HIV WGS - 400bp Amplicon Tiling - Oxford Nanopore Technology Protocol

Reverse Transcription

The LunaScript® RT SuperMix Kit will be used to reverse transcribe HIV RNA for preparation to perform the Primer Pool PCR assay. NOTE: LunaScript® RT SuperMix will be kept in the PCR Clean room.

Place 96-well PCR plate on an Eppendorf® PCR cooler (stored at -20oC).

Using a repeater pipettor aliquot 4 μl of LunaScript® RT SuperMix into each sample well of a 96‑well PCR plate according to plate map.

Cover the plate and move the PCR plate containing master mix from the PCR Clean Room to the processing bench for sample addition.

Add 16 μl of RNA extract to the appropriate sample well and gently mix by pipetting up and down. If less than 16 μl is available, make up the volume with nuclease-free water. Total volume = 20 μl per well.

Seal the plate with MicroAmp™ 8-cap strips and briefly centrifuge to collect droplets.

Return the RT plate to the Eppendorf® PCR cooler until thermocycler reaches 25oC.

10. Load the plate into the SimpliAmp™ thermocycler, then run the “HIV REVERSE TRANSCRIPTION” method with the following parameters: Set heated lid to 105oC  / Runtime ~14 minutes

PCR Primer Pool Preparation

Combine the following components to prepare the master mix for each primer pool (A or B). (Allow for pipetting loss by adding ~ 4 reactions per 32 samples.)

PCR Primer Pool Preparation

Combine the following components to prepare the master mix for each primer pool (A or B). (Allow for pipetting loss by adding ~ 4 reactions per 32 samples.)

NOTE: Use the HIV-1 RT and Primer Pool Preparation Worksheet to calculate the required amount of master mix. This can be made at the same time as the LunaScript® SuperMix RT plate and stored at 4oC until use.

@10μM

Component Volume

Nuclease-free Water 8.90 μl

HIV-1 Primer Pool (A or B) 1.10 μl

Q5® Hot Start HF 2X Master Mix 12.5 μl

Total volume per sample 22.5 μl

15. Place 96-well PCR plate on an Eppendorf® PCR cooler (stored at -20oC).

16. Using a repeater pipettor aliquot 22.5 μl of each master mix into the appropriate wells of the 96-well plate.

17. Cover and store at 4oC until reverse transcription protocol is complete.

18. Remove the RT plate from the thermocycler and centrifuge.

19. Carefully remove the 8-cap strips and using a multi-channel pipettor transfer 2.5 μl of the RT reactions into the appropriate wells of the PCR plate containing the primer pools. Gently mix by pipetting the contents of each well up and down. Total volume = 25 μl per well.

NOTE: It is useful to separate the plate in half with primer pool A in columns 1-6 and primer pool B in columns 7-12.

21. Seal the plate with 8-cap strips and briefly centrifuge.

22. Load the plate into the SimpliAmp™ thermocycler, then run the “HIV PCR PRIMERS” method with the following parameters: Set heated lid to 105oC   /   Runtime ~5 Hours

24. Repeat the same primer pool preparation from above (Step 1) and aliquot it into the same wells as in Step 1 into a new PCR plate in the same wells as the previous plate.

25. NOTE: Use the HIV-1 RT and Primer Pool Preparation Worksheet to calculate the required amount of master mix.

@10μM

Component Volume

Nuclease-free Water 8.90 μl

HIV-1 Primer Pool (A or B) 1.10 μl

Q5® Hot Start HF 2X Master Mix 12.5 μl

Total volume per sample 22.5 μl

26. Remove the cDNA plate from the thermocycler and centrifuge.

27. Carefully remove the 8-cap strips and using a multi-channel pipettor transfer 2.5 μl of the 1st cDNA products into the corresponding wells of the new PCR plate containing the 2nd set of primer pools.

(2.5 μl of 1st Pool A or B goes into corresponding 2nd Pool A or B of new plate.)

29. Gently mix by pipetting the contents of each well up and down. Total volume = 25 μl per well.

30. Seal the plate with 8-cap strips and briefly centrifuge.

31. Load the plate into the SimpliAmp™ thermocycler, then run the “HIV PCR PRIMERS” method again. Same parameters as above.

Set heated lid to 105oC   /   Runtime ~5 Hours

Addition of rapid barcodes

Thaw Rapid Barcode Plate and bring SPRI beads to room temperature ~ 1 hour before PCR is complete.

35. Spin down the Rapid Barcode Plate.

36. Remove the primer-specific cDNA plate from the thermocycler and centrifuge briefly to collect the contents at the bottom of the wells.

37. Place the plate on an Eppendorf® PCR cooler.

38. Carefully remove the 8-cap strips from the cDNA plate to prevent any splashing.

39. Place a clean 96-well PCR plate on a separate Eppendorf® PCR cooler (stored at -20oC) and combine each 25 μl cDNA reaction into a single well in the plate. Mix by pipetting up and down and set aside. Total volume per sample = 50 μl.

40. Place another clean PCR 96-well sample plate on a separate Eppendorf® PCR cooler and using a repeater pipettor aliquot 2.5 μl of nuclease-free water into corresponding sample wells.  This is the Barcode Attachment Plate (BAP).

41. Using a multichannel pipette, transfer 5 μl of the pooled cDNA products to the corresponding well of the Barcode Attachment Plate (BAP) and mix by pipetting.

42. Using a multichannel pipette, transfer 2.5 μl of the appropriate Rapid Barcodes to the corresponding well of the Barcode Attachment Plate (BAP) and mix by pipetting. Be careful not to cross-contaminate the different wells. Total volume per sample = 10 μl.

43. Seal the plate with 8-cap strips and briefly centrifuge.

44. Incubate the plate on the thermocycler using the “RAPID BARCODE ATTACHMENT” protocol with the following parameters:

Pooling Samples and Clean-up

Remove the Barcode Attachment Plate (BAP) from the thermocycler and centrifuge briefly to collect the contents at the bottom of the wells.

47. Carefully remove the 8-cap strips from the BAP plate to prevent any splashing.

48. Pool the barcoded samples into a 5 ml Eppendorf DNA LoBind tube. ~10 μl per sample

Pool the barcoded samples into a 5 ml Eppendorf DNA LoBind tube. ~10 μl per sample

For example:

50. Resuspend the SPRI beads by vortexing.

Add an equal volume of resuspended SPRI beads to the pooled samples and mix by flicking the tube.

For example:

53. Incubate at room temperature on a gentle rotator mixer (~700 rpm) for 5 minutes.

54. While incubating, prepare 3 ml of fresh 80% ethanol in nuclease-free water (2400 μl ethanol 600 μl water).

55. Spin down the sample tube briefly to bring any liquid down from the cap and place it on the magnet. A pellet will form on the side wall of the tube.

56. When the solution clears, keep the tube on the magnet and pipette off the supernatant and discard.

57. Without disturbing the beads, add 1.5 ml of the freshly prepared 80% ethanol. Rotate the tube on the magnet to move the beads front to back a few times to wash beads. Remove the ethanol using a pipette and discard.

58. Repeat the previous step.

59. Briefly spin down the tube and place it back on the magnet. Pipette off any residual ethanol. Allow to dry for 30 seconds. Do not over dry the pellet to the point of cracking.

60. Remove the tube from the magnet and resuspend the pellet in 30 μl Elution Buffer (EB). Gently mix by pipetting up and down.

61. Incubate for 10 minutes at room temperature.

62. Place the tube on the magnet until the eluate is clear and colorless.

63. Remove and transfer the 30 μl eluate (containing the DNA library) into a clean 1.5 ml Eppendorf DNA LoBind tube. Dispose of the pelleted beads. Be careful not to transfer any of the beads.

64. Proceed to quantifying the DNA concentration using the Qubit dsDNA HS Assy Kit. (Or any preferred quantification method)

Library quantification and normalization

Analyze 1 μl of the amplified library using the Qubit™ 4.0 Fluorometer and the Qubit™ dsDNA HS Assay Kit. For more information, see the Qubit™ dsDNA HS Assay Kits User Guide

Determine the amplified library concentration using Qubit™ 4.0 Fluorometer: .

68. Prepare the Qubit™ standards as directed in the user guide (10 μl standard  + 190 μl Qubit™ dsDNA HS Buffer ), mix well, and incubate for at least 2 minutes.

69. For each sample, combine 1 μl of the amplified library with 199 μl of Qubit™ dsDNA HS Buffer, mix well, and incubate for at least 2 minutes.

70. On the Qubit™ 4.0 Fluorometer home screen, select dsDNA then 1x dsDNA High Sensitivity.

71. Measure the prepared standards.

72. Select the sample concentration as “ng/μl” with an input value of 1 μl.

73. Measure and record the library concentration.

A total volume of 11 μl is required for the following step.

If concentration of the total pool is less than 600ng/μl, use library pool undiluted.

76. Add 1 μl of Rapid Adapter F (RAP F) to the 11 μl normalized barcoded DNA and mix gently.

77. Incubate at room temperature for 5 minutes.

78. After incubation, place the prepared library in a cold rack until ready to load onto the SpotON flowcell.

Priming and Loading the SpotON Flow Cell

Thaw the Sequencing Buffer II (SBII), Loading Beads II (LBII) or Loading Solution (LS, if using), Flush Tether (FLT) and Flush Buffer (FB) at room temperature.

81. Mix the SBII, FB and FLT tubes by vortexing. Spin down the SBII and FLT tubes.

82. Open the ONT sequencer lid and slide the flow cell under the clip. Press down firmly on the flow cell to ensure correct thermal and electrical contact.

83. QC the SpotON flowcell using the MinKNOW software before proceeding. QC passes if total active pores are >800.

84. After QC is complete, move the unit including the flowcell to the bench for priming and loading.

85. Slide the priming port cover clockwise to open the priming port.

Prime and load the SpotON Flow Cell

In a 1.5 ml Eppendorf LoBind tube, prepare the flow cell priming mix by adding 30 μl of Flush Tether (FLT) to 1.17 ml of Flush Buffer (FB) and vortex to mix.

88. Open the priming port and check for a small air bubble under the cover. Draw back a small volume to remove any bubbles (a few μl)

89. Set a P1000 to 200 μl

90. Insert the tip into the priming port

91. Turn the wheel until the dial shows 220-230 μl, or until you can see a small volume of buffer entering the pipette tip.

92. Visually inspect that there is continuous buffer from the priming port across the sensor array.

93. Load 800 ul of the priming mix into the flow cell via the priming port without introducing bubbles. Wait 5 minutes.

Prepare the library for loading

Thoroughly mix the contents of the Loading Beads II (LBII) by pipetting up and down. The LBII tube contains a suspension of beads that settle very quickly. It is vital that they are mixed immediately before use.

In a new tube, prepare the library for loading as follows:

Reagent Volume

Sequencing Buff II (SBII) 37.5 μl

Loading Beads II 22.5 μl

DNA Library 12 μl

Total volume 75 μl

Complete the flow cell priming

Gently lift the SpotON sample port cover to make the sample port accessible.

Slowly load 200 ul of the priming mix into the flow cell via the priming port (not the SpotON sample port), avoiding the instruction of bubbles.

Immediately, mix the prepared library by pipetting up and down gently.

Add 75 μl of sample to the flow cell via the SpotON port in a dropwise manner. Ensure each drop flows into the port before adding the next.

Replace the SpotON sample port cover, making sure it is seated correctly, close the priming port, and close the sequencer lid. Proceed to sequencing