1 Institut für Physiologie der Charité, Humboldt Universität, D 10117 Berlin, Germany; and 2 Department of Molecular Biophysics and Physiology, Rush Presbyterian St. Luke's Medical Center, Chicago, Illinois 60612
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ABSTRACT |
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Microglial activation is accompanied by changes in
K+ channel expression. Here we demonstrate that a
deactivating cytokine changes the electrophysiological properties of
microglial cells. Upregulation of delayed rectifier (DR) K+
channels was observed in microglia after exposure to transforming growth factor- (TGF-
) for 24 h. In contrast, inward
rectifier K+ channel expression was unchanged by TGF-
.
DR current density was more than sixfold larger in TGF-
-treated
microglia than in untreated microglia. DR currents of TGF-
-treated
cells exhibited the following properties: activation at potentials more
positive than
40 mV, half-maximal activation at
27 mV, half-maximal
inactivation at
38 mV, time dependent and strongly use-dependent
inactivation, and a single channel conductance of 13 pS in Ringer
solution. DR channels were highly sensitive to charybdotoxin (CTX) and
kaliotoxin (KTX), whereas
-dendrotoxin had little effect.
With RT-PCR, mRNA for Kv1.3 and Kir2.1 was detected in microglia. In
accordance with the observed changes in DR current density, the mRNA
level for Kv1.3 (assessed by competitive RT-PCR) increased fivefold after treatment of microglia with TGF-
.
brain macrophages; transforming growth factor-; inward rectifier
K+ current; delayed rectifier K+ current; reverse transcription-polymerase chain reaction; Kir2.1
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INTRODUCTION |
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MICROGLIA REPRESENT A POPULATION of resident macrophages of the brain. In addition to performing normal functions of macrophages in other tissues, microglia are highly specialized. Resting microglia are highly ramified, with extensive processes that presumably act as antennae, sensing inflammatory stimuli, tissue damage, cellular debris, etc. They are characterized by downregulated macrophage surface antigens and macrophage functions. On stimulation, microglia react by withdrawing their processes, becoming ameboid and macrophage-like. Activated microglia become capable of phagocytosis, proliferation, and chemotaxis and release various cytokines and cytotoxins (34, 39, 40). Recent interest in microglia has been piqued by the idea that hyperactive microglia may cause tissue damage, possibly contributing to lesions of human immunodeficiency virus-associated dementia (32), multiple sclerosis (42), Alzheimer's disease (28), and a variety of other neurodegenerative and autoimmune diseases (34, 39, 40). This background emphasizes the importance of understanding microglial deactivation, the reversal of microglial activation.
Electrophysiological studies in vitro have revealed dramatic changes in
the expression levels of K+ channels in microglia during
the process of activation (for review, see Ref. 10). Exposure to a
variety of activating stimuli produces a similar pattern of
electrophysiological changes in microglia. Lipopolysaccharide (LPS)
(31), granulocyte-macrophage colony-stimulating factor
(GM-CSF) (11, 13), or interferon- (IFN-
)
(13) induces upregulation of delayed rectifier (DR)
K+ currents, which is accompanied by simultaneous
downregulation of inward rectifier (IR) K+ currents. The DR
K+ currents of cytokine- or LPS-activated microglial cells
share many properties with cloned Kv1.3 channels, including an
activation threshold at about
40 mV, strongly use-dependent
inactivation, and a high sensitivity to the scorpion peptide toxins
CTX, KTX, and noxiustoxin (11, 31). On cellular
activation, upregulation of DR (Kv1.3) K+ currents also
occurs in other immune cells, such as macrophages and lymphocytes (for
review, see Ref. 7). Effects of deactivating cytokines on microglial
K+ channel expression have not yet been investigated.
Several studies have demonstrated that TGF- plays an important role
in deactivation of microglia. For example, TGF-
1 has been shown to
suppress superoxide production in LPS-activated microglial cells and to
reduce expression of Fc receptors and major histocompatibility complex
class II molecules in IFN-
-stimulated microglia (14, 23, 27,
29, 41). TGF-
is produced in the central nervous system.
Microglial cells express TGF-
receptors and are capable of releasing
TGF-
1 (5). In microglia, TGF-
reduces production of
various proinflammatory cytokines, such as interleukin (IL)-1, IL-6,
and tumor necrosis factor-
, and inhibits proliferation (20,
41).
In the present study we investigated the patterns of K+
channel expression in microglia before and after exposure to the
deactivating cytokine TGF-. The inward and outward rectifier
K+ currents of TGF-
-treated microglial cells were
characterized, and the molecular identities of the main K+
channels were determined. Preliminary accounts of this work have been
published in abstract form (35, 36).
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MATERIALS AND METHODS |
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Cell Culture
Primary and secondary microglial cultures.
Microglia were obtained from brain cell cultures of newborn
NMRI mice, supplied by Charles River (Sulzfeld, Germany). Mixed brain cell cultures were prepared as described previously
(12). Brain cortices were enzymatically dissociated (15 min at 37°C with 0.25% trypsin, type XI, Sigma), and a single cell
suspension was achieved by repeated triturations. Cells were seeded
into tissue culture flasks at a density of 2-4 × 106/10 ml in DMEM (GIBCO) supplemented with 10%
heat-inactivated FCS (GIBCO) and 30% supernatant of L-929 fibroblasts
as a source of macrophage CSF (M-CSF). After at least 10 days of
incubation, microglia were harvested by shaking the cultures (30 min,
300 rpm) to detach weakly adherent cells from the astrocytic monolayer. Isolated microglia were seeded on glass coverslips in 24-well Costar
plates (3 × 104/0.5 ml). These cells were cultured in
DMEM supplemented with 10% FCS. The culture medium did not contain
M-CSF. In secondary culture, untreated microglial cells were partially
activated, i.e., cells exhibited an ameboid morphology and the
expression of various macrophage surface antigens was upregulated
(12). One to two days after the isolation procedure,
microglial cells were treated with 10 ng/ml recombinant human TGF-1
or 10 ng/ml natural porcine TGF-
2 (both from Sigma) to induce
deactivation. Patch-clamp recordings were performed 24 h after the
TGF-
treatment.
BV-2 microglial cell line.
The immortalized mouse microglial cell line BV-2, which exhibited
properties similar to activated microglia (Refs. 1 and 2; kindly
provided by Dr. E. Blasi, Perugia, Italy), was used in some
experiments. BV-2 microglial cells were cultured permanently in DMEM
supplemented with 10% FCS and 2 mM L-glutamine. BV-2 cells were split twice a week and were plated on glass coverslips at a
density of 1-1.5 × 104/0.5 ml for subsequent
patch-clamp experiments. To induce deactivation, BV-2 cells were
treated for 24 h with 10 ng/ml TGF-1 or TGF-
2.
Electrophysiological Recordings
Whole cell measurements.
Whole cell membrane currents were measured using an EPC-9 patch-clamp
amplifier (HEKA, Lambrecht/Pfalz). The amplifier was interfaced to an
IBM computer for pulse application and data recording. Series
resistance compensation was routinely used to reduce the effective
series resistance by ~70%. Patch electrodes of 2-4 M were
fabricated on a two-stage puller (Narishige PP-83, Tokyo, Japan) from
borosilicate glass (outer diameter 1.5 mm and inner diameter 1 mm;
Hilgenberg, Malsfeld, Germany). The electrodes were filled with the
following solution (in mM): 120 KCl, 1 CaCl2, 2 MgCl2, 10 HEPES, and 11 EGTA. This solution was adjusted to pH 7.3 with KOH. The extracellular solution contained (in mM) 130 NaCl,
5 KCl, 2 CaCl2, 1 MgCl2, 10 HEPES, and 8 D-glucose. The pH of the extracellular solutions was
adjusted to 7.4 with NaOH. All recordings were done at room temperature
(20-23°C). Whole cell K+ currents were filtered at 3 kHz and stored on computer disk for subsequent analyses. Analyses were
performed on IBM computers with the Pulse/PulseFit program (HEKA,
Lambrecht/Pfalz). Data are presented as mean values ± SE. The
number of experiments is indicated. Membrane capacitances of the cells
analyzed in this study varied between 5 and 44 pF. In
morphologically complex cells such as microglia, long, narrow processes
may not be under voltage-clamp control. The cells studied here
typically had one or two short processes, probably comprising a small
fraction of the total membrane area. Although we did not observe
electrophysiological manifestations of significant space-clamp
problems, we cannot rule out the possibility that some small and
inaccessible regions of membrane were excluded electrically from
contributing to the whole cell currents. If not stated otherwise, leak
currents were subtracted from current records before further analysis.
Leak current was determined by measuring the current evoked by a
voltage step from
60 to
70 mV where no time-dependent current was
present. Leak current was subtracted from each K+ current
recording assuming a linear current-voltage relationship of
the leak currents.
Single channel recordings.
Single channel experiments were done at 20°C, with the bath
temperature controlled by Peltier devices. Micropipettes were pulled in
several stages using a Flaming Brown automatic pipette puller (Sutter
Instruments, San Rafael, CA) from 7052 or KG-12 glass (Garner Glass,
Claremont, CA), coated with Sylgard 184 (Dow Corning, Midland, MI), and
heat polished to a tip resistance usually 3-15 M. Electrical
contact with the pipette solution was achieved by a thin sintered
Ag-AgCl pellet (In Vivo Metric Systems, Healdsburg, CA) attached to a
silver wire covered by a Teflon tube or simply with a chlorided silver
wire. A reference electrode made from a Ag-AgCl pellet was connected to
the bath through an agar bridge made with Ringer solution. The current
signal from the patch clamp (List Electronic, Darmstadt, Germany,
Axopatch-1A or Axopatch 200B; Axon Instruments, Foster City, CA) was
digitized and stored in computer files for offline analysis using Indec
Laboratory Data Acquisition and Display Systems (Indec, Sunnyvale, CA)
or pCLAMP 6.0.3 (Axon Instruments). Data acquisition and analysis programs were written in BASIC-23 or FORTRAN. Data are presented without correction for liquid junction potentials.
Data analyses.
The steady-state activation curve was fitted using the following
Boltzmann equation
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Pharmacological Studies
For drug application, a four-barrel microperfusion pipette was positioned at a distance of ~30-50 µm from the recorded cell to permit a rapid exchange of solutions outside the cell, which was achieved in <1 s. The flow rate was adjusted by hydrostatic pressure.Effects of the following drugs were studied: CTX, -dendrotoxin
(
-DTX), and KTX (all from Latoxan, Rosans, France) and
BaCl2 and
-DTX (both from Sigma). Peptide toxins were
dissolved in 0.1% bovine serum albumin-containing solutions.
Statistics
The statistical significance of differences between experimental groups was evaluated by Student's t-test using the SPSS program. Data were considered to be statistically significant with P < 0.05.RT-PCR
RNA isolation. Total RNA was isolated from cultured BV-2 cells using a modification of the procedure given by Chomczynski (4). Approximately 2 × 106 cells were centrifuged and resuspended in a minimum volume of growth medium. One milliliter of Trizol (GIBCO-Life Technologies) reagent was added. The manufacturer of Trizol specified subsequent steps in the isolation procedure that were followed. RNA was suspended in diethylpyrocarbonate-treated water and quantified by ultraviolet (UV) spectroscopy.
Reaction conditions. Tth polymerase was used to convert RNA to cDNA as well as to amplify cDNA by the PCR in the same reaction tube. Each reaction tube contained 100 ng total RNA and 5 units of polymerase. The RT step was carried out at 60°C for 30 min in the presence of Mn and the downstream gene-specific primer. Before PCR, a buffer that chelated Mn and substituted Mg was added along with the upstream gene-specific primer at 90°C. Thirty cycles of PCR were then performed using a combined annealing and extension step at a temperature of 60°C. Reaction products were visualized by gel electrophoresis. The gel contained 3% agarose, was stained with 10 µg/ml ethidium bromide, and subsequently destained. Oligonucleotide primers used in this study included those for mouse Kv1.3 (GenBank M30441, upstream 5'-ATCTTCAAGCTCTCCCGCCA, downstream 5'-CGATCACCATATACTCCGAC), glyceraldehyde-3-phosphate-dehydrogenase (G3PDH, Genebank M32599, upstream 5'-TGATGACATCAAGAAGGTGGTGAAG, downstream 5'-TCCTTGGAGGCCATGTAGGCCAT), mouse Kv1.5 (GenBank L22218, upstream 5'-GCCATTGCCATCGTGTCGGT, downstream 5'-ACATGTGGTCTCCACGATGA), and mouse Kir2.1 (GenBank X73052, upstream 5'-CGACTGCCATGACAACTCAA, downstream 5'-CATATCTCCGATTCTCGCCT).
Competitive RT-PCR. The abundance of specific species of mRNA (target) was determined using a competitive RT-PCR assay. In this assay, parallel RT-PCR reactions are performed. Each reaction tube contained a constant amount of total RNA and a variable amount of artificial RNA (competitor). The ratio of target to competitor was measured after RT-PCR to infer the amount of target in the reaction (37). The competitor was a truncated form of the target but contained the same primer binding sites. The concentration of this RNA competitor was measured by UV spectrophotometry.
The ratio of amplified target and competitor was determined from measurements of the intensity of the corresponding bands seen in the electrophoresis gel after staining. The intensity was measured from images digitized using a low-light integrating charge-coupled device camera. The relative intensity of each band was calculated as the sum of the values of all pixels in the band after subtraction of the background. The ability of each competitor to measure a change in mRNA abundance was tested by applying the assay to known dilutions of total RNA.Restriction analysis. Products of RT-PCR were reamplified for restriction analysis and concentrated with a centrifugal filtration device. Digestion of the DNA was carried out in the appropriate buffer at 37°C for 1 h. Hph I was obtained from New England Biolabs. Rsa I and Sac I were obtained from Promega.
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RESULTS |
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IR K+ Currents in TGF--Treated
Microglia
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Single channel currents were recorded mainly from BV-2 cells. The
changes in K+ currents in BV-2 cells exposed to TGF- for
1 day were similar to those observed in microglia in primary culture.
When the pipette solution contained high extracellular K+
concentration, most patches (13/18) contained IR channels,
often at high density. Figure 1D illustrates ramp currents
in an inside-out patch with at least five active IR channels. During
voltage ramps, most of the IR channels were open at negative voltages,
indicating a high open probability. No outward current other than leak
was observed. Because of open-channel rectification, the value obtained for the conductance depended on the voltage range selected. Similar open-channel rectification has been observed in IR channels in endothelial cells (38). The single IR channel conductance
obtained by linear regression on the averaged leak-subtracted
open-channel current between
100 and
20 mV relative to the Nernst
potential for K+ was usually 25-30 pS.
In rat microglia, mRNA of both ROMK-1 and Kir2.1 channels was reported
(25). To identify the IR channels that are expressed in
TGF--treated murine microglia, we first investigated effects of
extracellular Ba2+ on whole cell IR currents, because
Kir2.1 channels are much more sensitive to Ba2+ than are
ROMK-1 channels (24, 43). Micromolar concentrations of
extracellular Ba2+ induced a voltage- and time-dependent
blockade of IR currents (data not shown), and 1 mM Ba2+
abolished IR currents in TGF-
-treated microglia (n = 10) (Fig. 2, A and
B). Next, we investigated effects of
-DTX, a peptide toxin that inhibits ROMK-1 but no other IR channels (19).
As shown in Fig. 2, C and D, neither the
amplitude nor the kinetics of IR currents was affected by 100 nM
-DTX (n = 6).
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Whole Cell DR K+ Currents in
TGF--Treated Microglia
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Single DR K+ Channel Currents
Figure 4A shows a single averaged leak-subtracted current in a patch with one active DR channel. The slope conductance of the open-channel current at the reversal potential was 26 pS. In other patches, the single channel DR conductance was typically 25-30 pS in symmetrical high K+ solutions. The properties used to identify the single channels as DR included all of the following: activation with voltage in the general vicinity of
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Figure 4B illustrates the leak-subtracted current elicited by a voltage ramp in a cell-attached patch containing a single active DR channel. The bath contained K-Ringer solution, intended to "clamp" the resting membrane potential near 0 mV, and the pipette contained Ringer solution. The channel displays typical flickery openings, as well as inward rectification at large positive voltages (i.e., positive to +80 mV). The slope conductance estimated from the linear portion of the current-voltage relationship (below +60 mV) was variable both within one patch and between patches and ranged 8-17 pS. The average value from the seven most reliable patches studied with Ringer solution in the pipette was 12.6 ± 0.9 pS. Under similar conditions, the conductance of Kv1.3 channels in human lymphocytes is 13 pS (3).
Voltage Dependence of Activation and Inactivation of DR K+ Currents
Kinetic and pharmacological properties of DR K+ currents in TGF-
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The "steady-state" voltage dependence of inactivation of DR
currents was studied by varying the holding potential between 90 and
+20 mV in 10-mV increments. After the holding potential was established
for at least 2 min, cells were pulsed to a test potential of +30 mV for
200 ms. When the holding potential was more positive than
60 mV, DR
currents during the test pulse decreased in amplitude. As shown for the
example of current recordings in Fig. 5B, DR current was not
activated during the test pulse when cells were held at potentials
positive to
20 mV. Peak amplitudes of the evoked currents were
measured, normalized, and then plotted as a function of the holding
potential (Fig. 5C). Fitted with a Boltzmann function,
half-maximal inactivation was at
38 mV, with k = 4.3 mV (n = 13).
Time-Dependent Activation and Inactivation of DR K+ Currents
The activation time constant was determined by fitting n4 kinetics (18) to the rising phase of DR current during voltage pulses to potentials between
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To characterize the time dependence of inactivation of DR currents,
2,000-ms voltage pulses were applied to potentials between 40 and +30
mV. At potentials near the activation threshold, inactivation became
faster with increasing depolarization. At potentials more positive than
0 mV, the inactivation rate did not change further (Fig. 6,
C and D). The time constant of inactivation was
581 ± 40 ms (n = 13) at +30 mV.
Use Dependence of DR K+ Currents
Pronounced use dependence of DR currents in TGF-
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Pharmacology of DR K+ Currents
We investigated the sensitivity of DR channels in TGF-
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Similar to the effect of CTX, extracellular KTX, applied at nanomolar
concentrations, blocked DR currents of TGF--treated microglial cells
(Fig. 8, B and D). When microglial cells were superfused with solutions containing 1 nM KTX, the mean amplitude of DR
currents evoked by voltage steps to +30 mV was reduced by 54 ± 10% (n = 8). At 100 nM extracellular KTX, DR currents
were almost completely blocked (by 93 ± 2%; n = 6) (Fig. 8D).
The snake toxin -DTX was tested at concentrations of 10 and 100 nM.
In contrast to the effects of CTX and KTX, both concentrations
-DTX
inhibited DR currents only slightly, as shown in Fig. 8, C
and D. At a test potential of +30 mV, DR currents were
reduced on average by 17 ± 8% (n = 7) at 10 nM
-DTX and by 28 ± 7% (n = 13) at 100 nM
-DTX.
Identification and Regulation of K+
Channel mRNA in TGF--Treated BV-2 Cells
The presence of Kv1.3 and Kir2.1 mRNA transcripts was tested with
RT-PCR. DNA products apparently corresponding to these species were
obtained after RT-PCR of total RNA when unique sets of primers were
used. An image of a gel of the PCR products from one RNA sample after
electrophoresis is shown in Fig.
9A. Also shown is the DNA band
corresponding to G3PDH after RT-PCR. The sizes of the bands can be
obtained by comparison with the molecular weight markers. When
amplified with the primer set used, Kv1.3 is expected to give a band
479-bp long. Kir2.1 should be 376 bp, and G3PDH, 240 bp. The bands
suggest that mRNA for these species are present. However, because a
band could theoretically also be obtained through amplification of
contaminating genomic DNA rather than mRNA, control reactions were
performed in which the RT step was omitted. In these cases, the
reaction remained at 4°C during the RT step. In the gel shown in Fig.
9A, the PCR product in the absence of RT was run in the lane
to the right of that in which RT was performed. No DNA was amplified in
the absence of RT, indicating that the amplified DNA originated from
mRNA.
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Restriction analysis was performed to determine whether the DNA obtained by RT-PCR had the base sequence expected for the Kv1.3 transcript. The DNA product was reamplified and subjected to digestion with either Rsa I, Sac I, or Hph I. A gel of the full-length DNA and products after digestion and electrophoresis is shown in Fig. 9B. Rsa I would be expected to cut the DNA into fragments of 376 and 103 bp. Exposure to Sac I would result in fragments of 425 and 54 bp. The fragments after digestion with Hph I should be 244 and 235 bp. In each case, the expected fragments were found. The most likely origin of the cDNA is, therefore, mouse Kv1.3.
The change in current density of the DR current in response to the
addition of TGF-1 to the growth media could be mediated by a change
in the abundance of Kv1.3 transcript. To test this idea, the abundance
of the transcript in the TGF-
-treated and untreated cells was
compared using a competitive RT-PCR assay. A typical experiment is
shown in Fig. 10. A sample of total RNA was isolated from each of the two cultures. RT-PCR was performed in a
set of four reaction tubes for each sample. Total RNA was fixed in each
reaction tube. However, a variable amount of competitor RNA was added
to different tubes in each set before RT. The image of the gel after
RT-PCR of Kv1.3, electrophoresis, and staining is shown in Fig.
10A. The amount of final product of the target or competitor
is proportional to the intensity of the band. The ratio of the target
to the competitor increased as the competitor in the reaction was
reduced. For any competitor concentration, the ratio of target to
competitor was higher in the reaction from the RNA sample obtained from
the cells treated with TGF-
1. These ratios are plotted in Fig.
10B as a function of the concentration of Kv1.3 competitor
added. The point of equal amplification of the two products occurs when
the ratio of the target to competitor equals one (ignoring the
difference in size). The amount of competitor added for this value is
the amount of target in the sample. From the graph it is apparent that
more competitor was added for equal amplification to the RT-PCR of the
sample of TGF-
-treated cells. In the example assay shown, Kv1.3
transcript was a factor of 4.60 higher in the TGF-
-treated cells
than in untreated microglial cells. To rule out the possibility that
there are differences in RNA integrity between samples, one test is to
determine whether there is a change in level of a species of mRNA not
expected to be altered by the TGF-
1. A similar analysis of G3PDH
abundance in the same samples (Fig. 10C) showed a small
(23%) decrease in the cells exposed to TGF-
1. In three separate
determinations, Kv1.3 mRNA abundance increased by an average factor of
4.67 ± 0.55. However, the abundance of the G3PDH transcript did
not change (1.03 ± 0.18).
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The competitive PCR method provides a good quantitative estimate of
relative amounts of mRNA in different cell populations. By making
several assumptions, it is possible to estimate the absolute numbers of
Kv1.3 mRNA molecules in each cell. First, the efficiency of mRNA
isolation is assumed to be 100%. Second, in contrast to difference
measurements, the efficiency of RT-PCR of the target is assumed to be
identical to that of the competitor. The control cells in the
experiment in Fig. 10B had 11 mRNA molecules/cell, and the
TGF-1-treated cells had 56 mRNA molecules/cell. If the actual
efficiency of isolation is <100% or if the efficiency of RT-PCR is
less than the competitor, the true numbers of Kv1.3 mRNA molecules/cell
would be proportionately higher.
Because immunofluorescence of Kv1.5 antibodies has been reported in rat
microglia (21, 33), we used RT-PCR to probe for Kv1.5 mRNA
in TGF--treated BV-2 cells. In three determinations, we could not
detect mRNA for Kv1.5, although we did amplify G3DPH (as a control),
and the primer set we used did amplify Kv1.5 from an identical amount
of total RNA from mouse brain (data not shown).
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DISCUSSION |
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In the present study we provide the first evidence that a
deactivating cytokine is capable of changing the pattern of
K+ channel expression in microglia. We demonstrate that
TGF- induces upregulation of Kv1.3 mRNA and expression of the
corresponding DR K+ channels in murine microglia. The
expression of IR channels was not affected by TGF-
.
Exposure of microglia to any of several different activating stimuli
results in a characteristic pattern of parallel downregulation of IR
channels and upregulation of DR channels. Thus large DR currents but no
IR K+ currents were detected in the majority of activated
microglial cells grown with GM-CSF or stimulated with IFN-
(13). A significantly reduced IR current density in
parallel with an increased DR current density was observed in
LPS-activated microglia (9, 31). The pattern of
K+ channel expression after treatment of microglia with the
deactivating cytokine TGF-
differs in that DR upregulation occurred
in the absence of any change in the expression of IR channels.
Therefore, upregulation of DR channels in microglia is not obligatorily
linked to downregulation of IR channels.
IR channels in TGF--treated microglia, similar to those of untreated
microglia (10), were highly sensitive to extracellular Ba2+ but unaffected by
-DTX, an inhibitor of ROMK-1
channels (19). The pharmacological properties together
with the single channel conductance of ~30 pS suggests that Kir2.1
channels are expressed in TGF-
-treated microglial cells. With
RT-PCR, we demonstrate the existence of Kir2.1 mRNA in control and
TGF-
-treated microglia. Küst et al. (25) detected
mRNA for ROMK-1 channels in rat microglia. On the basis of the
pharmacological profile of the IR currents, it can be excluded that
ROMK-1 channels are expressed to any significant extent in murine
microglial cells before or after TGF-
treatment.
With respect to the single channel conductance, kinetic and
pharmacological properties, DR K+ currents in
TGF--treated microglia closely resemble those in activated microglia
(10) and in other macrophages (7, 15). We
conclude that the same type of DR channel is upregulated in microglia
after exposure to either deactivating or activating substances. We used
RT-PCR to show that Kv1.3 mRNA is present in murine microglial cells
and propose that it codes for the DR channels in TGF-
-treated
microglia. This hypothesis is supported by our observation that the
levels of mRNA for Kv1.3 increased on average 4.7-fold in microglia
after treatment with TGF-
, i.e., changes in mRNA levels were similar
to those in DR current density. It has been reported that antibodies to
Kv1.5 as well as to Kv1.3 stained LPS-treated rat microglia
(33) and rat microglia in cultured tissue prints
(21). We found no electrophysiological evidence of
significant expression of Kv1.5 channels in TGF-
-treated murine
microglia. Both CTX and KTX, which inhibit Kv1.3 but not Kv1.5
(16), blocked nearly all of the outward current.
Furthermore, we could not detect mRNA for Kv1.5 using RT-PCR in BV-2
microglial cells treated with TGF-
. In contrast with rat microglia,
murine microglia evidently express Kv1.3 but not Kv1.5 K+ channels.
Increases in Kv1.3 mRNA levels were seen in rat microglia after stimulation of adenosine A2a receptors. Kv1.3 channel expression was detected by immunocytochemical methods, but no patch-clamp studies were performed, so a quantitative comparison of channel expression and mRNA levels is not possible (25). When THP-1 cells were induced to differentiate from monocytes into macrophages, downregulation of DR channels and upregulation of IR channels were paralleled by changes of the corresponding Kv1.3 and Kir2.1 mRNA levels (8). In rat microglia after stimulation with LPS, changes in Kv1.3 mRNA levels were not observed, although the proportion of cells expressing DR channels increased dramatically (30). When we used competitive PCR to examine mRNA levels more quantitatively, we found that mRNA levels for Kv1.3 in microglia changed in parallel with K+ channel expression.
Because DR channels are upregulated in microglia stimulated with LPS,
IFN-, or GM-CSF, it was proposed that the expression of DR channels
is a marker for activated microglia (13, 31). Because the
same Kv1.3 DR channel is also upregulated on deactivation by TGF-
,
expression of this channel is not an unambiguous marker of activation
of microglial cells. However, it is clear that the pattern of
K+ channel expression changes predictably with a variety of
stimuli. Perhaps DR channel expression is upregulated during changes
from one functional state to another rather than in a defined
functional state of microglial cells. This idea is supported by the
finding that ramified resting microglia upregulated DR channels
immediately after exposure to astrocyte-conditioned medium but no
longer expressed DR currents several days after treatment with
astrocyte-conditioned medium (12).
A specific role for K+ channels in microglial signaling
remains to be established. Pharmacological lesion experiments suggest that Kv1.3 channels may play a permissive role in proliferation of
microglia (21), similar to that in T lymphocytes
(6). One general function of K+ channels in
leukocytes is to maintain a large driving force for Ca2+
influx through calcium-release-activated Ca2+ channels
(26). When K+ channels open, they drive the
membrane potential toward the Nernst potential for K+,
which is usually negative to the resting membrane potential. Further
experiments are required to demonstrate a similar functional role of
Kv1.3 channels in microglia. Leukocytes expressing IR K+
channels have more negative resting membrane potentials than cells with
only Kv1.3 K+ channels, suggesting that cells express IR
K+ channels when they need a very negative resting
potential and DR K+ channels if they need a moderately
negative resting potential (7). The finding that resting
membrane potential and expression levels of IR channels remained
unchanged in microglia after exposure to TGF- (data not shown) or to
astrocyte-conditioned medium (12) indicates that IR
channels determine the resting membrane potential of untreated and
deactivated microglial cells. That DR K+ channels activate
with depolarization means that they have a much greater capacity than
IR K+ channels to resist depolarizing influences. The
change in expression of K+ channels in response to TGF-
is different from that with activating stimuli, because IR channels are
not downregulated. These cells may, therefore, have a rather negative
membrane potential and be resistant to depolarization. It should be
noted that both deactivating cytokines such as TGF-
and activating
stimuli such as LPS have several effects in common, including reduction
of proliferation. In addition, activating stimuli may lead to release
of TGF-
from microglial cells (22).
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ACKNOWLEDGEMENTS |
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The authors thank Sieglinde Latta for the preparation of cell cultures and Astrid Düerkop for technical assistance.
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FOOTNOTES |
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This work was supported in part by the Deutsche Forschungsgemeinschaft Grant SFB 507/C3 (to C. Eder and U. Heinemann), the Michael Reese Health Trust (to F. N. Quandt), Research Grant HL-52671 from the National Heart, Lung, and Blood Institute (NHLBI) (to T. E. DeCoursey), and NHLBI Training Grant HL-07692 (to W. Zhou).
Address for reprint requests and other correspondence: C. Eder, Institut für Physiologie der Charité, Humboldt Universität, Tucholskystr. 2, D 10117 Berlin, Germany (E-mail: claudia.eder{at}charite.de).
The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
Received 2 February 2000; accepted in final form 26 April 2000.
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