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Proc Natl Acad Sci U S A. Apr 15, 2008; 105(15): 5868–5873.
Published online Apr 8, 2008. doi:  10.1073/pnas.0801775105
PMCID: PMC2311380
Medical Sciences

Artificial miRNAs mitigate shRNA-mediated toxicity in the brain: Implications for the therapeutic development of RNAi


Huntington's disease (HD) is a fatal, dominant neurodegenerative disease caused by a polyglutamine repeat expansion in exon 1 of the HD gene, which encodes the huntingtin protein. We and others have shown that RNAi is a candidate therapy for HD because expression of inhibitory RNAs targeting mutant human HD transgenes improved neuropathology and behavioral deficits in HD mouse models. Here, we developed shRNAs targeting conserved sequences in human HD and mouse HD homolog (HDh) mRNAs to initiate preclinical testing in a knockin mouse model of HD. We screened 35 shRNAs in vitro and subsequently narrowed our focus to three candidates for in vivo testing. Unexpectedly, two active shRNAs induced significant neurotoxicity in mouse striatum, although HDh mRNA expression was reduced to similar levels by all three. Additionally, a control shRNA containing mismatches also induced toxicity, although it did not reduce HDh mRNA expression. Interestingly, the toxic shRNAs generated higher antisense RNA levels, compared with the nontoxic shRNA. These results demonstrate that the robust levels of antisense RNAs emerging from shRNA expression systems can be problematic in the mouse brain. Importantly, when sequences that were toxic in the context of shRNAs were placed into artificial microRNA (miRNA) expression systems, molecular and neuropathological readouts of neurotoxicity were significantly attenuated without compromising mouse HDh silencing efficacy. Thus, miRNA-based approaches may provide more appropriate biological tools for expressing inhibitory RNAs in the brain, the implications of which are crucial to the development of RNAi for both basic biological and therapeutic applications.

Keywords: gene therapy, Huntington's disease, RNAi, AAV

The ability of siRNAs to silence target genes was first demonstrated in 1998 by Andrew Fire et al. (1) and has since emerged as a revolutionary strategy to reduce target gene expression. RNAi occurs naturally in cells as a posttranscriptional regulatory mechanism mediated by endogenous miRNAs (25). RNAi is hypothesized to have evolved as a cellular coping mechanism providing the cell a means to decrease the expression of various deleterious viruses and transposons (6, 7). In recent years, scientists have coopted this biological process to reduce the expression of target mRNAs by using exogenously applied siRNAs, shRNAs, or artificial miRNAs. Aside from the widespread basic biological applications of RNAi, the ability to reduce gene expression marks a major advance toward the development of disease therapies, particularly for dominantly inherited disorders.

Among the dominant diseases that may benefit from RNAi-based therapies is Huntington's disease (HD). Our laboratory (8) and others (9) have previously demonstrated that partial reduction of mutant huntingtin expression by viral delivery of shRNAs is efficacious in preventing the development of motor deficits and neuropathology in transgenic mouse models of HD. In these proof-of-principal studies, the therapeutic effect on disease phenotype was studied by knocking down a mutant human HD transgene in the setting of two normal mouse HD homolog (HDh) alleles. Although allele-specific targeting of disease transcripts for HD therapy would be ideal, to date no prevalent SNP residing on the mutant transcript has been identified. Therefore, we undertook studies to identify inhibitory RNAs that would target both mouse HDh and human HD transcripts, with the intention of testing the efficacy of reducing the expression of both alleles in a knockin model of HD (10). Here we describe the surprising finding of neurotoxicity in mouse brain caused by some, but not all, shRNA expression vectors screened in vivo and the notable reduction in toxicity after moving those toxic inhibitory RNAs into miRNA-based delivery systems.


shRNAs Cause Striatal Toxicity in Mice.

We first designed and screened shRNAs (driven by the mouse U6 promoter) that target conserved sequences spanning human HD and mouse HDh mRNAs [Fig. 1A and supporting information (SI) Table S1], taking into consideration the most recent siRNA design rules (1113). Silencing of HD mRNA measured by quantitative real-time PCR (QPCR) and dot blot analysis revealed a decrease in huntingtin protein expression after transfection of shRNA expression plasmids into mouse C2C12 and human-derived HEK 293 cell lines (data not shown). Of the 35 shRNAs tested, three were chosen for further study based on silencing efficacy. The shRNAs target sequences in exons 2, 8, and 30 of HD mRNAs and are henceforth referred to as sh2.4, sh8.2, and sh30.1, respectively (Fig. 1B). Western blot analysis demonstrated that these shRNAs, but not mismatch (mis) control shRNAs, reduce endogenous huntingtin protein expression in mouse C2C12 cells (Fig. 1C). Similar results were seen in human-derived HEK 293 cells.

Fig. 1.
In vitro screening of shRNAs targeting human HD and mouse HDh transcripts. (A) Thirty-five shRNAs (bars above cartoon) targeting conserved sequences (Table S1) spanning human HD and mouse HDh mRNAs were generated with consideration for sequences that ...

To examine the long-term effects of brain-delivered shRNAs in the CAG140 knockin mouse model of HD (10), U6-shRNA expression cassettes were cloned into adeno-associated viral vectors (AAV serotype 2/1) (Fig. 2A). AAVs also contained a humanized Renilla GFP (hrGFP) expression cassette to identify the distribution and types of cells transduced. Five-week-old CAG140 knockin mice were injected bilaterally into the striatum with AAVsh2.4-GFP, AAVsh8.2-GFP, AAVsh30.1-GFP, or AAV-GFP (viral control) and killed 15 weeks later. Robust expression of GFP was observed in cells throughout the rostral/caudal extent of the striatum and within fibers of the globus pallidus (Fig. 2B). Immunofluorescence analyses indicated that GFP-positive cells colocalized with a neuronal marker (NeuN), but not with markers for astrocytes (GFAP) or oligodendrocytes (RIP1) (Fig. S1). QPCR performed on RNA isolated from GFP-positive striatal tissue showed a significant and statistically similar reduction of HDh mRNA expression (≈60%) among the different active shRNA-expressing vectors, compared with mice injected with AAV-GFP [F(3, 11) = 32.3, P < 0.001 for post hoc analyses comparing each AAV-shRNA group to the AAV-GFP control] (Fig. 2C). Moreover, Western blot analysis demonstrated a significant reduction in huntingtin protein levels after AAVshRNA-GFP administration, compared with mismatch controls [t(8) = 3.9, P < 0.01] (Fig. S2).

Fig. 2.
HD shRNAs cause sequence-specific striatal toxicity in mice. (A) Diagram of the recombinant AAV2/1 viral vectors containing shRNA and hrGFP expression cassettes. (B) Photomicrographs represent the rostral-to-caudal distribution of hrGFP-positive cells ...

Unexpectedly, immunohistochemical analyses for dopamine- and cAMP-regulated protein (DARPP-32), a marker of medium-sized spiny projection neurons in the striatum, revealed striatal toxicity in mice injected with AAVsh2.4-GFP and AAVsh30.1-GFP (Fig. 2D Upper). Reduction in DARPP-32 immunoreactivity was largely confined to the transduced (GFP-positive) regions of the striatum. Interestingly, this toxicity was not seen in mice injected with AAVsh8.2-GFP (Fig. 2D Upper). Striata from these mice were similar to AAV-GFP-injected control mice.

To assess whether the observed loss of DARPP-32 staining was associated with microglial activation, tissue sections were stained with an anti-Iba1 antibody to identify both resting and reactive microglia throughout the brain. AAVsh2.4-GFP- and AAVsh30.1-GFP-injected striata demonstrated high Iba1 expression, whereas AAVsh8.2-GFP-injected striata were similar to control mice (Fig. 2D Lower). Moreover, AAVsh2.4-GFP- and AAVsh30.1-GFP-injected mice demonstrated dramatic reactive astrogliosis, compared with AAVsh8.2-GFP- and control-injected mice, as evidenced by robust GFAP staining in areas of the striatum corresponding to high GFP positivity (data not shown). Notably, a mismatch control for the HD2.4 sequence, AAVsh2.4mis-GFP, induced toxicity similar to sh2.4 and sh30.1 without reducing HDh mRNA expression. This, in addition to the sh8.2 data, indicates that three (two active, one inactive) of four shRNAs were toxic and that toxicity is not caused by silencing huntingtin.

Although all U6-shRNA expression cassettes were cloned into the same viral vector, we tested for the possibility that toxicity correlated with steady-state levels of the expressed products. RNA samples harvested from shRNA-treated striata were analyzed by small transcript Northern blot probing for the mature antisense (AS) and sense (S) RNAs generated by the respective shRNAs. Results demonstrate that sh2.4 AS RNA and sh30.1 AS RNA are expressed more robustly than sh8.2 AS RNA (Fig. 3), thus correlating toxicity with increased expression levels of the shRNAs in vivo. The disparity in expression levels is interesting, particularly given the fact that each shRNA was designed using the same rules, injected at the same viral dose, driven by the same Pol-III promoter, and silenced HDh mRNA to a similar degree. Notably, the processed sense strands and unprocessed shRNA transcripts were not detectable in brain lysates. This finding suggests that the toxicity is due, in part, to high levels of mature inhibitory RNAs, rather than inappropriate sense strand loading into the RNA-induced silencing complex (RISC) or saturation of endogenous RNAi export machinery.

Fig. 3.
The nontoxic sh8.2 generates lower levels of processed antisense RNA. (A) Small transcript Northern blot was performed to assess AS RNA levels present in mouse striata treated with the indicated AAVshRNA-GFP. (Left) Two separately treated striatal tissue ...

Antisense Sequence Levels Are Reduced by Using an Artificial miRNA.

Because the toxic shRNAs were expressed at higher levels than the nontoxic, active hairpin, an obvious approach to reduce toxicity would be to lower the viral titer injected. In the brain, decreasing the titers of AAVsh2.4-GFP by a half log (1e12) or a full log (5e11) achieved silencing of HDh mRNA (47% and 51%, respectively), but did not alleviate striatal toxicity (Fig. S3). Decreasing the titers even further (1e11 or 5e10) reduced the silencing efficacy to 15% of controls, an activity level possibly below therapeutic efficacy (Fig. S3). Thus, we tested whether levels of inhibitory RNAs could be minimized without compromising silencing efficacy by using an artificial miRNA as an siRNA shuttle (vs. an shRNA).

In corresponding work, we have found that artificial miRNAs effectively silence target gene expression relative to shRNAs without generating excessive levels of inhibitory RNAs (R.L.B. and B.L.D., unpublished data). Consequently, we cloned two of the toxic sequences (HD2.4 and HD2.4mis) into an artificial miRNA scaffold based on human miR-30 (14), thus creating mi2.4 and mi2.4mis (Fig. 4A). We first compared the expression levels of mi2.4 and sh2.4 by small transcript Northern blot analysis at 48 h after transfection of RNAi-expressing plasmids into HEK 293 cells. Probing for the HD2.4 antisense strand revealed that mi2.4 produces substantially lower levels of inhibitory RNAs relative to sh2.4. Notably, sh2.4 generates an abundance of precursor and processed RNAs even at a 10-fold lower dose (Fig. 4B). Despite the dramatic difference in expression levels, mi2.4 reduced endogenous HD transcripts almost as effectively as sh2.4 (50% and 60% silencing, respectively) in HEK 293 cells (Fig. 4C).

Fig. 4.
An artificial miRNA approach naturally reduces precursor and mature inhibitory RNAs. (A) Sequences and comparison of sh2.4 and mi2.4 containing the core HD2.4 sequence (shaded boxes). Each transcript starts with the +1-G nucleotide natural to the U6 promoter. ...

Artificial miRNAs Mitigate Striatal Toxicity in Mice.

We next generated AAV2/1-expressing mi2.4 or the mi2.4 mismatch control (Figs. 2B and and44A) to test whether the development of striatal toxicity could be prevented relative to AAVsh2.4-GFP. Because shRNA-induced toxicity was not dependent on the disease model, subsequent studies were performed in wild-type mice. Mice were injected into the right striatum with AAVsh2.4-GFP, AAVmi2.4-GFP, or AAVmi2.4mis-GFP and killed 4 months after injection. The time course, volume, and titer were identical to those used in our earlier shRNA studies (Fig. 2). QPCR performed on RNA isolated from mouse striata showed a statistically significant reduction of HDh mRNA (≈70%) after treatment with either sh2.4- or mi2.4-expressing vectors, compared with uninjected striata or striata treated with mi2.4mis [F(2, 8) = 77.6, P < 0.001 for post hoc analyses comparing sh2.4 and mi2.4 vs. uninjected and mi2.4mis] (Fig. 5A). Importantly, the degree of HDh mRNA silencing between sh2.4 and mi2.4 was similar and not significantly different (P > 0.05). Additional QPCR analyses were performed on these samples to measure CD11b mRNA, a readout for microglial activation, as an initial assessment for toxicity. Striata treated with sh2.4 showed nearly a 4-fold increase of CD11b mRNA relative to uninjected striata, whereas mi2.4- and mi2.4mis-treated striata showed only minimal induction [F(2, 8) = 23.6, P < 0.001 for post hoc analyses comparing sh2.4 to all other groups] (Fig. 5B). To determine whether these differences in toxicity could be attributed to levels of HD2.4 inhibitory RNAs, we performed Northern blot analysis on the same RNA samples used for the QPCR analyses. Although silencing efficacies between the sh2.4- and mi2.4-treated groups were comparable, Northern blot analysis, probing for the HD2.4 antisense strand, demonstrated considerably more mature HD2.4 antisense RNAs in sh2.4-treated mice relative to mi2.4-treated mice (Fig. 5C). These results corroborate our in vitro findings and correlate the improvement in toxicity with reduced levels of HD2.4 antisense RNA.

Fig. 5.
Artificial miRNAs mitigate striatal toxicity in mice. (A and B) QPCR analyses were performed to measure mouse HDh (A) and CD11b (B) mRNA levels in AAV-RNAi-injected striata harvested 4 months after treatment (NS, not significant). Samples were normalized ...

We further assessed striatal toxicity by histological analyses. Immunolabeling for DARPP-32 expression revealed significant attenuation of striatal toxicity in AAVmi2.4-GFP-injected cohorts relative to AAVsh2.4-GFP-injected mice (Fig. 5D Middle). Moreover, the intense microglial activation (Iba1-positive cells) seen in AAVsh2.4-GFP-injected mice was scarcely present in AAVmi2.4-GFP-injected mice (Fig. 5D Bottom and Fig. S4). Of note, mi2.4mis-treated brains also showed no apparent toxicity by these analyses, whereas HD2.4mis was toxic when delivered as an shRNA (data not shown). Thus, sequences encoding HD2.4 and HD2.4mis were toxic in the setting of an shRNA in the brain, but not in the context of a miRNA scaffold.


Here, we show that some shRNAs cause toxicity in mouse striatum independent of HDh mRNA silencing. Similar to our work, Grimm and colleagues (15) observed acute liver toxicity and mortality in mice after systemic shRNA delivery, which correlated with increased mature antisense RNA levels. However, there are important differences between our findings. First, Grimm et al. found that lowering the vector dose by ≈10-fold significantly improved the lethal effects of some shRNAs on liver function and animal viability. In our studies, reducing the dose led to lower transduction throughout the striatum, but did not abrogate toxicity. Second, the data by Grimm and colleagues show significant buildup of shRNA precursors in liver cells. They attributed the liver toxicity, in part, to the saturation of endogenous RNAi export machinery. In our work, we detected abundant levels of unprocessed shRNAs in vitro, but, interestingly, low to undetectable levels in vivo. This finding suggests that export was likely not limiting in our studies. Alternatively, the striatal toxicity may be caused by the buildup of antisense RNAs and subsequent off-target silencing of unintended mRNAs. Our data on sh8.2 also are consistent with this; sh8.2 was not toxic when delivered at the same dose as sh2.4 and sh30.1. Although silencing activity was similar among the three shRNAs, levels of mature product for sh8.2 were significantly lower.

We found that moving the HD2.4 and HD2.4mis sequences, both of which caused toxicity in the context of a shRNA, into a miRNA scaffold significantly reduced neurotoxicity within the striatum with no sacrifice in gene-silencing efficacy. We correlated this positive effect to lower steady-state levels of mature antisense RNAs processed from the artificial mi2.4 relative to sh2.4. Whether this disparity in expression levels results from the differences in transcription or the stability between shRNAs and artificial miRNAs remains unknown. However, the latter provides a more likely explanation because sh2.4 and mi2.4 are expressed from the same mouse U6 promoter and only differ in size by ≈100 nucleotides.

In addition to improved safety profiles, artificial miRNAs are amenable to Pol-II-mediated transcription. Conversely, shRNAs have limited spacing flexibility for expressing shRNAs from Pol-II-based promoters (16). This advantage of miRNA-based systems allows for regulated and cell-specific expression of inhibitory RNAs. These versatile expression strategies advance the application of artificial miRNAs as biological tools and may further limit potential toxicity in therapeutic applications.

In some diseases, it is possible to specifically target disease-linked SNPs that exist on the mutant transcript (17, 18). For HD, however, no prevalent SNP has been reported. Because earlier work showed that a minimum of 50% huntingtin expression is required to offset the embryonic lethality noted in huntingtin-null mice (19), knowing the consequences of reducing huntingtin expression in adult brain is important to moving non-allele-specific RNAi forward as a HD therapy. Our data with sh8.2 and mi2.4 are encouraging and suggest that the mammalian brain can tolerate >50% reduction in HD mRNA for 4 months, the last time point studied. The long-term safety and efficacy of sh8.2 is currently being tested in a study including histochemical, biochemical, and behavioral readouts in CAG140 HD mice.

In summary, we show that reducing HDh mRNA levels in adult mammalian brain is tolerated. We also make the important observation that the toxicity of shRNAs after their expression in brain could be alleviated by moving the inhibitory RNA sequences into an artificial miRNA scaffold. Thus, miRNA-based approaches are more suitable for achieving RNAi in the brain to address basic research questions or develop disease therapies.

Materials and Methods

Expression Vectors and AAV.

shRNA expression cassettes were generated by PCR as described (8) and cloned into pCR-Blunt-II TOPO vectors (Invitrogen). Each candidate shRNA expression cassette consisted of a mouse U6 promoter, an shRNA [targeting huntingtin sequences, mismatch control sequences (containing four base pair changes relative to the respective huntingtin shRNAs), or E. coli β-gal; shLacZ], and an RNA polymerase III termination sequence (six thymidine nucleotides). For artificial miRNAs, siRNA sequences based on HD2.4 or HD2.4mis were embedded into an artificial miRNA scaffold comparable to human miR-30 to generate mi2.4 and mi2.4mis (general structure shown in Fig. 4A). The artificial miRNA stem loops were cloned into a mouse U6 expression vector so that >30 nt (5′ and 3′) flank the stem loop in the transcribed product.

AAV shuttle plasmids pAAVsh2.4-GFP, pAAVsh2.4mis-GFP, pAAVsh8.2-GFP, pAAVsh30.1-GFP, pAAVmi2.4-GFP, and pAAVmi2.4mis-GFP contain the respective RNAi expression cassettes driven by the mouse U6 promoter. The AAV shuttles also contained a hrGFP gene under the control of the human cytomegalovirus immediate-early gene enhancer/promoter region, a chimeric human β-globin eGFP expression cassette followed by the splice donor/human Ig splice acceptor site, and a bovine growth hormone poly (A) signal. These transcriptional units are flanked at each end by AAV serotype 2 145-bp inverted terminal repeat sequences. The transpackaging plasmids, pBSHSPR2C1, were constructed as follows: genomic DNA was extracted from AAV1 (American Type Culture Collection), and the cap coding sequence was amplified by PCR using Pfx polymerase (Invitrogen). The AAV2 cap gene was excised from the AAV2 helper plasmid pBSHSPRC2.3 and replaced with the amplified AAV1 cap sequence by using a Swa I restriction site in the rep/cap intergenic junction and a BsrG I site engineered just upstream of the AAV2 poly(A) signal. The resulting transpackaging construct, pBSHSPR2C1, contains the AAV2 rep gene under the control of a minimal eukaryotic promoter and the AAV1 cap ORF positioned between the AAV2 rep/cap intergenic junction and the AAV2 poly(A) signal. The plasmid pAd Helper 4.1 expresses the E2a, E4-orf6, and VA genes of adenovirus type 5 (Ad5) for AAV amplification.

Recombinant AAV vectors were produced by a standard calcium phosphate transfection method in HEK 293 cells by using the Ad helper, transpackaging, and AAV shuttle plasmids as described (20). Vector titers were determined by real-time PCR and were between 5 and 20 × 1012 DNase-resistant particles per ml. Vector infectivity was assessed in a TCID50 assay by using the HeLa-based B50 cell line (21).


All animal protocols were approved by the Institutional Animal Care and Use Committee at the University of Iowa. CAG140 heterozygous knockin mice (10) and wild-type littermates were bred and maintained in the animal vivarium at the University of Iowa. Mice were genotyped and repeat length identified by separate PCRs using primers flanking the CAG repeat. Mice were housed in groups of either two or three per cage and in a controlled temperature environment on a 12-h light/dark cycle. Food and water were provided ad libitum.

AAV Injections.

CAG140 knockin or wild-type mice were injected with AAVsh RNAs or AAV-miRNAs (at the indicated titer) at 5 weeks of age and killed at 4 months after injection. Procedures were performed as reported previously (8) with the following exceptions. In the initial study, 5-μl injections of either AAVsh2.4GFP, AAV30.1sh-GFP, AAVsh8.2-GFP, or AAV-GFP were made bilaterally into striata (coordinates: 0.86 mm rostral to bregma, ±1.8 mm lateral to midline, 3.5 mm ventral to the skull surface). For the miRNA/shRNA comparison study, 5-μl injections of vector were injected unilaterally. Injection rates for all studies were 0.2 μl/min. Mice used in histological analyses were anesthetized with a ketamine/xylazine mix and transcardially perfused with 20 ml of 0.9% cold saline, followed by 20 ml of 4% paraformaldehyde in 0.1 M PO4 buffer. Brains were removed and postfixed overnight, and 40-μm thick sections were collected. Mice used for molecular analyses were perfused with 20 ml of 0.9% cold saline, and brain was removed and blocked into 1-mm-thick coronal slices. Tissue punches were taken by using a tissue corer (1.4 mm in diameter). All tissue punches were flash frozen in liquid nitrogen and stored at −80°C until used.

Molecular Studies.

For in vitro shRNA screening, shRNA expression plasmids were transfected (Lipofectamine 2000; Invitrogen) into human HEK 293 cells or mouse C2C12 cells, which naturally express full-length human or mouse huntingtin, respectively. Huntingtin levels were assessed by protein dot blot (anti-huntingtin primary antibody MAB2166, 1:5,000; Chemicon) or Western blot (protein loading control, anti-β-catenin, 1:4,000; AbCam). Knockdown also was assessed by QPCR using a human huntingtin-specific TaqMan primer/probe set with normalization to a human GAPDH primer/probe set. This QPCR strategy also was used to evaluate HD knockdown mediated by sh2.4 and mi2.4 in Fig. 4B.

For in vivo QPCR analyses, tissue was dissected from GFP-positive striatum, and relative gene expression was assessed by using TaqMan primer/probe sets for mouse HDh, CD11b, and β-actin. All values were quantified by using the ΔΔCT method (normalizing to β-actin) and calibrated to either AAV-GFP-injected striata (screening study) or uninjected striata (miRNA-shRNA comparison study).

For Northern blot analyses, tissue was dissected from GFP-positive striatum. RNA was harvested by TRIzol reagent and RNA (1–5 μg and 15 μg for in vivo and in vitro studies, respectively) was resolved on 15% polyacrylamide/urea gels, and RNA was visualized by ethidium bromide staining and UV exposure to assess loading and RNA quality. Samples were then transferred to Hybond-N+/XL membranes (Amersham Pharmacia) and UV cross-linked. Blots were probed with 32P-labeled oligonucleotides at 30–36°C overnight, washed in 2× SSC at 30–36°C, and exposed to film.

For in vivo Western blot analysis, tissue was dissected from GFP-positive striatum and lysed in 150 μl of lysis buffer, and protein level was quantified with the DC protein assay (Bio-Rad). Then 10 μg of total protein was separated on 8% SDS polyacrylamide gel before transferring to a 0.45-μm PVDF membrane. The membrane was blocked with 2% milk in PBS-Tween 20 (0.05%) and incubated with either an anti-huntingtin antibody (1:5,000; Chemicon) or an anti-β-actin antibody (1:10,000; Sigma), followed by a conjugated goat anti-mouse secondary antibody (1:10,000; Jackson ImmunoResearch) and an ECL-Plus substrate (Amersham Biosciences), and then exposed to film.

Immunohistochemical Analyses.

Briefly, 40-μm-thick, free-floating coronal brain sections were processed for immunohistochemical visualization of striatal neurons (DARPP-32, 1:100; Cell Signaling Technology) and microglia (Iba1, 1:1,000; WAKO) by using the biotin-labeled antibody procedure. Primary antibody incubations were carried out for 24 h at room temperature. Sections were incubated in goat anti-rabbit biotinylated IgG secondary antibodies (1:200; Vector Laboratories) for 1 h at room temperature. In all staining procedures, deletion of the primary antibody served as a control. Sections were mounted onto Superfrost Plus slides and coverslipped with Gelmount (Biomeda). Images were captured by using an Olympus BX60 light microscope and DP70 digital camera, along with an Olympus DP Controller software.

Statistical Analyses.

All statistical analyses were performed by using SigmaStat statistical software (SYSTAT). QPCR analyses for huntingtin and CD11b expression were performed by using a one-way ANOVA, as was Northern blot densitometry analysis. Upon a significant effect, Bonferroni post hoc analyses were performed to assess for significant differences between individual groups. Western blot densitometry analysis was performed by using a two-tailed Student's t test. In all cases, P < 0.05 was considered significant.

Figure Preparation.

All photographs were formatted with Adobe Photoshop software, all graphs were made with Prism Graph software, and all figures were constructed with Adobe Illustrator software.

Supplementary Material

Supporting Information:


We thank the B.L.D. and McCray laboratories for feedback and discussion. This work was supported by National Institutes of Health Grants NS-50210, HD-44093, DK-54759, and NS-592372; the Hereditary Disease Foundation; and the Roy J. Carver Trust.


Conflict of interest statement: B.L.D. was a consultant for Sirna Therapeutics, Inc.

This article contains supporting information online at www.pnas.org/cgi/content/full/0801775105/DCSupplemental.


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