Graduate lab manual
a practical guide
September 2008
Christianna Choulaki
1
INDEX
Glossary .............................................................................................................................. 3
Genomic DNA isolation by disruption method ..................................................................... 4
Remarks .......................................................................................................................... 4
Methods ........................................................................................................................... 4
1) from peripheral blood leukocytes ............................................................................. 4
2) from mouse tails ....................................................................................................... 5
PCR ..................................................................................................................................... 7
Procedure ........................................................................................................................ 8
To minimize contamination risk .................................................................................... 9
Agarose gel analysis of PCR products ....................................................................... 10
Practical modifications to the PCR technique............................................................. 10
Endonuclease digestion .................................................................................................... 11
Fragment complementarity and splicing ..................................................................... 11
Restriction enzymes as tools...................................................................................... 11
General protocol ......................................................................................................... 12
hands-on ........................................................................................................................ 14
PBMC’s isolation with Ficoll (Histopaque-1077, Sigma) .................................................... 17
Cell counting in Neubauer chamber ............................................................................... 18
FACS ................................................................................................................................. 20
Principle ......................................................................................................................... 20
Flow cytometers............................................................................................................. 20
Fluorescent labels .......................................................................................................... 21
Measurable parameters ................................................................................................. 21
Applications ................................................................................................................... 21
hands on ........................................................................................................................... 22
Surface staining ............................................................................................................. 22
FACS ANALYSIS-ELITE CYTOMETER ........................................................................ 22
Total RNA isolation............................................................................................................ 25
A) Using Tri reagent (Sigma, cat# T 9424) .................................................................... 25
B) Using columns........................................................................................................... 27
Determine concentration ................................................................................................ 27
cDNA preparation (Reverse Transcription, RT) ................................................................. 28
Western Blot ..........................................................................................................................
Tissue preparation ......................................................................................................... 46
Gel electrophoresis ........................................................................................................ 46
Transfer ......................................................................................................................... 47
Blocking ......................................................................................................................... 47
Detection........................................................................................................................ 47
Analysis .............................................................................................................................
Colorimetric detection.....................................................................................................
Chemiluminescence .......................................................................................................
Secondary probing ......................................................................................................... 48
2
Glossary
APS
DTT
ddH2O
dNTPs
EtOH
FCS
Ficoll
gDNA
HBSS
KO
PBMCs
PBS
qPCR
RT
RT-PCR
SDS
TBE
TBS
TBS-T
TE
TEMED
Tris
TSE
UV
WT
W/V
Ammonium PerSulfate
Di ThioTheitol
double distilled Water
deoxyNucleoTides Phosphate
Ethanol
Fetal Bovine Serum
Histopaque-1077, Sigma
genomic DNA
Hank’s Balanced Salt Solution
Knock Out
Peripheral Blood Mononuclear Cells
Phosphate Buffer Saline
Quantitative PCR (ral time PCR)
Room Temperature
Reverse Transcription PCR
S Dodecyl Sulfate
Tris Borate EDTA buffer
Tris Buffer Saline
Tris Buffer Saline Tween
Tis EDTA
N,N,N,N’,N’-teteramethylethylenediamine
hydroxymethyl aminomethane
Tris Saline EDTA
ultraviolet
Wild Type
weight per volume
3
Genomic DNA isolation by disruption method
Remarks
Eukaryotic DNA is a double stranded, relatively stable molecule. However,
introduction of nucleases to DNA solutions should be avoided as these enzymes will
degrade DNA. Genomic DNA consists of very large DNA molecules, which are fragile
and can break easily. To ensure the integrity of gDNA, excessive and rough pipeting
and vortexing should be avoided. DNA is subject to acid hydrolysis when stored in
water, and should therefore be stored in TE buffer. Complete disruption and lysis of
cell walls and plasma membranes of cells and organelles is an absolute requirement
for all genomic DNA isolation procedures. Incomplete disruption results in
significantly reduced yields
Disruption method is based on the fact that the lysis buffer contains SDS, a detergent
that breaks down cellular membranes and proteinase K, a protease that digests
protein cellular components, resulting in the liberation of nucleic acids in the solution.
RNase is degrading RNA without interfering with DNA. Phenol/chloroform treatment
sequestrates proteins in the interphase, while chloroform absorbs phenol remainings
from the aqueous phase. Isopropanol adsorbs water molecules, resulting in a
“concentration” of DNA molecules witch facilitates their precipitation. The ionic
strength of the buffer is also very important as it neutralizes negative charges of the
DNA molecules. Ethanol 70% wash replaces isopropanol molecules (which can
inhibit subsequent enzymatic processes) with the more volatile Ethanol, rehydrates
DNA making the pellet easier to redissolve and removes co-precipitated salts.
Methods
1) from peripheral blood leukocytes
5ml peripheral blood in 50 ml tube containing 200μl EDTA0.5M (to prevent
coagulation)
spin at 4000 rpm for 15min at 4°C.
Remove serum and collect the white phase (leucocytes+reticulocytes)
cell disruption
+ an equal volume of TSE buffer
NaCl 150mM
EDTA pH8 100mM
Tris pH8 20 mM
+ SDS at a final concentration of 1%w/v
+ proteinase K at a final concentration of 10μg/ml
incubate 65°C 45-60 min
RNaseA treatment
+RNAseI at a final concentration of 100μg/ml
incubate at 37°C for 30min
DNA extraction
+ an equal volume of phenol:chloroform (1:1)
mix gently for 5 min
spin at 4000 rpm for 15 min at RT
4
carefully aspirate aqueous phase (containing nucleic acids) without taking any
proteins from the interphase and put in a new tube
repeat the phenol extraction till the interphase becomes clear
repeat the extraction with chlorophorm only (to absorb phenol remnants)
DNA precipitation
+ 1/10 volumes NaAcetate 3M, pH5.4 and mix
+ 2,2 volumes Ethanol -20°C
gently mix, watch the DNA filament formation
spin it around a flame-rounded Pasteur pipette
wash in 70% ethanol -20°C (desalting)
leave on air for some minutes to evaporate ethanol
dissolve DNA by immersion of the edge of the Pasteur pipette in a 1.7ml tube
containing 0.5 ml TE
Determine concentration by photometry:
For double stranded DNA, a solution containing 50μg/ml produce an OD260nm =1 (in
conventional photometers)
OD260nm =1 C=50μg/ml gDNA
hands on
(for a single beem photometer)
Dilution: add 5μλ DNA suspension in 0.995 ml dd H20 (dilution factor=200)
insert quartz cuvette in the photometer, fill it with ddH20
Set BLANK
remove ddH20 from the cuvette, fill it with the sample
note the OD260nm, and the OD280nm
the concentration of the sample is given by the formula
C= OD260nmx dilution factor x 50μg/ml
2) from mouse tails
Put a piece of 5 mm mouse tail in eppendorf tube
Add 700μl proteinase K buffer
Tris pH8 50 mM
EDTA pH8 100mM
NaCl 100mM
SDS 1%
+ proteinase K at a final concentration of 360 μg/ml, mix well
o/n 55°C
Add RNase I at a final concentration of 140 μg/ml
37°C 1-2hours
Add 700μl phenol/chloroform (1:1), shake 15 min (NO VORTEX)
Spin 15min, RT, 13.000rpm
Collect aqueous (upper) phase in a fresh tube
Add 700μl chloroform
Shake 15 min
Spin 15min, RT, 13.000rpm
Collect aqueous (upper) phase in a fresh tube
5
Add 700μl isopropanol
Gentle mix
If a visible pellet is formed, fish it out with a sealed Pasteur pipette
If pellet not visible, spin 15min, RT, 13.000rpm, decant supernatant.
Pellet might have a glassy appearance, care should be taken when removing the
supernatant as isopropanol precipitation pellets are more loosely attached to the side
of the tube.
Wash with 1 ml EtOH70%, brief vortex, spin 5min at 4°C, decant supernatant, absorb
residual liquid with a pipette tip
Take care not to dislodge DNA pellet
Air dry 10-15min (avoid over drying as this make DNA difficult to redisolve
Resuspend in appropriate V of H2O or TE
Let stand 15 min at RT, o/n at 4°C or, if difficult to ressuspend, o/n at RT or even at
55°C for 1-2 hours.
Verify that DNA is dissolved
Determine concentration by spectrophotometry and keep at -20°C
Determine concentration by photometry:
Add 5 μl DNA suspension in 995 μl H20 (dilution factor=200)
for OD260nm =1 C=50μg/ml gDNA
the concentration of the sample is given by the formula:
C= OD260nm * dilution factor * 50 μg/ml
6
PCR
PCR is used to amplify specific regions of a DNA strand. This can be a single gene,
just a part of a gene, or a non-coding sequence.
PCR, as currently practiced, requires several basic components:

DNA template that contains the region of the DNA fragment to be amplified

primers, which are complementary to the DNA regions at the 5' and 3' ends of
the DNA region that is to be amplified.

a DNA polymerase (e.g. Taq polymerase or another DNA polymerase with a
temperature optimum at around 70°C), used to synthesize a DNA copy of the
region to be amplified

Deoxynucleotide triphosphates, (dNTPs) from which the DNA polymerase
builds the new DNA

Buffer solution, which provides a suitable chemical environment for optimum
activity and stability of the DNA polymerase

Divalent cation. generally Mg2+ is used (Mn2+ can be utilized for PCR-mediated
DNA mutagenesis, as higher Mn2+ concentration increases the error rate
during DNA synthesis)

Monovalent cation (potassium ions)
For a typical 20μl PCR reaction
Template
Polymerase buffer
MgCl2
dNTPs
Primers
Polymerase
Genomic DNA ≈100ng
or
cloned DNA ≈50ng
or
cDNA ≈1-2μl of the reverse transcription reaction
1X
1,5-2,5 mM
200 μM
0,25-0,5 μM each
0,3-0.5 units
Always keep in mind that PCR is a very sensitive technique

The PCR is carried out in small reaction tubes (0.2-0.5 ml volume), containing a
reaction volume typically of 15-100 μl, that are inserted into a thermal cycler. This is
an instrument that heats and cools the reaction tubes within it to the precise
temperature required for each step of the reaction. Most thermal cyclers have heated
lids to prevent condensation on the inside of the reaction tube caps. Alternatively, a
layer of oil may be placed on the reaction mixture to prevent evaporation.
7
Procedure
The PCR usually consists of a series of 20 to 35 cycles. Most commonly, PCR is
carried out in three steps, often preceded by one temperature hold at the start and
followed by one hold at the end.
1. Prior to the first cycle, during an initialization step, the PCR reaction is often
heated to a temperature of 94-96°C, and this temperature is then held for 1-10
minutes. This first hold is employed to ensure that most of the DNA template
and primers are denatured, i.e., that the DNA is melted by disrupting the
hydrogen bonds between complementary bases of the DNA strands. (Also,
some PCRs require this step for activation of hot-start polymerase). Following
this hold, cycling begins, with one step at 94-98°C for 20-30 seconds
(denaturation step).
2. The denaturation is followed by the annealing step. In this step the reaction
temperature is lowered so that the primers can anneal to the single-stranded
8
DNA template. Brownian motion causes the primers to move around, and
DNA-DNA hydrogen bonds are constantly formed and broken between primer
and template. Stable bonds are only formed when the primer sequence
exactly matches the template sequence, and to this short section of doublestranded DNA the polymerase attaches and begins DNA synthesis. The
temperature at this step depends on the melting temperature of the primers,
and is usually between 50-64°C for 20-40 seconds.
3. The annealing step is followed by an extension/elongation step during which
the DNA polymerase synthesizes new DNA strands complementary to the
DNA template strands. The temperature at this step depends on the DNA
polymerase used. Taq polymerase has a temperature optimum of 70-74°C;
thus, in most cases a temperature of 72°C is used. The hydrogen bonds
between the extended primer and the DNA template are now strong enough to
withstand forces breaking these attractions at the higher temperature. Primers
that have annealed to DNA regions with mismatching bases dissociate from
the template and are not extended. The polymerase adds dNTP's that are
complementary to the template in 5' to 3' direction, thus reading the template
in 3' to 5' direction. The extension time depends both on the DNA polymerase
used and on the length of the DNA fragment to be amplified. As a rule-ofthumb, at its optimum temperature, the DNA polymerase will polymerize a
thousand bases in one minute. A final elongation step of 5-15 minutes
(depending on the length of the DNA template) after the last cycle may be
used to ensure that any remaining single-stranded DNA is fully extended. A
final hold of 4-15°C for an indefinite time may be employed for short-term
storage of the reaction, e.g., if reactions are run overnight.
To minimize contamination risk
 work in a template free environment
clean you bench periodically with 10% chlorine
keep your pre-PCR reagent tubes capped as long as possible
prepare reaction mixes and always add template last
To check whether the PCR generated the anticipated DNA fragment (also sometimes
referred to as amplimer), agarose gel electrophoresis is commonly employed for size
separation of the PCR products. The size(s) of PCR products is thereby determined
by comparison with a DNA ladder, which contains DNA fragments of known size, ran
on the gel alongside the PCR products.
the agarose gel is a matrix the density of which is determined by the concentration of
agarose. Single stranded DNA is a negatively charged molecule that migrates
through the gel towards the + pole of the field (cathode) at a rate inversely
proportional to its size. That means that larger molecules migrate more slowly.
Ethidium Bromide is a fluorescent dye that intercalates between the bases. DNA
bands are visualised upon excitation with UV light.
9
Agarose gel analysis of PCR products
Ethidium bromide-stained PCR products
after gel electrophoresis. Two sets of
primers were used to amplify the IGF gene
from 3 different DNA samples. In sample
#1 the gene was not amplified by PCR,
whereas bands for tissue #2 and #3
indicate successful amplification. A positive
control, and a DNA ladder containing DNA
fragments of defined length (last lane to
the right) to estimate fragment sizes in the
experimental PCRs, were also ran.
Practical modifications to the PCR technique

RT-PCR - RT-PCR (Reverse Transcription PCR) is a method used to amplify, isolate
or identify a known sequence from a cellular or tissue RNA. The PCR reaction is
preceded by a reaction using reverse transcriptase to convert RNA to cDNA. RT-PCR
is widely used in expression profiling, to determine the expression of a gene.

Quantitative PCR - Q-PCR (Quantitative PCR) is used to measure the quantity of a
PCR product (preferably real-time). It is the method of choice to quantitatively
measure starting amounts of DNA, cDNA or RNA. Q-PCR is commonly used to
determine whether a DNA sequence is present in a sample and the number of its
copies in the sample. The method with currently the highest level of accuracy is
Quantitative real-time PCR. It is often confusingly known as RT-PCR (Real Time
PCR) or RQ-PCR. QRT-PCR or RTQ-PCR are more appropriate contractions. RTPCR commonly refers to reverse transcription PCR (see above), which is often used in
conjunction with Q-PCR. QRT-PCR methods use fluorescent dyes, such as Sybr
Green, or fluorophore-containing DNA probes, such as TaqMan, to measure the
amount of amplified product in real time.

Hot-start PCR is a technique that reduces non-specific amplification during the initial
set up stages of the PCR. The technique may be performed manually by simply
heating the reaction components briefly at the melting temperature (e.g., 95˚C) before
adding the polymerase. Specialized enzyme systems have been developed that inhibit
the polymerase's activity at ambient temperature, either by the binding of an antibody
or by the presence of covalently bound inhibitors that only dissociate after a hightemperature activation step. Hot-start/cold-finish PCR is achieved with new hybrid
polymerases that are inactive at ambient temperature and are instantly activated at
elongation temperature.

Multiplex-PCR - The use of multiple, unique primer sets within a single PCR reaction
to produce amplicons of varying sizes specific to different DNA sequences. By
targeting multiple genes at once, additional information may be gained from a single
test run that otherwise would require several times the reagents and more time to
perform. Annealing temperatures for each of the primer sets must be optimized to
work correctly within a single reaction, and amplicon sizes, i.e., their base pair length,
should be different enough to form distinct bands when visualized by gel
electrophoresis.
10
Endonuclease digestion
A restriction enzyme (or restriction endonuclease) is an enzyme that cuts doublestranded DNA. The enzyme makes two incisions, one through each of the sugar-phosphate
backbones (i.e., each strand) of the double helix without damaging the nitrogenous bases.
The chemical bonds that the enzymes cleave can be reformed by other enzymes known as
ligases, so that restriction fragments carved from different chromosomes or genes can be
spliced together, provided their ends are complementary (more below). Many of the
procedures of molecular biology and genetic engineering rely on restriction enzymes. The
term restriction comes from the fact that these enzymes were discovered in E. coli strains
that appeared to be restricting the infection by certain bacteriophages. Restriction enzymes
therefore are believed to be a mechanism evolved by bacteria to resist viral attack and to
help in the removal of viral sequences. They are part of what is called the restriction
modification system.
Fragment complementarity and splicing
EcoRI cleavage produces "sticky" ends
SmaI restriction enzyme cleavage produces "blunt" ends
Because recognition sequences and cleavage sites differ between restriction enzymes, the
length and the exact sequence of a sticky-end "overhang", as well as whether it is the 5' end
or the 3' end strand that overhangs, depends on which enzyme produced it. Base-pairing
between overhangs with complementary sequences enables two fragments to be joined or
"spliced" by a DNA ligase. A sticky-end fragment can be ligated not only to the fragment from
which it was originally cleaved, but also to any other fragment with a compatible sticky end.
The sticky end is also called a cohesive end or complementry end in some reference. If a
restriction enzyme has a non-degenerate palindromic cleavage site, all ends that it produces
are compatible. Ends produced by different enzymes may also be compatible. Knowledge of
cleavage sites allows molecular biologists to anticipate which fragments can be joined in
which ways, and to choose enzymes appropriately.
Restriction enzymes as tools
Recognition sequences typically are only four to twelve nucleotides long. Because there are
only so many ways to arrange the four nucleotides --A,C,G and T-- into a four or eight or
11
twelve nucleotide sequence, recognition sequences tend to "crop up" by chance in any long
sequence. Furthermore, restriction enzymes specific to hundreds of distinct sequences have
been identified and synthesized for sale to laboratories. As a result, potential "restriction
sites" appear in almost any gene or chromosome. Meanwhile, the sequences of some
artificial plasmids include a "linker" that contains dozens of restriction enzyme recognition
sequences within a very short segment of DNA. So no matter the context in which a gene
naturally appears, there is probably a pair of restriction enzymes that can snip it out, and
which will produce ends that enable the gene to be spliced into a "plasmid" (i.e. which will
enable what molecular biologists call "cloning" of the gene).
Another use of restriction enzymes can be to find specific SNPs. If a restriction enzyme can
be found such that it cuts only one possible allele of a section of DNA (that is, the alternate
nucleotide of the SNP causes the restriction site to no longer exist within the section of DNA),
this restriction enzyme can be used to genotype the sample without completely sequencing it.
The sample is first run in a restriction digest to cut the DNA, and then gel electrophoresis is
performed on this digest. If the sample is homozygous for the common allele, the result will
be two bands of DNA, because the cut will have occurred at the restriction site. If the sample
is homozygous for the rarer allele, the sample will show only one band, because it will not
have been cut. If the sample is heterozygous at that SNP, there will be three bands of DNA.
Many recognition sequences are palindromic
While recognition sequences vary widely, many of them are palindromic; that is, the
sequence on one strand reads the same in the same direction on the complementary strand.
The meaning of "palindromic" in this context is different from what one might expect from its
linguistic usage: GTAATG is not a palindromic DNA sequence, but GTATAC is (GTATAC is
complementary to CATATG).
General protocol
1. Combine the following in a microcentrifuge tube:
a typical 20 μl reaction contains:
2 μl of appropriate 10X Restriction Enzyme Buffer
0.1 to 5 μg of DNA for digest
sterile ddH2O to a final volume of 19 μl
2. Add 1 to 2 μl (3 to 20 Units) Restriction Enzyme and mix gently. (keep in mind that enzyme
solutions contain glycerol which can inhibit digestion. The enzyme volume should not exceed
1:10 of the total reaction volume). Centrifuge using a microcentrifuge at low speed for a
couple of seconds.
3. Incubate at the appropriate temperature (usually 37°C) for 1 to 2 hours.
4. If the DNA is to be used for another application proceed to either
Heat inactivation of the Restriction Enzyme by heating (usually approximately 70°C)
for 15 min (continue with Step #5)
OR
Phenol-cloroform extraction:
b. Add an equal volume of Phenol:Chloroform and mix well by inversion.
c. Centrifuge in a microcentrifuge at maximum speed for 10 min to separate the
phases.
d. Save the upper phase (aqueous phase) and repeat the Phenol:Chloroform
extraction one more time (Step C).
e. transfer the aqueous phase in a new tube, add 5 M NaCl to a final concentration of
0.1 M NaCl and add an 1 volume of ice-cold 100% Ethanol.
f. Mix well by inversion, centrifuge in a microcentrifuge at maximum speed for 10 min
to pellet the DNA and discard the supernatant
g. Allow the DNA to air dry and dissolve the DNA in the appropriate buffer (or ddH 2O).
12
5. Run an aliquot on an agarose gel to check for digestion.
Applications
A. Tpl2 ko mouse genotyping
In order to create a transgenic mouse line that lacks the PD-1 (PD-1 ko) gene, a mouse ovule
has been injected with an engineered PD-1 gene in which the main part of the sequence has
been substituted by the Neomycine Resistance gene (Neo). The transgene has been
inserted in the mouse genome by homologous recombination.
transgene
Tpl2
genome
Neo
Tpl2
Tpl2
Neo
WT
KO
primers
primers
We then crossed the mouse carrying the transgene (called “founder”) with a WT mice, and
the F1 descendants have been crossed between them giving rise to the F2 mice. We aim to
investigate the genotype of the F2. To discriminate the homozygous Tpl2 ko mice we perform
a PCR with primers that specifically recognize the endogenous WT Tpl2 and give rise to a
PCR product of 188bp length or the Neo gene (PCR product 400bp). After gel analysis, we
expect to have homo- and heterozygous mice for both WT and ko.
B. Genetic screen of TRAF1
There is a lot of papers published that associates discrete point mutations (Single Nucleotide
Polymorfisms) with specific diseases. There is a variety of methods to genotype such SNPs
ie gene sequence, real time PCR or RFLPs (Restriction Fragment Length Polymorphism, the
later is by far the most popular till now.
In order to screen genomic DNA of individuals for the TRAF1 gene rs10818488 A1087G
polymorphism, a PCR with primers flanking the region of interest is performed followed by a
restriction enzyme-analysis.
The G allele creates an SduI site and, hence, the possibility to discriminate between the two
genotypes.
SduI recognition site
GGGCCC
CCCGGG
A
G
226
57
|
.
169
.
13
hands-on
Mouse Tpl2 genotyping
label PCR tubes 0.2 ml 1,2,3,4
1-NTC WT
2-WT
3- NTC KO
4-KO
Prepare the mixes:
mix WT
H2O
Taq pol buffer10X
Mg2Cl
betaine
dNTPs10mM
Primer Tpl2 for
Primer Tpl2 rev
Taq pol 5u/μl
Per
reaction
9
2.5
2.5
5
0.5
1
1
0.5
Per 2
reactions
18
5
5
10
1
2
2
1
Per
reaction
9
2.5
2.5
5
0.5
1
1
0.5
Per 2
reactions
18
5
5
10
1
2
2
1
mix KO
H2O
Taq pol buffer10X
Mg2Cl
betaine
dNTPs10mM
Primer Tpl2 for
Primer Tpl2 KO
Taq pol 5u/μl
dispense 22 μl of
add 3 μl
WT mix in tubes 1, 2
KO mix in tubees 3, 4
H2O in 1 and 3
mu tailDNA in 2 and 4
Hu TRAF1 SNP
label PCR tubes 0.2 ml 5,6
H2O
Taq pol buffer10X
Mg2Cl
dNTPs10mM
Primer for
SNP
18
2
3
1.5
1
X2
36
4
6
3
2
14
Primer rev
Taq pol 5u/μl
1
0.5
2
1
dispense 27 μl of mix SNP in tubes 5 an 6
add 3 μl
H2O in 5
hu gDNA in 6
spin briefly
insert in thermal cycler
lower heated lid
run programmes
PD-1
5' 94°C
(40" 94°C; 40'' 56°C; 40'' 72°C) 35 cycles
10' 72°C
∞4°C
TRAF-1
5' 94°C
(15" 94°C; 15'' 60°C; 30'' 72°C) 35 cycles
7' 72°C
∞4°C
TRAF-1 PCR product digest
label 2 tubes “6-“ and “6+”
in tube “-“ (undigested)
add
7.5 µl
PCR product
2.5 µl 6x gel loading buffer
keep at 4°C
in tube “+“ (digested)
add
10 µl PCR product
16 µl H2O
3,0 µl 10x Ppu21I (Fermentas)
1 µl SduI (10U/µl; Fermentas)
incubate for at least 2 hours at 37°C, then add 10μl 6x gel loading buffer.
Gel analysis
Prepare a 2% agarose gel
Weight 2.4 gr agarose
in a conical flask put 108 ml H2O
add agarose
swirl to mix
slowly melt by heating in a microwave and mix in-between
(be cautious, heat for short period at a time, don’t boil it, avoid proximity to
you face)
add 12 ml TBE 5X while swirling (to avoid local cooling of the agarose)
swirl to mix
+2 μl Ethidium bromide
swirl to mix
poor the melted agarose on a horizontally placed tray and insert the combs
let cool down to harden
place tray+gel in the electrophoresis apparatus
15
add 700 ml 0.5 X electrophoresis buffer (TBE)
remove combs carefully to preserve well integrity
loading
lane
tube
1
2
3
4
5
6
7
8
1
2
3
4
5
66+
Molecular
weight
marker


PCR
product
20 μl
20
20
20
10
7.5
30
Loading buffer
LB6x
6
6
6
6
3
2.5
10
run at 120V for about 1 hour.
Visualize the DNA fragments under UV-ligh.
16
PBMC’s isolation
Ficoll gradient (Histopaque-1077, Sigma)
Remarks : this method is based on the fact that ficoll has a grater density than PBMC’s but
lower than Red Blood Cells (RBCs). During centrifugation, RBCs sediment in the bottom of
the tube, PBMCs cannot pass through ficoll, so they accumulate in the interphase of
serum/PBS phase and ficoll phase.
Method
Collect blood in heparinized syringe (to prevent cagulation):
Load syringe (10ml) with ~200μl heparine (ex Leo 5.000iu/ml), make sure to imbibe the
walls, then poor off excessive heparine. For 10 ml blood, leave ~50μl in the syringe tip.
Change needle to proceed to the vain puncture.
Dilute heparinized peripheral blood with 1 volume PBS or HBSS
(usually 20 ml blood+20 ml PBS)
in 15 ml tubes, aliquot 5 ml ficoll and carefully (inclined position) overlay 10 ml diluted blood
centrifuge for 30 min, at 1,800 rpm, 22°C NO BRAKE
carefully aspirate the white interphase (PBMCs) with a Pasteur pipette without taking any
ficoll
washes:
qsp with PBS or HBSS at 50ml, centrifuge 10min, at 1200-1700 rpm
decant and resuspend pellet in 1 ml PBS or HBSS by gentle pipetting, qsp with PBS or
HBSS at 25 ml, centrifuge 10min, at 1200-1700 rpm
decant and resuspend pellet in 1 ml RPMI/10% FCS by repeated gentle pipetting,, add 4 to 9
ml RPMI/10% FCS depending on pellet size
check for aggregates. If needed, use a 5 ml or 10 ml syringe (G>21) to homogenize (no more
than 4 passes, to preserve cell integrity)
17
Cell counting in Neubauer chamber
Place slide cover on the Neubauer chamber
mix by pipetting 1V cell suspension+1V Trypan blue vital dye (usually 15 μl+15 μl)
Carefully fill in the chamber. (Take care to avoid under- or over-filling).
Count live cells in the entire 25 large squares area. For cells overlapping the border line,
count only cells in the upper and left side border lines. Dead cells appear light blue, live cells
are colorless.
Cell concentration: number of cells in the 25 large squares area x dilution factor( =2) x 10 4
cells/ml
C = n x 2*104 cells/ml
aliquot cells:
PBMCs
RIA tubes 1, 2, 3, 4
FACS
300-500*103 cells/tube
tube 4
RNA
3*106 cells
add RPMI/FCS to 1.5ml
tube 5
cell extracts
3*106 cells
PMA/ionomycine stimulation of PBMCs
PMA (Phorbol Myristate Acetate) is a potent pan-leucocyte stimulator that induces
downstream signaling pathways leading to cell activation.
in tube 4: Add appropriate volume of RPMI/FCS to obtain a concentration of 2*10 6 cells/ml in
a total volume of 1.5 ml
PMA/ionomycine stocks are concentrated. In order to deal with accurate volumes we need to
predilute the stock solutions.
working dilutions
PMA1
Iono1
stock
PMA (100μg/μl)
Iono (500μg/ml)
1μl
4
PBS
99μl
36
Dilution factor
100
10
C
1μg/μl
50μg/ml
18
In tube 4 add
- 37.5 μl PMA11mg/ml to obtain a final concentration of 25ng/ml
- 30 μl ionomycine1 50μg/ml to obtain a final concentration of 1μg/ml
incubate cell suspension at 37°C , 5%CO2 (incubator) for 2 hours
proceed to total RNA isolation (see below).
Cell extract preparation
PBMC tube 5:
centrifuge at 1.200 rpm 5-10min. Decant supernatant
wash with 2 ml cold PBS
spin 10.000 1 min Decant supernatant completely
dislock pellet
add 130μl cell extract lysis buffer
vortex 20sec
put on ice for 15 min
centrifuge 10 min at 10.000 rpm at 4°C
keep supernatant in a fresh tube , store at -80°C
determine concentration by Bradford method
FACS Staining
see below
19
FACS
Flow cytometry is a technique for counting, examining and sorting microscopic particles
suspended in a stream of fluid. It allows simultaneous multiparametric analysis of the
physical and/or chemical characteristics of single cells flowing through an optical and/or
electronic detection apparatus.
Principle
A beam of light (usually laser light) of a single wavelength is directed onto a hydrodynamically focused stream of fluid. A number of detectors are aimed at the point where the
stream passes through the light beam; one in line with the light beam (Forward Scatter or
FSC) and several perpendicular to it (Side Scatter (SSC) and one or more fluorescent
detectors). Each suspended particle passing through the beam scatters the light in some
way, and fluorescent chemicals found in the particle or attached to the particle may be
excited into emitting light at a lower frequency than the light source. This combination of
scattered and fluorescent light is picked up by the detectors, and by analysing fluctuations in
brightness at each detector (one for each fluorescent emission peak) it is then possible to
extrapolate various types of information about the physical and chemical structure of each
individual particle.
FSC correlates with the cell volume and SSC depends on the inner complexity of the particle
(i.e. shape of the nucleus, the amount and type of cytoplasmic granules or the membrane
roughness).
Some flow cytometers on the market have eliminated the need for fluorescence and use only
light scatter for measurement. Other flow cytometers form images of each cell's fluorescence,
scattered light, and transmitted light.
Flow cytometers
Modern flow cytometers are able to analyse several thousand particles every second, in "real
time", and can actively separate and isolate particles having specified properties. A flow
cytometer is similar to a microscope, except that instead of producing an image of the cell,
flow cytometry offers "high-throughput" (for a large number of cells) automated quantification
of set parameters. To analyze solid tissues single-cell suspension must first be prepared.
A flow cytometer has 5 main components:

a flow cell - liquid stream (sheath fluid) carries and aligns the cells so that they pass
single file through the light beam for sensing.

a light source - commonly used are lamps (mercury, xenon); high power water-cooled
lasers (argon, krypton, dye laser); low power air-cooled lasers (argon (488nm), redHeNe (633nm), green-HeNe, HeCd (UV)); diode lasers (blue, green, red, violet).

a detector and Analogue to Digital Conversion (ADC) system - generating FSC and
SSC as well as fluorescence signals.

an amplification system - linear or logarithmic.

a computer for analysis of the signals.
The data generated by flow-cytometers can be plotted in a single dimension, to produce a
histogram, or in two dimensional dot plots (or even in three dimension). The regions on these
plots can be sequentially separated, based on fluorescence intensity, by creating a series of
subset extractions, termed "gates". The plots are often made on logarithmic scales. Because
20
different fluorescent dyes' emission spectra overlap, signals at the detectors have to be
compensated electronically as well as computationally.
Fluorescent labels
The fluorescence labels that can be used, will depend on the lamp or laser used to excite the
fluorochromes and on the detectors available
blue argon laser (488 nm)
This is an air cooled laser and therefore cheaper to set up and run. It is the most commonly
available laser on single laser machines.
The most common fluorescence labels are

Green (usually labelled FL1): FITC, GFP, CFSE, CFDA-SE

Orange (usually FL2): PE

Red channel (FL3): PerCP, PE-Cy5, PE-Cy5.5, PI

Infra-red (FL4): PE-Cy7
Measurable parameters
This list is very long and constantly expanding.
Most importantly

cell surface antigens (Cluster of differentiation (CD) markers)

intracellular antigens (various cytokines, secondary mediators etc.)

DNA (cell cycle analysis, cell kinetics, proliferation etc.)

apoptosis (quantification, measurement of DNA degradation, mitochondrial membrane
potential, permeability changes, caspase activity)

cell viability

volume and morphological complexity of cells
also
Applications
The technology has applications in a number of fields, including molecular biology, pathology,
immunology, plant biology and marine biology. In the field of molecular biology it is especially
useful when used with fluorescence tagged antibodies. These specific antibodies bind to
antigens on the target cells and help to give information on specific characteristics of the cells
being studied in the cytometer. It has broad application in medicine (especially in
transplantation, hematology, tumor immunology and chemotherapy, genetics and sperm
sorting in IVF).
NB: For more informations refer to the relevant course
21
hands on
Surface staining
After PBMC isolation and cell counting, use 300.000-1.000.000 cells / staining (tube)
Label RIA tubes (1,2,3), add cell suspension,
+2 ml PBS/FCS2.5%
centrifuge at 1.200 rpm 5-10min. Decant supernatant, leave approx 100μl
Add appropriate volume of labeled Ab depending on cell number (see manufacturer
instructions)
panel
add 20 μl of the indicated Ab
Tube#
IgG PE
IgG FITC
IgG PC5
1
2
3
4
+
+
+
CD3 PE
CD4 FITC
CD8 PC5
+
+
+
CD19FITC
CD16PE
+
+
Mix by tapping, incubate 15 min at RT or 20 min at 4°C
Wash two times by adding 2.5 ml PBS/FCS2.5% and centrifuge 1.200 rpm 5-10min. Decant
supernatant
Add 0.1 ml PBS/PFA2%, keep at 4°C until analysis (max 2 days)
FACS ANALYSIS-ELITE CYTOMETER
1) Open side door, check
isoton II (upper bottle), if needed, add isoton II
(don’t remove bottle)
Waste bottle, if needed, throw away, add 3-4 cm
Chlorine
waste out of order, put a plastic bottle in the upper corner
door
2) MAIN POWER ON
AUXILIARRY POWER ON (for PC)
Right screen:
control screen,
argon laser ON (be careful for the finger inclination),
wait 20min to warm up
4) main screen
graph screen
4 graphs FS/SS PMT1 (Forward/ Side Scatter)
PMT2
PMT3
PMT4
for 3 colors
For all the graphs, make Y axe to have size of cells (FS) (P1)
5)PC
START, disconnect, restart in DOS
I read C:\WINDOWS>_
22
write C:\ELITE\CYTOMETER, 
6) acquisition protocol
file open
select: alignement check 100
&
desired protocol(s)
OK
To send settings to FACS:
F9 or Aquire
Start
F10 to move mouse
Screen protocol
Click on function :cytosettings received
On FACS: seath run
Beads in the frig door, or 2-3 drops in a RIA tube +WFI,
Vortex, put beads on FACS, use button no5 (near laser, side door) to have cells as “spot”
When 2000 events, it stops
Data rate 125, if >125, diminish pressure (sample flow)
If “horseshoe”-like image, there are debris in the channels, more wash
If OK, press F9 (start counting cells)
Screen analysis
Check half CV
It must be <2,5%
Aquire
new protocol twice to bring the desired protocol
“sample name” “, on screen, click to avoid
F9 (or aquire, start) to send protocol
F10 for mouse
Screen protocol
Cytosettings received
list mode save
screen
sample info
click on file name, always start with BUB
ex BUB120704
list mode: .LMD make .L1
sample name ex papadakis PB control fitc/pe
sample no :#1
start measures with control
SEATH RUN, vortex tube, put on FACS
F9
Modify voltage to have all cells in the scatter
F10 to stop
Surround cell population from EDIT and mouse left button
Mouse right button to 
replay start
modify length of cursor (can go in background  2% in theory, 1% in practice
note: some Ab like CD25 at 2%
23
Adjust PE and E
create  Quadr
For next sample press
F12
Sample name etc
F9
F10
Etc
WASHES
Blue detergent in RIA tube
VACUUM
control screen
Argon lase OFF
Valves screen
3rd:OFF
4rth ON
5th ON
absorb
2 tubes detergent
1 tube WFI
control screen
shut down valves
internal wash and message :valves action have completed
application
exit or WIN
windows screen on PC, start, shut down
on FACS
auxiliary power OFF
main power OFF
24
Total RNA isolation
Remarks: RNA is single stranded and hence very sensitive to hydrolysis. Ribonucleases
are extremely difficult to inactivate. Care should be taken to avoid inadvertently introducing
RNase activity into your RNA during or after the isolation procedure. This is especially
important if the starting material has been difficult to obtain or is irreplaceable.
Two of the most common sources of RNase contamination are the user’s hands and bacteria
that may be present on airborne dust particles. To prevent contamination from these sources,
use sterile technique. Gloves should be worn at all times.
Whenever possible, sterile disposable plastic ware should be used for handling RNA.
Treat nondisposable glassware and plasticware before use to ensure that it is RNase-free.
Bake glassware at 200°C o/n, and thoroughly rinse plasticware with 0.1N NaOH, 1mMEDTA.
Rinse with RNase free water
Treat solutions with DEPC water.1% o/n and then autoclave for 30min to remove any trace of
DEPC.
Make sure that the samples are always kept on ice after final dilution. Periodically
decontaminate the bench with RNase away or other commercially available nucleaseinhibiting agents or chlorine 10%
The successful isolation of RNA requires 4 essential steps:
1) effective disruption of cells or tissue,
2) denaturation of nucleoprotein complexes,
3) inactivation of endogenous ribonuclease activity and
4) removal of contaminating DNA and proteins.
The most important step is the immediate inactivation of endogenous RNases that are
released from membrane-bound organelles upon cell disruption.
Most commercial preparations for RNA isolation contain guanidine thiocyanate (GTC) to
inactivate the ribonucleases present in cell extracts. GTC, in association with SDS
(detergent), acts to disrupt nucleoprotein complexes, allowing the RNA to be released into
solution and isolated free of protein.
In the case of chemical purification (TriReagent, Trizol etc) proteins are sequestrated by
organic solvents (phenol) and the RNA-containing aqueous phase is subjected to chloroform
treatment in order to eliminate phenol contamination which interferes with subsequent
enzymatic procedures. RNA is then precipitated with ethanol or isopropanol.
In silica-column isolation methods, dilution of cell extracts in the presence of high
concentrations of GTC causes selective precipitation of cellular proteins to occur, while the
RNA remains in solution. After centrifugation to clear the lysate of precipitated proteins and
cellular debris, the RNA is selectively precipitated out of solution with ethanol and bound
electrostatically to the silica surface or the glass fibers of the column. DNA contaminants are
digested by DNaseI treatment. The bound RNA is purified from contaminating salts, proteins
and cellular impurities by washing. Finally, the RNA is eluted from the membrane by addition
of nuclease-free water
A) Chemical purification using Tri reagent (Sigma, cat# T 9424)
I. Sample Preparation
Suspension cells:
1 - centrifuge PBMC tube 4 at 1.700rpm for 10 min
decant supernatant
wash with 1 ml PBS
centrifuge 10.000 rpm 1min
dissociate pellet by tapping
25
2 - add 1 ml of TRI REAGENT, mix by repeated pipeting. (1 ml of the reagent is sufficient to
lyse 5-10 x 106 animal, plant or yeast cells or 107 bacterial cells).
Notes:
a. Some yeast and bacterial cells may require a homogenizer.
b. After the cells have been homogenized or lysed in TRI REAGENT, samples can be
stored at -70 °C for several months.
c. If samples are to be used for PCR or have a high content of fat, protein,
polysaccharides or extracellular material such as muscle, fat tissue and tuberous parts
of plants an additional step may be needed.
3 - After homogenization, centrifuge the homogenate at 12,000 x g for 10 minutes at 4
°C to remove the insoluble material (extracellular membranes, polysaccharides, and
high molecular weight DNA). The supernatant contains RNA and protein. (If the sample
had a high fat content there will be a layer of fatty material on the surface of the aqueous
phase that should be removed.)
4- Transfer the clear supernatant to a fresh tube.
let stand for 5 minutes at room temperature (To ensure complete dissociation of
nucleoprotein complexes)
5 - Add 0.2 ml of chloroform per ml of TRI REAGENT used.
Cover the sample tightly, shake vigorously for 15 seconds and allow to stand for 2-15
minutes at room temperature.
6 - Centrifuge the resulting mixture at 12,000 x g for 15 minutes at 4 °C. Centrifugation
separates the mixture into 3 phases: a red organic phase (containing protein), an interphase
(containing DNA), and a colourless upper aqueous phase (containing RNA). Note: The
chloroform used for phase separation should not contain isoamyl alcohol or other additives.
II. RNA Isolation
7 - Transfer the aqueous phase to a fresh tube
add 0.5 ml of isopropanol per ml of TRI REAGENT used in Sample Preparation (Step 1) and
mix.
let stand for 5-10 minutes at room temperature.
8 - Centrifuge at 12,000 g for 10 minutes at 4 °C. The RNA precipitate will form a pellet on
the side and bottom of the tube.
For use in PCR:. After transfer of aqueous phase to a fresh tube (Step 1 of RNA Isolation),
mix aqueous phase with 1/10 volume of isopropanol, store samples at room temperature for
5 minutes, and centrifuge at 12,000 x g for 10 minutes at 4 °C.
Transfer supernatant to a fresh tube and precipitate RNA by adding remaining amount of
isopropanol specified in Step 1 of RNA Isolation.
9 - Remove the supernatant and wash the RNA pellet by adding 1 ml (minimum) of 75%
ethanol per 1 ml of TRI REAGENT used in Sample Preparation, Step 1.
Vortex the sample and then centrifuge at 7,500 x g for 5 minutes at 4 °C.
Notes:
a. If the RNA pellets float, perform the wash in 75% ethanol at 12,000 x g.
b. Samples can be stored in ethanol at 4 °C for at least 1 week and up to 1 year at –20 °C.
10- Briefly dry the RNA pellet for 5-10 minutes by air drying or under a vacuum.
26
Do not let the RNA pellet dry completely, as this will greatly decrease its solubility. Do
not dry the RNA pellet by centrifugation under vacuum (Speed- Vac).
Add an appropriate volume of formamide, water or a 0.5% SDS solution to the RNA pellet. To
facilitate dissolution, mix by repeated pipetting with a micropipette at 55-60 °C for 10-15
minutes.
For long term storage, keep RNA at -80ºC
Notes:
a. Final preparation of RNA is free of DNA and proteins. It should have a 260/280 ratio of
≥1.7.
b. Typical yields from tissues (μg RNA/mg tissue): liver, spleen, 6-10 μg; kidney, 3-4 μg;
skeletal muscle, brain, 1-1.5 μg; placenta, 1-4 μg.
c. Typical yields from cultured cells
(μg RNA/106 cells): epithelial cells, 8-15 μg; fibroblasts, 5-7 μg.
Results
Troubleshooting Guide
I. RNA Isolation:
A. Low yield may be due to:
incomplete homogenization or lysis of samples.
the final RNA pellet may not have been completely dissolved.
B. If the A260/A280 ratio is <1.65:
-the amount of sample used for homogenization may have been too small.
-samples may not have been allowed to stand at room temperature for 5 minutes after
homogenization
-there may have been contamination of the aqueous phase with the phenol phase.
-the final RNA pellet may not have been completely dissolved.
If there is degradation of the RNA:
-the tissues may not have been immediately processed or frozen after removing from
the animal.
-the samples used for isolation or the isolated RNA preparations may have been stored
at –20 °C instead of –70 °C as specified in the procedure.
-cells may have been dispersed by trypsin digestion.
-aqueous solutions or tubes used for procedure may not have been RNAse-free.
-formaldehyde used for the agarose gel electrophoresis may have had a pH value
below 3.5.
If there is DNA contamination:
-the volume of reagent used for the sample homogenization may have been too small.
-samples used for the isolation may have contained organic solvents (ethanol,
DMSO), strong buffers or alkaline solution.
B) Using columns
Follow manufacturer instructions
Determine concentration
5μλ sample + 995μl ddH2O (dilution factor = 200), mix
count absorbance at 260/280nm, ratio should be ≥1,7, ideally 2
concentration (μg/ml) = OD260nm x dilution factor x 40
27
Expression analysis by Real-time PCR
cDNA preparation (Reverse Transcription, RT)
Remarks: reverse polymerase use mRNA as template to incorporate dNTPs resulting in the
formation of a single strand complementary DNA molecule (this single strand molecule will
serve as template during the first cycle of subsequent PCR, resulting in double stranded
cDNA).
Three types of primers are used in RT:
-random hexamers: mix of synthetic hexameric nucleotides with random sequences
that theoretically binds any possible sequence on RNA.
-oligodT primers: synthetic polyT nucleotides complementary to the polyA tail of
mRNAs, avoiding amplification of tRNA, rRNAs and RNA-constituents of ribonucleoproteins.
-specific primers: designed to be complementary with the gene of interest
Using thermoscript kit (invitrogen, cat#11146-016)
see also product data sheet
≤5μg total RNA in 9 μl RNAse free water
+primers 1μl
+dNTPs 2 μl
incubate for 5 min at 65°C (to denature secondary structures)
place on ice immediately (to avoid renaturation)
per reaction add
RT buffer5X 4 μl
DTT
1 μl
RNaseOUT 1 μl
H2O
1 μl
Reverse transcriptase 1 μl
Incubate 10 min @ 25°C
50
50°C
5
85°C
add Rnase H 1 μl
incubate
20min
37°C
store at -20°C
Real time PCR
Overview
Fluorescent-Based Chemistries
A. TaqMan probes (5’ nuclease assay)
This chemistry exploits the 5’ nuclease activity of AmpliTaq Gold DNA pol to cleave a
TaqMan probe during PCR.The TaqMan probe contains a reporter dye at the 5’ end and a
quencher dye at the 3’ of the probe. The latter suppresses the fluorescence emitted by
the reporter as long as they stay at proximity. During the reaction, cleavage of the probe
separates the reporter dye and the quencher dye which results in increased fluorescence
of the reporter. Accumulation of PCR products is detected directly by monitoring the
increase in fluorescence of the reporter dye.
The increase in fluorescence signal is detected only if the target séquence is
complementary to the probe an dis amplified during PCR, hence any nonspecific
amplification is not detected. This method is higly specific but yields signals with lower
intensity, requires optimisation and is not cost effective.
28
A. TaqMan probes
B. SYBR Green I
The SYBR Green I double-stranded Binding dye is used for the fluorescent detection of
double-stranded DNA ((dsDNA) generated during PCR. It binds non-specifically only to
dsDNA
29
During the exponentional growth phase the relaship of amplified PCR product to initial
template can be described by the equation:
Nn=N0 (1+E)n
Nn : quantity at cycle n
E : PCR efficiency 0  E 1
Methods of Quantification:
There are two basic quantification methods, and each is suitable for different applications:
absolute quantification and relative quantification.
Absolute quantification: The most direct and precise approach for analyzing quantitative data
is to use a standard curve that is prepared from a dilution series of control template of known
concentration. This is known as “standard curve” or “absolute” quantification. The absolute
quantification approach is used when it is important to the experimental design and objective
of the project to measure the exact level of template in the samples (e.g. monitoring the viral
load in a sample). A variety of sources can be used as standard templates. Examples include
a plasmid containing a cloned gene of interest (GOI or target), genomic DNA, cDNA,
synthetic oligos, in vitro transcripts, or commercially available total RNA.
Figure 8 describes a basic setup for standard curve quantification. Keep in mind that
selection of template is dependent upon the application being pursued. The most critical
consideration is that the primer set be optimized to work efficiently with the standards and the
experimental source material or tissue. Following amplification of the standard dilution series,
the standard curve is generated by plotting the log of the initial template copy number against
the Ct generated for each dilution. If the aliquoting was accurate and the efficiency of the
amplification does not change over the range of template concentrations being used, the plot
of these points should generate a straight line. This line is the standard curve. Comparing the
Ct values of the unknown samples to this standard curve allows the quantification of initial
copy numbers
Experimental setup for standard curve quantification.
Using a known starting concentration of template from one of a variety of sources, a dilution
series is performed. These samples are run under the standard well type on the same plate
as your unknowns. By comparing the Ct values of the unknowns to the Ct values of the
standards, the starting template quantities for the unknown samples can be calculated.
30
Figure 1 Ideally, a standard curve
will consist of at least 4 points, and
each concentration should be run at
least in duplicate (the more points
the
better).
The
range
of
concentrations in the standard curve
must cover the entire range of
concentrations that will be measured
in the assay (this may be several
orders of magnitude). Conclusions
cannot be drawn from samples
whose calculated initial quantity
exceeds the range of the curve.
In addition, the curve must be linear over the whole concentration range. The linearity is
denoted by the R squared (Rsq) value (R2 or Pearson Correlation Coefficient) and should be
very close to 1 (> 0.985). A linear standard curve also implies that the efficiency of
amplification is consistent at varying template concentrations. If the standard curve becomes
non-linear at very low template concentration, it is probably approaching the limit of detection
for that assay. Unknown samples that have Ct values that fall within a non-linear section of
the standard curve cannot be accurately quantified. Ideally, the efficiency of both the
standard curve and sample reactions should be between 90 and 110%. One hundred percent
efficiency implies perfect doubling of amplicon each cycle. If the efficiency is significantly
less, this implies the reaction is being slowed in some way, either from inhibitors present in
the reaction mix or suboptimal primer sets or reaction conditions. Efficiencies significantly
above 100% typically indicate experimenter error (e.g. miscalibrated pipettors, PCR
inhibitors, probe degradation, formation of non-specific products, and formation of primer
dimers). Primer dimer formation is typically of greatest concern with SYBR Green I assays
where any double-stranded product will be detected. Deviations in efficiency can also be due
to poor serial dilution preparation as well as extreme ranges of concentrations that either
inhibit PCR (high template amounts) or exceed the sensitivity of that particular assay (very
low amounts). The most important aspect is to have the efficiencies of standards and targets
within about 5% of each other if possible, with both near 100%. Once the reactions for the
standard curve and the samples have been optimized, Ct values can be compared to each
other and an initial template quantity can be estimated. It is important to remember that for
this type of quantification a standard curve must be run on the same plate as the unknown
samples. Replicates can vary in Ct when run at different times or on different plates, and thus
are not directly comparable to other runs. Also keep in mind that the “absolute” quantity
obtained from the standard curve is only as good as the DNA/RNA quantification methods
used to measure the standards, so you must take care to use very clean template and to
perform replicate measurements (whether using UV spectrophotometry or nucleic acid
binding dyes). There should also be at least 2–3 no template control (NTC) wells and for
QRT-PCR runs at least 2–3 no reverse transcriptase control wells. A more detailed
description of standard curve analysis will follow in the Data Analysis section.
Relative Quantification: Although standard curve (or Absolute) quantification can be useful in
determining absolute quantities of target, the majority of scientific questions regarding gene
expression can be accurately and reproducibly answered by measuring the relative
concentration of the gene of interest (target) in unknown samples compared to a calibrator, or
31
control sample. Here, the calibrator is a baseline for the expression of a given target gene.
This can be a zero time point in a time-course experiment or an untreated sample that will
serve as a benchmark to which the other samples can be compared. Using this approach,
differences in Ct value between an unknown sample and calibrator are expressed as foldchanges (i.e. up or down regulated) relative to the calibrator sample. In addition to comparing
the expression of the target gene alone in a control versus experimental sample, it is always
a good idea to normalize the results with a normalizing reference, typically a gene whose
expression is constant in both the control (calibrator) and experimental samples. This
normalization controls for differences in RNA isolation and in the efficiency of the reverse
transcription reaction arising from sample to sample and experiment to experiment.
Normalizers are explained in more detail in the following section. When designing a
comparative quantification experiment, it is not necessary to run a standard curve on every
plate as you would for absolute quantification. Rather the results are expressed as the fold
difference between the target and normalizer in experimental versus calibrator samples.
However, it is usually not accurate to assume that the amplification efficiency in any reaction
is going to be 100%, or that the same concentrations of template molecules will be detected
at a given Ct value each time the assay is run. Actual amplification efficiency values for a
particular reaction can be established via a standard curve measurement during assay
design, and multiple standard curves should be run to verify that this efficiency measurement
is reproducible (typical run-to-run variability is in the 5% range).
Controls
One of the most important considerations in a QPCR experiment is appropriate controls. The
specific controls that are needed will vary somewhat according to the experiment type, but
there are certain controls, such as No Template Controls (NTC), that should be included in
every run.
In QRT-PCR experiments, especially those based on comparative quantification, it is
important to include a reference gene (also called a normalizer gene). In order to generate
meaningful data that can be compared from run to run, sample to sample, and lab to lab, it is
essential to quantify the reference side by side with the Gene of Interest (target gene). The
reference gene is typically a “housekeeping” gene (HKG) whose expression should be
constant under the experimental conditions of the assay. This constant level of expression
must be verified experimentally, as the expression of housekeeping genes can vary under
certain conditions. The most common housekeeping genes used are GAPDH and β-actin,
which are ubiquitously expressed, but there is evidence that their level of expression can vary
considerably (Radonic, Thulke et al. 2004) (Bustin 2002). Alternative references like 18S or
28S rRNA have also been shown to be up and down regulated under different conditions
(Radonic, Thulke et al. 2004) and may not be applicable when poly A(+) RNA is used as the
template source. When working with a whole animal, it may be useful to normalize to total
cell number as well. In any case, it is crucial to select a reference or even multiple references
that have been empirically tested to be consistent across all experimental conditions in your
assay. You can find initial data in the literature or from microarray data (genomewww5.stanford.edu/).
Because the expression level of the reference is constant, any variation in the Ct of the
reference can be attributed to other sources of variation, such as efficiency of the reverse
transcription reaction, yield of the RNA purification, or variations in the number of cells from
which the RNA was isolated. These sources of variation will affect the Reference and the
Target genes equally, so differences in the Ct of the reference from sample to sample can be
used to correct for any variation in the Ct of the target that is not due to changes in
expression level. The most essential characteristic for successful reference genes is that they
are not affected (induced or suppressed) by the changing experimental conditions. It is also
important to choose a reference that has an expression level and an amplification efficiency
that is similar to that of the target. During assay design, it should also be confirmed that these
32
amplification efficiency values are reproducible. If they are not, the normalization results
cannot be considered reliable.
Any of the references above (housekeeping genes or rRNA) are also known as endogenous
references because they are part of the RNA pool. Because it can be difficult to find a truly
constant reference, an alternative is to use an external or exogenous reference. An
exogenous reference would be an RNA spike (in vitro transcript for example), that can be
added in a defined amount to the extracted RNA. This has the advantage that reference gene
expression levels are no longer a concern, but RNA isolation variances must still be
controlled for. For greatest control, endogenous and exogenous references can be combined
in a single assay.
Positive controls can be used to provide consistent positive reference data points in a given
experiment. The positive control material can also be used to create a standard curve.
Passive Reference Dye: Although it is not an amplification control, it is common practice
when performing QPCR to include a reference dye in the reaction mixture. The reference dye
is not linked to any amplification effect. Therefore, the fluorescence from this dye should be
constant throughout the amplification reaction. Provided concentration and volume are equal
in every well of the reaction, theoretically the fluorescence intensity for the reference dye
should be the same in every sample. The fluorescence signal for the fluorophores in the
reaction can be normalized to the reference dye by dividing the raw fluorescence intensity at
each cycle for the dye of interest by the fluorescence intensity from the reference dye at the
same cycle in the same tube. This will act to correct or “normalize” any signal level
differences (e.g. those caused by differences in plasticware transparency and reflectivity, or
volume differences due to aliquoting errors). Corrected data are designated as Rn or dRn in
the amplification plots and Report. The most commonly used reference dye is ROX.
Ultimately, the objective of using real-time quantitative PCR experiments is to determine the
absolute quantity of the target sequence present in the sample or to monitor the fold changes
of genes in response to experimental conditions. For accurate data analysis and meaningful
statistics using either of these approaches, the appropriate positive and negative controls
must be included with each real-time assay.
Primer and Probe Design
Primer and probe design is viewed as the most challenging step of setting up a new QPCR
experiment. However, the availability of numerous primer and probe design software
programs coupled with a set of easy to follow design rules makes the process relatively
simple and reliable. The first step in primer/probe design is to acquire the sequence of your
gene of interest. Numerous publicly available sequences can be found in open access
databases such as NCBI (www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=Nucleotide). After the
sequence is obtained, a primer/probe design software program should be used in order to
simplify and maximize success for the design process. (If you are requested to enter buffer
conditions by the design software, use 100 mM monovalent cation and 5 mM Mg++.)
Designer software packages are available both as freeware on the internet and through most
oligonucleotide vendors.
Standard Curves
After determining optimal primer and probe concentrations for the assay, it is recommended
to test the overall performance of the QPCR reaction in terms of efficiency, precision, and
sensitivity. Data generated from a serial dilution of a positive control template (standard
curve) are an excellent means of determining the overall performance of a QPCR assay. The
dilution series should encompass a large range of concentrations to ensure the reaction
performs at equal efficiency for high and low concentrations of starting template, ideally
encompassing the expected levels of target to be encountered with the experimental
samples. To accomplish this objective, a three-fold to ten-fold dilution series over several
33
orders of magnitude should be generated in triplicate. For example, for gene expression
experiments, a typical serial dilution would consist of five points of a five-fold serial dilution,
starting with 100 ng of total RNA per reaction (or the cDNA equivalent amount).
PCR Reaction Efficiency
The slope of the standard curve is used to determine reaction efficiency. Since the PCR
reaction is based on exponential amplification, if the efficiency of PCR amplification is 100%
the amount of template will double with each cycle, and the standard curve plot of the log of
starting template vs. PCR cycles which generate a linear fit with a slope between
approximately –3.1 and –3.6 are typically acceptable for most applications requiring accurate
quantification (90–110% reaction efficiency). If the amplification reaction is not efficient at the
point being used to extrapolate back to the amount of starting material (usually the Ct is used
for this purpose), then the calculated quantities may not be accurate.
Precision
The standard curve should be run in triplicate (or at least duplicate) so that it is possible to
determine the precision of pipetting, the reproducibility, and the overall sensitivity of an
assay. R2 is the fit of all data to the standard curve plot and can be influenced by accuracy of
the dilution series, and overall assay sensitivity. If all the data lie perfectly on the line, the R 2
will be 1.00. As the data fall further from the line, the Rsq decreases. As the R 2 decreases it
is more difficult to determine the exact location of the standard curve plot thus decreasing the
accuracy of quantification. An R2 value>0.985 is acceptable for most assays.
Sensitivity
The slope and R2 values of the standard curve help determine the sensitivity of a given
assay. If the slope of the standard curve is lower than –3.322, the R2 is below 0.985, and the
data points indicate an upward trend in the standard curve plot at the lower starting template
concentrations, this may indicate the reaction is reaching the threshold of sensitivity. In this
case, further assay optimization or even redesign of the primers and probe may be
necessary to extend the linear range.
Alternatively, the points outside the linear range can be culled from the standard curve.
However, unknown samples in that concentration range may not be trusted to give
quantitative results.
Standard Curve Examples
Figure 2 and Figure 3 illustrate a four-fold dilution series standard curve over three orders of
magnitude. In this example the data generate a linear standard curve with a slope of –3.401
which is well within the acceptable range of –3.1 to –3.6
and an amplification efficiency value (R2) of 98.6%, again, within the acceptable parameters
described above.
Figure 2
Amplification plots of standards in a fourfold dilution series over three
orders of magnitude.
34
Figure 3
Standard curve generated with data from
Figure 2, with slope and R2 indicated.
Further Optimization
If the assay is still not performing well after the probe and primer concentrations are
optimized, you can try altering the Mg++ concentration within the range of 3.5–5.5 mM for
TaqMan or Molecular Beacons reactions or in the range of 1.5–3.5 mM
for Scorpions or SYBR Green I reactions. If the reaction still does not work well after
complete optimization is performed, it may be necessary to redesign the primers and/or the
probe.
35
The ABI7000 prism
Double click on the programme icon
To create a new file click the “new”
button or go to menu→new, then click
OK in the dialog box
A new plate document pops up. To
set-up a new plate you have to
determine first which primer pair
you will use
Click tools→detector manager
Select the appropriate primer pair. If a
new pair is going to be used, click file
→new and determine the new primer
pair characteristics, then click OK.
If primers for SyBRGreen are used set
the reporter dye “SyBRGreen” and the
quencher dye “none” The new primer
pair is automatically incorporated in the
list, select and click “add to plate
document”.
36
Click the “well inspector” button
Select the du- or tri-plicates and specify
-which detector (primer) is used
-the task (NTC for negative control ie no
template samples, unknown for samples
to test and “standard” for the standard
curve points)
-if the task is standard, then fill the
quantity area. Attention! Use”.” And not “,”
as decimal separator
Keep in mind that the instrument will
present the results in a list going from A1
to A12, B1 to B12 etc. Load your samples
in that order (not A1 to H1).
Select the passive reference
Once you filled up the whole plate, move
to the “Instrument” tab
You can now specify the cycling
conditions
The stage 1 is optional. Use it only if you
follow the Amp erase procedure. You can
erase it by “shift clicking” on the stage 1
area, then “backspace”.
Stage 2: initial dissociation and polymerase activation.
Stage 3: modify the cycling conditions by
clicking on the corresponding areas
(NB. 0:15 =15 sec, 1:00= 1min)
Specify sample volume.
If SyBRGreen is used, don’t forget to click on the “dissociation protocol”. This will produce a”
melting curve” which will aloud you to see if there are any primer-dimers or secondary
products in your reaction. (see below)
37
To run a document, you have to save it first as an *sds extended file
Click file→save as
If you want to run the plate later, save as
an *sdt extended file .This template
document can be opened by selecting file
new→browse, select your template
document from the appropriate file, OK.
Again, if you want to run this template, you
have to save it as an *sds extended file.
Then ready click on the “Start” button in the
“instrument” tab.
The instrument will give you the remaining time
till the end of the experiment. (24:59 will show
up till the lid reaches 100oC)
Dissociation Curves (Only for SYBR® Green I)
As mentioned previously, when the detection chemistry is based on dsDNA detection, such as SYBR
Green I, you should run a melting (dissociation) curve at the end of your amplification reaction known
as a dissociation curve. The purpose of the dissociation curve is to determine if anything other than
the gene of interest was amplified in the QPCR reaction. Because SYBR green I will bind any double
stranded product, any non-specific amplification in your unknown wells will artificially increase
fluorescence and make it impossible to accurately quantitate your sample.
To view the SYBR Green I dissociation
curve, select the Results tab, and go to
‘Dissociation curve’ tab. The best way to
analyze the dissociation curve results is to
set the fluorescence to derivative. In this
view, every peak in the curve indicates a
specific product is melting.
Most QPCR products will melt somewhere in the range of 80–90°C, although this can vary
given the size and sequence of your specific target. Ideally, you should see a single peak
within this temperature range, and the melting temperature should be the same in all the
reactions where you have amplified the same sample. If any secondary peaks or shoulders
are seen on the peak of interest, it indicates that something other than your gene of interest
is present among the reaction products. Since there is no accurate way to determine how
much the amplified signal from each product is contributing to the Ct, if any secondary peaks
are observed the Ct value from that well should not be considered accurate. If secondary
peaks are seen, other controls run in the reaction may give you an indication of what was
causing this problem and how it can be prevented in the future. If these same secondary
peaks are present in your NTC wells, it may indicate primer dimer formation or the presence
of contamination by a sequence that was also amplified during the reaction. In the case of
primer dimers, re-optimizing the reaction conditions may be necessary. On occasion, it may
be necessary to re-design the primers. If the secondary peaks are not seen in the NTC wells,
38
it could indicate non-specific primer binding or the presence of differentially spliced products.
Performing a BLAST search following primer design may help decrease the incidence of this
type of problems.
Setting the Baseline
Fluorescence intensity data (Amplification plots) can be described as a two-component
function:
- a linear component or background and
- an exponential component that contains the relevant information.
To isolate the exponential component, the linear contributions to fluorescence can be
estimated and subtracted. This is the “baseline correction”: it can be performed either
automatically or manually.
On the results tab, amplification plot, select all wells by clicking on the upper left corner of the
plate setup.
For automated baseline click on the “auto baseline” button.
For manual baseline subtraction:
Double clic the Y axis.
1. Identify the range of cycles during which all contributions to fluorescence are strictly linear
(no exponential increase in fluorescence).
2. fill in the baseline start and end cycle boxes . The baseline end cycle is set at a cycle
around 3-5 cycles before the cycle at which an amplification is observed.
click “analyse”
However, there are a few options for determining which cycles to use to estimate the
contribution from the background fluorescence:
Automatic baseline (default method): When this method of baseline correction is selected,
the software will automatically select the appropriate cycles for each plot
Figure 4
An example of an amplification plot where
the baseline range is set incorrectly. In this
case, the baseline range for the orange
plot has been set to begin at cycle 3 and
end at cycle 15, and this range includes
the first part of the fluorescence shift.
Figure 5
The Amplification plot from Figure 4 after
the baseline range has been corrected to
the cycle range 3–10.
39
Setting the Threshold
The basic principle used in the analysis of real-time PCR data is that the number of cycles
necessary to reach a fixed concentration of amplicon in the reaction is an accurate estimator
of the initial target concentration at the beginning of the reaction.
Therefore, the number of cycles required to reach arbitrary fluorescence intensity should
correlate well with initial target concentration, as fluorescence intensity values correlate with
the concentration of the PCR products. This fluorescence value is referred to as the
“threshold fluorescence”, and the number of cycles required for any one reaction to reach it is
the “threshold cycle” or “Ct”. Ct values correlate very well with initial target concentration as
long as some assumptions are satisfied. Namely, that the kinetics of the reaction is
approximately constant throughout the reaction and that they are also similar between any
samples that are being compared to each other (e.g. standards and unknowns).
To satisfy these conditions, the threshold value has to be set at a point where all samples
being analyzed display the same rate of increase in the fluorescence intensity, and ideally
this increase responds to an exponential function. In addition, valid quantitative comparisons
can only be done between PCR reactions that amplify the same target (i.e. use the same
primer set).
There are different ways of setting the threshold value, a software algorithm and a manually
set threshold.
Amplification-Based Threshold:
This algorithm first determines the portion of the amplification plots where all of the data
curves display an exponential increase in fluorescence. To do this, the software looks at the
shift in fluorescence for each baseline-corrected curve and sets a point just above the
baseline at 0% and the maximum of the first derivative as 100% amplification. As a default,
the search range for the algorithm falls within 5–60% of this fluorescence shift for all the
curves. This range can be manually adjusted based on personal preferences, by accessing
the Analysis Settings, Threshold
Once the search range for the amplification-based threshold is established, the threshold
value is set based on one of two different criteria. In experiments where there are at least two
wells for each replicate, the algorithm calculates the threshold value that minimizes the
standard deviation (σ) in Ct values for each replicate set. If there are no replicate wells, the
algorithm will instead use a fixed amplification position. In such cases, the software sets the
threshold at the midpoint of the Search Range. If the default search range of 5–60% is used,
the threshold will be set at 32.5%.
Manually-Set Threshold:
Normally the software based methods will select a good threshold, but in cases where the
curves do not conform to the assumptions made by the algorithm, an incorrect threshold may
be calculated. Good indicators of improperly-set threshold values are false positives (Ct
values obtained from negative control wells), known positive samples giving very late Cts or
no Cts at all, or non-linear standard curves. There are other possible causes of all these
results which will be discussed later, but manually adjusting the threshold is one way to
correct these errors.
When manually adjusting the threshold, it is best to view the amplification plots in a semi-log
scale. To do this, double click the Y axis, and under the section Post run Settings, Y-axis
select the button for Log and click on the OK button at the bottom of the window.
In the log scale, the amplification plots will normally appear rather noisy during the baseline
cycles, due to the log scale. Following the baseline cycles, relatively straight lines rise
upward in the region where amplification begins. These plots will eventually reach a plateau
(Figure 5).
40
To adjust the threshold for each dye
collected, click and drug the horizontal
threshold line up or down to the desired
position.
Figure 5
Amplification Plots viewed with the Y Axis
set to a log scale. The optimal setting for
the threshold is the point where all the log
plots are linear and parallel, as shown in
where the threshold is set here.
Alternately, on the screen to the right of the amplification plots the threshold is listed on the
screen. A numerical value for the threshold can be entered there.
Ideally, the threshold should be set in the region where the plots are all linear and where they
are all as close as possible to parallel to one another. The threshold should not be so high
that it crosses any of the plots where they are starting to plateau and are no longer linear. If
possible, the threshold line should be placed above the highest points of the fluorescent plots
in the early (background fluorescence) cycles.
Check all ractions for normal amplification signal and dissociation. If there are reactions that
clearly did not work, you can omit them by clicking the “omit well” button in the well inspector
window.
Controls
Negative controls must be like Ceasar’s
wife
Above suspicion
Prior to moving on to analysis of the results, it is important to verify that the controls are
behaving as expected. If this is not the case, the quantitative results may not be accurate,
and further troubleshooting may be necessary.
Ideally, none of the negative control wells should cross the threshold, although it is not
uncommon to see the negative controls drift across the threshold during late cycles. If the
negative controls are displaying sigmoid-shaped amplification curves, the fact that real
amplification of the negative control is taking place would be indicated. This may be due to
template contamination or excessive primer dimmer formation.
41
NTC plot
Whether this will affect the Cts of the unknown samples will depend on the level of the signal
in your negative controls. If the Cts of the negative control wells are ten cycles higher than
the Cts of any of the unknown wells, it is safe to assume that these results are accurate. If
the Cts in the negative control wells are within five cycles of any of the unknowns, this may
call the validity of the results into question. Under these circumstances it may be necessary
to troubleshoot the reaction to determine the source of signal in the negative control wells.
The type of negative control well from which the signal was detected can provide an
important indication of the source of the trouble. A shift in the No RT controls would indicate
possible genomic DNA contamination. A shift in the NAC control wells could indicate probe
degradation and a shift in the NTC wells may indicate primer dimer formation (when
performing a SYBR Green I assay), or contamination. If the shift in the negative control wells
is due to primer dimers, you can determine if the primer dimers are also forming in the
unknown wells by looking at the dissociation curves.
If the positive control wells are not showing amplification, it will call into question whether any
of the unknown wells that did not amplify are actually negative samples or whether this is due
to non-specific failure of the PCR reaction (e.g. the presence of an amplification inhibitor). In
this case, it may be necessary to troubleshoot the reaction conditions (e.g. different water
and/or primer sources). The presence of PCR inhibitors in the template can also be identified
by decreasing the amount of template used. If the Ct values tend to decrease or remain
constant in the presence of lower amounts of template, this usually indicates the presence of
an amplification inhibitor.
Standard Curve Quantification
After amplification, given that both the standards and experimental samples are amplifying
efficiently, the Cts for each standard dilution can be determined and plotted against the initial
template quantity. Sample Ct values can be used to estimate template quantity by comparing
them to the standard curve. For this estimate to be accurate, the standard curve must be
linear across the whole range of template concentrations in your assay and the measured
efficiency of amplification near 100%.
The Ct values from each standard well will be used to create a standard curve. Figure 31
represents a typical standard curve constructed over three orders of magnitude (40 copies to
20,000 copies).
Data from a standard curve run can be viewed in multiple formats including: standard curve,
initial template quantity, and plate sample values.
42
In the standard curve view, as seen in
Figure 6, the efficiency and linearity will
automatically be displayed by the software
using the equation:
Xn = X0(1+E)n
Xn = amplified target amount (target
quantity at cycle n)
X0 = starting quantity
E = efficiency of amplification
n = number of cycles
When the efficiency is perfect (100% or 1), there is a perfect doubling of target amplicon
every cycle; a 10–fold amplification should take 3.32 cycles In a plot of Ct versus the log of
initial template, the slope should therefore be close to –3.32 (negative because a higher Ct
means lower template amount). Because of this relationship, you can calculate the efficiency
directly from the slope using the equation below:
Efficiency = 10 –1(–1/slope)
Relative or Comparative Quantification
In a typical plate setup for a comparative quantitation reaction, the reference and target
genes are run in separate wells, although they can be multiplexed in the same well. There
will be at least three different well types used for analysis: the unknowns (experimental
samples), the standards (controls), and negative controls (e.g. NTC wells). If a reference
gene is included in the experiment, then there will be unknown wells that have the target
gene of interest and others that have the reference (these can be run in the same tube if you
are multiplexing). The same is true of the calibrator wells.
Viewing the report
After setting the threshold, select all the
wells
by clicking on the upper left square of the
plate setup.
Click on the result tab, Report
To export the data click on menu File,
Export, Results. A dial box will appear.
Save the results and move to another
computer to analyze them.
The results are saved in a Comma Separated file *.CSV). Open it, select menu Data,
transform, text to table. Specify that the separator is comma, save XL file.
Select all columns, find and replace . by ,.
Analyse data
In order to obtain the “expression level” of each sample, the quantity of the target has to be
normalized (divided) by the quantity of the reference gene in the same sample.
43
If a calibrator is used (ie untreated sample) the fold induction is given by the ratio between
the expression level of a sample and the expression level of the calibrator.
fold induction =
expression level sample
expression level calibrator
44
hands on
plate setup
GRP
A
B
C
D
E
Standard
target
Standard
ref
A
1
2
3
4
5
6
7
NTC NTC target target NTC NTC ref
target target
ref
ref
8
ref
9
B
C
D
E
F
100
50
25
12.5
NTC
G
100
50
25
12.5
NTC
10
11
12
prepare mixes
mix target gene
H2O
Buffer 2X
primer target F10μM
primer target R10μM
2.6
10
1.2
1.2
X4
10.4
40
4.8
4.8
mix reference gene
H2O
buffer2X
primer ref F5μM
primer ref R5μM
3
10
1
1
X4
12
40
4
4
dispense 15 μl of appropriate mix in the correspondent wells
ie. group A will load well A1A4 with 15 μl of target mix and well A5A8 with 15 μl of ref
mix
group B will load well B1B4 with 15 μl of target mix and well B5B8 with 15 μl of ref
mix
add 5μl H2O in NTC wells (1 ,2, 5 &6 of you group lane)
template cDNA diluted 1/5 in test wells (3, 4, 7 & 8)
seal the plate
specify programme
95°C 10min
95°C 15sec
60°C 1 min
40 cycles
place in the cycler & run
45
western blot
Method overview
A western blot (alternately, immunoblot) is a method to detect a specific protein in a given
sample of tissue homogenate or extract. It uses gel electrophoresis to separate native or
denatured proteins by the length of the polypeptide (denaturing conditions) or by the 3-D
structure of the protein (native/ non-denaturing conditions). The proteins are then transferred
to a membrane (typically nitrocellulose or PVDF), where they are probed (detected) using
antibodies specific to the target protein. There are now many reagent companies that
specialise in providing antibodies (both monoclonal and polyclonal antibodies) against many
thousands of different proteins. Commercial antibodies are expensive, though can be re-used
(unbound antibody) between experiments.
o
Tissue preparation
Samples may be taken from whole tissue or from cell culture. In most cases, solid tissues are
first broken down mechanically using a blender (for larger sample volumes), using a
homogenizer (smaller volumes), or by sonication. Cells may also be broken open by one of
the above mechanical methods.
Assorted detergents, salts, and buffers may be employed to encourage lysis of cells and to
solubilize proteins. Protease inhibitors are often added to prevent the digestion of the sample
by its own enzymes.
A combination of biochemical and mechanical techniques – including various types of
filtration and centrifugation – can be used to separate different cell compartments and
organelles.
Gel electrophoresis
The proteins of the sample are separated using gel electrophoresis. Separation of proteins
may be by isoelectric point (pI), molecular weight, electric charge, or a combination of these
factors. The nature of the separation depends on the treatment of the sample and the nature
of the gel.
By far the most common type of gel electrophoresis employs polyacrylamide gels and buffers
loaded with (SDS) sodium dodecyl sulfate. SDS-PAGE (SDS polyacrylamide gel
electrophoresis) maintains polypeptides in a denatured state once they have been treated
with strong reducing agents to remove secondary and tertiary structure (e.g. S-S disulphide
bonds to SH and SH) and thus allows separation of proteins by their molecular weight.
Sampled proteins become covered in the negatively charged SDS and move to the positively
charged electrode through the acrylamide mesh of the gel. Smaller proteins migrate faster
through this mesh and the proteins are thus separated according to size (usually measured in
kilo Daltons, kD). The concentration of acrylamide determines the resolution of the gel - the
greater the acrylamide concentration the better the resolution of higher molecular weight
proteins. Proteins travel only in one dimension along the gel for most blots.
Samples are loaded into wells in the gel. One lane is usually reserved for a marker or ladder,
a commercially available mixture of proteins having defined molecular weights, typically
stained so as to form visible, coloured bands. When voltage is applied along the gel, proteins
migrate into it at different speeds. These different rates of advancement (different
electrophoretic mobilities) separate into bands within each lane.
It is also possible to use a two-dimensional (2-D) gel which spreads the proteins from a single
sample out in two dimensions. Proteins are separated according to isoelectric point (pH at
46
which they have neutral net charge) in the first dimension, and according to their molecular
weight in the second dimension.
Transfer
In order to make the proteins accessible to antibody detection, they are moved from within
the gel onto a membrane made of nitrocellulose or PVDF. The membrane is placed on top of
the gel, and a stack of tissue papers placed on top of that. The entire stack is placed in a
buffer solution which moves up the paper by capillary action, bringing the proteins with it.
Another method for transferring the proteins is called electroblotting and uses an electric
current to pull proteins from the gel into the PVDF or nitrocellulose membrane. The proteins
have now moved from within the gel onto the membrane while maintaining the organization
they had within the gel. As a result of this "blotting" process, the proteins are exposed on a
thin surface layer for detection (see below). Both varieties of membrane are chosen for their
non-specific protein binding properties (i.e. binds all proteins equally well). Protein binding is
based upon hydrophobic interactions, as well as charged interactions between the
membrane and protein. Nitrocellulose membranes are cheaper than PVDF, but are far more
fragile and do not stand up well to repeated probings.
Blocking
Since the membrane has been chosen for its ability to bind protein, and both antibodies and
the target are proteins, steps must be taken to prevent interactions between the membrane
and the antibody used for detection of the target protein. Blocking of non-specific binding is
achieved by placing the membrane in a blocking solution of protein - typically Bovine serum
albumin (BSA) or non-fat dry milk (both are inexpensive), with a minute percentage of
detergent such as Tween 20. The protein in the blocking solution attaches to the membrane
in all places where the target proteins have not attached. Thus, when the antibody is added,
there is no room on the membrane for it to attach other than on the binding sites of the
specific target protein. This reduces "noise" in the final product of the Western blot, leading to
clearer results, and eliminates false positives.
Detection
During the detection process the membrane is "probed" for the protein of interest with a
modified antibody which is linked to a reporter enzyme, which when exposed to an
appropriate substrate drives a colorimetric reaction and produces a colour. For a variety of
reasons, this traditionally takes place in a two-step process, although there are now one-step
detection methods available for certain applications.
Two step

Primary antibody
Antibodies are generated when a host species or immune cell culture is exposed to the
protein of interest (or a part thereof). Normally, this is part of the immune response, whereas
here they are harvested and used as sensitive and specific detection tools that bind the
protein directly.
After blocking, a dilute solution of primary antibody (generally between 0.5 and 5
micrograms/ml) is incubated with the membrane under gentle agitation. Typically, the solution
is comprised of buffered saline solution with a small percentage of detergent, and sometimes
with powdered milk or BSA. The antibody solution and the membrane can be sealed and
incubated together for anywhere from 30 minutes to overnight. It can also be incubated at
different temperatures, with warmer temperatures being associated with more binding, both
specific (to the target protein, the "signal") and non-specific ("noise").
47

Secondary antibody
After rinsing the membrane to remove unbound primary antibody, the membrane is exposed
to another antibody, directed at a species-specific portion of the primary antibody. This is
known as a secondary antibody, and due to its targeting properties, tends to be referred to as
"anti-mouse," "anti-goat," etc. Antibodies come from animal sources (or animal sourced
hybridoma cultures); an anti-mouse secondary will bind to just about any mouse-sourced
primary antibody. This allows some cost savings by allowing an entire lab to share a single
source of mass-produced antibody, and provides for more consistent results. The secondary
antibody is usually linked to biotin or to a reporter enzyme such as alkaline phosphatase or
horseradish peroxidase. This means that several secondary antibodies will bind to one
primary antibody and enhances the signal.
Most commonly, a horseradish peroxidase-linked secondary is used in conjunction with a
chemiluminescent agent, and the reaction product produces luminescence in proportion to
the amount of protein. A sensitive sheet of photographic film is placed against the membrane,
and exposure to the light from the reaction creates an image of the antibodies bound to the
blot.
Analysis
After the unbound probes are washed away, the western blot is ready for detection of the
probes that are labeled and bound to the protein of interest. In practical terms, not all
westerns reveal protein only at one band in a membrane. Size approximations are taken by
comparing the stained bands to that of the marker or ladder loaded during electrophoresis.
The process is repeated for a structural protein, such as actin or tubulin, that should not
change between samples. The amount of target protein is indexed to the structural protein to
control between groups. This practice ensures correction for the amount of total protein on
the membrane in case of errors or incomplete transfers.
colorimetric detection
The colorimetric detection method depends on incubation of the western blot with a substrate
that reacts with the reporter enzyme (such as peroxidase) that is bound to the secondary
antibody. This converts the soluble dye into an insoluble form of a different color that
precipitates next to the enzyme and thereby stains the nitrocellulose membrane.
Development of the blot is then stopped by washing away the soluble dye. Protein levels are
evaluated through densitometry (how intense the stain is) or spectrophotometry.
chemiluminescence
Chemiluminescent detection methods depend on incubation of the western blot with a
substrate that will luminesce when exposed to the reporter on the secondary antibody. The
light is then detected by photographic film, and more recently by CCD cameras which
captures a digital image of the western blot. The image is analysed by densitometry, which
evaluates the relative amount of protein staining and quantifies the results in terms of optical
density. Newer software allows further data analysis such as molecular weight analysis if
appropriate standards are used. So-called "enhanced chemiluminescent" (ECL) detection is
considered to be among the most sensitive detection methods for blotting analysis.
Secondary probing
One major difference between nitrocellulose and PVDF membranes relates to the ability of
each to support "stripping" antibodies off and reusing the membrane for subsequent antibody
probes. While there are well-established protocols available for stripping nitrocellulose
membranes, the sturdier PVDF allows for easier stripping, and for more reuse before
background noise limits experiments. Another difference is that, unlike nitrocellulose, PVDF
must be soaked in 95% ethanol, isopropanol or methanol before use. PVDF membranes also
tend to be thicker and more resistant to damage during use.
48
hands on
Important remark!!! all buffers and samples should be kept at 4ºC (ice)
N.B. there are slight modifications of the protocol from lab to lab.
sample preparation (see above)
in 1.7 ml tube put the appropriate amount of protein extracts ( usually 10-30μg)
add appropriate volume of LB4x ( ie 12μl extract+4μl LB4x)
heat at 100°C for 5 min, place on ice for 1-2 min, spin briefly, keep on ice until loading
12% acrylamide gel
place clean glasses in the casting device
prepare running gel
H2O
4.84 ml
Acrylamide 29%-Bis1% 9 ml
Tris pH8.8 1M
8.44 ml
SDS10%
225 μl
APS10% (add just
112.5 μl
before pooring)
Temed (add just
18 μl
before pooring)
prepare stacking gel
H2O
Acrylamide 29%-Bis1%
Tris pH6.8 1M
SDS10%
APS10% (add just
before pooring)
Temed (add just
before pooring)
5.99ml
1 ml
0.95
75 μl
75 μl
7.5 μl
mix & poor running gel with a Pasteur pipette till 1cm below the short glass plate
overlay Isopropanol
let polymerise for 20 min, check for polymerization
poor off isopropanol and poor stacking gel, place comb
let polymerise 15 min
place glass/gel sandwich in the electrophoresis device
fill the tank with running buffer
take off combs
wash wells
load marker and samples
run at 100V for 15min until the front reach the running gel
run at 150V45min
transfer
in a backet full of transfer buffer place:
the transfer frame with the black side down (negative pole)
fiber pad
2 whatman papers
49
the gel
a nitrocellulose sheet of the same size as the gel
AVOID AIR BUBBLES roll the air bubbles out
cover with
2 whatman papers
fiber pad
close the transfer frame and place in the transfer device with the black side toward the
negative pole
fill in the transfer buffer
put a magnetic bar at the bottom
place on magnetic stirrer
run the transfer
at 350 mA for 1 hour
take the membrane out, put it in a plastic container with face up (usually a tip box lid)
wash briefly in TBS
Block
incubate in TBS-T-5% non fat milk for 1 hour at RT on a rocker
wash briefly in TBS
primary antibody
make the desired dilution of the I Ab in TBS-T-1% non fat milk (usually 8 ml, or 2ml if plastic
bags are used)
incubate for 1 hour at RT or overnight at 4°C on a rocker
wash briefly in TBS-T
2*wash TBS-T-5% milk 15 min on a rocker
2*wash TBS-T- 15 min
secondary antibody
make the desired dilution of the II Ab in TBS-T-1% non fat milk (usually 8 ml, or 2ml if plastic
bags are used)
incubate for 1 hour at RT on a rocker
wash briefly in TBS-T
wash TBS-T-5% milk 15 min
3*wash TBS-T- 15 min
reveal
place the membrane, face-up, on a saran
following the manufacturer instructions, mix 1ml ECLreagent A+1ml ECLreagent B
poor the mix on the membrane
incubate I min,
decant
wrap in saran
use adhesive tape to immobilise it in a cassette
In the dark roo,
place a film on top of the membrane
(use the edge of the cassette as a grid
expose initially for 1 min.
Adjust the exposure time according to the signal
stripping
incubate mb face up 55ºC for 10-15min (covered by stripping buffer)
2Xquick washes in TBS-T
2X wash for 10min
50
blocking with TBS-T/milk 1hour
alternatively
wash 30min-1hr,in TBS-T 1-2%Tween at RT
procced with the normal washes in TBS-T & blocking
buffers
Lysis buffer
H2O
9.475 μl
Tris pH8 1M
100 μl
NaCl 5M
300 μl
EGTA pH8 0.4M
25 μl
Triton X100
100 μl
Protease inhibitors
1tb
keep up to 1mth at 4°C
Cfinale
10mM
150mM
16mM
0.01%
Electrophoresis running buffer 1X
Cfinal
Tris
25mM
Glycine
192 mM
SDS
0.1%
Transfer buffer 1X
Cfinal
Tris
25mM
Glycine
192 mM
SDS
0.02%
Methanol
20%
TBS
Tris
NaCl
12,1 gr
40 gr
Cfinal
10mM
15mM
TBS-T
add appropriate volume of Tween 100 according to the specificity of the Ab. Generally, a
range from 0.1 to 0.05% is used. Tween100 is a detergent, the more we put, the more the
conditions are stringent
Stripping buffer 1X
Tris pH6,8
SDS20%
Β-mercaptoethanol
(add prior to use)
Cfinal
62,5 mM
2%
10mM
51
References
Molecular Cloning
alaboratory manual
Sambrook, Fritsch, Maniatis
CSH editions
52