1. Preparation of target-specific primer working stocks


For multiplexed Illumina sequencing-by-synthesis, each sample must be uniquely tagged with molecular barcodes to extract the reads per sample out of the pool of sequence reads. These tags are added using a 4-primer PCR system developed by Fluidigm. We order fusion primers containing our MHC class I and class II sequence-specific PCR primers prefaced with Fluidigm Consensus Sequence (CS) linkers. Barcodes are then added via commercially available plates of outer primers ordered from Fluidigm that contain the complements to the CS linkers on 384 different barcode sequences. Prepare 6 total 8-well PCR tube strips with the following primers (Excel version of primer layout can be found here):

Pasted Graphic
Combine primers into FXR pairs shown above and dilute to 1 µM working stocks:
From 100 µM stocks – 2 µl forward and 2 µl reverse primers into 196 µl 0.1X TE buffer (or prepare a large stock of each of the 10 FXR primer pairs, then distribute across the appropriate wells of the PCR tube strips).

2. PCR setup with the Fluidigm Access Array system


2.1 Prepare template gDNA


(a) Normalize gDNA to 60 ng/µl.

2.2 Prime the 48.48 Access Array IFC


(a) Use a closed tip syringe to press down on accumulator openings to loosen springs.
(b) Hold the chip at a 45° angle, with accumulator being injected pointed towards the ground.
(c) Pull the syringe plunger back and hold in position to retain negative pressure. Remove the cap.
(d) Push the tip through the accumulator opening and ensure black o-ring has moved to the side so tip is between the o-ring and the tank; inject ~300 µl Control Line Fluid into accumulator.
(e) Turn chip so other accumulator is pointed downwards and repeat injection process.
(f) Add 500 µl of 1X Harvest Solution into the H1-H4 wells on the chip.
(g) Remove blue protective film from bottom of the chip; load chip into the IFC Controller (press “Eject” to open tray, put chip onto tray with notched corner of chip aligned with A1 mark, press “Load Chip” to close tray/register barcode/activate script selection).
(h) Select “Prime (151x)” and “Run Script” to prime the chip.

2.3 Prepare the primer and sample mixes


(a) Prepare working inner primer dilutions in a 96-well plate according to the Fluidigm mixes sheet instructions (per primer well: 9.5 µl of 1 µM target-specific primer mix (F+R), 0.5 µl 20X Loading Reagent - put ~5.5 µl/tube (8 tubes) Loading Reagent into strip-cap tubes to multichannel).
(b) Prepare sample master mix in a 1.5 ml tube according to the mixes sheet instructions (per sample: 3.75 µl Phusion HS Master Mix, 0.375 µl Loading Reagent).
(c) Vortex mix for 20 s, then spin in a microcentrifuge for 30 s.
(d) Add 4.1 µl of sample master mix to each sample well of the 96-well plate.
(e) Add 1.5 µl of Barcode Library outer primer to each sample well of plate, following sample layout in mixes sheet.
(f) Add 1.9 µl of DNA to each sample well of the plate, following sample layout in the mixes sheet.
(g) Seal sample/primer plate.
(h) Vortex plate for 20 s, then centrifuge for 30 s.

2.4 Load the 48.48 Access Array IFC


(a) Pipet 4 µl of primer mix into each primer inlet (right side of array – primers pivot).
(b) Pipet 4 µl of sample mix into each sample inlet (left side of array – samples straight).
(c) Load the chip into the IFC Controller (press “Eject” to open tray, put chip onto tray with notched corner of chip aligned with A1 mark, press “Load Chip” to close tray/register barcode/activate script selection).
(d) Select “Load Mix (151x)” and “Run Script” to load samples and primers into the chip.
(e) Once the load script is complete, press “Eject” to remove chip and take it to post-PCR to cycle.

2.5 Thermal Cycling the 48.48 Access Array IFC


Thermal Mix:
50°C for 120 s
70°C for 1200 s
Hot Start Denaturation:
98°C for 120 s
PCR Cycle (10X):
98°C for 10 s
60°C for 30 s
72°C for 20 s
C0t Cycle (2X):
98°C for 10 s
80°C for 30 s
60°C for 30 s
72°C for 20 s
PCR Cycle (8X):
98°C for 10 s
60°C for 30 s
72°C for 20 s
C0t Cycle (2X):
98°C for 10 s
80°C for 30 s
60°C for 30 s
72°C for 20 s
PCR Cycle (8X):
98°C for 10 s
60°C for 30 s
72°C for 20 s
C0t Cycle (5X):
98°C for 10 s
80°C for 30 s
60°C for 30 s
72°C for 20 s
Final Extension:
72°C for 300 s
Cool Down:
4°C for 10 s

2.6 Harvest the 48.48 Access Array IFC


(a) Remove the remaining Harvest Reagent from H1-H4 wells.
(b) Add 600 µl fresh Harvest Reagent into H1-H4 wells.
(c) Pipet 2 µl of Harvest Reagent into each sample inlet on chip.
(d) Load the chip into the IFC Controller (press “Eject” to open tray, put chip onto tray with notched corner of chip aligned with A1 mark, press “Load Chip” to close tray/register barcode/activate script selection).
(e) Select “Harvest (151x)” and “Run Script” to harvest samples into sample inlets.
(f) Once the script is complete, press “Eject” to remove chip.
(g) Transfer 10 µl of harvested PCR product from each sample inlet into 6 columns of a 96-well plate.
(h) Optional: check a subset of samples for primer dimer on a Bioanalyzer.
(i) Combine equal volumes of each sample into a 1.5 ml tube to create a pool for purification and sequencing.

3. Ampure XP (Agencourt) bead purification of amplicon pool


We use two rounds of bead cleanup to remove short product contaminants from our amplicons:

(a) Calculate the total volume of the amplicon pool. Into the tube containing the pool, pipet 1.3X the pool volume of Ampure XP beads (for example, add 124.8 µl of Ampure XP beads to a 96 µl pool). Pipet up and down ten times to mix.
(b) Incubate for 5 min at room temperature.
(c) Place tube on a tube magnet stand for 2 min.
(d) Being careful to avoid disrupting the bead pellets, remove all supernatant and discard.
(e) Rinse bead pellets with 950 µl 70% ethanol.
(f) Incubate for 30 s, discard supernatant.
(g) Repeat (f) for a second wash with 950 µl of 70% ethanol.
(h) After completely removing supernatant, remove plate from magnet and allow 2-4 min to dry (it's okay if it doesn't dry completely).
(i) Add 100 µl 1X TE buffer to elute purified product and pipet mix 10 times. OPTIONAL: during the second elution, you can concentrate the pool by eluting in <100 µl 1X TE. This is useful when one pool is expected to be much less concentrated than other pools in the same MiSeq run.
(j) Incubate for 3 min.
(k) Place tube on magnet for 2 min (on second clean-up, proceed to step (n) after beads are pelleted).
(l) Pipet (100 µl*1.3=) 130 µl of Ampure XP beads into a new 1.5 ml tube. Transfer the eluted, purified product from the first tube into this tube and pipet mix 10 times.
(m) Repeat steps (b)-(k) for second clean-up.
(n) Transfer eluted products to a new 1.5 ml tube for storage/sequencing.

4. Quantification of amplicon pool concentration


(a) Use the Qubit dsDNA High Sensitivity Assay Kit (Life Technologies).
(b) Make a master mix containing:
   199 µl Qubit HS buffer * (# of pools + 2 standards)
   1 µl Qubit HS dye * (# of pools + 2 standards)
(c) Vortex and aliquot 190 µl of master mix into two Qubit tubes and 198 µl of master mix into each sample Qubit tube.
(d) Add 2 µl of pool into each sample tube, and 10 µl of the standards (10 ng/µl and 0 ng/µl) into the standard tubes.
(e) Vortex the tubes for 5 s. Tap/flick tubes to allow any bubbles to escape.
(f) Measure the concentration of the sample tubes in a Qubit fluorimeter. Convert to sample concentrations multiplying by the conversion factor 0.1 (as per the this calculation):
   conc (ng/ml) * 200 µl/2 µl * 1 ml/1000 µl = conc * 0.1 ng/µl

5. Sequencing on the Illumina MiSeq


(a) Thaw a MiSeq reagent cartridge at RT.
(b) Generate a 2 nM stock of each sample using the Illumina copy number spreadsheet, along with the Qubit concentration and a DNA fragment size of 386bp.
(c) Pipet 2 µl of 2 nM stock from each sample into a 1.5 ml tube to create an undiluted amplicon pool.
(d) Pipet 10 µl of the DNA pool into a fresh 1.5 ml tube. Add 10 µl of 0.2 N NaOH solution to the DNA.
(e) Vortex briefly, then centrifuge for 1 min at 280 x g.
(f) Incubate for 5 min at RT to denature the DNA.
(g) Add 980 µl pre-chilled HT1 to the denatured DNA to generate a 20 pM denatured library.
(h) Transfer 600 µl of the denatured library to a new 1.5 ml tube, then add 400 µl of pre-chilled HT1 to dilute the denatured library to 12 pM.
(i) Invert several times to mix, then centrifuge briefly to get any droplets off of the cap. Place on ice.
(j) Pipet 2 µl of 10 nM PhiX library into a fresh 1.5 ml tube. Add 8 µl of 10 mM Tris-Cl (pH 8.5 with 0.1% Tween 20) to create a 2 nM dilution of PhiX library. PhiX is used as an internal control for sequencing.
(k) Combine 10 µl of 2 nM PhiX library with 10 µl of 0.2 N NaOH solution in a 1.5 ml tube to create a 1 nM PhiX library.
(l) Vortex briefly, then centrifuge for 1 min at 280 x g.
(m) Incubate for 5 min at RT to denature the PhiX library.
(n) Add 980 µl pre-chilled HT1 to the denatured PhiX library to generate a 20 pM PhiX library. Note: The denatured 20 pM PhiX library can be stored up to 3 weeks at -20°C.
(o) Transfer 12 µl of 20 pM PhiX library to a new 1.5 ml tube, then add 8 µl of pre-chilled HT1 to dilute the PhiX library to 12 pM.
(p) Combine 990 µl of denatured library (12 pM) and 10 µl of PhiX library (12 pM) to create a final pool for sequencing. Place on ice.
(q) Use an empty 1 ml pipet tip to pierce the foil seal of the MiSeq reagent cartridge over the reservoir labeled “Load Samples”.
(r) Pipet 600 µl of the final pool from step 16 into the “Load Samples” reservoir. Avoid touching the foil seal while dispensing the sample.
(s) Follow the onscreen prompts on the Illumina MiSeq instrument to set up and perform the sequencing run.