Biology 380
Cell Biology
Laboratory Manual
Prepared by The CSULA Department of Biological Sciences
Edited by Dr. Robert M. Nissen
TABLE OF CONTENTS
HOW TO KEEP YOUR LAB NOTEBOOK
3
1. INTRODUCTION TO BIOLOGICAL INSTRUMENTATION
4
2. SUBCELLULAR FRACTIONATION OF ORGANELLES
10
2A. CELLULAR FRACTIONATION
10
2B. PROTEIN QUANTIFICATION OF CELLULAR FRACTIONS
15
2C. ENZYME MARKERS TO ESTABLISH FRACTIONAL PURITY
19
3. PHOTOSYNTHESIS: THE HILL REACTION
23
4. RT-PCR ANALYSIS OF GENE EXPRESSION
28
4A. RNA ISOLATION
30
4B. RT-PCR
32
4C. AGAROSE GEL ELECTROPHORESIS
34
2
HOW TO KEEP YOUR CELL AND MOLECULAR BIOLOGY LAB NOTEBOOK
1. The notebook must be either spiral bound or hardcover.
2. The first page should contain the Table of Contents.
All pages should be numbered.
3. The notebook should be kept in continuous chronological order. The date should
be written on the upper left hand side of each page.
4. No pages or lines are to be skipped (left blank). If you want to leave a space
between entries, you must “X” out the page or line.
5. All data must be placed in the notebook in tables that are clearly titled. Your lab
instructor must be able to tell by looking at the data where the numbers came
from, i.e. OD400 readings or Apparent Molecular Mass. Numbers without units
are meaningless. Tables from the lab protocol handouts may be taped or pasted
into the lab notebook.
6. In general, materials and methods need not be written out extensively in the
notebook unless we deviate from the methods described herein. In addition to
providing the place for your data, the purpose of the notebook is to document
what you actually did. Therefore, using the lab manual and the notebook, another
scientist should be able to reproduce your work.
7. In addition to quantitative data, any observations of interest should be noted.
8. At the end of each experiment you should discuss the data with respect to relevant
aspects of Cell Biology theory as covered in the lecture and the lab. In addition,
you should note any errors or deviations in the outcome of your experiment from
the expected and discuss the potential sources for the error or deviation.
9. Entries in the notebook should be done in lab or immediately thereafter.
Notebooks must be legible. However, excessive neatness often indicates a
notebook done well after the fact! You need to write things down as they occur
because important facts or events are quickly forgotten or become unclear in
memory. The purpose of your notebook is to faithfully describe what you actually
did. It does not need to be a work of beauty ready for display on a museum shelf!
3
INTRODUCTION TO BIOLOGICAL INSTRUMENTATION…….1
INTRODUCTION
The spectrophotometer is an instrument commonly used by biologists. The
spectrophotometer enables biologists to investigate many chemical and physiological
processes that characterize the living cells. The purpose of the first exercise is to describe
in simple terms how this instrument works and to provide an exercise directly utilizing
the spectrophotometer to demonstrate the Beer-Lambert Law and prepare a standard
concentration curve for a colored solution used in future laboratory exercises (2,6dichlorophenol inophenol, or DCPIP). Your ability to understand the concepts
introduced in this exercise will be essential for the remaining laboratory exercises in this
course. In addition, you are given instructions for the use of automatic micropipetters,
which are used throughout this course.
RECORD ALL YOUR DATA AND RESPONSES IN YOUR LABORATORY
NOTEBOOK AND TURN YOUR COMPLETED WORK IN TO THE LABORATORY
INSTRUCTOR DURING THE NEXT CLASS PERIOD.
YOUR WORK WILL BE CORRECTED AND RETURNED TO YOU DURING THE
NEXT LABORATORY SESSION.
4
PIPETTER OPERATION
1. The range of volume dispensed by each pipette is indicated on the side of the
instrument and is in µl increments (2-20, 20-200, 200-1000 µl). Hold the pipette in
one hand. With the other hand turn the volume adjustment knob until the required
volume shows on the digital indicator. If you pass the desired setting, turn the dial a
couple of revolutions above the desired setting and reset the volume. Always DIAL
DOWN to the desired volume for maximum accuracy. Use the lock ring to lock the
volume selector in place.
2. Attach a new disposable tip to the pipette shaft. Press firmly to ensure a positive
airtight seal.
3. Depress the plunger to the first stop. This part of the stroke is the calibrated volume
displayed on the digital volume indicator.
4. Holding the pipette vertically, immerse the disposable tip into the sample liquid.
5. Allow the push-button to return slowly to the UP position. Never let it snap up.
6. Wait a moment to ensure that the full volume of the sample is drawn into the tip.
Withdraw the tip from the sample liquid. Should any liquid remain on the outside of
the tip, touch it to the inside edge of the vessel to remove the excess liquid.
7. To dispense sample, touch the tip end against the inside wall of the receiving vessel
and depress the plunger slowly to the FIRST STOP. Wait about two seconds (longer
for viscous solutions), then press the plunger to the SECOND STOP (bottom of
stroke), expelling any residual liquid in the tip.
8. With the plunger fully depressed, withdraw the pipette from the vessel carefully,
with the tip sliding along the wall of the vessel. Allow the plunger to return to the
UP position.
9. Discard the tip by depressing the tip ejector button. A fresh tip should be used for
each sample to prevent sample carryover.
TEST OF PIPETTING ACCURACY
Pipette 100 µl, 500 µl, and 50 µl of water onto three separate tared weigh boats.
Make sure you use the appropriate pipette. Measure the weight of the water. Record
your results. Calculate the % error in your pipetting. It is important that you weigh
directly after pipetting due to inaccuracies caused by evaporation.
5
INTRODUCTION TO SPECTROSCOPY
References

Pavia, D.L., Lampman, G. M., Kriz, G. S. Introduction to Spectroscopy. Saunder’s
College Publishing: Florida; 1996.

Mathew, C.K. &Van Holde K. E. Biochemistry. The Benjamin/Cummings
Publishing company, INC.: Menlo Park, California: 1996.
The spectroscopic measurement of protein concentration uses a
spectrophotometer. A cuvette of sample cell length (thickness) 1 cm containing a solution
of the protein is placed in a beam of monochromatic radiation of intensity Io. The
intensity of the emerging beam will be decreased to a value I because the solution
absorbs some of the radiation.
The greater the number of molecules capable of absorbing light of a given
wavelength (), the greater the extent of absorption. Furthermore, the more effectively a
molecule absorbs light of a given wavelength (), the greater the extent of light
absorption. From these ideas the following equation, known as the Beer-Lambert’s
Law may be formulated:
A = log (Io/I) =  . c . l
A
Io
I
c
l

=
=
=
=
=
=
absorbance
intensity of light incident upon the same cell
intensity of light leaving the sample cell
molar concentration of the solute
length of the sample cell (cm)
molar absorptivity (extinction coefficient)
The dimensions of the extinction coefficient depend on the concentration units
employed. If protein concentration is measured in molarity, the unit for  has to be M-1
cm -1 since the absorption is a dimensionless quantity.
A plot of the absorbance of a substance in solution at a given wavelength, as a
function of the molar amount of the substance present, is called a calibration plot or
standard curve. If the length of the sample cell is 1cm, the slope of a standard curve is the
extinction coefficient. An absorbance spectrum of a given compound is a graphical
representation of the absorbance at each wavelength over a given wavelength range
(usually UV-visible), that is, A vs. . Chemically different compounds have their own
distinctive absorbance spectrum. You will be determining the absorbance spectrum for
the electron acceptor 2,6-dichlorophenol inophenol (DCPIP) and will also plot
absorbance spectra in a future laboratory exercise on plant pigments.
6
SPECTROSCOPY: The Instrument
The typical UV-visible spectrophotometer consists of a light source, a
monochromator, and a detector. The light source is usually a deuterium lamp, which
emits electromagnetic radiation in the UV region of the spectrum. A second light source,
a tungsten lamp, is used for the wavelengths in the visible region of the spectrum. The
monochromator is a diffraction grating; its role is to spread the beam of light into its
component wavelengths. A system of slits focuses the desired wavelength on the sample
cell. The light, which passes through the sample cell, reaches the multiplier tube. In a
typical double beam instrument, the light emanating from the light source is split into two
beams, the sample beam and the reference beam. When there is no sample cell in the
reference beam, the detected light is taken to be equal to the intensity of light entering the
sample.
The sample cell must be constructed of a material that is transparent to the
electromagnetic radiation being used in the experiment. For measurements in the visible
range of the spectrum, cells composed of glass or plastic are generally suitable. For
measurements in the ultraviolet region of the spectrum, however, glass and plastic cannot
be used because they absorb ultraviolet radiation. Instead, cells made of quartz must be
used, since quartz does not absorb radiation in this region.
The instrument design just described is quite suitable for measurement at only one
wavelength. If a complete spectrum is desired, this type of instrument has some
deficiencies. A mechanical system is required to rotate the monochromator and provide a
scan of all desired wavelengths.
Using the spectrophotometer
1. The machine needs to warm up prior to use. Turn on the power switch and allow the
machine to warm up for at least 15 minutes.
2. Using the A/T/C option on the spectrophotometer, select the absorbance mode.
3. Using the nm button, select the appropriate wavelength.
4. Insert your cuvette by positioning it firmly in the opening provided.
5. Insert your blank and press OABS/100%T to calibrate the machine with your blank.
6. Remove the blank and insert your samples one by one and record the absorbance.
You do not need to blank between measurements.
7
PREPARING A STANDARD CURVE and CALCULATING THE MOLAR
EXTINCTION COEFFICIENT
1. Create a 100M stock solution of DCPIP by making a 1:10 dilution (10ml of 1mM
DCPIP + 90ml of DW).
2. Using the following table, create different concentrations of DCPIP by making serial
dilutions. Remember to change pipette tips between dilutions.
Tube Absorbance
(@ max. absorbance)
#
1
2
3
4
5
6
7
8
9
0.0
ml of
DCPIP
Stock
solution
0.0
ml of
distilled
water
Total volume
(ml)
Final DCPIP
concentration
(M)
5.0
5.0
5.0
5.0
5.0
5.0
5.0
5.0
5.0
5.0
0.0
0.2
0.5
1.0
5.0
10.0
20.0
50.0
100.0
Tube #9 should be the deepest in color.
EXERCISE
1. Set the spectrophotometer to the wavelength (595nm) at which DCPIP maximally
absorbs light.
2. Insert tube#1 and zero the spectrophotometer by pressing OABS/100%T to calibrate
the machine with your blank. Determine the absorbance of tubes #2-9 at this
wavelength and construct a standard curve on graph paper in your notebook.
3. Determine the molar extinction coefficient for the dye from your standard curve:
Molar extinction coefficient DCPIP @ _____nm = _____.
4. Calculate the molarity of the unknown solution (provided for you by the lab
instructor) using the Beer-Lambert law. Use the molar extinction coefficient you
have obtained from your curve.
8
ADDITIONAL PRACTICE PROBLEM
UV spectroscopy of phenol red resulted in the following data:
ml of 10-4 phenol red
M phenol red
A 560
0 (Blank)
0.05
0.10
0.20
0.30
0.40
0.50
Unknown
0
1 x 10-6
2.0 x 10-6
4.0 x 10-6
6.0 x 10-6
8.0 x 10-6
1.0 x 10-5
0
0.06
0.12
0.26
0.34
0.47
0.59
0.45
___________
1. Construct a standard curve in the provided space below.
2. Obtain the concentration of the unknown using the Beer-Lambert law.
ANSWER TO THE PRACTICE PROBLEM
The slope of the standard curve or the extinction coefficient will be 5.9 x 104 M-1 cm-1.
c unknown = Aunknown/(  l )
Answer: c unknown = 0.45/(5.9 x 104 M-1 cm-1 * 1 cm) = 7.6 x 10-6 M
9
CELLULAR FRACTIONATION…….2A
INTRODUCTION
All eukaryotic cells contain organelles that perform specific functions. Cellular
fractionation is a technique that enables the researcher to separate and isolate bulk
quantities of organelles and other cellular components. Once isolated, the functioning of
the cellular constituents can be studied more easily than within the intact cell.
Fractionation begins with homogenization, which involves the disruption of the
cells by mechanical shearing, ultrasound, or osmotic shock. This process releases the
cellular organelles, which can then be separated from each other by centrifugation.
Centrifugation at low speeds causes the sedimentation of only the largest and densest of
components. These components can be collected and the “unpelleted” portion, or
supernatant, can be centrifuged again at higher speeds and for longer durations to “pellet”
successively smaller components. Separation of organelles and other particles in solution
can be described mathematically using Stokes’ Law.
 = 2cr2 (d-d0/ 9)
Where;
 = velocity of sedimentation of the particle
c = centrifugal acceleration and is proportional to revolutions per minute of the
centrifuge
r = radius of the organelle
d and d0 = the density of the organelle and medium respectively
 = viscosity of the medium
Therefore, the rate at which an organelle settles is proportional to its radius and density.
Once isolated, the components can be assessed for purity by light or electron
microscopy and their biochemical activities assayed to enable structure/function
relationships to be determined.
In the following exercise, you will fractionate tissue from Mung Bean (Phaseolus
mungo aureus) seedlings to isolate cellular functions enriched in nuclei, mitochondria,
and cytoplasm. An overview of the procedures is provided in figure 1.1.
10
PRODEDURE FOR CELLULAR FRACTIONATION
A.
HOMOGENIZATION OF SEEDLINGS
Steps 1 though 3, part A should be performed per lab bench (2 groups). All
solutions and glassware should be maintained on ice.
1.
Chop 20 g of etiolated seedlings and add to 80 ml of ice-cold isolation medium.
Remember to keep the tissue cold.
2.
Pour the suspended tissue into a blender and homogenize the tissue with several
short bursts (10 seconds) of the blender. The tissue suspensions from different
labs benches may be pooled to save time. This blended solution will be referred to
as the homogenate.
3.
Place 2-4 layers of cheesecloth, pre-wetted with isolation medium, over a clean
400 ml beaker. Pour the homogenate though the cheesecloth into the beaker
maintained on ice. Rinse the blender with a further 10 ml of ice-cold isolation
medium and pass the wash through the cheesecloth into the beaker. Gently
squeeze the cloth to remove the remaining fluid from the tissue fragments. You
should have approximately 90 ml of tissue homogenate with a tissue: buffer ratio
of 1:4.5 (weight to volume). At this point, we will split the homogenate between
the two groups.
4.
Split the homogenate in half by pouring equal amounts into each of two 50 ml
cold, polypropylene centrifuge tubes. You must balance the tubes at this point.
Each group will now work with one of the tubes for the remainder of the
procedure. Keep the tubes on ice.
5.
Aliquot two 1.5 ml samples of the homogenate into clean, labeled microfuge
tubes. Label the tubes with your group name, the volume and sample type (in this
case homogenate) and place on ice. These samples will be used for later assays.
B.
DIFFERENTIAL CENTIFUGATION
1.
Take balanced pairs of centrifuge tubes from step A4 and centrifuge in a precooled centrifuge (4o C) for 5 min. at 500 g. Pour or pipette the supernatant
(taking care not to resuspend the pellet) into a cold, clean 50 ml centrifuge tube.
Discard the pellet, which consists mainly of whole cells and cell wall debris.
2.
Centrifuge the supernatant from step B1 at 600 g for 10 min. Make sure tubes are
balanced. Pour or pipette the supernatant (taking care not to re-suspend the pellet)
into a cold, clean 50 ml centrifuge tube and maintain on ice.
3.
Re-suspend the pellet in the centrifuge tube using 20 ml of isolation medium and
gently agitating with a paint brush or Pasteur pipette. Recentrifuge at 800 g for 10
11
min. Discard the supernatant and resuspend the pellet in 5 ml of ice-cold isolation
medium. This fraction is the nuclear fraction.
4.
Centrifuge the supernatant obtained during step B2 for 15 min. at 15,000 g.
Transfer the supernatant into a cold, clean centrifuge tube. This is the cytosolic
fraction.
5.
Resuspend the pellet obtained from Step B4 in 5 ml of ice-cold isolation medium.
This sample is the mitochondrial fraction.
5. You should produce 4 samples: the crude homogenate (two x 1.5 ml), nuclear
(approx. 5 ml), mitochondrial (5 ml) and cytosolic fractions (approx. 40 ml.). Keep
these samples on ice.
Figure 1.1. Overview of Cellular Fractionation Protocol.
1. Etiolated seedlings
2. Chop with razor
3. Blend with ice-cold isolation medium (1:4.5 tissue: buffer)
4. Strain through cheesecloth into 400 ml beaker
5. Transfer homogenate into 50 ml centrifuge tubes
6. Keep 3 ml homogenate
for future labs
7. Balance tubes.
8. Centrifuge at 500 g, 5 min.
9. Centrifuge at 600 g, 10 min.
10. Pellet
11. Resuspend in 20 ml isolation
medium
14. Supernatant (store on ice)
15. Balance and centrifuge 15,000 g, 15 min.
16. Pellet
17. Supernatant
12. Centrifuge 800g, 10 min.
13. Nuclear pellet/fraction
(resuspend in 5 ml buffer)
Mitochondrial Fraction
Cytosol fraction
(resuspend in 5 ml buffer)
12
D. STORAGE OF CELLULAR FRACTIONS
Divide the remainder of each of your four samples as shown below in table 1.1.
Use microfuge tubes and label them carefully using indelible markers. Include the
fraction name, volume, and group name on the label. Place samples in the -80o C freezer.
These samples will be used in the later exercises. The crude homogenate samples were
prepared in step A5.
Table 1.1
Sample
Aliquot Size
Tube 1
Tube 2
Crude Homogenate
1.5 ml
1.5 ml
Nuclear Fraction
1.5 ml
1.5 ml
Mitochondrial
1.5 ml
1.5 ml
Cytosolic
1.5 ml
1.5 ml
13
METHODS OVERVIEW (for quick reference):
Preparation of Bean Sprouts
1. Weigh out 20 grams (2 groups share)
2. Chop (crudely) with razor
3. Add ice cold isolation medium (80 ml)
4. Blender (short bursts!)
5. Filter through cheesecloth
Add extra 10ml of isolation medium to
“wash” blender (total volume = 90 ml)
Homogenate
Reserve 2x 1.5ml samples*
6. Split in half for the 2 groups (2 x 50 ml tubes)
Centrifugation
1. Balance tubes
2. Centrifuge @ 500xg “Junk Step” - Discard pellet
3. Centrifuge supernatant from step 2 @ 600xg for 10 min
resuspend pellet in 20ml isolation
medium, re-spin (centrifuge) @ 800xg for
10 min, resuspend pellet in 5ml isolation
medium (nuclear)
* reserve 2 x 1.5ml
4. Centrifuge supernatant from step 3 @ 15,000xg for 15 min
Resuspend pellet in 12ml isolation
medium (mitochondrial)
* reserve 2x 1.5ml
Supernatant = cytosol
* reserve 2 x 1.5ml
14
PROTEIN QUANTIFICATION OF CELLULAR FRACTIONS……2B
INTRODUCTION
The biochemical activity of a cellular protein is often expressed in the form of
activity mg-1. This expression requires that the protein content of the sample be
quantified. In this exercise, you will determine the protein content of the cellular fractions
obtained in the previous exercise using the Coomassie Blue method. You will be required
to prepare a standard curve by making serial dilutions of bovine serum albumen (BSA)
from a concentrated solution (2.0 mg ml-1). The standard curve can be used to calculate
the protein content of each cellular fraction.
A. PROCEDURE FOR PREPARATION OF A PROTEIN STANDARDIZATION
CURVE
1. You are provided with an ampule containing 2.0 mg ml-1 BSA. Prepare 4 ml of BSA
solution at a concentration of 333.3 µg ml-1 by diluting the BSA solution in the
ampule with the appropriate amount of distilled water.
2. Label 7 cuvettes and prepare a dilution series of BSA at the concentrations listed in
table 2.1. Start by adding 4ml of Coomassie Blue to each cuvette (the final total
volume will be 5 ml --see Table 2.1). The reagent contains the dye Coomassie
Brilliant Blue G-250, which binds to proteins to form a blue complex whose
absorbance is directly proportional to protein concentration.
3. Remember to calculate the volume first (see tables), then prepare all of the solutions
for the standard curve together (add appropriate volumes of BSA to the Coomassie
Blue). Mix the tubes thoroughly immediately after the dye has been added. Wait five
minutes, then read absorbance for each sample on the spectrophotometer.
4. Set the spectrophotometer wavelength to 595nm. Use tube#1 as a blank and set the
spectrophotometer to zero. Using this wavelength, measure the remaining tubes.
5. Plot the results on graph paper, plotting the absorbance of each tube on the y-axis
(vertical), and the protein concentration (g/ml) on the x-axis (horizontal).
15
Table 2.1
Tube #
1
2
3
4
5
6
7
BSA added
(Solution 1)
(ml)
0
Distilled
water added
(ml)
1
1.0
0.0
Coomassie
Reagent added
(ml)
4
4
4
4
4
4
4
Protein
content (g)
0
33.3
66.7
133.3
200.0
266.6
333.3
Protein
concentration
(g/ml)
0
6.66
13.32
26.64
39.96
53.28
66.66
Absorbance at
max
B. CALCULATION OF PROTEIN CONTENT OF CELLULAR FRACTIONS
COLLECTED EARLIER
1. Defrost one tube of each of your refrigerated sample types prepared during cellular
fractionation and pipette 500 l from each into four separate glass test tubes. Lyse
the samples by the addition of 2000 l of 0.1%SDS in 1mM EDTA (prepared for you
by the technician) and immerse the tubes in boiling water for two minutes. Remove
the tubes with the tongs provided – take care not to get burned.
2. Dilute 200 l of each of our boiled samples with 1.8ml of distilled water in a clean
cuvette. Mix the contents thoroughly using a new pipette tip. You should have two
tubes for each fraction type with concentrations of protein differing by an order of
magnitude. This is referred to as a serial dilution. Transfer 1ml of each sample to a
new tube and add 4 ml of the Coomassie reagent. Once again, gently, mix the
contents thoroughly by inverting a number of times. Measure the absorbance at the
595nm and record the data in Table 2.2. This assay is temperature dependent.
Make sure your samples are at room temperature before reading.
3. Determine the concentration of protein in the original sample held in the freezer from
the BSA protein standard curve. Make sure to account for the dilutions. Which
fraction contains the most protein? These values will be used in the following
exercise.
16
Outline of Experiment
A. PREPARE PROTEIN STANDARD CURVE
Ampule contains 1ml of 2mg/ml solution (BSA is a protein standard)
1. 2mg/ml (1ml) = 2mg
0.333mg/ml (xml) = 2 mg
x = 2mg/0.333mg/ml = 6ml
(1ml ampule & 5 ml H2O)
this is solution (1)
solution (1) will be used to make most concentrated
BSA solution for standard curve
will be: 1ml solution (1)
4ml coomassie blue reagent
final conc. = 333.3/5 = 66.6 g/ml
2. Calculate volumes required to make all solutions of the standard curve
3. Assemble all blue solutions at once, wait 5 mins, then read at 595nm (abs. Max)
B. PREPARE SAMPLES (from previous lab)
1. Remove one each of your 1.5ml aliquots from freezer, thaw
2. Prepare boiling water bath (600ml beaker on hotplate)
3. Take 500 l from each sample (above), add 2000 l SDS solution (prepared by
technician)
Boil x 2 min
Take care (use tongs)!
This is the 1:5 dilution
4. Take 200 l from step 3 above, add 1.8ml water. This is the 1:50 dilution
5. Take 1ml from each of these diluted, boiled samples, add 4.0ml Coomassie Blue,
wait 5 min., read absorbance.
6. Will extrapolate from standard curve to get unknown concentration.
*Remember to multiply by 5 or 50 to get actual protein concentration
Take the average of your 2 answers to get the protein concentration.
Alt. : use the reading obtained from 1:5 dilution
17
only (if @ 600 Abs. Range, this is most accurate)
18
TABLE 2.2
Sample
Absorbance
Protein Concentration
(g/ml) Corrected for
Dilution
Homogenate
1/5 Dilution*
1/50 Dilution
Cytosol Fraction
1/5 Dilution
1/50 Dilution
Mitochondrial Fraction
1/5 Dilution
1/50 Dilution
Nuclear Fraction
1/5 Dilution
1/50 Dilution
19
ENZYME MARKERS TO ESTABLISH FRACTIONAL
PURITY…….2C
INTRODUCTION
Enzymes are proteins that act as catalysts to accelerate biochemical reactions.
Although many chemical reactions are thermodynamically feasible, they are often not
spontaneous and require energy, termed “activation energy”, which is the minimum
amount of energy that two molecules must have in order for them to start to react. In the
laboratory, activation energy can be applied in the form of heat to increase the kinetic
energy of the molecules to initiate the reaction. Clearly, thermal activation is not a
feasible mode of initiating reactions within cells. So, the biological solution is to use
enzymes as catalysts to lower the activation energy such that reactions can proceed at
temperatures and pressures that are compatible with cellular processes.
One of the major objectives of a previous laboratory was to use centrifugation to
separate and isolate different components within the cell. The success of the separation
procedure is commonly assessed by monitoring the enzymatic activity of enzymes that
are characteristic of components expected within each faction. For example, succinic
dehydrogenase (SDH) activity is commonly used as an indicator of mitochondrial purity:
This inner mitochondrial membrane bound enzyme catalyzes the oxidation of succinate
to fumarate within the Krebs cycle. SDH covalently binds the coenzyme FAD which
accepts electrons during the oxidation of the succinate. In the cell, the reduced form of
FAD, FADH, normally becomes reoxidized by iron-sulfur proteins in the electron
transport chain to complete the catalytic cycle. The activity of this electron cascade can
be monitored in vitro by the addition of artificial electron acceptors of higher reduction
potential than FAD that show changes in absorption characteristics upon reduction. One
useful artificial electron acceptor is 2,6-dichlorophenol indophenol (DCPIP) which shows
a decrease in absorption at 600nm upon reduction and can therefore be used to measure
the rate of the enzyme catalyzed reaction.
A.
PROCEDURE FOR THE QUANTIFICATION OF SUCCINIC
DEHYDROGENASE ACTIVITY
1. Set the spectrophotometer to 600 nm and zero it using distilled water as a blank.
2. Pipette 2.5ml of the blue reaction mixture provided into a clean cuvette and preincubate the tube at 37oC for 10 min. using the water bath provided.
The reaction mixture contains a pre-filtered solution of the following: 10mM
succinate substrate; 0.025mg/ml DCPIP, electron receptor indicator dye; 10mM K2HPO4
buffer, pH 7.4, physiological pH buffer; 2mM KCN, respiratory poison to block electron
transport chain; 1mM salicylhydroxamic acid (SHAM), respiratory poison for cyanide
insensitive alternate oxidases in electron transport chain; 1mM CaCl2, to increase
20
mitochondrial membrane permeability; and 0.5 mg/ml bovine serum albumen (BSA), to
maintain osmotic balance.
(Thaw your remaining small cellular samples at this point)
3. Initiate the reaction in the cuvette by the ADDITION of 1.0ml of isolation medium
(used in exercise 1 for isolating your fractions). Place a parafilm seal over the mouth
of the tube and mix thoroughly by inverting 3 times. Be careful not to introduce too
many air bubbles in your sample. IMMEDIATELY take an absorbance reading at
600nm. Record the value. The initial absorbance reading should be around 1.0.
This will be your t = 0 time point. IMMEDIATELY transfer the cuvette back into
the water bath.
4. Take absorbance reading at t = 1, 3, 5, 10, and 15 min taking care to mix the contents
by inverting the tube before taking the measurement. This sample is your control
and you should observe little or no change in absorbance over time.
5. Defrost one 1.5 ml tube of each of your 4 cellular samples; homogenate, nuclear,
mitochondrial, and cytosolic fractions. Conduct the procedure described above for
the homogenate and all the subcellular fractions, substituting 1.0 ml of the fraction
for the isolation medium. Record your data in Table 3.1:
TABLE 3.1
Time (min.)
Control
(Blank)
Homogenate
Cytosol
Fraction
Mitochondrial
Fraction
Nuclear
Fraction
0
1
3
5
10
15
Do the blank first, then “tag team” your other four fractions with your group (preincubate 4 sample of blue mix), add the 1.0ml of cellular sample to each at time 0 (one
after the other at 1-5 minute intervals)
21
Outline of Experiment:
1. Defrost your remaining small tubes from lab (2)
2. ‘Incubation’ of “BLANK”
a. Set spec. to 600nm
b. 2.5ml reaction mix (Blue)
into cuvette, pre- incubate at 37oC x 10 min
(water bath)
c. Add 1.0 ml (1000 l) plain isolation medium, mix,  immediately take
absorbance reading = To
d. Take reading at 1, 3, 5, 10, 15 minutes (mix before each reading)
3. Incubation of your samples
a. Set up and pre-incubate 4 tubes with reaction mix (Abs. above) x 10 min
b. Add 1.0 ml (1000 l) of your cellular fraction to each, mix, take To reading
c. Return to water bath, take reading at 1, 3, 5, 10, 15
22
DATA ANALYSIS FOR EACH FRACTION:
Amax
∆ Abs. over 15 min.
Abs.
= Amax - Amin
Amin
- - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - - -- -
Time (min)
1.
To calculate the concentration of DCPIP reduced/min, use
BEER’S LAW: A = c .  . l
l = path lenth (1cm)
 = molar extinction coefficient for DCPIP = 0.014 m-1 cm-1
c/min = ∆ Abs. /  . time
2. Express final answer as concentration of DCPIP reduced per minute per mg protein
in your sample (determined in last week’s lab)
c/ mg protein = moles per min per mg protein
23
PHOTOSYNTHESIS: THE HILL REACTION…….3
INTRODUCTION
In 1905, Blackman suggested that photosynthesis consisted of two major series of
reactions. The first series is referred to as the “light reactions” because they are light
dependent and are responsible for trapping radiant energy. The second reaction series
does not require light, but makes use of the energy captured by the light reactions for the
synthesis of organic molecules. The fact that a plant in flashing light has essentially the
same photosynthetic rate as a comparable plant in constant light, provided that the flashes
are spaced at very close intervals, is evidence that photosynthesis consists of two major
reactions. The objective of this experiment is to test the hypothesis that the light reaction
does in fact result in the formation of compounds that contain the energy of light.
Robert Hill first demonstrated that isolated chloroplasts, in the presence of light,
produced oxygen and reducing power. The latter is especially significant because it
represents a form of chemical energy. Hill’s experiments, concerning the reactions which
now bear his name, involve only the light-dependent or energy capture phase of
photosynthesis. No carbon dioxide is fixed, and no carbohydrate is formed.
A general equation for this process may be written:
A + H2O  A2e- + 2H+ + 1/2O2
(1)
Here, A stands for any suitable electron acceptor. While NADP+ may be the
ultimate electron acceptor of the energy capture process in the chloroplasts of living plant
cells, many additional electron acceptors are sequentially reduced before this final
reduction step and the subsequent transfer of the chemical energy to the carbon
assimilation process of the Calvin cycle.
The Hill reaction procedure requires the introduction of a substitute electron
acceptor which captures electrons early in the series of electron transfers. We will use
dichlorophenol indophenol (DCPIP) as a convenient substitute acceptor. This organic
compound, when in solution appears blue in the normal oxidized form but becomes
colorless when reduced, that is, after accepting electrons. Progress of the reduction
reaction will be followed by a decrease in optical density or absorbance by DCPIP at
600nm.
The progress of the reduction reaction in the chloroplast is light dependent. As
light intensities increase, the rate of the reaction increases to the point where the light
gathering mechanism is saturated. Today, you will examine the effect of light intensity
(expressed as moles of photons per unit area per second) on the reducing power of
24
chloroplasts. Light saturation of the photosynthetic mechanisms in the chloroplast will
be demonstrated.
The Hill reaction illustrates an important methodology in determining the function
of an organelle: removal of the organelle from the cytoplasm, and preparation of a
“clean” dense suspension of these minute bodies to determine their metabolic
capabilities. Preparation of active organelle suspensions requires care. To maintain the
organelles in an active condition, one must prevent osmotic injury, bursting of the
membrane-bounded organelles, and inactivation of enzyme proteins. One must also
provide the optimum pH for the reaction, and may have to supply any inorganic ions
required by the enzyme system.
During the chloroplast isolation procedure, all solutions and glassware should be
maintained on ice. The chloroplast suspension should be kept at low temperature at all
times. The initial grinding of leaves should be performed by two groups, and the
material subsequently divided into four or six portions, equally weighted to correspond to
the tube capacity of the refrigerated centrifuge.
PROCEDURE
A. ISOLATION OF CHLOROPLASTS
Isolate the chloroplasts in the following manner:
1. Grind 50 grams of fresh spinach leaves in a laboratory blender for one minute, with
150 ml of ice-cold 0.40 M sodium chloride solution (NaCl).
2. Filter the material through several layers of cheesecloth, and divide the cell
suspension into four equally weighted, cold centrifuge tubes.
3. Using the Sorvall® RC 5B centrifuge, spin the material for three minutes at 100 x g.
4. Remove the tubes from the centrifuge and decant the suspension into additional cold
centrifuge tubes. Discard the “pellet” at the bottom of the tube; this consists only of
cell wall fragments and nuclei.
5. Centrifuge the suspension for 10 minutes at 1400 x g. The increased gravitational
force should bring a large number of chloroplasts to the bottom of the tube.
6. Remove the tubes from the centrifuge and discard the supernatant. Resuspend the
chloroplasts in approximately 20ml of ice-cold 0.40 M NaCl solution.
7. Centrifuge the chloroplasts once again at the same relative gravitational force
(revolutions per minute) as step 5 for 10 minutes.
8. Remove the centrifuge tubes, discard the liquid, and resuspend the “pellet” of
chloroplasts, this time in 20ml of 0.05 M sodium phosphate buffer at pH 6.4.
25
Make as uniform a suspension of chloroplasts as you can, and use this suspension for
your active chloroplasts preparation.
B. THE REDUCING POWER OF CHLOROPLASTS
To demonstrate the light-dependent reducing power of isolated chloroplasts, proceed as
follows:
1. Obtain 9 spectrophotometer cuvettes and label them according to table 5.1. The last
cuvette will be used as your blank. Pipette into each cuvette 4.0 ml of Na2HPO4
buffer solution and 1.0 ml of the blue indicator dye, DCPIP. Tightly wrap all but one
of the cuvettes with aluminum foil to exclude light. Prepare a foil cap for the top of
each tube as well.
2. In order to create a blank, pipette 1 ml of the chloroplast suspension into the
uncovered cuvette, and then add a small amount of solid ascorbic acid. Mix well.
The color should turn from blue to a pale green color. Ascorbic acid is an electron
donor (reducing agent). This will serve as the spectrophotometer blank, with which
the instrument will be adjusted to zero absorbance. This cuvette may remain in the
light. Maintain all cuvettes in a beaker of ice at all times.
3. Set the spectrophotometer to 600 nm, and adjust it to zero absorbance using the
decolorized blank. The lights in the laboratory will be switched off at this point to
conduct the experiment.
4. Working with one cuvette at time, pipette 1ml of the chloroplast suspension into a
foil-covered cuvette and place the cuvette inside an ice filled beaker positioned such
that light exposure of the cuvette is maximized.
NOTE: Dry and wipe the tubes clean before introducing them into the
spectrophotometer.
5. Remove the foil wrap from your cuvette and determine the absorbance at time zero
of exposure to light. Return this tube, uncovered, to the ice-water bath, and place it
in front of the light source, starting at the lowest intensity (5.0 moles m-2 sec-1).
6. Take absorbance readings on the same tube at two minutes intervals of light exposure
for eight minutes or until a decrease of > 0.5 absorbance reading. You can do this by
pushing and holding down the select button, which is located on top of the slide
projector.
7. Repeat the procedure at each light intensity. At light intensities of 100 moles m-2
sec-1 and higher, expose the cuvette at one minute intervals for a total of 4 minutes or
until a decrease of > 0.5 absorbance units has occurred. The treatment at zero light
intensity (wrapped in foil for the entire exposure duration) can be run parallel with
any of the other treatments and checked at two minute intervals. No change in
26
absorbance should occur in this cuvette. Note: the spectrophotometer should be
periodically re-zeroed with the blank.
8. Using a molar extinction coefficient of 0.014 M-1cm-1, calculate the rate of
reduction of DCPIP per chloroplast (expressed as moles DCPIP reduced min.-1
chloroplast-1) at each light intensity. Typically this preparation yields approximately
5 x 107 chloroplasts ml-1. Two electrons reduce one molecule of DCPIP. Plot
electrons transferred min.-1 chloroplast-1 vs. light intensity. Saturation of the
photosynthetic apparatus by photons of light should be apparent when you plot
electron transfer vs. light intensity.
Table 5.1
Light Intensity
(moles m-2sec.-1)
0
Absorbance at T = 0 Subsequent absorbance reading at 2 min. intervals
10
25
50
Subsequent absorbance readings at 1 min. intervals
100
150
200
250
27
Additional Notes:
Use Beer’s law to calculate conc. of DCPIP remaining at each time point
A= .c.l
c = A/  . l
Example: at A = 1.64, c = 1.64/(0.014M -1 cm-1) (1cm) = 117.1 M = 0.00117 M
Find multiple values of c to determine dc/dt, keep in mind that formation of reduced
DCPIP is indirectly measured as the loss of the oxidized blue form. Correct your
numbers for the volume assayed as well (6mL) in order to yield units of moles per
minute.
0.00117 x 6mL x 1L/1000mL = 0.00000702 moles
If dc/dt for reduction of DCPIP were to equal 0.00000702 moles/min,
Then to find # of e- transferred per minute, per chloroplast:
(0.00000702 moles/min) x (6.023 x 1023 molecules/mole) x (2 e-/molecule)
/ (5x107 chloroplasts) = 1.7 x 1011 e- transferred/chloroplast/min
28
RT-PCR ANALYSIS OF GENE EXPRESSION…….4
INTRODUCTION
In developmental biology, cellular differentiation is the process by which a less
specialized cell becomes a more specialized cell type. Differentiation occurs numerous
times during the development of a multi-cellular organism as the organism changes from
a single zygote to a complex system of organs and tissues. Differentiation is a common
process in adults as well: adult stem cells yield differentiated daughter cells during tissue
repair and during normal cell turnover. Differentiation can dramatically change cell size,
shape, metabolic activities, and responsiveness to other signals. These changes are largely
due to modifications in the pattern of gene expression within the differentiating cell. With
a few exceptions, cellular differentiation almost never involves a change in the genomic
DNA sequence itself. Thus, in spite of having identical genomic DNA sequences,
different cells within an individual can exhibit remarkably different physical properties.
Cell differentiation is the transition of a cell from one cell type to another and
involves a switch from one pattern of gene expression to another. In many cases, the
‘switch’ comes in the form of exposure to an extracellular signal molecule. Many of the
signal molecules that convey information from cell to cell during the control of cellular
differentiation are called growth factors. Although the details of specific signal
transduction pathways vary, these pathways often share the following general steps. A
ligand (signal) produced by one cell binds to the extracellular domain of a
transmembrane receptor embedded in the plasma membrane of another cell. Binding of
the ligand to the extracellular domain induces a conformational change in the cytoplasmic
domain of the receptor. The conformational change in the cytoplasmic region allows the
receptor to acquire an enzymatic activity. The active receptor then triggers a cascade of
phosphorylation reactions that eventually activates a dormant transcription factor. The
transcription factor then binds to the promoter regions of particular genes, activating or
repressing their expression. Thus, the transcription factor changes the pattern of gene
expression within the cell.
During skeletal myogenesis, cell cycle withdrawal accompanies the expression of
the contractile phenotype. The mouse C2C12 myoblast cell line can be triggered to
undergo differentiation into myotubes by withdrawal from growth factor-rich serum.
Withdrawal of serum facilitates cell cycle arrest and other changes in signal transduction
pathways within C2C12 cells. One of the early events triggered by these changes in
signal transduction is the transcriptional upregulation of Myogenin gene expression by
the transcription factors MyoD and Myf-5. Myogenin is a DNA binding transcriptional
regulator that, once expressed, activates expression of additional genes involved in
skeletal muscle development and function. The critical role played by Myogenin is
highlighted by the observation that knockout mice lacking the Myogenin gene form
myoblasts but fail to form fully differentiated skeletal muscle.
29
In this lab exercise, you will examine the change in gene expression for the
Myogenin gene in differentiated C2C12 myotubes (DCM sample) relative to its
expression level in undifferentiated C2C12 myoblasts (UCM sample). In order to do this,
you will first isolate RNA from one plate each of UCM and DCM cells. In the second
week, you will then use the isolated RNA as template in a Reverse TranscriptasePolymerase Chain Reaction (RT-PCR) assay. You will ‘program’ the assay with short
DNA primers specific to either the Myogenin gene or to a control gene Glyceraldehyde
3-Phosphate Dehydrogenase (GAPDH). Reverse Transcriptase is an RNA templatedependent DNA polymerase that will use the provided DNA primers to synthesize a
DNA strand complimentary to the RNA strand to which the DNA primer can bind.
Following the Reverse Transcriptase step, a thermostable DNA polymerase enzyme will
be used to perform traditional PCR amplification. After performing the RT-PCR steps, in
the third week you will then separate the reaction products on an agarose gel and examine
it for the potential presence of DNA bands representing either the Myogenin or GAPDH
RNA transcripts.
For additional information on the PCR technique, see Chapter 18.14 of the lecture
textbook (Karp, Cell and Molecular Biology, 5th ed).
30
RNA ISOLATION……..4A
1. Each student group will receive one plate of undifferentiated C2C12 myoblasts
(UDM) and one plate of differentiated C2C12 myotubes (DCM).
***NOTE: Students handling samples should wear gloves at all
times. Not only is this important for protecting you, but also for
protecting the samples from becoming contaminated by molecules
present on the surface of your skin.
2. Harvest total RNA from the C2C12 cells using the RNeasy Mini Kit and provided
protocol (summarized in the following steps).
3. Aspirate the growth media from both plates.
4. Gently rinse each plate with 3ml of 1xPBS and aspirate again.
5. Add 350uL of Buffer RLT to each plate and collect the resulting lysates with a
cell scraper. Transfer each lysate into separate labeled 1.5ml tubes.
***NOTE: Use a different cell scraper for each sample. Be very
careful not to contaminate one sample with the other sample.
6. Thoroughly homogenize the lysates by vortexing.
7. Next, pass the lysate 5 times through a 20 gauge syringe.
***NOTE: Use a different needle and syringe for each sample. Be
very careful not to contaminate one sample with the other sample.
8. Add 350uL of 70% EtOH to each sample.
9. Mix thoroughly but gently and then transfer up to 700uL of each sample into
separate labeled RNeasy spin columns mounted into labeled 2.0ml collection
tubes.
10. Centrifuge the spin columns mounted into the collection tubes at room
temperature for 15sec at 10,000 rpm.
11. Remove the RNeasy spin columns from the collection tubes and discard the flowthrough solution from the collection tubes. Return the RNeasy spin columns to
their original matching collection tube.
12. Add 700uL of Buffer RW1 to each RNeasy spin column. Repeat steps (10-11).
31
13. Add 500uL of Buffer RPE to each RNeasy spin column. Centrifuge for 2min at
10,000 rpm. Discard the flow-through.
14. Add 500uL of Buffer RPE to each RNeasy spin column. Centrifuge for 4min at
10,000 rpm. Discard the flow-through.
***NOTE: After centrifugation, carefully remove the RNeasy spin
column so that the column does not contact the flow-through.
15. Place each RNeasy spin column into a new labeled 1.5mL eppendorf tube.
16. Add 30uL of RNase-free water directly onto each of the spin column membranes.
Be careful not to damage the column material with the pipette tip.
17. Incubate at room temperature for 2 minutes. Centrifuge for 1min at 10,000 rpm to
elute total RNA from the spin column.
18. Using a fresh pipette tip, transfer 1 uL from each sample into separate new
labeled 1.5mL eppendorf tubes containing 499 uL sterile H2O. Mix well. These
are the 1:500 dilutions of your samples.
19. Measure the A260 and A260/A280 ratios for your 1:500 dilutions. An A260/A280
ratio of 1.8-2.0 indicates a high purity nucleic acid. An A260/280 ratio of 0.5-0.7
would be expected from a high purity protein sample.
20. Calculate the concentrations of your undiluted RNA samples using Beer’s law.
(for RNA,  = 25 ul/ug/cm)
21. Store the undiluted RNA samples in -80C
Questions:
How pure are your RNA samples?
Assuming cells grow to ~105 cells per square cm and contain ~10pg RNA per cell, what
is your theoretical yield of RNA from one of your samples? How does your actual yield
compare to the theoretical yield?
END OF TODAY’S ACTIVITY
32
REVERSE TRANSCRIPTASE – POLYMERASE CHAIN REACTION……..4B
Reverse Transcriptase (RT) is an RNA template-dependent DNA polymerase that
can be used to synthesize DNA strands that are complementary to a provided RNA
sample. DNA strands generated in this manner are often referred to as cDNA.
The basic equation for PCR states that the amount of amplified DNA after x
cycles (Nx) is the starting amount of the target DNA (N0) times the amplification
efficiency (E) to the power x. The PCR efficiency in this equation is a number between 1
and 2 (2 indicates 100% efficiency).
Basic equation for PCR:
Nx = N0 * Ex
If two samples contain different levels of a target cDNA (different N0 values),
this difference should theoretically be maintained throughout amplification. Thus, if
sample #1 contained ten times as much cDNA as sample #2 at the start of the reaction, it
should contain ten times as much cDNA at the end of the reaction as well. This principle
allows PCR, when coupled with reverse transcription, to be used as a method for
quantifying the relative levels of gene expression under various conditions.
RT-PCR method:
1. Prepare and label tubes for each sample in the table below:
Sample Name
MyoG
UCM
MyoG
DCM
GAPDH
UCM
GAPDH
DCM
Negative
control
2. Thaw your UCM and DCM RNA samples on ice.
3. A master mix stock containing most of the reagents for your reactions will be
provided to you in a single tube that must be kept on ice.
a. The master mix contains enough reagent for all 5 samples above plus an
additional sample in case any pipetting errors occur.
b. The table below shows the contents of the master mix.
Master mix (volume per 1 rxn):
for 6 reactions:
14.25ul RNase-free water
85.5ul
2.5 ul 10x RT-PCR Buffer
15.0ul
0.5 ul dNTP mix (40mM)
3.0ul
0.5 ul Reverse Transcriptase enzyme
3.0ul
0.25ul Thermostable DNA polymerase (Taq) 1.5ul
33
4. You will also be provided with the pre-mixed DNA oligonucleotide primer stocks
for Myogenin and GAPDH in separate tubes (1 tube each per group).
c. The Myogenin tube contains the following primers at 1.25uM each
i. Primer 1: 5’-GAGCGCGATCTCCGCTACAGAGG-3’
ii. Primer 2: 5’-CTGGCTTGTGGCAGCCCAGG-3’
iii. Below is the 380 nucleotides length region of the Myogenin
mRNA that will be amplified as cDNA by the Primer 1, 2 pair:
5’-GAGCGCGATCTCCGCTACAGAGGcgggggcgggccccagcccatggtgcccagtgaatgcaactc
ccacagcgcctcctgcagtccggagtggggcaatgcactggagttcggtcccaacccaggagatcatttgctcgcggctgacc
ctacagacgcccacaatctgcactcccttacgtccatcgtggacagcatcacggtggaggatatgtctgttgccttcccagacga
aaccatgcccaactgagattgtctgtcaggctgggtgtgcatgtgagcccccaagttggtgtcaaaagccatcacttctgtagcag
ggggcttttaagtggggctgtcctgatgtccagaaaacagcCCTGGGCTGCCACAAGCCAG-3’
d. The GAPDH tube contains the following primers at 1.25uM each
i. Primer 3: 5’-ACCCAGAAGACTGTGGATGG-3’
ii. Primer 4: 5’-CCCTGTTGCTGTAGCCGTAT-3’
iii. Below is the 421 nucleotides length region of the GAPDH mRNA
that will be amplified as cDNA by the Primer 3,4 pair:
5’-ACCCAGAAGACTGTGGATGGcccctctggaaagctgtggcgtgatggtcgtggggctgcccagaacat
catccctgcatccactggtgctgccaaggctgtgggcaaggtcatcccagagctgaacgggaagctcactggcatggccttcgtgttcctac
ccccaatgtgtccgtcgtggatctgacgtgccgcctggagaaacctgccaagtatgatgacatcaagaaggtggtgaagcaggcatctgag
ggcccactgaagggcatcttgggctacactgaggaccaggttgtctcctgcaacttcaacagcaactcccactcttccaccttcgatgccggg
gctggcattgctctcaatgacaactttgtcaagctcatttcctggtatgacaatgaATACGGCTACAGCAACAGGG-3’
5. Using the provided master mix and primer stocks, setup the following reactions
keeping all tubes on ice as much as possible:
Reagent / Samples
Master mix
MyoG
UCM
18 ul
MyoG
DCM
18 ul
GAPDH
UCM
18 ul
GAPDH
DCM
18 ul
Negative
control
18 ul
Myogenin primers
2 ul
2 ul
N/A
N/A
2 ul
GAPDH primers
N/A
N/A
2 ul
2 ul
2 ul
UCM RNA template
1 ul
N/A
1 ul
N/A
N/A
DCM RNA template
N/A
1 ul
N/A
1ul
N/A
water
4 ul
4 ul
4 ul
4 ul
1 ul
6. Return samples to TA to be placed in the thermocycler.
34
e. Samples will be incubated following the parameters below:
Number of Cycles
1
1
40
1
1
Temp *C
45
95
95
60
68
68
4
Time
15 min
1min
30sec
30sec
2min
5min
hold
Questions:
How many moles of DNA product would you have at the end of the reaction if you start
with ten molecules of target DNA and amplify for 40 cycles?
If our target DNA is a 500 base pair long molecule, how many micrograms of DNA
product would you get above? (the average molecular weight of a base pair is 660g/mol)
Which components of the PCR reaction limit the total amount of DNA product that can
possibly be obtained? What is the maximum yield, in moles, under the conditions
provided?
END OF TODAY’S ACTIVITY
35
AGAROSE GEL ELECTROPHORESIS……..4C
1. A 2% agarose gel containing Ethidium Bromide will be provided. Ethidium
Bromide is a mutagen and suspected carcinogen, wear gloves at all times.
2. Once the gel has solidified (this will take about 30 minutes cooling at room
temperature), remove the gel comb gently, place the gel into the gel running
apparatus and cover it with 1xTBE
3. Add 5ul of the 5x Sample Loading Dye to each of your RT-PCR samples.
4. Load 2ul of 100bp DNA ladder containing 0.5ug-1ug DNA as the molecular
weight (MW) markers into the left-most lane of the agarose gel.
5. Load 10ul of sample into the next adjacent lane. Repeat until all samples are
loaded into the gel similar to the sketch below:
1 = Myogenin UCM
2 = Myogenin DCM
3 = GAPDH UCM
4 = GAPDH DCM
5 = negative control
6. Run the gel at 80-90 volts for 30-45 minutes.
7. Turn off the power source and visualize the gel under UV light to detect RT-PCR
products. The 500bp MW marker is double intensity to aid ladder orientation.
Questions:
What are the expected lengths of the Myogenin and GAPDH PCR products? What are the
expected levels of gene expression for Myogenin and GAPDH?
If you substitute Primer 1 for the sequence 5’-CCACAGCGCCTCCTGCAGTC-3’, what
size PCR product would you expect to observe at the end of the experiment?
If you detected any PCR products with sizes other than the expected size, why might that
have happened? What variable(s) do you think could influence their presence or absence?
What consequence would contamination of the DCM RNA sample with the UCM RNA
sample have on your results? What about contamination of the UCM RNA sample with
the DCM RNA sample?
36