Major version notes

This is a protocol based on Kapa LTP Preparation Kit manual KR0453 - v3.13
The main difference centers around the use of 1/2 volume in each reaction so as to get twice as many sample preps for the same price.
This was validated on 2 extremely similar samples from Gabo by DED in July of 2014, and the final data was indistinguishable between the 1x volume sample and the 0.5x volume sample.
This has subsequently been used in more than a dozen (4-15-15) other preps without incident.
The default fragmentation protocol is now NEB fragmentase rather than Covaris.
This protocol is for standard applications. Please click here for rare variant within a population protocol.

Before Beginning and Important Notes

1. Low retention (maximum recovery) tubes and filter tips should be used at every step. Regular tips commonly used for EtOH removal during wash steps.
2. Before starting make sure ligated adapters are available in cardboard box labeled 'Anealed adapters'.
   Importance of this box/these tubes never being warmed above RT cannot be underscored enough
3. WFI water should be used for all steps where water is needed. Aliquots of water should be taken from bottle to avoid contamination.
4. A 2 barcode system comprising internal and external barcodes is currently being employed (it allows to mix different samples in a single tube for multiplexing the sequencing, thus economical; it is different than dual barcoding strategies which use 2 different external barcodes). There are 8 annealed adapters, each corresponding to 1 of 8 internal barcodes (IBC). External barcode(s) (EBC) are added to samples in the PCR step. Identical EBC can and should be pooled together to meet the sequencing core's minimum requirement of 10million reads per sample this should give ~70-80x coverage ("single-end" reads; see Coverage calculator) of E. coli genomes which is the current standard.
   NEVER pool identical IBC as this type of pooling can not be reassigned at the data analysis step.
5. Sample should never be vortexed, always thoroughly mixed by pipetting. This is done to avoid sample getting near the top of the tube and increasing contamination risk during magnetic bead clean up.
6. "Freshly prepared 80% EtOH" for cleanup reactions should be prepared daily, and be made combining 100% EtOH with WFI water.
7. Always have the next master mix prepared to add to the tubes after wash. Never leave the beads in tubes without liquid on them for very long (< 3 min) as they will dry and ruin your prep.
8. Several different alternative options are available, steps which have alternative strategies are marked as such.

Adapter Anealation

Note, this step is not necessary each time. Check the C lab -20, NGS 2 rack, "EBC + IBC Design Anealed Adapter" paper box for annealed adapters (small tubes not IDT tubes). If annealed primers are old, best to trash and remake. Do not trash IDT tubes.

1. Mix the following in PCR tube for a 50µM adapter concentrated stock:
   1. 10µl 100µM IBCF#
   2. 10µl 100µM IBCR#
   • Using mismatched identifier numbers will cause complete failure of ligation, or terrible data.
2. Use thermocycler with following conditions:
   1. 97C 2minutes
   2. 97C 1minute
      1. Repeat previous step for 72 total cycles at -1C per cycle
   3. 25C 5 minutes
      After this step, adapters should NEVER be warmed above 25C.
3. Alliquot 2µl to 10 PRECHILLED 600µl low retention eppendorf tubes. Make sure tubes are labeled.
4. Store barcodes 'anealed adpaters' box.

Harvest DNA and determine Concentration

1. Harvest DNA from sample using Invitrogen PureLink Genomic DNA kit using standard protocol.
2. It is important to elute in elution buffer rather than water.
3. Use Qbit to determine concentration.

Fragment DNA

• The goal of this step is to transfer 500ng of sheared DNA in 25µl to a low retention eppendorf tube.

NEB Fragmentase®

* Protocol employed by Mike due to large quantity of samples. Has since been used by iGEM, Dacia and Gabo (contributor to the protocol below). * Subsequent steps will assume your samples were fragmented with Fragmentase. If this is not the case, other changes are needed

1. Pull Ampure beads to allow to warm to room temperature.
2. Set Fragmentase on ice and set Fragmentase 10X buffer at room temp to allow to thaw. For reproducible results, pre-vortex (~2 seconds) all reactants prior to use.
3. Set heat block (with water on the wells) to 37°C and corroborate temperature with a thermometer. *Beware some heat-blocks may have a timer and will stop prematurely if not set properly.
4. Combine the following in a PCR tube:
   

Item	Amount
DNA	500 ng (minimal) - 1 µg (ideal)
Fragmentase 10x buffer	2 µl
Fragmentase enzyme mix (last ingredient to add)	2 µl
diH2O	complete to 20µL

*Important:* Fragmentase is viscous, so pay attention you are not adding extra and remove excess by lightly tapping the pippette tip against the inside of Fragmentase tube. When pippetting into the reaction mix, pipette up and down several times making sure complete transfer has been achieved.

1. Vortex the rxn mix for ~2 sec
2. Incubate at 37°C for ~27 minutes.
   Note: A typical 20uL reaction with 1ug gDNA will take ~27 minutes for ~300bp fragments (tested with DNA eluted in Tris pH 8.5 with ~400ng/mL concentration). A test reaction should be performed with a sample that has been extracted with exactly the same conditions (i.e., same method, reactants and elution buffer) as the samples that will undergo sequencing prep. This is critical, since small variations in buffers or DNA extraction methods can dramatically affect digestion times required for the desired fragment size. Run 500ng-1ug from these test reactions on a gel to determine results. Preferably, fragments will be anywhere from 300-500bp; fragmentation to smaller than 300bp should be avoided.
3. STOP rxn: Fragmentase reactions must be stopped (5µL of EDTA 0.5 M pH 8.0) to prevent further digestion of your product. Best way (fast and precise) to do this is to have another set of PCR tubes ready with 5µL EDTA pre-chilled on ice and a promptly transferring the completed reactions into them (you can use a P100 pipette set to 30µL) making sure to pipette up and down to mix well.

Other notes:
The Fragmentase reaction is not linear, but its efficiency can be roughly estimated by plugging time (t) or base pairs (size) in the following formulas:
52000*(t^-1.55) = estimated average fragment size
10^(LOG(size/52000)/-1.55) = time in minutes

You can estimate digestion time by entering desired fragment size in the box below:

=

estimated minutes

Half-Fragmentase
Using half-amount Fragmentase (1uL in 20uL rxn) can be performed at a little over double the time (2.4X). For example, for half-fragmentase reactions with 1ug gDNA in 20uL, we've seen that it took 32 minutes to fragment to ~900bp fragments and 62-72 minutes to achieve ~300 bp size.
Don't worry if you get a large smear, as long as enough material is observed <500bp, next steps will narrow fragment size range with size selection.

Post FRAGMENTASE clean up.

1. Add 60µl of room temperature Ampure beads to the stopped 25µL Fragmentase rxn
2. Mix by pipetting up and down till homogeneous mixture.
3. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
4. Place tubes on magnet until liquid is clear (usually ~5 minutes).
5. Carefully remove and discard supernatant without disturbing the beads.
6. Add 200 µl of 80% EtOH.
7. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
8. Repeat for a 2nd wash.
9. Allow beads to dry at room temperature for all ethanol to evaporate.
   1. ethanol carryover will interfere with subsequent steps, and over-drying of beads may damage them (typically characterized by cracks appearing in the beads)
10. Remove beads from magnet.
11. Add 25µL of WFI Water
12. Make sure all beads come off the wall and into solution.

End Repair DNA

1. Devin and Tyler of the iGEM team have confirmed that this reaction CAN be completed in the presence of beads.
2. Move blue heat block to -4C, and set to 20C before starting so heat block will adjust temperature accordingly.
3. Place PEG on benchtop to warm to room temperature depending on if beads are already in your reaction.
   OTHER fragmentation methods may require use of ampure beads not PEG
   
4. To each sample tube add:
   

Item	Amount
WFI water	4µl
10x KAPA End Repair Buffer	3.5µl
KAPA End Repair Enzyme Mix	2.5µl

1. Pipette to mix.
2. Incubate 20C for 30 minutes
3. Remove tubes from block, increase temperature to 30C (necessary for A-tailing steps below)

Wash

OTHER fragmentation methods may require use of ampure beads not PEG

1. To each 35µl reaction, add 60µl of room temperature PEG/NaCl Solution.
   There should be beads in your tube
   
2. Mix by pipetting up and down till homogeneous mixture.
3. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
4. Place tubes on magnet until liquid is clear (usually ~5 minutes).
5. Carefully remove and discard supernatant without disturbing the beads.
6. Add 200 µl of 80% EtOH.
7. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
8. Repeat for a 2nd wash.
9. Allow beads to dry at room temperature for all ethanol to evaporate.
   1. ethanol carryover will interfere with subsequent steps, and over-drying of beads (typically characterized by cracks appearing in the beads)
10. Remove beads from magnet.

A-Tailing Reaction

1. Make sure heat block is set to 30C and holding.
2. Pull PEG/NaCL SPRI Solution and place on bench to warm to room temperature
3. To each tube add the following:
   

Item	Amount
WFI water	21µl
10x KAPA A-Tailing Buffer	2.5µl
KAPA A-Tailing Enzyme	1.5µl

1. Mix by pipetting till homogeneous solution
2. Incubate 30 minutes at 30C.
3. Remove tubes from block.
4. Remove block from base, and place on bench in cold room. Set block to 20C. Check back periodically (at least 15 minutes rxn time needed) for block to stay at or below 20C once back in base. Ice buckets can be used instead of cold room bench for better heat dispersal.

Wash

1. To each 25µl reaction, add 45µl PEG/NaCl Solution.
2. Mix by pipetting up and down till homogeneous mixture.
3. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
4. Place tubes on magnet until liquid is clear (usually ~5 minutes).
5. Carefully remove and discard supernatant without disturbing the beads.
6. Add 200 µl of 80% EtOH.
7. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
8. Repeat for a 2nd wash.
9. Allow beads to dry at room temperature for all ethanol to evaporate.
   1. ethanol carryover will interfere with subsequent steps, and over-drying of beads (typically characterized by cracks appearing in the beads)
10. Remove beads from magnet.

Adapter Ligation

1. Make sure heat block is holding at 20C.
2. Protocol assumes ~300bp shearing, if different sized inserts, adjust Adapter amounts.
3. Pull Ampure beads from 4C and PEG/NaCl solution from -20C and set on bench to warm to room temperature.
4. To each tube add the following:
   

Item	Amount
WFI water	16µl
5x KAPA Ligation Buffer	5µl
KAPA t4 DNA Ligase	2.5µl
Adapter ( NOT IN Master Mix)	1.5µl

1. Mix by pipetting to homogeneous mixture.
2. Incubate 20C for 30 minutes.
3. Remove tubes from heat block, turn heat block off, bring back into lab.

Wash

1. The ligation buffer interferes with the normal precipitation of DNA, therefore must be washed before size selection.
2. To each 25µl reaction, add 25µl PEG/NaCl Solution.
3. Mix by pipetting up and down till homogeneous mixture.
4. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
5. Place tubes on magnet until liquid is clear (usually ~5 minutes).
6. Carefully remove and discard supernatant without disturbing the beads.
7. Add 200 µl of 80% EtOH.
8. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
9. Repeat for a 2nd wash.
10. Allow beads to dry at room temperature for all ethanol to evaporate.
    1. ethanol carryover will interfere with subsequent steps, and over-drying of beads (typically characterized by cracks appearing in the beads)
11. Remove beads from magnet.
12. Resuspend in 100µl of WFI water.

Size Selection

1. To each 100µl reaction, add 60µl PEG/NaCl Solution.
2. Mix by pipetting up and down till homogeneous mixture.
3. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
   • This will bind DNA fragments of ~450bp
4. Place tubes on magnet until liquid is clear (usually ~5 minutes).
5. Carefully transfer ~155µl of supernatant to a new tube. It is critical that no beads be transfered with supernatant.
6. To each new tube containing 155µl of DNA smaller than 450bp add 20µl of Ampure beads.
7. Mix by pipetting up and down till homogeneous mixture.
8. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
9. Place tubes on magnet until liquid is clear (usually ~5 minutes).
10. Carefully remove and discard supernatant without disturbing the beads.
11. Add 200 µl of 80% EtOH.
12. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
13. Repeat for a 2nd wash.
14. Allow beads to dry at room temperature for all ethanol to evaporate.
    1. ethanol carryover will interfere with subsequent steps, and over-drying of beads (typically characterized by cracks appearing in the beads)
15. Remove beads from magnet.
16. Resuspend in 50µl of WFI water.

PCR Addition of EBC

• Take note of which external barcode is being added on this step for the purpose of pooling multiple samples.
• Primers are located in C lab -20, NGS 2 rack, EBC + IBC Design Anealed Adapter paper box.
• EBC Options are as follows. the UTBC# is what is entered on the GSAF website for barcode:

• EBC_UTBC74_for_IBC

  EBC_UTBC75_for_IBC

  EBC_UTBC76_for_IBC

  EBC_UTBC77_for_IBC

1. Pull Ampure beads to allow to warm to room temperature.
2. Size selcted DNA added to PCR reaction should not include beads.
   1. Make sure beads are resuspended well before pulling beads to magnet and taking 10µl supernatant for PCR reaction
3. Combine the following in a PCR tube:
   

Item	Amount
Size Selected DNA	10 µl
EBC Barcode mix (10µM)	2.5 µl
2x KAPA HiFi HotStart Ready Mix	12.5 µl

PCR Conditions
Step	Temp	Time
1	98	45 sec
2	98	15 sec
3	60	30 sec
4	72	30 sec
5	go to	Step 2
6	72	1 min

• Store remaining DNA at -20C in labeled tube. This will be used if the PCR is unsuccessful, or if further optimization is required. Beads can be left in this DNA.
• Minimum number of cycles should be preformed to maximize diversity (complexity). This is especially important for mixed population sequencings. Typically 5-7 Cycles is sufficient (for 1µg of amplified library generated).
• Optional: it's ok at this point to use 1µl direct from the PCR products to measure DNA concentrations with Qubit. Adding one more cycle won't generally double the DNA concentration, but two more cycles likely would if there are already signs of at least some amplification (e.g., >15ng/uL).
• Safe Stop: It is ok to pause procedure at this step, store the PCR tubes with product at -20C and continue another day. Furthermore, even more PCR cycles can be added later, since PCR mix components remain active for at least 2 weeks in freezer.

Wash

1. To each 25µl reaction, add 25µl of room temperature Ampure beads.
2. Mix by pipetting up and down till homogeneous mixture.
3. Incubate at room temperature for 10 minutes for DNA to precipitate onto beads.
4. Place tubes on magnet until liquid is clear (usually ~5 minutes).
5. Carefully remove and discard supernatant without disturbing the beads.
6. Add 200 µl of 80% EtOH.
7. Allow to incubate more than 30 seconds at room temperature. It is not necessary to resuspend the beads.
8. Repeat for a 2nd wash.
9. Allow beads to dry at room temperature for all ethanol to evaporate.
   1. ethanol carryover will interfere with subsequent steps, and over-drying of beads (typically characterized by cracks appearing in the beads)
10. Remove beads from magnet.
11. Resuspend in 25µl WFI water or TE.

QC

1. Use 1 µl of supernatant sample (mixed with 3µl of the corresponding TapeStation buffer) for TapeStation; reagents (D1000 ladder and D1000 buffer for DNA) and gel strips located in Deli fridge Lab A, top shelf.
   • TapeStation tips (black box) and 8-tube-strips are kept at room temp in Lab A. A multipippetor to load the special tips on the TapeStation is available at the instrument's site.
   • If you haven't used the TapeStation before, you must first check-in for training with Elizabeth Kahane or Courtney Bridges; the instrument is currently MBB 1.124 (Harris Lab).
   • Schedule use (free of cost) for the Tape Station Agilent 2200 at GSAF sign-up
   • Do not put beads into tape station as they can cause things to run funny.
   • You will use 1 of the gel strip channels (usually the first lane) for ladder (1µl Ladder + 3µl TapeStation buffer); if done correctly, the program will automatically label each band's size.
   • For convenience, you may download a free copy of the TapeStation software (for Windows)
   • If substantial dimer contamination, rewash, possibly adding slightly less PEG/NaCl solution than a 1:1 ratio.
   • If product not visible, toubleshoot PCR step first, possibly by increasing 2-3 cycles as this is the cheapest and most likely failure step.
2. Use 2 µl for Qbit.
3. Pool samples with different IBC and identical EBC into a single submission tube for Core. This should be done based either on the Qbit readings, or on the fraction of correctly sized product from the tape station if different amounts of Adapter contamination detected (requires that correctly sized peak be significantly greater than the off product).
   • Aim for ~25µL total volume in a single tube containing equimolar amounts of each of the samples pooled together (see Sample Input Guidelines and Ready to submit samples? for more information)
4. See Dan for current information about submitting for sequencing.

Alternative non-standard methods

DNA Fragmentation

The current standard method of DNA fragmentation is Fragmentase. Prior to July of 2016 Covaris shearing was the standard method. The shift was precipitated by the cost discrepancy and increasing number of samples being analyzed. Each Covaris tube is $5 and can not be reliably reused for both cross contamination and structural integrity issues.

Covaris shearing

• Successfully done with 500ng of sheared DNA in 25µl.
  During ligation 0.4µl of adapter should be used, NOT 1µl using 500ng in the library prep

1. Place between 1µg and 4 µg of DNA in a covaris microtube (total µg can be increased to 10, but should never be necessary, or decreased below 1µg if necessary).
2. Bring total volume up to 50 or 130µl using Invitrogen Elution Buffer.
   • Covaris only recommends 130 and 50 µl with the microtubes as other volumes can create air bubbles and lead to poor shearing.
3. Signup for an appropriate amount of time in the GSAF core, and take all samples to core.
4. Use appropriate shearing conditions for the Covaris instrument either from Barrick lab folder, or GSAF folder (if GSAF protocol used, save copy to Barrick folder).
   1. For standard sequencing projects, ~300bp is a good target size (if other size used adapter concentrations need to be adjusted).
   2. Consult Covaris manual for other size distributions.
5. Spin tubes down in black plates on plate centrifuge (plates have shallow wells that cause the bottom of the Covaris tube to reach the bottom of the plate so there is no force on the lid)
6. Transfer 1µg of sheared DNA to low retention eppendorf tube.
7. Adjust volume to 25 µl using speedvac if necessary.
8. Volume must be 25 µl, different amounts of DNA can be used with subsequent adjustments to adapter amounts. If attempted, update appropriate sections with working concentrations.
9. Store remaining sheared DNA at -20C at a minimum until sequencing data is back, it can be used to save a Covaris tube if sample fails, or for other experiments.

ERROR rates

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Topic reads

Improved Protocols for Illumina Sequencing

-- Main.DanielDeatherage - 09 Jan 2015
-- Gabriel Suarez - 28 Ago 2018