CSI TRU SAMPLE PLATE PREPARATION
Using the Quantifiler® Human DNA Quantification Kit
You will need:
Your samples
Quantifiler® Human DNA Quantification Kit
Set of 8 serially diluted standards (prepared and in freezer)
T10 E0.1 Buffer (your Elution Buffer)
One 96-Well Plate
Optical Adhesive Seal
Adhesive Roller
Preparation:
Vortex your samples, standards and Quantifiler® Human DNA primer mix
Flick the Quantifiler® reaction mix and the T10 E0.1 Buffer
1.
2.
3.
4.
5.
6.
Remove your DNA extracts, standards and the Quantifiler® Human DNA
Quantification Kit from the freezer to thaw.
While thawing, load the protocol and plate files onto the Bio-Rad operating
system and rename the samples as outlined in the Bio-Rad Operation Protocol.
Use fresh gloves through the rest of the process.
Prepare your PCR Master Solution as follows (include additional reactions in
your calculations to allow excess volume for the loss that occurs due to
reagent transfers):
a. In labeled 1.5 microtube(s) aliquot as follows:
i. Quantifiler® Human Primer Mix = (#of samples + 3) x 10.5 µL
ii. Quantifiler® Reaction Mix
= (#of samples + 3) x 12.5 µL
iii. Flick the PCR Master Solution to mix
iv. If the final volume of your PCR Master Solution will be greater
than 1.5 mL use two 1.5 mL microtubes and split the volumes
evenly while preparing the solution.
v. Centrifuge the tube toward the end dispensing to maximize reagent
recovery and consolidate the tubes.
With gloved hands lay down clean bench paper and place your sample plate
on the table with well #A-1 in the upper left hand corner – Handle the plate
using the lipped edges to avoid putting any kind of mark or film on the wells
that may interfere with the optical detector.
Your plate should be set up as follows to exactly reflect the BioRad plate
setup:
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A
1
2
3
4
5
6
7
8
9
10
11
12
Empty
STD 1
STD 2
STD 3
STD 4
STD 5
STD 6
STD 7
STD 8
NC
Blank
Empty
B
001 S
010 S
C
011 S
020 S
D
021 S
E
026 S
035 S
F
036 S
045 S
G
046 S
025 S
050 S
H
a.
b.
c.
d.
7.
8.
Standards in wells A2 through A9
Negative Control (Quantifiler® plus buffer) in well A10
Blank (just buffer) in well A11
Samples in the rest of the wells as shown in the example (two people per
plate)
e. Adjust the sample wells accordingly for the number of donors you will be
running (20 or 25 donors) and be sure that your sample plate exactly
matches your BioRad plate setup (see Tips Section for suggestions on how
to avoid mistakes). Since two students will run samples on one plate pay
close attention to loading the plate so you do not cross contaminate
samples.
Your wells will contain a total volume of 25 µL. Load your plate as follows
(see Tips section on how to avoid pipetting errors and bubbles):
a. Aliquot 23 µL of the PCR Master Solution into
i. each of the eight standard wells
ii. the negative control well
iii. each of the sample wells
iv. DO NOT aliquot any master mix into the Blank well
b. Aliquot 2 µL of
i. each of the standards into the corresponding wells (change tips)
ii. T10E0.1 buffer into the negative control
iii. each sample into the corresponding wells
c. Aliquot 25 µL of T10E0.1 buffer into the Blank well
Seal the well plate with the optical adhesive using the roller to ensure a proper
seal. DO NOT use a bare or gloved hand to press the optical adhesive to the
plate. This may put marks on the optical seal that will interfere with the
optical detector response.
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9.
10.
Load your plate into the BioRad iCycler making sure that well A1 is in the
upper left hand corner. Note: the lower left hand corner of the plate is
notched.
Operate the iCycler according to the written protocol.
TIPS:
Plate Set Up:
1.
2.
3.
4.
5.
Graph your plate setup on paper according to the BioRad plate setup.
Double check the well designation before you aliquot the standard or sample
to ensure your actual plate will exactly match your BioRad plate setup.
As you aliquot each well cross it off from the graph so you know the well is
complete.
Use two of the sample racks that are labeled similarly to the plate.
a. In the first rack, set your standard and sample tubes up to match your plate
setup.
b. As you aliquot into the wells move the used standard/sample to the second
rack.
c. This will help you identify which standards/samples have been aliquoted.
The key to avoiding mistakes is to TAKE YOUR TIME.
Pipetting:
These pipettes are very touchy. The trick to improved accuracy is to pipette slowly and
gently.
1.
2.
3.
4.
Always double check the volume on your pipette before placing a tip on it.
Always change your tip between standards and samples to avoid cross
contamination.
Very lightly, gently and slowly depress/release the plunger on the pipette
(especially on the smaller pipettes) to ensure consistent volume draw and
delivery.
To avoid bubbles (interferes with the reaction and the optical detector):
a. Slowly deliver the Master Mix starting at the bottom-side of the well and
sliding the tip up the well as you deliver. However, before aspirating the
pipette raise the tip to just above the surface of the total volume in the
well, then aspirate.
b. Deliver the 2 µL of standard/sample placing the pipette tip just above the
volume of Master Mix and depress the plunger to the second stop (fully
aspirate). If the 2 µL of standard/sample hangs on the end of the tip, touch
it to the Master Mix to get it to release.
c. If you still get bubbles:
i. Set the pipette to draw the entire contents of the well.
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ii. Very slowly redeliver the total volume into the same well while
bringing the tip up the side of the well
iii. We do not have a plate centrifuge so you will have to repeat this
process until no bubbles appear in the well.
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Updated: 19JAN08