Authors: Greenfield Sluder, Joshua J. Nordberg, Frederick J. Miller and Edward H. Hinchcliffe
This protocol was adapted from “A Sealed Preparation for Long-Term Observations of Cultured Cells,” Chapter 18, in Live Cell Imaging (eds. Goldman and Spector). Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY, USA, 2005.
The continuous long-term observation of cultured cells on the microscope has always been a technically demanding undertaking. This protocol describes a sealed preparation that allows the continuous long-term observation of cultured mammalian cells on upright or inverted microscopes without environmental CO2 control. The preparation allows for optical conditions consistent with high-quality imaging and good cell viability for at least 100 hours. The preparation is an aluminum support slide with a square aperture cut in its center. The coverslip bearing the cells is attached to the top of the slide with a thin layer of silicone grease, and the bottom of the slide is similarly covered with a clean coverslip of the same size. The thickness of the slide is intended to coordinately maximize the volume of the medium while maintaining optical properties that allow Koehler illumination with standard condensers. The chamber is filled in equal parts with HEPES-buffered media containing fetal calf serum and a low-viscosity fluorocarbon oil. These oils have a high solubility for atmospheric gases. The inclusion of the oil in the preparations is intended to provide a source of oxygen and perhaps a sink for some of the CO2 produced by the cells. Although the inclusion of fluorocarbon oil in the preparation may not be necessary for short-term (~24 hr) observations, particularly with cells that are sparsely plated, long-term cell viability is ensured when the oil is present.
Support slides should be prepared as described in A Sealed Preparation for Long-Term Observations of Cultured Cells: Details of Support Slide Construction.
- Cells for analysis
- This protocol has been used extensively for BSC-1, CV-1, COS-7, CHO, mouse embryo fibroblasts, and hTERT RPE1 cells. The FC47, FC40, FC43, and FC77 oils (see below) all work well for these cells. Most cells appear viable and have normal interphase morphology at 250 hours. Normally, observations of cells stop after 70-120 hours because by that time the cells have become confluent. Over the course of 100 hours, all of the cell types show constant motility, and mitoses continue for the duration of the observations with no noticeable prolongation of the cell cycle at later times. Note, however, that at later times, cells often develop small spherical inclusions that are phase-bright. The identity of these inclusions remains obscure, but they may be large pinocytotic vesicles. In any case, they do not appear to have an adverse impact on cell motility, mitosis, or gross morphology.
- Culture medium
- This protocol uses culture medium appropriate for each cell type supplemented with 12.5 mM HEPES, 10% fetal calf serum, and a 1:100 dilution of the antibiotic/antifungal reagent sold by GIBCO (catalog no. 15240-062).
- Ethanol (70%, 95%)
- Fluorocarbon oil
- The oils we use are manufactured by 3M Corp. and are part of its Fluorinert series of performance liquids (3M Fluorinert product guide, 1997). These are short (primarily eight-carbon) hydrocarbons that are fully substituted with fluorine. They are extremely inert and immiscible with water, and they do not present any recognized significant health hazards. However, it is recommended that all users consult the Material Safety Data Sheets provided by 3M Corp. for potential health hazards before using these oils. These oils are available from three manufacturers’ representatives in three-quarter gallon (or 11 pound) amounts. 3M Corp. does not directly sell anything but truckload quantities of these oils. Small sample quantities are available from Acuity Technical Sales (New Hampshire), AMS Materials (Florida), and Semitorr (Oregon).
- Vaseline:lanolin:paraffin, melted and combined 1:1:1 (VALAP)
- Alcohol burner
- Aluminum support slide
- Beaker, 1 liter
- Controlled-temperature environment
- Since this preparation is sealed, control of environmental CO2 is not needed. However, the cells under observation need to be maintained at 37°C. This can be achieved by enclosing the entire microscope in a box and using a proportional feedback control apparatus to blow warmed air into the enclosure. Cardboard boxes work well when configured so that the video camera and the mercury arc lamp (when present) are external to the enclosure. The oculars should project from the top edge of the box. A more elegant and user-friendly setup is a Plexiglas enclosure with sliding doors custom-fabricated for the particular microscope. Again, the oculars, camera, and arc lamp housing should be located outside of the box. Three alternative heating strategies include placing the microscope in a 37°C room, enclosing the volume around the stage with a custom-built Plexiglas box, and warming the preparation alone with a temperature-controlled support apparatus on the stage. This last strategy suffers because there will be a temperature gradient from the margins of the preparation to the cells under observation. The gradient is particularly severe when a water or oil immersion objective is used, unless the objective is equipped with a heated collar or other heating device.
- Coverslip jars
- Coverslips, size 1.5
- Culture dish, 100 × 20 mm
- Pipettes, 1 ml
- Silicone vacuum grease (High Vacuum Grease; Dow Corning)
- Squirt bottle containing distilled H2O
- Tissue culture hood
- UV light source
- Water bath, preset to 37°C
- Clean the coverslips used in the construction of this preparation prior to use by sonicating them in distilled H2O containing a small amount of detergent and then rinsing the coverslips several times in distilled H2O. Store cleaned coverslips in jars containing 95% ethanol.
- Although the utility of these preparations is not sensitive to coverslip thickness, use number 1.5 (i.e., 0.17 mm thick) coverslips, because microscope objectives (without coverslip correction collars) are designed for this thickness of glass. Use of number 1 or number 2 coverslips introduces spherical aberration that degrades image quality.
- To prepare the coverslips for use in growing cells, pass each coverslip through the flame of an alcohol burner to burn off excess ethanol.
- To ensure a uniform and lasting seal, do not apply silicone grease to the margins of wet coverslips coming out of a tissue culture dish. Instead, grease the margins of the coverslips before the cells are grown on them by applying a thin and uniform coating of silicone vacuum grease to the margins of the coverslip with a small spatula.
- Place the coverslips in a 100 × 20-mm tissue culture dish. To ensure sterility, place the dish with coverslips in a tissue culture hood, and expose it to UV light for 10 minutes.
- Following sterilization, plate the cells onto coverslips and culture them in ~10 ml of media.
- For our applications, we use the media appropriate for each cell type supplemented with 12.5 mM HEPES, 10% fetal calf serum, and a 1:100 dilution of the antibiotic/antifungal reagent sold by GIBCO.
- Wipe the aluminum support slide with a tissue soaked with 70% ethanol, and then briefly pass the slide through a flame to remove residual alcohol.
- Use a small spatula to apply a thin rim of silicone grease around the top and bottom margins of the opening.
- Use a thin, even layer of silicone grease. Thicker layers, although they may work well, can lead to dimensional instability as the preparation “settles” at 37°C. This can lead to a constantly changing focus for the first few hours, even though the microscope may have a stable focus mechanism.
- Flame a cleaned blank coverslip (from Step 1) to remove the alcohol, and attach the coverslip to the bottom of the support slide. Use the back of a pair of curved forceps to gently tamp the coverslip to ensure a good seal.
- Place the prepared slide in a plastic culture dish, and expose it to UV light for 10 minutes in a tissue culture hood.
- Warm the media and fluorocarbon oil to 37°C in a water bath.
- Use a sterile 1-ml pipette to fill the chamber halfway with oil (~350 μl).
- Use a fresh sterile 1-ml pipette to add medium until the oil at the margins of the opening just barely overflows from the chamber (again, ~350 μl).
- Remove a coverslip with cells from the culture dish, and aspirate off excess medium. Quickly place the coverslip, silicone-grease-side down, on the chamber. Tamp the coverslip with forceps to ensure a good seal. Aspirate off any excess media and oil that may have flowed over onto the top of the coverslip.
- Wash the top of the preparation prior to use to prevent salts from the medium forming crystals on the coverslips once they air dry.
- Fill a 1-liter beaker with 37°C water, and place a squirt bottle of distilled H2O in it to warm.
- When the water is warm, take the assembled chamber out of the incubator. Use a small spatula to apply a small amount of melted 1:1:1 vaseline:lanolin:paraffin (VALAP) to the edges of the top coverslip to provide an extra seal. Use the squirt bottle to gently wash the top coverslip, and aspirate off any excess water.
- Keep the amount of VALAP used to a minimum to reduce the chance that any will catch on the objective when the preparation is in use. This soft, waxy material is difficult to remove from optical surfaces.
- Proceed with microscopy--the culture preparation is now ready for observation.
See Movie 1, which illustrates BSC-1 (monkey kidney epithelial) cells imaged by time-lapse video microscopy.
Movie 1. BSC-1 (monkey kidney epithelial) cells imaged by time-lapse video microscopy. The sequence shows an individual cell and its subsequent daughter cells undergoing three rounds of cell division (mitosis). The cells are imaged by phase contrast microscopy. Frames are captured every 3 minutes using a CCD camera coupled to a personal computer.
- 3M Corp. (1997) Fluorinert liquids, product and contact guide (Engineering Fluids and Systems, 3M Specialty Chemicals Division, St. Paul, MN.)