Integrated Methodologies for Subcellular Protein Localization and Interaction Analysis in Eukaryotic Systems

This document details various advanced experimental protocols employed for the quantitative analysis of protein localization, interactions, and modifications within eukaryotic cells, leveraging a combination of high-resolution microscopy and sophisticated biochemical techniques.

Pericentrosomal Protein Localization and Quantification

Quantitative analysis of protein localization was performed using advanced microscopy techniques. For instance, the total fluorescence intensity of RTN4 or Calnexin within two 6-µm-diameter circular pericentrosomal regions (centered on the two mitotic centrosomes) was quantified manually using Fiji software. This intensity was then expressed as the ratio to the total cellular fluorescence intensity. Similarly, to quantify ER accumulation around the centrosome in mitotic C. elegans embryos, the fluorescence intensity of SP12 within an 8-µm-diameter circular pericentrosomal region (centered on the mitotic centrosome at a relatively low z-position in the AB cell) was manually measured using Fiji. This measurement was expressed as the ratio to that of an equivalent cortical area (a circular area of equal size at the AB cell cortex on the side away from the P1 cell). Maximum-intensity z-projections were generated from acquired three-dimensional image stacks before quantification, typically obtained using a 0.4 NA oil-immersion objective.

For further guidance on cellular quantification techniques, including principles applicable to fluorescence analysis, one might consult resources such as the Flow Cytometry Guides .

Protein Purification and Interaction Assays

GST Pull-down and In Vitro Binding Assays

For GST pull-down assays, Escherichia coli Rosetta (DE3) cells expressing GST-tagged proteins were harvested and lysed by ultrasonication in pull-down buffer (see Table 1) containing protease inhibitors on ice. Following centrifugation, the supernatant was incubated with Glutathione Sepharose 4B beads (Cytiva, 17075601) for 2 hours at 4°C. After extensive washing, GST-tagged proteins were eluted with 10 mM reduced glutathione (Amresco, 0399).

The Strep-tagged fusion protein 2×Strep-HA-RTN4B was stably expressed in HeLa cells. Harvested cells were lysed in a cell lysis buffer (see Table 2) containing protease and phosphatase inhibitors. After centrifugation, the supernatant was incubated with Strep-Tactin XT 4Flow resins for 2 hours at 4°C. Following extensive washing, the fusion protein-bound beads were utilized for in vitro binding assays.

For in vitro binding assays, beads coated with Strep-tagged proteins were incubated with purified GST-tagged proteins for 2 hours at 4°C. After extensive washing with lysis buffer, bound proteins were eluted using Strep-Tactin XT elution buffer (IBA Lifesciences, 2-1042-025). Eluted samples were subsequently analyzed by Western blotting or Coomassie Brilliant Blue staining.

In GST pull-down assays, asynchronous and STLC-arrested mitotic HeLa cells stably expressing 2×Strep-HA-RTN4B were lysed in cell lysis buffer containing protease and phosphatase inhibitor cocktails. After centrifugation, the supernatant was incubated with GST-fusion protein-bound beads for 2 hours at 4°C. The beads were washed extensively with cell lysis buffer and boiled in sample loading buffer. Samples were then analyzed by Western blotting or Coomassie Brilliant Blue staining.

Immunoprecipitation and Mass Spectrometry Sample Preparation

For mass spectrometry sample preparation, cells were collected and lysed in lysis buffer. After centrifugation, the supernatants were incubated with either protein G-Sepharose beads (Cytiva, 17061805) prebound with antibodies or anti-Flag M2 affinity gels (Millipore, A2220) for 2 hours at 4°C. Following four washes with lysis buffer, the beads were boiled in protein SDS loading buffer. Samples were separated by SDS-PAGE and stained with Coomassie Brilliant Blue. After tryptic digestion, proteins were analyzed using an Orbitrap Fusion Lumos mass spectrometer (Thermo Fisher Scientific) to identify protein post-translational modifications.

Kinase Assays

HeLa cells stably expressing 3×Flag-mEmerald-RTN4B WT or 6A mutant were harvested and lysed in a specific lysis buffer (see Table 3) containing protease and phosphatase inhibitors. After centrifugation, the supernatant was incubated with anti-Flag M2 affinity gels for 2 hours at 4°C. Flag-tagged fusion proteins were eluted with 100 µg/mL 3×FLAG peptide. The purified 3×Flag-mEmerald-RTN4B WT or 6A mutant was then incubated with active CDK1/Cyclin B1 complex (Abcam, ab271456) in 1× kinase buffer (see Table 4) containing phosphatase inhibitors and 500 µM ATP (NEB, P0756S) at 30°C for 45 minutes. The reaction was terminated by adding SDS sample loading buffer, and the samples were analyzed by Western blotting using the appropriate antibodies.

Buffer Compositions

Electron Microscopy Sample Preparation and Imaging

Transmission Electron Microscopy (TEM)

Wild-type and RTN4 knockout HeLa cells were plated on ACLAR 33C film (Electron Microscopy Sciences, 50425) and fixed with 2.5% (v/v) glutaraldehyde (Sigma-Aldrich, G5882) in 0.1 M phosphate buffer (PB, pH 7.4). After fixation, samples were extensively washed with 0.1 M PB and post-fixed for 30 minutes at room temperature in the dark using a mixture of 1% osmium tetroxide and 0.8% potassium ferrocyanide (Sigma-Aldrich, 60279). Samples were subsequently incubated in a 1% aqueous solution of uranyl acetate and then washed with double-distilled water (ddH ₂ O). Dehydration was performed through a graded ethanol series (30%, 50%, 70%, 85%, 95%, and 100%; 6 minutes each), followed by two 6-minute changes in 100% acetone. The cells were gradually infiltrated with Embed 812 resin (Electron Microscopy Sciences, 14120) and polymerized at 65°C for 24 hours. After polymerization, the ACLAR 33C film was detached from the polymerized resin blocks before trimming. Ultrathin sections (70 nm thick) were cut using an ultramicrotome (Leica Microsystems, UC7) fitted with a diamond knife (Diatome, Ultra 35°). Serial sections were mounted on single-slot copper grids and stained with uranyl acetate and lead citrate. Grids were examined using a transmission electron microscope (Thermo Fisher Scientific, Tecnai G2 Spirit BioTWIN) operated at an accelerating voltage of 120 kV. Micrographs were acquired using a digital camera (Gatan, Orius 832).

Focused Ion Beam-Scanning Electron Microscopy (FIB-SEM)

HeLa cells stably expressing 3×Flag-mEmerald-RTN4B WT were synchronized with thymidine for 20 hours, washed three times, and released into fresh medium for 7–8 hours. Samples were immediately fixed with 4% (w/v) PFA. Mitotic HeLa cells from prophase to prometaphase were selected based on rounded morphology and RTN4 fluorescence intensity using a Zeiss LSM980 confocal microscope equipped with a 20×/0.8 NA air objective. Cells were subsequently fixed with 2.5% (v/v) glutaraldehyde in 0.1 M PB for 1 hour at room temperature. After four washes with 0.1 M PB, samples were post-fixed with 1% osmium tetroxide and 0.8% potassium ferrocyanide for 1 hour at room temperature and washed with ddH ₂ O. The samples were then treated with 1% thiocarbohydrazide (Sigma-Aldrich, 223220) for 20 minutes at room temperature (protected from light), followed by washing with ddH ₂ O. Samples were then treated again with 1% osmium tetroxide for 30 minutes at room temperature. After rinsing again with ddH ₂ O, the samples were stained overnight with 1% uranyl acetate at 4°C. Following a final rinse with ddH ₂ O, samples were dehydrated through a graded series of ethanol (30%, 50%, 70%, 85%, 95%, and three changes of 100%; 6 minutes each) and embedded in Embed 812 resin. The cured resin blocks were trimmed and sectioned using an ultramicrotome (Leica Microsystems, UC7) until the selected mitotic cells were tangentially exposed for FIB-SEM imaging. FIB-SEM imaging was performed using a Helios Nanolab G3 UC (Thermo Fisher Scientific). Images were acquired at a 10.74 nm pixel size with a 2 kV accelerating voltage and 0.2 nA beam current using the in-column energy-selective backscattered electron detector (ICD). A 10 nm FIB step size (z-thickness) was obtained with a 30 kV, 2.5 nA gallium beam.

Image Segmentation and Analysis

Instance segmentation of intracellular structures was performed using the empanada-napari software package, following a previously described pipeline for applying a generalist deep learning model to instance segmentation in electron microscopy images [56]. Initially, a subset of the FIB-SEM images was manually annotated to generate preliminary training labels. These annotated datasets were then used to fine-tune the generalist deep learning model integrated within the empanada-napari platform. The model was iteratively refined by adjusting inference parameters and retraining with updated annotations until satisfactory 2D segmentation performance was achieved. This optimized model was subsequently applied to generate 3D segmentations.