Pannexin1 Drives Multicellular Aggregate Compaction via a Signaling Cascade That Remodels the Actin Cytoskeleton^*

3D Multicellular Scaffold-free Assays

A modified version of our 3D non-adhesive hydrogel system (20) was used to examine C6 proliferation and multicellular aggregate assembly. Briefly, agarose hydrogels were produced by pouring a sterile 3% molten agarose solution (500 μl) into 3D PetriDish^TM micromolds (MicroTissues, Inc.) designed for 24-well tissue culture plates. After allowing the agarose to set for 10 min, the gels were separated from the micromolds according to the manufacturer's protocol and transferred to the tissue culture plates with the seeding chambers facing upwards (1 gel/well). The gels were briefly degassed and then equilibrated overnight in the appropriate culture medium. For the proliferation assay, we used DMEM with 10% FBS and 1% penicillin/streptomycin, and for aggregate assembly assays, we used serum-free DMEM with 1% penicillin/streptomycin and, if applicable, the drug of interest: carbenoxolone (CBX) (50–100 μm), probenecid (PBN) (200 μm), ATP (500 μm), suramin (100 μm), 2′,3′-(benzoyl-4-benzoyl)-ATP (BzATP) (200–400 μm), oxidized ATP (oATP) (200 μm), brilliant blue G (1–5 μm), 2-methylthio-ADP (MeSADP) (250 μm), pyridoxal-phosphate-6-axophenyl-2′,4′-disulphonic acid (PPADS) (100 μm), and UTP (100 μm). Stock solutions of drugs were prepared according to the manufacturers' protocols, and appropriate vehicle controls were used. Spheroid micromolds with a rectangular array of 96 (8 × 12) cylindrical round-bottom recesses (400-μm diameter, 800-μm depth) were used to study proliferation and for assembly assays requiring fluorescent protein visualization. Rod micromolds containing an array of 24 (3 × 8) trough-shaped round-bottom recesses (2200-μm length, 400-μm width, 800-μm depth) were used for assembly assays requiring quantification of compaction kinetics.

Seeding of cells into gels, considered 0 h, was as follows. C6 cells grown to near confluence in DMEM with 10% FBS and 1% penicillin/streptomycin in 37 °C and 5% CO₂ were trypsinized for 5 min, counted, and resuspended to the desired seeding density: 100 cells/60 μl for the proliferation assay, 0.12 × 10⁶ cells/60 μl for spheroid-based assembly assays, and 0.6 × 10⁶ cells/60 μl for rod-based assembly assays. Cells were seeded by pipetting 60 μl of cell suspension into the seeding chamber and allowed to settle for 10 min before they began to spontaneously assemble into aggregates. 1 ml of additional culture medium was added before time-lapse microscopy began.

Microscopy and Image Analysis

Proliferation and aggregate compaction kinetics were quantified by analyzing time-lapse phase contrast images taken with an Axio Observer Z1 microscope equipped with an AxioCam mRm camera and a humidified chamber with temperature (37 °C) and CO₂ (5%) control (Carl Zeiss MicroImaging). To quantify proliferation kinetics, images were taken every 24 h for 13 days. The fit ellipse tool in ImageJ (National Institutes of Health) was then used to measure the area of the growing tumor spheroid. To quantify rod aggregate compaction, images were taken every hour for 9 to 24 h. The line tool in ImageJ was used to measure rod length, defined as the long axis (x axis) length normalized to the 0 h length. Percent inhibition by drug treatment was defined as the percent difference between the drug-treated and control rod lengths. Visualization of specific proteins (Panx1-eGFP, calcein acetoxymethyl, F-actin) within aggregates was achieved by harvesting the aggregates via gel inversion and brief centrifugation followed by imaging with a LSM 510 meta confocal laser scanning microscope (Zeiss).

Protein Visualization and Immunostaining

Panx1-eGFP was visualized within homocellular aggregates directly without fixation. For live-cell visualization of Panx1-eGFP and Panx2-eGFP in heterocellular aggregates, Panx2 cells were first stained with 5 μm CellTracker Blue 7-amino-4-chloromethylcoumarin (Invitrogen) for 45min before being mixed with Panx1 cells and seeded into gels. For immunostaining of F-actin, aggregates seeded 12 h earlier were washed in PBS, fixed in 4% paraformaldehyde for 1 h, and washed again. Samples were then permeabilized with 0.5% Triton X-100 for 30 min, followed by additional washes with PBS. Aggregates were then stained with Alexa Fluor 568 phalloidin (Invitrogen) for 45 min. After a final wash, aggregates were imaged. Aggregate viability with drug treatment was determined using the live/dead viability/cytotoxicity kit (Invitrogen) as described previously (21).

Panx1 Channels Accelerate Compaction of Multicellular C6 Glioma Aggregates

To determine whether our 3D system can recapitulate tumor behavior in vivo, we developed an assay investigating the role of Panx1 on C6 glioma tumorigenicity to compare with our previous in vivo findings (18). Briefly, wild-type C6 cells and those stably transfected with Panx1-eGFP were trypsinized and seeded into micromolded agarose gels with circular recesses so that there was only 1 cell/recess initially (100 cells into 96 recesses/gel). The area of the growing tumor spheroids was measured over 13 days (Fig. 1A). Mirroring our in vivo results, there was minimal growth within 48 h, but from day 7 onwards, Panx1 spheroids were significantly smaller than their wild-type counterparts (day 13, 23,983 ± 9851 μm² versus 60,706 ± 17,810 μm²) (n = 20).

To quantify the role of Panx1 in 3D biomechanics and the assembly of large multicellular structures, an alternate assay was employed. Mono-dispersed wild-type and Panx1 cells, as well as C6 cells expressing Panx2-eGFP, were seeded into gels with rod-shaped recesses so that there were ∼25,000 cells/recess (0.6 × 10⁶ cells into 24 recesses/gel). Panx2 cells were chosen as an additional control, given their intracellular localization (19) and subsequent lack of channel activity (22). Aggregation and compaction kinetics were quantified by measuring the length of the contracting rod aggregates and normalizing this value to its initial length over 24 h, prior to any significant proliferation in our 3D system (Fig. 1B). Panx1 multicellular rods displayed significantly accelerated compaction compared with wild-type and Panx2 rods, whose kinetics were comparable. Panx1 aggregate compaction was most dynamic from 3–15 h and slowed toward equilibrium.

To determine whether the acceleration in 3D assembly was directly correlated with Panx1 channel activity, cells were seeded into gels pre-equilibrated with CBX (50–100 μm) or PBN (200 μm). CBX is a potent gap junction inhibitor that directly blocks Panx1 channels (23) in a dose-dependent manner, whereas PBN is specific for Panx1 (24). The percent inhibition of rod contraction with drug treatment was determined at 12 h, within the dynamic range of assembly, and shows that spontaneous Panx1 rod contraction was inhibited by both PBN (10.7 ± 2.1%) and CBX, with the latter displaying a dose-dependent relationship (16.8 ± 7.0% for 50 μm and 34.5 ± 2.1% for 100 μm) (n = 10) (Fig. 1C). As expected, wild-type and Panx2 rods were unaffected by drug treatment. Drug effects were observed in Panx1 aggregate compaction as early as 3 h and maintained through 24 h, with live/dead analysis confirming that viability was not compromised (data not shown). Interestingly, we observed that CBX caused an internalization of Panx1-eGFP, normally localized in a uniform manner on the cell membrane, in cells within aggregates (Fig. 1D).

Panx1 within the Heterocellular Microenvironment

Having examined Panx1 in only homocellular aggregates, it was important to extend our investigation to include a heterocellular environment. To this end, we observed the behavior of heterocellular Panx1/Panx2 aggregates by titrating the proportion of Panx1 cells in 10% increments while keeping the total cell number constant. The normalized rod lengths at four different times over 24 h (6 h, 12 h, 18 h, 24 h) were measured as a function of the proportion of Panx1 cells (Fig. 2A). For all times measured, we found that increasing the Panx1 cell percentage resulted in a linear shortening in rod length (6 h r²0.949, 12 h r²0.967, 18 h r²0.980, 24 h r²0.991). The slopes of these four curves were nearly constant (−0.0013, −0.0015, −0.0015, and −0.0014, respectively), suggesting that heterocellular Panx1/Panx2 aggregate compaction is dependent on the proportion of the two constituent cell types.

To determine whether localization of either Panx1 or Panx2 proteins had been altered, unlabeled Panx1 cells were mixed with Panx2 cells labeled with CMAC (blue) at 50%:50% (Fig. 2B). Confocal images at 24 h confirm that trafficking of Panx1-eGFP and Panx2-eGFP were unchanged in the heterocellular aggregates. Interestingly, we also observed a tendency of Panx1 cells to compartmentalize to the center of these heterocellular aggregates.

Given reports of possible heteromeric Panx1/Panx2 interactions when coexpressed within the same cell (25), we compared the assembly kinetics of C6 cells expressing both Panx1 and Panx2 to C6 rods expressing Panx1 alone (Fig. 2C). The Panx1 + 2 rods compacted as quickly as the Panx1 rods, both significantly faster than the wild-type control.

Panx1-mediated ATP Release Drives Compaction via a Signal Cascade That Up-regulates Cytoskeletal Function

Rather than forming gap junctions, recent evidence suggests that Panx1 channels release ATP into the extracellular space, which then binds to cell surface purinergic receptors (26). A modified dye transfer assay confirmed the absence of gap junction-mediated coupling within Panx1 aggregates (supplemental Fig. 1). Having established the relationship between Panx1 channel activity and aggregate assembly, we then determined if this effect was facilitated by the specific release of ATP. We found that addition of exogenous ATP significantly, although not completely, reverses the inhibitive effects of CBX on Panx1 aggregate compaction (Fig. 3A). Panx1 cells in gels pre-equilibrated with ATP (500 μm) and CBX (50 μm) formed more compact aggregates, resembling the control rather than the loose aggregates seen with CBX treatment alone. Live (green)/dead (red) images at 24 h confirm that viability was not compromised by treatment with CBX and/or ATP. Furthermore, the addition of exogenous ATP (500 μm) alone could accelerate compaction of both wild-type and Panx1 aggregates (Fig. 3B). Although the effect on wild-type aggregates was immediate and consistent through 24 h, its onset was delayed but more pronounced for Panx1 aggregates.

Given that extracellular ATP can bind to P₂X₇ cationic channel (27) and P₂Y₁/P₂Y₂ G-protein coupled (28) purinergic receptors, both shown to be expressed in C6 glioma cells (29, 30), we then determined which receptor pathway is dominant in aggregate compaction. Wild-type and Panx1 rods were first treated with suramin (100 μm), a nonspecific purinergic antagonist. Although wild-type rods were unaffected, suramin immediately inhibited Panx1 rod compaction (Fig. 4A). To differentiate between the P₂X₇ and P₂Y₁/P₂Y₂ pathways, samples were then treated with a complimentary series of agonists/antagonists specific for P₂X₇ or P₂Y₁/P₂Y₂. When treated with the P₂X₇ agonist BzATP (200–400 μm), Panx1 rods displayed a significant acceleration in assembly that was dose-dependent (Fig. 4B) and more pronounced than that with ATP (500 μm) at 12 h. When concomitantly treated with BzATP (200 μm) and the P₂X₇ antagonist oATP (200 μm), the BzATP-induced effect was mitigated as these Panx1 rods compacted similarly to the untreated control. Wild-type aggregates were unaffected by treatment with BzATP and/or oATP. Interestingly, treatment with the P₂X₇ antagonist BBG (1–5 μm) did not affect Panx1 rod compaction (supplemental Fig. 2). Wild-type and Panx1 rods were then treated with reagents specific for P₂Y₁ (agonist MeSADP, antagonist PPADS) and P₂Y₂ (agonist UTP) (Fig. 4C). Neither wild-type nor Panx1 aggregates were affected. Taken together, these findings implicate the involvement of Panx1-mediated ATP release and subsequent P₂X₇ purinergic receptor activation in aggregate assembly.

The actomyosin cytoskeleton system is involved in aggregate compaction upon initial cell-cell adhesion (11). We therefore investigated whether this Panx1/ATP/P₂X₇ cascade modulated the F-actin cytoskeletal network (Fig. 5A). In comparison to wild-type aggregates, which possess a moderate cortical F-actin network and spherical cell morphology, we found that Panx1 aggregates possessed a significantly more robust F-actin cytoskeleton. In addition, the F-actin distribution extended to non-cortical regions, whereas cell morphology became flattened. Not surprisingly, these same changes became even more pronounced with the addition of exogenous ATP. Although CBX reduced F-actin expression, the addition of exogenous ATP to CBX-treated aggregates reverted the cytoskeletal phenotype to resemble that of the untreated control. Within heterocellular Panx1/Panx2 aggregates, the F-actin network of Panx1 cells is noticeably more developed than that of Panx2 cells, which remains mostly cortical (Fig. 6A). To further explore this relationship, wild-type and Panx1 aggregates were treated with cytochalasin B (5 μm), a potent inhibitor of actin polymerization (Fig. 5B). Although assembly was nearly arrested by drug treatment in both samples, Panx1 aggregates alone were able to resume compaction upon drug washout at 3 h. Confocal images at 12 h confirm that cytochalasin B disrupted the F-actin network within Panx1 aggregates and that this effect was reversed with washout (Fig. 5C).