Preparation of spinal cord slices
The University of Newcastle Animal Care and Ethics Committee approved all procedures used in this study and they comply with the policies and regulations of The Journal of Physiology as outlined by Drummond (2009). Mice (C57Bl/6, both sexes) were divided into two age groups, P0–5 (hereafter termed neonates) and P24–45 (adults). These ages were selected as they represent developmental stages either side of a critical period in the development of electrophysiological properties in lumbar SDH neurons (Walsh et al. 2009). Approximately equal numbers of male and female mice were used in our experiments (46%vs. 54%, respectively).
Neonatal mice were immersed in ice to induce hypothermia and adult animals were anaesthetized with ketamine (100 mg kg−1i.p.). Once deep hypothermia or anaesthesia was achieved, animals were decapitated and the vertebral column and posterior thoracic wall were rapidly isolated and immersed in ice-cold oxygenated sucrose substituted artificial cerebrospinal fluid (S-ACSF) containing (in mm): 250 sucrose, 25 NaHCO2, 10 glucose, 2.5 KCl, 1 NaH2PO4, 1 MgCl2 and 2.5 CaCl2. The S-ACSF was continually bubbled with 95% O2–5% CO2 to maintain a pH of 7.3–7.4. Lengths of vertebral column containing upper cervical (C2–4), thoracic (T8–10) or lumbar (L3–5) spinal cord segments were isolated and the corresponding spinal cord region was removed. Transverse slices were prepared using previously described techniques (Graham et al. 2008). The isolated spinal cord region was placed on a Styrofoam support-block. The block and spinal cord were then glued to a cutting stage with cyanoacrylate glue (Loctite 454, Loctite, Caringbah, Australia) and transverse slices (400 μm thick for neonates; 300 μm for adults) were obtained using a vibrating blade microtome (Microm HM650, Microm International GmbH, Germany). Slices were transferred to an interface storage chamber containing oxygenated ACSF (118 mm NaCl substituted for sucrose in S-ACSF) and allowed to recover for 1 h at room temperature (22–24°C) before recording commenced.
Slices were transferred to a recording chamber (volume 0.4 ml) and continually superfused (4–6 chamber volumes/min) with ACSF. Recording temperature was maintained at near-physiological temperature (32°C) using an in-line temperature control unit (Model TC324B, Warner Instruments, Hamden, CT, USA). Whole cell patch clamp recordings were obtained from SDH neurons using an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA, USA). Individual neurons were visualized using infrared differential interference contrast optics and an infrared-sensitive camera (Hamamastu C2400-79C, Hamamatsu City, Japan). In adult mice, the substantia gelatinosa (lamina II) appears as a translucent band and recordings were made from neurons located within or dorsal to this region. In neonatal mice this translucent band is not apparent, due to the lack of myelination at this age, so recordings in neonates were restricted to within 140 μm of dorsal surface of the slice as this corresponds to the limit of lamina II in Nissl stained slices of P2 animals (Walsh et al. 2009).
Patch pipettes (2–5 MΩ resistance) were filled with a potassium methylsulphate-based internal solution containing (in mm): 135 KCH3SO4, 6 NaCl, 2 MgCl2, 10 Hepes, 0.1 EGTA, 2 MgATP and 0.3 NaGTP (pH adjusted to 7.3 with KOH). The whole cell recording mode was first established in voltage clamp (holding potential −60 mV, series resistance <20 MΩ). Input resistance (RIN) was measured from the averaged response (5 trials) to a 5 mV hyperpolarizing step. This value was examined at the beginning and end of each recording session and data were rejected if it changed by >20%. We next tested for the presence of the four major subthreshold currents known to exist in rodent SDH neurons (Yoshimura & Jessell, 1989a; Ruscheweyh & Sandkuhler, 2002; Graham et al. 2007c). This was achieved by delivering a hyperpolarizing pulse (to −90 mV, 1 s duration), followed by a depolarizing step (to −40 mV, 200 ms duration). The protocol was repeated five times to obtain an average for analysis. This protocol identified the four major subthreshold currents described previously, including the outward potassium currents rapid A (IAr) and slow A (IAs), the inward T-type calcium current (ICa), and the non-specific cationic H current (IH). In a recent study, we have confirmed the identity of these currents in mouse SDH neurons according to their sensitivity to 4AP, nickel and caesium, respectively (see Graham et al. 2007c). We restricted the depolarizing step to −40 mV to avoid activation of tetrodotoxin-sensitive Na+ and delayed rectifier channels (Safronov, 1999). Capacitive and leakage currents were removed via the automated P/N leak subtraction method within the Axograph software.
For neurons displaying IAr, voltage-dependent activation and steady state inactivation were further assessed using two protocols. Voltage-dependent activation was measured by applying a hyperpolarizing pulse to −90 mV (1 s duration), followed by a series of depolarizing voltage steps of increasing amplitude (from −85 mV to −40 mV, 5 mV increments, 200 ms duration). IAr steady state inactivation was assessed by application of a series of hyperpolarizing prepulses (from −90 mV to −40 mV, 5 mV increments, 1 s duration), followed by a depolarizing voltage step to −40 mV (200 ms duration).
After running the above protocols, the recording mode was switched to current clamp. The membrane potential observed ∼15 s after this switch was considered to be resting membrane potential (RMP) and all current clamp recordings were made from this potential. All reported membrane potentials were corrected for a 10 mV junction potential (Barry & Lynch, 1991). Individual action potential (AP) properties and discharge characteristics were examined by injecting a series of depolarizing and hyperpolarizing current steps (800 ms duration, 20 pA increments, delivered every 8 s).
At the end of each recording session, the location of the recorded neuron was mapped as described previously (Graham et al. 2007c). Briefly, digital images of the spinal cord slice with the electrode still attached to the neuron were captured using a Rolera-XR digital camera, and QCapture Pro software (Spectra Services, Rochester, NY, USA). The image was then imported into Adobe Illustrator and manipulated so the dorsal horns aligned. A standardized template of the relevant spinal cord segment was then resized until it could be overlaid on the dorsal grey matter borders of the imported image. The location of the recorded neuron was plotted on these standardized templates. For neonates, templates were from the spinal cord of a P4 mouse (courtesy of Charles Watson and Gulgun Kayalioglu). Adult templates were from Watson et al. (2009).
Data capture and analysis
Data were digitized online (sampled at 10 kHz, filtered at 5 kHz) via an ITC-16 A/D board (Instrutech, Long Island, NY, USA) and stored on a Macintosh G4 computer running Axograph v4.6 software (Axon Instruments, Union City, CA, USA). All data were analysed offline using the Axograph software. Subthreshold currents were classified as previously described (Graham et al. 2007c; Walsh et al. 2009). Because the rapid A-type potassium current (IAr) is the dominant subthreshold current at all developmental stages in mouse lumbar SDH neurons (Walsh et al. 2009), its features were analysed further for neonatal and adult mice. IAr amplitude was measured by subtracting the amplitude of any steady state current component (in the last 50 ms of the step to −40 mV) from the maximal IAr current peak (Graham et al. 2008). Activation and steady state inactivation curves were generated and subsequently fit with Boltzmann equation: g/gmax= 1/[1 + exp((V−V50)/k)] where g/gmax is normalized conductance, V is membrane potential, V50 is voltage at half-maximal activation (or inactivation) for membrane potentials −90 to −40 mV, and k is the slope factor. Under these conditions IAr is never fully activated, so we could not compare half-activation in the classic sense. We therefore compare data points at each membrane potential (5 mV increments over −90 to −40 mV). The decay phase of the IAr response was fit with a single exponential (over 20–80% of its falling phase).
Individual APs were captured using a derivative threshold method, with the threshold ranging from 10 to 15 mV ms−1 for neonatal neurons, and 18–20 mV ms−1 for adult neurons. AP threshold was defined as the inflection point during spike initiation. Rheobase current was defined as the smallest step-current that elicited at least one AP. The amplitude of each AP was measured as the difference between threshold and its maximum positive peak. AP half-width was calculated at 50% of AP amplitude. AP afterhyperpolarization (AHP) amplitude was measured as the difference between AP threshold and its maximum negative peak.
The SPSS v16 software package (SPSS, Chicago, IL, USA) was used for statistical analysis. One-way ANOVA was used to compare means of passive and active properties between/across the three regions of the cord and between neonatal and adult mice. Scheffe's post hoc tests were used to determine where data differed. Data that failed Levene's test of homogeneity of variance were compared using the non-parametric Kruskal–Wallace test, followed by Tamhane's T2 post hoc test. G tests, with Williams's correction, were used to determine whether the prevalence of discharge categories, responses to hyperpolarizing steps, and subthreshold currents differed between regions of the cord and between neonatal and adult mice. Statistical significance was set at P < 0.05 and all data are presented as means ± SEM.