Notes

• The 2023 MIT iGEM team has successfully used this protocol to assemble Yeast Toolkit and Mammalian Toolkit plasmids, both pL1s (single transcriptional units) and pL2s (multiple transcriptional units).
• This protocol follows the NEB provided protocol exactly, although we always use the NEB thermocycler program designed for 5 to 10 inserts.
• Pipetting volumes less than 1 uL is unreliable. Create 75 ng/uL stock solutions of your part plasmid minipreps if their concentrations are above 100 ng/uL.

Materials

• NEBridge Golden Gate Assembly Kit
• BsaI-HFv2 (NEB #E1601) or BsmBI-v2 (NEB #E1602) depending on assembly
• 75 ng of each part plasmid (the 75 ng is considering the size of the whole plasmid, not just the fragment that is to be cloned together)
• 200 uL PCR tube strip

Protocol

1. Set up the assembly reaction in a 200 uL PCR tube in the following order:
• x uL nuclease-free water, with x = 17-N uL where N is the number of inserts. This volume is calculated by taking the 20 uL total reaction volume and subtracting 2 uL for ligase buffer, 1 uL for NEB Golden Gate Assembly Mix, and N uL for total part plasmid volume, assuming 1 uL per part, i.e. miniprep concentration of 75 ng/uL
• 1 uL each of 75 ng/uL part plasmid stock solutions
• 2 uL 10X T4 DNA ligase buffer
• 1 uL NEB Golden Gate Assembly Mix
2. Run the following thermocycler program:
1. Repeat 30 cycles: 37ºC for 1 minute (digestion) then 16ºC for 1 minute (ligation)
2. 60ºC for 5 minutes (final digestion and heat inactivation of enzymes)
3. 4ºC infinite hold
3. Transform immediately into competent E. coli or store at 4ºC overnight and use the next day.

Materials

• 1 LB+antibiotic plate per transformation
• 1 LB+Amp plate for pUC19 control
• 50 µL competent cells per transformation
• ~50 ng plasmid to transform (stored at -20ºC)

Protocol

1. Fill heat block wells with dH2O and turn on the heat block to 42ºC.
2. Thaw a 50 µL tube of NEB 5-alpha or 10-beta competent cells for each reaction on ice for 10 minutes.
3. Use 10-beta cells for larger plasmids (>10kb), or plasmids with more repeats. Otherwise, 5-alpha cells should suffice.
4. Warm a LB+antibiotic agar plate for each transformation to room temperature. Prepare an LB+Amp agar plate for pUC19 transformation control.
5. Add the DNA you wish to transform into the bacteria. Be sure to gently mix your plasmid stock by pipetting up and down before adding it (otherwise the plasmid can settle to the bottom of your microfuge tube):
6. If transforming a plasmid: Add 1 µL of each plasmid to each cell aliquot. Gently mix by flicking the tube 4-5 times.
7. If transforming a MoClo reaction: Add 2 µL of MoClo reaction mix to each cell aliquot.
8. Incubate on ice for 30 minutes.
9. Heat shock at 42ºC for 30 seconds (exactly).
10. Quickly place the cells back on ice for 5 minutes.
11. Add 950 µL of room temperature growth media without antibiotics. Incubate at 37ºC for 60 minutes, shaking vigorously (250 rpm).
12. Use Stable Outgrowth Media if using NEB 10-beta or NEB Stable Cells.
13. Use Super Optimal broth with Catabolite repression (SOC) if using NEB 5-alpha cells.
14. Mix cells thoroughly by flicking the tube and inverting, then spread 50 µL using glass beads onto LB+antibiotic agar plate.
15. Spread a larger amount if transformation efficiency is expected to be low (e.g. many inserts, large plasmid).
16. Alternatively, perform a four-quadrant streak dilution on the LB+antibiotic agar plate.
17. Incubate the plate overnight at 37ºC. Keep the competent cell tube in 4ºC overnight.
18. Let the plate grow for no more than 16 hours. Parafilm and store in 4ºC.
19. Bleach remaining competent cell contents with 20% bleach in a 1:1 volume ratio and dispose.

Materials

• Glycerol stock of bacteria with desired plasmid OR plate containing colonies of bacteria transformed with desired plasmid
• Disposable culture tubes

Protocol

1. Per plate or glycerol stock, label a culture tube with identifying information for the plasmid.
2. Add 5 mL of LB media into each 14 mL tube.
3. Add 5 uL of appropriate 1000x antibiotic solution.
4. Make sure you pipet the antibiotic solution into the liquid and NOT the side of the tube. Also make sure you do not lower the tip too far below the surface of liquid to avoid contaminating the pipet.
5. If using plates, they may have a mix of colored and white colonies. Pick two colonies per plate to inoculate, determining the appropriate color to pick based on the construct (e.g. a GFP dropout plasmid will stay green if incorrectly constructed and will become white when correctly constructed).
6. For each tube: Using a P200, obtain a new tip, open the petri dish or glycerol stock, and dip the pipette tip into the colony.
7. For plates, it is important to avoid colony cross contamination — more important than getting a lot of bacteria on your pipette tip.
8. Take the tip and lower it into the liquid culture. Discard the tip into the biowaste bin.
9. Gather tubes and grow on a shaker for no more than 16 hours at 37ºC at 280 rpm.

Materials

• QIAprep Spin Miniprep kit
• 1.5 mL microcentrifuge tubes
• Tabletop centrifuge
• Cryovials
• 40-50% glycerol

Protocol

1. If making a glycerol stock is desired (always the case unless the inoculation is already from a glycerol stock), add 900 uL of 40-50% glycerol and 900 uL culture to a cryovial. Label tubes with part ID, colony #, date, and miniprep number (if prepping more than one miniprep of the same colony). Transfer to -80ºC freezer.
2. Transfer 1 mL of bacterial overnight culture into a microcentrifuge tube. Centrifuge the culture at 6800g for 3 minutes. Discard the supernatant.
3. Obtain 31n µL of nfH2O in a microcentrifuge tube, where n is the number of minipreps you are performing.
4. Heat the nfH2O to 50ºC on a heat block as you complete the following steps.
5. Shake RNAse A + Buffer P1 vigorously to ensure all LyseBlue particles (if present) are dissolved. Resuspend pelleted bacterial cells in 250 µL of RNAse A + Buffer P1 and transfer the solution to a microcentrifuge tube. (After making glycerol stocks, it’s okay to pellet all remaining culture.)
6. No cell clumps should be visible after resuspension of the pellet.
7. Add 250 µL of Buffer P2 and mix thoroughly by gently inverting the tube 4–6 times. Continue inverting the tube until the solution becomes viscous and slightly translucent. Do not allow the lysis reaction to proceed for more than 5 min. Do not vortex the solution and do not pipette up and down aggressively, because this will result in shearing of genomic DNA and contamination of plasmid.
8. If LyseBlue has been added to Buffer P1, the cell suspension will turn blue after addition of Buffer P2.
9. Inverting the tube should result in a homogeneously colored mixture (blue if LyseBlue has been added).
10. Add 350 µL of Buffer N3. Mix immediately and thoroughly by inverting the tube 4–6 times. The solution should become cloudy, and small particles of “precipitate” should become visible.
11. Large culture volumes (>5 ml) may require inverting up to 10 times.
12. If LyseBlue reagent has been used, the suspension should be mixed until all traces of blue are gone and the suspension is colorless.
13. A homogeneous colorless suspension (translucent liquid with white, floating precipitate) indicates that the SDS has been effectively precipitated.
14. Centrifuge at 17,900g for 10 minutes in a table-top microcentrifuge. A white pellet will form spread along one face of the microfuge tube (the face that was most distal from the center of the centrifuge).
15. Transfer 800 µL of the supernatant from step 4 to the QIAprep 2.0 Spin Column. Avoid touching the pellet at all costs, even if it means not getting all 800 µL.
16. Transferring 750 µL works reliably in avoiding the pellet.
17. Centrifuge for 60 seconds at 17,900g. Discard the flow through.
18. If purifying a low-copy number plasmid OR using an endA+ strain of E. coli, wash the QIAprep 2.0 Spin Column by adding 500 µL Buffer PB and centrifuging for 60 seconds at 17,900g. Discard the flow through.
19. NEB 10-beta and NEB 5-alpha are both endA1 (endA deficient) and thus this step may be skipped.
20. Add 750 µL of Buffer PE to the QIAprep 2.0 Spin Column.
21. Centrifuge at 17,900g for 60 seconds. Discard the flow-through.
22. Centrifuge at 17,900g for an additional 60 seconds to remove residual wash buffer. Discard the flow-through.
23. Place the QIAprep 2.0 Spin Column in a clean 1.5 mL microcentrifuge tube. To elute the DNA off the column, add 50 µL of 50ºC-heated nfH2O to the center of each QIAprep 2.0 Spin Column.
24. Let stand for 1 min. This step is critical for releasing the plasmid DNA from the column.
25. Centrifuge at 17,900g for 60 seconds. Do not discard the flow-through. The elution in your clean 1.5 mL microcentrifuge tube is your miniprepped sample(s).
26. Quantify the plasmid concentration for each miniprepped sample via Nanodrop.
27. Keep the DNA elution product at -20ºC for long-term storage.

Materials

• Complete media (DMEM + 10% FBS + 1x NEAA)
• PBS
• 0.05%–0.25% trypsin
• 10 cm dish (protocol can be modified to accommodate other sizes of dishes and flasks)

Protocol

1. Warm aliquots of complete media, PBS, and trypsin in a 37°C bead bath for 30 minutes.
2. Aspirate media from the 10 cm dish, being careful not to touch the monolayer of adherent cells.
3. Rinse the dish with 5 mL warmed PBS, then aspirate.
4. Add 1 mL trypsin to the dish and gently swirl.
5. Place the dish in the incubator for 3-5 minutes then examine under a microscope to see cells detached.
6. Add 9 mL of warmed complete media to the 10 cm dish. Pipette up and down several times in a "waterfall pattern" to ensure the cells are in single cell suspension.
7. Create dilutions (e.g. 1:5 and 1:20) and label tops of dishes with name, cell type, passage number, and date.
8. To create dilutions: for a 1:5 dilution in a 10 cm dish, place 8 ml prewarmed media and add 2 ml cells. Similarly, for a 1:20 dilution in a 10 cm dish, place 9.5 mL prewarmed media and add 0.5 mL cells.
9. Place the dish in the incubator.

Notes

• 1:5 dilution will typically allow you to passage every 2-3 days while 1:20 will allow you to passage every 5-7 days.
• Always make two dishes at two different dilutions! Ensures you have cells ready almost any day and if you make a mistake, don’t lose the cells.

Materials

• Lipofectamine 2000
• Opti-MEM
• Complete media (DMEM + 10% FBS + 1x NEAA)
• PBS
• 0.05%–0.25% trypsin
• 24-well plate (this protocol can be modified for other sizes)
• 1.5 mL microcentrifuge tube

Protocol

Day 0 (seeding well plate)

1. Trypsinize the 10 cm plate and count cells. Passage as needed.
2. Seed cells at appropriate concentration based on ThermoFisher's Useful Numbers for Cell Culture, e.g. 0.05x10^6 cells in 0.5 mL media without antibiotics for a 24-well plate.
3. After counting, pellet cells then resuspend in the appropriate amount of media and divide into wells. For example, if you are seeding 24 wells each with 0.5 mL media containing 10^5 cells, you need to pellet 26 * 10^5 cells, and then add enough media to bring the concentration to 2 * 10^5 cells/mL
4. Grow cells for 24 hours. Cells should be at 70-90% confluence on the day of transfection.
5. Prepare the plasmid DNA desired for transfection to concentrations of 50 ng/uL.

Day 1 (transfection)

1. Warm aliquot of Opti-MEM in a 37°C bead bath for 30 minutes.
2. Create reaction mix A by combining, in a 1.5 mL microcentrifuge tube, N * (50 uL Opti-MEM + 1.8 uL Lipofectamine 2000)
3. For each of the N wells being transfected, create reaction mix B in N 1.5 mL microcentrifuge tubes: 40 uL Opti-MEM + 12 uL plasmid DNA (diluted to 50 ng/uL)
4. Incubate reaction mix A and B at room temperature for 5 minutes.
5. Combine reaction mix A (51.8 uL) and B (52 uL), resulting in N microcentrifuge tubes with ~103.8 uL total volume.
6. Mix gently and incubate at room temperature for 20 minutes.
7. Add each microcentrifuge tube mix dropwise to appropriate corresponding well.
8. Place the well plate in the incubator.

Materials

• 5 mL polystyrene tube with cell strainer
• Complete media (DMEM + 10% FBS + 1x NEAA)
• PBS
• 0.05%–0.25% trypsin
• 15 mL conical tube

Protocol

1. Warm aliquots of complete media, PBS, and trypsin in a 37°C bead bath for 30 minutes.
2. Aspirate off media and wash cells with 500 uL PBS.
3. Add 500 uL trypsin to each well and incubate for 5 mins at 37°C until the cells come off.
4. Add 500 uL warmed complete media to each well
5. Transfer everything to a conical tube and centrifuge for 5 min at 250 g
6. Aspirate media and resuspend in 500 uL warmed complete media.
7. Pipet cells up and down to break up aggregates and create a single cell suspension.
8. Transfer cells to a 5 mL polystyrene tube; cells should be passed through a cell strainer to remove clumps.
9. Place cells on ice until ready to do flow cytometry.

Materials

• ThermoFisher’s Pierce Anti-DYKDDDDK Magnetic Agarose beads
• PBS
• Milli-Q water
• 1.5 mL microcentrifuge tubes
• M-PER reagent
• End-over-end microcentrifuge tube spinner
• Magnetic stand that can hold microcentrifuge tubes

Protocol

1. Equilibrate magnetic agarose to room temperature and invert to mix.
2. Place 50 uL slurry into a 1.5 mL microcentrifuge tube. Add 450 uL of M-PER reagent and gently vortex to mix.
3. Place the tube into a magnetic stand to collect the beads against the side of the tube. Remove and discard the supernatant.
4. Add 500 uL of M-PER reagent to the tube, mix well, collect the beads with a magnetic stand, and remove the supernatant. Repeat this once more for a total of two washes.
5. Add the sample (at least 300 uL; dilute with M-PER if necessary to reach this volume) containing 3xFLAG-tagged protein to the washed magnetic agarose and gently vortex to mix.
6. Incubate the sample at room temperature with constant end-over-end mixing for 20 minutes.
7. Collect the beads with a magnetic stand, then remove the supernatant.
8. Add 500 uL PBS to the tube, mix well, collect the magnetic agarose with a magnetic stand, and remove the supernatant. Repeat this once more for a total of two washes.
9. Add 500 uL of milli-Q water to the tube, collect the beads with a magnetic stands, and remove the supernatant.
10. Elute with one of the following protocols below, acid elution or sample buffer elution (our team has successfully tried both protocols, with similar yield).

Materials

• 0.1 M glycine (pH 2.8)
• 1 M tris (pH 8.5)

Protocol

1. Add 100 uL of 0.1 M glycine (pH 2.8) to the tube, mix well, and incubate for 5 minutes at room temperature with frequent vortexing.
2. Collect the magnetic agarose with a magnetic stand and save the supernatant, which contains the eluted protein.
3. To neutralize the acid, add 15 uL 1 M tris (pH 8.5) per 100 uL eluate.

Materials

• 1x Laemmli sample buffer

Protocol

1. Add 100 uL 1x Laemmli sample buffer to the tube, mix well, and incubate at 95C for 10 minutes.
2. Collect the magnetic agarose with a magnetic stand and save the supernatant, which contains the eluted protein.

Materials

• Fume hood
• SDS-PAGE gel mold (glass plates, spacers, gasket, well comb
• Serological pipette gun and 10 mL pipettes
• 12% resolving gel
• 4% stacking gel
• 10% w/v ammonium persulfate (APS)
• TEMED
• Isopropyl alcohol
• Sharpie
• 15 mL conical tube

Protocol

1. Degas the resolving and stacking gel solutions in a vacuum desiccator for 15 minutes.
2. Assemble the SDS-PAGE gel mold following the manufacturer's instructions.
3. Mark with Sharpie the level that is 1 cm below the teeth of the well comb.
4. Place a comb into the empty gel mold and mark with Sharpie the locations of each of your wells.
5. Set the resolving gel: add 15 mL of 12% resolving gel solution, 75 uL of fresh 10% w/v APS, and 7.5 uL of fresh TEMED to a 15 mL conical tube to initiate polymerization.
6. Gently invert the tube multiple times to ensure the TEMED and APS are evenly mixed.
7. Use a serological pipette to quickly transfer the solution to the casting mold, filling up to the line you marked earlier (~1 cm below the wells).
8. Add a layer of isopropyl alcohol to prevent exposure to oxygen. Allow the gel to polymerize for 90 minutes.
9. After the gel has polymerized, use Kimwipes to soak up the layer of isopropyl alcohol.
10. Place the gel comb into the gel mold with the teeth at a 15º angle (this prevents air from becoming trapped under the comb while the acrylamide solution is being poured). Allow the gel to polymerize for 30 minutes.
11. Set the stacking gel: add 5 mL of 4% stacking gel solution, 25 uL of fresh 10% w/v APS, and 5 uL of fresh TEMED to a 15 mL conical tube to initiate polymerization.
12. Gently invert the tube multiple times.
13. Use a serological pipette to transfer the stacking gel solution until the casting mold is filled.
14. Align the comb in the gel mold. Allow the gel to polymerase for 90 minutes.
15. Remove the gasket from the gel.

Materials

Protocol

1. Assemble the gel box according to the manufacturer's instructions. If only running one gel, insert empty gel glass plates on the side that is not being used to seal the inner chamber.
2. Place a magnetic stir bar at the bottom of the outer chamber.
3. Remove the gaskets and spacers from the gel mold. Place the gel mold with the gel in it into the gel box, following the manufacturer's instructions. The plate with the indentation should face inward.
4. Fill the inner chamber with running buffer to the top of the gel, so that buffer fills the wells.
5. Remove the comb by pulling it straight up slowly and gently. If the wells appear to have pieces of broken polyacrylamide in them, pipette Milli-Q water into the wells to rinse them.
6. Add ice packs to the outer chamber. The goal is to maintain an outer chamber temperature of 15ºC.
7. Fill the outer chamber with running buffer until the buffer reaches just below the sample wells.
8. Load your samples. For a 14-well, 1.5 mm gel, the maximum recommended loading volume is 30 uL. A good rule of thumb is to load 2 ug of purified protein or 200 ug protein of cell lysate
9. Load your protein ladder on the far left, middle, and far right, based on the number of samples you are loading.
10. Load unused lanes with 5 uL of 1x Laemmli sample buffer
11. Turn on the stirrer (130 rpm).
12. Run at 80 V (constant voltage) until samples have fully entered the stacking gel (around 30 minutes) then 120V (constant voltage) until the dye is near the bottom of the gel, typically around 3 hours.

Materials

• Coomassie blue staining solution
• Destaining solution
• Tupperware
• Large KimWipes or paper towels

Protocol

1. Rinse gel under tap water a few times to remove residual SDS.
2. Add 50 mL Coomassie blue staining solution to tupperware.
3. Place gel in the tupperware then microwave for 1 minute.
4. Place tupperware on shaking table for 10 minutes at moderate speed to constantly wash gel with staining solution.
5. Dispose of staining solution.
6. Rinse with Milli-Q water, used destaining solution. or used rinsing solution twice, each time pouring out the diluted solution into a bottle for rinsed solution.
7. Transfer the gel to a new tupperware and add 75 mL destaining solution.
8. Tear up a large KimWipse and surround gel with wipes, being careful not to allow wipes to lay over the gel (this causes uneven destaining).
9. Microwave for 1 minute or until solution boils.
10. Place on shaking table for 10 minutes at moderate speed.
11. Discard kimwipes and add another 25 mL destaining solution.
12. Prepare another 4-kimwipe masterpiece and surround gel with wipes, being careful not to allow wipes to lay over the gel.
13. Place on shaking table overnight at moderate speed.

Materials

• Solution containing IL-6, whether that may be after protein purification or directly from a HEK293 whole cell freeze-thaw lysate
• 12-well plate

Protocol

1. Add 100-115 uL of the IL-6 solution or up to 1 mL of the whole cell lysate containing IL-6 to the well.
2. Let the cells incubate at 37 degrees Celsius and 5% CO2 for 6 hours.
3. Prepare the cell samples for flow cytometry and proceed with analysis.