TITLE: GT-seq protocol EFGL
AUTHORS: EagleFish GeneticsLab

[DESCRIPTION]
The purpose of this protocol is to prepare extracted samples of DNA to be read on a NextSeq 2000. This will be achieved through  SNP-PCR, barcoding, SequalPrep Normalization, Ampure bead size selection, and Qubit quantification. This will result in a single pooled vial of DNA ready for the NextSeq 2000 EFGL Loading Protocol. 


[STEPS]
SECTION: SNP-PCR
1. Prepare SNP-PCR master mix using species specific primer panel. 


 

SECTION: SNP-PCR
1.1. Vortex and spin down aliquots of Qiagen Multiplex Master Mix and panel specific GT-seq primers, keep them in benchtop coolers. The table has been formulated to account for pipette errors. 

 

Combine the ingredients in a non- Lo-bind tube. Vortex and spin down tube and keep in benchtop cooler till ready to dispense. 
SECTION: SNP-PCR
1.2. Label unskirted PCR plate(s): SNP-PCR, date, tray#, project initials, and your initials.
SECTION: SNP-PCR
2. Remove the DNA tray(s) from fridge. Briefly vortex and spin down   DNA trays. 
SECTION: SNP-PCR
2.2. Using a p10 multichannel pipette, add 2 µL of DNA into corresponding well. Discard tips after each use.
SECTION: SNP-PCR
3. Run the SNP-PCR plate(s) on a thermalcycler using the following program. (In our lab, the program can be found using the menu options GTseq --> gs-fxtd+10c-65)

1. 95 °C for 15 min
2. 95 °C for 30 s
3. 57 °C for 30 s
4. 72 °C for 2 min
Repeat 2-4 for 5 cycles, with a 5% ramp
5. 95 °C for 30 s
6. 65 °C for 30 s
7. 72 °C for 30 s
Repeat 5-7 for 10 cycles
8. 4 °C indefinitely

SECTION: SNP-PCR
3.1. Heat seal the DNA tray(s), and stored them back in the fridge. 
SECTION: Barcoding
4. Barcoding amplicons with i5s and i7s. 


 

SECTION: Barcoding
4.2. Pour lab-grade water into a 50mL reservoir. Using a p200 multichannel pipette, dispense 133ul of lab grade water into each well. 
 

SECTION: Barcoding
4.3. Heat seal diluted SNP-PCR tray(s). Vortex well, and centrifuge 3000 rpm, 30 s 

SECTION: Barcoding
5. Make barcoding master mix. 

IMPORTANT! Each DNA tray needs to have an unique i7, if they will be loaded on the same library. 

SECTION: SNP-PCR
2.1. Using a 0.1mL combi-tip, dispense 5 µL of PCR master mix into each well.
SECTION: SNP-PCR
2.3. Heat seal the SNP-PCR plate(s). Vortex and briefly spin down   SNP-PCR plate(s). 
SECTION: Barcoding
4.1. After SNP-PCR thermalcycler program has completed, remove SNP-PCR plate(s). Briefly vortex and spin down   . 
SECTION: Barcoding
5.1. Check that i7s have been diluted to working stock of 11uM, and proceed with preparing barcoding mix. If not, add 961 µL of lab-grade water to 119 µL of 100uM i7 to make working stock.

Vortex Hotstart MM and i7(s), spin down and store in benchtop cooler.

 

 Combine ingredients in a 1.5mL non- Lo-bind tube(s). Vortex, spin down and store in benchtop cooler.
SECTION: Barcoding
5.2. Label unskirted PCR plate: BARCODE, Date, Tray#, project initials and your initials.
SECTION: Barcoding
5.4. Using a 0.1mL combi-tip, dispense 6 µL of barcoding mix into each well of the Barcode plate(s).

SECTION: Barcoding
5.3. Remove the pre-aliquot i5 trays. Briefly vortex, and spin down   .

SECTION: Barcoding
5.5. Using a p10 multi-channel pipette, add 2 µL of i5 barcode into each corresponding well of the Barcode plate(s). Discard tips after each use. 
SECTION: Barcoding
5.6. Once the i5s have been added, add 3 µL of the diluted SNP-PCR into each corresponding well of the Barcode plate(s). Discard tips after each use. 
SECTION: Barcoding
5.7. Heat seal the Barcode plate(s). Gently vortex, and spin down   .
SECTION: Barcoding
6. Run the Barcode plate(s) on a thermalcycler using the following program. (In our lab, the program can be found using the menu options GTseq --> RAD-amp)

1. 95 °C for 15 min
2. 95 °C for 10 s
3. 65 °C for 30 s
4. 72 °C for 30 s
Repeat 2-4 for 10 cycles
5. 97 °C for 5 min
6. 4 °C indefinitely

SECTION: Barcoding
6.1. Heat seal the i5 tray, and diluted SNP-PCR plate(s). Store them in fridge.
SECTION: Sequalprep Normalization
7. Normalize the barcoded samples with SequalPrep kit. 


 

SECTION: Sequalprep Normalization
7.1. After Barcode thermalcycler program has completed, remove Barcode plate(s). Briefly vortex and spin down   . 
SECTION: Sequalprep Normalization
7.2. Label a new SequalPrep Normalization plate: Date, Tray#, project initials and your initials.
SECTION: Sequalprep Normalization
7.3. Using a 1mL combi-tip, dispense 10 µL of Binding Buffer into each well of the SequalPrep plate(s). If you have a partial tray, only dispense binding buffer that contains barcoded samples.
SECTION: Sequalprep Normalization
7.4. Using a p10 multi-channel pipette, transfer all 11 µL of barcoded samples into corresponding well of the SequalPrep plate(s). Discard tips after each use.
SECTION: Sequalprep Normalization
7.5. Heat seal SequalPrep plate(s). Vortex, and briefly spin down   .

SECTION: Sequalprep Normalization
7.6. Incubate SequalPrep plate(s) at room temperature for 60 min. (OK to leave plate(s) at room temperature overnight if necessary)
SECTION: Sequalprep Normalization
8. After incubation, empty out all contents from the SequalPrep plate(s) into sink. 
SECTION: Sequalprep Normalization
8.1. Tap plate(s) on paper towels to remove any residual binding buffer/barcoded samples.
SECTION: Sequalprep Normalization
8.2. Using a 5mL combi-tip, dispense 50ul of Wash Buffer into each well of the SequalPrep plate(s). Avoid touching the side of the wells, change tip if you think there might be contamination.
SECTION: Sequalprep Normalization
8.3. Empty out all wash buffer from SequalPrep plate(s) into sink. 
SECTION: Sequalprep Normalization
8.4. Tap plate(s) on paper towels to remove any residual wash buffer in wells.
SECTION: Sequalprep Normalization
9. Using a 1mL combi-tip, dispense 20 µL of Elution Buffer into each well of the SequalPrep plate(s). Avoid touching the side of the wells, change tip if you think there might be contamination.

SECTION: Sequalprep Normalization
9.1. Heat seal SequalPrep plate(s). Vortex, and spin down   . 

SECTION: Sequalprep Normalization
9.2. Incubate SequalPrep plate(s) at room temperature for 5 min. (OK to store plate(s) in fridge  overnight before pooling if necessary)
SECTION: Sequalprep Normalization
10. Using a p10 multi-channel pipette, pool 10 µL of each sample into a 25mL reservoir. OK to reuse tips. 

SECTION: Sequalprep Normalization
10.1. Transfer all pooled samples from reservoir into a 1.5mL Lo-bind tube. Label tube(s): NP# (tray#), project initials. Store in fridge. 
SECTION: Sequalprep Normalization
10.2. Heat seal SequalPrep plate(s), and store in fridge. 
SECTION: Beads Size Selection
11. Invitrogen‱ DynaMag‱- Spin Magnet
 

SECTION: Beads Size Selection
11.1. Vortex Ampure XP beads well, and briefly spin down to avoid beads getting stuck on lid. Keep in benchtop cooler. 


 

SECTION: Beads Size Selection
11.2. Vortex NP tube(s), and spin down. Keep in benchtop cooler. 
SECTION: Beads Size Selection
11.3. Label 1.5mL Lo-bind tube(s) with the corresponding NP tube number, if performing multiple bead size selection simultaneously. 
SECTION: Beads Size Selection
11.4. Add 25 µL of Ampure XP beads into each labeled 1.5mL Lo-Bind tube. 

SECTION: Beads Size Selection
11.5. Add 50 µL of NP tube into the corresponding labeled 1.5mL Lo-bind tube containing Ampure XP beads. 

SECTION: Beads Size Selection
11.6. Gently vortex, and quick spin down. Incubate tube(s) at room temperature for 5 min. 

Meanwhile, store the NP tube(s) back into fridge. 

SECTION: Beads Size Selection
11.7. Place tube(s) on magnetic stand for 5 min. 

SECTION: Beads Size Selection
11.8. Carefully transfer supernatant (~75 µL) into a new 1.5mL Lo-bind tube. Try to avoid disturbing the bead pellet, and be sure to label the new tube(s). 

SECTION: Beads Size Selection
11.9. To the supernatant, add 35 µL of Ampure XP beads. Repeat step if having multiple tube(s). 

SECTION: Beads Size Selection
11.10. Gently vortex, and quick spin down. Incubate tube(s) at room temperature for 5 min. 

In the meantime, prep the 80% ethanol wash solution. Each tube will need a total of 400 µL 80% ethanol wash solution. 
Add 400 µL of 100% ethanol with 100 µL of lab-grade water, multiply volume according to the total number of tubes you'll be doing. Vortex, spin down and set aside on benchtop.

SECTION: Beads Size Selection
11.11. Place tube(s) on magnetic stand for 5 min.

If time allows, store Ampure XP beads back in fridge. 
SECTION: Beads Size Selection
11.12. Transfer and discard all supernatant (~110 µL) without disturbing the bead pellet. 

SECTION: Beads Size Selection
11.13. While on magnetic stand, gently add 200 µL of 80% ethanol wash solution to each tube(s). 

SECTION: Beads Size Selection
11.14. Incubate for 30 s, and discard supernatant. Repeat wash step one more time.

SECTION: Beads Size Selection
11.15. While on magnetic stand, air dry beads for 5 min. Avoid overdrying beads. 

SECTION: Beads Size Selection
11.16. Remove tube(s) from magnetic stand, and add 17 µLlow TE. Resuspend beads by pipetting or gently vortex. 
SECTION: Beads Size Selection
11.17. Incubate tube(s) at room temperature for 2 min. 
SECTION: Beads Size Selection
11.18. Place tube(s) on magnetic stand for 5 min.
SECTION: Beads Size Selection
11.19. Carefully transfer all 17 µL supernatant into a new 1.5mL Lo-bind tube(s), labeled BSS#. 

This is now your final, undiluted GT-seq library(s). 
SECTION: Quantify library
12. Qubit GT-seq library


 

SECTION: Quantify library
12.1. If BSS tube(s) were stored in fridge, allow tube(s) to come to room temperature prior to qubit. 
SECTION: Quantify library
12.2. Make 1:200 diluted dye mix in a 2.0mL microcentrifuge tube. 

 

Label each qubit assay tube(s). Only label on the lid, never the side of the tube(s).
SECTION: Quantify library
12.3. Pipette 198 µL of diluted dye mix into each qubit assay tube(s). Add 2 µL of corresponding BSS to labeled qubit assay tube(s).

Vortex thoroughly, quick spin. 
SECTION: Quantify library
12.4. Incubate tube(s) at room temperature for 3 min. 

SECTION: Quantify library
12.5. Put qubit assay tube in Qubit Fluorometer and take 2 readings per sample and enter them into Library Prep sheet in PROJECT CHECKLIST_v2.4 spreadsheet. The spreadsheet has a column that will take the average, and make the calculations in the Tray-Quant tab to dilute each library down to 1nM.
SECTION: Quantify library
12.6. In a new 1.5mL Lo-bind tube, transfer 10 µL of BSS and dilute it with the calculated amount with 1X TE. Label tube as D#, include your initials and project initials. 

SECTION: Quantify library
12.7. Store diluted BSS tube(s) in GT-seq run box in fridge. Discard the undiluted BSS tube(s). Print out the  Library Prep sheet when all tube(s) are ready for the run.