TITLE: FLASH v2.0
AUTHORS: Amy Lyden, Gloria Castañeda, Emily Crawford, Saharai Caldera, Jenai Quan, Katherine Travisano

[DESCRIPTION]
<div class = "text-blocks"><div class = "text-block"><a href="#" style = "text-decoration:underline;color:blue;cursor:pointer;"><span style = ":;"> </span></a><span> FLASH is a crispr-cas9 technology that enriches for targeted sequences in sequencing libraries. The initial DNA sample undergoes a blocking step that removes of the 5’ phosphoryl groups of the DNA fragments, resulting in a product that is not amendable for downstream adaptor ligation or amplification via standard Illumina-based library preparation. The subsequent incorporation of targeted CRISPR-cas9 library exposes the desired regions of interest, allowing them to be processed into a library.  For more information on methods and results, please see the </span><a href="https://doi.org/10.1093/nar/gkz418" style = "text-decoration:underline;color:blue;cursor:pointer;"><span style = ":;">FLASH paper</span></a><span>. For FLASH guide RNA design help, please see our </span><a href="https://github.com/czbiohub/flash" style = "text-decoration:underline;color:blue;cursor:pointer;"><span style = ":;">github</span></a><span>.</span></div><div style = "text-align :; float : ;"><img style = "" src = "https://s3.amazonaws.com/pr-journal/bi423q5w.png" /></div></div>

[STEPS]
?. [Dephosphorylation]
Prepare a reaction for each cDNA or gDNA sample and mix well. Add the components in the order specified below. You can make a master mix (MM) of the rapid alkaline phosphatase (RAP) buffer and enzyme. If you are using the RAP MM, mix  of MM with each sample. Mix thoroughly with a pipette or by tapping to avoid shearing.  AB1  Component  1X2  DNA, 10pg - 100ng    x μL  3  rAPid Alkaline Phosphatase Buffer    2 μL  4  rAPid Alkaline Phosphatase    1 μL  5  H2O    up to 20 μL
3 µl
AB1  Component  1X2  DNA, 10pg - 100ng    x μL  3  rAPid Alkaline Phosphatase Buffer    2 μL  4  rAPid Alkaline Phosphatase    1 μL  5  H2O    up to 20 μL
?. [Cas9 Treatment]
Prepare dual-guide RNAs by annealing crRNA and tracrRNA at an equimolar amount at  for  then allowing the mixture to cool to room temperature on the bench. (Note. If crRNA and tracrRNA have been previously annealed and stored, it is recommended to re-anneal the dual-guide RNAs under the above conditions)
95 °C
?. [Cas9 Treatment]
If your starting stock of Cas9 is more than 4μM, you must dilute your stock of Cas9 to 4μM by using 1X Cas9 activity buffer.
?. [Cas9 Treatment]
Make a Cas9 master mix as described below. Add the components in the order specified to prevent precipitation.   ABC1  Component    1X  ____ X2  10x Cas9 Activity buffer    3 μL    ____ μL  3  Cas9 4μM*    2.5 μL    ____ μL  4  dgRNAs 4μM**    3 μL    ____ μL  5  H2O    0.5 μL    ____ μL  6  Total  9
  μL  ____
  μL  *Remember to dilute your Cas9 stock to 4uM if not already at 4uM. Use 1x Cas9 activity buffer to dilute your Cas9 enzyme if you do need to dilute your stock.**Remember to dilute your dgRNAs to 4uM using water if needed. Note: Most experiments for the 2018 FLASH paper were performed at this Cas9 concentration. However, we have demonstrated that lower concentrations work equally well on bacterial isolate DNA. Consult the manuscript for more details.
ABC1  Component    1X  ____ X2  10x Cas9 Activity buffer    3 μL    ____ μL  3  Cas9 4μM*    2.5 μL    ____ μL  4  dgRNAs 4μM**    3 μL    ____ μL  5  H2O    0.5 μL    ____ μL  6  Total  9
  μL  ____
  μL
?. [Cas9 Treatment]
Add  of the master mix to each of your  blocked DNA samples.
9 µl
21 µl
?. [Cas9 Treatment]
Mix well by pipetting or tapping the PCR tubes and incubate the reaction at  for .
37 °C
?. [Cas9 Treatment]
Deactivate the Cas9 by adding  of Proteinase K to each of your sample tubes, mixing again by pipetting or tapping. Incubate at  for .
1 µl
37 °C
?. [SPRI Clean-up at 1.7X]
Equilibrate clean SPRI beads to room temperature and vortex well to mix.
?. [SPRI Clean-up at 1.7X]
Add beads equivalent to 1.7X the sample volume to each sample tube (for  of sample, add  beads).
31 µl
53 µl
?. [SPRI Clean-up at 1.7X]
Mix well by pipetting or tapping the tubes. Pulse-spin in a picofuge for no more than 2 seconds.
?. [SPRI Clean-up at 1.7X]
Incubate for  at room temperature, then put the tubes on the magnetic rack. Allow beads to separate on the magnet for 3-5 minutes, or until the supernatant is clear.
?. [SPRI Clean-up at 1.7X]
Keeping the tubes on the magnet, carefully remove and discard the supernatant.
?. [dA tailing]
Prepare the following mixture for each sample. You can prepare the dA-tailing buffer and Klenow fragment ahead of time as a MM. If you prepared dA-tailing MM, add 9.6uL of MM to each sample.  AB1  Component    1X  2  FLASHed sample    50.4 μL  3  dA-Tailing buffer    6 μL  4  Klenow fragment    3.6 μL  5  Total    60
  μL
AB1  Component    1X  2  FLASHed sample    50.4 μL  3  dA-Tailing buffer    6 μL  4  Klenow fragment    3.6 μL  5  Total    60
  μL
?. [dA tailing]
Mix well by pipetting up and down several times with a P200 set to 40uL.
?. [dA tailing]
Incubate all tubes at  for  with heated lid OFF
37 °C
?. [dA tailing]
Cool all tubes to  and proceed with the next part as soon as possible.
4 °C
?. [Adaptor Ligation]
! The reagents used in this step are very viscous and must be mixed well before using. ! Do NOT make a master mix for this step, although the Ligation Master Mix and Ligation Enhancer may be mixed up to 4 hours before and kept at 4°C.
?. [Adaptor Ligation]
Prepare the following mixture for each sample. Alternatively, if you prepared an adaptor MM with Ligation MM and Ligation Enhancer, add 31uL of MM to each sample and then add 2.5uL of adaptor to each sample.  AB1  Component    1X  2  dA-tailed sample from part IV    60 μL  3  NEB Ultra II Ligation Master Mix    30 μL  4  NEBNext Ligation Enhancer    1 μL  5  NEBNext Adaptor 1:100 or 1:300
  dilution*    2.5 μL<i style="mso-bidi-font-style:
  normal">  6  Total    93.5
  μL  Note: Adaptor dilution of 1:100 works for an initial DNA input up to 100 ng. A 1:300 dilution is recommended for an initial DNA input of under 10 ng. The subsequent cleanup step should remove all extra adaptor and adaptor dimers.
AB1  Component    1X  2  dA-tailed sample from part IV    60 μL  3  NEB Ultra II Ligation Master Mix    30 μL  4  NEBNext Ligation Enhancer    1 μL  5  NEBNext Adaptor 1:100 or 1:300
  dilution*    2.5 μL<i style="mso-bidi-font-style:
  normal">  6  Total    93.5
  μL
?. [Adaptor Ligation]
Prepare the above mixture and mix well by pipetting up and down several times with a P200 set to .
50 µl
?. [Adaptor Ligation]
Incubate at  for  in a thermocycler with the heated lid OFF.
20 °C
?. [SPRI Clean-up at 1.7X]
Add  ethanol (prepared fresh). Incubate beads for  and then remove the ethanol
200 µl
?. [SPRI Clean-up at 1.7X]
Repeat the above ethanol wash step.
?. [SPRI Clean-up at 1.7X]
Allow the beads to air dry for . Do not overdry. Dry beads should appear matte (rather than glossy), but should not have a cracked appearance. Overdried beads, as indicated by a cracked appearance, may not resuspend or elute well.
?. [SPRI Clean-up at 1.7X]
Remove tubes from magnet and resuspend in  nuclease-free H2O by pipetting up and down.
53 µl
?. [SPRI Clean-up at 1.7X]
Continue to resuspend by tapping the tubes and then spin down briefly in a picofuge.
?. [SPRI Clean-up at 1.7X]
Allow  for DNA to elute from beads, then transfer tubes back to magnet
?. [SPRI Clean-up at 1.7X]
Allow the beads to separate for at least 2 minutes.
?. [SPRI Clean-up at 1.7X]
Collect  of supernatant to clean PCR tubes.
50.4 µl
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Thaw a TruSeq i5/i7 barcode plate or other TruSeq primers, and choose barcodes for each sample. Take note of plate color/barcodes to be used. DO NOT use the same barcode for more than one sample in a sequencing run. Dual unique TruSeq barcodes are preferable.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Equilibrate clean SPRI beads to room temperature and vortex well to mix.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Add beads equivalent to 1X the sample volume to each sample tube (for  of sample, add  beads).
93.5 µl
93.5 µl
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Mix well by pipetting or tapping the tubes. Pulse-spin in a picofuge for no more than 2 seconds.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Incubate for  at room temperature, then put the tubes on the magnetic rack. Allow beads to separate on the magnet for 3-5 minutes, or until the supernatant is clear.
Longer incubation time during SPRI addtion is because it may be helpful in removing unwanted fragments/adapter dimers
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Keeping the tubes on the magnet, carefully remove and discard the supernatant.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Add  ethanol (prepared fresh). Incubate beads for  and then remove the ethanol
200 µl
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Repeat the above ethanol wash step.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Allow the beads to air dry for . Do not overdry. Dry beads should appear matte (rather than glossy), but should not have a cracked appearance. Overdried beads may not resuspend or elute well. Ensure the beads are fully dry before eluting.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Remove tubes from magnet and resuspend in  nuclease-free H2O.
17 µl
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Resuspend well by tapping the tubes and spin down briefly in a picofuge.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Allow  for DNA to elute from beads, then transfer tubes back to magnet
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Allow the beads to separate for at least 2 minutes.
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Collect  of elution.
15 µl
?. [SPRI Clean Up 1X + Addition of TruSeq Indexing Primers]
Mix  of the elution with the appropriate TruSeq indexing primer barcodes as planned.
10 µl
STOPPING POINT: If necessary, samples may be stored at -20C.
?. [USER Enzyme + Indexing PCR]
Spin down plate or tube strips briefly to collect liquid to bottom of the well.
?. [USER Enzyme + Indexing PCR]
Prepare the following mixture for each sample. You can make a master mix of USER enzyme and Q5. If doing this, add 28µL of MM to your 25µL sample.  AB1  Component    1X  2Ligated sample + TruSeq indexing primers from part VI25 μL3  NEBNext Ultra II 2X Q5 PCR Master Mix    25 μL  4  NEB USER Enzyme    3 uL  5  Total    53
  μL
AB1  Component    1X  2Ligated sample + TruSeq indexing primers from part VI25 μL3  NEBNext Ultra II 2X Q5 PCR Master Mix    25 μL  4  NEB USER Enzyme    3 uL  5  Total    53
  μL
?. [USER Enzyme + Indexing PCR]
Set up the following cycling conditions for USER enzyme cutting and indexing PCR in a post-PCR room:  ABC1  Temperature    Time    Cycles  2  37°C    15mins    1  3  98°C    30 sec    1  4  98°C    10 sec    12 cycles  5  65°C    75 sec  6  65°C    5 min    1  7  4°C    -    -  Set the lid heat to ON at 105°C.
ABC1  Temperature    Time    Cycles  2  37°C    15mins    1  3  98°C    30 sec    1  4  98°C    10 sec    12 cycles  5  65°C    75 sec  6  65°C    5 min    1  7  4°C    -    -
?. [SPRI clean-up 0.9X]
Equilibrate clean SPRI beads to room temperature and vortex well to mix.
?. [SPRI clean-up 0.9X]
Add beads equivalent to 1:0.9x the sample volume to each sample tube (for  of sample, add  beads).
53 µl
47.7 µl
?. [SPRI clean-up 0.9X]
Mix well by pipetting or tapping the tubes. Pulse-spin in a picofuge for no more than 2 seconds.
?. [SPRI clean-up 0.9X]
Incubate for  at room temperature, then put the tubes on the magnetic rack. Allow beads to separate on the magnet for 3-5 minutes, or until the supernatant is clear.
?. [SPRI clean-up 0.9X]
Keeping the tubes on the magnet, carefully remove and discard the supernatant.
?. [SPRI clean-up 0.9X]
Add  ethanol (prepared fresh). Incubate beads for and then remove the ethanol
200 µl
?. [SPRI clean-up 0.9X]
Repeat the above ethanol wash step.
?. [SPRI clean-up 0.9X]
Allow the beads to air dry for . Do not overdry. Dry beads should appear matte (rather than glossy), but should not have a cracked appearance. Overdried beads may not resuspend or elute well.
?. [SPRI clean-up 0.9X]
Remove tubes from magnet and resuspend in  nuclease-free H2O.
53 µl
?. [SPRI clean-up 0.9X]
Resuspend well by tapping the tubes and spin down briefly in a picofuge.
?. [SPRI clean-up 0.9X]
Allow  for DNA to elute from beads, then transfer tubes back to magnet
?. [SPRI clean-up 0.9X]
Allow the beads to separate for at least 2 minutes.
?. [SPRI clean-up 0.9X]
Collect  of supernatant to clean PCR tubes.
23 µl
?. [KAPA Fluorescence-guided Amplification]
Using optical PCR strip tubes which are separated from each other so that they can be removed from the thermocycler one at a time, add KAPA amplification MM and Illumina P7 and P5 (5sol-20 and 5sol-21) primers at 5µM to your samples as below. You can make a master mix of KAPA master mix and primers.  AB1  Component    1X  2  Amplified and indexed DNA    23 μL  3  Kapa amplification master mix    25 μL  4Illumina P5 and P7 5uM primers  2 μL  5  Total    50
  μL
AB1  Component    1X  2  Amplified and indexed DNA    23 μL  3  Kapa amplification master mix    25 μL  4Illumina P5 and P7 5uM primers  2 μL  5  Total    50
  μL
?. [KAPA Fluorescence-guided Amplification]
Cap all tubes with optical caps. Do not write on the caps.
?. [KAPA Fluorescence-guided Amplification]
Set up the following PCR conditions:  ABC1  Temperature    Time    Cycles  2  98°C    45 sec    1  3  98°C    15 sec  204  60°C    30 sec  5  72°C    1 min 30 sec  6  Plate read  7  72°C    30 sec  Run the program, and watch until your sample either:1. Crosses the standard (STD) 2 threshold 2. Starts to plataeuThen pull your sample out DURING THE 72C 30sec INCUBATION after the plate read. It is critical to pull it out during this step, and not when it is denaturing or annealing.If your sample still has not reached the STD 2 by 20 cycles, let the program finish.Alternatively, if you have many samples, you can simply apply 10 cycles to all, and evaluate if you need further amplification later.You may want to take note of the unique number of cycles each sample needed to reach the STD 2 threshold.
ABC1  Temperature    Time    Cycles  2  98°C    45 sec    1  3  98°C    15 sec  204  60°C    30 sec  5  72°C    1 min 30 sec  6  Plate read  7  72°C    30 sec
?. [KAPA Fluorescence-guided Amplification]
Place your labeled samples in the RT-PCR thermocycler.
?. [KAPA Fluorescence-guided Amplification]
Add  of STD 2 to a clean optical PCR tube.
50 µl
?. [SPRI Clean-up 1X]
Equilibrate clean SPRI beads to room temperature and vortex well to mix.
?. [SPRI Clean-up 1X]
Add beads equivalent to 1X the sample volume to each sample tube (for  of sample, add  beads).
50 µl
50 µl
?. [SPRI Clean-up 1X]
Mix well by pipetting or tapping the tubes. Pulse-spin in a picofuge for no more than 2 seconds.
?. [SPRI Clean-up 1X]
Incubate for  at room temperature, then put the tubes on the magnetic rack. Allow beads to separate on the magnet for 3-5 minutes, or until the supernatant is clear.
?. [SPRI Clean-up 1X]
Keeping the tubes on the magnet, carefully remove and discard the supernatant.
?. [SPRI Clean-up 1X]
Add  ethanol (prepared fresh). Incubate beads for  and then remove the ethanol
200 µl
?. [SPRI Clean-up 1X]
Repeat the above ethanol wash step.
?. [SPRI Clean-up 1X]
Allow the beads to air dry for . Do not overdry. Dry beads should appear matte (rather than glossy) but should not have a cracked appearance. Overdried beads may not resuspend or elute well.
?. [SPRI Clean-up 1X]
Remove tubes from magnet and resuspend in  nuclease-free H2O.
27 µl
?. [SPRI Clean-up 1X]
Resuspend well by tapping the tubes and spin down briefly in a picofuge.
?. [SPRI Clean-up 1X]
Allowfor DNA to elute from beads, then transfer tubes back to magnet
?. [SPRI Clean-up 1X]
Allow the beads to separate for at least 2 minutes.
?. [SPRI Clean-up 1X]
Collect  of supernatant to clean PCR tubes.
25 µl
?. [Library Analysis]
Quantify by HS DNA Qubit
?. [Library Analysis]
Run a fragment analysis, such as with the HS DNA Bioanalyzer. You are expecting a fragment trace that is characterized by sharp peaks in the 250-650 range. A successful FLASHed sample trace looks approximately like this: (lower marker in green, upper marker in purple)
?. [Library Analysis]
If there is a large spike at ~138 bp, this is indicative of adaptor dimer in the sample. We recommend: 1) Additional SPRI clean-ups with a sample:bead volume ratio of 1X (as many as necessary for removal of dimers), or 2) size selection to 250-650bp with the BluePippin on a 2% gel.
?. [Library Analysis]
If pooling multiple samples, use the concentration of DNA between 250-650bp for normalization.
?. [Library Analysis]
When satisfied with the quality of your pooled/individual library, proceed with quantitative PCR and sequencing.
?. [Dephosphorylation]
Normalize your cDNA or gDNA to anywhere between 10pg-100ng. For most samples, we recommend 5-10ng input.
?. [Dephosphorylation]
Incubate at  for  with heated lid OFF.
37 °C
?. [Dephosphorylation]
Add  sodium orthovanadate (competitive inhibitor of phosphatases) to the quench the reaction and mix well with a pipette or by tapping.
1 µl