A role for ion channels in glioma cell invasion

Abstract

Many cells, including neuronal and glial progenitor cells, stem cells and microglial cells, have the capacity to move through the extracellular spaces of the developing and mature brain. This is particularly pronounced in astrocyte-derived tumors, gliomas, which diffusely infiltrate the normal brain. Although a significant body of literature exists regarding signals that are involved in the guidance of cells and their processes, little attention has been paid to cell-shape and cell-volume changes of migratory cells. However, extracellular spaces in the brain are very narrow and represent a major obstacle that requires cells to dynamically regulate their volume. Recent studies in glioma cells show that this involves the secretion of Cl(-) and K(+) with water. Pharmacological inhibition of Cl(-) channels impairs their ability to migrate and limits tumor progression in experimental tumor models. One Cl(-)-channel inhibitor, chlorotoxin, is currently in Phase II clinical trials to treat malignant glioma. This article reviews our current knowledge of cell-volume changes and the role of ion channels during the migration of glioma cells. It also discusses evidence that supports the importance of channel-mediated cell-volume changes in the migration of immature neurons and progenitor cells during development. New unpublished data is presented, which demonstrates that Cl(-) and K(+) channels involved in cell shrinkage localize to lipid-raft domains on the invadipodia of glioma cells and that their presence might be regulated by trafficking of these proteins in and out of lipid rafts.

Fig. 1. K+ and Cl− efflux aids glioma-cell shrinkage

It is thought that cells must shrink as they invade the brain. To reduce their cytosplasmic content, cells release Cl⁻ and K⁺ through ion channels, and water follows passively through water channels or aquaporins. For this mechanism to function, cells must accumulate Cl⁻ and K⁺ above their respective electrochemical gradients. This is accomplished by the combined activity of the NKCC Cl⁻ transporter and the Na⁺–K⁺ ATPase for K⁺ ions. Modified, with permission, from Soroceanu et al. (1999)

Fig. 2. Glioma cells shrink as they invade

Images obtained by either light (A) or electron (B) microscopy. (A) To demonstrate the polarized, wedge-shape of invading glioma cells, which indicates cell shrinkage, we stably transfected D54-MG glioma cells with EGFP. These cells were placed on the surface of a 400-μm thick slice from rat brain and allowed to invade for 6 hours in a fully oxygenated chamber at 37°C. A series of confocal images was obtained (400 nm optical sections), which allows complete, 3-D reproduction of invading cells. Blood vessels are stained with CD31-Abs conjugated to phycoerythrin. (B) For electronmicroscopy of invading cells, D54-MG cells were grown in a spheroid and confronted with a spheroid of fetal rat brain cells. 20-nm sections are shown at 12 000× magnification. Images in (B) are reproduced, with permission, from Soroceanu et al. 1999.

Fig. 3. Glioma cells take advantage of a resting Cl⁻ conductance as they invade

(A) To demonstrate the presence of a resting Cl⁻ conductance, recordings were made from glioma cells using amphotericin-perforated, whole-cell patch-clamp recordings. A holding current of −200 pA maintained the cell at −40 mV (left axis). Cells had input resistances of ~100 MΩ (right axis). Application of the Cl⁻-channel inhibitor NPPB inhibited the holding current and increased cell resistance to ~200 MΩ, which is consistent with inhibition of a resting Cl⁻ conductance. By contrast, TEA did not alter the input resistance or the holding current. (B) To show that a NPPB-sensitive Cl⁻ conductance is required for successful migration across a spatial barrier, D54-MG glioma cells were plated on the upper surface of a Transwell insert with 8 μm pores and allowed to migrate for 4 hours towards vitronectin coated on the bottom of the filter insert (top left). Under control conditions, most cells migrated successfully, indicated by crystal violet staining of cells at the bottom (top right). In the presence of 30 μM NPPB only few cell migrated to the bottom of the filter (bottom right). Instead, most cells extend a process through the filter, but failed to move the entire cell through (bottom left). Modified, with permission, from Ransom et al. (2001).

Fig. 4. Glioma cells express functional ClC-3 Cl⁻ channels

(A) D54-MG glioma cells in culture labeled using antibodies to ClC-3. Channels were identified with a secondary antibody conjugated to 6-nm immuno-gold particles. (B) Lysates of cultured D54-MG cells contain ClC-3. To suppress expression of this protein, sister cultures were treated for 48 hours with antisense oligonucleotides against ClC-3. This suppressed the concentration of ClC-3 significantly but did not affect expression of ClC-, another Cl⁻ channel. (C) Whole-cell patch clamp recordings reveal the presence of outwardly rectifying Cl⁻ currents with time-dependent inactivation in control D54-MG cells. These currents are sensitive to NPPB, DIDS and Cltx. (D) Sister cultures treated with ClC-3 antisense show a significant (>50%) suppression of these currents. Modified, with permission, from Olsen et al. (2003).

Fig. 5. ClC-3 and BK channels localize to lipid raft domains on invadipodia of glioma cells

(A) Fluorescein-conjugated cholera toxin B subunit was used to label lipid-raft domains of D54-MG cells grown on glass coverslips. Cells were fixed and labeled with polyclonal, rabbit anti-BK K⁺ primary antibodies followed by secondary labeling with Alexa 546 goat anti-rabbit antibodies to illustrate the localization of BK K⁺ to lipid-raft domains. (B) Sister cultures were labeled with fluorescein-conjugated cholera toxin B subunit followed by labeling with polyclonal anti-ClC-3 antibodies, which exhibited similar localization patterns. (C) Lipid rafts were isolated from D54-MG cells by subcellular fractionation followed by density-gradient centrifugation. Proteins in each fraction were separated by SDS-PAGE and Western blotted for BK channels, ClC-3 channels and the caveolar lipid raft marker caveolin-1. Fractions W and D represent water-soluble and detergent-soluble fractions, respectively. Fractions 1–9 represent the fractions from the top to the bottom following density-gradient centrifugation (5–40% Optiprep) of the detergent insoluble fraction. Fraction 2 contains the buoyant lipid-raft fraction, as evidenced by caveolin-1 controls. BK and ClC-3 channels partition into this lipid-raft fraction.

Fig. 6. Cltx inhibits glioma Cl⁻ currents by inducing endocytosis of channels into caveoli

(A) Outwardly rectifying, inactivating Cl⁻ currents recorded by whole-cell patch clamp recording in D54-MG glioma cells using voltage steps ranging from −120−160 mV (20 mV increments). Application of Cltx causes a slow and irreversible reduction of measurable currents. The example illustrated was recorded after 15 minutes. (B) Recombinant 6×-His-Cltx peptide was used to affinity purify interacting proteins from glioma lysates. The affinity-purified fraction was run on SDS-PAGE and probed with antibodies to ClC-3 to detect the channel in lysates from glioma cells but not from astrocytes. (C) A surface biotinylation approach, previously described by Ye et al. (1999) was used to assess the surface expression of ClC-3 protein. Separation over avidin beads isolated the cell-surface proteins that were accessible to the biotinylation reagent at the time of exposure. This fraction was probed by Western blot with antibodies to ClC-3 and anti-His to detect bound His-Cltx. ClC-3 protein was present in the membrane of untreated glioma cells. Treatment with Cltx for 30 minutes before exposure to the biotinylation reagent reduced the amount of ClC-3 on the membrane surface. Disrupting caveolar endocytosis with 5 μg ml⁻¹ Filipin (a sterol-binding drug) prevents Cltx-mediated reduction in surface expression of ClC-3 in these cells, which indicates that Cltx causes endocytosis of ClC-3 into caveoli.