Gilbert Lab:Protocols

Plate–DNeasy PowerSoil HTP 96 Kit

Notes before starting

• Perform all centrifugation steps at room temperature (15–25°C).
• All steps, save for vacuum or centrifuge steps, should be done under sterile conditions, ideally in a BSC.
• Use a multichannel pipette for all steps

Steps

1. Prep swabs
   • Get a new bead plate (large sealed plates) and label it. Make a mark on the corner of the mat and the corner of the plate to ensure the two can be matched up later. Remove the rubber well mat from a bead plate. Save the well mat for later
     • Tip: An easy way to keep the mat sterile is to adhere a plate seal to it.
   • Cut off the tips of the swabs (right above the cotton tip) using cutters
     • Tip: Try not to cut right through the tip--the force of the cut will propel the tip into the well, sending liquid flying out. Instead, bear down with the cutters, and then wiggle the rest of the swab back and forth to break off the tip.
     • Note: Sterilize cutters between samples by dipping the cutters into a beaker with ethanol; use a kimwipe to remove excess ethanol to avoid depositing ethanol on tips.
   • Record sample location using a plate map; leave at least 3 wells blank to monitor contamination.
   • Add 750 ul of PowerBead Solution into each bead plate well. The easiest way to do this is by pouring the solution into a reservoir and distributing it using a multichannel.
2. Heat solution C1 at 60 C in a water bath for 5 minutes. Add 60 ul of Solution C1 to each well.
3. After the samples and the two reagents have been added, tightly secure the the well mat back onto the bead plate by using a plate roller. Put the entire plate into the water bath for 20 min at 60 C, making sure that the water level in the bath does not reach too high (up to the seal) on the plate.
4. Remove the plate from the water bath, dry it off, and press the mat into each well to ensure each sample is firmly sealed. Sandwich the plate in between the two aluminum adapter plates for the shaker.
5. Secure the plate into the shaker by inserting it so the short end of the plate is parallel to the bench top and twisting the knob until the armature of the shaker clamps onto the plate (check the sides of the plate to make sure the couplings are seated correctly).
   • Once the plate is held in, but not yet tightened down, twist the metal pin atop the knob so it slots into the base it sits on. This locks the knob from loosening once the machine is on. Continue tightening until you feel firm resistance, but don’t put everything you’ve got into it.
   • If you’re only doing one plate, insert a balance plate into the shaker as well, following the instructions above.
   • Set the shaker for 10 mins at 20 Hz, close the cover, and turn it on.
   • After ten minutes, flip plate so that the backside of the plate is now the frontside. Shake again for 10 mins at 20 Hz. This ensures all samples receive an even treatment of mechanical lysis.
6. Centrifuge at room temperature for 30 mins at 2500 x g.
7. While the plate is in the centrifuge begin preparing your C2 and C3 plates.
   • If you’re processing low biomass samples, add 150 ul of Solution C2 AND 150 ul of C3 to a new collection plate.
   • If you’re processing standard samples, add 250ul of C2 to a new plate, and then 200ul of C3 to another new plate. Seal with aluminum plate seals until used.
     • Note, these don’t have to be done ahead of time, but doing so can speed up the extraction process.
8. When the centrifugation is done, discard the well cover. Transfer all of supernatant to the plate containing either your C2 or you C2 and C3 mix.
   • The best way to do this is by inserting the pipette tips directly into the garnets at the bottom of the bead plate and then drawing up.
9. Apply sealing tape to collection plate. Invert the WELL SEALED plate a few times to mix. Incubate for 10 mins at 4° C (fridge).
10. Centrifuge the plate at room temperature for 12 mins at 2500 x g. Discard sealing tape.
11. Now transfer the entire volume of the supernatant from your C2 plate to your C3 plate.
    • It is hard to see the pellet when doing a plate extraction, so insert the tips of your multichannel just until the line on the tip (where the taper of the tip meets the more cylindrical portion) is even with the top of the well, then draw up.
    • You will skip this step if you are doing the low biomass protocol.
12. Apply sealing tape to collection plate. Invert the WELL SEALED plate a few times to mix. Incubate for 10 mins at 4° C (fridge)
13. Remove the plate from the fridge and centrifuge at room temperature for 12 min at 2500 x g.
14. While the plate is in the centrifuge, add 1300 ul of Solution C4 to each well of a new 2ml collection plate.
15. Once the centrifugation is done, transfer 650 ul of supernatant to the plate containing C4.
    • Following the same supernatant avoidance procedure as mentioned above.
    • Mix samples by pipetting up and down.
    • Note: this good place to stop and store plate (sealed at 4C) if necessary.
16. Place a spin plate (96 filter plate) onto an S-block.
17. Load approximately 900 ul of each sample into the corresponding wells on the spin plate. Seal plate with an AirPore Tape Sheet.
18. Place the setup into the vacuum block (you’ll need to remove the lid, place the plate in, and then replace the lid). Turn the vacuum on for 3 min. After 3 minutes check the filters to see how much of the liquid has travelled through. If not all the liquid has travelled through, continue vacuuming. It is possible that contaminated wells may never vacuum completely if they become clogged.
    • Discard the flow-through and AirPore Tape Sheet. Place the spin plate back on the same S-block.
19. Repeat steps 18 and 19 until all supernatant has been processed. Discard final flow through.
20. Place the spin plate back on the same S-block.
21. Add 500 ul of Solution C5-D to each well of the spin plate. Seal each plate with an AirPore Tape Sheet. To prepare Solution C5-D, add equal parts 100% ethanol to Solution C5-D and mix well.
    • Vacuum for 3 min. Again, if samples have not moved through vacuuming longer is okay.
22. Place the spin plate onto a sterile PCR plate. Seal the filters with an Airpore Tape Sheet. Tape both the PCR plate and filter together (only at the edges; do not cover wells). Centrifuge at room temperature for 1 min at 2500 x g.
    • This step just helps to remove any lingering ethanol. While the vacuum block is great at pulling the supernatant through, it doesn't do an excellent job of pulling ethanol out of the filter plate.
23. Discard the PCR plate. Move the filter and plate back to the BSC and allow the plate to air dry for 20 mins on a new PCR plate at room temperature.
24. Now add 100 ul of Solution C6 to the center of each well. Seal the filters with an Airpore Tape Sheet. Tape both the PCR plate and filter together (only at the edges; do not cover wells).
25. Centrifuge at room temperature for 5 mins at 2500 x g.
26. Remove and discard the filter. Seal the PCR plate with an aluminum plate seal and store your eluted DNA, which now resides in the PCR plate, in a -20 or -80 freezer.

Single Tube-DNeasy PowerSoil Kit

Notes before starting

• Perform all centrifugation steps at room temperature (15–25°C).
• If Solution C1 has precipitated, heat at 60°C until precipitate dissolves.
• 2 ml collection tubes are provided.

Steps

1. Add 0.25 g of soil sample to the PowerBead Tube provided. Gently vortex to mix.
2. Add 60 μl of Solution C1 and invert several times or vortex briefly.
   • Note: Solution C1 may be added to the PowerBead tube before adding soil sample
3. Heat the tubes containing the samples and reagents in a water bath at 60 C for 10 minutes.
4. Place in the shaker for 10 minutes at 20Hz.
   • Secure PowerBead Tubes horizontally using a Vortex Adapter tube holder (cat. No. 13000–V1–24).
     • More detail on this procedure can be gained by looking at the plate extraction protocol.
   • If you are only processing a few samples, place the tubes into the tube holder so that they will sit in the spots furthest away from the pivot point of the vortexer’s arms.
   • If you are processing more than a few samples (so that more than one row in the tube holder needs to be used), you will need to flip the block around its Y-axis and vortex it for another 10 minutes (after the initial 10 minutes) so that all tubes are lysed equally.
5. While shaker is running prepare your tubes containing C2, C3, and C4.
   • While this is not necessary, it will speed up the process considerably.
   • Add 250 μl of Solution C2 to new 2ml tubes, so that each sample has its own tube
   • Add 200 μl of Solution C3 to new 2ml tubes, so that each sample has its own tube
   • Shake first, and then add 1200 μl of Solution C4 to new 2ml tubes, so that each sample has its own tube
6. After the shaker has finished, centrifuge tubes at 10,000 x g for 30 s.
7. Transfer 400-500ul of supernatant to your tubes containing solution C2.
   • Vortex briefly then incubate at 2–8°C for 5 min.
8. Centrifuge the tubes for 1 min at 10,000 x g.
9. Transfer 600ul of supernatant to your tubes containing solution C3.
   • Vortex briefly then incubate at 2–8°C for 5 min.
10. Centrifuge the tubes for 1 min at 10,000 x g.
11. Avoiding the pellet, transfer up to 750 μl of supernatant to your tubes containing C4.
    • Note: when transferring the supernatant to the tubes containing C4, instead of depositing the supernatant into the tubes and then vortexing, a faster alternative is to transfer the supernatant, pipette up and down 5-10 times, and then transfer that volume to the spin filter
12. Load 675 μl onto an MB Spin Column and centrifuge at 10,000 x g for 1 min. Discard flow through.
13. Repeat step 14 twice, until all of the sample has been processed.
14. Add 500 μl of Solution C5 to the filter. Centrifuge for 30 s at 10,000 x g.
15. Discard the flow through. Centrifuge again for 1 min at 10,000 x g.
16. Carefully place the MB Spin Column into a clean 2 ml collection tube. Avoid splashing any Solution C5 onto the column.
17. Add 100 μl of Solution C6 to the center of the white filter membrane. Alternatively, you can use sterile DNA-Free PCR Grade Water for this step (cat. no. 17000–10).
18. Centrifuge at room temperature for 30 s at 10,000 x g. Discard the MB Spin Column. We recommend storing DNA frozen (–20° to –80°C).

PCR (no blockers)

Reagent	Per Well (ul)	Per Reservoir (ul)
Water	9.5	950
Unbarcoded primer	1	100
Master Mix	12.5	1250
Sample	1	-
Barcoded primer	1	-

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Notes before starting

• Use a plate spinner (or centrifuge) to spin down your sample plates and barcoded primer plates after they defrost
• Use a centrifuge to spin down your mastermix and unbarcoded primer tubes after they defrost
• Let the BSC run for 30 minutes with UV on to sterilize it
• Wash the BSC (and ideally your gloves before you start) with ethanol (ideally use a spray bottle or dampen some kimwipes)
• Get all your materials into hood before starting

Materials

• Mastermix (or MM, this contains your Taq polymerase, dNTPs, MgCl2 and reaction buffers)
• Unbarcoded primers
  • Depending on your application, these could either be reverse (16S) or forward (ITS, 18S) primers
• HyClone water (or another PCR-grade equivalent)
• One of each pipette and tips (you'll need 2 boxes of 10ul tips for each sample plate)
• Sterile PCR plates
• Reservoirs (white plastic trough) and plate seals (either aluminum or plastic)

Steps

1. In a reservoir , add (in order):
   • Water (950ul), unbarcoded primer (100ul), and one tube of MM (1250 ul)
   • Pipet up and down a few times to mix the reagents thoroughly
   • If you're doing more than one plate at a time, you can add double or triple these amounts to the same reservoir.
     • This is most easily done by adding 950ulx3 of water, then 100ulx3 of unbarcoded primer, then 1250ulx3 of MM.
2. Now grab sterile PCR plate by the edges and label the front of it (ITS/16S-#_____Plate name___Date).
3. Using a multichannel, pipette 23ul of the water-primer-MM mix out of the reservoir and into the wells of the new plate you just labelled.
   • Minimize touching and passovers, but you can use the same tips multiple times to load all rows of the plate.
4. Now get your plate with the barcoded primers, remove the foil by firmly holding the base of the plate and peeling the foil from one edge, and add 1ul of barcoded primer into each well using multichannel
   • The plate seals are sticky, so when you're peeling the seal off you really want to make sure you hold the plate down well, or the plate may hop (as the seal peels of in fits and spurts) and mix all your primers (or samples) together. This is bad news.
   • Row A from the primer plate goes to row A of the new plate, row B goes to row B, and so on.
     • Change tips each row.
   • Cover the primer plate with a plate seal once done (peel the paper backing of the seal and apply it as you would a bandaid)
5. Now remove the seal from your sample plate and add 1ul of DNA sample into each plate using a multichannel
   • Again A to A, B to B, and so forth.
   • Cover your sample plate with a plate seal once done.
   • Again, change tips each row
6. Cover your newly filled PCR plate(s) with a seal
   • After everything is covered, use a plate roller (like a rolling pin) on every plate you've just covered to ensure each well is sealed.
7. Spin down the plates that are destined for the thermocyler
8. Bring the plates over to the thermocycler
   • Turn it on using the switch in the back (Cycler 1 should always be used first), hit enter twice to bypass the login screens.
   • Gilbert Lab>Standard Programs>16S and ITS> Cycler 1
   • Exit and repeat for multiple thermocyclers, selecting different cyclers each time.

PCR (with blockers)

Reagent	Per Well (ul)	Per Reservoir (ul)
Water	7.5	750
Primer	1	100
Master Mix	12.5	1250
Sample	1	-
Barcode	1	-
mPNA	1	100
pPNA	1	100

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Notes before starting

• Use a plate spinner (or centrifuge) to spin down your sample plates and barcoded primer plates
• Use a centrifuge to spin down your mastermix and unbarcoded primer tubes after they defrost
• Let the BSC run for 30 minutes with UV on to sterilize it
• Wash the BSC and gloves with ethanol (ideally use a spray bottle or dampen some kimwipes)
• Get all materials into hood before starting

Materials

• Mastermix (Taq polymerase, dNTPs, MgCl2 and reaction buffers)
• Forward (16S) or reverse (ITS) primers
• mPNA and/or pPNA
• HyClone water
• One of each pipette and tips
• Reservoir and plate seals

Steps

1. Heat the mPNA and/or pPNA in a waterbath for 5min at 55C to ensure a complete resuspension of the primers prior to use
   • Give it a quick vortex after the heat, then spin it down
2. In reservoir (white plastic trough), mix (in order):
   • Water (850ul for one blockers, 750ul for two), primer (100ul), blocker (100ul for each blocker used), and one tube of MM (1250 ul)
   • Pipet up and down a few times to mix
   • If you're doing more than one plate at a time, you can add double or triple these amounts to the same reservoir.
3. Using a mulitchannel, pipette 23ul of the water-primer-MM mix into each well using
   • Minimize touching and passovers
4. Add 1ul of barcoded primer into each well using multichannel
   • Change tips each row
   • Cover the primer plate with a plate seal once done
5. Add 1ul of DNA sample into each plate using a multichannel
   • Cover your sample plate with a plate seal once done
   • Again, change tips each row
6. Cover your PCR plate(s) with a seal and then spin it down plate
7. Bring the plates over to the thermocycler
   • Turn it on using the switch in the back (Cycler 1 should always be used first), hit enter twice to bypass the login screens.
   • Gilbert Lab>Standard Programs>16S and ITS> Cycler 1
   • Exit and repeat for multiple thermocyclers, selecting different cyclers each time.

Picogreen Assay

Reagent	Per Well (ul)	Per Reservoir
Dye	-	50ul
Buffer	-	20ml
Sample	2	-
Dye+Buffer	198	-

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Materials

• TAE Buffer
• Reservoir
• Picogreen dye
• Resevoirs
• One black plate for each PCR plate
• Plate seals
• One box of 10ul tips for each PCR plate
• 50ml pipette

Steps

1. Get the picogreen dye from 4C fridge, let it defrost in an area away from bright light
   • Your palm is a good place
2. Get your PCR sample plates and spin them down
3. Dilute the dye with Tris EDTA in a reservoir (cover when not using)
   • We’re aiming for a 1:400 ratio
     • 50ul dye : 20 ml Tris EDTA for 96 well
     • 100ul dye : 40 ml Tris EDTA for 2 plates
   • Add the dye to the reservoir first, followed by the TE buffer, as that will allow for better mixing of the dye and buffer
     • As you add the TE move the pipette left and right to aid in mixing
4. Using a multichannel, add 198 ul of dye mix into each well of a black plate
   • It is not necessary to change tips in between each row
   • If you’re doing multiple plates at one time, it is easiest to allot the dye mix all at once
     • For the plates you temporarily set aside, lay a plate seal over them to protect them from light
5. Remove the foil from the PCR plates and, using a multichannel, transfer 2ul of PCR from the sample plate into the wells of the blank plate (do change tips each row here)
   • Once you complete a plate, replace the plate seal on the sample plate and use the leftover backing from the seal to cover the completed black plate
   • Keep track of which black plate refers to which sample plate
6. Run through the plate reader
   • Protocols>Nucleic acids>Picogreen Fluorescence
   • Open the plate reader, insert the plate with well A1 in the top left corner, close the plate reader and hit read
   • Once the plate reader returns the values, copy and paste the output to a labelled word document

Pool clean-up

Reagent	Per Well (ul)
Beads	100
Pooled Samples	100
70% Ethanol	200(x2)
HyClone	100

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Materials

• AMPure beads
• 70% molecular grade ethanol
• Sterile micro-centrifuge tubes
• HyClone water
• Magnetic rack
• Reservoir and plate seals

Steps

1. Clean your pipettes with ethanol dampened kimwipes
2. Mix together a 1:1 ratio of beads (4C, shake THOROUGHLY first) and pooled DNA in a new tube
   • 100ul:100ul is the standard ratio
   • Vortex to mix, then spin down very briefly
     • You want to consolidate the liquid, but if you spin it too hard you'll form a bead pellet (which is no good). An easy way to walk this fine line is to use a mini-centrifuge and switch it on and off briefly, assessing as you go.
3. Incubate the mix for 5 minutes away from the magnetic rack
4. Then place the sample tubes onto the magnetic rack and let them sit there for 2 minutes
   • A brown pellet should form on the wall of the tube
   • DNA binds to iron oxide when salt concentration is high, like after PCR with MgCl2
5. Using a pipette, remove and discard the liquid surrounding the pellet
   • Keep the tubes in the magnetic rack while doing this
6. Add 200ul of 70% ethanol to the tube (70% ethanol and 30% HyClone water)
7. Wait for 1 minutes and then remove and discard the ethanol wash
8. Repeat the ethanol wash step, then leave the tubes open in the BSC to let ethanol evaporate
   • Try to prevent pellet from dying and cracking, but the goal here is to evaporate as much ethanol as possible
   • The pellet should look slightly damp, but matte, when it is ready
   • If persistent ethanol beads cling to the sides of the tube, use a pipette tip to smear them around in order to encourage evaporation
9. Once the ethanol has been dealt with, add 100ul of HyClone water and resuspend pellet using pipette (sucking up and down) or shaker, wait a few minutes
   • This should be done off of the magnetic rack
10. Put the tubes back on the magnetic rack and let the pellet reform (now DNA has been eluted and is in the water), pipet out the liquid into new tubes
    • Avoid sucking up magnetic beads like the plague; any brown or red in your final elution means the run will likely fail

Qubit quantification

Reagent	Per Qubit Tube (ul)	Per 15ml Tube
Dye	-	10ul
Buffer	-	2ml
Sample	2	-
Standards	10	-
Dye+Buffer	198/190	-

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Steps

1. The Qubit dye and standards (1 and 2) are stored at room temperature
2. In a 15 ml tube, mix together dye and buffer in a 1:200 ratio
   • 10ul dye and 2 ml of buffer for 10 samples, but this can be scaled up or down
3. In individually labelled qubit tubes, add:
   • 2ul sample : 198 ul dye mix
   • 10ul standard : 190 dye mix
     • You should prepare one tube for standard 1 and one tube for standard 2
4. Vortex and spin down each of the tubes
   • The dye is light sensitive, so protect your samples from bright light
5. Run through the Qubit
   • dsDNA> high sensitivity> read standards>run samples
     • Generally the qubit should be set to a 2ul sample volume, and it double-checks this with you before each run
   • Standard 1 should be around 30, and standard 2 should be anywhere from 12,000-19,000
     • If either of these numbers are far off, prepare a new batch of dye-buffer reagent
6. The Qubit will read concentration down to about 0.1 ng/ul. While it will temporarily store data, you should write all value down

Gel Electrophoresis

So you want to run a gel, huh? You really think you can handle it? Yeah, actually it’s not that hard; you can definitely handle it.

Reagent	Per Gel	Per Lane
Agarose	0.75g	-
TAE	50ml	-
SYBR Safe	5ul	-
Loading Dye	-	1ul
Ladder	-	5ul

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Steps

1. Mix up agarose and TAE buffer in an Erlenmeyer flask (these following ratios are for a small gel; triple them if you want to run a large gel)
   • 1% (0.50g agarose:50ml TAE)
   • 1.5% (0.75g agarose:50ml TAE)
   • 2% (1.00g agarose:50ml TAE)
2. Microwave the mix in the microwave until agarose is fully dissolved
   • Start out with 15 or 30 second bursts and watch to make sure it doesn’t bubble over
3. Place inner well perpendicular to walls of the outer well so the rubber seals create a closed compartment
4. After donning protective gloves, remove the hot agarose-TAE mix from the microwave
5. Add 5ul of sybr safe for every 50 ml of volume directly to flask with the agarose-TAE and swirl it about
6. Pour the dissolved agarose into the well, checking to avoid leaks and overflows as you do so
   • After you’ve poured your gel is a great time to wash the flask, otherwise the remaining gel with set in it and be hard to clean. Rinse it out in the sink, making sure you’re properly diluting the agarose with lots of water to avoid clogs.
7. Now grab your favorite gel comb and place it into one of the slots (usually the one at the end unless you’re running two rows on the same gel)
   • Watch out for bubbles forming on the teeth of the comb
   • You don’t want to let things get heated in the lab, so let that gel cool off for 10 minutes or so
8. Once the gel is cool (it will now be opaque), pick up that inner container and flip it so the inner and outer reservoirs are parallel (and the comb is closer to the negative terminal)
9. Add enough TAE to the outer reservoir so that the gel is covered by about half a centimeter of liquid
10. Now you can release the beast and take that comb out of the gel by pulling upwards on it
    • It’s best to remove the comb after the TAE has been added to prevent the wells from suctioning shut when you remove the comb
11. Get some Parafilm, a true lab staple, and cut yourself a piece
12. Using a pipette, make 1ul dots of the loading dye at even intervals (so that there is 1 dot for each sample you want to run)
    • To each of those dots add 4ul of your sample; mix by pipetting up and down
13. Now comes the time to prove your worth: loading the wells
    • Suck up all 5ul of your dye-sample solution and load them into the wells by approaching the wells at a shallow angle with the pipette held parallel to the wells
    • Dispense the liquid into the wells but don’t blow out the tip
14. Finally, add 5ul of the ladder to an open well
15. Place the top on the contraption, making sure red goes to red and black goes to black
16. Plug in the leads to the power supply
17. Last chance to check that you have the positive (red) lead furthest away from your samples, so that the DNA (- charge) runs through the gel and not off the short end
18. Set the power supply at 80-100 V and run that gel like you’d run after the ice cream truck during a mid-August heatwave
19. Let the gel run for ~45 minutes, or until the yellow band of the ladder is about halfway or more down the gel
20. When you’re satisfied with your masterpiece, turn off the power supply and unplug the leads for extra safety points
21. Slide the gel out of the inner reservoir onto a clean surface or container
22. Remove the protective cover from the Invitrogen gel reader, place the gel directly onto the glass, and then put the orange cover back on (that is, you know, if you want to ever see again)
23. Turn on the machine and check out those bands