How to prepare samples for the Cell Cycle Assay on NC-3000

Background

The cell cycle represents the most fundamental and important process in eukaryotic cells. Being an ordered set of events, culminating in cell growth and division into two daughter cells, the cell cycle is tightly regulated by defined temporal and spatial expression, localization and destruction of several cell cycle regulators. Cyclins and cyclin dependent kinases (CDK) are major control switches for the cell cycle, causing the cell to move from G1 to S or from G2 to M phases. 

In a given population, cells will be distributed among three major phases of cell cycle: G1 /G0 phase (one set of paired chromosomes per cell), S phase (DNA synthesis with variable amount of DNA), and G2/M phase (two sets of paired chromosomes per cell, prior to cell division). 

DNA content can be measured using fluorescent, DNA-selective stains that exhibit emission signals proportional to DNA mass. This analysis is typically performed on permeabilized or fixed cells using a cell-impermeant nucleic acid stain but is also possible using live cells and a cell-permeant nucleic acid stain. Because cell cycle dysregulation is such a common occurrence in neoplasia, the opportunity to discover new targets for anticancer agents and improved therapeutics has been the focus of intense interest.

The cell cycle assay has applicability to a variety of areas of life science research and drug development, including cancer biology, apoptosis analysis, drug screening and measuring health status of cell cultures, e.g. in bioreactors.

How to prepare samples for the Cell Cycle Assay on NC-3000

In most cases, cells must be fixed or permeabilized to allow entry of the dye which is otherwise actively pumped out by living cells.  For fixation, alcohol or aldehyde are commonly used. Alcohol is a dehydrating fixative which also permeabilizes. This will allow easy access of the dye to the DNA and gives good profiles (low coefficient of variation, CV). The disadvantage is that it is often incompatible with fluorescent proteins and some surface markers.

Note: It is recommended that cells in each sample be synchronized to the same cell cycle phase; this can be achieved by culturing the cells in serum -free or serum-depleted media for 24-48 hours prior to treatment, depending on cell type.

Protocol for two-step Cell Cycle Analysis

Protocol for cells in suspension:

For proper staining it is crucial to keep the cell density within the range of 1×106 to 2×106 cells/mL. In case of limited amounts of cells the procedure can be scaled down, e.g for 2×105 to 4×105 cells use 100 μL of Solution 10 and Solution 11 in steps 1 and 2.

  1. Harvest 5×105 to 1×106 cells by centrifuging 5 minutes at 400 g at room temperature. Wash once with PBS, remove PBS completely and thoroughly resuspend cells in 250 μL Solution 10 supplemented with 10 μg/mL DAPI.
  2. Incubate cells at 37° C for 5 minutes, add 250 μL Solution 11. Cells are now ready for acquisition.

Protocol for cells attached to T-fasks, cell culture plates or micro-carriers:

The number of seeded cells needs to be optimized for each cell type to have a sufficient number of cells for the analysis and at the same time avoiding the cells to arrest in G1 due to high confluences of the cells. A starting point could be to seed out a total of 2.5×105 to 5×105 cells/well in a 6-well culture plate the day before the experiment.

For 6-well culture plates (for other culture plates, see table 1):

  1. Remove culture medium, wash once with 3 ml of PBS, remove PBS completely and add 250 μL of Solution 10 supplemented with 10 μg/mL DAPI.
  2. Incubate cells at 37° C for 5 minutes, resuspend cells thoroughly by pipetting, add 250 μL Solution 11.
  3. Sample is ready to run.

Protocol for fixed-cell Cell Cycle Analysis

For proper staining it is crucial to keep the cell density within the range of 2×106 to 4×106 cells/ml. In case of limited amounts of cells, the procedure can be scaled down, e.g use 2×105 to 4×10cells in 100 mL PBS in step 1. 

  1. Collect cells for fixation  a. For cells growing in suspension or hematologic samples. Harvest cells by centrifuging 5 minutes at 500 at room temperature. Wash once with PBS. Count cells (e.g. by using a Via1-Cassette™) and thoroughly resuspend 1×106 to 2×106 cells in 500 mL PBS.  b. For adherent cells. Harvest cells by trypsinization and pool the trypsinized cells with cells floating in the medium (latter consist of detached mitotic, apoptotic and dead cells). Centrifuge cells for 5 minutes at 500 at room temperature. Wash once with PBS. Count cells (e.g. by using a Via1-Cassette™) and thoroughly resuspend 1×106 to 2×106 cells in 500 mL PBS. 
  2. Add 4.5 mL of 70% ethanol to each of an appropriate number of 10-15 mL centrifuge tubes. Keep on ice.
  3. Transfer the cells suspensions (prepared in step 1) into the appropriate tubes containing ice-cold 70% ethanol, vortex rigorously, and keep the cells in the fixative for at least 2 hours.  a. Important: it is essential to have a single-cell suspension at the time that cells are mixed with ethanol.  b. Cells can be stored in 70% ethanol for several months at 0-4⁰ C. 
  4. Centrifuge ethanol-suspended cells for 5 minutes at 500 g. Decant ethanol thoroughly.  a. Note: Cell pellet may be loose. Make sure that no cells are lost in this and subsequent washing steps. b. Suspend cell pellet in 5 mL PBS, leave for 50 sec, and centrifuge 5 minutes at 500 g.
  5. Resuspend cell pellet in 500 mL Solution 3 and incubate for 5 minutes at 37⁰ C. 
  6. Sample is ready to run.
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