p53 tumor suppressor protein regulates the levels of ... - Nature

Oncogene (2006) 25, 1–7

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ORIGINAL ARTICLE

p53 tumor suppressor protein regulates the levels of huntingtin gene expression Z Feng1,7, S Jin1,2,7, A Zupnick3, J Hoh4, E de Stanchina5, S Lowe5, C Prives3 and AJ Levine1,6 1 Cancer Institute of New Jersey, University of Medicine and Dentistry of New Jersey, New Brunswick, NJ, USA; 2Department of Pharmacology, University of Medicine and Dentistry of New Jersey, Piscataway, NJ, USA; 3Department of Biological Sciences, Columbia University, New York, NY, USA; 4Department of Epidemiology and Public Health, Yale University, New Haven, CT, USA; 5Cold Spring Harbor Laboratories, Cold Spring Harbor, NY, USA and 6School of Natural Sciences, Institute for Advance Study, Princeton, NJ, USA

The p53 protein is a transcription factor that integrates various cellular stress signals. The accumulation of the mutant huntingtin protein with an expanded polyglutamine tract plays a central role in the pathology of human Huntington’s disease. We found that the huntingtin gene contains multiple putative p53-responsive elements and p53 binds to these elements both in vivo and in vitro. p53 activation in cultured human cells, either by a temperature-sensitive mutant p53 protein or by gamma-irradiation (c-irradiation), increases huntingtin mRNA and protein expression. Similarly, murine huntingtin also contains multiple putative p53-responsive elements and its expression is induced by p53 activation in cultured cells. Moreover, c-irradiation, which activates p53, increases huntingtin gene expression in the striatum and cortex of mouse brain, the major pathological sites for Huntington’s disease, in p53 þ / þ but not the isogenic p53/ mice. These results demonstrate that p53 protein can regulate huntingtin expression at transcriptional level, and suggest that a p53 stress response could be a modulator of the process of Huntington’s disease. Oncogene (2006) 25, 1–7. doi:10.1038/sj.onc.1209021; published online 7 November 2005 Keywords: p53; hungtingtin; Huntington’s disease; p53-responsive element; transcription regulation; gene expression

Introduction Huntington’s disease is a progressive neurodegenerative disorder characterized by selective loss of neurons (Huntington’s Disease Collaborative Research Group, 1993). The degeneration preferentially occurs in the striatum and the deep layers of the cortex and, during Correspondence: Dr AJ Levine, School of Natural Sciences, Institute for advanced study, Einstein Drive, Princeton, NJ 8540, USA. E-mail: [email protected] 7 These two authors contributed equally to this work Received 17 May 2005; revised 7 July 2005; accepted 7 July 2005; published online 7 November 2005

the later stages of the disease, extends to a variety of brain regions (Huntington’s Disease Collaborative Research Group, 1993; Li and Li, 2004). This disease is caused by expansion of a polymorphic trinucleotide repeat (CAG)n located in the coding region of the huntingtin gene, which translates as a polyglutamine repeat in the protein product. Accumulation of cytoplasmic and nuclear aggregates of the mutant huntingtin protein with an expanded polyglutamine tract is thought to play a central role in the pathology of Huntington’s disease (DiFiglia et al., 1997). The huntingtin gene is expressed in most tissues and it is required for normal development (Nasir et al., 1995). The normal function of the huntingtin protein is not fully understood. In addition to its possible functions in the cell nucleus, recent studies indicate that normal huntingtin protein is required for fast axonal trafficking (Trushina et al., 2004). Reduction of normal huntingtin expression, or expression of the polyglutamine track expanded mutant huntingtin proteins, which interact with the normal huntingtin proteins, impairs vesicular and mitochondrial trafficking in mammalian neurons, leading to mitochondrial dysfunction and toxicity (Trushina et al., 2004). Although the regulation of huntingtin gene expression has not been well studied, both genetic and environmental stress factors could affect huntingtin gene expression (Dixon et al., 2004), and that in turn could impact upon the onset and prognosis of Huntington’s disease (Georgiou et al., 1999; Anca et al., 2004). p53 is a tumor suppressor gene that is mutated in more than 50% of human cancer (Levine, 1997; Vogelstein et al., 2000). It encodes a transcription factor that plays a central role in integrating various stress signals, in particular, genotoxic stresses such as DNA damage, hypoxia, and oncogene activation (Levine, 1997; Vogelstein et al., 2000; Jin and Levine, 2001). These stresses trigger signal transduction pathways leading to the activation of the p53 protein, which in turn binds to the p53-responsive elements (p53 REs), in its downstream target genes and alters their rates of transcription. The transcriptional activation of downstream target genes appears to account for most of the mechanism by which p53 regulates its downstream cellular programs, including cell cycle arrest and

p53 regulates huntingtin Z Feng et al

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programmed cell death. The consensus p53-responsive elements contain a repeat of PuPuPuC(A/T) (T/A)GPyPyPy (Pu, purine; Py, pyramidine) with a spacer ranging from 0 to 14 base pair (el-Deiry et al., 1992; Hoh et al., 2002). Recently a novel algorithm (p53MH) was developed for detecting the p53-responsive genes (Hoh et al., 2002). A genome wide search for p53 target genes in human and mouse was carried out using this algorithm. Interestingly, the huntingtin gene was found to contain a number of putative p53 binding sites scattered throughout the gene (Table 1, Supplementary Information; and Figure 1a). Since p53 is a central integrator for stress signals that is involved in regulating cell death and senescence, and a mutant form of the huntingtin is responsible for an age related neurodegenerative disease, Huntington’s disease, we investigated the regulation of the huntingtin gene by p53. Here we demonstrate that the p53 protein interacts with the putative p53-responsive elements in the huntingtin gene both in vivo and in vitro. Moreover, in both cultured cells and mouse brain tissues, p53 mediates the regulation of the huntingtin gene that is triggered after DNA damage. These results demonstrate that the p53 protein regulates the huntingtin gene.

Results Human huntingtin gene contains putative p53-responsive elements The tumor suppressor p53 protein is a transcription factor that specifically recognizes a degenerate DNA responsive element loosely defined by PuPuPuC(A/T) (T/A)GPyPyPy (N)0–14 PuPuPuC(A/T) (T/A)GPyPyPy, where Pu stands for purine, Py stands for pyramidine and N stands for any nucleotide (el-Deiry et al., 1992). Hoh et al. (2002) developed a novel algorithm, which employs a weighted matrix to score the likelihood of p53 binding, and a program for genome-wide scanning for putative p53-responsive elements. The human genome was scanned for putative p53 sites and the genes that contain high-score putative p53 RE sites were identified and compiled (Hoh et al., 2002). Interestingly, huntingtin was one of the genes that contained high-score putative p53 RE sites (Supplementary Information, Table 1). As shown in Figure 1a there are three potential p53 RE sites, including one in the promoter region, one in intron 2, and one in intron 3. p53 binds to human huntingtin p53-responsive elements in vivo and in vitro The ability of these sites to interact with the p53 protein was examined both in vivo and in vitro. First, a chromatin immunoprecipitation (ChIP) assay was performed in a human cell line, H1299/V138, to test the in vivo interactions between p53 and these three potential p53-responsive elements. H1299/V138 was established by stably transfecting a temperature-sensitive mutant form of p53 (alanine 138 to valine) into H1299, a human lung cancer cell line that does not contain endogenous Oncogene

Figure 1 The Putative human huntingtin p53 REs interact with p53 in in vivo and in vitro. (a) The potential p53 REs in human huntingtin gene. Number indicates the nucleotide position relative to the transcription initiation site. Pu, purine and Py, pyramidine. (b) Chromatin immunoprecipitation (ChIP) assay of interactions between p53 and chromatin regions (as indicated) containing p53 REs in H1299/V138 cells and the parental p53 null H1299 cells. H1299/V138 and H1299 cells were cultured at 321C for 18 h before collecting. DO-1 p53 antibody (lanes 4 and 5), no antibody (lane 2) or nonspecific mouse IgG (lane 3) were used for ChIP assay. 1/20 DNA of ChIP was used for PCR as Input DNA (lane 1). MDM2 promoter p53 RE region was used as a positive control. (c) Gel mobility shift and supershift analysis. Reaction mixtures containing 32 P-labeled GADD45, human huntingtin intron 2, or intron 3 oligonucleotides were incubated with 40, 60, or 80 ng of bacterial His-p53 protein (lanes 1–3, 7–9, and 13–15). Unlabeled DNA (50 ) of the corresponding sequence was added as a specific competitor (lanes 4, 10, and 16). p53 specific antibody PAb1801 was added to supershift p53-DNA complexes (lanes 5, 11, and 17). Mutant GADD45 oligonucleotide was used to indicate nonspecific binding (lane 19). Migration positions of free probe, p53-oligo, and 1801-p53-oligo complexes are shown.

p53 (Pochampally et al., 1999). By shifting the culture temperature from 37 to 321C, p53 is stabilized, binds to p53 REs and activates transcription of its downstream genes. As shown in Figure 1b, the H1299/V138 cells were shifted from 37 to 321C and cultured for another 18 h, and the ChIP assay performed with an anti-p53 antibody, DO-1, detected the interactions between p53


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and the chromatin regions containing the putative p53 REs in huntingtin protomer region, intron 2 and intron 3, respectively (Figure 1b, lane 4). These interactions are comparable to that between p53 and the MDM2 gene p53 RE located in the promoter region. As negative controls, these chromatin fragments could not be co-immunoprecipitated with the DO-1 antibody in the parental H1299 cells cultured at 321C, which does not contain any endogenous p53 protein (Figure 1b, lane 5); and a nonspecific mouse IgG antibody could not precipitate these fragments in H1299/V138 cells shifted to 321C (Figure 1b, lane 3). These results clearly show that p53 interacts with all these three putative p53 REs in the cells. To further confirm the direct interactions between p53 and these putative p53 REs, gel mobility shift and supershift analyses were performed. The oligonucleotides corresponding to the putative p53 REs and their endogenous flanking sequences in intron 2 and intron 3 of the human huntingtin gene were 32P-labeled and incubated with the p53 protein. As shown in Figure 1c, these two oligonucleotides bind to the p53 protein in a dose-dependent manner (lanes 6–9 and lanes 12–15, respectively), and the interactions between 32P-labelled oligonucleotides and p53 were disrupted by an excess amount of nonradio-labelled oligonucleotides (lanes 10, 16). Moreover, the identities of the p53-oligonucleotide complexes were further confirmed by mobility supershift assay with a p53 antibody, Pab1801 (lanes 11 and 17). Taken together, these results indicate that p53 specifically interacts with these putative p53 REs, although the affinities between these elements and p53 are somewhat lower than that between p53 and the well-characterized p53 RE in GADD45 gene (compare lanes 6–11, lanes 12–17, to lanes 1–5). Activation of p53 in cultured cells increases huntingtin mRNA and protein levels To analyse the functional significance of the interactions between p53 and the p53 REs in the huntingtin gene, the ability of p53 to regulate the expression of the huntingtin gene was examined. p53 activity was induced by shifting the culture temperature of the H1299/V138 cells from 37 to 321C, and the mRNA levels of huntingtin in these cells were measured by real-time PCR. As shown in Figure 2a (left panel), upon p53 activation, the huntingtin mRNA levels increased in a time dependent manner. By 24 h, huntingtin mRNA increased by B2.8-fold as normalized against actin mRNA. As a control, exactly the same temperature shift had little effect on huntingtin expression in the parental H1299 cells (Figure 2a, right panel), which lacks endogenous p53 protein. Similarly, the huntingtin protein levels increased by B3-fold in H1299/V138 cells upon temperature shift to 321C, while little changes in huntingtin protein levels were observed in the parental H1299 cells upon the same treatment (Figure 2b). DNA damage is among the most important physiological signal that activates the p53 protein (Levine, 1997; Vogelstein et al., 2000; Jin and Levine, 2001). To determine whether activation of p53 by this physio-

Figure 2 p53 activation increases huntingtin gene expression in cultured human cells. (a) Quantitative real-time PCR analysis of huntingtin mRNA levels after temperature shift from 37 to 321C for indicated amount of time in H1299/V138 cells and in H1299 cells. The levels of huntingtin mRNA were normalized against the levels of actin. (b) Western blot analysis of huntingtin protein in H1299/V138 cells and in H1299 cells after growing at 321C for indicated amount of time. The protein levels of actin were determined as control. (c) Western blot analysis of huntingtin protein in HCT116 p53 þ / þ cells and HCT116 p53/ cells after g-irradiation (IR). HCT116 p53 þ / þ and HCT116 p53/ cells at 50% confluence were treated with g-irradiation (5 Gy) and cultured for indicated amount of time before harvesting. Proteins were extracted and Western blot analysis was performed to determine the levels of huntingtin protein. The protein levels of GAPDH were determined as control. The data represent three independent experiments, and the error bar represents standard deviation.

logical stress induces huntingtin gene expression, we measured huntingtin mRNA and protein levels in HCT116 cells upon gamma(g)-irradiation. HCT116 p53 þ / þ is a human colon carcinoma cell line, which contains a wild-type p53 gene, whereas the HCT116 p53/ is a p53 knockout cell line derived from HCT116 p53 þ / þ by homologus recombination (Bunz et al., 1998). As shown in Figure 2c, g-irradiation (5 Gy), which induces double-stranded DNA breaks and activates p53, increased the huntingtin protein levels by B2–3-fold in HCT116 p53 þ / þ cells. It is apparent that the huntingtin upregulation is p53-dependent, since g-irradiation had no effect on huntingtin expression in HCT116 p53/ cells (Figure 2c, right panel). The mRNA levels of huntingtin were also determined by real-time PCR analysis and g-irradiation increased huntingtin mRNA levels by B2-fold in HCT116 p53 þ / þ cells but had little effect in HCT116 p53/ cells (data not shown). The same p53-dependent upregulation of huntingtin gene expression after g-irradiation was also observed in human lung epithelial Oncogene


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H460 cells containing wild-type p53 while not in p53 null human lung epithelial H1299 cells (data not shown). Taken together, these results demonstrated that activation of p53 indeed upregulates the expression the huntingtin gene in cultured cells. p53 mediates the upregulation of huntingtin upon DNA damage by g-irradiation in mouse striatum and cortex Although huntingtin gene is expressed in most tissue types, the major targets affected by the mutant (trinucleotide expanded) huntingtin protein are striatum and deep layers of cortex neurons (Huntington’s Disease Collaborative Research Group, 1993; Sharp and Ross, 1996; Li and Li, 2004). Apparently, accumulation of the mutant huntingtin protein aggregates contributes to neuron degeneration in the pathological regions, which causes most symptoms in Huntington’s disease. Therefore, it is of interest to examine the effect of p53 activation on huntingtin gene expression in these tissues in animals. As shown in Supplementary Table 1, similar to the human huntingtin gene, the mouse huntingtin gene also contains multiple high-score putative p53 REs. Thus, the ability of p53 in regulating huntingtin gene was tested in murine cells, Vm10 cells, the fibroblast cells expressing a temperature-sensitive mutant p53 protein (alanine 135 to valine) (Wu and Levine, 1994). As shown in Figure 3a, upon a temperature shift from 39 to 321C, huntingtin mRNA levels increased in a time dependent manner and reached B5-fold increased levels by 24 h, as measured by Northern blot analysis with a murine huntingtin specific probe. Similarly, the huntingtin protein levels increased in these cells and by 24 h they reached B4-fold increased levels (Figure 3b). These results demonstrated that p53 induces huntingtin gene expression in cultured murine cells, as p53 does in human cells. To directly test whether a similar regulation occurs in mouse brain tissues, both wild-type mice and the sex-

and age-matched isogenic p53 knockout mice were exposed to 6 Gy of g-irradiation. Various parts of the mouse brain tissues were dissected out and the effect of p53 activation on huntingtin gene expression was examined in cortex, cerebellum, and striatum tissues by real-time PCR. The p53 activation in these tissues upon g-irradiation was confirmed by measuring mRNA levels of the p21 gene, a known p53 downstream target gene. As shown in Figure 4b (left panel), p21 mRNA increased by B2-fold in cortex and cerebellum, and B4-fold in striatum tissues in wild-type mice. The p21 mRNA increase was clearly mediated by p53, since no increase in p21 mRNA levels were observed in p53 knockout mice (Figure 4b, right panel). The expression patterns of the huntingtin gene were determined in the same mice brain tissues. As shown in Figure 4a, huntingtin mRNA levels increased by B2.1-fold in wild-type mice cortex tissues, B2.7-fold in striatum 24 h after g-irradiation, while huntingtin mRNA levels changed little in tissues of the p53/ mice. The p53-dependent activation of huntingtin is similar in magnitude compared to that of the p21 gene in these tissues. However, the p53-dependent regulation of the huntingtin gene was not observed in cerebellum tissue (Figure 4a, left panel), which may be due to a tissuespecific difference in transcriptional regulation. Consistent with the increase of the huntingtin mRNA, the huntingtin protein levels increased by B2.4-fold in the cortex and B3.2-fold in striatum tissues of the wild-type mice 24 h postirradiation, as detected by Western blot analysis with an anti-huntingtin antibody (Figure 4c), whereas there were little change in huntingtin protein levels in the age- and sex-matched isogenic p53 knockout mice (Figure 4d). The p53-dependent upregulation of huntingtin gene by g-irradiation and its correlation with p21 transactivation in the striatum and cortex strongly suggest that p53 can regulate huntingtin gene transcription in these tissues.

Discussion

Figure 3 p53 activation increases huntingtin gene expression in cultured murine cells. Vm10 cells were grown to 50% confluence at 391C and transferred to a 321C incubator and cultured for indicated amount of time. Total RNA and proteins were extracted from each sample. Northern blot analysis (a) or Western blot analysis (b) were performed to determine the levels of huntingtin mRNA and protein. The mRNA levels of GAPDH and the protein levels of RAN were also determined as control. The data represent three independent experiments, and the error bar represents standard deviation. Oncogene

Hungtington’s disease is one of the most prevalent neurodegenerative diseases caused by expansion of polyglutamine tract in disease-associated huntingtin protein. Although the huntingtin protein was discovered over 10 years ago, its function, regulation, and the mechanisms in pathogenesis are still largely unknown. However, accumulation of cytoplasmic and nuclear aggregates of the mutant huntingtin protein in pathological sites of the brain is believed to play a central role in the pathology of Huntington’s disease (DiFiglia et al., 1997); and it has been suggested that both genetic and environmental stress factors could affect huntingtin gene expression, which in turn could impact upon the onset and prognosis of Huntington’s disease (Georgiou et al., 1999; Anca et al., 2004; Dixon et al., 2004). The tumor suppressor gene p53 encodes a transcription factor that represents a central integrator of various stress signals, such as cellular DNA damage, hypoxia,


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Figure 4 Huntingtin gene expression is upregulated by g-irradiation in a p53-dependent manner in mouse cortex and striatum. p53 þ / þ or p53/ mice were exposed to g-irradiation (IR, 6 Gy) and killed at 24 h after irradiation. Total RNA and proteins were extracted and from the cerebellum, cortex, and striatum. The levels of huntingtin mRNA and p21 mRNA were determined by quantitative real-time PCR and normalized against the levels of GAPDH. The levels of huntingtin protein were analyzed by Western blot assay and normalized against the levels of GAPDH. (a) huntingtin mRNA levels in p53 þ / þ and p53/ mice; (b) p21 mRNA levels in p53 þ / þ and p53/ mice. (c) and (d), Western blot analysis with anti-huntingtin antibody (left panels) and quantification of huntingtin protein levels (right panels) in p53 þ / þ and p53/ mice, respectively. The data represent three independent experiments, and the error bar represents standard deviation.

and oncogene activation (Levine, 1997; Vogelstein et al., 2000; Jin and Levine, 2001). In this study, we have identified multiple potential p53-REs in both mouse and human huntingtin genes. The interactions between human huntingtin p53 REs and p53 have been tested and they are capable of interacting with p53 protein both in vitro and in cells. In both cultured murine and human cells, p53 activation is capable of increasing huntingtin gene expression. Moreover, p53 mediates the increasing of huntingtin gene expression in mouse tissues after g-irradiation. This study revealed a novel regulation of the huntingtin gene expression at transcriptional level by the tumor suppressor p53. Although we only tested the expression regulation of the normal huntingtin gene by p53, it is expected that the mutant huntingtin gene is regulated in a similar way by p53 since the trinucleotide repeat expansion occurs in the coding region while not in the regulatory regions or p53 REs in the huntingtin gene. It is notable that p53

activation by DNA damage increases huntingtin expression in the striatum and cortex tissues of mouse, the two major pathological sites for Huntington’s disease. As an important signal integrator, p53 responds to a number of stress signals. It is expected that a variety of environmental factors that can alter p53 activity could then increase the mutant huntingtin protein expression levels. An accumulation of mutant huntingtin is deleterious, therefore the activation of p53 would lead to increased expression of mutant huntingtin, thus increase toxicity and alter the time of onset and the developmental course of this disease. Since mutant huntingtin can accumulate in the nucleus, where it may then bind to p53 (Steffan et al., 2000), it is not clear whether an altered response to p53 might be observed in neurons of Huntington’s disease. This will be of interest to determine and could be performed readily in the cell lines used for this study. In addition, experiments and epidemiological analyses should now be carried out to Oncogene


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determine the impact of p53 activity (DNA damage, hypoxia, chemical stresses, etc) upon the development of Huntington’s disease in both Huntington’s disease mouse models and in human patients.

Materials and methods Cells and cell culture The human lung epithelial H1299 cells (American Type Culture Collection, ATCC) were cultured in DMEM medium supplemented with 10% fetal bovine serum. H1299/V138 cells (a generous gift from Jiandong Chen at H Lee Moffitt Cancer Center, Tampa, FL, USA), established by stably transfecting a temperature-sensitive mutant form of p53 (alanine 138 to valine) into H1299 cells (Pochampally et al., 1999), were cultured in DMEM medium supplemented with 10% fetal bovine serum and 500 mg/ml G418. Human colon epithelial HCT116 p53 þ / þ , HCT116 p53/ cells (generous gifts from Bert Vogelstein at Johns Hopkins Medical Institutions, Baltimore, MD, USA) were cultured in McCoy’s 5A medium supplemented with 10% fetal bovine serum (Bunz et al., 1998). Vm10 cells, the murine fibroblast cells expressing a temperature-sensitive mutant p53 protein (alanine 135 to valine), were established in this lab as previously described (Wu and Levine, 1994), and were cultured in DMEM medium supplemented with 10% fetal bovine serum and 500 mg/ml G418. Mice and treatment C57BL/6 p53 knockout mice (4–6-week-old) (Lowe et al., 1993) and age- and sex-matched C57BL/6 wild-type controls (Jackson Laboratories) were subject to 6 Gy of total body irradiation with a 137Cs gamma source. Mice were killed at different time after irradiation and different tissues were harvested for further experiments. Chromatin immunoprecipitation (ChIP) assay ChIP assays were performed using Upstate ChIP Assay Kit (Lake Placid, NY, USA) according to the manufacturer’s instructions. In brief, cells were fixed with 1% formaldehyde for 10 min at room temperature. After washing with cold-PBS, cells were lysed in SDS lysis buffer, and sonicated to shear DNA to an average fragment size of 500 bp. Anti-p53 DO-1 antibody (SC-126, Santa Cruz, CA, USA) or normal mouse IgG was added. After overnight incubation at 41C, immune complexes were collected with salmon sperm DNA/protein A agarose-50% sulrry (Upstate) for 1 h, and then extensively washed. Samples were extracted with elution buffer (1% SDS, 0.1 M NaHCO3), and heated at 651C overnight to reverse crosslinks. DNA was purified and used for PCR. The primer sets were designed to encompass the potential p53 RE sites in the human huntingtin gene. The sequences are as follows: For promoter p53 RE site: 50 -GGGACTACAGGCATGCA CCACTAC-30 and 50 -AAAAATTAAGTTCCAGCGAGG TG-30 For intron 2 p53 RE site: 50 -GCCTGGGTGACTGAGCG AGAC-30 and 50 -CTTGTTCTCCGTGCCTGCTGAT-30 For intron 3 p53 RE site: 50 -GGAAACAGCGATGAGCA ATAAG-30 and 50 -AATAGCTGTTAATGGTTTTCAC-30

For Mdm2 promoter p53 RE site: 50 -GGTTGACTCAGC TTTTCCTCTTG-30 and 50 -GGAAAATGCATGGTTTAAA TAGCC-30 (Jin et al., 2002). Electromobility shift assays Bacterially expressed 6 His-p53 protein was purified by NiNTA-agarose chromatography. Sequences of the linear, blunt oligonucleotides used were: GADD45, 50 -GTAGCTGATATC GAATTCTCGAGCAGAACATGTCTAAGCATGCTGGGC TCGAGAATTCCTGCAGCC-30 ; Huntingtin intron 2, 50 -CT CGAGGCTGAGAGTTCATTATGCTTGTTCTACAGAA GAGCATGTTAAAAGGAGTTTTTGAGATCT-30 ; Huntingtin intron 3, 50 -CTCGAGCATGGTGCCATTTAGGGCCTGCT TCCAGTTAAGCTTGCTTCTCCACAGGCCTAAAGAT CT-30 ; mutant GADD45, 50 -GTAGCTGATATCGAATTCT CGAGCAGAAAATTTCTAAGAATTCTGGGCTCGAGA ATTCCTGCAGCG-30 . DNA probes were labeled using [g-32P]ATP and T4 polynucleotide kinase. Reaction mixtures (20 ml) contained 20 mM HEPES (pH 7.9), 25 mM KCl, 0.1 mM EDTA (pH 8.0), 2 mM MgCl2, 10% glycerol, 0.025% NP-40, 4 mM spermidine, 2 mM DTT, 0.2 mg BSA, 25 mutant GADD45 oligo, and 4 ng 32P-labeled oligonucleotide. After 30 min of incubation at room temperature, the reaction mixtures were subjected to electrophoresis on a 4% polyacrylamide gel (29:1 acrylamide/bisacrylamide) containing 0.5 Tris-borate-EDTA buffer at 165 V for 1.25 h at room temperature. The gel was dried and exposed to Kodax Biomax MR film. Northern and Western blot analysis Standard Northern and Western blot analysis were used to analyse RNA and protein expression, respectively. Anti-huntingtin (SC-8767), GAPDH (SC-20357) and RAN (SC-1156) antibodies were purchused from Santa Cruz Biotechnology (Santa Cruz, CA, USA). Anti-b-actin (A5441) was purchused from Sigma (Saint Louis, MO, USA). The RNA and protein levels were quantified by digitalization of the X-ray film and analysed with Scion Image software (Scion Corporation, Maryland). Quantitative real-time PCR Total RNA was prepared from cells or mouse brain tissues with the RNeasy kit (Qiagen, Valencia, CA, USA) and treated with the DNase I to remove residual genomic DNA. The cDNA was prepared with random primers using Taqman Reverse Transcription kit (Applied Biosystems, Foster city, CA, USA). Real-time PCR was performed in triplicate with Taqman PCR Mix (Applied Biosystems) for 15 min at 951C for initial denaturing, followed by 40 cycles of 951C for 30 s and 601C for 30 s in the 7000 ABI sequence Detection System. Assay-on-demand for human huntingtin (catalogue number Hs00169273_m1), human GAPDH (catalogue number Hs99999905_m1), mouse p21 (catalogue number Mm00432448_m1), mouse GAPDH (catalogue number Mm99999915_g1), mouse b-actin (catalogue number Mm00607939_s1) and assay-on-design for mouse huntingtin (HD-EXN3-211888), were purchased from Applied Biosystems. The expression of genes was normalized to expression of housekeeping gene, GAPDH and b-actin.

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