| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
School of Biomedical Sciences, Faculty of Health and Hunter Medical Research Institute, The University of Newcastle, Callaghan, New South Wales, Australia
Submitted 22 October 2007; accepted in final form 18 February 2008
 |
ABSTRACT |
|---|
 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
INTRODUCTION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
,b). Despite this, much of the data generated over the past two decades on neuronal excitability in the mammalian nervous system using in vitro preparations have been acquired at room temperature (21–25°C). In cases where in vitro experiments have been repeated at higher temperatures, many phenomena are markedly different (Cao and Oertel 2005; Lee et al. 2005; Micheva and Smith 2005). For example, neurons in the brain stem (Borst and Sakmann 1998; Cao and Oertel 2005), hypothalamus (Griffin and Boulant 1995), hippocampus (Thompson et al. 1985), and cortex (Lee et al. 2005; Volgushev et al. 2000) have altered input resistance, action potential (AP) amplitude, and AP width at elevated temperature (
32°C). Such differences have important implications for extrapolation of data collected in vitro at room temperature, to the in vivo situation where neurons operate at physiological temperatures (
37°C). Alternatively, observations made in vivo can be further examined in vitro with the proviso that the influence of factors, such as recording temperature, are fully appreciated (e.g., Margrie et al. 2001).
Temperature considerations are especially relevant when studying heterogeneous neuron populations where differential expression of various temperature-sensitive voltage-gated conductances shapes neuronal discharge (Hille 2001). Superficial dorsal horn (SDH) neurons in the spinal cord are one such example of a highly heterogeneous neuron population (Melnick et al. 2004a,b; Ruscheweyh and Sandkühler 2002; Ruscheweyh et al. 2004; Yoshimura and Jessell 1989). These central neurons play important roles in processing noxious, thermal, itch, and innocuous tactile stimuli transmitted by A
and C-fiber primary afferents (Christensen and Perl 1970; Sugiura et al. 1986; Tuckett and Wei 1987; Vallbo et al. 1999). They can be divided into various categories based on their AP discharge during depolarizing current injection. For example, some SDH neurons discharge APs tonically, others display prominent spike frequency adaptation, and others exhibit delayed AP discharge. Progress has been made toward identifying the ionic mechanisms underlying these discharge categories. Specifically, the relative levels of tetrodotoxin-sensitive Na+ current and a delayed rectifier K+ current are thought to underlie the tonic and adaptive AP discharge categories (Melnick et al. 2004a,b). The fast activating and inactivating potassium current, termed rapid A-type (IAr), has been shown to delay AP discharge (Ruscheweyh and Sandkühler 2002; Ruscheweyh et al. 2004; Yoshimura and Jessell 1989).
To date, in vitro studies investigating the discharge properties of SDH neurons and their underlying conductances have been carried out at room temperature. No study has comprehensively examined how elevating temperature to more biologically relevant levels affects AP discharge properties in SDH neurons, or how this might affect our understanding of nociceptive processing in the SDH. Therefore we have assessed the in vitro membrane and AP discharge properties of SDH neurons at room temperature (RT, 22°C) and at elevated, near-physiological temperature (PT, 32°C). Comparison of these results with the in vivo behavior of SDH neurons suggests that, on balance, in vitro data collected at elevated temperature more closely resemble data collected under in vivo conditions.
|
METHODS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
). In vitro spinal cord slice preparation
Mice (C57Bl/6, both sexes: 17–69 days) were anesthetized with ketamine [100 mg/kg, administered intraperitoneally (ip)] and decapitated. The vertebral column, attached ribs, and soft tissue were surgically isolated and immersed in ice-cold oxygenated sucrose-substituted artificial cerebrospinal fluid (S-ACSF). The S-ACSF contained (in mM): 250 sucrose, 25 NaHCO2, 10 glucose, 2.5 KCl, 1 NaH2PO4, 1 MgCl2, and 2.5 CaCl2, and was continually bubbled with carbogen (95% O2-5% CO2) to achieve a pH of 7.3. Slices were prepared as described previously (Graham et al. 2007a,b). Briefly, the lumbosacral enlargement of the spinal cord (L3–L5) was dissected free of the vertebrae under a dissecting microscope using a ventral approach. The isolated cord was placed against a Styrofoam support block and glued (rostral end down) to a cutting platform. The block and tissue were placed in a chamber containing oxygenated S-ACSF and transverse slices (300 µm thick) were obtained using a vibrating microtome (VT-1000S, Leica, Heidelberg, Germany). Slices were transferred to a storage chamber containing oxygenated ACSF (118 mM NaCl substituted for sucrose in S-ACSF) and allowed to incubate for 1 h before recording.
In vitro electrophysiology
Spinal cord slices were continually superperfused with oxygenated ACSF in a recording chamber (chamber volume 0.4 ml; exchange rate 4–6 bath volumes/min). Patch-clamp recordings were made using an Axopatch 200B amplifier (Molecular Devices, Sunnyvale, CA). SDH neurons were visualized using infrared differential interference contrast (IR-DIC) optics (Stuart et al. 1993). Under IR-DIC visualization lamina II appears as a translucent band: recordings were restricted to neurons in or dorsal to this region and 20–100 µm below the slice surface. Patch pipettes (2- to 5-M
resistance) were filled with a K+-based internal solution containing (in mM): 135 KCH3SO4, 6 NaCl, 2 MgCl2, 10 HEPES, 0.1 EGTA, 2 MgATP, and 0.3 NaGTP (pH 7.3 with KOH). The whole cell recording configuration was established in voltage clamp (holding potential –60 mV, series resistance <20 M). Input resistance was calculated according to the average response to a 5-mV hyperpolarizing step (10-ms duration, 30 repetitions). In most experiments a number of protocols were run in voltage clamp before the amplifier was switched to current clamp. The membrane potential observed about 15 s after the switch to current clamp was designated as resting membrane potential (RMP). All reported membrane potentials were corrected for a calculated liquid junction potential of 10 mV (Barry and Lynch 1991). All signals were amplified, filtered at 10 kHz, and digitized at 10 or 20 kHz via an ITC-16 A/D interface (InstruTECH, Port Washington, NY), connected to an Apple Macintosh G4 computer running AxoGraph software (v4.8; Axon Instruments, Foster City, CA).
After electrophysiological characterization the location of each recorded neuron within the SDH was mapped as described previously (Graham et al. 2007b). Briefly, we photographed the dorsal horn while the electrode was still attached to the recorded neuron with a digital camera and Viewfinder Lite software (Olympus, Tokyo, Japan). Images were imported into Adobe Illustrator and manipulated so the location of each neuron could be plotted on a standardized template of the appropriate segment. Templates of the gray and white matter borders for L3, L4, and L5 segments were generated using Franklin and Paxinos (1997).
In vivo preparation
Details of the in vivo mouse spinal cord preparation have been described previously (Graham et al. 2004a). Briefly, mice (26–42 days, both sexes) were anesthetized with urethane (2.2 g/kg ip). After reaching a deep level of anesthesia, animals were transferred to a customized frame and stabilized with ear and palate bars. A thermal pad placed under the animal maintained body temperature between 34 and 37°C and humidified 100% O2 was continuously blown over the animals' nostrils. The vertebral column was stabilized with custom-made clamps and a laminectomy (at L1) exposed the widest point of the lumbosacral enlargement (L4). The dura was reflected and a small incision made in the pia to allow penetration of the underlying dorsal horn with a recording pipette. Throughout the experiment the surface of the spinal cord was irrigated with ACSF (as used for in vitro experiments), maintained at 37°C. At the completion of experiments animals were overdosed with Nembutal (100 mg/kg ip).
In vivo electrophysiology
Recording pipettes (8–12 M) were fabricated from thick-wall (OD 1.5 mm, ID 0.86 mm) borosilicate glass capillaries and filled with the same K+-based internal solution used for in vitro experiments. Patch-clamp recordings were made using an Axoclamp 2B amplifier (Molecular Devices). Pipettes were first advanced through the white matter of the spinal cord to a depth of about 100 µm, while positive pressure (0.5 bar) was applied to the pipette tip. The pressure on the pipette was reduced to 0.1 bar and we searched for neurons by advancing a further 250 µm (3-µm steps) into the dorsal horn. After a tight seal (>1 G) was obtained on a SDH neuron the membrane patch was ruptured using gentle suction to establish the whole cell recording configuration (holding potential –60 mV, series resistance <50 M). As with in vitro experiments, when the amplifier was switched to current-clamp mode the membrane potential observed about 15 s after this switch was designated as RMP. In these experiments all protocols were run in current clamp (bridge mode). Data were amplified, filtered, digitized, and stored as for in vitro experiments. All membrane potential values were corrected for a 10-mV calculated liquid junction potential (Barry and Lynch 1991). We note that the different amplifiers used for our in vitro and in vivo experiments have been reported to filter and distort voltage signals differently (Magistretti et al. 1996). The extent of this distortion was quantified using the same model cell (500 M, 16.5-ms time constant, 10-M series resistance) on the Axopatch 200B and Axoclamp 2B amplifiers. The Axoclamp 200B and Axopatch 2B amplifiers reduced theoretically calculated values for peak amplitude and time constant by approximately 4 and 7%, respectively. Thus we may have slightly overestimated spike height and underestimated spike width in our in vitro experiments.
Experimental protocols
All voltage-clamp protocols, run in vitro, were made from a holding potential of –60 mV and used standard P/N leak subtraction protocols to remove capacitive and leakage currents and to isolate whole cell subthreshold ionic currents (Sontheimer and Ransom 2002) (semiautomated procedure, Axograph 4.6 software). The first protocol tested for the presence of a transient, rapidly decaying potassium current (termed IAr) by delivering a hyperpolarizing prepulse to –90 mV (1-s duration), followed by a depolarizing step to –40 mV (200-ms duration). The second protocol assessed steady-state inactivation of IAr by delivering a series of prepulses from –90 to –40 mV in 5-mV increments, followed by a depolarizing voltage step to –40 mV (200-ms duration). The third protocol assessed the voltage-dependent activation of IAr by applying a hyperpolarizing prepulse to –90 mV (1-s duration) followed by voltage steps of increasing amplitude from –85 to –40 mV in 5-mV increments. A common set of current-clamp protocols were run for both in vitro and in vivo experiments. Depolarizing and hyperpolarizing current steps (800-ms duration, 20-pA increments, delivered every 8 s) were injected to determine each neuron's voltage response. During these protocols, sustained membrane deflections were limited to –20 mV during depolarizing steps and –100 mV during hyperpolarizing steps to minimize neuronal damage.
Data analysis
Data analysis was performed off-line using semiautomated procedures within Axograph v4.8 and Igor Pro software v5 (WaveMetrics, Lake Oswego, OR). Individual APs elicited by depolarizing current injection were captured using a derivative threshold method (threshold set at dV/dt = 15–20 V/s). The inflection point during spike initiation was defined as AP threshold. Rheobase current was defined as the smallest current step that elicited at least one AP. Individual AP properties for all SDH neurons were measured from the rheobase response. AP amplitude was measured as the difference between AP threshold and its maximum positive peak. AP base width was measured at AP threshold. AP afterhyperpolarisation (AHP) amplitude was measured as the difference between AP threshold and the maximum negative peak following the falling phase of the AP. Several parameters were measured to describe AP discharge during depolarizing current injections. For responses that contained multiple APs, mean frequency was calculated as the average of all instantaneous AP frequencies.
Activation and steady-state inactivation curves for the IAr current were fit with the Boltzmann equation, g/gmax = 1 – {1/[1 + exp (V – V1/2)/
]}, where g/gmax = normalized conductance, V = membrane potential, V1/2 = voltage at half-maximal activation (or inactivation), and is the slope factor. The temperature sensitivity of measured parameters was expressed as Q10 values (the proportionate change for a 10°C change in temperature). Q10 values were calculated using the equation Q10 = (X1/X2)10/t2–t1, where t2 is 32°C and t1 is 22°C and X2 and X1 are the corresponding parameters measured at the two temperatures. Because comparisons are made between 22 and 32°C, a Q10 value close to 1 for a given property indicates little or no temperature dependence. A Q10 value <1 indicates that a property will decrease as temperature is elevated, whereas a Q10 value >1 indicates a property will increase as temperature is elevated.
SPSS v10 software package (SPSS, Chicago, IL) was used for most statistical analyses. One-way ANOVA was used to compare variables between/across discharge categories. Student–Neuman–Keuls post hoc tests were used to determine where data differed. Data that failed Levene's test of homogeneity of variance were compared using the nonparametric Kruskal–Wallace test. G-tests, with Williams' correction, were used to determine whether discharge patterns differed at RT and PT recording temperatures (Sokal and Chapman 2003). Separate post hoc Pearson's chi-squared tests were subsequently applied to compare the proportions of each discharge category observed under the two temperature conditions (tonic firing, initial bursting, delayed firing, single spiking, and reluctant firing). All values are presented as means ± SE. All comparisons are described as significant when P < 0.05, unless otherwise stated.
|
RESULTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Data from 93 SDH neurons, previously collected using an in vivo (IV) mouse spinal cord preparation (Graham et al. 2004a), were compared with data obtained at the two in vitro temperatures (RT and PT). IV results were included to assess how closely elevating temperature in an in vitro preparation reproduces the active and passive properties of SDH neurons recorded in vivo.
Temperature effects on passive and active membrane properties
The locations of recorded neurons across the three spinal segments for the two recording temperatures are summarized in Fig. 1. Neurons were similarly distributed across the rostrocaudal, mediolateral, and dorsoventral extent of the SDH, suggesting any observed differences are not due to a bias in recording location. Temperature influenced most passive and active membrane properties in SDH neurons. For example, input resistance and RMP are altered when recordings are made at the three temperatures. Input resistance was significantly higher at RT compared with PT and IV (510 ± 26 vs. 370 ± 14 and 361 ± 21 M, n = 106, n = 105, n = 93, respectively). RMP at RT was more hyperpolarized than at PT (–69. 4 ± 0.7 vs. –67.1 ± 0.8 mV, n = 113, n = 106, respectively). These in vitro values, however, were almost 10 mV more hyperpolarized than the in vivo values we have previously reported for SDH neurons (–58.1 ± 0.7 mV, n = 93) (Graham et al. 2004a).
|
|
; Ruscheweyh and Sandkühler 2002). Within-cell temperature effects were assessed during heating (RT to PT, n = 12) and cooling (PT to RT, n = 13). AP threshold was stable when neurons were heated from RT to PT (–32.7 ± 1.3 vs. –33.0 ± 2.0 mV) or cooled from PT to RT (–30.7 ± 1.4 vs. –32.2 ± 1.3 mV). Heating from RT to PT significantly reduced AP amplitude (79.6 ± 4.7 vs. 54.8 ± 3.7 mV) and AP base width (2.9 ± 0.3 vs. 1.7 ± 0.1 ms). Likewise, cooling from PT to RT significantly increased AP amplitude (57.5 ± 4.1 vs. 78.0 ± 4.0 mV) and AP base width (1.6 ± 0.1 vs. 2.8 ± 0.1 ms). Changing bath temperature had variable effects on AHP amplitude, although the mean differences during heating from RT to PT (–29.8 ± 1.6 vs. –34.8 ± 1.9 mV) and cooling from PT to RT (–35.5 ± 1.4 vs. –32.7 ± 1.2 mV) were not significantly different. Q10 values were used to compare "within-cell" versus "between-cell" temperature effects on membrane and AP properties (Table 1). Similar conclusions are reached irrespective of the way temperature effects are examined. In summary, the effect of temperature on most membrane and AP properties is modest apart from those on AP amplitude and base width, which are highly temperature sensitive.
|
Neurons in the SDH are a heterogeneous population with several different categories described according to AP discharge patterns during depolarizing current injection (Graham et al. 2007a; Grudt and Perl 2002; Hu and Gereau 2003; Lopez-Garcia and King 1994; Lu et al. 2006; Ruscheweyh and Sandkühler 2002; Thomson et al. 1989; Yoshimura and Jessell 1989). We identified five AP discharge categories in this study under RT, PT, and IV conditions (Fig. 3A). Tonic firing was characterized by persistent AP discharge that lasted for the duration of the current injection. Initial bursting was characterized by AP discharge limited to the beginning of the current injection. These neurons were the most likely to exhibit rebound depolarization, or occasional APs, after release from hyperpolarizing current injection. Delayed firing featured a prominent delay between the onset of the current injection and the initiation of AP discharge. Single spiking was characterized by the discharge of one or two APs at the onset of the current injection. Finally, reluctant firing neurons did not discharge APs despite the delivery of depolarizing current injections that moved membrane potential well above Na+ current activation thresholds. These reluctant firing neurons had input resistance similar to that of neurons in the other categories (372 ± 39 vs. 370 ± 15 M, n = 15, n = 90, respectively in PT recordings); however, their RMPs were more hyperpolarized (–74 ± 2 vs. –66 ± 1 mV, n = 15, n = 91, respectively). Together, these two measurements suggest reluctant firing neurons were neither damaged nor unhealthy and were therefore included in our analysis.
|
), both RT and PT proportions showed some similarities. For example, the prevalence of initial bursting neurons at RT is similar to the proportion exhibiting this discharge pattern in vivo. On the other hand, a greater representation of reluctant firing neurons and reduced delayed firing neurons at PT resemble our previous in vivo findings. Temperature effects between AP discharge categories
We investigated whether the different temperature conditions affected membrane and AP properties (summarized in Fig. 2 and Table 1 for all SDH neurons) within each of the four discharge categories featuring AP discharge (Table 2). At elevated temperature, input resistance was decreased in initial bursting and delayed firing neurons, and RMP was more depolarized in tonic firing and delayed firing neurons. Rheobase current was temperature sensitive only in single spiking neurons. Several AP features within discharge categories also exhibited different temperature sensitivities. For example, elevated temperature significantly depolarized AP threshold for tonic firing neurons only. AP amplitude decreased at elevated temperature in all categories except delayed firing neurons. AP base width was consistently reduced at elevated temperature for all discharge categories, whereas AHP amplitude was temperature sensitive only in single spiking neurons. Thus membrane and AP properties between the four discharge categories exhibit complex temperature sensitivities. This precludes the use of simple extrapolation when comparing data acquired at different temperatures.
|
|
|
Since it is well established that IAr underlies delayed firing in SDH neurons (Ruscheweyh and Sandkühler 2002; Ruscheweyh et al. 2004), and because we have demonstrated a relationship between delayed and reluctant firing (Fig. 5), we next investigated the role IAr plays in reluctant firing (Fig. 6). Depolarizing current injections were repeated in reluctant firing neurons while they were held at more depolarized membrane potentials (Fig. 6A). Because IAr is voltage sensitive, we predicted this procedure would diminish the ability of this current to inhibit AP discharge. All reluctant firing neurons subjected to current injections from more depolarized membrane potentials exhibited AP discharge (RT, n = 2; PT, n = 3). It is important to note that our classification of reluctant firing neurons was based on current injections that were well above AP threshold. Thus the initial absence of AP discharge in neurons classified as reluctant firing cannot be explained simply by a failure to reach AP threshold.
|
|
DISCUSSION |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Membrane properties
Reduced input resistance at elevated temperature is a consistent finding in a variety of neuronal types (Cao and Oertel 2005; Griffin and Boulant 1995; Klee et al. 1974; Lee et al. 2005; Thompson et al. 1985; Volgushev et al. 2000). This is also the case for SDH neurons in general (Table 1); however, when neurons were separated by discharge category the reduction in input resistance was not uniform. For example, the effect of elevated temperature on input resistance is greatest in delayed firing neurons (37% reduction) and least in tonic firing neurons (11% reduction). The mechanisms underlying these changes have not been directly assessed in this study, although, in cortical neurons, the greater temperature sensitivity of potassium versus sodium conductances is considered responsible (Lee et al. 2005; Volgushev et al. 2000).
The influence of temperature on RMP is equivocal across studies. In cat motoneurons and rat visual cortex neurons cooling shifts RMP to more depolarized levels (Klee et al. 1974; Volgushev et al. 2000). In contrast, cooling mouse cochlear neurons leads to hyperpolarization and heating leads to depolarization (Cao and Oertel 2005). Finally, studies in hippocampal CA1, hypothalamic, and neocortical pyramidal neurons suggest temperature has little or no effect on RMP (Griffin and Boulant 1995; Lee et al. 2005; Thompson et al. 1985). In mouse SDH neurons we observed a modest depolarization of RMP (3 mV) at elevated temperature (Table 1) in population comparisons. Unlike input resistance, however, this effect was not consistent across discharge categories. The RMPs of initial bursting and single spiking neurons were unaffected by elevated temperature (<2-mV shift). Conversely, the RMPs of tonic firing and delayed firing neurons were depolarized by elevated temperature (>5-mV shift). Because RMP is set by several voltage-sensitive conductances, it is not surprising that in a heterogeneous population, like SDH neurons, temperature effects are variable.
AP properties
Elevating temperature caused a marked reduction in AP amplitude and base width in SDH neurons. These findings are strikingly similar to previous temperature-sensitivity studies across several neuronal populations (Cao and Oertel 2005; Hodgkin and Huxley 1952a,b; Joyner 1981; Klee et al. 1974; Lee et al. 2005; Volgushev et al. 2000). In some of these studies the differing temperature sensitivity has been attributed to the two major currents underlying AP generation: voltage-sensitive sodium and potassium currents. Cao and Oertel (2005) suggested slowed activation of the repolarizing potassium current at room temperature allows greater depolarization during sodium current activation. This effect is compounded by slowed sodium current inactivation. Alternatively, Volgushev et al. (2000) attributed an increased activation threshold of the delayed rectifier potassium current (with little effect on sodium current) as the underlying mechanism for enhanced AP amplitude and width at room temperature. From our current data, we are unable to differentiate between these two possible mechanisms in SDH neurons.
There was a trend toward increased AHP amplitude at elevated temperature; however, this was significant only in single spiking neurons. Studies in different neuronal populations have noted that elevated temperature has little effect on AHPs. Lee et al. (2005), however, suggested the onset and kinetics of Ca+-dependent slow AHPs was delayed by cooling in neocortical pyramidal neurons. Little is known about the prevalence of Ca+-dependent AHPs in SDH neurons (Safronov 1999) and it remains to be determined whether a similar mechanism operates in the SDH.
AP discharge properties
The discharge properties of SDH neurons have been examined extensively using in vitro preparations at room temperature (Grudt and Perl 2002; Melnick et al. 2004a,b; Prescott and De Koninck 2002; Ruscheweyh and Sandkühler 2002; Thomson et al. 1989; Yoshimura and Jessell 1989). At RT, injection of depolarizing current steps reveals four major discharge categories: tonic firing, initial bursting, delayed firing, and single spiking. Likewise, at RT we also observed four main discharge patterns, although we also found a small proportion of neurons that do not discharge APs during depolarizing current steps. Such neurons are rarely described in vitro, except for one study where they were classified together with single spiking neurons (Prescott and De Koninck 2002). At elevated temperature the prevalence of these reluctant firing neurons increased dramatically (Fig. 5A). We propose elevated temperature enhances IAr (see Fig. 6C) and "converts" some delayed firing neurons into the reluctant firing state. This notion is supported by five observations: 1) a reduced prevalence of delayed firing neurons at elevated temperature, and a concomitant increase in reluctant firing neurons (Fig. 5A); 2) conversion of delayed firing responses to reluctant firing by elevating temperature in vitro (Fig. 5B); 3) significantly larger IAr amplitude in reluctant firing neurons (Fig. 6C); 4) the capacity of reluctant firing neurons to discharge spikes when IAr is either partially inactivated or pharmacologically blocked (Fig. 6, A and B); and 5) the increased prevalence of reluctant firing SDH neurons in our in vivo recordings (Fig. 3B). It should be noted, however, that the proportion of initial bursting neurons also decreased at elevated temperature (Fig. 3B) and their possible conversion to reluctant firing neurons cannot be discounted.
The role of IAr has been studied extensively throughout the nervous system and its predominant function is to provide shunting inhibition (for recent review see Jerng et al. 2004). At the soma, IAr reduces the effect of injected current, thus larger currents are required to reach AP threshold. In dendrites IAr attenuates backpropagating APs. Numerous studies have shown that during depolarizing current injections the shunting inhibition provided by IAr increases rheobase, delays the onset of AP discharge, and increases interspike interval (Mitterdorfer and Bean 2002; Molineux et al. 2005; Russier et al. 2003; Varga et al. 2004; Vydyanathan et al. 2005). Our data suggest an additional role for IAr, where the shunting inhibition actually prevents AP discharge in reluctant firing neurons altogether. This particular role becomes significant at elevated in vitro and in vivo temperatures because of the temperature sensitivity of this potassium current. Thus the presence or absence of IAr has functional relevance for signal processing in the SDH in vivo.
In terms of AP discharge rates, both tonic firing and initial bursting neurons fired at significantly increased frequency at PT (Fig. 4). This is in stark contrast to many delayed firing neurons, which were silenced when temperature was raised (Fig. 5). Thus elevating temperature in slices has differing effects on AP discharge in the various SDH neuron subpopulations. The excitability of some neurons is enhanced (tonic firing and initial bursting), whereas in others excitability is reduced (delayed firing). Consequently, models of SDH neuron circuitry and function, developed using data collected at room temperature, will necessarily behave differently at physiological temperature.
Comparison of in vitro and in vivo AP discharge
The question of how well any experimental preparation reflects the in vivo state is of great importance. A unique feature of this study is that a single laboratory has made comparable recordings from SDH neurons at two in vitro temperatures and at body temperature (in vivo). Comparison of data collected at the two in vitro temperatures (RT and PT) with our in vivo experiments provides a new perspective on the behavior of SDH neurons. In particular, mean values for input resistance, AP amplitude, and AP base width are almost identical at elevated temperature and in vivo, although they differ significantly from values obtained at room temperature.
The impact of larger, broader APs at room temperature has been shown to enhance Ca2+ entry during AP discharge in a number of studies (Borst and Sakmann 1998; Lee et al. 2005; Markram et al. 1995). In SDH neurons this could influence Ca2+-dependent mechanisms such as neurotransmitter release and long-term potentiation, which have so far been studied only at room temperature in the SDH (Ikeda et al. 2003; Liu and Sandkühler 1995, 1997). The effects of lowered temperature, however, may be countered by the slowing of biochemical reactions and intracellular processes that follow Ca2+ influx. For example, electrogenic pumps (e.g., Na+/K+-ATPase) are highly temperature sensitive (Q10 >2), and show reduced activity as temperature decreases (Thompson et al. 1985). Future studies, at elevated temperatures, are required to fully understand the net result of altered Ca2+ entry, electrogenic pump activity, and the downstream effects, in vivo.
We expected all SDH neuron properties assessed at elevated in vitro temperature to approach those in vivo values we have previously reported. RMP, AP threshold, and AHP amplitude, however, did not respond as predicted. This result highlights that, although in vitro temperature was elevated to near physiological levels, other important differences still exist between the in vitro tissue slice and an intact "living" nervous system. One major difference is the reduced connectivity in a 300-µm-thick spinal slice where many neurons are disconnected from primary afferent and descending inputs. This would remove tonic facilitatory and/or inhibitory drive to the SDH (Mason 2005). The tissue slicing procedure could also truncate the dendrites of recorded neurons. This will influence net synaptic connectivity and membrane properties of neurons in vitro. These factors might be expected to alter the level of spontaneous synaptic activity between preparations and may contribute to differences in RMP and AP threshold. The effect of such disconnection and truncation on SDH neuronal membrane and AP properties in slices is unclear at this stage.
Our in vivo recordings were all made under urethane anesthesia and the actions of this drug are not fully understood. Studies using recombinant expression of the major fast excitatory and inhibitory ligand-gated ion channels suggest its action is broad, influencing all receptors studied (Hara and Harris 2002). More recently, the effect of urethane on cortical neurons was studied in brain slices. This study suggested urethane had little effect on receptors underlying synaptic transmission, but instead activated a potassium leak conductance that diminished neuronal excitability and AP discharge (Sceniak and MacIver 2006). Thus although its actions are still under debate, we cannot exclude a contribution of urethane anesthesia to our in vivo data.
In summary, the results from this study indicate that in vitro experiments completed at elevated temperature, on balance, more accurately reflect SDH neuron properties recorded in vivo than experiments carried out at room temperature. Extrapolating in vivo functions from data collected at room temperature has proved problematic not only for neuronal excitability (Cao and Oertel 2005; Griffin and Boulant 1995; Lee et al. 2005; Thomson et al. 1989; Volgushev et al. 2000), but also for processes underlying synaptic transmission (Micheva and Smith 2005; Thomson et al. 1989; Volgushev et al. 2000). Our findings therefore provide a basis for comparing room-temperature recordings with those made at elevated temperature or in vivo. Moreover, they argue that future in vitro experiments investigating SDH neuron function be undertaken at more physiologically relevant temperatures, whenever possible.
|
GRANTS |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
|
FOOTNOTES |
|---|
1 The online version of this article contains supplemental data. 
Address for reprint requests and other correspondence: R. J. Callister, School of Biomedical Sciences, Faculty of Health, The University of Newcastle, Callaghan, NSW 2308, Australia (E-mail: robert.callister{at}newcastle.edu.au)
|
REFERENCES |
|---|
|
TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSREFERENCES |
|---|
Borst JGG, Sakmann B. Calcium current during a single action potential in a large presynaptic terminal of the rat brainstem. J Physiol 506: 143–157, 1998.
Cao X-J, Oertel D. Temperature affects voltage-sensitive conductances differentially in octopus cells of the mammalian cochlear nucleus. J Neurophysiol 94: 821–832, 2005.
Christensen BN, Perl ER. Spinal neurons specifically excited by noxious or thermal stimuli: marginal zone of the dorsal horn. J Neurophysiol 33: 293–307, 1970.
Franklin KBJ, Paxinos G. The Mouse Brain in Stereotaxic Coordinates. San Diego, CA: Academic Press, 1997.
Graham BA, Brichta AM, Callister RJ. An in vivo mouse spinal cord preparation for patch-clamp analysis of nociceptive processing. J Neurosci Methods 136: 221–228, 2004a.[CrossRef][Web of Science][Medline]
Graham BA, Brichta AM, Callister RJ. In vivo responses of mouse superficial dorsal horn neurones to both current injection and peripheral cutaneous stimulation. J Physiol 561: 749–763, 2004b.
Graham BA, Brichta AM, Callister RJ. Pinch-current injection defines two discharge profiles in mouse superficial dorsal horn neurones, in vitro. J Physiol 578: 787–798, 2007a.
Graham BA, Brichta AM, Schofield PR, Callister RJ. Altered potassium channel function in the superficial dorsal horn of the spastic mouse. J Physiol 584: 121–136, 2007b.
Griffin J, Boulant J. Temperature effects on membrane potential and input resistance in rat hypothalamic neurones. J Physiol 488: 407–418, 1995.
Grudt TJ, Perl ER. Correlations between neuronal morphology and electrophysiological features in the rodent superficial dorsal horn. J Physiol 540: 189–207, 2002.
Hara K, Harris RA. The anesthetic mechanism of urethane: the effects on neurotransmitter-gated ion channels. Anesth Analg 94: 313–318, 2002.
Hille B. Ion Channels of Excitable Membranes. Sunderland, MA: Sinauer, 2001.
Hodgkin AL, Huxley AF. The components of membrane conductance in the giant axon of Loligo. J Physiol 116: 473–496, 1952a.
Hodgkin AL, Huxley AF. A quantitative description of membrane current and its application to conduction and excitation in nerve. J Physiol 117: 500–544, 1952b.
Hu H-J, Gereau RW 4th. ERK integrates PKA and PKC signaling in superficial dorsal horn neurons. II. Modulation of neuronal excitability. J Neurophysiol 90: 1680–1688, 2003.
Ikeda H, Heinke B, Ruscheweyh R, Sandkühler J. Synaptic plasticity in spinal lamina I projection neurons that mediate hyperalgesia. Science 299: 1237–1240, 2003.
Jerng HH, Pfaffinger PJ, Covarrubias M. Molecular physiology and modulation of somatodendritic A-type potassium channels. Mol Cell Neurosci 27: 343–369, 2004.[CrossRef][Web of Science][Medline]
Joyner RW. Temperature effects on neuronal elements. Fed Proc 40: 2814–2818, 1981.[Web of Science][Medline]
Klee MR, Pierau FK, Faber DS. Temperature effects on resting potential and spike parameters of cat motoneurons. Exp Brain Res 19: 478–492, 1974.[Web of Science][Medline]
Lee JCF, Callaway JC, Foehring RC. Effects of temperature on calcium transients and Ca2+-dependent afterhyperpolarizations in neocortical pyramidal neurons. J Neurophysiol 93: 2012–2020, 2005.
Liu X-G, Sandkühler J. Long-term potentiation of C-fiber-evoked potentials in the rat spinal dorsal horn is prevented by spinal N-methyl-D-aspartic acid receptor blockage. Neurosci Lett 191: 43–46, 1995.[CrossRef][Web of Science][Medline]
Liu X-G, Sandkühler J. Characterization of long-term potentiation of C-fiber-evoked potentials in spinal dorsal horn of adult rat: essential role of NK1 and NK2 receptors. J Neurophysiol 78: 1973–1982, 1997.
Lopez-Garcia JA, King AE. Membrane properties of physiologically classified rat dorsal horn neurons in vitro: correlation with cutaneous sensory afferent input. Eur J Neurosci 6: 998–1007, 1994.[CrossRef][Web of Science][Medline]
Lu VB, Moran TD, Balasubramanyan S, Alier KA, Dryden WF, Colmers WF, Smith PA. Substantia gelatinosa neurons in defined-medium organotypic slice culture are similar to those in acute slices from young adult rats. Pain 121: 261–275, 2006.[CrossRef][Web of Science][Medline]
Magistretti J, Mantegazza M, Guatteo E, Wanke E. Action potentials recorded with patch-clamp amplifiers: are they genuine? Trends Neurosci 19: 530–534, 1996.[CrossRef][Web of Science][Medline]
Margrie TW, Sakmann B, Urban NN. Action potential propagation in mitral cell lateral dendrites is decremental and controls recurrent and lateral inhibition in the mammalian olfactory bulb. Proc Natl Acad Sci USA 98: 319–324, 2001.
Markram H, Helm PJ, Sakmann B. Dendritic calcium transients evoked by single back-propagating action potentials in rat neocortical pyramidal neurons. J Physiol 485: 1–20, 1995.
Mason P. Deconstructing endogenous pain modulations. J Neurophysiol 94: 1659–1663, 2005.
Melnick IV, Santos SFA, Safronov BV. Mechanism of spike frequency adaptation in substantia gelatinosa neurones of rat. J Physiol 559: 383–395, 2004a.
Melnick IV, Santos SFA, Szokol K, Szucs P, Safronov BV. Ionic basis of tonic firing in spinal substantia gelatinosa neurons of rat. J Neurophysiol 91: 646–655, 2004b.
Micheva KD, Smith SJ. Strong effects of subphysiological temperature on the function and plasticity of mammalian presynaptic terminals. J Neurosci 25: 7481–7488, 2005.
Mitterdorfer J, Bean BP. Potassium currents during the action potential of hippocampal CA3 neurons. J Neurosci 22: 10106–10115, 2002.
Molineux ML, Fernandez FR, Mehaffey WH, Turner RW. A-type and T-type currents interact to produce a novel spike latency-voltage relationship in cerebellar stellate cells. J Neurosci 25: 10863–10873, 2005.
Prescott SA, De Koninck Y. Four cell types with distinctive membrane properties and morphologies in lamina I of the spinal dorsal horn of the adult rat. J Physiol 539: 817–836, 2002.
Ruscheweyh R, Ikeda H, Heinke B, Sandkühler J. Distinctive membrane and discharge properties of rat spinal lamina I projection neurones in vitro. J Physiol 555: 527–543, 2004.
Ruscheweyh R, Sandkühler J. Lamina-specific membrane and discharge properties of rat spinal dorsal horn neurones in vitro. J Physiol 541: 231–244, 2002.
Russier M, Carlier E, Ankri N, Fronzaroli L, Debanne D. A-, T-, and H-type currents shape intrinsic firing of developing rat abducens motoneurons. J Physiol 549: 21–36, 2003.
Safronov BV. Spatial distribution of NA+ and K+ channels in spinal dorsal horn neurones: role of the soma, axon and dendrites in spike generation. Prog Neurobiol 59: 217–241, 1999.[CrossRef][Web of Science][Medline]
Sceniak MP, MacIver MB. Cellular actions of urethane on rat visual cortical neurons in vitro. J Neurophysiol 95: 3865–3874, 2006.
Sokal DM, Chapman V. Effects of spinal administration of muscimol on C- and A-fibre evoked neuronal responses of spinal dorsal horn neurones in control and nerve injured rats. Brain Res 962: 213–220, 2003.[CrossRef][Web of Science][Medline]
Sontheimer H, Ransom CB. Whole-cell patch-clamp recordings. In: Patch-Clamp Analysis Advanced Techniques, edited by Waltz W, Boulton AA, Barker GB. Totowa, NJ: Humana Press, 2002, p. 35–68.
Stuart GJ, Dodt H-U, Sakmann B. Patch clamp recordings from the soma and dendrites of neurones in brain slices using infrared video microscopy. Pflügers Arch 423: 511–518, 1993.[CrossRef][Web of Science][Medline]
Sugiura Y, Lee CL, Perl ER. Central projections of identified, unmyelinated (C) afferent fibers innervating mammalian skin. Science 234: 358–361, 1986.
Thompson SM, Masukawa LM, Prince DA. Temperature dependence of intrinsic membrane properties and synaptic potentials in hippocampal CA1 neurons in vitro. J Neurosci 5: 817–824, 1985.[Abstract]
Thomson AM, West DC, Headley PM. Membrane characteristics and synaptic responsiveness of superficial dorsal horn neurons in a slice preparation of adult rat spinal cord. Eur J Neurosci 1: 479–488, 1989.[CrossRef][Web of Science][Medline]
Tuckett RP, Wei JY. Response to an itch-producing substance in cat. II. Cutaneous receptor populations with unmyelinated axons. Brain Res 413: 95–103, 1987.[CrossRef][Web of Science][Medline]
Vallbo AB, Olausson H, Wessberg J. Unmyelinated afferents constitute a second system coding tactile stimuli of the human hairy skin. J Neurophysiol 81: 2753–2763, 1999.
Varga AW, Yuan L-L, Anderson AE, Schrader LA, Wu G-Y, Gatchel JR, Johnston D, Sweatt JD. Calcium-calmodulin-dependent kinase II modulates Kv4.2 channel expression and upregulates neuronal A-type potassium currents. J Neurosci 24: 3643–3654, 2004.
Volgushev M, Vidyasagar TR, Chistiakova M, Yousef T, Eysel UT. Membrane properties and spike generation in rat visual cortical cells during reversible cooling. J Physiol 522: 59–76, 2000.
Vydyanathan A, Wu ZZ, Chen SR, Pan HL. A-type voltage-gated K+ currents influence firing properties of isolectin B4-positive but not isolectin B4-negative primary sensory neurons. J Neurophysiol 93: 3401–3409, 2005.
Yoshimura M, Jessell TM. Membrane properties of rat substantia gelatinosa neurons in vitro. J Neurophysiol 62: 109–118, 1989.
This article has been cited by other articles:
|
|
 |
|
  M. A. Walsh, B. A. Graham, A. M. Brichta, and R. J. Callister Evidence for a Critical Period in the Development of Excitability and Potassium Currents in Mouse Lumbar Superficial Dorsal Horn Neurons J Neurophysiol, April 1, 2009; 101(4): 1800 - 1812. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
80 pA above Rh in 20-pA increments (i.e., Rh20, Rh40, Rh60, and Rh80). The slope, or gain, of this relationship is increased at PT (i.e., higher firing frequencies are achieved for the same current injection). B: stimulus current/AP-frequency relationship for initial bursting neurons at RT and PT. Left traces show examples of AP discharge in initial bursting neurons at RT and PT during injection of depolarizing current steps of increasing magnitude (bottom-most traces). Right plot shows stimulus current/AP-frequency plots of group data at RT (black squares n = 15) and PT (gray squares, n = 10). Firing frequency at rheobase is elevated at PT and the slope, or gain, of this relationship is also increased.
