DNA sequencing

Overview

Options

• Sanger sequencing
• Illumina – for short reads. Can be done either at our core facility or sent to EMBL's Genecore. Options:
  • NextSeq: 2 x 100 bp reads (most common)
  • MiSeq: 2 x 150 bp reads, but fewer samples and reads than NextSeq
• Oxford Nanopore – for long reads (but few reads overall). Typically done for whole plasmids and amplicons via SeqVision.
• PacBio – for long reads. Can be done at out core facility.

Illumina sequencing preparation via PCR

An easy way to prepare samples for NGS is to amplify your DNA with primers that already contain the required adaptors and barcodes.

Forward primer

It should look like this:

5' - P5 - SP1 - Plasmid site 2' - 3',

where:

• P5: AAT GAT ACG GCG ACC ACC GAG ATC T – attaches to the flowcell during sequencing;
• SP1: A CAC TCT TTC CCT ACA CGA CGC TCT TCC GAT CT (Read 1 Sequencing Primer);
• Plasmid site 2': A reverse complement of the site on the plasmid where the primer attaches.

Thus, the sequence is:

AAT GAT ACG GCG ACC ACC GAG ATC T - A CAC TCT TTC CCT ACA CGA CGC TCT TCC GAT CT - ...

Reverse primer

It should look like this:

5' - P7 - Barcode - SP2' - Plasmid site 1 - 3',

where:

• P7: CAA GCA GAA GAC GGC ATA CGA GAT – attaches to the flowcell during sequencing;
• Barcode: A unique identifier to know which sample was yours in a pooled sequencing; must be unique across all samples that are submitted for sequencing;
• GT: optional barcode extension to have an 8-letter barcode;
• SP2': G ACT GGA GTT CAG ACG TGT GCT C (Read 2 Sequencing Primer) – a reverse complement of the SP2 sequence.

Thus, the sequence is (assuming the barcode is TGT ATT):

CAA GCA GAA GAC GGC ATA CGA GAT - TGT ATT - GT - G ACT GGA GTT CAG ACG TGT GCT C - ...

Design details

• You want to have primers end with one or two G's or C's, but not more.
• Check in Benchling for primer dimers. If delta G < -7 kcal/mol, this is already becoming risky.
• Sequence diversity will be low at the beginning for each read. Consider adding PhiX to resolve chip failures.

NEBNext Ultra II DNA Library Prep Kit for Illumina

Overview

NEBNext Adaptor for Illumina sequence:

5´-/5Phos/ GAT CGG AAG AGC ACA CGT CTG AAC TCC AGT C dU A CAC TCT TTC CCT ACA CGA CGC TCT TCC GAT C-s-T-3´

Final sequences that we submit for sequencing:

5'-P5-SP1-Sequence-SP2-Barcode'-P7'-3' 5'-P5-SP2-Sequence'-SP1-Barcode'-P7'-3'

where:

• P5: AAT GAT ACG GCG ACC ACC GAG ATC T – attaches to the flowcell during sequencing;
• SP1: A CAC TCT TTC CCT ACA CGA CGC TCT TCC GAT CT (Read 1 Sequencing Primer);
• SP2: GAC TGG AGT TCA GAC GTG TGC TCT TCC GAT CT (Read 2 Sequencing Primer);
• P7: CAA GCA GAA GAC GGC ATA CGA GAT – attaches to the flowcell during sequencing.

The following sequences are used for adaptor trimming of NEBNext adaptors for Illumina:

• Read 1: A GAT CGG AAG AGC ACA CGT CTG AAC TCC AGT CA - SP2 plus an additional A from somewhere;
• Read 2: AG ATC GGA AGA GCG TCG TGT AGG GAA AGA GTG T - SP1'.

A schematic tracking all sequences and their orientations:

General guidelines

• All pipetting up and down must be done at least 10 times to mix very well. The presence of a small amount of bubbles will not interfere with performance.
• Always spin down PCR tubes after incubation in a thermocycler for 1 min. Use the fixed-speed microcentrifuge that you typically use to spin down tubes.

Sample Tracking Sheet

Step	Qubit concentration (ng/uL)	Vol. (uL)	Milli-Q	Amount (ng)	Notes
End repair	[C1]	V	to 50 uL	[C1] x V	Ideally 1 ug, but no less than .5 ng
Barcoding	[C2]	15	-	[C2] x 15	Barcode:
Final preparation	[C3]	19		[C3] x 19

End repair (90 min)

1. Get sample concentrations using Qubit High Sensitivity kit and record them in the Sample Tracking Sheet.
2. Transfer between .5 ng and 1 ug (ideal) of DNA to PCR tubes and adjust volume to 50 uL with Milli-Q.
3. Prepare master mix on ice in a .5 mL LoBind tube and pipette it gently up and down:

Component	Per reaction (uL)	Per reaction + 5% (uL)
End Prep Reaction Buffer	7	7.35
End Prep Enzyme Mix	3	3.15
Total	10	10.5

1. Add 10 uL of master mix to each DNA sample and pipette up and down.
2. Incubate in a thermocycler (60 uL):

Step	Temperature	Time (min)
Incubation	20	30
Enzyme inactivation	65	30
Hold	4	Hold

1. Spin down for 1 min.

Adaptor ligation (45 min)

1. Thaw NEBNext Adaptor Dilution Buffer.
2. In the meantime, prepare Master Mix on ice in a .5 mL LoBind tube:

Component	Per reaction + 10%(uL)
NEBU2 Ligation Master Mix	15.4
NEBU2 Ligation Enhancer	1.1

1. Add 15 uL of Master Mix to each DNA sample.
2. You'll need 2.3 uL per sample of diluted NEBNext Adaptor. Dilute NEBNext Adaptor (from the Index Kit) with NEBNext Adaptor Dilution Buffer according to the DNA amount you have, making sure the final diluted volume is sufficient for all your samples.

DNA amount (ng)	Dilution
<5	25X
5-100	10X
101-1000	No dilution

1. Add 2.3 uL of NEBNext Adaptor (diluted, Step 4) to each sample, pipette up and down gently and spin down.
2. Incubate for 15 min at room temperature or an open-lid thermocycler set to 20C.
3. Add 3 uL of USER enzyme and gently pipette.
4. Incubate in a thermocycler (80.3 uL):

Step	Temperature	Time (min)	Lid temperature
Incubation	37	15	47
Hold	4	Hold

1. Spin down for 1 min.
   • You may take a break here and store samples at -20C until ready.

Magnetic bead purification (45 min)

1. Let the magnetic beads thaw to room temperature, then vortex.
2. Transfer PCR products to a new 1.5 mL tube.
3. Choose bead/sample ratio based on the your fragment size in the chart below.
   • After adaptor ligation, the fragment size increases by 66 nt
   • After barcoding, another 66 nt are added
   • NEB uses a rough estimate of 120 nt being added
   • For AMPure XP beads:
   • For Zymo Select-a-size MagBeads:

Round	Input from	Input vol (uL)	Bead vol. (uL)	Amount of water for resuspention (uL)	Amount of sample to transfer (uL)
1	Adaptor ligation	80.3	80.3 x ratio	23	20
2	Barcoding	50	50 x ratio	53	50
3	Magnetic bead purification, Round 2	50	50 x ratio	23	20

1. Add the computed bead amount to each sample and pipette up and down.
2. Incubate at room temperature for 5 min.
3. In the meantime, make fresh 80% ethanol (need 200 uL x 2 times x 3 rounds = 1.2 mL per sample):

Component	Per reaction + 5% (mL)
100% ethanol	1.01
Milli-Q	.25

1. Place the strip tube inside the magnet rack until supernatant is clear (~5 min), then remove and discard the supernatant, ensuring no beads are discarded.
2. Keeping the plate on the magnet, add 200 uL freshly prepared 80% ethanol to each sample, wait 30 seconds, and discard ethanol. 
3. Repeat the previous step. 
4. Draw out any remaining ethanol with a 10 uL pipette.
5. Air-dry beads for 7 minutes at room temperature on the magnet with the lids open, but do not let the beads dry out.
   • The beads should look brown and glossy. If they look light brown and cracked, you've dried them for too long. This may result in the loss of the recovered DNA amount.
6. Remove the strip of tubes from the magnet and add 23 uL (or 53 uL in Round 2) of water to each sample to re-suspend the beads, pipette, and incubate at room temperature for 2 minutes.
7. Place the strip to the magnet until supernatant is clear (~5 minutes). 
8. Transfer 20 uL (or 50 uL in Round 2) of each sample to new tubes (PCR tubes for barcoding or 1.5 mL tubes for purification rounds 2 and 3), making sure not to transfer any beads. 

Barcoding (45 min)

1. Use Qubit High Sensitivity kit to determine DNA concentration, using 5 uL of each sample and a high sensitivity dye.
   • 15 uL remain for barcoding.
   • Record concentrations in the Sample Tracking Sheet.
2. Do one of the following:
   1. If using ==96 Index Primers== (from a 96-well plate):
      1. Thaw the NEBNext Index/Universal Primer Mix plate for 10-15 min at room temperature.
      2. Remove the hard plastic plate cover. Briefly centrifuge the plate (280 × g for ~1 min) to collect all of the primer at the bottom of each well.
      3. Using a pipette, add 25 µL of NEBNext Ultra II Q5 Master Mix into each sample.
      4. Add 10 uL of indexed primer from the plate to each sample.
         • The mix contains both a universal i5 primer and a specific i7 primer.
         • Note down the i7 barcodes used in the Sample Tracking Sheet.
   2. If using an ==Index Primer Set== (from individual tubes containing a single index):
      1. Prepare a Master Mix.
      2. Add 30 uL of the Master Mix into each sample.
      3. Choose your Index (i7) primers from an Index Primer Set, note them down, add 5 uL to each sample, and pipette up and down.

Component	Per reaction (uL)	Per reaction + 5% (uL)
NEBNext Ultra II Q5 Master Mix	25	26.25
Universal (i5) primer	5	5.25
Total	30	31.5

1. Incubate in a thermocycler (50 uL):

Stage	Name	Temperature	Duration (sec)	Number of cycles
1	Initial Denaturation	98	30	1
2	Denaturation	98	10	See table below
2	Annealing	65	75
3	Final Extension	65	300	1
3	Hold	4	Hold	1

Total DNA amount (ng)	Number of PCR Cycles
0.5-10	12
10-99	8
100-499	6
500-1000	4

1. Spin down for 1 min.
   • You may take a break here and store samples at -20C until ready.

Final preparation

1. Purify with magnetic beads again twice (Rounds 2 and 3)
2. Check the concentration with the Qubit High Sensitivity kit high sensitivity dye before proceeding to Bioanalyzer.
   • Record concentrations in the Sample Tracking Sheet.
3. Dilute samples to about 1 ng/uL.
4. Run 1 uL of each sample on an Agilent Bioanalyzer High Sensitivity DNA chip. After the run, export the ladder and each of your samples as a csv file in order to produce a plot with the measured peaks.
5. Complete information in the Sequencing projects tab and in the Genecore submission form (where Server name = Enzyme_library_S1_time)
6. Provide about 2 ng of each of your sample for sequencing (using Bioanalyzer's estimated concentration of everything but the markers).
7. Store the final samples at -20C.

Resources

• Source protocol
• Improved Protocols for Illumina Sequencing - PMC
• Illumina_Sequencing_Overview_15045845_D.pdf
• Index primers
• NEBNext® Multiplex Oligos for Illumina® (96 Index Primers)