Protocols

Protocol

1. In a conical flask, fill to 50% of the total volume with DI water.
2. Add LB broth powder to specifications on the side of the bottle (current LB is 20 g/L).
3. Clean spatula with DI water before & after use.
4. Lightly cover with aluminum foil (allows steam to get in but no bacteria).
5. Add autoclave tape.
6. Autoclave on the LIQUID45 cycle.
7. Take out and let cool. Careful not to contaminate with non-sterile equipment or by leaving the top uncovered.
8. Label flask with “LB Broth,” initials, and date. Leave in the fume hood.
9. Dispose when stock becomes cloudy.

Protocol

1. The day before, make LB per LB broth protocol.
1. Note: Do not breathe over an open LB stock — keep closed at all times and only open minimally when extracting a portion from the stock.
2. Add 5 mL of LB to a culture tube, and sterilize:
1. Spray down counter and your gloves with ethanol.
2. Once ethanol is dry, strike bunsen burner.
3. Remove foil from erlenmeyer flask and set on area of counter that was sterilized with ethanol.
4. Run tip of neck of erlenmeyer flask above the flame (not the actual flame, but the hot air above) to sterilize.
5. Use a serological pipette to transfer LB to culture tube.
6. Hold foil cap over flame to sterilize, and recap LB flask.
3. Add 5 µL Ampicillin Sodium Salt solution to LB in the culture tube, only keeping the foil open for a bit. Gently vortex or swirl to mix (do not want liquid to get onto cap).
1. If Ampicillin Sodium Salt solution is not made: Add 100 mg of Ampicillin Sodium Salt to 1 mL DI H₂O. Store in −20 °C, for no longer than 3 months.
4. Poke BL21 glycerol stock with sterile pipette tip, get a visible bit of stock on tip and dip in culture tube.
5. Set cap on tube and add tape to secure lid. Do not screw on lid; the goal is sterile aerobic growth.
6. Put the culture tube in the shaker at ~200 RPM, and turn the temperature to 37 °C.

Protocol

Making the Gel:

1. Add 50 mL of 1x TBE buffer to a 250 mL Erlenmeyer flask:
1. Make 1x TBE buffer by diluting 10x TBE buffer in DI water (100 mL 10x TBE in 900 mL water).
2. For the % agarose gel you want to run, add the appropriate amount of Agarose powder into the flask (1% = 1 g agarose per 100 mL TBE). Microwave flask on high for 30 second intervals until the agarose has dissolved (60-90 seconds total). The powder should no longer be visible in the flask. The flask will be hot, handle carefully!
3. Set up gel mold with appropriate well combs in the meantime.
4. When the solution is cool enough to touch the flask (but should still be hot), add 5 µL of SYBR stain to the solution. Swirl to mix.
5. Immediately pour the solution from the flask into the gel casting tray.
6. Wait 20 minutes for the gel to solidify.

Running the Gel:

1. Once the gel has solidified, carefully remove the comb, loosen the gel tray, and place the gel (in the casting tray) into the gel box.
2. Pour 0.5x TBE buffer into the gel rig until the gel is completely submerged.
3. Prepare all samples in separate PCR tubes:
1. Dilute 1 µL of 1 kb DNA ladder in 1 µL of loading dye and 4 µL Millipore water.
2. Dilute 6 µL of each sample in 1 µL of loading dye.
4. Use a p20 or p10 pipette to quickly load the ladder and each sample into separate wells of the gel.
5. Once all samples have been loaded, attach the lid to the gel box in the orientation which allows the (negatively charged) DNA to run towards the positively charged (red) cathode:



6. Attach the lid to the power supply. Make sure leads are color matched.
1. Always make sure that the current is off or paused before inserting or removing cords from the power supply.
7. Set the voltage to 100V and run the gel for about 30-45 minutes. Check on the gel from time to time to make sure that it is proceeding normally and ensure the dye does not run too far off the gel.
1. Be sure to hit “Run” on the voltage machine to begin the electrophoresis.
8. When the dye is about 75% of the way through the gel, pause/stop the voltage, disconnect the cords from the lid, remove the lid, and take the gel in its tray to be imaged.
9. Visualize the stained gel using a standard transilluminator (302 or 312 nm) and image the gel.

Protocol

(Choose one lysis method)

Heat Lysis Method:

1. Grow a 5 mL E coli culture overnight as above.
2. The next day, split culture into 1mL aliquots in Eppendorf tubes.
3. Spin down E coli at 14000 RPM for 5 min. If the supernatant still looks cloudy afterwards, repeat centrifugation.
1. From this point onwards, keep everything on ice at all times.
4. Remove supernatant using a 1 mL pipette. Be carefµL not to disturb the pellet.
5. Gently add 500µL of Millipore water (ddH2O 50 mL conical tube in cabinet), being sure to keep the cell pellet intact. Do not resuspend. Gently remove the 500µL of Millipore water. (This is intended to wash the cell pellet before PCR)
6. Resuspend the pellet in 250 µL Millipore water until suspension is homogenous.
7. In the specified order, add the following to a PCR tube (25µL total reaction volume):
1. 8 µL Nuclease-free water.
2. 2 µL of cell solution.
3. 1.25µL of 10 uM gapA Forward Primer
1. If 10 uM solution is not made: Dilute 100 µL of 100 uM primer in 900 µL Nuclease-free water, store at 4 °C. Mix well, sometimes they stick to the bottom!
4. 1.25 µL of 10 uM gapA Reverse Primer.
5. 12.5µL Q5 Master Mix.
1. Keep Q5 on ice at all times, thaw on ice. Do not hold in your hands.
8. Place PCR tube in thermocycler, and set the following cycle:
1. 5 min & 30 secs at 98 °C (cell lysis and denaturation)
1. If using the alternative lysis method with lysozyme, change this step to 30 secs.
2. 30 cycles of: (annealing)
1. 98 °C for 10 secs
2. 62 °C for 30 secs
3. 72 °C for 1 min.
3. 72 °C for 2 mins (extension).
4. Hold at 4 °C.
9. Using the product, run an agarose gel to confirm results.
1. Use a 1 kb ladder.
2. We are hoping to see the gapA gene amplified, which should show a band around 1 kb.

Alternative Lysis Method:

1. Repeat above protocol, and stop after step 5 to perform this alternative lysis method.
2. Resuspend pellet in 450 µL cell lysis buffer.
1. Cell lysis buffer = 100 mM NaCl + 20 mM TRIS at pH 7.5.
3. Add 50µL 0.1 mg/mL lysozyme to cell solution.
1. Lysozyme stock solution is 10 mg/mL. Make a 100 µL dilution to 0.1 mg/mL by adding 1 µL lysozyme to 99 µL cell lysis buffer. Throw away remaining 0.1 mg/mL dilution.
4. Incubate lysozyme + cell solution on ice for 30 minutes.
5. cWhile incubating, add water to a beaker and heat on the hot plate to 75 ºC.
1. Use a thermometer from the prep room to measure temperature.
6. When incubation on ice is complete, heat samples in 75 °C water bath for 20 mins.
1. Can do this by attaching sample lids to a piece of tape, then stringing the tape along the top of the beaker.
7. At this point, split the sample into two halves.
1. With one half, use this directly as our DNA template for PCR. Proceed with step 8 from heat lysis protocol.
2. With the other half, spin the sample down at 14000 RPM for 5 min. If the supernatant still looks cloudy afterwards, repeat centrifugation. Take from supernatant for DNA template. Proceed with step 8 from heat lysis protocol.

Protocol

Creating Blanks:

1. Combine 95 g of Paraplast Plus wax chips with a 0.67 g block of colored candle-making dye in a glass flask.
2. Heat the wax over a Bunsen burner or hot plate until all the wax has melted.
3. Pour the wax into a disposable 120 × 120 mm plastic Petri dish.
4. Holding the 75 × 100 mm cellulose filter sheet by the top two corners (along its long edge), lower the sheet into the wax, starting from the bottom and allowing the wax to impregnate the filter paper.
5. Once approximately two-thirds of the cellulose filter is submerged in the wax, allow the wax to wick up the filter paper by capillary action so that it forms a straight line parallel with the top of the paper, leaving ~20 mm of paper uncovered.
6. When removing the filter paper from the wax, wipe it along the side of the Petri dish to remove excess wax.
7. Set the wax-soaked filter paper on aluminum foil to allow the wax to harden for at least 1 min.
8. Draw a pencil line parallel to the wax edge across the unwaxed portion 6 mm away from the wax edge. This line sets the length of the nucleic acid binding area on the dipstick.

Preparation of Dipsticks from Blanks:

1. Fold a 105 × 50 mm sheet of colored photocopy paper in half along its long axis to create a 105 × 25 mm folded sheet.
2. Place the folded sheet of colored paper into the pasta maker, with the folded edge against the 2-mm wide cutters and the open edge up.
3. Hold the colored paper in one hand while turning the handle of the pasta maker with the other hand to engage the paper in the cutting wheels.
4. Place the unwaxed portion of the dipstick blank between the two edges of the folded sheet of colored paper that has been inserted into the pasta maker.
5. Hold the dipstick blank in one hand while slowly turning the handle of the pasta maker with the other hand.
6. When the cut dipsticks have protruded from the underside of the pasta cutter by ~30–40 mm, gently pull down on them with one hand while continuing to turn the handle with the other to help guide them out of the cutter.
7. Once they are fully released from the pasta maker, gently bend the dipsticks to allow the colored paper to peel away from the dipsticks so that it can be easily removed.

Reagent Preparation

Before sample processing, prepare extraction buffer, wash buffer and DNA amplification reaction mix and divide the reagents into aliquots (typically, 1.5 mL tubes are used for extraction and wash buffers, and a 0.2 mL tube is used for the DNA amplification reaction mix). One set is needed for each sample to be processed.

Lysis Buffer

20 mM Tris (pH 8), 25 mM NaCl, 2.5 mM EDTA, 0.05% (wt/vol) SDS

• Prepare 1 liter of extraction buffer by dissolving 2.42 g of Tris base, 1.46 g NaCl in water. Add 5 ml of EDTA (pH 8.0, 500 mM) and 5 ml SDS (10% (wt/vol)). Adjust the pH to 8 with HCl and make the volume up to 1 liter with water.
• The extraction buffer does not need to be sterilized and can be stored at room temperature (18–25 °C) for at least 1 year.

Wash buffer

10 mM TRIS pH 8

• Prepare 1 liter of dipstick wash buffer (10 mM Tris (pH 8)) by dissolving 1.21 g of Tris base in ~900 mL of water. Adjust the pH to 8 with HCl (use 0.42ml HCl) and make the volume up to 1 liter with water.
• The dipstick wash buffer does not need to be sterilized and can be stored at room temperature for at least 1 year.

Elution buffer

LAMP reagent buffer (see LAMP protocol for recipe)

Timing

• Reagent preparation: 10 min
• Sample preparation and cell lysis: 10 seconds to 30 minutes
• Nucleic acid purification using the dipsticks: ~ 30 seconds
• DNA amplification with LAMP

Protocol

1. Resuspend the cell pellet in lysis buffer, pipette vigorously, let sit for two minutes at room temperature.
2. Use 2 µL of the lysate to start the LAMP reaction.
3. Run LAMP reaction for 30 min at 65 °C in the thermocycler. Mix components by pipetting or vortexing in a PCR tube, then briefly pulse-spin in a microcentrifuge.
1. Components: Isothermal Amplification Buffer, dNTPs, MgSO₄. Primers, Bst 2.0, sample, H₂O. Mix as in table above.
2. Run a no-template control to ensure amplification specificity. (All the same reagents without DNA)
4. Heat-inactivate polymerase for 2 minutes at 95 °C.
5. Perform endpoint detection with half of LAMP product to assess for presence of product.
1. Run agarose gel electrophoresis.

Protocol

1. After preparing the actual dipsticks, collect overnight suspension cultures.
2. Centrifuge 2 mL of the culture sample centrifuge for 30 seconds in a 2 mL Eppendorf tube. Dump the supernatant, repeat once. The result is a cell pellet.
3. Resuspend cell pellet in 500 µL of 100 mM NaCl 20 mM TRIS at pH 7.5, which is the cell lysis buffer.
4. Add 0.1 mg/mL (50 µL of the 10 mg/mL stock) lysozyme and incubate for 5 minutes at 98 °C.
5. Incubate on ice for 30 minutes after adding 0.5 mg (10 µL of 50 mg/mL) of cell lysis agent.
6. To test optimal dipstick efficiency, spin down cell debris and then do dipstick protocol on the supernatant (as that holds the genomic DNA).
7. For actual application, capture nucleic acids by dipping the dipstick into the sample until the nucleic acid binding zone is completely soaked ( ~5 seconds).
8. Gently dip the dipstick into 800 µL wash buffer five times ( ~5 seconds total).
9. Dip the dipstick into 20–50 µl DNA amplification reaction mix 15 times ( ~10 seconds total).
1. NOTE: Maximize the elution of nucleic acids from the dipstick by encouraging liquid movement through the nucleic acid binding zone of the dipstick by pushing the dipstick into the bottom of the tube with each dip, causing the cellulose to bend and compress.
10. After elution, wipe the dipstick on the edge of the amplification tube to ensure there are no large droplets of DNA amplification reaction mix remaining on the dipstick.
11. With the mastermix now containing template DNA, run PCR amplification using a preprogrammed thermocycler.
12. Run our sample on 1.0% agarose gel alongside a DNA marker, observing if a band is visible at the length of our target site. (Add 1µL EZ view to sample and marker for observation).

Protocol

Primer Preparation

For 100 µl of 10× LAMP primers, add:

• 16 µl 100 µM FIP
• 16 µl 100 µM BIP
• 2 µl 100 µM F3
• 2 µl 100 µM B3
• 4 µl 100 µM Loop F
• 4 µl 100 µM Loop B
• 56 (add 64 not 56 if no Loop F or Loop B) µL H₂O

Amplification

1. Prepare DNA — Dilute to 1 ng/µL.
1. Run this reaction in a PCR Tube.
2. Run LAMP reaction for 30 min at 65 °C in the PCR machine. Mix components by pipetting or vortexing in a PCR tube, then briefly pulse-spin in a microcentrifuge.
1. Components: Isothermal Amplification Buffer, dNTPs, MgSO₄. Primers, Bst 2.0, sample, H₂O. Mix as in table above.
2. Run a no-template control to ensure amplification specificity. (All the same reagents without DNA).
3. For specificity and amplification efficiency, you can denature a sample for 5 min at 95 °C with just template and primers and then place it on ice to add enzyme.
3. For regular LAMP or a two pot reaction for fluorescence:
1. Incubate for 60 min at 65 °C degrees in PCR machine
2. Heat-inactivate polymerase for 2 minutes at 95 °C.
4. Perform endpoint detection with half of LAMP product to assess for presence of product
1. Run agarose gel electrophoresis

Fluorescence

1. Dilute ProbeF, ProbeQ, Sink1, and Sink2 to 8 uM.
1. Dilute probes and sinks to 100 uM.
2. Then, take 10 µL of each diluted stock solution and add 115 µL of water to get to 8 uM.
2. Add 2.5 µL of each of the 4 probes to each 10 µL reaction mixture (Hyman paper recommends 20 µL reaction total). This should get us to 1 µM per probe/sink.
3. If one pot reaction (if two pot reaction, go look at regular LAMP steps for how to set the thermocycler):
1. Heat to 95 °C degrees in the qPCR machine.
2. Decrease temp by 2 °C/min in qPCR machine all the way to 21 °C and monitor for binding. Verify by confirming fluorescence (caused by binding of fluorescent dye).
4. Run a no template control (step 2) as a blank relative to the fluorescing sample.

Recommendations

1. If optimization is desired, try titrating concentration of Mg (4–10 mM final) or Bst 2.0 (0.04-0.32 U/µL), or changing reaction temperature (50–72 °C).
1. What we currently use is 0.32 U/µL of Bst 2.0 and 8 mM of MgSO₄
2. We could also try varying the temperature of amplification.
2. For improved specificity and amplification efficiency with some templates, denature the template or sample prior to amplification by heating it to 95 °C for 5 min and then placing it on ice.

Protocol

1. Spray gloved hands and 1X hood down with 70% ethanol.
2. Take out T75 culture flask with 293T cells from incubator and aspirate the spent media.
3. Pipette 2 mL of PBS and shake the flask to cover the entire surface area.
1. 1 mL for the T25.
4. Aspirate the PBS from the flask.
1. Use a clean 1000uL (no filter) pipette tip each time.
5. Add 2 mL of 0.25% Trypsin to the flask (turn flask upside down for this step)
1. 1 mL for the T25.
6. Place the flask back in the incubator for about 5 minutes.
1. During this incubation step, label a 15 mL conical cell line name.
7. Check on cells during incubation to determine if cells are detached:
1. Should be rounded in apperance under brightfield microscope.
2. Strike flask against the palm of hand to dislodge cells.
8. Remove the flask from the incubator following appropriate detachment.
9. Into the flask with trypsinized cells, add 3X the amount of media as the amount of trypsin previously added (i.e. 6 mL to the flask to neutralize/stop trypsin activity).
1. Use a 5 mL serological pipette since this fits the best in the T75 flask.
10. Wash flask by pipetting up and down to remove any adherent cells.
11. Transfer the cell suspended in media into a labeled 15 mL conical tube.
12. Place the conical tube in the centrifuge, balance with conical tube, and then spin on program 1 (5 mins, 1400rpm, 25 ºC).
13. While the centrifuge is working, label new flask/petri dish with cell line/date/passage #/ and your initials.
1. Use T25 or petri dish, depending on what the previous passage used.
14. Remove the 15 mL flask from the centrifuge and aspirate the spent media from the flask.
15. Dispense 2 mL of fresh media into the 15mL conical containing the cell pellet and gently pipette up and down to break it up.
16. Add an additional 3 mL-8 mL of media (depending on how large the pellet is).
1. Count cells to know the true number of cells you are seeding into your new flask if needed during this step.
2. 4 mL into T25 and 9 mL into petri dish.
17. Pipette 1 mL of cell suspension and then (in a fan motion) dispense in the flask and petri dish.
18. Place the flasks in the incubator.

Protocol

1. Aspirate, rinse, trypsinize, and count cells.
2. Pellet the cells, and resuspend cells in PBS to rinse
3. Add enough PBS (with cells in it) to 1.5 mL Eppendorf tubes such that you have tubes with 1,000; 2,000; 4,000; 8,000; 16,000 cells each.
4. Spin these tubes in the 1.5 mL-tube centrifuge at 1,500 RPM for 5 min
5. Decant liquid from these tubes, preserving pellets. When you have done as much as you dare to with a pipette, tap the remaining liquid out on a paper towel (which then goes in the biohazard bin).
6. Resuspend the cell pellet in each of the tubes in 50 µL of lysis buffer: it should have Tween20, Triton, and guanidinium HCl written on the side. Let these sit for 10 min on ice.
7. In the meantime, prep 5 tubes of 200 µL wash buffer. This is the wash buffer with Tween in it, not the one with just TRIS.
1. Take 5 dipsticks from the bin, and cut off the wax part with scissors, preserving the tip.
2. Obtain a clean pair of needle nose tweezers and clean them well. They need to have a straight point and fit to the bottom of the Eppendorf tubes for dipsticking.
8. After the 10 min incubation, take the tip of a dipstick that you just cut off, and use the tweezers to swish it gently in the cell lysate for 5 seconds. Then dip 5 times in the wash buffer (use a fresh wash buffer for each one). Then swish 15 times gently in the LAMP mix.
1. Make enough LAMP mix for six 20 µL reactions, aliquot into 6 PCR tubes, and then proceed with dipsticking.
2. You will have a negative control, along with 5 conditions of increasing cell concentration.
9. Carefully add 2 µL of 0.5:1000 SYBR to each tube.
10. Pipette 12 µL or so of each condition into the qPCR plate, and put the lid on. Be carefµL not to contaminate the negative control.
11. Spin the plate for 3 minutes at 2500 RPM in the plate centrifuge, and run on qPCR machine.

Protocol

1. Create two 50 µL reactions — one with each variant of the F2RL3 template — by mixing the reagents below in a PCR tube.
1. The primers used were the NEB2 F3 & NEB3 B3.
2. Dilute reverse primer 1:10 before placing in reaction mix.
2. Place PCR tube in thermocycler, and set the following cycle:
1. 98 ºC for 30 seconds.
2. 60 cycles of
1. 98 ºC for 7 seconds.
2. 66 ºC for 20 seconds.
3. 72 ºC for 20 seconds.
3. 72 ºC for 2 mins (extension).
4. Hold at 4 ºC.
3. Run agarose gel to confirm presence of product.

Protocol

1. Prep 35 µL of LAMP mix.
2. Dilute 30 µL of this to 150 µL in nuclease-free water.
3. Take 50 µL and add 20 units of Ale1 restriction enzyme and 1X rCutSmart buffer.
4. With the next 50 µL, add 20 units of Ale1 and 1X isothermal amplification buffer (came with BST 2.0 LAMP Polymerase)
5. With the last 50 µL, do not add anything else.
6. Incubate each sample overnight at 37 ºC in a water bath.
7. Heat inactivate samples the next day for 20 min at 65 ºC.
8. Run each sample on agarose gel.