This protocol is used for the detection of critically informative gene fusions for patient tumor samples. The fusion status of dozens of genes is assessed simultaneously in a single assay. The advantage of this technique is that I can identify gene fusions regardless of the identity of the fusion partner from tumor samples that have been processed by formulative fixation.
The detection of certain gene fusion in clinical tumor sample directly informs diagnosis, prognosis and treatment selection in many cancer types. It is critical to understand that many fusions called by the analysis algorithm are artifacts. The most difficult task when analyzing data is distinguishing between artifact and real fusion calls.
Begin by diluting total nucleic acid to achieve the desired concentration of RNA. For each sample, transfer 20 microliters of the dilution into the random priming reagent strip tubes in a pre-chilled aluminum block and mix by pipetting up and down six to eight times. Briefly spin down the samples, transfer the entire volume to a 96-well PCR plate, and seal with plate sealer film.
Insert the plate into a thermocycler, cover with the compression pad, and close the lid. Incubate at 65 degree Celsius for five minutes. After the incubation, transfer the entire volume of the random priming product in the first strand reagent strip tubes.
Mix by pipetting up and down and briefly spin down. Transfer the entire volume to a 96-well PCR plate and seal with RT film. Insert the plate into the thermocycler and run the reaction according to the manuscript directions.
Perform second strand cDNA synthesis, and repair, and ligation step one according to the manuscript directions and proceed to the second ligation step. To number the molecular barcode or MBC adapter strip tubes, position the tubes horizontally with hinges to the back and use a permanent marker. Take the bead purification plate from the first ligation step and transfer 40 microliters of each sample into the MBC adapter strip tubes, taking care not to disturb the bead pellet.
Mix the reagents by pipetting, spin down the tubes, and transfer the entire volume to the ligation step two reagent strip tubes. Mix the samples, spin them down, and place in a thermocycler block. Leave the heated lid off and run the thermocycler at 22 degrees Celsius for five minutes, followed by 4 degrees Celsius hold.
Next, prepare the ligation cleanup beads by vortexing them and adding 50 microliters to a new set of PCR strip tubes. Incubate on the magnet for one minute and discard the supernatant. Remove the strip tubes from the magnet and resuspend the beads in 50 microliters of ligation cleanup buffer by pipetting up and down.
Transfer the entire sample volume from the second ligation step into the ligation cleanup bead strip. Vortex the samples to mix and leave them at room temperature for 10 minutes. Vortex the sample once halfway through the incubation.
Afterwards, vortex the samples, spin them down, and incubate on the magnet for one minute. Discard the supernatant and add 200 microliters of fresh ligation cleanup buffer. Vortex to resuspend, spin down, and place on the magnet for one minute.
After the two washes, perform a third wash using ultrapure water instead of buffer. Resuspend the beads in 20 microliters of 5 millimeters sodium hydroxide and transfer the samples to a 96-well PCR plate. Place the plate into a thermocycler with a compression pad and run it at 75 degrees Celsius for 10 minutes, followed by a 4 degrees Celsius hold.
Once the samples have cooled to 4 degrees Celsius, place the plate on a magnet for at least three minutes and proceed with the first PCR. Prepare the PCR reactions by adding two microliters of GSP1 primers to each well of the first PCR reagent strip. Transfer 18 microliters of the second ligation cleanup product to the first PCR reagent strip tubes and mix by pipetting up and down.
Spin down the samples and transfer them to a 96-well PCR plate. Place them into the thermocycler and run PCR according to the manuscript directions. Proceed with bead purification by adding 20 microliters of the PCR product to a U-bottom plate filled with 24 microliters of purification beads per well.
Pipette up and down to mix and leave the plate at room temperature for five minutes, then incubate the plate on the magnet for two minutes. Discard the supernantant from the beads and wash them twice with 200 microliters of 70%ethanol. After the final wash, remove all of the ethanol and let the sample air dry for two minutes.
Remove the plate from the magnet and resuspend the beads in 24 microliters of 10 millimeter Tris-HCl and pH 8.0. Incubate off the magnet for three minutes and then place the plate back on the magnet for two minutes. Begin library quantitation by preparing one to five dilutions of the second PCR product in 10 millimeter Tris-HCl.
Follow this by a serial dilution of 1:199, 1:199, and 20:80 in 10 millimeter Tris-HCl with 05%polysorbate. Set up key PCR by adding six microliters of master mix to each well of a optical 96-well plate, followed by four microliters of the appropriate dilution or standard. Spin down the plate and load it into the qPCR instrument.
Perform qPCR according to the manuscript directions. After library quantification is complete, dilute all libraries to two nanomolar with 10 millimeter Tris-HCl. Make a library pool by combining 10 microliters of each normalized library into one 1.5 milliliter microcentrifuge tube.
Next, prepare the denatured amplicon library, or DAL pool, by combining 10 microliters of the library pool with 10 microliters of 0.2 normal sodium hydroxide and incubating the mixture for five minutes at room temperature. After the incubation, add 10 microliters of 200 millimeter Tris-HCl at pH 7.0, followed by 970 microliters of HT1 Hybridization Buffer. Make the final load tube by combining 300 microliters of HT1, 25 microliters of 20 picomolar PhiX, and 675 microliters of the DAL pool.
Add the entire volume of the load tube to the sample well of the sequencer reagent cartridge and load the cartridge in to the sequencer. Start by selecting the positive control sample and ensure that all expected fusions and oncogenic isoforms have been detected and are listed in the strong evidence tab. For each sample, inspect the average unique start sites per GSP2 control value, then visualize the supporting reads for each potential fusion by clicking the Visualize link which takes the user to a web-based JBrowse view of pileups of individual fusion supporting reads.
Confirm that the reads are mostly free of mismatch, but more than 30 bases of the reads align with the fusion partner, and that sequences adjacent to the breakpoint between the genes and primer binding sites are free of insertions or deletions. It is critical to ensure that every call fusion is generally free of misalignment and that a significant portion of the reads align to the fusion partner. This protocol has been used to investigate gene fusion status in a lung adenocarcinoma sample.
The summary shows strong evidence fusions and the Read Statistics page shows metrics of the sample. This sample exhibits good RNA quality so negative results would be reported if no fusions were found. A legitimate fusion call has a high number of supporting reads, a high percent of reads from the primer supporting the fusion, and a high number of start sites.
Visualization of the reads demonstrates good alignment to large regions of the fusion partner. Conversely, an artifactual fusion call has lower supporting metrics and a high error rate when mapping to the partner. Artifactual fusion calls made by the analysis algorithm are common.
It is critical to manually inspect every call to ensure that it represents a real gene fusion in the patient sample.
