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Department of Neurobiology, Center for Glial Biology in Medicine, University of Alabama, Birmingham, Alabama
Submitted 27 March 2007; accepted in final form 30 May 2007
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ABSTRACT |
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INTRODUCTION |
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). However, these values fluctuate due to K+ release into the extracellular space during neuronal activity. Because of the small volume of the extracellular space and low baseline levels of [K+]o, the release of even small amounts of K+ can lead to dramatic increases in extracellular K+ concentrations (Van Harreveld and Malhotra 1967). It has been demonstrated in the rat spinal cord that a single stimulus can increase extracellular K+ by as much as 5 mM (Walton and Chesler 1988). Sustained increases in [K+]o above baseline levels leads to hyperexcitability and affects the integrity of synaptic transmission (Walz 2000). Imperatively, several mechanisms are in place that aid in extracellular K+ clearance including simple diffusion and energy-dependent mechanisms such as glial and neuronal Na+-K+ pumps (Amedee et al. 1997; Kofuji and Newman 2004). Additionally, K+-selective channels, largely in glial cells, assist in K+ clearance by moving K+ ions across their membrane when extracellular K+ concentrations increase. Potassium spatial buffering, as is termed, is thought to redistribute K+ to surrounding glia cells via gap junctions where it is released at sites of lower [K+]o. Potassium spatial buffering is an attractive mechanism for K+ clearance as it is energy efficient and sequesters potassium in the intracellular space. However, energy-dependent mechanisms of K+ clearance have been implicated in potassium uptake as well (D'Ambrosio et al. 2002; MacVicar et al. 2002; Ransom et al. 1995; Xiong and Stringer 1999). It is likely that all three mechanisms contribute to some extent to K+ clearance after neuronal activity.
Glial cell membranes are endowed with K+ channels perfectly suited for the task of K+ clearance. Kir4.1, an inwardly rectifying K+ channel, has garnered much attention. This channel is expressed in glial cells throughout the CNS (Kofuji et al. 2000; Martin-Caraballo and Dryer 2002; Olsen et al. 2006; Poopalasundaram et al. 2000). Kir4.1 channels have a high open probability at rest (Ransom and Sontheimer 1994), therefore contributing to the high K+ permeability and negative resting membrane potential of astrocytes. Importantly, channel conductance increases with increasing extracellular K+ (Hagiwara and Takahashi 1974; Newman 1993; Sakmann and Trube 1984), making Kir4.1 ideally suited for K+ clearance. The channel's significance in the context of extracellular K+ regulation and its contribution to the hyperpolarized resting membrane potential, a hallmark of mature astrocytes, has been conclusively demonstrated in animals where Kir4.1 has been genetically inactivated. In Kir4.1 knock-out animals, Muller cells (Kofuji et al. 2000), astrocytes of the ventral respiratory group (Neusch et al. 2006), and spinal cord astrocytes (Olsen et al. 2006) lack inwardly rectifying K+ currents. Furthermore, K+ uptake, or clearance capabilities, is decreased, the resting membrane potential is depolarized, and input resistance is markedly increased.
In spinal cord, the dynamics of [K+]o are well documented (Frankenhauser and Hodgkin 1956; Jendelova and Sykova 1991; Walton and Chesler 1988). Importantly, striking regional differences in K+ accumulation were seen after neuronal activity (Walton and Chesler 1988). Specifically, neuronal activity in the dorsal horn causes larger increases in [K+]o than in the ventral horn. We demonstrate here that Kir4.1 is more prominently expressed in the ventral horn and after K+ stimulation gives rise to much enhanced uptake currents. Hence changes in the expression of Kir4.1 appear to translate into marked differences in extracellular K+ homeostasis.
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METHODS |
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Experiments were performed on either sex of Sprague-Dawley rats and were approved by the University of Alabama Institutional Animal Care and Use Facility. For slice experiments, P12–P20 pups were anesthetized with CO2 and decapitated, and the spinal cord was removed and placed in ice-cold calcium-free artificial cerebral spinal fluid (ACSF, containing in mM, 116 NaCl, 4.5 KCl, 0.8 MgCl2, 26.2 NaHCO3, 11.1 glucose, and 5.0 HEPES) for 30 s. The cord was then placed in liquid low-melt agar at 
30°C, which was quickly solidified by placing on ice. Sections from the cervical, thoracic, and lumbar regions were cut at 250 µm using a Vibratome 3000 (Ted Pella, Redding, CA) in calcium-free ACSF. Before recording, slices were allowed to recover for 
1 h at room temperature in calcium-free ACSF, which was continuously bubbled with 5% CO2-95%O2.
Slice electrophysiology
Whole cell voltage-clamp recordings were made as described previously (Olsen et al. 2006). Patch pipettes were made from thin-walled (outer diameter 1.5 mm, inner diameter 1.12 mm) borosilicate glass (TW150F-4) WPI, Sarasota, FL) and had resistances of 4–6 M
. Slices were transferred after the recovery period to either a Zeiss Axiskop FS microscope (Zeiss, Thornwood, NY) or Leica DMLFSA microscope equipped with Nomarski optics; a x40 water-immersion lens and infrared illumination was used to visualize glia cells. Signals were acquired using an Axopatch 700B (Molecular Devices, Sunnyvale, CA) or a Axopatch 200B amplifier, both controlled by Clampex 9.0 software via a Digidata 1200B interface (Axon Instruments). Signals were filtered at 2 kHz and digitized at 5 kHz. Data acquisition and storage were conducted with the use of pClamp 9.0 (Axon Instruments). Resting membrane potentials were measured directly from the amplifier in I = 0 mode 1 min after whole cell access was obtained. Where described in the text whole cell capacitance and series resistances were also measured directly from the amplifier, with the upper limit for series resistance being 10 M and series resistance compensation adjusted to 80% to reduce voltage errors. Slices were continuously superfused with ACSF with the addition of 2.0 mM CaCl2.
The standard KCl pipette solution contained (in mM) 145 KCl, 1 MgCl2, 10 EGTA, and 10 HEPES sodium salt, pH adjusted to 7.3 with Tris-base. CaCl2 (0.2 mM) was added to the pipette solution just before recording, resulting in a free calcium concentration of 1.9 nM. In some experiments, biocytin (0.5 mg/ml) was added to the pipette solution for posthoc identification of recorded cells. Cells were continuously superfused at room temperature with oxygenated ACSF containing 2 mM CaCl2. Drugs were added directly to these solutions. Unless stated otherwise, all drugs were purchased from Sigma (St. Louis, MO).
Western blot analysis
Thick spinal cord slices (1–2 mm) were obtained as described in the preceding text and placed in ice-cold ACSF supplemented with protease inhibitors. Slices were cooled to aid in the removal of the white matter. Gray matter tissue from the ventral or dorsal horn from several serial sections of the spinal cord were placed in lysis buffer (100 mM Tris, pH 7.5, 1% SDS supplemented with protease inhibitors). The tissue was mechanically homogenized and then sonicated for 10 s. Tissue homogenates were centrifuged for 5 min at 12,000 g at 4°C. Protein quantification was performed on the supernatant using a DC protein assay kit from Bio-Rad (Hercules, CA). Protein was heated to 60°C for 15 min in an equal volume of 2x sample buffer (100 mM Tris, pH 6.8, 10% SDS in Laemmli-sodium dodecylsulfate, 600 mM β-mercaptoethanol). Equal amounts of protein were loaded into each lane of a 4–20% gradient precast sodium dodecylsulfate polyacrilamide gel electrophoresis (Bio-Rad). Gels were transferred onto PVDF paper (Millipore, Bedford, MA) at 200 mA constant for 2 h at room temperature and membranes were blocked in blocking buffer (5% dried milk, 2%BSA and 2% goat-serum). The Kir4.1antibody was obtained from Alomone (Jeruselum, Israel) and diluted according to the manufacturer's instructions. Blots were incubated for 90 min at room temperature. The membranes were then rinsed three times for 15 min and incubated with horseradish peroxidase-conjugated secondary antibody for 60 min. Blots were once again washed three times for 10 min and developed with enhanced chemiluminesence (Amersham, Arlington Heights, IL) on Hyperfilm (Amersham).
Immunocytochemistry
Animals (P12-20) were killed with a peritoneal injection of ketamine (100 mg/kg) and were perfused with a 4% paraformaldehyde solution for 15 min. The whole spinal column was removed and stored in 4% paraformaldehyde overnight. After washing in PBS, the cord was removed from the column, 50-µm sections were cut using a Vibratome (Oxford Instruments) from the mid-cervical, thoracic, and lumbar regions. Sections were blocked for 1 h in 10% horse serum and 0.3% Triton-X100 in PBS. Primary antibodies were diluted according to manufacturer's instructions in blocking buffer diluted 1:3 and incubated with slices overnight. Antibodies against GLT-1, GFAP, Neu-N, and MAP2 were obtained from Chemicon (Temecula, CA). We used GFAP as a positive marker for astrocytes, recognizing that GFAP does not label all astrocytes. For this reason, we also chose to label astrocytic membrane with the astrocyte-specific membrane marker GLT-1. The following day, the sections were washed two times with PBS and two times with diluted blocking buffer before incubating with FITC or TRITC-conjugated secondary antibodies obtained from Molecular Probes for 60 min at room temperature. Slices were then washed two times with diluted blocking buffer and two times with PBS before being mounted onto glass coverslips. Fluorescent images were acquired with a Zeiss Axiovert 200M (München, Germany). To quantify Kir4.1 expression low-magnification (x5) images were imported into the National Institutes of Health imaging software program ImageJ1.37C. Intensity measurements were made in selected regions of interest corresponding to Rexed's Laminae IX (region1), Rexed's Laminae VII (region 2), Rexed's Laminae V (region 3), and Rexed's Laminae I/II (region 4). Comparisons were matched as we were able to visualize all four regions in each image at one time controlling for exposure times within each individual image.
Statistical analysis
Current responses to varied voltage steps and ramps were analyzed and measured in Clampfit (Molecular Devices); the resulting raw data were graphed and plotted in Origin 6.0 (MicroCal, Northampton, MA). Two-tailed t-test and Tukey-Kramer Multiple Comparisons Test were performed using Graphpad software (San Diego, CA) and P values are reported in the text. Unless otherwise stated, all values are reported as means ± SE with n indicating the number of cells sampled.
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RESULTS |
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; Neusch et al. 2001; Olsen et al. 2006). Higher-magnification images demonstrate Kir4.1 reactivity in the ventral horn was the most intense in regions surrounding neuronal cell bodies and along axons (arrowheads, Fig. 1G) as well as along capillaries (not shown). These data demonstrate marked heterogeneity of Kir4.1 in astrocytes of the spinal cord with highest expression of Kir4.1 in the ventral horn. In an attempt to quantify Kir4.1 in transverse sections of the spinal cord, we investigated 21 low-magnification images in which we were able to visualize both horns in the same image (Fig. 1E). We focused on four particular regions Rexed's Laminae IX (region1), Rexed's Laminae VII (region 2), Rexed's Laminae V (region 3), and Rexed's Laminae I/II (region 4). Mean data from 21 images demonstrate significant differences in Kir4.1 expression between Rexed's Laminae 1/II and all other regions of the spinal cord examined (P < 0.001, Tukey-Kramer multiple comparisons test, Fig. 1F). Kir 4.1 expression in region 3 or Rexed's Laminae V is also significantly different from Rexed's Laminae 9 (n = 21, P < 0.001). These data suggest that Kir4.1 exhibits a gradient of immunoreactivity in neonatal rat spinal cord.
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50 and 200 kD corresponding to a monomer and tetramer of Kir4.1 (Fig. 2). Consistent with the immunohistochemistry results, less Kir4.1 immunoreactivity was detected in the dorsal lanes from two P16 rats. Re-probing the same blot with two astrocytic markers GLT-1 and GFAP indicate that astrocytic number or percentage of astrocytic membrane/milligram of total protein were the same in the dorsal and ventral regions; indeed GLT-1 expression appears slightly higher in the dorsal regions. Hence the differences in Kir4.1 expression cannot be due to an overall difference in astrocyte number in these two regions. To demonstrate that equal concentrations of protein were loaded, this blot was probed with actin. Human embryonic kidney (HEK) cells, which do not express Kir4.1, served as a negative control. We presume the band at 100 kD to be nonspecific binding as it is also present in the HEK cell lane.
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). Of the six cells that were biocytin filled from the ventral horn, only one cell demonstrated significant cell coupling. In the dorsal horn, 8 of the 11 cells visualized by biocytin labeling demonstrated extensive cell coupling. Representative examples of biocytin-filled cells are illustrated in Fig. 3C.
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; Ransom and Sontheimer 1995; Sontheimer et al. 1989; Williamson et al. 1997). Consistent with these observations, mean data demonstrate astrocytes from the dorsal horn are slightly, but significantly, more depolarized relative to astrocytes from the ventral horn (–67.4 ± 2.4, n = 33, and –76.8 ± 1.1, n = 34, P = 0.0008, Fig. 5A). Furthermore, the input resistance (Fig. 5B) of cells from the dorsal horn was significantly higher (760.5 ± 171.9, n = 18) than astrocytes from the ventral horn (279.1 ± 60.9, n = 18, P = 0.0153). Kir channels have a high open probability at rest and therefore contribute to the relatively low input resistance seen in astrocytes. The higher input resistance observed in the dorsal horn astrocytes is likely a direct consequence of fewer Kir channels in these astrocytes.
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25 µm) from the cell of interest at the same focal plane and using pipette tips with similar diameters. We intentially placed the puffing pipette very close in proximity and pulsed with saturating concentrations of K+ to maximize the response in the target cell. Whole cell recordings were used to measure the K+-induced inward currents under voltage clamp at –80 mV. Representative examples are illustrated in Fig. 6A and mean data, normalized to cell size, is shown in Fig. 6B. Inward currents in response to a K+ puff were nearly twice the amplitude in the ventral horn when compared with the dorsal horn. In both regions, this response was almost completely eliminated in Ba2+ (not shown). As a means to quantify this data, we integrated the area under the curve before and after Ba2+ application. The subtracted area represents the Ba2+-sensitive charge flow, i.e., K+ ions fluxing through Kir channels during the pulse. These data were then normalized to whole cell capacitance to yield specific K+ flow/unit membrane. Mean data from 10 ventral horn and 5 dorsal horn cells examined this way are illustrated in Fig. 6B. These data demonstrate that on average, equal K+ challenges resulted in a 60% smaller K+ ion influx in the dorsal horn as compared with the ventral horn (31.9 ± 6.3 pC/pF, n = 10, and 11.9 ± 2.3 n = 5 pC/pF, respectively, P = 0.049). Taken together, these data suggest regional differences in astrocytic Kir4.1 expression leads to a differential ability to respond to extracellular [K+].
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DISCUSSION |
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; Neusch et al. 2006; Olsen et al. 2006) suggest Kir 4.1 is important for K+ homeostasis. A high open probability at rest (Po 0.9, (Ransom and Sontheimer 1995) and an increased conductance with increased extracellular K+ make Kir4.1 an ideal pathway for K+ clearance. In this study, we demonstrate that astrocytes from two different regions of the spinal cord, regions that have clear distinctions in terms of function, vary in regard to Kir4.1 channel expression. Our immunohistochemical studies demonstrate highest Kir4.1 expression in the ventral lateral edge of the ventral horn (Rexed Laminae 9) and show decreasing expression as toward the superficial layers of the spinal cord (Rexed Laminae I/II). Western blots confirm our immunohistochemical results demonstrating enhanced expression of Kir4.1 in the ventral horn of the spinal cord. Importantly, our recordings from astrocytes in acute spinal cord slices confirm this difference is reflected in function, i.e., a more than twofold difference in the magnitude of Ba2+-sensitive inwardly rectifying currents. This finding is significant as it correlates directly with, and may indeed explain previously observed differences in, extracellular K+ homeostasis in these regions of the spinal cord. Specifically, in neonatal rat a single electrical stimulus to the dorsal root evokes significant increases in [K+]o in the dorsal horn, whereas the same stimulus fails to produce noticeable [K+]o changes in the ventral horn (Walton and Chesler 1988). Furthermore, in the dorsal horn, a single orthodromic stimulation caused [K+]o increases by as much as 4–5 mM. Additionally, baseline extracellular K+ levels vary in this structure (Svoboda et al. 1988). Our data suggest this phenomenon may be directly explained by the reduced expression of Kir4.1 in astrocytes of the dorsal horn, and we show experimentally that these astrocytes have a significantly reduced ability to take up K+. As in the preceding studies, our experiments were performed in neonatal animals, therefore it remains to be shown whether similar differences in Kir4.1 channel expression would be observed in adult animals. Importantly, the preceding findings were not due to overall differences in astrocyte number as GFAP expression did not differ between these regions. Astrocytic membrane as detected by GLT-1 staining on our Western blot appeared slightly higher in the dorsal horn of the spinal cord. Moreover, this difference in Kir current amplitude was not due to an overall reduction of K+ channels in dorsal horn astrocytes as the relative expression of delayed rectifying and fast-inactivating K+ channels was identical in astrocytes from both regions.
These differences in K+ uptake may reflect differing demands on K+ homeostasis in these two regions of the spinal cord, and hence Kir4.1 expression may simply be tuned to the relative demand for K+ clearance. The ventral horn contains primarily motor pathways in which high-frequency electrical discharge is the norm. Hence K+ release by motor neurons would be substantial. The dorsal horn, on the other hand, contains primarily sensory pathways many of which contain slowly conducting unmyelinated fibers with comparably slower discharge rates. It should be noted that Kir4.1 is not the only protein showing regionally differing expression within the spinal cord. Aquaporin1 (Oshio et al. 2006), Aquaporin4 (Nesic et al. 2006; Vitellaro-Zuccarello et al. 2005), GLT-1, and GLAST (Rothstein et al. 1994) are each differentially expressed, showing higher levels of expression in the superficial layers of the dorsal horn.
In addition to its role in K+ clearance, Kir4.1 has been demonstrated to contribute to the negative resting membrane potential typical of most astrocytes. Astrocytes in which Kir4.1 is blocked with exogenously applied Ba2+ or astrocytes from Kir4.1 knock-out animals each rest at more depolarized potentials relative to normal astrocytes (Kofuji et al. 2000; Svoboda et al. 1988). A negative RMP is crucial for many transport systems operating in astrocytes including, glucose and neurotransmitter transport. A more negative resting membrane potential translates in a greater Na+ gradient across the membrane that is used as the energy source to drive these systems. Indeed, it was recently demonstrated that glutamate uptake in cultured cortical astrocytes decreased by >50% when cells were treated with siRNA to Kir4.1 (Kucheryavykh et al. 2007). In our study, astrocytes from the dorsal horn were slightly but significantly depolarized relative to ventral horn astrocytes. Hence glutamate uptake would be expected to be reduced unless cells showed a compensatory upregulation of transporter expression.
Recently, it was demonstrated that a progressive loss of Kir4.1 from astrocytes in the ventral horn parallels disease progression in an ALS mouse model (Kaiser et al. 2006). Moreover, when motor neurons were exposed to modest elevations in K+ (10 mM) for 120 h, significant neuronal cell death occurred. The high expression of Kir4.1 in this region suggest that neurons from the ventral horn may be particularly sensitive to elevated extracellular K+, and efficient clearance mechanisms are necessary for neuronal cell survival, again explaining the observed increases in Kir4.1 expression. In contrast, the reduced expression of Kir4.1 in the dorsal horn, the region of the spinal cord responsible for pain perception, may be of functional significance. Here it may indeed be beneficial to have a less effective system for K+ clearance, which allows K+ to increase during a painful or noxious stimulus. The increased K+ load resulting from increased "painful" neuronal activity in this region may amplify the signal by slightly depolarizing neurons in this region, hence, increasing their firing frequency proportional to the nociceptive stimulus. Prolonged subtle increases in K+ have been reported that are suggestive of lessened K+ regulation in the dorsal horn. In a recent study, after being applied to the hind paw of rats, it was demonstrated a sustained increase in [K+]o, by 
3 mM above baseline, in the lower dorsal horn that persisted for >2 h (Svoboda et al. 1988). This elevation was due to self-sustained neuronal firing induced by the injury. This [K+]o increase was graded with the intensity of the nociceptive stimulus. Clearly, these experiments demonstrate the spinal cord may indeed be a highly specialized structure regarding neuronal transmission and the physiological relevance of K+ homeostasis warrants further examination in this regard.
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GRANTS |
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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Address for reprint requests and other correspondence: H. Sontheimer, 1719 6thAve. S., CIRC 425, Birmingham, AL 35294 (E-mail: sontheimer{at}uab.edu)
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ABSTRACT
INTRODUCTION
