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REPORT
Department of Physiology and Biophysics, School of Medicine, University of Washington, Seattle, Washington
Submitted 11 December 2006; accepted in final form 17 May 2007
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ABSTRACT |
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 TOP ABSTRACT INTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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INTRODUCTION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). PICs appear to be mediated by membrane channels located primarily on the dendrites as evidenced by a clockwise hysteresis in the whole cell inward currents recorded in response to depolarizing then repolarizing voltage ramp commands, i.e., the deactivation of the inward current on the repolarizing ramp occurs at a lower somatic voltage than activation on the depolarizing ramp (Carlin et al. 2000; Lee and Heckman 1998; Powers and Binder 2003; Svirskis and Hounsgaard 1997). This phenomenon is thought to arise from the fact that the voltage-dependent channels carrying the inward current are electrically distant from the soma and not under voltage-clamp control. As a result, the distal channels are activated at relatively high somatic voltages during the depolarizing voltage ramp command but then continue to supply current to the soma during the repolarizing ramp command (cf. Booth et al. 1997; Carlin et al. 2000; Lee and Heckman 1998). Other signs that distal dendritic channels may make a significant contribution to inward voltage-clamp currents are a delayed current onset in response to a depolarizing voltage-clamp step (Booth et al. 1997; Carlin et al. 2000; Muller and Lux 1993; Powers and Binder 2003) and a prolonged decay following the termination of the step (Carlin et al. 2000; Powers and Binder 2003).
It is also possible that many of the features of PICs might result from a depolarization-induced facilitation of Cav1 channels that is independent of their location as reported in hippocampal neurons (Kavalali and Plummer 1994), hippocampal pyramidal neurons (Cloues et al. 1997), embryonic spinal motoneurons (Hivert et al. 1999), and cerebellar granule cells (Koschak et al. 2007). The "anomalous gating" of the Cav1 channels, consisting of prolonged channel reopenings during repolarizations after strong depolarizations, has been well characterized (Forti and Pietrobon 1993; Hivert et al. 1999), but the molecular mechanisms remain unresolved (see Koschak et al. 2007; Perrier et al. 2002 for discussion).
The goal of this study was to determine the extent to which the kinetics of calcium channels in isolated somata might account for some of the characteristics of the PICs observed in rat hypoglossal motoneurons (Powers and Binder 2003). To do so, we used the "nucleated patch" technique (Martina and Jonas 1997) to isolate a sphere (
20 µm) of somatic membrane from the rest of the cell. Our novel finding was that, contrary to the "remote dendritic hypothesis," nucleated patch recordings can also display a clockwise hysteresis in the whole cell calcium currents recorded in response to voltage ramp commands as well as delayed onsets and prolonged tail currents after voltage-clamp steps. Further, these somatic PICs display "facilitation" in response to conditioning depolarization as previously observed in whole cell recording from intact neurons (Svirskis and Hounsgaard 1997). These results suggest that many of the characteristics of PICs recorded in rat hypoglossal motoneurons may be mediated by the intrinsic properties of the Cav1 channels in these cells and may not be dependent on their spatial distribution.
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METHODS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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The general experimental and surgical procedures used here have been detailed in recent publications from our laboratory (Powers and Binder 2003; Zeng et al. 2005). Experiments were carried out in accordance with the animal-welfare guidelines at the University of Washington. Sprague-Dawley rats (15–18 days old) of either sex were anesthetized by injection of 1.8 ml/kg im of a 5:1.6:6.6 solution of ketamine:xylazine:saline. When fully anesthetized, the animals were decapitated. To study hypoglossal motoneurons, a section of brain stem was removed and glued to a Plexiglas tray filled with cooled, modified artificial cerebrospinal fluid (ACSF). A series of transverse slices 250 µm thick were cut throughout the length of the hypoglossal nucleus, transferred to a holding chamber and incubated at room temperature (19–21°C) for 30 min in the modified ACSF followed by incubation in standard ACSF.
Solutions and chemicals
The modified ACSF solution contained (in mM) 220 sucrose, 3 KCl, 1.25 NaH2PO4, 26 NaHCO3, 2 MgCl2, 2 CaCl2, and 10 D-glucose. Kynurenic acid (1 mM) and sodium lactate (4 mM) are added to improve cell viability. The standard ASCF is identical to that of the modified ACSF except that 120 mM NaCl is substituted for sucrose, and kynurenic acid is omitted.
Various channel blockers or agonists were added to the standard ACSF. In all experiments, potassium channel blockers (4 mM 4-AP; 10 mM TEACl) were used. The following were applied from stock solution aliquots: FPL 64176 (10 µM prepared in DMSO; Sigma), nifedipine (10 µM prepared in absolute ethanol; Sigma), TTX (1 µM prepared in Pi-free recording ACSF; Molecular Probes), 
-agatoxin IVA (480 nM; Alomone Labs), and -conotoxin GIVA (4 µM; Bachem). These toxin concentrations are saturating in this preparation (Umemiya and Berger 1994). To study isolated inward currents, the pipette solution was composed of (in mM) 100 CsCl, 20 TEACl, 5 MgCl2, 2 bis-(o-aminophenoxy)-N,N,N',N' tetraacetic acid (BAPTA), 10 HEPES, 5 Na2ATP, 0.5 Na3GTP, and CsOH/HCl for pH 7.3.
Measurement and recording techniques
Whole cell and nucleated patch recordings were obtained from the somata of rat hypoglossal motoneurons using a Zeiss Axioskop equipped with Nomarski optics for differential interference contrast and infrared video recording. Motoneuron identity was based on anatomical location and the similarity of intrinsic properties to our previous samples (Powers and Binder 2003; Sawczuk et al. 1997; Zeng et al. 2005). The patch electrodes were glass pipettes with tip diameters of 1–2 µM and resistances of 3–6 M
positioned with a Sutter MP-285 micromanipulator. Electrical recordings were made with an Axon Instruments Multiclamp 700A amplifier, digitized at 10 kHz using an Instrutech A/D board connected to a Macintosh PowerPC. Data acquisition and voltage-clamp commands were controlled by custom software routines running in Igor (WaveMetrics).
After the establishment of whole cell recording, the membrane potential was clamped at –70 mV. Whole cell currents were measured in response to triangular voltage-clamp commands (28 mV/s) from –70 to 0 mV and back and to a series of 1-s voltage-clamp steps delivered at 4-s intervals, each increasing in amplitude by 10 mV beginning at –70 mV.
After the whole cell measurements had been made, the transition to nucleated patch was attempted. To do so, a light negative pressure was applied to the recording pipette, and simultaneously the pipette was retracted slowly (20 µ/s) from the soma at an angle of 37°. Once the pipette and attached somatic membrane neared the surface of the slice, the pipette was retracted vertically (100 µ/s) out of the plane of the slice. Success was assessed visually by the appearance of a sphere of membrane on the end of the pipette, which could be moved freely above the slice. Isolation was confirmed electrophysiologically by a dramatic decrease in input conductance. Once the nucleated patch recording configuration was established, another set of voltage-clamp triangular ramps and a family of 1-s voltage steps were obtained as in whole cell mode. As the nucleated patches were typically 10–20 µ in diameter and the series resistances of the patch electrodes ranged from 4.4 to 10.6 M, the voltage-clamp control was excellent, with discrepancies between the command and the actual applied voltages of <5 mV.
After these basic measurements had been made, various other tests were performed. In some instances, a set of steps with a prepulse to –50 mV were performed to inactivate Cav3 channels (Umemiya and Berger 1994). Tests for facilitation were performed by varying the amplitude and duration of the prepulses, with 10- to 15-s intervals between trials.
All records were leak-subtracted off-line based on scaling the responses to voltage changes within 10 mV of holding potential. The liquid junction potential was calculated using the Henderson liquid junction potential equation and mobilities (Barry and Lynch 1991) to be 3.2 mV, so that a command potential of –70 mV, for example, would have actually been –73.2 mV. The reported values are the potentials used during experiments and do not reflect the correction for the junction potential. Mean values for tail currents are reported for trials in which a response occurred. The durations of the tail currents were quantified by first filtering the records using a fourth order Butterworth filter with a cutoff frequency of 50 Hz, then finding the times at which the derivatives of the smoothed signals exceeded a threshold value of 40 pA/s. In the recordings that did not have step-like tails, we set the threshold at the point when the current relaxed to 63% of its maximum value following the voltage step. Data were analyzed using Matlab (The Mathworks, Natick MA).
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RESULTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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) in the amplitude and time course of the inward current.
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During the application of 1-s voltage-clamp steps, both the whole cell and nucleated patch recordings displayed sustained inward calcium currents as previously described (Powers and Binder 2003). In many instances, at the end of the voltage step, there was a prolonged tail current in both whole cell and nucleated patch recordings that gradually dissipated. Moreover, in eight of the nucleated patches, after the voltage step was completed and the holding potential had returned to –70 mV, there was a pronounced step-like tail current that remained relatively stable for a prolonged duration followed by an abrupt cessation (Fig. 2B). The threshold for activation of these tail currents varied by cell, with the average being –27.1 mV, ranging from –50 to –20 mV. The duration of the tail currents often increased with the amplitude of the step, although the largest amplitudes of step currents often occurred near threshold for activation. The area of the inward current during the step was associated positively with the mean amplitude of the tail currents (r = 0.73; P < 0.02).
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). The –50-mV prepulses had no significant effect on the time course of the tail currents although in this example the delayed current onset in response to the test pulse was somewhat more pronounced (Fig. 2, D vs. B). However, delayed current onsets were also seen in the absence of a conditioning prepulse (e.g., upper black trace in Fig. 3A).
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In several experiments, we obtained data in the presence of various calcium channel agonists and antagonists. We made four nucleated patch recordings during the application of the Cav1 channel antagonist nifedipine. We also recorded the calcium currents in four cases before and after the addition of -conotoxin and -agatoxin to block Cav2.2 and Cav2.1 calcium channels, respectively. In five experiments, we examined the effects of adding the Cav1 channel agonist FPL 64176 to the bath. The results, as shown in Fig. 3, suggest that the prominent inward currents and prolonged tail currents observed in nucleated patches are carried primarily by somatic Cav1 channels. In all cases, the tail currents and the clockwise hysteresis evident in triangular voltage-clamp ramps (e.g., Fig. 1C) were obliterated or significantly reduced after addition of nifedipine (Fig. 3A, red: nifedipine, black: control). In the presence of FPL 64176, voltage steps that failed to evoke tail currents under control conditions (data not shown) evoked tail currents lasting 
9 s (Fig. 3C; cf. Fig. 2B, recorded the same nucleated patch before the addition of FPL) and the hysteresis was enhanced. The addition of Cav2.1 and Cav2.2 channel blockers produced a minor reduction in net inward current but did not eliminate the tail currents (Fig. 4B, red: conotoxin + agatoxin, black: control) or the hysteresis.
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We studied the facilitation of calcium tail currents after prepulses of varying durations and amplitudes in three of the nucleated patch experiments (cf. Cloues et al. 1997; Hivert et al. 1999; Koschak et al. 2007). We define facilitation here as the presence of a tail current after a step to a voltage that did not exhibit tail currents in the absence of a conditioning prepulse. Variations in the amplitude of short (200 ms) prepulses to +10, 0, –10, or –20 mV altered the response during a test pulse to –30 mV for 500 ms. The test pulses after the prepulses to +10, 0, and –10 mV were all followed by tail currents, whereas the prepulse to –20 mV did not evoke a tail current (Fig. 4A, black trace). The prepulse to +10 mV evoked the largest amplitude tail current, but the duration was longest for the prepulse to 0 mV (Fig. 4A, blue and red, respectively). To examine the effects of prepulse duration, a conditioning voltage step to –30 mV for 1 s, 500 ms, or 200 ms was followed by a subsequent step to –20 mV for 500 ms (Fig. 4B). After the 1-s and 500-ms steps (red and blue traces) there was an appreciable inward current during the step to –20 mV, which was not the case following the 200-ms prepulse (black trace). The inward current during the test step to –20 mV was similar for the 1-s and 500-ms conditioning pulses. Interestingly, the tail currents after the 1-s prepulse test were longer than after the 500-ms prepulse.
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DISCUSSION |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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). Thus it was assumed that PICs with prolonged tails could not be generated by somatic channels under voltage-clamp because these proximal channels would be held below threshold for activation on repolarization. Instead PICs were assumed to be generated by current flow through dendritic channels that are too far from the electrode to be under perfect voltage-clamp control. Second, the slow onset of the inward current suggested that time was required for voltage to spread to the site of the channels (dendrites) before they were activated (cf. Carlin et al. 2000). Third, model motoneurons can be made to exhibit inward currents during voltage clamp by placing "hot spots" of calcium channels 400 µ from the soma (Bui et al. 2006; Elbasiouny et al. 2005; Taylor and Enoka 2004).
The results reported here suggest the possibility that many of the features of PICs in rat hypoglossal motoneurons (Powers and Binder 2003) might result from a depolarization-induced facilitation of voltage-gated calcium channels that is independent of their location as reported in other types of neurons (Cloues et al. 1997; Hivert et al. 1999; Kavalali and Plummer 1994; Koschak et al. 2007). The anomalous gating of the Cav1 channels, consisting of prolonged channel reopenings during repolarizations after strong depolarizations has been well characterized (Forti and Pietrobon 1993; Hivert et al. 1999) and could account for both the hysteresis and long tail currents. However, the tail currents and facilitation we observed in our nucleated patches could be induced with depolarizing pulses of significantly lower amplitude than those reported previously (vide supra).
The identity of the calcium channels responsible for the observed currents was also of interest to us. Our lab previously found that the calcium current in rat hypoglossal motoneurons seems to be carried predominately by Cav2.1 and 2.2 channels, with a smaller contribution from Cav1 channels (Powers and Binder 2003). Yet the present data suggest that the tail currents we observed were primarily carried by Cav1 calcium channels. The tail currents were nearly completely blocked by nifedipine and greatly enhanced by FPL, and the addition of Cav2.1 and 2.2 channel blockers did not have a significant effect on the tail currents. Furthermore we have immunohistochemical evidence for the presence of both Cav1.2 and 1.3 channels on the somata of rat hypoglossal motoneurons (Westenbroek et al. 2005). One explanation for the difference may be that in whole cell mode, when the current being measured is influenced by both somatic channels and dendritic channels, a larger proportion of dendritic Cav2.2 and 2.2 calcium channels such as we observed previously would mask the influence of somatic Cav1 channels. If this was true, only in isolated patches of somatic membrane could we observe the activity of Cav1 channels as reported here. The fact that we did not observe the step-like tails in four cells suggests that not all hypoglossal motoneurons have the same distribution of somatic Cav1 channels, consistent with our preliminary immunocyctochemical data (Westenbroek et al. 2005).
Although our data clearly show that PICs can be generated at the channel level and do not require a dendritic source, dendritic calcium channels are clearly important nonetheless. By virtue of their sheer number, dendritic calcium channels must be the primary source of PICs in motoneurons. The most parsimonious assumption would be that dendritic calcium channels exhibit the same properties observed here for the somatic channels. Thus we would speculate that the increased efficacy of synaptic input over injected somatic current in activating PICs (Bennett et al. 1998; Lee and Heckman 2000) reflects the greater surface area of the dendrites and the associated larger number of dendritic calcium channels and excitatory synaptic inputs (Cushing et al. 2005; Powers and Binder 2001).
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GRANTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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ACKNOWLEDGMENTS |
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TOPABSTRACTINTRODUCTIONMETHODSRESULTSDISCUSSIONGRANTSACKNOWLEDGMENTSREFERENCES |
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FOOTNOTES |
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Address for reprint requests and other correspondence: M. D. Binder, Dept. of Physiology and Biophysics, University of Washington School of Medicine, Box 357290, Seattle, WA 98195-7290 WA (E-mail: mdbinder{at}u.washington.edu)
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REFERENCES |
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Bennett DJ, Hultborn H, Fedirchuk B, Gorassini M. Synaptic acyivation of plateaus in hindlimb motoneurons of the decerebrate cat. J Neurophysiol 80: 2023–2037, 1998.
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Cushing S, Bui T, Rose PK. Effect of nonlinear summation of synaptic currents on the input-output properties of spinal motoneurons. J Neurophysiol 94: 3465–3478, 2005.
Elbasiouny SM, Bennett DJ, Mushahwar VK. Simulation of dendritic CaV1.3 channels in cat lumbar motoneurons: spatial distribution. J Neurophysiol 94: 3961–3974, 2005.
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Kavalali ET, Plummer MR. Selective potentiation of a novel calcium channel in rat hippocampal neurones. J Physol 480: 475–784, 1994.
Koschak A, Obermair GJ, Pivotto F, Sinnegger-Brauns MJ, Striessnig J, Pietrobon D. Molecular nature of anomalous L-type calcium channels in mouse cerebellar granule cells. J Neurosci 27: 3855–3863, 2007.
Lee RH, Heckman CJ. Bistability in spinal motoneurons in vivo: systematic variations in persistent inward currents. J Neurophysiol 80: 583–593, 1998.
Lee RH, Heckman CJ. Adjustable amplification of synaptic input in the dendrites of spinal motoneuron in vivo. J Neurosci 20: 6734–6740, 2000.
Martina M, Jonas P. Functional differences in Na+ channel gating between fast-spiking interneurons and principal neurons of rat hippocampus. J Physiol 505: 593–603, 1997.
Muller W, Lux HD. Analysis of voltage-dependent membrane currents in spatially extended neurons from point-clamp data. J Neurophysiol 69: 241–247, 1993.
Perrier JF, Alaburda A, Hounsgaard J. Spinal plasticity mediated by postsynaptic L-type Ca2+ channels. Brain Res Brain Res Rev 40: 223–229, 2002.[CrossRef][Medline]
Powers RK, Binder MD. Input-output functions of mammalian motoneurons. Rev Physiol Biochem Pharmacol 143: 137–263, 2001.[Web of Science][Medline]
Powers RK, Binder MD. Persistent sodium and calcium currents in rat hypoglossal motoneurons. J Neurophysiol 89: 615–624, 2003.
Sawczuk A, Powers RK, Binder MD. Contribution of outward currents to spike-frequency adaptation in hypoglossal motoneurons of the rat. J Neurophysiol 78: 2246–2253, 1997.
Soong TW, DeMaria CD, Alvania RS, Zweifel LS, Liang MC, Mittman S, Agnew WS, Yue DT. Systematic identification of splice variants in human P/Q-type channel alpha1(2.1) subunits: implications for current density and Ca2+-dependent inactivation. J Neurosci 22: 10142–10152, 2002.
Svirskis G, Hounsgaard J. Depolarization-induced facilitation of a plateau-generating current in ventral horn neurons in the turtle spinal cord. J Neurophysiol 78: 1740–1742, 1997.
Taylor AM, Enoka RM. Quantification of the factors that influence discharge correlation in model motor neurons. J Neurophysiol 91: 796–814, 2004.
Umemiya M, Berger AJ. Properties and function of low- and high-voltage-activated Ca2+ channels in hypoglossal motoneurons. J Neurosci 14: 5652–5660, 1994.[Abstract]
Westenbroek RE, Newkirk GS, Powers RK, Binder MD. Distribution of Cav1.2, Cav1.3, Cav2.1, and Cav2.2 channels on the somata and dendrites of rat hypoglossal motoneurons. Soc Neurosci Abstr 31: 750.11, 2005.
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