Microtubule Staining in S2 Cells

For microtubule staining, S2 cells were rinsed in BRB80 (80 mM Pipes, pH 6.9, 1 mM MgCl₂, 1 mM EGTA) and fixed in the same buffer containing 0.5% glutaraldehyde (EM Sciences), 3% formaldehyde (EM Sciences), and 1 mg/ml saponin for 10 min. The cells were then permeabilized in PBS containing 0.5% SDS, treated with sodium borohydride, and blocked with 5% normal goat serum in PBS/0.1% Triton X-100. In experiments examining the localization of Dm EB1, cells were fixed for 10 min by immersion in a solution of 90% methanol, 3% formaldehyde, 5 mM sodium carbonate (pH 9) chilled to −80°C. Samples were then rehydrated into PBS/0.1% Triton X-100 and blocked as above. All antibodies were diluted into 5% normal goat serum in PBS/Triton (DM1α, 1:500; rabbit anti-EB1, 1:1,000) and applied to the fixed cells for 1 h followed by extensive washing with PBS/Triton X-100. Fluorescent secondary antibodies (Cy2-conjugated anti–rabbit and rhodamine-X–conjugated anti–mouse; Jackson ImmunoResearch Laboratories) were used at a final dilution of 1:300. After antibody staining, cells were treated with DAPI (0.5 μg/ml in PBS) for 10 min, briefly rinsed with distilled water, and mounted in 90% glycerol, 10% 0.1 M borate, pH 9.0, plus 5% n-propyl gallate. Specimens were imaged by confocal microscopy (TCS; Leica) and presented as maximum intensity projections.