Sork Lab:Protocols

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DNA Extractions

We have found CTAB with a pre-wash to be the best method for extracting DNA in Oak. The Qiagen Plant-Mini prep is also useful if using the CTAB pre-wash method first before beginning the Qiagen protocol. For PacBio and whole-genome sequencing, a nuclei-isolation protocol was ideal (see below), however it requires 10g of plant tissue per prep.

CTAB DNA Extraction for Oak

CTAB DNA Extraction Protocol with Pre-Wash Sork Lab, Henriquez updated 02/2018

This protocol can take 3.5-4 hours for Pre-Wash and Extraction, plus extra time the next day or following week for quality assessment with Nanodrop & Qubit and recording of data.


  • This original Pre-Wash concept was found by Keith in Li 2007, “An optimized mini-preparation method to obtain high-quality genomic DNA from mature leaves of sunflower”. It has been optimized for our needs with Oak by Keith Gaddis.

Step 1- Mixing the Buffers:

  • All buffer and chemical information can be found at the end of this document

Prep: After the pre-wash, you’ll continue immediately with the regular extraction procedure. You should make both the Pre-Wash Buffer and CTAB Extraction Buffer at the same time. The Pre-Wash Buffer mix above can be made and stored at room temperature for a few weeks. If you know you will use less, scale it down so that you make new buffer as often as possible. Obviously if you have the time, it’s better to make it fresh every single time you do the extraction.

1) Label both the top and side of two 50mL falcon tubes: one for Pre-Wash Buffer and one for CTAB Extraction Buffer. 2) Add the dry chemicals; measure the necessary PVP into each tube, then using a weigh paper measure the necessary CTAB and add to the CTAB extraction tube on top of the PVP. 3) For the Pre-Wash Buffer, add 1mL of the pre-made buffer per individual to be extracted and add one extra mL to account for pipette error. 4) Calculate the amount of reagents necessary to add to the CTAB Extraction Buffer plus one extra to account for pipette error and add to that falcon tube. 5) Vortex both tubes of buffers until the solids dissolve fully into solution. You may need to poke it with a weigh spoon. 6) Take the tubes to the hood and add the necessary 2-mercaptoethanol. Shake/vortex the tubes to fully mix the buffers.

Step 2- Grinding the Tissue and Pre-Wash:

  • Make sure you have liquid nitrogen. The UCLA storeroom is open 8-noon & 1-5pm only

1) Stuff 2mL grinding tubes (with the flatter bottom) with tissue (usually ~100mg) and 2 beads per tube. This is best done ahead of time when the leaves are fresh then stored in the -80 to avoid any freeze-thaw cycles 2) Place the tubes into liquid nitrogen immediately when removed from the -80 freezer 3) Freeze the grinding mill adapters in liquid nitrogen and if possible their lids as well 4) Moving quickly, place tubes in chilled grinding adapters, turn on the tissue homogenizer once your fingers are free of the grinding area, and grind at 25-30Hz for 1 minute. 5) Re-freeze the tubes and adapters and grind a second time, 1 min at 25-30Hz. 6) Check that all the tissue has been ground (it will be light in color), place tubes a tube rack and quickly move to the fume hood where your buffers are waiting 7) Add 1mL of the Pre-Wash Buffer to each time, inverting/mixing tubes as quickly as possible in order to get all the tissue into contact with the buffer while the tissue is still cold. Make sure the lids are closed tightly. 8) Move back to the tissue homogenizer and grind again, 25-30Hz for 30 seconds 9) Check that the tissue has completely dissolved into solution. If there are dry bits stuck to the bottom, move back to the hood and poke it with a clean pipette tip until it is dislodged, then vortex of homogenize again until very well mixed. 10) Spin tubes in centrifuge for 10 min at 10,000 rpm (check ‘Before you begin…’ below to make sure you are set to continue with the extractions during this wait time) 11) Pipet off supernatant and keep the pellet. 12) Proceed with regular extraction protocol immediately (below)

DNA Extractions:

  • This protocol has been modified by Beckley from the Soltis Lab CTAB DNA Extraction Protocol, Reference: Doyle & Doyle, 1987; and Cullings 1992.

Before you begin, make sure the incubator is set to 55°C. Also, place the chemicals we need to cold in the -20°C freezer: 7.5M ammonium acetate, isopropanol, 70% ethanol, and 95% ethanol.

1. Add 500 uL CTAB Extraction Buffer to pre-washed samples. Mix thoroughly- you can either grind them again at 25-30Hz or 30 seconds or vortex until completely mixed. If you need to, poke stubborn pellets with pipette tips to dislodge and then vortex. 2. Incubate samples at 55°C for 45 minutes. Make sure the incubator is slowly moving at 300rpm, to keep the samples from browning on the bottom- but not too fast as to break DNA. *The original protocol said you could leave samples overnight, but this did not work well for oak tissue. 3. Add 3uL RNase per sample (stored in the fridge), carefully invert tubes to mix and then continue to incubate at 55°C for 15 minutes 4. Move to the fume hood. 5. Add 500 uL of 24:1 Chloroform: Isoamyl Alcohol and mix well by shaking/vortexing for a minimum of 30 seconds per sample. Use filter tips. The chloroform should be no more than 3 months old. 6. Centrifuge for 5-10 min at maximum speed. a. Following centrifugation, you should have 3 layers: aqueous phase (top), debris and proteins (middle), chloroform (bottom) b. Go onto next step quickly so the phases do not remix c. While waiting, label your new (final) set of tubes (1.5mL tubes) with sample name on the top and side, and your initials and date on the side as well. d. While waiting, grab the 7.5M ammonium acetate and isopropanol from the -20°C freezer and place on ice in the hood. 7. Pipette off aqueous phase taking care not to suck up any of the middle or chloroform phases. Pipetting slowly helps with this. You can always re-centrifuge. If I am doing 10 I usually do 6 then re-centrifuge (4+ min) for the remainder. 8. Place the aqueous phase into your new (final) labeled eppendorf tube. 9. Estimate the volume of the aqueous phase. ~425-450 uL- see attached table under ‘Buffers, Reagents, & Tables’. 10. Add 0.08 volumes of cold 7.5 M ammonium acetate- see attached table. 11. Add 0.54 volumes (using the combined volume of aqueous phase and added AmAc) of cold isopropanol (=2-propanol)- see attached table. 12. Mix well. 13. Let sit in freezer for 30 min to overnight. a. Longer times will tend to yield more DNA, but also more contaminants. b. 30min-2hours seems to be ideal for oak. c. DNA will start to clump and look like thick mucous. 14. Move back to the hood, and centrifuge for 3 min at maximum speed. a. While waiting, grab the 70% and 95% ethanol from the -20°C freezer and place on ice in the hood. 15. Pipette off the liquid slowly, being careful not to lose the pellet with your DNA. 16. Add 700 uL of cold 70% Ethanol and invert several times to mix. 17. Centrifuge for 1 min at maximum speed. 18. Pipette off the liquid, being careful not to lose the pellet with your DNA. 19. Add 700 uL of cold 95% Ethanol and invert several times to mix. 20. Centrifuge for 1 min at maximum speed. 21. Pipette off the liquid, being careful not to lose the pellet with your DNA. 22. Dry the pellet by leaving it in the fume hood for 10-15 min. a. Longer dry times could damage DNA b. You’re looking for a dry white or opaque pellet, and no ethanol stuck to the side or any droplets. If droplets are stubborn and won’t dry, try to smear them with a pipette tip to speed up dry time. 23. Re-suspend samples with 30-40 uL of TE buffer. Allow to re-suspend overnight in the fridge (Or if you’re in a hurry, for 1 hr at 55°C).

Checking Sample Quality Nanodrop: 1) Place samples in tube rack (they are OK at room temp for awhile as long as we’re not freeze-thawing) and get your notebook ready with a column for ng/uL, 280/260 ratio, 260/230 ratio, and notes 2) Grab your re-suspension buffer used (here we used TE), a 2uL pipette, tips, and water. 3) Double click on the black/blue hourglass icon on the desktop computer and open the Nanodrop software (ND-1000 V3.8.1). Currently (2016) we are using the Shaffer Lab’s Dell with password: herplab. 4) Choose ‘Nucleic Acid’ (if you’re doing RNA extractions, once you open the program and add water you can choose ‘RNA’ (in pink) from the drop down menu on the right of the screen that says ‘Sample Type’ 5) Add 1.5-2uL water when it prompts you. 6) Wipe the plate off with a kim wipe and add 1.5-2uL TE (or whatever your elution/re-suspension buffer was) and click ‘Blank’ 7) Before analyzing each sample, make sure to kim-wipe. 8) Vortex each sample thoroughly before placing on nanodrop plate (right in the middle on top of the little black dot is where you pipette the product!) 9) Re-blank occasionally, every 20 samples is a good amount. 10) Write down all information and when finished, save as follows: click ‘Show Report’. On the drop down menu on the new screen under ‘Reports’ click ‘Save Report’ and make sure to save BOTH ‘Full Report’ & ‘Export Report & Standards Tables’ to your folder on the desktop. 11) If the samples fit the critera for what we need (i.e. rations of 2+ are ideal, but sometimes the ratios can fall to 1.5 and be OK- ask the designer of the experiment for their suggestions) then move on to Qubit.

Qubit: We use Qubit for a better concentration calculation. The nanodrop’s assessment of ng/uL is usually off. Once made, the samples should incubate for about 2-3 minutes before running, though it shouldn’t take more than 10 minutes as the samples will be sensitive to light. Use the dsDNABR program. We use 2ul DNA because there is less pipette error, and because this provides us a ratio of 1/100 DNA/buffer, so the concentration the Qubit measures can easily be multiplied by 100 to give us our exact ng/ul concentration amount. If for some reason you do not have enough Dye Buffer Mix to finish all your samples, always prepare fresh standards for each Dye Buffer Mix you end up using for consistency.

1) Remove standards from the fridge and let come to room temp (10-30min) 2) Label tubes- samples + 2 standards. 3) Make your calculations (see below). Make sure to account for pipette error and make 1-2 more reactions than necessary 4) Mix the Dye Buffer Mix. Quickly place the dye back into the foil sheath to avoid light when finish. 5) Pipette out the buffer mix into sample tubes (198uL) and standards tubes (190uL each) 6) Add 2uL of DNA to each sample tube. 7) Add 10uL of each standard to their correct tube 8) Vortex thoroughly and let stand 2-3 minutes. Pop bubbles if any form. 9) Turn on Qubit 10) Select dsDNA 11) Select dsDNA- Broad Range (or whatever kit you use) 12) Select ‘Read Standards’ *this needs to be done every time you make a new batch. 13) Follow the prompts to each both standards (It is OK if it’s not exactly 0 on standard 1) 14) Select 2uL sample before running your samples 15) Record all info into your lab notebook

The following reaction conditions are generally followed by the Sork Lab: 1) Dye Buffer Mix: 1ul dye + 199ul buffer for standards and sample 2) 10ul Standard1 + 190ul Dye Buffer Mix= total Standard 1, 200ul 10ul Standard2 + 190ul Dye Buffer Mix= total Standard 2, 200ul 3) 2ul DNA + 198ul Dye Buffer Mix= total each sample, 200uL Storing Samples Samples once nanodroped and the good ones run through the Qubit should be stored in the -20°C freezer for long term storage in top and side and bottom labeled white or blue tube boxes. For short term storage or if needing to transport the DNA, leave the extractions in the fridge so that the DNA goes through as few freeze-thaw cycles as possible. Make sure the project name, your name, and the approximate date ranges are all visible on the top and bottom of the boxes (in case the lids get mixed up). The goal here is to avoid freeze-thaw cycles, so if you know where everything is and it is labeled well we won’t have to go back and re-organize it and subject the samples to freeze-thaw which damages the DNA. Make sure you add the box information for each sample to the excel spreadsheet so we know where to find them!

Sample quality information should be recording in: • Your lab notebook that is dated for each day you do the work • On the computer spreadsheet for the particular project • Backed up on SorkLab3 or another server when possible.

Buffers, Reagents, & Tables Reagents and solution for 500mL Pre-Wash Buffer:

  • This is enough for ~500 extractions!

• Washing buffer: o 50 mL 1M Tris o 50 mL 0.5M EDTA o 100 mL 5M NaCl o 300 mL H2O • 2-mercaptoethanol (fumehood); polyvinylpyrrolidone (PVP) (k-30, SABC)

Reagents and solution for 1L CTAB Extraction Buffer:

  • This is enough for ~2,000 extractions!

• Extraction Buffer: o 100 mL of 1 M Tris, pH 8.0 o 40 mL of 0.5M EDTA o 280 mL of 5M NaCl o 20 g of CTAB (Cetyltrimethyl ammonium bromide, Amresco cat# 0833-1Kg) o Add H2O to 1L (~580mL) • Add PVP and 2-mercaptoethanol as necessary for amount of extractions

Buffer Calculations for 1 sample:

  • Calculations for 10+1 and 20+1 samples can be found on the solutions page after the protocol

Pre-Wash Buffer: measure 0.01g PVP, then add 100uL Tris, 100uL EDTA, 200uL NaCl, 600uL H2O, and in the hood add 4.5uL 2-mercaptoethanol

CTAB Extraction Buffer: measure 0.02g PVP and 0.01g CTAB, then add 50uL Tris, 20uL EDTA, 140uL 5M NaCL, 290uL H2O, and in the hood add 2.5uL 2-mercaptoethanol

For 10 Extractions (calculated to 11 for pipette error): Pre-Wash Buffer:

  • This should equal 11,000uL, or 1,000uL (1mL) per extraction

• Measure 0.11g PVP into a falcon tube • Add 1,100uL Tris, • Add 1,100uL EDTA • Add 2,200uL NaCl • Add 6,600uL H2O • In the hood add 49.5 uL 2-mercaptoethanol CTAB Extraction Buffer

  • This should equal 5,500uL, or 500uL (0.5mL) per extraction

• Measure 0.22g PVP into a falcon tube • Measure 0.11g CTAB onto weigh paper and add to falcon tube • Add 550uL Tris • Add 220uL EDTA • Add 1,540uL 5M NaCL • Add 3,190uL H2O • In the hood add 27.5uL 2-mercaptoethanol

For 20 Extractions (calculated to 21 for pipette error): Pre-Wash Buffer:

  • This should equal 21,000uL, or 1,000uL (1mL) per extraction

• Measure 0.21g PVP into a falcon tube • Add 2,100uL Tris, • Add 2,100uL EDTA • Add 4,200uL NaCl • Add 12,600uL H2O • In the hood add 94.5 uL 2-mercaptoethanol CTAB Extraction Buffer

  • This should equal 10,500uL, or 500uL (0.5mL) per extraction

• Measure 0.42g PVP into a falcon tube • Measure 0.21g CTAB onto weigh paper and add to falcon tube • Add 1,050uL Tris • Add 420uL EDTA • Add 2,940uL 5M NaCL • Add 6,090uL H2O • In the hood add 52.5uL 2-mercaptoethanol

TE buffer [Final] for 1L use: 10 mM -10 mL of 1 M Tris, pH 8.0 1 mM- 2 mL of 0.5 M EDTA

1 M Tris, pH 8.0: for 1 L 121.1 g Tris (Fish Cat#: BP152-5) 700 mL ddH2O Dissolve tris and bring to 900 mL pH to 8.0 with concentrated HCl (will need ~50mL) Bring to 1 L

0.5 M EDTA pH 8.0: for 1 L 186.12 g of EDTA (Fish Cat#: BP120-1) 750 mL ddH2O Add about 20 g of NaOH pellets Slowly add more NaOH until pH is 8.0 EDTA will not dissolve until the pH is 8.0

5 M NaCl: for 1 L 292.2 g of NaCl (Fish Cat# BP358-10) 700 mL ddH2O Dissolve and bring to 1 L

Ammonium Acetate and Isopropanol Addition Table

  • For Steps 9-11 in DNA Extraction Protocol

Aqueous Phase Recovered 7.5M AmAc to Add Isopropanol to Add 400uL 32uL 233uL 425uL 34uL 248uL 450uL 36uL 262uL 475uL 38uL 277uL 500uL 40uL 292uL

Additional Resources

Min Deng Lab Genomic DNA Extraction Protocol on Fagaceae Samples [1]

Nuclei-Isolation (for PacBio) for Oak

Nuceli Isolation Protocol- for HMW DNA for PacBio Krista’s Version for Sork Lab- with ALL chemical recipes included Based off Sean Gordon & PacBio protocol. 03/2016

  • This protocol should produce >50ug purified genomic DNA.
    • For PacBio 60SMRT cells for whole-genome sequencing, you would need 2 isolations for 100ug total DNA (=100,000ng)).

To Do/Make Ahead of Time: 1. For each 10g isolation: -Autoclave spoons, 3 600mL beakers, 2 100mL or more graduated cylinder, magnetic stir bar, scissors (to cut pipette tips), a funnel for a small flask, and 2mL tubes. 2. TKE- Tris, KCL, EDTA- pH 9.4-9.5- Stored at 4’C- used in sucrose buffer This is 0.1M Tris Base, 1.0 M KCL, and 0.1M EDTA -For 100mL 1.0M Tris Base: 12.11g Tris base in 70mL ddH20, bring volume to 100mL. To make 0.1M Tris base, use 10mL 1M Tris and add ddH20 to a total of 100mL. -For 100mL 1.0M KCL: 7.455 KCL and add 90mL ddH20. Make up final volume of 100mL. Autoclave 20min on liquid cycle (or just autoclave the combo TKE). Store at room temp. Ideally, divide into small 100uL aliquots in sterile tubes and use each aliquot only once. -0.1M EDTA I bought it already at a 0.1M concentration. -Once all three are prepared, add 10mL each to a sterile container. Adjust the pH to 9.4-9.5 (HCL for more acidic, NaOH for more basic) and autoclave on liquid setting. When you add ddH20 in the SEB, this becomes ~pH 8.0 -Store at 4’C.

3. 1M KOH-used to adjust pH of sucrose buffer before adding β-mercaptoethanol You can buy this pre-made from Sigma, or...

    • This is to be done with full PPE in the fume hood!

-For 10ml 10M KOH stock solution: 5.6g KOH and gradually add 8mL ddH20. Stir constantly, and place beaker in ice bath. Do not add more KOH into the water before confirming the added pellets are completely dissolved and the water temp is cooling down. After all the pellets are dissolved, fill up solution to 10mL. This must be stored in a plastic container as KOH can slowly dissolve glass. -For 1M KOH stock solution: Add 90mL ddH20 to a graduated cylinder. Add 10mL 10M KOH stock. Mix. Store in labeled sterile plastic container (such as two 50mL falcon tubes) in cool, dry, well-ventilated area away from water, strong acids, metals, and flammable liquids.

4. 10% Triton This is a surfactant used to open cell walls -For 10% Triton: 10mL Triton X-100 to a total of 100mL ddH20, store at 4’C and on ice while performing extraction.

5. 10% SDS This digests proteins. -For 10mL 10% SDS Wear protective mask to avoid inhaling SDS powder or do in fume hood. 1g SDS in 250mL flask/beaker. Add 8mL ddH20. You can use a magnetic stir bar and increase temp to 68’C or a water bath. Adjust volume to total of 10mL. Autoclaving is not required. This can be stored for several months at RT, storage at any lower temp will cause precipitate- which can be re-dissolved by warming to 68’C for 10 minutes.

Protocol- Day of: For one nuclei isolation of 10g of plant tissue, you need ~260mL of the sucrose based extraction buffer (SEB). For each 10g isolation, make 300mL of the extraction buffer FRESH the DAY OF the extraction. Directions below chart. Chemicals for SBE buffer= 300mL... 2.0% w/v PVP (MW 40,000)=6g, 10% v/v TKE-made ahead and stored in fridge (2-8’C)=30mL, 500mM sucrose= 51.3g, 4mM spermidine trihydrochloride -stored in fridge (2-8’C)= 0.3g, 1mM spermine tetrahydrochloride= 0.105g, 0.1% w/v ascorbic acid= 0.3g, 0.13% w/v sodium diethyldithiocarbamate- in flame hood= 0.39g, Adjust to pH=9.0-9.1 with 1M KOH- made ahead and stored in plastic at RT. Day of- add β-mercaptoethanol- in fridge (2-8’C) (=BME)=600uL

Directions to make SEB+BME: 1- Combine everything before the KOH in a beaker with a magnetic stir bar in a 600mL beaker. 2- Add ddH20 to ~280mL 3- Add KOH testing with pH strips. For PacBio, DNA should not be subjected to a pH above 9.0, so make sure the pH is between 8.5 and 9.0. I added ~1mL of KOH, but add ~200uL at a time, the pH rises quickly. 4- Adjust to a total of 300mL solution 5- Add the β-mercaptoethanol 6- Make sure everything is kept as cold as possible and place on ice immediately after combining chemicals. It must be ice cold to use, which is >1 hour on ice.


1. Room and machine prep:

  • These steps can be done the day before

-Clean the fume hood. Remove any unnecessary objects. Have a new bag for waste materials ready. Remember to only use filter tips and only autoclaved beakers, spoons, etc. -Make sure to wear a lab coat and never bring anything out of the hood unless completely sealed. -Place the magnetic stir plate in the hood. -Set heating block to 65’C (then 45’C). -Set big centrifuge for 50mL Falcon tubes to 4’C -Set small 2mL centrifuge to 4’C -Place isopropanol in -20’C freezer -Make 6mL 70% ethanol for 6 tubes x2 washes, which is 4,200uL ethanol to 1,800 uL ddH20-keep in -20’C freezer. -Cut the tips off of 1000uL pipette filter tips with autoclaved scissors. -Clean mortars and pestles ethanol and let dry completely (I use at least 3 to grind 10mg tissue)

2. Make 300mL of the SEB(sucrose based extraction buffer) + BME (β-mercaptoethanol) in the fume hood in a 600mL beaker. Make sure the TKE, spermidine trihydrochloride, and β-mercaptoethanol are kept on ice.

    • The SEB+BME is made on ice, and it must be kept on ice at all times. Before plant tissue can be added, make sure the buffer is ice-cold (kept on ice, or 0’C for >1 hour). Keep it in the hood while you move onto step 3.

3. Freeze 10g of fresh plant tissue in liquid nitrogen. In our case, the plant tissue was collected with the mid-rib cut out at the time of sampling, and weighed into 10g bags that were immediately placed on dry ice (flash frozen). They were then put into a -80’C freezer. Prepare an Erlenmeyer flask (~300mL) in a beaker on/surrounded by ice. You’ll place the ground tissue in this flask. Get out the autoclaved spoons.

4. Place the frozen tissue in a mortar containing liquid nitrogen. Grind the tissue to a powder with the mortar and pestle (Grinding a small amount each time will improve grinding and increase yield- try keeping the leaves on dry ice while you grind). When the liquid nitrogen has evaporated in each grinding, grind for approximately 30 more seconds to a flour-like fineness. All more liquid nitrogen and grind again for 30 seconds after liquid nitrogen has evaporated. Repeat for a total of 3 grindings. Transfer the power to the chilled Erlenmeyer flask (on ice).

5. Take the flask on ice to the hood. With an autoclaved graduated cylinder, add 185ml ice-cold SEB + BME to the chilled flask.

6. Place the mixture on ice for 12-20 minutes. During this time, continuously and SLOWLY swirl the contents of the container until powder is dissolved.  

  • Note: Nuclei in sucrose-based buffers must be handled with extreme care. The absence of divalent cations coupled with the extreme osmotic conditions in the SEB+BME make the nuclei extremely fragile. If roughly agitated, the nuclei will break.

7. Filter the homogenate through 2 layers of cheesecloth into a clean 500mL beaker. I used grade 50 cheesecloth from Fisher.

8. Add 15mL SEB+BME (total volume 200mL) to rinse remaining powder from flask and filter homogenate through the same cheesecloth again. Do not force anything through. (PacBio protocol uses nylon mesh filters, first with 200um mesh, then wash, then filtered through 100um then 40um nylon mesh filters.)

9. Add 10ml of 10%Triton (keep on ice if not in fridge) to the beaker(s) slowly along side of beaker over the course of 2 min, while gently stirring with a magnetic stir bar (or gently swirling the flash). Then leave it on ice for 8 min. During this time, swirl the contents of the beaker for 20-30 seconds every two minutes.

10. Transfer the mixture into 4 * 50 mL polypropylene Falcon tubes, and spin the tubes in a centrifuge at 650 x g (1970rpm) for 15 min (4ºC).

11. Very gently decant and discard the supernatants. Add 10 ml of cold SEB+BME to each pellet, and gently resuspend nuclei with pipette tip with pointy tip cut off.

12. Consolidate the nuclear suspensions into 2 * 50ml centrifuge tubes (same tubes OK). Add SEB+BME to final volume of 30mL in each tube, and centrifuge tubes at 650 x g (1970rpm) for 15 min (4ºC).

13. Decant and discard the supernatants, and resuspend the nuclei in 1,440ul TE- this is for each of the tubes (again with pipette tips with the pointy end cut off (with autoclaved scissors of course). At this step, transfer half (~850uL) of each of the 2 falcon tubes into two 4 * 2mL tubes.

14. Into each 2mL tube, add 95ul of 1M NaCl and 24ul of 10 mg/ml RNase A (kept in fridge, keep on ice) into ~850ul nuclei; incubate at 65ºC for 30 min (to digest RNA). *When finished, change the heating block to 45’C.

  • Here, set the big centrifuge to RT and change fitting for 15mL falcon tubes.

15. Into each 2mL tube, add 24ul of 10mg/ml Proteinase K (kept in fridge, keep on ice), cap tube and invert gently 2x.

16. Into each 2mL tube, add 95ul of room-temperature 10% SDS, cap tube and invert gently 2x. Incubate at 45ºC for 60 min (to digest protein).  Cool tubes to RT.

  • Take bottle of phenol:chloroform:isoamyl alcohol out of fridge and warm to room temperature while on the heating block.

17. Combine samples from 2 mL tubes into 2 * 15mL falcon tube. Add 2178ul (or 1 volume) of phenol:chloroform:isoamyl alcohol and invert tube very gently.  Vortex 2s. (Vortexing too long can break the DNA.) Centrifuge for 5 min. at RT at 1500 x g (2,300rpm).

18. Using 1000uL pipette tips with pointy-tip end cut off, suck up only the top layer of the extraction and repeat with 1 volume phenol:chloroform:isoamyl (~2000uL).

  • Repeat this until the interface is clear, placing the top layer into new 15mL falcon tubes each time.

19. Pipette cleared extraction (using cut-tip pipettes) 6 new 2mL tubes, mix the cleared top layer of DNA (~670uL in each tube) with 70uL (or ~1/10 volume) of 3M NaOAc (ph 5.2).

20. In each tube 2mL tube, add 750uL (or ~1 volume) COLD isopropanol. Let the tubes sit in the -20’C freezer for 30-60 minutes (or, it can sit at 4’C overnight).

21. Centrifuge 30 min. at 13,000 rpm at 4ºC. Discard supernatant.

22. Wash with 500ul 70% ethanol; centrifuge >10min at 13,000 rpm and 4ºC. Discard supernatant (That is 6mL 70% ethanol for 6 tubes x2 washes, which is 4,200uL ethanol to 1,800 uL ddH20, kept in the -20’C freezer).

23. Repeat step 22

24. Spin for 2 min at 13,000rpm at 4ºC and decant ethanol with pipette. (Careful, pellet can easily fall out).

25. Air dry the pellet(s) at room temperature for 10min.

26. Resuspend in 30 - 50ul TE per tube. Flick tube or vortex briefly.  Centrifuge for 1 sec to get TE to bottom of tube.

27. Allow to rest in 2-8’C fridge overnight to elute DNA. Many days in the fridge and even over the weekend is better.

28. Test on nanodrop for quality. 260/280 and 260/230 need to be above 2.0. Peak should be clear and smooth.

29. Test concentration using Qubit (see Sork Lab Qubit protocol for specifics).

30. Run on 0.8% agarose gel with 1Kb plus ladder.

31. If sending for PacBio sequencing, cover tubes in parafilm and send overnight on ice (not dry ice, do not want to freeze/thaw just keep cool). Make sure to fill out their form and attach a hard copy to the inside of the package.

AMPure Clean-up for DNA/RNA Extractions

AMPure Clean-Up for DNA/RNA Extractions Updated 07/2016- KLB When do you use this? The AMPure beads utilize size selection to remove unwanted contaminants from DNA extractions. It can also be utilized in PCR cleanup (see AMPure website for that particular protocol) or in GBS clean-up (see ‘Genotyping by Sequencing_Modified for Sork Lab’ Protocol in the Sork Lab protocols binder) among other uses. In this particular modified protocol, contaminants under ~280bp are removed. Specifically, in the Sork Lab, in order to have a useful GBS sequencing run, the best quality extractions must be used. We can see if there are contaminants that would affect sequencing such as polyphenols and proteins by analyzing the 260/230 and 260/280 ratios with the Nanodrop. In order to run through GBS sequencing, samples should have a 260/230 ratio, which shows the presence of polyphenols (the lower the ratio the more polyphenols) above 1.4; we should also have a 260/280 ratio about 1.5, which shows the presence of proteins (the lower the ratio the more proteins). For samples that are OK- have a clear peak at 260- but whose ratios are below the stated amount, the AMPure cleanup can make a significant difference for down-stream applications such as GBS. For example, a sample which had a 260/280 ratio below 1.4 and a 260/230 ratio below 1.35 was found to, when reanalyzed after AMPure bead cleanup, to have both ratios close to 2. The quantity (concentration) of DNA was slightly lower, but with a much cleaner sample there is a much higher probability of a clean GBS run. It is important to note that if the sample flat out did not work in the extraction or the peaks look funny, this protocol is not a magic bullet that will totally fix it. It just makes OK samples spectacular.

Protocol: Note 1: These DNA extractions were run first with the Pre-Wash Protocol for CTAB (PVP+B-Mercaptoethanol+CTAB Buffer) before extraction (See ‘DNA Extractions in the Sork Lab’ or ‘CTAB DNA Extraction Protocol 2016’ protocol in the Sork Lab protocols binder. Note 2: These DNA extractions were prepared with the QIAGEN DNeasy Plant-Mini Kit. This always works for CTAB extractions. Note 3: The FINAL elution was a total of 50ul (two elutions at 25ul with 5 minutes at room temperature before centrifugation).For CTAB it is usually 30-40ul. Note 4: This protocol is a 70% AMPure bead to sample ratio. It was modified from a higher initial amount of product. Any amount should work as long as it stays at 70%- the reason for a lower initial amount of the DNA extraction utilized is because the AMPure beads are very expensive, so I took into account needing this cleaned solution to have enough volume for Qubit analysis, Nanodrop analysis, GBS sequencing, and other possible uses- it can always be repeated on the remaining normal DNA extracted sample As an example: 50ul Product+ 35ul beads= 50ul final recovered product.

Example 1: 25ul Product +17.5ul AMPure Beads= Final 25ul product Example 2: 30ul Product +21ul AMPure Beads= Final 30ul product Example 3: 35ul Product +24.5ul AMPure Beads= Final 35ul product Example 4: 40ul Product +28ul AMPure Beads= Final 40ul product Example 5: 45ul Product +31.5ul AMPure Beads= Final 45ul product Example 6: 50ul Product +35ul AMPure Beads= Final 50ul product

1) Warm AMPure beads to room temperature- 30 minutes minimum (they live in the fridge) 2) Re-suspend beads by vortexing VERY WELL 3) Make sure you know exactly how much sample you have to start with. Usually this means pipetting the sample into a new tube to test the amount. This needs to be exact as this clean-up works by exact ratios. 4) Mix 70% AMPure beads by pipetting up and down 10 times (For example 17.5ul AMPure beads with 25ul sample). 5) Incubate 15 minutes off the magnetic rack at room temperature. 6) Place tubes on rack for 5 minutes. During this time make enough 80% ethanol for steps 7 & 8 (200ul x2 for each samples + extra for pipette error). 7) Aspirate sample VERY SLOWLY. Up to 5ul can remain in tube. If any beads come up while pipetting, pipette them all back out and let the tube sit again 5 minutes before trying again. 8) Add 200ul FRESH 80% ethanol for 30 seconds- no need to pipette up and down or invert tubes, just pipette the ethanol over the beads and that’s it. Remove ethanol. 9) Repeat step 7. Remove any residual ethanol with a fine pipette tip. 10) Dry beads for 10 minutes on bench at room temperature on the rack- drying too long can cause a significant decrease in final DNA concentration. When the ethanol is evaporated, you are good to go. 11) Elute with 25ul (or whatever amount of product you started with) QIAGEN AE elution buffer or RNAse/DNAse free water (or TE if you are cleaning a CTAB extraction) 12) Vortex 15-30 seconds until well combined and beads are in solution. 13) Incubate 2 minutes at room temperature off the magnetic rack. 14) Place tubes on rack for 5 minutes. 15) Transfer solution to new (final) tube, properly labeled that it is the cleaned version. Aspirate VERY SLOWLY. No beads should be picked up during this process. If the elution is not clear, pipette it back out and wait 5 minutes before trying again.

So, how do Agencourt’s Ampure XP or SPRIselect bead precipitate DNA? (From genohub)

The answer has to do with the chemical properties of DNA, polyethylene glycol (PEG), the beads being used and water.  Polystyrene – magnetite beads (Ampure) are coated with a layer of negatively charged carboxyl  groups. DNA’s highly charged phosphate backbone makes it polar, allowing it to readily dissolve in water (also polar). When PEG [ H-(O-CH2-CH2)n-OH ] is added to a DNA solution in saturating condition, it forms large random coils in water. Adding this hydrophilic molecule with the right concentration of salt (Na+) causes DNA to aggregate and precipitate out of solution from lack of solvation (1, 2). Too much salt and you’ll have a lot of salty DNA, too little will result in poor recovery. The Na+ ions shield the negative phosphate backbones causing DNA to stick together and anything else that’s in near vicinity (including carboxylated beads). Once you’re ready to elute your DNA and put it back into solution (after you’ve done your size selection or removal of enzymes, nucleotides, etc.) an aqueous solution is added back (TE or water) fully hydrating the DNA and moving it from an aggregated state back into solution. The negative charge of the carboxyl beads now repel DNA, allowing the user to extract it in the supernatant. Changing the amount of PEG and salt concentration can aid in size selecting DNA (2). This is a common method in NGS library preparation where the user is interested in size selecting a fragment of particular size It’s often used to replace gel steps in NGS library prep.

RNA Sequencing

RNA Extraction for Oak

Oak RNA Prep Protocol Stephanie Steele and Krista Beckley Sork Lab, UCLA

Updated May 17, 2016 Pre-wash adapted from Cronn Lab:

RNA Extraction Buffer (50 mL) (store in 4C fridge)

  • Make the buffer fresh for each new set of extractions if possible.

Ð 18.75 mL 8 M LiCl solution Ð 24 g Urea Ð 8 mL 11% PVP K-60 solution Ð The above reagents should add up to 45 mL. Just before use, add: Ð 0.5 mL 1 M Dithiothreitol (DTT)

Chemical Mix Notes: -To make 8M LiCl: MW is 42.392g/mol. For 40mL (enough for 50 extractions), add 30mL RNase-free water to 13.568g LiCl, dissolve, and bring to 40mL with more water. The solution actually feels very warm to the touch- don’t be alarmed! Autoclave and store in the 2-8C fridge for up to 6 months. -To make 11% PVP K-60: For enough stock for 50 extractions, make 20mL. Add 15.1 mL RNase-free water to 4.9 mL 45% PVP K-60. Approximate with the falcon tube, because the PVP K-60 is too difficult to pipette accurately, even with reverse pipetting. -To make 1M DTT: MW is 154.25 g/mol. For 10mL add 8mL RNase-free water to 1.5g DTT, dissolve, and bring to 10mL with more water. Make 1mL aliquots, cover in aluminum foil, and store in the -20 freezer indefinitely. SAFETY NOTE: Wear safety goggles, close-toed shoes, and lab coat at all times! Use cryogen gloves when handling liquid nitrogen.

Time notes: Day one takes 1-2 hours. Day two takes 3-3.5 hours. -It is better to do fewer samples at a time, and focus on good sterile technique then try to get it all done in one go.


Day 1 First: Turn on cold centrifuge and set to 4C, for a minimum of 20 minutes. 1) Clean mortars, tools, and surfaces with 10% bleach and RNase away. 2) Label 2 mL grinding tubes, add 2 metal beads, and freeze in trays in -80C freezer (At least 15 minutes). It is best to have only a few samples per tray, as the trays will defrost when in use. 3) Add dithiothreitol to working stock of RNA extraction buffer and keep on ice. 4) Fill 2 small dewars with liquid nitrogen (approximately 1 L and 2 L) and two mini coolers with 2 sheets of dry ice each. 5) Place zip-locked samples to be extracted in mini-cooler between dry ice sheets. 6) Remove tray of tubes from -80C freezer. 7) Using a dewar with a spout, pour liquid nitrogen into mortar until it is ¼ full. 8) Remove first sample from mini-cooler, cut approximately 50 mg of leaf tissue, and place in liquid nitrogen in mortar. Replace unused sample in second mini-cooler. 9) Cut sample into strips in liquid nitrogen. Use tweezers to place strips in 2 mL grinding tube, and cap tube once liquid nitrogen gas has visibly evaporated. Throw tube in 1 L dewar with liquid nitrogen. Repeat for all samples using clean mortars. 10) Freeze grinding adapters in liquid nitrogen for approximately 30 seconds. Use tongs to briefly submerge 2 mL tubes in liquid nitrogen before placing in adapters. 11) Grind samples for 1 minute at 30 Hz. 12) Briefly refreeze adapters with tubes in liquid nitrogen. 13) Grind samples for another 1 minute at 30 Hz, or until samples are ground into a fine powder. 14) Quick spin samples and place on ice. 15) Add 1.8 mL cold (4C) RNA extraction buffer to each tube and shake by hand immediately to ensure all tissue is in contact with the buffer. 16) Vortex tissue vigorously and use a pipet tip to dislodge tissue stuck to sides or lid of tubes. 17) Grind samples 10 seconds at 30 Hz. Continue to vortex samples until ground tissue is fully resuspended in buffer. May not fully resuspend- test lower amounts of vortexing to see if it improves RIN score. 18) Spin 10 minutes, 1000 rcf at 4C. 19) Transfer 1.4 mL supernatant to a clean RNase-free tube without disturbing the pellet. 20) Place on ice and keep at 4C overnight. Day 2 First: Turn on cold centrifuge and set to 4C, for a minimum of 20 minutes. 1) Spin tubes 30 minutes, 20,000 rcf at 4C. 2) Discard supernatant carefully. 3) Add 500 uL fresh 70% ethanol. Finger flick to mix and ensure all debris is removed from the sides of the tubes. Spin 5 minutes, 5,000 rcf at RT. Discard supernatant. 4) Repeat step 3. 5) Air dry in fume hood for 10 minutes. While waiting, turn on the heat block to 56C. 6) Proceed to QIAGEN RNeasy Mini Kit Protocol.

QIAGEN PROTOCOL Step numbers correspond to the Quick-Start Protocol and the DNase step in Appendix D of the RNeasy Mini Handbook.

If you’re starting a new box-kit from Qiagen, make sure to add 450mL of 2-Mercaptoethanol to the RLT buffer (this can stay at RT for 1 month, after that move it to the 2-8C fridge. Also, make sure ethanol has been added to the wash buffer, RPE.

2) Add 450 uL Buffer RLT to the pellet. Vortex vigorously. Incubate 2 min at 56C on the Thermo Heat Block, spinning at 300 rpm. Test effect on RIN score. 3) Transfer the lysate to a QIAshredder spin column (lilac) placed in a 2 mL collection tube. Centrifuge 2 min at 20,000 rcf. Transfer the supernatant of the flow-through to a new RNase-free microcentrifuge tube (not supplied) without disturbing the cell-debris pellet.

4) Add 0.5 volume 100% ethanol to the cleared lysate and mix immediately by pipetting. Do not centrifuge. Proceed immediately to step 5.

5) Transfer the sample to an RNeasy Mini spin column (pink) in a 2 mL collection tube. Centrifuge 30 sec at 8,000 rcf. Discard the flow-through.

6) Skip step 6 and proceed to DNase 1 protocol in Appendix D.

D1) Add 350 uL Buffer RW1 to the RNeasy spin column. Centrifuge 30 sec at 8,000 rcf to wash the spin column membrane. Discard the flow-through.

D2) Add 10 uL DNase 1 stock solution to 70 uL Buffer RDD. Mix by gently inverting the tube (DO NOT VORTEX!) and spin briefly.

D3) Add 80 uL DNase 1 incubation mix directly to the RNeasy spin column membrane, and place on the benchtop (20-30C) for 15 min.

D4) Add 350 uL Buffer RW1 to the RNeasy spin column. Centrifuge 30 sec at 8,000 rcf. Discard the flow-through.

7) Add 500 uL Buffer RPE to the RNeasy spin column. Centrifuge 30 sec at 8,000 rpm. Discard the flow-through. Repeat wash.

8) Add 500 uL Buffer RPE to the RNeasy spin column. Centrifuge 2 min at 8,000 rcf. Place the RNeasy spin column in a new 2 mL collection tube. Centrifuge at 20,000 rcf for 1 min to dry the membrane.

9) Place the RNeasy spin column in a new 1.5 mL collection tube (supplied). Add 25 – 30 uL RNase-free water directly to the spin column membrane. Centrifuge 1 min at 8,000g to elute the RNA.

10) Repeat step 9 reusing the collection tube. Discard spin column.

Nanodrop samples and submit to the Bioanalyzer on an RNA Nano chip.

Genotype by Sequencing GBS

Genotyping by Sequencing (GBS) Protocol Sork Lab Modified, 06/2016

OVERVIEW: DAY 1: ~1 hour -Prepare samples to 100ng concentration, 15ul total per PCR tube. Half plate (48 samples) per procedure works to produce best quality. Store in -20 freezer.

DAY 2: ~6.5 hours -Digest samples -Ligate adapters -Pool to create library -Clean-up with Qiagen PCR cleanup kit -Clean-up with Bead Size Selection Protocol.

DAY 3 (Or, can all be done on Day 2): ~2.5 hours -PCR Amplification -Cleanup with Qiagen PCR cleanup kit *this step may not be necessary (test it out). -Clean-up with Bead Size Selection Protocol.

Validation: -Quality assessment with BioRad spectroscopy/Bioanalyzer. -Quantification is measured using Qubit.

Sequencing: -Samples are run on the Illumina sequencer. This occurs when enough pooled samples (7-8) have been collected to justify the cost of a run.

Preparing Adapters: 48 adapters were prepared in house (in the past from a collaborator). We order single strand adapters and anneal them together. Adapters are plated out in a 96 well plate left in the refrigerator (FYI I cut the plate in half, so there are two aliquots of ready to go adapters, as well as much more undiluted adapter in the fridge). They are prepared so that each well’s adapter has a 25% likelihood of beginning with an A, G, T, or C. We have chosen to use the longer adapters that we call the ‘Hardeep ApeK1 adapters’ from the person who developed them. I’ve attached the information for each well/adapter on the last page of this document. The adapters (each well) include: 600ng total adapters (300ng barcoded adapter & 300ng common adapter= 600ng/200ul=3ng/ml concentration), 1X TE (to make 200ul total). This is, according to the Buckler protocol, the concentrated stock. We work directly with this stock, no dilution is necessary.

Sample Preparation: The original protocol involved plating the sample DNA out and drying the samples out. We noticed no change in quality if instead we took the water used to make the equally concentrated DNA samples (all diluted to 100ng DNA concentration), and subtracted that water used from the total water in the plating for the digest. The day before the digest (DAY 1- can be a few days before), ligation, first cleanup, PCR, and second cleanup you can set up your samples. Using the data from Qubit analysis of DNA concentration in extracted individuals, bring the sample to 100ng DNA in a total of 15ul water per sample. Samples should be prepared in 8-strip thin-walled DNase, DNA, RNase, PCR inhibitor free 0.2mL PCR tubes with dome caps. We have found that a run of 48 samples is ideal for Quercus studies and most plant studies. A run of 96 does not produce enough reads per individual for the analysis we need to do at this time in the lab (as of 2016).

Digest: The following day (DAY 2) the first step is digestion by restriction enzymes. The enzyme we use is called ApeK1 (Reorder: NEB R0643L). For ApeK1, the temperature we use during incubation in this protocol provides maximal digestion. This ideal temperature is different for other adapters/enzymes- always check the literature for ideal temperatures with new reagents. When preparing the Digestion Master Mix (MM) (as with all Master Mixes) follow this order: water + buffer + enzyme last. This MM should be prepared over ice. The reagents are stored in the -20 freezer.

For ½ PLATE (48)...NEB Buffer 3=110ul, ApeK1=50ul, H2O (RNase free)=110ul...TOTAL per tube=5ul

In a clean 8-well PCR tube strip, pipette out the total/8 (Half Plate= 270/8= 33ul per well). Then, use the multi-channel pipetter (Ranin 12 channel up to 20ul, use the center 8). Use the VWR purple-box filter tips for best fit here using the 20ul tips. • Pipette 5ul of the Digestion MM into each well • Close strip caps securely & spin down each strip • Incubate in the thermocycler for 2 hours at 75 degrees C. • Finish at 4 degrees C to hold (Or, place plate on ice- digestion must stop before ligation can occur).

Ligation: This is the step where the adapters are added to the samples. Our Ligation MM here differs from the Buckler Protocol because we did not dilute the plated adapters. We make up for that here by just adding less H2O. This makes no difference to the quality of the final product. There is also no clean-up between digestion and ligation- this is OK because although the restriction enzyme is still there, it is not active in ligation because the temperature is not high enough and it is also not as concentrated at the other reagents. You can create this MM before the end of the incubation from the digestion step. First you are going to add the Ligation MM then you will add the correct adapter to each tube using the multi-channel pipeter (I started using just a single channel later because I felt like our multi-channels were not totally accurate across the row). The Ligation MM also must be prepared on ice. (For re-ordering, T4 DNA Ligase: NEB MK0202L). The barcoded adapters are stored in the refrigerator in the hallway. Our protocol here differs here from the Buckler Protocol because we first add the Ligation MM to each cell, then we add each adapter directly into the liquid. That is, 20ul from the digestion + 28.8ul from the Ligation MM + 1.2ul each adapter= 50ul total in each sample tube

The calculations here for the half plate is assuming ~55 samples for overflow. REAGENT ½ PLATE (48)=10x T4 DNA Ligase Reaction Buffer=275ul, T4 DNA Ligase=88ul, H2O=1,221ul...TOTAL per tube=28.8ul

In a clean 8-well PCR tube strip, pipette out the total/8 (Half Plate= 1,584/8= 198ul per well). Then, use the multi-channel pipetter borrowed from the Wayne Lab (It is slate gray GeneMate 8 channel up to 50ul). Use the VWR purple-box filter 50ul tips for best fit. For adding the adapters, use the Ranin 12 channel center 8 tips. Again, use the VWR purple-box filter 20ul tips for the best fit. For the missing wells, note which wells are missing and pipette adapters into those wells one at a time. Pipette adapters directly into the other liquid. The heating step at the end of the ligation kills the enzymes. The actual protocol only requires 10 minutes for heating, but more time (up to 20-30 minutes) will not affect quality. Make sure to check the literature as the time period needed to destroy the enzymes varies for different enzymes. Before you pipette in the adapters, spin down the adapter plates. Use the Smith Lab plate spinner. To use it, you need to only select ‘run’ and allow it to spin for only a few seconds. • Pipette 28.8ul of the Ligation MM into each well • Pipette 1.2ul of each adapter into each well. • Close strip caps securely & spin down each strip • Ligate in the thermocycler for 1 hour at 22 degrees C. • Heat at 65 degrees C for 15-20 minutes • Finish at 4 degrees C to hold (Or, place plate on ice).

Pooling: Because we use a half plate for optimal sequence reads per individual, as we pool we need to add 10ul of each sample to the 1-tube library so that our final concentration is the same as if we did 1 full plate (if doing a full plate, you would use 5ul per sample well) for which the protocol was developed. • Using the Ranin multi-channel pipeter (or a single channel), pipette 10ul from each 8 tube-strip into a clean 8-tube-strip. Then, pipette each of those 8 tubes into a single 1.5mL eppendorf tube. Pipette up and down gently to mix.

Cleanup with Qiagen: Before you start this step, remove the AMPure XP beads from the fridge to allow to come to room temperature. This step removes junk and tiny fragments such as primers and enzymes. Follow the procedures for the kit (All of the pooled library is run through one Qiagen lilac-spin column). We use the following conditions in addition to the directions: spin down for 60sec each step (not 30sec). The amount you pool is ~480ul. For Step 1, x5 total amount of PB means you need to mix 2,400 PB buffer to pooled samples. (Or two tubes with 240ul well-mixed samples and 1,200 PB buffer per tube). Final elution is 102ul EB. This is because we need exactly 100ul for the Bead Size Selection clean-up protocol, and 102ul accounts for pipette error.

Clean-up with Bead Size Selection: We use AMPureXP beads which are stored in the fridge as an aliquot. We use this to select ~200-500bp fragments (Size selection is 0.6:1 then 1:1). Changing the buffer ratio changes the size of DNA fragments recovered. First, only the biggest fragments bind at the 0.6:1 ratio- then, at 1:1 smaller fragments bind. This process is cleaner and more efficient then running a gel.

Before using the AMPureXP beads, they must be well-suspended in solution and at room temperature (vortex well).

  • Use filter tips for every step of this process!

1. You should have exactly 100 μL sample in Qiagen EB buffer from the last step. 2. Add 60 µL of well-resuspended AMPureXP beads (produces 1:0.6 DNA:bead buffer ratio).Mix by pipetting. 3. Mix well and incubate 5 min at room temperature. 4. Incubate 5 min on a magnetic stand (the Wayne Lab owns this rack). When on the stand, you can leave the tubes open a crack so when you move on to step 5, you don’t jostle the beads. 5. While on the magnetic stand, carefully transfer the supernatant to a new tube (do not touch the beads). (If this is continually a problem, you can try to use the magnetic PCR plates in the RNA room- though this becomes a bit like playing ‘Operation’). 6. Add 40 µL of AMPureXP beads to the newly transferred solution (brings DNA:bead buffer ratio to 1:1), mix well and incubate 5 min at room temperature (not on the magnetic stand). 7. Incubate 5 min on the magnetic stand and discard supernatant. 8. While on the magnetic stand, add 800 µL of freshly prepared 80% ethanol. Just to wash over, be gentle. 9. Incubate 30 sec and remove the supernatant. 10. Repeat the steps 8-9 once. 11. Completely remove the ethanol and air dry for ~10 minutes on the magnetic stand. 12. Remove from the magnetic stand and add 52 µL (after GBS ligation) of Qiagen EB buffer (or 32 µL after GBS PCR), mix well and incubate 15 min at room temperature. 13. Place the tube on the magnetic stand and incubate 5 minutes at room temperature. 14. Transfer 50 µL (this step) (or 30 µL (the second time we do this after PCR) the supernatant to a new tube.

                                              • OPTIONAL BREAK HERE

PCR: The 2x Taq Master Mix is from New England Biolabs (NEB #M0270S).

Primers are kept in the GBS glass dish in the fridge- if you run out they are cheap to re-order.

You can do 6 or 8, it doesn’t matter. You should store any left-over pooled samples in the -20’ and label as ‘cleaned- Pre-PCR’ in the off-chance you may want to re-do this step later due to a failure down the line instead of having to start day 1 over.

The original PCR Protocol from the Buckler Lab Protocol is as follows: • 5 minutes at 72deg C. • 30 seconds at 98deg C. • 18 cycles of: • 10 seconds at 98deg C. • 30 seconds at 65deg C. • 30 seconds at 72deg C. • 5 minutes at 72deg C. • Hold at 4degC.

REAGENT uL PER TUBE...FOR 6= 12ul PCR library, NEB 2x Taq MM=150ul, PCR Primer 1=6ul, PCR Primer 2=6ul, RNase free H2O=126ul...TOTAL per tube=50ul

Cleanup with Qiagen: Follow the exact same instructions as the first Qiagen clean-up.

Clean-up with Bead Size Selection: Follow the exact same instructions as the first Bead-Size Selection clean-up, except the final elution here (Step 14) is going to be 30ul.

Storing Samples: The finished cleaned up elution is to be stored in the -20’ freezer along with the first clean-up-before-PCR-leftover-elution.

DNA Quality Assessment and Quantification: The Buckler Protocol Method relies on the proper ratio of adapters to genomic DNA sticky-ends. According to the protocol, factors that can affect this radio include accurate DNA quantification and variation in DNA quality. Poor quality DNA samples will not digest completely. Differentially complete digestion across the samples in the 96plex and/or differing amounts of sample DNA can cause varying ratios of adapters to sticky-ends and therefore variation in the number of reads from any given sample.

Quality Assessment with BioRad: According to the Buckler Protocol, protein contamination is assessed by the 260/280 ratio. A 260/280 ratio greater than 1.7 indicates the sample is free from protein contamination. Polysaccharide contamination is assessed by the 260/230 ratio. A 260/230 radio greater than 1.7 indicates that the sample is free from polysaccharide contamination.

In the results you are looking for 200-500bp with a peak at ~300bp (in particular for the Quercus lobata study).

Quantification with Qubit: The Qubit is more reliable for assessing concentration of the final product than the Nanodrop. Keep in mind that the Qubit reactions should be done quickly as the reagents are sensitive to light. Use the dsDNABR program.

The following reaction conditions are followed by the Sork Lab: 1) Dye Buffer Mix: 1ul dye + 199ul buffer for standards and sample 2) 10ul Standard1 + 190ul Dye Buffer Mix= total Standard 1, 200ul 10ul Standard2 + 190ul Dye Buffer Mix= total Standard 2, 200ul 3) 2ul DNA + 198ul Dye Buffer Mix= total each sample, 200ul

We use 2ul DNA because there is less pipette error, and because this provides us a ratio of 1/100 DNA/buffer, so the concentration the Qubit measures can easily be multiplied by 100 to give us our exact ng/ul concentration amount. If for some reason you do not have enough Dye Buffer Mix to finish all your samples, always prepare fresh standards for each Dye Buffer Mix you end up using for consistency.

Sequencing: In order to set up the sample for sequencing, you need to know: 1) Concentration (from the Qubit) & 2) Size fragment from the bioanalyzer peak and a few calculations which can be done at

We need a final concentration of 10nM (which is # of molecules per unit volume NOT mass) diluted in EB buffer from the Qiagen cleanup kit. Because of the randomness of adapter ligation where some will have the same barcode on both ends, this means that half of the reads don’t really count. We have talked about this with Suhua and we tell them when running through the Illumina Sequencing Facility (on the third floor, done through Suhua in the lab across the hall) to ‘Run the Illumina like it is doing a 10nM read, but our concentration is going to be 20nM’. This is different for other sequencing protocols, so always ask the facility. *NOTE: for one run we did in late 2016 this was NOT the case, the we had half the amount of reads. Before proceeding, talk to whoever will running your library for their recommendations.

From the Qubit we know mass unit/volume. From the bioanalyzer we pick the best peak (zoom in- for this study it is usually between 290 and 300). Using the online calculator, we plug in dsDNA with a size of (for example 0.295 would be 295nm peak). We then calculate the molar concentration by plugging in the concentration we got from the Qubit. This will give you the M concentration. -We need to dilute to 20nM with 20ul (0.1%) tween in a total of 25ul. -Using C1V1=C2V2, solve for V1. For example: a 100nM library would be…100nM(V1)=(20nM)(25ul), where V1=5uL...=5uL of library + (25-5) 20uL of EB+0.1% tween