FEM model: mouse spinal cord.

A three-dimensional finite element model (FEM) of both the in vivo mouse spinal cord and the ex vivo mouse spinal cord recording setup was created in COMSOL Multiphysics (version 5.1; COMSOL, Burlington, MA) to compare the electric fields (and thereby currents) generated by the scaled-down electrodes in the experimental preparation with those expected to be generated by the same electrode in an in vivo mouse and by a commercial epidural lead in a human. For the in vivo FEM, the dimensions of the cerebrospinal fluid (CSF) space, dura, epidural fat, and vertebral bony layers were generated by downscaling the x and y dimensions of the corresponding anatomical features from a previously published FEM of the human thoracic spinal cord (Lee et al. 2011) by factors of 4 and 4.5, respectively, derived from the previously described scaling procedure. Dirichlet boundary conditions (V=0) were used at the lateral and dorsoventral boundaries of the model, and Neumann boundary conditions (n·J=0) were used at the rostrocaudal boundaries of the model. For the ex vivo FEM, the model was scaled according to the actual experimental setup and consisted of an insulated rectangular chamber measuring 76 mm in length, 45 mm in width, and 8 mm in depth filled with artificial CSF (aCSF) and a 1-mm-diameter ground wire placed 10 mm from the center of the stimulation lead. In the ex vivo model, the faces of the ground wire were set to Dirichlet boundary conditions (V=0), and the other outer boundaries of the model were set to Neumann boundary conditions (n·J=0).

An extruded mouse spinal cord 35 mm in length and a model of the experimental stimulation lead were preserved across both the in vivo and ex vivo models. The dimensions of the gray and white matter of the mouse spinal cord were obtained by directly tracing the left half of a slice image of the mouse T10 spinal cord (Watson et al. 2009) and reflecting the tracing over the midline using MATLAB (version R2015a; The MathWorks, Natick, MA) before being imported into COMSOL. Furthermore, the gray matter was displaced 20 µm ventrally from the original tracing to allow the spacing between the dorsal gray matter and white matter boundary to fulfill the minimum mesh element size requirement of the finest mesh setting in COMSOL (~15.2 µm). Stimulating leads were modeled as conductive thin cubic domains with a side length of 0.3 mm surrounded by a rectangular insulating substrate with outer dimensions of 2.4 mm × 0.6 mm × 0.5 mm and suspended from an insulating cylindrical electrode mount 0.4 mm in diameter and centered on the rectangular substrate. The ventral surfaces of each contact on each lead were each set to a uniform current density with Neumann boundary conditions corresponding to the total stimulation amplitude (50, 200 µA) divided by the surface area of each lead (Pelot et al. 2018). Stimulation currents from each of the two contacts were set to the same amplitude at opposite polarities to depict the electric field generated by bipolar stimulation in the spinal cord proximal to the leads. Domain conductivities representing tissues and materials of interest are shown in Table 1, and domain permittivities were set equal to the permittivity of free space, consistent with the quasi-static assumption (Bossetti et al. 2007). Results for each model were then obtained by solving for Laplace’s equation (·σV=0) over an adaptively scaled tetrahedral mesh.

Constant-current stimulators provide compliance voltages that automatically adjust to the tissue impedance. Boundary conditions of the model were set with this in mind. Because significant clinical variability exists in patient perception thresholds, the theoretical outcomes of the model are representative of electric field strengths expected. The amplitude of current delivered to the mouse model was scaled using the intended experimental geometry for the voltages and electric fields to conform to the clinical ranges without severe discrepancies. One clear limitation of the mouse model is that it represents a scaling from this normalized, exemplary range.

Spinal cord isolation.

All procedures were approved by the Emory University Institutional Animal Care and Use Committee: C57/Bl6 mice (both sexes, n = 35, postnatal day 60 and older) were anesthetized with ketamine (100 mg/kg) and xylazine (10 mg/kg) mix by intraperitoneal injection following light anesthetization in an isoflurane chamber. To induce tissue hypothermia, the dorsal skin overlying the vertebral column was removed and mice were placed in an ice bath until there was a noticeable slowing of the respiration rate (2–3 min) before decapitation and isolation of spinal cord in a dish containing ice-cold, oxygenated (95% O₂-5% CO₂), low-Ca²⁺ aCSF containing (in mM) 128 NaCl, 1.9 KCl, 13.3 MgSO₄, 1.1 CaCl₂, 1.2 KH₂PO₄, 10 glucose, and 26 NaHCO₃. The isolated cord was equilibrated to room temperature for 1 h and then pinned dorsal side up in a Sylgard-lined recording chamber while superfused with an oxygenated aCSF containing (in mM) 128 NaCl, 1.9 KCl, 1.3 MgSO₄, 2.4 CaCl₂, 1.2 KH₂PO₄, 10 glucose, and 26 NaHCO₃ at ~40 ml/min. All experiments were undertaken at room temperature.

Histological test of spinal cord viability.

In four experiments, 8 h following spinal cord isolation and experimentation, the spinal cord was fixed in 2% paraformaldehyde for 2 h, rinsed with phosphate buffer solution (PBS), sectioned with a vibrating blade microtome (Leica VT1000S) in 300-µm-thick slices, stained 30–60 min in 1:250 Neurotrace (N-21482; Molecular Probes) mixed in PBS with 0.3% Triton, and then washed twice in PBS without Triton. Captured images of stained slices allowed assessment of neuronal anatomical integrity (Nikon Eclipse E800 microscope with DXM1200 camera).

Electrophysiology.

The DC largely comprises myelinated Aαβ primary afferents fibers that transmit proprioceptive and mechanosensitive information but also includes ascending projections from PSDC tract neurons and unmyelinated axons, whereas the neighboring LT mostly consists of C fibers with short-range projections and Aδ nociceptors with long-range projections (Lidierth 2007) (Fig. 2A).

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Fig. 2.

Ex vivo adult mouse preparation: experimental setup and viability. A: schematic of gross anatomy of spinal cord from dorsal surface and simplified organization of axon fiber populations subject to recruitment by spinal cord stimulation (SCS; inset). Axonal recruitment with SCS was assessed in dorsal roots (DR), dorsal column (DC), and Lissauer tract (LT). Whereas the DC predominately carries nonnociceptive sensory information, the neighboring LT contains projection from nociception transmitting primary afferents. PSDC, postsynaptic dorsal column. B: schematic of general experimental paradigm. Two electrodes (~1.5 mm apart) are placed on or above the medial DC in a region between T9 and L3. This is used to simulate a bipolar SCS system. Responses to SCS are recorded via glass suction electrodes on the DC and DRs and by a microelectrode on LT. Inset: orientation of bipole (cathode, black; anode, red) and direction of current flow for cathodic and anodic stimulation used for experimental studies. R and C denote rostral and caudal, respectively. C: postexperimentation Nissl staining revealed persistent cellular viability 8 h after cord isolation (top) but dramatic deterioration when superfused for 2 h in the absence of oxygenation (bottom).

Experimental characterization of axonal recruitment was conducted with clinically analogous SCS by using a bipolar electrode constructed of two glass electrodes (300-μm tip diameters, separation 1.5–2 mm) filled with aCSF and positioned (rostrocaudally) medial over the DC, typically in caudal thoracic segments (T9–T11; Fig. 2B). Circular glass electrodes were used for experimental studies due to the repeatability of the construction process relative to manual cutting of 300 μm × 300 μm electrodes. A constant-current stimulator (Hammar et al. 2011) was used to deliver a single monophasic pulse of bipolar SCS while contacting (0 μm) or above the spinal cord DC (200 or 600 μm). Stimulation amplitudes were incrementally increased from 1 to 500 μA with 50-, 200-, and 500-μs pulse durations for assessments of recruitment threshold. The advantage of a constant-current stimulation protocol, based on modeling studies, is that resistivity values normally seen for dura and arachnoid do not significantly affect spinal cord current densities (Coburn and Sin 1985). Thus our experimental arrangement excludes the presence of dura but still is expected to realistically appraise neural system recruitment at defined electrode configurations and distances from the spinal cord at the stimulus intensities applied.

Glass recording suction electrodes were positioned in the lumbar spinal cord on the DC (tip diameter 200 μm), DR (tip diameter 200–250 μm), and LT (tip diameter 1–2 μm, 3–5 MΩ). Targeted recording of LT was accomplished by positioning a glass microelectrode lateral to the DC, in between dorsal root entry zones. Appropriate positioning was confirmed by stimulating the ipsilateral spinal cord one to three spinal segments rostral to the recording site and observing that the conduction velocity of evoked response was slower than that recorded in DR and DC of the same segment. Recruitment threshold was defined as the lowest stimulus amplitude producing a >50% response rate in any recording location. Since threshold was typically lowest for DC axonal recruitment (see results), relative thresholds for recruitment were standardized as the multiple of DC recruitment threshold (T_DC) necessary to elicit responses in LT and DR. All recorded data were digitized at 50 kHz (Digidata 1322A 16-bit data acquisition system; Molecular Devices) with pClamp acquisition software (v. 10.7; Molecular Devices). Recorded signals were amplified (5,000 times) and low-pass filtered at 3 kHz using in-house amplifiers. Unless otherwise stated, reported results are for cathodic stimulation 200 μm above the cord with 200-μs pulse duration. Recruitment order trends for cathodic and anodic stimulation were similar, despite differences in polarity. Cathodic stimulation, in some cases, recruited populations at a similar or lower threshold, and SCS experimental studies utilizing monophasic stimulation frequently report observations based on cathodic stimulation (Song et al. 2009; Stiller et al. 1996; Yakhnitsa et al. 1999; Yuan et al. 2014). This is likely due to knowledge that cathodic stimulation recruits neural tissue at lower amplitudes than anodic stimulation (McIntyre and Grill 1999; Ranck 1975; Rattay 1998). For these reasons, we chose to focus our reporting on cathodic stimulation. In all data presented, the number of animals utilized for analysis is represented by the noted n value. For each animal, a representative value was determined by averaging the response from multiple sweeps/trials within that animal (a minimum of 5).

Preclinical finite element analysis model.

Preclinical stimulation was simulated using a bipolar electrode consisting of two square contacts, dimensions of which are noted in materials and methods, with an edge-to-edge spacing of 1.2 mm and positioned epidurally over the dorsal surface at the midline of the spinal cord, for both the ex vivo and in vivo models described in materials and methods. The clinical standard for SCS electrode placement involving SCS at conventional frequencies associated with paresthesia-based SCS involves implantation of the electrode at the T7–T8 vertebral segments, with the intent of targeting the dorsal column fibers coursing through the T9–T10 spinal cord level. Using a mouse model of the T9–T10 spinal cord is consistent with this therapeutic approach and with prior literature (Canbay et al. 2014). The electric field magnitudes generated during ex vivo preclinical SCS (50 to 200 µA), with a CSF fluid layer of 0, 200, or 600 µm, were compared with those generated in the clinical model with a typical range of SCS amplitudes (3 and 10 mA). Results from analysis were used to guide experimental investigations in the ex vivo adult mouse spinal cord. Qualitatively, preclinical SCS positioned 200 μm (50–200 μA) above the spinal cord in the ex vivo model generated electric field distributions comparable to those generated by clinical SCS (Fig. 3). Preclinical SCS 0 μm above the spinal cord with the same amplitudes (50–200 μA) suggests broader recruitment of populations in the DC and dorsal horn than would be observed in the clinical SCS, indicating that lower stimulation amplitudes than those modeled would be advised for analogous recruitment, whereas stimulation 600 μm above suggests that higher amplitudes of stimulation would be necessary to capture the range of electric field distributions observed in the clinical case. Comparisons between ex vivo and in vivo preclinical cases suggest that although 200 µm is suitable for the ex vivo preclinical SCS, 600 µm in vivo most closely mimics the relative CSF thickness in human and that the bone, fat, and dura contribute to the generation of comparable electric field distributions with stimulation this distance above the cord (Supplemental Fig. S1; all Supplemental Material is available at https://doi.org/10.6084/m9.figshare.8323535).

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Fig. 3.

Finite element modeling of electric field magnitudes identifies stimulation amplitudes for clinically analogous spinal cord stimulation (SCS) in the adult mouse. A: rostrocaudal orientation of bipole for clinical (left) and preclinical (right) SCS. The anode (red) is rostral over the spinal cord (T9), and the cathode (blue) is caudal over the spinal cord (T10). Scales are capped at±0.4 V; scale bars are 5 (left) and 0.5 mm (right). B: clinical SDS. Electric field magnitudes in the human spinal cord during stimulation were generated using a finite element model of clinical bipolar SCS with a clinical 1 × 8 percutaneous lead with amplitudes in the typical range. C: preclinical ex vivo SCS. Electric field magnitudes in the spinal cord during SCS were generated using a finite element model of the ex vivo mouse spinal cord (absence of dura, epidural fat, and bone) in a rectangular bath of cerebrospinal fluid, equivalent to experimental conditions. Properties, mesh settings, and boundary conditions of the finite element model are consistent with those of the clinical model except that all dimensions have been scaled down by a factor of 4.5. Stimulation was simulated using a bipolar electrode consisting of 2 square contacts of side length 0.3 mm with an edge-to-edge spacing of 1.2 mm and positioned epidurally 0, 200, or 600 μm over the dorsal surface of the midline of the spinal cord. The fields generated at 200 μm above the spinal cord are most comparable to those generated in clinical conditions.

SCS recruits afferents that invade multiple segmental dorsal roots and antidromically conduct at velocities consistent with all classes of primary afferents.

Multisegmental recruitment was directly assessed with bipolar SCS (50 and 200 µA, 200 µs) positioned 0 (n = 5) or 200 µm (n = 3) above the DC. Afferent recruitment was seen in all caudal DRs sampled, the most caudal being eight spinal segments away (Fig. 4). In the example shown, T9/T10 SCS recruitment was abolished following DC lesion above T11, demonstrating that stimulation recruited axons at the stimulation site with subsequent antidromic propagation to DRs. CVs were divided into three categories approximating reported peripheral CVs of Aαβ (>4.2 m/s), Aδ (0.56–4.2 m/s), and C fibers (<0.56 m/s) at room temperature (Pinto et al. 2008b).

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Fig. 4.

Spinal cord stimulation (SCS) recruits primary afferents that invade multiple segmental dorsal roots and conduct at velocities consistent with all classes of sensory fibers. A: SCS directly activates afferent axons that project to multiple caudal spinal segments. A single pulse of SCS (200 μA, 200 μs) was applied 200 μm above T9/T10, and antidromic responses in select caudal dorsal roots (DRs; T11–L5) were recorded before (black) and after (red) dorsal column (DC) lesioning above T11 with iridectomy scissors. Lesion of the DC blocked all stimulus-evoked responses in DRs. Each trace is the average response of 10 sweeps. B: distance-dependent recruitment of afferent fiber populations can be identified by differences in peripheral CV. In the experiment shown, bipolar SCS was applied 200 μm above T11/T12 or L2/L3 at amplitudes of 50 and 200 µA (500 μs). T11/T12 SCS led to the recruitment of A fibers in the L5 DR, but only at the higher stimulus amplitude (arrow). In comparison, SCS at a closer segment (L2/L3) recruited A fibers at the lower intensity with higher magnitude stimulation leading to recruitment of slower conducting C fibers (arrows). Each trace is the average of 6 sweeps. The peripheral conduction velocities estimated using this configuration are summarized in Table 2. Shaded box identifies the location of stimulation artifact.

To accurately identify afferent classes on the basis of peripheral CV, we used the two-point recording method in DRs (Fig. 4B; Table 2). We observed a rostrocaudal distance dependence of electrode position on afferent recruitment. By reducing stimulus amplitude (50 μA), we also observed that low-amplitude SCS only recruited A fibers in DRs from nearby spinal segments (0–3 segments away; n = 8) and that higher amplitude SCS (200 μA) was required to recruit A fibers in more distant DRs (n = 7; Fig. 4B; Table 2). Higher amplitude and longer pulse duration SCS also recruits slower conducting Aδ and C fibers in DRs of nearby segments (Fig. 4B; Table 2). Although recruitment of C fibers was not observed in DRs more distal to SCS, we cannot exclude the possibility that multisegmentally projecting unmyelinated C fibers (Mandadi et al. 2013) were undetectable due to dispersion of the volley.

SCS evokes synaptically mediated recruitment of primary afferents at thresholds comparable to direct antidromic activation.

Dorsal root potentials (DRPs) and dorsal root reflexes (DRRs), resulting from primary afferent depolarization (PAD), were observed with SCS. These longer latency, synaptically mediated events had variable onset latency and amplitude (observed in 13 of 23 animals). Recordings at the dorsal root entry zone (DREZ) permitted observation of both events, with arrival of suprathreshold DRRs coinciding with the presence of the DRP (Fig. 5, A and Bi). Recordings away from the DREZ led to observations of only DRRs. The jitter and sensitivity of this activity to bicuculline (10 μM; n = 4) confirmed it as synaptically mediated (Fig. 5B).

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Fig. 5.

Spinal cord stimulation (SCS) recruited GABAergic presynaptic inhibitory pathways observed as bicuculline-sensitive dorsal root potentials (DRPs) and dorsal root reflexes (DRRs). A: SCS-generated DRP with suprathreshold DRRs recorded at the L5 dorsal root entry zone. By convention, negativity is presented upwards to convey the DRP as primary afferent depolarization. Shaded box indicates the location of stimulus artifact and truncated direct antidromic axonal volley preceding the slow DRP. The average response (black) is presented overlying 10 individual events (gray) to demonstrate variability in timing of individual DRRs. B: SCS evokes DRP and DRRs that are blocked following application of the GABA_A receptor antagonist bicuculline (Bic; 10 μM). i: averages of 10 sweeps from the T13 dorsal root entry zone. Both DRP and DRRs are blocked after the addition of Bic (blue). ii: Bic-sensitive DRRs are shown in the absence of an underlying DRP. Shown are 6 events overlaid from an L2 DR recording obtained farther from the root entry zone. In both i and ii, shaded box indicates the location of stimulation artifact and direct primary afferent recruitment. C: schematic depicting possible circuitry responsible for GABAergic actions on afferent axons leading to a form of presynaptic inhibition called primary afferent depolarization (PAD), experimentally observed as DRPs and DRRs. SCS antidromically recruits Aαβ afferent axons responsible for recruiting the spinal circuit leading to PAD. Aαβ afferents synaptically recruit last-order GABAergic interneurons directly or via interposed interneurons (not shown). GABAergic axo-axonic synapses on primary afferents activate Bic-sensitive GABA_A receptors that mediate Cl⁻ efflux to produce PAD. Because these experiments cannot identify the afferent axons generating PAD, possible actions on all afferent classes are shown.

Two distinct populations of DRRs were commonly observed in lumbar DRs, the earliest event being 9.0±1.6 ms after the onset of the direct antidromic response, with the second DRR event occurring 9.2±1.2 ms later. Thresholds for direct antidromic axonal recruitment and synaptically mediated (indirect) DRRs were comparable, with mean threshold for DRR recruitment being numerically only 3.8±2.2 μA higher than the antidromic event (n = 5; P > 0.05, paired t test; Fig. 6). This suggests that the indirect recruitment is activated by excitation of low-threshold primary afferents. The population(s) of axons receiving primary afferent depolarization was not identified, but the presence of two different populations of DRRs suggests that one population may be initiating PAD, whereas two separate populations could be receiving PAD (Fig. 5C), although it is also possible that the second DRR event reflects recurrent activity initiated by the first DRR event or doublet firing within the same recruited axons.

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Fig. 6.

Spinal cord stimulation (SCS) evokes direct and indirect primary afferent recruitment at comparable thresholds. SCS was applied contacting (0 μm; A) or 200 µm above (B) T9/T10 during recording of a lumbar dorsal root (DR). Stimulus amplitude was incrementally increased (1–200 µA, 200 µs; i), and the recruitment threshold for direct primary afferent recruitment and synaptically mediated primary afferent activity (indirect) was identified post hoc (ii). Mean threshold () for indirect recruitment was not significantly higher than that observed for direct recruitment when SCS was applied contacting (0 μm; n = 4 animals, 15.2±10.1 vs. 13.0±6.9 μA, respectively) or 200 µm above (n = 5 animals; 49.8±48.3 vs. 48.0±52.3 μA, respectively; Aii and Bii, paired t test). Arrows indicate direct recruitment, dashed boxes denote indirect recruitment, and shaded boxes depict stimulation artifact. Black lines are the average of 6 traces.

Characterization of dorsal column recruitment with SCS.

DC recruitment was also characterized in the adult mouse spinal cord as depicted in Fig. 7A, left. Recruitment of axons in DR, DC, and LT was compared when SCS was contacting (0 μm), 200 μm, or 600 µm above the spinal cord (n = 12). Direct recruitment of axons in LT was identified on the basis of a lack of variability in onset latency and amplitude (Fig. 7A, bottom right). Recruitment order and relative thresholds of recruitment was then determined by increasing the magnitude of constant-current SCS (200-μs pulse width; Fig. 7A). Interestingly, when SCS electrodes were contacting or above the DC, axonal recruitment was first seen in the DC regardless of stimulation polarity (n = 12). The mean threshold current for recruitment with SCS contacting the DC was lower for cathodic (6.3±4.3 µA) than anodic (9.1±3.7 µA) stimuli (P < 0.05, n = 12). When stimulation was positioned 200 or 600 µm above the spinal cord, recruitment thresholds were similar at 30.5±26.6 (cathodic) vs. 26.3±13.3 µA (anodic; P = 0.213, n = 12) or 231.9±127.5 (cathodic) vs. 278.5±106.0 µA (anodic; P = 0.125, n = 10), respectively. Compared with thresholds observed with SCS contacting the spinal cord DC, threshold values were ~3–5 times higher at 200 µm above (P < 0.05) and 31–37 times higher at 600 µm above the DC (P < 0.001; Fig. 7B). Thresholds for SCS positioned 600 µm above the DC were 8–11 times that seen with placement 200 µm above (P < 0.001).

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Fig. 7.

Spinal cord stimulation (SCS) recruitment thresholds are lower for dorsal column (DC) axons. A: example configuration for investigations of recruitment threshold and order. To identify the recruitment threshold of dorsal axon tracts, bipolar SCS (T9/T10, 1–600 µA, 200 µs) was applied contacting or above the spinal cord, with simultaneous recordings of dorsal root (DR), Lissauer tract (LT), and DC at the same segmental level. In the example, DC (purple) is recruited first at threshold (T_DC), with DR (green) recruitment observed at 1.7× T_DC and overt LT (blue) recruitment observed at 2.3× T_DC. Shaded box identifies the location of stimulation artifact; arrows point to dashed boxes denoting the presence of indirect recruitment/reflexes. The colored trace is an average of 6 sweeps. B: Threshold_DC was determined with stimulation 0, 200, or 600 µm above T9/T10 (n = 12 animals). T_DC significantly increased at 200 µm above and 600 µm above the spinal cord. *P < 0.05; ***P < 0.001 (matched one-way ANOVA, Tukey’s multiple comparison). C: DC recruitment was observed at the lowest threshold and significantly lower than threshold for DR and LT with SCS at 0 (n = 12) or 200 µm (n = 12) above the spinal cord. With SCS 600 µm above, recruitment beyond DC was observed in only n = 2 of 10 cases. Values are mean and SD for the test conditions. *P < 0.05; ***P < 0.001; ****P < 0.0001 (Friedman test with Dunn’s multiple comparisons).

Relative thresholds for dorsal root and Lissauer tract recruitment.

We also compared the relative recruitment of antidromic events in DR and LT relative to the threshold seen in dorsal column (T_DC) when SCS was contacting or positioned above the DC (n = 12; Fig. 7, A and C). When SCS was contacting the DC, events in DR and LT were recruited at 3.8±2.4 (P < 0.05) and 5.0±4.5 (P < 0.001) times T_DC, respectively. At 200 µm above the DC, events in DR and LT were recruited at 2.6±1.0 (P < 0.05) and 4.4±3.1 (P < 0.0001) times T_DC, respectively. In comparison, there were no differences in relative recruitment threshold between DR and LT when SCS was touching (P > 0.9) or 200 μm above the DC (P = 0.26). At 600 µm above, only DC recruitment was seen in most experiments (n = 7/10), even at maximal current amplitudes (>200 μA). However, when DR and LT recruitment was seen, both thresholds were similar to T_DC, being 1.5±0.4 (n = 5) and 1.5±0.1 (n = 2) times T_DC. No axonal recruitment was elicited in 3 of 10 experiments with placement 600 µm above the DC.