TITLE: Section 2: NGS library preparation for sequencing
AUTHORS: Ester Kalef-Ezra, Ben Harvey, Katherine Roper, Christos Proukakis

[DESCRIPTION]
This protocol details NGS library preparation for sequencing and should be performed after Section 1: Enzymatic DNA Fragmentation (Manually).

[STEPS]
SECTION: Section 2: NGS library preparation for sequencing
1. NGS library preparation for sequencing can be done in two ways.
SECTION: Prepare the ligation master mix
2. Remove the AMPure XP beads from cold storage and equilibrate to Room temperature for at least 30 min prior use.
SECTION: Prepare the ligation master mix
3. Thaw on ice End Repair-A Tailing Buffer (yellow cap or bottle, to be used for End Repair-A Tailing master mix) and Ligation Buffer (purple cap or bottle, to be used for Ligation master mix). They may require >20 minutes). In the meantime, proceed to the next steps.
SECTION: Prepare the ligation master mix
4. Vortex the thawed vial of Ligation Buffer for 15 s at high speed to ensure homogeneity.

 


SECTION: Prepare the ligation master mix
5. Place T4 DNA Ligase (blue cap) on ice and mix by inversion.
SECTION: Prepare the ligation master mix
6. Calculate Ligation master mix as on calculations, as on Table 4.

Table 4. Ligation master mix. 
        Reagent  Volume for 1 reaction    Ligation Buffer (purple cap or bottle)  23 μl    T4 DNA Ligase (blue cap)  2 μl    Total  25 μl  
 
SECTION: Prepare the ligation master mix
7. Prepare Ligation master mix:
SECTION: Prepare the ligation master mix
7.1. Slowly pipette the Ligation Buffer into a 1.5-ml Eppendorf tube, ensuring that the full volume is dispensed.
SECTION: Prepare the ligation master mix
7.2. Slowly add the T4 DNA Ligase, rinsing the enzyme tip with buffer solution after addition.
SECTION: Prepare the ligation master mix
7.3. Mix well by slowly pipetting up and down 15–20 times or seal the tube and vortex at high speed for 10–20 seconds.
SECTION: Prepare the ligation master mix
7.4. Spin briefly to collect the liquid.
SECTION: Prepare the ligation master mix
8. Keep at Room temperature the Ligation master mix for 30–45 minutes before use on step on Sub-section 3 step 2.
 

SECTION: Repair and dA-Tail the DNA ends
9. Pre-program a thermal cycler as on Table 5. If required, use a reaction volume setting of 70 µL and lid temperature @65 °C.

Table 5. Thermal cycler program for End Repair/dA-Tailing.  
          Step  Temperature (°C)      1  20  15    2  72  15    3  4  Hold  

SECTION: Repair and dA-Tail the DNA ends
10. Vortex the thawed vial of End Repair-A Tailing Buffer for 15 s at high speed to ensure homogeneity. Visually inspect the solution; if any solids are observed, continue vortexing until all solids are dissolved.
 

SECTION: Repair and dA-Tail the DNA ends
11. Place End Repair-A Tailing Enzyme Mix (orange cap) on ice and mix by inversion.

SECTION: Repair and dA-Tail the DNA ends
12. Calculate End Repair/dA-Tailing master mix as on Table 6.

Table 6. End Repair/dA-Tailing mix. 
        Reagent  Volume for 1 reaction    End Repair-A Tailing Buffer (yellow cap
  or bottle)  16 μl    End Repair-A Tailing Enzyme Mix (orange
  cap)  4 μl    Total  20 μl  

SECTION: Repair and dA-Tail the DNA ends
13. Prepare End Repair/dA-Tailing master mix:
SECTION: Repair and dA-Tail the DNA ends
14. Add 20 µL of the End Repair/dA-Tailing master mix to each sample well containing approximately 50 µL of fragmented DNA. 
SECTION: Repair and dA-Tail the DNA ends
13.1. Slowly pipette the End Repair-A Tailing Buffer into a 1.5-ml Eppendorf tube, ensuring that the full volume is dispensed.
SECTION: Repair and dA-Tail the DNA ends
13.2. Slowly add the End Repair-A Tailing Enzyme Mix, rinsing the enzyme tip with buffer solution after addition.
SECTION: Repair and dA-Tail the DNA ends
13.3. Mix well by pipetting up and down 15–20 times or seal the tube and vortex at high speed for 5–10 seconds.
SECTION: Repair and dA-Tail the DNA ends
13.4. Spin briefly to collect the liquid and keep on ice.
SECTION: Repair and dA-Tail the DNA ends
15. Mix by pipetting up and down 15–20 times using a pipette set to 50 µL or cap the wells and vortex at high speed for 5–10 seconds.
SECTION: Repair and dA-Tail the DNA ends
16. Briefly spin the samples.
SECTION: Repair and dA-Tail the DNA ends
17. Immediately place the plate or strip tube in the thermal cycler and start the ‘End Repair/dA-Tailing’ program.
SECTION: Repair and dA-Tail the DNA ends
18. Once the thermal cycler reaches the 4 °C hold step, centrifuge the samples briefly.
SECTION: Repair and dA-Tail the DNA ends
19. Transfer the samples to ice and proceed to Ligate the molecular-barcoded adaptor.
SECTION: Ligate the molecular-barcoded adaptor
20. Pre-program a thermal cycler as on Table 7. If required, use a reaction volume setting of 100 µL and turn the lid temperature OFF.
 

Table 7. Thermal cycler program for Ligation.
 
          Step  Temperature (°C)      1  20  30    2  4  Hold  
 
SECTION: Ligate the molecular-barcoded adaptor
21. Thaw SureSelect XT HS2 Adaptor Oligo Mix (clear cap) on ice.
SECTION: Ligate the molecular-barcoded adaptor
22. To each end-repaired/dA-tailed DNA sample (approximately 70 µL), add 25 µL of the Ligation master mix that was prepared on section Prepare the ligation master mix and kept at Room temperature.
SECTION: Ligate the molecular-barcoded adaptor
23. Mix by pipetting up and down at least 10 times using a pipette set to 70 µL or cap the wells and vortex at high speed for 5–10 seconds.
SECTION: Ligate the molecular-barcoded adaptor
24. Briefly spin the samples.
SECTION: Ligate the molecular-barcoded adaptor
25. Add 5 µL of SureSelect XT HS2 Adaptor Oligo Mix (clear-capped tube) to each sample.
SECTION: Ligate the molecular-barcoded adaptor
26. Mix by pipetting up and down 15–20 times using a pipette set to 70 µL or cap the wells and vortex at high speed for 5–10 seconds. 
 

SECTION: Ligate the molecular-barcoded adaptor
27. Briefly spin the samples.
SECTION: Ligate the molecular-barcoded adaptor
28. Immediately place the plate or strip tube in the thermal cycler and start ‘Ligation’ program.

 

SECTION: Purify the sample using AMPure XP beads
29. Verify that the AMPure XP beads were held at Room temperature for at least 30 min.
SECTION: Purify the sample using AMPure XP beads
30. Prepare 1000 µL of 70% ethanol per sample.
 

SECTION: Purify the sample using AMPure XP beads
31. Mix the AMPure XP bead suspension well so that the reagent appears homogeneous and consistent in color.
SECTION: Purify the sample using AMPure XP beads
32. Add 80 µL of homogeneous AMPure XP beads to each DNA sample (approximately 100 μl) in the PCR plate or strip tube.
SECTION: Purify the sample using AMPure XP beads
33. Pipette up and down 15–20 times or cap the wells and vortex at high speed for 5–10 seconds to mix.
SECTION: Purify the sample using AMPure XP beads
34. Incubate samples for 5 min at Room temperature.
SECTION: Purify the sample using AMPure XP beads
35. Put the plate or strip tube into a magnetic separation device.
SECTION: Purify the sample using AMPure XP beads
36. Wait for the solution to clear (approximately 5 to 10 minutes).
SECTION: Purify the sample using AMPure XP beads
37. Keep the plate or strip tube in the magnetic stand. Carefully remove and discard the cleared solution from each well. 
 

SECTION: Purify the sample using AMPure XP beads
38. Continue to keep the plate or strip tube in the magnetic stand while you dispense 200 µL of freshly prepared 70% ethanol in each sample well.
SECTION: Purify the sample using AMPure XP beads
39. Wait for 1 min to allow any disturbed beads to settle, then remove the ethanol.
SECTION: Purify the sample using AMPure XP beads
40. Repeat ethanol wash by adding another 200 µL of freshly prepared 70% ethanol in each sample well.
SECTION: Purify the sample using AMPure XP beads
41. Wait for 1 min to allow any disturbed beads to settle, then remove the ethanol.
SECTION: Purify the sample using AMPure XP beads
42. Seal the wells with strip caps, then briefly spin the samples to collect the residual ethanol.
SECTION: Purify the sample using AMPure XP beads
43. Return the plate or strip tube to the magnetic stand for 30 s.
SECTION: Purify the sample using AMPure XP beads
44. Remove the residual ethanol with a P20 pipette. 
 

SECTION: Purify the sample using AMPure XP beads
45. Add 35 µL nuclease-free water to each sample well.
SECTION: Purify the sample using AMPure XP beads
46. Seal the wells with strip caps, then mix well on a vortex mixer and briefly spin the plate or strip tube to collect the liquid.
SECTION: Purify the sample using AMPure XP beads
47. Incubate for 2 min at Room temperature.
SECTION: Purify the sample using AMPure XP beads
48. Put the plate or strip tube in the magnetic stand and leave for approximately 5 min, until the solution is clear.
SECTION: Purify the sample using AMPure XP beads
49. Remove the cleared supernatant using a P20 twice (pipette set at 34 µL) to a fresh PCR plate or strip tube sample well and keep on ice. You can discard the beads at this time. 
 

SECTION: Amplify the adaptor-ligated library
50. Determine the appropriate index pair assignment for each sample. See Table 51 through Table 58 in the “Reference” chapter (see original SureSelect XT HS2 DNA System protocol) for sequences of the 8 bp index portion of the primers used to amplify the DNA libraries.
 

SECTION: Amplify the adaptor-ligated library
51. Thaw the following reagents for PCR amplification on ice: 
Herculase II Fusion DNA Polymerase (red cap)
5× Herculase II Buffer with dNTPs (clear cap)
SureSelect XT HS2 Index Primer Pairs
SECTION: Amplify the adaptor-ligated library
52. Mix my pipetting up and down 20 times Herculase II Fusion DNA Polymerase (red cap).
SECTION: Amplify the adaptor-ligated library
53. Mix by vortexing Herculase II Buffer with dNTPs (clear cap) and SureSelect XT HS2 Index Primer Pairs.
SECTION: Amplify the adaptor-ligated library
54. Pre-program a thermal cycler as on Table 8. If required, use a reaction volume setting of 50 µL.
SECTION: Amplify the adaptor-ligated library
55. Turn the lid temperature ON @105 °C and pre-heat the lid.

Table 8. Thermal cycler program for Amplification PCR. 
 
            Step  Number of Cycles  Temperature (°C)      Time (min)    1  1  98  2    2  5-9*  98  0.5        60  0.5        72  1    3  1  72  5    4  1  4  Hold  
 
SECTION: Amplify the adaptor-ligated library
56. Prepare the appropriate volume of Pre-Capture PCR reaction mix as on Table 9.
 

Table 9. Amplification PCR mix.  
        Reagent  Volume for 1 reaction    5× Herculase II Buffer with dNTPs (clear
  cap)  10 μl    Herculase II Fusion DNA Polymerase (red
  cap)  1 μl    Total  11 μl  


SECTION: Amplify the adaptor-ligated library
57. Add 11 µL of the PCR reaction mixture to each purified DNA library sample (∼34 µL) in the PCR plate wells.
SECTION: Amplify the adaptor-ligated library
58. Add 5 µL of the appropriate SureSelect XT HS2 Index Primer Pair to each reaction.
SECTION: Amplify the adaptor-ligated library
59. Cap the wells then vortex at high speed for 5 s.
SECTION: Amplify the adaptor-ligated library
60. Spin the plate or strip tube briefly to collect the liquid and release any bubbles.
SECTION: Amplify the adaptor-ligated library
61. Immediately place the plate or strip tube in the thermal cycler and start ‘Amplification PCR’ program.
 

SECTION: Purify the sample using AMPure XP beads
62. Verify that the AMPure XP beads were held at Room temperature for at least 30 min.
SECTION: Purify the sample using AMPure XP beads
63. Prepare 400 µL of 70% ethanol per sample, if not have done already on the same day.
SECTION: Purify the sample using AMPure XP beads
64. Mix the AMPure XP bead suspension well so that the reagent appears homogeneous and consistent in color.
SECTION: Purify the sample using AMPure XP beads
65. Add 50 µL of homogeneous AMPure XP beads to each amplification reaction sample (approximately 50 µL) in the PCR plate or strip tube.
SECTION: Purify the sample using AMPure XP beads
66. Pipette up and down 15–20 times or cap the wells and vortex at high speed for 5–10 seconds to mix.
SECTION: Purify the sample using AMPure XP beads
67. Incubate samples for 5 min at Room temperature.
SECTION: Purify the sample using AMPure XP beads
68. Put the plate or strip tube into a magnetic separation device.
SECTION: Purify the sample using AMPure XP beads
69. Wait for the solution to clear (approximately 5 min).
SECTION: Purify the sample using AMPure XP beads
70. Keep the plate or strip tube in the magnetic stand. Carefully remove and discard the cleared solution from each well. 
 

SECTION: Purify the sample using AMPure XP beads
71. Continue to keep the plate or strip tube in the magnetic stand while you dispense 200 µL of freshly prepared 70% ethanol in each sample well.
SECTION: Purify the sample using AMPure XP beads
72. Wait for 1 min to allow any disturbed beads to settle, then remove the ethanol.
SECTION: Purify the sample using AMPure XP beads
73. Repeat ethanol wash by adding another 200 µL of freshly prepared 70% ethanol in each sample well.
SECTION: Purify the sample using AMPure XP beads
74. Wait for 1 min to allow any disturbed beads to settle, then remove the ethanol.
SECTION: Purify the sample using AMPure XP beads
75. Seal the wells with strip caps, then briefly spin the samples to collect the residual ethanol.
SECTION: Purify the sample using AMPure XP beads
76. Return the plate or strip tube to the magnetic stand for 30 s.
SECTION: Purify the sample using AMPure XP beads
77. Remove the residual ethanol with a P20 pipette. 
 

SECTION: Purify the sample using AMPure XP beads
78. Add 15 µL nuclease-free water to each sample well.
SECTION: Purify the sample using AMPure XP beads
79. Seal the wells with strip caps, then mix well on a vortex mixer and briefly spin the plate or strip tube to collect the liquid.
SECTION: Purify the sample using AMPure XP beads
80. Incubate for 2 min at Room temperature.
SECTION: Purify the sample using AMPure XP beads
81. Put the plate or strip tube in the magnetic stand and leave for 2 to 3 minutes, until the solution is clear.
SECTION: Purify the sample using AMPure XP beads
82. Remove the cleared supernatant (approximately 15 µL) to a fresh PCR plate or strip tube sample well and keep on ice. You can discard the beads at this time.
 

SECTION: Purify the sample using AMPure XP beads
83. If you do not continue to the next step, seal the sample wells and store at 4 °C 2 min or at -20 °C for prolonged storage.
SECTION: Preparation of AMPure XP Bead Plates
2. 
 
In each case, follow these instructions:

Clear the Bravo deck of all plates and tip boxes, then wipe it down with DNA Away.
SECTION: Preparation of AMPure XP Bead Plates
3. Switch on Bravo computer, VWorks software, Bravo robot, Inheco Multi TEC control and Thermo Cube.
SECTION: Preparation of AMPure XP Bead Plates
4. Place AMPure XP beads for at least 30 min @Room temperature, mix well and verify that they are in suspension.
SECTION: Preparation of AMPure XP Bead Plates
5. On the VWorks software SureSelect setup form, under Select protocol to execute, select the ‘AMPureXP_Aliquot_LibPrep’ or ‘AMPureXP_Aliquot_PreCap’ protocol and label a deep well plate accordingly.
SECTION: Preparation of AMPure XP Bead Plates
6. Select the number of columns of samples to be processed.
SECTION: Preparation of AMPure XP Bead Plates
7. Click Display Initial Bravo Deck Setup.
SECTION: Preparation of AMPure XP Bead Plates
8. Pour the AMPure XP beads in suspension into the Reservoir and place the reservoir with the beads at Bravo deck 6 position as on Figure 1.
 
SECTION: Preparation of AMPure XP Bead Plates
9. Place on Bravo a new tip box without the cap (deck 2), empty tip box (deck 8) and empty deep well plate (deck 5) as on Figure 1.
SECTION: Preparation of AMPure XP Bead Plates
10. Verify that the Bravo deck has been set up as displayed on the right side of the form.
SECTION: Preparation of AMPure XP Bead Plates
11. Verify that the Current Tip State indicator on the form matches the configuration of unused and used tips in the tip boxes at Bravo Deck positions 2 and 8, respectively.
SECTION: Preparation of AMPure XP Bead Plates
12. When verification is complete, click Run Selected Protocol. 
 

SECTION: Preparation of AMPure XP Bead Plates
13. When verification is complete, click Run Selected Protocol.
SECTION: Preparation of AMPure XP Bead Plates
14. When the protocol is complete, remove the Agilent Deep Well plate containing the AMPure XP bead suspension from position 5 of the Bravo deck.
 

SECTION: Prepare adaptor-ligated libraries
15. Clear the Bravo deck of all plates and tip boxes, then wipe it down with DNA Away. 
SECTION: Prepare adaptor-ligated libraries
16. Switch on Bravo computer, VWorks software, Bravo robot, Inheco Multi TEC control and Thermo Cube.
SECTION: Prepare adaptor-ligated libraries
17. Set Thermo Cube to 0 °C at position 9 of the Bravo deck. Add 300 mL 25% EtOH to Thermo Cube if necessary (this will be indicated on Thermo Cube screen).
SECTION: Prepare adaptor-ligated libraries
18. Select protocol for Bravo to execute ‘01 LibraryPrep_XT_HS2_ILM’ program.
SECTION: Prepare adaptor-ligated libraries
19. Verify that the Processing Plate selection is set to the correct plate type.
SECTION: Prepare adaptor-ligated libraries
20. Select the number of columns of samples to be processed.
SECTION: Prepare adaptor-ligated libraries
21. Click Display Initial Bravo Deck Setup.
SECTION: Prepare adaptor-ligated libraries
22. Pre-set the temperature of Bravo deck position 4 (CPAC 2 position 1) to 79 °C and deck position 6 to 20 °C (CPAC 2 position 2) using the Inheco Multi TEC control touchscreen.
SECTION: Prepare adaptor-ligated libraries
23. Thaw on ice End Repair-A Tailing Buffer and Ligation Buffer (purple cap or bottle, to be used for Ligation master mix). They may require >20 minutes). Then vortex at high speed for 15 s them to mix well.
SECTION: Prepare adaptor-ligated libraries
24. Place on ice and mix by inversion End Repair-A Tailing Enzyme Mix (orange cap), T4 DNA Ligase (blue cap).
SECTION: Prepare adaptor-ligated libraries
25. Place on ice and mix by vortexing SureSelect XT HS2 Adaptor Oligo Mix (white cap).
SECTION: Prepare adaptor-ligated libraries
26. Prepare the DNA End-Repair/dA-Tailing master mix (Table 10), Ligation master mix (Table 11) and Adaptor Oligo Mix (Table 12) based on the number of columns needed. Store on ice until use. 
 

Table 10. Preparation of End Repair/dA-Tailing master mix.
   Table 11. Preparation of Ligation master mix.
  Table 12. Preparation of Adaptor Oligo Mix dilution.
  

SECTION: Prepare adaptor-ligated libraries
27. Prepare the master mix source plate containing the mixtures prepared on step 4 as shown on Table 13 based on the number of columns needed. Distribute them as on Figure 2.

Table 13. Preparation of the master mix source plate for 01 LibraryPrep_XT_HS2_ILM protocol.
    

SECTION: Prepare adaptor-ligated libraries
28. Optional: Seal the plate and centrifuge for 30 s. Then remove the plate sealing. Alternatively, inspect visually and remove the bubbles with a pipette tip.
 

SECTION: Prepare adaptor-ligated libraries
29. Place the source plate on Deck position 9.
SECTION: Prepare adaptor-ligated libraries
30. Place all materials as on the right side of the Bravo form as on Figure 3.
  
SECTION: Prepare adaptor-ligated libraries
31. Verify that the Bravo deck has been set up as displayed on the right side of the form.
SECTION: Prepare adaptor-ligated libraries
32. Verify that the Current Tip State indicator on the form matches the configuration of unused and used tips in the tip boxes at Bravo Deck positions 2 and 8, respectively.
SECTION: Prepare adaptor-ligated libraries
33. When verification is complete, click Run Selected Protocol.
 

SECTION: Purify adaptor-ligated DNA using AMPure XP beads
34. Clear the Bravo deck of all plates and tip boxes, then wipe it down with DNA Away. 
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
35. If not already, switch on Bravo computer, VWorks software, Bravo robot and Inheco Multi TEC control.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
36. Switch OFF Thermo Cube.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
37. Select protocol for Bravo to execute ‘02 Cleanup_LibPrep_XT_HS2_ILM’ program.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
38. Pre-set the temperature of Bravo deck position 4 (CPAC 2 position 1) to 45 °C and deck position 6 to 20 °C (CPAC 2 position 2) using the Inheco Multi TEC control touchscreen.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
39. Prepare 2 Agilent deep well reservoirs containing: 30 mL of nuclease-free water and with 50 mL 70% EtOH (freshly made).
 

SECTION: Purify adaptor-ligated DNA using AMPure XP beads
40. Select the number of columns of samples to be processed.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
41. Verify that the Processing Plate selection is set to the correct plate type. 
 

SECTION: Purify adaptor-ligated DNA using AMPure XP beads
42. Click Display Initial Bravo Deck Setup and place all plates and tips needed.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
43. Verify that the Bravo deck has been set up as displayed on the right side of the form, as on Figure 4.  
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
44. Verify that the Current Tip State indicator on the form matches the configuration of unused and used tips in the tip boxes at Bravo Deck positions 2 and 8, respectively.
SECTION: Purify adaptor-ligated DNA using AMPure XP beads
45. When setup and verification is complete, click Run Selected Protocol.
 
  

SECTION: Amplify adaptor-ligated libraries
46. Clear the Bravo deck of all plates and tip boxes, then wipe it down with DNA Away.
SECTION: Amplify adaptor-ligated libraries
47. Switch ON Thermo Cube.
 

SECTION: Amplify adaptor-ligated libraries
48. Select protocol for Bravo to execute ‘03 Pre-CapPCR_XT_HS2_ILM’ program.
SECTION: Amplify adaptor-ligated libraries
49. Pre-set the temperature of Bravo deck position 6 to 4 °C (CPAC 2 position 2) using the Inheco Multi TEC control touchscreen.
SECTION: Amplify adaptor-ligated libraries
50. Select the number of columns of samples to be processed.
SECTION: Amplify adaptor-ligated libraries
51. Verify that the Processing Plate selection is set to the correct plate type. 
 

SECTION: Amplify adaptor-ligated libraries
52. Click Display Initial Bravo Deck Setup.
SECTION: Amplify adaptor-ligated libraries
53. Thaw on ice and mix by vortexing 5× Herculase II Reaction Buffer (clear cap) and SureSelect XT HS2 Index Primer Pairs to be used.
 

SECTION: Amplify adaptor-ligated libraries
54. Thaw on ice and mix by pipetting up and down 15-20 times Herculase II Fusion DNA Polymerase (red cap).
SECTION: Amplify adaptor-ligated libraries
55. Centrifuge briefly 5× Herculase II Reaction Buffer (clear cap), SureSelect XT HS2 Index Primer Pairs and Herculase II Fusion DNA Polymerase (red cap).
SECTION: Amplify adaptor-ligated libraries
56. Pre-program a thermal cycler as on Table 14. If required, use a reaction volume setting of 50 µL and lid temperature @105 °C.

Table 14. Thermal cycler program for Pre-Capture PCR. 
          Step  Temperature (ºC)  Time (min)    1  98  2    2  98  0.5    3  60  0.5    4  72  1    5  Return back to step 2 until 8
  cycles      in total      6  72  5    7  4  Hold  

SECTION: Amplify adaptor-ligated libraries
57. Using a multichannel pipette, transfer 5 µL of each SureSelect XT HS2 Index Primer Pair from the 96-well plate in which the primer pairs are provided into the PCR plate to be used for the pre-capture PCR thermal cycling.
 

SECTION: Amplify adaptor-ligated libraries
58. Prepare the appropriate volume of pre-capture PCR master mix, according to Table 15. Vortex at medium speed for 15–20 seconds and keep TemperatureOn ice 
 .
Table 15. Preparation of the Pre-Capture master mix for 03 Pre-CapPCR_XT_HS2_ILM protocol.
  

SECTION: Amplify adaptor-ligated libraries
59. Distribute the Pre-Capture master mix in Column 2 of a full skirt source plate (Eppendorf twin.tec or 
Armadillo plate) as on Table 16.

Table 16. Volumes for Pre-Capture master mix source plate for 03 Pre-CapPCR_XT_HS2_ILM protocol.
  

SECTION: Amplify adaptor-ligated libraries
60. Cap the Pre-Capture master mix source plate and centrifuge for 30 s to minimize bubbles. 
SECTION: Amplify adaptor-ligated libraries
61. Uncap the plate and place it on Bravo position deck 9.
SECTION: Amplify adaptor-ligated libraries
62. Place a full tip box, empty tip box and waste plate as needed.
SECTION: Amplify adaptor-ligated libraries
63. Verify that the Bravo deck has been set up as displayed on the right side of the form, as on Figure 6.  
SECTION: Amplify adaptor-ligated libraries
64. Verify that the Bravo deck has been set up as displayed on the right side of the form, as on Figure 6.
SECTION: Amplify adaptor-ligated libraries
65. Verify that the Current Tip State indicator on the form matches the configuration of unused and used tips in the tip boxes at Bravo Deck positions 2 and 8, respectively.
SECTION: Amplify adaptor-ligated libraries
66. When verification is complete, click Run Selected Protocol.
 

SECTION: Amplify adaptor-ligated libraries
67. When the Bravo program is complete, collect the PCR plate at position 6 of the Bravo deck.
SECTION: Amplify adaptor-ligated libraries
68. Cap the plate, centrifuge briefly and initiate Pre-Capture PCR.
SECTION: Purify amplified DNA using AMPure XP beads
69. Clear the Bravo deck of all plates and tip boxes, then wipe it down with DNA Away. 
SECTION: Purify amplified DNA using AMPure XP beads
70. If not already, switch on Bravo computer, VWorks software, Bravo robot and Inheco Multi TEC control.
SECTION: Purify amplified DNA using AMPure XP beads
71. Switch OFF Thermo Cube.
SECTION: Purify amplified DNA using AMPure XP beads
72. Select protocol for Bravo to execute 04a Cleanup_Pre-CapPCR_SP_XT_HS2_ILM.
SECTION: Purify amplified DNA using AMPure XP beads
73. Pre-set the temperature of Bravo deck position 4 (CPAC 2 position 1) to 45 °C and deck position 6 to 20 °C (CPAC 2 position 2) using the Inheco Multi TEC control touchscreen.
SECTION: Purify amplified DNA using AMPure XP beads
74. Prepare 2 Agilent deep well reservoirs containing: 30 mL of nuclease-free water and with 50 mL 70% EtOH.
 

SECTION: Purify amplified DNA using AMPure XP beads
75. Select the number of columns of samples to be processed.
SECTION: Purify amplified DNA using AMPure XP beads
76. Verify that the Processing Plate selection is set to the correct plate type. 
 

SECTION: Purify amplified DNA using AMPure XP beads
77. Click Display Initial Bravo Deck Setup and place all plates and tips needed.
SECTION: Purify amplified DNA using AMPure XP beads
78. Verify that the Bravo deck has been set up as displayed on the right side of the form, as on Figure 7.
 
SECTION: Purify amplified DNA using AMPure XP beads
79. Verify that the Current Tip State indicator on the form matches the configuration of unused and used tips in the tip boxes at Bravo Deck positions 2 and 8, respectively.
SECTION: Purify amplified DNA using AMPure XP beads
80. When setup and verification is complete, click Run Selected Protocol.
 

SECTION: Purify amplified DNA using AMPure XP beads
81. Cap and centrifuge briefly the purified libraries.