Video: Workflow for subculture of adherent cells
Once a cell culture has been started, it cannot be grown indefinitely due to the increase in cell number, consumption of nutrients and increase in toxic metabolites which eventually results in cell death. Moreover researchers usually want to perform experiments on their cells several times, and therefore do not want to use up all of the cells at once. Subculturing, or splitting the cells, produces new cultures with lower cell density than the original culture. By removing the medium and transferring the cells into fresh growth medium, the cells are given fresh nutrients and toxic metabolites are removed, allowing long-term maintenance of the culture.
After initially seeding the cells, growth starts with a lag phase and proceeds to a log phase, where the cells proliferate exponentially followed by a stationary phase where growth rate and death rate are equal (Fig. 1). In the death phase, cells die due to lack of nutrients and inadequate living conditions.
In order to keep cells healthy and actively growing it is necessary to renew the growth medium and to subculture them at regular intervals. Change of culture medium can take place several times in the log phase dependent on the cell type. The best time to subculture cells is between the log phase and the stationary phase, before the cells reach confluence.
It is important to examine the cell culture every day and immediately prior to subculturing to monitor cell health, check for contamination and determine when to split the cells.
A first examination of the culture for fungal contamination, turbidity and particles in the medium as well as unexpected pH shifts, indicated by color change of the medium, can be done at the macroscopic level, by eye. After this, a closer check of the general cellular morphology and growth patterns should be examined using an inverted microscope. The optics of an inverted microscope are located below the specimen. Since the cells are attached to the bottom of the dish, they can be viewed easily from this perspective. Observation should take place with a total magnification of 100 – 200x and with phase contrast, because most cells are difficult to observe in normal bright field illumination.
There are many variations in mammalian cell morphology, but most mammalian cells in culture can be divided into three categories: fibroblastic cells (Chinese Hamster Ovary cells (CHO)), epithelial-cell-like (human cervix cells (HeLa)) and lymphoblast-like cells (human leukaemia cells (HL60)). In addition to this certain cell lines can have specific morphological characteristics, e.g. neurons (SH-SY5Y) which have very long dendritic processes. Cell morphology is also affected by events in the cell lifecycle. During mitosis many cells round up, forming very refractile bright spheres that may float around in the medium. Dead cells often round up and become detached also but are usually not bright and refractile.
Various cell lines not only differ in size and shape, they also differ in their growth behaviour. They either growing adherent (fibroblastic and epithelial cells) or in suspension (lymphoblast-like cells). Most adherent cell lines grow as a single cell layer (monolayer) attached to glass or treated plastic substrates (coated with poly-lysine, fibronectin, collagen or gelatine).
The most common method to prepare cells for subculture is by breaking the intercellular and cell-to-substrate connections with proteolytic enzymes like trypsin. Trypsin in combination with Ethylenediaminetetraacetic acid (EDTA) causes cells to detach from the growth surface. Trypsin cuts away the focal adhesions that anchor the cell to the culture dish and EDTA acts as a calcium chelator.
By removing calcium, cadherins which are involved in cell-cell interactions, are broken and cells separate from one another. Once separated from the growth surface and the surrounding cells, they can be easily separated and grown in new cell culture dishes.
Cell culture conditions and subculture methods vary for each cell type. Figure 2 describes the basic steps in the subculture workflow. During the whole subculture process it is important to work in a contamination-free environment. Examination of the cells at the beginning, during trypsination, cell counting, and after splitting is essential. For consistent results, maintaining good records and documentation is also important.
The following protocol describes the basic principles of the subculture routine for Madin Darby Canine Kidney Cells (MDCK cells) grown in a 90 mm Petri dish. These are epithelial cells isolated from the distal tubules of a dog. In culture, they grow adherently and form a monolayer of polygonal cells after they have reached confluence.
The following material and equipment is needed for subculture:
- Pre-warmed medium to 37° C (for MDCK cells: MEM with 5% FCS, 2 mM Glutamine, 100 U/mL Penicillin, 100 mg/mL Streptomycin)
- Pre-warmed PBS without Ca2+ /Mg2+
- Pre-warmed 0.05% Trypsin, 0.02% EDTA in D-PBS
- Trypan Blue (vital stain)
- 70% ethanol or isopropanol for disinfection
- Cell counting chamber
- Pipette/ micropipette with disposable tips
- Phase contrast capable inverted microscope (Leica DMi1, for example)
- Personal protective equipment
- Water bath set to appropriate temperature
- Incubator at 37°C with 5% CO2 and high humidity
- Pre-labelled dishes
The splitting ratio is 1:10.
Step 1: Cell examination
Take cells out of the incubator and put them under the microscope for quick cell check. Cells should be checked microscopically every day to ensure they are healthy (no contamination, few dead cells) and growing as expected.
Cells should be mainly attached to the bottom of the dish or flask and the media should be pink-orange in color. The pH indicator phenol red turns yellow upon acidification as a result of metabolism products from the cells (or contaminations!) in the media. MDCK cells are polygonal in shape when in the log phase and grow as a monolayer. They are difficult to see with normal brightfield illumination. By switching to phase contrast the cells can be identified more easily. On the Leica DMi1 this can be done by simply moving the slider housing the condenser annulus, as shown.
Documentation of the cell status is very important to ensure uniform results across experiments. The Leica DMi1 can be equipped with a camera and screen which allows easy imaging and saving via remote control.
Comparison of brightfield and phase contrast images of MDCK cells. The phase contrast image gives a better overview and makes inspection of the cell morphology and cell counting easier.
Step 2: Cell harvesting
Pipet off the medium from cells into a waste container.
Carefully wash cells up to three times with 5 ml pre-warmed PBS without calcium and magnesium to get rid of the fetal bovine serum (FBS) in residual culture media. FBS will inhibit the trypsin. Add 3 ml pre-warmed Trypsin/EDTA and swirl gently to cover all cells at the bottom of the dish. Incubate cells for a few minutes at 37°C to detach them.
Different cell lines require different trypsin incubation times. To avoid over-trypsination which can damage the cells, check them every few minutes under the microscope.
Gently rinse plate and transfer cell suspension into a 50 ml tube and spin them down for 5 min at 800 rpm. Aspirate the supernatant and resuspend the cells in 10 ml fresh medium to fully remove the trypsin.
Detached cells should be round shaped and free floating in the trypsin solution. As soon as cells have detached, add 5 ml culture medium to the dish to inactivate the trypsin.
Step 3: Cell counting
Mix 100 µL cell suspension with an equal amount of 0.4% Trypan Blue solution. Trypan Blue selectively penetrates cell membranes of dead cells and stains them blue, but is not absorbed by living cells. Prepare the haemocytometer by placing the coverslip over the counting surface.
Load the counting chamber with the cell suspension (~ 4 µl per counting area) by placing the pipette tip at the edge of the cover slip and gently expel the cell suspension. The area under the cover slip fills by capillary action. In most cases the chamber has two counting areas that can be loaded independently.
Place the chamber on the microscope stage and focus the cells. The square pattern of the counting grid differs depending on the chamber type. The Fuchs-Rosenthal counting chamber you can see here has a pattern of 16 areas of one square millimeter each bordered by triple lines. Each square is subdivided into 16 smaller squares. Count all the cells in one 16 square region, as shown in the figure. To avoid counting cells twice at the edges of the area, count only those cells on the lines of two sides of a square. In this example, the cells touching the upper and left limits should be counted (thicker red lines). Cells touching the lower and right limits should be not taken into account. Count the live and dead cells in 5 one square millimeter rectangles of the counting chamber. For calculation, you will need to combine the counting results from all 5 squares. For greater accuracy in the measurement, additional squares of the counting chamber can also be counted.
Step 4: Plating
Pipette the required volume of cells (appropriate number of cells) into new dishes at the required split ratio (here 1:10) and top up with culture medium to the required final volume in each dish (10 ml). Note cell type, day of cell splitting and passage number on the lid of the dish. Place cells back in the incubator at 37° C.
Leave cells overnight to recover and settle. Check cells 24 hours later under the microscope for shape, adhesion and contamination.
Cells should be attached to the bottom of the dish and starting to grow and divide. Change the media to get rid of any residual trypsin. Grow cells until they are confluent and ready for your experiment or next subculture.