
38
.
8.
Glover
DM
,
Leibowitz
MH
,
McLean
DA
,
Parry
H
. 
Mutations in aurora prevent centrosome separation leading to the formation of monopolar spindles
.
Cell
1995
;
81
:
95
105
.
9.
Hannak
E
,
Kirkham
M
,
Hyman
AA
,
Oegema
K
. 
Aurora-A kinase is required for centrosome maturation in Caenorhabditis elegans
.
J Cell Biol
2001
;
155
:
1109
16
.
10.
Schumacher
JM
,
Ashcroft
N
,
Donovan
PJ
,
Golden
A
. 
A highly conserved centrosomal kinase, AIR-1, is required for accurate cell cycle progression and segregation of developmental factors in Caenorhabditis elegans embryos
.
Development
1998
;
125
:
4391
402
.
11.
Giet
R
,
Prigent
C
. 
The Xenopus laevis aurora/Ip11p-related kinase pEg2 participates in the stability of the bipolar mitotic spindle
.
Exp Cell Res
2000
;
258
:
145
51
.
12.
Liu
Q
,
Ruderman
JV
. 
Aurora A, mitotic entry, and spindle bipolarity
.
Proc Natl Acad Sci U S A
2006
;
103
:
5811
6
.
13.
Roghi
C
,
Giet
R
,
Uzbekov
R
,
Morin
N
,
Chartrain
I
,
Le Guellec
R
, et al
The Xenopus protein kinase pEg2 associates with the centrosome in a cell cycle-dependent manner, binds to the spindle microtubules and is involved in bipolar mitotic spindle assembly
.
J Cell Sci
1998
;
111
(
Pt 5
):
557
72
.
14.
Hirota
T
,
Kunitoku
N
,
Sasayama
T
,
Marumoto
T
,
Zhang
D
,
Nitta
M
, et al
Aurora-A and an interacting activator, the LIM protein Ajuba, are required for mitotic commitment in human cells
.
Cell
2003
;
114
:
585
98
.
15.
Kunitoku
N
,
Sasayama
T
,
Marumoto
T
,
Zhang
D
,
Honda
S
,
Kobayashi
O
, et al
CENP-A phosphorylation by Aurora-A in prophase is required for enrichment of Aurora-B at inner centromeres and for kinetochore function
.
Dev Cell
2003
;
5
:
853
64
.
16.
Du
J
,
Hannon
GJ
. 
Suppression of p160ROCK bypasses cell cycle arrest after Aurora-A/STK15 depletion
.
Proc Natl Acad Sci U S A
2004
;
101
:
8975
80
.
17.
Girdler
F
,
Gascoigne
KE
,
Eyers
PA
,
Hartmuth
S
,
Crafter
C
,
Foote
KM
, et al
Validating Aurora B as an anti-cancer drug target
.
J Cell Sci
2006
;
119
:
3664
75
.
18.
Yang
H
,
Burke
T
,
Dempsey
J
,
Diaz
B
,
Collins
E
,
Toth
J
, et al
Mitotic requirement for aurora A kinase is bypassed in the absence of aurora B kinase
.
FEBS Lett
2005
;
579
:
3385
91
.
19.
Cowley
DO
,
Rivera-Pérez
JA
,
Schliekelman
M
,
He
YJ
,
Oliver
TG
,
Lu
L
, et al
Aurora-A kinase is essential for bipolar spindle formation and early development
.
Mol Cell Biol
2009
;
29
:
1059
71
.
20.
Lu
LY
,
Wood
JL
,
Ye
L
,
Minter-Dykhouse
K
,
Saunders
TL
,
Yu
X
, et al
Aurora A is essential for early embryonic development and tumor suppression
.
J Biol Chem
2008
;
283
:
31785
90
.
21.
Sasai
K
,
Parant
JM
,
Brandt
ME
,
Carter
J
,
Adams
HP
,
Stass
SA
, et al
Targeted disruption of Aurora A causes abnormal mitotic spindle assembly, chromosome misalignment and embryonic lethality
.
Oncogene
2008
;
27
:
4122
7
.
22.
Gautschi
O
,
Heighway
J
,
Mack
PC
,
Purnell
PR
,
Lara
PN
,
Gandara
DR
. 
Aurora kinases as anticancer drug targets
.
Clin Cancer Res
2008
;
14
:
1639
48
.
23.
Carter
SL
,
Eklund
AC
,
Kohane
IS
,
Harris
LN
,
Szallasi
Z
. 
A signature of chromosomal instability inferred from gene expression profiles predicts clinical outcome in multiple human cancers
.
Nat Genet
2006
;
38
:
1043
8
.
24.
Dar
AA
,
Goff
LW
,
Majid
S
,
Berlin
J
,
El-Rifai
W
. 
Aurora kinase inhibitors–rising stars in cancer therapeutics?
Mol Cancer Ther
2010
;
9
:
268
78
.
25.
Doménech
E
,
Malumbres
M
. 
Mitosis-targeting therapies: a troubleshooting guide
.
Curr Opin Pharmacol
2013
;
13
:
519
28
.
26.
Guerra
C
,
Mijimolle
N
,
Dhawahir
A
,
Dubus
P
,
Barradas
M
,
Serrano
M
, et al
Tumor induction by an endogenous K-ras oncogene is highly dependent on cellular context
.
Cancer Cell
2003
;
4
:
111
20
.
27.
Indra
AK
,
Li
M
,
Brocard
J
,
Warot
X
,
Bornert
JM
,
Gérard
C
, et al
Targeted somatic mutagenesis in mouse epidermis
.
Horm Res
2000
;
54
:
296
300
.
28.
Guy
CT
,
Cardiff
RD
,
Muller
WJ
. 
Induction of mammary tumors by expression of polyomavirus middle T oncogene: a transgenic mouse model for metastatic disease
.
Mol Cell Biol
1992
;
12
:
954
61
.
29.
Ruzankina
Y
,
Pinzon-Guzman
C
,
Asare
A
,
Ong
T
,
Pontano
L
,
Cotsarelis
G
, et al
Deletion of the developmentally essential gene ATR in adult mice leads to age-related phenotypes and stem cell loss
.
Cell Stem Cell
2007
;
1
:
113
26
.
30.
Braun
KM
,
Niemann
C
,
Jensen
UB
,
Sundberg
JP
,
Silva-Vargas
V
,
Watt
FM
. 
Manipulation of stem cell proliferation and lineage commitment: visualisation of label-retaining cells in wholemounts of mouse epidermis
.
Development
2003
;
130
:
5241
55
.
31.
Sotillo
R
,
García
JF
,
Ortega
S
,
Martin
J
,
Dubus
P
,
Barbacid
M
, et al
Invasive melanoma in Cdk4-targeted mice
.
Proc Natl Acad Sci U S A
2001
;
98
:
13312
7
.
32.
García-Higuera
I
,
Manchado
E
,
Dubus
P
,
Cañamero
M
,
Méndez
J
,
Moreno
S
, et al
Genomic stability and tumour suppression by the APC/C cofactor Cdh1
.
Nat Cell Biol
2008
;
10
:
802
11
.
33.
Zanet
J
,
Freije
A
,
Ruiz
M
,
Coulon
V
,
Sanz
JR
,
Chiesa
J
, et al
A mitosis block links active cell cycle with human epidermal differentiation and results in endoreplication
.
PLoS ONE
2010
;
5
:
e15701
.
34.
Yoon
Y
,
Cowley
DO
,
Gallant
J
,
Jones
SN
,
Van Dyke
T
,
Rivera-Pérez
JA
. 
Conditional Aurora A deficiency differentially affects early mouse embryo patterning
.
Dev Biol
2012
;
371
:
77
85
.
35.
Torchia
EC
,
Zhang
L
,
Huebner
AJ
,
Sen
S
,
Roop
DR
. 
Aurora kinase-A deficiency during skin development impairs cell division and stratification
.
J Invest Dermatol
2013
;
133
:
78
86
.
36.
Sasai
K
,
Katayama
H
,
Stenoien
DL
,
Fujii
S
,
Honda
R
,
Kimura
M
, et al
Aurora-C kinase is a novel chromosomal passenger protein that can complement Aurora-B kinase function in mitotic cells
.
Cell Motil Cytoskeleton
2004
;
59
:
249
63
.
37.
Bayliss
R
,
Sardon
T
,
Ebert
J
,
Lindner
D
,
Vernos
I
,
Conti
E
. 
Determinants for Aurora-A activation and Aurora-B discrimination by TPX2
.
Cell Cycle
2004
;
3
:
404
7
.
38.
Fu
J
,
Bian
M
,
Liu
J
,
Jiang
Q
,
Zhang
C
. 
A single amino acid change converts Aurora-A into Aurora-B-like kinase in terms of partner specificity and cellular function
.
Proc Natl Acad Sci U S A
2009
;
106
:
6939
44
.
39.
Hans
F
,
Skoufias
DA
,
Dimitrov
S
,
Margolis
RL
. 
Molecular distinctions between Aurora A and B: a single residue change transforms Aurora A into correctly localized and functional Aurora B
.
Mol Biol Cell
2009
;
20
:
3491
502
.
40.
Bischoff
JR
,
Anderson
L
,
Zhu
Y
,
Mossie
K
,
Ng
L
,
Souza
B
, et al
A homologue of Drosophila aurora kinase is oncogenic and amplified in human colorectal cancers
.
EMBO J
1998
;
17
:
3052
65
.
41.
Wang
X
,
Zhou
YX
,
Qiao
W
,
Tominaga
Y
,
Ouchi
M
,
Ouchi
T
, et al
Overexpression of aurora kinase A in mouse mammary epithelium induces genetic instability preceding mammary tumor formation
.
Oncogene
2006
;
25
:
7148
58
.
42.
Manchado
E
,
Guillamot
M
,
Malumbres
M
. 
Killing cells by targeting mitosis
.
Cell Death Differ
2012
;
19
:
369
77
.
43.
Storchová
Z
,
Breneman
A
,
Cande
J
,
Dunn
J
,
Burbank
K
,
O'Toole
E
, et al
Genome-wide genetic analysis of polyploidy in yeast
.
Nature
2006
;
443
:
541
7
.
44.
Quignon
F
,
Rozier
L
,
Lachages
AM
,
Bieth
A
,
Simili
M
,
Debatisse
M
. 
Sustained mitotic block elicits DNA breaks: one-step alteration of ploidy and chromosome integrity in mammalian cells
.
Oncogene
2007
;
26
:
165
72
.
45.
Paulsen
RD
,
Soni
DV
,
Wollman
R
,
Hahn
AT
,
Yee
MC
,
Guan
A
, et al
A genome-wide siRNA screen reveals diverse cellular processes and pathways that mediate genome stability
.
Mol Cell
2009
;
35
:
228
39
.
46.
Huck
JJ
,
Zhang
M
,
McDonald
A
,
Bowman
D
,
Hoar
KM
,
Stringer
B
, et al
MLN8054, an inhibitor of Aurora A kinase, induces senescence in human tumor cells both in vitro and in vivo
.
Mol Cancer Res
2010
;
8
:
373
84
.
47.
Görgün
G
,
Calabrese
E
,
Hideshima
T
,
Ecsedy
J
,
Perrone
G
,
Mani
M
, et al
A novel Aurora-A kinase inhibitor MLN8237 induces cytotoxicity and cell-cycle arrest in multiple myeloma
.
Blood
2010
;
115
:
5202
13
.
48.
Liu
Y
,
Hawkins
OE
,
Su
Y
,
Vilgelm
AE
,
Sobolik
T
,
Thu
YM
, et al
Targeting aurora kinases limits tumour growth through DNA damage-mediated senescence and blockade of NF-κB impairs this drug-induced senescence
.
EMBO Mol Med
2013
;
5
:
149
66
.
49.
Tsunematsu
T
,
Takihara
Y
,
Ishimaru
N
,
Pagano
M
,
Takata
T
,
Kudo
Y
. 
Aurora-A controls pre-replicative complex assembly and DNA replication by stabilizing geminin in mitosis
.
Nat Commun
2013
;
4
:
1885
.

Supplementary data