Human Molecular Genetics, 2002, Vol. 11, No. 19 2233-2241
© 2002 Oxford University Press
Identification of a presymptomatic molecular phenotype in Hdh CAG knock-in mice
Molecular Neurogenetics Unit, Massachusetts General Hospital, Building 149, 13th Street, Charlestown, MA 02129, USA
Received April 10, 2002; Accepted July 18, 2002
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSLimited variability of the...DISCUSSIONMATERIALS AND METHODSREFERENCES |
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The hallmark striatal neurodegeneration of Huntington's disease (HD) is first triggered by a dominant property of the expanded glutamine tract in mutant huntingtin that increases in severity with glutamine size. Indeed 111-glutamine murine huntingtin leads to a dominant cascade of phenotypes in HdhQ111 mice, although these abnormalities are not manifest in HdhQ50 mice, with 50-glutamine mutant protein. Therefore, to identify phenotypes that might reflect events closer to the fundamental trigger mechanism, and that can be measured as a consequence of adult-onset HD mutant huntingtin, we have screened for altered expression of genes conserved in evolution, which are likely to encode essential proteins. Probes generated from HdhQ111 homozygote and wild-type striatal RNAs were hybridized to human gene segments on filter arrays, disclosing a mutant-specific increase in hybridization to Rrs1, encoding a ribosomal protein. Subsequent, quantitative RT–PCR assays demonstrated increased Rrs1 mRNA from 3 weeks of age in homozygous and heterozygous HdhQ111 striatum and increased Rrs1 mRNA expression with a single copy's worth of 50-glutamine mutant huntingtin in HdhQ50 striatum. Moreover, quantitative RT–PCR assays for the human homologue demonstrated elevated Rrs1 mRNA in HD compared with control postmortem brain. These findings, therefore, support a chronic impact of mutant huntingtin on an essential ribosomal regulatory gene to be investigated for its role very early in HD pathogenesis.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSLimited variability of the...DISCUSSIONMATERIALS AND METHODSREFERENCES |
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Polyglutamine segments encoded by expanded HD CAG repeats endow mutant versions of huntingtin with a new property that leads to the loss of neurons in the striatum and to the chorea that characterizes Huntington's disease (HD) (1). Genetic studies in HD patients have shown that this novel trigger property increases in severity with increased polyglutamine size in the mutant protein and is dominant over the normal protein, while the context of huntingtin determines the special vulnerability of striatal neurons (2).
We have hypothesized that the expanded polyglutamine tract in mutant huntingtin may interact inappropriately with a cellular target that is essential to neuronal cells, triggering a cascade of secondary and tertiary consequences that broadens with time, as the cell becomes progressively sicker (2,3). Furthermore, consistent with mutant huntingtin expression at all ages, correlations with HD brain pathology suggest that the initial trigger mechanism may operate through the lifetime of the cell, perhaps from birth (4–6).
In vitro assays with N-terminal mutant huntingtin fragments have shown that long polyglutamine tracts promote the formation of an aberrant structure and aggregation in a manner that conforms to the HD genetic criteria, implicating this structural property in triggering the human disease (7,8). Indeed, expanded polyglutamine segments in mutant fragment can perturb CREB- (9–12) and SP1- (13) mediated gene transcription, disrupt glutamate uptake by synaptic vesicles (14) and sensitize cells to apoptotic insult (reviewed in 15). However, in HD patients, the polyglutamine tract may first trigger the disease cascade from the context of the full-length mutant protein.
Studies in precise genetic HD mouse models, Hdh CAG knock-in mice, expressing murine huntingtin with juvenile-onset HD glutamine tracts (
82–150 residues) have defined a temporal cascade of dominant phenotypes that exhibit striatal specificity and whose timing is hastened by increased CAG size (reviewed in 16). The earliest change is nuclear accumulation of full-length mutant huntingtin, implicating the 350 kDa protein in the HD trigger mechanism, followed by somatic CAG repeat instability (17,18), changes in glutamate receptor sensitivity (19), formation of insoluble N-terminal product and intranuclear inclusions (14,20), behavioral abnormalities, and neurodegenerative changes (21,22). However, while full-length nuclear mutant protein is detected in HdhQ111 and HdhQ92 striatal neurons at 6 weeks and 3.5 months of age, respectively, this abnormality is not found in HdhQ50 mice, expressing 50-glutamine mutant huntingtin (16,20). Thus, for mutant huntingtin bearing a polyglutamine tract that triggers most HD cases, the earliest phenotype in the cascade is not sufficiently early to be manifest within the mouse's short 3-year lifespan.
Therefore, to reveal phenotypes that may reflect disease events earlier than those currently known, we set out to identify genes involved in fundamental cellular processes that exhibit altered mRNA expression from a young age in HdhQ111 striatum. These candidates would then be placed in the temporal cascade by assessing HdhQ50 mice. To screen for changes in fundamental genes, we have used cross-hybridization of homozygous HdhQ111 and wild-type murine mRNAs to filter arrays of human gene segments, because the primary DNA sequence of genes that perform essential functions tends to be conserved in evolution. This strategy has revealed a dominant impact of the HD mutation on the expression of a conserved gene encoding a ribosomal protein, Rrs1, that is detected early in the disease cascade in Hdh CAG knock-in mice, and which is evident in postmortem HD brain. Thus, as Rrs1 is essential for normal ribosome biogenesis in yeast (23), our findings have disclosed an impact of mutant huntingtin on a fundamental cellular pathway that merits investigation for its role in the disease process in other HD model systems and in at-risk HD patient samples.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSLimited variability of the...DISCUSSIONMATERIALS AND METHODSREFERENCES |
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Identification of Rrs1 with Human GENEFILTERS
We sought to delineate early molecular consequences of the HD mutation on conserved and therefore fundamental genes. Thus, we used HdhQ111 homozygote and wild-type striatal mRNAs to probe filter arrays of human gene segments, in order to reveal changes in the expression of conserved genes at both 2.5 and 5.0 months of age. These ages are just after nuclear mutant huntingtin accumulation is detected, but >7 months before nuclear-inclusion formation and almost 2 years before neurodegeneration (22). To minimize potential animal-to-animal variability, pooled striata dissected from HdhQ111 homozygote mice or age-matched wild-type mice were processed to produce batches of mutant or wild-type striatal mRNA. These pools were then used to prepare [33P]dCTP labeled cDNA probes. For each age, the HdhQ111 homozygote probe and the wild-type probe were hybridized to filters bearing a total of 31 104 human cDNAs arrayed on Research Genetics Human GENEFILTERS, using stringent temperature and salt conditions. The data, collected by phosphoimager, were analyzed with Pathways software, using recommended criteria to reveal significant differences in the hybridization patterns of the HdhQ111 compared with the cognate wild-type probe.
These analyses revealed a single human cDNA, Rrs1 (GenBank accession no. NM_015169) that was differentially detected at both ages. Compared with the 2.5- and 5.0-month wild-type probes, the HdhQ111 homozygote probes yielded increased signal intensities of 5.5- and 3.8-fold, respectively. Rrs1, first studied in Saccharomyces cerevisiae (GenBank accession no. NP_014937), encodes a protein essential for ribosome biogenesis (23) that has been conserved through evolution, with homologues for example in Caenorhabditis elegans (GenBank accession no. NP_506573) and Drosophila melanogaster (GenBank accession no. AAM52679). Database searches (BLAST2) revealed that the human Rrs1 cDNA (1718 bp) and mouse Rrs1 cDNA (GenBank accession no. XM_123481) (2009 bp) encoded protein products of 394 and 364 amino acids, respectively, that exhibited 90% similarity. Moreover, these homologues were 88% identical at the nucleotide level.
Early dominantly increased Rrs1 mRNA in HdhQ111 striatum
To determine whether Rrs1 might be differentially expressed in HdhQ111 homozygote and HdhQ111 heterozygote striatum, we performed quantitative RT–PCR assays specific for Rrs1 and for a control mRNA, ß-actin, at ages ranging from 3 weeks to 5 months. The results, summarized in Figure 1A, revealed that Rrs1 mRNA was significantly increased in both heterozygous and homozygous mutant striatum compared with wild-type striatum. The increase was 1.4-fold at 3 weeks, 1.4-fold at 2.5 months and 1.6-fold at 5.0 months, indicating an early dominantly inherited molecular phenotype. Furthermore, assays of Rrs1 mRNA in homozygous HdhQ111 and wild-type striatum at 10.0 months revealed an 1.6-fold (P<0.001) increase in the mutant striatum, indicating an ongoing impact of mutant huntingtin on Rrs1 mRNA expression (data not shown).
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To assess the striatal specificity of the Rrs1 mRNA increase, we performed quantitative RT–PCR assays with mRNAs isolated from other brain regions of homozygous and heterozygous HdhQ111 mice. Cerebral cortex and cerebellum have been shown previously to exhibit nuclear mutant huntingtin at ages after this phenotype is detected in striatum, with a delay in heterozygotes compared with homozygotes (20). The results of the RT–PCR assays at 5 months, shown in Figure 1B, disclosed elevated Rrs1 mRNA in homozygous mutant cortex and cerebellum of 1.2- and 1.6-fold respectively, although no significant change was detected in tissue from HdhQ111 heterozygotes. Thus, in contrast to striatum, a single copy's worth of mutant huntingtin was not sufficient to yield increased Rrs1 mRNA in these brain regions at a young age, indicating that the underlying process exhibits striatal specificity.
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Limited variability of the Rrs1 phenotype in single HdhQ111 mice and age of onset |
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TOPABSTRACTINTRODUCTIONRESULTSLimited variability of the...DISCUSSIONMATERIALS AND METHODSREFERENCES |
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The RT–PCR results described above revealed increased Rrs1 mRNA in striatum with a single HdhQ111 allele carried on an outbred (CD1) genetic background. To determine whether the Rrs1 phenotype might be influenced by factors other than the mutant allele, we used the quantitative RT–PCR assay to assess striata from single wild-type and homozygous HdhQ111 mice at 5 months of age. Moreover, to more precisely assess the timing of onset of increased Rrs1 mRNA, we assayed at 3 weeks of age the individual striata of 22 mice: 14 littermates born to heterozygous HdhQ111 parents, 4 siblings born to wild-type parents and 4 littermates from homozygous HdhQ111 parents.
To provide a meaningful baseline for this study, we first examined the variability inherent in the Rrs1 mRNA assay. Within a comparative experiment, separate Rrs1 and control ß-actin RT–PCR reactions are each performed in triplicate, on the same RNA dilution using the same reaction mix, providing the ratio of Rrs1 to ß-actin mRNA, in assays that exhibit very little deviation. However, as shown in Figure 2A, the ratios of Rrs1 to ß-actin mRNA exhibited slight variation when determined in three experiments to compare HdhQ111 and wild-type striatal RNAs as performed on different iCycler machines by different investigators on three different occasions. The ratio for wild-type mRNA averaged 2.32±0.09, with each determination not significantly different from any other (P>0.1). Similarly, the ratios for mutant HdhQ111 mRNA averaged 3.47±0.13, and were not significantly different from each other (P>0.05). This limited variability may reflect differences in the efficiencies of reverse transcription and PCR amplification for the test mRNA or the ß-actin mRNA control, from experiment to experiment. Importantly, the biological effect of the HdhQ111 allele masks the relatively minor interexperimental variability, since any given wild-type Rrs1/ß-actin ratio, when compared with any mutant Rrs1/ß-actin ratio, was significantly different, yielding an 1.5-fold increase (P<0.0005). Thus, the ratio of Rrs1 to ß-actin mRNA provided a reliable measure of the Rrs1 mRNA fold change, although determination of Rrs1 mRNA level in different samples should be done within the same experiment.
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Figures 2B and C present the results of RT–PCR experiments to determine the variation in Rrs1 mRNA measured in striata of single mice at 5 months and at 3 weeks of age, respectively. The data revealed a tight distribution of Rrs1/ß-actin mRNA ratios, whose relative levels mirrored the Hdh CAG genotype. At 5 months, all of the wild-type striata were readily distinguished from HdhQ111 homozygotes. At 3 weeks of age, the lower range of HdhQ111 homozygotes overlapped with that of a single wild-type sample, and HdhQ111 heterozygotes were clustered at the low end of the mutant homozygote range. These data, therefore, indicated a relatively uniform onset at around 3 weeks of age as a consequence of even a single HdhQ111 allele, although the Rrs1 mRNA phenotype had progressed by 5 months, permitting mutant mice to be reliably distinguished from wild-type mice.
Rrs1 mRNA phenotype in HdhQ92 and HdhQ50 striatum
To determine whether increased Rrs1 mRNA was also a consequence of HD CAG repeat expansions carried in other lines of Hdh CAG knock-in mice, we performed quantitative RT–PCR assays to assess striata from homozygous and heterozygous HdhQ92 mice, with a 92-glutamine tract. As shown in Figure 3A, a significant increase compared with wild-type of 1.5-fold was observed, consistent with a molecular phenotype that reflects a dominant polyglutamine-dependent mechanism. Furthermore, given the early timing of onset at 3 weeks of age due to the HdhQ111 allele, we monitored Rrs1 mRNA in striata of heterozygous HdhQ50 mice, which express a single allele's worth of mutant huntingtin with 50 glutamines. The results of these RT–PCR analyses at 14 months of age are shown in Figure 3B. On average, Rrs1 mRNA was elevated 2-fold in HdhQ50 heterozygote striata compared with levels in wild-type striata. These data, therefore, demonstrated onset of the Rrs1 molecular phenotype in a precise genetic model of adult-onset HD, consistent with an early event in the disease process.
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Rrs1 mRNA is increased in HD postmortem brain
Fulfillment of the HD genetic criteria in Hdh CAG knock-in mice strongly implicated elevated Rrs1 mRNA in the human disease. To test this possibility, we performed quantitative RT–PCR assays for the human Rrs1 homologue, measuring Rrs1 mRNA in parietal cortex dissected from age-matched control and HD postmortem brains (Vonsattel grade 3 neuropathology) (24). For the latter, the expanded HD CAG repeats on the disease chromosome encoded polyglutamine tracts in mutant huntingtin that ranged from 43 to 45 residues. The results shown in the scattergraph in Figure 4 revealed an increase of Rrs1 mRNA in four of the five HD samples, while the level of Rrs1 mRNA for the fifth overlapped with the high end of the control sample range. On average, Rrs1 mRNA was significantly increased
2.4-fold in HD compared with control brains (P<0.0005), demonstrating that expression of this conserved gene is increased as a consequence of mutant huntingtin in the human disease.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSLimited variability of the...DISCUSSIONMATERIALS AND METHODSREFERENCES |
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We have postulated that the novel property conferred on mutant huntingtin by the expanded polyglutamine tract may alter a fundamental cellular process, triggering a disease cascade that ultimately leads to the demise of striatal neurons. To investigate this hypothesis, we have been studying Hdh CAG knock-in mice to identify abnormalities that stem from processes with the genetic attributes of the HD trigger mechanism: striatal specificity, dominant inheritance, and earlier onset with increased expanded polyglutamine length. We and others have previously demonstrated a temporal cascade that is evident with long juvenile-onset HD glutamine tracts. This cascade first features accumulation of full-length mutant huntingtin in the nucleus of striatal neurons that precedes neurodegenerative changes by
1–2 years. However, mutant huntingtin with 50 glutamines does not produce these phenotypes in HdhQ50 striatum, implying that the underlying events are not sufficiently early in the disease cascade to produce effects within the mouse's short 3-year lifespan. By seeking changes in fundamental processes orchestrated by genes conserved through evolution, we have now identified a new pathway to be investigated in Hdh CAG knock-in mice, in other HD model systems and in HD patients. Our data demonstrate an impact of mutant huntingtin on the expression of Rrs1 mRNA, encoding a ribosomal regulatory protein, which is likely to be very early in the disease cascade. This conclusion is supported by several findings. Firstly, increased Rrs1 mRNA was correlated with dominant inheritance of mutant huntingtin with juvenile-onset HD glutamine tracts, and was manifest at an age that precedes all other reported phenotypes. Secondly, elevated Rrs1 mRNA was persistent, slightly worsening with age, consistent with a chronic phenotype that reflects a subtle ongoing disease process. Thirdly, increased Rrs1 mRNA was correlated with dominant inheritance of a single allele's worth of mutant huntingtin bearing a polyglutamine tract of 50 residues. Moreover, the increase was detected in HdhQ50 striatum by 14 months of age, providing the first abnormal phenotype in an accurate genetic model of adult-onset HD and implying an event that may be evident in very young presymptomatic HD individuals. Fourthly, as suggested by fulfillment of the HD genetic criteria in Hdh knock-in mice, Rrs1 mRNA was found to be increased in HD postmortem brain, directly implicating the underlying process in the human disorder.
Rrs1 protein has been shown in yeast to be required for the coordinated regulation of ribosomal protein and rRNA synthesis in response to varying physiologic conditions (23,25,26). Thus, our data provide strong support for investigating the impact of mutant huntingtin on ribosome function as a very early consequence of the disease process in Hdh knock-in mice and in the human disorder. Indeed, previous reports are consistent with altered ribosome function in HD. Wyttenbach et al. (12) noted an impact of overexpressed mutant huntingtin fragment on genes encoding ribosomal proteins in an inducible cell system. Moreover, Iqbal et al. (27), in a 1974 study, noted ribosomal protein abnormalities in subcellular fractions of neuronal cells prepared from cerebral cortex of HD postmortem brain.
While indicating that increased Rrs1 mRNA is likely to reflect an early disease event, our study does not address whether this phenotype is a side-effect of the disease mechanism or whether altered ribosome function may be a step on the main pathway that leads to neurodegeneration. It is noteworthy that normal huntingtin is a nuclear and a cytoplasmic protein (28–32) that has been implicated in RNA biogenesis. Its deficiency effects nucleolar morphology and perinuclear membrane function (33), and normal and mutant huntingtin bind proteins involved in transcription and mRNA processing (10,13,32,34–36). It will be important, therefore, to determine whether in the course of its intrinsic huntingtin activities, the mutant protein may directly impact Rrs1 expression, via an effect on transcription or mRNA stability. It will also be important to test whether increased Rrs1 product or altered ribosome function amplifies the disease cascade, yielding phenotypes immediate to neuronal cell death that have been implicated by studies in HD model systems and HD patients. These include alterations in CREB (10–12,37) and SP1 (13) transcriptional targets, decreased levels of neuronal pro-survival proteins (38–42), altered neurotransmitter receptors (9,19,43), abnormal mitochondrial function (44,45) and direct effects on apoptotic pathways (46–49).
The ability to assay for increased Rrs1 mRNA also provides a unique opportunity to test whether an early molecular HD phenotype occurs in other inherited polyglutamine diseases that target different neuronal cells. Altered Rrs1 mRNA was not reported in a study of mRNA changes in a transgenic mouse model of spinocerebellar ataxia 1 (50). However, the phenotype has not been directly monitored in this transgenic SCA1 model (51) or in a Sca1 knock-in model (52). In addition, mouse models are now being characterized for Machado–Joseph disease (53), spinocerebellar ataxia 7 (54), dentatorubro and pallidoluysian atrophy (55), and Kennedy disease (56,57). The results may reveal whether expanded polyglutamine tracts in different protein contexts perturb the pathway that gives rise to the Rrs1 mRNA phenotype, perhaps providing evidence for a common trigger mechanism as suggested by genetic data in the human polyglutamine disorders (2).
Finally, our findings in Hdh CAG knock-in mice suggest monitoring Rrs1 mRNA levels as a quantitative measurement for a molecular disease phenotype that might be detectable in the peripheral tissues of individuals which carry the HD mutation but who are not yet symptomatic. In a preliminary experiment, we have noted increased Rrs1 mRNA in liver of the single homozygous HdhQ111 mouse tested (data not shown), suggesting that additional experiments with homozygous and heterozygous mutant mice may provide evidence for the phenotype in the periphery. An outcome that indicated altered Rrs1 expression in peripheral tissue would provide the impetus to develop and test a reliable biomarker for the early disease process, which could be important for monitoring the effects of rational therapeutics in presymptomatic and symptomatic HD individuals.
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MATERIALS AND METHODS |
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Wild-type and Hdh knock-in mice
HdhQ92 and HdhQ111 knock-in mice with 90 and 109 CAG repeats, respectively, inserted at the appropriate location in the mouse HD gene, Hdh, have been described previously (18,20), and have been maintained on an outbred CD1 genetic background. HdhQ50 mice, with 48 CAG repeats (58), initially CD1 background, were backcrossed for 10 generations to C57BL/6J, giving a mixed CD1/C57BL/6J background. Genotyping was performed by Southern blot analyses and PCR assay for CAG repeat size as described in previous reports (18,20).
Postmortem HD brain tissue
Control and HD postmortem brain tissue was collected by the Harvard Brain Tissue Resource Center, McLean Hospital (Belmont, MA) and used pursuant to the guidelines of the MGH IRB for discarded human tissue. Sections at the level of the caudate and putamen were evaluated at the Harvard Brain Tissue Resource Center for HD neuropathology by microscopic examination, and all of the HD brains used in this study were assigned Vonsattel grade 3 pathology (24). Postmortem intervals were from 12 to 32 h for control and from 8 to 30 h for HD brains. Control brains were from three males (ages 72, 74 and 79 years) and one female (age 76 years). HD brains were from five males (ages 64, 70, 70, 71 and 75 years). Death for controls was myocardial infarct and for HD end-stage disease. The presence of expanded and normal CAG alleles was assessed using PCR as previously described (59). Control brain exhibited HD allele sizes of 25/17, 20/18, 18/18 and 21/21. HD brain allele sizes were 41/18, 41/37, 42/23, 43/17 and 43/17.
RNA extraction
Mouse brains were dissected into striatum, cortex and cerebellum, immediately frozen in dry ice, and stored at -70°C. Small blocks of parietal cortex were dissected from frozen coronal slices of the control and HD postmortem brains. Total RNA was isolated with TRIZOL reagent (Invitrogen/Life Technologies, Carlsbad, CA) according to manufacturer's instructions, quantified spectrophotometrically at 260 nm, and analyzed by agarose gel electrophoresis. Total RNA used for quantitative RT–PCR assays was pretreated with DNase I using a DNA-free kit (Ambion Inc., Austin, TX). For microarray analyses, a single pool of total RNA was generated from RNA isolated from the striata of 100 HdhQ111/HdhQ111 homozygotes and 100 wild-type mice at 2.5 and 5.0 months of age.
Human GENEFILTERS microarray hybridization and analyses
Human GENEFILTERS microarrays (Research Genetics Inc., Huntsville, AL) used in the present study were GF200 Release I, GF201 Release II, GF202 Release III, GF203 Release IV, GF204 Release V, and GF211 Known genes, Release I. Each filter contains the 3'-UTR (1 kb in length) of 5184 cDNAs from the IMAGE/LLNL collection. Generation of labeled probe, filter pre-hybridization, filter hybridization and washes were performed according to Research Genetics protocols. Briefly, probe (750 ng total RNA), was labeled with [33P]dCTP using Superscript II (Invitrogen/Life Technologies, Carlsbad, CA) for 90 min at 37°C, and purified by passage through a Bio-Spin column (Bio-Rad Laboratories, Hercules, CA). GENEFILTERS membranes were prehybridized for 2 h at 42°C in MicroHyb solution (Research Genetics Inc., Huntsville, AL) with 5 µg of Cot-1 (Invitrogen/Life Technologies, Carlsbad, CA) and 5 µg of Poly dA (Research Genetics Inc., Huntsville, AL). Hybridization was overnight at 42°C in MicroHyb solution (Research Genetics Inc., Huntsville, AL). Filters were washed twice at 50°C in 2x SSC, 1% SDS for 20 min and once at room temperature in 0.5x SSC, 1% SDS for 15 min. After hybridization, filters were stripped in heated 0.5% SDS solution, as recommended by the manufacturer, and reprobed.
Exposed phosphoimaging screens were acquired through a Cyclone Phosphor system (Packard Instrument Co., Meriden, CT). Images were imported into the database of Pathways 2.01 software (Research Genetics Inc., Huntsville, AL), aligned using the 16 control points, and analyzed according to the internal software protocol. Normalized images of each hybridization were compared with each other according to the software protocol. The change in expression level for each gene was calculated as the ratio between the normalized intensity values. To minimize the number of false positives, we sorted the genes whose intensity values were 50-fold higher than the background intensity. Those that met this criterion were sorted by ‘fold change’ of mutant compared with wild-type. mRNA changes that were 
3.0, in either direction, were judged to be significant.
TaqMan real-time quantitative RT–PCR
Quantitative RT–PCR reactions were performed with an iCycler iQ instrument (Bio-Rad Laboratories, Hercules, CA), using the 5'-nuclease TaqMan assay. These RT–PCR reactions were performed in 50 µl reaction volumes containing 200 ng of DNase-treated total RNA, 200 nM of each primer and probe, 2x reaction mix, and 1 µl of Superscript II RT/Platinum Taq mix (Superscript One-Step RT–PCR with PlatinumR Taq, Invitrogen). Each reaction was performed in triplicate. Cycle parameters were 15 min at 50°C and 5 min at 95°C (30 s at 95°C, 1 min at the Tm specific for each primer set) for 45 cycles.
To quantify the mRNA levels for each sample and primer set, a standard curve was generated with known dilutions of total mouse brain RNA (Clontech, Palo Alto, CA). The relative value for each unknown sample was then calculated from its respective standard curve using linear regression analysis. To normalize the differences in the amount of total RNA added to each reaction, we used ß-actin as endogenous control. The normalized expression value was then calculated by dividing the relative quantitation value of each sample and primer set by the relative quantitation value of ß-actin. Primers and probe were designed for each gene using Primer Express V1.5 software (ABI, Foster City, CA) and selected to contain minimal internal structure. Probes were selected to have a Tm 10°C higher than the matching primers. All probes were dual-labeled with 5'-FAM fluorophore and BHQ-1-3' quencher. Specific forward and reverse primers (Invitrogen/Life Technologies, Carlsbad, CA) and probe sets (Biosearch Technologies, Novato, CA) were as follows: murine Rrs1 (GenBank accession no. XM_123481), 5'-GCA AGT ATT GTA AAT GGA TGC AG-3', 5'-TCT CAA GCA CAC TCC ATA TTG-3', 5'-FAM-CGG TAG AGG TTG CAT GTG AAG CCA GT-BHQ-1-3'; murine ß-actin (GenBank accession no. NM_007393), 5'-AGA GGG AAA TCG TGC GTG AC-3', 5'-CAA TAG TGA TGA CCT GGC CGT-3', 5'-FAM-CAC TGC CGC ATC CTC TTC CTC CC-BHQ-1-3'; human Rrs1 (GenBank accession no. NM_015169), 5'-CGT GGT TAT GCT GCC G-3', 5'-CTC GGC TCA CTC GCT T-3', 5'-FAM-AGT TTG GGC CGC CAC TGT AGG AA-BHQ-1-3'; human ß-actin (GenBank accession no. NM_001101), 5'-CTG GAA CGG TGA AGG TGA-3', 5'-TCA AAG TCC TCG GCC ACA-3'; 5'-FAM-AGC AGT CGG TTG GAG CGA GCA TCC-BHQ-1-3'.
Experiments in Figure 1A were with RNA isolated from pooled striata dissected from mice as follows: at 3 weeks of age, 7 HdhQ111 heterozygotes, 6 HdhQ111 homozygotes and 9 wild-type mice; at 2.5 and 5.0 months of age, 5 HdhQ111 heterozygotes, 100 HdhQ111 homozygotes and 100 wild-type mice. Experiments in Figure 1B at 5 months of age were with pooled tissue (cortex and cerebellum) from 25 HdhQ111 homozygotes, 5 HdhQ111 heterozygotes and 25 wild-type mice. Experiments in Figure 2A at 5 months were with RNA generated from pooled striata from 100 HdhQ111 homozygotes and 100 wild-type mice. Experiments in Figure 3A at 8.5 months were with RNA isolated from pooled striata from 6 HdhQ92 mice (2 homozygotes and 4 heterozygotes) and 2 wild-type mice. Experiments in Figure 3B at 14 months were with RNA prepared from pooled striata dissected from 5 HdhQ50 heterzygotes and 7 wild-type mice.
Statistical analyses
Statistical analyses were Student's t-test, choosing P<0.05 as significant. Weighted means were calculated for data produced in replica experiments. Calculated means and standard deviation were plotted using the graph tool of Microsoft Excel.
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ACKNOWLEDGEMENTS |
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We are grateful to Drs Weining Lu and David Beier (The Collis Genome Laboratory, Brigham and Women's Hospital) for assistance with GENEFILTER hybridization and analyses. This work was supported by NINDS Grants NS16367 (HD Center Without Walls), NS32765, an anonymous donor, the Hereditary Disease Foundation and the Huntington's Disease Society of America (Coalition for the Cure).
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FOOTNOTES |
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* To whom correspondence should be addressed. Tel: +1 6177265726; Fax: +1 6177265736; Email: persiche{at}helix.mgh.harvard.edu
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REFERENCES |
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1 Huntington's Disease Collaborative Research Group (1993) A novel gene containing a trinucleotide repeat that is expanded and unstable on Huntington's disease chromosomes. Cell, 72, 971–983.[Web of Science][Medline]
2 Gusella, J.F. and MacDonald, M.E. (2000) Molecular genetics: unmasking polyglutamine triggers in neurodegenerative disease. Nat. Rev. Neurosci., 1, 109–115.[Web of Science][Medline]
3 Gusella, J. and MacDonald, M. (2002) No post-genetics era in human disease research. Nat. Rev. Genet., 3, 72–79.[Web of Science][Medline]
4 Myers, R.H., Vonsattel, J.P., Paskevich, P.A., Kiely, D.K., Stevens, T.J., Cupples, L.A., Richardson, E.P., Jr, and Bird, E.D. (1991) Decreased neuronal and increased oligodendroglial densities in Huntington's disease caudate nucleus. J. Neuropathol. Exp. Neurol., 50, 729–742.[Web of Science][Medline]
5 Furtado, S., Suchowersky, O., Rewcastle, B., Graham, L., Klimek, M.L. and Garber, A. (1996) Relationship between trinucleotide repeats and neuropathological changes in Huntington's disease. Ann. Neurol., 39, 132–136.[Web of Science][Medline]
6 Penney, J.B., Jr., Vonsattel, J.P., MacDonald, M.E., Gusella, J.F. and Myers, R.H. (1997) CAG repeat number governs the development rate of pathology in Huntington's disease. Ann. Neurol., 41, 689–692.[Web of Science][Medline]
7 Huang, C.C., Faber, P.W., Persichetti, F., Mittal, V., Vonsattel, J.P., MacDonald, M.E. and Gusella, J.F. (1998) Amyloid formation by mutant huntingtin: threshold, progressivity and recruitment of normal polyglutamine proteins. Somat. Cell Mol. Genet., 24, 217–233.[Web of Science][Medline]
8 Wanker, E.E. (2000) Protein aggregation and pathogenesis of Huntington's disease: mechanisms and correlations. Biol Chem., 381, 937–942.[Web of Science][Medline]
9
Luthi-Carter, R., Strand, A., Peters, N.L., Solano, S.M., Hollingsworth, Z.R., Menon, A.S., Frey, A.S., Spektor, B.S., Penney, E.B., Schilling, G. et al. (2000) Decreased expression of striatal signaling genes in a mouse model of Huntington's disease. Hum. Mol. Genet., 9, 1259–1271.
10
Steffan, J.S., Kazantsev, A., Spasic-Boskovic, O., Greenwald, M., Zhu, Y.Z., Gohler, H., Wanker, E.E., Bates, G.P., Housman, D.E. and Thompson, L.M. (2000) The Huntington's disease protein interacts with p53 and CREB-binding protein and represses transcription. Proc. Natl Acad. Sci. USA, 97, 6763–6768.
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Nucifora, F.C., Jr, Sasaki, M., Peters, M.F., Huang, H., Cooper, J.K., Yamada, M., Takahashi, H., Tsuji, S., Troncoso, J., Dawson, V.L. et al. (2001) Interference by huntingtin and atrophin-1 with cbp-mediated transcription leading to cellular toxicity. Science, 291, 2423–2428.
12
Wyttenbach, A., Swartz, J., Kita, H., Thykjaer, T., Carmichael, J., Bradley, J., Brown, R., Maxwell, M., Schapira, A., Orntoft, T.F. et al. (2001) Polyglutamine expansions cause decreased CRE-mediated transcription and early gene expression changes prior to cell death in an inducible cell model of Huntington's disease. Hum. Mol. Genet., 10, 1829–1845.
13
Li, S.H., Cheng, A.L., Zhou, H., Lam, S., Rao, M., Li, H. and Li, X.J. (2002) Interaction of Huntington disease protein with transcriptional activator Sp1. Mol. Cell. Biol., 22, 1277–1287.
14 Li, H., Li, S.H., Johnston, H., Shelbourne, P.F. and Li, X.J. (2000) Amino-terminal fragments of mutant huntingtin show selective accumulation in striatal neurons and synaptic toxicity. Nat. Genet., 25, 385–389.[Web of Science][Medline]
15 Evert, B.O., Wullner, U. and Klockgether, T. (2000) Cell death in polyglutamine diseases. Cell. Tissue Res., 301, 189–204.[Web of Science][Medline]
16 MacDonald, M.E. and Wheeler, V.C. (2000) Huntington's disease pathogenesis: Insights from HD mouse models. NeuroScience News, 3, 38–44.
17
Shelbourne, P.F., Killeen, N., Hevner, R.F., Johnston, H.M., Tecott, L., Lewandoski, M., Ennis, M., Ramirez, L., Li, Z., Iannicola, C. et al. (1999) A Huntington's disease CAG expansion at the murine Hdh locus is unstable and associated with behavioural abnormalities in mice. Hum. Mol. Genet., 8, 763–774.
18
Wheeler, V.C., Auerbach, W., White, J.K., Srinidhi. J., Auerbach, A., Ryan, A., Duyao, M.P., Vrbanac, V., Weaver, M., Gusella, J.F. et al. (1999) Length-dependent gametic CAG repeat instability in the Huntington's disease knock-in mouse. Hum. Mol. Genet., 8, 115–122.
19 Levine, M.S., Klapstein, G.J., Koppel, A., Gruen, E., Cepeda, C., Vargas, M.E., Jokel, E.S., Carpenter, E.M., Zanjani, H., Hurst, R.S. et al. (1999) Enhanced sensitivity to N-methyl-D-aspartate receptor activation in transgenic and knockin mouse models of Huntington's disease. J. Neurosci. Res., 58, 515–532.[Web of Science][Medline]
20
Wheeler, V.C., White, J.K., Gutekunst, C.A., Vrbanac, V., Weaver, M., Li, X.J., Li, S.H., Yi, H., Vonsattel, J.P., Gusella, J.F. et al. (2000) Long glutamine tracts cause nuclear localization of a novel form of huntingtin in medium spiny striatal neurons in HdhQ92 and HdhQ111 knock-in mice. Hum. Mol. Genet., 9, 503–513.
21
Lin, C.H., Tallaksen-Greene, S., Chien, W.M., Cearley, J.A., Jackson, W.S., Crouse, A.B., Ren, S., Li, X.J., Albin, R.L. and Detloff, P.J. (2001) Neurological abnormalities in a knock-in mouse model of Huntington's disease. Hum. Mol. Genet., 10, 137–144.
22
Wheeler, V.C., Gutekunst, C.A., Vrbanac, V., Lebel, L.A., Schilling, G., Hersch, S., Friedlander, R.M., Gusella, J.F., Vonsattel, J.P., Borchelt, D.R. and MacDonald, M.E. (2002) Early phenotypes that presage late-onset neurodegenerative disease allow testing of modifiers in Hdh CAG knock-in mice. Hum. Mol. Genet., 11, 633–640.
23
Tsuno, A., Miyoshi, K., Tsujii, R., Miyakawa, T. and Mizuta, K. (2000) RRS1, a conserved essential gene, encodes a novel regulatory protein required for ribosome biogenesis in Saccharomyces cerevisiae. Mol. Cell. Biol., 20, 2066–2074.
24 Vonsattel, J.P., Myers, R.H., Stevens, T.J., Ferrante, R.J., Bird, E.D. and Richardson, E.P., Jr. (1985) Neuropathological classification of Huntington's disease. J. Neuropathol. Exp. Neurol., 44, 559–577.[Web of Science][Medline]
25
Miyoshi, K., Tsujii, R., Yoshida, H., Maki, Y., Wada, A., Matsui, Y., Toh, E.A. and Mizuta, K. (2002) Normal assembly of 60S ribosomal subunits is required for the signaling in response to a secretory defect in Saccharomyces cerevisiae. J. Biol. Chem., 277, 18334–18339.
26 Tsujii, R., Miyoshi, K., Tsuno, A., Matsui, Y., Toh-e, A., Miyakawa, T. and Mizuta, K. (2000) Ebp2p, yeast homologue of a human protein that interacts with Epstein–Barr virus nuclear antigen 1, is required for pre-rRNA processing and ribosomal subunit assembly. Genes Cells, 5, 543–553.[Abstract]
27 Iqbal, K., Tellez-Nagel, I. and Grundke-Iqbal, I. (1974) Protein abnormalities in Huntington's chorea. Brain Res., 76, 178–184.[Web of Science][Medline]
28
Hoogeveen, A.T., Willemsen, R., Meyer, N., de Rooij, K.E., Roos, R.A., van Ommen, G.J. and Galjaard, H. (1993) Characterization and localization of the Huntington disease gene product. Hum. Mol. Genet., 2, 2069–2073.
29
De Rooij, K.E., Dorsman, J.C., Smoor, M.A., Den Dunnen, J.T. and Van Ommen, G.J. (1996) Subcellular localization of the Huntington's disease gene product in cell lines by immunofluorescence and biochemical subcellular fractionation. Hum. Mol. Genet., 5, 1093–1099.
30 Wilkinson, F.L., Nguyen, T.M., Manilal, S.B., Thomas, P., Neal, J.W., Harper, P.S., Jones, A.L. and Morris, G.E. (1999) Localization of rabbit huntingtin using a new panel of monoclonal antibodies. Brain Res. Mol. Brain Res., 69, 10–20.[Medline]
31
Trettel, F., Rigamonti, D., Hilditch-Maguire, P., Wheeler, V.C., Sharp, A.H., Persichetti, F., Cattaneo, E. and MacDonald, M.E. (2000) Dominant phenotypes produced by the HD mutation in STHdh(Q111) striatal cells. Hum. Mol. Genet., 9, 2799–2809.
32
Kegel, K.B., Meloni, A.R., Yi, Y., Kim, Y.J., Doyle, E., Cuiffo, B.G., Sapp, E., Wang, Y., Qin, Z.H., Chen, J.D. et al. (2000) Huntingtin is present in the nucleus, interacts with the transcriptional corepressor C-terminal binding protein, and represses transcription. J. Biol. Chem., 277, 7466–7476.
33
Hilditch-Maguire, P., Trettel, F., Passani, L.A., Auerbach, A., Persichetti, F. and MacDonald, M.E. (2000) Huntingtin: an iron-regulated protein essential for normal nuclear and perinuclear organelles. Hum. Mol. Genet., 9, 2789–2797.
34
Faber, P.W., Barnes, G.T., Srinidhi, J., Chen, J., Gusella, J.F. and MacDonald, M.E. (1998) Huntingtin interacts with a family of WW domain proteins. Hum. Mol. Genet., 7, 1463–1474.
35
Boutell, J.M., Thomas. P., Neal, J.W., Weston, V.J., Duce, J., Harper, P.S. and Jones, A.L. (1999) Aberrant interactions of transcriptional repressor proteins with the Huntington's disease gene product, huntingtin. Hum. Mol. Genet., 8, 1647–1655.
36 Rega, S., Stiewe, T., Chang, D.I., Pollmeier, B., Esche, H., Bardenheuer, W., Marquitan, G. and Putzer, B.M. (2001) Identification of the full-length huntingtin-interacting protein p231HBP/HYPB as a DNA-binding factor. Mol. Cell. Neurosci., 18, 68–79.[Web of Science][Medline]
37
McCampbell, A., Taylor, J.P., Taye, A.A. Robitschek, J., Li, M., Walcott, J., Merry, D., Chai, Y., Paulson, H., Sobue, G. and Fischbeck, K.H. (2000) CREB-binding protein sequestration by expanded polyglutamine. Hum. Mol. Genet., 9, 2197–2202.
38 Dragatsis, I., Levine, M.S. and Zeitlin, S. (2000) Inactivation of Hdh in the brain and testis results in progressive neurodegeneration and sterility in mice. Nat. Genet., 26, 300–306.[Web of Science][Medline]
39
Zuccato, C., Ciammola, A., Rigamonti, D., Leavitt, B.R., Goffredo, D., Conti, L., MacDonald, M.E., Friedlander, R.M., Silani, V., Hayden, M.R. et al. (2001) Loss of huntingtin-mediated BDNF gene transcription in Huntington's disease. Science, 293, 493–498.
40 Cattaneo, E., Rigamonti, D., Goffredo, D., Zuccato, C., Squitieri, F. and Sipione, S. (2001) Loss of normal huntingtin function: new developments in Huntington's disease research. Trends Neurosci., 24, 182–188.[Web of Science][Medline]
41 Leavitt, B.R., Guttman, J.A., Hodgson, J.G., Kimel, G.H., Singaraja, R., Vogl, A.W. and Hayden, M.R. (2001) Wild-type huntingtin reduces the cellular toxicity of mutant huntingtin in vivo. Am. J. Hum. Genet., 68, 313–324.[Web of Science][Medline]
42
Song, C., Perides, G. and Liu, Y.F. (2002) Expression of full-length polyglutamine-expanded Huntingtin disrupts growth factor receptor signaling in rat pheochromocytoma (PC12) cells. J. Biol. Chem., 277, 6703–6707.
43 Cha, J.H., Frey, A.S., Alsdorf, S.A., Kerner, J.A., Kosinski, C.M., Mangiarini, L., Penney, J.B., Jr, Davies, S.W., Bates, G.P. and Young, A.B. (1999) Altered neurotransmitter receptor expression in transgenic mouse models of Huntington's disease. Phil. Trans. R. Soc. Lond. B Biol. Sci., 354, 981–989.[Web of Science][Medline]
44 Browne, S.E., Bowling, A.C., MacGarvey, U., Baik, M.J. Berger, S.C., Muqit, M.M., Bird, E.D. and Beal, M.F. (1997) Oxidative damage and metabolic dysfunction in Huntington's disease: selective vulnerability of the basal ganglia. Ann. Neurol., 41, 646–653.[Web of Science][Medline]
45 Panov, A.V., Gutekunst, C.A., Leavitt, B.R., Hayden, M.R., Burke, J.R., Strittmatter, W.J. and Greenamyre, J.T. (2002) Early mitochondrial calcium defects in Huntington's disease are a direct effect of polyglutamines. Nat. Neurosci., 5, 731–736.[Web of Science][Medline]
46
Wellington, C.L., Singaraja, R., Ellerby, L., Savill, J., Roy, S., Leavitt, B., Cattaneo, E., Hackam, A., Sharp, A., Thornberry, N. et al. (2000) Inhibiting caspase cleavage of huntingtin reduces toxicity and aggregate formation in neuronal and nonneuronal cells. J. Biol. Chem., 275, 19831–19838.
47 Ona, V.O., Li, M., Vonsattel, J.P., Andrews, L.J., Khan, S.Q., Chung, W.M., Frey, A.S., Menon, A.S., Li, X.J., Stieg, P.E. et al. (1999) Inhibition of caspase-1 slows disease progression in a mouse model of Huntington's disease. Nature. 399, 263–267.[Medline]
48
Rigamonti, D., Sipione, S., Goffredo, D., Zuccato, C., Fossale, E. and Cattaneo, E. (2001) Huntingtin's neuroprotective activity occurs via inhibition of procaspase-9 processing. J. Biol. Chem., 276, 14545–14548.
49 Humbert, S., Bryson, E.A., Cordelieres, F.P., Connors, N.C., Datta, S.R., Finkbeiner, S., Greenberg, M.E. and Saudou, F. (2002) The IGF-1/Akt pathway is neuroprotective in Huntington's disease and involves huntingtin phosphorylation by Akt. Dev. Cell, 2, 831–837.[Web of Science][Medline]
50 Lin, X., Antalffy, B., Kang, D., Orr, H.T. and Zoghbi, H.Y. (2000) Polyglutamine expansion down-regulates specific neuronal genes before pathologic changes in SCA1, Nat. Neurosci., 3, 157–163.[Web of Science][Medline]
51 Burright, E.N., Clark, H.B., Servadio, A., Matilla, T., Feddersen, R.M., Yunis, W.S., Duvick, L.A., Zoghbi, H.Y. and Orr, H.T. (1995) SCA1 transgenic mice: a model for neurodegeneration caused by an expanded CAG trinucleotide repeat. Cell, 82, 937–948.[Web of Science][Medline]
52 Watase, K., Weeber, E.J., Xu, B., Antalffy, B., Yuva-Paylor, L., Hashimoto, K., Kano, M., Atkinson, R., Sun, Y., Armstrong, D.L. et al. (2002) A long CAG repeat in the mouse Sca1 locus replicates SCA1 features and reveals the impact of protein solubility on selective neurodegeneration. Neuron. 34, 905–919.[Web of Science][Medline]
53 Ikeda, H., Yamaguchi, M., Sugai, S., Aze, Y., Narumiya, S. and Kakizuka, A. (1996) Expanded polyglutamine in the Machado–Joseph disease protein induces cell death in vitro and in vivo. Nat. Genet., 13, 196–202.[Web of Science][Medline]
54
Garden, G.A., Libby, R.T., Fu, Y.H., Kinoshita, Y., Huang, J., Possin, D.E. Smith, A.C., Martinez, R.A., Fine, G.C., Grote, S.K. et al. (2002) Polyglutamine-expanded ataxin-7 promotes non-cell-autonomous purkinje cell degeneration and displays proteolytic cleavage in ataxic transgenic mice. J. Neurosci., 22, 4897–4905.
55 Schilling, G., Wood, J.D., Duan, K., Slunt, H.H., Gonzalez, V., Yamada, M., Cooper, J.K., Margolis, R.L., Jenkins, N.A., Copeland, N.G. et al. (1999) Nuclear accumulation of truncated atrophin 1 fragments in a transgenic mouse model of DRPLA. Neuron, 24, 275–286.[Web of Science][Medline]
56 Bingham, P.M., Scott, M.O., Wang, S., McPhaul, M.J., Wilson, E.M., Garbern, J.Y., Merry, D.E. and Fischbeck, K.H. (1995) Stability of an expanded trinucleotide repeat in the androgen receptor gene in transgenic mic. Nat. Genet., 9, 191–196.[Web of Science][Medline]
57
La Spada, A.R., Peterson, K.R., Meadows, S.A., McClain, M.E., Jeng, G., Chmelar, R.S., Haugen, H.A., Chen, K., Singer, M.J., Moore, D. et al. (1998) Androgen receptor YAC transgenic mice carrying CAG 45 alleles show trinucleotide repeat instability. Hum. Mol. Genet., 7, 959–967.
58 White, J.K., Auerbach, W., Duyao, M.P., Vonsattel, J.P., Gusella, J.F., Joyner, A.L. and MacDonald, M.E. (1997) Huntingtin is required for neurogenesis and is not impaired by the Huntington's disease CAG expansion. Nat. Genet., 17, 404–410.[Web of Science][Medline]
59 Warner, J.P., Barron, L.H. and Brock, D.J. (1993) A new polymerase chain reaction (PCR) assay for the trinucleotide repeat that is unstable and expanded on Huntington's disease chromosomes. Mol. Cell. Probes, 7, 235–239.[Web of Science][Medline]
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