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In This Article

  • Summary
  • Abstract
  • Introduction
  • Protocol
  • Results
  • Discussion
  • Disclosures
  • Acknowledgements
  • Materials
  • References
  • Reprints and Permissions

Summary

Here, a novel quantitative fluorescence assay is developed to measure changes in the level of a protein specifically at centrosomes by normalizing that protein’s fluorescence intensity to that of an appropriate internal standard.

Abstract

Centrosomes are small but important organelles that serve as the poles of mitotic spindle to maintain genomic integrity or assemble primary cilia to facilitate sensory functions in cells. The level of a protein may be regulated differently at centrosomes than at other .cellular locations, and the variation in the centrosomal level of several proteins at different points of the cell cycle appears to be crucial for the proper regulation of centriole assembly. We developed a quantitative fluorescence microscopy assay that measures relative changes in the level of a protein at centrosomes in fixed cells from different samples, such as at different phases of the cell cycle or after treatment with various reagents. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region, and normalize that measurement against the same for another protein that does not vary under the chosen experimental condition. Utilizing this assay in combination with BrdU pulse and chase strategy to study unperturbed cell cycles, we have quantitatively validated our recent observation that the centrosomal pool of VDAC3 is regulated at centrosomes during the cell cycle, likely by proteasome-mediated degradation specifically at centrosomes.

Introduction

Centrosomes consist of a pair of centrioles surrounded by pericentriolar material (PCM). Being the major microtubule organizing centers (MTOCs) in mammalian cells, centrosomes serve as the two poles of mitotic spindles in dividing cells, and thus help maintain genomic integrity1. In quiescent cells (e.g., during G0 phase), one of the two centrioles of the centrosome, namely the mother centriole, is transformed into a basal body to assemble the primary cilium, a sensory organelle protruding out from the cell surface2. Once the cells re-enter the cell cycle, primary cilia are disassembled and each centriole directs the assembly of a procentriole at its proximal end that gradually elongates to form a mature centriole3. At the onset of S-phase, a cartwheel-like structure that provides the 9-fold symmetry to the centriole is formed on the surface of each existing centriole and will become the base of each procentriole. Sas6 that is indispensable for centriole assembly is recruited to form the hub of the cartwheel4-6. Other centriolar proteins are then assembled onto the cartwheel in a highly regulated, proximal to distal manner7. After precisely completing centriole duplication, cells assemble additional pericentriolar materials to build two functional centrosomes by the end of G2 phase8. In addition to the core centriolar components9-11, several other proteins including kinases, phosphatases, chaperones, scaffold components, membrane associated proteins and degradation machinery are associated with centrioles, basal bodies and PCM at different times of the cell cycle12-16. It is often noted that the centrosomal levels of many proteins are temporally regulated by centrosomal targeting mechanisms and/or proteasomal degradation at centrosomes. Importantly, the fluctuations in the centrosomal level of several proteins such as Plk4, Mps1, Sas6, and CP110 at different points of the cell cycle appears to be crucial to regulate centriole assembly5,17-22, and in the case of Mps1 preventing this centrosomal degradation leads to the formation of excess centrioles19. On the other hand, the centrosomal fractions of several proteins are less labile compared to cytosolic pools. For example siRNA-mediated down-regulation of Centrin 2 (Cetn2) led to only a moderate decrease of the protein level at the centrioles despite great reduction in its whole cell levels23. It is therefore crucial to measure the changes in the level of centrosomal proteins at the centrosome rather than measuring the whole cell protein levels when assessing their centrosome-specific functions.

In this study we have developed an assay using indirect immunofluorescence (IIF) to quantify the relative level of a protein at centrosomes. This assay is developed particularly to analyze cells that are from different samples and thus can not be imaged at the same time. These samples can be cells that were treated with different reagents (i.e., drug versus control), collected at different time points (i.e., pulse versus chase), or are in different phases of the cell cycle. The principle of this assay lies in measuring the background corrected fluorescent intensity corresponding to a protein at a small region and to normalize that value against the same for another protein whose levels do not vary under the chosen experimental conditions. Several studies in centrosome biology have recently utilized various quantitative microscopy techniques, in both live or fixed cells, to determine the centrosome-specific function of candidate proteins24-27. Similar to those assays, the present technique also measures the background corrected fluorescence intensity of the test protein. However, the inclusion of the normalization using an internal standard in this assay would likely offer greater accuracy and confidence in analyzing two different samples that are on two different coverslips. Moreover, in addition to examining the protein level at centrosomes, with minor adjustments this method can be applied to a diverse set of experimental conditions or at other cellular sites.

Here, we combine our quantitative microscopy assay with a BrdU pulse-chase strategy to compare cells from different cell cycle stages. Instead of using standard cell cycle arrest and release techniques to study various cell cycle time points, asynchronously growing cells are incubated with BrdU to label cells in S-phase, and the labeled cells are chased for various times (typically 4-6 hr). Most of the labeled cells will be in S-phase immediately after the pulse. The length of the chase is chosen so that after the chase, labeled cells will be in late S, G2, or mitosis, which can be verified by morphological characteristics such as- position of centrosomes with respect to nuclei, distance between centrosomes, condensation of chromosomes etc. Thus, the length of the chase depends on the average duration of S, G2 and M phase of a particular cell type. Since this approach avoids cell cycle inhibitors such as hydroxyurea, aphidicolin, nocodazole, etc., it allows a more physiologically relevant cell cycle analysis.

Thus, we demonstrate here that the quantitative fluorescence microscopy assay alone, or in combination with BrdU pulse-chase assay, is a simple yet powerful technique to accurately measure the relative changes in the centrosomal level of a candidate protein during an unperturbed cell cycle. We measured the centrosomal level of VDAC3, a protein that we recently identified at centrosomes in addition to mitochondria16,28, using these assays. Results obtained here verify our previous observation that the centrosomal pool of VDAC3 is regulated by degradation, and also varies in a cell cycle dependent manner16, furthermore validating the applicability of this method.

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Protocol

1. Cell Culture

  1. Use human telomerase reverse transcriptase (hTERT) immortalized retinal pigment epithelial cells (hTERT-RPE1; referred to here as RPE1).
    NOTE: RPE1 cells are near-diploid, non-transformed human cells that are commonly used to study centriole assembly and ciliogenesis. These cells follow a normal centriole duplication cycle coordinated with a regulated cell cycle.
  2. Passage a near-confluent 100 mm cell culture dish of RPE1 cells at 1:5 dilution of the original culture into a fresh 100 mm dish containing 10 ml of DME/F-12 (1:1) medium supplemented with 10% Fetal Bovine Serum (FBS), 100 U/ml penicillin G and 100 µg/ml streptomycin (the complete medium is referred as DME/F-12 medium). Grow cells at 37 °C in the presence of 5% CO2.
    1. Incubate 12 mm round glass coverslips in 50 ml of 1 N HCl in a covered glass beaker, O/N without shaking, at 60 °C in a water bath or incubator.
    2. After discarding the acid solution, wash the coverslips three times in 100 ml distilled water by incubating for 15 min with occasional shaking. Repeat the washing similarly in 70% Ethanol and 95% Ethanol.
    3. Dry the coverslips by individually spreading them out on a laboratory blotting paper in a biosafety cabinet, followed by sterilization with UV radiation for 60 min.
  3. Place 2-4 dry coverslips in a 35 mm cell culture dish. Dilute the solution of fibronectin or fibronectin like engineered polymer (stock concentration of 1 mg/ml) to a concentration of 25 µg/ml in cell culture PBS.
    1. Place 80-100 µl of the diluted solution onto each coverslip and incubate for 60 min to coat the top side of the coverslip. Wash the coverslips thrice using PBS before adding complete medium.

2. Growing Cells and Treating Cells with Proteasome Inhibitors

  1. Passage 2 x 105 asynchronously growing RPE1 cells in each 35 mm dish containing coverslips and grow the cells in 2 ml complete medium. Replace the culture medium every 24 hr with fresh pre-warmed complete medium.
    1. To analyze the effect of proteasome inhibition on the centrosomal level of the test protein (here VDAC3 or γ-tubulin), grow the cells in two 35 mm dishes for 44 hr. Replace the culture medium with complete medium and add either MG115 or DMSO as the control solvent at a final concentration of 5 µM or 0.05% respectively. At the same time, add BrdU to the cells at a final concentration of 40 µM and incubate cells for 4 hr.
    2. Transfer each coverslip in a 24-well plate. Add 500 µl chilled methanol to each well and incubate the plate at -20 °C for 10 min to fix the cells on the coverslips. Immediately wash the coverslips three times with 500 µl wash buffer (1x PBS containing 0.5 mM MgCl2 and 0.05% Triton-X 100).
  2. Harvest the residual cells from the 35 mm dish and centrifuge the cells at 1,000 x g for 5 min. Use the cell pellet to analyze the total protein by western blotting (step 6).

3. BrdU Pulse and Chase Assay to Analyze Protein Levels in Different Cell Cycle Phases

  1. To analyze the variation of the level of a protein (here VDAC3, Sas6 or Cep135) at centrosomes at different phases of the cell cycle, grow cells in two 35 mm dishes containing coverslips for 44 hr. Replace the culture medium with complete medium containing 40 µM BrdU and incubate the cells for 4 hr in the cell culture incubator.
  2. From one dish, transfer the coverslips to a 24-well plate to fix the cells using chilled methanol as described in step 2.1.2.
  3. After the 4 hr BrdU pulse, remove the BrdU-containing medium, wash the cells once with PBS and once with complete medium, add fresh medium, and then grow the cells in the absence of BrdU for various times (typically another 4 hr for RPE1 cells) before fixing the cells as described in step 2.1.2.
    NOTE: For RPE1 cells, the majority of the BrdU-positive cells are in late S or G2-phase after a 4 hr chase. This must be determined independently for each cell type.

4. Immunostaining

  1. Incubate the fixed cells on coverslips in 200 µl blocking buffer (2% BSA and 0.1% TritonX-100 in 1x PBS) for 30 min.
    1. Incubate the cells with a mix of primary antibodies (typically one raised in rabbit and another raised in mouse) diluted in blocking buffer O/N at 4 °C in a humidified chamber.
    2. To make a humidified chamber, put a wet paper towel in the bottom half of an empty 1,000 µl pipette tip box. Lay a strip of Parafilm on the rack surface, and spot a droplet (15-20 µl) of the antibody solution onto the Parafilm for each coverslip to be incubated. Invert a coverslip onto a droplet of antibody solution (so that cells are immersed) and close the lid of the tip box.
    3. Invert coverslips again and return them back to the 24-well dish. Wash coverslips and then incubate them in 150 µl secondary antibody mixture (here green-fluorescent dye conjugated anti-rabbit and red-fluorescent dye conjugated anti-mouse diluted in blocking buffer) for 1 hr at RT. Wash the coverslips three times.
      NOTE: The fluorophore-conjugated Secondary antibodies are light sensitive. Protect the samples from light during steps 4.1.3-5.1.2.
  2. Prepare for anti-BrdU staining by fixing the stained RPE1 cells again with 500 µl chilled methanol at -20 °C for 10 min, as described in step 2.1.2. Wash the coverslips thrice.
    NOTE: This fixation secures the primary and secondary antibody labeling of the protein(s) of interest during the acid hydrolysis process in the following step.
    1. Incubate the cells in 200 µl of 2 N HCl for 30 min at RT. Neutralize with 300 µl of 1 M Tris-Cl, pH 8 and wash the cells three times using wash buffer.
    2. Block the cells again using 200 µl blocking buffer for 30 min at RT.
    3. Incubate the cells with rat anti-BrdU antibody (diluted in blocking buffer) for 45 min at 37 °C in a humidified chamber. Return the coverslips back to the 24-well dish, wash them and then incubate with blue-fluorescent dye conjugated anti-rat secondary antibody (diluted in 150 µl blocking buffer in 24-well dish for 1 hr at RT.
  3. Wash the coverslips three times. Spot a droplet (roughly 3-6 µl) of a mounting solution containing antifade reagent on a glass microscope slide. Invert a coverslip, with the cell-side facing down, onto the mounting solution. Wipe excess liquid by gently pressing the coverslip against the slide using soft cleaning tissue. Apply transparent nail polish along the edge of the coverslip to seal it onto the microscope slide.

5. Immunofluorescence Image Acquisition and Analysis

  1. Use a 100X Plan Apo oil immersion objective (with a 1.4 numerical aperture) to acquire the images of BrdU-positive RPE1 cells at ambient temperature.
    1. Put a slide on the microscope (see equipment) attached with a camera capable of digital imaging. For each fluorophore, determine the appropriate exposure time (typically between 300-1,000 msec) by manually examining all samples of an experiment. Before acquiring images of a cell, manually determine the appropriate top and bottom focal plane along the Z-axis (typically 0.2 µm step size). Acquire images of different samples that are on separate coverslips using identical exposure time for different fluorophores along the Z-axis using a digital microscopy imaging software package.
  2. Perform deconvolution (in these cases using No neighbor algorithm) of all image stacks acquired along the Z-axis.
    1. Obtain the total intensity projection of each image stack along the Z-axis.
  3. In cells where centrosomes are close (distance between two centrosomes is less than 2 µm), draw a small square (typically 20-30 pixels per side) around both centrosomes and mark the selected area.
    1. Draw a larger square (typically 24-35 pixels per side) surrounding the first square and mark the selected area of the large square.
    2. Obtain the area (A) and the total fluorescence intensity (F) of each fluorophore in each box (S denotes small box and L denotes large box).
    3. Analyze the background corrected fluorescence intensity of each fluorophore using the formula described by Howell et al.29: F= FS – [( FL– FS) x AS /( AL – AS)].
    4. Obtain the normalized fluorescence intensity of the protein of interest (here VDAC3 or Sas6) by calculating the ratio of the background-corrected fluorescence intensity of its fluorophore to that of the fluorophore used for the chosen internal standard (here γ-tubulin, Sas6 or Cep135).
  4. In the case of analyzing cells where centrosomes are well separated (distance between two centrosomes is greater than 2 µm), analyze each centrosome separately by drawing two separate sets of boxes. Draw a small square (typically 15-25 pixels per side) around each centrosome and mark the selected area. Draw a larger square (20-30 pixels per side) surrounding the first square and mark the selected area.
    1. Obtain the area (A) and the total fluorescence intensity (F) of each fluorophore in each box, and calculate the background corrected fluorescence intensities of each fluorophore, separately for each centrosome, as described in step 5.3.3.
    2. Combine the background corrected fluorescence intensities of each protein from the two centrosomes to obtain the total centrosomal level of that protein in that cell.
    3. Obtain the normalized intensity for the protein of interest by calculating the ratio of its total centrosomal intensity to the total centrosomal intensity of the internal standard.
  5. Analyze at least 15-25 cells for each experimental condition and plot the graph in a spreadsheet.

6. Analyzing the Total Protein Using Western Blotting

  1. Resuspend the cells in radioimmunoprecipitation assay (RIPA) buffer containing 50 mM Tris-HCl pH 8, 150 mM NaCl, 1% Nonidet P-40, 1% sodium deoxycholate, 0.1% sodium dodecyl sulphate (SDS) and incubate for 10 min on ice to lyse the cells.
    1. Centrifuge the mixture at 10,000 x g for 10 min, separate the lysate from the pellet fraction and measure the total protein concentration of each lysate using a bicinchoninic acid (BCA) assay kit.
  2. Mix 40 µg of lysate with 4x SDS-PAGE loading buffer (50 mM Tris-Cl, pH 6.8, 2% SDS, 10% glycerol, 100 mM Dithiothreitol, 0.1% Bromophenol blue).
    1. Boil the samples for 10 min and run the samples on a 12.5% denaturing SDS-PAGE at a constant voltage of 200 V.
  3. Transfer the proteins separated on SDS-PAGE onto a nitrocellulose membrane using western blotting technique (at a constant voltage of 90 V for 1 hr in cold).
    1. Block the membrane with 3% non-fat milk in 1x PBS, and then incubate the membrane with solutions of diluted primary antibodies (in this case rabbit anti-VDAC3, rabbit anti-γ-tubulin and mouse anti-α-tubulin) for 1 hr at RT.
    2. After washing the membrane three times (each 5 min incubation under shaking) using PBS-T buffer (1x PBS and 0.2% Tween-20), incubate the membrane in a mixture of near-Infrared (IR) fluorescent dye conjugated anti-rabbit and IR fluorescent dye conjugated anti-mouse secondary antibodies (diluted in PBS-T) for 1 hr.
    3. Wash the membrane three times and then scan the membrane to analyze the bands using an infrared fluorescence imaging system suitable for immunoblot detection.

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Results

Our recent studies identified novel centrosomal localization and function of VDAC3, one of the mitochondrial porins16,28. Immunostaining of several mammalian cells including RPE1 cells using a VDAC3-specific antibody showed prominent centrosomal staining and comparatively weak mitochondrial staining. We also showed that centrosomal VDAC3 is preferentially associated with the mother centriole, and the centrosomal pool of both the endogenous and ectopically expressed VDAC3 is regulated by degradation16

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Discussion

Quantitative microscopy in cell biology is commonly associated with live-cell imaging assays such as Fluorescence Resonance Energy Transfer (FRET), Fluorescence Recovery After Photobleaching (FRAP), etc. However, there are growing examples of cell biologists developing different quantitative microscopy assays for fixed cells in recent years27,34-36. Importantly, progress in understanding centrosome biology often requires understanding of the centrosome-specific function of proteins whose centrosomal p...

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Disclosures

The authors declare that they have no competing financial interests.

Acknowledgements

This work was supported by a National Institutes of Health grant (GM77311) and a seed grant from The Ohio Cancer Research Associates (to H.A.F.). SM was partially supported by an Up on the Roof fellowship from the Human Cancer Genetics Program of The Ohio State University Comprehensive Cancer Center.

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Materials

NameCompanyCatalog NumberComments
FibronectinSigmaF8141Stock of 1 mg/ml in water
DMSOSigmaD2650
MG115SigmaSCP0005Stock of 10 mM in DMSO
BrdUSigmaB5002Stock of 10 mM in DMSO
Anti-γ-tubulin (mouse monoclonal, clone GTU 88)SigmaT 65571:200 in IIF blocking buffer 
Anti-VDAC3 (rabbit polyclonal) Aviva Systems BiologyARP35180-P0501:50 in IIF blocking buffer, 1:1,000 in WB blocking buffer
Anti-Sas6 (mouse monoclonal) Santa cruz biotechnologysc-814311:100 in IIF blocking buffer
Anti-Cep135 (rabbit polyclonal)Abcamab-750051:500 in IIF blocking buffer
Anti-BrdU (rat monoclonal) Abcamab63261:250 in IIF blocking buffer
Alexa Fluor 350 Goat Anti-Rat IgG (H+L)Life technologiesA210931:200 in IIF blocking buffer
Alexa Fluor 488 Donkey Anti-Mouse IgG (H+L) AntibodyLife technologiesA212021:1,000 in IIF blocking buffer
Alexa Fluor 594 Donkey Anti-Mouse IgG (H+L) AntibodyLife technologiesA212031:1,000 in IIF blocking buffer
Alexa Fluor 488 Donkey Anti-Rabbit IgG (H+L) AntibodyLife technologiesA212061:1,000 in IIF blocking buffer
Alexa Fluor 594 Donkey Anti-Rabbit IgG (H+L) AntibodyLife technologiesA212071:1,000 in IIF blocking buffer
Anti-γ-tubulin (rabbit polyclonal)SigmaT51921:1,000 in WB blocking buffer
Anti-α-tubulin (mouse monoclonal, DM1A)SigmaT90261:20,000 in WB blocking buffer
Alexa Fluor 680 Donkey Anti-Rabbit IgG (H+L)Life technologiesA100431:10,000 in WB blocking buffer
Mouse IgG (H&L) Antibody IRDye800CW ConjugatedRockland antibodies610-731-0021:10,000 in WB blocking buffer
SlowFade Gold Antifade Reagent Life technologiesS36936Mounting media
Round coverslips 12CIR.-1Fisherbrand12-545-80
Olympus IX-81 microscopeOlympus
Retiga ExiFAST 1394 IR cameraQImaging 32-0082B-238
100X Plan Apo oil immersion objectiveOlympus1.4 numerical aperture
Slidebook software packageIntelligent Imaging Innovations
Odyssey IR Imaging SystemLi-cor Biosciences
Bicinchoninic acid (BCA) assayThermo Scientific23227
U-MNU2 Narrow UV cubeOlympusU-M622Filter
U-MNU2 Narrow Blue cubeOlympusU-M643Filter
U-MNU2 Narrow Green cubeOlympusU-M663Filter

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Keywords Quantitative ImmunofluorescenceProtein LevelsCentrosomesMitotic SpindlePrimary CiliaCell CycleCentriole AssemblyFluorescence MicroscopyVDAC3Proteasome mediated Degradation

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