Quantitative RT-PCR
8/31/10
1. RNA preparation
We use the Absolutely RNA kit from Stratagene (cat #: 400800). I like this kit
because we have gotten consistent results with the DNAse treatment, and it is simple to
use. We have used it for RNA preparation from as little as 10mg of tissue and about
125,000 cultured neurons. We follow the protocol with the following adaptations:
For small samples, we use the small sample appendix information which has you
lyse the cells in a smaller volume, spin them on the first column for a short time, and then
add more lysis buffer which you spin through.
Because there is never enough DNAse with the kit, we make 50uL DNAse
digestion buffer with 5uL DNAse for EXACTLY the number of samples you are
processing, then use 50uL of this mix (not 55) per sample and you will have enough.
Elute 1 x 30uL elution buffer.
Always spec the RNA, diluting 2uL of the RNA in 100uL ddH2O. You expect to
have an OD260 of 0.037 or higher and if the samples were originally similar, the OD260
values should be within 2 fold of each other. The 260:280 ratio is ideally about 1.8 and if
it is significantly off the RNA may not be good.
Store the RNA at -80 for up to several years if all is truly RNAse free and freezethaws are limited. Of course all rules for safe RNA work must be followed – filter tips
only, GLOVES always, DEPC water and solutions only, all solutions must be specifically
labeled for RNA work, and don’t spit in the samples (i.e. don’t talk while you pipette).
2. cDNA synthesis
We use the Superscript II cDNA synthesis kit from Invitrogen (cat #:11904-018).
I like this kit because it has been reliable and because most of the key reagents can be
replenished from the Jones Cell Culture stockroom if they run out. We follow the
protocol with the following adaptations:
Assuming the RNA meets the criteria described above, (OD260>0.037, no more
than 2X variation across samples) we use 8uL of the RNA for the cDNA synthesis. The
kit can accommodate up to 5ug RNA which with the dilutions above in 8uL would be an
OD260 of about 0.3, so with anything below that you can put 8uL in the cDNA synthesis
(usually our OD260 values range from 0.03 to 0.15 depending on the quantity of the
starting material).
You can use either the oligo dT or the random hexamer primers in the synthesis
reaction, but it is probably best to be consistent. I have used both and noticed no obvious
differences. We have used oligo dT the most recently.
Use the PCR machine and the PCR tubes for the incubations to ensure accurate
heating, and use the 2uL pipettes for pipetting. Make sure you put the caps on the PCR
tubes and then pull them off to loosen before you put actual samples in there. The caps
can be hard to get off as they are so tight and sometimes the samples fly.
Make sure you use 25mM MgCl2 – there are a lot of 50mM tubes floating around
in the freezer, so check the label.
ONLY use solutions intended for cDNA synthesis, and all RNA rules (as above)
apply.
Here is the protocol:
1) Make a primer/dNTP master mix for n+2 samples. For each:
1µL dNTPs
1µL oligodT or random hexamers
vortex, spin down, and aliquot 2uL in each tube
2) Add up to 8uL RNA (and if needed at DEPC water up to 8uL)
3) Vortex to mix, spin down, put at 65C for 5’, then chill on ice for 1’
4) Make buffer master mix for n+2 samples. For each:
2µL 10X RT buffer
4µL 25mM MgCl2
2µL 0.1mM DTT
1µL RNAseOUT
vortex, spin down, and add 9µL to each of the samples
5) For oligo dT:
incubate at 42C for 2’, then add 1µL Superscript II to each tube
Vortex and spin down, then go to step 6
For random hexamers:
Incubate at 25 for 2’, then add 1µL Superscript II to each tube
Vortex and spin down, incubate 10’ at 25C, then go to step 6
6) Run the PCR program AW-RT
50’ at 42C, 15’ at 70C, hold at 4C
7) Add 0.5µL RNAseH/sample, vortex, spin down
Incubate 37C for 20’
At the end of the synthesis, take ~2 µL from each sample and pool them together
in a common well to serve the standards for upcoming PCR reactions. Then dilute all
other samples 5 fold with ddH2O. So for example, you will have 20uL of each cDNA
reaction at the end of the protocol. If you take 2µL from each for the standards, and have
18µL left, add 72µL ddH2O to each cDNA sample. This will allow you to have enough
sample for as many PCR reactions as you need. If you anticipate using the cDNA
samples for multiple PCR reactions (requiring multiple freeze thaws or over a long time)
aliquot the cDNA in multiple tubes and store the samples not in current use at -80. Store
the samples in current use at -20.
For the standards, make 1:10 and 1:100 dilutions of your pooled sample. Mix
VERY well (vortex and spin down) and pipette carefully to ensure they are accurate.
3. Quantitative PCR
We use Power SYBR green from ABI (cat #: 4368706), an ABI 7300 cycler,
plates and optical covers from ABI (cat #s: 4306737 for plates, 4311971 for covers) and
primers from IDT-DNA (www.idtdna.com/duke). You can use any SYBR green PCR
enzyme system that has Rox in it, but we have had the best results with ABI and rather
inconsistent results with the AbGene enzyme which ended our desire to test others. Our
machine can also be used for Taqman PCR in up to 4 colors.
Make sure to sign up for a time to use the machine in the red book near the cycler.
You should sign up for about 4 hours because it takes a while to get the machine set up at
the start and to analyze your data at the end, and the program runs for about 3 hours.
Common primers are listed at the end of this protocol. In every case for cDNA
synthesis, the first step is to check the RNA quality by PCR for the housekeeping enzyme
Gapdh. We use this enzyme because it is not so highly expressed as some other
housekeeping enzymes and therefore is closer in level to the other genes we usually
study. Gapdh has been reported to vary between tissues and individuals, so in some cases
you may choose to normalize to another housekeeping gene such as actin or the 18S
ribosomal subunit. (Realize that since 18S isn’t a PolII transcript, you can’t use this
primer pair for oligo dT primed cDNA.)
Make a 12x8 grid and note which samples are in which wells!!!! All samples must
be run in triplicate, plus you need four samples for the standards and no template control
for each primer pair if you want absolute values (which is easier than doing ddCt). On a
96 well plate that means you can do 28 samples with one primer pair, 12 samples with 2
primer pairs, 6 samples with three primer pairs, or 4 samples with 4 primer pairs.
Arrange eppendorf tubes in a 12x8 eppendorf tube rack in the order you will place
the samples on the plate. I usually do sample numbers top to bottom and triplicates L to
R, but you can do it any way that works for you.
Before starting, mix up the primers. We store our primer stocks at 1ug/uL. You
will use them at 10ng/uL. So start by making a working dilution of the primers by mixing
6uL F and 6uL R of the stock solutions in 108µL ddH2O (makes enough for 16 samples,
or half a plate – increase or decrease as needed depending on the sample number).
We make enough mix for 3.25 wells for each sample, then separate it into the
three wells for the triplicate read. That means you put in each tube:
22.75µL ddH2O (FRESH daily from the white tap)
32.5 µL 2X SYBR green Master Mix (current at 4 degrees, extra at -20)
6.5µL Working dilution primer mix
3.25µL cDNA
Add the solutions in this order to minimize tip switching. You can use one tip for
all the water. Then switch tips and use one for all the master mix (move an aliquot of
master mix from the stock vial to a tube for your own use to avoid contaminating the
stock). Note the master mix is thick with glycerol, so pipette slowly to dispense the
correct amount. Then add primers using a single tip for all samples that get the sample
primer mix. Finally the cDNA goes in last. PIPETTE CAREFULLY! This is the smallest
volume, and pipetting is the largest source of error in this protocol. No talking while
pipetting. (Seriously – this will help.)
For the standards, use the dilutions you made of the cDNA for 1, 1:10, and 1:100.
In the last tube, use 3.25µL ddH2O instead of cDNA as a no template control. This well
will show you the non-specific primer dimer signal and will also ensure that you are not
contaminating your reaction with cDNA from somewhere other than your samples (like
plasmid DNA that is on your pipetters).
IF you cannot use exon skipping primers or were unable to treat the samples with
DNAse, an additional control is to run noRT controls. In these wells, instead of using
cDNA, you simply add RNA at the dilution that matches how much is in the cDNA
sample (for example, in the protocol above you took 8uL RNA and it wound up in 90uL
of cDNA. So to mimic that, take 1uL RNA and add 10uL ddH2O, then use 3.25 of that
mix in the PCR reaction. If the DNAse treatment worked, the signal in that sample should
be MORE THAN 3 cycles (preferable more than 6 cycles) below the cDNA sample - so
the DNA contamination accounts for <10% of the signal and preferably <1%.
Once all the tubes are prepared, cap them, number them, vortex briefly, and spin
down. Then put a realtime 96 well plate on the black scaffold and CAREFULLY pipette
20µL of each sample into each of three wells.
When pipetting is done, cover with optical tape (careful to get NO
FINGERPRINTS on it) and burnish with the square tape sticky down thing.
Spin down in the plate rack of the table tap centrifuge 1’ at 1K. Make sure to
balance the rotor with all four buckets and put plates in each.
Set up the machine: make sure the PCR machine is turned on before you open the
program so the machine will initiate. If the program has already been running, shut it
down and reopen it to get it to run the initiation. Open ABI7300 and choose new plate.
Give it a name that includes the date (083108 CarfCocaine). Choose your detectors (these
are the genes you are PCRing for) and mark where they will go on the plate. If you want
you can choose “done” at this point.
When the next plate page opens, double click a well to open the “well detector”.
You can hold shift and highlight the triplicate wells to label them at once. Name all the
wells NOW!!! You will not want to do this later and if you don’t it makes it hard to go
back and make sense of the data. Mark the wells that are standards and the no template
control. For values call the standards 1, 0.1 and 0.01. Note that these are therefore relative
absolute values.
Now click the “instrument” tab and set the program. Click the first 50 degree step
and hit delete – we don’t use that one. Place the cursor after the second 95 degree step
and do “add step”. They preset a single annealing/extension step, but we do them
separately. In the newly added step (which is not the yellow step…) set the annealing
temp you have determined for your primers (usually about 60 if you have to guess), and
make the time 0:45. In the next step (the extension step, yellow because it is where the
photo is taken) make the temp 72 and the time 1:00. Now place your cursor after the last
step and click “add dissociation”. A bunch of steps will appear that will allow you to
ensure you have a single PCR product. Finally change the volume to 20uL. Then click
the “save” button at the top and navigate to your folder.
Once the file is saved you are ready to click “start”. Make sure the program starts
to run. It will tell you the estimated time for the run. The data will be automatically saved
at the end. Now pull the paper over the screen to discourage others from using the
machine during your run. Occasionally the machine will freeze up, and if it does your
experiment will in fact be ruined.
When the run is done, click on the cycles tab and then click “analyze”. You rarely
need to change the baseline or the threshold manually. All the curves should now appear.
Highlight your standards and make sure 1) the triplicates are close, 2) the dilutions are
3.3 cycles apart, 3) the NTC is way below the 1:100 standard. If your data passes this,
highlight all the wells and go to the “standard curves” tab. Make sure the line has a slope
close to -3.3 and your samples are within the standard range. Now click on “(is it
dissociation?)” tab and make sure there is only one peak. The NTC may give other peaks,
but the samples should not.
Finally highlight all the wells and go to “results”. You should save the data at this
point and you can export to Excel as a .csv file (that can be converted to an excel spread
sheet in Excel). Make sure to print out a copy of the results file to save for future
reference.
For Gapdh you want to see two three things – your triplicate samples should be
very close to one another, overall your samples should vary by no more than 2 fold if
possible, and finally, for neurons treated as described above, you want the Ct values for
Gapdh to be about 18-20. If the values are much lower (mid-20s) or if they vary a lot, it is
likely either the RNA was degraded or the cDNA synthesis failed. Both can happen, so
you must troubleshoot. Check the RNA by running it on a gel to see what it looks like,
and replace all possible bad reagents from the cDNA synthesis kit if you plan to try
again.
If the read is good, then go ahead and run your other PCRs! You will normalize
each value for any other gene by the Gapdh value by dividing through in excel. The
triplicates from the PCR plate are just technical replicates and therefore get pooled
without worrying about calculating errors. The error comes from sample replicates.
Primers:
Fos m/r
Arc m
Junb m
Bdnf4 m/r
TTTATCCCCACGGTGACAGC
GAGCCTACAGAGCCAGGAGA
AAACTCCTGAAACCCACCTTAGC
CGCCATGCAATTTCCACTATCAATAA
CTGCTCTACTTTGCCCCTTCT
TGCCTTGAAAGTGTCTTGGA
CTGATCCCTGACCCGAAAAGT
GCCTTCATGCAACCGAAGTATG
Gapdh m/r
CATGGCCTTCCGTGTTCCT
TGATGTCATCATACTTGGCAGG
TT