Tuesday, November 18, 2014. St. Amand
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Genotyping Lab ABI Protocols
Overview
The ABI 3730 is a 48 capillary electrophoresis system that uses fluorescently
labeled dyes for detection of DNA. Typically, a PCR reaction that will use the
ABI will have 3 primers in the PCR mix: a "tailed" forward primer, a reverse
primer, and an M13 Dye labeled forward primer. Conversely, primers may be
"directly labeled" and in that case, only two primers are used just like a standard
PCR. Up to 4 dyes (4 different PCR reactions) can be pooled into a single ABI
capillary and run at the same time.
Primers
Modify the FORWARD primer sequence by adding an "M13" tail to the 5' end of
the primer. We use an 18 base tail. For example:
5'- ACG ACG TTG TAA AAC GAC + primer sequence -3'
Do not modify the reverse sequence. Resuspend new lyophilized primers in Te
(10mM TRIS, 1mM EDTA) to a stock concentration of 100 pm/ul and working
concentration of 10 pm/ul. Most of the SSR primers in our lab are already
"tailed". Before using a primer, find out if it is tailed or not.
M13 primers are labeled with fluorescent dyes.
Dye Set "G5"
FAM (Blue)
VIC (Green)
NED (Yellow)
PET (Red)
5'5'5'5'-
FAM-ACG
VIC-ACG
NED-ACG
PET-ACG
ACG
ACG
ACG
ACG
TTG
TTG
TTG
TTG
TAA
TAA
TAA
TAA
AAC
AAC
AAC
AAC
GAC
GAC
GAC
GAC
-3'
-3'
-3'
-3'
Dye Set "D"
FAM (Blue)
5'- FAM-ACG ACG TTG TAA AAC GAC -3'
HEX (Green) 5'- HEX-ACG ACG TTG TAA AAC GAC -3'
NED (Yellow) 5'- NED-ACG ACG TTG TAA AAC GAC -3'
We keep dye labeled M13 primers in stock in TH3401. Certain dyes can be
substituted for others with fair results. VIC and HEX are interchangeable. NED
and TAM are interchangeable. Use these same dye sets if you are directly
labeling a primer. VIC, NED, and PET dye labeled Oligos must be ordered from
https://www.lifetechnologies.com/order/custom-oligo/fluorescent-labeledprimers. FAM can also be ordered there or from any other oligo maker.
Tuesday, November 18, 2014. St. Amand
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PCR
The following master mix works well for most SSR and STS primers, but may
need to be modified for specific primers. Tween-20 at a final concentration of
0.5% is often helpful. Create a master mix by adding the following reagents (in
this order):
Reagent (Stock Conc.)
Autoclaved Nanopure Water
Ammonium Sulfate Buffer Stock (10.00 X)
MgCl2 (25.00 mM)
dNTP mix (5.00 mM each)
Forward Tailed Primer (10.00 uM)
Reverse Primer (10.00 uM)
M13 Dye labeled Primer (10.00 uM)
Home-made Polymerase (5.00 U/ul)
Master Mix total =
DNA (10 to 100 ng/reaction) =
PCR reaction total volume =
Vol (ul/reaction)
6.472
1.300
1.300
0.520
0.065
0.104
0.039
0.200
10.00
3.00
13.00
Final Conc.
1X
2.50 mM
200.00 uM each
50.00 nM
80.00 nM
30.00 nM
1.00 U/Rxn
Thermocycler profiles
The following profiles work well for SSR primers and many STS primers, but
may need to be modified for specific primers. The Q1SSR profile differs from the
LicorSSR profile only in step 11 and in denaturing, but is about 45 minutes
quicker per run. That allows the same PCR block to be used 4 to 5 times per day
as opposed to 2 or 3 times per day. The F50SSR profile takes about as long as
Q2SSR but has more final cycles for increased amplification.
LicorSSR
(Calculated profile,
~4 hours per run)
1. 95C, 5 min
2. 95C, 45 sec
3. 68C, 5 min, -2.0C/cycle
4. 72C, 1 min
5. Goto step 2, 4 more
times
6. 95C, 45 sec
7. 58C, 2 min, -2.0C/cycle
8. 72C, 1 min
9. Goto step 6, 4 more
times
10. 95C, 45 sec
11. 50C, 2 min
12. 72C, 1 min
13. Goto step 10, 24 more
times
14. 72C, 5 min
15. 4C, 15 min
Q2SSR
(Quicker LicorSSR,
Calculated profile,
~3 hours per run)
1. 95C, 5 min
2. 96C, 1 min
3. 68C, 5 min, -2.0C/cycle
4. 72C, 1 min
5. Goto step 2, 4 more
times
6. 96C, 1 min
7. 58C, 2 min, -2.0C/cycle
8. 72C, 1 min
9. Goto step 6, 4 more
times
10. 96C, 1 min
11. 50C, 1 min
12. 72C, 1 min
13. Goto step 10, 24 more
times
14. 72C, 5 min
15. 4C, 5 min
F50SSR
(Calculated profile, 3:13h)
Tuesday, November 18, 2014. St. Amand
1. 95C, 5 min
2. 96C, 1 min
3. 68C, 3 min, -2.0C/cycle
4. 72C, 1 min
5. Goto step 2, 4 more
times
6. 96C, 1 min
7. 58C, 2 min, -2.0C/cycle
8. 72C, 1 min
9. Goto step 6, 4 more
times
10. 96C, 20 sec
11. 50C, 20 sec
12. 72C, 30 sec
13. Goto step 10, 39 more
times
14. 72C, 5 min
15. 4C, 5 min
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Tuesday, November 18, 2014. St. Amand
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Pooling PCR Reactions
Up to 4 separate PCR reactions can be pooled into one ABI sample if each PCR
uses a different M13 dye. Pooling is not necessary but greatly increases ABI
throughput and lowers costs. We use dye set "G5" which has 4 dyes for samples
and a 5th dye (LIZ) for the standard. Each of the 4 sample dyes will fluoresce at
different wavelengths (colors) and at different intensities. Intensity differences
require specific pooling ratios for the samples. Robot scripts are available for
pooling using 384 and 96 well plates. We can also use dye set "D" which has 3
dyes for samples and a 4th dye (ROX) for the standard.
For dye set "G5", vortex and centrifuge each PCR plate and add:
3.0 ul FAM, 2.5 ul VIC (or HEX), 3.5 ul NED (or TAM), & 3.5 ul PET into 7.5 ul
water. Keep the total volume of the pool at 20 ul even if you are not pooling all 4
colors.
For dye set "D", vortex and centrifuge each PCR plate and add:
3.0 ul FAM: 3.0 ul HEX (or VIC): 4.0 ul NED (or TAM) into 10 ul water. Keep the
total volume of the pool at 20 ul even if you are not pooling all 3 colors.
The pooled plate can be stored in the dark at -20°C until needed, but avoid
freeze-thaw cycles.
Preparing Samples for an ABI run
A size standard must be added to each sample. The standard is labeled with an
orange dye (LIZ) for dye set "G5" or a red dye (ROX) for dye set "D". The samples
must also be suspended in formamide to denature the DNA. Prepare the
formamide-standard (FS) mix on ice. Add 0.15 ul GS500 LIZ (or ROX) to 9.85 ul
Hi-Di Formamide per sample, vortex. For CASSUL600, use 0.075 ul per sample
(or a ratio of 0.06, STD:Sample). For CASSUL1200, use 0.10 ul per sample (or a
ratio of 0.08, STD:Sample). (ALWAYS examine the peak heights of your size
standard. It should never be below 200 RFU. It should average about 400 RFU. If
it is above 800 RFU, you are wasting size standard and you are LOWERING the
sample peak heights because of competition between the standard and your
sample.) Pipette 10 ul of the FS into each well of a new, empty plate. Vortex and
centrifuge the pooled plate. Add 2.0 ul from the pooled plate to the FS plate for a
total volume of 12.0 ul per sample. Once the sample is mixed with formamide, it
is best to run the plate on the ABI within 48 hours. However, we have frozen
plates at -80 for a week or two and found them to be fine. Denature the FS plate
at 95°C for 5 minutes, place on ice for 5 minutes, centrifuge, and then place an
ABI rubber septa on the FS plate. Load the plate into a carrier and place it in the
ABI 3730. Purchase size standards and Hi-Di Formamide from Lifetechnologies
(www.lifetechnologies.com). Get ABI3730 certified plates from LabSource and
MidSci.
Tuesday, November 18, 2014. St. Amand
GeneScan™ 400HD ROX™ Size Standard
GeneScan™ 500 LIZ™ Size Standard
GeneScan™ 500 ROX™ Size Standard
GeneScan™ 600 LIZ™ Size Standard
GeneScan™ 1200 LIZ™ Size Standard
Hi-Di™ Formamide
ABI3730 96 well plates, www.clpdirect.com
ABI3730 96 well plates, LabSource
ABI3730 384 well plates, MidSci
ABI3730 96 well plates, bioexpress.com
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Part #402985
Part #4322682
Part #401734
Part #4366589
Part #4379950
Part #4311320
Part #3442
Part #T53-407
Part #AV384
Part #T-3149-1
User profiles on the ABI 3730
Before any runs can be started, each user must have their own user profile
(results group) stored in the ABI software. Ask for help in setting up your own
results group on the lab computer. This only needs to be done once.
Plate Records
Prepare a new plate record for each plate to be run. Records differ slightly for 96
vs. 384 well plates. Get an example plate record from Paul or Amy. The
"container name" must be a new, unique name or the record will fail to import.
The plate record stores the name of each sample, the instrument protocol, and
the well position of each sample. See the example below. Sample names can not
contain spaces, tabs, periods, slashes, or unusual characters. Use Excel or a word
processor to prepare the plate record and save the file as a tab-delimited textonly file. Use your own name in the "owner" and "results group" fields. The
"instrument protocol" must be listed exactly as "gene_mapper_50_DS-33-600bp"
for dye set "G5" or "gene_mapper_50_DS-30" for dye set "D". For longer sized
fragments, use the G5 dye set and "gene_mapper_50_DS-33-1200bp". From the
"Plate Record" section of the ABI software, import your plate record.
Container Name
Plate ID
PlateNameHere
PlateNameHere
AppServer
AppInstance
GeneMapper
GeneMapper_Generic_Instance
Well
Sample Name
A01
Description
ContainerType
AppType
Owner
Operator
PlateSealing
96-Well
Regular
Paul
Paul
Septa
Size
Standard
Results Group
1
Instrument Protocol 1
Sample1
CASSUL5
Paul
gene_mapper_50_DS-30
B01
Sample2
CASSUL5
Paul
gene_mapper_50_DS-30
C01
Sample3
CASSUL5
Paul
gene_mapper_50_DS-30
D01
Sample4
CASSUL5
Paul
gene_mapper_50_DS-30
E01
Sample5
CASSUL5
Paul
gene_mapper_50_DS-30
F01
Sample6
CASSUL5
Paul
gene_mapper_50_DS-30
G01
Sample7
CASSUL5
Paul
gene_mapper_50_DS-30
H01
Sample8
CASSUL5
Paul
gene_mapper_50_DS-30
A02
Sample9
CASSUL5
Paul
gene_mapper_50_DS-30
B02
Sample10
CASSUL5
Paul
gene_mapper_50_DS-30
…
…
…
…
…
H12
SampleN
CASSUL5
Paul
gene_mapper_50_DS-30
SchedulingPref
1234
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Data Files
The data for each individual well in the plate will be output as a separate file. For
example, a 384 well plate will create 384 separate files. Each file contains the data
for all 5 dye colors for a given well. The files are named using the sample name
(as it appears in the plate record), the well position, the capillary, and the run
number. All files are placed on drive E:/ in a new folder named with the date
and the "container name" as listed in the plate record. It is very helpful to name
samples using the plant name, the primer used, and the dye or color used for
each primer.
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Runs and Well positions
The array has 48 capillaries. For a 96-well plate, injections are made from every
well in a column and skips 1 column. A full 96-well plate requires 2 runs (loads)
to inject all samples. See figure. A minimum number of samples in a plate is 48
and must be loaded in odd columns only. If you have fewer samples than 48, put
ddH20 in the empty wells.
For a 384-well plate, injections are made from every other well in a column and
skips 3 columns. See figure. A full plate of 384 sample requires 8 runs (loads) to
inject all samples once. A minimum number of samples in a plate is 48 and must
be loaded as below in the "First Quadrant, First Load" pattern. If you have fewer
samples than 48, put ddH20 in the empty wells.
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Tuesday, November 18, 2014. St. Amand
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Starting an ABI Run
Do not load a plate or plate record without assistance from either Amy or Paul.
Place your samples in the proper size carrier and place the carriers on the tray.
Insure that the carriers are fully seated and that all septa are in place. Close the
ABI tray stacker. In the "Run Scheduler" panel of the ABI software, press the
"Find All" button to locate your plate record. Click on the plate record to select it
then click on "Add". The run can then be started. A 96-well genotyping plate
takes 1.5 hours to run. A 384-well plate takes 6 hours to run. A 384-well plate
with GS1200 or CASSUL1200 takes 12 hours to run. A plate must be run within
48 hours once the sample is added to the formamide.
Errors or Problems
If the ABI stops during a run and presents an error message, use the "Print
Screen" button on the keyboard to copy the screen. Open the "Paint" application
and paste the graphic image in the file and save the file to the desktop. That way,
we can send the exact error message to the ABI technicians. You should then
alert Paul or Amy to the problem. If it is necessary to restart the computer or the
ABI, the proper procedure is:
1. Quit all 4 programs of the Data Collection Software
2. Shut down the computer
3. Press the "On" button on the front of the ABI to switch it off.
4. Restart or reboot the PC.
5. Log-in as "Administrator" with the password "WheatDNA"
6. Once log-in is complete, press the "On" button on the ABI.
7. Once the ABI has started and is showing a steady green light, start the "Data
Collection" Software.
Data Backup
All of your run data will be stored in your own run folder. It is YOUR
responsibility to copy each batch of data to your own folder on the back-up drive
(H:/). Your data may also be emailed to you. You should also copy the data to a
portable USB key drive and move it to another computer. Do NOT store data on
the computer connected to the ABI 3730. For safety reasons, the ABI 3730 is NOT
connected to the network or to the Internet. Do your data analysis on another
computer, not on the ABI 3730. Do not quit the ABI Data Collection Software. Do
not run any other software on the ABI 3730. Do not install any other software on
the ABI 3730. Do not insert disks or USB devices in the ABI computer.
ABI Maintenance
ABI maintenance is done by Amy or Paul. Once a week the PC and the ABI
should be rebooted, the buffer and water changed, and the water jacket flushed.
Once a month, the polymer bottle should be replaced with new POP-7. Every 500
to 1000 runs, the array should be changed. Every 4 to 6 months the air filter
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should be changed. If specific capillaries seem to be plugged, have high
background florescence, or lower signal intensity, then the capillary likely should
be changed. Every capillary change requires that a spatial calibration and a
matrix calibration be done.
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Data Analysis
We use "Gene Marker" software to analyze ABI data. This software can be
installed on any computer, but requires a USB license key in order to run. We
own 5 keys. The basic steps required to use this software are:
1. Add data files (File menu > Open Data). Add only those files produced using
the same set of primers.
2. Process files (Project menu > Run). Select "ABI-5-Color" template, the
appropriate size standard (usually GS500 or CASSUL), and "Fragment
Analysis(Plant)". Press the "Next" button and insure that the following items are
"checked" and on: Auto Range, Smooth, Peak Saturation, Baseline Subtraction,
Pull-up Correction, Spike Removal, and Local Southern. The following are useful
first-time options and may need to be adjusted for your data: Allele Call
start=50bp, end=600bp, Intensity=200, Global Percentage=0, Local
Percentage=25, Stutter Peak Filter=on, left=95, right=95. Press the "Next" button,
set Reject=0 and Check=0. Then press the "OK" button.
3. Save the project (File menu > Save Project). Save your project now and after
other steps because the program will occasionally lock-up and become
unresponsive.
4. Check the calibration. From the button bar choose "Size Calibration". Each
sample file will have a score. Those above 90 are usually sized properly. You
should examine each file and make sure that all of the expected size peaks are
properly marked with a green triangle. If they are not, control-click on the
triangle and move the triangle to the proper peak. Once all peaks have been
adjusted for that file, right-click and choose update calibration. Then move on to
other sample files. Close the calibration window and re-save the project.
5. View the data. Double-click on a file to view a trace of the data or press the
"Gel Image" button to see a representation of all of the samples.
6. View the report. Press the "Show Report" button to see which peaks were
identified. Change report options using the "Report Settings" button and the
"Show Color" button. The report only shows the currently selected colors.
7. Change allele calling options. If the report shows peaks that are misidentified
you will need to change the allele calling options and "re-call" the alleles. Press
the "Call Allele" button. Change the allele range and detection threshold options
to increase or decrease the allele calling sensitivity based on your data. You will
likely need to do this several times and then examine each peak and manually
correct some of the calls. After saving a report and PDF for each completed
primer, you will likely need to re-call alleles using different settings for other
primers. Re-calling alleles removes all previous calls for all primers.
8. Create a Panel. This is an optional, but usually necessary step. Open the panel
editor and create a new panel using the manual options. Create a new marker for
each color. Insert new alleles at the desired locations for each marker. Save the
panel. You must then "re-call" the alleles using this panel. The report will now
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only show defined alleles based on the panel used. You may also need to change
the allele calling options to refine called bands.
9. Output a report. Once the report is correct. Press the "Save Report" button to
export a text-only version of the report.
10. Print trace files. To save the graphic images of the trace data, choose the
printer icon and change the settings and print. We have a PDF printer installed
so that you may save PDF versions of the print file. Simply choose "Cute PDF
Writer" for the printer model.
11. Save gel image. To save the graphic gel image of the data, press the "Show
Gel Image" button and then the small "=>" icon near the image. You can then
export a graphic image of the gel.
12. Save your project and create a back-up to another location to prevent dataloss.
13. Evaluating peak data. When using fluorescent detection, the best results are
obtained when the allele peak heights are above 1,000 RFU and below 15,000
RFU. Peaks below 1,000 RFU may be usable, but the software has a harder time
doing auto scoring and small amounts of DNA contamination can produce
background peaks. Peaks below 200 RFU should NOT be used. Peaks larger than
15,000 RFU cause “dye pull-up” problems. This produces a false peak in the
other color channels. Also, the software may not score very large peaks well. If
your peaks are too large, load less of the pool into the formamide solution. If
your peaks are too small, try loading 50% more of the pool and also try loading
50% less of the pool. Too little sample can produce small peaks, but too much
sample can also produce small peaks.
14. Evaluating size standard peaks. ALWAYS examine the peak heights of your
size standard. It should never be below 100 RFU. It should average about 400
RFU. If it is above 1000 RFU, you are wasting size standard and you are
LOWERING the sample peak heights because of competition between the
standard and your sample.