| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
|
|
|
|||||||
|
|||||||||
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
Neural and Vascular Biology Theme, Ordway Research Institute, Albany, New York
Submitted 28 April 2006; accepted in final form 7 June 2006
 |
ABSTRACT |
|---|
|
|
|
INTRODUCTION |
|---|
|
1 kDa (reviewed in Theis et al. 2005
). This interastrocyte coupling is thought to serve several of the many proposed functions of astrocytes, such as potassium buffering (Holthoff and Witte 2000; Kofuji and Newman 2004; Orkand et al. 1966; except also see Barres et al. 1990), calcium waves (Venance et al. 1995), and metabolic support to neurons (Farahani et al. 2005). However, little is known about the developmental time course of gap junction coupling and how it relates to the electrophysiological properties of astrocytes. We recently found, using whole cell in situ voltage-clamp recording and immunohistochemistry for the astrocyte marker GLAST, that the rat hippocampal astrocyte population shows changes in voltage-gated ion channel expression in the first 3 wk of postnatal development and ultimately appears exclusively as having a low membrane resistance and a nearly linear current–voltage (I–V) relationship (passive) after 3 postnatal weeks of development (Zhou et al. 2006). Because the passive astrocytes become the majority of GLAST positive cells, the other two types, the outwardly rectifying and variably rectifying astrocytes, characterized by their distinct I–V relationships (Zhou et al. 2006), become less abundant. During this same time period the interastrocytic coupling in the visual cortex, as measured by dye diffusion, increases (Binmoller and Muller 1992) and the astrocyte-expressed connexin 30 also increases (Nagy et al. 1999). If interastrocyte gap junctions conduct significant currents between cells and development of the syncytium is coincident with the observed electrophysiological changes, then it could be argued that the degree of cell coupling determines the passive electrophysiological phenotype by masking voltage-gated ion currents. In the present study we used patch-clamp electrophysiology and dye filling to measure the precise time course of the development of astrocyte dye coupling, the changes in expression of voltage gated ion channels, and apparent membrane resistances in GLAST(+) astrocytes observed from postnatal days 1 through 43. We also describe the electrophysiological phenotypes of excised patches from the different cells and the effect of the gap junction blocker meclofenamic acid (MFA). We conclude that the passive phenotype of mature astrocytes does not arise from the development of the astrocyte syncytium.
|
|
METHODS |
|---|
|
Experiments performed with rats were done in accordance with the regulations of the Wadsworth Center's Institutional Animal Care and Use Committee. Sprague–Dawley rats (Taconic, Germantown, NY) were fed ad libitum and housed in 12 h light/12 h dark cycles. All chemicals were purchased from Sigma (St. Louis, MO) except where noted otherwise. Rats of appropriate postnatal age (P1–P43) were anesthetized with CO2 and decapitated with a guillotine. Brains were quickly removed and adhered to the pan of a Pelco 1500 vibratome (Ted Pella, Redding, CA). Coronal 300-µm-thick slices were cut in ice-cold pH 7.4 slice preparation solution (containing, in mM: 26 NaHCO3, 1.25 NaH2PO4, 2.5 KCl, 10 MgCl2, 10 glucose, 0.5 CaCl2, 240 sucrose) and transferred to artificial cerebrospinal fluid (aCSF, containing in mM: 125 NaCl, 25 NaHCO3, 10 glucose, 3.5 KCl, 1.25 NaH2PO4, 2 CaCl2,1 MgCl2) at room temperature, and bubbled with 95% O2-5% CO2. Slices were allowed to recover for at least 1 h at room temperature in aCSF. Individual slices were transferred to a perfusion chamber at room temperature on an upright Olympus model BX51W1 microscope equipped with infrared differential interference contrast (DIC) optics and x10 and x40 objectives. Using the principal cell layers of the hippocampus for orientation, cells were selected in the CA1 stratum radiatum, stratum oriens, and stratum lacunosum-moleculare. Typically, cells with small (diameter <10 µm, as previously described in Zhou et al. 2006) cell bodies were patched. The electrode internal solution contained (in mM): 140 KCl, 0.5 Ca2Cl, 1 MgCl2, 5 EGTA, 10 HEPES, 3 Mg-ATP, and 0.3 Na-GTP (pH = 7.3, 280 ± 5 mOsm) and included 0.1–0.5% biocytin for locating the recorded cell and its associated syncytium for postrecording analyses. For electrode solution containing low intracellular chloride, 137 mM KCl was replaced by equimolar K-gluconate that resulted in a 6 mM total chloride inside the electrode solution. A MultiClamp 700A amplifier was used for all whole cell and patch recordings. The whole cell seals obtained were of 1–4 G
and all acquired data were handled by the PClamp9.0 program (Axon Instruments, Forster City, CA) running on a personal computer. Recordings were done at room temperature and each cell was patched for 
10 min. For glial electrophysiological phenotype identification, the whole cell recording was held at –70 mV. Before the delivery of each test voltage pulse, the cell was clamped to –110 mV for 300 ms. This prepulse allows maximal activation of voltage-gated transient outward K+ current (IKa) as well as inward Na+ current (INa+) (Zhou and Kimelberg 2000). The test pulses were of 50-ms duration and 10-mV increment, ranging from –180 to +40 mV, separated by a 1-s interval. Membrane resistance (Rm) as reported here refers to the Rm given by the "membrane test" function of the PClamp9.0 program. The cell was held at –70 mV, then a 10-ms pulse to –60 mV was applied. The membrane resistance was calculated from the current measured near the end of the 10-ms voltage command. At the end of each recording care was taken to not pull out the recorded cell or spill excessive amounts of biocytin in the tissue. When biocytin did leak into the tissue there were conspicuous strong puncta near the electrode mark, usually with nearby pyramidal neurons labeled. Cell-coupling data were not obtained from these slices. At around P30 and later nearly all slices displayed a biocytin leak, possibly arising from the novel expression of an as yet unidentified channel. For excised outside-out patch recording, the cell phenotype was first determined in whole cell mode, then the patches were pulled gradually from the cell while the cell was still held at –70 mV. The completion of patch separation was indicated by the visible separation of the patch from the whole cell soma. At completion of whole cell or patch recording, slices were removed from the recording chamber and immediately fixed in phosphate-buffered 4% formaldehyde pH 7.4 for 45–60 min at room temperature, washed in phosphate-buffered saline (PBS), and then stored at 4°C in PBS with 0.01% sodium azide until histochemistry and immunohistochemistry were performed.
To inhibit gap junction communication between astrocytes in acute slices we pretreated the slices with 100 µM MFA for 15–25 min before whole cell patch-clamp recording. Cells were patch-clamp recorded to confirm their passive nature. Some slices were fixed after recording and the biocytin was labeled and imaged as described below.
Histochemistry and immunohistochemistry
All steps were performed at room temperature and all incubations were done with gentle agitation in solutions containing 0.01% sodium azide. Slices were permeabilized for 30 min in 1% Triton X-100 in PBS. After a brief wash in PBS, slices were incubated for 4 h in 1:1,200 Cy2-conjugated streptavidin (Jackson ImmunoResearch Labs, West Grove, PA). Slices were washed with PBS and stored at 4°C in PBS also containing 0.01% sodium azide.
Before immunohistochemistry, slices were previewed for quality of tracer fill on a wide-field fluorescence microscope (Leica Microsystems, Wetzlar, Germany) or a confocal microscope (details below). Biocytin fills in immature animals were almost always found, whereas those in animals older than about 3 wk were more variable. Slices in which filled cells could not be found were used as negative controls for the immunohistochemical procedures. The blocking solution and the diluent for antisera contained 3% normal goat serum, 0.1% Triton X-100, and 0.01% sodium azide in PBS (NGS/TX). Nonspecific binding was blocked with a 3- to 4-h incubation in NGS/TX. Anti-GLAST (raised in guinea pig, Chemicon International) and anti-NG2 (raised in rabbit, Chemicon) were diluted to 1:4,000 and 1:600, respectively. Diluted antibodies were applied twice, 1 h apart, and then allowed to incubate overnight. NGS/TX alone was applied to negative control slices. Slices were washed once in NGS/TX and then three to four times in PBS/TX (0.1% Triton X-100 in PBS) over a 3- to 4-h period. The slices were reblocked with NGS/TX, then incubated with 1:1,000 Cy3-anti-Guinea pig and 1:600 Cy5-anti-rabbit IgG (each from Jackson ImmunoResearch). After the overnight secondary antibody incubation the slices were washed the same as was done after primary antibody incubation. Slices were then stored in PBS with sodium azide.
For microscopy, slices were moved to a glass-bottom chamber containing PBS. A Carl Zeiss LSM510 META (Oberkochen, Germany) was used to acquire images of single confocal planes or stacks of planes. For each image or image stack the gain and detector offset were adjusted to minimize saturated pixels, yet still detect weakly fluorescent coupled cells. In nearly all cases the pinhole was set to 1 Airy unit. Each channel was acquired separately. The presence of biocytin and a cell type marker in individual cells was assessed using the LSM510 software or the "Colocalization" plugin (Bourdoncle 2003) for ImageJ (Rasband 1997). Cells were counted with the aid of the "pointpicker" of ImageJ. Image cropping was performed in Adobe Photoshop CS. Transparent and maximum projection images were made with Zeiss LSM Image Browser and ImageJ, respectively.
|
|
RESULTS |
|---|
|
GLAST is expressed in several brain regions from neonate to adult, and astrocytes are specifically labeled with anti-GLAST antibodies (Chaudhry et al. 1995; Lehre et al. 1995) and (Furuta et al. 1997). In other studies we found that the GLAST(+) population included >90% of all of the GFAP(+) population in the CA1 region (Zhou et al. 2006). Another major constituent glia of the hippocampus, the NG2(+) cells, have been found to be negative for GLAST in freshly isolated cells from the hippocampus and in situ (Schools et al. 2003; Zhou et al. 2006). Therefore GLAST is a good membrane protein marker to label astrocytes after electrophysiological recording. Slices were also stained for the oligodendrocyte precursor cell marker NG2 and the recorded GLAST(+) cells were never NG2(+) (not shown).
The three fluorescent labels in each slice were imaged with confocal laser scanning microscopy. For each recorded cell enough z-planes were acquired to include the whole dye-coupled syncytium. GLAST(+) recorded cells were coupled 78% of the time (n = 74, P1–P43), but there was large variation in the degree of coupling. Figure 1 shows examples of noncoupled (A), moderately coupled (D), and highly coupled (G) GLAST(+) cells. The smaller panels to the right show biocytin diffusion (top) and GLAST immunoreactivity (bottom) in single confocal planes containing the recorded cells. The coupled GLAST(+) recorded cells are coupled almost exclusively to other GLAST(+) cells (e.g., see arrowheads in Fig. 1, E and F).
|
and Fig. 7 for examples). Outwardly rectifying glia (ORG) show strong outward rectification in their current–voltage relationships (I–V plot) with negligible currents inducible with hyperpolarizing voltage steps. Variably rectifying glia (VRG) show both outward and inward currents with multiple rectification points in their I–V plots, and passive glia (PG) show no apparent voltage- and time-dependent ion currents resulting in linear whole cell I–V plots. However, judging from the slow decay in capacitance current and noticeably low membrane resistance, it was impossible to voltage clamp the passive glia, which is a prerequisite to resolve and quantify voltage-gated ion conductance. We previously reported differences in ion current profiles showing different channels among the three astrocyte types described above from either freshly isolated astrocytes or astrocytes in situ (Zhou and Kimelberg 2000; Zhou et al. 2006; also shown in Fig. 7). Each of these cell types differed in the frequency of cell coupling. Among the GLAST(+) cells the passive astrocytes (PAs) are more frequently coupled than the outwardly rectifying astrocytes (ORAs) and variably rectifying astrocytes (VRAs) (Fig. 2A).
|
|
). The mean syncytium size reaches 50% of the maximum significantly earlier than the passive phenotype is 50% of maximum.
|
, 2001). However, these studies did not assess the intercellular diffusion of dye. Evidence that electrophysiological phenotype is independent of cell coupling
The above-presented results suggest an inverse relationship between cell coupling and the presence of voltage-gated ion currents. Stated differently, the presence of coupling or the degree of coupling can, in theory, contribute significantly to the measured electrophysiological properties of individual glia. For example, the mean syncytium size of GLAST(+) PAs is higher than that of ORAs and VRAs (Fig. 2B) and all PAs were coupled (Fig. 2A). However, on closer examination it appears that the coupling does not directly correlate with the presence of voltage-gated ion currents.
If increased syncytium size caused the observed transition from ORA to VRA to PA then we would expect to see coincidence between the increase in syncytium size and percentage of PA observed, although this does not occur. Rather, the increase in mean syncytium size of GLAST(+) glia precedes the transition from those types expressing a complex electrophysiological phenotype and those expressing passive phenotypes. Figure 3 shows that the coupling increase peaks at least a week before the PA type reaches its highest frequencies.
If the degree of cell coupling determines the whole cell electrophysiological profile then there should be minimal overlap in range of syncytium sizes between the different types. However, we observed overlap of syncytium size between ORA/VRA, ORA/PA, and VRA/PA (Fig. 4). Figure 5 shows an example of immunohistochemistry of a VRA with approximately the same size syncytium as that of a PA.
|
|
) we showed a difference in Rm values between each of the cell types, with ORA having the highest and PA the lowest. Again, it can be argued that the membrane resistances of the PAs are the lowest because these cells have the largest syncytia. However, the PA with the smallest observed syncytium was coupled to five cells so that the membrane resistance for VRAs and PAs coupled to five or more cells were compared, thus correcting for the amount of coupling. VRAs coupled to similar numbers of cells as PAs had significantly higher membrane resistances than that of the PAs. This shows that in these cases the membrane resistance is not a reflection of the number of cells it is coupled to and therefore implies that it is the result of an intrinsic difference in the recorded cells' expression of leak channels between the two cell types. Effect of the gap junction inhibitor meclofenamic acid
To look for further evidence supporting the idea that the passivity of the mature astrocytes reflects the nature of the intrinsic ion channel expression, we next investigated whether blocking gap junction channels between astrocytes by applying the reversible connexin 43 blocker, meclofenamic acid (MFA) (Harks et al. 2001), alters either passive current profile or passive membrane properties. MFA (100 µM) was applied to P18 hippocampal slices for 15–25 min before establishing whole cell recordings from cells with astrocyte morphology. When the glia (n = 11) were brought to 10-mV steps between –180 and +40 mV no voltage-dependent currents were uncovered in MFA-pretreated slices (Fig. 6A). The mean resting membrane potential was –76.5 mV for MFA-pretreated cells and –77.6 mV for control P18 cells (n = 5). Imaging of the biocytin that filled the recorded cells revealed a significant reduction in syncytium size as a result of the MFA (Fig. 6, B and C; Student's t-test, P = 0.026). Also indicating a reduction in syncytium size, the capacitance decreased and the membrane resistance increased relative to control (Fig. 6D). The MFA-induced changes in these passive membrane parameters also indicate that there is some amount of gap junction current contribution to the passive conductance because the membrane resistance shows a 12-fold increase in MFA-treated cells.
|
), this raises the possibility that the MFA effects we observed might partially be attributable to the MFA effects on passive astrocytic chloride channels. To this end, we investigated the potential presence of chloride conductance in the astrocyte passive conductance by replacing intracellular KCl by K-gluconate (see METHODS). This resulted in a reduced intracellular chloride to 6 mM and a calculated chloride equilibrium potential (ECl) shift from 0 to –83 mV. If chloride channel–mediated currents are involved in the passive conductance, the reduction in electrode chloride concentration should result in a negative shift in membrane potential. However, we found no significant change in resting membrane potential between MFA-treated (–76.3 ± 3.0 mV, n = 7) and untreated (–76.5 ± 2.2 mV, n = 11) passive astrocytes, showing no evidence that Cl– conductance is contributing to passive conductance. Therefore the MFA effects on membrane capacitance and resistance should not be derived from an effect on intrinsic Cl– channels.
MFA has also been shown to inhibit junctional conductance through connexin 50 (Srinivas and Spray 2003). However, there is no evidence for the expression of this connexin isoform in astrocytes in situ. Therefore the MFA effect in our experiments should most likely be through the inhibition of connexin 43.
Excised patches
We also made recordings from outside-out membrane patches excised from cells that were previously whole cell recorded. This recording configuration allows for determination of membrane ion channel expression without influence from syncytia and significantly improved voltage-clamp quality. If the syncytium has a substantial effect on the whole cell profile, then we would expect to see a transition from passive to nonpassive electrophysiological type when the patch recording is compared with its whole cell recording. As the patches were pulled from the cell bodies, a decrease in membrane capacitance (patch Cm = 6.2 ± 3.3 pF, n = 17) was observed, although there was no significant drop in membrane potential as the transmembrane ion gradients are maintained (patch Vm = –61 ± 13, n = 23). Figure 7, A–C shows representative whole cell and patch recordings from ORG, VRG, and PG cells. (Note: these are denoted ORG, VRG, and PG because immunohistochemistry for GLAST was not performed.) In each example the outside-out membrane patches predominantly resemble the electrophysiological profiles of the whole cell from which it was pulled. More patches reflected their parent cells than did not (summarized in Fig. 7D) and it can be seen that the patches from ORGs most frequently (92%) yielded the parent cell character. When VRG and ORG yielded a different profile from that of the parent cell it was always passive; >60% of the patches excised from PG cells yielded passive patches (n = 16) and those that did not were VRG (n = 9). Considering all three cell types, the concordance between the whole cell electrophysiological type and the corresponding excised membrane patches was significant (Goodman–Kruskal's Gamma = 0.92, P << 0.001). Thus the overall electrophysiological phenotype of hippocampal glia is likely attributable to channels in the membrane of the recorded cell.
|
|
DISCUSSION |
|---|
|
The results presented here extend our understanding of the relationship between cell coupling and the electrical properties of the membranes in astrocytes measured from individual cell bodies in situ. It is assumed that if a tracer molecule, such as Lucifer yellow or biocytin, can pass between gap junction–connected astrocytes, the electrical current will also, although the converse need not be the case because of the large difference in the timescales over which these two phenomena are observed, 50 ms for current and 10 min for dye transfer. A relative abundance of leak type channels in the cell soma ensures that most of the injected current will pass through these rather than the gap junction channels, especially if the latter are located on the ends of the thin processes with higher cross-sectional resistance as they are in the cerebellum (Muller et al. 1996). This is different from the situation in primary astrocyte cultures where there are no fine cell processes and gap junctions are spread around the entire rim of the flat cells (Dermietzel et al. 1991). In the mature rat hippocampus individual astrocytes occupy domains with little overlap at the distal ends of their processes (Bushong et al. 2002), so points of contact between cells are likely small relative to those seen in culture. The whole cell electrophysiological type is not determined by gap junction communication for the following additional reasons: All three electrophysiological types were found to be coupled, albeit at different frequencies (Fig. 2A) and with different syncytium sizes (Fig. 2B). The membrane resistances of VRAs are different from those of PAs, even when syncytium size is taken into consideration (Fig. 5). Excised plasma membrane patches most frequently displayed currents similar to those of the parent cell (Fig. 7).
Further evidence that the cell coupling is not a primary determinant of electrophysiological type comes from the developmental increase in syncytium size, which reaches a maximum before the transition to the passive phenotype. If the high degree of coupling is the basis of the passive phenotype then one might expect the syncytium size to increase coincidentally with the transition to the passive phenotype.
Three lines of evidence are generated from the experiments with the gap junction blocker MFA. First, the overall passive current pattern did not change on MFA treatment, which is consistent with previous studies by others using two other gap junction blockers (Wallfaff et al. 2004) and our outside-out patch data analysis, indicating that the characteristic passive current pattern is attributed to the ion channel properties of each individual cell instead of being formed by the junctional currents flowing to coupled astrocytes. MFA, like all gap junction blockers, is not selective and has been shown to inhibit and activate several ion channels (Lee and Wang 1999; Peretz et al. 2005). However, in our experimental conditions it does not alter the resting membrane potential and therefore does not obviously affect ion channel activity. Additionally, when we decreased the Cl– concentration in the recording pipette we observed no change in resting membrane potential. Therefore MFA is not likely to be acting on Cl– channels in astrocytes in rat hippocampal slices.
If the cell coupling was the cause of ohmic whole cell responses to voltage command, then we would not expect to see ORAs and VRAs coupled because coupling to even a few cells with free current access should greatly decrease Rm and convert these voltage- and time-dependent currents to more passive-appearing currents. However, both of these electrophysiological types are also coupled (see Fig. 2). There is no apparent threshold in syncytium size, beyond which an ORA becomes a VRA or a VRA becomes a PA. It is most significant that there are examples of all three electrophysiological types that are coupled in similar-sized syncytia (Fig. 4). Thus although we cannot exclude some contribution of the gap junctions, the simplest explanation from both our studies and those of other investigators (Barres et al. 1990; Muller et al. 1996; Wallraff et al. 2004) is that the passive or nonpassive electrophysiological phenotypes of glial cells reflect their intrinsic membrane channels, rather than the effect of intercellular current spread through gap junctions, leading to attenuated voltage clamping, and preventing the observation of voltage-gated and time-dependent channels.
Data in the literature also argue against gap junction coupling as a major determinant of individual astrocyte electrophysiological phenotype. Wallraff et al. (2004) used other gap junction blockers (octanol and carbenoxolone) and found no effect on input resistance measured from glia in the 9- to 65-day-old hippocampal slice. When dye-coupled pairs of Bergmann glial cells of the mouse cerebellum were voltage clamped, junctional conductance was low compared with current through the membrane of the current-injected cell (Muller et al. 1996). Finally, Barres et al. (1990) calculated a length constant of around 100 µm from voltage-clamped freshly isolated P50 white matter astrocytes and that it was too small to support the significant transfer of current between coupled astrocytes.
However, we have an apparent discrepancy with the MFA data where this inhibitor causes a decrease in capacitance and an increase in the apparent membrane resistance (obtained from the PClamp program), as seen when the individual values are plotted (Fig. 6D). Dye transfer was also reduced (Fig. 6, B and C). Thus on an individual cell basis, gap junctions in part determined the Cm and Rm but had no effect on the passive profile (Fig. 6A).
Development of astrocyte dye coupling
Coupled GLAST(+) astrocytes first appear at P1 and their frequency rapidly increases thereafter. We thus found dye coupling between astrocytes at an earlier age than was reported for astrocytes in the rat visual cortex (Binmoller and Muller 1992) or in Bergman glia of the mouse cerebellum (Muller et al. 1996). The former study found no coupling before P11 using Lucifer yellow as a fluorescent dye and the latter found coupling at P20–P24, but not P5–P7. This could be a result of differences between brain regions.
All of the GLAST(+) passive astrocytes in the mature animal were coupled. This contradicts the results presented recently, that there are coupled and uncoupled cells with astrocyte properties (Wallraff et al. 2004). In that study cells were selected based on human GFAP promoter–driven EGFP expression in transgenic mice, where the more highly fluorescent cells are glutamate transporting cells and weakly fluorescent cells are glutamate receptor expressing cells. An earlier report from the same group found that about a third of the green fluorescent cells express the mouse homologue of NG2, AN2 (Mattias et al. 2003). Therefore it is likely that some of the noncoupled cells they observed (Walraff et al. 2004) are NG2(+) cells.
Properties of excised patches
The passive cell behaves as if it were almost invisible in the whole cell patch-clamp configuration. Its influence is clearly seen only in the reversal potential of around –72 mV compared with the calculated EK of –93 mV. Because the considerable voltage drop over the electrode resistance of about 5 M is on the same scale as the cell membrane resistance (Rm), measured as best as can be done from the resistance and capacitance change by the amplifier, the passivity can be partly attributed to the passive resistance of the electrode as well as the membrane conductance. This is why others (Bordey and Sontheimer 1997) selected cells with higher input resistance, but as we have seen (Zhou et al. 2006) the low-resistance cells are the major constituent of glial cells in the adult hippocampus and, by GLAST immunoreactivity, are the major population of astrocytes, at least in the mature hippocampus. In their seminal study on glial cell electrophysiology, Kuffler et al. (1966) impaled the glial cell bodies of an amphibian optic nerve with sharp electrodes and obtained resistances of <1 to 7 M. This prepatch-clamp technique involves injecting current with one electrode and recording the transmembrane voltage with the other and thus is not subject to the series resistance limitation of the single electrode voltage clamp.
Membrane patches excised from cells most often reflected the same electrophysiological type as the cell from which they were pulled. However, each cell type did yield some patches of a different type. The causes for the discrepancy between whole cell and membrane patch may be ascribed to uneven distribution of channels in the soma and the localization of different currents in the processes or even reliable voltage clamping in the patches that allows a minority of voltage- and time-dependent K+ currents to be activated.
In conclusion we have found that 1) glial cell coupling in the hippocampus occurs earlier than the complete transition to the apparent passive phenotype and 2) all electrophysiological types of GLAST(+) glia, which include those showing voltage-gated and time-dependent currents, may be coupled. Thus in the hippocampus the cells coupled to astrocytes do not determine whether voltage-gated and time-dependent ion channels are seen. Further work with RT-PCR and staining, combined with electrophysiology of membrane patches, is required to determine the molecular nature of ion channels that make the majority of mature astrocytes passive. These findings have important implications for the characterization of mature astrocytes and their functional roles, such as voltage-driven potassium distribution and diffusion of other compounds in the astrocyte syncytium.
|
|
GRANTS |
|---|
|
|
|
NOTE ADDED IN PROOF |
|---|
|
|
|
ACKNOWLEDGMENTS |
|---|
|
|
|
FOOTNOTES |
|---|
Address for reprint requests and other correspondence: G. P. Schools, Ordway Research Institute, Inc., 150 New Scotland Avenue, Albany, NY 12208 (E-mail: gschools{at}ordwayresearch.org)
|
|
REFERENCES |
|---|
|
Binmoller FJ and Muller CM. Postnatal development of dye-coupling among astrocytes in rat visual cortex. Glia 6: 127–137, 1992.[CrossRef][Web of Science][Medline]
Bordey A and Sontheimer H. Postnatal development of ionic currents in rat hippocampal astrocytes in situ. J Neurophysiol 78: 461–477, 1997.
Bourdoncle P. Colocalization Plugin in Java. Paris, France: Institut Jacques Monod, Service Imagerie, 2003.
Bushong EA, Martone ME, Jones YZ, and Ellisman MH. Protoplasmic astrocytes in CA1 stratum radiatum occupy separate anatomical domains. J Neurosci 22: 183–192, 2002.
Chaudhry FA, Lehre KP, van Lookeren Campagne M, Ottersen OP, Danbolt NC, and Storm-Mathisen J. Glutamate transporters in glial plasma membranes: highly differentiated localizations revealed by quantitative ultrastructural immunocytochemistry. Neuron 15: 711–720, 1995.[CrossRef][Web of Science][Medline]
Dermietzel R, Hertberg EL, Kessler JA, and Spray DC. Gap junctions between cultured astrocytes: immunocytochemical, molecular, and electrophysiological analysis. J Neurosci 11: 1421–1432, 1991.[Abstract]
Farahani R, Pina-Benabou MH, Kyrozis A, Siddiq A, Barradas PC, Chiu FC, Cavalcante LA, Lai JC, Stanton PK, and Rozental R. Alterations in metabolism and gap junction expression may determine the role of astrocytes as "good Samaritans" or executioners. Glia 50: 351–361, 2005.[CrossRef][Web of Science][Medline]
Furuta A, Rothstein JD, and Martin LJ. Glutamate transporter protein subtypes are expressed differentially during rat CNS development. J Neurosci 17: 8363–8375, 1997.
Harks EG, de Roos AD, Peters PH, de Haan LH, Brouwer A, Ypey DL, van Zoelen EJ, and Theuvenet AP. Fenamates: a novel class of reversible gap junction blockers. J Pharmacol Exp Ther 298: 1033–1041, 2001.
Holthoff K and Witte OW. Directed spatial potassium redistribution in rat neocortex. Glia 29: 288–292, 2000.[CrossRef][Web of Science][Medline]
Kofuji P and Newman EA. Potassium buffering in the central nervous system. Neuroscience 129: 1045–1056, 2004.[CrossRef][Web of Science][Medline]
Kuffler SW, Nicholls JG, and Orkand RK. Physiological properties of glial cells in the central nervous system of amphibia. J Neurophysiol 29: 768–787, 1966.
Lee YT and Wang Q. Inhibition of hKv2.1, a major human neuronal voltage-gated K+ channel, by meclofenamic acid. Eur J Pharmacol 378: 349–356, 1999.[CrossRef][Web of Science][Medline]
Lehre KP, Levy LM, Ottersen OP, Storm-Mathisen J, and Danbolt NC. Differential expression of two glial glutamate transporters in the rat brain: quantitative and immunocytochemical observations. J Neurosci 15: 1835–1853, 1995.[Abstract]
Matthias K, Kirchhoff F, Seifert G, Huttmann K, Matyash M, Kettenmann H, and Steinhauser C. Segregated expression of AMPA-type glutamate receptors and glutamate transporters defines distinct astrocyte populations in the mouse hippocampus. J Neurosci 23: 1750–1758, 2003.
Muller T, Moller T, Neuhaus J, and Kettenmann H. Electrical coupling among Bergmann glial cells and its modulation by glutamate receptor activation. Glia 17: 274–284, 1996.[CrossRef][Web of Science][Medline]
Nagy JI, Patel D, Ochalski PA, and Stelmack GL. Connexin30 in rodent, cat and human brain: selective expression in gray matter astrocytes, co-localization with connexin43 at gap junctions and late developmental appearance. Neuroscience 88: 447–468, 1999.[CrossRef][Web of Science][Medline]
Orkand RK, Nicholls JG, and Kuffler SW. Effect of nerve impulses on the membrane potential of glial cells in the central nervous system of amphibia. J Neurophysiol 29: 788–806, 1966.
Peretz A, Degani N, Nachman R, Uziyel Y, Gibor G, Shabat D, and Attali B. Meclofenamic acid and diclofenac, novel templates of KCNQ2/Q3 potassium channel openers, depress cortical neuron activity and exhibit anticonvulsant properties. Mol Pharmacol 67: 1053–1066, 2005.
Rasband WS. ImageJ. Bethesda, MD: National Institutes of Health, 1997–2004. http://rsb.info.nih.gov/ij/
Rash JE, Duffy HS, Dudek FE, Bilhartz BL, Whalen LR, and Yasumura T. Grid-mapped freeze-fracture analysis of gap junctions in gray and white matter of adult rat central nervous system, with evidence for a "panglial syncytium" that is not coupled to neurons. J Comp Neurol 388: 265–292, 1997.[CrossRef][Web of Science][Medline]
Rash JE, Yasumura T, Dudek FE, and Nagy JI. Cell-specific expression of connexins and evidence of restricted gap junctional coupling between glial cells and between neurons. J Neurosci 21: 1983–2000, 2001.
Schools GP, Zhou M, and Kimelberg HK. Electrophysiologically "complex" glial cells freshly isolated from the hippocampus are immunopositive for the chondroitin sulfate proteoglycan NG2. J Neurosci Res 73: 765–777, 2003.[CrossRef][Web of Science][Medline]
Srinivas M and Spray DC. Closure of gap junction channels by arylaminobenzoates. Mol Pharmacol 63: 1389–1397, 2003.
Theis M, Sohl G, Eiberger J, and Willecke K. Emerging complexities in identity and function of glial connexins. Trends Neurosci 28: 188–195, 2005.[CrossRef][Web of Science][Medline]
Venance L, Piomelli D, Glowinski J, and Giaume C. Inhibition by anandamide of gap junctions and intercellular calcium signaling in striatal astrocytes. Nature 376: 590–594, 1995.[CrossRef][Medline]
Wallraff A, Köhling R, Heinemann U, Theis M, Willecke K, and Steinhäuser C. The impact of astrocyte gap junctional coupling on potassium buffering in the hippocampus. J Neurosci 26: 5438–5447, 2006.
Wallraff A, Odermatt B, Willecke K, and Steinhauser C. Distinct types of astroglial cells in the hippocampus differ in gap junction coupling. Glia 48: 36–43, 2004.[CrossRef][Web of Science][Medline]
White MM and Aylwin M. Niflumic and flufenamic acids are potent reversible blockers of Ca2(+)-activated Cl– channels in Xenopus oocytes. Mol Pharmacol 37: 720–724, 1990.[Abstract]
Zhou M and Kimelberg HK. Freshly isolated astrocytes from rat hippocampus show two distinct current patterns and different [K+]o uptake capabilities. J Neurophys 84: 2746–2757, 2000.
Zhou M, Schools GP, and Kimelberg HK. Development of GLAST(+) astrocytes and NG2(+) glia in rat hippocampus CA1: mature astrocytes are electrophysiologically passive. J Neurophysiol 95: 134–143, 2006.
This article has been cited by other articles:
|
|
 |
|
  M. Zhou, G. Xu, M. Xie, X. Zhang, G. P. Schools, L. Ma, H. K. Kimelberg, and H. Chen TWIK-1 and TREK-1 Are Potassium Channels Contributing Significantly to Astrocyte Passive Conductance in Rat Hippocampal Slices J. Neurosci., July 1, 2009; 29(26): 8551 - 8564. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
V. Houades, A. Koulakoff, P. Ezan, I. Seif, and C. Giaume Gap Junction-Mediated Astrocytic Networks in the Mouse Barrel Cortex J. Neurosci., May 14, 2008; 28(20): 5207 - 5217. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. Andersson, F. Blomstrand, and E. Hanse Astrocytes play a critical role in transient heterosynaptic depression in the rat hippocampal CA1 region J. Physiol., December 15, 2007; 585(3): 843 - 852. [Abstract] [Full Text] [PDF]
|
|
|
|
|
|
J. C. Nawroth, C. A. Greer, W. R. Chen, S. B. Laughlin, and G. M. Shepherd An Energy Budget for the Olfactory Glomerulus J. Neurosci., September 5, 2007; 27(36): 9790 - 9800. [Abstract] [Full Text] [PDF]
|
|
|
|
 |
|
 M. L. Olsen, S. L. Campbell, and H. Sontheimer Differential Distribution of Kir4.1 in Spinal Cord Astrocytes Suggests Regional Differences in K+ Homeostasis J Neurophysiol, August 1, 2007; 98(2): 786 - 793. [Abstract] [Full Text] [PDF]
|
|
| ||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||||
| HOME | HELP | FEEDBACK | SUBSCRIPTIONS | ARCHIVE | SEARCH | TABLE OF CONTENTS |
| Visit Other APS Journals Online |
TOP
ABSTRACT
INTRODUCTION
