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Genetics
Mitosis Lab
INTRODUCTION
When you look at cells using regular brightfield microscopy, details appear as
differences in light intensity (light versus dark regions) or as different colors.
Because most unstained cells have little contrast, they appear transparent. Staining
enhances details by adding color to the cell. However, most dyes are toxic to cells
limiting their use to fixed (dead) cells.
Another technique used to increase contrast is fluorescence microscopy. With
regular light microscopy, when you look at a sample, in addition to the sample, you
are also observing the background illuminating light. This light can make it difficult
to see weakly stained cellular components. With fluorescence microscopy, the
background light is eliminated so you see the fluorescently stained object
contrasted against a black background. It is like looking at a dim light outside on a
sunny day versus in a dark room. Fluorescent molecules, also called fluorophores
or fluorochromes, absorb light of one particular wavelength (the excitation
wavelength) and then re-emit light at a longer, lower energy wavelength (the
emission wavelength). This process is known as fluorescence. Fluorochromes are
characterized by their absorption and emission spectra. The most intense
fluorescence (emission) occurs when the molecule is irradiated with wavelengths
close to the peak of the absorption (excitation) curve.
Absorption and emission spectra for one compound, fluorescein isothiocyanate
(FITC), are shown in the figure below. The excitation curve shows that FITC
absorbs maximally near 488 nm, in the blue light range of the visible light
spectrum. The emission spectrum shows that when FITC is irradiated at 488 nm, it
fluoresces maximally at 520 nm, emitting a greenish-yellow glow.
Other fluorochromes that you will use, such as tetramethylrhodamine
isothiocyanate (TRITC), are excited by light in the yellow-green range and emit in
the red wavelengths.
Because the wavelength of the emitted light is different than that of the absorbed
light, it is possible to filter out the excitation light so that the viewer only sees the
emitted fluorescent light. The fluorescence microscope has been designed to
separate these two wavelengths. As a result, the excitation illumination is invisible
to the viewer, who sees only glowing fluorescent structures against a black
background. Note that although the resulting image is strikingly intense, this is by
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contrast to the black background. The actual amount of fluorescence emitted is
very small. This signal is easily swamped, so conventional microscope illumination
must be switched off and stray light reduced to a minimum by working in a
darkened room.
In a fluorescence microscope, the exciting beam does not shine up through the
specimen as in conventional microscopy but instead impinges down on it ("epi" or
incident illumination). The excitation light (L) first passes through a filter (labeled
E) to select light of the optimum exciting wavelength. This light is reflected down
onto the sample. Fluorochromes in the sample emit fluorescent light, which is then
filtered again (labeled F) before observation. The heart of the incident fluorescence
microscope is the chromatic beam splitter (CBS). This is a dichroic mirror that
reflects light shorter than a certain wavelength and transmits light of longer
wavelengths. Thus, the CBS reflects the shorter wavelengths of the exciting light
onto the specimen and transmits the emitted fluorescence towards the eyepieces.
Different fluorochromes have their own absorption and emission maxima. This
requires filters specific to each fluorochrome. The microscope you will be using
today can accommodate up to four different filter cubes that are selected by sliding
the filter selector. The filter cubes are labeled: UV-2E/C DAPI for excitation in the
Ultraviolet (330-385 nm) range which is useful for DAPI; B-2E/C FITC for
excitation in the Blue (460-490 nm) range which is useful for fluorochromes like
FITC; and G-2E/C TRITC for excitation in the Green (520-550 nm) range which is
useful for TRITC.
The intense light beams used to excite fluorescence generate oxygen radicals that
alter the resonance properties of a molecule. As a result, fluorescence is destroyed
and the image fades irreversibly. Some fluorophores fade very quickly. We use an
"antifade" compound (Prolong mounting medium) that scavenges oxygen radicals
and protects the fluorescence signal.
To minimize bleaching you should pay attention to the following guidelines,
especially the first one:
 High energy UV light (used with the DAPI stain and filter) causes the
most bleaching. Keep its use to a minimum. Slide in the shutter when
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not looking down the microscope, even for a moment (but DO NOT
switch off the power supply!).
 Reduce the area of slide illuminated by the beam to the smallest possible, so
that only a small patch of cells gets bleached at a time, i.e. do not look at the
samples very long when using lower power objectives (e.g. the 20X
objective).
FLUORESCENCE MICROSCOPY STAINING METHODS
Because relatively few biological molecules fluoresce in the visible range, various
fluorescence staining methods have been developed. Three basic staining methods
are typically used.
(1) Direct stains
Some fluorescent chemicals have a high affinity for a particular cellular
component so the stain can be used as a marker for that structure. For
example, in today's lab, the fluorescent molecule DAPI has a high affinity for
DNA. Since DNA is found predominantly in the nucleus, DAPI serves as useful
marker for the nucleus.
(2) Conjugated (indirect) stains:
In other cases, a combinatorial method is used. Here, a non-fluorescent
molecule that binds specifically to some part of the cell is covalently linked to
a fluorochrome. By itself, the fluorochrome would not bind to the cell, but by
"piggy-backing" on the specificity-conferring molecule, it is possible to
indirectly label cellular structures. For example, in today's lab, FITC is
conjugated to phalloidin to stain the actin filaments. Phalloidin binds to Factin and the attached FITC molecule fluoresces revealing the location of the
actin filaments. Since the fluorochrome itself has no specificity, the same
fluorescing moiety can be conjugated to any choice of detector molecule.
Moreover, by coupling each detector molecule to a different colored
fluorophore it is possible to distinguish multiple structures in the same cell by
their color of fluorescence. For example, in today’s lab you will triple stain
cells with three differently colored fluorochromes. This will allow you to
compare the relative orientation of the different structures in the same cell.
(3) Immunofluorescence (antibody dependent stains)
A specialized case of the combinatorial method is immunofluorescence. In
this technique, the cell is incubated with an antibody that reacts with a
specific cellular antigen (e.g. tubulin). Antibodies are particularly useful tools
because they typically bind to antigens with high affinity and specificity. The
antibody molecule by itself is not fluorescent, however, a fluorochrome can
be covalently linked (conjugated) to the antibody. For example, an FITCconjugated anti-tubulin antibody will reveal the intracellular location of
microtubules. The microtubules will glow green because they are coated with
the FITC-conjugated antibody. While it is possible to use fluorescently
conjugated antibodies directly, it is more common to use a combination of
antibodies. In this technique, known as indirect immunofluorescence, an
unlabeled primary antibody is first bound to the cellular antigen. Then, a
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Mitosis Lab
fluorochrome-conjugated secondary antibody that binds specifically to the
primary antibody is added. As with the direct method, the staining pattern
reveals the distribution of the antigen within the cell.
In today's lab, you will be using a number of fluorescent stains to reveal different
parts of the cell.
ACTIN
Microfilaments, composed of actin, will be stained green with FITC
conjugated phalloidin. Phallotoxins, originally isolated from poisonous
mushrooms, bind relatively tightly (Kd ~ 20 nM) to F-actin, the polymerized
Filamentous actin in microfilaments. They exhibit no binding to G-actin, the
cytosolic globular actin protein.
TUBULIN
Microtubules (composed of tubulin) will be stained red by indirect
immunofluorescence. The cells will first be treated with a mouse antibody
that is specific for tubulin. They are then treated with a TRITC-conjugated
secondary antibody that is specific for the primary antibody.
DNA/NUCLEI
DAPI is a small water-soluble fluorescent molecule (MW 350d) with extreme
avidity and specificity for DNA, preferentially binding to the A:T rich regions
of DNA. DAPI will also reveal the state of the DNA at different stages of the
cell cycle. When cells undergo mitosis, the DNA condenses into brightly
staining figures – the characteristic chromosome shapes. Review your
textbook for pictures of cells undergoing mitosis. During the rest of the cell
cycle, when, the DNA is not so tightly packaged, the nucleus appears as a
diffuse blue oval.
Experimental Protocol: Triple stain - actin, tubulin, DNA
For this experiment, you will be using three different fluorochromes simultaneously
to stain the cytoskeleton and DNA of PtK2 cells. PtK2 cells are cells derived from
the kidney of a potoroo, or a rat kangaroo. Prior to lab, the cells (grown on a cover
slip) were fixed in a 3.7% formaldehyde solution in PBS (phosphate-buffered saline)
for 7 minutes at room temperature, then treated with 0.1% sodium azide for 7
minutes, and finally treated with 0.5% Triton-X 100 for 7 minutes to permeabilize
the cells. The Triton-X treatment allows the antibodies to penetrate the cell's
membranes. The cells have already been incubated with a mouse antibody directed
against tubulin. This is the primary (1°) antibody.
You will treat the cover slips with the secondary (2°) antibody - a TRITC-conjugated
anti-mouse antibody. The 2° ab staining solution also contains FITC -phalloidin,
which will bind to and stain F-actin green, and DAPI to stain the DNA in the cell
blue. Finally, the cover slips will be mounted with Prolong Antifade mounting
solution.
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1. Wear gloves for this part of the lab. Cut a piece of Parafilm into a square
roughly 3 inches by 3 inches (approx. 8 cm x 8 cm). Tape it as flat as possible on
your bench. Fill up the beakers labeled PBS with 1X PBS (Phosphate Buffered
Saline solution).
2. Get a tube of stain from your instructor. The tube will contain a mixture of FITCphalloidin, the TRITC secondary antibody, and DAPI. Gently pipet (~ 140 µl) the
staining solution onto the piece of Parafilm, forming a little puddle. Try not to
create air bubbles. If the bubbles are trapped under the cover slip, the cells under
the bubble will not stain well.
3. The PtK2 cells have been grown on one side of a glass cover slip, so you need to
pay attention to the orientation of the cover slip. All the cover slips will be in the
petri dish with the cells facing up. Using a pair of forceps, remove a cover slip
from the petri dish with fixed PtK2 cells. Keep track of which side the cells are on.
Dip the cover slip in PBS (phosphate-buffered saline) a couple of times to rinse it.
Be very careful not to drop the cover slip and keep track of which side the
cells are on. If you drop the cover slip, you will not know which side has the cells
(you have a 50-50 chance of guessing the right side).
4. Touch the edge of the cover slip to a Kimwipe briefly (no more than 2-3 seconds)
to remove excess liquid. You do not want the cells to dry out on the cover slip. With
the forceps, gently place the cover slip CELL SIDE DOWN onto the drop of staining
solution. Place a box over the Parafilm/cover slip to protect the cells from any stray
light. Stain the cells at least for 45-60 minutes.
5. Take a glass slide, wipe it clean with a Kimwipe and lay it down on the benchtop.
Use a marker to label the edge with your initials and the slide number (=#1,
TRIPLE).
6. Just before the staining is finished, put 25 µl of Prolong mounting medium onto
the center of your labeled slide. Do not put the Prolong on the cover slip.
7. When the staining period is done, pipet 200 µl PBS onto the Parafilm, just
adjacent to the cover slip. This will raise the cover slip somewhat. Carefully, slide
your forceps under the cover slip and pick up the cover slip.
8. Wash the cells twice in PBS, for about a minute each. During the wash steps,
remain aware of which side of the cover slip the cells are on. You have two
beakers of PBS. Dip the cover slip quickly in beaker PBS #1 and then repeatedly for
about 30-60 seconds in beaker PBS #2. You do not have to worry about over
washing - the antibody complexes are quite stable.
9. Drain the PBS off of the cover slip by holding the cover slip at an angle with the
lower edge briefly touching a Kimwipe. Don't rub the cells off and don't let the cells
dry out. Try to get most of the PBS off of the cover slip without drying out the cells.
10. Mount the cover slip onto the slide, lowering the cover slip gradually so that the
Prolong mounting medium spreads under the cover slip without creating bubbles.
Carefully slide out the forceps. Ideally there should be no excess liquid around the
cover slip. Gently, use a Kimwipe to blot off any excess liquid. Soak up the staining
solution on the Parafilm with a Kimwipe and put the Parafilm and Kimwipe into the
STAIN WASTE beaker at your station.
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VIEWING THE CELLS
11. Allow the Prolong mounting medium to dry (several hours). Before viewing the
slide, rinse off the cover slip with deionized water. Gently blot the top of the
slide dry with a Kimwipe. Thoroughly dry the underside of the slide with a Kimwipe.
12. Observe the cells using the fluorescence microscope. Focus on the cells using
the 20X objective and the PH1 phase contrast setting.
13. Once the cells are in focus, turn off the visible light source (the switch is located
on the front base of the microscope). You will not be able to see the fluorescent
signal if the visible light is left on. Select the appropriate filter cube for the
fluorochrome you want to observe.
14. Turn the filter cube to the "DAPI" position (second position from the left for
filter sliders) to examine the DAPI (DNA) stain. The filter cube slider is just below
the eyepieces. Open the shutter by pushing in the shutter slider (right side of
scope). You should see a white coming out of the objective, but avoid looking at the
UV light directly bouncing off the slide - the UV light can damage your eyes. When
you look at the cells through the microscope, you should see blue fluorescence.
Scan the slide, looking for mitotic figures. In interphase, the DNA in the nucleus will
appear as a diffuse blue spot. During mitosis, DNA (chromosomes) will be
condensed into discrete figures. Scan the field to find a nice set of cells to examine
further.
LOOK AT THE STAINING PATTERNS
15. Next, rotate the filter cube turret to the "TRITC" position (fourth position from
the left for filter sliders) to examine the TRITC fluorochrome. This will reveal the
microtubule pattern. Try not to move the stage when switching from DAPI to
TRITC. This allows you to examine the different structures in the same set of cells.
Make a sketch of the stained cells. Describe the distribution, size, and shape of
microtubules and describe their arrangement relative to the nucleus.
16. Finally, switch to the “FITC” cube (third position on the left for filter sliders) to
examine the FITC-phalloidin staining pattern. Make a sketch of the stained cells.
Describe the distribution, size, and shape of microfilaments.
Try to find cells in different stages of mitosis: prophase, metaphase, anaphase,
telophase. Scanning the slide may be easier using the 20X objective. Then go back
and examine the actin and tubulin patterns in these mitotic cells.
Compare the appearance of microfilaments and microtubules in mitotic cells versus
interphase cells.
Does the distribution of the microfilaments/microtubules change?
Are there any differences in the structure of the microfilaments or the
microtubules?
Why does the distribution and/or structure of the microfilaments/microtubules
change during different stages of mitosis?
Compare the cell shape of interphase and mitotic cells.