Unit I – HLA-DQ Alpha Assay
Reading:
Handouts; Online Protocols and Lectures
March 30
Class basics
Lecture:
Background for HLA DQα
Lecture:
Cell extract
Lab check-in
Lecture:
Basic concepts of molecular assays
Preparation of your genomic DNA from buccal cells
April 1
Lecture: PCR
Set up and run PCR
Lecture: Immunology I
A fraction of your PCR reaction will be run on a gel and documented by the Instructional Support
Technician, Daisy Chau, for us to analyze. You will get to run your own agarose gels for Units III
and IV.
April 6
Analysis of gel results
Lecture: column cleanup of PCR products
Lecture: Spectrophotometric analysis to determine DNA concentration
Column cleanup of PCR products and spectrophotometric determination of
concentration
Lecture: Immunology II
April 8
Lecture: Sequencing
Set up and run sequencing reactions
Lecture: Continue Immunology II
April 13
Dry down samples and load on sequencer
Lecture: Immunology III
Closing lectures
April 15
Analyze sequencing results
Pointers on writing papers
Begin Unit II – Mock HIV ELISA
Introduction of SOP for ELISA
Protocols:
Preparation of Extract Containing Genomic DNA from Cheek Cells
HLA-DQalpha Assay
PCR amplification
Check gel: Agarose Gel Electrophoresis: Casting and Running the Gel
Column clean-up of PCR products
Spectrophotometric determination of PCR product concentration
Sequencing
Analysis of Sequencing Results
Presentation of Sequencing Results
Sequencing alignment
Chromatogram – location of sites with two base calls
Comparison of your sequence to haplotype chart
Unit I: HLA-DQα Assay
Preparation of Extract Containing Genomic DNA from Buccal (Cheek) Cells
First and foremost, it is necessary to have a sample of your nuclear DNA, which contains
within it the sequence of DNA you wish to amplify. Your nuclear DNA is the template for
the PCR reaction because it provides the pattern of base sequence to be duplicated. The
source of DNA is a sample of several thousand cells from your cheek, obtained by saline
mouthwash. The cells are collected by centrifugation and resuspended in a solution
containing the resin "Chelex," which binds metal ions that inhibit the PCR reaction. The
cells are lysed by boiling and centrifuged to remove cell debris and the Chelex beads. A
sample of the supernatant containing DNA from the nucleus will be mixed with the other
reagents for the PCR reaction.
Materials to be provided
Microcentrifuge for 1.5 ml tubes and with adaptors for PCR-sized tubes
Thermocycler set at 99oC
Thermocycler position grid sheet
Vortexers
Labeled storage box for storage of collected, labeled, individual DNA samples.
For each student:
200 microliters 5% Chelex in a PCR tube (the suspension must be visually inspected after
the beads have settled to be sure each student has the same total volume and
volume of beads)
0.9% saline – 10 ml in a sterile paper cup
1.5 ml microfuge tubes (2)
For each group of 4 students:
150 ul 0.9% saline in a microfuge tube
Permanent marker
Procedures
PCR Amplification
Materials
Thermocycler
Barrier pipet tips
PCR tubes
Hot Start Taq
Master Mix
Primer Mix
DNA samples from previous period
Ultrapure or distilled water
These instructions are written for one group of four students.
Use barrier tips and wear gloves for all pipetting related to PCR.
1.
Label six 0.2 ml reaction tubes numbers 1-6.
They will be used for the following six reactions:
1) Student #1’s DNA
2) Student #2’s DNA
3) Student #3’s DNA
4) Student #4’s DNA
5) Positive Control DNA (commercially prepared human genomic DNA)
6) Negative Control – ultrapure or double distilled water
Be sure to make a record of who within your group of 4 is student #1, #2, etc.
Use barrier pipet tips for the following steps.
2.
Pipette 25μl of Master Mix (2X) into each tube.
3.
Pipette 15μl of Primer/MgCl2 Mix into each tube. (Primers 3L and 2R)
4.
Pipette 10μl of each template into correspondingly labeled tube.
Note: final concentrations of most reaction components are found on a lecture slide.
PCR Program
Step 1: 95oC for 2 minutes (to activate the Taq)
Step 2: 94oC for 30sec.
Step 3: 56oC for 30sec.
Step 4: 72oC for 30sec.
Repeat step 2 to step 4 for total of 40 cycles.
Step 5: 72oC for 10 minutes
Hold at 4oC forever.
Agarose Gel Electrophoresis
(to be done for you)
Supplies
gel casting tray
tape
gel comb
electrophoresis unit
power supply
microwave oven
125 ml flasks
p10 or p20, p100 or p200 pipettors
pipet tips
25 or 50 ml graduated cylinder
gloves
VersaDoc imager
Reagents
1.
2.
3.
4.
5.
ethidium bromide (10 mg/ml) Wear gloves. This reagent is a mutagen.
agarose – 2 kinds – Low Melt NuSeive and High Melt SeaKem (or equivalent)
1X SB electrophoresis buffer: 5 mM sodium boric acid, pH 8.5
5X gel loading buffer
1.5 g Ficoll 400
25 mg bromphenol blue
25 mg xylene cyanol
dH20 to a final volume of 10 ml
molecular weight marker (100 bp ladder), aliquotted for each team
Procedures
Pouring the gel
Before starting, be sure all parts of your gel unit fit together properly.
1.
Prepare the gel casting tray by sealing the ends with tape.
Position the comb at one end of the tray.
2.
Measure 50 ml 1X SB into a 125 ml flask.
3.
Weigh out and add 0.75g NuSieve agarose and 0.25 g SeaKem agarose to the SB
in the flask. Set the flask on a level surface and carefully mark the level of the buffer
with a black marking pen.
4.
Microwave the flask until the SB is boiling and the agarose is completely dissolved
(about 2 minutes). After the liquid first boils, you will need to swirl and reheat several
times before all the agarose is in solution. When all the agarose is in solution,
check the level of liquid. If necessary, add additional dH20 to return the solution to
the level you marked on the flask.
5.
Cool the agarose solution by holding the flask under running water and swirling. The
flask should be just cool enough that you can hold the bottom in your hand without
severe discomfort.
6.
Add 2.5 ul of the ethidium bromide solution (10 mg/ml) to the agarose solution.
Because ethidium bromide is a mutagen, wear gloves in this and in all subsequent
steps in which you touch the gel. Swirl the flask gently to avoid forming bubbles.
7.
Pour the agarose solution into the gel casting tray. Allow the gel at least 30 minutes
to cool and harden.
Running the gel
Be sure to wear gloves. The ethidium bromide in the gel is a mutagen!
1.
Remove and add to fresh, appropriately labeled tubes, 10 µl of each of your PCR
reactions. Freeze the remainder of each of the reaction volumes for cleanup to use
for sequencing.
2.
Add 2 µl gel loading buffer to each of the 10 µl samples and to the molecular weight
marker. Mix well.
3.
Set your electrophoresis apparatus up over absorbent paper, absorbent side up.
Pour a little running buffer (without ethidium bromide!) over the hardened gel.
Remove the comb from the hardened gel by pulling slowly, straight up. Position the
gel in the gel box so that the wells are at the black electrode. (DNA has a negative
charge and will move towards the positive, red electrode during electrophoresis.)
4.
Check with your instructor before continuing. Add running buffer (1X SB + ethidium
bromide, 5 µl EtBr/100 ml buffer) to the gel box, covering the gel to a depth of
about 2 mm. Be careful not to overfill the gel box.
5.
For each of your samples, load as much as you can into one well. Be sure to note
the order in which you have loaded your samples.
6.
In the last lane, load the molecular weight marker.
7.
Connect the lid and the power leads to the power supply. Make sure that the DNA
will be running toward the positive (red) electrode by connecting the lead at the
bottom of the gel to the outlet marked positive. If you have followed the color
coding, everything red will be connected and everything black will be connected,
and your wells will be near the “black” electrode.
8.
Set the power supply for 150 volts and run the electrophoresis until the darker
dye is about 1 inch or 2 cm from the bottom of the gel. Don’t let the gel run
too long.
9.
After the gel has run, remove the gel tray, slide the gel onto the VersaDoc, and
make a digital image of the gel.
Column Cleanup of PCR Products
The protocol below is adapted from the instructions provided by QIAgen, Valencia, CA, the
manufacturer of the kit.
QIAquick PCR Purification Kit Protocol
using a microcentrifuge
This protocol is designed to purify single- or double-stranded DNA fragments from PCR and other enzymatic reactions.
Fragments ranging from 100 bp to 10 kb are purified from primers, nucleotides, polymerases, and salts using QIAquick
spin columns in a microcentrifuge.
Important points before starting
■
The yellow color of Buffer PBI indicates a pH ≤7.5.
■
Add ethanol (96–100%) to Buffer PE before use (see bottle label for volume).
■
All centrifugation steps are carried out at 17,900 x g (13,000 rpm) in a conventional tabletop microcentrifuge at
room temperature.
Procedure
1.
Add 5 volumes of Buffer PBI to 1 volume of the PCR sample and mix.
For example, add 500 μl of Buffer PBI to 100 μl PCR sample.
2.
Check that the color of the mixture is yellow (similar to Buffer PBI without the PCR sample).
If the color of the mixture is orange or violet, add 10 μl of 3 M sodium acetate, pH 5.0, and mix. The color of the
mixture will turn to yellow.
3.
Place a QIAquick spin column in a provided 2 ml collection tube.
4.
To bind DNA, apply the sample to the QIAquick column and centrifuge for 60 s.
To maximize binding of DNA to the column, take the flow-through, reapply it to the column, and
centrifuge again for 60 s.
5.
Discard flow-through. Place the QIAquick column back into the same tube.
Collection tubes are re-used to reduce plastic waste.
6.
To wash, add 0.75 ml Buffer PE to the QIAquick column and centrifuge for 60 s.
7.
Discard flow-through and place the QIAquick column back in the same tube.
Centrifuge the column for an additional 1 min.
IMPORTANT: Residual ethanol from Buffer PE will not be completely removed unless the flow-through is
discarded before this additional centrifugation.
8.
Place QIAquick column in a clean 1.5 ml microcentrifuge tube.
9.
To elute DNA, add 30 μl elution buffer (Buffer EB,10 mM Tris·Cl, pH 8.5) to the very center of the
QIAquick membrane, let the column stand for 1 min, and then centrifuge.
IMPORTANT: Ensure that the elution buffer is dispensed directly onto the QIAquick membrane for complete
elution of bound DNA. The average eluate volume is 28 μl from 30 μl elution buffer. Elution efficiency is
dependent on pH. The maximum elution efficiency is achieved between pH 7.0 and 8.5.
How to Measure DNA Concentration Using the NanoDrop Spectrophotometer
1) Log into Windows (L: nanodrop / PW: bs365)
2) Double click the icon labeled “ND-1000 3.3.0” on the desktop
3) Login under your Lab
4) Lift the sampling arm up and remove the foam spacer
* DO NOT LIFT THE ARM BY THE FIBER OPTIC CABLE *
5) Moisten a Kim-Wipe with a small amount of double distilled water and wipe the upper (on the
bottom of the arm) and the lower (on the nanodrop) pedestals.
6) On the main menu, click on the upper left button labeled Nucleic Acid.
7) You should see the following message:
8) Load 2µl of double distilled water onto the lower measurement pedestal, close the sampling
arm, and press down the sampling arm lightly to make water(blank or sample-later) have
connection in between. Then click OK.
9) When the message Initializing Spectrophotometer – Please Wait disappears, the NanoDrop
is ready for use.
10) Using a dry Kim Wipe, wipe the lower pedestal.
11) On the right hand side of the monitor display, make sure the sample type is set to DNA-50.
12) Load 2µl of your blank onto the lower pedestal and click on the button labeled Blank. A
straight line will appear on the screen.
13) Wipe the upper and lower pedestals with a Kim Wipe.
14) You are ready to measure your DNA sample.
15) Load 2µl of your sample onto the lower pedestal and type in a sample name inside the box
labeled Sample ID.
Make sure the sample has formed a bridge between the lower pedestal and the sampling
arm before measurement!
And click Measure
16) If you have more than one sample, repeat steps 13 – 15.
17) When you’ve measured all your samples, you can view your results by pressing Show Report
or printing your results by pressing Print Screen or Print Report. You should print the scan result
for each sample immediately after you have measured it.
18) When all measurements are complete, wipe the upper and lower measurement pedestals with
a Kim-Wipe moistened with double distilled water.
19) Replace the foam spacer, lower the sampling arm, exit the ND-1000 program, and log out of
Windows.
Sequencing
Each individual will sequence his/her own PCR product in both directions (2 reactions per student),
using both a 5’ primer and a 3’ primer in separate reactions.
One member of the class will also sequence one of the control DNA PCR products.
The instructions below are adapted from the instructions for the Applied Biosystems Big Dye
Terminator v3.1 Cycle Sequencing kit.
You will set up your sequencing reactions in PCR tubes. Before you start, make the following
changes in the table below:
Terminator Ready Reaction Mix, Quantity 4.0 μl
Enter the volume for the Primer, to be given in class.
Total Volume 10 μl
Enter a volume of template that will be within the correct range of mass, and will not result in a
Total Volume of more than 10 μl.
If necessary, use ultrapure or double distilled water to make a total volume of 10 μl.
The thermocycler will be set to:
96C – 3 min
33 cycles of:
96C – 10 secs
50C – 15 secs
60C – 4 minutes
4C forever.
Cleanup of DNA sequencing products.
We will be cleaning up our DNA using DyeEx columns from QIAgen.
The samples will be dried down in a SpeedVac, redissolved in 10μl formamide, and loaded on the
sequencing analyzer.
We will be analyzing the sequencing reaction on an ABI 3100 Avant Sequencer by capillary
electrophoresis.
If you go to the URL below and click in the right column on the link under “Demos and Tutorials,”
you will see a video on the analyzer.
https://products.appliedbiosystems.com/ab/en/US/adirect/ab?cmd=catNavigate2&catID=600527&t
ab=Literature