Spinal Laminae I-II Neurons in Rat Recorded In Vivo in Whole Cell, Tight Seal Configuration: Properties and Opioid Responses

Alan R. Light and Helen H. Willcockson

Department of Cell and Molecular Physiology and Curriculum in Neurobiology, University of North Carolina, Chapel Hill, North Carolina 27599-7545


    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

Light, Alan R. and Helen H. Willcockson. Spinal Laminae I-II Neurons in Rat Recorded In Vivo in Whole Cell, Tight Seal Configuration: Properties and Opioid Responses. J. Neurophysiol. 82: 3316-3326, 1999. Using the in vivo whole cell recording procedure described previously, we recorded 73 neurons in laminae I and II in the lumbar spinal cord of the rat. Input impedances averaged 332 MOmega , which indicated that prior sharp electrode recordings contained a significant current shunt. Characterization of the adequate stimuli from the excitatory hindlimb receptive field indicated that 39 of 73 neurons were nociceptive, 6 were innocuous cooling cells, 20 responded maximally to brush, and 8 cells were not excited by stimulation of the skin of the hindlimb. The locations of 15 neurons were marked with biocytin. Nociceptive neurons were mostly found in lamina I and outer II, cooling cells in lamina I, and innocuous mechanoreceptive cells were mostly found in inner II or in the overlying white matter. The µ-opioid agonist [D-Ala2, N-Me-Phe4, Gly5-ol]-Enkephalin (DAMGO) hyperpolarized 7 of 19 tested neurons with a conductance increase. This hyperpolarization was reversed by naloxone in the neurons in which it was applied. DAMGO also decreased the frequency of spontaneous PSPs in 13 neurons, 7 of which were also hyperpolarized by DAMGO. Five of the seven hyperpolarized neurons were nociceptive, responding to both heat and mechanically noxious stimuli, whereas two responded to slow, innocuous brush. These results indicate that whole cell, tight seal recordings sample a similar population of lamina I and II neurons in the rat as those found with sharp electrode recordings in cat and monkey. They further indicate that DAMGO hyperpolarizes a subset of the nociceptive neurons that have input from both heat and mechanical nociceptors and that presynaptic DAMGO effects can be observed in nociceptive neurons that are not hyperpolarized by DAMGO.


    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

Laminae I and II (the marginal zone and substantia gelatinosa, respectively) of the spinal cord have been implicated as a primary integration center for pain processing and potent sites at which opioid analgesics act. These laminae receive input from both Adelta and C primary afferent nociceptors and from Adelta and C innocuous thermal receptors and innocuous mechanoreceptors (e.g., reviewed in Light 1992; Light and Perl 1979; Réthelyi et al. 1982; Sugiura et al. 1986, 1993).

The inputs that activate laminae I and II neurons and the anatomy of the dendrites and axons of these neurons have been well defined in the cat and the monkey (Bennett et al. 1980, 1981; Christensen and Perl 1970; Han et al. 1998; Jones et al. 1990; Kumazawa et al. 1975; Light and Kavookjian 1988; Light et al. 1979, 1993; Randic and Miletic 1978; Réthelyi et al. 1989). Thus neurons in lamina I and outer lamina II (IIo) are usually excited by noxious stimuli applied to the appropriate receptive field, by innocuous thermal stimuli, or by both types of inputs. Neurons in inner lamina II (IIi) are usually excited by innocuous mechanical stimuli. However, this differentiation has not been documented in the rat.

Only a few intracellular and extracellular studies have documented the input properties of neurons in lamina I and II in vivo in the rat with most recording only from lamina I. Furthermore, the results in the rat are inconsistent. Some investigators report neurons located above lamina I that respond only to innocuous mechanoreceptive stimulation. Some report neurons in and above lamina I that respond only to innocuous cooling. Others report mostly multireceptive neurons in lamina I and II and others describe mostly nociceptive specific neurons in lamina I and II (Hope et al. 1990; Hylden et al. 1989; McMahon and Wall 1985, 1988; McMahon et al. 1984; Mokha et al. 1987; Woolf and Fitzgerald 1983). In most cases neurons were not intracellularly labeled, complicating localization of the neurons. This is largely because of the difficulty of recording and labeling a population of the very small neurons in laminae I and II of the rat.

In the rat the biophysical properties of neurons in the superficial laminae and their response to opioids have been studied intensively in vitro (Baba et al. 1994; Glaum et al. 1994; Jeftinija 1988; Jeftinija and Urban 1994; Magnuson and Dickenson 1991; Miletic and Randic 1981; North and Yoshimura 1984; Rusin et al. 1993; Yoshimura and Jessell 1989; Yoshimura and Nishi 1993). These studies have documented both pre- and postsynaptic mechanisms for the effects of opioids on laminae I and II neurons. The postsynaptic hyperpolarization in response to opioids is caused by the activation of a G protein-coupled, inward-rectifying potassium channel. The observed presynaptic effect is a decrease in spontaneous excitatory postsynaptic potentials (EPSPs), presumably mediated by a decrease in free intracellular calcium in presynaptic terminals.

The relevance of the in vitro effects to clinically effective doses of opioids and the specificity of opioid inhibition for nociception are still questioned. Spinal neurons studied in vitro have no peripheral inputs and thus cannot be functionally identified as nociceptive using adequate stimuli applied to their "receptive fields." Both intracellular and extracellular recording experiments on functionally defined neurons in cats and monkeys have suggested that opioids produce both excitatory and inhibitory effects on nociceptive as well as nonnociceptive neurons (Craig and Hunsley 1991; Craig and Serrano 1994; Jones et al. 1990; Willcockson et al. 1986). However, in vitro experiments in rats indicate that opioids produce a predominantly inhibitory effect both pre- and postsynaptically (e.g., Glaum et al. 1994; Schneider et al. 1998).

To overcome some of these difficulties, we have developed a procedure in rat that relies on whole cell, tight-seal recordings of spinal neurons in laminae I and II. This technique allows recording in the whole cell mode for extended periods of time and allows for the recorded neuron to be labeled. Recently, similar recordings have been made in the kitten cortex (Nelson et al. 1994), the bat inferior colliculus (Covey et al. 1996), and the rat cortex (Moore and Nelson 1998), demonstrating the utility of this recording mode both for recording and for manipulating the internal environment of recorded neurons.

Our results demonstrate that the same functional types of neurons (defined by stimulation of the receptive field) are found in the rat in approximately the same proportions as in monkey (Kumazawa and Perl 1978; Light et al. 1979). The membrane and synaptic properties of these types of neurons recorded in vivo are similar to those found in vitro. Furthermore, the responses to opioids are similar to those reported in vitro with the additional finding that most of the affected neurons are nociceptive. Finally, the low noise of the patch pipettes and the high-input impedances of the recorded neurons allow for detailed analyses of both normal and opioid affected subthreshold synaptic inputs, which have not been resolved in vivo with sharp electrode recordings in the past. Preliminary reports of some of these findings have appeared (Light et al. 1997; Light and Willcockson 1996).


    METHODS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

The guidelines on ethical standards for the investigation of experimental pain in animals were strictly followed and the following experimental protocol was approved by the Institutional Animal Care and Use Committee of the University of North Carolina. Fifty adult female Long-Evans rats (Charles River, 225-350 g) were used in these studies. Long-Evans rats were used because their survival is more robust under our experimental conditions and they routinely maintained adequate physiology following unilateral pneumothorax, which was necessary to achieve adequate stability of the preparation. Bilateral pneumothorax further decreased the survival of the animals. In addition, we have been using this strain in behavioral experiments because they exhibit better performance on a variety of behavioral tasks (e.g., Vierck et al. 1995).

The rats initially were anesthetized deeply and maintained in this state with sodium pentobarbital and chloral hydrate (50 mg/kg and 125 mg/kg, respectively) delivered i.p. Catheters were inserted in the jugular vein and carotid artery for administering drugs and measurement of blood pressure, respectively. The rats were placed in a head holder and stabilized in a spinal frame with hip pins and vertebral clamps attached laterally. A laminectomy exposed ~7 mm of lumbar spinal cord. A small reservoir (~150 µL) was formed with dental impression material (CutterSil Light; Heraeus Kulzer, South Bend, IN) and filled with oxygenated, artificial cerebral spinal fluid (ACSF) containing (in mM) 120 NaCl, 2.5 KCl, 2.5 CaCl2, 1.5 MgSO4, 1.25 NaH2PO4 · H2O, 26 NaHCO3, and 10 glucose at 37°C (pH 7.35-7.45, 305-310 mOsm).

After a unilateral pneumothorax, the rat was paralyzed with pancuronium bromide (4-8 mg/kg) and respirated with 100% O2. To assess adequate depth of anesthesia the animal was monitored for respiratory CO2, rectal temperature, heart rate, and blood pressure and kept within normal physiological limits with supplemental doses of anesthetic (sodium pentobarbital, 4-8 mg/kg). In addition, paralysis was occasionally allowed to wear off to assess that the rats were areflexic. The dura over the lumbar spinal cord was carefully removed and the arachnoid between the dorsal roots was dissected away over a small section of the lateral spinal cord with care being taken not to compress the cord or damage small blood vessels. Initially in some experiments, needle electrodes were inserted into the receptive field locations to electrically activate primary afferents.

Microelectrodes

Whole cell, patch pipettes (N-51A custom borosilicate glass; Drummond Scientific, Broomall, PA) with a DC resistance of 6-8 MOmega were fabricated on a Model P-87 Flaming-Brown puller (Sutter Instrument Co., Novato, CA) and the tip was filled with an internal solution containing (in mM) 130 K-gluconate, 5 NaCl, 1 CaCl2, 1 MgCl2, 11 EGTA, 10 HEPES, 0.1 GTP, and 2 Mg-ATP (pH 7.30, 280-290 mOsm) (Schneider et al. 1998). Some electrodes were then backfilled with 0.1 to 1% biocytin (free base, FW = 372.5; Sigma, St. Louis, MO) dissolved in the internal solution (Horikawa and Armstrong 1988). Because biocytin labeling of unrecorded neurons could occur inadvertently if multiple attempts were made with a single electrode, and because we had observed some possible effects of biocytin on patch sealing and on opioid responsiveness in our in vitro studies, later experiments used a reduced concentration of biocytin (0.1%) in the recording pipette.

Recording procedure

Patch electrodes were lowered into the spinal cord via a hydraulic microdrive (Model 607-C; David Kopf Instruments, Tujunga, CA) attached to an electrode holder with a pressure port (Axon Instruments, Foster City, CA). Pulses of hyperpolarizing current (0.1 nA, 60 ms, 1 Hz) were used to continuously monitor series resistance. Recordings were made with an AxoClamp 2 electrometer (Axon Instruments) in bridge mode.

When approaching the surface of the cord, positive pressure was applied to the pipette to prevent plugging the tip of the electrode with pia or overlying white matter. This pressure was released following penetration of the cord surface. Seals were formed by initially reducing the pressure to atmospheric and applying gentle suction to the pipette. Neurons were recognized by negatively directed action potentials in response to electrical stimulation of the periphery, spontaneous action potentials, or simply by the increase in impedance observed when forming seals. Seals often formed on glial cells or on unidentified debris. Seals on nonneuronal elements were broken by applying positive pressure to the pipette. After an unsuccessful attempt to attain a neuron, electrodes were replaced to prevent leakage of biocytin into surrounding tissue and the subsequent undesired labeling of neuropil. Whereas neurons deeper in the dorsal horn could be "patched" in this same manner, we restricted our attempts to the first 500 µm to more thoroughly sample laminae I and II.

After acquiring a seal >1GOmega (usually 2-10 GOmega ), the membrane patch was ruptured by further gentle suction to the electrode, establishing a whole cell recording configuration and the pressure was immediately returned to atmospheric. Electrode signals were displayed on a storage oscilloscope and also digitized and saved on videotape for later analysis using Axotape and pClamp software (Axon Instruments). Using chart recordings at moderate speeds with high resolution, we measured ~5 consecutive conductance pulses (occasionally contamination by action potentials made it possible to obtain only 4 consecutive pulses) from before and during the ([D-Ala2, N-Me-Phe4, Gly5-ol]-Enkephalin (DAMGO) applications (see Fig. 6C). Measurements were made from the mid-baseline to the middle of the noise in the pulse. A t-test (P < 0.05) was used to determine significant changes in conductance during DAMGO application. Only units with significant conductance changes were classified as affected postsynaptically by DAMGO.

A neuron was characterized according to the type of natural stimuli that evoked action potentials (i.e., brush, pinch, cool, heat, or combinations of each) to allow comparisons with previous samples from our lab (Light 1992; Light et al. 1993, 1997) and others that have used extracellular recordings to characterize neurons (see INTRODUCTION and DISCUSSION for references). We often observed subliminal PSPs [EPSPs, inhibitory postsynaptic potentials (IPSPs), and slow potentials] from other inputs that did not activate action potentials. These PSPs appeared to be variable in their appearance, and could be affected by a number of factors. However a thorough characterization of these inputs under various conditions is beyond the scope of this paper. We used our standard tests to determine whether the cell had specific innocuous mechanical input (responded only to very slowly moving stimuli), innocuous thermal inputs (<43°C for heat and >20°C for cooling, using a feedback controlled thermal stimulator), and nociceptive input (responded only to intense mechanical and/or intense thermal stimulation, >45°C for heat and <10°C for cooling). Timings of stimuli applications were marked on recordings with a foot switch. A strip chart recorder (TA-2000; Gould, Valley View, Ohio) was used to monitor long-term fluctuations in Vm and RN, the latter using 600 ms negative pulses (0.05 nA) through the pipette.

Drug application

In some experiments, the selective µ-opioid receptor agonist DAMGO, (free base, MW = 513.6; Sigma) was applied by placing 0.5 µl of a 1 µg/µl solution directly into the ASCF reservoir overlying the spinal cord (ASCF bath volume ~150 µl). The final concentration in the reservoir was estimated to be ~5 µM. Drugs could not be removed from the reservoir so only cumulative applications of drugs were performed. The opioid receptor antagonist, naloxone hydrochloride (FW = 363.8; Sigma) was applied in a similar fashion (0.5 µl of 0.1 µg/µl) for an estimated final concentration of ~0.5 µM. The drug application was later modified and applied to the reservoir by continuous flow similar to the in vitro studies of Schneider et al. (1998). Flow application had the advantage of allowing application of known concentrations of drugs that could be removed quickly. This allowed before and after drug comparisons and applications of several different drug concentrations. Because we could not determine the actual drug concentration at the membrane, which is affected by clearance of the drug via tissue dilution and the vascular uptake, the final concentration is not known. However, it was always less than the maximum applied concentration of 5 µM. In addition, because of the close proximity of the recorded cells to the spinal cord surface, the concentration at the membrane was probably relatively close to that in the bath.

Histochemistry

After the neurons were characterized, the animals were perfused through the heart with 0.01 M phosphate buffered saline (pH 7.4) followed by cold 4% paraformaldehyde and 10% sucrose in 0.1 M phosphate buffered saline (pH 7.4). The lumbar spinal cord was removed, postfixed (4% paraformaldehyde with 30% sucrose), and sectioned on a cryostat at 60 µm. Free-floating sections were collected, rinsed in 0.05 M tris-buffer (pH 7.6) with 0.3% Triton-X and 2.7% NaCl (TBS/TX buffer, pH 7.6), and pretreated with graded alcohols to eliminate red blood cells. The sections were rinsed with TBS/TX buffer and 2% normal goat serum, and incubated in avidin-biotin HRP complex (Vectastain Elite ABC Kit, PK-6100; Vector Laboratories, Burlingame, CA) for 1 h. After rinsing with TBS/TX, the tissue was incubated in diaminobenzidine (DAB, 0.2%) and nickel ammonium sulfate (0.14%) with 7 µl of 30% hydrogen peroxide. The sections were rinsed with TBS/TX, mounted, air-dried, and coverslipped with DPX. On examination, biocytin labeled neurons were revealed as densely stained, gray-black cell bodies with heavy to diffuse stained dendrites and axons. Cells were magnified (×400-1000), photographed, and reconstructed using a Nikon Optiphot microscope equipped with a drawing tube. The laminar locations were designated based on the relative density and orientation of myelinated fibers (Light 1992) and verified using dark field illumination.


    RESULTS
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

The data presented are from 73 neurons. Recording times varied from 10 to 90 min with half being >= 30 min. The input impedances ranged from 100 to 700 MOmega [332 ± 132 (SD) MOmega , n = 60]. The membrane time constants were 4-25 ms (13.1 ± 4.8 ms, n = 29). The resting membrane potentials were between -24 and -80 mV (-52 ± 12 ms, n = 65). The size of action potentials ranged 15-80 mV (52 ± 17 mV, n = 40). Small action potentials were not related to less negative resting membrane potentials, lower inputs impedances, or spontaneous action potentials, suggesting that they were not the result of injury. Furthermore, when action potentials were less than the membrane potential (not overshooting), the PSPs were often quite large (5-15 mV). The lack of signs of injury and the nonovershooting action potentials combined with anecdotal evidence from biocytin labeling suggested that these recordings were from dendrites at a distance from the action potential initiation site. Furthermore, these data indicate that the dendrites of some neurons recorded here did not back-propagate action potentials to the dendritic recording sites (see Johnston et al. 1996 for review of this topic).

Activating stimuli

Receptive fields were most often located on the lateral portion of the proximal hindlimb (see examples in Fig. 1A). Using natural stimuli, we classified neurons (n = 73) according to action potential firing. Thirty-nine neurons were nociceptive. More specifically, the nociceptive neurons were fairly evenly divided by response such that six responded only to noxious pinch; five to pinch and rapid cooling; eight to pinch, cold, and noxious heat; and seven to pinch, cold, and heat, but demonstrating some responses to innocuous brush as well. Five neurons responded to noxious heat and pinch; three to brush and pinch; and two to brush, pinch, and cooling, but not to heat. Three nociceptive neurons were not held long enough to complete their characterization in detail. Six neurons were innocuous cooling cells, 20 responded maximally to brush, however in 8 neurons we could not find excitatory receptive fields. Whereas this was tested in all cases, only one unit, held briefly and not characterized fully enough to be included in this sample, appeared to respond to innocuous warming. The types of input to these neurons are very similar to the percentages and types of neurons found in spinal laminae I and II in cats and monkeys (e.g., Jones et al. 1990; Light and Kavookjian 1988; Light et al. 1979; Réthelyi et al. 1989), and similar to the description of rat neurons recorded intracellularly (Woolf and Fitzgerald 1983). Examples of some of the responses of these neurons are shown in Figs. 2, 5, and 6. PSPs evoked by natural stimuli are shown in Fig. 7.



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Fig. 1. Location of hindlimb receptive fields and labeled neurons in the lumbar spinal cord. A: lateral aspect of the hindlimb shows representative receptive fields from 3 neurons responsive to brush, 2 to nociceptive stimuli, and 1 cooling cell. Neurons labeled with biocytin and characterized according to response to stimuli and application of DAMGO are located in the dorsal horn shown in B.



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Fig. 2. Low gain, DC patch-clamp recordings in vivo from the most common types of responsive neurons encountered in the substantia gelatinosa of the rat. Examples of slow brush (A), pinch (B), and innocuous cooling (ethyl chloride spray applied to brush and brushed on skin) (C) responsive neurons.

The high resolution of the recordings (caused by very high-input impedance and very low noise of patch-clamp electrodes) allowed us to observe PSPs evoked by stimulating various types of afferent input (see Figs. 2 and 7). These potentials varied in size from 0.5 mV to 8 mV, depending on the input impedance and intrinsic properties of the neuron. In many instances, both spontaneous and evoked PSPs appeared to have a unitary nature, i.e., a PSP of a constant size and shape that decayed exponentially appeared to sum with others of the same size to produce larger events (see Fig. 7). Similar, apparent unitary events have been observed in other patch clamp experiments in vitro (McQuiston and Colmers 1996) and have also been observed in the spinal cord (Bao et al. 1998). However, we interpret most if not all of our observations of unitary PSPs in vivo as representing summed PSPs from the activation of primary afferent axons or spinal interneurons (much like those observed in motoneurons with spike triggered averaging from Ia axons e.g., Mendell and Henneman 1971) unlike the "true" miniature events observed in in vitro studies.

Biocytin labeling

The locations of 15 recording sites were marked with biocytin (see Fig. 1B). The two neurons responding only to innocuous cooling were found in lamina I and in Lissauer's tract. The one well labeled cooling cell (Fig. 3E), may represent a pyramidal lamina I cell as suggested for cooling cells in other species (Han et al. 1998). Neurons responding to brush were found in lamina I, the overlying white matter and lamina IIi (Fig. 3, F and G, and 4). Innocuous mechanoreceptive neurons in lamina IIi were both islet and stalked cell types (Fig. 3G and 4). Neurons responding to noxious inputs were found in both lamina I and lamina IIo. Nociceptive lamina I neurons were either fusiform (Fig. 3B) or multipolar in shape (Fig. 3, C and D). Nociceptive lamina II neurons were either similar to islet cells (Fig. 3A) or to stalked cells. Examples of several of these neurons are reconstructed in the parasagittal plane and shown in Fig. 3. Photomicrographs of a labeled cell that responded to brush are seen in Fig. 4, A and B.



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Fig. 3. Drawing tube reconstructions from rat substantia gelatinosa cells labeled with biocytin. A: nociceptive islet cell located in outer lamina II, responded to noxious pinch, noxious heat, and noxious cooling and was hyperpolarized by DAMGO, recording shown in Fig. 5. B: nociceptive lamina I islet cell responded to noxious pinch. C: multipolar, triangular-shaped cell located in the outer SG responded to noxious pinch and was classified as nociceptive specific, nonresponsive to DAMGO. D: multipolar, pyramidal cell located in lamina I classified as nociceptive (responded to noxious pinch, noxious heat, and noxious cooling) with excitatory responses to DAMGO. E: pyramidal lamina I cell responsive to noxious cooling. F: nonnociceptive cell responding only to slow brush, classified as innocuous mechanoreceptive and hyperpolarized by DAMGO, located above lamina I. G: islet cell located in inner lamina II responded to brush. E and F: dashed line indicates lamina I-II border. Because of possible errors in parallax from the reconstruction of cells from multiple sections, laminar borders are not indicated for all neurons.



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Fig. 4. Photomicrographs (A and B) of an innocuous mechanoreceptive cell labeled with biocytin and located in inner lamina II. This stalked cell was responsive to brush and was hyperpolarized by the application of DAMGO. A darkfield image is shown in B to demonstrate the location of the cell near the inner-outer lamina II border. C: drawing tube reconstruction demonstrates extensive dendritic arborization into lamina III and axon collaterals into lamina IIo and III.

DAMGO effects

Fifteen of 19 neurons were affected by adding DAMGO to the recording reservoir (Table 1). In 7 of 19 tested neurons, DAMGO caused a hyperpolarization of 7.0-25 mV (11.9 ± 6.5 mV) accompanied by a significant (P < 0.05) conductance increase (7-38%, 23.3 ± 11.2%; see Figs. 5 and 6). The locations of five of these neurons are shown with filled symbols in Fig. 1. All but one of these neurons were found in lamina II. Naloxone was applied to four of these seven neurons. In all four, naloxone clearly reversed the hyperpolarization and conductance increases evoked by DAMGO.


                              
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Table 1. Effects of DAMGO on classified cells



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Fig. 5. Low gain DC recordings from the same neuron in Fig. 3A. Before DAMGO (A-G) indicates responses of the neuron before the application of DAMGO. Note brisk responses to noxious stimulation, including brush (A), pinch (C), noxious heat (E), and noxious cooling (G). During DAMGO, B-H demonstrates low gain DC recordings 70 s after the application of DAMGO (final concentration in ACSF pool ~5 µM). Responses to brush (B) increased whereas pinch (D) was reduced. Note the reduction in response to noxious heat (F) and cooling (H) such that no action potentials were observed. I: low-gain DC chart recording of the membrane potential from this same neuron. The neuron was classified as nociceptive specific and fired action potentials only to pinch, noxious heat, and noxious cooling. DAMGO was applied at arrow to the stationary ACSF bath over the exposed spinal cord (final concentration in bath ~5 µM). Downward deflections are 200 ms current pulses to test membrane conductance. Note the conductance increase as indicated by reduction in the size of these pulses following application of DAMGO (actual hyperpolarization 7 mV, conductance increase 14%, P < 0.03). Upward deflections are PSPs and truncated action potentials evoked by various forms of natural stimulation (indicated below by brackets). In the stationary bath, the DAMGO remained on the cell causing the cell to desensitize and thus showed no effect to the second application of DAMGO. The resting membrane potential before drug application was -58 mV.



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Fig. 6. In vivo whole cell recording from lamina II cell classified as nociceptive specific, firing action potentials only to pinch and heat. Upper chart record (A) shows membrane and action potential alterations evoked by indicated stimuli (scale as in B). The chart recording (B) shows the membrane potential following application of 5 µM DAMGO and 2 applications of 0.5 µM Naloxone to the artificial cerebral spinal fluid (ACSF) bath. Downward deflections are 200 ms current pulses to test membrane conductance. ; pulses which are shown enlarged in C. Five consecutive pulses are shown (C) for time periods before DAMGO, during DAMGO, and during Naloxone. Note the increase in conductance as indicated by the reduction in size of these pulses following the application of DAMGO (actual hyperpolarization 14.4 mV, conductance increase 33%, P < 0.002, and increases following first Naloxone. Increases in pulses during second Naloxone (B) cannot be seen because of action potential contamination. The upward deflections are PSPs and truncated action potentials evoked by various forms of natural stimulation (indicated by brackets below). Resting membrane potential before application of drugs was -58 mV.

In addition, DAMGO decreased the frequency of some evoked and spontaneous PSPs in seven of the hyperpolarized neurons and in seven additional neurons in which hyperpolarization or significant conductance increases were not observed. In some cases, it appeared that larger PSPs were lost following DAMGO application, but there was an increase in very small PSPs. The locations of three of these latter neurons are shown with stars inside open symbols in Fig. 1B. Naloxone restored the spontaneous PSPs and evoked action potential responses toward pre-DAMGO levels (see Fig. 6). The reduction by DAMGO of responses to noxious cooling and heating is demonstrated in Fig. 7, C and D. The reductions in evoked action potentials shown in Figs. 5 and 6 resulted both from hyperpolarization of the recorded neuron and reductions in the frequency of spontaneous and evoked PSPs.



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Fig. 7. In vivo patch clamp recording from nociceptive specific neuron in lamina II (from same neuron with record shown in Fig. 5). Continuous, sequential AC recordings of PSPs evoked by the indicated stimuli. Arrows indicate onset of stimuli in A and B. C and D: stimuli were begun just before this section of recording. *; onset and offset of 0.05 nA current pulses used to determine conductance. A: PSPs evoked by brush are much smaller with slower rise times. B: PSPs evoked by pinch are intermediate in size. C: note the fast rise time and fast decay of apparent unitary PSPs evoked by noxious cooling. After DAMGO (5 µM) record was taken ~80 s after application of drug. Downward pulses, 200 ms long are current pulses to determine conductance changes. Note fewer PSPs following DAMGO application. D: PSPs evoked by noxious heat were much smaller with slower rise times. After DAMGO record taken about 100 s after application of drug. Downward pulses 200 ms long were current pulses to determine conductance changes. Note the truncated action potentials in the before DAMGO traces and the many small PSPs following stimulation. Two stimulus applications are in both before and after DAMGO. Note fewer small PSPs following DAMGO application. Action potentials were evoked by both heat and pinch and are truncated because of their large size in these recordings (). These did not occur after DAMGO.

Of the 19 tested neurons, 1 was nociceptive and 5 were hyperpolarized by DAMGO. All of these five responded to noxious thermal stimuli. The morphology of one of these is shown in Fig. 3A. Of the nine nociceptive neurons that were not hyperpolarized, five demonstrated pronounced reductions in spontaneous postsynaptic potentials because of DAMGO application, whereas three showed no effect and one was depolarized by DAMGO application and demonstrated a 28% conductance decrease (located in lamina I with an asterisk in Fig. 1B and shown morphologically in Fig. 3D). The depolarization and conductance decreases were blocked by the coadministration of naloxone with DAMGO (this neuron was tested with the circulating drug system described in the METHODS section). The three nociceptive neurons that demonstrated no effect did not respond to noxious thermal stimuli and the five that demonstrated only effects on PSPs demonstrated either responses to noxious thermal stimuli (3) or only responded to noxious mechanical stimuli (2).

Of the five nonnociceptive neurons tested, four were mechanoreceptive. Of these four neurons, two were hyperpolarized by DAMGO (shown morphologically in Fig. 3F and 4), one showed reductions in spontaneous PSPs because of DAMGO, and one showed no effect of DAMGO application. The remaining tested neuron responded only to innocuous cooling and demonstrated no membrane effect but demonstrated a reduction in spontaneous PSPs when DAMGO was applied.


    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

The present study used whole cell, tight-seal in vivo recordings in laminae I and II neurons in the spinal cord and examined the response properties of these neurons in the presence of opioids. In vivo recordings of this type have been obtained successfully from the visual cortex of kittens (Nelson et al. 1994), the somatosensory cortex of rats (Moore and Nelson 1998), and the inferior colliculus of the bat (Covey et al. 1996). However, this is the first demonstration of the application of this technology in the spinal cord. Moreover, this study 1) documents the classes of neurons in laminae I and II in the rat in vivo, both physiologically and anatomically, using whole cell recording techniques; 2) documents the hyperpolarization of identified nociceptive lamina I and II neurons by DAMGO in vivo in the rat; 3) demonstrates the decrease in frequency of PSPs evoked by noxious thermal stimuli by DAMGO in lamina I and II neurons in the rat; and 4) compares some of the biophysical properties of lamina I and II neurons recorded by patch clamp in vivo with those in vitro using the same recording procedures.

Methodological considerations

Whole cell, tight-seal recordings have several advantages over sharp electrode, intracellular recordings. The pipettes have very low impedances, which allows for very low noise recordings that are much more stable. Whereas we and others have been able to stabilize cat and monkey spinal cord sufficiently to obtain satisfactory sharp electrode recordings (Iggo et al. 1988; Jones et al. 1990; Light et al. 1979, 1986), we have been largely unsuccessful in obtaining stable recordings from the small cells of laminae I and II in previous attempts to record with sharp electrodes in rats.

The quality of the in vivo recordings was very good. The cell input impedances corresponded well with those obtained from similar cells recorded in vitro (Schneider et al. 1998). Both ongoing and evoked PSPs were observed with high resolution. The enhanced resolution allowed for analyses of the characteristics of evoked PSPs and consequent changes induced by opioid drugs. Conversely, these data help to verify results from in vitro studies, demonstrating similar properties (Schneider et al. 1998). In previous in vitro studies in rat, the input impedance of neurons recorded in the whole cell mode was considerably higher than that reported in sharp electrode studies (North and Yoshimura 1984; Yoshimura and North 1983). These values are much higher than sharp electrode studies conducted on laminae I and II neurons in vivo in the cat (Iggo et al. 1988; Jones et al. 1990; Light et al. 1979, 1986). This study makes it clear that the high values of input impedances are not a result of preparing slices for in vitro recording or bathing solutions, but rather a property of the whole cell recording configuration which presumably has less current shunt than sharp electrode recording techniques.

Physiological properties of lamina I and II neurons

The present results demonstrate that the organization of the rat spinal cord marginal zone and substantia gelatinosa is quite similar to that of the monkey spinal cord (Kumazawa and Perl 1978; Light et al. 1979). Marginal zone neurons had synaptic inputs and fired action potentials in a manner consistent with nociceptors or innocuous cooling afferent inputs. Occasionally, slow brush inputs were observed in neurons in the overlying white matter similar to previous observations by others in the rat (Woolf and Fitzgerald 1983) or deeper in the substantia gelatinosa as previously found in the rat, the cat, and the monkey (Light et al. 1979; Réthelyi et al. 1989; Woolf and Fitzgerald 1983). Neurons in the outer substantia gelatinosa appeared to receive inputs from both mechanoreceptive and polymodal nociceptors; however, cells with input from innocuous mechanoreceptors were located in the inner SG. These recordings confirm the importance of the substantia gelatinosa in nociceptive processing and demonstrate its similarity to the same region in cats and monkeys. This study indicates that most of the neurons in the inner substantia gelatinosa in all three species are nonnociceptive. These neurons appear to have inputs dominated by primary afferents that respond best to gentle, slowly moving stimuli. The function of these neurons is unknown, but they may play a role in modulating neurons in other laminae as we have observed that at least some of these neurons have axons with terminal collaterals in laminae I, III, and IV (Light and Kavookjian 1988). Either by affecting neurons in other laminae or by alternative projections to higher centers, some of these neurons may be involved in sensations such as "tickle" (Zotterman 1939).

Anatomic appearance of recorded neurons

In addition to noting the location of some of the recorded neurons, biocytin labeling allowed us to determine the types of laminae I and II neurons recorded with the patch-clamp technique. A few neurons lying in Lissauer's tract, immediately above lamina I and some lamina I neurons were innocuous cooling input cells. One well-labeled cooling cell may be of the same category (pyramidal) as cooling neurons described from the cat dorsal horn (Han et al. 1998). Other lamina I neurons fall into the categories of fusiform or multipolar, as suggested by these same authors, and all were nociceptive. The lamina II neurons labeled here can be described as islet cells, stalked cells, or stellate cells. All morphological types were nociceptive as well as innocuous mechanoreceptive with the only distinction being that the innocuous mechanoreceptive cells were located in IIi or overlying white matter, similar to the findings of Woolf and Fitzgerald (1983).

Response to opioids

Our data indicate that evoked action potentials in a subset of nociceptive neurons are reduced by opioids both by hyperpolarization and by reductions in spontaneous and evoked PSPs. This results in a powerful overall inhibition of transmission from these neurons.

DAMGO, a selective µ-opioid receptor agonist, hyperpolarized (with a conductance increase) about 37% of the neurons to which it was applied in these experiments. This percentage is somewhat lower than found in sharp electrode in vitro experiments (Grudt and Williams 1994; Jeftinija 1988; Miletic and Randic 1981; Yoshimura and North 1983), but about the same as similar in vitro patch-clamp experiments (Schneider et al. 1998). As in previous in vitro experiments, the predominant postsynaptic effect of DAMGO application in vivo was hyperpolarization with a conductance increase. In addition, we have shown that many of these neurons could be classified as nociceptive cells. However, one neuron was depolarized and appeared to be excited and demonstrated a 28% decrease in conductance. This is not entirely consistent with previous in vivo experiments in cats in which we used sharp electrode recordings. In these studies many neurons were excited by morphine, a µ-opioid agonist (Jones et al. 1990). It also is not entirely consistent with other results in the cat (Craig and Hunsley 1991; Craig and Serrano 1994) that demonstrated that innocuous cooling neurons in lamina I were excited by morphine. The reasons for these discrepancies are unclear, but may be caused by a species difference, because behavioral responses to opioid administration are different in cats compared with rats. However, others have observed excitatory effects in rat spinal cord with extracellular recording techniques both in vitro (Magnuson and Dickenson 1991) and in vivo (Sastry and Goh 1983, Woolf and Fitzgerald 1981).

The majority of neurons hyperpolarized by DAMGO were nociceptive neurons that appeared to receive inputs from both mechanical and thermal nociceptors. DAMGO also provoked a profound decrease in the frequency of PSPs evoked by noxious heat and cooling after the application of DAMGO in many nociceptive neurons. PSPs evoked by noxious mechanical stimulation were affected to a lesser degree. Previous studies using whole cell recording methods have consistently reported that opioids caused decreases in the frequency of PSPs of unknown origin (Glaum et al. 1994; Hori et al. 1992; Schneider et al. 1998). This study suggests that many of these PSPs are from nociceptive primary afferent neurons or interneurons.

All but one of the labeled neurons that were hyperpolarized by DAMGO were found in lamina II, with neurons demonstrating only presynaptic responses being found mostly in lamina I. The small numbers reported here make the significance of this separation unknown, but could reflect the presumed axonal projection of lamina II cells to lamina I.


    SUMMARY
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

This study documents the physiological and anatomic types of neurons using whole cell, tight-seal recordings in lamina I and II of the lumbar spinal cord of the adult rat in vivo. It also documents the relative selectivity of µ-opioid effects on C fiber evoked nociceptive inputs, both presynaptically and postsynaptically. Furthermore, it validates in vitro studies in neurons of lamina I and II in the rat for recording the biophysical properties and effects of opioids.


    ACKNOWLEDGMENTS

The authors acknowledge K. McNaughton for expertise with the histological processing, B. Taylor-Blake and M. Roberts for technical assistance, and T. Grudt and P. Dougherty for reading earlier versions of this manuscript.

This work was supported by National Institute of Neurological Disorders and Stroke Grants R01-NS-16433 and P01-NS-14899.


    FOOTNOTES

Address for reprint requests: A. R. Light, Dept. of Cell and Molecular Physiology, CB#7545 Medical Sciences Research Building, University of North Carolina, Chapel Hill, NC 27599-7545.

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

Received 24 May 1999; accepted in final form 17 August 1999.


    REFERENCES
TOP
ABSTRACT
INTRODUCTION
METHODS
RESULTS
DISCUSSION
SUMMARY
REFERENCES

0022-3077/99 $5.00 Copyright © 1999 The American Physiological Society