TITLE: Combining NexteraXT and Native BK to Identify Gene Expression Differences in Staphylococcus aureus Strains
AUTHORS: Katelyn Roberts

[DESCRIPTION]
The purpose of these two protocols combined is to improve the accuracy of genome assembly and gene expression analysis, especially when investigating the mechanisms of antibiotic resistance. Four samples are chosen including MSSA, MRSA, VSSA, and VRSA, with MSSA/VSSA acting as genomic controls and the other two samples acting as the antibiotic resistant genomes. 

Long reads provide the ability to span large, complex, and repetitive regions of the genome, which is critical when identifying structural variations and fully assembling genomic regions like those harboring antibiotic resistance genes. Short reads, on the other hand, offer high base-level accuracy and depth, which is important for identifying small mutations or variants, such as SNPs or indels, that may contribute to antibiotic resistance or virulence.

Once both sequencing runs are complete, a workflow aligning both kits results could reveal variations in resistance gene loci and differences in gene expression that influence pathogenicity, with short read sequencing filling in the gaps of information that long read sequencing may have left.  

[GUIDELINES]
Epi2me Workflow Guidelines:
Native Barcoding: AMR Gene Analysis

ARIBA Workflow:
Germline Pipeline 

Comparing Datae:
Combine files using SamTools
Cross Validate findings from Gene resistance results with ARIBA and AMR Gene 

[STEPS]
SECTION: Lysis and Confirmation
2. Measure Optical Density at 600nm and record the value as a reference. 
SECTION: Lysis and Confirmation
3. Dilute lysostaphin to a final concentration of 100 µg/µL  in nuclease-free water.

SECTION: Lysis and Confirmation
4. Store the lysostaphin solution on ice until ready for use.
SECTION: Lysis and Confirmation
5. Add 2 µL2 µL  of lysostaphin to 198 µL of the bacterial suspension in nuclease-free water (for a total volume of 200200 µL  µL).

SECTION: Lysis and Confirmation
6. Incubate the mixture at 37°37 °C 30 min  to allow efficient lysis of the Staphylococcus aureus cells.

SECTION: Lysis and Confirmation
7. Record another set of OD600 values to confirm successful lysis after incubation. 
Alternatively, take a small aliquot and observe the cell structure under a microscope to confirm lysis.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
9. Thaw the AMPure Beads and mix by vortexing. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
10. Prepare the NEBNext FFPE DNA Repair Mix and NEBNext Ultra II End Repair / dAtailing Module reagents in accordance with manufacturer’s instructions, and place on ice.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
11. In clean 0.2 ml thin-walled PCR tubes (or a clean 96-well plate), prepare your DNA samples. For this experiment and 4 barcodes, 1000ng of sample should be aliquoted. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
12. Make up each sample to 11 µl using nuclease-free water. Mix gently by pipetting and spin down
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
13. Combine the following components per tube/well


      



Between each addition, pipette mix 10 - 20 times. 
Reagent Volume DNA sample 12 µl  
NEBNext FFPE DNA Repair Buffer 0.875 µl 
Ultra II End-prep Reaction Buffer 0.875 µl 
Ultra II End-prep Enzyme Mix 0.75 µl 
NEBNext FFPE DNA Repair Mix 0.5 µl 
Total 15ul
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
14. Ensure the components are thoroughly mixed by pipetting and spin down in a centrifuge. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
15. Using a thermal cycler, incubate at 20°C for 5 minutes and 65°C for 5 minutes.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
16. Transfer each sample into a clean 1.5 ml Eppendorf DNA LoBind tube. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
17. Resuspend the AMPure XP beads (AXP) by vortexing.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
18. Add 15 µl of resuspended AMPure XP Beads (AXP) to each end-prep reaction and mix by flicking the tube. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
19. Incubate on a Hula mixer (rotator mixer) for 5 minutes at room temperature. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
20. Prepare sufficient fresh 80% ethanol in nuclease-free water for all of your samples. Allow enough for 400 µl per sample, with some excess. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
21. Spin down the samples and pellet the beads on a magnet until the eluate is clear and colourless. Keep the tubes on the magnet and pipette off the supernatant. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
22. Keep the tube on the magnet and wash the beads with 200 µl of freshly prepared 80% ethanol without disturbing the pellet. Remove the ethanol using a pipette and discard. If the pellet was disturbed, wait for beads to pellet again before removing the ethanol.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
23. Repeat the previous step. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
24. Briefly spin down and place the tubes back on the magnet for the beads to pellet. Pipette off any residual ethanol. Allow to dry for 30 seconds, but do not dry the pellets to the point of cracking. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
25. Remove the tubes from the magnetic rack and resuspend the pellet in 10 µl nuclease free water. Spin down and incubate for 2 minutes at room temperature.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
26. Pellet the beads on a magnet until the eluate is clear and colourless.
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
27. Remove and retain 10 µl of eluate into a clean 1.5 ml Eppendorf DNA LoBind tube. Dispose of the pelleted beads
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
28. Quantify 1 µl of each eluted sample using a Qubit fluorometer
SECTION: Barcode Ligation
30. Prepare the NEB Blunt/TA Ligase Master Mix according to the manufacturer's instructions, and place on ice: 
SECTION: Barcode Ligation
31. Thaw the reagents at room temperature.
SECTION: Barcode Ligation
32. Spin down the reagent tubes for 5 seconds. 
SECTION: Barcode Ligation
33. Ensure the reagents are fully mixed by performing 10 full volume pipette mixes.
SECTION: Barcode Ligation
34. Thaw the EDTA at room temperature and mix by vortexing. Then spin down and place on ice. 
SECTION: Barcode Ligation
35. Thaw the Native Barcodes (NB01-24) required for your number of samples at room temperature. Individually mix the barcodes by pipetting, spin down, and place them on ice. 
SECTION: Barcode Ligation
36. Select a unique barcode for each sample to be run together on the same flow cell. Up to 24 samples can be barcoded and combined in one experiment. Please note: Only use one barcode per sample.
SECTION: Barcode Ligation
37. In clean 0.2 ml PCR-tubes or a 96-well plate, add the reagents in the following order per well: Between each addition, pipette mix 10 - 20 times.
7.5ul of Reagent Volume 
2.5ul of Native Barcode (NB01-24) 
10ul of Blunt/TA Ligase Master

SECTION: Barcode Ligation
38. Thoroughly mix the reaction by gently pipetting and briefly spinning down. 
SECTION: Barcode Ligation
39. Incubate for 20 minutes at room temperature. 
SECTION: Barcode Ligation
40. Add the following volume of EDTA to each PCR tube or well and mix thoroughly 
EDTA clear cap: 2ul
EDTA blue cap: 4ul
SECTION: Barcode Ligation
41. Pool all the barcoded samples in a 1.5 ml Eppendorf DNA LoBind tube

Total Volume for preps:
EDTA Clear Cap - 22ul,     For 6 samples: 132ul
EDTA Blue Cap - 24 ul,      For 6 samples: 144 ul

SECTION: Barcode Ligation
42. Resuspend the AMPure XP Beads (AXP) by vortexing.
SECTION: Barcode Ligation
43. Add 0.4X AMPure XP Beads (AXP) to the pooled reaction, and mix by pipetting.

Volume of AXP for preps:
EDTA Clear Cap - 9ul,  For 6 samples: 53ul
EDTA Blue Cap - 10ul,  For 6 samples: 60ul
SECTION: Barcode Ligation
44. Incubate on a Hula mixer (rotator mixer) for 10 minutes at room temperature. 
SECTION: Barcode Ligation
45. Prepare 2 ml of fresh 80% ethanol in nuclease-free water.
SECTION: Barcode Ligation
46. Spin down the sample and pellet on a magnet for 5 minutes. Keep the tube on the magnetic rack until the eluate is clear and colourless, and pipette off the supernatant.
SECTION: Barcode Ligation
47. Keep the tube on the magnetic rack and wash the beads with 700 µl of freshly prepared 80% ethanol without disturbing the pellet. Remove the ethanol using a pipette and discard. 
SECTION: Barcode Ligation
48. Repeat the previous step. 
SECTION: Barcode Ligation
49. Spin down and place the tube back on the magnetic rack. Pipette off any residual ethanol. Allow the pellet to dry for ~30 seconds, but do not dry the pellet to the point of cracking. 
SECTION: Barcode Ligation
50. Remove the tube from the magnetic rack and resuspend the pellet in 35 µl nucleasefree water by gently flicking. 
SECTION: Barcode Ligation
51. Incubate for 10 minutes at 37°C. Every 2 minutes, agitate the sample by gently flicking for 10 seconds to encourage DNA elution. 
SECTION: Barcode Ligation
52. Pellet the beads on a magnetic rack until the eluate is clear and colourless.
SECTION: Barcode Ligation
53. Remove and retain 35 µl of eluate into a clean 1.5 ml Eppendorf DNA LoBind tube. 
SECTION: Barcode Ligation
54. Quantify 1ul of eluted sample using a Qubit fluorometer. 
SECTION: Adapter Ligation and Cleanup
56. Prepare the NEBNext Quick Ligation Reaction Module according to the manufacturer's instructions, and place on ice:
SECTION: Adapter Ligation and Cleanup
57. Thaw the reagents at room temperature. 
SECTION: Adapter Ligation and Cleanup
58. Spin down the reagent tubes for 5 seconds. 
SECTION: Adapter Ligation and Cleanup
59. Ensure the reagents are fully mixed by performing 10 full volume pipette mixes. 

Note: Do NOT vortex the Quick T4 DNA Ligase. The NEBNext Quick Ligation Reaction Buffer (5x) may have a little precipitate. Allow the mixture to come to room temperature and pipette the buffer up and down several times to break up the precipitate, followed by vortexing the tube for several seconds to ensure the reagent is thoroughly mixed.
SECTION: Adapter Ligation and Cleanup
60. Spin down the Native Adapter (NA) and Quick T4 DNA Ligase, pipette mix and place on ice. 
SECTION: Adapter Ligation and Cleanup
61. Thaw the Elution Buffer (EB) at room temperature and mix by vortexing. Then spin down and place on ice.
SECTION: Adapter Ligation and Cleanup
62. Thaw either Long Fragment Buffer (LFB) or Short Fragment Buffer (SFB) at room temperature and mix by vortexing. Then spin down and place on ice. 
SECTION: Adapter Ligation and Cleanup
63. In a 1.5 ml Eppendorf LoBind tube, mix in the following order

30 ul of Reagent Volume Pooled barcoded sample
5ul of Native Adapter (NA)
10ul of NEBNext Quick Ligation Reaction Buffer (5X)
5ul of Quick T4 DNA Ligase
For a total volume of 50ul
SECTION: Adapter Ligation and Cleanup
64. Thoroughly mix the reaction by gently pipetting and briefly spinning down.
SECTION: Adapter Ligation and Cleanup
65. Incubate the reaction for 20 minutes at room temperature.
SECTION: Adapter Ligation and Cleanup
66. Resuspend the AMPure XP Beads (AXP) by vortexing. 
SECTION: Adapter Ligation and Cleanup
67. Add 20 µl of resuspended AMPure XP Beads (AXP) to the reaction and mix by pipetting. 
SECTION: Adapter Ligation and Cleanup
68. Incubate on a Hula mixer (rotator mixer) for 10 minutes at room temperature.
SECTION: Adapter Ligation and Cleanup
69. Spin down the sample and pellet on the magnetic rack. Keep the tube on the magnet and pipette off the supernatant. 
SECTION: Adapter Ligation and Cleanup
70. Wash the beads by adding either 125 μl Long Fragment Buffer (LFB) or Short Fragment Buffer (SFB). Flick the beads to resuspend, spin down, then return the tube to the magnetic rack and allow the beads to pellet. Remove the supernatant using a pipette and discard. 
SECTION: Adapter Ligation and Cleanup
71. Repeat the previous step. 
SECTION: Adapter Ligation and Cleanup
72. Spin down and place the tube back on the magnet. Pipette off any residual supernatant. 
SECTION: Adapter Ligation and Cleanup
73. Remove the tube from the magnetic rack and resuspend pellet in 15 µl Elution Buffer (EB). 
SECTION: Adapter Ligation and Cleanup
74. Spin down and incubate for 10 minutes at 37°C. Every 2 minutes, agitate the sample by gently flicking for 10 seconds to encourage DNA elution.
SECTION: Adapter Ligation and Cleanup
75. Pellet the beads on a magnet until the eluate is clear and colourless, for at least 1 minute.
SECTION: Adapter Ligation and Cleanup
76. Remove and retain 15 µl of eluate containing the DNA library into a clean 1.5 ml Eppendorf DNA LoBind tube.
SECTION: Adapter Ligation and Cleanup
77. Depending on your DNA library fragment size, prepare your final library in 12 µl of Elution Buffer (EB). 
Fragment library length:
Flow cell loading amount: Very short (<1 kb) 100 fmol 
Short (1-10 kb) 35–50 fmol
 Long (>10 kb) 300 ng
SECTION: Lysis and Confirmation
1. The purpose of the lysis addition and modification is due to the thickness of Staphylococcus aureus's cell wall. In order to achiever desired results for the following library kits, samples should have high molecular weight DNA, the only way to achieve this is by lysing the cells using lysostaphin. Lysostaphin ensures the cells are lysed efficiently for gDNA isolation (whereas normal gram-negative techniques will not have).  Quantification is used to ensure successful lysis and the ability to proceed onto extraction and library prep. 
SECTION: Loading and Sequencing Settings
78. Loading should be done on a MinION flow cell according to these protocols (Ligation sequencing gDNA - Native Barcoding Kit 24 V14 (SQK-NBD114.24) (NBE_9169_v114_revR_15Sep2022) (nanoporetech.com) and sequencing should be done on a GridION device. 
SECTION: Loading and Sequencing Settings
79. Using the MinKNOW software, select Super Accurate Basecalling, the Native Barcoding Kit, 72 hour run time, and enable quality score filtering with a threshold of Phred > 9
SECTION: Nextera XT: Tagmenting DNA
81. Prepare the following consumables:

 

SECTION: Nextera XT: Tagmenting DNA
81.1. Save the following tag program on the thermocycler:
Choose the preheat lid option and set to 100C
Set the reaction volume to 50ul
55C for 5 minutes
Hold at 10C
SECTION: Nextera XT: Tagmenting DNA
82. Add the following volumes in the order listed to each well of a new 96-well PCR plate.
•	TD (10 µl) 
•	1 ng DNA (5 µl)
SECTION: Nextera XT: Tagmenting DNA
83. Pipette to mix.
SECTION: Nextera XT: Tagmenting DNA
84. Add 5 µl ATM to each well.
SECTION: Nextera XT: Tagmenting DNA
85. Pipette 10 times to mix, and then seal the plate.
SECTION: Nextera XT: Tagmenting DNA
86. Centrifuge at 280 × g at 20°C for 1 minute.
SECTION: Nextera XT: Tagmenting DNA
87. Place on the preprogrammed thermal cycler and run the TAG program. When the program reaches 10°C, immediately proceed to step 7 because the transposome is still active.
SECTION: Nextera XT: Tagmenting DNA
88. Add 5 µl NT to each well.
SECTION: Nextera XT: Tagmenting DNA
89. Pipette 10 times to mix, and then seal the plate.
SECTION: Nextera XT: Tagmenting DNA
90. Centrifuge at 280 × g at 20°C for 1 minute.
SECTION: Nextera XT: Tagmenting DNA
91. Incubate at room temperature for 5 minutes.
SECTION: Amplify Libraries
92. This step amplifies the tagmented DNA using a limited-cycle PCR program. The PCR step adds the i7 adapters, i5 adapters, and sequences required for sequencing cluster generation. This is also a safe stopping point, the plate can be sealed and stored at 2C-8C for up to 2 days. 
SECTION: Amplify Libraries
93. Prepare the following consumables:

 

SECTION: Amplify Libraries
93.1. Save the following NXT PCR program on a thermal cycler:
Choose the preheat lid option and set to 100C
Set the reaction volume to 50ul
72C for 3 minutes
95C for 30 minutes
12 cycles of:
(95C for 10 seconds )
(55C for 30 seconds )
(72C for 30 seconds )
72C for 5 minutes
Hold at 10C
SECTION: Amplify Libraries
94. Add the following index adapter volumes per sample according to your index adapter kit type.

 

SECTION: Amplify Libraries
95. Add 15 µl NPM to each well.
SECTION: Amplify Libraries
96. Pipette 10 times to mix, and then seal the plate.
SECTION: Amplify Libraries
97. Centrifuge at 280 × g at 20°C for 1 minute.
SECTION: Amplify Libraries
98. Place on the preprogrammed thermal cycler and run the NXT PCR program.
SECTION: Clean Up Libraries
99. This section uses single-sided bead purification to purify amplified libraries.
SECTION: Clean Up Libraries
100. Prepare the following consumables: 

 

SECTION: Clean Up Libraries
100.1. Prepare fresh 80% EtOH from absolute ethanol. 
SECTION: Clean Up Libraries
101. Centrifuge at 280 × g at 20°C for 1 minute to collect contents at the bottom of the well.
SECTION: Clean Up Libraries
102. Transfer 50 µl supernatant from each well of the PCR plate to corresponding wells of a new MIDI plate. 

Note: The ratio of supernatant to volume of IPB is 3:2. If you transfer less than 50 µl supernatant, adjust the volume of IPB accordingly.
SECTION: Clean Up Libraries
103. If you are using standard DNA input, add 30 µl IPB to each well containing supernatant.
SECTION: Clean Up Libraries
104. Seal the plate, and then use a plate shaker at 1800 rpm for 2 minutes.
SECTION: Clean Up Libraries
105. Incubate at room temperature for 5 minutes.
SECTION: Clean Up Libraries
106. Place on the magnetic stand and wait until the liquid is clear (~2 minutes).
SECTION: Clean Up Libraries
107. Without disturbing the beads, remove and discard all supernatant.
SECTION: Clean Up Libraries
108. Wash two times with 200 µl 80% EtOH as follows. 
a. With the plate on the magnetic stand, add 200 µl fresh 80% EtOH without mixing. 
b. Incubate for 30 seconds. 
c. Without disturbing the beads, remove and discard all supernatant.
SECTION: Clean Up Libraries
109. Use a 20 µl pipette to remove and discard residual EtOH.
SECTION: Clean Up Libraries
110. Air-dry on the magnetic stand for 15 minutes.
SECTION: Clean Up Libraries
111. Remove from the magnetic stand.
SECTION: Clean Up Libraries
112. Add 52.5 µl RSB to the beads.
SECTION: Clean Up Libraries
113. Seal the plate, and then use a plate shaker at 1800 rpm for 2 minutes.
SECTION: Clean Up Libraries
114. Incubate at room temperature for 2 minutes.
SECTION: Clean Up Libraries
115. Place on the magnetic stand and wait until the liquid is clear (~2 minutes).
SECTION: Clean Up Libraries
116. Transfer 50 µl supernatant to a new 96-well PCR plate.
SECTION: Check Library Quality
117. Library quality should be checked to determine if PCR was successful and if there were any contaminnats in the sample. 
SECTION: Check Library Quality
118. Run 1 µl undiluted library on an Agilent Technology 2100 Bioanalyzer using a High Sensitivity DNA kit.
Typical libraries show a broad size distribution of ~250–1000 bp, as shown in the top panel. Various libraries can be sequenced with average fragment sizes as small as 250 bp or as large as 1500 bp.
SECTION: Check Library Quality
118.1. 
 

SECTION: Quality Metrics
119. This section is to troubleshoot library size and concentration inconsistencies. 
SECTION: Quality Metrics
119.1. Checking Library Size:
Larger molecules cluster less efficiently than smaller molecules. If the fragment size after tagmentation is larger than expected, low cluster numbers are possible. The inverse is also true. The average expected library size after tagmentation is between 400 bp and 1.2 kb.
Check the library size with a high sensitivity Bioanalyzer trace after the PCR cleanup step. Look for a long low plateau. Alternatively, PCR-amplify the library with qPCR primers and run the product on an agarose gel. The sequence for these primers is available in the Sequencing Library qPCR Quantification Guide (document # 11322363).

Short libraries indicate too little input DNA: Requantify the input DNA with a fluorometric method. Start with 10%–25% more input DNA. If the library peak is below 400 bp and you want to continue with this library, dilute the library further.

Long libraries indicate too much input DNA or inhibitor presence: Start with less input DNA, make sure that the input DNA is free from inhibitors, and repeat the quantification step.
SECTION: Quality Metrics
119.2. Check library concentration: 
Bead-based normalization is most efficient when the library yield after amplification is 10–15 nM, or higher. Measure library concentration using high sensitivity dsDNA Qubit after library cleanup, and measure library size with a Bioanalyzer to calculate molarity.
If you are starting with high-quality DNA and see low yield after library cleanup, there are possible issues with IPB cleanup or the amplification step. If results show either condition, confirm proper storage of the PCR master mix at ‑25°C to ‑15°C in a no-frost freezer. Confirm minimal freeze-thaw cycles.
SECTION: Normalize Libraries
120. This step normalizes the quantity of each library made with Nextera XT Index v2 or Nextera XT Index Kits to ensure more equal library representation in the pooled library.
SECTION: Normalize Libraries
121. Prepare the following consumables:
 

SECTION: Normalize Libraries
122. Transfer 20 µl supernatant from each well of the PCR plate to the corresponding well of a new MIDI plate.
SECTION: Normalize Libraries
123. Combine the following volumes in a 15 ml conical tube to prepare the LN master mix. Multiply each volume by the number of samples being processed. 
•	LNA1 (46 µl) 
•	LNB1 (8 µl) 
Reagent coverage is included in the volume to ensure accurate pipetting.
SECTION: Normalize Libraries
124. Pipette 10 times to mix.
SECTION: Normalize Libraries
125. Pour the LN master mix into a trough.
SECTION: Normalize Libraries
126. Use a 200 µl multichannel pipette to transfer 45 µl LN master mix to each well.
SECTION: Normalize Libraries
127. Seal the plate, and then use a plate shaker at 1800 rpm for 30 minutes.
SECTION: Normalize Libraries
128. Place on the magnetic stand and wait until the liquid is clear (~2 minutes).
SECTION: Normalize Libraries
129. Without disturbing the beads, remove and discard all supernatant.
SECTION: Normalize Libraries
130. Wash two times with 45 µl LNW1 as follows. 
a.	Add 45 µl LNW1 to each well. 
b.	Seal the plate, and then use a plate shaker at 1800 rpm for 5 minutes. 
c.	Place on the magnetic stand and wait until the liquid is clear (~2 minutes). 
d.	Without disturbing the beads, remove and discard all supernatant.
SECTION: Normalize Libraries
131. Add 30 µl 0.1 N NaOH to each well.
SECTION: Normalize Libraries
132. Seal the plate, and then use a plate shaker at 1800 rpm for 5 minutes.
SECTION: Normalize Libraries
133. Add 30 µl LNS1 to each well of a new 96-well PCR plate labeled SGP.
SECTION: Normalize Libraries
134. After the 5 minute elution completes, make sure that all samples in the MIDI plate are resuspended. If they are not, resuspend as follows.  
a.	Pipette 10 times to mix or lightly tap the sample plate on the bench. 
b.	Seal the plate, and then use a plate shaker at 1800 rpm for 5 minutes.
SECTION: Normalize Libraries
135. Place on the magnetic stand and wait until the liquid is clear (~2 minutes).
SECTION: Normalize Libraries
136. Transfer 30 µl supernatant from each well of the MIDI plate to the corresponding well of the SGP plate.
SECTION: Normalize Libraries
137. Seal the sample plate, and then centrifuge at 1000 × g for 1 minute. At this point, the libraries are single-stranded DNA, which resolves poorly on an agarose gel or Bioanalyzer chip. For quality control, use the double-stranded DNA saved from step 17 of the cleanup procedure.
SECTION: Loading and Sequencing
138. Load libraries according to loading protocols for the MiSeq platform.  (https://support.illumina.com/content/dam/illumina-support/documents/documentation/system_documentation/miseq/miseq-system-guide-15027617-06.pdf). 
Sequencing setting sshould be set to paired end reads and a target of 30x-100x coverage. 
SECTION: Nextera XT: Tagmenting DNA
80. This sections fragments and tags DNA with adaptor sequences. 
SECTION: Adapter Ligation and Cleanup
55. This sections is meant to ligate adaptors onto the ends of DNA sequences and enrich for DNA fragments larger than 3kb
SECTION: Barcode Ligation
29. Barcodes are ligated onto the end prepped DNA sequences. 
SECTION: DNA Repair and End Prep (Native Barcoding Kit)
8. This sections ensures DNA is repaired and prepares the DNA ends by creating blunt ends or A-tails, making the fragments compatible for ligation with sequencing adapters.