Compositions and Methods for the Treatment of Diseases Associated with Aberrant Cilia Assembly and Regulation
Compositions and methods are provided for identifying agents which have efficacy for the treatment of disorders related to aberrant cilial structure and function, including polycystic kidney disease.
This application is a Continuation-in-Part Application of U.S. patent application Ser. No. 12/374,209 filed Apr. 2, 2009, which is a §371 Application of PCT/US07/73722, filed Jul. 17, 2007, which claims priority to U.S. Provisional Applications 60/831,479 and 60/925,272 filed Jul. 17, 2006 and Apr. 19, 2007. This CIP application also claims priority to U.S. Provisional Application, 61/122,303 filed Dec. 12, 2008. The entire disclosures of each of the foregoing application are incorporated herein by reference.
Pursuant to 35 U.S.C. §202(c), it is acknowledged that the U.S. Government has certain rights in the invention described, which was made in part with funds from the National Institutes of Health, Grant Numbers RO1 CA63366, CA-06927, DOD W81XWH-07-1-0676 and RO1 CA-113342.
FIELD OF THE INVENTIONThe present invention relates to the fields of molecular biology and cilia-associated structural and cellular signal transduction. More specifically, the invention provides methods for identifying compounds which modulate cilia assembly and disassembly, thereby providing treatment for disorders associated with aberrant coordination of cilia function, including, for example, polycystic kidney disease, renal cysts, cancer, hypertension and infertility.
BACKGROUND OF THE INVENTIONSeveral publications and patent documents are cited throughout the specification in order to describe the state of the art to which this invention pertains. Each of these citations is incorporated by reference herein as though set forth in full.
In polycystic kidney disease (PKD), Bardet-Biedl Syndrome (BBS), and other disorders, mutations in cilia-associated structural or signaling proteins cause insensitivity to external mechanical and diffusible signaling cues, resulting in disorganized, hyperplastic cell growth (Benzing and Walz, 2006; Pan et al., 2005; Singla and Reiter, 2006). On the organismal level, ciliary defects produce renal cysts, infertility, respiratory disorders, situs inversus, and predisposition to obesity, diabetes, and hypertension. Notably, recent studies have shown that the Hedgehog, Wnt, PDGFαα, and other signaling cascades are coordinated at cilia (Cano et al., 2004; Huangfu and Anderson, 2005; Liu et al., 2005; Schneider et al., 2005; Simons et al., 2005; Tanaka et al., 2005). The frequent deregulation of these pathways during cell transformation, together with the common disappearance of cilia in transformed cells, raises the possibility that defective ciliary signaling may promote cancer.
Although an increasing number of proteins are being defined as ciliary structural components or cilia-associated signaling proteins, very little is currently known about the cellular machinery controlling the formation and resorption of cilia. It has long been known that cilia are regulated dynamically throughout the cell cycle. In many cells, resorption occurs at mitotic entry, and reappearance after progression into G1. However, resorption is not solely linked to mitotic entry, with some cells undergoing waves of resorption at different points in cell cycle: for example, Tucker et al. have noted ciliary resorption as cells emerge from quiescence, prior to S-phase (Quarmby and Parker, 2005; Rieder et al., 1979; Tucker et al., 1979). Given their increasingly apparent role in detecting and transmitting extracellular signals, regulated formation, disassembly, or shortening of cilia may play an important role in cellular growth controls, serving as a rheostat to limit response to overly persistent or abnormal cell growth cues in the extracellular environment.
A cilium arises from a basal body, a structure that differentiates from one of the centrioles in the centrosome in non-proliferating cells and organizes the microtubule bundles that constitute the ciliary axoneme. Cilia are evolutionarily related to the motile flagella of lower eukaryotes, such as the green algae Chlamydomonas. Genetic studies in Chlamydomonas have recently begun to dissect the process of flagellar resorption (Bradley and Quarmby, 2005; Marshall et al., 2005; Pan and Snell, 2005; Quarmby, 2004). These studies have identified altered functionality of the intraflagellar transport (IFT) machinery and destabilization of the axoneme as hallmarks of disassembly, and implicated CALK and other kinases as regulators of disassembly. The means by which CALK becomes activated at initiation of disassembly and the critical CALK effectors in the disassembly process remain unknown, as does the relevance of these observations to higher eukaryotes.
CALK is very distantly related to the human Aurora A (AurA) kinase, with 55% similarity centered on the protein catalytic domain. In humans, Aurora A (AurA) is a centrosomal kinase that regulates mitotic entry through activation of Cdk1-cyclin B and other substrates that organize the mitotic spindle (Bischoff et al., 1998; Marumoto et al., 2005). AurA amplification or activation is common in many cancers characterized by centrosomal amplification and genomic instability (Anand et al., 2003; Goepfert et al., 2002; Gritsko et al., 2003). In the past year, altered expression of the HEF1 (Law et al., 1996; O'Neill et al., 2000) scaffolding protein has recently been identified as part of a pro-metastatic signature in breast cancer (Minn et al., 2005), shown to contribute to the aggressiveness of glioblastomas (Natarajan et al., 2006), and found to be critical for progression to metastasis in melanomas (Kim et al., 2006). HEF1 is best known as a transducer of integrin-initiated attachment, migration, and anti-apoptotic signals at focal adhesions (O'Neill et al., 2000).
SUMMARY OF THE INVENTIONIn accordance with the present invention, methods for identifying agents which modulate ciliary function and assembly/disassembly are provided. An exemplary method entails providing cells which express AurA and HEF1 and incubating the cells in the presence and absence of the agent. Following treatment, the cilia present on the cells are assessed for alterations which occur in the presence but not the absence of the agent, agents which cause alterations being identified as modulators of ciliary function and assembly.
In yet another aspect of the invention, an in vivo model for assessing agents which modulate kidney cyst formation is provided. In the model, a first strain of mice in which Pkd1 is conditionally inactivated is provided and the mice crossed with each of the following mice, i) a transgenic HEF-1 knock out mouse; ii) a mouse expressing functional HEF-1 and iii) a third strain of mice which are heterozygous for HEF-1 expression. After crossing, Pkd1 is inactivated and a test agent administered to each of the newly created strains of mice. After a suitable time period of administration, the mice are assessed to determine whether the agent modulates cyst formation relative to untreated mice.
In yet another aspect the invention provides a method for identifying agents which modulate calmodulin-Aurora A complex formation having efficacy for the treatment of polycystic kidney disease (PKD). An exemplary method entails providing kidney cells which express Aurora A and calmodulin and incubating cells in the presence and absence of said agent and in the presence of a molecule which induces release of Ca2+. The extent to which the agent disrupts calmodulin-Aurora A complex formation following release of calcium is then determined, agents which disrupt complex formation identified by the aforementioned method should have efficacy for the treatment of PKD. In a preferred embodiment, the agent inhibits activation of aurora A kinase and includes, without limitation, siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, MLN8054, VX-680 and hesparadin. In further embodiments of the invention, the effect of the agent on disruption of aurora A binding to PC2 and/or cellular localization of PC2 and/or Aurora A phosphorylation of PC2 may be determined.
In yet another embodiment of the invention, a method for inhibiting progression of polycystic kidney disease in a patient in need thereof is provided. In one aspect the method entails administration of an effective amount of an aurora kinase inhibitor the patient in an amount effective to inhibit cyst formation in the kidney. The inhibitor may inhibit aurora kinase A or aurora kinase B or it may inhibit both.
The mammalian cilium protrudes from the apical/lumenal surface of polarized cells, and acts as a sensor of environmental cues. Numerous developmental disorders and pathological conditions have been shown to arise from defects in cilia-associated signaling proteins. Despite mounting evidence that cilia are essential sites for coordination of cell signaling, little is known about the cellular mechanisms controlling their formation and disassembly. Here we show that defined interactions between the pro-metastatic scaffolding protein HEF1/Cas-L/NEDD9 and the oncogenic Aurora A (AurA) kinase at the basal body of cilia causes phosphorylation and activation of HDAC6, a tubulin deacetylase, promoting ciliary disassembly. We show that this pathway is both necessary and sufficient for ciliary resorption, and constitutes an unexpected non-mitotic activity of AurA in vertebrates. Moreover, we demonstrate that small molecule inhibitors of AurA and HDAC6 selectively stabilize cilia from regulated resorption cues, suggesting a novel mode of action for these clinical agents.
Oncogenic hyperactivation of the mitotic kinase Aurora-A (AurA) in cancer is associated with genomic instability. Increasing evidence indicates AurA also regulates critical processes in normal interphase cells, but the source of such activity has been obscure. We report that multiple stimuli causing release of Ca2+ from intracellular endoplasmic reticulum (ER) stores activate AurA by inducing direct Ca2+-dependent calmodulin binding to AurA. Subsequently, activated AurA binds, phosphorylates, and limits the activity of the Ca2+-permeable nonselective cation channel polycystin 2 (PC2/TRPP2, encoded by the polycystic kidney disease-associated gene PKD2), limiting the amplitude of Ca2+ release from the ER. Active AurA is abundant in non-mitotic cells in normal kidneys and elevated in cells lining PKD-associated renal cysts, and inhibitors of AurA significantly enhance PC2-dependent Ca2+-release. These and other findings provide a new context for evaluating AurA function in normal cells and cancer, and suggest AurA may be a relevant new modifier gene for PKD.
The following definitions are provided to facilitate an understanding of the present invention:
Disorders associated with aberrant cilia function and regulation include, without limitation, polycystic kidney disease (PKD), Bardet-Biedl Syndrome (BBS), renal cysts, infertility, respiratory disorders, situs inversus, and predisposition to obesity, diabetes, and hypertension.
The phrase “aurora kinase inhibitor” refers to any agent which functions to inhibit or down regulate aurora kinase A and/or aurora kinase B. Such agents include, without limitation, small molecules, chemical compounds and nucleic acid molecules which function to down regulate expression of target genes. Exemplary agents include C1368, PHA-680632, hesparadin, MLN 8327, VX-680, MLN8054, and siRNA which hybridize selectively to aurora kinase encoding mRNA and down regulate expression of the aurora kinase protein product. Exemplary siRNAs that target aurora kinase have the following sequence: Hs_AURKA—1 TCCCAGCGCATTCCTTTGCAA and Hs_STK6—5 CACCTTCGGCATCCTAATATT.
The terms “transform”, “transfect”, “transduce”, shall refer to any method or means by which a nucleic acid is introduced into a cell or host organism and may be used interchangeably to convey the same meaning. Such methods include, but are not limited to, transfection, electroporation, microinjection, PEG-fusion and the like. The introduced nucleic acid may or may not be integrated (covalently linked) into nucleic acid of the recipient cell or organism. In bacterial, yeast, plant and mammalian cells, for example, the introduced nucleic acid may be maintained as an episomal element or independent replicon such as a plasmid. Alternatively, the introduced nucleic acid may become integrated into the nucleic acid of the recipient cell or organism and be stably maintained in that cell or organism and further passed on or inherited to progeny cells or organisms of the recipient cell or organism. Finally, the introduced nucleic acid may exist in the recipient cell or host organism only transiently.
The term “selectable marker gene” refers to a gene that when expressed confers a selectable phenotype, such as antibiotic resistance, on a transformed cell.
The term “operably linked” means that the regulatory sequences necessary for expression of the coding sequence are placed in the DNA molecule in the appropriate positions relative to the coding sequence so as to effect expression of the coding sequence. This same definition is sometimes applied to the arrangement of transcription units and other transcription control elements (e.g. enhancers) in an expression vector.
“Native” refers to a naturally occurring (“wild-type”) nucleic acid sequence.
“Heterologous” sequence refers to a sequence which originates from a foreign source or species or, if from the same source, is modified from its original form.
A “coding sequence” or “coding region” refers to a nucleic acid molecule having sequence information necessary to produce a gene product, when the sequence is expressed.
The term “animal” is used herein to include all vertebrate animals, except humans. It also includes an individual animal in all stages of development, including embryonic and fetal stages. A “transgenic animal” is any animal containing one or more cells bearing genetic information altered or received, directly or indirectly, by deliberate genetic manipulation at the subcellular level, such as by targeted recombination or microinjection or infection with recombinant virus. The term “transgenic animal” is not meant to encompass classical cross-breeding or in vitro fertilization, but rather is meant to encompass animals in which one or more cells are altered by or receive a recombinant DNA molecule. This molecule may be specifically targeted to defined genetic locus, be randomly integrated within a chromosome, or it may be extrachromosomally replicating DNA. The term “germ cell line transgenic animal” refers to a transgenic animal in which the genetic alteration or genetic information was introduced into a germ line cell, thereby conferring the ability to transfer the genetic information to offspring. If such offspring in fact, possess some or all of that alteration or genetic information, then they, too, are transgenic animals.
The alteration or genetic information may be foreign to the species of animal to which the recipient belongs, or foreign only to the particular individual recipient, or may be genetic information already possessed by the recipient. In the last case, the altered or introduced gene may be expressed differently than the native gene.
The DNA used for altering a target gene may be obtained by a wide variety of techniques that include, but are not limited to, isolation from genomic sources, preparation of cDNAs from isolated mRNA templates, direct synthesis, or a combination thereof.
Methods for Treating PKDAs explained above, the present invention is directed towards methods for modulating or alleviating the symptoms of polycystic kidney disease comprising, the administration of an Aur-A inhibitor such as those listed above. Administration can be achieved by any suitable route, such as parenterally, transmucosally, e.g., orally, nasally, or rectally, or transdermally. Preferably, administration is parenteral, e.g., via intravenous injection. Alternative means of administration also include, but are not limited to, intra-arteriole, intramuscular, intradermal, subcutaneous, intraperitoneal, intraventricular, and intracranial administration, or by injection into the renal cyst being treated.
The Aur-A inhibitor may be employed in any suitable pharmaceutical formulation, as described above, including in a vesicle, such as a liposome [see Langer, Science 249:1527-1533 (1990); Treat et al., in Liposomes in the Therapy of Infectious Disease and Cancer, Lopez-Berestein and Fidler (eds.), Liss: New York, pp. 317-327, see generally, ibid] Preferably, administration of liposomes containing the agents of the invention is parenteral, e.g., via intravenous injection, but also may include, without limitation, intra-arteriole, intramuscular, intradermal, subcutaneous, intraperitoneal, intraventricular, and intracranial administration, or by injection into the renal cyst being treated.
In yet another embodiment, a pharmaceutical composition of the present invention can be delivered in a controlled release system, such as using an intravenous infusion, an implantable osmotic pump, a transdermal patch, liposomes, or other modes of administration. In a particular embodiment, a pump may be used [see Langer, supra; Sefton, CRC Crit. Ref. Biomed. Eng. 14:201 (1987); Buchwald et al., Surgery 88:507 (1980); Saudek et al., N. Engl. J. Med. 321:574 (1989)]. In another embodiment, polymeric materials can be used [see Medical Applications of Controlled Release, Langer and Wise (eds.), CRC Press: Boca Raton, Fla. (1974); Controlled Drug Bioavailability, Drug Product Design and Performance, Smolen and Ball (eds.), Wiley: New York (1984); Ranger and Peppas, J. Macromol. Sci. Rev. Macromol. Chem. 23:61 (1983); see also Levy et al., Science 228:190 (1985); During et al., Ann. Neurol. 25:351 (1989); Howard et al., J. Neurosurg. 71:105 (1989)]. In yet another embodiment, a controlled release system can be placed in proximity of the target tissues of the animal, thus requiring only a fraction of the systemic dose [see, e.g., Goodson, in Medical Applications of Controlled Release, supra, vol. 2, pp. 115-138 (1984)]. In particular, a controlled release device can be introduced into an animal in proximity of the site of inappropriate immune activation or a tumor. Other controlled release systems are discussed in the review by Langer [Science 249:1527-1533 (1990)].
The examples set forth below are provided to exemplify certain embodiments of the invention. They are not intended to limit the invention in any way.
Example IWe demonstrate that an association between AurA and HEF1 at cilia in response to extracellular cues is required for ciliary disassembly. We also show that AurA activation is independently sufficient to induce rapid ciliary resorption, and that AurA acts in this process through phosphorylating HDAC6, thus stimulating HDAC6-dependent tubulin deacetylation (Hubbert et al., 2002) and destabilizing the ciliary axoneme. Importantly, our identification of a spatiotemporally restricted action of AurA at the ciliary basal body in cells emerging from G0 demonstrates an unexpected non-mitotic activity for AurA in vertebrate cells. We also determine that small molecule inhibitors of AurA and HDAC6 reduce regulated disassembly of cilia, which may have important implications for the action of these drugs in the clinic. Together, these data reveal important activities for HEF1, AurA, and HDAC6 in regulation of ciliary resorption, which should also inform the actions of these proteins in cell cycle and cancer (Hideshima et al., 2005; Kim et al., 2006; Marumoto et al., 2005; Pugacheva and Golemis, 2005).
The following materials and methods are provided to facilitate the practice of the present example.
Cell culture and siRNA. hTERT-RPE1 cells were grown in DMEM with 10% fetal bovine serum (FBS). For analysis of ciliary disassembly, cells were plated at 30% confluence in plates containing glass cover slips, and starved for 48 hours (in Opti-MEM or regular DMEM, without added serum) to induce cilia formation, followed by treatments described hereinbelow. For siRNA treatment, cells were initially plated in DMEM/10% FBS in plates containing cover slips, and 12 hours later siRNA transfection was performed in Opti-MEM with Oligofectamine (Invitrogen) according to manufacturer recommendations, and fixed 48 hours after transfection, following treatments indicated in Results. The remaining cells on plate were lysed, then either directly analyzed by Western blot analysis, or used for immunoprecipitation (IP)-kinase reaction to measure AurA activity.
For RNA interference (RNAi)-induced depletion of HEF1 and AurA, 2 independent, synthetic duplex siRNAs were used for each gene: 1) Ambion, cat#16704, NEDD9, ID:17729. sense 5′GGUAUAUCAGGUGCCACCAtt3′; 2) Dharmacon, custom, sense: 5′ AAGGGGUAUAUGCCAUUCCGCdTd T 3′57. Non-specific control siRNAs including scrambled (Dharmacon, cat#D-001206-13-05) and GFP-directed sequences (Dharmacon, cat#D-001300-01-20) were used for reference.
Drug inhibition experiments. The Aurora kinase inhibitor PHA-680632, GSK3β-inhibitor 1 (Calbiochem), FTI-277 (Calbiochem), Tubacin, Niltubacin or DMSO vehicle were added to hTERT-RPE1 cells 2 hours prior to the initiation of ciliary disassembly. After initial titration experiments to establish effective range, PHA-680632 was used at 0.5 μM, Tubacin and Niltubacin at 2 μM, GSK3β-inhibitor 1 at 2 μM, FTI-277 at 50 nM concentration for the experiments described.
Protein expression, Western blotting, and immunoprecipitation. For microinjection, recombinant glutathione-S-transferase (GST), GST fused AurA mutants T288A and D274N) produced from BL21 (DE3) bacteria were purified using the MicroSpin GST Purification Module (Amersham Biotech.). Purified recombinant AurA was purchased from Upstate; this AurA was pre-activated based on incubation with ATP. Mutationally inactive AurA (T288A,) was also made using a baculoviral expression system (Invitrogen), and was purified by Ni-Sepharose 6FF (Amersham).
To prepare lysates for Western blotting and IP, mammalian cells were disrupted by M-PER lysis buffer (Pierce) supplemented with EDTA-free protease inhibitor cocktail (Roche). Lysates used for IP were incubated overnight with antibody at 4° C., subsequently incubated for 2 hours with protein A/G-sepharose (Pierce), washed, and resolved by SDS-PAGE. Western blotting was performed using standard procedures and proteins visualized using the West-Pico system (Pierce). Antibodies used included mouse monoclonal antibody (mAb) anti-HEF1 2G9 (Pugacheva and Golemis, 2005), anti-α-tubulin mAb (Sigma), anti-AurA (BD Bioscience) for Western blotting, anti-AurA rabbit polyclonal (Cell Signaling) for IP, anti-Phospho-AurA/T288 (BioLegend), anti-Phospho-AurA/T288 (Cell Signaling), anti-HDAC6 rabbit polyclonal (Upstate; 1:5000), anti-HDAC2 rabbit polyclonal (Invitrogen) and mAb anti-β-actin (AC15, Sigma), anti-IFT88 and anti-IFT20. Secondary horseradish peroxidase (HRP)-conjugated antibodies were from Amersham Biotech.
Immunofluorescence. Cells were fixed with 4% paraformaldehyde (10 min) then methanol (5 min), permeabilized with 1% Triton-X100 in PBS, blocked in 1×PBS, 3% BSA, and incubated with antibodies using standard protocols. Primary antibodies included rabbit polyclonal anti-Aurora A and anti-phospho-AuroraA/T288, (Cell Signaling), mouse mAb anti-HEF1 (14A11), polyclonal anti-γ-tubulin (Sigma), anti-α-tubulin mAb (Sigma), anti-acetylated α-tubulin mAb (clone 6-11B-1, Sigma, and clone K(Ac)40 Biomol), anti-IFT88 and anti-IFT20 (gifts of G. Pazour), mouse anti-glutamylated tubulin (Sigma), and anti-HDAC6 (Upstate). Secondary antibodies labeled with Alexa-488, Alexa-568, and Alexa-633, and TOTO-3 dye to stain DNA, were from Molecular Probes/Invitrogen. DNA was co-stained in some experiments by propidium iodine (Sigma) or Draq5 (Alexis). Confocal microscopy was performed using a Radiance 2000 laser scanning confocal microscope ((Carl Zeiss, Thornwood, N.Y.) coupled to a Nikon Eclipse E800 upright microscope (Nikon). Statistical analysis of data by one-way ANOVA was performed using GraphPad Instat 3.0 (San Diego, Calif.).
Microinjection. Microinjections were performed on a Nikon TE300 Microscope (Nikon, Melville, N.Y.) that was equipped with an Eppendorf Transjector 5246 semi-automatic microinjector and micromanipulator (Eppendorf, Westbury, N.Y.). Cells were plated on gridded coverslips (Belco) and starved for 48 hours before cytoplasmic microinjection of 0.05 μM pre-activated AurA (Upstate), inactive AurA (T288A) and (D274N), GST protein, or buffer. Proteins were pre-filtered through a 0.2-μm Milliopore membrane and mixed with Dextran Green488 (Molecular Probes) to mark injected cells. Injected cells were incubated at 37° C. before fixation. Typically, 150 cells were microinjected in each of 3 experiments.
Kinase and tubulin deacetylation assays. In vitro kinase assays were performed using recombinant active AurA (Upstate), mutationally inactive AurA purified from baculovirus and BL21 (DE3) bacteria, or endogenous AurA immunoprecipitated from mammalian cells. A standard kinase reaction with γ-32P(ATP) and histone H3 and MBP (Upstate) substrates was done as in (Pugacheva and Golemis, 2005). For deacetylase assays, HDAC6 and HDAC2 were in vitro translated using a TnT-Coupled Reticulocyte Lysate System (Promega), immunoprecipitated, and incubated with/without active AurA(Upstate) in the presence of (25 μg) stabilized microtubules prepared from purified bovine brain tubulin (Cytoskeleton) to measure deacetylase activity (as in (Hubbert et al., 2002)) and with γ-32P-ATP (Perkin-Elmer) in AurA reaction buffer. 1/10 volume of samples were reserved for Western blotting.
We established a system to study ciliary dynamics in the hTERT-RPE1 cell line. 48 hours after plating cells at 50-70% confluence in Opti-MEM medium without serum, >80% of hTERT-RPE1 cells had clearly visible cilia (
Dynamic regulation of HEF1 and AurA at the basal body during ciliary disassembly. AurA (
If AurA were functionally important for ciliary disassembly, we would expect changes in the activity of AurA 1-2 hours after serum treatment, potentially accompanied by changes in the AurA activator HEF1. Indeed, HEF1 expression increased at 1-2 hours after serum stimulation, dropped, and peaked again at ˜18-24 hours after serum stimulation (
HEF1-dependent activation of AurA induces ciliary disassembly. We used two complementary approaches to establish that AurA activation is necessary and sufficient for induction of ciliary disassembly, and that HEF1 is likely to contribute to this process.
First, exponentially growing hTERT-RPE1 cells were treated with siRNA targeting AurA or HEF1, or with control siRNA, plated for 2 days in OptiMEM to allow cilia formation, then treated with serum to induce ciliary disassembly. Immunoblotting confirmed siRNA treatment efficiently depleted AurA and HEF1 (
Second, we used the small molecule AurA kinase inhibitor PHA-680632 (Nerviano Medical Sciences (Soncini et al., 2006)) to inactivate AurA kinase (FIGS. 3D,E). Disassembly of cilia was strongly reduced in cells pre-treated for 3 hours with 500 nM PHA-680632 (
Together, these data imply that activation of AurA by HEF1 contributes to resorption of cilia at 2 and 18 hours following serum stimulation (
AurA activation is sufficient to induce rapid disassembly of cilia. Next, as a direct approach to establish sufficiency of active AurA to induce disassembly, we microinjected pre-activated wild type AurA (aAurA), T288A AurA (a hypomorphic mutant, (Satinover et al., 2004)), D274N AurA (an inactive mutant), GST, or buffer alone, together with fluorescent marker dye, into hTERT-RPE1 cells with pre-formed cilia. Microinjection of aAurA rapidly induced the disappearance of cilia from cells maintained in low serum medium: Essentially as soon as cells could be fixed after microinjection, more than 80% of injected cells lacked cilia (
HDAC6 is required for ciliary disassembly. Little is known about the cellular machinery necessary for disassembling cilia. In seeking targets of AurA phosphorylation that might be relevant to this process, we considered the possibility that the acetylated α-tubulin commonly used to visualize cilia might play an active role in stabilizing the ciliary axoneme, based on reports that α-tubulin deacetylation promoted the in vivo destabilization of microtubules (Matsuyama et al., 2002). In particular, histone deacetylase 6 (HDAC6) has been identified as an important cytoplasmic tubulin deacetylase that influences mitosis and chemotaxis through regulating tubulin stability (Hubbert et al., 2002).
To assess whether altered regulation of tubulin acetylation might mediate HEF1/AurA signaling, we treated ciliated hTERT-RPE1 cells with small molecule deacetylase inhibitors, and established the ciliary disassembly profile (
To further confirm a specific requirement for HDAC6, we next established that cilia do not disassemble in serum-treated cells with siRNA-depleted HDAC6 (
AurA phosphorylates HDAC6 to activate tubulin deacetylase activity. Taken together, our data suggested that the mechanism of ciliary disassembly by AurA requires intact HDAC6 deacetylation activity, to destabilize microtubules. AurA-dependent regulation of tubulin deacetylation may be direct or indirect. Importantly, although microinjection of AurA induced loss of ciliary α-acetylated tubulin as cilia disassemble, the non-ciliary α-acetylation of cytoplasmic microtubule networks were unaffected, suggesting a specific action of AurA and HDAC6 at the cilia (
To directly determine whether HDAC6 might be an AurA substrate, recombinant activated AurA was used in an in vitro kinase assay with purified HDAC6, HDAC2, or GST, as in (Pugacheva and Golemis, 2005). AurA phosphorylated HDAC6, but not HDAC2 or the GST negative control (
Ciliary disassembly and intraflagellar transport (IFT). Intraflagellar transport proteins perform important roles in mediating transport of proteins to and from the apical tip of cilia, and in many cases mutations in IFT proteins have been linked to ciliary dysfunction, loss of cilia, and pathological conditions (Sloboda, 2005). In contrast to depletion of HEF1 or AurA, depletion of representative IFT proteins IFT88 (
Cilia and flagella have been described as cellular “antennas”, sensing a multiplicity of extracellular stimuli to induce an intracellular response (Singla and Reiter, 2006). In addition to undergoing regulated resorption induced by extracellular cues, for over four decades cilia have been known to be dynamically resorbed and resynthesized throughout the cell cycle. Taken in sum, our data suggest a model (
An important finding of this work is the novel connection between AurA and HDAC6. HDAC6 tightly interacts with α and β tubulins through its HDAC domain, which may restrict its enzymatic activity, based on reports that taxol treatment causes HDAC6 to accumulate on microtubules, and is accompanied by increased tubulin acetylation (Zhang et al., 2003). Localized phosphorylation by AurA may increase the turnover of HDAC6 at microtubules, thus increasing the active pool of HDAC6 at cilia. Interestingly, studies in Chlamydomonas indicate that an important element of flagellar resorption is destabilization of the microtubule-based axoneme, suggesting this signaling cascade may be evolutionarily conserved (Pan and Snell, 2005; Pan et al., 2004). Further supporting the idea of conservation, the C. elegans gene MEC-12 encodes an α-tubulin variant that is specifically required only in mechanosensing neurons, which depend on intact cilia: MEC-12 is the only α-tubulin in this species with a conserved site for acetylation (Fukushige et al., 1999). Interestingly, HDAC6 has been reported to associate with protein phosphatase 1 (PP1) (Brush et al., 2004), which binds microtubules (Liao et al., 1998), and dephosphorylates and inactivates AurA kinase. Such feedback may limit AurA activation at cilia.
A number of growth stimuli induce HEF1 expression and phosphorylation, influencing its protein interactions. These include PDGF, which is here shown to partially induce ciliary disassembly (Natarajan et al., 2006). Intriguingly, recent studies of p130Cas, a protein structurally similar to HEF1, indicate that p130Cas acts as a stretch sensor; HEF1 contains all sequence motifs necessary for similar function (Kostic and Sheetz, 2006). As one major function of cilium is to sense fluid flow, and overly persistent flow has been reported to induce ciliary disassembly (Iomini et al., 2004), stretch sensation may be an important action of HEF1. Our data suggest that HEF1 both activates AurA and stabilizes the protein from degradation; it will be interesting to determine if the HEF1 scaffolding activity also contributes to AurA interaction with its effector HDAC6. Our data also indicate that AurA activity influences IFT88 localization during disassembly, and suggest integrity of the IFT system is important for the disassembly process in animals, as in Chlamydomonas (Pan and Snell, 2005).
Our establishment of a HEF1-AurA-HDAC6 cascade at cilia also informs the understanding of the mitotic activities of these proteins. Dynamic changes in microtubule acetylation and deacetylation characterize the stages of mitosis, and HDAC inhibitors that inhibit family members with microtubule deacetylase activity induce mitotic arrest (Blagosklonny et al., 2002). The identification here of HDAC6 as an AurA target suggests that HEF1-AurA regulation of tubulin deacetylation at mitosis through HDAC6 might offer a mechanism to fine-tune the mechanical properties of the mitotic spindle. This signaling cascade may also influence re-establishment of focal adhesions at and following cytokinesis, given the growing appreciation of the role of microtubules in guiding the formation of these structures (Ezratty et al., 2005; Strickland et al., 2005). Further, one intriguing possibility is that the common use of an AurA-HEF1-HDAC6 switch at the basal body of quiescent cells and the centrosome of G2/M cells may serve as part of a checkpoint mechanism coordinating responsiveness to extracellular cues at different points in cell cycle. In this context, our observation that inhibition of AurA causes appearance of mitotically arrested cells possessing both spindles and cilia (results not shown) may reflect triggering of such a centrosomally based checkpoint.
These results also have implications for the understanding and treatment of cancer. Tumor cells commonly do not have cilia, and both HEF1 and AurA are often upregulated in cancer. The roles for these proteins at the centrosome and focal adhesions described earlier already offer two mechanisms by which these proteins may promote tumor initiation and progression. The current study indicates a third mechanism, in which elevation of HEF1 or AurA in tumors may destabilize cilia, thus conditioning cellular response to external cues and impacting multiple signaling pathways. Further, AurA is regarded as a promising chemotherapeutic target, with agents inhibiting this protein currently in clinical trials (Andrews, 2005). TSA and other broad-spectrum agents targeting HDACs are used in the clinic (Vanhaecke et al., 2004), with more focused agents such as tubacin in preclinical development (Hideshima et al., 2005). Our data suggest that AurA- or HDAC-targeted drugs may have previously unappreciated in vivo effects involving cilia, that may contribute to the observed efficacy and/or side effects of these agents.
PKD is one of the best-described cilia-related diseases (Wilson, 2001), with mutation of the cilia-localized polycystin proteins 1 and 2 (PKD1 and PKD2) responsible for the significant majority of PKD patients. p130Cas interacts directly with complexes containing PKD1 and PKD2, and also with nephrocystins, cilia-associated proteins that are mutated in a second renal cystic syndrome, nephronophthisis (Benzing et al., 2001). Although an association of HEF1 with these proteins has never been assessed, HEF1 is abundant in the kidney and conserves many protein interaction sequences with p130Cas. It is also tantalizing to consider that closer connections exist between dysplastic disorders leading to cysts and cancer than have previously been appreciated. One of the surprising results of a recent large study to analyze the cancer genome was the identification of the PKHD1 protein, a ciliary protein which is mutant in autosomal recessive PKD, as commonly mutated in colorectal cancer (Sjoblom et al., 2006). Overall, deregulated AurA/HEF1/HDAC6 signaling appears to have broad implications for studies of human development and disease.
Example 2 Generation of a Mouse Model to Study PKDAt present, there are numerous competing models to explain the basis for cyst formation, and the differences between the various syndromes associated with kidney cysts. Studies of the signaling changes that occur in PKD have identified anomalous function of pathways that affect proliferation, cell cycle, and apoptosis (Edelstein, C. L. (2005) Cell Cycle 4:1550-4). Downstream elements of these signaling pathways include the tumor suppressors PTEN, TSC2, and p53, the oncogenes Bcl-2 and Akt, and other important growth regulators such as mTOR (Shillingford, J. M. et al. (2006) PNAS 103: 5466-71). Current therapeutic strategies are attempting to exploit this information by using drugs that target the relevant processes and pathways, such as the use of caspase inhibitors to reduce apoptosis, and mTOR inhibitors to block cell proliferation (Tao Y., et al. (2005) PNAS 102: 6954-9). In some cases, these approaches are alleviating symptoms and slowing cyst growth. However, no highly effective disease management strategy currently exists.
In the past several years, new insights into cyst pathogenesis have come from the consideration of the possible role of defects in renal cilia (Benzing et al., 2006; Snell, W. J., et al. Cell (2004) 117:693-7)). This “ciliary hypothesis” is based on the recognition that the protein products of genes mutated in PKD (and other pleiotropic syndromes involving cyst formation) both localize to cilia and impact ciliary function. Understanding the regulation of cilia in PKD, should provide the basis for novel therapeutic approaches to PKD.
In the past two years, increasing attention has focused on the identification of other structural and signaling proteins associated with the cilium, the basal body, or the adjacent plasma membrane. Importantly, many proteins that have been identified as the genetic cause of human developmental defects associated with polycystic kidney disease, including polycystins 1 and 2 (encoded by PKD1, PKD2), fibrocystin (PKHD1), nephrocystins (NPHP1,3-5), and inversin (NPHP2), in each case localize to cilia. Additional disease-associated proteins localizing to cilia or basal bodies include the Bardet-Biedl Syndrome (BBS) proteins. Defects in BBS genes lead to kidney failure associated with renal cysts, and also loss of eyesight, obesity, and diabetes. Kartagener syndrome, characterized by reversed left-right symmetry (“situs inversus”) of the heart, stomach and liver, as well as additional defects, arises from ciliary dyskinesia (Carlen B., et al., (2005) Ultrastruct. Pathol. (2005) 29:217-20). Finally, targeted or spontaneous mutation in mice of other cilia-associated proteins including the kinesin motor KIF3A or of Tg737/polaris, both involved in IFT, results in similar syndromes (Siroky, B. J., et al., (2006) Am J Physiol Renal Physiol.; Cano, D. A., et al., (2004) Development 131:3457-67; Yoder, B. K., et al., (2002) Am J Physiol Renal Physiol. 282:F541-52; Nishimura, T., et al., (2004) Nat Cell Biol. 6:328-34).
These and other studies make it clear that defects in proteins affecting ciliary functions are a major cause of renal cysts in general, and PKD in particular, as well as other serious diseases.
Mechanistically, the role of cilia in development and in disease is not yet well-defined, although the field is advancing rapidly (
While many of the downstream signaling components activated by Ca2+ are not yet well defined, the functional consequences of cilial bending for embryonic development are clearly profound. In development, cilial bending by extracellular fluid flow sends a polarity cue that conditions the future direction of cellular propagation. Inability to sense such flow is likely to underly the situs inversus observed in individuals mutant for some ciliary proteins such as inversin. In cell migration, fluid flow over cilia has been shown in some cases to contribute a polarity cue providing a direction for cell migration in development (Sawamoto K., et al. (2006) Science 311:629-32; Ciruna B., et al., (2006) Nature 439: 220-4). Within kidneys, flow sensing is thought to regulate proliferative response, such that defective sensing may cause overproliferation and cyst development. Separately, recently described connections between BBS proteins and the planar cell polarity (PCP) machinery imply that altered polarity of the cell division plane may cause dysplastic growth during maintenance of renal tubules, again leading to cyst formation (Fischer, E., et al., (2006) Nat. Genet. 38:21-3; Ross, A. J., et al., (2005) Nat. Genet. 37:1135-40).
An important point to consider is that as quiescent cells are induced to cycle, the cilium is reabsorbed, and the basal body returns to function within the centrosome, which includes action as microtubule organizing center (MTOC) for the bipolar spindle in mitosis. Although there are some differences among the cell systems used to study cell cycle regulation of cilia, most studies agree that cilia are reabsorbed by the time a cell enters mitosis, then re-form at several hours after the completion of cytokinesis (Quarmby et al., (2005). There are several implications of these findings. As discussed below, in addition to BBS, a number of other proteins have been identified at both basal bodies and centrosomes, and may act at both structures. These proteins may act directly at cilia and/or at centrosomes to orient the mitotic division plane of renal cells, allowing normal formation and maintenance of renal tubules. Conversely, as we demonstrate herein proteins that function at the centrosome to govern entry and exit from mitosis are ideally positioned to influence the formation and disassembly of cilia, and hence altered regulation of these proteins may contribute to the pathogenesis of PKD and related cilia-based syndromes.
Accordingly, the in vivo roles of AurA and HEF1 in cyst formation will be examined using mouse models. Appropriate models are available for this purpose. Notably, in late 2005, Seo et al. first described a HEF1 knockout mouse (Seo et al, (2005) J. Immunol. 175:3492-501). This knockout is viable and able to reproduce as a homozygote. To date, based on the interests of Sachiko Seo and her colleagues, characterization of these mice has been limited to the hematopoietic system. Even from this very limited analysis, it is clear that elimination of HEF1 has at least some phenotypic consequences, as B-cell maturation is defective. These mice will be utilized to explore the role of HEF1 in cell cycle and centrosome functions, and we have recently established a colony. Models for study of overexpressed AurA also exist (Fukuda et al., (2005) Mol. Cell Biol. 25:5270-81). However, in vivo overexpression of AurA has been reported as technically challenging, in part because of efficient proteasomal degradation of the protein. Further, AurA overexpression causes secondary phenotypes related to the failed cytokinesis seen in cells with too much AurA (Warner et al., (2003) Mol. Cancer Ther. 2:589-95). Instead, we will use small molecule inhibitors of AurA and HDACs to perform in vivo manipulation of these proteins.
Numerous mouse models have been developed for the study of PKD. Particularly because of the reported interaction of the HEF1-related protein p130Cas with polycystin, we wished to first explore the consequences of HEF1 and AurA in modulation of PKD1-associated PKD. A particularly attractive model is the conditional floxed Pkd1 model developed by the Germino group (Piontek K. B., et al. (2004) J. Am. Soc. Nephol. 15:3035-43). Mating of these animals with mice expressing Cre recombinase causes somatic loss of Pkd1, and leads to formation of renal and hepatic cysts by 10 weeks of age.
Taken as a whole, our results clearly indicate that HEF1 and AurA are regulated in time and space in a manner compatible with a controlling role in ciliary disassembly. They also demonstrate that increased activity and/or expression of HEF1 and AurA actively promote ciliary disassembly, and that clinical agents that block AurA stabilize cilia. Importantly, these results validate genetic predictions from Chlamydomonas CALK, demonstrating evolutionary conservation of AurA regulation of cilia and flagella. This conservation allows us to exploit ongoing discoveries regarding CALK in Chlamydomonas to guide our future studies. In the context of these and other published works, we theorize that a basal body-associated complex including HEF1 and polycystin (and potentially other proteins) comprises a stretch- and growth factor-responsive sensor at the cilium. In normal cells, AurA localizes to the basal body, but is only activated following receipt of signals through the HEF1-containing stretch complex. We will analyze the factors contributing to AurA activation. We also hypothesize that upon activation, AurA phosphorylates substrates located in the basal body and cilia. We also expect to observe moderately elevated expression of wild type or even inactive AurA influences the activity of effectors regulating ciliary disassembly, as well as cilia-associated signaling proteins. Methods are also provided to elucidate the mechanisms by which AurA and HEF1 condition cilia-associated signaling responses, and promote ciliary disassembly. Finally, as mentioned above, the consequences of modulating HEF1, AurA, and HDAC6 upon cyst formation in a PKD mouse model system will also be assessed.
To generate the required mouse strains for the experiments, we will first take the HEF1−/− mice we have received from Seo and coworkers and backcrossed to a C57/B16 background. Our first step will be to create a series of mice that are heterozygous or homozygous null for HEF1 on a homozygous Pkd1cond/cond background. See Piontek et al. In parallel, we will cross the HEF1 null mice to the nestinCre strain, ultimately creating mice heterozygous or homozygous null for HEF1, and hemizygously bearing the nestinCre gene. Next, we will combine these strains of mice to allow us to examine the consequences of HEF1 status on cyst formation in Pkd1cond/condnestinCre mice. These same mice can also be used for evaluation of the AurA and HDAC inhibitors. We emphasize, we do not view null status for HEF1 as independently likely to generate kidney cysts; rather, we expect the most likely activity will be to modify cyst formation induced by somatic mutation of Pkd1. See
The central component of these experiments will be to compare rate and degree of cyst formation in HEF1−/−Pkd1cond/condnestinCre, HEF1wt/wtPkd1cond/condnestinCre, and HEF1−/−Pkd1wt/wtnestinCre mice. We anticipate that most HEF1wt/wtPkd1cond/condnestinCre will have extensive cyst formation at 8-12 weeks of age. For each of the three strains of mice, we will sacrifice 10 animals at 8 and 16 weeks of age, and will use standard approaches to analyze timing, number, size, and pathological features of cyst formation. This will allow us to determine whether loss of HEF1 independently promotes cyst formation in the kidney, and whether lack of HEF1 positively or negatively regulates cyst formation dependent on defects in Pkd. We note, our crosses will also generate HEF1−/wtPkd1cond/condnestinCre animals: if a significant effect is seen with HEF1 null status, we will then determine whether HEF1 heterozygous status has an intermediate phenotype.
We note that there is (at present) no evidence directly implicating mutation of HEF1, its family member p130Cas, or Aurora in hereditary PKD in humans. This may still emerge. However, we view it as more likely that mutation of these genes would have a broader effect: for instance, mice with loss of p130Cas are embryonic lethal. By contrast, elevated levels of p130Cas, AurA, and HEF1 (unpublished) are found in numerous cancers, and thought to promote deregulated cell growth. We believe similar conditioning of cell growth by some of these proteins occurs in PKD. The methods disclosed herein will lead to the creation of new therapeutic strategies to treat PKD, by providing the scientific basis to apply existing compounds already in use for treatment of cancer to this disease. Moreover, the mice described herein will provide an in vivo model to assess the efficacy of various agents that may be useful for the treatment of diseases associated with aberrant cilia formation, such as PKD In addition, by improving our understanding of the molecular basis of PKD, we may be able to better predict disease progression and severity. Overall, the goal is to reduce the incidence and/or severity of PKD in those genetically prone to the disease, alleviate the symptoms of PKD in early stage patients, and limit the number of PKD patients that progress to end stage renal failure.
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The Aurora-A (AurA) kinase is a member of the evolutionarily conserved Ipl family of kinases (reviewed in (Marumoto et al., 2005)). AurA is most abundant at the centrosome in G2 to M phase (Fukuda et al., 2005), and in studies performed in mammals and model organisms including Drosophila, AurA has been shown to be activated at mitotic entry, and perform critical functions in regulating entry into and passage through mitosis (Marumoto et al., 2005; Pugacheva and Golemis, 2006). In the past several years, AurA has attracted increasing attention because it has been found to be overexpressed in a high percentage of tumors arising in breast, colon, ovary, and other tissues (Bischoff et al., 1998; Goepfert et al., 2002; Tanaka et al., 1999; Tanner et al., 2000; Zhou et al., 1998), and because it has been shown to function as an oncogene when exogenously expressed in various cell line models (Anand et al., 2003; Meraldi et al., 2002; Tatsuka et al., 1998; Zhang et al., 2004). AurA is now being actively exploited as a target for development of new anti-cancer agents (reviewed in (Andrews, 2005)), based on its known role as a mitotic regulator.
Intriguingly, a number of studies have emerged in recent years to challenge the idea that AurA is solely a mitotic kinase. Serum induces AurA activation at the basal body of the cell cilium in non-cycling G0/G1 phase mammalian cells, causing AurA-dependent ciliary resorption (Pugacheva et al., 2007), and hence indirectly impacting the functionality of the cilia-dependent and cancer-relevant Hedgehog signaling cascades (Wong et al., 2009). AurA directly phosphorylates and regulates the activity of the RalA GTPase, an important EGFR/Ras effector important in many cancers (Wu et al., 2005), with this activity observed in interphase cells. AurA also has been reported to regulate microtubule dynamics in interphase cells (Lorenzo et al., 2009). All of these studies strongly imply a non-mitotic activity for AurA.
In a series of studies, we here demonstrate that interactions between the NEDD9 (also known as HEF1 and CAS-L) scaffolding protein and AurA are important for AurA activation at mitotic entry (Example I) and prior to ciliary resorption in interphase cells (Pugacheva et al., 2007). Interestingly, based on mRNA and protein analysis, NEDD9 (Law et al., 1996; Law et al., 1998) and AurA (Kurahashi et al., 2007) are predicted to be abundant in kidney. Pathologic conditions of the kidney include renal cell carcinoma (RCC), which has been linked to elevated AurA expression (Kurahashi et al., 2007). Abnormal cell division associated with formation of renal cysts is also very strongly linked to the changes in Ca2+ signaling induced by autosomal dominant polycystic kidney disease (ADPKD)-associated mutations in the PKD1 and PKD2 genes, encoding the PC1 transmembrane flow receptor and the PC2 calcium channel (Benzing and Walz, 2006; Pan et al., 2005; Wilson, 2004). Interestingly, an antibody cross-reactive with NEDD9 and its paralog BCAR1/p130Cas detected one of these proteins in a complex with PC2 (Geng et al., 2000). NEDD9 is also known to bind directly to the differentiation regulatory protein Id2 (Law et al., 1999), which in turn has been reported to bind directly to PC2 and mediate proliferative signals in PKD (Li et al., 2005). Further, recent studies of a PC2 ortholog in Chlamydomonas suggested a role for this protein in increasing intraflagellar calcium concentrations during the mating response (Huang et al., 2007), shortly prior to the activation of the CALK kinase. Separately, a recent study of Xenopus oocyte maturation indicated that inhibition of Ca2+ signaling led to eventual failure to accumulate and activate AurA (Sun et al., 2008). Cumulatively, these studies led us to hypothesize that NEDD9 and AurA might physically or functionally interact with PC2 to regulate cellular calcium signaling under normal or pathological conditions.
Based on these and other studies, we have investigated the role of AurA in calcium signaling responses. Our data here demonstrate that elevated cytoplasmic calcium signals rapidly and transiently through calmodulin to activate AurA, and that active AurA directly binds and phosphorylates PC2 to limit its Ca2+ channel activity, providing the first clear mechanism for AurA activation in non-mitotic cells. Our work also demonstrates AurA is abundant and frequently active in normal renal tissue, and hyperactivated in renal cysts associated with PKD. Finally, ADPKD affects as many as 1 in 500 individuals, and currently has few viable treatment options. We show that low concentrations of drugs that inhibit AurA activity augment PC2-dependent Ca2+ release, suggesting potential clinical applications in renal proliferation-associated pathologies.
The following materials and methods are provided to facilitate the practice of example 3.
Plasmids and constructs. Lentiviral constructs were obtained by cloning full-length PKD2 into the pLV-CMV-H4-puro-vector (provided by P. Chumakov and A. Ivanov). PKD2 cloned in pcDNA3.1-myc was provided by Dr. Stefan Somlo. The PKD2 CT fragment (amino acids 779-968) was cloned into the pEGFP (Clontech, Mountain View, Calif.) and pGEX-6P1 vectors (Millipore, Billerica, Mass.). Amino acid substitution mutations were introduced into wild type human PKD2 cDNA by site-direct mutagenesis, using the QuikChange XL Site-Directed Mutagenesis Kit (Stratagene, La Jolla, Calif.). Primer sequences are available on request. The FLAG-fused C-terminal domain of PKD1 (aa 4191-4302), containing the PC1-PC2 interaction site, was cloned in pcDNA3.1(+) vector (Invitrogen, Carlsbad, Calif.). FLAG- and GST-fused HEF1 were expressed from the vectors pCatch-FLAG (O'Neill and Golemis, 2001) and pGEX-2T (Law et al., 1998), respectively. AurA and derivatives were expressed from pCMV-SPORT6-C6 (OpenBiosystems, Huntsville, Ala.) and pcDNA3.1-mRFP vectors. A PCR product of mRFP1 was ligated into pcDNA3.1(+) (Invitrogen, Carlsbad, Calif.) to create pcDNA3.1-mRFP. pLV-CMV-H4-puro-vector, pEFGP and pcDNA3 were used for negative controls.
Cell culture and transfection. HEK293 cells were maintained in DMEM with 10% FBS, plus penicillin/streptomycin. The immortalized human kidney proximal tubular cell line (HK-2, ATCC, catalog #CRL-219) was grown to subconfluence in keratinocyte serum-free media (Invitrogen, Carlsbad, Calif.). We transiently infected HEK293 cells with expression constructs for PKD2, HEF1 and AurA using Lipofectamine and Plus reagent (Invitrogen, Carlsbad, Calif.), according to the manufacturer's instructions. Cells were used for electrophysiological studies 24-48 hours post-transfection. For lentiviral infection, pLV constructs were co-transfected with pVSV-G and psPAX2 into the packaging cell line 293-T. After 24 h, media was collected, filtered through a 0.45-μm PVDF filter (Millipore, Billerica, Mass.), and applied to HK-2 cells with 8 μg/μl polybrene (Sigma, St Louis, Mo.) for 2 days, with fresh viral supernatant added every 12 h. After 48 h, cells were lysed, analyzed by western blot analysis, and used for further experiments. HK-2 cells stably expressing PC2 were obtained by infecting the HK-2 cell line with the pLV-PKD2 lentiviral vector, then selecting for 6-10 days with 1 mg/ml puromycin to produce a mass culture, as in (Pugacheva and Golemis, 2005). PC2 expression was verified by immunoblot and immunofluorescence analyses. Transient transfection of siRNAs was carried out using RNAi max transfection reagent (Invitrogen, Carlsbad, Calif.). Cells were assayed after 48 h of transfection. IC50 determinations with the AurA kinase inhibitor PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) were performed as described in (Skobeleva et al., 2007).
Immunofluorescence. Cells growing on coverslips were fixed with 4% paraformaldehyde (10 min) then methanol (5 min), permeabilized with 1% Triton-X100 in PBS, blocked in 1×PBS with 3% BSA, and incubated with antibodies using standard protocols. Primary antibodies included mouse anti-AurA (BD Biosciences, San Jose, Calif.) and rabbit polyclonal anti-phospho-AuroraA/T288, (Cell Signaling, Beverly, Mass.), polyclonal anti-α-tubulin (Sigma), anti-α-tubulin mAb (Sigma), anti-acetylated α-tubulin mAb (clone 6-11B-1, Sigma, St Louis, Mo. and clone K(Ac)40 Biomol), anti-PC2 (G20, Santa Cruz Biotechnology, Santa Cruz, Calif., and YCC2 (a gift of S. Somlo), and mouse anti-protein disulfide isomerase (PDI) mAb (Abcam, Cambridge, Mass.). Secondary antibodies labeled with Alexa-488, Alexa-568, and DAPI to stain DNA, were from Molecular Probes/Invitrogen, Carlsbad, Calif. Confocal microscopy was performed using a Nikon C1 Spectral confocal microscope (Nikon).
Protein expression, western blotting and immunoprecipitation. Recombinant glutathione-S-transferase (GST), GST-fused to amino acids 779-968 of the PC2 carboxy-terminus (GST-PC2779-968) and NEDD9 (aa 1-363, previously shown to bind and activate AurA, (Pugacheva and Golemis, 2005)) were expressed in BL21 (DE3) bacteria, induced with IPTG, and purified using the MicroSpin GST Purification module (GE Healthcare, Piscataway, N.J.). Purified recombinant AurA was purchased from Upstate (Charlottesville, Va.). For western blotting and immunoprecipitation, mammalian cells were disrupted in CelLytic M lysis buffer (Sigma, St. Louis, Mo.) supplemented with a protease and phosphates inhibitor cocktails (Roche, Basel Switzerland). Whole-cell lysates were used either directly for SDS polyacrylamide gel electrophoresis (SDS PAGE), or for immunoprecipitation. Immunoprecipitation samples were incubated overnight with antibody at 4° C., and subsequently incubated for 2 h with protein A/G-sepharose (Pierce, Rockford, Ill.), washed and resolved by SDS-PAGE. For analysis of PC2 glycosylation, cell lysates were treated with endoglycosidase H (Endo H) (New England Biolabs, Beverly, Mass., USA) and analyzed by SDS-PAGE followed by immunoblotting as described previously (Cai et al., 1999; Koulen et al., 2002). GST-pulldown assays used wild type AurA (Upstate, Charlottesville, Va.) mixed with titrated quantities of GST and GST-PC2779-968.
Western blotting was done using standard procedures, and developed by chemoluminescence using the West-Pico system (Pierce, Rockford, Ill.). Primary antibodies included mouse anti-NEDD9 mAb (clone 2G9, (Pugacheva and Golemis, 2005)), anti-AurA (BD Biosciences, San Jose, Calif.), anti-phospho-AurA-T288 (Cell Signaling, Beverly, Mass.), anti-Myc- and anti-VlaR (Santa Cruz Biotechnology, Santa Cruz, Calif.), anti-β-actin mAb (AC15, Sigma, St Louis, Mo.), and anti-PC2 (G20, Santa Cruz Biotechnology, Santa Cruz, Calif., and YCC2, a gift of S. Somlo). Rabbit anti-GFP (Abcam, Cambridge, Mass.; ab290) was used for immunoprecipitation, and mouse anti-GFP (JL-8; BD Biosciences, San Jose, Calif.) was used for western blotting. Anti-GST mAb (Cell Signaling, Beverly, Mass.), polyclonal EZview Red anti-Flag M2 affinity gel (Sigma, St Louis, Mo.) and polyclonal anti-AurA agarose immobilized conjugate (Bethyl, Montgomery, Tex.) were used for immunoprecipitations. Secondary anti-mouse and anti-rabbit HPR conjugated antibodies (GE Healthcare, Piscataway, N.J.) were used at a dilution of 1:10,000 for visualization of western blots. Image analysis was done using NIH ImageJ—Image Processing and Analysis software (National Institutes of Health, Bethesda, Md.), with signal intensity normalized to β-actin or total AurA level.
To assess AurA phosphorylation of PC2 in vivo, myc-tagged PC2 was transiently expressed alone or with AurA or T288D-AurA in HEK293 cells and then immunoprecipitated with anti-myc antibody (Santa Cruz Biotechnology, Santa Cruz, Calif.). Phosphorylation of the S829 site was assessed by western blot with Phospho-(Ser/Thr) PKA Substrate Antibody (Cell Signaling, Beverly, Mass.). 500 nM PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) was used to inhibit AurA and 10 μM H89 PKA inhibitor (Calbiochem) were used to inhibit phosphorylation.
Calmodulin pull-down assay and Immunoprecipitation. Cell lysates in lysis buffer (PBS with 1% Triton X-100) or purified GST fusion protein diluted in binding buffer (50 mM Tris-HCl, pH7.6; 120 mM NaCl; 1% Brij) were incubated with Calmodulin-Sepharose® 4B (GE Healthcare, Piscataway, N.J.) or control Sepharose for 1-2 h at 4° C. as indicated in the figure legends. After washing, beads were boiled in SDS sample buffer and separated by SDS-PAGE followed by Western blotting.
Kinase assays. To assess phosphorylation of PC2 by AurA, an in vitro kinase assay was performed using bacterially expressed GST-fused PC2 CT and recombinant active AurA (Upstate, Charlottesville, Va.) or overexpressed AurA immunoprecipitated from mammalian cells. in standard kinase buffer with addition of an Mg/ATP cocktail (Upstate, Charlottesville, Va.). MBP (Upstate, Charlottesville, Va.) and histone H1 (Upstate, Charlottesville, Va.) were used as positive and negative controls for AurA phosphorylation, using standard methods. Parallel aliquots without γ-32P(ATP) were processed for SDS-PAGE/Coomassie staining (Invitrogen, Carlsbad, Calif.). To assess CaM-dependent AurA activation in vitro kinase assay was performed using AurA purified from baculovirus or according the protocol described above in the presence of 1 μM CaM (Calbiochem) and 1 mM Ca2+. For kinase assay without Ca2+, 1 mM EGTA was substituted for 1 mM Ca2+ in the kinase buffer.
Mass spectrometry. After an in vitro kinase reaction with AurA produced from baculovirus in the presence or absence of CaM plus 1 mM CaCl2, gels were stained with Coomassie blue, and phosphorylated AurA bands were excised and sequenced at the Taplin Biological Mass Spectrometry Facility at Harvard Medical School, by using microcapillary reverse-phase HPLC nanoelectrospray tandem mass spectrometry on a Finnigan LCQ DECA quadrupole ion trap mass spectrometer).
Cytosolic Ca2+ measurements. Cells expressing PC2 constructs were plated on glass coverslips and grown to ˜80% subconfluence. The coverslips were rinsed in Hanks' balanced salt solution (HBSS), and incubated with Fluo-4AM (5 μM in HBSS) (Invitrogen, Carlsbad, Calif.) in the presence of 0.02% pluronic acid (Invitrogen, Carlsbad, Calif.), and 2.5 mM probenecid (Invitrogen, Carlsbad, Calif.) for 15-30 minutes at RT. The coverslips were washed twice in HBSS, mounted in a perfusion chamber (FC2, Bioptechs, Butler, Pa.), and analyzed with a Nikon C1 Spectral confocal microscope (Nikon). Cytosolic Ca2+ measurements done in the absence of extracellular Ca2+ were performed on cells washed and assayed in the HBSS described above except that CaCl2 was omitted and 0.5 M EGTA was added. In experiments involving AurA inhibition, cells were treated for 2 h with 500 nM PHA-680632 (Nerviano Medical Sciences, Nerviano, IT) before calcium measurement. Fluo-4 was excited at 488 nm and emission was time-lapse recorded at 522 nm. Cells were individually selected, and their fluorescence intensities (F) were normalized to baseline (F0), and analyzed with Metamorph and Metafluor softwares (Molecular Devices, Union City, Calif.).
To compare cellular responses, we examined differences in intensity over time using a generalized linear model assuming Gamma distribution and log link. We fit the models by generalized estimating equations assuming an autoregressive correlation structure to account for correlation of observations over time. We included baseline intensity, group, time, and the interaction between time and group in the models. To allow for flexible effects over time, we entered time and the related interactions in the model using restricted cubic splines assuming 5 knots (Harrell, 2001). We used Wald tests to assess p-values of group effects at each time point.
To induce ciliary formation in the HK2 cell line, cells were plated at 50%-70% confluence and grown for 3-4 days in medium with 10% serum, then grown for 2 days in Opti-MEM medium without serum. Immunofluorescence was used to confirm >80% of HK2 cells were ciliated before use in experiments.
Immunohistochemistry. All tissue samples examined were IRB-consented, 10-20 mm sections of formalin-fixed, paraffin-embedded tissues representing either normal human kidney tissue, or kidney tissue from patients diagnosed with PKD. A standard two-stage indirect immunoperoxidase staining protocol was used for all tissues (Vectastain ABC System; Vector Laboratories), with antigen retrieval buffer from BD Biosciences, San Jose, Calif. As controls, sections were stained with diluent alone (5% goat serum in Tris-buffered saline) and antibody pre-absorbed with the immunizing peptide. Incubations with tissue sections were carried out at room temperature for 1 h or at 4° C. overnight, and subsequent steps were carried out at room temperature. Staining was visualized with either diaminobenzidine (brown) or VIP (purple) (Vector Laboratories), and sections were counterstained with hematoxylin (Sigma, St Louis, Mo.). Antibody to total AurA (anti BTAK ab, provided by T. Gritsko) was used at a dilution of 1:500, and to phospho-T288 AurA at 1:100 (Bethyl, Montgomery, Tex.). Images were acquired at 10× and 40× using a Nikon Eclipse E600 microscope.
Statistical Analysis. Statistical comparisons were made using a two-tailed Student's t test. Experimental values were reported as the means±S.E. Differences in mean values were considered significant at p<0.05. All calculations of statistical significance were made using the GraphPad InStat software package (San Diego, Calif.).
Rapid transient activation of AurA accompanies stimuli for Ca2+ release from ER stores in kidney cells. Polycystin-2 (PC2) Ca2+ channel activity is stimulated by treatment of kidney cells with the peptide hormone and anti-diuretic arginine vasopressin (AVP) (Cai et al., 2004; Koulen et al., 2002), which signals through the AVPR1 receptor to initiate InsP3 and other second messenger cascades that initiate Ca2+ release from the ER. To assess whether activation of AurA occurred in this physiological context, we treated the HK-2 proximal tubule-derived human kidney cell line (Ryan et al., 1994) with AVP, and used immunofluorescence to measure AurA activation (
The rapid AurA activation following treatment might be induced by the second messengers transmitting signals from membrane localized receptors, or alternatively, the elevated cytoplasmic Ca2+ levels resulting from ER-release might more directly regulate AurA activation, with no second messengers required. To distinguish between these mechanisms, we treated AurA-transfected HEK293 cells with the Ca2+ uptake inhibitor thapsigargin, which blocks the recycling of cytoplasmic Ca2+ into ER stores (Sagara and Inesi, 1991), thus increasing cytoplasmic Ca2+ levels. Thapsigargin treatment caused a rapid (<1 minute) increase in levels of activated T288-AurA, and increased the in vitro kinase activity of immunoprecipitated AurA against defined substrates (
Calcium induces calmodulin to bind and activate AurA. Increased cytoplasmic Ca2+ might induce AurA activation by direct binding, or through inducing conformational changes in AurA by triggering its binding to a Ca2+-binding effector such as calmodulin (CaM), or by activating of an intermediate kinase or phosphatase that targets AurA, In an in vitro kinase assay, titration of Ca2+ into the reaction did not affect AurA autophosphorylation or activity towards substrates (not shown). However, addition of Ca2+ with CaM strongly induced AurA autophosphorylation (
Aurora A interacts directly with PC2. To assess the role of Ca-dependent AurA activation, and identify its physiological substrates, we considered a possible connection to PC2, based on the reported association between polycystins and Cas proteins (p130Cas and NEDD9). The bulk (>95%) of intracellular PC2 associates with the ER, and mediates Ca2+ release to the cytoplasm (Cai et al., 1999). To assess whether there might be direct interactions between AurA, its previously defined partner and activator NEDD9 (Pugacheva and Golemis, 2005), and PC2, we first established that endogenous AurA and PC2 coimmunoprecipitated from HK-2 cells (
AurA phosphorylates PC2 on C-terminal residue S829. We next asked if PC2 might be a substrate of AurA. Besides the PC1 interaction motif, the PC2 C-terminus (
To investigate the in vivo phosphorylation of the S829 site, we exploited the fact that this site is quite similar to the general PKA substrate consensus (RRxS). Phospho-PKA substrate antibody recognized PC2 but not S829A-mutated PC2 in transiently transfected HEK293 cells (
AurA negatively regulates PC2 Ca2+ channel activity. We next asked if AurA might play a role in regulating the PC2 Ca2+ channel. We transiently transfected HEK293 cells with PC2 together with RFP-AurA or an RFP negative control, and measured fluorescence of the cytoplasmic Ca2+-binding dye Fluo-4 following AVP treatment, comparing the amplitude and duration of Ca2+ release (
A cytoplasmic pool of AurA regulates ER-localized PC2. AurA is typically thought of as localized to the centrosome or centrosomally-derived ciliary basal body, which would limit access to an ER-localized pool of PC2. In contrast, cancerous cells with overexpressed AurA have an extensive diffuse pool of cytoplasmic AurA. However, AurA is intrinsically abundant in kidney cells, and indeed, siRNA depletion experiments followed by detection with two different antibodies to AurA (
PC2 activity can be controlled by direct changes in the activity of the ER-localized protein, or alternatively, based on phosphorylation-mediated relocalization of PC2 between ER and other cellular compartments such as the cilium or plasma membrane, for example via CK2 phosphorylation at S812 (Kottgen et al., 2005). The experiments described above were performed in cycling, predominantly unciliated cells, leading us to favor the former mechanism; however, to specifically assess whether the presence of intact cilia influence AurA regulation of PC2 activity, we directly compared the ability of PHA-680632 to increase AVP-induced PC2 channel activity in predominantly non-ciliated (<15%), cycling versus ciliated (>80%), non-cycling HK-2 cells. These results showed no significant difference in the degree of Ca2+ release under the two growth conditions (
Abundant AurA expression and activity in normal and cystic kidney tissue. If AurA has in vivo, non-cell cycle function relevant to PC2, AurA should be detectable and potentially active in non-dividing renal tissue. Immunohistochemical analysis of primary human kidney specimens readily detected AurA in multiple sub-structures (
PHA-680632 enhances PC2 activity at low IC values. There is a considerable need for novel therapeutics to treat PKD. Given that one consequence of mutations in PKD1 is to reduce PC2 activation (Nauli et al., 2003), if AurA inhibition enhances PC2 activation, this might lead to therapeutic benefits. In vivo, Aurora kinase inhibitors have marked effects as cell cycle inhibitors (Gautschi et al., 2008), relevant to their action in cancer therapy, raising the possibility of toxic side effects if these agents used in PKD. However, some recent studies have suggested that the cytotoxic effects of Aurora kinase inhibitors used in vivo at least partially reflects their cross-reactive inhibition of Aurora-B rather than AurA, which occurs at higher concentrations (discussed in (Gautschi et al., 2008)).
We have compared the doses of PHA-680632 required to inhibit cell growth with those required to enhance PC2 signals. In HK-2 cells, the IC50 value for PHA-680632 is 3.25 μM, while the 500 nM concentration used for the experiments described above represents an IC50 value (
The work presented here reveals and mechanistically defines a novel activity of AurA, as a direct target of calcium activation, and in direct control of cellular homeostasis for calcium.
Although to our knowledge a direct AurA-calcium connection has never previously been investigated, there are some suggestions that a connection between AurA and calcium signaling may be evolutionarily conserved. Flagellar disassembly in Chlamydomonas is induced by stimuli including gamete fusion following mating (Cavalier-Smith, 1974). Intriguingly, recent studies of a PC2 ortholog in Chlamydomonas suggested a role for this protein in increasing intraflagellar calcium concentrations during the mating response (Huang et al., 2007), prior to the activation of the CALK kinase. Separately, a recent study of Xenopus oocyte maturation indicated that inhibition of Ca2+ signaling led to eventual failure to accumulate and activate AurA, and subsequently a defective phenotype in meiosis (Sun et al., 2008). Dynamic changes in calcium signaling play a key role in meiosis, have been implicated in action at spindle-associated microdomains in mitosis (Parry et al., 2005), and may both regulate and be regulated by AurA. An attractive feature of Ca2+-dependent AurA regulation is that it offers a potential mechanism to help explain the rapid, timed activation of AurA at cell cycle transitions. At present, although multiple proteins have been demonstrated to bind and support AurA activation at the mitotic boundary, most of these also interact with AurA in G2, implying the existence of a triggering event at the actual transition point. The transient increase of cytoplasmic Ca2+ could provide a suitable mitotic trigger. Such Ca2+- and CaM-dependent regulation of the centrosomal proteins centrin and CP110 has been shown to be critical for the action of these proteins in supporting cytokinesis. In addition, reports that PC2 itself localizes to the mitotic spindle in dividing cells (Rundle et al., 2004) supports the idea that the mechanism of PC2-dependent Ca2+ release and calmodulin binding may be a physiological trigger for AurA during cell cycle, causing conformational changes that are permissive for other AurA interacting proteins to sustain its activity. However, the extremely transient nature of the Ca2+-dependent activation we observe makes it difficult to test this idea directly.
Interestingly, published studies of AurA in cancer often suggest the pro-oncogenic activity of AurA may arise because its overexpression allows a normally centrosomal protein access to inappropriate substrates. However, our results suggest changes in AurA expression in cancer may promote quantitative changes in AurA activity more than qualitative phosphorylation of novel substrates, as based on the data presented here, AurA has a broader sub-cellular localization profile in untransformed cells than is commonly appreciated, with the centrosome acting as a concentration point at which the protein is most readily visualized. Our data indicate that not only kidney cells, but also primary kidney tissue express significant quantities of AurA in the cytoplasm and nucleus, and that some of this AurA is activated in non-cycling cells in normal kidney tissue, particularly in cells of the distal convoluted tubules and collecting ducts from which cysts arise. This broader view of AurA activity is compatible with recent reports that AurA phosphorylates RalA (Wu et al., 2005) (a protein not known for either centrosomal localization or mitotic-specific action, but which has itself been described as calmodulin-binding (Clough et al., 2002)).
In general, calcium signaling differs significantly in cancerous (Roderick and Cook, 2008) and cystic (Harris and Torres, 2008) cells versus normal cells, promoting increased cell proliferation through the abnormal activation of numerous calcium-responsive signaling pathways. The fact that AurA activation was elevated in PKD-associated cysts is interesting, and may reflect paradoxical activation in the context of mutated PKD1 and PKD2, analogous to the overexpression of growth inhibitory proteins in tumors that have eliminated partners in a feedback loop. Based on our results, it appears that inappropriately activated AurA may act as an intermediate in some of signaling processes relevant to PKD. For example, besides binding the PC2 partner Id2, NEDD9 directly binds and is both target and activator of Src kinase (recently discussed in (Singh et al., 2007)). Src signaling is abnormal in PKD, and a recent study has indicated that inhibition of Src produces clinical benefits in PKD (Sweeney et al., 2008). Through interactions with NEDD9, AurA may influence the activity of Src and Id2 in either normal renal tissue or in cysts. These close physical interactions suggest further topics of study not only in renal cysts but also in cancer, where NEDD9, Src, and Id2 all have oncogenic function.
There is potential therapeutic benefit to identifying ways to stimulate PC2 channel activity, and there is an urgent need to develop effective therapies for PKD. At present, a number of targeted therapeutic agents are moving through pre-clinical development and clinical trials. Besides c-Src, these include agents targeting mTOR, HER2, and others. These studies provide a precedent for adapting drugs originally developed as cancer therapeutics in PKD. An obvious concern is that given the chronic but survivable nature of PKD, it is necessary to be extremely cautious in using powerful compounds that may themselves ultimately select for oncogenic changes. However, our data suggest that very low doses of an AurA-targeting inhibitor are able to enhance PC2 activity, suggesting a basis for further investigation of such agents in cases of PKD linked to PKD1 mutation, where PC2 is insufficiently activated but structurally intact. It is also fascinating to note that defects in PKD1 and PKD2 have recently been linked to centrosomal amplification in both animal models and human patients (Battini et al., 2008; Burtey et al., 2008), reducing the separation between cystic syndromes and cancer, and supporting the idea that calcium-dependent activation of AurA is relevant to the severity of PKD presentation.
It is clear that mutations that cause polycystic kidney disease (PKD) reduce the functionality of two proteins, PC1 and PC2, encoded by the PKD1 and PKD2 genes. Therapies that help increase PC1 and/or PC2 activity are predicted to help reverse the consequences of having PKD1 or PKD2 mutations. Our group has for the first time identified a role for the Aurora-A kinase in negatively regulating PC2 activity. Because Aurora-A has attracted interest as an oncogene important for some cancers, drugs have been developed and are being evaluated in clinical trials that can block Aurora-A activity. The data provided herein indicate that these drugs increase PC2 function, thus supporting their use as new therapeutics for PKD. See
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While certain preferred embodiments of the present invention have been described and specifically exemplified above, it is not intended that the invention be limited to such embodiments. Various modifications may be made to the invention without departing from the scope and spirit thereof as set forth in the following claims.
Claims
1. A method for identifying agents which modulate calmodulin-Aurora A complex formation having efficacy for the treatment of polycystic kidney disease (PKD), comprising:
- a) providing kidney cells which express Aurora A and calmodulin and incubating cells in the presence and absence of said agent and in the presence of a molecule which induces release of Ca2+;
- b) determining the extent said agent disrupts calmodulin-Aurora A complex formation following release of calcium, agents which disrupt complex formation having efficacy for the treatment of PKD.
2. The method of claim 1, wherein said agent inhibits activation of aurora A kinase.
3. The method of claim 2, wherein said aurora A kinase inhibitor is selected from the group consisting of siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, MLN 8054, VX-680 and hesparadin.
4. The method of claim 1, comprising determining whether said agent disrupts aurora A binding to PC2.
5. The method of claim 1, comprising determining whether said agent alters cellular localization of PC2.
6. The method of claim 1, comprising determining whether said agent alters Aurora A phosphorylation of PC2.
7. A method for inhibiting progression of polycystic kidney disease in a patient in need thereof, comprising administration of an effective amount of an aurora kinase inhibitor, said inhibitor being effective to inhibit cyst formation in the kidney.
8. The method of claim 7, wherein said kinase inhibitor is effective to inhbit aurora kinase A and aurora kinase B.
9. The method of claim 7, wherein said kinase inhibitor is effective to inhibit aurora kinase A or aurora kinase B.
10. The method of claim 7, wherein said aurora kinase inhibitor is selected from the group consisting of siRNA which specifically down modulate Aurora A expression, C1368, PHA-680632, MK-0457, ZM447439, MLN 8237, VX-680 and hesparadin.
Type: Application
Filed: Dec 14, 2009
Publication Date: Jun 3, 2010
Inventors: Olga V. Plotnikova (Philadelphia, PA), Elena N. Pugacheva (Morgantown, WV), Erica A. Golemis (Oreland, PA)
Application Number: 12/637,669
International Classification: A61K 31/7088 (20060101); C12Q 1/48 (20060101); A61P 13/12 (20060101); C12Q 1/68 (20060101);
