Download Culturing and Fluorescent Staining of B16 Melanoma Cells

Culturing and Fluorescent Staining
of B16 Melanoma Cells
In this lab exercise you will learn how animal cells can be grown in culture and used to
analyze the cytosolic arrangement of actin filaments. Cell culture is a widely used methodology
for studying the behavior of cells independent of the variables existing within the whole
organisms. The technique allows investigation of intracellular processes for both normal and
abnormal cells (such as cancerous cells), while allowing analysis of cellular responses to
experimental modifications. Animal cell culture is one of the most important approaches to the
study cell structure and is widely used in applied applications such as in the production of
vaccines, antibodies and other commercially important biological products.
A primary culture is created when the cells placed into culture are taken directly from
an animal source, typically by mechanically disrupting a tissue source and allowing the cells to
disperse onto the culture medium. Although sometimes a suspension culture may be produced,
in which the cells remain free-floating in the medium, more commonly the culture will
eventually form a monolayer of cells attached to lower surface of the culture flask.
Animal cells are typically cultured using specially designed culture flasks in a solution
containing a complex mixture of nutrients and supplemented with additional hormonal
substances. For example B16 cells are cultured in CDMEM – which consists of DMEM,
Dulbecco’s Modified Eagle’s Medium (see Table 1) to which is added FBS (fetal bovine serum),
to provide essential growth hormones, and the antibiotics penicillin and streptomycin, to
discourage bacterial contamination. Fortunately many types of cell culture media, such as
DMEM, are commercially available as a premixed formulation. Every several days some of the
cells must be caused to detach and subcultured (“passaged”) to fresh medium, yielding a
secondary culture. Some animal cell lines, such as human HeLa cells and mouse B16 have
been maintained in culture for many decades.
Normal cells generally display contact inhibition when grown in culture, meaning that
when the cells have become a confluent monolayer (and in direct contact with other cells) they
stop dividing. Furthermore, normal cells (other than stem cells) will divide only for a limited
number of times before ceasing to grow. Unlike normal cells, some cultured cells are said to be
immortal, i.e., they will grow continuously in culture and often lack contact inhibition. A
transformed state is typical of many types of cancerous cells, such as HeLa and B16.
Objectives
The objectives of this lab exercise are for you to learn:
 basic principles of animal cell culture.
 aseptic culture technique to maintain a continuous cell line over several weeks.
 how fluorescent labeling can be used to study distribution of cytoskeletal filaments.
Cell Culture and Fluorescent Staining
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How are animal cells cultured?
The culture conditions we will use are typical for many different types of cultured cells.
The cells are grown in plates containing CDMEM medium. The cultures are maintained in an
incubator at 37OC supplemented with 5% CO2 in the atmosphere and containing a tray of water
to maintain humidity.
“Passaging” refers to the transfer of cells from one culture plate to another. This is
performed by causing the cells to detach from the culture plate and then transferring a sample to
a new culture well with fresh medium. The wells to be used for each round of subculturing are
shown in Figure 1. The cells are caused to detach with a solution of trypsin-EDTA. Trypsin is a
protease that gently cleaves cell surface proteins adhere the cells to the plate surface; EDTA is a
chelating compound that binds to calcium, which also promotes cell release.
One of the challenges to performing cell culture is the need to prevent bacterial or fungal
contamination of the culture medium, and you will need to take many precautions when the cells
are being passaged. We will culture B16 cells in 6 well culture plates.
Supplies needed include:
CDMEM – screw-capped bottles
B16-F10 cells (ATCC #CRL-6475)
12 well culture plate (Corning #3336)
Trypsin/EDTA– in 13 mm brown cap tube
-- thaw before using
PBS – small bottle
P-1000 pipetter & sterile tips
sterile Petri plate
100 ml beaker – for solution waste discard
Inverted microscopes
CDMEM composition
DMEM (Sigma-Aldrich, D6429)
10% FBS (PAA labs, A15-507)
1% Penicillin/Streptomycin
(Sigma-Aldrich, P4333)
PBS (Sigma-Aldrich, D8537)
2.7 mM KCl
8.1 mM Na2HPO4
1.5 mM KH2PO4 137 mM NaCl
Trypsin/EDTA (Sigma-Aldrich, T4174)
0.1% Trypsin
0.4% EDTA•4Na
0.9% NaCl
General procedures for maintaining sterile cultures
Before performing any procedure with cell culture the workspace must be carefully cleaned.
1. Wipe down your workspace with disinfectant.
2. Wipe shaft of micropipetter with disinfectant.
3. Have all supplies available on your bench before beginning.
4. Wash your hands and arms.
To avoid contamination when working with cultured cells, good aseptic technique must be
applied, which includes:
1. Do not open your culture plate more than necessary to add or remove samples.
2. Immediately recover the culture plate between procedural steps.
3. Immediately dispose of a pipet or pipet tip that touches an unintended surface.
4. Dispose of pipets or pipet tips between each operation.
5. Store lids of tubes and vials face down in the sterile Petri plate.
6. After removing the cap of a tube or vial, always hold it at an angle.
7. Never touch the lip of a tube or vial.
Cell Culture and Fluorescent Staining
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Procedures for passaging the cells
Passaging of cells is not to be performed during regular lab periods. Cells should be
transferred on an as-needed basis (approximately every 3 days) by monitoring cell density with
the inverted microscope. Cells should be transferred when they reach approximately “80%
confluence”.
*** Keep a full record all of your culturing activities in your lab notebook. ***
1. Trypsinizing to release cells from culture well
1. Prep the work space as described above; be sure to wash your hands and arms. Remove
CDMEM from refrigerator, and thaw Trypsin/EDTA.
2. Remove the media from the well with the cells.
3. Add 1 ml of PBS, and gently swirl. Remove and discard the PBS.
4. Repeat step 3 again.
5. Add 0.25 ml of trypsin/EDTA solution, and incubate the plate at 37OC for 5 minutes.
Return the trypsin/EDTA to freezer.
6. Swirl the plate and observe that the cells with the inverted microscope. If the cells have not
detached, return the plate to the 37OC incubator for another 5 minutes.
7. When the cells have been observed to detach, add 0.5 ml of CDMEM to quench the
trypsin.
8. Slowly and gently suck the cell suspension into and out of the pipet tip 2 -3 times to fully
separate and suspend the cells
9. Transfer to a sterile 13 mm test tube.
2. Doing a cell count
Determine the number of cells in the suspension as described on the next page. Calculate the
volume needed to passage approximately 5000 cells to the next well.
3. Starting the next cell culture
Figure 1. Passaging cells to new culture wells.
1. Making sure the cells are well
suspended, and avoiding any cell
clumps, transfer the appropriate volume
to the next culture well, as indicated in
Figure 1. Draw cells gently into the
pipet tip, and discharge them gently.
2. Add 1 ml of CDMEM to the well to
receive the passaged cells, and swirl to
distribute the cells.
3. Remove and discard remaining media
from the old culture well.
4. Return plate to the CO2 incubator.
Return unused PBS and CDMEM to
refrigerator.
 The day before the actin staining lab exercise, transfer your cell culture to the two
wells containing cover slides. The reason for this is described in the section of fluorescence
staining. 
Cell Culture and Fluorescent Staining
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How are cells counted?
In order to maintain a cell culture under steady
conditions, a consistent number of cells should be
transferred with each passaging cycle. This is
accomplished by determining the cell concentration in the
suspension released from the plate, and then transferring a
specific number of cells to the next culture. Counting is
done using a hemocytometer type counting chamber
(Figure 2).
As demonstrated in lab, a small volume of the
trypsinized cell suspension is introduced in the loading
ports on either side of counting chamber. On each side of
the hemocytometer the cell suspension will fill a space
under the cover slide and over a grid.
Figure 2. Hemocytometer
Hemocytometer grids have several grid areas, designed for counting cells of different sizes and
densities (Figure 3). We will be counting cells in the 5 outer + center area grid. The volume
contained in each of these areas is:
1mm x 1 mm x 0.1 mm = 0.1 mm3
The total volume of the 5 grids is:
0.1 mm3 x 5 = 0.5 mm3
Since 1 ml volume = 1000 mm3, the volume counted =
1/2000 of 1 ml (1000 mm3  0.5 mm3 = 2000)
Thus, cells / ml = cell count x 2000
It is critical to be systematic when counting cells. Follow a
standard pattern as you survey the grid area. By
convention, cells that within the grid area or on a left or top
side boundary line are counted; As shown in the figure,
cells touching a bottom or right side line are excluded.
Example:
5 cell counts: 10 + 5 + 8 + 12 + 3 = 38 total
Cell density = 38 x 2000 = 1.14 x 105 cells/ml
If 5000 cells are to be passaged to a new culture well:
5000 cells / 1.14 x 105 cells/ml = 0.088 ml (88 μl)
88 μl of the trypsinized cells would be passaged to the
next well.
Cell Culture and Fluorescent Staining
Figure 3. Cell counting
with hemocytometer
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How are actin filaments fluorescently stained?
Actin filaments can be stained using phalloidin bound to a
fluorescent tag. Phalloidin is a toxin produced by the death cap
mushroom, Amanita phalloides, that binds to f-actin (actin
associated into filaments) and prevents their depolymerization.
Phalloidin can be covalently bound to a fluorescent molecule such
as rhodamine (Figure 4), and used to probe for actin filaments
within cells. Before doing so, the cells must be “fixed”, which
means that the cells are treated with chemicals that stabilize their
cellular components (which also kills the cells). The cell membrane
is then “permealized” with a mild detergent (Triton X-100) to allow
entry of the staining reagents into the cell.
Figure 4. Structure of
The cells will be mounted in a medium that reduces
rhodamine-phalloidin.
photobleaching (fading) of the fluorescence and which
contains a fluorescent molecule called DAPI (Figure 5), which
binds DNA to stain the nuclei.
In order to stain cultured cells and view them using a
compound microscope, they must be grown on a surface other
Figure 5. Structure of DAPI.
than culture flasks. Thus, for the final passaging, the cells will
be transferred to sterile coverslips, placed in two of the culture
wells of your plate.
 Transfer your cell culture to the two wells containing cover slides
the day before the actin filament staining lab. 
What are the principles of epifluorescence microscopy?
Epifluorescence microscopy is the most widely used form of microscopy used in for
biological research. Typically, the specimen being viewed has been treated with a fluorescent
stain (fluorochrome) that attaches to a particular protein or other cellular structure. In our
experiment the fluorochromes are rhodamine attached to
phalloidin, which binds to f-actin, and DAPI, which
binds to DNA.
A fluorochrome absorbs light and transmits light
at a longer wavelength. Figure 4 shows the path of light
in an epifluorescence microscope. A very high intensity
light source is needed, such as a xenon arc lamp or
mercury-vapor lamp. An excitation filter restricts the
wavelengths of light to only those absorbed by the
fluorochrome. The excitation light passes through the
objective lens and then onto the specimen A special
dichroic mirror, set at a 45O angle, directs the
Figure 4. Light path in a
excitation light to the specimen. The light released
fluorescence microscope. Adapted
from the specimen, the emitted light, passes back
from http://www.ncbi.nlm.nih.
through the objective lens. However, the fluorescent
gov/bookshelf/br.fcgi?book=mcb&part=
light emitted by the fluorochrome passes through the
dichroic mirror to the ocular lens. Although most of the excitatory light passes through the
Cell Culture and Fluorescent Staining
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specimen unabsorbed, an emission (barrier) filter also is present to block transmission of
unwanted excitation light.
For each type of fluorochrome, a
particular set of filters and dichroic mirror must
be selected to accommodate its particular light
absorbance and emission characteristics. The
absorbance and emission spectra for rhodamine
are shown in Figure 5. Notice that the excitation
filter only transmits wavelengths that rhodamine
absorbs, and that the barrier filter only transmits
of wavelengths that rhodamine emits. The
dichroic mirror only reflects wavelengths below
about 570 nm.
You can visit the Nikon website for further
discussion of fluorescence microscopy at:
http://www.microscopyu.com/articles/
fluorescence/fluorescenceintro.html
Figure 5. Characteristics of rhodamine
fluorescence.
Rhodamine-Phalloidin Staining Procedure
Supplies needed include:
a-d are all components of the ‘F-actin visualization kit’ (Cytoskeleton, #BK005)
a) Rhodamine-phalloidin (red cap vial)
P-200 pipetters
b) Wash buffer (blue cap test tubes)
black plastic box
c) Fixative (green cap vial)
clear nail polish
d) Permealization buffer (yellow cap vial) mounting medium
100 ml solution discard beaker
tissue cassette
Mounting medium with DAPI
You must wear gloves as you perform this procedure.
Phalloidin is very toxic through skin exposure and inhalation.
The fixative contains 10% formaldehyde, a suspected carcinogen.
It is critical that you know and maintain the orientation of the cover slide during this procedure.
Each student should process a separate coverslip.
1. Place a piece of damp filter paper in the plastic box. This creates a humid environment.
2. Carefully remove the coverslips from culture wells using tweezers, and place them cell
side up in a tissue cassette -- the cassette will make it easier to work with the cover slide.
Blot from the bottom or side any medium that flows off the cover slide, and then place
the cassettes in the plastic box.
Cell Culture and Fluorescent Staining
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Perform the following procedures for each of the coverslips.
3. Cover the coverslip with 200 µl of Wash Buffer for 30 seconds.
4. Remove the Wash Buffer by holding a paper towel under the cassette and tipping it until
the fluid flows off the coverslip; you can then also touch the edge of the coverslip with
the paper towel to remove any remaining liquid. Do not blot the surface of the
coverslip as this will damage the cells.
5. Add 200 µl of Fixative solution to the coverslip and incubate for 10 minutes, and then
remove the Fixative with dry filter paper as in step 4.
6. Wash the coverslip again with 200 µl of Wash Buffer for 30 seconds, and then remove
the Wash Buffer as in step 4.
7. Permealize the cells by adding 200 µl of Permeabilization Buffer for 5 minutes, and then
remove the Permabilization Buffer as in step 4.
8. Wash the coverslip again with 200 µl of Wash Buffer for 30 seconds, and then remove
the Wash Buffer as in step 4.
9. Add 200 µl of the Rhodamine Phalloidin solution and incubate for 15 minutes. This step
is carried out in the dark so close the Dark Box lid. Remove the Rhodamine Phalloidin as
in step 4.
10. Wash three times with 200 µl of Wash Buffer for 30 seconds each, removing the Wash
Buffer each time as in step 4. Blot the edge of the coverslip well after the third wash.
Mounting onto a microscope slide
11. Add one drop of mounting medium onto a microscope slid. Carefully pickup the
coverslip with forceps and carefully invert onto a microscope slide.
12. Seal the edges of the coverslip with clear nail polish and allow to dry for 5-10 minutes in
the dark. The slide should be stored at 4ºC in the dark until it is viewed.
Table 1. Dulbecco’s Modified Eagle’s Medium (DMEM)
Inorganic Salts
CaCl2 (anhydrous)
Fe(NO3)3·9H2O
MgSO4 (anhydrous)
KCl
NaHCO3
NaCl
NaH2PO4·H2O
(g/liter)
0.20000
0.00010
0.09770
0.40000
1.50000
6.40000
0.12500
Vitamins
Choline Chloride
Folic Acid
myo-Inositol
Nicotinamide
D-Pantothenic Acid
Pyridoxine·HCl
Riboflavin
Thiamine·HCl
(g/liter)
0.00400
0.00400
0.00720
0.00400
0.00400
0.00400
0.00040
0.00400
Amino Acids
L-Arginine·HCl
L-Glutamine
L-Histidine·HCl·H2O
L-Leucine
L-Methionine
L-Serine
L-Tryptophan
L-Valine
(g/liter)
0.08400
0.58400
0.04200
0.10500
0.03000
0.04200
0.01600
0.09400
Amino Acids
L-Cystine·2HCl
Glycine
L-Isoleucine
L-Lysine·HCl
l-Phenylalanine
L-Threonine
L-Tyrosine·2Na·2H2O
(g/liter)
0.06260
0.03000
0.10500
0.14600
0.06600
0.09500
0.10379
Cell Culture and Fluorescent Staining
Other
(g/liter)
D-Glucose
4.50000
Phenol Red
0.01500
Sodium Pyruvate 0.11000
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Cell Culture and Fluorescent Staining
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