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Keywords:

  • Basal ganglia;
  • cerebral cortex stereology;
  • Huntington's mouse model;
  • neurodegeneration;
  • neurotrophin;
  • striatum;
  • thalamus

Abstract

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information

Patients with Huntington's disease (HD) and transgenic mouse models of HD show neuronal loss in the striatum as a major feature, which contributes to cognitive and motor manifestations. Reduced expression of the neurotrophin brain-derived neurotrophic factor (BDNF) in striatal afferents may play a role in neuronal loss. How progressive loss of BDNF expression in different cortical or subcortical afferents contributes to striatal atrophy and behavioral dysfunction in HD is not known, and may best be determined in animal models. We compared age-dependent alterations of BDNF mRNA expression in major striatal afferents from the cerebral cortex, thalamus and midbrain in the R6/2 transgenic mouse model of HD. Corresponding changes in striatal morphology were quantified using unbiased stereology. Changes in motor behavior were measured using an open field, grip strength monitor, limb clasping and a rotarod apparatus. BDNF expression in cortical limbic and midbrain striatal afferents is reduced by age 4 weeks, prior to onset of motor abnormalities. BDNF expression in motor cortex and thalamic afferents is reduced by 6 weeks, coinciding with early motor dysfunction and reduced striatum volume. BDNF loss in afferents progresses until death at 13–15 weeks, correlating with progressive striatal neuronal loss and motor abnormalities. Mutant huntingtin protein expression in R6/2 mice results in progressive loss of BDNF in both cortical and subcortical striatal afferents. BDNF loss in limbic and dopaminergic striatal inputs may contribute to cognitive/psychiatric dysfunction in HD. Subsequent BDNF loss in cortical motor and thalamic afferents may accelerate striatal degeneration, resulting in progressive involuntary movements.

Huntington's disease (HD) is a progressive autosomal dominant neurodegenerative disorder primarily affecting basal ganglia function. The latter consist of several functionally related subcortical nuclei, which interact with the cerebral cortex, thalamus and brainstem to modulate motor, cognitive and emotional behaviors (Tepper et al. 2007). HD gene located on the chromosome 4 contains an expansion of the normal number of cytosine-adenine-guanine (CAG; glutamine) triplet repeats (generally >35) resulting in production of a mutant form of the huntingtin protein (htt) (Huntington's disease Collaborative Research Group 1993; Reiner et al. 2011). Increased CAG repeat size is associated with more severe disease and earlier age of onset (Duyao et al. 1993). Motor disturbances are characterized by uncontrolled hyperkinetic movements, which can evolve to severe hypokinesia at terminal stages. Cognitive and emotional manifestations such as deficits in frontostriatal executive function, anxiety and depression may precede overt manifestations (Paulsen 2011). Although mutant htt protein is expressed ubiquitously throughout the brain, the most striking neurodegenerative changes are preferentially observed in the medium spiny projection neurons of the striatum (Reiner et al. 1988; Rosas et al. 2003; Stack et al. 2005; Vonsattel et al. 1985). The cause of this predilection for striatal medium spiny neurons is unknown, but excitotoxic and anterograde neurotrophic mechanisms are proposed (Kim et al. 2011; Raymond et al. 2011; Zuccato et al. 2010).

The neurotrophic hypothesis of HD proposes impaired production of brain-derived neurotrophic factor (BDNF), a member of the neurotrophin family of growth factors, contributes to degeneration of neurons in the striatum (Saudou & Humbert 2008; Zuccato et al. 2010). The striatum contains few if any BDNF mRNA-expressing neurons, but contains high levels of BDNF protein (Altar et al. 1997; Conner et al. 1997; Hofer et al. 1990). In addition to corticostriatal glutamatergic afferents, the striatum also receives massive glutamatergic inputs from the parafascicular (Pf) thalamic nucleus (Berendse & Groenewegen 1990; Sadikot et al. 1992a,1992b; Smith et al. 2009) and dopaminergic (DA) inputs from the substantia nigra pars compacta (SNc) (Parent & Hazrati 1995; Parent & Parent 2010; Smith et al. 1994). Interestingly, the cerebral cortex, Pf and SNc all express high levels of BDNF mRNA (Conner et al. 1997). Lesions in cortical, nigral or thalamic afferents reduce BDNF protein concentrations in the striatum, suggesting anterograde transport of BDNF in striatal afferents (Altar et al. 1997; Conner et al. 1997; Sadikot et al. 2005). BDNF protein content in the striatum is reduced in both HD (Ferrer et al. 2000; Gauthier et al. 2004) and in animal models of HD (DeMarch et al. 2008; Giralt et al. 2011; Peng et al. 2008; Spires et al. 2004). Reports of BDNF levels in cerebral cortex of HD patients vary from no significant loss in sampled temporal or parietal cortices to significant reduction in both mRNA and protein (Ferrer et al. 2000; Gauthier et al. 2004; Zuccato et al. 2008, 2010). A reduction in cortical BDNF mRNA is detected in transgenic mouse models of HD (Huntington's disease Collaborative Research Group 1993; Luthi-Carter et al. 2002; Zuccato & Cattaneo 2007; Zuccato et al. 2001), although a systematic regional assessment in motor- and limbic-related areas has not been performed. Furthermore, little information is available regarding the levels of BDNF mRNA expression in other major sources of striatal afferents in HD or in animal models. Exploring BDNF expression in striatal afferents, motor behavior and changes in striatal morphology in the same animal model of HD would allow for better understanding of how neurotrophins may participate in mechanisms of neuronal degeneration, further insight into the relationship between morphology and behavior, and provide important normative data for preclinical therapeutic strategies.

The objective of this study is to characterize in detail the time course of neostriatal neuronal degeneration in the R6/2 mouse model of HD, and determine its relationship to motor impairment and reduced BDNF synthesis in major striatal afferents. The R6/2 mouse is the most widely used animal model for studies of the biology and treatment of HD (Gil & Rego 2009; Kim et al. 2011). Initially produced to include exon 1 of the human HD gene with ∼150 CAG repeats (Mangiarini et al. 1996), several lines with higher or lower numbers of CAG repeats are now in use (Cummings et al. 2012; Menalled et al. 2009). Here, we take advantage of a widely available R6/2 line with ∼110 CAG repeats.

We show a close temporal relationship between onset of loss of BDNF expression in motor cortices and the Pf, striatal degeneration and deterioration in motor behavior. Loss of BDNF mRNA in limbic cortices and the SNc precedes decreases seen in motor areas, and may correspond to early deficits in learning and memory. We suggest that the R6/2 mouse models many of the cardinal motor manifestations and morphological features of HD. Our results also indicate that in addition to corticostriatal afferents, thalamostriatal and nigrostriatal projections may play an important role in maintaining trophic support to striatal neurons in HD, and contribute to the pathophysiology of progressive motor abnormalities.

Materials and methods

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information

Animals

The experiments were performed using male carrier (R6/2) and noncarrier (WT) mice from the same breeding colony (B6CBA-Tg(HDexon1)62Gpb/3J) maintained at the Montreal Neurological Institute facility in accordance with the standards of the Canadian Council on Animal Care. The breeders, males and ovary-transplanted females, were from the same strain, obtained from a line maintained at The Jackson Laboratory (Bar Harbor, ME, USA) involving a C57BL/6 and CBA background (stock number 006494). Offspring were genotyped by Laragen (Culver City, CA, USA), CAG repeat lengths of 105–112 were confirmed and cohorts of R6/2 and WT littermates were evaluated. All testing was performed during the light phase of a 12-h light–dark cycle. Weight gain and survival of female R6/2 mice are significantly less than male R6/2 mice (Stack et al. 2005; Wood et al. 2011). Only males were used in order to reduce variability related to sex (Menalled et al. 2009). Animal numbers used for different behavioral tests, history of reuse after another behavioral test and the number of animals used in morphological analysis are summarized in Tables S1 and S2, Supporting Information.

Evaluation of motor behavior

Spontaneous activity in an open field

Spontaneous locomotor activity was measured in an open field using a video tracking system (Videotrack, Viewpoint Life Sciences, Montreal, Canada) with infrared backlighting (Bailoo et al. 2010). Locomotor activity measured was categorized as follows: inactivity or nonambulatory movements (<5 cm/second); moderate movement speed (between 5 and 20 cm/second) or fast movements (>20 cm/second). Distance refers to the total distance traveled by the animal during the testing period, and was measured separately for moderate and fast activity periods. Duration is the total duration spent in inactivity or nonambulatory movements, moderate activity or fast activity. Locomotor activities were monitored continuously during the 2-h testing period, with output intervals of 3 min.

Grip strength

Grip strength (Columbus Instruments, Columbus, OH, USA) determines the peak tension (T-PK) and peak compression (C-PK) developed by a rodent as it instinctively grips a wire mesh grid. The examiner attempts to overcome the grip by gently pulling at the base of the tail (Hockly et al. 2002). The device provides a digital readout of maximal force generated, expressed in grams (g) or Newton. Seven consecutive trials were performed with each mouse, separated by a rest period on the table of 5 seconds. The observer was blind to genotype. The mean of at least five successful trials was taken for analysis. Peak tension forces (T-PK) generated by the forelimbs, or all limbs as mice pulled on the wire mesh, or peak compression forces (C-PK) generated by the hindlimbs were assessed.

Clasping score

In this test, mice were suspended by the tail at a height of at least 30 cm, for two 20-second intervals, and limb movements were videotaped. Clasping was defined as a retraction of a limb toward the body, and rated by an observer blind to genotype in order to provide a semiquantitative index of abnormal involuntary movements. Clasping was rated as mild when the forelimb or hindlimb retracted toward the midline but did not reaching the midline, and the contraction was not sustained. Moderate clasping was a high-amplitude limb retraction to or beyond the midline, but not sustained. Severe clasping was a high-amplitude limb retraction sustained for more than 15 seconds. Quality of clasping at each limb was graded as: none = 0, mild = 0.25, moderate = 0.5, severe and constant = 0.75, for a maximal clasping score of 3. The score from the two tail suspension trials was averaged and recorded.

Rotarod test

A rotarod apparatus (Columbus Instruments) was used to measure limb coordination and balance using an incremental fixed speed protocol (Carter et al. 1999; Monville et al. 2006). At the beginning of the experiments, mice were trained on the rotarod during 3 consecutive days. Each day mice had four trials at a constant speed (24 r.p.m.) for a maximum of 60 seconds, to obtain a training baseline. On the 4th day mice received three to four trials at nine increasing speeds (5–44 r.p.m.) for a maximum of 5 min each, with a 20-min rest period between each trial (Carter et al. 1999). The mean latency to fall at each speed, at different ages, was used in the analysis. The observer was blind to genotype.

Tissue preparation and stereology

Animals at different ages were deeply anesthetized and then perfused transcardially with 0.9% heparinized saline followed by 4% paraformaldehyde in phosphate buffer (0.1 m, pH 7.4). Brains were removed and immersed for 24 h in the same fixative and then passed through a series of graded concentrations of phosphate-buffered sucrose solutions to a final concentration of 30%. Brains were cut in the coronal plane at 40 µm using a freezing microtome and free-floating sections were collected serially in six vials containing phosphate-buffered saline (PBS, 0.1 m, pH 7.4). One of the six sets of sections was mounted out of distilled water onto slides, stained with 0.1% Cresyl Violet (Nissl stain) and coverslipped using Permount (Fisher, Fairlawn, NJ, USA).

Unbiased stereology was carried out on Nissl-stained coronal sections using the Stereo Investigator program (Microbrightfield, Willston, VT, USA) and an Olympus BX-40 microscope equipped with a motorized XYZ stage. The optical fractionator and nucleator were used as stereology probes to obtain unbiased estimates of the total number of neurons and estimates of cell size, respectively, as detailed in our previous work (Luk et al. 2003; Rymar et al. 2004). The neostriatum was delineated according to defined boundaries (Sadikot & Sasseville 1997) using a stereotaxic atlas of the mouse brain (Franklin & Paxinos 2008). Briefly, the selection includes levels throughout the neostriatum including regularly spaced sections caudal and rostral to the decussation of the anterior commissure. The dorsal, medial and lateral limits of the neostriatum are well defined (Franklin & Paxinos 2008). Ventrally, the neostriatum interfaces with the amygdala and substantia innominata in its postcommissural part and with the nucleus accumbens in its precommissural division. The ventral limit of the striatum at the postcommissural part is well delineated on Nissl stains. However, the ventral limit of the neostriatum at its precommissural part is an arbitrary interface. At the two precommissural levels analyzed, we therefore delimit the dorsal striatum from the nucleus accumbens with a line that extends from above the ventralmost part of the lateral ventricle medially, to the tapered external capsule laterally, at an angle of 25–30° below the axial plane. Section thickness was assessed and neuronal counts were performed under oil immersion using a 100× objective with a high aperture lens (NA 1.4). The systematic random sampling grid size was 500 × 500 µm, and mean section thickness was measured at every fifth sampling site. The Cavalieri estimator was used to measure the striatal reference volume. The optical fractionator brick size was 60 × 60 µm, with a mean section thickness of 12 µm, and guard zones of 1 µm at the top and bottom surface, resulting in a dissector height of 10 µm. Neurons were defined as Nissl-stained profiles measuring at least 7 µm in diameter, with a lighter cytoplasm containing organelles. The four-ray nucleator was used to estimate neuronal size (Gundersen et al. 1988). The observer was blind to genotype.

In situ hybridization

A second set of 1/6 sections from each animal was processed for in situ hybridization under RNAase-free conditions. [35S]UTP-labeled cRNA probes with either sense or antisense orientation were synthesized by in vitro transcription from a full-length cDNA clone encoding rat BDNF (MGC105254). The single-stranded cRNA probe was synthesized and labeled using Riboprobe Kit of Promega (Promega, Madison, WI, USA) with T7 RNA polymerase for antisense and [33S]UTP (Perkin Elmer Inc., Woodbridge, Ontario, Canada). The sense single-stranded RNA probe was synthesized with SP6 RNA polymerase and [33S]UTP. After short fixation of 5 min in 4% paraformaldehyde (PFA)/PBS, free-floating sections were washed in PBS and incubated for 10 min in proteinase K/PBS, followed by second fixation in 4% PFA/PBS during 10 min. After washing with PBS and ddH2O, sections were incubated for 10 min in 0.25% acetic anhydride in 2% triethanolamine and then washed in saline-sodium citrate for 5 min. In situ hybridization was performed at 58°C overnight in a standard hybridization buffer containing 50% formamide as previously described (Beaudry et al. 2000). Following stringency washes, sections were mounted onto superfrost plus slides, air-dried and dehydrated during 2 min in 30%, 60% and 100% ethanol. The slide-mounted tissue sections were then air-dried and exposed to radioactive-sensitive films (Kodak, Biomax MR, New Haven, CT, USA) for 5 days at room temperature. Films were scanned, and analysis of cerebral cortex, thalamus and midbrain sections in R6/2 and WT sections was performed using public domain National Institutes of Health image analysis software (NIH ImageJ, v1.45s, Bethesda, MD), normalizing optical density using the corpus callosum as a reference for background activity.

Statistical comparisons for differences in behavior and morphology between R6/2 carrier mice and WT mice at different ages were performed using analysis of variance (anova) followed by post hoc analysis using the Fisher PLSD test (Stat-View, Abacus Corporation, Baltimore MD, USA). Data are expressed as a mean ± standard error of the mean (SEM), and P < 0.05 is considered significant.

Results

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information

Motor behavior

Locomotor activity
Total distance traveled

When the total distance traveled during the 2-h test interval by R6/2 (n = 197) and WT (n = 135) mice was compared at different ages, significant differences were noted with respect to main effects and their interaction [F(genotype)1,310 = 49.19, P < 0.0001; F(age)10,310 = 3.04, P < 0.0001; F(genotype × age)10,310 = 2.185, P = 0.01]. Further analysis showed that the total distance traveled by R6/2 mice was comparable to WTs at 4 and 6 weeks, and R6/2 mice showed a trend to reduced activity of marginal significance at 7 weeks (F1,33 =3.82, P = 0.05; Fig. 1a). Motor activity in R6/2 mice declined progressively at 8–15 weeks (Fig. 1a) compared with age-matched WT controls, deteriorating to 58% by 15 weeks (F1,15 =18.32, P < 0.001).

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Figure 1. Spontaneous locomotor activity of R6/2 mice compared with WT controls at different ages in a video-tracked open field. (a) Total distance traveled by R6/2 mice compared with WTs. (b) Distance traveled at different movement speeds. (c) Duration of locomotor activities at different movement speeds or during nonambulatory behavior. (d) Sample video traces generated during 2-h sessions at 4, 8 and 12 weeks illustrate patterns of spontaneous locomotor activity in the open field. Trace colors code for different movement speeds. Although activity traces of 4-week-old R6/2 mice are virtually identical to age-matched WT controls, marked deterioration is seen by 8 and 12 weeks, with mice favoring movement close to the edges of the open field. *P < 0.05, **P < 0.01 or P < 0.005, ***P < 0.001, R6/2 vs. WT.

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Distance traveled at different movement speeds

The contribution of distances traveled at different movement speeds to the observed differences in total distance traveled was analyzed in R6/2 and WT mice. Results in R6/2 mice are expressed as a percentage of controls (Fig. 1b). There was no difference in distances traveled at different movement speeds in 4-week-old R6/2 mice (n = 6) compared with WT controls (n = 7) [F(genotype)1,11 = 0.004, P = 0.95; F(genotype × distance at different speeds)2,22 = 0.12, P = 0.88]. Six-week-old R6/2 mice (n = 38) showed no difference compared with WTs (n = 11) in total distance traveled at different movement speeds [F(genotype)1,47 = 2.15]; however, there was a significant interaction effect between genotype and distance traveled at different movement speeds [F(genotype × distance at different speeds)2,94 = 5.93, P = 0.003]. Post hoc analysis showed a significant reduction of fast activity at 6 weeks [50% of control (F1,47 = 8.28, P = 0.006)] (Fig. 1b), but there was no significant difference at other speeds. Seven-week-old R6/2 mice (n = 23) showed marginal significance for distance traveled compared with WT (n = 12) mice, but there was a significant interaction effect [F(genotype)1,33 = 3.82, P = 0.05; F(genotype × distance at different speeds)2,66 = 5.78, P = 0.005]. At this age, R6/2 mice show significant reduction in distance traveled during both fast and moderate locomotor activity [70% of control (F1,33 = 4.28, P = 0.04)] (Fig. 1b). Distances traveled by R6/2 mice during both moderate and fast activity declined progressively at each week from 9 (n + 26) to 15 (n = 7) weeks. For example, 9-week-old R6/2 mice (n = 26) showed a significant reduction in distance traveled during both moderate [72% of control (F1,35 = 5.27, P = 0.02)] and fast activity [63% of control (F1,35 = 4.35, P = 0.04)] compared with WT mice (n = 11). By 15 weeks (Fig. 1b), R6/2 mice (n = 7) showed severe reduction in distance traveled during both moderate activity [32% of control (F1,15 = 19.32, P = 0.0005)] and fast activity [5% of control (F1,15 = 11.28, P = 0.004)] compared with WT mice (n = 10).

Duration of locomotor activity at different movement speeds

The time spent in movement at fast, moderate and nonambulatory activities was compared in R6/2 and WT mice at different ages. There was no significant difference in time spent at different movement speeds in 4-week-old R6/2 (n = 6) compared with WT (n = 7) mice [F(genotype)1,11 = 0.85, P = 0.38; F(genotype × duration at different speeds)2,22 = 0.92, P = 0.42]. Although total distance traveled remained unchanged compared with controls (Fig. 1a), 6-week-old R6/2 mice (n = 38) showed a significant difference in time spent at moderate and fast movements speeds compared with WT controls (n = 11) [F(genotype)1,47 = 5.31, P = 0.02]. Interaction between genotype and duration of locomotor activity was not significant at this age [F(genotype × duration at different speeds)2,94 = 1.27, P = 0.28]. Reduced duration of fast activity was apparent in 6-week-old R6/2 mice (n = 38) compared with WT (n = 11) controls [50% of control (F1,47 = 7.18, P = 0.01)] (Fig. 1c). At 7 weeks, there was a significant interaction between genotype and duration of locomotor activity at different speeds [F(genotype × duration at different speeds)2,35 = 4.21, P = 0.01], an effect also noted at older ages up to 15 weeks [F(genotype × duration at different speeds)2,15 = 18.42, P < 0.0001]. Post hoc analysis at 7 weeks showed a marginal reduction in duration of moderate activity [75% (F1,33 = 4.08, P = 0.05)] in R6/2 mice (n = 23) compared with WT controls (n = 12; Fig. 1c), with concomitant reduction in total distance traveled (Fig. 1a). R6/2 mice older than 9 weeks showed a progressive reduction in duration of moderate and fast activity compared with WT controls. By 15 weeks, R6/2 mice (n = 7) spent significantly reduced times in moderate [47% of control (F1,15 = 18.39, P = 0.0006)] and fast activity [4% of control (F1,15 = 12.63, P = 0.002)] compared with WT mice (n = 10; Fig. 1c). The duration of time spent in nonambulatory movements is mildly increased in R6/2 mice (n = 23) from 7 weeks [110% of control: n = 12 (F1,33 = 4.29, P = 0.04)] onward, possibly suggesting hypokinesia, and increase in grooming or stereotyped motor behaviors (Fig. 1c). To illustrate the pattern of locomotor activity in the open field, video traces generated during sample 2-h sessions are shown at 4, 8 and 12 weeks (Fig. 1d). Although activity traces of 4-week-old R6/2 mice are virtually identical to age-matched WT controls, marked deterioration is seen by 8 and 12 weeks, with mice favoring movement close to the edges of the open field (Fig. 1d).

Grip strength

R6/2 mice showed progressive loss of grip strength compared with WT controls. When mice held the wire mesh grid using all limbs, a significant difference is detected between R6/2 (n = 129) and WT mice (n = 54) at different ages [F(genotype)1,163 = 180.79, P < 0.0001; F(age)9,163 = 3.68, P = 0.0003; F(genotype × age)9,163 = 10.68, P < 0.0001]. Impaired grip strength was first detected at 9 weeks of age in R6/2 mice (n = 12) compared with WT (n = 5) controls (F1,15 = 16.45, P = 0.001; Fig. 2a). Much of this loss is accounted for by impaired forelimb strength [F(genotype)1,163 = 76.97, P < 0.0001; F(age)9,163 = 6.14, P < 0.0001; F(genotype × age)9,163 = 3.02, P = 0.002], which was progressively lost beginning at 9 weeks (F1,15 = 7.254, P = 0.01; Fig. 2b). Hindlimb grip strength was also impaired [F(genotype)1,163 = 42.32, P < 0.0001; F(age)9,163 = 7.27, P < 0.0001; F(genotype × age)9,163 = 2.06, P = 0.03]. However, hindlimb grip strength was lost later at 10 weeks (R6/2: n = 14 and WT: n = 5; F1,17 = 9.14, P = 0.007; Fig. 2c).

image

Figure 2. Grip strength of R6/2 mice compared with WT controls at different ages. Grip strength generated when mice held the wire mesh grid using all limbs (a) or using only the forelimbs (b). Grip strength generated when mice held the wire mesh grid using the hindlimbs (c). (d) Body mass of male R6/2 compared with WT mice at different ages. The reduction of weight in male R6/2 mice was evident by age 9 weeks. When normalized for weight, muscular force reduction declined significantly after 10 weeks, when grip strength was measured with all limbs touching the grid (data not shown). Muscular force assessed by grip strength is presented as the mean of T-PK or C-PK for the force of forelimbs, hindlimbs or all limbs ± SEM. *P < 0.05, **P < 0.01 R6/2 vs. WT.

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R6/2 mice (n = 12) also showed significantly impaired weight gain [F(genotype)1,163 = 161.59, P < 0.0001; F(age)9,163 = 19.26, P < 0.0001; F(genotype × age)9,163 = 9.15, P < 0.0001] beginning at 9 weeks compared with WT controls (n = 5; F1,15 = 10.08, P = 0.006; Fig. 2d). A significant reduction of weight-normalized grip strength was first evident in 11-week-old R6/2 mice (n = 14) compared with WT controls (n = 5) [86% of control (F1,17 = 8.43, P = 0.009)], with all limbs pulling on the grid, and remained impaired until 15 weeks (data not shown). Weight-normalized forelimb grip strength of R6/2 mice was also virtually identical to WT controls at 10 weeks and significantly reduced at 11 weeks (R6/2: n = 14 and WT: n = 5) [84% of control (F1,17 = 4.18, P = 0.05)] and 13 weeks (R6/2: n = 15 and WT: n = 6) [85% of control (F1,19 = 6.54, P = 0.01)]. No significant difference was observed in weight-normalized hindlimb grip strength at any age in R6/2 mice compared with WT controls (4–15 weeks, P > 0.05). If normalized for weight, pulling using hindlimbs alone therefore appears to be a less sensitive indicator in R6/2 mice, but pulling using the forelimbs or all four limbs remain sensitive measures of grip strength abnormality.

Involuntary clasping movements

Involuntary clasping movement scores were significantly different in R6/2 mice (n = 77) compared with WT controls (n = 64) at different ages [F(genotype)1,119 = 1320.75, P < 0.0001; F(age)10,119 = 22.79, P < 0.0001; F(genotype × age)10,119 = 28.43, P < 0.0001]. Clasping movements were rare or absent in WT controls (Fig. 3). Mild dystonic movements (clasping score = 0.5) were detectable in R6/2 mice as early as age 4 weeks (R6/2: n = 7 and WT: n = 7; F1,12 = 15.00, P = 0.002) and at 6 weeks (R6/2: n = 22 and WT: n = 14; F1,34 = 3.82, P = 0.05). The severity of these dystonic movements increased progressively with each week in R6/2 mice, and by 9 weeks clasping involved both forelimbs and hindlimbs (clasping score = 2; R6/2: n = 28 and WT: n = 13; F1,39 = 110.84, P < 0.0001). Maximum clasping was detected by age 14 (R6/2: n = 7 and WT: n = 13; F1,18 = 13433.55, P < 0.0001) and 15 weeks (R6/2: n = 4 and WT: n = 13; F1,15 = 1.16E17, P < 0.0001), with severe retraction of all four limbs upon tail suspension (clasping score = 3).

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Figure 3. Clasping score in R6/2 mice at different ages. Clasping score ± SEM, *P < 0.05, **P < 0.0001, R6/2 vs. WT.

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Rotarod performance

A significant decline in rotarod performance was observed at all rotation speeds in 7- to 13-week-old R6/2 mice compared with their respective age-matched WT controls (P < 0.005–<0.0001; Fig. 4), except at 5 r.p.m. and 8 r.p.m. in 7- to 8-week-old mice. Thus, a significant decline in rotarod performance was observed at speeds as low as 5 r.p.m. [F(genotype)1,101 = 44.30, P < 0.0001; F(age)5,101 = 4.09, P = 0.002; F(genotype × age)5,101 = 4.09, P = 0.002] and as high as 44 r.p.m. [F(genotype)1,101 = 74.74, P < 0.0001; F(age)5,101 = 0.68, P = 0.64; F(genotype × age)5,101 = 0.92, P = 0.47] in R6/2 mice (n = 63) compared with their respective age-matched WT controls (n = 50; Fig. 4). However, post hoc analysis did not detect any difference between 7-week-old R6/2 mice (n = 20) and WT controls (n = 6) at 5 and 8 r.p.m. (5 r.p.m.: F1,24 = 1.82, P = 0.19; 8 r.p.m.: F1,24 = 3.81, P = 0.063).

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Figure 4. Incremental fixed speed rotarod performance in R6/2 mice and WT controls at different ages. Data are presented as the mean of latency to fall ± SEM. **P < 0.005, ***P < 0.0005, +<0.05 for 9-week-old animals and ns refers to 7- and 8-week-old animals at 5 and 8 r.p.m.

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When latency to fall of 9-week-old R6/2 mice (n = 14) is compared with 7- to 8-week-old mice (Fig. 4), progressive loss of performance is noted at lower rotation speeds (7 weeks: 5, 8, 15 and 24 r.p.m.; P < 0.05 to P < 0.001). At higher rotation speeds, curves of impaired latency of R6/2 mice are not significantly different at 7 (n = 20) and 9 (n = 14) weeks. No significant difference is observed in latency to fall at any speed when comparisons are made between 9- (n = 14), 11- (n = 13), 12- (n = 5) and 13- (n = 5) week-old R6/2 mice. Indeed, there is virtual overlap of latency to fall curves of R6/2 mice at 11, 12 and 13 weeks, especially at speeds greater than 8 r.p.m., indicating that performance deterioration on this test had reached a plateau. Therefore, a rotation speed of 15 r.p.m. appears to be the most useful in resolving progressive differences in motor performance in 7- to 13-week-old R6/2 mice during a 5-min testing session. Rotation speeds greater than 24 r.p.m. do not provide useful additional information regarding progressive motor loss in R6/2 mice.

Morphology of striatum quantified using unbiased stereology over the R6/2 mouse lifespan

Volume of neostriatum

A significant difference in striatal volume is detected in R6/2 mice (n = 19) compared with controls (n = 22) [F(genotype)1,31 = 31.41, P < 0.0001; F(age)4,31 = 3.79, P = 0.01; F(genotype × age)4,31 = 2.96, P = 0.03]. Post hoc analysis indicates that neostriatal volume was virtually identical in 4-week-old R6/2 (n = 4) and WT (n = 4) mice (F1,6 = 0.28, P = 0.62). However, striatal volume was significantly reduced in 6-week-old R6/2 mice (n = 3) compared with WT controls (n = 4; 88% of control, F1,5 = 10.69, P = 0.02), and decreased progressively as R6/2 animals aged. By 13 weeks, striatal volume in R6/2 mice (n = 4) was 71% of WT (n = 4) controls (F1,6 = 31.69, P = 0.001; Fig. 5a,d).

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Figure 5. Unbiased stereology for quantification of morphological changes in the neostriatum at different ages in R6/2 mice and WT controls. (a) Striatal volume at different ages. (b) Total number of neostriatal neurons. (c) Cross-sectional area of striatal neurons. (d) Photomicrographs of coronal sections of striatum from R6/2 mice and age-matched WT controls at 4, 6, 9 and 13 weeks. Note progressive enlargement of the lateral ventricle with reduced striatal volume. Scale bar: 1500 µm. Data are presented as the mean of values ± SEM. *P < 0.05, **P < 0.01 or P < 0.005, ***P < 0.001 vs. WT.

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The Gundersen coefficients of error (CEs) for the Cavalieri volume estimates (m = 1) at different ages in WT and R6/2 mice, respectively, were as follows: 0.009 ± 0 (mean ± SEM) vs. 0.014 ± 0.0033 at 4 weeks; 0.009 ± 0.0005 vs. 0.009 ± 0.0009 at 6 weeks; 0.008 ± 0.0005 vs. 0.009 ± 0.0003 at 9 weeks; 0.005 ± 0.0004 vs. 0.011 ± 0.0014 at 11 weeks and 0.009 ± 0.0007 vs. 0.009 ± 0.0006 at 13 weeks.

Number of neostriatal neurons

Quantification of the total number of neostriatal neurons in R6/2 mice (n = 19) using unbiased stereology (Fig. 5b) showed no significant difference compared with age-matched WT controls (n = 22) in 4- to 9-week-old animals, but significant differences were noted in older mice. On post hoc analysis a significant decrease in neuron number was first detected at 11 weeks in R6/2 mice (n = 4) compared with age-matched WT (n = 4) mice [87% of control; WT: 1 711 615 ± 71 470 (mean ± SEM) vs. R6/2: 1 502 640 ± 29 308; F1,6 = 7.31, P = 0.03]. The number of neostriatal neurons was further reduced in 13-week-old R6/2 mice (n = 4) compared with WT (n = 4) mice [78% of control; WT: 1 564 184 ± 72 986 vs. R6/2: 1 225 461 ± 41 816; F1,6 = 16.21, P = 0.006].

The Gundersen CEs for neuronal counts (m = 1) at different ages in WT and R6/2 mice, respectively, were as follows: 0.03 ± 0 (mean ± SEM) vs. 0.03 ± 0.003 at 4 weeks; 0.03 ± 0 vs. 0.03 ± 0 at 6 weeks; 0.02 ± 0.002 vs. 0.02 ± 0 at 9 weeks; 0.02 ± 0 vs. 0.03 ± 0.002 at 11 weeks and 0.03 ± 0 vs. 0.03 ± 0 at 13 weeks.

When WT mice were compared at 11 (n = 4) and 13 (n = 4) weeks, no significant difference in the number of striatal neurons is detected (F1,6 = 2.08, P = 0.20). However, a significant difference in the number of striatal neurons was detected between R6/2 mice at 11 (n = 4) and 13 (n = 4) weeks (F1,6 = 29.46, P = 0.001). In young WT mice, an increase in the number (F1,6 = 36.00, P = 0.001) of striatal neurons is noted between 4 (n = 4) and 6 (n = 4) weeks. A marginal increase in the number of striatal neurons is also noted when comparing 4- (n = 4) and 6- (n = 3) week-old R6/2 mice (F1,5 = 6.41, P = 0.05). Glial profiles were also counted at each age. There was no significant difference in glial number at earlier ages (data not shown), but 13-week-old animals showed a 26% increase (434 970 ± 58 505 in WTs vs. 587 292 ± 32 396 in R6/2 mice; F1,6 = 5.19, P = 0.041).

Cross-sectional area of neostriatal neurons

Interestingly, reduction in cross-sectional area of neostriatal neurons [F(genotype)1,31 = 23.20, P < 0.0001; F(age)4,31 = 14.15, P < 0.0001; F(genotype × age)4,31 = 2.85, P = 0.04] preceded neuronal loss (R6/2: n = 19 and WT: n = 22; Fig. 5b,c). The cross-sectional area of striatal neurons was not changed in R6/2 mice at 4 weeks (n = 4; F1,6 = 0.32, P = 0.59) and 6 weeks (n = 3; F1,5 = 5.28, P = 0.07) compared with WT controls (n = 4; Fig. 5c). Reduction in neuronal area was significant in 9- (R6/2: n = 4 and WT: n = 6; 80% of control; F1,8 = 7.69, P = 0.02), 11- (R6/2: n = 4 and WT: n = 4; 76% of control; F1,6 = 7.22, P = 0.03) and 13-week-old R6/2 mice (R6/2: n = 4 and WT: n = 4; 77% of control; F1,6 = 11.14, P = 0.01). As a caveat, nucleator-based estimates of cell size assume isotropic shape of the object measured. The results can therefore be biased if striatal neurons have an anisotropic orientation, unless randomly oriented sections are used (Gundersen et al. 1988). In the nervous system, analysis on independent uniform random sections may be difficult, especially when delineating an irregular reference space with complex boundaries, such as the striatum (Schmitz et al. 1999). As we performed measurements of cell size in conjunction with the optical fractionator, neurons were sampled in an unbiased manner. However, a bias could be introduced in estimating absolute size of medium spiny neurons if they were anisotropically oriented. As the presence of mutant huntingtin protein (mhtt) is not expected to alter the orientation of striatal neurons, we would expect that size comparisons between WT and R6/2 mice are still valid, although the absolute value may not be ‘unbiased’.

BDNF mRNA expression in major afferents to the neostriatum

To determine whether a neurotrophin with known anterograde action (Fawcett et al. 2000; von Bartheld et al. 1996) in mammals could help account for impaired motor behavior and morphological changes in the striatum of R6/2 mice, we analyzed the time course of expression of BDNF in the major sources of afferents to the neostriatum. These include glutamatergic inputs from limbic and motor areas of the cerebral cortex, glutamatergic inputs from the thalamus and dopaminergic (DA) inputs from the midbrain (Smith et al. 1994). Interestingly, these sources of major striatal afferents also show high levels of BDNF expression (Conner et al. 1997). Representative autoradiograms of hybridization to antisense BDNF probe (Fig. 6) show the anatomical distribution of BDNF mRNA expression in the forebrain (Bregma 1.70 and Bregma 1.18), thalamus (Bregma −2.18) and midbrain (Bregma −3.16) of 11-week-old R6/2 and WT mice (Franklin & Paxinos 2008). There is virtual absence of BDNF mRNA expression in the striatum (Str). In contrast, WT mice show moderate to dense expression in sources of major striatal afferents, including the cerebral cortex, the thalamic Pf nucleus and the SNc. R6/2 mice show reduced BDNF expression in the cerebral cortex, thalamus and midbrain (Fig. 6). The nonspecific control represents hybridization to sense BDNF probe (Fig. 6).

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Figure 6. Representative autoradiograms of BDNF mRNA showed by in situ hybridization. BDNF mRNA expression in different brain regions following hybridization to antisense probe is shown in paired coronal sections from R6/2 and WT mice. Nonspecific negative control represents hybridization to sense probe. Nuclear outlines are adapted from a stereotactic mouse brain atlas (Franklin & Paxinos 2008). PrL, prelimbic cortex; Cg1 and Cg2, cingulate cortices; M1, motor cortex; M2, premotor cortex; Pf, parafascicular nucleus of thalamus; fr, fasciculus retroflexus; STh, subthalamic nucleus; SNc and SNr, substantia nigra compacta and reticulata.

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Cerebral cortex

To compare changes in BDNF mRNA expression in motor and associational/limbic areas of the frontal lobe at different ages in R6/2 mice and WT controls, prelimbic, cingulate, primary motor (M1) and accessory motor (M2) areas were analyzed (Fig. 6). Two-way anova detected significant differences between R6/2 mice (n = 27) and their age-matched controls (n = 28) in concentrations of BDNF mRNA in prelimbic cortex [F(genotype)1,47 = 76.97, P < 0.0001; F(age)3,47 = 1.67, P = 0.18; F(genotype × age)3,44 = 0.63, P = 0.60]; cingulate cortex [F(genotype)1,47 = 55.97, P < 0.0001; F(age)3,47 = 1.05, P = 0.38; F(genotype × age)3,47 = 3.62, P = 0.02]; M1 cortex [F(genotype)1,47 = 52.59, P < 0.0001; F(age)3,47 = 0.62, P = 0.61; F(genotype × age)3,47 = 3.44, P = 0.02] and M2 cortex [F(genotype)1,47 = 52.59, P < 0.0001; F(age)3,47 = 0.62, P = 0.61; F(genotype × age)3,47 = 2.38, P = 0.08]. Post hoc analysis indicated that BDNF mRNA expression was significantly reduced as early as 4 weeks in R6/2 mice (n = 8) compared with WT controls (n = 8) in prelimbic [49% of control (F1,14 = 8.55, P = 0.01)] and cingulate cortices [71% of control (F1,14 = 4.38, P = 0.05)] (Fig. 7a,b). By 13 weeks, BDNF mRNA in markedly reduced in R6/2 (n = 6) compared with WT (n = 6) mice in both prelimbic [34% of control (F1,10 = 119.10, P < 0.0001)] and cingulate cortices [40% of control (F1,10 = 37.05, P < 0.0001)] (Fig. 7a,b).

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Figure 7. BDNF mRNA expression in cortical and subcortical areas of R6/2 mice and WT controls. BDNF mRNA expression in (a) prelimbic cortex, (b) cingulate cortex, (c) M1 motor cortex, (d) M2 motor cortex, (e) Pf nucleus of thalamus and (f) SNc in R6/2 mice compared with WT controls is compared at different ages. Values for [35S] BDNF mRNA are presented in µci/g of tissue as the mean for each region ± SEM. *P < 0.05, **P < 0.005, ***P < 0.0005, R6/2 vs. WT.

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BDNF mRNA expression is not significantly altered in motor cortical areas (M1 and M2) in 4-week-old R6/2 (n = 8) mice compared with WT (n = 8) controls (Fig. 7c,d). However, by 6 weeks, R6/2 mice (n = 7) show significantly reduced levels of BDNF mRNA compared with WT (n = 6) controls in both M1 [65% of control (F1,11 = 32.84, P = 0.0001)] and M2 [64% of control (F1,11 = 40.64, P < 0.0001)] (Fig. 7c,d). There is a progressive reduction in BDNF mRNA expression in primary and accessory motor cortices during the remainder of the R6/2 mouse lifespan. At 11 weeks, BDNF mRNA expression in R6/2 mice (n = 6) is reduced to 59% of WT controls (n = 8) in M1 (F1,12 = 14.39, P = 0.002) and to 47% of controls in M2 (F1,12 = 28.02, P = 0.0002). Finally, by 13 weeks, BDNF mRNA expression in R6/2 mice (n = 6) is reduced to 51% of WT mice (n = 6) in M1 (F1,10 = 22.93, P = 0.0007) and to 46% of control in M2 (F1,10 = 18.48, P = 0.001).

To determine whether the observed early changes in BDNF mRNA expression are related to neuronal loss, we performed a stereological analysis of the number of neurons in the frontal lobe in 6-week-old R6/2 and WT mice. There was no significant difference in the number of neurons in different areas of the frontal cortex. Specifically, the number of neurons in different cortical regions delineated according to the atlas of Franklin and Paxinos (2008) in R6/2 and WT mice, respectively, was as follows: primary motor cortex (M1): 480 170 ± 40 317 (mean ± SEM) vs. 447 861 ± 22 840 (P = 0.51); secondary motor cortex (M2): 482 158 + 47 617 vs. 468 083 + 10 164 (P = 0.78); cingulate cortex: 350 440 + 30 537 vs. 354 771 + 16 008 (P = 0.90) and prelimbic/infralimbic cortex: 136 187 ± 13 893 vs. 121 732 ± 17 444 (P = 0.54).

Parafascicular nucleus of the thalamus

BDNF mRNA levels in Pf were significantly reduced in R6/2 (n = 27) mice compared with WT (n = 28) controls [F(genotype)1,48 = 43.85, P < 0.0001; F(age)3,48 = 8.31, P = 0.0001; F(genotype × age)3,48 = 7.38, P = 0.0004]. As in the motor cortex, post hoc analysis indicated no significant difference in the level of BDNF mRNA in the Pf nucleus of 4-week-old R6/2 mice (n = 8) compared with WT controls (n = 8; F1,14 = 0.15, P = 0.90). However, by 6 weeks, BDNF mRNA expression in Pf was significantly reduced in R6/2 (n = 7) compared with WT (n = 6) mice [73% of control (F1,12 = 5.76, P = 0.03)]. There was progressive decrease in BDNF expression during the remainder of the R6/2 lifespan. By 11 weeks, BDNF mRNA levels in R6/2 mice (n = 6) were reduced to 56% of WT (n = 8) controls (F1,12 = 12.76, P = 0.004); BDNF mRNA levels in 13-week-old R6/2 mice (n = 6) were reduced to 44% of age-matched WT (n = 6) controls (F1,10 = 84.25, P < 0.0001; Fig. 7e).

Substantia nigra pars compacta

BDNF mRNA levels in the SNc were significantly reduced in R6/2 (n = 27) mice compared with WT (n = 28) controls [F(genotype)1,49 = 110.48, P < 0.0001; F(age)3,47 = 3.03, P = 0.03; F(genotype × age)3,49 = 3.78, P = 0.01]. Post hoc analysis shows a robust reduction of BDNF mRNA expression in the SNc as early as 4 weeks in R6/2 mice (n = 8) compared with WT (n = 8) controls [68% of controls (F1,13 = 17.51, P = 0.001)]. BDNF mRNA expression in DA neurons is reduced progressively during the remainder of the R6/2 mouse lifespan. By 11 weeks, R6/2 mice (n = 6) show a marked reduction in BDNF mRNA levels compared with WT (n = 8) mice [42% of control (F1,12 = 30.43, P = 0.0001)]. By 13 weeks, R6/2 mice (n = 6) show further reduction in BDNF mRNA levels in the SNc [33% of control (F1,10 = 91.57, P < 0.0001)] (Fig. 7f).

Discussion

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information

Our results confirm and extend previous work demonstrating striatal neuronal atrophy and neuronal loss are associated with impaired motor coordination in the R6/2 mouse, a widely used model of morphological and behavioral features of HD (Carter et al. 1999; Cummings et al. 2012; Hickey et al. 2005; Kim et al. 2011; Mangiarini et al. 1996; Stack et al. 2005; Sun et al. 2002; Zuccato et al. 2010). R6/2 mice also show progressive loss of the neurotrophin BDNF in sources of afferents to the striatum. Progressive reduction of BDNF expression is noted not only in motor and limbic cortices but also in neurons of origin of the massive nigrostriatal and thalamostriatal tracts. Severity and time course of phenotypic changes in R6/2 mice can vary with gender (Cowin et al. 2012; Cummings et al. 2012; Menalled et al. 2009; Stack et al. 2005; Wood et al. 2011), with the number of CAG repeats expressed in a given strain (Cowin et al. 2012; Cummings et al. 2012; Menalled et al. 2009; Stack et al. 2005) and with genetic background (Cowin et al. 2012). Our analysis in strain and sex-matched animals with a uniform number of CAG repeats, therefore, provides a unique opportunity for rational interpretation of the relationship between motor behavior, morphology and gene expression.

Evolution of motor behavior in R6/2 mice

Amongst a wide variety of HD models described in different host species, R6 mouse lines are the most widely used (Kim et al. 2011). R6/2 mice carry about 1 kb of the human mutant htt promoter, driving expression of exon 1 of the htt protein gene with ∼110 to ∼320 CAG triplet repeats, resulting in abnormally long polyglutamine tracts (Cowin et al. 2012; Cummings et al. 2012; Mangiarini et al. 1996; Menalled et al. 2010). R6/2 mice show more rapidly evolving behavioral changes than other commonly used mouse models, and may represent a juvenile form of HD (Mangiarini et al. 1996; Nance & Myers 2001; Robitaille et al. 1997).

Progressive motor abnormalities are detected in R6/2 mice (Carter et al. 1999; Hickey et al. 2008; Mangiarini et al. 1996; Stack et al. 2005). Abnormal limb clasping, a measure of dystonia, is an early motor sign. R6/2 mice in this study show a clear weekly progression in clasping score, with mild limb dystonia (score: 0.5–1.5) at 4–8 weeks, moderate dystonia by 9–11 weeks (score: 1.5–2.5) and severe dystonia (score >2.5) involving all limbs in 12- to 15-week-old mice. Reports in mice with larger CAG repeat lengths in the 125–150 range show some variability, with onset of clasping either at a later age (8 weeks) without clear progression or at 4–6 weeks with progression (Carter et al. 1999; Cummings et al. 2012; Hickey et al. 2008; Mangiarini et al. 1996; Stack et al. 2005). This dystonic phenomenon is reduced or absent in R6/2 mouse lines with high CAG repeat lengths (∼200–300 CAG), which show milder and more variable phenotypes, limiting utility of these mice in therapeutic trials (Cummings et al. 2012; Kim et al. 2011; Stack et al. 2005).

CAG repeat size, sex and light–dark conditions can influence locomotor activity (Cummings et al. 2012; Dunnett et al. 1998; Hickey et al. 2008; Menalled et al. 2009). In R6/2 mice, spontaneous locomotor activity varies from hyperactivity detected at 3 weeks, followed by several weeks of activity similar to WT controls, and reduced activity after 8 weeks (Luesse et al. 2001). Male R6/2 mice with CAG repeat sizes in the 125–150 range show mild reduction in total distance traveled by 4–5 weeks during the dark phase of the diurnal cycle and by 6 weeks during the light phase (Menalled et al. 2009). Reduced locomotion is more apparent throughout the diurnal cycle in both sexes by 8 weeks (Carter et al. 1999; Hickey et al. 2008; Menalled et al. 2009, 2010).

In this study using male R6/2 mice with ∼110 CAG repeats, locomotor activity was measured over a prolonged period to reduce effects related to initial measures of novelty, anxiety and habituation (Bolivar et al. 2003; File et al. 1998; O'Keefe et al. 1979). A trend to reduced travel distance is first noted at 6 weeks, followed by significant and progressive reduction after 7 weeks. As significant reduction in fast activity is noted by 6 weeks, movement speed appears to be a more sensitive measure of early decline in locomotor activity compared with distance traveled alone. As mice were not tested earlier than 4 weeks, we cannot exclude the period of hyperactivity detected at 3 weeks in previous studies (Luesse et al. 2001).

Latency to fall at different speeds on a rotarod apparatus is a measure of balance and coordination (Dunham & Miya 1957; Jones & Roberts 1968; Monville et al. 2006; Rustay et al. 2003). Mild impairment in performance is seen as early as 6–7 weeks in R6/2 mice (Carter et al. 1999; Cummings et al. 2012; Hickey et al. 2008; Stack et al. 2005), and more significant declines are noted by age 9–11 weeks, followed by a plateau. We obtained similar results using an incremental fixed speed rotarod protocol (Carter et al. 1999; Cummings et al. 2012; Monville et al. 2006). Optimal rotation speeds for evaluating motor performance using this protocol in R6/2 mice are in the 5–24 r.p.m. range, with little advantage at higher speeds. Abnormal grip strength is a later motor sign. Previous studies of R6/2 mice have indicated variable loss of grip strength ranging from 7 to 12 weeks (Hickey et al. 2005; Menalled et al. 2009; Stack et al. 2005). In this study, loss of grip strength is first detected at 9 weeks when mice pull on a wire mesh either with their forelimbs alone or with all four limbs. Loss of strength in the hindlimbs is a later sign.

Evolution of morphological changes in the neostriatum in R6/2 mice

A major hallmark of HD is loss of projection neurons in the neostriatum, with relative sparing of interneuron subpopulations (Ferrante et al. 1985; Reiner et al. 1988; Vonsattel et al. 1985). Amongst striatal projection neurons, there is a predilection for loss of enkephalin-expressing cells of the indirect pathway at early stages of HD, with relative sparing of projection neurons of the direct pathway, which express substance P (Reiner et al. 1988; Richfield et al. 1995). Initial studies in R6/2 mice detected striatal atrophy (Gil & Rego 2009; Mangiarini et al. 1996; Sun et al. 2002; Turmaine et al. 2000). Gene expression studies suggest a decrease in the number of enkephalin-positive neurons (Luthi-Carter et al. 2000; Menalled et al. 2000; Sun et al. 2002). Unbiased stereological analysis of the precommissural striatum in an R6/2 line with ∼150 CAG repeats documents atrophy and progressive loss of projection neurons (Stack et al. 2005), and Golgi analysis shows loss of dendritic spines (Klapstein et al. 2001).

In our analysis, neostriatal volume loss is first detected in R6/2 mice at age 6 weeks, prior to loss of striatal neuronal volume. This time point corresponds to mild early dystonic movements, decreased latency to fall on a rotarod especially at higher speeds and mild reduction in spontaneous locomotor activity. Neuronal atrophy does not occur until 9 weeks and precedes neuronal loss, which is first detected at 11 weeks. The early phase of abnormal motor behavior with striatal volume loss may therefore correspond to loss or atrophy of striatal afferents or efferents, and these degenerative changes in neuropil may progress throughout the R6/2 lifespan.

Moderate limb clasping abnormalities, increasingly poor balance on a rotarod, further loss of spontaneous activity and onset of grip strength abnormalities correlate best with striatal neuronal atrophy without cell loss at 9 weeks. Finally, severe abnormalities on motor tests are noted after 11 weeks, corresponding to the onset of progressive neuronal loss and further neuronal atrophy.

BDNF expression in frontal cortices and their relationship to striatal morphology and behavior

Corticostriatal abnormalities may precede striatal dysfunction and morphological abnormalities in HD (Coyle & Schwarcz 1976; Laforet et al. 2001; Sapp et al. 1999; Stack et al. 2007; Zuccato et al. 2001). BDNF expression can be regulated by htt, which has an important role in intracellular transport, synaptic function, neurogenesis and neuronal gene transcription (Baquet et al. 2004; Gauthier et al. 2004; Raymond et al. 2011; Wu et al. 2010; Zuccato et al. 2001, 2010). Wild-type but not mutant htt increases BDNF production by cytosolic sequestration of the transcriptional repressor, RE1-Silencing Transcription factor/Neuron-Restrictive Silencer Factor (REST/NSRF), preventing binding to its nuclear target neuron-restrictive silencer element (NSRE), which has a negative regulatory effect on the BDNF gene (Zuccato et al. 2003). Loss of normal htt function in HD may lead to reduced BDNF gene transcription (Cattaneo et al. 2005; Zuccato et al. 2003). Polyglutamine expansion in HD also alters the interaction between huntingtin-associated protein 1 (HAP1) and BDNF leading to impaired fast axonal anterograde transport of the neurotrophin along microtubules (Gauthier et al. 2004; Wu et al. 2010).

The striatum expresses abundant BDNF protein and trkB receptors, but contains virtually no BDNF mRNA, except for sparse expression in a few interneuron subpopulations (Conner et al. 1997; Fryer et al. 1996; Radka et al. 1996). BDNF expression in the corticostriatal pathway exerts an anterograde trophic effect on GABAergic medium spiny neurons and interneurons (Altar et al. 1997; Baquet et al. 2004; Xie et al. 2010). Overexpression of BDNF in forebrain of HD mouse models at least partially rescues striatal neurons and improves motor function (Gharami et al. 2008; Xie et al. 2010). Conversely, reduced striatal BDNF expression accelerates motor dysfunction and loss of enkephalin-containing striatal neurons in R6/1 mice (Canals et al. 2004).

Studies of BNDF in cerebral cortex from HD patients vary from no mRNA loss (Ferrer et al. 2000; Gauthier et al. 2004) to a decrease of both message and protein in samples of parietal cortex without a correlation with disease state (Zuccato et al. 2008). It is proposed that reduced BDNF protein consistently noted in HD striatum (Gauthier et al. 2004; Seo et al. 2004), and in animal models (DeMarch et al. 2008; Giralt et al. 2011; Peng et al. 2008; Simmons et al. 2011) may reflect reduced cortical BDNF production (Zuccato & Cattaneo 2009). Mutant htt may impair kinesin motors resulting in reduced axonal transport or terminal release of BDNF in striatal afferents and contribute to loss of neurotrophic support (Borrell-Pages et al. 2006; Gauthier et al. 2004). In contrast to limited studies possible on human tissue, reverse transcriptase-polymerase chain reaction shows normal cortical BDNF mRNA levels in R6/2 mice in presymptomatic 4-week-old animals, but progressive reduction after 6 weeks (Apostol et al. 2008; Zuccato et al. 2005). A systematic evaluation of regional cortical loss of BDNF gene expression has not been performed in HD or in animal models.

On the basis of regional anatomical analysis using in situ hybridization, we provide evidence for progressive reduction in BDNF mRNA expression in different frontal areas in R6/2 mice. BDNF expression is downregulated in limbic frontal cortices in 4-week-old R6/2 mice, preceding loss in motor cortices. Interestingly, time course analysis of a vast array of genes in the R6/2 mouse cortex and striatum shows first alterations at 4 weeks (Luthi-Carter et al. 2002). BDNF gene expression changes in limbic frontal cortices may therefore reflect an early pathological change resulting in loss of corticostriatal integrity and correlating with early learning and memory deficits (Lione et al. 1999; Ruskin et al. 2011). If documented in HD patients, this early reduction in limbic cortical BDNF may contribute to psychiatric and cognitive manifestations, which can precede overt motor signs (Stout et al. 2011).

Reduction of BDNF mRNA expression in premotor and motor cortices follows early loss in limbic cortices and best correlates with the onset of motor dysfunction in 6-week-old R6/2 mice. Interestingly, the number of cortical neurons remains unaltered in both limbic and motor cortices at 6 weeks, indicating that the early loss of BDNF mRNA expression in these areas is not a reflection of neuronal loss. As striatal volume loss noted at 6 weeks precedes striatal neuronal atrophy, atrophy of corticostriatal afferents may occur as cortical BDNF expression is reduced. Further declines in BDNF in cortical and subcortical afferents, especially when combined, may contribute to striatal neuronal loss in older R6/2 mice.

Altered performance on the open field and rotarod tests, and worsened clasping scores at 6–9 weeks correlate with onset of BDNF gene expression changes in the motor cortex and the Pf, which are major sources of glutamatergic afferents to the striatum. Conditional knockout of BDNF mainly in the cerebral cortex of Emx-BDNF mice results in mild clasping abnormalities, unaltered rotarod performance and loss of striatal dendritic spines but only mild neuronal loss in aging animals (Baquet et al. 2004). It is, therefore, possible that loss of anterograde delivery of BDNF from other sources, such as the thalamus and midbrain, may also be important to generation of the striatal HD phenotype. Numerous other factors related to striatal expression of mutant htt, including increased neuronal vulnerability and excitotoxicity, likely also contribute to neuronal degeneration (Kim et al. 2011; Raymond et al. 2011; Saudou & Humbert 2008; Zuccato & Cattaneo 2009).

Time course of changes in BDNF expression in striatal afferents from the thalamus and midbrain and their relationship to behavior

In addition to forebrain cortical areas, R6/2 mice show progressive downregulation of BDNF mRNA expression in two other major sources of striatal afferents, the thalamus and the ventral midbrain. Dopaminergic neurons localized in the SNc (Bjorklund & Dunnett 2007; Hillarp et al. 1966; Sourkes & Poirier 1965) are the source of a massive nigrostriatal projection, express high levels of BDNF (Conner et al. 1997; Seroogy et al. 1994) and depend on the neurotrophin for survival (Alonso-Vanegas et al. 1999; Baquet et al. 2004, 2005; Hyman et al. 1991). Changes in BDNF expression in the SNc of R6/2 mice are detectable by 4 weeks of age and correlate with onset of dystonic clasping and with discriminant learning deficits sensitive to frontostriatal dysfunction (Lione et al. 1999). Interestingly, WNT-BDNF conditional knockout mice with reduced brainstem BDNF expression also exhibit hindlimb clasping and poor rotarod performance (Baquet et al. 2005), suggesting a potential functional role for BDNF in the nigrostriatal system. R6/1 HD mice may show reduced anterograde transport of BDNF in the nigrostriatal system (Pineda et al. 2005) Previous work indicates no significant loss of DA neurons in HD except at advanced stages (Robitaille et al. 1997). Our results suggest that reduced nigrostriatal BDNF anterograde support may contribute to motor disturbance and striatal neuronal degeneration in HD.

Besides cortical inputs, the neostriatum also receives massive glutamatergic afferents from the intralaminar thalamic nuclei. In primates, the caudal intralaminar complex comprises the centromedian (CM) and Pf nuclei, which provide massive inputs to the sensorimotor and associative striatal territories, respectively (Berendse & Groenewegen 1990; Sadikot et al. 1992a,b; Smith et al. 2009). In HD, the thalamus appears normal at initial stages, but neuronal loss is detected in the CM at later stages (Sadikot & Rymar 2009; Vonsattel et al. 2011). In rodents, the CM and PF are not clearly distinct on Nissl stain, and the lateral part of the Pf is considered the homolog of the primate CM. The Pf expresses one of the highest levels of BDNF mRNA in the brain (Conner et al. 1997), and BDNF in the thalamostriatal system may play an important role in survival of vulnerable striatal neurons during developmental apoptosis (Sadikot et al. 2005). Our results show an important reduction of BDNF mRNA expression in Pf of R6/2 mice at intermediate and later stages of the disease, corresponding to significant atrophy and reduced number of striatal neurons. In addition to corticostriatal afferents, anterograde thalamostriatal BDNF trophic support may therefore also play an important role in survival of striatal neurons in HD.

In conclusion, increased vulnerability of striatal projection neurons in HD results from multiple cell autonomous and extracellular factors. Current theories implicate excitotoxic and neurotrophic mechanisms (Raymond et al. 2011; Saudou & Humbert 2008; Zuccato et al. 2010). We provide evidence for an age-dependent decrease in BDNF expression in major sources of afferents to the striatum in the R6/2 mouse model of HD. BDNF mRNA is progressively reduced not only in the cerebral cortex but also in subcortical sources of striatal afferents, including inputs from the thalamus and the midbrain. Analysis of behavior and morphology during the R6/2 mouse lifespan suggests that loss of BDNF plays an important role in motor and nonmotor abnormalities in HD, and contributes to striatal neurodegeneration. Restoring neurotrophic support to striatal neurons may therefore be a potential therapeutic strategy to be explored for patients with HD.

References

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information
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Acknowledgments

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information

This work was supported by grants from the CIHR and NSERC to A.F.S. P.S. held the Tomlinson Fellowship from McGill University, and Jeanne Timmins Costello Fellowship from MNI.

Supporting Information

  1. Top of page
  2. Abstract
  3. Materials and methods
  4. Results
  5. Discussion
  6. References
  7. Acknowledgments
  8. Supporting Information
FilenameFormatSizeDescription
gbb858-sup-0001-TableS1.docxWord 2007 document287K Table S1: Test history and number of mice for behavioral test at different ages. Note1: The majority of animals used for clasping scores were used in other behavioral experiments. Details are not tabulated because of lack of test history in some mice. As with other behavioral tests, the number of R6/2 mice is reduced at later ages because of mortality.
gbb858-sup-0002-TableS2.docxWord 2007 document108K Table S2: Number of mice used for anatomical analysis at different ages.

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