Muscle afferent excitability testing in spinal root-intact rats: dissociating peripheral afferent and efferent volleys generated by intraspinal microstimulation

Excitability testing of primary afferents has been used to evaluate presynaptic modulation of synaptic transmission in experiments conducted in vivo. However, to apply this method to muscle afferents of animals with intact spinal roots, it is crucial to dissociate antidromic and orthodromic volleys induced by spinal microstimulation. We propose a new method to make this dissociation possible without cutting spinal roots and demonstrate that it facilitates excitability testing of muscle afferents.

Abstract

Presynaptic inhibition of the sensory input from the periphery to the spinal cord can be evaluated directly by intra-axonal recording of primary afferent depolarization (PAD) or indirectly by intraspinal microstimulation (excitability testing). Excitability testing is superior for use in normal behaving animals, because this methodology bypasses the technically challenging intra-axonal recording. However, use of excitability testing on the muscle or joint afferent in intact animals presents its own technical challenges. Because these afferents, in many cases, are mixed with motor axons in the peripheral nervous system, it is crucial to dissociate antidromic volleys in the primary afferents from orthodromic volleys in the motor axon, both of which are evoked by intraspinal microstimulation. We have demonstrated in rats that application of a paired stimulation protocol with a short interstimulus interval (ISI) successfully dissociated the antidromic volley in the nerve innervating the medial gastrocnemius muscle. By using a 2-ms ISI, the amplitude of the volleys evoked by the second stimulation was decreased in dorsal root-sectioned rats, but the amplitude did not change or was slightly increased in ventral root-sectioned rats. Excitability testing in rats with intact spinal roots indicated that the putative antidromic volleys exhibited dominant primary afferent depolarization, which was reasonably induced from the more dorsal side of the spinal cord. We concluded that excitability testing with a paired-pulse protocol can be used for studying presynaptic inhibition of somatosensory afferents in animals with intact spinal roots.

NEW & NOTEWORTHY Excitability testing of primary afferents has been used to evaluate presynaptic modulation of synaptic transmission in experiments conducted in vivo. However, to apply this method to muscle afferents of animals with intact spinal roots, it is crucial to dissociate antidromic and orthodromic volleys induced by spinal microstimulation. We propose a new method to make this dissociation possible without cutting spinal roots and demonstrate that it facilitates excitability testing of muscle afferents.

originally discovered by Eccles and collaborators (Eccles et al. 1961, 1962b), as well as by Frank and Fuortes (1957), presynaptic inhibition is now known to function broadly within the central nervous system (CNS) and plays a role in the hippocampus (Laviv et al. 2011) and in the olfactory (Olsen and Wilson 2008), visual (Schubert et al. 2013), and corticothalamic systems (Alexander and Godwin 2005). Presynaptic inhibition not only regulates sensory input into specific modalities (Janig et al. 1967) or projections (Chavez et al. 2012; Contreras-Hernandez et al. 2015; Lomeli et al. 1998), it also may be crucial for higher behavioral control, such as long-term memory (Cullen et al. 2014), behavioral choices (Gaudry and Kristan 2009), sense of taste (Chu et al. 2014), and reward expectation (Jiménez-Rivera et al. 2012). Recent discoveries of spinal neuron molecular markers in mice (for summary, see Alaynick et al. 2011) provide further evidence for the importance of presynaptic inhibition in controlling normal behavior. For example, Fink et al. (2014) ablated GABA neurons that project to primary afferents in transgenic mice, which results in deficits in smooth arm movement and induces ataxic movement. However, cell-targeted genetic ablation or chemical inactivation abolishes presynaptic inhibition either chronically or semichronically. Therefore, time-varying changes in the level of presynaptic inhibition in behaving animals cannot be analyzed using this approach. A new method that can measure the level of presynaptic inhibition in the context of a behavioral experiment is needed for understanding the role of presynaptic inhibition in controlling normal movements.

Presynaptic inhibition is induced by different types of presynaptic receptors (Miller 1998), with that induced by GABA_A receptors the most extensively studied (for references, see Rudomin and Schmidt 1999). In the spinal cord, activation of GABA_A receptors on the intraspinal terminals of afferent axons produces so-called primary afferent depolarization (PAD) by opening chloride channels and allowing the efflux of Cl⁻ ions from the terminals (Alvarez-Leefmans et al. 1988; Gallagher et al. 1978). This depolarization increases the conductance of the terminal membrane (Curtis et al. 1995) and inactivates both Na⁺ (Graham and Redman 1994) and voltage-gated Ca²⁺ channels (Graham and Redman 1994; Walmsley et al. 1995), which may reduce neurotransmitter release. Therefore, GABA_A-mediated presynaptic inhibition can be estimated with PAD (Eccles et al. 1962a, 1963) by recording intra-axonal potentials from the terminals of primary afferents with intracellular electrodes (Brink et al. 1984; Gossard et al. 1989). However, intracellular recordings are very sensitive to movement of the targeted cell or axon; therefore, although not impossible (Chen and Fetz 2005; Lee et al. 2009), it is difficult to apply in a study using behaving animals.

Excitability testing of primary afferent terminals, which was introduced by Wall (1958) as a method for evaluating PAD, may be an alternative to intracellular recording. PAD increases the probability of inducing antidromic volleys when the corresponding terminals are electrically stimulated; therefore, the size of the antidromic volleys recorded at the peripheral nerve reflects the extent of PAD at the time of stimulation. The advantages of this method are that the PAD level can be estimated without measuring intra-axonal potentials, and the PAD level can be measured in a variety of experimental preparations, including acute experiments using anesthetized animals (Harrison and Jankowska 1989; Riddell et al. 1995) or decerebrated animals exhibiting fictive movements (Baev 1980; Duenas and Rudomin 1988). Previous studies, including our own (Ghez and Pisa 1972; Seki et al. 2003, 2009), have applied this technique in awake, behaving animals in the context of a motor task and have reported that the PAD size was larger during active movement of the cutaneous afferent from the forearm (i.e., the superficial radial nerve), suggesting that input from the cutaneous afferent was suppressed by presynaptic inhibition. This suppression may have been induced by descending commands, because it began before the onset of muscle activity. These reports demonstrate that excitability testing can be used to measure presynaptic inhibition of cutaneous afferents in behaving animals, thereby documenting the importance of presynaptic inhibition in behavioral control.

Given the importance of proprioceptive input from muscles and joints for movement control (e.g., Prochazka 2011), the next conceivable step was to apply excitability testing to muscle and joint afferents in normal, behaving animals. However, the anatomical constraints of proprioceptive afferents in limbs appeared to be prohibitive. Most afferents from muscle and tendon receptors merge into mixed nerves (i.e., nerves containing both afferent and efferent fibers). During excitability testing, therefore, an electrode implanted in a mixed nerve records both the antidromic and orthodromic volleys. Thus, in previous studies, the ventral root was transected to isolate the antidromic volleys from the recorded nerve potentials (Harrison and Jankowska 1989; Riddell et al. 1995; Wall 1958). However, this technique is not appropriate for measuring PAD in animals performing normal behaviors, because transecting ventral roots also abolishes muscle activity. This limitation is applicable only to the muscle and joint nerves; therefore, it has been possible to measure antidromic volleys in the cutaneous afferents of behaving animals without cutting ventral roots (Ghez and Pisa 1972; Seki et al. 2009). When measuring PAD in muscle and joint afferents of normal behaving animals, consequently, it is crucial to establish a method that identifies antidromic volleys with intact peripheral nerves and ventral roots.

In this study, we propose electrophysiological criteria that distinguish antidromic from orthodromic volleys using a paired-pulse protocol for excitability testing. Specifically, in peripheral nerves, we measured the amplitude and latency of antidromic and orthodromic volleys evoked by intraspinal paired microstimulation in rats with transected ventral and dorsal roots, respectively. After establishing the stimulation parameters that successfully dissociated antidromic and orthodromic volleys, we tested whether the antidromic volley classified by this method could exhibit PAD by applying conditioning stimuli to other muscle afferents in animals with intact spinal roots.

MATERIALS AND METHODS

Ethics statement.

All experimental procedures were approved by the local ethics committee for animal research at the National Institute of Neuroscience.

Animals.

Experiments were performed using 20 male Jcl:Wistar rats (3–8 mo of age; body weight 437.1 ± 64.4 g) that were raised under specific pathogen-free conditions with ad libitum access to food.

Surgical procedures.

Rats were anesthetized with pentobarbital sodium (initial dose: 50 mg/kg ip). Adequate anesthesia depth was monitored frequently by checking the pupil size and flexion reflex to paw pinch. Supplementary injections (10 mg/kg iv) were administered when necessary. The trachea, common carotid artery, and external jugular vein were cannulated. The tibial nerve was isolated from the surrounding tissues and exposed from the calf to the popliteal fossa in the left leg. Subsequently, the tibial nerve was transected just distal to its branching point at the medial and lateral gastrocnemius muscles, with the branches to the medial gastrocnemius (MG nerve) and lateral gastrocnemius kept intact. For recording and stimulation, the MG nerve was used. For the experiments evoking primary afferent depolarization, the nerve projecting to the posterior biceps semitendonitis (PBST) muscle was isolated as well as the MG nerve. The lumbar spinal cord was exposed by performing a laminectomy (from the 12th thoracic vertebra to the 2nd lumbar vertebra), and the lumbosacral spine and tail were immobilized in a metal frame using clamps.

To maintain better circulation, both hind limbs were raised and also fixed to the metal frame. For the experiment involving PBST nerve stimulation, the hind limbs were not fixed to prevent potential confound by its conditioning effect on the PAD of the MG nerve. The skin flaps and muscle surrounding the exposed tissues were raised and tied to form a pool, which was filled with mineral oil to protect the exposed cord or nerve. The exposed MG nerve was mounted on an Ag-AgCl bipolar hook electrode. For the experiment evoking PAD, the PBST nerve was mounted on another hook electrode.

Body temperature and arterial blood pressure were maintained within their appropriate physiological ranges. After the surgical procedure, gallamine triethiodide (20 mg iv) was administered to achieve neuromuscular blockade, and the rats were artificially ventilated.

Recording procedures.

First, electrical stimulation was applied to the MG nerve, and the target segments of the lumbar spinal cord (L4–L6) were identified in each animal. To do so, cord dorsum potentials were detected using a silver ball electrode that moved over the rostrocaudal axis of the dorsal root entry zone. L4–L6 ventral or dorsal roots were then transected for the experiment such that pure evoked antidromic volleys (Fig. 1A) or orthodromic volleys (Fig. 1B), respectively, could be recorded from the MG nerve following intraspinal paired microstimulation.

Fig. 1.

Recording of antidromic and orthodromic volleys in peripheral nerves. A: preparation used to record antidromic volleys. All ventral roots within the segments that exhibited responses evoked by the MG nerve were transected. When stimuli are applied within the spinal cord, all volleys recorded in the periphery can be identified as antidromic volleys through the dorsal root. ISI, interstimulus interval; Stim, timing of the stimulation; ADVs, antidromic volleys. Paired stimuli (0.1-ms single pulses, 10–80 μA) were delivered with an ISI between 2 and 90 ms. B: preparation used to record orthodromic volleys. All dorsal roots within the segments that exhibited responses evoked by the MG nerve were transected, whereas the ventral roots were left intact. All volleys recorded in the periphery can be identified as orthodromic volleys through motor axons. ODVs, orthodromic volleys. C: measurement of the onset latency and amplitude of the volleys induced by the first and second pulses.

In experiments in which the ventral root (19 recorded points) or dorsal root (23 recorded points) was transected, a tungsten microelectrode (tip exposure 30–50 μm, 1 MΩ; Alpha Omega, Alpharetta, GA) was placed in the intermediate nucleus (mean depth from the cord surface: 1,695 ± 809 μm; Fig. 1A) or ventral horn (mean depth: 2,331 ± 686 μm; Fig. 1B) of the spinal cord by monitoring field potentials evoked by stimulation of the MG nerve. Once the intraspinal site in each penetration track that exhibited the largest field potential was identified, the same microelectrode was used to deliver intraspinal paired microstimulation, aiming to activate the afferent terminals (Fig. 1A) or motoneuron pool (Fig. 1B) of the MG nerve. The location of the intraspinal paired microstimulation within the track was further adjusted by determining where the volleys were evoked at the lowest stimulus intensity. For each stimulation point (unique to each penetration), a paired stimulation pulse (pulse width: 0.1 ms; strength: 2–5 times the threshold value, ∼10–100 μA) was delivered repeatedly (10 times) with an interval of 1 s. The interval between the two pulses was changed from 10 to 90 ms using 10-ms steps, and from 2 to 10 ms using 1-ms steps. In some cases (10 recorded points), these experiments were repeated in root-intact animals, that is, without transection of the ventral or dorsal root.

To evaluate if the antidromic or orthodromic volleys identified using the aforementioned method would exhibit primary afferent depolarization or hyperpolarization, the following study was performed. In these experiments, the characteristics of the evoked volleys at different depth within each penetration track were compared using borosilicate glass microelectrodes (tip diameter: 1.0–2.5 μm; impedance: 0.5–1.5 MΩ) filled with 2 M potassium citrate (pH 7.4). At each recording point (200- or 300-μm step from the cord surface), the following were recorded: 1) local field potentials evoked in the spinal cord by single-pulse stimulation to the MG nerve (2–5 times the threshold, 20–50 μA; pulse width; 0.1 ms); 2) volleys at the MG nerve evoked by single-pulse intraspinal microstimulation; 3) volleys at the MG nerve evoked by intraspinal paired microstimulation [interstimulus interval (ISI): 2, 3, and 4 ms]; and 4) volleys at the MG nerve evoked by single-pulse intraspinal microstimulation accompanied by preceding conditioning stimulation of the PBST nerve (excitability testing).

For paired-pulse stimulation, two stimulus pulses (pulse width: 0.1 ms; strength: 1–3 times the threshold value, ∼5–50 μA) were applied repetitively, 60 times for each interpulse interval (2, 3, and 4 ms). This sequence was repeated for each intraspinal point.

The excitability testing was performed using the following stimulation sequences. First, four pulses (2–3 times the threshold, 25–125 μA; pulse width: 0.1 ms; interval: 2 ms) were delivered as a conditioning stimulus to the PBST nerve. Subsequently, with a various delay (12, 17, 22, 27, 32, 37, and 42 ms), intraspinal microstimulation (single pulse; pulse width: 0.1 ms; strength: 1–3 times the threshold value, ∼5–50 μA) was delivered as a test stimulus. This sequence was repeated 60 times for each condition-test interval and averaged. To compare the effect of conditioning stimulation on the PBST nerve, a single test stimulus without preceding conditioning pulses was applied between each conditioning test sequence (n = 60).

The volleys at the MG nerve and the field potentials in the spinal cord were amplified (1,000× or 20×), bandpass filtered (15 Hz to 3 kHz), and digitized at 10 or 44 kHz for subsequent analysis.

Data analysis.

Volleys recorded from the MG nerve were averaged over 10 trials for each ISI, and the size and latency of the two volleys evoked by the first and second stimuli, respectively, were measured. The size was measured as the peak-to-peak amplitude, and the latency was measured from the onset of the first or second stimulus to the first peak of the corresponding volley (Fig. 1C). The amplitudes and latencies of responses to the second stimulus were then compared with those to the first stimulus. Data from the second amplitude (percentage of first amplitude) and second latency (percentage of first latency) were separated into antidromic volleys and orthodromic volleys at different ISIs. Receiver operating characteristic (ROC) analysis (Fawcett 2006) was used to calculate a cutoff value to find a point to dissociate antidromic and orthodromic volleys in the second amplitude and latency values.

Histological procedures.

Near the end of the experiments, small electrolytic lesions were created in the spinal cord by passing a 30-μA direct current through the stimulating electrode for 30 s. At the end of each experiment, animals were euthanized by administering an overdose of pentobarbital sodium (50 mg/kg iv) and perfused with 10% formalin. After fixation, the spinal cord was removed and immersed in graded sucrose solutions (final strength: 30%). After the immersion, the spinal cord was cut into 50-μm sections on a freezing microtome and stained with cresyl violet. The histological slices were photographed, and the positions of the lesions were detected.

Statistical analysis.

To test the significance of the modulation of the second response compared with the first response, one-sample t-tests (vs. 100%) were used. Unpaired Student's t-tests were used to determine differences in the amplitude and latency of the second response at specific intervals of elicitation between antidromic and orthodromic volleys. All data are presented as means ± SD. To evaluate the effect of the conditioning stimuli on the PBST nerve, one-sample t-tests (vs. 100%) were performed for each depth. The criterion for accepting statistical significance was P < 0.05.

RESULTS

Characterization of antidromic and orthodromic volleys recorded in root-transected animals.

On average, we were able to record 3 tracks for each rat, and a total of 42 volleys were analyzed: 23 antidromic volleys in the dorsal root-sectioned rats and 19 orthodromic volleys in the ventral root-sectioned rats. Typical pure antidromic volleys and orthodromic volleys evoked by intraspinal paired microstimulation in peripheral nerves are shown in Fig. 2, A and B, respectively. In these examples, the amplitude of the antidromic volleys (Fig. 2A) evoked by the second stimulus was slightly facilitated at ISIs from 2 to 10 ms compared with those evoked by the first stimulus (2 ms: 106.9%; 4 ms: 108.3%; 6 ms: 102.4%; 8 ms: 103.3%; 10 ms: 108.5%). By contrast, the orthodromic volleys (Fig. 2B) evoked by the second stimulus were depressed (71.9%) at an ISI of 2 ms, with slight facilitations at ISIs of 4 to 10 ms. The second latencies in antidromic volleys were equivalent to the first latencies (100%) at ISIs of 4 to 10 ms and were prolonged only at the 2-ms ISI (105%). By contrast, the second latency in orthodromic volleys was prolonged at ISIs of 2 ms (115%) and 4 ms (105%).

Fig. 2.

Examples of pure antidromic volleys (A) and orthodromic volleys (B) in the MG nerve evoked by paired intraspinal microstimuli. Each waveform shows the average of 10 single sweeps. Volleys induced by paired-pulse stimulation with 6 different intervals are shown for each animal (A and B). Note that the size of antidromic volleys evoked by the second stimulus is relatively similar to those evoked by the first stimulus at each ISI (A), but the size of the orthodromic volley to the second stimulus is decreased at an ISI of 2 ms (B). The spinal cord schemes in the insets indicate the intraspinal microstimulation site (x, stimulation site).

Changes in the second amplitude (percentage of first amplitude) of the antidromic volleys (n = 23) and that of orthodromic volleys (n = 19) for different ISIs are summarized in Fig. 3. The changes in amplitude of the second antidromic and orthodromic volleys from 2 to 90 ms are shown in Fig. 3, A and B, respectively. Figure 3C shows data from Fig. 3, A and B, expanded for ISIs from 2 to 10 ms. At ISIs from 3 to 90 ms, the amplitudes of the second response were unchanged or increased compared with those of the first response in both antidromic and orthodromic volleys (Fig. 3, A and B). However, at the 2-ms ISI, the size of the second orthodromic volley was significantly depressed (70.0 ± 21.9%, P < 0.01), but that of the second antidromic volley did not change (100.9 ± 13.3%, P = 0.72), as shown in Fig. 3C. The ratio of the second amplitude to the first amplitude was significantly different between antidromic and orthodromic volleys (100.9 ± 13.3% vs. 70.0 ± 21.9%, P < 0.01) at an ISI of 2 ms; no differences were observed for the remaining ISIs.

Fig. 3.

Size of the responses evoked by the second pulse at different interstimulus intervals. A and B: size of the second response (A, antidromic volley; B, orthodromic volley) induced by paired stimuli is plotted against the ISI from 2 to 90 ms. C: the same data as in A and B, but the horizontal scale is expanded for ISIs from 2 to 10 ms. Values are means ± SD. Note that the size of the second response in both antidromic volleys (A) and orthodromic volleys (B) is not decreased compared with the size of the first response at ISIs from 3 to 90 ms. At an ISI of 2 ms, the size of the second response in the orthodromic volleys is significantly decreased (P < 0.01), whereas that for the second response in the antidromic volleys is not decreased (C). *P < 0.05.

Changes in the second latency (percentage of first latency) of antidromic (n = 23) and orthodromic volleys (n = 19) for different ISIs are summarized in Fig. 4. The latency of the second antidromic volley was unchanged from that of the first antidromic volley at ISIs of 4 to 90 ms but was significantly longer at ISIs of 2 ms (106.5 ± 5.9%, P < 0.01) and 3 ms (102.9 ± 3.1%, P < 0.01; Fig. 4, A and C). By contrast, the peak latency of the second orthodromic volley was significantly longer than that of the first orthodromic volley at ISIs of 2 ms (110.8 ± 7.1%, P < 0.01), 3 ms (104.8 ± 3.0%, P < 0.01), and 4 ms (102.2 ± 2.5%, P < 0.01; Fig. 4, B and C). The ratio of the second peak latency to the first peak latency significantly differed between antidromic and orthodromic volleys at ISIs of 2 to 4 ms (P values <0.05).

Fig. 4.

Latency of the responses evoked by the second pulse using different interstimulus intervals. A and B: peak latency of the second response (A, antidromic volley; B, orthodromic volley) induced by paired stimuli is plotted against ISIs from 2 to 90 ms. The same format is used as in Fig. 3. Note that the peak latency of the second response in the antidromic volleys is delayed beginning at an ISI of 3 ms (A). By contrast, the peak latency of the second response in the orthodromic volleys is delayed beginning at an ISI of 4 ms (B). At an ISI of 2–3 ms, the peak latency in the orthodromic volleys is significantly later than that in the antidromic volleys (C). *P < 0.05.

Classification of volleys in sensory and motor fibers in an intact preparation.

As shown in Figs. 3 and 4, the difference between antidromic and orthodromic volleys was most evident in their second volley at the shortest ISI, that is, at 2 ms; the second orthodromic volley was smaller and later than the second antidromic volley. We postulated that the second response evoked at an ISI of 2 ms could be used to dissociate antidromic volleys in the nerve-intact animals. Therefore, we compared the distributions of the amplitude and latency of the second antidromic and orthodromic volleys. As shown in Fig. 5A, the relative amplitude (left) of the antidromic volley evoked by the second pulse was distributed between 90% and 120% of that evoked by the first pulse (20/23), and the majority of the orthodromic volleys by the second pulse were distributed from 20% to 80% (11/19) of that by the first pulse. However, the latency of the second response (right) was mostly distributed from 100% to 120% of that of the first response, irrespective of whether it was an antidromic or orthodromic volley. Therefore, although the distributions of the antidromic and orthodromic volleys evoked by the second stimulus were overlapped both in their amplitude and latency, the extent of the overlap was much smaller for the amplitude measurement. These results led us to propose that the amplitude of the second response at ISI of 2 ms could be used to dissociate antidromic volleys, which are further examined in Fig. 5B.

Fig. 5.

Discrimination of antidromic volleys from orthodromic volleys using paired intraspinal microstimuli. A: distributions of the amplitude (left) and latency (right) of the second response relative to the first response at an ISI of 2 ms. Red bars represent antidromic volley, and blue bars represent orthodromic volley data. Note that there are significant differences between antidromic and orthodromic volleys in both amplitude and latency. B: ROC curves for amplitude (left) and latency (right) of the second response to paired intraspinal microstimuli at an ISI of 2 ms. Dotted black lines correspond to the expected chance prediction. The best cutoff value is obtained at the shortest distance from the point (0, 1), the most ideal point, represented by dotted red lines. For amplitude, the cutoff value is 95.3% of the first amplitude (accuracy of prediction is 88.1%), and for latency, the cutoff value is 108.3% of the first latency (accuracy is 66.7%). C: application of the proposed criterion for nerves with intact dorsal and ventral roots. Five putative antidromic volleys (red) and five putative orthodromic volleys (blue) are dissociated using the criterion of 95.3%.

We plotted ROC curves in Fig. 5B to assess the predictive power of the second amplitude (left) and the second latency (right). The area under the curve, which indicates the probability of assignment, was 0.91 for the amplitude and 0.69 for the latency, suggesting that amplitude was the better dissociation index. The maximum correct dissociation ratio (88.1%) was achieved with an ISI of 2 ms and with a cutoff value at the second amplitude of 95.3% of the first amplitude. By contrast, a comparable dissociation ratio was not achieved using any latency as the threshold (maximum = 66.7% at 108.3%). Therefore, we concluded that the best dissociation of antidromic and orthodromic volleys was achieved when the amplitude of the second response exceeded 93.5% of that for the first response. Using this criterion in nerve-intact animals (10 recorded points; Fig. 5C), we demonstrated that half of the volleys could be identified as antidromic (red) and the other half as orthodromic (blue).

Excitability testing in nerve-intact animals.

Thus we found that the amplitude of the second response induced by intraspinal paired microstimulation at an ISI of 2 ms could be used as a criterion to dissociate antidromic from orthodromic volleys: an amplitude of the second response comparable to or larger than 95.3% of that for the first response identified antidromic volleys in the muscle's nerve. Applying this method along with the excitability testing of muscle afferents for evaluating PAD (Seki et al. 2009; Wall 1958) in the nerve-intact animal could be advantageous because it would dissociate the antidromic volleys in the muscle afferent (target for excitability testing) from the orthodromic volleys in the motor axons (confounds for excitability testing). To demonstrate this point, we examined the effect of conditioning stimuli applied to another muscle nerve (PBST) before the intraspinal microstimulation evoking antidromic or orthodromic volleys. It is well established that primary afferent depolarization is evoked by the incoming volleys from other muscle afferents (Eccles et al. 1962a). More specifically, the volleys from the PBST nerves would evoke PAD at the MG afferents (Jiménez et al. 1988). Our results are summarized in Fig. 6.

Fig. 6.

Relationship between responses to paired intraspinal microstimuli and depth of recording sites or the PBST conditioning effect in animals with intact muscle nerves. A, left: spinal field potentials evoked by stimulating the MG nerve. The intensity of nerve stimulation was constant (20 μA). Middle, volleys in the MG nerve evoked by intraspinal microstimulation. The intensity of intraspinal microstimulation is shown [in both μA and threshold current (T)]. Threshold current at each depth is also shown (right). B: examples of responses of the intact MG nerve evoked by paired intraspinal microstimulation at two depths. Note that at 800 μm, the amplitude of the second response is increased relative to that of the first response (filled arrow); however, at 2,000 μm, it is decreased (open arrow). C: examples of responses in the intact MG nerve evoked by a single intraspinal microstimulation after PBST conditioning. Recording sites are the same as those in B. Intervals between the first PBST stimulation and the single intraspinal microstimulation were 12, 17, 22, 27, 32, 37, and 42 ms. Magenta-colored traces indicate the control response (without PBST stimulation), and black traces indicate conditioned responses (with PBST stimulation). Note that at 800 μm black traces always show larger responses than magenta traces; however, at 2,000 μm, no difference is observed. For A–C, each waveform shows the average of 60 single sweeps. D: summary of the results of the PBST conditioning effect and the paired stimulation effect at each depth and each ISI. Triangles and dots indicate averaged values of all ISIs of PBST conditioning for each depth. Depth of each intraspinal site is labeled by different colors. A diamond indicates a significant difference with a value of 100% (P < 0.05, t-test), and a dot indicates no significant difference. E: average values for the PBST conditioning effect (left) and paired stimulation effect (right) are compared between the dorsal (surface to 1,200 μm) and ventral portions (1,400 to 2,200 μm) of the spinal cord. *P < 0.05; **P < 0.01 indicate significant difference between dorsal and ventral portions (t-test).

Spinal field potentials evoked at different depths (using a 200-μm step) by MG nerve stimulation [20 μA (1.4 threshold current)] are shown in Fig. 6A, left. The negative peak at 1.6 ms (dashed line: presumed group I field potential from MG afferent) at a superficial depth reversed to become positive at a depth of 1,600 to 1,800 μm. In addition, earlier small positive peaks (presumed antidromic field potential of MG motoneurons) changed to larger negative peaks at a depth of 1,200 to 1,600 μm, suggesting that the motoneuron pool of the MG muscle existed below a depth of 1,200 μm in this track. To each intraspinal point, then, we applied intraspinal microstimulation to evoke volleys at the MG nerve (Fig. 6A, right). The threshold current required to evoke these responses was also plotted as a function of depth (Fig. 6A, right). The threshold current was increased at depths of about 600 to 1,200 μm and began decreasing at 1,400 μm (8 μA) before reaching a nadir at 1,600 μm (2 μA). This result suggested that the motoneuron pool existed below a depth of 1,400 μm. Together, these results indicated that for this electrode track it was likely that the MG volley induced by intraspinal stimulation to a depth of 1,400 μm or shallower was likely antidromic, whereas that induced at a deeper position was likely orthodromic.

The effect of paired-pulse stimulation and that of the conditioning stimuli on the PBST in the same intraspinal locations as those in Fig. 6A are shown in Fig. 6, B and C. Representative results of paired pulse stimulation (Fig. 6B) and conditioning stimuli (Fig. 6C) applied to shallower (800 μm) and deeper (2,000 μm) areas are shown as examples. At an ISI of 2 ms (Fig. 6B), the amplitude of the second volley was markedly increased (161.3% of the first volley) for the volley induced at a depth of 800 μm (see filled arrow in Fig. 6B, left), whereas it was decreased (85.7% of the first volley) for that induced at 2,000 μm (see open arrow in Fig. 6B, right). According to the criterion proposed above, the former was categorized as an antidromic volley and the latter as an orthodromic volley. This result was consistent with the characteristics of the depth profile for this recording track (Fig. 6A), because a depth of 800 μm was deemed to be dorsal to the motoneuron pool, whereas that of 2,000 μm was considered the ventral portion of the motoneuron pool of the MG. Therefore, this result also validated our proposed criterion.

The effect of stimulating the PBST nerve on the MG volleys induced from these intraspinal points (excitability testing) suggested that the volley induced at a depth of 800 μm was facilitated by the preceding PBST stimuli (Fig. 6C, top, and Fig. 6D, red), especially at an ISI of 22 ms (287.3% of control). By contrast, no dominant modulation was observed at the volley induced at 2,000 μm (Fig. 6C, bottom, and Fig. 6D, sky blue). These results demonstrate that the intraspinal terminal of the MG afferent generating an antidromic volley in the MG nerve was depolarized by another muscle afferent (PAD) (Jiménez et al. 1988). By contrast, the volley categorized as orthodromic showed little sign of PAD, as expected.

The same recordings as exemplified in Fig. 6, B and C, were performed at all intraspinal sites in this track (Fig. 6A). The contrasting results between the volleys induced at superficial and deeper locations were well reproduced (Fig. 6D). At each location from the surface to 1,200 μm (yellow to reddish purple), we found significant conditioning effects by stimulating the PBST nerve (diamonds in Fig. 6D; P values <0.05). At deeper recording points (purple to blue), on the other hand, the conditioning effect was not significant (dots; P values >0.05). As for the paired-pulse response (Fig. 6D, right), second volleys with amplitudes larger than 95.3% of the first volleys were found in 6 of 7 points in the shallower positions (depth <1,400 μm), in contrast to 2 of 5 points in the deeper positions (depth ≥1,400 μm). These results are summarized in Fig. 6E. The conditioning effect of the PBST nerve (Fig. 6E, left) was significantly larger in the shallower positions (open bars), in which a larger second volley was also recorded (Fig. 6E, right). Overall, the results of this representative track demonstrated that volleys evoked from the shallower positions of spinal cord showed larger (≥95.3%) second amplitudes; that is, they were putative antidromic volleys and also exhibited PAD induced by another nerve. However, volleys evoked from deeper positions did not exhibit these larger second amplitudes; that is, they were putative orthodromic volleys and did not show significant PAD. These results strongly suggested that our criterion could dissociate the nature of the volleys. This conclusion was confirmed in two animals (4 electrode penetrations), as shown in Fig. 7. Putative antidromic volleys (second amplitude ≥95.3%) exhibited significant PAD (P values <0.05), whereas putative orthodromic volleys (second amplitude <95.3%) did not. Therefore, we can conclude that our proposed criterion for dissociating antidromic volleys is valuable for use with the excitability testing of muscle afferents that is needed for conducting experiments in a nerve-intact preparation.

Fig. 7.

Putative antidromic volley also exhibits primary afferent depolarization (PAD). Open bars indicate putative antidromic volleys, that is, averaged data whose second volleys evoked by paired-pulse intraspinal microstimulation are 95.3% above those of the first amplitude at ISI of 2 ms. Filled bars indicate putative orthodromic volleys, that is, second responses below 95.3% of the first response. *P < 0.05 indicate significant difference from 100% (t-test). At ISIs of 17, 22, 27, and 42 ms, differences between putative antidromic and putative orthodromic volleys are significant (P < 0.05, t-test).

However, one caution should be applied when the proposed method is used to identify antidromic volleys induced from deeper in the ventral horn of spinal cord (Fig. 8). In the example shown, we stimulated the bottom of ventral gray matter (depth: 2,400 μm) and the ventral funiculus (3,000 μm) in the same recording track using the same recording protocol as that used for data in Fig. 6, B–E. At both sites, no PAD was observed by stimulating the PBST nerve (open bars). This result was not surprising because intraspinal microstimulation to these sites should mainly activate the motor neuron or its axon, and thus both should be categorized as orthodromic volleys. However, the criterion for antidromic volleys (95.3% or more of the first response in paired stimuli) incorrectly categorized the volley induced from the ventral funiculus as antidromic (97.7% of the first response), although the volleys induced from the motoneuron pool were appropriately categorized as orthodromic (77.3% of first response). These results suggested that excitability testing may not accurately evaluate PAD when it is applied to the bottom of the ventral horn in nerve-intact animals, because the proposed criterion may not accurately dissociate antidromic volleys induced from this area of the spinal cord.

Fig. 8.

Characteristics of volleys evoked deeper in the ventral horn. A: 2 intraspinal sites (2,400 and 3,000 μm from the surface) deep in the ventral horn (black dots) overlaid with a cross-sectional image corresponding to the stimulated site. B: characterization of volleys of the MG nerve evoked from the 2 ventral horn sites. Near the motor nuclei (2,400 μm), paired stimuli reduce the amplitude of the second response, but no change is observed following PBST conditioning stimulation. In the white matter (3,000 μm), the effect of paired stimulation exceeds the criterion (95.3%), whereas PBST conditioning stimulation does not alter the amplitude.

DISCUSSION

We have demonstrated that a short-ISI, paired stimulation protocol could be used to classify volleys in peripheral muscle nerves as either antidromic or orthodromic. At an ISI of 2 ms, the amplitude of the response to the second stimulation was decreased in orthodromic volleys of dorsal root-sectioned rats but unchanged or increased in antidromic volleys of ventral root-sectioned rats (Fig. 3C). We found a criterion of 95.3% of the second response amplitude relative to the first response could dissociate the antidromic volleys at a selectivity of 88.1% (Fig. 5B). Volleys identified as antidromic using this method showed primary afferent depolarization that was induced by conditioning stimuli applied to other muscle afferents (Fig. 7). We concluded that excitability testing accompanied by a paired-pulse stimulus protocol is a promising tool for studying presynaptic inhibition of somatosensory afferents in normal behaving rats.

Effect of paired-pulse stimulation on the properties of the second volley.

In the present experiments, the orthodromic volleys in peripheral nerves of animals with transected dorsal roots were presumably evoked by stimulation to motor nuclei and cell bodies of motoneurons. The neuronal cell body becomes momentarily unexcitable, a refractory period cause by afterhyperpolarization (AHP), following activation by a stimulus to subsequently generate an action potential. Furthermore, the AHP duration is long-lasting in motoneurons, 30–116 ms in rats (Bakels and Kernell 1993a, 1993b) or 50–200 ms in cats (Botterman and Cope 1986; Eccles et al. 1958). By contrast, the discharge rate of Ia afferents in the cat soleus muscle increases linearly with the frequency of vibration, up to 500 Hz (Brown et al. 1967), suggesting negligible AHP in primary afferents. These reports predict that the second orthodromic volley following paired stimuli at very short ISIs can exhibit a reduction in amplitude. In fact, the present study found that the majority of second orthodromic volleys displayed smaller amplitudes, but the majority of second antidromic volleys did not when the ISI was 2 ms (Fig. 5). Therefore, the distinctly different responses observed to the second stimuli in antidromic and orthodromic volleys could be ascribed, at least in part, to the different refractory periods between afferent terminals and motoneurons to the second stimulus. Suppression of the second volleys could also be achieved if the first pulse activated last-order inhibitory interneurons (for review, see Jankowska 1992) or their axons to MG motoneurons, and inhibitory postsynaptic potentials from these interneurons could increase the threshold for the second stimulus to elicit motoneuron action potentials.

By contrast, antidromic volleys in peripheral nerves of animals with transected ventral roots were presumably evoked by stimulation of afferent terminals, because we applied stimulation to the intermediate zone. The unchanged or slightly increased amplitude (Fig. 5) of the second response may represent negligible AHP in primary afferents, or it also may be explained by a mechanism similar to that for the dorsal root reflex (Brooks and Koizumi 1956; Eccles et al. 1961). Namely, the first stimulation may activate the GABAergic interneurons projecting to the terminals of the MG afferent (PAD interneuron), either directly or indirectly, and it may decrease the threshold current to evoke the second response by temporal summation.

Criterion for identifying antidromic volleys in nerve-intact animals.

In this study, we proposed a new criterion that could be used to dissociate antidromic from orthodromic volleys in excitability testing using nerve-intact animals. Namely, volleys evoked by paired intraspinal microstimulation at a short ISI are antidromic when the amplitude of second response exceeds 95.3% of the first response. As shown in Fig. 5, this proposed criterion was able to successfully dissociate antidromic from orthodromic volleys with a high dissociation ratio (88.1%). As shown in Fig. 6, putative antidromic volleys could be attributed to the dorsal and intermediate portion of spinal cord, suggesting involvement of both Ia and Ib afferents. As shown in Fig. 7, the terminals of the primary afferents that were involved in generating the putative antidromic volleys showed PAD induced from other muscle nerves. These results support our proposal that the size of the second response evoked by intraspinal paired microstimuli is useful for identifying antidromic volleys in intact muscle nerves. This criterion could be especially useful for future studies by applying it in parallel with excitability testing for investigating presynaptic inhibition.

However, the use of this criterion is limited when examining the ventral portion of spinal cord. As shown in Fig. 8, obvious orthodromic volleys induced by stimulation of the axon, not the cell body, of the motoneuron were erroneously classified as antidromic volleys using our criterion. During the excitability testing at variable depths in the spinal cord, the disadvantage of this limitation can be minimized if the threshold current to evoke a single volley is also taken into account. As shown in Fig. 6, the threshold current to evoke a single volley in an intact animal was lowest in the ventral horn, probably representing the lowest threshold to activate the initial segment of the motoneuron (Gustafsson and Jankowska 1976). In the representative penetration shown in Fig. 6, the depth was ∼1,600 μm from the surface (threshold current was 2 μA). The erroneous classification was evident only at a depth of 2,200 μm (Fig. 6D, deep blue), where the amplitude of the second volley was not smaller than that of the first volley at an ISI of 2 ms (126%). In addition, the threshold current to induce a single volley was gradually increased below a depth of 1,600 μm. Therefore, we suggest that the proposed criterion is applicable up to the intraspinal depth at which the lowest threshold for inducing a single volley is observed, and we do not recommend applying our criterion to deeper areas.

Potential application to other experimental preparations.

We developed this method to estimate the effect of presynaptic inhibition on muscle afferent control in normal animals without having to transect their ventral spinal roots. This method can be applied to behaving rodents and thus will advance the understanding of presynaptic inhibition during volitional motor control. A recent study described the process of generating a GABAergic presynaptic inhibitory circuit by using synaptic expression of the GABA synthetic enzyme glutamic acid decarboxylase (GAD65) in the spinal cord of developing mice (Betley et al. 2009). They found that the connectivity and synaptic features of GABAergic interneurons were selectively directed by their sensory terminal targets. Subsequent studies using a mouse model (Fink et al. 2014) also demonstrated that presynaptic inhibition appears to be necessary to stabilize motor output, as determined by observing oscillatory limb movement in mice without these GABAergic interneurons. However, further evaluation of these arguments is needed using excitability testing on an epoch-by-epoch basis during various behaviors. Intraspinal microstimulation of the spinal cord is now feasible in freely moving rodents (Fuentes et al. 2009; Minev et al. 2015), and the technology for manufacturing miniature cuff electrodes has recently been markedly advanced (Ordonez et al. 2014). Therefore, our method can be practically applied in behaving rodents to characterize developmental mechanisms as well as functional roles of presynaptic inhibition.

In monkeys, presynaptic inhibition of cutaneous afferents has been studied during wrist movement (Seki et al. 2003, 2009); however, presynaptic inhibition of muscle afferents has not been examined. Our method, which discriminates antidromic from orthodromic volleys, may allow the investigation of how presynaptic inhibition of muscle afferent input is modulated during voluntary movement. Specifically, our method will make it possible to examine presynaptic inhibition of Ia afferent terminals that monosynaptically project to motoneurons in awake, behaving animals. Ia presynaptic inhibition has been estimated in humans by assessing changes in the effectiveness of the Ia reflex loop in leg muscles during various locomotor activities (Capaday and Stein 1986; Faist et al. 1996; Zehr and Stein 1999). However, these noninvasive approaches used in human studies do not permit a distinction between changes in postsynaptic excitability and presynaptic inhibition from the afferents to the interneurons or motoneurons. By applying the proposed approach to awake, behaving monkeys, further understanding of the function of presynaptic inhibition on volitional movement will be ensured.

GRANTS

This work was partly supported by Grants-in-Aid 18020030 and 18047027 for Scientific Research on Priority Areas “Mobilligence” (Grant 18020030) and “System Study on Higher-Order Brain Function” (Grant 18047027) and innovative area “Understanding Brain Plasticity on Body Representations to Promote Their Adaptive Functions” (Grant 26120003) from the Ministry of Education, Culture, Sports, Science and Technology of Japan (to K. Seki) and the Japan Science and Technology Agency Precursory Research for Embryonic Science and Technology program (to K. Seki).

DISCLOSURES

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

AUTHOR CONTRIBUTIONS

S.T., G.K., J.C., and K.S. performed experiments; S.T. and G.K. analyzed data; S.T., G.K., and K.S. interpreted results of experiments; S.T. and G.K. prepared figures; S.T. and G.K. drafted manuscript; S.T. and K.S. edited and revised manuscript; S.T., G.K., J.C., and K.S. approved final version of manuscript; K.S. conceived and designed research.