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DENDRITIC SPINE DYSGENESIS CONTRIBUTES TO HYPERREFLEXIA AFTER 1 SPINAL CORD INJURY 2
3 Abbreviated title: Dendritic Spine Dysgenesis in Hyperreflexia after SCI 4
5 Samira P. Bandaru, Shujun Liu, Stephen G. Waxman, and Andrew M. Tan 6
7 Department of Neurology and Center for Neuroscience and Regeneration Research, 8 Yale University School of Medicine, New Haven, CT 06510, and Rehabilitation 9 Research Center, Veterans Affairs Connecticut Healthcare System, West Haven, CT 10 06516 11 12 Corresponding author: 13 Andrew M. Tan, Ph.D. 14 Center for Neuroscience and Regeneration Research (127A) 15 950 Campbell Avenue, Building 34 16 West Haven, CT 06516 17 Tel: 203-932-5711 x3663 18 Fax: 203-937-3801 19 e-mail: [email protected] 20 21 22 23 24 25 26 Conflict of Interest: None 27 28 29 30 31 32 33 34
Articles in PresS. J Neurophysiol (December 10, 2014). doi:10.1152/jn.00566.2014
Copyright © 2014 by the American Physiological Society.
35
Abstract 36 37 Hyperreflexia and spasticity are chronic complications in spinal cord injury (SCI), with 38
limited options for safe and effective treatment. A central mechanism in spasticity is 39
hyperexcitability of the spinal stretch reflex, which presents symptomatically as a 40
velocity-dependent increase in tonic stretch reflexes and exaggerated tendon jerks. 41
Here we test the hypothesis that dendritic spine remodeling within motor reflex 42
pathways in the spinal cord contributes to H-reflex dysfunction indicative of spasticity 43
after contusion SCI. Six-weeks after SCI in adult Spague-Dawley rats, we observed 44
changes in dendritic spine morphology on α-motor neurons below the level of injury, 45
including increased density, altered spine shape, and redistribution along dendritic 46
branches. These abnormal spine morphologies accompanied the loss of H-reflex rate-47
dependent depression (RDD) and increased H/M ratio. Above the level of injury, spine 48
density decreased compared with below-injury spine profiles and spine distributions 49
were similar to uninjured controls. As expected, there was no H-reflex hyperexcitability 50
above the level of injury in forelimb H-reflex testing. Treatment with NSC23766, a Rac1-51
specific inhibitor, decreased the presence of abnormal dendritic spine profiles below the 52
level of injury, restored RDD of the H-reflex, and decreased H/M ratios in SCI animals. 53
These findings provide evidence for a novel mechanistic relationship between abnormal 54
dendritic spine remodeling in the spinal cord motor system and reflex dysfunction in SCI. 55
56
57
58
59
Introduction 60
61
Hyperreflexia and spasticity, which arise in up to 60% of patients with spinal cord injury 62
(SCI), can severely affect quality of life, contribute to chronic pain, and lead to 63
musculoskeletal deformity (Skold et al., 1999; Walter et al., 2002). Although currently 64
available drugs, such as baclofen, can provide some relief, these drugs have limited 65
therapeutic utility and effectiveness. Thus, there is a significant need for a more 66
complete understanding of spasticity, and for more effective treatment after SCI. 67
68
Central mechanisms that underlie pathological reflex control after injury or disease 69
include the loss of cortical and local spinal inhibition, injury-induced plasticity, and 70
increased motor neuron excitability (Bennett et al., 2001a; Boulenguez et al., 2009; 71
Hultborn et al., 2007; Hunanyan et al., 2013). While plasticity between Ia afferents and 72
α-motor neurons shapes the H-reflex response in an activity-dependent manner in 73
human and rodent (Raisman, 1994; Thompson et al., 2009), maladaptive changes can 74
also contribute to pathological H-reflex function associated with spasticity—clinically 75
defined as a velocity-dependent increase in tonic stretch reflexes with exaggerated 76
tendon jerks, resulting from hyperexcitability of the spinal stretch reflex (Ashby et al., 77
1987; Lance, 1980; Nielsen et al., 2007). 78
79
Dendritic spines, micron-sized structures that are sites of postsynaptic activity, regulate 80
the efficacy of synaptic transmission and can thereby alter the electrical information 81
passing through circuit pathways (Bourne et al., 2007; Calabrese et al., 2006; Pongracz, 82
1985; Segev et al., 1988; Tan et al., 2009). Localized increases in synaptic strength 83
through the de novo formation and development of postsynaptic dendritic spines 84
constitute a persistent structural basis for learning and memory in the CNS (Xu et al., 85
2009; Yuste et al., 2001a). In the present study, we assess the possibility that 86
abnormalities in dendritic spine morphology on α-motor neurons contribute to the 87
persistent dysfunctional state within the spinal motor reflex pathway after SCI. 88
89
Our previous studies and evidence in the literature demonstrate that dendritic spine 90
morphology can change following disease or injury (Kim et al., 2006; Tan et al., 2012b; 91
Tan et al., 2013; Tan et al., 2008). Importantly, adverse changes in spine morphology 92
including (1) the elaboration from thin, filopodia-like spines to a mushroom shape, a 93
morphology associated with increased synaptic strength and stability (Yuste et al., 94
2001b); (2) an increase in spine density along dendrites, which provides more sites for 95
postsynaptic connections (Bonhoeffer et al., 2002), and a spatial redistribution of spines 96
along dendrites to locations closer to the cell body (Kim et al., 2006; Ruiz-Marcos et al., 97
1969) have been shown to contribute to neuronal hyperexcitability (Tan et al., 2009). 98
Although dendritic spine remodeling occurs in the motor cortex after SCI (Kim et al., 99
2006), no study has reported on dendritic spines located on spinal α-motor neurons. 100
Moreover, it is unknown whether SCI-induced changes in dendritic spine morphologies 101
can contribute to spasticity. 102
103
The activity of small GTP-binding protein Rac1 governs actin cytoskeleton 104
reorganization to regulate dendritic spine morphology (Tashiro et al., 2000; Tashiro et 105
al., 2004). Constitutively activated Rac1 increases the rate of dendritic spine turnover, 106
spine density and stability, and spine volume (Nakayama et al., 2000). In contrast, 107
dominant negative Rac1 expression or administration of a Rac1-specific inhibitor 108
NSC23766 disrupt dendritic spine formation and development (Tan et al., 2011; Tan et 109
al., 2012c; Tashiro et al., 2000; Tolias et al., 2007). Importantly, SCI increases Rac1 110
mRNA expression, with levels that can remain elevated for up to three months or more 111
(Dubreuil et al., 2003; Erschbamer et al., 2005). It is not known, however, if Rac1 112
activity contributes to dendritic spine remodeling and reflex dysfunction following SCI. 113
114
Here we provide the first evidence of dendritic spine plasticity on α-motor neurons after 115
SCI and demonstrate a structure-function relationship between dendritic spine 116
dysgenesis and exaggerated spinal motor reflexes associated with spasticity. Six weeks 117
after contusion SCI, animals exhibited increased H-reflex responsiveness (i.e., shown 118
by reduced rate-dependent depression or RDD). Histological assessment in these 119
animals demonstrated that α-motor neurons located below the level of injury within the 120
L4-L5 spinal segments had an increase in dendritic spine density, a significant 121
redistribution of spines along dendrites, and increased dendritic spine head-diameter—122
morphological profiles consistent with those shown to contribute to increased neuronal 123
excitability (Pongracz, 1985; Tan et al., 2009). In contrast, above the level of injury, we 124
observed an absence of exaggerated H-reflex response, reduced dendritic spine 125
densities, a close-to-normal distribution of spines, and normal dendritic spine length and 126
head diameter. Inhibition of Rac1 disrupted SCI-induced dendritic spine profiles on α-127
motor neurons below the injury site, reduced VGluT1 expression (a marker for 128
excitatory primary afferent terminals), and decreased H-reflex responsiveness. Taken 129
together, these observations provide evidence for a new perspective into mechanisms 130
of neuroplasticity within the spinal reflex pathway, and demonstrate a relationship 131
between dendritic spine remodeling and reflex dysfunction after SCI. Targeting of 132
molecular-pathways that regulate spine structure could represent a novel avenue for 133
managing spasticity after SCI. 134
135
Materials and Methods 136
137
Animals and spinal cord injury. Experiments were performed in accordance with the 138
National Institutes of Health Guidelines for the Care and Use of Laboratory Animals. All 139
animal protocols were approved by the Yale University Institutional Animal Use 140
Committee. Animals were housed under a 12 h light/dark cycle in a pathogen-free area 141
with water and food provided ad libitum. A total of 32 adult male Sprague-Dawley rats 142
(175-200 g; Harlan, Indianapolis, IN) underwent procedures to produce each treatment 143
group (Fig. 1, Study Design: Sham n = 11; SCI + Vehicle n = 11; SCI + anti-Rac n = 144
10). Animals were first divided into two treatment arms (Fig. 1). The first group received 145
a contusive spinal cord injury at the 2nd lumbar spinal segment (L2): animals were 146
anesthetized with a mixture of ketamine and xylazine (80/5 mg/kg, i.p.). A small 147
laminectomy was carefully performed at the 12th thoracic vertebra (T12), which exposed 148
the dorsal L2 spinal cord surface (Hebel, 1976). We stabilized the spinal cord in an 149
Infinite Horizon (IH) Impactor device (Precision Systems and Instrumentation [PSI], 150
Lexington KY) by clamping the rostral T11 and caudal T13 vertebral bodies with Adson 151
stabilizing forceps attached to the IH stage (Scheff et al., 2003). The spinal contusion 152
injury was performed with a metal rod (tip diameter of 2.5mm) that was applied to the 153
spinal cord surface with an impact force of 170 kdyn (Rabchevsky et al., 2003; Scheff et 154
al., 2003) (data shown in Fig. 2). For Sham animals (without SCI), the same surgical 155
procedure was followed; including placement of the animal within the IH stabilizing 156
forceps, except no contusion injury was performed. Following all surgical procedures, 157
muscle, fascia, and skin were sutured in sequential layers with 4-0 monofilament 158
sutures. Postoperative treatments included twice daily injections of 0.9% saline solution 159
for rehydration (3.0 cc/s.c.) and Baytril (0.3cc; 3.5 mg/kg b.w., sc, twice daily for 3 days) 160
to prevent urinary tract infection. 161
162
Behavior. Two experimenters blinded to group assignment evaluated animals using the 163
Basso, Beattie, Bresnahan (BBB) locomotor rating scale (Basso et al., 1995) for 164
validation of injury equivalency across SCI animals, as well as to determine whether 165
treatments had an effect on overall locomotor ability,. The BBB score (1 worst to 21 166
best) consists of a combination of hind-limb movements, trunk position and stability, 167
hind limb stepping and coordination, paw placement, and tail position. Behavioral 168
testing was performed at three time-points (Fig. 1): 1) on naïve animals prior to any 169
surgical procedures, 2) within one-week after catheter implantation (and before any 170
drug infusions at ~5-weeks post-SCI), and 3) immediately prior to experimental endpoint 171
(6-weeks post-SCI and Sham surgeries). Prior to any testing, animals were allowed to 172
acclimatize to the testing area for 60-90 minutes. During an experimental trial, animals 173
were allowed to roam freely in the test field (enclosed 3’ x 3’ flat surface) and a similar 174
four minute timeframe of movement was assessed by the experimenters using the BBB 175
scale, as previously described (Basso et al., 1995). After each trial, the surface was 176
cleaned with soap and water and dried. For analysis, BBB scores from both right and 177
left sides of animals were averaged and data from the two experimenters were 178
averaged within groups, and then statistically compared across groups. 179
180
Intrathecal catheter implantation and drug delivery. Five weeks after SCI or Sham 181
surgeries, all animals received ketamine/xylazine anesthesia (80/5 mg/kg, i.p.). As 182
described previously (Tan et al., 2012b), a small craniotomy was performed to expose 183
the atlanto-occiptal membrane (between the occipital bone and vertebral column 184
C1/atlas). A sterile 32G catheter (Recath Co., PA) was carefully inserted through a slit 185
in the membrane and threaded intrathecally until the tip of the catheter reached the 186
lumbar enlargement. The catheter was secured near the base of the skull with sutures 187
placed through overlying muscle and skin. To prevent leakage and infection, the 188
exposed rostral tip of the catheter was heat-sealed by pinching the end with a 189
sufficiently heated and sterilized forceps. The location of the caudal end of the catheter 190
was validated at the experimental endpoint after animals were sacrificed. Animals were 191
allowed to recover for 2-3 days after catheter implantation and then received one of two 192
infusions through the catheter: i) drug vehicle (0.9% sterile saline, 10μl volume, twice 193
daily for 3 days) and ii) NSC23766, a target-specific Rac1-GTPase inhibitor (EMD 194
Chemicals, Darmstadt, Germany), at 2.65 μg/μl (5μl volume, twice daily for 3 days) 195
followed by a 5μl sterile 0.9% saline flush. To measure the maximal affect of treatments, 196
experimental assessments were performed within 1-2 days following the last infusion of 197
vehicle or drug solution. We did not infuse NSC23766 drug in Sham animals in this 198
study, since we had previously already established that NSC23766 does not 199
significantly affect higher-order electrophysiological or behavioral function in uninjured, 200
control animals (Tan et al., 2012b). At the end of the study, this study design produced 201
four comparator arms (Fig. 1, gray): Sham, SCI + Vehicle (includes SCI (above) and 202
SCI (below)), and SCI + anti-Rac. 203
204
Histology. For Golgi-cox staining using a commercial kit and according to 205
manufacturer’s instructions (FD Neurotechnologies; Ellicot, MD) (Sham, n = 5; SCI + 206
Vehicle, n = 4; SCI + anti-Rac, n = 5), a subpopulation of rats from terminal 207
electrophysiological recordings (see below) under ketamine/xylazine anesthesia were 208
killed and processed. Spinal cord tissue (from the cervical enlargement, C4-C5, and 209
lumbar enlargement, L4-L5) was quickly removed (<5 minutes), rinsed in distilled water, 210
and immersed in the kit’s impregnation solution. After the incubation period (~3 weeks), 211
200μm-thick sections were cut on a vibratome (DTK-1000 microslicer; Ted Pella) and 212
mounted on gelatinized glass slides. Sections were stained, rinsed in distilled water, 213
dehydrated, cleared, and coverslipped with Permount medium. For 214
immunohistochemistry, remaining rats were deeply anesthetized with ketamine-xylazine 215
and transcardially perfused with 250 ml of 0.1M phosphate buffer (PB) at 37°C followed 216
by 300 ml of freshly prepared cold paraformaldehyde solution (4% in 0.1M PB). The 217
spinal cord was removed and post-fixed for 2 hours at room temperature, cryoprotected 218
by immersion in 30% sucrose, 0.1M PB at 4°C. Frozen coronal sections from C4-C5, 219
the injury site at L2, and L4-L5, were cut at 20μm thickness using a cryostat (Leica; 220
Bannockburn, IL). Sections were collected onto Superfrost plus slides (Fischer 221
Scientific; Pittsburgh, PA). Immunofluorescence staining methods are described 222
previously (Tan et al., 2006). Sections were washed in blocking solution (0.1 M PBS, 223
0.1% Triton X-100, and 4% normal donkey serum) and incubated overnight at 4°C in 224
mouse anti-VGlut antibody (University of California at Davis/NIH NeuroMab facility 225
1:1000), rabbit anti-GFAP (1:2000 Abcam) or rabbit anti- PKC-γ (Santa Cruz 226
Biotechnology 1:1000). After washing in blocking solution, sections were incubated in 227
fluorescent secondary antibodies, CY3 donkey anti-mouse (Jackson ImmunoResearch 228
Laboratories 1:500) or Alexa Fluor 488 donkey anti-rabbit (Invitrogen 1:2000). Sections 229
were visualized and digitally imaged using a Nikon Eclipse 80i fluorescence microscope 230
equipped with an HQ Coolsnap camera (Roper Scientific; Tucson, Arizona) or Nikon D-231
Eclipse C1 confocal microscopy system. Multi-capture mosaic images were digitally 232
stitched with NIS Elements software (Nikon Instruments, Inc.). 233
234
Dendritic spine visualization on motor neurons and analysis. Investigators blinded 235
to treatment conditions performed all imaging studies and analyses. To visualize 236
neurons and ultra-fine processes, we used a Golgi-staining method as previously 237
described (Tan et al., 2008). For our purpose, we required the ability to fully reconstruct 238
neuronal structure, which required that tissue be exposed to high-intensity light for long 239
periods of time (up for 4 hours per imaging session), which can quickly bleach or 240
diminish other visualization tools, e.g., fluorophores. Golgi staining permits the 241
identification and sampling of a relatively large number of neurons from cervical and 242
lumbar levels within the same animal, and provides robust visualization of the entire 243
neuronal structure, including detailed resolution of dendritic spines. We were specifically 244
interested in motor neuron pools that innervated muscle groups in the forelimb, i.e., 245
extensor carpi radialis, and hindlimb, i.e., plantar muscle. To identify these α-motor 246
neurons, we followed a screening workflow based on data from our previous study (Tan 247
et al., 2012a) and those previously validated in rats (Crockett et al., 1987; Hashizume et 248
al., 1988; Jacob, 1998). We began with a broad sample population of neurons by 249
identifying Golgi-stained α-motor neurons located in the ventral spinal cord in Rexed 250
lamina IX and with soma diameters >25μm (Hashizume et al., 1988; Jacob, 1998). 251
Above the injury site (C4-C5), we narrowed candidate neurons for analysis by selecting 252
neurons from motor pools located in similar dorsolateral coordinates of α-motor neurons 253
shown to innervate the extensor carpi radialis muscle group (~1.5-2 mm deep; 1.5-2 254
mm lateral from midline), as we and others have demonstrated through intramuscular 255
retrograde tracing studies (Sunshine et al., 2013; Tan et al., 2012a; Tosolini et al., 2013). 256
Below the injury site (L4-L5), we narrowed our sampled α-motor neurons to those 257
located in ventral motor pools with similar dorsolateral coordinates of motor pools 258
known to innervate the plantar muscle (~1.5-2.5 mm deep; 1.5 mm-2.2 mm lateral from 259
midline) as shown by retrograde tracing (Crockett et al., 1987; Jacob, 1998). As a 260
refinement step for analysis a priori, we only included α-motor neurons for analysis that 261
had 1) dendrites and dendritic spines that were clearly and completely impregnated, 2) 262
neurons that had dendritic branches appearing as a continuous length for at least 350 263
μm within the tissue slice, and 3) with at least one-half of the primary dendritic branches 264
that remained within the thickness of the tissue section, such that their endings were not 265
cut and appeared to taper into a complete ending (see representative neuron in Fig. 3). 266
To determine if there were any morphological differences across our sample neurons, 267
we used Neuroexplorer software (MicroBrightfield, Williston, VT) to measure maximum 268
cell diameter, aspect ratio (feret max/feret min), form factor (4π x area/perimeter2), 269
number of primary dendrites, and the total dendritic branch length of each treatment 270
arm and compared these morphometry values across treatment groups (Table 1). To 271
refine our identification and measurements of dendritic spines, specific morphological 272
characteristics were used (Kim et al., 2006; Tan et al., 2012b): we defined a spine neck 273
as the structure between the base of the spine and the interface with the parent dendrite 274
branch, and the base of the spine head where the appearance of the spine swells 275
distally into a bulb-like structure. Thin- and mushroom-shaped spines were classified as 276
follows: thin spines had head diameters that were less than, or equal to the length of the 277
spine neck, whereas mushroom spines had head diameters that were greater than the 278
length of the spine neck. These criteria for two spine geometric categories were used 279
because classification into only two spine shapes allowed us to use simple, but very 280
strict rules in classifying spine morphology. Although this approach prevented the 281
discrimination of subtle variations in spine shape, it allowed the collection of a very large 282
sample size and others and we have described the physiological characteristics of thin 283
and mushroom spine shapes on neuronal and circuit function (Bourne et al., 2007; 284
Holmes, 1990; Tan et al., 2009). Note that these criteria do not imply the physiological 285
characterization of the neurons we analyzed, but rather control for morphological 286
diversity within the sampled spinal motor neuron population (Kitzman, 2005; Tashiro et 287
al., 2003). 288
289
To digitally reconstruct motor neurons, we used a Neurolucida software suite (version 290
9.0; MicroBrightfield, Williston, VT) and a pen tablet (Intuos 5 touch, Wacom). We 291
analyzed the completed three-dimensional reconstructions of motor neurons for spine 292
density and distribution. Each imaging session consisted of a contour map (outline of 293
the spinal cord section with location of identified neuron) and the motor neuron, which 294
was traced in the X-, Y-, and Z-axis. Dendritic spine type were located and marked on 295
each reconstructed dendritic branch (thin spines, blue; and mushroom spines, red). 296
297
length. To determine any changes in spatial distribution of dendritic spines relative to 298
the cell body, we used a Sholl’s analysis (Tan et al., 2008). 299
bins were formed around each cell body and spine density within each bin was 300
averaged within each treatment group. For statistical comparison, spine density at 301
dendrite branch locations within 50-150 μm (proximal bins) and 200- μm (distal 302
bins) from the cell body were pooled and compared across treatment groups. 303
304
To determine changes in spine dimensions, five neurons were arbitrarily chosen from 305
each treatment group and visible spines were measured for spine length and spine 306
head diameter (Kim et al., 2006). Spine length was defined as the distance from the tip 307
of the spine to the junction of the spine to the main dendrite branch. Spine head 308
diameter was defined as the longest line drawn normal to the length of the parent 309
dendrite branch. A total of 411 dendrites from 82 identified α-motor neurons (in 4-5 310
animals per treatment group) were included in our analyses (Sham = 118; SCI (below 311
injury) = 116; SCI (above injury) = 90; SCI + anti-Rac = 87). 312
313
Areal density of VGluT1 labeling. To examine changes in the number of VGluT1-314
expressing synaptic terminals, we assessed areal density of VGluT1 expression using a 315
modified approach described previously (Tan et al., 2012a). High-resolution digital 316
photographs were taken at 10x magnification and combined into a single mosaic image 317
of the entire spinal cord at L4-L5. We photographed 10 sections per animal. Sections for 318
analysis were chosen on the basis of tissue integrity (e.g., no major tears) and 319
equivalent loss of PKC-γ immunoreactivity in the dorsal CST of the spinal cord dorsal 320
columns. Sections were aligned according to the point of intersection between the gray 321
matter above the central canal and the dorsal median septum. All images underwent 322
threshold adjustments using equivalent contrast/brightness levels to highlight only 323
VGluT1 expressing puncta (Photoshop, Adobe, San Jose, CA). Images were binarized 324
and color-inverted for analysis. For color-coded heat maps in Fig. 9, binarized images 325
were exported into MATLAB (Mathworks, Natick, MA) and averaged using custom 326
scripts, as described previously (Brus-Ramer et al., 2007; Friel et al., 2007). For image 327
analysis, mosaic images of each spinal cord coronal section were divided into three 328
dorso-ventral regions corresponding to the dorsal zone (~lamina I-III), the intermediate 329
zone (~lamina IV-VI), and the remaining ventral horn of the gray matter (Tan et al., 330
2012a). Because we were only interested in VGluT1 in the gray matter, the white matter 331
areas were digitally removed prior to analyses. Because VGluT1 expressing puncta 332
were visually distinct from each other, without overlap, the number of VGluT1 puncta 333
were easily counted in each region using ImageJ software 334
(http://rsb.info.nih.gov/ij/index.html). Data from gray matter regions were pooled within 335
groups and compared across experimental treatment groups. 336
337
H-reflex testing. Terminal electrophysiological experiments were performed 6 weeks 338
after SCI or Sham surgeries. Because H-reflex responses in rats under ketamine 339
anesthesia resemble those seen in unanesthetized humans (Ho et al., 2002) and does 340
not alter the time course of presynaptic inhibition, which may occur with other 341
anesthetics (e.g., sodium pentobarbital) (Tang et al., 1973), we anesthetized animals 342
with an induction dose of ketamine and xylazine (80/5mg/kg, i.p.) and maintained on 343
ketamine alone (20 mg/kg, i.p.) (Ho et al., 2002; Hosoido et al., 2009). Core body 344
temperature was monitored with a rectal thermometer and maintained at 38 ± 1°C with 345
a circulating water heating pad placed under an absorbent pad. To record 346
electromyogram (EMG) data, which included the muscle response (M wave) and the 347
monosynaptic reflex response (H reflex) above and below the injury site in SCI animals, 348
we used an established percutaneous needle preparation (Boulenguez et al., 2009; Lee 349
et al., 2009; Schieppati, 1987; Thompson et al., 1992a; Valero-Cabre et al., 2004). We 350
chose to use this minimally invasive procedure because it is analogous to methods 351
used to evoke and record H-reflex in humans (Palmieri et al., 2004; Schieppati, 1987) 352
and provides the opportunity to stimulate and record at all four limbs within the same 353
animal without perturbing muscle or nerve tissue that would otherwise be disrupted from 354
more invasive surgical electrode placement, e.g., cuff electrode implants. In addition, 355
this recording approach maintains the integrity of the vascular system and optimizes the 356
tissue preservation and collection methods for histological study (see above) that we 357
performed following electrophysiological experiments. For stimulation, a pair of Teflon 358
insulated stainless steel fine wire electrodes (0.002” bare metal diameter; A-M Systems, 359
Inc., Carlsborg, WA) were threaded into a 32 G syringe needle. The wire ends were 360
carefully bent into sharp barbs, the insulation was removed with heat (to expose tips 361
~1mm), and the needle and wire was then transcutaneously inserted until the wire tip 362
was in close proximity to the mixed nerves of the deep radial nerve or tibial nerve, 363
above or below the injury site in SCI animals, respectively. The needle was retracted 364
and the wire remained in place. The second electrode was inserted similarly, spaced 365
~2mm apart from the first electrode. Stimulating electrode placement was adjusted until 366
the intensity of square wave stimulating pulses (0.2 ms duration continuously given at a 367
rate of 1 every 3 seconds) required to induce subtle visible motor twitch responses, i.e., 368
wrist extension/radial abduction or plantar flexion was below 1mA (Lee et al., 2009; 369
Valero-Cabre et al., 2004). For recording electrodes, we used insulated wire electrodes 370
made of similar materials and exposed ~2 mm of the wire tips using heat. To record 371
EMG data from the forelimb (e.g., brachioradialis reflex), an electrode was inserted into 372
the interosseous muscles between the fourth and fifth digit and a reference electrode 373
was placed subcutaneously in the dorsolateral surface of the paw. To record EMG data 374
from the hindlimb (e.g., plantar reflex), an electrode was inserted into the plantar 375
muscles within the palmar/ventral surface of the hindpaw and proximal to the ankle 376
region. A reference electrode was placed subcutaneously within the dorsolateral surface 377
of the hindpaw. These reflexes were chosen based on our pilot experiments and 378
previous work that demonstrated EMG reflex response evoked in these muscles could 379
be reproducibly recorded after SCI (Boulenguez et al., 2010; Kim et al., 2009; Valero-380
Cabre et al., 2004). Note that SCI-induced changes in the plantar reflex, primarily 381
innervated by motor pools in L5, less from L4 (Crockett et al., 1987), have been shown 382
to be similar to changes in reflexes elicited in other hindlimb muscles, i.e., tibialis 383
anterior and gastrocnemius, which are also innervated by L4 and L5 (Lee et al., 2009; 384
Valero-Cabre et al., 2004). EMG responses were filtered (10-1000 hz), amplified, and 385
recorded for offline analysis using Spike 2 (version 7.08; CED Software, Cambridge, 386
England). To identify optimal stimulation intensity for activating stable M-wave and H-387
reflex responses, square wave pulses (0.2 ms duration) were applied at a rate of 1 388
every 3 seconds. The intensity of electrical stimulation was first adjusted to determine 389
the minimum intensity to evoke an M-wave response ~50% of the time and 390
progressively increased until a stable M-wave and maximal H-reflex response could be 391
observed. 392
393
To measure rate-dependent depression (RDD) of the H-reflex response, we performed 394
a paired-pulse stimulation paradigm with a conditioning and test pulse that we applied at 395
a range of interpulse intervals (10 ms to 2000 ms). Three trials (10 sweeps/trial) with at 396
least 30 seconds between trials were recorded for each interpulse interval. The M and H 397
response wave amplitudes were quantified from averaged and rectified waveforms 398
within each animal (Boulenguez et al., 2010; Tan et al., 2012a). For comparison across 399
treatment groups, the maximum waveform amplitudes of the H and M response to the 400
test pulse were converted into a percentage of the maximum amplitude response to the 401
conditioning pulse. M and H waveform amplitudes were measured from baseline to 402
peak amplitude. To determine trial-to-trial consistency, the coefficient of variation (CoV) 403
from the M and H waves were calculated by dividing the standard deviation with the 404
mean maximum amplitude. Following recording experiments, animals were sacrificed 405
for Golgi-staining or immunohistological study as described above. 406
407
A number of studies have shown that dendritic spine morphology can change within 408
minutes following cortical injury or activity (Majewska et al., 2003; Mizrahi et al., 2004; 409
Zhang et al., 2005). Moreover, others have shown that exogenous electrical stimulation 410
can induce plasticity of the H-reflex, which can persistent for many hours or days (Chen 411
et al., 2003; Chen et al., 2006). Although in this study our H-reflex testing was 412
performed acutely (i.e., lasting ~1 hour per animal), we ensured that all animals, 413
including both spinal cord injured and uninjured, Sham animals, underwent similar H-414
reflex testing protocols to control for potential confounds of direct nerve stimulation. 415
416
Statistical analysis. All statistical tests were performed at the alpha-level of 417
significance of 0.05 by two-tailed analyses using parametric or non-parametric tests, as 418
appropriate. For comparisons of anatomical and functional changes above and below 419
the injury site after SCI, we compared multiple comparisons of data collected within the 420
same animals and with Sham animals. To determine the appropriate statistical model to 421
apply to these specific datasets, we incorporated two assumptions: 1) that the above 422
and below datasets were dependent variables affected by the application of the 423
independent treatment variable, SCI, and 2) as a consequence of SCI, a putative 424
secondary pathway interaction arises between above and below injury spinal segments 425
that can affect the above or below datasets (e.g., an emergent interaction following SCI 426
that leads to differential affects on spinal cord tissue located above or below the SCI 427
injury site). Given these assumptions, our datasets fit most closely with a one-way (or 428
one-factor) ANOVA statistical model, where SCI is the treatment factor applied to two 429
dependent variables, above and below. Moreover, to control for multiple comparison 430
errors with the additional comparison against the Sham group, we applied a post-hoc 431
repeated measures correction. While we considered the application of a split-plot 432
analytical model, this approach falsely assumes that the secondary, post-SCI interaction 433
is a factor that is independent of the first-cause treatment factor, SCI. In summary, for 434
comparing datasets gathered from above and below the injury site, we applied 435
repeated-measures corrections (Dunn or Bonferonni post hoc tests) following ANOVA 436
and Kruskal-Wallis one-way ANOVA on ranks analyses. As a note, previous reports 437
have similarly applied ANOVA repeated measures design to compare ipsilateral and 438
contralateral sides of the spinal cord following unilateral nerve trauma, or compare 439
regions above and below the injury site after SCI. The statistical design of these studies 440
encompassed the assumed effect of a post-injury secondary, bidirectional interaction, 441
(Anderson et al., 1998; Chang et al., 2010; Tal et al., 1994; Tan et al., 2012a; Tan et al., 442
2007), which may have also emerged within our experimental SCI model. Data 443
management and statistical analyses were performed using SigmaPlot (version 12.5; 444
Systat Software Inc.) and Microsoft Office Excel (2011). Data in the text are described 445
as mean ± SD. All graphs are plotted as mean ± SEM using SigmaPlot. 446
447
448
449
Results 450
451
Contusion SCI disrupts the dorsal corticospinal tract 452
In SCI animals, contusion injury at spinal cord segment L2 resulted in severe damage of 453
the dorsal columns and gray matter (Fig. 2A), in contrast with Sham (Fig. 2B). 454
Histological examination of caudal spinal cord tissue (segments below L3) showed no 455
visible tissue damage or glial scar tissue (not shown). The IH Impactor device provided 456
biomechanical measurements during each SCI procedure (Fig. 2C and Fig. 2D). The 457
cord surface displacements upon rod impact were not significantly different across both 458
SCI groups, SCI + Veh and SCI + anti-Rac (Fig. 2C; 1419.8 ± 293.3 vs. 1394.9 ± 266.2 459
μm, p>0.05, t test). Similarly, there was no difference in the actual applied force 460
between the SCI groups (Fig. 2D; 172.4 ± 12.4 vs. 181.4 ± 19.7 kdyn, p>0.05, ANOVA 461
on ranks with Dunn’s post hoc). Applied impact force with the IH device predicts the 462
amount of tissue sparing, which correlates closely with locomotor functional outcome 463
(Scheff et al., 2003). PKC-γ immunoreactivity served as an anatomical marker to help 464
confirm the injury magnitude of our SCI model (Bradbury et al., 2002; Sasaki et al., 465
2009; Tan et al., 2012a). In coronal spinal cord sections above the injury in the cervical 466
enlargement, PKC-γ immunoreactivity symmetrically labeled the dorsal corticospinal 467
tract (dCST) and small-diameter cells located in laminae I/II (Fig. 2E) (Mori et al., 1990). 468
Six weeks after SCI, below the injury level in the lumbar enlargement, PKC-γ staining of 469
the dCST was bilaterally eliminated from the dorsal columns (Fig. 2F). PKC-γ staining 470
profiles of the dCST and superficial laminae remained intact (data not shown) in cervical 471
and lumbar enlargement tissues in Sham animals. 472
473
Dendritic spine density changes on motor neurons after SCI 474
Dendritic spines remodel in the motor cortex after SCI (Kim et al., 2008; Kim et al., 475
2006); however, it is not known whether SCI-induced dendritic spine dysgenesis occurs 476
on α-motor neurons within the spinal cord. Injury-induced changes in dendritic spine 477
morphology on nociceptive neurons in the dorsal horn have been shown to contribute to 478
increased excitability associated with neuropathic pain (Tan et al., 2009; Tan et al., 479
2012b; Tan et al., 2008). To determine whether dendritic spine remodeling occurs on 480
spinal cord α-motor neurons, we identified α-motor neurons (see Materials and 481
Methods) and performed a morphological comparison of α-motor neurons across 482
treatment groups (Fig. 3). We identified α-motor neurons located in lamina IX and within 483
motor pools in the lateral regions of the ventral horn (Fig. 3A and Fig. 3B). Six-weeks 484
after SCI, motor neurons had widely projecting dendritic trees containing numerous 485
spines. Qualitative observations demonstrated marked differences in spine number 486
across treatment arms (Fig. 3C-F). To ensure equivalent sampling across groups, we 487
assessed several morphological criteria and compared these values across treatment 488
groups (Table 1). There were no statistically significant differences in maximum cell 489
diameter, aspect ratio, form factor, number of primary dendrites, or total dendrite branch 490
lengths (for all comparisons: p>0.05). We therefore interpreted any differences in 491
dendritic spine profiles across groups as not due to variations in neuronal sampling and 492
rather an effect of experimental treatments. As a note, the values for maximum cell 493
diameter, form factor, and dendritic branch lengths were similar to measurements for α-494
motor neurons that were labeled by intramuscular injected retrograde tracers (Bose et 495
al., 2005; Crockett et al., 1987; Hashizume et al., 1988; Jacob, 1998). 496
497
To obtain an accurate measure of dendritic spine profiles from ventral spinal cord tissue, 498
we digitally reconstructed α-motor neurons using Neurolucida software (Fig. 4). We 499
marked the location of sample neurons on a contour map of the spinal cord gray matter 500
(Tan et al., 2008). α-motor neurons from each treatment group (Fig. 4A-D; red dots; 501
n=20-21 cells/group) were located in the ventrolateral regions of the gray matter, shown 502
as a single representative contour from segmental level C5 (above injury) or L5 (below 503
injury). Dendritic spines on traced motor neurons were marked along dendritic branches 504
and color-coded with thin-shaped (blue) or mushroom-shaped (red) spines (Fig. 4A’-D’). 505
506
A main objective of this study was to assess the contribution of SCI-induced changes in 507
dendritic spines to reflex dysfunction; we therefore measured three morphological 508
profiles of spines that have been associated with injury-induced neuronal 509
hyperexcitability: 1) increased density of dendritic spines, particularly mature 510
mushroom-spine spines, 2) redistribution of spines toward dendritic branch locations 511
close to the cell body, and 3) enlargement of the spine head diameter. Because 512
spasticity often presents below the injury site following SCI, and less commonly above 513
(Skold et al., 1999), we measured dendritic spines on α-motor neurons in motor pools of 514
the cervical (C4-C5; above injury) and lumbar (L4-L5; below injury) spinal segments that 515
innervate forelimb and hindlimb musculature, respectively (see Methods and Materials). 516
517
As shown in Fig. 5A, six weeks after SCI, total dendritic spine density on motor neurons 518
below the injury site increased compared with Sham control and neurons above the 519
injury site (p<0.05; 2.80 ± 0.78 vs. 1.82 ± 0.52 vs. 1.10 ± 0.54 spines/10 μm dendrite; 520
ANOVA on ranks with Dunn’s post hoc). In contrast, motor neurons above the injury site 521
in the cervical enlargement had dendritic spine densities that decreased compared with 522
Sham (p<0.05). A similar profile of dendritic spine density was also observed for thin-523
shaped dendritic spines (Fig. 5B): below the injury site there was a significant increase 524
in thin spines compared with above the injury and Sham Sham (p<0.05; 2.42 ± 0.63 vs. 525
1.03 ± 0.54 vs. 1.61 ± 0.40 spines/10 μm dendrite; ANOVA on ranks with Dunn’s post 526
hoc). Importantly, there was a significant increase in the density of mature, mushroom-527
shaped spines located on α-motor neurons below the injury site compared with Sham 528
and above the injury (p<0.05; 0.45 ± 0.33 vs. 0.20 ± 0.26 vs. 0.06 ± 0.06 spines/10 μm 529
dendrite; ANOVA on ranks with Dunn’s post hoc) (Fig. 5C). Note that the mushroom-530
spine density observed below the injury site after SCI represents a more than 200-700% 531
increase compared with mature-shaped spine densities above the injury site and Sham 532
control. 533
534
Dendritic spines redistribute toward proximal branches on motor neurons after 535
SCI 536
Excitatory afferent inputs located closer to the neuronal cell body can have a greater 537
weighted impact upon the overall electrical output of a neuron due to the closer 538
proximity to the axon hillock (Pongracz, 1985; Tan et al., 2009; Yuste et al., 2004). To 539
profile changes in dendritic spine distribution along motor neuron branch processes, we 540
applied a Sholl’s analysis and pooled spine densities within proximal regions close to 541
the cell body (50-150 μm) and distal regions (200-350 μm) (see Methods and Materials) 542
(Fig. 5D-F). 543
544
On motor neurons below the injury site in SCI animals, total spines, thin-shaped, and 545
mushroom-shaped dendritic spines increased on proximal dendrite branches compared 546
with equivalent regions in Sham and SCI neurons above the injury site (p<0.05; for total 547
spines: 3.1 ± 1.1 vs. 2.1 ± 1.1 vs. 1.2 ± 0.69; for thin spines: 2.7 ± 0.16 vs. 1.9 ± 0.05 vs. 548
1.1 ± 0.05; for mushroom spines: 0.39 ± 0.04 vs. 0.17 ± 0.06 vs. 0.08 ± 0.01 spines/10 549
μm dendrite; one-way ANOVA with Bonferroni’s post hoc) (Fig. 5D-F). At distal regions, 550
SCI did not change spine density of any category on motor neurons from below the 551
injury compared with Sham (p>0.05). On the other hand, motor neurons below the injury 552
had increased spine density for all categories as compared with above the injury at 553
distal regions (p<0.05; for total spines: 3.2 ± 1.4 vs. 1.4 ± 0.9; for thin spines: 2.3 ± 1.4 554
vs. 1.4 ± 0.9; for mushroom spines: 0.85 ± 0.9 vs. 0.08 ± 0.16 spines/10 μm dendrite; 555
ANOVA on ranks with Dunn’s post hoc). There was no difference in any spine densities 556
at distal regions on neurons from Sham compared with neurons above the injury in SCI 557
animals (p>0.05). 558
559
Dendritic spine dimensions change on motor neurons after SCI 560
To quantify the effects of SCI on spine length and spine head diameter, we analyzed 561
880 to 1,305 dendritic spines that were sampled from 4-7 motor neurons per group (see 562
Materials and Methods; Sham n = 3 animals/5 neurons; above SCI level n = 3 animals/7 563
neurons; below SCI level n = 3 animals/4 neurons). As shown in Fig. 5G, dendritic 564
spines below the injury site in SCI animals decreased in length compared with above 565
the injury site and Sham (p<0.05; 1.38 ± 0.79 μm vs. 1.67 ± 0.97 μm vs. 1.60 ± 0.81 μm, 566
one-way ANOVA with Bonferroni’s post hoc). In contrast, spine-head diameter 567
increased after SCI below the injury site compared with above the injury and Sham 568
(p<0.05; 1.22 ± 0.65 μm vs. 0.97 ± 0.57 μm and 0.95 ± 0.56 μm, one-way ANOVA with 569
Bonferroni’s post hoc) (Fig. 5H). Motor neurons above the injury site after SCI did not 570
differ compared with Sham in any dimension measured (p>0.05). 571
572
H-reflex response increases below the injury site after SCI 573
Dendritic spine morphology significantly influences synaptic function, i.e., in a structure-574
function relationship (Pongracz, 1985; Segev et al., 1988; Segev et al., 1998). As we 575
and others have shown (Leuner et al., 2004; Majewska et al., 2000; Tan et al., 2009; 576
Zhou et al., 2004), increased dendritic spine density, mature dendritic spine 577
morphologies, e.g., mushroom-shapes, and proximal redistribution of spine synapses 578
can amplify neuronal excitability, enhance frequency-following ability, reduce noise-579
filtering capabilities, and attenuate inhibitory input. To determine the effect of SCI on 580
reflex function in association with dendritic spine remodeling on α-motor neurons, we 581
measured the H-reflex response in uninjured Sham, i.e., in the hindlimb, and SCI 582
animals above and below the injury in the forelimb and hindlimb, respectively. 583
We also measured the M-wave, which reveals the electrical responsiveness of motor 584
axons, its ability to conduct an action potential, and the electrochemical coupling of the 585
efferent and muscle tissue, e.g., neuromuscular junction (Hultborn et al., 1995). In 586
normal animals, the H-reflex undergoes activity rate-dependent depression (RDD). A 587
reduction in H-reflex RDD is a physiological indicator of spasticity (Boulenguez et al., 588
2010; Ho et al., 2002; Lee et al., 2009; Taylor et al., 1984). To determine H-reflex and 589
M-wave response in SCI animals, we electrically stimulated the deep radial nerve or 590
tibial nerves and recorded reflex response from muscle in the forelimb, extensor carpi 591
radialis, and hindlimb, plantar muscle. As a comparison, we measured evoked H- and 592
M-response from hindlimb muscle in uninjured Sham animals. We chose these reflexes 593
based on preliminary studies and previously published work that demonstrated that 594
evoked reflex responses for these muscles could be reproducibly produced in adult rats 595
after SCI (Boulenguez et al., 2010; Kim et al., 2009; Valero-Cabre et al., 2004). 596
Importantly, previous studies have shown that changes in plantar reflex after SCI are 597
similar to changes in reflexes elicited for other hindlimb muscles, i.e., tibialis anterior 598
and gastrocnemius, which are also innervated by motor pools in L4/L5 (Lee et al., 2009; 599
Valero-Cabre et al., 2004). 600
601
We used a paired-pulse stimulation paradigm: a control and test pulse, separated by a 602
range of interpulse intervals from 2000 ms to 10 ms (Fig. 6). A representative trace in 603
Sham produced from recordings of plantar muscle shows two evoked EMG waves: the 604
M-response and the H-reflex (central loop pathway) (Fig. 6A). As the interval between 605
the control and test pulse shortened from 2000 ms to 10 ms, there was a marked 606
depression of the H-reflex. The M-wave amplitude also decreased with shortening 607
interpulse intervals, demonstrating rate-dependent depression of motor neuron-to-608
muscle response. Fig. 6B and Fig. 6C shows a qualitative example of SCI-induced 609
reductions in H- and M-wave RDD from muscle recordings above and below the injury 610
at 100ms and 10ms interpulse intervals. In SCI animals, the M-wave appeared to 611
maintain stable amplitude even at shorter interpulse intervals. 612
613
We quantified the percentage change in H-reflex for uninjured Sham (n = 7) and SCI 614
animals (n = 6) over the range of interpulse intervals (Fig. 6D). In Sham animals, the H-615
reflex maintained stable amplitude between 2000ms and 300 ms and with a steady 616
decline at shorter interpulse intervals, similar to that observed in previous studies (Ho et 617
al., 2002; Hosoido et al., 2009; Tan et al., 2012a). The M-wave amplitude in Sham 618
animals remained stable through a wider range of interpulse intervals, 2000 ms and 100 619
ms (%M-wave amplitude 2000 vs. 50ms: p<0.05, one-way ANOVA with Bonferroni’s 620
post hoc) (Fig. 6E). Therefore, the H-reflex depression response in sham animals is not 621
due to the inability of muscle to respond to repeated stimulus activity. At shorter 622
interpulse intervals (i.e., 3-5 ms), both the H-reflex and M-wave responses depressed at 623
a much greater rate and, in most stimulus-recording trials, failed to appear in sufficient 624
number for analysis (data not shown). 625
626
Six-weeks after SCI, H-reflex measurements from evoked hindlimb reflex revealed a 627
significant reduction in RDD, i.e., H-reflex amplitude stabilized, through the entire range 628
of interpulse intervals tested (Fig. 6D). Between interpulse intervals 50 to 150 ms, there 629
was a significant increase in H-reflex response in SCI below the injury site compared 630
with Sham (p<0.05; one-way ANOVA with Bonferroni’s post hoc). Notably, SCI 631
appeared to amplify the reflex response below the injury site at the shortest interpulse 632
interval at 10 ms compared with Sham (p<0.05; one-way ANOVA with Bonferroni’s post 633
hoc), with amplitude responses greater than 100% of control amplitude. In contrast, 634
within the same SCI animal, there continued to be significant RDD in recordings above 635
the injury at 500 ms, 50 ms, and 10 ms (p<0.05; one-way ANOVA with Bonferroni’s post 636
hoc) (Fig. 6D). H-reflex RDD above the injury was not significantly different to Sham 637
across all interpulse intervals (p > 0.05). These findings indicate that SCI-induced 638
increases in H-reflex response only occurred below the level of the injury and from 639
muscle innervated primarily by motor pools in spinal segment L4-L5. Over the range of 640
interpulse intervals tested from 2000 ms to 100 ms, the %M-wave amplitude remained 641
close to 100% across all comparator groups (Fig. 6E). However, there was a significant 642
increase in %M-wave amplitude above 100% of control in SCI above or below injury 643
compared with Sham, which depressed at 50ms and 10ms (p<0.05; ANOVA on ranks 644
with Dunn’s post hoc). The H/M ratio calculated from reflex data in SCI below the injury 645
was larger than Sham or above injury at all interpulse intervals (Fig. 6F). SCI below the 646
injury site resulted in a more stabilized rate of decay for H/M ratio values, an indication 647
of hyperreflexia and spasticity (Little et al., 1985; Matthews, 1966; Nielsen et al., 2007). 648
649
To assess changes to H-reflex response fidelity, we calculated the coefficient of 650
variation (CoV = StDev/mean of %H-reflex amplitude) for 50 ms and 10 ms for Sham 651
and SCI animals. At the 50 ms interpulse interval, the CoV after SCI below the injury 652
was nearly 30-50% smaller than for SCI above the injury or Sham (SCI below injury, 653
0.20; SCI above injury, 0.58; Sham, 0.85). Similarly, the CoV for SCI below the injury 654
site was almost 40-60% smaller than Sham (SCI below injury, 0.37; SCI above injury, 655
0.66; Sham, 0.92). Taken together, these values show that in addition to increasing H-656
reflex amplitude, SCI also increases the reliability of reflex activation below the injury 657
site. 658
659
Inhibition of Rac1 disrupts dendritic spine remodeling 660
We reasoned that if abnormal dendritic spine profiles after SCI contributes to increased 661
reflex excitability, then disruption of dendritic spine remodeling would reduce signs of 662
spasticity. To determine whether disruption of dendritic spine remodeling on α-motor 663
neurons in L4-L5 after SCI reduces spasticity, we assessed the effects of administering 664
NSC23766, a specific Rac1-GTPase inhibitor. Treatment with NSC23766 resulted in a 665
decrease in total, thin-, and mushroom-shaped spine density compared with SCI + Veh 666
(p<0.05, SCI + anti-Rac vs. SCI + Veh; 0.95 ± 0.24 vs. 2.8 ± 0.78 total spines/10 μm 667
dendrite; 0.92 ± 0.23 vs. 2.4 ± 0.63 thin spines/10 μm dendrite; 0.03 ± 0.03 vs. 0.44 ± 668
0.33 mushroom spines/10 μm dendrite, ANOVA on ranks with Dunn’s post hoc) (Fig. 669
7A-C). We also determined the effect of NSC23766 treatment on dendritic spine 670
distribution by measuring spine density in proximal and distal locations along dendrites 671
of α-motor neurons after SCI (Fig. 7D-F). NSC23766 treatment significantly decreased 672
spine density in proximal and distal regions and for all spine categories, including total 673
spines, thin-shaped, and mushroom-shaped dendritic spines (p<0.05; for proximal total 674
spines: 3.1 ± 0.2 vs. 0.9 ± 0.4; for proximal thin spines: 2.7 ± 0.16 vs. 0.9 ± 0.4; for 675
proximal mushroom spines: 0.4 ± 0.4 vs. 0.02 ± 0.02; for distal total spines: 3.2 ± 0.5 vs. 676
1.2 ± 0.5; for distal thin spines: 2.4 ± 0.5 vs. 1.1 ± 0.5; for distal mushroom spines: 0.8 ± 677
0.9 vs. 0.04 ± 0.07 spines/10 μm dendrite; one-way ANOVA with Bonferroni’s post hoc). 678
679
Dendritic spines on α-motor neurons in SCI animals that were treated with NSC23766 680
decreased in spine length and head diameter compared with SCI + Veh (p<0.05; length, 681
0.7 ± 0.42 μm vs. 1.38 ± 0.79 μm; head diameter, 0.93 ± 0.51 μm vs. 1.23 ± 0.66 μm, 682
ANOVA on ranks with Dunn’s post hoc) (Fig. 7G-H). Together, these findings show that 683
Rac1-inhibitor NSC23766 treatment can effectively disrupt SCI-induced dendritic spine 684
remodeling on α-motor neurons. 685
686
Inhibition of dendritic spine remodeling reduces H-reflex excitability after SCI 687
Previous work has demonstrated that intrathecal infusion of NSC23766 is efficacious in 688
restoring close-to-normal dendritic spine profiles on nociceptive neurons within the 689
dorsal horn after SCI and peripheral nerve injury (Tan et al., 2011; Tan et al., 2008). In 690
these studies, treatment with NSC23766 also reduced neuronal hyperexcitability 691
associated with central sensitization, demonstrating that Rac1-regulated dendritic spine 692
remodeling can contribute to mechanisms underlying neuropathic pain (Tan et al., 693
2012c). To determine whether disruption of Rac1-regulated dendritic spine profiles on α-694
motor neurons attenuates exaggerated H-reflex responsiveness after SCI, we used a 695
paired-pulse stimulation paradigm as described above (Hultborn et al., 1995; Tan et al., 696
2012a) (see Fig. 6). Representative EMG traces in SCI animals below the injury in the 697
hindlimb demonstrates that stimulation produced both M-wave and H-reflex responses 698
(Fig. 8). At the shortest interpulse interval at 10 ms, there was a notable reduction of the 699
RDD. At the 10 ms interpulse interval, treatment with the Rac1-inhibitor NSC23766 700
appeared to restore RDD of the H-reflex in hindlimb EMG recordings (e.g., reduced H-701
reflex amplitude in response to the test pulse) (Fig. 8B). 702
703
Fig. 8C shows the quantified changes in the H-reflex response in SCI + Vehicle (n = 6) 704
and SCI + anti-Rac1 (n = 5). Six weeks after SCI, in animals treated with control vehicle, 705
the hindlimb H-reflex response demonstrated almost no RDD, with %H-reflex response 706
remaining stable (e.g., close to 100%) across the range of interpulse intervals from 707
2000 ms to 10 ms (for a comparison against Sham, see Fig. 6). Treatment of SCI 708
animals with the Rac1-inhibitor resulted in a restoration of RDD at the three shortest 709
interpulse intervals at 100 ms, 50 ms, and 10 ms compared with SCI + Vehicle (p<0.05; 710
at 100 ms, 56.9 ± 29.9% vs. 91.6 ± 8.5%; at 50 ms, 34.5 ± 35.3% vs. 92.4 ± 19.3%; at 711
10 ms, 13.3 ± 21.4% vs. 88.9 ± 32.9%, one-way ANOVA with Bonferroni’s post hoc) 712
(Fig. 8C). In comparisons across SCI animal groups, the %M-wave amplitude remained 713
close to 100% between interpulse intervals 2000 ms and 100 ms (group means: for SCI 714
+ Veh, 98.6 ± 1.5%; for SCI + anti-Rac1, 107.3 ± 11.4%) (Fig. 8E). The %M-wave 715
response at shorter interpulse intervals, 50 ms and 10 ms, exhibited greater variability 716
compared with longer stimulus intervals, but was not statistically significant (p > 0.05). 717
718
We calculated the coefficient of variation of the %H-reflex at 50ms and 10ms across 719
SCI animal groups treated with vehicle or the Rac1-inhibitor. At the 50ms and 10ms 720
interpulse intervals, treatment with the Rac1-inhibitor in SCI animals resulted in a CoV 721
that was nearly 4-fold greater than vehicle treated SCI animals (SCI + Veh, 0.27 – 0.37; 722
SCI + anti-Rac, 0.9 – 1.6). Thus, in addition to restoring RDD, Rac1-inhibition also 723
increased the variability of the reflex response. Fig. 8E shows the plot for the H/M ratio. 724
Treatment with the Rac1-inhibitor decreased the overall H/M ratio across all interpulse 725
intervals tested between 2000 ms and 10 ms, as shown by a downward shift in trend 726
line slope. 727
728
VGluT1 bouton areal density in the gray matter does not increase after injury 729
Synapse-associated protein markers (i.e., synaptophysin and PSD-95) increase after 730
SCI, demonstrating the presence of injury-induced synaptic plasticity (Tan et al., 2008; 731
Tan et al., 2012c). Because upper motor tract injury and SCI can increase the 732
excitability of spinal reflex pathways below the injury (Baastrup et al., 2010; Little et al., 733
1985; Tan et al., 2012a), we next determined if excitatory inputs, particularly those of IA 734
afferents, change after SCI. Vesicular glutamate transporter-1 (VGluT1) is a widely used 735
marker for excitatory Ia afferent terminations in the spinal cord (Alvarez et al., 2011; 736
Alvarez et al., 2004; Kitzman, 2007). 737
738
As shown in representative images in Fig. 9A-D, immunopositive VGluT1 puncta were 739
distributed throughout the spinal cord gray matter of the lumbar enlargement, L4-L5, for 740
each analyzed treatment group (Fig. 9A-D, left panels). To visualize the distribution of 741
VGluT1 puncta in the gray matter, we compiled the VGluT1-staining profiles from 742
multiple tissue sections from each treatment group and produced spatial heat maps (Fig. 743
9A-C, right panels). Although the highest concentration of VGluT1 positive boutons 744
appeared to correspond with Rexed laminae 5-6 (Hantman et al., 2010; LaMotte et al., 745
1991), VGluT1 puncta were distributed throughout all laminae. The areal densities of 746
VGluT1 boutons were calculated in the total gray matter (Fig. 9D), and within three 747
regions: the dorsal horn, the intermediate zone, and the ventral horn (Fig. 9E-G; also 748
see insets). Six weeks after SCI, we observed no significant difference in the areal 749
density of VGLUT boutons in the total gray matter compared with uninjured Sham 750
animals (p>0.05). Similarly, there was no statistical difference following SCI + Veh in the 751
other three gray matter regions analyzed, as compared with uninjured Sham (p>0.05). 752
In contrast, treatment with NSC23766 significantly decreased the areal density of 753
VGluT1 compared with SCI + Veh in the total gray matter, in the intermediate zone, and 754
the ventral horn (for total gray matter: 63.6 ± 37.1 vs. 124.5 ± 57.7; for intermediate 755
zone: 72.4 ± 25.2 vs. 138.7 ± 51.3; for ventral horn: 58.1 ± 48.4 vs. 129.8 ± 55.9 puncta; 756
ANOVA on ranks with Dunn’s post hoc). There was no significant change in areal 757
density of VGluT1 in the superficial dorsal horn following Rac1-inhibitor treatment in SCI 758
animals as compared with SCI + Veh (p > 0.05). 759
760
Rac1 inhibition does not affect locomotor behavior 761
To rule out any differences in gross locomotor ability in SCI animals, we assessed post-762
injury locomotor behavior using the Basso, Beattie, and Bresnahan (BBB) locomotor 763
scale (Basso et al., 1995; Basso et al., 1996) (Fig. 10). Blinded observers performed 764
behavioral testing at three time-points: on naïve animals prior to any surgical 765
procedures, within one-week after catheter implantation and before drug treatment, and 766
at the six week post-SCI endpoint (also see Fig. 1). All naive animals exhibited a 767
baseline locomotor score of 21 (1 worst to 21 best). Five weeks after SCI and catheter 768
implantation, before any treatments, animals exhibited a mean BBB score of 13.6 ± 3.8, 769
demonstrating the expected locomotor ability in the late SCI recovery phase (Basso et 770
al., 1995). BBB testing of SCI animals after vehicle or drug delivery demonstrated no 771
significant affect on locomotor ability with scores remaining unchanged between SCI 772
animals treated with (n = 10) or without Rac1 inhibitor (n = 11) (p >0.05, 14.2 ± 1.6 vs. 773
13.8 ± 2; ANOVA on ranks). 774
775
Discussion 776
777
Spinal cord circuits can reorganize, changing in structure and function after injury 778
(Raisman, 1991). Our present findings demonstrate robust changes in dendritic spine 779
morphology on α-motor neurons after SCI, including an increase in dendritic spine 780
density, a distribution of spines closer to the cell body, and the presence of more mature 781
dendritic spines. These postsynaptic dendritic changes have been shown to accompany 782
increased neuronal excitability after SCI (Rall et al., 1992; Segev et al., 1998; Tan et al., 783
2009). In agreement, we observed a significant loss of H-reflex RDD below the injury 784
(i.e., increased H/M ratio), indicative of spasticity (Boulenguez et al., 2010; Matthews, 785
1966; Nielsen et al., 2007). Importantly, dendritic spines above the injury exhibited a 786
nearly opposite morphological profile with decreased spine density, and distribution and 787
shape that were more similar to control. As expected, there was no change in H-reflex 788
excitability above the level of injury. Overall, these results demonstrate that abnormal 789
dendritic spine profiles below the level of injury accompany spasticity after SCI, and 790
conversely, the lack of such spine profiles above the injury correspond with a lack of 791
spinal reflex hyperexcitability. 792
793
To further elucidate the structure-function link between dendritic spine dysgenesis and 794
hyperreflexia, we disrupted dendritic spine remodeling by targeting Rac1-signaling in 795
SCI animals. We have previously shown that Rac1 inhibition disrupts dendritic spine 796
remodeling in dorsal horn sensory neuron after SCI, nerve injury, and diabetes mellitus 797
(Tan et al., 2011; Tan et al., 2012b; Tan et al., 2008). In the present study, we observed 798
a decrease in spine density on α-motor neurons and a closer-to-normal distribution of 799
dendritic spines following treatment with NSC23766, a Rac1-inhibitor. NSC23766 800
treatment also decreased spine length and head diameter, and partially restored normal 801
H-reflex activity (i.e., increased RDD). Taken together, our study is the first to 802
demonstrate robust dendritic spine reorganization on α-motor neurons in the ventral 803
horn, which accompanies spasticity after SCI. We implicate Rac1-signaling as an 804
important mediator in both the structural and functional changes within the spinal reflex 805
pathway after injury. 806
807
Spasticity after SCI has been attributed to a variety of mechanisms within the spinal 808
reflex arc (Nielsen et al., 2007; Roy et al., 2012). Muscle spindle afferents may lose 809
either presynaptic inhibition or reciprocal inhibition. Alternatively, loss of Renshaw 810
interneuron activity, thought to mediate reciprocal inhibition, can trigger spasticity 811
(Nielsen et al., 2007). Evidence obtained from intracellular recordings of spinal 812
motoneurons also demonstrates increased motoneuron excitability, i.e., the ability to 813
generate action potentials, including the appearance of plateau potentials and persistent 814
inward sodium and calcium currents in rat motor neurons after SCI (Bennett et al., 815
2001b; Heckmann et al., 2005; Li et al., 2004), KCC2 downregulation (Boulenguez et al., 816
2010; Vinay et al., 2008), and sodium channel misexpression (Harvey et al., 2006; Li et 817
al., 2003). Inflammation, e.g., microgliosis, occurs in a number of nervous system injury 818
models, including SCI (Craner et al., 2005; Hains et al., 2006), and may contribute to 819
increasing excitability of neuronal populations within spinal circuits (Gwak et al., 2009; 820
Zhao et al., 2007b). Astrocyte activation after injury can potentially maintain 821
hyperexcitability (Scholz et al., 2007). Finally, maladaptive plasticity such as ‘collateral’ 822
or ‘reactive’ sprouting may contribute to altered spinal motor control (Boulenguez et al., 823
2010; Krenz et al., 1998; Nielsen et al., 2007; Raisman, 1994). Others have shown 824
altered dendrite branch length on motor neurons in the spinal cord that accompanies 825
spasticity after SCI (Kitzman, 2005). Although dendritic spine morphologies change on 826
pyramidal neurons in the motor cortex after SCI (Kim et al., 2006), the functional role for 827
these spine alterations is not firmly understood. Computer simulations have attempted 828
to predict the physiological contribution of dendritic spines on motor neurons (Rall et al., 829
1967); however, in vivo changes in dendritic spine structure have not been reported in 830
spinal cord motor pools. 831
832
Dendritic spine morphology partly determines synaptic function, and therefore provides 833
a visual clue into how neural networks function (Calabrese et al., 2006; Segev et al., 834
1998). Dendritic spines can reorganize rapidly following synaptic activity, e.g., activity-835
dependent plasticity, and increase in density which provides new or stronger synapses 836
(Halpain, 2000). Abnormal dendritic spine morphologies have been reported in a wide 837
spectrum of neuropsychiatric diseases, including post-traumatic stress disorder, 838
substance dependence and addiction, autism spectrum disorders, and mental 839
retardation (Halpain et al., 2005; Purpura, 1974). Although adaptive plasticity between 840
Ia afferents and spinal motor neurons can shape H-reflex response in both humans and 841
rodents (Thompson et al., 2009; Wolpaw, 1994), maladaptive plasticity can contribute to 842
pathological H-reflex function associated with hyperreflexia and spasticity (Lance, 1980; 843
Nielsen et al., 2007). In chronic SCI, hyperexcitability of the spinal stretch reflex, e.g., H-844
reflex, is thought to underlie spasticity, which manifests as a velocity-dependent 845
increase in tonic stretch reflexes, with uncontrollable “jerking” movement and abnormal 846
muscle tone, whereby muscle continually contract (Ashby et al., 1987; Lance, 1980; 847
Nielsen et al., 2007; Skold et al., 1999). In our study, we observed significant SCI-848
induced changes in dendritic spine morphologies on α-motor neurons below the injury 849
site, which accompanied a loss of RDD and a stabilization of the H/M ratio over a broad 850
range of nerve stimulation rates. Importantly, we observed only minor changes in M-851
wave response after SCI, indicating that changes in RDD and H/M ratio were primarily 852
due to mechanistic changes within the spinal cord monosynaptic circuit. 853
854
Although sacrocaudal injuries might better replicate some aspects of clinical spasticity 855
(Li et al., 2003; Ritz et al., 1992), these SCI models only allow studies of neurological 856
deficits in tail musculature, which are absent in human. To permit sufficient locomotor 857
ability for open-field behavioral assessment and H-reflex testing of hindlimb musculature, 858
a parallel of leg muscle groups in human, we performed contusion SCI at spinal 859
segment L2. As with all SCI animal studies, however, we encountered an observation 860
suggesting that our injury model also cannot entirely reflect the human SCI condition. In 861
contrast with human SCI at lower-thoracic or upper lumbar segments, which generally 862
produces some chronic negative motor signs, including flaccidity or lower limb 863
weakness (Doherty et al., 2002), we observed increased spinal motor reflex activity 864
associated with increase muscle tone six weeks after injury. Thus, it is important to 865
mention that our contusion SCI model was utilized as a compromise to study spinal 866
reflex function in hindlimb musculature. 867
We noted in our study that the RDD of the H-reflex did not exhibit depression at a 868
similar rate compared with an earlier report of the effect of spinal contusion on RDD 869
(Thompson et al., 1992b). Whereas Thompson et al. observed activity-rate depression 870
of approximately 85% of control at 5 Hz (Thompson et al., 1992a); we observed a 871
similar loss of H-reflex activity at 10-20 Hz (i.e., 50-100ms interpulse interval). A 872
probable explanation for this discrepancy is due to the additional procedures that 873
animals underwent prior to reflex testing in our study, including surgical implantation of 874
intrathecal catheters and infusions of vehicle or drug solutions. Although no effect of 875
catheter implantation and drug infusion have been observed in previous nociceptive 876
testing in control animals (Tan et al., 2008), it is possible that these additional 877
experimental procedures could have led to sensitization of afferents within the spinal 878
reflex circuit. Nonetheless, the magnitude of activity-rate depression in Sham and SCI 879
animals in our study fell within other documented ranges (Hosoido et al., 2009; Tan et 880
al., 2012a). 881
882
Exogenous electrical stimulation of the H-reflex circuit can directly induce changes in H-883
reflex response, which can persist for many hours or days (Chen et al., 2003; Chen et 884
al., 2006). These previous studies demonstrate the presence of activity-dependent 885
plasticity within the monosynaptic reflex system. In our current study, to establish a 886
structure-function relationship between dendritic spine remodeling and H-reflex 887
dysfunction after SCI, we were required to assess spine changes and H-reflex function 888
within the same animals. This acute H-reflex function and dendritic spine assessment 889
approach limited our ability to control for the possible confound that dendritic spine 890
changes could have also resulted from EMG reflex testing, which required direct 891
stimulation of muscle-sensory nerves. Although H-reflex testing only lasted about an 892
hour per animal and we ensured that all animals underwent similar testing protocols, we 893
cannot exclude the possibility that each treatment group could have had a different 894
capacity to respond to potential EMG testing-induced dendritic spine changes. We note 895
that in uninjured Sham animals, there were a greater proportion of thin-shaped dendritic 896
spines compared with mushroom-shaped dendritic spines (see Fig. 4 and 5). Thin, 897
filopodia-like dendritic spines are thought to represent newly formed or more plastic 898
dendritic spines; whereas mushroom shaped spines may represent more stable, mature 899
spines (Bourne et al., 2007; Bourne et al., 2008). This observation in Sham animals 900
does suggest the possibility that EMG testing could have resulted in the de novo 901
presence of more structurally responsive thin-shaped dendritic spines. Our current 902
experiments, however, do not permit us to determine whether spine changes are a sole 903
result of treatments, i.e., SCI, drug intervention, etc., or a combination of treatments and 904
the potential direct affect of EMG electrophysiological assessment, which could have 905
also influenced spine morphologies. 906
907
Our findings raise the question of how altered dendritic spine morphologies below the 908
level of injury contribute to hyperexcitable spinal reflex function. The dendritic spine 909
profiles we observed after SCI below the injury can have direct biophysical effects on 910
motor neuron excitability (Rall et al., 1967; Segev et al., 1988; Tan et al., 2009). The 911
distribution of spine synapses closer to motor neuron cell bodies can increase the 912
overall impact of excitatory input and transmission. The increased head volume of 913
mature, mushroom-shaped dendritic spines permits increased clustering of excitatory 914
membrane receptors, i.e., AMPA receptors (Wiens et al., 2005). These properties can 915
amplify synaptic transduction. Importantly, we observed an increase in mature spine 916
geometries after SCI, which could increase input discretization (e.g., narrow EPSP 917
waveforms) and allow trains of suprathreshold potentials to propagate at higher rates of 918
activity with greater fidelity (Rall, 1967; Rall et al., 1967). This transmission property 919
could contribute to excessive firing in motor neurons (Tan et al., 2009; Yuste et al., 920
2004) and facilitate supra- and sub-threshold temporal summation (Carter et al., 2007; 921
Gold et al., 1994; Martiel et al., 1994). In our study, temporal summation would most 922
likely have occurred at the shorter interpulse intervals during H-reflex testing. 923
Consecutive transcutaneous electrical stimulations to facilitate withdrawal reflex have 924
demonstrated maximal temporal facilitation at the 10-20 Hz range (Arendt-Nielsen et al., 925
1997; Arendt-Nielsen et al., 2000), suggesting the presence of a temporally dependent 926
integration mechanism within reflex pathways. Thus, it is interesting to consider that 927
SCI-induced dendritic spine changes leads to an amplification of presynaptic excitatory 928
input and enhancement of a temporally dependent integration mechanism to increase 929
the gain of H-reflex function after SCI. 930
931
We have implicated Rac1 in the pathological changes that contribute to chronic 932
nociceptive hyperexcitability after SCI (Tan et al., 2008). Rac1 is a small molecule 933
GTPase that is involved in regulating dendritic spine morphology (Tashiro et al., 2008). 934
In a mouse model of Fragile X syndrome (FMR knockout), Rac1 dysfunction is 935
associated with abnormal dendritic spine morphology and decreased pain (Chen et al., 936
2010; Lee et al., 2003; Price et al., 2007). In our model of SCI, we administered 937
NSC23766, a Rac1-specific inhibitor that does not affect cdc42 or Rho GTPases, or 938
affect the interaction of Rac1 to its downstream effector PAK1 (Gao et al., 2004). 939
Although our current study does not preclude off-target effects, we have previously 940
shown that the Rac1 inhibitor disrupts the development and appearance of dendritic 941
spines in vitro (Tan et al., 2011) and that NSC23766 treatment is effective in attenuating 942
dendritic spine remodeling in the dorsal horn after SCI (Tan et al., 2008). Moreover, 943
NSC23766 treatment does not significantly affect electrophysiological or behavioral 944
signs of pain-reflex withdrawal function in normal, uninjured animals (Tan et al., 2012b). 945
Although microglial activation has been implicated in altering the excitability of sensory 946
neurons after injury (Tan et al., 2012a; Zhao et al., 2007a), administration of NSC23766 947
does not appear to affect the activation of microglia after nerve injury (Tan et al., 2011). 948
In agreement with previous studies (Beauparlant et al., 2013; Kitzman, 2007), we report 949
that VGluT1 expression did not change after SCI, suggesting that the presence of H-950
reflex hyperexcitability is not due to increased excitatory presynaptic afferent inputs 951
(Kitzman, 2007). Although it is possible that Rac1-inhibition may have affected 952
presynaptic elements, NSC23766 treatment only decreased VGluT1 areal density in the 953
intermediate zone and ventral horn. This suggests that the Rac1-inhibitor had a 954
topographically specific effect on deeper lamina only, where sensory-motor neuron 955
populations exhibit dendritic spine plasticity after injury (Tan et al., 2011; Tan et al., 956
2008). 957
958
A major challenge in SCI research has been identifying the mechanisms that contribute 959
to secondary complications such as spasticity and pain. Our previous work has 960
demonstrated that dendritic spine remodeling in the dorsal horn contributes to 961
hyperexcitability associated with neuropathic pain after SCI (Tan et al., 2008; Tan et al., 962
2012c). Here we show that dendritic spine dysgenesis on α-motor neurons also 963
contribute to the development of spasticity. Our data further show that Rac1 signaling 964
participates in this dendritic spine remodeling and may provide a novel opportunity to 965
specifically address the underlying pathophysiology of spasticity after SCI. 966
967
968
969
Figure Legends 970
971
Fig. 1. Study design. All weight-matched animals underwent BBB locomotor testing to 972
obtain baseline behavioral data. Animals (n values) in each group are shown. In week 1, 973
animals were randomly assigned to receive Sham or SCI surgical procedures. In week 974
5, animals received intrathecal catheter implants. After 2-3 days of recovery, we 975
performed pre-treatment BBB testing and immediately administered intrathecal 976
infusions of control vehicle or NSC23766 (twice a day for 3 days). At experimental 977
endpoint at week 6, these treatments produced four comparator groups (gray shade). 978
Note that within SCI animals treated with vehicle we assessed and compared data 979
outcomes from above or below the injury site, i.e., forelimb and hindlimb. At endpoint, 980
we also performed post-treatment BBB testing, H-reflex assessment, and histological 981
analysis. 982
983
Fig. 2. Spinal cord injury. (A) Contusion injury at L2 resulted in severe damage of the 984
dorsal columns and gray matter, as shown by GFAP staining in coronal spinal cord 985
tissue sections. Asterisks (*) denotes lesion epicenter. (B) Intact spinal cord tissue from 986
Sham. (C-D) Biomechanical data provided by the IH impactor demonstrated no 987
difference between vehicle and Rac1-inhibitor treated SCI groups. (E) Six weeks after 988
SCI, PKC-γ) staining produced bilateral labeling of the dorsal corticospinal tract (dCST) 989
and lamina I/II. (F) At the lumbar level, L5, (below the injury), the absence of PKC-γ 990
immunoreactivity in the dorsal columns white matter tracts demonstrated significant 991
disruption of the dCST. SCI did not affect PKC-γ staining in superficial laminae. Scale 992
bars = 500 μm 993
994
Fig. 3. Golgi staining of spinal cord tissue reveals dendritic spines on motor 995
neurons in the ventral gray matter. (A) Image of the ventral gray matter with an 996
identified α-motor neuron located in Rexed lamina IX (arrow and white box). (B) High 997
power field of motor neuron shown in the inset in panel A. Six weeks after Sham and 998
SCI, representative images of dendritic branches show apparent differences in dendritic 999
spine profiles from (C) Sham, (D) SCI + Vehicle above the injury, (E) SCI + Vehicle 1000
below the injury, and (F) SCI with NSC23766, Rac1-inhibitor, treatment groups. (C’, D’, 1001
E’, F’) High magnification of selected dendrite regions from panels C-F (red boxes). 1002
Scale bar for A = 500 μm; B = 100 μm; C-F = 10 μm; C’-F’ = 2 μm 1003
1004
Fig. 4. Digital reconstructions of spinal cord motor neurons. To obtain an accurate 1005
profile of dendritic spines in motor neurons, we digitally reconstructed the entire branch 1006
structure of sampled neurons. (A, B, C, D) Contour traces from each group show the 1007
locations of all sampled motor neurons (red dots) within the gray matter (representative 1008
black trace). Density and distribution were measured from three-dimensional neuron 1009
reconstructions from (A’) Sham, (B’) SCI + Vehicle above the injury, (C’) SCI + Vehicle 1010
below the injury, and (D’) SCI + Rac1-inhibitor treatment. (A”, B”, C”, D”) An ~50 μm 1011
length of dendrite from neurons shown in panels A’-D’ (gray shaded region) show thin-1012
shaped (blue dots) and mushroom-shaped spines (red dots). Scale bar in A, B, C, D = 1013
500μm; A’, B’, C’, D’ = 50μm, A”, B”, C”, D” = 10μm 1014
1015
Fig. 5. Quantitative analysis of dendritic spine profiles between Sham and SCI 1016
animals above or below the injury. Analysis of dendritic spine profiles reveals 1017
differences in dendritic spine density (top row), distribution (middle row), and shape 1018
(bottom row). (A) Total dendritic spine density, which includes all spine shapes, (B) thin-1019
spines, and (C) mushroom spines decreased on motor neurons located above the injury 1020
after SCI compared with Sham (* = p<0.05). In contrast, total spine density increased on 1021
motor neurons located below the injury compared with either Sham or above the injury 1022
after SCI (* = p<0.05). Dendritic spine distribution for (D) total, (E) thin, and (F) 1023
mushroom spines differed across the comparator groups. At proximal regions in SCI 1024
animals, all spine densities increased below the injury as compared with Sham and 1025
motor neurons above the injury (* = p<0.05). In contrast, neurons above the injury had 1026
less total and thin-shaped spine density at proximal regions compared with Sham and 1027
below the injury (* = p<0.05). (F) Although mushroom-spine density on motor neurons 1028
above the injury did not differ from Sham at proximal regions, these neurons had 1029
significantly less mushroom spine density compared with below the injury. At distal 1030
regions, motor neurons below the injury had greater spine density in all categories 1031
compared with motor neurons above the injury (* = p<0.05). There was no difference in 1032
any spine densities at distal regions on neurons from Sham and above the injury in SCI 1033
animals. Dendritic spine shape analysis revealed no change in (G) spine length or (H) 1034
spine head diameter on motor neurons located above the injury in SCI animals 1035
compared with Sham (* = p<0.05). Below the injury, these measurements demonstrated 1036
a decrease in spine length, and an increase in spine head diameter compared with 1037
neurons in Sham and above the injury (* = p<0.05). 1038
1039
Fig. 6. Rate-dependent depression of the H- and M- responses above and below 1040
SCI. As a physiological assessment of the monosynaptic H-reflex, we performed a 1041
paired-pulse stimulation protocol. Representative traces (averaged 10-20 traces) of the 1042
M- and H-responses to control (first) and test (second) pulse in (A) Sham, (B) SCI 1043
above the injury, and (C) SCI below the injury. The control and test pulses were 1044
separated with a range of interpulse latencies between 2000 – 10ms. Note that in Sham 1045
animals, as the interpulse intervals decreased (e.g., increasing the rate of activity) 1046
between the test and control pulse, the amplitude of the M- and H-response decreased. 1047
As shown in panel C, in SCI below the injury, rate-dependent depression in amplitude of 1048
either the M- or H-response failed to appear. (D) %H-reflex and (E) %M-wave 1049
amplitudes are normalized values of the evoked stimulus-response of the test and 1050
control pulse. (D) After SCI, there was no significant difference in % H-reflex in SCI 1051
above the injury compared with Sham at any interpulse interval. In contrast, in SCI 1052
animals below the injury, the %H-reflex significantly increased compared with Sham at 1053
the shortest interpulse intervals between 100-10ms (* = p<0.05), demonstrating a loss 1054
of RDD and increased excitability of the H-reflex. Similarly, %H-reflex below the injury 1055
was significantly greater than above the injury in SCI animals at 500, 50, and 10ms 1056
interpulse intervals (§ = p<0.05). (E) % M-wave demonstrated a significantly increased 1057
response below the injury compared with both Sham (* = p<0.05) and above the injury 1058
(# = p<0.05) response. (F) The H/M ratio was calculated from M-wave and H-wave 1059
responses. 1060
1061
Fig. 7. Rac1-inhibitory treatment disrupts dendritic spine morphology on motor 1062
neurons in the ventral horn after SCI. Treatment with NSC23766 in SCI animals (SCI 1063
+ anti-Rac) resulted in a significant decrease in (A) total, (B) thin, and (C) mushroom-1064
spine density compared with SCI + Veh (* = p<0.05). (D, E, F) Assessment of dendritic 1065
spine distribution on motor neurons showed that NSC23766 treatment in SCI animals 1066
resulted in decreased spine density for all spine categories at both proximal and distal 1067
branch regions (* = p<0.05). NSC23766 treatment decreased SCI-induced (G) spine 1068
length and (H) spine head diameter compared with SCI + Veh (* = p<0.05). 1069
1070
Fig. 8. Disruption of Rac1-regulated dendritic spines reduces SCI-induced H-1071
reflex hyperexcitability. Representative traces show the M- and H-responses from 1072
paired-pulse testing in (A) SCI + Veh, below the injury and (B) SCI + anti-Rac treatment. 1073
(C) Quantification of the %H-reflex response demonstrated that the H-reflex response in 1074
SCI + Veh animals exhibited reduced RDD (also see Fig. 6). Rac1-inhibitor treatment in 1075
SCI animals reduced the %H-reflex at 100, 50, and 10 ms as compared with SCI + Veh 1076
(* = p<0.05). (D) There was no significant difference in the %M-wave between SCI + 1077
Veh and SCI + anti-Rac. (E) Treatment with the Rac1-inhibitor in SCI animals 1078
decreased the H/M ratio compared with SCI + Veh, as demonstrated by a steeper 1079
downward trend line. 1080
1081
Fig. 9. Excitatory terminals in the spinal cord gray matter. VGluT1-immunopositive 1082
puncta appeared throughout all laminae of the spinal cord gray matter in the lumbar 1083
enlargement, L4-L5 (left panels in A-C). Spatial heat maps (right panels in A-C; red 1084
highest density, blue lowest density) shows the overall areal density of VGluT1 1085
expression in (A) Sham, (B) SCI + Veh, and (C) SCI + anti-Rac treatment. 1086
Quantification of the VGluT1 puncta within the (D) total gray matter region, (E) the 1087
dorsal horn, (F) the intermediate zone, and (G) the ventral horn (panel insets and gray 1088
shade) demonstrated no significant change in SCI + Veh compared with Sham. 1089
Treatment with the Rac1-inhibitor in SCI animals decreased VGluT1 areal density 1090
compared with SCI + Veh in the total gray matter, intermediate zone, and ventral horn 1091
only (* = p<0.05) with no significant change in the dorsal horn (p > 0.05). The areal 1092
density of VGluT1 decreased in SCI + anti-Rac1 compared with Sham in the dorsal horn 1093
(* = p<0.05). Scale bar for A, B, C = 500 μm. 1094
1095
Fig. 10. Locomotor testing. Blinded observers performed BBB testing on animals at 1096
three time points: before any procedure, before treatment, and after treatment. All naive 1097
animals exhibited a baseline locomotor score of 21. There were no significant 1098
differences in BBB scores across group (p>0.05). 1099
1100
1101
1102
1103
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Acknowledgements: The work is supported by grants from the Paralyzed Veterans of 1445
America (PVA) and the Department of Veterans Affairs (VA) Medical Research Service 1446
and Rehabilitation Research Service. Andrew M. Tan is funded by the PVA Research 1447
Foundation and a VA Career Development Award (1 IK2 RX001123-01A2). The Center 1448
for Neuroscience and Regeneration Research is a Collaboration of the Paralyzed 1449
Veterans of America with Yale University. We thank Pamela Zwinger and Peng Zhao for 1450
their excellent technical assistance. 1451
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Table 1. Spinal cord motor neuron morphometry
Maximum cell
diameter (um) Aspect Ratio Form Factor
Number of
primary
dendrites
Total dendrite
length (m)
Sham 54.6 ± 19.4 0.57 ± 0.17 0.49 ± 0.17 8.16 ± 3.69 1073.0 ± 671.4
SCI (above injury) 46.7 ± 11.7 0.51 ± 0.16 0.50 ± 0.16 5.73 ± 1.95 815.10 ± 451.9
SCI (below injury) 51.4 ± 17.4 0.64 ± 0.13 0.62 ± 0.09 4.80 ± 1.23 739.30 ± 280.6
SCI + anti-Rac 46.1 ± 8.7 0.63 ± 0.11 0.60 ± 0.10 4.38 ± 1.15 1085.8 ± 307.9 Data are shown as mean ± SD