Dendritic spine dysgenesis contributes to hyperreflexia after spinal

Dendritic spine dysgenesis contributes to hyperreflexia after spinal cord injury

Samira P. Bandaru,1,2 Shujun Liu,1,2 Stephen G. Waxman,1,2 and Andrew M. Tan1,2

1Department of Neurology and Center for Neuroscience and Regeneration Research, Yale University School of Medicine,New Haven, Connecticut; and 2Rehabilitation Research Center, Veterans Affairs Connecticut Healthcare System,West Haven, Connecticut

Submitted 30 July 2014; accepted in final form 5 December 2014

Bandaru SP, Liu S, Waxman SG, Tan AM. Dendritic spinedysgenesis contributes to hyperreflexia after spinal cord injury. JNeurophysiol 113: 1598–1615, 2015. First published December 10,2014; doi:10.1152/jn.00566.2014.—Hyperreflexia and spasticity arechronic complications in spinal cord injury (SCI), with limited optionsfor safe and effective treatment. A central mechanism in spasticity ishyperexcitability of the spinal stretch reflex, which presents symp-tomatically as a velocity-dependent increase in tonic stretch reflexesand exaggerated tendon jerks. In this study we tested the hypothesisthat dendritic spine remodeling within motor reflex pathways in thespinal cord contributes to H-reflex dysfunction indicative of spasticityafter contusion SCI. Six weeks after SCI in adult Sprague-Dawleyrats, we observed changes in dendritic spine morphology on �-motorneurons below the level of injury, including increased density, alteredspine shape, and redistribution along dendritic branches. These ab-normal spine morphologies accompanied the loss of H-reflex rate-dependent depression (RDD) and increased ratio of H-reflex to M-wave responses (H/M ratio). Above the level of injury, spine densitydecreased compared with below-injury spine profiles and spine dis-tributions were similar to those for uninjured controls. As expected,there was no H-reflex hyperexcitability above the level of injury inforelimb H-reflex testing. Treatment with NSC23766, a Rac1-specificinhibitor, decreased the presence of abnormal dendritic spine profilesbelow the level of injury, restored RDD of the H-reflex, and decreasedH/M ratios in SCI animals. These findings provide evidence for anovel mechanistic relationship between abnormal dendritic spineremodeling in the spinal cord motor system and reflex dysfunction inSCI.

H-reflex; hyperreflexia; Rac1; spasticity; spinal cord injury

HYPERREFLEXIA AND SPASTICITY, which arise in up to 60% ofpatients with spinal cord injury (SCI), can severely affectquality of life, contribute to chronic pain, and lead to muscu-loskeletal deformity (Skold et al. 1999; Walter et al. 2002).Although currently available drugs, such as baclofen, canprovide some relief, these drugs have limited therapeutic utilityand effectiveness. Thus there is a significant need for a morecomplete understanding of spasticity and for more effectivetreatment after SCI.

Central mechanisms that underlie pathological reflex controlafter injury or disease include the loss of cortical and localspinal inhibition, injury-induced plasticity, and increased mo-tor neuron excitability (Bennett et al. 2001a; Boulenguez andVinay 2009; Hultborn and Nielsen 2007; Hunanyan et al.2013). Whereas plasticity between Ia afferents and �-motorneurons shapes the H-reflex response in an activity-dependentmanner in human and rodent (Raisman 1994; Thompson et al.

2009), maladaptive changes can also contribute to pathologicalH-reflex function associated with spasticity, clinically definedas a velocity-dependent increase in tonic stretch reflexes withexaggerated tendon jerks, resulting from hyperexcitability ofthe spinal stretch reflex (Ashby et al. 1987; Lance 1980;Nielsen et al. 2007).

Dendritic spines, micron-sized structures that are sites ofpostsynaptic activity, regulate the efficacy of synaptic trans-mission and can thereby alter the electrical information passingthrough circuit pathways (Bourne and Harris 2007; Calabreseet al. 2006; Pongracz 1985; Segev and Rall 1988; Tan et al.2009). Localized increases in synaptic strength through the denovo formation and development of postsynaptic dendriticspines constitute a persistent structural basis for learning andmemory in the central nervous system (Xu et al. 2009; Yusteand Bonhoeffer 2001). In the present study, we assess thepossibility that abnormalities in dendritic spine morphology on�-motor neurons contribute to the persistent dysfunctional statewithin the spinal motor reflex pathway after SCI.

Our previous studies and evidence in the literature demon-strate that dendritic spine morphology can change followingdisease or injury (Kim et al. 2006; Tan et al. 2008, 2012b,2013). Importantly, adverse changes in spine morphologyincluding 1) the elaboration from thin, filopodia-like spines toa mushroom shape, a morphology associated with increasedsynaptic strength and stability (Yuste and Majewska 2001),and 2) an increase in spine density along dendrites, whichprovides more sites for postsynaptic connections (Bonhoefferand Yuste 2002), and a spatial redistribution of spines alongdendrites to locations closer to the cell body (Kim et al. 2006;Ruiz-Marcos and Valverde 1969) have been shown to contrib-ute to neuronal hyperexcitability (Tan et al. 2009). Althoughdendritic spine remodeling occurs in the motor cortex after SCI(Kim et al. 2006), no study has reported on dendritic spineslocated on spinal �-motor neurons. Moreover, it is unknownwhether SCI-induced changes in dendritic spine morphologiescan contribute to spasticity.

The activity of small GTP-binding protein Rac1 governsactin cytoskeleton reorganization to regulate dendritic spinemorphology (Tashiro et al. 2000; Tashiro and Yuste 2004).Constitutively activated Rac1 increases the rate of dendriticspine turnover, spine density and stability, and spine volume(Nakayama and Luo 2000). In contrast, dominant negativeRac1 expression or administration of a Rac1-specific inhibitorNSC23766 disrupts dendritic spine formation and development(Tan et al. 2011; Tan and Waxman 2012; Tashiro et al. 2000;Tolias et al. 2007). Importantly, SCI increases Rac1 mRNAexpression, with levels that can remain elevated for up to 3 moor more (Dubreuil et al. 2003; Erschbamer et al. 2005). It is not

Address for reprint requests and other correspondence: A. M. Tan, Centerfor Neuroscience and Regeneration Research (127A), 950 Campbell Ave.,Bldg. 34, West Haven, CT 06516 (e-mail: [email protected]).

J Neurophysiol 113: 1598–1615, 2015.First published December 10, 2014; doi:10.1152/jn.00566.2014.

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known, however, if Rac1 activity contributes to dendritic spineremodeling and reflex dysfunction following SCI.

In this article we provide the first evidence of dendritic spineplasticity on �-motor neurons after SCI and demonstrate astructure-function relationship between dendritic spine dysgen-esis and exaggerated spinal motor reflexes associated withspasticity. Six weeks after contusion SCI, animals exhibitedincreased H-reflex responsiveness (i.e., shown by reducedrate-dependent depression, or RDD). Histological assessmentin these animals demonstrated that �-motor neurons locatedbelow the level of injury within the L4–L5 spinal segments hadan increase in dendritic spine density, a significant redistribu-tion of spines along dendrites, and increased dendritic spinehead diameter, morphological profiles consistent with thoseshown to contribute to increased neuronal excitability (Pon-gracz 1985; Tan et al. 2009). In contrast, above the level ofinjury, we observed an absence of exaggerated H-reflex re-sponse, reduced dendritic spine densities, a close-to-normaldistribution of spines, and normal dendritic spine length andhead diameter. Inhibition of Rac1 disrupted SCI-induced den-dritic spine profiles on �-motor neurons below the injury site,reduced vesicular glutamate transporter 1 (VGluT1) expression(a marker for excitatory primary afferent terminals), and de-creased H-reflex responsiveness. Taken together, these obser-vations provide evidence for a new perspective into mecha-nisms of neuroplasticity within the spinal reflex pathway anddemonstrate a relationship between dendritic spine remodelingand reflex dysfunction after SCI. Targeting of molecular path-

ways that regulate spine structure could represent a novelavenue for managing spasticity after SCI.

MATERIALS AND METHODS

Animals and spinal cord injury. Experiments were performed inaccordance with the National Institutes of Health Guidelines for theCare and Use of Laboratory Animals. All animal protocols wereapproved by the Yale University Institutional Animal Use Committee.Animals were housed under a 12:12-h light-dark cycle in a pathogen-free area with water and food provided ad libitum. A total of 32 adultmale Sprague-Dawley rats (175–200 g; Harlan, Indianapolis, IN)underwent procedures to produce each treatment group (Fig. 1; studydesign: sham, n � 11; SCI � vehicle, n � 11; SCI � anti-Rac, n �10). Animals were first divided into two treatment arms (Fig. 1). Thefirst group received a contusive spinal cord injury at the 2nd lumbarspinal segment (L2): animals were anesthetized with a mixture ofketamine (80 mg/kg ip) and xylazine (5 mg/kg ip). A small laminec-tomy was carefully performed at the 12th thoracic vertebra (T12),which exposed the dorsal L2 spinal cord surface (Hebel and Strom-berg 1976). We stabilized the spinal cord in an Infinite Horizon (IH)impactor device (Precision Systems and Instrumentation, LexingtonKY) by clamping the rostral T11 and caudal T13 vertebral bodies withAdson stabilizing forceps attached to the IH stage (Scheff et al. 2003).The spinal contusion injury was performed with a metal rod (tipdiameter 2.5 mm) that was applied to the spinal cord surface with animpact force of 170 kdyn (Rabchevsky et al. 2003; Scheff et al. 2003)(data shown in Fig. 2). For sham animals (without SCI), the samesurgical procedure was followed, including placement of the animalwithin the IH stabilizing forceps, except no contusion injury wasperformed. Following all surgical procedures, the muscle, fascia, and

Fig. 1. Study design. All weight-matched animals underwent Basso, Beattie, and Bresnahan (BBB) locomotor testing to obtain baseline behavioral data. Thenumber of animals (n values) in each group are shown. In week 1, animals were randomly assigned to receive sham or spinal cord injury (SCI) surgicalprocedures. In week 5, animals received intrathecal catheter implants. After 2–3 days of recovery, we performed pretreatment BBB testing and immediatelyadministered intrathecal infusions of control vehicle or NSC23766 (twice a day for 3 days). At experimental endpoint at week 6, these treatments produced 4comparator groups (gray-shaded boxes). Note that within SCI animals treated with vehicle (SCI � Veh), we assessed and compared data outcomes from aboveor below the injury site (i.e., forelimb and hindlimb, respectively). At endpoint, we also performed posttreatment BBB testing, H-reflex assessment, andhistological analysis.

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skin were sutured in sequential layers with 4-0 monofilament sutures.Postoperative treatments included twice daily injections of 0.9%saline solution for rehydration (3.0 ml sc) and Baytril (0.3 ml, 3.5mg/kg body wt sc, twice daily for 3 days) to prevent urinary tractinfection.

Behavior. Two experimenters blinded to group assignment evalu-ated animals using the Basso, Beattie, and Bresnahan (BBB) locomo-tor rating scale (Basso et al. 1995) for validation of injury equivalencyacross SCI animals, as well as to determine whether treatments had aneffect on overall locomotor ability. The BBB score (1 � worst to 21 �best) consists of a combination of hindlimb movements, trunk positionand stability, hindlimb stepping and coordination, paw placement, andtail position. Behavioral testing was performed at three time points(Fig. 1): 1) on naive animals before any surgical procedures, 2) within1 wk after catheter implantation (and before any drug infusions at �5wk post-SCI), and 3) immediately before experimental endpoint (6 wkpost-SCI and sham surgeries). Before undergoing any testing, animalswere allowed to acclimatize to the testing area for 60–90 min. Duringan experimental trial, animals were allowed to roam freely in the testfield (enclosed 3 � 3-ft. flat surface), and a similar 4-min time frameof movement was assessed by the experimenters using the BBB scale,as previously described (Basso et al. 1995). After each trial, thesurface was cleaned with soap and water and dried. For analysis, BBBscores from both right and left sides of animals were averaged, anddata from the two experimenters were averaged within groups andthen statistically compared across groups.

Intrathecal catheter implantation and drug delivery. Five weeksafter SCI or sham surgeries, all animals received ketamine-xylazineanesthesia (80 and 5 mg/kg ip, respectively). As described previously(Tan et al. 2012b), a small craniotomy was performed to expose theatlanto-occiptal membrane (between the occipital bone and vertebralcolumn C1/atlas). A sterile 32-gauge catheter (ReCathCo, AllisonPark, PA) was carefully inserted through a slit in the membrane andthreaded intrathecally until the tip of the catheter reached the lumbarenlargement. The catheter was secured near the base of the skull withsutures placed through overlying muscle and skin. To prevent leakageand infection, we heat-sealed the exposed rostral tip of the catheter bypinching the end with a sufficiently heated and sterilized forceps. Thelocation of the caudal end of the catheter was validated at theexperimental endpoint after animals were killed. Animals were al-lowed to recover for 2–3 days after catheter implantation and thenreceived one of two infusions through the catheter: 1) drug vehicle(0.9% sterile saline, 10-�l volume, twice daily for 3 days) or 2)NSC23766, a target-specific Rac1-GTPase inhibitor (EMD Chemi-cals, Darmstadt, Germany), at 2.65 �g/�l (5-�l volume, twice dailyfor 3 days) followed by a 5-�l sterile 0.9% saline flush. To measurethe maximal effect of treatments, we performed experimental assess-ments within 1–2 days following the last infusion of vehicle or drugsolution. We did not infuse NSC23766 drug in sham animals in thisstudy, because we had previously already established that NSC23766does not significantly affect higher-order electrophysiological or be-havioral function in uninjured, control animals (Tan et al. 2012b). Atthe end of the study, this study design produced four comparator arms(Fig. 1, gray-shaded boxes): sham, SCI � vehicle [includes “SCI(above injury)” and “SCI (below injury)”], and SCI � anti-Rac.

Histology. For Golgi-Cox staining with the use of a commercial kitand according to the manufacturer’s instructions (FD Neurotechnolo-gies, Ellicott, MD), a subpopulation of rats (sham, n � 5; SCI �vehicle, n � 4; SCI � anti-Rac, n � 5) from terminal electrophysi-ological recordings (see below) under ketamine-xylazine anesthesiawere killed and processed. Spinal cord tissue (from the cervicalenlargement, C4–C5, and lumbar enlargement, L4–L5) was quicklyremoved (�5 min), rinsed in distilled water, and immersed in the kit’simpregnation solution. After the incubation period (�3 wk), 200-�m-thick sections were cut on a vibratome (DTK-1000 microslicer; TedPella) and mounted on gelatinized glass slides. Sections were stained,rinsed in distilled water, dehydrated, cleared, and coverslipped with

Permount medium. For immunohistochemistry, remaining rats weredeeply anesthetized with ketamine-xylazine and transcardially per-fused with 250 ml of 0.1 M phosphate buffer (PB) at 37°C followedby 300 ml of freshly prepared cold paraformaldehyde solution (4% in0.1 M PB). The spinal cord was removed, postfixed for 2 h at roomtemperature, and cryoprotected by immersion in 30% sucrose in 0.1 MPB at 4°C. Frozen coronal sections from C4–C5, the injury site at L2,and L4–L5 were cut at 20-�m thickness using a cryostat (Leica,Bannockburn, IL). Sections were collected onto Superfrost Plus slides(Fischer Scientific, Pittsburgh, PA). Immunofluorescence stainingmethods were described previously (Tan et al. 2006). Sections werewashed in blocking solution (0.1 M PBS, 0.1% Triton X-100, and 4%normal donkey serum) and incubated overnight at 4°C in mouseanti-VGluT (1:1,000; UC Davis/NIH NeuroMab facility), rabbit anti-glial fibrillary acidic protein (1:2,000; Abcam) or rabbit anti-PKC-�antibody (Santa Cruz Biotechnology 1:1,000). After being washed inblocking solution, sections were incubated in the fluorescent secondaryantibodies CY3 donkey anti-mouse (1:500; Jackson ImmunoResearchLaboratories) or Alexa Fluor 488 donkey anti-rabbit (1:2,000;Invitrogen). Sections were visualized and digitally imaged using aNikon Eclipse 80i fluorescence microscope equipped with an HQCoolSNAP camera (Roper Scientific, Tucson, Arizona) or a NikonD-Eclipse C1 confocal microscopy system. MultiCapture mosaicimages were digitally stitched with NIS Elements software (NikonInstruments).

Dendritic spine visualization on motor neurons and analysis. In-vestigators blinded to treatment conditions performed all imagingstudies and analyses. To visualize neurons and ultrafine processes, weused a Golgi-staining method as previously described (Tan et al.2008). For our purpose, we required the ability to fully reconstructneuronal structure, which required that tissue be exposed to high-intensity light for long periods of time (up to 4 h per imaging session),which can quickly bleach or diminish other visualization tools (e.g.,fluorophores). Golgi staining permits the identification and samplingof a relatively large number of neurons from cervical and lumbarlevels within the same animal and provides robust visualization of theentire neuronal structure, including detailed resolution of dendriticspines. We were specifically interested in motor neuron pools thatinnervated muscle groups in the forelimb (i.e., extensor carpi radialis)and hindlimb (i.e., plantar muscle). To identify these �-motor neu-rons, we followed a screening workflow based on data from ourprevious study (Tan et al. 2012a) and those previously validated inrats (Crockett et al. 1987; Hashizume et al. 1988; Jacob 1998). Webegan with a broad sample population of neurons by identifyingGolgi-stained �-motor neurons located in the ventral spinal cord inRexed lamina IX and with soma diameters �25 �m (Hashizume et al.1988; Jacob 1998). Above the injury site (C4–C5), we narrowedcandidate neurons for analysis by selecting neurons from motor poolslocated in similar dorsolateral coordinates of �-motor neurons shownto innervate the extensor carpi radialis muscle group (�1.5–2 mmdeep, 1.5–2 mm lateral from midline), as we and others have dem-onstrated through intramuscular retrograde tracing studies (Sunshineet al. 2013; Tan et al. 2012a; Tosolini et al. 2013). Below the injurysite (L4–L5), we narrowed our sampled �-motor neurons to thoselocated in ventral motor pools with similar dorsolateral coordinates ofmotor pools known to innervate the plantar muscle (�1.5–2.5 mmdeep, 1.5–2.2 mm lateral from midline) as shown by retrogradetracing (Crockett et al. 1987; Jacob 1998). As a refinement step foranalysis a priori, we only included �-motor neurons for analysis thathad 1) dendrites and dendritic spines that were clearly and completelyimpregnated, 2) dendritic branches appearing as a continuous lengthfor at least 350 �m within the tissue slice, and 3) at least one-half ofthe primary dendritic branches remaining within the thickness of thetissue section such that their endings were not cut and appeared totaper into a complete ending (see representative neuron in Fig. 3). Todetermine if there were any morphological differences across oursample neurons, we used NeuroExplorer software (MicroBrightField,

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Williston, VT) to measure maximum cell diameter, aspect ratio (Feretmaximum/Feret minimum), form factor [(4� � area)/(perimeter)2],number of primary dendrites, and total dendritic branch length of eachtreatment arm and compared these morphometry values across treat-ment groups (Table 1). To refine our identification and measurementsof dendritic spines, we used specific morphological characteristics(Kim et al. 2006; Tan et al. 2012b): we defined a spine neck asthe structure between the base of the spine and the interface with theparent dendrite branch, and the base of the spine head where theappearance of the spine swells distally into a bulblike structure. Thinand mushroom-shaped spines were classified as follows: thin spineshad head diameters that were less than or equal to the length of thespine neck, whereas mushroom spines had head diameters that weregreater than the length of the spine neck. These criteria for two spinegeometric categories were used because classification into only twospine shapes allowed us to use simple but very strict rules in classi-fying spine morphology. Although this approach prevented the dis-crimination of subtle variations in spine shape, it allowed the collec-tion of a very large sample size, and we and others have described thephysiological characteristics of thin and mushroom spine shapes onneuronal and circuit function (Bourne and Harris 2007; Holmes 1990;Tan et al. 2009). Note that these criteria do not imply the physiolog-ical characterization of the neurons we analyzed, but rather control formorphological diversity within the sampled spinal motor neuronpopulation (Kitzman 2005; Tashiro and Yuste 2003).

To digitally reconstruct motor neurons, we used a Neurolucidasoftware suite (version 9.0; MicroBrightField) and a pen tablet (In-tuos5 Touch; Wacom). We analyzed the completed three-dimensionalreconstructions of motor neurons for spine density and distribution.Each imaging session consisted of a contour map (outline of the spinalcord section with location of identified neuron) and the motor neuron,which was traced in the X-, Y-, and Z-axes. Dendritic spine type werelocated and marked on each reconstructed dendritic branch (thinspines, blue; mushroom spines, red). Dendritic spine density wasexpressed as dendritic spine number per 10-�m dendrite length. Todetermine any changes in spatial distribution of dendritic spinesrelative to the cell body, we used a Sholl’s analysis (Tan et al. 2008).Seven 50-�m-wide spherical bins were formed around each cell body,and spine density within each bin was averaged within each treatmentgroup. For statistical comparison, spine density at dendrite branchlocations within 50–150 �m (proximal bins) and 200–350 �m (distalbins) from the cell body were pooled and compared across treatmentgroups.

To determine changes in spine dimensions, five neurons werearbitrarily chosen from each treatment group and visible spines weremeasured for spine length and spine head diameter (Kim et al. 2006).Spine length was defined as the distance from the tip of the spine tothe junction of the spine at the main dendrite branch. Spine headdiameter was defined as the longest line drawn normal to the length ofthe parent dendrite branch. A total of 411 dendrites from 82 identified�-motor neurons (in 4–5 animals per treatment group) were includedin our analyses [sham, 118 dendrites; SCI (below injury), 116 den-drites; SCI (above injury), 90 dendrites; SCI � anti-Rac, 87dendrites].

Areal density of VGluT1 labeling. To examine changes in thenumber of VGluT1-expressing synaptic terminals, we assessed areal

density of VGluT1 expression using a modified approach describedpreviously (Tan et al. 2012a). High-resolution digital photographswere taken at �10 magnification and combined into a single mosaicimage of the entire spinal cord at L4–L5. We photographed 10sections per animal. Sections for analysis were chosen on the basis oftissue integrity (e.g., no major tears) and equivalent loss of PKC-�immunoreactivity in the dorsal corticospinal tract (dCST) of the spinalcord dorsal columns. Sections were aligned according to the point ofintersection between the gray matter above the central canal and thedorsal median septum. All images underwent threshold adjustmentsusing equivalent contrast/brightness levels to highlight only VGluT1-expressing puncta (Photoshop; Adobe, San Jose, CA). Images werebinarized and color-inverted for analysis. For color-coded heat mapsin Fig. 9, binarized images were exported into MATLAB (TheMathWorks, Natick, MA) and averaged using custom scripts, asdescribed previously (Brus-Ramer et al. 2007; Friel and Martin 2007).For image analysis, mosaic images of each spinal cord coronal sectionwere divided into three dorsoventral regions corresponding to thedorsal zone (�lamina I–III), the intermediate zone (�lamina IV–VI),and the remaining ventral horn of the gray matter (Tan et al. 2012a).Because we were only interested in VGluT1 in the gray matter, thewhite matter areas were digitally removed before analyses. BecauseVGluT1-expressing puncta were visually distinct from each other,without overlap, the number of VGluT1 puncta was easily countedin each region using ImageJ software (http://rsb.info.nih.gov/ij/index.html). Data from gray matter regions were pooled within groupsand compared across experimental treatment groups.

H-reflex testing. Terminal electrophysiological experiments wereperformed 6 wk after SCI or sham surgeries. Because H-reflexresponses in rats under ketamine anesthesia resemble those seen inunanesthetized humans (Ho and Waite 2002) and do not alter the timecourse of presynaptic inhibition, which may occur with other anes-thetics (e.g., pentobarbital sodium) (Tang and Schroeder 1973), weanesthetized animals with an induction dose of ketamine (80 mg/kgip) and xylazine (5 mg/kg ip) and maintained on ketamine alone (20mg/kg ip) (Ho and Waite 2002; Hosoido et al. 2009). Core bodytemperature was monitored with a rectal thermometer and maintainedat 38 � 1°C with a circulating water heating pad placed under anabsorbent pad. To record electromyogram (EMG) data, which in-cluded the muscle response (M-wave) and the monosynaptic reflexresponse (H-reflex) above and below the injury site in SCI animals,we used an established percutaneous needle preparation (Boulenguezand Vinay 2009; Lee et al. 2009; Schieppati 1987; Thompson et al.1992a; Valero-Cabre et al. 2004). We chose to use this minimallyinvasive procedure because it is analogous to methods used to evokeand record H-reflex in humans (Palmieri et al. 2004; Schieppati 1987)and provides the opportunity to stimulate and record at all four limbswithin the same animal without perturbing muscle or nerve tissue thatwould otherwise be disrupted from more invasive surgical electrodeplacement (e.g., cuff electrode implants). In addition, this recordingapproach maintains the integrity of the vascular system and optimizesthe tissue preservation and collection methods for histological study(see above) that we performed following electrophysiological exper-iments. For stimulation, a pair of Teflon-insulated stainless steelfine-wire electrodes (0.002-in. bare metal diameter; A-M Systems,Carlsborg, WA) were threaded into a 32-gauge syringe needle. The

Table 1. Spinal cord motor neuron morphometry

Maximum CellDiameter, �m Aspect Ratio Form Factor

No. of PrimaryDendrites

Total DendriteLength, �m

Sham 54.6 � 19.4 0.57 � 0.17 0.49 � 0.17 8.16 � 3.69 1,073.0 � 671.4SCI (above injury) 46.7 � 11.7 0.51 � 0.16 0.50 � 0.16 5.73 � 1.95 815.10 � 451.9SCI (below injury) 51.4 � 17.4 0.64 � 0.13 0.62 � 0.09 4.80 � 1.23 739.30 � 280.6SCI � anti-Rac 46.1 � 8.7 0.63 � 0.11 0.60 � 0.10 4.38 � 1.15 1,085.8 � 307.9

Values are means � SD. SCI, spinal cord injury.



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wire ends were carefully bent into sharp barbs, the insulation wasremoved with heat (to expose tips �1 mm), and the needle and wirewere then transcutaneously inserted until the wire tip was in closeproximity to the mixed nerves of the deep radial nerve or tibial nerve,above or below the injury site in SCI animals, respectively. Theneedle was retracted and the wire remained in place. The secondelectrode was inserted similarly, spaced �2 mm apart from the firstelectrode. Stimulating electrode placement was adjusted until theintensity of square-wave stimulating pulses (0.2-ms duration contin-uously given at a rate of 1 every 3 s) required to induce subtle visiblemotor twitch responses (i.e., wrist extension/radial abduction or plan-tar flexion) was �1 mA (Lee et al. 2009; Valero-Cabre et al. 2004).For recording electrodes, we used insulated wire electrodes made ofsimilar materials and exposed �2 mm of the wire tips using heat. Torecord EMG data from the forelimb (e.g., brachioradialis reflex), anelectrode was inserted into the interosseous muscles between thefourth and fifth digit and a reference electrode was placed subcuta-neously in the dorsolateral surface of the paw. To record EMG datafrom the hindlimb (e.g., plantar reflex), an electrode was inserted intothe plantar muscles within the palmar/ventral surface of the hindpawand proximal to the ankle region. A reference electrode was placedsubcutaneously within the dorsolateral surface of the hindpaw. Thesereflexes were chosen on the basis of our pilot experiments andprevious work that demonstrated EMG reflex response evoked inthese muscles could be reproducibly recorded after SCI (Boulenguezet al. 2010; Kim et al. 2009; Valero-Cabre et al. 2004). Note thatSCI-induced changes in the plantar reflex, primarily innervated bymotor pools in L5, less from L4 (Crockett et al. 1987), have beenshown to be similar to changes in reflexes elicited in other hindlimbmuscles, i.e., tibialis anterior and gastrocnemius, which are alsoinnervated by L4 and L5 (Lee et al. 2009; Valero-Cabre et al. 2004).EMG responses were filtered (10–1,000 Hz), amplified, and recordedfor offline analysis using Spike2 (version 7.08; CED Software, Cam-bridge, UK). To identify optimal stimulation intensity for activatingstable M-wave and H-reflex responses, square-wave pulses (0.2-msduration) were applied at a rate of one every 3 s. The intensity ofelectrical stimulation was first adjusted to determine the minimumintensity to evoke an M-wave response �50% of the time andprogressively increased until a stable M-wave and maximal H-reflexresponse could be observed.

To measure RDD of the H-reflex response, we performed a paired-pulse stimulation paradigm with a conditioning and test pulse that weapplied at a range of interpulse intervals (10 to 2,000 ms). Three trials(10 sweeps/trial) with at least 30 s between trials were recorded foreach interpulse interval. The M and H response wave amplitudes werequantified from averaged and rectified waveforms within each animal(Boulenguez et al. 2010; Tan et al. 2012a). For comparison acrosstreatment groups, the maximum waveform amplitudes of the H and Mresponses to the test pulse were converted into a percentage of themaximum amplitude response to the conditioning pulse. M and Hwaveform amplitudes were measured from baseline to peak ampli-tude. To determine trial-to-trial consistency, the coefficients of vari-ation (CoV) from the M and H waves were calculated by dividing thestandard deviation by the mean maximum amplitude. After recordingexperiments, animals were killed for Golgi staining or immunohisto-logical study as described above.

A number of studies have shown that dendritic spine morphologycan change within minutes following cortical injury or activity (Ma-jewska and Sur 2003; Mizrahi et al. 2004; Zhang et al. 2005).Moreover, others have shown that exogenous electrical stimulationcan induce plasticity of the H-reflex, which can persistent for manyhours or days (Chen et al. 2003, 2006). Although in this study ourH-reflex testing was performed acutely (i.e., lasting �1 h per animal),we ensured that all animals, including both spinal cord-injured anduninjured sham animals, underwent similar H-reflex testing protocolsto control for potential confounds of direct nerve stimulation.

Statistical analysis. All statistical tests were performed at the �level of significance of 0.05 with two-tailed analyses using parametricor nonparametric tests, as appropriate. For comparisons of anatomicaland functional changes above and below the injury site after SCI, wecompared multiple comparisons of data collected within the sameanimals and with sham animals. To determine the appropriate statis-tical model to apply to these specific datasets, we incorporated twoassumptions: 1) that the above- and below-injury data sets weredependent variables affected by the application of the independenttreatment variable, SCI, and 2) as a consequence of SCI, a putativesecondary pathway interaction arises between above- and below-injury spinal segments that can affect the above- or below-injury datasets (e.g., an emergent interaction following SCI that leads to differ-ential effects on spinal cord tissue located above or below the SCIinjury site). Given these assumptions, our data sets fit most closelywith a one-way (or one factor) ANOVA statistical model, where SCIis the treatment factor applied to two dependent variables, above andbelow injury. Moreover, to control for multiple comparison errorswith the additional comparison against the sham group, we applied apost hoc repeated-measures correction. Although we considered theapplication of a split-plot analytical model, this approach falselyassumes that the secondary, post-SCI interaction is a factor that isindependent of the first-cause treatment factor, SCI. In summary, forcomparing data sets gathered from above and below the injury site, weapplied repeated-measures corrections (Dunn or Bonferroni post hoctests) following ANOVA and Kruskal-Wallis one-way ANOVA onranks analyses. As a note, previous reports have similarly appliedANOVA repeated-measures design to compare ipsilateral and con-tralateral sides of the spinal cord following unilateral nerve trauma orto compare regions above and below the injury site after SCI. Thestatistical design of these studies encompassed the assumed effect ofa postinjury secondary, bidirectional interaction (Anderson et al.1998; Chang et al. 2010; Tal and Bennett 1994; Tan et al. 2007,2012a), which also may have emerged within our experimental SCImodel. Data management and statistical analyses were performedusing SigmaPlot (version 12.5; Systat Software) and Microsoft OfficeExcel (2011). Data in the text are means � SD; data in graphs areplotted as means � SE using SigmaPlot.

RESULTS

Contusion SCI disrupts the dorsal corticospinal tract. In SCIanimals, contusion injury at spinal cord segment L2 resulted insevere damage of the dorsal columns and gray matter (Fig. 2A),in contrast with sham animals (Fig. 2B). Histological exami-nation of caudal spinal cord tissue (segments below L3)showed no visible tissue damage or glial scar tissue (notshown). The IH impactor device provided biomechanical mea-surements during each SCI procedure (Fig. 2, C and D). Thecord surface displacements upon rod impact were not signifi-cantly different across both SCI groups, SCI � vehicle and SCI �anti-Rac (Fig. 2C; 1,419.8 � 293.3 vs. 1,394.9 � 266.2 �m,P � 0.05, t-test). Similarly, there was no difference in the actualapplied force between the respective SCI groups (Fig. 2D;172.4 � 12.4 vs. 181.4 � 19.7 kdyn, P � 0.05, ANOVA onranks with Dunn’s post hoc test). Applied impact force with theIH device predicts the amount of tissue sparing, which corre-lates closely with locomotor functional outcome (Scheff et al.2003). PKC-� immunoreactivity served as an anatomic markerto help confirm the injury magnitude of our SCI model (Brad-bury et al. 2002; Sasaki et al. 2009; Tan et al. 2012a). Incoronal spinal cord sections above the injury in the cervicalenlargement, PKC-� immunoreactivity symmetrically labeledthe dCST and small-diameter cells located in laminae I/II



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(Fig. 2E) (Mori et al. 1990). Six weeks after SCI, below theinjury level in the lumbar enlargement, PKC-� staining ofthe dCST was bilaterally eliminated from the dorsal col-umns (Fig. 2F). PKC-� staining profiles of the dCST andsuperficial laminae remained intact (data not shown) incervical and lumbar enlargement tissues in sham animals.

Dendritic spine density changes on motor neurons after SCI.Dendritic spines remodel in the motor cortex after SCI (Kim etal. 2006, 2008); however, it is not known whether SCI-induceddendritic spine dysgenesis occurs on �-motor neurons withinthe spinal cord. Injury-induced changes in dendritic spinemorphology on nociceptive neurons in the dorsal horn havebeen shown to contribute to increased excitability associatedwith neuropathic pain (Tan et al. 2008, 2009, 2012b). Todetermine whether dendritic spine remodeling occurs on spinalcord �-motor neurons, we identified �-motor neurons (seeMATERIALS AND METHODS) and performed a morphological com-parison of �-motor neurons across treatment groups (Fig. 3).We identified �-motor neurons located in lamina IX and withinmotor pools in the lateral regions of the ventral horn (Fig. 3, Aand B). Six weeks after SCI, motor neurons had widelyprojecting dendritic trees containing numerous spines. Quali-tative observations demonstrated marked differences in spinenumber across treatment arms (Fig. 3, C–F). To ensure equiv-alent sampling across groups, we assessed several morpholog-ical criteria and compared these values across treatment groups(Table 1). There were no statistically significant differences inmaximum cell diameter, aspect ratio, form factor, number ofprimary dendrites, or total dendrite branch lengths (for allcomparisons: P � 0.05). We therefore interpreted any differ-ences in dendritic spine profiles across groups as not due tovariations in neuronal sampling, but rather an effect of exper-imental treatments. As a note, the values for maximum celldiameter, form factor, and dendritic branch lengths were sim-ilar to measurements for �-motor neurons that were labeled by

intramuscularly injected retrograde tracers (Bose et al. 2005;Crockett et al. 1987; Hashizume et al. 1988; Jacob 1998).

To obtain an accurate measure of dendritic spine profilesfrom ventral spinal cord tissue, we digitally reconstructed�-motor neurons using Neurolucida software (Fig. 4). Wemarked the location of sample neurons on a contour map of thespinal cord gray matter (Tan et al. 2008). �-Motor neuronsfrom each treatment group (Fig. 4, A–D, red dots; n � 20–21cells/group) were located in the ventrolateral regions of thegray matter, shown as a single representative contour fromsegmental level C5 (above injury) or L5 (below injury). Den-dritic spines on traced motor neurons were marked alongdendritic branches and color-coded with thin-shaped (blue) ormushroom-shaped (red) spines (Fig. 4, A=–D=).

A main objective of this study was to assess the contributionof SCI-induced changes in dendritic spines to reflex dysfunc-tion; we therefore measured three morphological profiles ofspines that have been associated with injury-induced neuronalhyperexcitability: 1) increased density of dendritic spines,particularly mature mushroom-shaped spines, 2) redistributionof spines toward dendritic branch locations close to the cellbody, and 3) enlargement of the spine head diameter. Becausespasticity often presents below the injury site following SCIand less commonly above (Skold et al. 1999), we measureddendritic spines on �-motor neurons in motor pools of thecervical (C4–C5; above injury) and lumbar (L4–L5; belowinjury) spinal segments that innervate forelimb and hindlimbmusculature, respectively (see METHODS AND MATERIALS).

As shown in Fig. 5A, 6 wk after SCI, total dendritic spinedensity on motor neurons below the injury site increasedcompared with neurons from sham control and neurons abovethe injury site (P � 0.05; 2.80 � 0.78 vs. 1.82 � 0.52 vs. 1.10 �0.54 spines/10-�m dendrite, respectively; ANOVA on rankswith Dunn’s post hoc test). In contrast, motor neurons abovethe injury site in the cervical enlargement had dendritic

Fig. 2. Spinal cord injury. A: contusion injury at L2 resulted in severe damage of the dorsal columns and gray matter, as shown by glial fibrillary acidic protein(GFAP) staining in coronal spinal cord tissue sections. Asterisk denotes lesion epicenter. B: intact spinal cord tissue from sham animal. C and D: biomechanicaldata provided by the Infinite Horizon (IH) impactor demonstrated no difference between vehicle (SCI � Veh)- and Rac1 inhibitor NSC23766-treated (SCI �anti-Rac) SCI groups. E: 6 wk after SCI (above injury), PKC-� staining produced bilateral labeling of the dorsal corticospinal tract (dCST) and lamina I/II. F:at the lumbar level L5 (below injury), the absence of PKC-� immunoreactivity in the dorsal column white matter tracts demonstrates significant disruption ofthe dCST. SCI did not affect PKC-� staining in superficial laminae. Scale bars, 500 �m.



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spine densities that decreased compared with neurons fromsham (P � 0.05). A similar profile of dendritic spine densitywas also observed for thin-shaped dendritic spines (Fig. 5B):below the injury site there was a significant increase in thinspines compared with above the injury and sham (P � 0.05;2.42 � 0.63 vs. 1.03 � 0.54 vs. 1.61 � 0.40 spines/10-�mdendrite, respectively; ANOVA on ranks with Dunn’s posthoc test). Importantly, there was a significant increase in thedensity of mature, mushroom-shaped spines located on�-motor neurons below the injury site compared with neu-rons from sham and neurons above the injury (P � 0.05;0.45 � 0.33 vs. 0.20 � 0.26 vs. 0.06 � 0.06 spines/10-�mdendrite, respectively; ANOVA on ranks with Dunn’s posthoc test) (Fig. 5C). Note that the mushroom-shaped spine densityobserved below the injury site after SCI represents a more than200–700% increase compared with mature-shaped spine densitiesabove the injury site and sham control.

Dendritic spines redistribute toward proximal branches onmotor neurons after SCI. Excitatory afferent inputs locatedcloser to the neuronal cell body can have a greater weightedimpact on the overall electrical output of a neuron because ofthe closer proximity to the axon hillock (Pongracz 1985; Tan etal. 2009; Yuste and Urban 2004). To profile changes indendritic spine distribution along motor neuron branch pro-cesses, we applied a Sholl’s analysis and pooled spine densitieswithin proximal regions close to the cell body (50–150 �m)and distal regions (200–350 �m) (see METHODS AND MATERIALS)(Fig. 5, D–F).

On motor neurons below the injury site in SCI animals, total,thin-shaped, and mushroom-shaped dendritic spines increasedon proximal dendrite branches compared with equivalent re-

gions in sham and SCI neurons above the injury site (P � 0.05;for total spines: 3.1 � 1.1 vs. 2.1 � 1.1 vs. 1.2 � 0.69, for thinspines: 2.7 � 0.16 vs. 1.9 � 0.05 vs. 1.1 � 0.05, and formushroom spines: 0.39 � 0.04 vs. 0.17 � 0.06 vs. 0.08 � 0.01spines/10-�m dendrite, respectively; 1-way ANOVA withBonferroni’s post hoc test) (Fig. 5, D–F). At distal regions, SCIdid not change spine density of any category on motor neuronsfrom below the injury compared with sham (P � 0.05). On theother hand, motor neurons below the injury had increased spinedensity for all categories compared with above the injury atdistal regions (P � 0.05; for total spines: 3.2 � 1.4 vs. 1.4 �0.9, for thin spines: 2.3 � 1.4 vs. 1.4 � 0.9, and for mushroomspines: 0.85 � 0.9 vs. 0.08 � 0.16 spines/10-�m dendrite,respectively; ANOVA on ranks with Dunn’s post hoc test).There were no differences in any spine densities at distalregions on neurons from sham animals compared with neuronsabove the injury in SCI animals (P � 0.05).

Dendritic spine dimensions change on motor neurons afterSCI. To quantify the effects of SCI on spine length and spinehead diameter, we analyzed 880–1,305 dendritic spines thatwere sampled from 4–7 motor neurons per group (see MATE-RIALS AND METHODS; sham, n � 3 animals/5 neurons; above SCIlevel, n � 3 animals/7 neurons; below SCI level, n � 3animals/4 neurons). As shown in Fig. 5G, dendritic spinesbelow the injury site in SCI animals decreased in lengthcompared with those above the injury site and in sham animals(P � 0.05; 1.38 � 0.79 vs. 1.67 � 0.97 vs. 1.60 � 0.81 �m,respectively; 1-way ANOVA with Bonferroni’s post hoc test).In contrast, spine head diameter increased after SCI below theinjury site compared with that above the injury and in shamanimals (P � 0.05; 1.22 � 0.65 vs. 0.97 � 0.57 vs. 0.95 �

Fig. 3. Golgi staining of spinal cord tissuereveals dendritic spines on motor neurons inthe ventral gray matter. A: image of ventralgray matter with an identified �-motor neu-ron located in Rexed lamina IX (arrow andwhite box). B: high-power field of motorneuron shown in inset in A. Six weeks aftersham and SCI procedures, representative im-ages of dendritic branches show apparentdifferences in dendritic spine profiles fromsham (C), SCI � Veh above the injury (D),SCI � Veh below the injury (E), and SCI �anti-Rac (F) treatment groups. C=–F=: highmagnification of selected dendrite regionsfrom C–F (red boxes). Scale bars: A, 500�m; B, 100 �m; C–F, 10 �m; C=–F=, 2 �m.



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0.56 �m, respectively; 1-way ANOVA with Bonferroni’s posthoc test) (Fig. 5H). Motor neurons above the injury site afterSCI did not differ compared with sham in any dimensionmeasured (P � 0.05).

H-reflex response increases below the injury site after SCI.Dendritic spine morphology significantly influences synapticfunction (i.e., in a structure-function relationship) (Pongracz1985; Segev and Rall 1988, 1998). As we and others haveshown (Leuner and Shors 2004; Majewska et al. 2000; Tan etal. 2009; Zhou et al. 2004), increased dendritic spine density,mature dendritic spine morphologies (e.g., mushroom shapes),and proximal redistribution of spine synapses can amplifyneuronal excitability, enhance frequency-following ability, re-duce noise-filtering capabilities, and attenuate inhibitory input.To determine the effect of SCI on reflex function in associationwith dendritic spine remodeling on �-motor neurons, we mea-sured the H-reflex response in uninjured sham animals (i.e., inthe hindlimb) and in SCI animals above and below the injuryin the forelimb and hindlimb, respectively.

We also measured the M-wave, which reveals the elec-trical responsiveness of motor axons, its ability to conductan action potential, and the electrochemical coupling of theefferent and muscle tissue (e.g., neuromuscular junction)(Hultborn and Nielsen 1995). In normal animals, the H-re-flex undergoes activity RDD. A reduction in H-reflex RDDis a physiological indicator of spasticity (Boulenguez et al.2010; Ho and Waite 2002; Lee et al. 2009; Taylor et al.1984). To determine H-reflex and M-wave response in SCIanimals, we electrically stimulated the deep radial nerve ortibial nerves and recorded reflex response from muscle inthe forelimb (extensor carpi radialis) and hindlimb (plantar

muscle). As a comparison, we measured evoked H- andM-responses from hindlimb muscle in uninjured sham ani-mals. We chose these reflexes on the basis of preliminarystudies and previously published work demonstrating thatevoked reflex responses for these muscles could be repro-ducibly produced in adult rats after SCI (Boulenguez et al.2010; Kim et al. 2009; Valero-Cabre et al. 2004). Impor-tantly, previous studies have shown that changes in plantarreflex after SCI are similar to changes in reflexes elicited forother hindlimb muscles, i.e., tibialis anterior and gastrocne-mius, which are also innervated by motor pools in L4 –L5(Lee et al. 2009; Valero-Cabre et al. 2004).

We used a paired-pulse stimulation paradigm: a control andtest pulse, separated by a range of interpulse intervals from2,000 to 10 ms (Fig. 6). A representative trace in shamproduced from recordings of plantar muscle shows two evokedEMG waves: the M-response and the H-reflex (central looppathway) (Fig. 6A). As the interval between the control and testpulse shortened from 2,000 to 10 ms, there was a markeddepression of the H-reflex. The M-wave amplitude also de-creased with shortening interpulse intervals, demonstratingRDD of motor neuron-to-muscle response. Figure 6, B and C,shows a qualitative example of SCI-induced reductions in H-and M-wave RDD from muscle recordings above and belowthe injury at 100- and 10-ms interpulse intervals. In SCIanimals, the M-wave appeared to maintain stable amplitudeeven at shorter interpulse intervals.

We quantified the percentage change in H-reflex for unin-jured sham (n � 7) and SCI animals (n � 6) over the range ofinterpulse intervals (Fig. 6D). In sham animals, the H-reflexmaintained stable amplitude between 2,000 and 300 ms and

Fig. 4. Digital reconstructions of spinal cordmotor neurons. To obtain an accurate profileof dendritic spines in motor neurons, wedigitally reconstructed the entire branchstructure of sampled neurons. A–D: contourtraces from each group as indicated show thelocations of all sampled motor neurons (reddots) within the gray matter (representativeblack trace). Density and distribution weremeasured from 3-dimensional neuron recon-structions from sham (A=), SCI � Veh abovethe injury (B=), SCI � Veh below the injury(C=), and SCI � anti-Rac (D=) treatmentgroups. A–D: �50-�m lengths of den-drites from neurons shown in A=–D= (gray-shaded regions) show thin-shaped (bluedots) and mushroom-shaped spines (reddots). Scale bars: A–D, 500 �m; A=–D=, 50�m; A–D, 10 �m.



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with a steady decline at shorter interpulse intervals, similar tothat observed in previous studies (Ho and Waite 2002; Hosoidoet al. 2009; Tan et al. 2012a). The M-wave amplitude in shamanimals remained stable through a wider range of interpulseintervals, 2,000 and 100 ms (%M-wave amplitude, 2,000 vs. 50ms: P � 0.05; 1-way ANOVA with Bonferroni’s post hoc test)(Fig. 6E). Therefore, the H-reflex depression response in shamanimals is not due to the inability of muscle to respond torepeated stimulus activity. At shorter interpulse intervals (i.e.,3–5 ms), both the H-reflex and M-wave responses depressed ata much greater rate and, in most stimulus-recording trials,failed to appear in sufficient number for analysis (data notshown).

Six weeks after SCI, H-reflex measurements from evokedhindlimb reflex revealed a significant reduction in RDD (i.e.,H-reflex amplitude stabilized) through the entire range ofinterpulse intervals tested (Fig. 6D). Between interpulse inter-vals from 50 to 150 ms, there was a significant increase inH-reflex response in SCI below the injury site compared withsham (P � 0.05; 1-way ANOVA with Bonferroni’s post hoctest). Notably, SCI appeared to amplify the reflex responsebelow the injury site at the shortest interpulse interval at 10 mscompared with sham (P � 0.05; 1-way ANOVA with Bonfer-roni’s post hoc test), with amplitude responses greater than100% of control amplitude. In contrast, within the same SCIanimal, there continued to be significant RDD in recordings

Fig. 5. Quantitative analysis of dendritic spine profiles between sham and SCI animals above or below the injury. Analysis of dendritic spine profiles revealsdifferences in dendritic spine density (A–C), distribution (D–F), and shape (G and H). Total dendritic spine density (A), which includes all spine shapes, thinspine density (B) and mushroom spine density (C) decreased on motor neurons located above the injury after SCI compared with neurons from sham (*P � 0.05).In contrast, total spine density increased on motor neurons located below the injury compared with neurons from either sham or above the injury after SCI(*P � 0.05). Dendritic spine distribution for total (D), thin (E), and mushroom spines (F) differed across the comparator groups. At proximal regions in SCIanimals, all spine densities increased below the injury compared with neurons from sham and above the injury (*P � 0.05). In contrast, neurons above the injuryhad lower total and thin spine density at proximal regions compared with neurons from sham and below the injury (*P � 0.05). Although mushroom spine densityon motor neurons above the injury did not differ from that on neurons from sham at proximal regions (F), these neurons had significantly lower mushroom spinedensity than below the injury. At distal regions, motor neurons below the injury had greater spine density in all categories compared with motor neurons abovethe injury (*P � 0.05). There was no difference in any spine densities at distal regions on neurons from sham and above the injury in SCI animals. Dendriticspine shape analysis revealed no change in spine length (G) or spine head diameter (H) on motor neurons located above the injury in SCI animals compared withneurons from sham (*P � 0.05). Below the injury, these measurements demonstrated a decrease in spine length and an increase in spine head diameter comparedwith neurons from sham and above the injury (*P � 0.05).



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above the injury at 500, 50, and 10 ms (P � 0.05; 1-wayANOVA with Bonferroni’s post hoc test) (Fig. 6D). H-reflexRDD above the injury was not significantly different from thatin sham across all interpulse intervals (P � 0.05). Thesefindings indicate that SCI-induced increases in H-reflex re-sponse only occurred below the level of the injury and frommuscle innervated primarily by motor pools in spinal segmentL4–L5. Over the range of interpulse intervals tested from2,000 to 100 ms, the %M-wave amplitude remained close to100% across all comparator groups (Fig. 6E). However, therewas a significant increase in %M-wave amplitude above 100%of control in SCI above or below injury compared with sham,which depressed at 50 and 10 ms (P � 0.05; ANOVA on rankswith Dunn’s post hoc test). The ratio of H-reflex to M-waveresponses (H/M ratio) calculated from reflex data in SCI belowthe injury was larger than that in sham or in SCI above injuryat all interpulse intervals (Fig. 6F). SCI below the injury siteresulted in a more stabilized rate of decay for H/M ratio values,an indication of hyperreflexia and spasticity (Little and Halar1985; Matthews, 1966; Nielsen et al. 2007).

To assess changes to H-reflex response fidelity, we calcu-lated the CoV (SD/mean %H-reflex amplitude) for 50 and 10ms for sham and SCI animals. At the 50-ms interpulse interval,the CoV after SCI below the injury was nearly 30–50% smaller

than for SCI above the injury or for sham (SCI below injury,0.20; SCI above injury, 0.58; sham, 0.85). Similarly, the CoVfor SCI below the injury site was almost 40–60% smaller thanfor sham (SCI below injury, 0.37; SCI above injury, 0.66;sham, 0.92). Taken together, these values show that in additionto increasing H-reflex amplitude, SCI also increases the reli-ability of reflex activation below the injury site.

Inhibition of Rac1 disrupts dendritic spine remodeling. Wereasoned that if abnormal dendritic spine profiles after SCIcontributes to increased reflex excitability, then disruption ofdendritic spine remodeling would reduce signs of spasticity. Todetermine whether disruption of dendritic spine remodeling on�-motor neurons in L4–L5 after SCI reduces spasticity, weassessed the effects of administering NSC23766, a specificRac1-GTPase inhibitor. Treatment with NSC23766 resulted ina decrease in total, thin, and mushroom-shaped spine densitycompared with SCI � vehicle (P � 0.05; SCI � anti-Rac vs.SCI � vehicle: 0.95 � 0.24 vs. 2.8 � 0.78 total spines/10-�mdendrite, 0.92 � 0.23 vs. 2.4 � 0.63 thin spines/10-�mdendrite, and 0.03 � 0.03 vs. 0.44 � 0.33 mushroom spines/10-�m dendrite; ANOVA on ranks with Dunn’s post hoc test)(Fig. 7, A–C). We also determined the effect of NSC23766treatment on dendritic spine distribution by measuring spinedensity in proximal and distal locations along dendrites of

Fig. 6. Rate-dependent depression (RDD) of the H-reflexand M-wave responses above and below SCI. As a phys-iological assessment of the monosynaptic H-reflex, weperformed a paired-pulse stimulation protocol. Represen-tative traces (averaged 10–20 traces) of the M and Hresponses to control (first) and test (second) pulse in sham(A), SCI above the injury (B), and SCI below the injury(C). The control and test pulses were separated with arange of interpulse latencies between 2,000 and 10 ms.Note that in sham animals, as the interpulse intervalsdecreased (e.g., increasing the rate of activity) between thetest and control pulse, the amplitude of the M and Hresponses decreased. As shown in C, in SCI below theinjury, RDD in amplitude of either the M or H responsefailed to appear. D and E: %H-reflex and %M-waveamplitudes are normalized values of the evoked stimulusresponse of the test and control pulse. D: after SCI, therewas no significant difference in %H-reflex in SCI abovethe injury compared with sham at any interpulse interval.In contrast, in SCI animals below the injury, the %H-reflex significantly increased compared with sham at theshortest interpulse intervals between 100 and 10 ms (*P �0.05), demonstrating a loss of RDD and increased excit-ability of the H-reflex. Similarly, %H-reflex below theinjury was significantly greater than above the injury inSCI animals at 500-, 50-, and 10-ms interpulse intervals(§P � 0.05). E: % M-wave demonstrated a significantlyincreased response below the injury compared with theresponse in sham animals (*P � 0.05) and above theinjury in SCI animals (#P � 0.05). F: the H/M ratio wascalculated from M-wave and H-wave responses.



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�-motor neurons after SCI (Fig. 7, D–F). NSC23766 treatmentsignificantly decreased spine density in proximal and distalregions and for all spine categories, including total, thin-shaped, and mushroom-shaped dendritic spines (P � 0.05; forproximal total spines: 3.1 � 0.2 vs. 0.9 � 0.4, for proximal thinspines: 2.7 � 0.16 vs. 0.9 � 0.4, for proximal mushroomspines: 0.4 � 0.4 vs. 0.02 � 0.02, for distal total spines: 3.2 �0.5 vs. 1.2 � 0.5, for distal thin spines: 2.4 � 0.5 vs. 1.1 � 0.5,and for distal mushroom spines: 0.8 � 0.9 vs. 0.04 � 0.07spines/10-�m dendrite; 1-way ANOVA with Bonferroni’s posthoc test).

Dendritic spines on �-motor neurons in SCI animals thatwere treated with NSC23766 decreased in spine length andhead diameter compared with neurons in SCI � vehicle (P �0.05; length, 0.7 � 0.42 vs. 1.38 � 0.79 �m; head diameter,0.93 � 0.51 vs. 1.23 � 0.66 �m; ANOVA on ranks withDunn’s post hoc test) (Fig. 7, G and H). Together, thesefindings show that Rac1-inhibitor NSC23766 treatment caneffectively disrupt SCI-induced dendritic spine remodeling on�-motor neurons.

Inhibition of dendritic spine remodeling reduces H-reflexexcitability after SCI. Previous work has demonstrated thatintrathecal infusion of NSC23766 is efficacious in restoringclose-to-normal dendritic spine profiles on nociceptive neuronswithin the dorsal horn after SCI and peripheral nerve injury(Tan et al. 2008, 2011). In these studies, treatment withNSC23766 also reduced neuronal hyperexcitability associatedwith central sensitization, demonstrating that Rac1-regulateddendritic spine remodeling can contribute to mechanisms un-derlying neuropathic pain (Tan and Waxman 2012). To deter-mine whether disruption of Rac1-regulated dendritic spineprofiles on �-motor neurons attenuates exaggerated H-reflexresponsiveness after SCI, we used a paired-pulse stimulationparadigm as described above (Hultborn and Nielsen 1995; Tanet al. 2012a) (see Fig. 6). Representative EMG traces in SCIanimals below the injury in the hindlimb demonstrates thatstimulation produced both M-wave and H-reflex responses(Fig. 8). At the shortest interpulse interval of 10 ms, there wasa notable reduction of the RDD. At the 10-ms interpulseinterval, treatment with the Rac1 inhibitor NSC23766 appeared

Fig. 7. Rac1 inhibitory treatment disrupts dendritic spine morphology on motor neurons in the ventral horn after SCI. Treatment with NSC23766 in SCI animals(SCI � anti-Rac) resulted in a significant decrease in total (A), thin (B), and mushroom spine density (C) compared with SCI � Veh (*P � 0.05). D–F:assessment of dendritic spine distribution on motor neurons showed that NSC23766 treatment in SCI animals resulted in decreased spine density for all spinecategories at both proximal and distal branch regions (*P � 0.05). NSC23766 treatment decreased SCI-induced spine length (G) and spine head diameter (H)compared with SCI � Veh (*P � 0.05).



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to restore RDD of the H-reflex in hindlimb EMG recordings(e.g., reduced H-reflex amplitude in response to the test pulse)(Fig. 8B).

Figure 8C shows the quantified changes in the H-reflexresponse in SCI � vehicle (n � 6) and SCI � anti-Rac1 (n �5). Six weeks after SCI, in animals treated with control vehicle,the hindlimb H-reflex response demonstrated almost no RDD,with %H-reflex response remaining stable (e.g., close to 100%)across the range of interpulse intervals from 2,000 to 10 ms(for a comparison with sham, see Fig. 6). Treatment of SCIanimals with the Rac1 inhibitor resulted in a restoration ofRDD at the three shortest interpulse intervals of 100, 50, and10 ms compared with SCI � vehicle (P � 0.05; 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 10 ms, 13.3 � 21.4% vs. 88.9 � 32.9%;1-way ANOVA with Bonferroni’s post hoc test) (Fig. 8C). Incomparisons across SCI animal groups, the %M-wave ampli-tude remained close to 100% between interpulse intervals of2,000 and 100 ms (group means: for SCI � vehicle, 98.6 �1.5%; for SCI � anti-Rac1, 107.3 � 11.4%) (Fig. 8E). The%M-wave response at shorter interpulse intervals, 50 and 10ms, exhibited greater variability compared with that at longerstimulus intervals, but the difference was not statisticallysignificant (P � 0.05).

We calculated the CoV of the %H-reflex at 50 and 10 msacross SCI animal groups treated with vehicle or the Rac1inhibitor. At the 50- and 10-ms interpulse intervals, treatmentwith the Rac1 inhibitor in SCI animals resulted in a CoV thatwas nearly fourfold greater than in vehicle-treated SCI animals(SCI � vehicle, 0.27 and 0.37; SCI � anti-Rac, 0.9 and 1.6).Thus, in addition to restoring RDD, Rac1 inhibition alsoincreased the variability of the reflex response. Figure 8Eshows the plot for the H/M ratio. Treatment with the Rac1inhibitor decreased the overall H/M ratio across all interpulseintervals tested between 2,000 and 10 ms, as shown by adownward shift in trend line slope.

VGluT1 bouton areal density in the gray matter does notincrease after injury. Synapse-associated protein markers (e.g.,synaptophysin and PSD-95) increase after SCI, demonstratingthe presence of injury-induced synaptic plasticity (Tan et al.2008; Tan and Waxman 2012). Because upper motor tractinjury and SCI can increase the excitability of spinal reflexpathways below the injury (Baastrup et al. 2010; Little andHalar 1985; Tan et al. 2012a), we next determined if excitatoryinputs, particularly those of Ia afferents, change after SCI.VGluT1 is a widely used marker for excitatory Ia afferentterminations in the spinal cord (Alvarez et al. 2004, 2011;Kitzman 2007).

Fig. 8. Disruption of Rac1-regulated dendritic spinesreduces SCI-induced H-reflex hyperexcitability. Rep-resentative traces show the M and H responses frompaired-pulse testing in SCI � Veh below the injury(A) and SCI � anti-Rac (B) treatment groups. C:quantification of the %H-reflex response demonstratesthat the H-reflex response in SCI � Veh animalsexhibited reduced RDD (also see Fig. 6). Rac1 inhib-itor treatment in SCI animals reduced the %H-reflex at100, 50, and 10 ms compared with SCI � Veh (*P �0.05). D: there was no significant difference in the%M-wave between SCI � Veh and SCI � anti-Rac.E: treatment with Rac1 inhibitor in SCI animals de-creased the H/M ratio compared with SCI � Veh, asdemonstrated by a steeper downward trend line.



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As shown in representative images in Fig. 9, A–D, immu-nopositive VGluT1 puncta were distributed throughout thespinal cord gray matter of the lumbar enlargement, L4–L5, foreach analyzed treatment group (Fig. 9, A–D, left). To visualizethe distribution of VGluT1 puncta in the gray matter, wecompiled the VGluT1 staining profiles from multiple tissuesections from each treatment group and produced spatial heatmaps (Fig. 9, A–C, right). Although the highest concentrationof VGluT1-positive boutons appeared to correspond withRexed laminae V/VI (Hantman and Jessell 2010; LaMotte etal. 1991), VGluT1 puncta were distributed throughout alllaminae. The areal densities of VGluT1 boutons were calcu-lated in the total gray matter (Fig. 9D) and within three regions:the dorsal horn, intermediate zone, and ventral horn (Fig. 9,

E–G; also see insets). Six weeks after SCI, we observed nosignificant difference in the areal density of VGLUT boutons inthe total gray matter compared with that in uninjured shamanimals (P � 0.05). Similarly, there was no statistical differ-ence following SCI � vehicle in the other three gray matterregions analyzed compared with uninjured sham (P � 0.05). Incontrast, treatment with NSC23766 significantly decreased theareal density of VGluT1 compared with SCI � vehicle in thetotal gray matter, intermediate zone, and ventral horn (for totalgray matter: 63.6 � 37.1 vs. 124.5 � 57.7, for intermediatezone: 72.4 � 25.2 vs. 138.7 � 51.3, and for ventral horn: 58.1 �48.4 vs. 129.8 � 55.9 puncta; ANOVA on ranks with Dunn’spost hoc test). There was no significant change in areal densityof VGluT1 in the superficial dorsal horn following Rac1

Fig. 9. Excitatory terminals in the spinal cord gray matter. Vesicular glutamate transporter 1 (VGluT1)-immunopositive puncta appeared throughout all laminaeof the spinal cord gray matter in the lumbar enlargement L4–L5 (A–C, left). Spatial heat maps (A–C, right; red � highest density, blue � lowest density) showsthe overall areal density of VGluT1 expression in sham (A), SCI � Veh (B), and SCI � anti-Rac (C) treatment groups. Quantification of the VGluT1 punctawithin the total gray matter region (D), dorsal horn (E), intermediate zone (F), and ventral horn (G) as represented in insets as gray shading demonstrates nosignificant change in SCI � Veh compared with sham. Treatment with the Rac1 inhibitor in SCI animals decreased VGluT1 areal density compared with SCI � Vehin the total gray matter, intermediate zone, and ventral horn only (*P � 0.05) with no significant change in the dorsal horn (P � 0.05). The areal density ofVGluT1 decreased in SCI � anti-Rac1 compared with sham in the dorsal horn (*P � 0.05). Scale bar for A–C, 500 �m.



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inhibitor treatment in SCI animals compared with SCI �vehicle (P � 0.05).

Rac1 inhibition does not affect locomotor behavior. To ruleout any differences in gross locomotor ability in SCI animals,we assessed postinjury locomotor behavior using the BBBlocomotor scale (Basso et al. 1995, 1996) (Fig. 10). Blindedobservers performed behavioral testing at three time points: onnaive animals before any surgical procedures, within 1 wk aftercatheter implantation and before drug treatment, and at the6-wk post-SCI endpoint (also see Fig. 1). All naive animalsexhibited a baseline locomotor score of 21 (1 worst to 21 best).Five weeks after SCI and catheter implantation, before anytreatments, animals exhibited a mean BBB score of 13.6 � 3.8,demonstrating the expected locomotor ability in the late-SCIrecovery phase (Basso et al. 1995). BBB testing of SCI animalsafter vehicle or drug delivery demonstrated no significanteffect on locomotor ability with scores remaining unchangedbetween SCI animals treated with (n � 10) or without Rac1inhibitor (n � 11) (P � 0.05, 14.2 � 1.6 vs. 13.8 � 2;ANOVA on ranks).

DISCUSSION

Spinal cord circuits can reorganize, changing in structureand function after injury (Raisman 1991). Our present findingsdemonstrate robust changes in dendritic spine morphology on�-motor neurons after SCI, including an increase in dendriticspine density, a distribution of spines closer to the cell body,and the presence of more mature dendritic spines. Thesepostsynaptic dendritic changes have been shown to accompanyincreased neuronal excitability after SCI (Rall et al. 1992;Segev and Rall 1998; Tan et al. 2009). In agreement, weobserved a significant loss of H-reflex RDD below the injury(i.e., increased H/M ratio), indicative of spasticity (Boulenguezet al. 2010; Matthews 1966; Nielsen et al. 2007). Importantly,dendritic spines above the injury exhibited a nearly oppositemorphological profile with decreased spine density and withdistribution and shape that were more similar to control pro-files. As expected, there was no change in H-reflex excitabilityabove the level of injury. Overall, these results demonstrate

that abnormal dendritic spine profiles below the level of injuryaccompany spasticity after SCI, and conversely, the lack ofsuch spine profiles above the injury correspond with a lack ofspinal reflex hyperexcitability.

To further elucidate the structure-function link betweendendritic spine dysgenesis and hyperreflexia, we disrupteddendritic spine remodeling by targeting Rac1 signaling in SCIanimals. We have previously shown that Rac1 inhibition dis-rupts dendritic spine remodeling in dorsal horn sensory neuronafter SCI, nerve injury, and diabetes mellitus (Tan et al. 2008,2011, 2012b). In the present study, we observed a decrease inspine density on �-motor neurons and a closer-to-normaldistribution of dendritic spines following treatment withNSC23766, a Rac1 inhibitor. NSC23766 treatment also de-creased spine length and head diameter, and partially restorednormal H-reflex activity (i.e., increased RDD). With these resultstaken together, our study is the first to demonstrate robust den-dritic spine reorganization on �-motor neurons in the ventral horn,which accompanies spasticity after SCI. We implicate Rac1signaling as an important mediator in both the structural andfunctional changes within the spinal reflex pathway afterinjury.

Spasticity after SCI has been attributed to a variety ofmechanisms within the spinal reflex arc (Nielsen et al. 2007;Roy and Edgerton 2012). Muscle spindle afferents may loseeither presynaptic inhibition or reciprocal inhibition. Alterna-tively, loss of Renshaw interneuron activity, thought to mediatereciprocal inhibition, can trigger spasticity (Nielsen et al.2007). Evidence obtained from intracellular recordings of spi-nal motoneurons also demonstrates increased motoneuron ex-citability, i.e., the ability to generate action potentials, includ-ing the appearance of plateau potentials and persistent inwardsodium and calcium currents in rat motor neurons after SCI(Bennett et al. 2001b; Heckmann et al. 2005; Li et al. 2004),potassium chloride cotransporter KCC2 downregulation (Bou-lenguez et al. 2010; Vinay and Jean-Xavier 2008), and sodiumchannel misexpression (Harvey et al. 2006; Li and Bennett2003). Inflammation (e.g., microgliosis) occurs in a number ofnervous system injury models, including SCI (Craner et al.2005; Hains and Waxman 2006), and may contribute to in-creasing excitability of neuronal populations within spinalcircuits (Gwak and Hulsebosch 2009; Zhao et al. 2007b).Astrocyte activation after injury can potentially maintain hy-perexcitability (Scholz and Woolf 2007). Finally, maladaptiveplasticity such as “collateral” or “reactive” sprouting maycontribute to altered spinal motor control (Boulenguez et al.2010; Krenz and Weaver 1998; Nielsen et al. 2007; Raisman,1994). Others have shown altered dendrite branch length onmotor neurons in the spinal cord that accompanies spasticityafter SCI (Kitzman 2005). Although dendritic spine morphol-ogies change on pyramidal neurons in the motor cortex afterSCI (Kim et al. 2006), the functional role for these spinealterations is not firmly understood. Computer simulationshave attempted to predict the physiological contribution ofdendritic spines on motor neurons (Rall et al. 1967); however,in vivo changes in dendritic spine structure have not beenreported in spinal cord motor pools.

Dendritic spine morphology partly determines synapticfunction and therefore provides a visual clue into how neuralnetworks function (Calabrese et al. 2006; Segev and Rall1998). Dendritic spines can reorganize rapidly following syn-

Fig. 10. Locomotor testing. Blinded observers performed BBB testing on SCIanimals at 3 time points: before any procedure (baseline), before treatment, andafter treatment (SCI � Veh and SCI � anti-Rac). All naive animals exhibiteda baseline locomotor score of 21. There were no significant differences in BBBscores across groups (P � 0.05).



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aptic activity (e.g., activity-dependent plasticity) and increasein density, which provides new or stronger synapses (Halpain2000). Abnormal dendritic spine morphologies have been re-ported in a wide spectrum of neuropsychiatric diseases, includ-ing posttraumatic stress disorder, substance dependence andaddiction, autism spectrum disorders, and mental retardation(Halpain et al. 2005; Purpura 1974). Although adaptive plas-ticity between Ia afferents and spinal motor neurons can shapeH-reflex response in both humans and rodents (Thompson et al.2009; Wolpaw 1994), maladaptive plasticity can contribute topathological H-reflex function associated with hyperreflexiaand spasticity (Lance 1980; Nielsen et al. 2007). In chronicSCI, hyperexcitability of the spinal stretch reflex (e.g., H-re-flex) is thought to underlie spasticity, which manifests as avelocity-dependent increase in tonic stretch reflexes, with un-controllable “jerking” movement and abnormal muscle tone,whereby muscle continually contract (Ashby et al. 1987; Lance1980; Nielsen et al. 2007; Skold and Woolf 1999). In ourstudy, we observed significant SCI-induced changes in den-dritic spine morphologies on �-motor neurons below the injurysite that accompanied a loss of RDD and a stabilization of theH/M ratio over a broad range of nerve stimulation rates.Importantly, we observed only minor changes in M-waveresponse after SCI, indicating that changes in RDD and H/Mratio were primarily due to mechanistic changes within thespinal cord monosynaptic circuit.

Although sacrocaudal injuries might better replicate someaspects of clinical spasticity (Li and Bennett 2003; Ritz et al.1992), these SCI models only allow studies of neurologicaldeficits in tail musculature, which are absent in human. Topermit sufficient locomotor ability for open-field behavioralassessment and H-reflex testing of hindlimb musculature, aparallel of leg muscle groups in human, we performed contu-sion SCI at spinal segment L2. As with all SCI animal studies,however, we encountered an observation suggesting that ourinjury model also cannot entirely reflect the human SCI con-dition. In contrast with human SCI at lower thoracic or upperlumbar segments, which generally produces some chronicnegative motor signs, including flaccidity or lower limb weak-ness (Doherty et al. 2002), we observed increased spinal motorreflex activity associated with increase muscle tone 6 wk afterinjury. Thus it is important to mention that our contusion SCImodel was utilized as a compromise to study spinal reflexfunction in hindlimb musculature.

We noted in our study that the RDD of the H-reflex did notexhibit depression at a similar rate compared with an earlierreport of the effect of spinal contusion on RDD (Thompson etal. 1992b). Whereas Thompson et al. observed activity-ratedepression of �85% of control at 5 Hz (Thompson et al.1992a), we observed a similar loss of H-reflex activity at10–20 Hz (i.e., 50- to 100-ms interpulse interval). A probableexplanation for this discrepancy is due to the additional pro-cedures that animals underwent before reflex testing in ourstudy, including surgical implantation of intrathecal cathetersand infusions of vehicle or drug solutions. Although no effectof catheter implantation and drug infusion has been observed inprevious nociceptive testing in control animals (Tan et al.2008), it is possible that these additional experimental proce-dures could have led to sensitization of afferents within thespinal reflex circuit. Nonetheless, the magnitude of activity-rate depression in sham and SCI animals in our study fell

within other documented ranges (Hosoido et al. 2009; Tan etal. 2012a).

Exogenous electrical stimulation of the H-reflex circuit candirectly induce changes in H-reflex response, which can persistfor many hours or days (Chen et al. 2003, 2006). Theseprevious studies demonstrate the presence of activity-depen-dent plasticity within the monosynaptic reflex system. In ourcurrent study, to establish a structure-function relationshipbetween dendritic spine remodeling and H-reflex dysfunctionafter SCI, we were required to assess spine changes andH-reflex function within the same animals. This acute H-reflexfunction and dendritic spine assessment approach limited ourability to control for the possible confound that dendritic spinechanges could also have resulted from EMG reflex testing,which required direct stimulation of muscle-sensory nerves.Although H-reflex testing only lasted about 1 h per animal andwe ensured that all animals underwent similar testing proto-cols, we cannot exclude the possibility that each treatmentgroup could have had a different capacity to respond topotential EMG testing-induced dendritic spine changes. Wenote that in uninjured sham animals, there was a greaterproportion of thin-shaped dendritic spines compared withmushroom-shaped dendritic spines (see Figs. 4 and 5). Thin,filopodia-like dendritic spines are thought to represent newlyformed or more plastic dendritic spines, whereas mushroom-shaped spines may represent more stable, mature spines(Bourne and Harris 2007, 2008). This observation in shamanimals does suggest the possibility that EMG testing couldhave resulted in the de novo presence of more structurallyresponsive thin-shaped dendritic spines. Our current experi-ments, however, do not permit us to determine whether spinechanges are a sole result of treatments (i.e., SCI, drug inter-vention, etc.) or a combination of treatments and the potentialdirect effect of EMG electrophysiological assessment, whichcould also have influenced spine morphologies.

Our findings raise the question of how altered dendritic spinemorphologies below the level of injury contribute to hyperex-citable spinal reflex function. The dendritic spine profiles weobserved after SCI below the injury can have direct biophysicaleffects on motor neuron excitability (Rall et al. 1967; Segevand Rall 1988; Tan et al. 2009). The distribution of spinesynapses closer to motor neuron cell bodies can increase theoverall impact of excitatory input and transmission. The in-creased head volume of mature, mushroom-shaped dendriticspines permits increased clustering of excitatory membranereceptors (e.g., AMPA receptors; Wiens et al. 2005). Theseproperties can amplify synaptic transduction. Importantly, weobserved an increase in mature spine geometries after SCI,which could increase input discretization (e.g., narrow excit-atory postsynaptic potential waveforms) and allow trains ofsuprathreshold potentials to propagate at higher rates of activ-ity with greater fidelity (Rall 1967; Rall et al. 1967). Thistransmission property could contribute to excessive firing inmotor neurons (Tan et al. 2009; Yuste and Urban 2004) andfacilitate supra- and subthreshold temporal summation (Carteret al. 2007; Gold and Bear 1994; Martiel et al. 1994). In ourstudy, temporal summation would most likely have occurred atthe shorter interpulse intervals during H-reflex testing. Con-secutive transcutaneous electrical stimulations to facilitatewithdrawal reflex have demonstrated maximal temporal facil-itation at the 10- to 20-Hz range (Arendt-Nielsen et al. 1997,



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2000), suggesting the presence of a temporally dependentintegration mechanism within reflex pathways. Thus it is in-teresting to consider that SCI-induced dendritic spine changeslead to an amplification of presynaptic excitatory input andenhancement of a temporally dependent integration mechanismto increase the gain of H-reflex function after SCI.

We have implicated Rac1 in the pathological changes thatcontribute to chronic nociceptive hyperexcitability after SCI(Tan et al. 2008). Rac1 is a small-molecule GTPase that isinvolved in regulating dendritic spine morphology (Tashiroand Yuste 2008). In a mouse model of fragile X syndrome(FMR knockout), Rac1 dysfunction is associated with abnor-mal dendritic spine morphology and decreased pain (Chen etal. 2010; Lee et al. 2003; Price et al. 2007). In our model ofSCI, we administered NSC23766, a Rac1-specific inhibitor thatdoes not affect Cdc42 or Rho GTPases or affect the interactionof Rac1 to its downstream effector, PAK1 (Gao et al. 2004).Although our current study does not preclude off-target effects,we have previously shown that the Rac1 inhibitor disrupts thedevelopment and appearance of dendritic spines in vitro (Tanet al. 2011) and that NSC23766 treatment is effective inattenuating dendritic spine remodeling in the dorsal horn afterSCI (Tan et al. 2008). Moreover, NSC23766 treatment doesnot significantly affect electrophysiological or behavioral signsof pain-reflex withdrawal function in normal, uninjured ani-mals (Tan et al. 2012b). Although microglial activation hasbeen implicated in altering the excitability of sensory neuronsafter injury (Tan et al. 2012a; Zhao et al. 2007a), administra-tion of NSC23766 does not appear to affect the activation ofmicroglia after nerve injury (Tan et al. 2011). In agreementwith previous studies (Beauparlant et al. 2013; Kitzman 2007),we report that VGluT1 expression did not change after SCI,suggesting that the presence of H-reflex hyperexcitability is notdue to increased excitatory presynaptic afferent inputs (Kitz-man 2007). Although it is possible that Rac1 inhibition mayhave affected presynaptic elements, NSC23766 treatment onlydecreased VGluT1 areal density in the intermediate zone andventral horn. This suggests that the Rac1-inhibitor had atopographically specific effect on deeper lamina only, wheresensory-motor neuron populations exhibit dendritic spine plas-ticity after injury (Tan et al. 2008, 2011).

A major challenge in SCI research has been identifying themechanisms that contribute to secondary complications such asspasticity and pain. Our previous work has demonstrated thatdendritic spine remodeling in the dorsal horn contributes tohyperexcitability associated with neuropathic pain after SCI(Tan et al. 2008; Tan and Waxman 2012). In the present workwe show that dendritic spine dysgenesis on �-motor neuronsalso contribute to the development of spasticity. Our datafurther show that Rac1 signaling participates in this dendriticspine remodeling and may provide a novel opportunity tospecifically address the underlying pathophysiology of spastic-ity after SCI.

ACKNOWLEDGMENTS

We thank Pamela Zwinger and Peng Zhao for excellent technical assis-tance.

GRANTS

This work was supported by grants from Paralyzed Veterans of America(PVA) and the Department of Veterans Affairs (VA) Medical Research

Service and Rehabilitation Research Service. A. M. Tan is funded by the PVAResearch Foundation and VA Career Development Award 1 IK2 RX001123-01A2. The Center for Neuroscience and Regeneration Research is a Collabo-ration of PVA with Yale University.

DISCLOSURES

No conflicts of interest, financial or otherwise, are declared by the authors.

AUTHOR CONTRIBUTIONS

S.P.B., S.L., and A.M.T. performed experiments; S.P.B. and A.M.T. ana-lyzed data; S.P.B. and A.M.T. prepared figures; S.P.B., S.L., S.G.W., andA.M.T. approved final version of manuscript; S.G.W. and A.M.T. interpretedresults of experiments; S.G.W. and A.M.T. edited and revised manuscript;A.M.T. conception and design of research; A.M.T. drafted manuscript.

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Dendritic spine dysgenesis contributes to hyperreflexia after spinal