Human Molecular Genetics Advance Access originally published online on December 22, 2006
Human Molecular Genetics 2007 16(4):391-409; doi:10.1093/hmg/ddl467
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Wild-type huntingtin participates in protein trafficking between the Golgi and the extracellular space
1 Department of Genetics, Stanford University School of Medicine, Stanford, CA 94305-5120, USA and 2 Stanford Human Genome Center, 975 California Avenue, Palo Alto, CA 94304, USA
* To whom correspondence should be addressed at: Department of Genetics, M344, Stanford University School of Medicine, Stanford, CA 94305-5120, USA Tel: +1 6507259687; Fax: +1 6507259689; myers{at}shgc.stanford.edu
Received October 9, 2006; Accepted December 9, 2006
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Huntington disease (HD) is an autosomal dominant neurodegenerative disease caused by an expanded CAG trinucleotide repeat in the first exon of the HD gene, which results in a toxic polyglutamine stretch within huntingtin, the protein it encodes. Understanding the normal function of this essential protein is vital to understanding the root of the disease, yet despite more than a decade of investigation, its role in the cell remains elusive. Identifying the subcellular localization of huntingtin and understanding its effects on global gene expression are critical to this endeavor. While most reports agree that huntingtin is predominantly a cytoplasmic protein, conflicting distribution patterns have been demonstrated at the subcellular level. Here, we examine wild-type huntingtin's localization in cultured cells by expressing the full-length human protein tagged with enhanced green fluorescent protein (EGFP) within its unspliced genomic context. In fibrosarcoma and neuroblastoma cells, huntingtin shows discrete punctate, perinuclear localization overlapping largely with the trans-Golgi and cytoplasmic clathrin-coated vesicles, implicating huntingtin in vesicle trafficking. To determine whether huntingtin is involved in trafficking a specific subset of proteins, we measured changes in global transcription levels in embryonic stem cells and neurons lacking huntingtin. Huntingtin null neurons exhibit a significant reduction in transcripts encoding proteins destined for the extracellular space, many of which are components of the extracellular matrix or involved in cellular adhesion, receptor binding and hormone activity. Together, these findings support a role for huntingtin in the intracellular trafficking of proteins required for the construction of the extracellular matrix.
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Huntington disease (HD) is a progressive neurodegenerative disorder characterized by progressive motor, cognitive and behavioral dysfunction caused by the inheritance of a single mutated copy of the HD gene (1,2). The disease phenotype ensues when the naturally polymorphic CAG repeat within the first exon of the HD gene is larger than 36 residues, translating a toxic polyglutamine stretch within the protein it encodes, huntingtin (2). In humans, the length of the CAG repeat region is inversely correlated to the age-at-onset of the disease as well as its progression (3,4). Both normal and mutant huntingtins are widely expressed throughout the adult brain and peripheral tissues, yet selective degeneration of GABAergic medium spiny projection neurons occurs in the caudate and putamen of HD patients (2,5–7). Huntingtin also plays a crucial and indispensable role during embryogenesis, as demonstrated by the finding from multiple groups that mouse embryos lacking huntingtin die at approximately embryonic day 7.5 (8–10). Although the initiation of gastrulation is not inhibited, neurulation fails and the somites, nodes and notochord are not formed (8).
Despite its identification more than a decade ago, the function of huntingtin remains largely unclear. Insights have come from studies investigating huntingtin's interacting partners, subcellular localization and effects on gene expression. Dozens of proteins have been shown to interact with huntingtin, the bulk of which exhibit enhanced binding to the mutant version of the protein. Only a handful prefer associating the wild-type huntingtin (11–23). Many residents of the nucleus, including nearly a dozen transcription factors, such as CBP [cAMP response element-binding protein (CREB)-binding protein] and Sp1, are among the demonstrated binding partners of huntingtin (24,25). These associations appear to greatly affect the transcriptional profile of HD-afflicted cells and brain tissue, evidenced by altered global gene expression on microarrays (24,26–29). However, interactions between mutant huntingtin and ubiquitous transcription factors cannot account for gene- and tissue-specific effects of mutant huntingtin. Thus, it is possible that the specific vulnerability of the striatum in HD results from the depletion of wild-type huntingtin or loss of a normal interaction or activity.
Analysis of both the subcellular localization of wild-type huntingtin and its impact on gene expression can provide clues to its potential function. Biochemical and immunohistochemical evidence has shown that wild-type huntingtin in neurons is primarily in the cytoplasm (7,30,31). However, some support exists for huntingtin's presence within the nucleus (21,32–34). Conflicting associations between huntingtin and a variety of organelles within the cell have been shown: microtubules, Golgi complex, vesicles, axon terminals, endoplasmic reticulum, centrosomes and mitochondria (7,30–32,35–38). The degree of association between the protein and each of these cellular substructures varies widely between studies, as disparate cellular and animal model systems, antibodies and experimental methods have yielded disparate results. The absence of a firm consensus makes accurately predicting huntingtin's activity in the cell challenging.
We investigated the discrete localization of wild-type huntingtin protein with fluorescently labeled protein expressed from its intact gene on a bacterial artificial chromosome (BAC) transfected into cells. By exploiting a homologous recombination technique with a BAC clone spanning the entire human HD locus, we were able to express full-length wild-type huntingtin tagged with enhanced green fluorescent protein (EGFP) within its natural genomic context. As the endogenous promoter, regulatory elements and splice sites directing the protein's expression may be crucial in influencing its proper subcellular location, this approach enabled such regulation. As we show here, fluorescent microscopy and immunocytochemistry in various cell types define a punctate, perinuclear localization pattern. The protein's distribution overlaps largely with the trans-Golgi and cytoplasmic clathrin-coated vesicles, implicating huntingtin in vesicle trafficking between the Golgi complex and plasma membrane.
Applying genome-wide gene expression arrays, we sought to test further whether huntingtin is involved in trafficking a specific subset of proteins from the Golgi. By taking advantage of the viability of both Hdh null mouse embryonic stem (ES) cells and neurons that are formed from these cells by differentiation in culture, we were able to examine the effect of huntingtin-deficiency on global gene expression throughout the differentiation process from ES cells to neurons, up to 10 days post-differentiation, and identify pathways disrupted by the absence of huntingtin. We found that cells lacking huntingtin exhibit a significant reduction in transcripts encoding proteins destined for the extracellular space, many of which are components of the extracellular matrix or involved in cellular adhesion, receptor binding and hormone activity. Furthermore, lysosomal activity and apoptosis are increased, suggestive of elevated cellular stress in Hdh null cells. Together, these findings further support a role for huntingtin in intracellular trafficking, more specifically, transport of a specific subset of proteins required for the construction of the extracellular matrix.
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RESULTS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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The general strategy for targeted BAC modification
BACs are useful tools for examining genes within their genomic context. Spanning hundreds of base pairs in length, BACs often include generous upstream and downstream flanking sequences in addition to the gene/genes of interest being studied (39). Innovative approaches have been devised for altering BACs in Escherichia coli by using electroporated linear DNA. The homologous recombination system employs a defective
prophage, which both protects and recombines a linear DNA substrate within the bacterial cell, allowing the DNA to be efficiently recombined in vivo (40–42). We obtained the BAC (RPCI-11 399e10 human male) spanning the human HD locus located at 4p16.3 from the Children's Hospital Oakland Research Institute (CHORI) (GenBank accession no. AC005516). The HD-containing BAC (HDBAC) contains 166 113 bp of HD sequence from the transcription start site through exon 67; 3268 bp upstream genomic sequence; 30 967 bp downstream genomic sequence and 8823 bp of pBACe3.6 backbone vector sequence (43). We confirmed the integrity of the HDBAC by restriction enzyme analysis (data not shown).
We generated a panel of BACs for investigating huntingtin localization and expression by inserting several cassettes into the HDBAC, within the open reading frame as well as the flanking sequence. To visualize huntingtin by fluorescence, we targeted EGFP to both the 5'-and 3'-ends of HD just following the initiating methionine codon and immediately prior to the transcription stop codon, respectively. To enable the selection of stable transformants in bacterial cells, we inserted a zeocin cassette by targeting nearly 30 kb downstream of the coding region of the HD transcript. The most distant location was chosen to avoid the interruption of any functional regulatory elements immediately distal to HD. As huntingtin expression is modest in most cell types, we replaced huntingtin's endogenous promoter in one construct with elongation factor I (EF1) (Fig. 1).
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Homologous recombination in the BAC requires two successive insertion events. The primary event involves the insertion of a tetracycline resistance (Tet®) cassette in the location where the desired final cassette (EGFP, zeocin and EF1) will ultimately reside. We identified recombinants by their ability to grow in the presence of tetracycline. The secondary event involves the replacement of the Tet® cassette with the desired cassette (EGFP, zeocin and EF1), resulting in tetracycline-sensitive recombinants (40,42).
To enable homologous recombination, we incorporated 42 bp of target sequence at the 5' end of each forward and reverse primer. For each cassette to be inserted in the HDBAC, we used two sets of primers, one set for the primary insertion of the Tet® cassette and a second set for the successive insertion of the EGFP, zeocin or EF1 cassette. Following each recombination event, we confirmed integration by PCR, XhoI restriction digest and sequencing of the purified BAC. Restriction digests verified that rearrangements had not taken place during the modification process (Fig. 2C).
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Wild-type huntingtin exhibits punctate, perinuclear localization polarized to a single side of the nucleus
We analyzed the subcellular distribution of huntingtin following transient transfection of the HDBACs into human HT1080 fibrosarcoma and U87 neuronal/gliablastoma/astrosarcoma cells. We used fluorescent microscopy to visualize the subcellular distribution of wild-type huntingtin 24, 48, 72 and 96 h post-transfection, enabling adequate time for sufficient expression off the BAC.
Huntingtin expression was variable, but generally low, even after 96 h post-transfection. Given the increased size of the vector, splicing requirements and less-robust promoter activity, we expected and witnessed considerably lower expression off the BAC compared to the positive EGFP plasmid control (44). While no expression was visible at 24 h post-transfection, faint fluorescence gradually became discernable at 48 h, with 72 and 96 h post-transfection yielding the brightest cells. Few of the cells fluoresced as brightly as cells transfected with an EGFP positive control vector. In more modestly fluorescing HT1080 cells, huntingtin-EGFP expression localized around the perimeter of the nucleus in small, punctated structures (Fig. 3A and C). It also appeared to be polarized to a single side of the nucleus. Huntingtin disseminated throughout the cytoplasm in more brightly fluorescing cells, yet remained more highly concentrated around the nucleus with a gradual lessening of expression towards the plasma membrane. In all cases, we noted an absence of full-length wild-type huntingtin in the nucleus, evidenced by a void in the region of nuclear DAPI staining. We observed the same expression pattern in the U87 neuroblastoma cells (Fig. 3B and D).
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Transiently transfected HT1080 cells expressing huntingtin from the EF1 high-expressing mammalian promoter on HDBAC/EF1/EGFP(C)/Zeo exhibited considerably higher expression than those expressing the protein from the endogenous promoter. We noted modest expression visible as early as 24 h post-transfection, with a gradual intensifying of fluorescence in the following days. Similar to expression from the endogenous promoter, huntingtin localization was enhanced in the perinuclear region, slanted to a single side of the nucleus (Fig. 3E). However, at all four time-points, we observed signal throughout the entire cytoplasm, but specifically lacking in the nucleus. The lack of expression restricted around the nucleus may simply be due to EF1-regulated over-expression of the protein, saturating the perinuclear region and causing huntingtin to disseminate more quickly away from the cell's hub. Again, we saw the same expression pattern in the U87 neuroblastoma cells (Fig. 3F).
Given the identical expression patterns for both the N-terminally and C-terminally tagged huntingtin, we propose that the EGFP tag does not interfere with the protein localization or function, and thus represents accurate assessment of its location within the cell. Fluorescent protein expression did not affect cell viability.
Huntingtin co-localizes with the Golgi and clathrin-coated vesicles in HT1080 and U87 cells
To determine whether huntingtin's punctate, perinuclear expression is consistent with the distribution of other subcellular structures, we performed immunocytofluorescence assays on HT1080 and U87 cells transiently transfected with HDBAC/EGFP(C)/Zeo by using a panel of antibodies that recognize discrete substructures in the cell: microtubules, the Golgi and clathrin-coated vesicles.
Staining for the Golgi zone in both U87 and HT1080 cells expressing fluorescently tagged huntingtin indicated that both huntingtin and the Golgi have similar punctate, perinuclear-enhanced expression co-localizing to a single side of the nucleus (Fig. 4A–D, M–P). In all cases, the protein localization was skewed to the same side of the nucleus. In higher fluorescing cells, the regions of most intense huntingtin expression correlated with the regions of Golgi distribution. Co-localization was also apparent between huntingtin and clathrin-coated vesicles. Anti-clathrin showed an enhanced juxtanuclear distribution, with gradually lessening expression towards the plasma membrane, similar to cells highly fluorescent for huntingtin (Fig. 4E–H, Q–T). In cells expressing moderate levels of huntingtin, the biased perinuclear huntingtin fluorescence matches the most intense staining of clathrin.
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Staining for tubulin in U87 cells expressing EGFP-tagged huntingtin indicated that huntingtin's perinuclear-enhanced expression does not strictly co-localize with tubulin's spindly star-like distribution (Fig. 4I–L). We found that regions of highest huntingtin expression near the nucleus are the regions relatively devoid of tubulin labeling. As huntingtin overlaps with the Golgi, some of this may be due to the Golgi's location alongside the centrosome, the organizing center from which microtubules polymerize outward (45). Therefore, the void in tubulin staining overlapping the dense Golgi complex is consistent with intense huntingtin fluorescence.
The demonstrated codistribution of full-length wild-type huntingtin expressed from the human HDBAC with both the Golgi complex and clathrin-coated vesicles in both transiently transfected fibrosarcoma cells and neuroblastoma cells support a role for wild-type huntingtin in the endocytotic pathway.
Hdh null neurons exhibit more transcriptional changes than do Hdh null ES cells
As subcellular localization hinted at a role for huntingtin in intracellular transport, we sought to determine whether huntingtin is involved in trafficking a specific subset of proteins. To test this hypothesis, we analyzed changes in global gene expression in Hdh null ES cells and Hdh null differentiated neurons—cultured cell lines that lack huntingtin expression—at 6, 8 and 10 days post-differentiation compared to similar wild-type control cells (these cells are described in Materials and Methods). For each time-point, three biological replicates were subjected to separate culturing, differentiation and total RNA extraction. We reserved aliquots of the RNA samples for quantitative PCR (qPCR) confirmation of a select number of transcripts (see Confirmation of Bead Chip Array Findings below), using the remainder for hybridization to Illumina Sentrix® MouseRef-8 BeadChip expression arrays. We compared average normalized fluorescence measurements for the three Hdh null replicates to the average normalized fluorescence measurements for the three wild-type replicates at each time-point. We assessed gene expression differences by using the paired Student's t test statistic, enabling the fold-change difference between the two measurements to be calculated. This method allowed us to identify mRNAs that were comparatively increased or decreased in the Hdh null ES cells and differentiated neurons relative to controls. mRNAs that met more than 2-fold expression elevation or repression for at least one time-point at P 
0.05 were explored further. Using the 2-fold stringency cutoff, a cumulative total of 472 transcripts on the array platform demonstrated a significant difference between Hdh null and wild-type cells across all four time-points, representing ~2% of the probes included on the array. The number of genes with altered expression in cells lacking huntingtin was comparatively fewer (80) in the ES cells compared to the differentiated neuronal cells. The number of mRNAs exhibiting more than 2-fold change gradually increased as the neuronal differentiation process progressed, with the array detecting 199, 207 and 249 transcripts at day 6, 8 and 10, respectively.
Global assessment of gene expression changes in Hdh null ES cells and neurons
We observed temporal patterns of gene expression among the 24 045 transcripts on the array. Clustering the transcripts by related expression with Cluster and TreeView software generated large blocks of genes following similar profiles in undifferentiated ES cells and differentiated neurons (Fig. 5A). Comparing the number of transcripts with differential expression across all four time-points, this global perspective highlights the relative paucity of mRNAs in Hdh null ES cells (UD) with altered expression, suggesting that at this very early stage the cells are less affected—at the level of transcription—by the absence of huntingtin than are differentiated neurons. This is in agreement with the known expression levels of huntingtin, which indicate that ES cells express low, but detectable, levels of huntingtin compared to the relatively high concentrations observed in the brain (6,8,9,46,47). It is also apparent that although many genes exhibit an elevation or repression in expression across several time-points or throughout the entire time-course, evidenced by flanking time-points exhibiting similar trends, subsets of genes appear to be vital to discrete stages, spiking or falling rapidly. These same patterns are mirrored in the cluster diagram representing only the 472 transcripts that comply with our 2-fold stringency cutoff (Fig. 5B).
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Hdh null ES cells exhibit diminished expression of vital patterning genes and increased lysosomal activity
Eighty transcripts fit the criterion of at least a 2-fold change in expression with P
0.05 [91% of these 80 transcripts (75) have P 0.01 and 83% (66) have P 0.001] in Hdh null ES cells compared to wild-type controls. A quarter of these transcripts represent genes of unknown or inferred function based on homology, with the majority of this subset identified as RIKEN cDNA clones. The remaining genes fall into several functional categories: patterning, extracellular matrix, protein degradation and cell cycle (Table 1).
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Three genes with key roles in patterning the developing embryo exhibited differential expression in Hdh null ES cells compared to wild-type controls. Lefty1 (LeftB), a morphogen required for proper left-right axis asymmetry of developing organ systems, was down-regulated. Lefty1 is also known to be involved in restricting Nodal localization in the developing embryo without affecting its transcript levels. In accordance, no change in Nodal mRNA expression was detected on our array. Similarly, Otx2, involved in anterior-posterior patterning and neuronal development, and Pem (Rhox5), a marker of extraembryonic tissue, were both repressed. Our findings are in agreement with a recently published non-quantitative in situ hybridization approach, which demonstrated a down-regulation of Lefty, Otx2 and Pem mRNA in E7.5 Hdh knock-out mice. The same study also indicated that Nodal was dispersed away from its normal restricted region of localization, though its expression level remained unchanged (48). The observed dysregulation of these transcripts is intriguing. Although their skewed expression has little effect on the viability of Hdh ES cells that remain healthy in culture after many successive splittings, the absence of huntingtin in the early stages of differentiation and development is devastating to the embryo, ultimately resulting in early prenatal lethality (8–10).
We noted an elevation in transcript levels of several genes involved in protein degradation or lysosomal function in Hdh null ES cells. Cops8 and Psmd8, components of the 26S proteasome involved in the degradation of ubiquitinated proteins, and Ubei21, a ubiquitin-conjugating enzyme, were increased several-fold over control cells. Glycoprotein catabolism and disulfide oxidoreductase activity within the lysosome may also be elevated, evidenced by the increased mRNA levels of Manba and Ifi30, respectively.
Several genes (Plk1, Ccdc5, Aurkc and Pttg1) with known involvement in cell cycle regulation exhibited altered expression in Hdh null ES cells, suggestive of impaired cell division. Once again, misregulation may have little influence on cells in the undifferentiated state, but more serious repercussions may result during the repetitive cell cycling required during development, perhaps indicated by the sustained elevation of Plk1 and Ccdc5 throughout the entire neuronal differentiation process.
We also identified four transcripts with involvement in the formation of the extracellular matrix: two co-regulated sheet-forming collagens (Col4a1 and Col4a2) found in basement membranes, Adam23, a metallopeptidase involved in cell-matrix interactions, and B3galt6, required for the synthesis of proteins essential to the basement membrane. Although these functional categories do not represent statistically significant gene ontology (GO) groupings, likely due to the limited number of transcripts with differential expression in Hdh null undifferentiated cells, their early misregulation in ES cells suggests they function as precursors or instigators of increased cellular dysfunction in more specialized cell types. It may be important to note that altered transcript levels do not necessarily imply altered protein levels due to possible compensatory mechanisms by the cell.
Deficiencies in extracellular proteins in Hdh null neurons suggests faulty formation of the extracellular matrix and interrupted receptor binding and signaling
Hundreds of genes exhibited differential gene expression between Hdh null and wild-type cells through the progression from ES cells into neuronal cells. A large subset of the transcripts remained down-regulated throughout the entire time-course with a lesser number exhibiting sustained elevation from day 6 to day 10 post-differentiation. Several significant GO categories can be pulled from the data, indicating the possible disruption of several functional pathways in the absence of huntingtin. Using GoStat software to identify biological, molecular and physiological classes of genes, we found that several over-represented gene groups: extracellular space (P = 1.77e–13), receptor binding (P = 0.000291), hormone activity (P = 0.00101), enzyme inhibitor activity (P = 0.00101), extracellular matrix (P = 0.0025), structural components conferring tensile strength (P = 0.0326), retinol/retinal/retinoid/isoprenoid binding (P = 0.0114, P = 0.0363, P = 0.0435, P = 0.0435 respectively) and lysosome activity (P = 0.0379). By further investigating the function of detected transcripts via NCBI and Expasy, several genes involved in cell adhesion, axonogenesis, action within the synaptic region, transport and apoptosis were readily apparent (Fig. 6).
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Extracellular space
The most striking observation is the widespread down-regulation of transcripts whose protein products make their way to the extracellular space (see Fig. 6 for the complete list). Many of the transcripts encode for secreted proteins that function outside the cell's interior, such as ApoA1 (cholesterol transport), Sfrp1 (Wnt signaling) and Ltf (iron binding and transport), and proteins integral to the plasma membrane, including Fxyd6 and Prom1. A large subset of this broader class includes genes involved in the construction of the extracellular matrix, scaffolding to the plasma membrane and cell adhesion. Six transcripts from the collagen family (Col4a1, Col4a2, Col6a1, Col6a3, Col16a1 and Col18a1) were down-regulated in Hdh null neurons. As collagen fibers are crucial for strengthening and organizing the matrix, their diminished presence is suggestive of a deficit in structural architecture and integrity of the cell's exterior. Likewise, the transcripts of proteins (Adamts2, Mmp2, Adamts4 and Dcn) required for the cleavage of pro-collagen prior to fibril assembly and matrix degradation, thus mediating its organization and plasticity, are similarly decreased. Perhaps in response to the down-regulation of matrix metalloproteinases, the repression of several metalloproteinase inhibitors (Serpina3n, Serpinb9f and Timp3) was also detected on the arrays. The array also detected diminished expression for a dozen adhesion molecules (Tgfbi, Mfap4, Adam23, Tgm2, Emp3, Spp1, Cldn6, Thbs2, Epb4.1l3, Tpm2, Mucdhl and Prelp) involved in mediating cell–cell and cell–matrix interactions in differentiated neurons lacking huntingtin. Insufficient amounts of these binding proteins hint at a weakened ability to anchor cells to one another or tether them to the extracellular meshwork in the absence of huntingtin.
Interestingly, Hs3st3a1, B3galt6, Chst1 and Mgat3, enzymes that function in the Golgi to synthesize carbohydrate moieties for proteins essential to basement membranes, proteoglycans and other components of the extracellular matrix, showed elevated expression in Hdh null cells. Increased attempts at Golgi glycosylation for hyaluronan and proteoglycans (Hapln3, Dcn, Agrin, Gpc1, Thbs2)—despite their reduced expression—may be indicative of a compensatory mechanism by the cell in an attempt to increase production of these vital structural components.
Receptor binding, hormone activity and signaling
The down-regulation of transcripts involved in cellular adherence and the construction and binding activities of the extracellular matrix may also interfere with proper receptor function, thus interrupting signaling and hormonal pathways. The vast majority of transcripts involved in receptor binding and hormonal activity detected on the arrays showed diminished expression in Hdh null neurons. In addition, six genes involved in the Wnt signaling pathway [En1, Sfrp1, Dkk3, Wisp1, IGF-II and Csnk1e (CK1e)], including several Wnt target genes, demonstrated similar down-regulation (Fig. 6) (49–52). The Wnt signaling molecules play an important role in early brain development, and recent evidence has demonstrated that these molecules require the presence of heparan sulfate proteoglycans (HSPGs) to regulate their pathways (53,54). The depletion or disruption of HSPGs may be responsible for initiating secondary signal transduction defects in Hdh null neuronal cells.
Axonogenesis and activity with the synapse
mRNAs with roles in axon guidance and neurite extension are subject to dysregulation in Hdh null neurons (Fig. 6). A large set of transcripts [Sox21, Nav1, Nfatc4, Ptn (HB-GAM), Lgals1, St6gal1, Sema3f, Srgap2] is indicative of diminished or aberrant outgrowth and extension. The arrays detected a similar dysregulation of transcripts encoding for proteins functioning within the synapse, specifically the down-regulation of mRNAs (Adra2b, Rab17, Kcnc4, Csnk1e) involved in the regulation of neurotransmitter release or uptake at the synaptic terminals. While no single neurotransmitter signaling pathway was over-represented on the arrays in the absence of huntingtin, the altered transcription of a group of synapse-enriched transcripts (Clic6, Pde1b, Slc1a1, Kcnc4, Csnk1e) indicates interruption of dopamine signaling or potential excitotoxicity via increased activity of glutamate receptors. Furthermore, Pde1b and Csnk1e are enriched within the striatum, the brain region most susceptible to damage in HD, suggesting that the loss of huntingtin function may contribute to the excitotoxic damage observed in HD patients. In response to the potential oxidative injury, Gsta1 and Gsta2, detoxification proteins that protect cells following excess glutamate exposure, showed up-regulation in Hdh null neurons.
Increased lysosomal and apoptotic activity and decreased cell growth and proliferation in Hdh null neurons
Lysosome activity
A subset of proteins misregulated in Hdh null neuronal cells has known involvement in lysosomal function and protein degradation. Four genes (Cops8, Psmd8, Ube2i and Fbxo44) encoding participants in the ubiquitin–proteasome pathway maintained elevated transcript levels throughout the neuronal differentiation. Likewise, Gaa, Manba and Fuca1, all involved in the breakdown of glycoproteins within the lysosome, showed an enhanced expression. Increased lysosomal activity may be indicative of an over-representation of misdirected or misfolded proteins within the cell's cytoplasm, which are ultimately targeted for destruction.
Apoptosis
Cells expressing the polyglutamine-expanded version of huntingtin have been shown to be more susceptible to cell death, evidenced by massive neurodegeneration in later stages of Huntingtin disease and transcriptional activation of apoptosis-inducing genes in affected tissues and cells (5,55,56). Embryos lacking huntingtin also exhibit pyknotic nuclei and enhanced apoptosis prior to death at embryonic day E7.5, suggesting that both presence of polyglutamine-expanded huntingtin and the loss of wild-type huntingtin lead to cell death at the organismal level (9,10). We noted a similar presence of apoptosis-related transcripts on our array, detecting elevated expression of several cell death-inducing transcripts [Txnl1, Prdx2, Casp3, Pdcd8, Ern2 (Ire2)]. Although arrays are not strictly quantitative, the transcripts showed a general rise in expression in Hdh null cells over the length of the time-course, culminating in their highest levels at day 10 post-differentiation. Furthermore, the arrays detected a repression of transcripts encoding protective proteins, such as Gpx3, that guards against oxidative damage, and the cathepsin proteinase inhibitor, Serpina3g.
Growth, proliferation and differentiation
Despite the ability to maintain Hdh null ES cells and differentiated neurons in culture, huntingtin is vital for growth and development given that mouse embryos lacking the protein are underdeveloped and smaller in size (9,10). We confirmed this essential role for huntingtin on our arrays, detecting the general down-regulation of genes involved in growth, proliferation and cellular differentiation (Fig. 6). A subset of these transcripts appear to function specifically within the brain: Serpinf1, Emp3, Tcfap2c, Dlx3 and Elavl4. Many general growth-related or mitogenic mRNAs (Ghrh, Scgb3a1, Cdkn1c, Sp6, Gkn1, Igf2) also exhibited reduced expression. Conversely, Hdh null neurons consistently overexpressed Ccng2, a negative regulator of cell proliferation and cell cycle progression.
Iron regulation and intracellular transport disrupted in Hdh null neurons
Transport
Iron transport has been previously shown to be askew in Hdh null cells and mouse embryos (57,58). We also detected transcripts hinting at aberrant metal ion transport in cells lacking huntingtin (Fig. 6). We observed increases in two transcripts (Picalm, Cubn) involved in iron uptake at the plasma membrane, as well as repression of three others (Slc40a1, Hamp, Ltf) responsible for the regulation of iron export and binding. The increased uptake of iron into the cells coinciding with the diminished ability to expel it may result in iron overload and toxicity in Hdh null cells. Increased amounts of iron in the brain has been linked to the occurrence of neurodegenerative disease (59,60). This is in agreement with the reported neurodegeneration observed adult mice lacking huntingtin strictly within the forebrain (61). The arrays also indicated the misregulation of transcripts involved in zinc and copper transport.
In addition to the transport of iron across the plasma membrane, we noted the dysregulation of several transcripts involved in intracellular trafficking in Hdh null cells. Although our localization results hint at a role for huntingtin in intracellular transport, the absence of the wild-type protein was not assumed to affect the transcription of transport-related proteins. However, both Dnaic1, a component of the microtubule-based dynein motor protein, and Rab17, a regulator of receptor-mediated transcytosis and endosome recycling, showed reduced expression throughout the differentiation process. With increased expression in Hdh, null neurons were Abca5, a lysosomal ATPase involved in transport, and Gdi3, an inhibitor of vesicle docking on the Golgi. Collectively, these findings indicate a possible impairment of vesicle transport throughout the cell.
Confirmation of microarray findings
To confirm the changes in expression detected by the arrays, we performed qPCR analyses on 12 transcripts, for genes that were both up-regulated and down-regulated to varying degrees (Fig. 7). Like the RNA samples hybridized to the arrays, the measurements from qPCR were average values from the three original biological replicates and a pooled sample. The expression of each of the 12 transcripts were elevated or repressed in the same manner detected on the array, although the fold-change difference between Hdh null and wild-type cells was often higher with qPCR. We observed a similar fold-change discrepancy in previous experiments within the lab, and it is likely attributable to binding saturation on the array and/or increased sensitivity and specificity of qPCR. The Pearson's r-value was measured at 0.98 between the two methods, thus confirming the accuracy and specificity of the array data with regard to differences in transcript level.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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By using novel approaches, we linked huntingtin's perinuclear subcellular localization with a potential role in trafficking a discrete set of proteins between the Golgi and the extracellular space. We homologously recombined an EGFP fluorescent tag into a BAC carrying the chromosomal region spanning the entire human HD gene. Expression of huntingtin-EGFP in cultured fibrosarcoma and neuroblastoma cells indicates that full-length, wild-type huntingtin localizes to the perinuclear region in a punctate vesicular-like pattern polarized to a single side of the nucleus. Immuncytochemistry demonstrates that this pattern mimics the distribution of the Golgi and clathrin-coated cytoplasmic vesicles, which also show enhanced localization around the nucleus. We did not observe fluorescence within the nucleus. Our results fit well with a subset of previously published reports of both the localization of huntingtin itself, as well as the distribution of several of the protein's binding partners. Early reports using immunocytochemistry detected huntingtin diffusely throughout the cytoplasm, with evidence of its association with a variety of intracellular organelles, including the microtubules, endoplasmic reticulum, mitochondria, vesicular structures and Golgi complex (7,31,35). Several reports also identified huntingtin within the nucleus, with one report suggesting that huntingtin may shuttle between the nucleus and the cytoplasm (32,33,62). Follow-up studies suggest that huntingtin's localization is more restricted, focused in a discrete region encircling the nucleus. However, only selective antibodies have detected this pattern (38,63). As different antibodies have elicited a range of localization patterns, likely due to cross-reactivity and their unique affinities for huntingtin, our use of a fluorescent tag bypassed the requirement for antibodies. With this alternative approach, our localization studies of full-length, wild-type huntingtin under the control of its endogenous local regulatory elements corroborate the latter perinuclear findings.
Analysis of the subcellular localization of huntingtin provides some clues to its potential function. The combined distribution pattern from the earlier reports prompted speculation of huntingtin's role in vesicle trafficking of the secretory and endocytic pathways (7,31,35,38,63). Our observed perinuclear co-localization of huntingtin with the Golgi and clathrin-coated vesicles meld well with several others lines of evidence involving huntingtin in vesicular transport. Huntingtin has been shown to associate with various proteins that play a role in intracellular trafficking, including HAP1, HIP1, HIP14, 
-adaptin C (HYPJ), SH3-containing GRB-like protein 3 (SH3GL3), Fip2, PACSIN1 and Cdc42-interacting protein 4 (CIP4) (16,18,19,64–68).
To determine if huntingtin may be involved in trafficking a specific subset of proteins, we evaluated changes in global transcription levels in ES cells and neurons lacking huntingtin. We identified genes with aberrant expression using by Illumina BeadChip arrays, which contain probes for more than 24 000 human transcripts, enabling us to identify classes of genes that fit into distinct functional networks and that require huntingtin for normal activity. Our data suggest that the disruption in transcripts encoding crucial components of the extracellular space, primarily structural building blocks and modifiers of the extracellular matrix, serve as the primary cellular disturbance in Hdh null cells. The down-regulation of many collagens, adhesion molecules and metalloproteinases may ultimately contribute to the downstream interference of receptor binding and signaling pathways, most of which were also decreased in transcript levels in the mutant cells. Likewise, we detected misregulation of many transcripts with roles in axon or dendrite extension or that function within the synapse in Hdh null cells compared to control cells. We also noted an increase in transcript levels of genes involved in lysosomal and apoptotic-related activity. These results fall in line with reports of increased cell death and neurodegeneration in mouse embryos and adult brains lacking huntingtin expression (8–10,61,69).
Gene expression of Hdh null ES cells appeared minimally affected by the absence of huntingtin. This is not wholly unexpected given their prolonged viability in culture (8–10). However, we detected a down-regulation of several transcripts involved in early patterning of the developing embryo and reconfirmed this observation by qPCR. These results are in accordance with recent in situ hybridization results demonstrating misregulation of these same mRNAs (48). Although it has been shown previously that Hdh null ES cells can be differentiated into largely functional neurons, we indicate here that these neurons exhibit a more definitive dysfunction at the transcriptional level, made apparent by the altered expression of many classes of genes (46).
This study provides the first evidence that the absence of huntingtin expression results in a disruption of many components of the extracellular matrix and basal lamina. These supporting structures play pivotal roles in the maintenance and activity of the cell, influencing localized activation of signaling pathways, internal cytoskeletal rearrangements and organization of the matrix into specialized basement membranes. Depending on the cell type and the context, these interactions drive cell fate, cell migration and cellular architecture, such as polarization and process extension (70). Thus, the down-regulation of key components of the matrix may ultimately affect the cell's overall stability and its ability to interact with neighboring cells. Such intercellular communication is particularly important for the activity and viability of neurons.
As the matrix plays a supportive role in signal transduction, the repressed expression of many matrix components may contribute to the observed interference of receptor binding and signaling pathways, most of which demonstrated a similar repression. Of notable interest, several mRNAs of the Wnt signaling pathway showed diminished expression in Hdh null neuronal cells. Accumulating evidence demonstrates that the signaling activity (ligand levels, distribution, receptor-interactions and expression of downstream target genes) of the Wnts rely upon the presence of HSPGs, cell surface and extracellular matrix macromolecules that consist of long, sulfated sugar moieties attached to a protein core (53,54). Furthermore, just as HSPGs are major components of the mammalian central nervous system, Wnts have been shown to play important roles in early brain development (71,72). While these signaling disruptions may not represent primary defects of huntingtin loss, they may be suggestive of the secondary effects of a weakened extracellular matrix and our observed repressed expression of agrin and glypican, two key HSPGs. These pathways have not been previously implicated in HD or huntingtin-related activity, and thus provide new avenues for future research.
In addition to their role in signal transduction, proteoglycans have been implicated in many important neuronal functions, such as synaptic remodeling, neural plasticity, axon pathfinding and neurite outgrowth (71). Likewise, brain matrix metalloproteinases, which are involved in the modification of the extracellular matrix, have also been shown to be important regulators of many of these same biological processes (73). We detected many dysregulated transcripts involved in synaptic activity (endoscytosis and neurotransmitter release) and axon and neurite extension on our arrays. The differential expression of decorin (chondroitin sulphate proteoglycan) and agrin and glypican (HSPGs), as well as many proteoglycan-binding proteins (i.e. Prolargin, HB-CAM, Collagen XVIII) indicates that repression of these cell-surface and extracellular matrix proteins, as well as its remodeling, may negatively influence these neuronal processes in Hdh null differentiated neurons. Although differentiated neurons lacking huntingtin appear normal, small morphological variances are possible and may not affect their short-term viability in culture. Given the critical role axonogenesis and neurite outgrowth play in the development of the brain, the dysregulation of many of the brain-enriched transcripts may help explain the early neurological defects and disorganization observed in mice embryos lacking huntingtin.
The statistically significant decreased expression of proteins whose actions are in the extracellular space in Hdh null differentiated neurons suggest that there is a defect in intracellular transport in the absence of huntingtin. Our results coincide with a role for huntingtin in vesicular trafficking and intracellular transport. More specifically, huntingtin may participate in trafficking a subset of proteins destined for the extracellular matrix, transporting them from the Golgi, where they are modified, to the plasma membrane via clathrin-coated vesicles. We hypothesize that inability of these proteins to reach the cell's exterior could result in their build-up within the cytoplasm, and in the cell's attempt to achieve homeostasis, their expression is down-regulated. The wayward proteins are eventually targeted for degradation in the lysosome, accounting for the increase in transcripts involved in lysosomal activity and protein degradation.
This study cannot rule out the possibility that huntingtin functions as a transcription factor or indirectly modulates transcription. Our localization studies suggest that the native protein is restricted to the cytoplasm, a finding that conflicts with a handful of other reports (32,33). We attribute this incongruity to the specificity of the two differing techniques. Directly observing EGFP-tagged huntingtin off the BAC maintains its endogenous regulation and eliminates variable binding capabilities and cross-reactivity of unique antibody epitopes. Thus, given its localization within the cytoplasm, huntingtin would have to mediate gene expression outside the nucleus, possibly by sequestering transcription factors within the cytoplasm. While huntingtin has been shown to bind to a variety of known transcription factors—NcoR, CBP, Sp1, TAFII130, TBP, RAP30, p231HBP/HYPB, REST, CtBP—the vast majority of these associated factors exhibit enhanced binding to the mutant version of huntingtin, bringing into question whether these associations are functional with wild-type huntingtin in vivo under normal physiological conditions (7,11,74). Wild-type huntingtin appears to bind to the transcriptional co-repressor REST in the cytoplasm of neuronal cells, possibly preventing REST from entering the nucleus and restricting expression of its target genes, including transcription of bone-derived neurotrophic factor (BDNF) (23,75). One might expect the sequestration of REST within the cytoplasm to be lost in neurons lacking huntingtin, resulting in a down-regulation of BDNF. However, we did not detect any change in the mRNA levels of BDNF at day 6, 8 or 10 post-differentiation. No change would be expected in Hdh null ES cells, as REST localizes to the nucleus in non-neuronal cells. It may be important to note that the array probe spanned a region of the 5' untranslated region upstream of Exon II, where REST regulation of the neurotrophin occurs.
This work refines the role of wild-type huntingtin within the cell. Our results suggest its specific involvement in the transport of extracellular proteins to the cell's exterior, indicating that many of the effects of huntingtin's absence may be secondary to this disruption. Our findings open the door for future investigation into the link between huntingtin and proteoglycan distribution, as well as its role in signal transduction.
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MATERIALS AND METHODS |
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TOPABSTRACTINTRODUCTIONRESULTSDISCUSSIONMATERIALS AND METHODSREFERENCES |
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Bacterial cells and BACs
A BAC—clone 399e10 from the RPCI-11 Human Male BAC Library—spanning the entire human HD gene as well as 3268 bp upstream and 31 009 bp downstream was acquired from CHORI (76). The human DNA is cloned in the pBACe3.6 (8823 bp) backbone and selected in a growth media containing 20 µg/ml chloramphenicol (Cam) (Sigma, Cat. no. C-4881).
A temperature-sensitive recombination-proficient E. coli strain, EL250, was used for all BAC modification steps. The EL250 strain carries a defective prophage that enables targeted recombination while protecting the electroporated linear DNA substrate (42).
BAC DNA preparation
For transfection in EL250 E. coli, BAC DNA was prepared using the Qiagen Plasmid Mega Kit according to the standard protocol (Qiagen, Cat. no. 12183). One liter BAC cultures were grown overnight at 37°C in LB medium containing 20 µg/ml chloramphenicol. The BAC DNA pellet was air dried for 
15 min before being dissolved in 500 µl TE-buffer, pH 8.0 for 20 min. Samples were store at –30°C till use. XhoI digests (New England BioLabs, Cat. no. R0146S) were performed to confirm the integrity and purity of the BAC DNA using 400 ng purified BAC DNA in 25 µl total reaction volume incubated at 37°C for 4 h. Gels were electrophoresed overnight at 25 V in 0.5% agarose.
Transforming EL250 cells with the BAC clone
In order to modify the BAC, it was first transformed into the EL250 bacterial cell line to enable temperature-sensitive homologous recombination. Overnight, 3 ml cultures were grown from fresh single EL250 colonies at 32°C. Starter cultures were diluted 50-fold in 100 ml LB medium and grown at 32°C until OD (A600) reached mid-log phase (0.4). The culture was incubated on ice for 20 min, divided into five prechilled 50 ml tubes and centrifuged at 5000 r.p.m. for 10 min at room temperature (Beckman JA10 rotor). The supernatant was removed, and cells were resuspended in 3 ml cold, sterile Millipore water before being aliquoted into two 1.5 ml microfuge tubes on ice. Tubes were centrifuged at full-speed for 5 min at 4°C using a benchtop centrifuge. After rinsing and spinning the pellets two additional times with 1.5 ml cold, sterile water, a final spin was preformed to remove all traces of supernatant. The pellets were resuspended in 35 µl 10% glycerol, snap-frozen on dry ice and stored at –80°C until needed. This large culture prepared a total of 10 reactions.
Transformation of the BAC was performed by electroporation of 500 ng circular, supercoiled BAC into 35 µl electrocompetent cells thawed on ice in pre-chilled 0.1 cm gap cuvettes (BIORAD, Cat. no. 165–2089) using a Bio-Rag Gene Pulser set at 1.75 kV, 25 µF and 200 ohms. One ml of SOC media (Invitrogen, Cat. no. 1229974) was added to the cells following electroporation. Cells were grown at 32°C for 90 min while shaking and plated on LB agar media supplemented with chloramphenicol (final concentration 20 µg/ml). Individual colonies were selected and grown, the DNA prepped as described above, and XhoI digested for confirmation. The above procedure was modified from Lee et al. (42).
Preparation of linear DNA cassettes with homology ends
A series of linear cassettes were created for recombination into the HDBAC: EGFP for both the 5'-end and 3'-end of the HD gene, zeocin-resistance (Zeo®) selectable marker, EF1 mammalian promoter and the tetracycline-resistance (Tet®) selectable marker. Standard PCR conditions were used to amplify linear DNA fragments using the Finnzyme Polymerase system (DyNAzyme EXT DNA Polymerase, Cat. no. F-505L).
A series of Tet® cassettes 2,451 bp in length were made for the initial ‘Positive Selection’ homologous recombination events, in which the gene encoding tetracycline-resistance was inserted into locations where it would ultimately be replaced by EGFP, Zeo®, or EF1. The secondary ‘negative selection’ homologous recombination events replacing the Tet® cassette with EGFP, Zeo® or EF1were selected for by tetracycline-sensitivity. Chimeric primers were designed containing two domains: 42 bp homologous to flanking regions on the target BAC where the cassette was to be inserted and 21 bp for amplifying the cassette from plasmid template. Therefore, two sets of chimeric primers were designed for the insertion of EGFP, SV40-Zeo® and EF1. Whereas the BAC homology arms are identical in both primer pairs for a single cassette, one pair contains 21 bp for amplifying Tet® and one pair contains 21 bp for amplifying the replacement cassette. The linear DNA products were amplified using standard PCR conditions with 3.75 mM Mg2+ and 5% DMSO.
For all primers listed, nucleotides in italics are homologous to the targeted BAC sequence and those in roman type are homologous to insertion cassettes. For each positive selection event, the tetracycline-resistant cassette was amplified using Tn10 primers 5'AGATCTATGATTCCCTTTGTCAACAG and 5'AAGCTTATGATGATGATGTGCT TAAAAAC. The SV40-driven zeocin-resistant (Zeo®) cassette (1042 bp) was amplified from pcDNA3.1/Zeo (Invitrogen, Cat. no. V860–20) with primers 5'TTGGTTGTAGATCCTGGCAGCACGGGTGAGTAAGGGAAGGGCCCAGGCAGGCAGAAGTATGCA and 5'TAGGGCTGAGCAAGACCTGAGCCCTGCCCTCCCCTCCTCAGGGAGGTCGACGGTATACAGACA for insertion 28 549 bp downstream of the HD open reading frame. The EGFP tags (792 bp) were amplified from the pLP-EGFP-C1 vector (Clontech, Cat. no. 6342-1) using two sets of primers, one set for insertion into the N-terminus and one set for insertion into the C-terminus. For recombination into the N-terminal of HD just following the initiating ATG, primers 5'GAGGCCTCCGGGGACTGCCGTGCCGGGCGGGAGACCGCCATGGTGAGCAAGGGCGAGGAGCTG and 5'CTTGAGGGACTCGAAGGCCTTCATCAGCTTTTCCAGGGTCGCTCTAGATCCGGTGGATCCCGG were used. For insertion into the C-terminal of HD just prior to the stop codon, primers 5'CTGCTGACTTGTTTACGAAATGTCCACAAGGTCACCACCTGCGT GAGCAAGGGCGAGGAGCTG and 5'GCCCCAGCTGCCGCCTCACAGTCTCTCCCACCATGGCGCTCATCTAGATCCGGTGGATCCCGG were used. The EF1 promoter (1248 bp) was amplified from pEF1/V5-His (Invitrogen, Cat. no. V920-20) with primers 5'CTTTCTAGAAGTACTACTCATTACTTCTGCTTGCATCTCCCTAGGAATCTTTGCAGCTAATGG and 5'GCAGGTTCTGCCTCACACAGCAAGGCCGCTGACAGCGCAGCGGTACCAAGCTAATTCCTCACG. The amplified linear PCR products were purified using QIAquick PCR purification kits (Qiagen, Cat. no. 28104) following the standard protocol, suspended in sterile water and quantified by spectroscopy.
Preparation of recombination-proficient, electroporation-competent, EL250 cells
Overnight cultures of the EL250 bacterial cell line carrying the HDBAC were grown from fresh single colonies at 32°C in an LB medium supplemented with 20 µg/ml Cam, diluted 50-fold in 100 ml LB (20 µg/ml Cam) and grown at 32°C until OD (A600) reached mid-log phase (0.3–0.4). The cultures were immediately transferred to a 43°C shaking waterbath for 20 min to induce expression of recombination proteins: Beta, Exo and Gam. The culture was swirled in ice water for 20 min to cool the culture quickly, divided into five prechilled 50 ml tubes and centrifuged at 5000 r.p.m. for 10 min at 4°C (Beckman JA10 rotor). The supernatant was removed and the cells were resuspend in 3 ml cold, sterile Millipore water before being aliquoted into two 1.5 ml microfuge tubes on ice. Tubes were centrifuged at full-speed for 5 min at 4°C in a benchtop centrifuge. After rinsing and spinning the pellets two additional times with 1.5 m cold, sterile water, a final spin was preformed to remove all traces of supernatant. The pellets were resuspended in 35 µl 10% glycerol, snap-froze on dry ice, and stored at –80°C until needed. This protocol was adapted from Yu et al. (77). For BACs with an integrated Zeo® cassette, 25 µg/ml zeocin (Invitrogen, Cat. no. 46-0509) was added to the medium.
Cell transformation with linear cassettes for homologous recombination
Step I—Positive selection
Transformation of the linear DNA cassettes were performed by electroporation using 100–300 ng linear Tet® cassette DNA into 35 µl of heat-induced EL250 electrocompetent cells thawed on ice. The cells were transferred into pre-chilled 0.1 cm gap cuvettes and electroporated with a BioRad gene pulser set at 1.75 kV, 25 µF and 200 ohms. One ml of SOC medium was added immediately following electroporation. Cells were incubated at 32°C for 90 min with shaking and 500 µl of cell culture was spread on two large LB agar plates supplemented with Cam (20 µg/ml) and Tet (7.5 µg/ml). This protocol was adapted from Lee et al. (42). Tetracycline-resistant colonies are picked 36 h post-transformation for confirmation of cassette integration.
Step II—Negative selection
Transformation of replacement cassettes (EGFP, Zeo® or EF1) was performed by electroporation using 100–300 ng linear replacement cassette DNA in 35 µl of heat-induced EL250 competent cells carrying the Tet®-BAC thawed on ice. Following electroporation as described above, cells were incubated at 32°C for 90 min with shaking and 500 µl were spread on two large LB agar plates (9 g agar, 5 g tryptone, 2.5 g yeast extract, 5 g sodium chloride, 5 g NaH2PO4 H2O, 8 ml chlorotetracycline (6.3 mg/ml), 156 µl Cam (20 mg/ml), 125 µl Zeo (100 mg/ml) and 500 ml water) and incubated at room temperature overnight. The media was supplemented with zeocin (25 µg/ml) if a zeocin cassette had been previously recombined. The faint lawn was gently scraped from the surface of the agar into 10 ml LB broth ( 10–1 dilution) using a disposable cell scraper (Costar, Cat. no. 3010). Using glass beads, 100 µl of 10–3 and 10–4 dilutions were plated on several similar LB agar plates supplemented with Cam (6.25 µg/ml), chlorotetracyline (50 µg/ml) and fresh fusaric acid (12 or 24 µg/ml) (Sigma, Cat. no. F-6513). As a control to estimate titer of viable cells, 100 µl of a 10–7 dilution was plated on LB agar plates supplemented with Cam (6.25 µg/ml) only. The plates were incubated at 32°C for 48 h. Individual tetracycline-sensitive colonies were picked for screening. This protocol was adapted from Yang et al.(40) and Lee et al. (42).
Confirmation of recombinants
After both Steps I and II, recombinants were screened and verified by three methods: PCR amplification around the region of cassette insertion, XhoI restriction digest and sequencing.
To PCR around the integration site, the potential-positive BACs were purified using the Qiagen R.E.A.L. Prep 96 Plasmid kit (QIAGEN, Cat. no. 26171). Two ml LB cultures supplemented with 20 µg/ml Cam were inoculated with single Tet® colonies and incubated for 16 h at 32°C with shaking at 150 r.p.m. in a Precision shallow form shaking bath. Instructions were followed precisely, with the exception that all inversion steps were eliminated to avoid cross-contamination between neighboring colonies. DNA pellets were air-dried for 20 min and redissolved in 35 µl 10 mM Tris-Cl, pH 8.5 by incubating overnight at room temperature. PCR was carried out with 3.75 mM Mg2, 5% DMSO and 0.75 µl purified BAC DNA template from 96-well BAC prep using primers flanking the insertion site.
Clones with proper integration were prepped using the Qiagen Mega Prep Kit as described above. To ensure that the BAC did not undergo rearrangement during recombination, a XhoI restriction digest using 300–400 ng purified BAC DNA in 25 µl total reaction volume confirmed the expected digest pattern. Rearrangement-free clones were sequenced around the region of the BAC encompassing the inserted cassette using the ABI 377 Sequencer.
Cultured cell lines
Human U87 neuronal/gliablastoma/astrosarcoma cells were grown in Dulbecco's Modified Eagle's minimum essential media (Gibco, Cat. no. 10569–010) with GlutaMax I, high Glucose, 110 mg/L sodium pyruvate and pyridoxine-HCl, supplemented with 10% fetal bovine serum (HyClone, Cat. no. SH30071.03) and 1% penicillin/streptomycin (Gibco, Cat. no. 15140–122). Human HT1080 fibrosarcoma cells were grown in Eagle's DMEM supplemented with 4 mM L-glutamine and Earle's BSS, 1.5 g/l sodium bicarbonate, 0.1 mM non-essential amino acids, 4.5 g/l L-Glucose, 10% fetal bovine serum and 1% penicillin/streptomycin. Cells were grown at 37°C in 5% CO2 in 25 cc culture flasks and split 1 : 4 (U87) or 1 : 20 (Ht1080) upon 85% confluency using 0.25% Trypsin-EDTA (Gibco, Cat. no. 25200–056). Cells were fed every 2 days following split.
Liposomal transfection of eukaryotic cells with BAC DNA
Chamber slides were coated with 700 µl Poly-d-Lysine (Invitrogen, Cat. no. P-6407) diluted to 0.1 mg/ml and incubated for 24 h at room temperature. Prior to plating, the Poly-d-Lysine was removed and slides were washed three times with tissue culture-grade distilled water. Cells were trypsinized at 80% confluency from one 25 ml flask with 0.25% Trypsin/EDTA and harvested by centrifugation. The cells were resuspended in 5 ml culture media. Cell density was counted using Trypan Blue and 2.5 x 10–4 to 7.5 x 10–4 cells were plated per well in four-well chamber slides. Additional media was added for a total of 800 µl, and cells were grown at 37° in 5% CO2 overnight. Depending on the number of cells plated, cell density was approximately 30–50% confluent after an overnight incubation. The following day, 3 : 2 FuGene 6 Reagent : DNA complex reactions were prepared using 3 µl FuGene (Roche, Cat. no. 1814075) : 1 µg PBS negative control/1 µg EGFP-C1 positive control/2 µg BAC DNA (HDBAC/EGFP(C)/Zeocin, HDBAC/EGFP(N)/Zeocin or HDBAC/EF1/EGFP(C)/Zeocin) in a total reaction volume of 100 µl OPTI-MEM lacking serum (Gibco, Cat. no. 31985-070). Preparation of complexes followed the standard protocol. After a 30–40 min incubation, the complex mixture was added drop-wise into each well and gently swirled to mix. Media was changed daily. Cells were incubated for 24–96 h before fluorescence was captured.
Fluorescent microscopy
Cells were visualized under the fluorescent microscope (Zeiss AxioSkop2) at 400x magnification. To fix cells, chamber slides were washed three times with room temperature PBS and incubated in 750 µl 4% paraformaldehyde for 30 min at room temperature in the dark. Paraformaldehyde was removed, the slides were washed three times with PBS, and cells were incubated with DAPI diluted 1 : 1000 in PBS for 5 min. Vectashield mounting media (Vector Labs, Cat. no. H-1000) and cover slips were applied.
Fluorescent immunocytochemistry
Cells were fixed with paraformaldehyde as described above. Paraformaldehyde was removed, the slides were washed three times with PBS, and the cells were incubated in 500 µl 5% goat serum blocking buffer for 1 h. Blocking buffer was removed and cells were incubated in 500 µl primary antibody diluted in PBS for 1 h: anti-ß-tubulin (final concentration diluted 1 : 500 in PBS, 2 µg/ml) (monoclonal, clone KMX-1, Chemicon, Cat. no. MAB3408), anti-Golgi zone (final concentration diluted 1 : 30 in PBS) (monoclonal, clone 371-4, Chemicon, Cat. no. MAB1271) and anti-clathrin heavy chain (final concentration diluted 1 : 100 in PBS, 2.5 µg/ml) (BD Biosciences, Cat. no. 610499). Primary antibody was removed, cells were washed four times with PBS and 500 µl Alexa Fluor 555 goat anti-mouse IgG secondary antibody (final concentration diluted 1 : 400 in PBS, 5 µg/ml) (Molecular Probes, Cat. no. A-21422) diluted in PBS was added to each well. Cells were incubated for 1 h. Secondary antibody was removed, cells were washed four times with PBS and incubated with DAPI (final concentration diluted 1 : 1000 in PBS) for 5 min. Following four washes with PBS, mounting media and coverslips were applied. All incubations were performed in the dark at room temperature.
Growth of ES cells
Mouse ES cells were grown on inactivated hygromycin-resistant 129 mouse embryonic fibroblasts (MEFs) [Cell and Molecular Technology (Specialty Media), Cat. no. PMEF-H]. One day prior to plating or splitting the ES cells, a vial of MEFs was thawed and plated into four 25 cc tissue culture flasks (1.3 x 106 cells/flask) coated with 0.2% gelatin (Sigma, Cat. no. G1393) and allowed to settle overnight in Dulbecco's minimum essential media (Gibco, Cat. no. 11960-069) supplemented with high glucose, lacking pyridoxine hydrochloride, L-Glutamine and sodium pyruvate, 10% fetal bovine serum (Gibco, Cat. no. 16000-044), 1% 100x penicillin/streptomycin (Gibco, Cat. no. 15140-122) and 1% 100x GlutaMax (Gibco, Cat. no. 35050-061).
Hdh null mouse ES cells (courtesy of Scott Zeitlin) and wild-type EmbryoMax Strain 129/SVEV ES cells [Cell and Molecular Technology (Specialty Media), Cat. no. CMTI-1] were plated in the MEF-containing flasks in Dulbecco's minimum essential media supplemented with high glucose, lacking pyridoxine hydrochloride, L-Glutamine and sodium pyruvate, 10% fetal bovine serum, 10% neonatal calf serum (Gibco, Cat. no. 16010-159), 1% 100x penicillin/streptomycin, 0.1% ESGRO Leukemia Inhibitory Factor (LIF) (Chemicon, Cat. no. ESG1106) and 7 x 10–6% ß-mercaptoethanol (Sigma, Cat. no. M7522). Upon 85% confluency, the media was removed from flasks and cells were washed with warm PBS before incubating in 1 ml warm 0.05% Trypsin-EDTA (Gibco, Cat. no. 25300-054) at 37°C for 10 min. Cells were removed by pipetting with 5 ml media and centrifuged to pellet cells and remove Trypsin-EDTA. After removal of the supernatant, cells were resuspended in 15 ml warm growth media and each 25 cc flask was replated into a single new gelatin-coated 75 cc flask. Splitting reduced MEFs to 4.6 x 105 cells/flask. ES cells were not fed the following day post-plating, after which the media was replaced daily. ES cells were either collected at the undifferentiated state for array analysis or differentiated into neurons (see below).
To collect cells in the undifferentiated state, media and cells were removed from three 75 cc flasks upon 85% confluency for both Hdh null and wild-type ES cells, as described above, and replated in 0.2% gelatin-coated flasks for 1 h to remove feeder cells. The supernatant containing the stem cells was siphoned off and spun down. One ml TRIzol® Reagent (Gibco, Cat. no. 15596-018) was added to the cell pellets and allowed to incubate for 5 min before dislodging and homogenizing the cells by pipetting. Cell suspension was collected and transferred into a QIAshredder (Qiagen, Cat. no. 79654) in 600 µl aliquots. After spinning for 2 min, the eluate was collected in a 1.7 ml tube and stored at –80°C until RNA preparation.
Neuronal differentiation of ES cells
For differentiation into neurons, cells were prepared as described above. For each time-point, the media, ES cells and MEFs were removed from three 75 cc flasks each for both Hdh null and wild-type cell cultures, as described above. The cell suspension was replated in 0.2% gelatin-coated flasks for 1 h to remove feeder cells, the supernatant was siphoned off and 1.5 x 106 cells were transferred in single-cell suspension onto non-coated bacteriologic dishes, cultured in the presence of differentiation medium containing Dulbecco's minimum essential media, 10% fetal bovine serum, 10% neonatal calf serum, 2 mM glutamine, 1 mM sodium pyruvate, 0.1 mM non-essential amino acids, 0.1 mM ß-mercaptoethanol and penicillin/streptomycin. Under these conditions, embroid bodies were formed and cultured for 4 days in the absence of, then followed for 4 days in the presence of 0.5 x 10–6M all-trans-retinoic acid (Sigma, Cat. no. R2625). Subsequently, 8-day EBs were transferred onto 0.1% gelatinized tissue culture plates and cultured for an additional 6, 8 and 10 days (three plates each) prior to collection. Dishes were washed with PBS to remove the media and 1 ml of Trizol was added to each dish. Cells incubated in the TRIzol® reagent for 5 min to enable the dislodging and homogenization of the cells by pipetting. Cell suspension was collected and transferred into a QIAshredder in 600 µl aliquots. After spinning for 2 min, the eluate was collected in a 1.7 mL tube and stored at –80°C until RNA preparation.
RNA extraction from mouse ES cells and neuronal cells for microarray analysis
The RNA preparation for all samples—undifferentiated ES cells and differentiated neurons—were completed together to avoid variation in sample preparation. TRIzol®-homogenized samples were thawed on ice before adding 200 µl of chloroform per 1 ml of TRIzol® to each tube (240 µl/tube). Samples were mixed thoroughly and allowed to settle for 5 min at room temperature before centrifuging at 3500 r.p.m. for 15 min at 4ºC in a tabletop centrifuge (IEC Centra CL3). The aqueous phase was transferred to a new 14 ml round-bottom tube. Total RNA was collected according to manufacturer's instructions using the RNeasy Mini Kit (Qiagen, Cat. no. 74104). RNA was eluted in 150 µl RNase-free water. The concentrations were calculated before storing RNA at –80°C until RNA amplification step. A total of 24 samples (six undifferentiated ES cells and 18 differentiated neurons) were prepared.
Ilumina chip labeling and hybridization
We used the Illumina Sentrix® MouseRef-8 Expression BeadChips (Illumina, Cat. no. BD-26–201), covering over 24 000 well-annotated mouse RefSeq transcripts, for our gene expression comparisons. A single round of RNA amplification is required prior to hybridization of the RNA to the array. The Illumina TotalPrep RNA Amplification Kit (Ambion, Cat. no. IL1791-1) generated biotinylated, amplified RNA. The manufacturer's recommended instructions were followed with only slight deviations. To synthesize First Strand cDNA by reverse transcription, we used 250 ng total RNA from each sample collected above. Following the Second Strand cDNA synthesis and cDNA purification steps, the in vitro transcription to synthesize cRNA was prepared and allowed to incubate overnight for 15 h. The cRNA was purified according to instructions and eluted in 100 µl nuclease-free water. A total of 850 ng of RNA was aliquoted into tubes and brought up in a total volume of 42.5 µl before concentrating samples by vacuum centrifugation for 1 h. Samples were resuspended in 11.3 µl RNase-free water and kept on ice until hybridization. For hybridization to three Illumina Sentrix® MouseRef-8 BeadChips, the protocol using included reagents were followed precisely. Samples were arranged on the chips so that the three biological replicates of each time point were spread across three separate arrays, thus controlling for possible chip–chip variation. Chips were read by the Illumina BeadStation 500x and collected data was reviewed for analysis.
Statistical analysis
The raw data were processed by the Illumina BeadStudio Data Analysis Software, which generated the averaged intensity signals across 30 beads for each transcript. We imported the intensity data for all 24 000 transcripts across 24 samples into R-values, log transformed with base of 2, and quantile normalized the data to generate an equal intensity distribution across the 24 samples (78). Using the normalized data, we calculated a Two-way ANOVA with Strain (Hdh null, wild-type) and Stage (undifferentiated, neuronal day 6, neuronal day 8, neuronal day 10) as factors, to identify transcripts showing strain-dependent or stage-dependent expression patterns. Two-group comparisons using the Student's t test identified, separately, transcripts that were differentially expressed between Hdh null cells and 129 wild-type cells at each of the four stages: undifferentiated ES cells, neuronal day 6, neuronal day 8 and neuronal day 10. For each of the four comparisons, we recorded the t scores, P-values and fold-change (expressed in log scale). This data was further filtered, with a stringency of 2-fold elevation or repression in Hdh null cells compared to wild-type cells at P 0.05. Up-regulated expression was color-coded yellow, and down-regulated expression was color-coded blue.
qPCR confirmation of differentially expressed transcripts
Select transcripts demonstrating at least a 2-fold increase or 2-fold decrease in gene expression in Hdh null ES cells compared to wild-type ES cells were chosen for confirmation using real-time PCR analysis. Primers for qPCR were designed to exonic regions of each gene, generating amplicons 60–80 bp in length with similar melting temperatures (62°C). After confirming that the amplicons aligned to the proper genomic region and did not form secondary structure characteristics using the University of Santa Cruz In-Silico PCR program, primers were tested by Real-Time using mouse genomic DNA (50, 5 and 0.5 ng) to verify that they produced gene-specific products of an appropriate size and that proper standard curves were generated based on observed threshold cycle numbers (Tc). GAPDH and ß-actin housekeeping genes were used as positive controls.
qPCR used the same RNA preparations used for the original array experiments. For each sample, 4 µg of RNA was treated with amplification grade DNase I following manufacturer's protocol (Invitrogen, Cat. no. 18068-015). The treated RNA was then converted to cDNA using Invitrogen's SuperScript III First-Strand Synthesis System for RT-PCR (Invitrogen, Cat. no. 18080-051) following the manufacturer's protocol. Separate cDNA preparations amplified with oligo(dT)12–18 and random hexamers were combined prior to real-time PCR.
The following primers (Operon) were used for amplification of mouse cDNA (all primers are listed 5' to 3'): GAPDH—ACATCAAGAAGGTGGTGAAGCAG/GACAACCTGGTCCTCAGTGTAGC; ß-actin—CAAGTACTCTGTGTGGATCGGTG/ACATCTGCTGGAAGGTGGACAG; Mcm6 (accession no. NM_008567 [GenBank] .1)—TGAACAACCTTCTCAATGATCGTC/CCTGAAGGAAATCGAATCAGAGA; Card4 (accession no. NM_172729 [GenBank] .1)—GACTCCAAGTTCATGCTGTGCTA/TGTCCATATAGGTCTCCTCCAGC; Mest (accession no. NM_008590 [GenBank] .1)—ACCACATTAGCCACTACCCACAG/AGAAGGAGTTGATGAAGCCCATA; Tuba4 (accession no. NM_021344 [GenBank] .2)—ATTGAGCGTCCAACCTATACCAA/GAAGCTGTGATGGAGGAGACAAT; Mt2 (accession no. NM_008630 [GenBank] .1)—GGGTCCCCACATCTGTGTAAATA/GTGGAGAACGAGTCAGGGTTGTA; Dab2 (accession no. NM_023118 [GenBank] .1)—GAGACAAGGTGTCCTCTTGGGTA/GGAGAAAGGGTTCTCGAGTGAAT; Camk2b (accession no. NM_007595 [GenBank] .2)—TACATCCGCCTCACACAGTACAT/GTCTCTTCGGACTGGCTGGTA; Clu (accession no. NM_013492 [GenBank] .1)—GTGACCACCCATTCCTCTGACT/TCAGAGTCAAACAGCTTCACCAC; Leftb (Lefty1) (accession no. NM_010094 [GenBank] .2)—GATGAAGTGGGCCGAGAACT/CCCACACATTCATATGTCAGGAA; Col4a1 (accession no. NM_009931 [GenBank] .1)—GTCTGGAAGAGTTTAGAAGCGCC/GCGTAGTAATTGCACGTTCCTCT; Otx2 (accession no. NM_144841 [GenBank] .1)—GCTATGCTGGCTCAACTTCCTAC/GGTGATGCATAGGGGTCAAATAA; Rhox5 (Pem) (accession no. NM_008818 [GenBank] .1)—CGACACCGAGATTTTGATTTGAT/GCTGAATTCTTGAAAAGTAAGGGC. A 0.6 µl aliquot of the cDNA samples was used as template for each reaction containing AmpliTaq DNA Polymerase Stoffel Fragment solution (Applied Biosystems, Cat. no. N808-0038) and SYBR Green (Molecular Probes, Cat. no. S-7563) with 1 µl each of the left and right primers (10 mM). Real-time thermal cycling was performed using a BioRad iCycler with continuous SYBR Green monitoring according to the manufacturer's recommendations, using iCycler software. Cycling parameters for all amplifications were as follows: initial melt at 95°C for 10 min followed by 40 cycles of 95°C for 20 s, 63°C for 20 s and 72°C for 20 s. All real-time PCR reactions were performed in triplicate, including negative controls (no DNA and no reverse transcriptase) and positive controls (serial dilutions of known amounts of genomic DNA).
Target DNA transcript quantities were estimated from Tc using iCycler software. The Tc value is determined as the cycle at which fluorescence (measured in relative fluorescence units, RFU) rises above a baseline threshold. To achieve uniformity across all experiments, PCR baseline thresholds were set at 50 RFU; this fluorescence threshold was empirically chosen because at this point PCR reactions were in the linear range. For every gene sequence studied, the Tc for each sample was normalized to the ß-actin and GAPDH Tc values to normalize for differences in cDNA sample aliquots. To calculate the corresponding fold-change between the HD null and wild-type expression levels for each transcript, the DTc value was determined by subtracting the average normalized Tc value for the HD null samples from the average normalized Tc value for the corresponding wild-type samples. DNA quantities were expressed using the following equation. Fold-change = 2(DTc). A Pearson's r-value was calculated to determine the degree of correlation between the fold-change values measured using real-time analysis to those measured from the Illumina array platform.
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ACKNOWLEDGEMENTS |
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The authors thank Scott Zeitlin for the generous gift of the Hdh null ES cells, Doug Mortlock and Greg Barsh for providing the EL250 recombination-proficient cells and TetBH2.4 #5 plasmid, respectively, Greg Barsh, Patrick Brown and Devin Absher for helpful discussions, and Larissa Tsavaler, Patrick Collins, Nathan Trinklein and Loan Nguyen for technical assistance and support. This research was supported by an award from the Wills Foundation and Training Grant T32 GM 07790-25 from the General Medical Sciences Institute at the National Institutes of Health.
Conflict of interest statement. The authors declare that they have had no involvements that might raise the question of bias in the work reported or in the conclusions, implications or opinions stated.
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