Objective

Author(s): Pattarin Blanchard

Prepare Blue-Green 11 (BG-11) nutrient medium to culture Synechococcus elongatus.

BG-11 Medium

For a final volume of 1 L:

1. Add ~500 mL of Milli-Q water to a clean 1 L media bottle (carboys by the lab coat rack).

2. Add the following stock components to the medium according to the table below.

   Component	Volume added (mL)	Final Conc.
   1000x MgSO₄	1	1x
   1000x K₂HPO₄	1	1x
   1000x CaCl₂	1	1x
   1000x Na₂CO₃	1	1x
   1000x ferric ammonium citrate + citric acid	1	1x
   1000x Na₂EDTA	1	1x
   1000x BG-11 trace metals	1	1x
   100x NaNO₃	10	1x

3. Top up with Milli-Q water to 1 L and autoclave. Label with iGEM, BG-11, date, and initials.

4. If necessary, add 1 mL of antibiotic stock under a flame, and add to the label.

BG-11 agar medium

For 1 L worth of agar medium:

2X BG-11 for agar plates

1. In a 1 L media bottle, add ~400 mL Milli-Q water.

2. Add the following components according to the table below:

   Component	Volume added (mL)	Final Conc.
   1000x MgSO₄	1	1x
   1000x K₂HPO₄	1	1x
   1000x CaCl₂	1	1x
   1000x Na₂CO₃	1	1x
   1000x ferric ammonium citrate + citric acid	1	1x
   1000x Na₂EDTA	1	1x
   1000x BG-11 trace metals	1	1x
   100x NaNO₃	10	1x

3. Top up with Milli-Q water to 500 mL and shake to mix well.

4. If only preparing stock, autoclave after the previous step, otherwise, continue to next step. ## 2X Agar solution

5. In a 500 mL bottle, add 15 g phytagel/gellan gum powder (if unavailable, regular bacto-agar) to 400 mL Milli-Q water.

6. Top up with Milli-Q water to 500 mL and shake to mix well.

Agar preparation

7. Autoclave both the 2x medium and agar solutions.

8. Aseptically add 1 mL 1000x sodium thiosulfate (Na₂S₂O₃) solution to the agar solution. This slows the solidification, making agar easier to work with.

9. Add the agar solution to the 1 L containing the 2x medium and mix well.

Pouring plates

10. If reusing prepared agar stock, melt agar by microwaving with the cap loosely on in 30-60s intervals to bring to a boil 3 times.

11. If adding antibiotic and/or inducers like theophylline, make an aliquot of the agar in a sterile flask. Decide on the number of plates beforehand; use 30 mL per 100mm plate, or 12 mL per 60mm plate.

12. For each pour, use a sterile 50 mL falcon tube to measure out the volume (30 mL for 100mm, 12 mL for 60mm). Hold the tube in the non-dominant hand, and on the dominant hand, alternate between picking up the bottle to measure out agar and picking up a plate lid to pour.

13. Store remaining agar medium at room temperature.

Objective

Author(s): Sarah Duan, Ada Jiang

This protocol describes heat-shock transformation of BL21(DE3) chemically competent E. coli (NEB C2527 series).

Notes:

Perform Steps 1—7 in the transformation tube (the tube the cells are thawed/aliquoted into). Keep cells cold until the heat-shock step. Avoid vortexing at all stages before outgrowth.

Materials

• BL21(DE3) Competent E. coli (C2527H)

• Plasmid DNA (1 pg—100 ng in 1—5 µL; supercoiled works best)

• SOC medium (or LB; SOC recommended)

• Prewarmed selective agar plates (e.g., LB-agar + appropriate antibiotic)

• Sterile 1.5 mL microcentrifuge tubes (for serial dilutions)

• Sterile pipette tips

• Optional: sterile water or SOC for dilutions

Equipment

• Ice bucket with crushed ice

• 42°C water bath or heat block (verified temperature)

• 37 °C shaking incubator (≈250 rpm) or rotator

• 37 °C incubator for plates (30°C or 25°C optional for slower growth)

• Pipettes (2—20 µL, 20—200 µL, 100—1000 µL)

• Sterile spreaders or glass beads

Protocol

Important: Do not vortex cells prior to outgrowth. Mix by gentle flicking or slow pipetting only.

1. Thaw cells: Place the 50 µL single-use tube on ice for 10 min.
2. Add DNA. Add 1—5 µL of plasmid DNA (total mass 1 pg—100 ng) to the 50 µL cells. Gently flick 4—5× to mix. Do not vortex.
3. Incubate on ice. Keep the cell—DNA mix on ice for 30 min. Do not mix during this period.
4. Heat shock. Transfer the tube to a 42°C water bath or heat block for exactly 10 s. Do not mix during heat shock.
5. Return to ice. Immediately place the tube on ice for 5 min. Do not mix.
6. Add SOC. Add 950 µL room-temperature SOC to the tube to bring the total volume to ~1 mL. Gently invert to mix.
7. Outgrowth. Incubate at 37 °C for 60 min with vigorous shaking (≈250 rpm) or tube rotation to allow expression of antibiotic resistance.
8. Prepare plates. Warm selective plates to 37°C (dry if overly wet).
9. Serial dilutions (recommended). Mix the culture thoroughly by flicking/inverting. Perform 10-fold serial dilutions in SOC (e.g., 10⁰, 10⁻¹, 10⁻², 10⁻³) in sterile 1.5 mL tubes.
10. Plate cells. Spread 50—100 µL of undiluted and/or diluted cultures onto selection plates. Allow liquid to absorb, then invert plates.
11. Incubate. Incubate plates overnight at 37 °C. Alternative growth: 30 °C for 24—36 h or 25 °C for 48 h for temperature-sensitive constructs.

Tips for Transformation

Thawing

• Best practice: thaw on ice and add DNA as soon as the last ice disappears. Hand-thawing is possible, but warming above 0°C lowers efficiency. Ice incubation (DNA + cells)

• 30 min on ice yields maximal efficiency. Expect ~2× loss for every 10 min you shorten this step. Heat shock

• 10 s at 42°C is optimal for a 1 mL total transformation in the supplied tube. Different volumes/vessels may require recalibration. Outgrowth

• 37°C, 60 min with shaking is optimal.

• Expect ~2× loss for every 15 min you shorten outgrowth.

• SOC typically gives ~2× higher efficiency than LB. Shaking/rotation also improves efficiency ~2× vs. static incubation. Plating

• Plates may be used warm or cold, wet or dry without large efficiency changes; however, warm, dry plates spread easiest and give faster colonies.

Objective

Author(s): Jessie Luo, Ada Jiang

While Caulobacter cannot be chemically transformed, it can be electroporated to introduce plasmids into the cells. This protocol describe and electroporation protocol adapted from iGEM British_Columbia 2016, Smit et al., 2000, and a protocol provided by our advisor, Beth.

Materials

• Electrocompetent cells
• PYE medium
• PYE Agar plates (with appropriate antibiotic - e.g. CM 2 µg/mL)
• Cuvettes (0.1cm interelectrodal gap)
• Electropulser (Eppendorf Eporator®)
• Shaking incubator at 30˚C
• Plasmid DNA (100-1000 ng/µL)
• 10 mL aeration/culture tubes (sterile)
• Glass beads
• Kimwipes / Paper towel
• Eppendorf tubes (sterile)

Set-Up

Preparation

1. Put cuvettes on ice for at least 20 mins to chill (or pre-chill by putting them in the fridge).
2. Thaw electrocompetent CB2A cells on ice (one tube per plasmid). Do not handle the cells vigorously, as this will reduce transformation efficiency.
3. (Optional) Thaw DNA samples. Put 1 µL of 100-1000 ng/µL undiluted DNA in new eppendorf tubes (labelled) on ice.
4. Pre-warm PYE medium by placing a 50 mL aliquot of medium in 30 or 37˚C for about 15 minutes.
5. Set up a sterile workspace (in the flow hood, BSC or under the flame).
6. Plug in, turn on and adjust the electropulser to the appropriate settings: 2500 V

Procedure

Electrotransformation

1. Transfer 50 µL electrocompetent cells to tube with 1 µL DNA on ice. Or directly add the appropriate amount of DNA to the cells.

2. Wait 1-2 minutes. In this time, dry a cuvette thoroughly with paper, remove its lid and place it in the cuvette holder such that the plastic nose points towards the back.

3. Transfer the 50 µL DNA/cell mix to the cuvette (careful to place in 1 mm slit) and slide the cuvette holder into the electropulser until you hear a click, confirming the holder is locked in place. Click START to pulse. Uptake 950 µL PYE medium into a p1000 pipette.

4. Take out the cuvette. Resuspend cells gently, but immediately in 950 µL PYE medium.

5. Transfer cell suspension to an aeration tube. Use a p200 with gel loading tips to extract remaining cells from the cuvette.

6. Incubate at 30°C, 200 rpm shaking for 2-4 hours.

7. Plate on PYE/CM and incubate 30°C for 2-7 days.

Notes on arcing:

• When pulsing, the reading gives a number between 1-5 - this is the time constant. 4.5-5 is best.
• If arc happens on pulsing (spark etc), then cells are useless - dispose.
• Arcing happens if salt concentration is too high. Can dilute the DNA used with water and try again with new cells if this happens.
• Ensure cuvettes are ice cold, and wiped properly before pulsing.

Objective

Author(s): Arda Sayla

This protocol describes how to prepare and preserve CB2A culture for long-term storage at -70 °C.

Procedure

1. Pipette 100 µL sterile glycerol into a sterile cryoculture or Eppendorf tube. Add 900 µL of resuspended pellet/culture (final glycerol concentration will be 10%).
2. Store the glycerol stocks at -70 °C.

This can also be done using 5% dimethyl sulfoxide (DMSO). Sterilization of freezer vials with about 50 µL of DMSO is recommended. Pipette 1 ml of fresh culture into the vial, mix briefly, and freeze using dry ice or liquid nitrogen.

This frozen stock should never be fully thawed. When cultures are to be inoculated, take out a stock on ice. Under the flame, scrape a little of the “ice” from the surface using a sterile tool and inoculate a plate or liquid culture.

Objective

Author(s): Ada Jiang

Clean and sterilise plating beads for reuse in bacterial transformation experiments.

Materials

• Waste beaker with dirty beads
• Beaker (250 mL or larger to catch beads easier)
• 10% bleach
• 70% EtOH
• Deionized (DI) / MilliQ water
• Aluminum foil
• Paper towels
• Autoclave bin
• Autoclave
• Metal spatula/spoon (or you can try mixing the beads around by swirling the container, not as effective as stirring though)
• Glass test tubes and tube rack Perform steps by the sink (or in the autoclave bin to avoid beads falling into the sink).

Procedure

1. When finished plating, discard used beads into a reserved 100 mL waste beaker. Once there are a lot of beads accumulated in the waste beaker (or stock of sterilized beads is running low), begin re-sterilization.
2. Fill a clean beaker with 10% bleach. Transfer dirty beads to the new beaker by flipping the sieve upside down and gently tapping on the side of the beaker. Let the beads soak for 30 mins to kill cells. Can skip if beads are already soaking in 10% bleach from waste container.
3. Decant bleach solution from the beaker, catching the beads with a sieve (found sitting at the top shelf at the left end of the bench, above the ice boxes).
   1. Transfer the beads back into the beaker.
   2. Fill up the beaker with tap water until beads are fully covered, liquid level should be at least 2 cm over the level of beads.
   3. Use a clean metal spatula/spoon to stir the beads around.
4. Repeat Step 3 three more times. Once more with tap water, then the last two washes with DI (use the white DI tap at the sink) or Milli-Q water.
5. Repeat Step 3 once with 70% EtOH.
6. Find a clean plastic container/any container with wide surface area. Line the container with paper towel, and cover with a clean sheet of aluminum foil. Gently shake the container a bit so that most of the beads are spread out and aren’t clumping. Let the beads dry overnight or put them in the 37C incubator for a few hours.
7. Check that the beads are dry. They should be rolling around without sticking to one another.
8. Carefully transfer the beads to a clean beaker, and aliquot the beads into clean glass test tubes (preferably the tubes have been sterilized beforehand). Fill up each tube to about 1/3 (or less) of the volume of the tube. Alternatively, you could autoclave the beads in the beaker first then aliquot into sterile tubes under the flame.
9. Autoclave the beads in the beaker/tubes with dry cycle (40ish mins).

Objective

Author(s): Ada Jiang

Screen bacterial colonies for presence of desired DNA construct by directly using cell lysate for PCR amplification.

Procedure

1. Prepare template material. There are several options:
   1. Pick a colony (ideally fresh) and mix directly in PCR reaction mixture (will look slightly cloudy, don’t pick up too much cell material)
   2. Option A but replace water content in PCR mix with TE buffer.
   3. Pick a colony and dissolve in 50-100 uL media / DMSO / water. Use 1 µL of the cell lysate for PCR.
   4. Use 1 uL of fresh overnight cultu
2. Run PCR cycle, using Polymerase Chain Reaction (PCR) with NEB Q5 High-Fidelity 2X Master Mix.
   1. Consider setting a longer initial denature step to help break up the cells.

Objective

Author(s): Pattarin Blanchard, ZiQi Chen

Prepare nutrient-rich medium for recovery of cyanobacteria after electroporation. This is a modified recipe of Castenholtz D Medium (DTN), based on Mühlenhoff⁩ and ⁨Chauvat⁩, 1996; Katoh et al., 2003; and ⁨Wiktelius⁩ and ⁨Stenberg⁩, 2007.

Procedure

Recipe is for 1 L medium.

1. If not already done, prepare 10 mL of the following 1000x trace metals solution in Milli-Q water:

   Component	Amount (g)	Notes
   FeCl₃·6H₂O	0.01
   NH₄Cl	0.05
   CuSO₄⋅5H₂O	0.25
   Na₂MoO₄⋅2H₂O	0.25
   CoCl₂⋅6H₂O	0.45
   NiSO₄⋅6H₂O	0.08	Original is 0.019 g of nickel ammonium sulfate in 1 L
   NaSeO₃	0.04	Original was 0.004g of NaSeO₄ in 1 L

2. Add ~500 mL of Milli-Q water to a 1 L media bottle. Weigh out and add the following components

   Component	Amount (g)	Notes
   Disodium EDTA	0.19
   MgSO₄	0.053
   CaCl₂·6H₂O	0.05	originally 0.042 g of CaCl₂·4H₂O
   KNO₃	0.1
   NaNO₃	0.7
   NaH₂PO₄·H₂O	0.11	originally 0.12 for NaH₂PO₄·2H₂O
   Na₂S₂O₃·5H₂O	0.157	originally 0.1 for anhydrous
   Na₂SO₃	0.2	per Mühlenhoff⁩ and ⁨Chauvat⁩, 1996 → 1.6 mM final conc

   | H₂SO₄ (pure) | 0.5 mL | | H₃BO₃ | 0.4 | originally 0.5 g of H₃BO₄ | MnSO₄⋅H₂O | 1.6 | Originally 2.28g of pentahydrate | ZnSO₄⋅7H₂O | 0.5 | | Tricine | 1.43 | 8 mM final concentration | Trace Elements Solution | 1 mL |

3. (Optional) adjust to pH 8 using 1 M NaOH.

4. Autoclave at LIQUID20.

Objective

Author(s): Pattarin Blanchard

Safely deactivate and dispose of bacterial culture.

Procedure

1. Add 10% bleach or 1 M NaOH solution to the culture in a 1:1 volume ratio.
2. Allow at least 30 mins contact time before pouring down the drain with running water.
   1. Leaving overnight in the 4°C room prior to disposal is preferred, but the above is sufficient.

Objective

Author(s) : Ada Jiang

Determine the size of DNA fragments and/or reaction products based on separation due to size and charge. Following this procedure, gel purification can be conducted to isolate and purify target DNA. This protocol was adapted from Mupid®-One and Thermo Scientific Owl™ EasyCast™ B1A Mini Gel user guides.

Materials

• 1xTAE buffer
• DNA ladder: FroggaBio 1 kB DNA Ladder or NEB 1kb Plus Ladder
• DNA loading buffer: Neb Gel Loading Dye, Purple (6X)
• SYBR Safe DNA Gel Stain
• DNA samples
• Agarose powder
• Gel electrophoresis equipment

Procedure

Melting agarose solution

1. Combine 1xTAE and agarose powder to create the desired thickness and concentration of gel for your application.
   1. Measure out the agarose powder in a dry 250 mL Erlenmeyer flask
   2. Fill the flask with 1xTAE to the needed volume
2. Microwave for 1:30-2:00 min.

• Check every 30s after the initial 1 min, to make sure no over spill due to bubbling
• Stop when the liquid has no or minimal visible translucent pellets

1. Add SYBR Safe into agarose when the liquid stops boiling (<60°C)

• 10,000 dilution (for example: 60 mL gel needs 6 µL SYBR safe)
• Swirl to mix well

Prepare loading samples

1. Add loading buffer to each sample for a final concentration of 1x dye.
2. Gently flick the tubes to mix, then spin down.
3. Consider boiling the samples if enzymes from an upstream reaction could inhibit movement though the gel.

Thermo Scientific™ Owl™ EasyCast™ B1A Mini Gel System

Casting the gel

1. Insert rubber strips (gaskets) into the grooves of a gel tray, then insert the tray into the buffer chamber so that the gaskets face the walls, sealing off the tray. This is the Casting position.
2. Insert the correct size comb (10-wells). Carefully pour the molten agarose, and wait until the agarose hardens (~15-20 min).

• Use a clean pipette tip to remove bubbles (push them out to the corners).

1. Once hardened, reorient the entire chamber such that the electrodes are pointing to the right. This is the Running position.
   1. Lift the tray up and place it back such that well side is closer to the wall of the chamber that is furthest from your body.
   2. If done correctly, the black/negative electrodes will be further from you than the red/positive electrodes.
   3. Remove the comb.

Running the gel

1. Fill up the chamber with 1X TAE buffer (can be reused) until the buffer fully covers the gel.

2. Load 3 µL of DNA ladder, and add DNA samples. To determine amount of sample to load, estimate the volume of sample needed to load at least 10 ng of DNA or for your intended application.

   Application	Volume of TAE	Max amount of sample
   Analysis (thin gel)	30-40 mL	15 µL
   Purification (thick gel)	60 mL	50 µL

3. Run the gel at 100-150V until desired distance.

4. Prepare a piece of saran wrap large enough to fold over the gel. Retrieve the gel by lifting up the tray and carefully sliding the gel into the saran wrap. Pour the TAE back into its bottle.

Mupid®-One Electrophoresis System

Casting the gel

1. Place the casting stand on a level surface. Insert the center partition if to make a small gel. Insert the gel tray(s).
2. Insert the correct size comb (13 or 26-wells). Carefully pour the molten agarose, and wait until the agarose hardens (~15-20 min).

• Use a clean pipette tip to remove bubbles (push them out to the corners).

1. Once hardened, transfer the gel trays to the electrophoresis chamber.

   1. Remove the comb.

2. Connect the power supply to the chamber. Running the gel

3. Fill up the chamber with 1X TAE buffer (can be reused) until the buffer fully covers the gel.

4. Load 3 µL of DNA ladder, and add DNA samples. To determine amount of sample to load, estimate the volume of sample needed to load at least 10 ng of DNA or for your intended application.

   Application	Volume of TAE	Max amount of sample
   13-well (thin gel)	30-40 mL	15 µL
   26-well (thick gel)	60 mL	50 µL

5. Run the gel at 50-135V until desired distance (generally 30 minutes).

6. Prepare a piece of saran wrap large enough to fold over the gel. Retrieve the gel by lifting up the tray and carefully sliding the gel into the saran wrap. Pour the TAE back into its bottle.

Visualizing the gel

7. Check the sample bands using the UV gel imager.

8. Use upstairs gel imaging devices to visualize the bands and print a copy of the gel.

Objective

Author(s) : Pattarin Blanchard

Purify DNA fragments for downstream cloning. This protocol is based on the EZ-10 Spin Column PCR Products Purification Kit.

Materials

• Bio Basic EZ-10 PCR purification kit
• 2x microcentrifuge tube per sample + 1 to warm elution buffer
• 1x EZ-10 column per sample
• Heat block set to 50C
• Centrifuge
• (Optional) PCR tubes

Procedure

1. Unless there is an existing aliquot of elution buffer, make a 1 mL aliquot of elution buffer in a microcentrifuge tube. Prewarm to 55-80C using a heat block
2. Each sample will have 2 microcentrifuge tubes; a mixing tube, and a purified product tube, as well as a column. Label the tubes accordingly.
3. Transfer PCR product to a microcentrifuge tube and add 5x volume of buffer B3 (e.g. 25 uL PCR product ---> add 125 uL buffer B3). Mix well.
4. Transfer contents to spin column and wait 2 minutes, then centrifuge at 10,000 RPM for 2 minutes.
5. Discard flow-through and add 750 uL wash solution. Centrifuge at 10,000 RPM for 2 minutes. Do this twice.
6. Discard the flow-through and centrifuge the empty column at 10,000 RPM for 1 minute to discard residual wash solution.
7. Transfer column to clean centrifuge tube (and optionally, place in heat block at 37-50C, for >10 kb fragments). Add 30-50 uL of elution buffer to the centre of the column (there is a visible raised ring on the edge). Incubate for 2 minutes.
   1. Selected volume should correspond as closely as possible to volume of PCR reaction - e.g. 25 uL PCR rxn ---> 30 uL elution buffer (minimum)
8. Centrifuge column and tube at 10,000 RPM for 2 minutes.
9. Optionally, transfer elute to PCR tubes.

Objective

Author(s): Jessie Luo, Ada Jiang

Preparation of CB2A cells to create electrocompetent cells, which are receptive to DNA uptake via electroporation for construct expression. This protocol is adapted from a protocol provided by our advisor, Beth.

Materials

Colony plates

• CB2A PYE plate colonies

Media/Solutions

• (550 mL) PYE liquid medium
• (1 L) Distilled or Milli-Q water, sterile
• (55 mL) 10% glycerol

Equipment

• 250 mL sterile flask with foil
• 2 L sterile flask with foil
• 10 sterile Falcon tubes
• 50 sterile 1.5 mL Eppendorf tubes
• 1 cm cuvettes for spectrometer
• Pipette boy and 10 mL sterile pipettes
• Low-speed centrifuge

Procedure

Set-up

Day 0 - Pre-culture

1. Inoculate 50 mL of PYE culture in a 250 mL flask with a CB2A colony.
2. Shake at 30˚C for ~48 hours. If incubating for over 48 hours, you can grow the culture on the bench at room temperature until the appropriate optical density at 600 nm (OD600) is reached.

Day 2 - Culture set up

1. Early morning: Inoculate culture with pre-culture.
   1. Use a spectrometer to find the pre-culture OD600.
   2. Fill a 2 L flask with 500 mL of PYE media.
   3. Add the correct volume of pre-culture to make OD600 0.1 in 500 mL of PYE. If pre-culture OD5, add 10 mL pre-culture to 480 mL fresh PYE culture for OD0.1 (1:50 dilution)
2. Shake at 30˚C until OD600 is 0.3-0.6. This should take 3-7 hours.

Electrocompetent cell preparation

Wash Steps

1. Pour culture into 50 mL Falcon tubes (10), in the flow hood or sterile environment.
2. Centrifuge the culture at 3700 xg for 20 min.
3. Pipette out the culture supernatant and discard (pouring may also work but may dislodge the pellet - be careful if doing this).
4. Add about 5 mL sterile cold water to each bottle and resuspend with a 10 mL pipette carefully.
5. Then, fill the bottles to the top with sterile cold water (“full volume wash”) and centrifuge at 3700 xg for 20 minutes.
6. Pipette off supernatant (note: pellets tend to be looser after the water washes).
7. Repeat the process for suspension of each pellet in 2 mL sterile cold water
8. Fill bottles approximately half full with sterile cold water (“half wash”).
9. Centrifuge at 3700 xg for 20 min
10. Pipette out and discard supernatant.

Glycerol Steps

1. Suspend pellets (10) in 5 mL 10% glycerol each.
2. Combine suspensions into 2 Falcon tubes and centrifuge at 3700 xg for 15 min.
3. Discard supernatant.
4. Resuspend pellets (2) in 0.75 mL 10% glycerol each.
5. Aliquot 50 μL portions into sterile 1.7 ml Eppendorf tubes and freeze in -70 to 80°C.

Objective

Author(s): Pattarin Blanchard

Isolate and purify DNA fragments from agarose gel for downstream cloning experiments.

Materials

• GeneJET gel extraction kit
• 2x Microcentrifuge tube per gel slice
• Scalpel or razor blade
• Water bath or heat block
• Purple purification column

Procedure

Gel extraction

1. Weigh and record a microcentrifuge tube for each extraction.
2. Cut away the desired gel slice using a clean scalpel or razor blade and place into the preweighed tube.
3. Add 1:1 volume (uL) : weight (mg) ratio of binding buffer (e.g. 100 uL binding buffer for 100 mg gel slice).
4. Incubate tube at 50-60 C for 10 minutes (until gel is completely dissolved). Invert the tube every few minutes to aid dissolution.
5. (OPTIONAL) if DNA is:

   1. <500 bp, add the same volume isopropanol as binding buffer in step 3 and mix thoroughly.

   2. 10 kb, add the same volume ultrapure water as binding buffer in step 3 and mix thoroughly.

Gel purification

1. Transfer up to 800 uL (at a time) of the dissolved gel to a purification column.
2. Centrifuge at 12,000 g or rcf (not rpm) for 1 minute. Discard flow-through and reuse collection tube.
   1. See additional step if preparing DNA for sequencing in the linked manual.
3. Add 700 uL wash buffer supplemented with ethanol (should have been done on the first use) to the purification column.
4. Centrifuge at 12,000 g or rcf for 1 minute. Discard flow-through and reuse collection tube.
5. Centrifuge again at 12,000 g or rcf for 1 minute to remove residual wash buffer.
6. Place purification column into a new microcentrifuge tube.
7. Add 30-50 uL elution buffer or ultrapure to the centre of the column membrane. Centrifuge at 12,000 g or rcf for 1 minute and discard the column.
   1. 50 uL is the standard volume, however if DNA weight is expected to be low, reduce down to 30 uL.
8. Nanodrop to determine concentration. See NanoDrop Spectrophotometer. Remember to blank with UPH2O.

Objective

Author(s): Pattarin Blanchard

This protocol will produce a curve that quantifies cell biomass at a given OD.

Materials

• Cuvette
• Kimwipe
• 10x Microcentrifuge tubes
• 15 mL falcon tube
• 50 mL falcon tube
• 20 mL Serological pipette
• PBS
• Liquid culture
• Milli-Q water
• Sterile pipette tips or serological pipette
• Aluminum drying dish
• Analytical balance
• Spectrophotometer
• Waste beaker

Procedure

OD measurements

1. For ease of working and to avoid contamination, aliquot just over 5 mL of liquid culture into a 15 mL falcon tube.

2. Under a flame, make the following dilutions of liquid culture in PBS in appropriately labelled microcentrifuge tubes. Use the aliquoted liquid culture.

   Tube	1	2	3	4	5	6	7	8	9	10
   Liquid culture (µL)	0	50	100	200	300	500	600	750	1000	1500
   PBS (µL)	1500	1450	1400	1300	1200	1000	900	750	500	0

3. Blank the spectrophotometer with Milli-Q water at the appropriate wavelength for the organism.

   1. E. coli, CB2A: 600 nm
   2. UTEX: 750 nm

4. Measure the OD of each tube starting from tube 1, wiping the cuvette with a kimwipe before each measurement. After each measurement, discard the contents in a waste beaker, rinse 3 times with water, then leave face-down on a paper towel soaking with ethanol or disinfectant if not reusing right away.

Dry cell weight

5. Aliquot 25 mL of liquid culture into a 50 mL centrifuge tube and centrifuge at 10000G for 5 mins.

6. In the meantime, weigh the drying dish in the analytical balance.

7. Remove supernatant and resuspend with 10 mL PBS and centrifuge again at 10000G for 5 mins.

8. Repeat step 3 once.

9. Remove supernatant and resuspend with 3-5 mL water, try to recover all of the pellet. Pour into the drying dish.

10. Place the drying dish in an oven and come back the next day to weigh the dish.

11. Calculate the dry cell “weight” (per mL) from the obtained measurements:

    $$\rho = \frac{m_{dry}-m_{empty}}{25 \text{ mL}}$$

Dry cell weight curve

1. Use the value from the previous step to calculate the dry cell weight at each dilution:

   $$DCW=\frac{\rho \cdot V_{culture}}{1.5 \text{ mL}}$$

2. Calculate true OD by subtracting each value with the OD of PBS only (tube 1).

3. Plot true OD vs. DCW, it should be approximately linear between OD 0.1-1.

4. Perform a linear regression on the data points in the linear region. The reciprocal of the slope is the desired quantity and has the units $\text{g/mL}\cdot \text{AU}$.

Objective

Author(s): Pattarin Blanchard

Monitor bacterial growth over time using spectrophotometric measurements of culture density. This protocol will produce a curve that can be used to determine the target organism’s rate of growth (doubling time).

Materials

• PBS (phosphate-buffered saline)
• Milli-Q water
• Sterile pipette tips or 1 mL serological pipette
• x2 disposable cuvettes
• Parafilm
• Kimwipes
• Spectrophotometer
• Liquid medium
• 500 mL culture flask
• Waste beaker with 10% bleach

Procedure

Inoculating a new culture

It is desired to have starting optical density (OD) of 0.1 (for the sake of time) in the new flask culture. Therefore, when possible, the volume of inoculum from the mother culture has to be calculated, and the OD of the mother culture has to be measured.

1. Using a new cuvette, blank the spectrophotometer with 600 µL water. Use a kimwipe to clean the outside of the cuvette prior to measuring. Choose an appropriate wavelength. Dump the contents into a waste beaker.
   1. E. coli: 600 nm
   2. C. crescentus (CB2A): 600 nm
   3. S. elongatus (UTEX): 750 nm
2. Using the same cuvette, add 600 µL growth medium or diluent and measure the OD. Record this value down, as this will be subtracted from every subsequent measurement. Dump the contents and rinse the cuvette with water using a wash bottle 3 times into the waste beaker.
   1. Alternatively skip step 1 and blank with the medium, but step 1 is standard practice.
3. Under a flame, take 1 mL of the mother culture, add it to a cuvette, and measure its OD. Note that this doesn’t measure the cell color, but rather turbidity, or light scattering.
4. Generally OD measurements are only accurate between 0.1 and 1. If the initial reading is above 1, dilute it as necessary with medium or PBS so that the absorbance is between 0.1 and 1. Record the value and dump the contents into the waste beaker. Rinse with water 3 times, then set it face down onto a paper towel soaking with ethanol or disinfectant.
   1. A cuvette has a max volume just over 3 mL, if up to a 3x dilution is needed, it is okay to dilute in the cuvette itself and mix by pipetting up and down.
5. Calculate the volume of inocculum needed from the mother culture. First subtract the diluent (PBS or medium) OD, then multiply by the dilution factor to get the “true” OD of the mother culture. $$OD_{true}=(OD_{measured}-OD_{diluent})\cdot DF$$

Then calculate the volume needed. The final volume shall be 250 mL + volume added, and the starting OD is 0.3. Note the final equation gives volume in mL, so make the appropriate conversion to uL if necessary.

$$OD\_{inoc}V\_{add}=OD\_{start}(250 + V\_{add})$$ $$V_{add}=\frac{250\text{ mL}\cdot OD_{start}}{OD_{inoc}-OD_{start}}$$ $$V_{add}=\frac{250 \text{ mL}\cdot 0.3}{OD_{inoc}-0.3}$$

1. Under a flame, add 250 mL of medium to a culture flask, then inocculate with $V_{add}$ calculated from the previous step.

2. Sample 1 mL from the new culture and measure its OD using the same cuvette from earlier. Dump cuvette contents into the waste beaker and rinse 3 times with water, then set face down on a paper towel soaked with ethanol or disinfectant.

Taking measurements

3. It is recommended a spreadsheet is used instead of a notion table because calculations will have to be performed. Set up a table like so in a spreadsheet, the initial readings should be recorded like so:

   Time (h)	Dilution factor	Measured OD	True OD	Initials
   PBS	1	x	0
   0	1	~0.3+x	~0.3

4. An appropriate interval should be selected for the organism. For example, UTEX has an expected doubling time of around 2 hours, so samples will be taken in 1 hr intervals.

5. For each subsequent OD measurement:

• Always take a sample under a flame with sterile tips or serological pipettes.
• Anticipate when a dilution should be increased, knowing that OD is generally accurate between 0.1 and 1.
• Ensure the spectrophotomer is set to the correct wavelength. If it is being used frequently by other people, keep a second cuvette with Milli-Q water to blank it as needed.
• For dilutions up to 3x, 1 mL of sample can be taken and can be diluted in the cuvette itself since the max working volume is 3 mL. If a higher dilution is anticipated, take a smaller volume of sample (e.g. 5x dilution: sample 0.6 mL, dilute with 2.4 mL).
• Wipe the cuvette with a Kimwipe to ensure an accurate reading.
• Always rinse the cuvette 3 times with water and set it face down on a paper towel soaking with ethanol or disinfectant. The paper towel needs to be resprayed every once in a while, make sure it does not dry.

1. Once finished, calculate the true OD using this formula: $$OD_{true}=(OD_{measured}-OD_{PBS})\cdot DF$$

Analysing the growth curve

1. Plotting true OD vs. time should give a curve that looks exponential or logistic. Plotting $\ln (\frac{OD_t}{OD_0})$ vs. time should give a linear section. Why this works is during the exponential growth phase, cell growth follows this differential equation:

   $$\frac{dX}{dt}=\mu X$$

   Which when solved yields:

   $$X(t)=X_0e^{\mu t}$$

   And can be linearized as the following. Here OD is a stand-in for cell concentration $X$

   $$\ln(\frac{X}{X_0})=\mu t$$

2. Select the log data points in the linear section and run a linear regression versus time. The resulting slope is the growth rate $\mu$. If using LINEST, the standard error of $\mu$, $\Delta \mu$ is the value just below it. Choose consecutive data points such that the $R^2$ is maximized, but prioritize as many data points as possible.

3. Calculate the doubling time $t_d$:

   $$t_d=\frac{\ln 2}{\mu}$$

   The error in doubling time can be propagated from the error in growth rate using a first-order approximation

   $$\Delta t_d=\bigg |\frac{\partial t_d}{\partial \mu}\bigg |\Delta \mu=\bigg |\frac{\ln 2}{-\mu^2} \bigg |\Delta \mu = \frac{\ln 2}{\mu^2}\Delta \mu$$

Objective

Author(s): Ada Jiang

Assemble multiple DNA fragments into a single construct using Gibson cloning method for seamless, directional ligation. This protocol is based off of the NEBuilder® HiFi DNA Assembly Reaction Protocol.

Materials

• DNA fragments
• NEBuilder HiFi DNA Assembly Master Mix (2X)
• Molecular grade water
• PCR tubes
• Thermocycler or heat block

Procedure

1. Calculate the number of pmols of DNA components using the NEBuilder Protocol Calculator.

• For 2-3 fragment assembly: 0.03 - 0.2 pmols of DNA
• For 4-6 fragment assembly: 0.2 - 0.5 pmols of DNA > Note: The efficiency of assembly decreases as the number or length of fragments increases.

1. On ice, add the following components to set up the reaction: (scale proportionally for larger reactions)

   Component	Volume (µL)	Final Concentration
   DNA fragments	X amount calculated from Step 1	0.03 - 0.5 pmols, in total
   NEBuilder
   HiFi DNA Assembly Master Mix (2X)	10	1X
   Water	up to 20 μl

• For 2-3 fragment assembly: vector:insert = 1:2
• For 4-6 fragment assembly: vector:insert = 1:2 > Note: Optimized cloning efficiency is 50 - 100 ng of vector with 2-fold excess of each insert. Use 5-fold molar excess of any insert(s) less than 200 bp. > Note: The total volume of unpurified PCR fragments in assembly reaction should not exceed 20%. Consider purifying fragments for optimal efficiency.

1. Incubate samples in a thermocycler at 50°C.

• For 2-3 fragment assembly: 15 minutes
• For 4-6 fragment assembly: 60 minutes

1. Store samples on ice, OR at -20°C if transformation will be carried out in a different lab session.

Objective

Author(s): Pattarin Blanchard

Assemble multiple DNA fragments into a single construct using Golden Gate cloning method for seamless, directional ligation. This protocol is based off of the Golden Gate Assembly Protocol for NEBridge® Ligase Master Mix (NEB #M1100).

Materials

Reagents

Keep the following on ice:

• NEBridge Ligase Master Mix

• DNA fragments

• BsaI-HF-v2 or SapI

• Molecular water

• PCR tubes

• PCR tube rack, pre-chilled Equipment

• Thermocycler

Procedure

1. Use NEBioCalculator 1.17.3 to calculate how much volume of each fragment is needed such that 0.05 pmol of each fragment is added. Use the Oligonucleotide/Gene Database to retrieve fragment lengths. It is recommended to use a table like so in Excel to calculate:

   Name	Label	Length (bp)	Mass (ng) for 0.05 pmol	Concentration (ng/ul)	Volume (uL)

   1. Recall $C=\frac{m}{V}$ so $V=\frac{m}{C}$
   2. Synthesized gene fragments are resuspended to a standard concentration of 10 mM unless otherwise stated.

2. On ice or a pre-chilled PCR tube rack, add the components into a PCR tube in the following amounts for each reaction:

   1. Add water, then the DNA fragments. Since volumes may be very small, make sure the tip touches the water when pipetting down.
   2. Then add the remaining components, and pipette up and down 3-5 times gently to mix.

For 2-6 fragments (including backbone):

| Component | Volume (µL) |

--- | --- | | NEBridge Ligase Master Mix 3X | 5 | DNA fragments | 0.05 pmol each (from previous step) | BsaI-HFv2 or SapI | 1 | Molecular water | top up to 15 µL | Total | 15

For 7+ fragments (including backbone)

| Component | Volume (µL) |

--- | --- | | NEBridge Ligase Master Mix 3X | 10 | DNA fragments | 0.05 pmol each (from previous step) | BsaI or SapI | 1 if BsaI, 2 if SapI | Molecular water | top up to 30 µL | Total | 30

1. Run the appropriate cycle in the thermocycler:
   1. 2 fragment assembly (single insertion):
      1. 37°C for 15 min
      2. 60°C for 5 min
      3. 4°C infinite hold for retrieval
   2. 3-6 fragment assembly:
      1. 30 cycles of [37°C for 1 min + 16°C for 1 min]
      2. 60°C for 5 min
      3. 4°C infinite hold for retrieval
   3. 7+ fragment assembly:
      1. 30 cycles of [37°C for 5 min + 16°C for 5 min]
      2. 60°C for 5 min
      3. 4°C infinite hold for retrieval
2. Proceed to transformation or agarose gel electrophoresis, or freeze in -20°C for later use.

Objective

Author(s): Pattarin Blanchard, Ada Jiang

Transform chemically competent E. coli cells to enable uptake of DNA plasmid for replication of genetic material and downstream protein expression. This protocol uses a thermocycler for the heat shock and incubation step.

Materials

Reagents

• Sterile PCR tubes
• Competent E. coli cells, kept on ice
• LB agar plates with appropriate antibiotic (and additives)
• Glass beads, glass spreader, or sterile loop
• Floating foam tube rack
• Sterile microcentrifuge tubes
• LB or SOC medium, room-temperature
• Plasmids/inserts, thawed on ice
• Parafilm strips

Equipment

• Thermocycler
• Bunsen burner / Flow hood / BSC
• Timer
• Ice bucket

Procedure

Set up the thermocycler program. These cycles can be combined to one program (set the volume to 52 uL) or they can be set up before the appropriate steps.

Heat shock cycle:

Set volume to 12 µL

1. 4°C for 30 mins
2. 42°C for 10-45 secs, depending on competent cell strain
3. 4°C for 5 mins (Optional) Add an additional 4°C infinite hold step to keep the cells in the thermocycler longer than 5 mins before recovery step.

Recovery cycle:

Set volume to 50-100 μL

1. 37°C for 1 hr
2. 4°C infinite hold

Transformation

1. Chill PCR tubes on ice and label appropriately. Place agar plates in the 37°C room to warm.
2. Aseptically add 10 µL competent E. coli to a sterile PCR tube for each transformation desired.
3. Add 2-5 µL DNA or assembly product for each transformation desired. Close tube and pipette up and down to mix.
   1. Consider having one tube with only bacteria (no DNA added) for a negative control.
   2. Consider spinning everything down.
4. Place PCR tubes in the thermocycler and run the heat shock cycle (see above).
5. Once complete, retrieve tubes and, under a flame, add 88 µL SOC (total volume 100µL) medium to each well. Flick to mix.
   1. Spin down the PCR tube before moving on, not enough to pellet bacteria.
6. Place PCR tubes in the thermocycler and run the recovery cycle (see above).
7. Under a flame or in a BSC, plate cells on agar plates and leave the lid off for ~5 minutes to dry
   1. For one transformation per plate, add 50 µL (Or everything, if possible) of transformed bacteria and spread with 7-10 glass beads. Once finished, carefully “pour” the beads into the dirty beads beaker.
   2. If splitting plates into sections (i.e. halves, quadrants, or sixths), add 20 µL of transformed bacteria per section and spread with a spreader or loop.
8. Once sufficiently dried, close and wrap plates, label appropriately, and place in the 37°C room. The label must have
   1. iGEM
   2. transformant genotype or shorthand e.g. pRep-HpCA. If a shorthand is used, make sure it is clearly commuicated in the corresponding lab notebook entry
   3. Antibiotic (and other additives)
   4. Date plated
   5. Initials

Objective

Author(s): Pattarin Blanchard, Ada Jiang

Transform chemically competent E. coli cells to enable uptake of DNA plasmid for replication of genetic material and downstream protein expression. This protocol uses a water bath for the heat shock step.

Materials

Reagents

• Competent E. coli cells, kept on ice
• LB agar plates with appropriate antibiotic (and additives)
• Glass beads, glass spreader, or inoculating loop
• Floating foam tube rack
• Sterile microcentrifuge tubes
• LB or SOC medium, room-temperature
• Plasmids/inserts, thawed on ice
• Parafilm strips

Equipment

• Water bath set to 42°C
• Bunsen burner
• Timer
• Ice bucket

Procedure

1. Label and pre-chill centrifuge tube(s) on ice. Ensure the water bath is ready at 42°C. Warm agar plates in the 37°C room.
2. Under sterile conditions, add 50 µL competent cells per tube and keep on ice.
3. Pipette 2-5 µL (higher end for larger plasmids) into a centrifuge tube with comp cells. Close tubes and flick gently to mix, but do not vortex.
4. Incubate tubes on ice for 30 mins.
5. Using a floating tube rack, heat shock tubes in the 42°C water bath for 30s, then rest tubes on ice for 5 mins.
6. Under sterile conditions, pipette 950 µL LB or SOC medium (no AB needed) into each tube. This volume can be reduced to concentrate transformants.
7. Incubate tubes in the 37°C room for 1 hour, preferably on a shaker or rotor. Use one of the designated 50 mL falcon tubes to place microcentrifuge tubes in for a shaker.
8. (Optional) For experimental tubes and positive controls, perform a 10x dilution in LB or SOC medium in a sterile microcentrifuge tube (100 µL product + 900 µL medium). For negative controls, no dilution is needed.
9. Under sterile conditions, pipette 100 µL diluted product onto a plate and spread. Keep the agar side down and lid side up.
   1. If spreading on the whole plate, use 7-10 glass beads. Carefully deposit the beads in an ethanol-filled beaker (dirty beads beaker).
   2. If spreading onto a section of a plate use a glass spreader or loop.
10. Keep the lid off and allow plates to dry under the flame for 5-10 minutes.
11. Close and wrap plates, and label appropriately. In addition to section labels, a plate label must contain:
    1. iGEM
    2. transformant genotype or shorthand e.g. pRep-HpCA. If a shorthand is used, make sure it is clearly commuicated in the corresponding lab notebook entry
    3. Antibiotic (and other additives)
    4. Date plated
    5. Initials
12. Dispose of culture products already inside tubes by closing them tightly and throwing them in the biohazard bin.

Objective

Author(s): Evan Wong, Yuki Guan, Matthew Chan

To identify the most efficient clones and optimal condition (pH, temperature, Ca²⁺⁺ bicarbonate) for CaCO₃ precipitation through carbonic anhydrase (CA).

• Phase I: Screen ALL candidates with a colorimetric assay (o-Cresolphthalein chromogenic reagent) ⇒ high-throughput
• Phase II: A gravimetric + XRD/SEM validation of the best-performing clones/conditions

PHASE I: High-Throughput Screening via Colorimetric Ca²⁺ Depletion

Materials Required:

• o-Cresolphthalein reagent

• Alkaline buffer (imidazole or Tris, pH ~10.5)

• Calcium calibrator (prepare 2.5 mM CaCl₂)

Calcium Assay Protocol Summary: (Following inoculation, induction, and CO₂ incubation)

1. Add samples (supernatant) and/or standards to wells
2. Add buffer, mix gently (no bubbles)
3. Add chromogenic reagent (o-Cresolphthalein reagent) and incubate at RT for 5 min
4. Read the OD absorbance at 570nm and 700nm
5. Analyze with microplate reader

Variable	Levels to Test on Each clone
pH	6.5, 7.0, 7.5, 8.0, 8.5
Temperature	20°C, 30°C, 37°C, 42°C
[CaCl₂]	5, 10, 25, 50 mM
[NaHCO₃]	10, 25, 50, 100 mM
Expression Time	24h, 48h, 72h (optional variable)

PHASE IIa: Gravimetric Validation and Mineral Characterization (Medium Throughput)

Materials Required:

• PBS
  • 750 mM phosphate buffer, pH 7 (tube)
  • 750 mM phosphate buffer, pH 8 (tube)
  • 30X 450 mM phosphate buffer, pH 7 (tube)
  • 30X 450 mM phosphate buffer, pH 8 (tube)

Gravimetric Analysis Protocol Summary: (Following identification of the top Calcium producing clones and conditions)

1. Harvest CaCO3 precipitate via centrifugation
2. Wash each precipitate with:

• 1 mL PBS (pH 8.0) → removes residual media and salts
• 1 mL distilled water → removes buffer salts

1. Drying precipitates at 60 °C overnight (12—16 h)
2. Determine each pellet’s mass
3. Store sample in desiccator if not actively in use

PHASE IIb: XRD+SEM Protocols: (Low Throughput)

SEM Protocol FEI Quanta 650 SEM Tools:

• Gloves
• Tweezers
• Small metal spatula
• SEM stub
• Stub sample box (as a stub holder)
• Double sided carbon or copper conductive tape
• Sputter Coater Sample prep:

1. Attach carbon tape to SEM stub
2. Scoop a small amount of powder and sprinkle a very THIN layer
3. Slightly spread sample on adhesive
4. Tilt and tap to get rid of loose particles
5. Load stub into sputter coater chamber and place on quartz crystal microbalance
6. Turn small black knob to get the stage rotating for better coverage
7. Vacuum the chamber and begin deposition roughly 2.0 nm thickness of coating is enough
8. Remove from sputter coater and place into microscope using gloves and tweezers

XRD:( prepping powdered sample, loaded sample holder)

1. Ground crystals into fine powder using a mortar and pestle

2. Take the top of the sample holder, orient it flat-side down and ensure that it clicks in place held by the 3 buttons

3. Fill well with sample and press down with pressing block

1. Remove excess with the edge of a glass slide or a brush

2. Take the bottom of the sample holder and press down until it clicks in place.

3. Flip entire sample holder over and click the button on the 2 slides to release.

Objective

Author(s): ZiQi Chen

Prepare Luria-Bertani (LB) medium for growth of E. coli strains.

Procedure

1. Fill up a glass media bottle with Milli-Q water to half its volume. Weigh out and add the the following reagents:

   Component	Amount	Final concentration
   LB	25g	25g/L
   Agar (for solid medium)	15g	15g/L
   Antibiotic	1mL	~1/1000th original conc.

2. Mix well to dissolve all components, using heat and a stir bar as needed. Top up to maximum capacity (500 or 1000 mL) with Milli-Q water.

3. Mix well to dissolve all solutes.

4. Loosely cap the bottle and apply autoclave tape.

5. Sterilize the media by autoclaving at LIQUID20.

6. After autoclaving, immediately tighten the cap. Note: If you notice that there is pressure build-up, loosen the cap and cool the bottle before re-tightening the cap.

7. Cool to room temperature before using/adding antibiotics.

To store, keep the bottle in a cool, dry place at room temperature.

Objective

Author(s): Ada Jiang

Measure optical density (ID) of bacterial culture.

Materials

• PYE media
• CB2A culture
• Cuvettes
• Spectrophotometer
• Parafilm

Protocol

1. Under sterile conditions, add 600 µL - 1 mL of fresh media (blank) and 600 µL - 1 mL of overnight culture into 2 clean cuvettes. Add parafilm to cover the cuvette containing the culture if needed.

2. Measure OD with spectrophotometer

   1. Turn on machine (takes 30 secs to turn on)
   2. Press Esc twice to get default screen
   3. In default screen: set nm> type in 600nm > enter
   4. Ensure cuvette is clean (use kimwipe)
   5. Place the cuvette into the carousel (make sure it is in correct orientation so that the light is passing through the clear ends of the cuvette)
   6. Measure blank (should be 0A)
   7. Measure sample (press 1)
   8. Do not need to turn off between samples

3. Rinse cuvettes with water (at least 3 times) and let them dry on a paper towel until they are ready to reuse.

Objective

Author(s): Matthew Chan

Extract, purify, and quantify plasmid DNA from bacterial cells. This protocol extracts more DNA than Miniprep.

Materials

• Culture
  • If high-copy: 25-50 mL
  • If low-copy: 50-100 mL
• Buffers
  • P1: kept cool (RNase added) (Resuspension Buffer)
  • P2: pre-warm to 37C (Lysis Buffer)
  • P3: pre-chill to 4C (Neutralization Buffer)
  • QBT
  • QC
  • QF
• Isopropanol
• 70% ethanol
• x2 sterile 50 mL round-bottom centrifuge tubes (can withstand high RPM) per prep
• x2 sterile 15mL high-performance centrifuge tube (>15,000g)
• Ice bucket
• 1x QIAGEN-tip 100 column

Procedure

Always inoculate cultures under sterile environment.

1. Pellet bacterial culture (in 50mL or 15mL conical tubes) by centrifuging for 15 minutes at 6000g. Refrigerate during centrifugation at 4C.
2. Resuspend in 4 mL buffer P1, vortex or pipette vigorously until no cell clumps remain. Transfer to a round-bottom centrifuge tube..
3. Add 4mL lysis buffer P2 and invert 4-6 times to mix. Incubate at room temp for 5 minutes.
   1. DO NOT LEAVE EXPOSED TO OPEN AIR, close the P2 bottle right away.
4. Add 4mL neutralization buffer P3 and invert 4-6 times to mix. incubate ON ICE for 15 minutes.
5. Centrifuge lysate at 20,000 g for 30 minutes at 4C.
   1. Ensure the mixture is in a round-bottom centrifuge tube; regular cone-bottom tubes will fail at 20,000 g
   2. Alternatively use QIAfilter kits or cartridges, but the tubes can be reused
6. Transfer supernatant to another 50mL round-bottom centrifuge tube. Centrifuge again for 15 minutes at 20,000 g at 4C.
7. Place QIAGEN-tip 100 column in a 15 mL tube and equilibriate/pre-load with 4 mL buffer QBT.
8. Transfer supernatant to the column and let it flow through by gravity.
9. Wash by adding 10 mL buffer QC, let it flow through by gravity. Discard flow-through. Do this twice.
10. Elute DNA with 5 mL buffer QF into 15mL centrifuge tube.
    1. Optionally pre-warm the buffer to 65C to increase yield.
    2. Can store elute at 4C to continue later
11. Add 3.5 mL room temperature isopropanol to precipitate DNA. Mix then centrifuge at 15,000g at 4C for 30 minutes. Carefully discard supernatant.
    1. a clear DNA pellet should be visible
12. Wash with 2 mL room temperature 70% ethanol. Mix and centrifuge at 15,000g for 10 minutes at 4C. Carefully discard supernatant without disturbing pellet.
    1. a clear DNA pellet should be visible
    2. when discarding the supernatant, consider grabbing a petri dish if the DNA pellet moves a lot > you can remove most of the supernatant and put the pellet on the petri dish
    3. use a P200 to remove large ethanol droplets
13. Air dry pellet for 5-10 min, redissolve in 500 uL ultrapure water or TE buffer.
14. Measure concentration NanoDrop Spectrophotometer
    1. 260/280 > 1.7
    2. 260/230 > 2

Objective

Author(s): Pattarin Blanchard

Adapted from Pronobis et al., 2016 supplement 1, this is a plasmid purification protocl that uses miniprep kits and columns and delivers midi/maxi-level DNA yields using a fraction of the reagents and time. Some sources say the name is short for Miracle Prep, but the first author’s first name is also Mira.

While the protocol describes using a large culture with miniprep columns, the key trick is the 1:1 ethanol addition to the lysate supernatant, which increases the DNA carrying capacity of the silica columns. Users report the ethanol addition getting them more than double the yield from a typical miniprep culture volume,.

The standard protocol is for 50 mL culture. See the toggle for a 10 mL version.

Materials

• 1x 50 mL centrifuge tube

• 1x 15 mL centrifuge tube

• Resuspension buffer, no RNase added

• RNase A, 10 mg/mL stock

• Miniprep kit components; lysis, neutralization, wash buffers

• 96% ethanol, molecular biology grade

• x1 2 mL microcentrifuge tube

• x9 1.5 mL microcentrifuge tubes

• x5 spin columns

• Ultrapure water

Equipment

• Low-speed centrifuge with refrigeration (preset to 4C)

• Microcentrifuge

• Pipettes

• Vortex

Procedure

1. Pellet a 50 mL overnight (12-16h) culture in a 50 mL centrifuge tube at 4000 g rcf at 4C for 10 minutes.
2. While waiting, prepare 2 mL resuspension buffer with RNase A to a working concentration of 50 ug/mL in a 2 mL tube.
   1. The Thermo Scientific RNase A stock in the -20 freezer is 10 mg/mL; use 10 uL of this stock
3. Discard supernatant and in the same tube, add prepared 2 mL resuspension buffer+RNase A to resuspend by vortexing.
4. Add 2 mL lysis buffer, invert 4 times and incubate for 3 minutes at room temperature.
5. Add 2 mL of neutralization buffer and invert 3-4 times.
6. Distribute the lysate into four 1.5 mL microcentrifuge tubes by pouring. Do not pipette to not disturb the lysate and debris.
7. Pellet debris at 13,200xg rcf for 10 minutes.
8. Collect supernatants into a 15 mL centrifuge tube and add a 1X volume of ethanol. Mix thoroughly by vortexing for 5 seconds.
9. Load the mix into 5 spin columns, 700 uL at a time. Spin for 30s at 13,200xg rcf and discard flow-through.
10. Repeat step 9 until all of the sample has been spun through.
11. Wash columns by adding 500 uL wash solution per column, spin at 13,200xg rcf for 30 seconds, and discard flow-through. Do this twice.
12. Dry the columns by spinning at 13,200xg for 1 minute 30 seconds.
13. Place the columns into 1.5 mL microcentrifuge tubes. For each column, add 30 uL ultrapure water to the centre, and wait 2 minutes.
14. Spin the columns at 13,200xg rcf for 2 minutes to elute DNA.
15. Combine the elutes into one tube for a final volume of 150-175 uL.
16. Measure DNA concentration, see NanoDrop Spectrophotometer.

10 mL version

Materials

• 2x 15 mL centrifuge tubes

• Resuspension buffer, no RNase added

• RNase A, 10 mg/mL stock

• Miniprep kit components; lysis, neutralization, wash buffers

• 96% ethanol, molecular biology grade

• x3 1.5 mL microcentrifuge tubes

• x2 spin columns

• Ultrapure water

Equipment

• Low-speed centrifuge with refrigeration (preset to 4C)

• Microcentrifuge

• Pipettes

• Vortex

Procedure

1. Pellet a 10 mL overnight (12-16h) culture in a 15 mL centrifuge tube at 4000xg rcf at 4C for 10 minutes.
2. While waiting, prepare 400 uL resuspension buffer with 2 uL RNase A for a working concentration of 50 ug/mL in a 1.5 mL tube.
3. Discard supernatant and in the same tube, add prepared 400 uL resuspension buffer+RNase A to resuspend by vortexing.
4. Add 400 uL lysis buffer, invert 4 times and incubate for 3 minutes at room temperature.
5. Add 400 uL of neutralization buffer and invert 3-4 times.
6. Pellet debris at 13,200xg rcf for 10 minutes. Transfer supernatant to a new 15 mL centrifuge tube.
7. Add a 1X volume of ethanol and mix thoroughly by vortexing for 5 seconds.
8. Load the mix into 2 spin columns, 600 uL at a time. Spin for 30s at 13,200xg rcf and discard flow-through.
9. Repeat step 8 until all of the mix has been spun through.
10. Wash columns by adding 500 uL wash solution per column, spin at 13,200xg rcf for 30 seconds, and discard flow-through. Do this twice.
11. Dry the columns by spinning at 13,200xg for 1 minute 30 seconds.
12. Place the columns into 1.5 mL microcentrifuge tubes. For each column, add 30 uL ultrapure water to the centre, and wait 2 minutes.
13. Spin the columns at 13,200xg rcf for 2 minutes to elute DNA.
14. Combine the elutes into one tube for a final volume of ~60 uL.
15. Measure DNA concentration, see NanoDrop Spectrophotometer.

Objective

Author(s): Alya Kaynak, Matthew Chan, ZiQi Chen

Preparation of competent E. coli cells.

Procedure

CCMB80 Preparation (100 mL)

Chemical	Final Molarity	Weight/Volume Needed
KOAc	10 mM	0.098 g
CaCl₂·2H₂O	80 mM	1.18 g
MnCl₂·4H₂O	20 mM	0.4 g
MgCl₂·6H₂O	10 mM	0.2 g
Glycerol*	10%	10 mL
Water		top-off until 100 mL

• for glycerol use pipette aid

1. weigh + add chemicals into 250 mL erlenmeyer flask
2. top-off with water, everything should dissolved with aggressive swirling of the erlenmeyer flask
3. syringe or vacuum filter it under a flame, store at 4C

SOB Media Preparation (250 mL)

Chemical	Final Molarity	Weight/Volume Needed
Yeast Extract	10 mM	1.25 g
Tryptone	80 mM	5 g
NaCl	20 mM	0.15 g
KCl	10 mM	0.05 g
MgSO₄	10%	0.6 g
Water		top-off until 250 mL

1. weigh + add chemicals into 250 mL erlenmeyer flask
2. top-off with water, everything should dissolved with aggressive swirling of the erlenmeyer flask
3. autoclave media before use

Procedure

Set-up

Day 0 - Plate Colony

1. plate desired E. coli strain to become competent

Day 1 - Mini-Culture

1. pick a colony and place it in 3 mL of SOB Media Day 2 - Culture set up

2. inoculate 1 mL of mini-culture into 250 mL of SOB

3. grow at 20°C until OD600 = 0.3

Pre-chill equipment needed the next day (ie tube + pipettes)

Making Competent Cells

Set the Av-Gay centrifuge to 4°C + close the lid so that it can reach the correct temperature

1. harvest cells by centrifugation at 3000 RPM and 4°C for 10 minutes
   1. split 250 mL into 6 50 mL tubes of ~ 42 mL
   2. pellet will be suprisingly small !
2. pour off the supernatant in one smooth movement, remove residual liquid with a pipette
3. gently resuspend the cells in 80 mL of ice cold CCMB80 buffer, incubate the resuspended cells on ice for 20 minutes
   1. swirl the cells in the buffer rather than shaking or pipetting vigorously, only pipette on the sides
   2. there will be 6 tubes in total, split them into 2 group
      1. within each group resuspend 1 of the 3 tubes with 10 mL then use the same 10 mL to resuspend + gather cells from 3 tubes into 1
      2. next use another 10 mL to wash the 3 tubes into the same final tube (there will be 2 main tubes with 20 mL each)
      3. combine the 2 20 mL, homogenize by mixing, split the 40 mL back into 20 mL
      4. top off each tube so it has a final 20 mL each
4. centrifuge at 3000 RPM and 4°C for 10 minutes and discard supernatant
5. resuspend cells in 10 mL of ice cold CCMB80 buffer
   1. resuspend + combine with 5 mL, then use other 5 mL for washing
6. test the oD600 of a mixture of 200 μl SOB and 50 μl of the resuspended cells
   1. add chilled CCMB80 to the cell slurry to yield a final OD of 1 .0 - 1 .5 in this test
7. aliquot into microcentrifuge tubes and freeze at -80ºC.

Objective

Author(s): ZiQi Chen

Purify DNA fragments for downstream cloning. This protocol is based on the Monarch® Spin PCR & DNA Cleanup Kit.

Materials

Item	Clarification
BZ	Binding
WZ	Wash buffer
EY	Elution buffer
Column	Top part of column assembly
Collection tube	Bottom part of column assembly

If kit is unopened, prepare buffers by following instructions on website:

Required materials:

• Monarch BZ buffer (large bottle)
• Monarch WZ buffer (medium bottle)
• Monarch EY buffer (tiny bottle)
• P200 and P20 pipette and tips

Procedure

BIND: Binding buffer precipitates DNA

1. Transfer DNA to a “mixing” tube. Adjust PCR results to 20 uL if below that. Mix 5:1 parts of 5 Monarch buffer BZ to 1 DNA PCR results. Mix by flicking or pipetting, DO NOT VORTEX, ensure DNA is suspended for pouring. For example:

   Name	Vol uL
   PCR DNA	20
   BZ buffer	100 (5*20)
   Total	120

2. Assemble the column and collection tube (provided in bags) into the full assembly. Pour in your PCR result and BZ buffer mix onto the column (top part). Spin for 1 min at 16,000g (~13000 rpm) then discard the FLOW-THROUGH (liquid) collected in the bottom part.

WASH: Clean bound DNA of impurities

1. Re-insert the column onto the collection tube. Wash by adding 200 uL of WZ wash buffer and spin at 16,000g for 1 minute.
2. Discard flow-through and repeat step 3.

ELUTE: Dissolve DNA buffer and collect.

1. Transfer the column (top part) onto a clean 1.5mL microfuge (Eppendorf) tube. This should be labelled “postwash” “washed” or similar for identification.

2. Add 5-20 uL of EY elution buffer to the center of the matrix (sieve part). If the DNA does not appear to be getting extracted, add more at expense of concentration. More elution buffer may increase yield.

3. Wait for 1 minute then spin at 16,000g. Remove column and collect flow-through for further use, this contains your DNA. Remember not to discard the column too early.

Objective

Author(s): Matthew Chan, Ada Jiang

Quantify amount of extracted and purified plasmid DNA from bacterial cells using NanoDrop spectrophotometer.

Materials

• NanoDrop 2000/2000c Spectrophotometer
• Eluted Samples
• Elution Buffer / Water as blank

Procedure

Protocol adapted from: NanoDrop 2000/2000c Spectrophotometer

DNA yield is determined by measuring the absorbance at 260 nm. The ratio of the readings at 260 nm and 280 nm. (A260/A280) provides an estimate of the purity of DNA with respect to UV absorbing contaminants such as protein.

Set-up Machine/Software

1. start up and log into the computer.

   1. enter our username and password (on sticky note by our bench)

      username	ajiang
      password	ubcigem$2025

   2. log on to MI, click on nd-1000 v3.8.1 program

2. in the nd-1000 v3.8.1 program, click User default > Nucleic acid > DNA-50
3. clean pedestal with sterile water or dH2O
   1. load in 1 ul of water sample > click initialize
   2. load 1 ul of blank (liquid used to elute DNA), then click BLANK

Take measurements

1. enter in sample ID

2. take a reading of your plasmid using 1.0 μl of your plasmid DNA. Record the results in a table like so:

   Construct Name	Concentration (ng/ul)	A260/A280	A260/A230

3. exit and close program

4. clean pedestal and gently lower the nanodrop arm onto clean folded kip wipe.

5. log off > turn off computer

Objective

Author(s): Jessie Luo, Ada Jiang

Prepare Peptone Yeast Extract (PYE) nutrient medium selectively culture Caulobacter crescentus. This protocol describes the preparation of 500 - 1000 mL of medium, adapted from both DSMZ 1462: CAULOBACTER PYE MEDIUM and a protocol provided by our advisor, Beth.

Procedure

Follow the same procedure as LB Medium Recipe, using the following reagents instead:

Version 1 (from standard)

Component	Amount (for 500 mL)	Final concentration
Peptone	1 g	2 g/L
Yeast extract	0.5 g	1 g/L
CaCa₂·2H₂O (Calcium chloride dihydrate)	0.05 g	0.1 g/L
MgSO₄·2H₂O (Magnesium sulfate heptahydrate)	0.1 g	0.2 g/L
Optional: Agar (for solid medium)	6 g	1.2%

Version 2 (from manuscript)

0.3% peptone, 0.2% yeast extract, 1.5 mM $MgSO_4$, 1 mM $CaCl_2$, 1.2% agar for solid medium

Component	Amount (for 1000 mL)	Final concentration
Peptone	3.00 g	3 g/L
Yeast extract	2.00 g	2 g/L
CaCa₂·2H₂O (Calcium chloride dihydrate)	219.08 mg	0.219 g/L or 1 mM
MgSO₄·2H₂O (Magnesium sulfate heptahydrate)	369.71 mg	0.370 g/L or 1.5 mM
Optional: Agar (for solid medium)	12.00 g	1.2%

Note that the concentration of agar can be increase to 1.5% for a denser gel.

Objective

Author(s): Matthew Chan, Ada Jiang

Extract, purify, and quantify plasmid DNA from bacterial cells. This 2-day procedure involves lysing bacterial cells, removal of impurities and plasmid purification techniques. The resulting purified DNA product can be quantified, sequenced and stored for further analysis or use in cloning experiments.

Materials

GeneJET Plasmid Miniprep Kit

• Resuspension Solution (Buffer 1)
• Lysis Solution (Buffer 2)
• Neutralization Solution (Buffer 3)
• Wash Solution
• RNase A
• Elution Buffer (10 mM Tris-HCL, pH 8.5)
• GeneJET Spin Columns
• Collection Tubes (2 mL)
• 1.5 mL or 2 mL Eppendorf tubes
• Molecular grade water (DNase/RNase-free deionized water)
• 96% Ethanol (only if you are opening a new kit)
• Culturing media, generally Luria Broth (LB)
• Antibiotic (if necessary)
• Shaker
• 37°C incubator/room

Procedure

Set-up

Day 0 - Transformation

1. Transform E. coli with desired construct

2. Streak on proper plate Day 1 - Culture set up

3. Pick colonies from E. coli bacterial colony plates

   Construct	# of colonies picked
   [insert names of plates]

4. Start a 1 - 5 mL culture and grow at 37°C for 12 - 16 hrs (add necessary antibiotics)

Miniprep

GeneJET / QIAGEN Plasmid Miniprep Kit Protocol

1. transfer 1.5 mL of bacterial culture using pipette
   1. into 1.5 mL microcentrifuge tube
   2. into 15 mL Falcon tube
2. pellet bacteria by centrifuging for 3 minutes @ 8000 RPM, decant supernatant
   1. can use pipette to remove supernatant if worried
3. resuspend in 250 µl resuspension Buffer P1
   1. transfer to microcentrifuge tube if you did 1. b. with 15 mL Falcon tube
4. add 250 µl lysis Buffer P2 to tube + invert 4-6 times gently to mix
   1. do not allow lysis reaction to occur for 5 minutes
   2. for small-medium pellets, can add Buffer N3 immediately
   3. for larger pellets, wait up to 5 minutes
5. add 350 µL neutralization Buffer N3 to tube + invert 4-6 times gently to mix
   1. allow neutralization process to take 9 minutes
6. centrifuge lysate for 1 minutes @ max. speed 13,000 RPM
   1. transfer the supernatant to a new microcentrifuge tube.
7. add 650 µL ethanol to the collected supernatant and pipette up and down to mix.
8. transfer 800 µL of the mix to a spin column and centrifuge Column for 1 minute, discard flow-through. Repeat until all of the mix has been spun through.
9. wash Column by adding 500 µL Buffer PB, centrifuge for 1 minute TWICE (13,000 RPM)
10. discard flow-through transfer to new tube, centrifuge for 2 minutes to dry
11. place Column in new 1.5 mL microcentrifuge tube
12. add 50 μL ULTRA PURE WATER, let stand for 1 min + centrifuge for 1 min
    1. Buffer EB can be used as well, however sometimes the reagents in the buffer can inhibit/interfere with downstream usage
    2. Pre-warm water to 50-60 °C for increased recovery
13. nanodrop final products
    1. 260/280 should be higher than 1.70
    2. 260/230 higher than 2.00
    3. A260 (absorbance) should be between 0.1 - 1.0 for a reliable reading. Anything higher indicates that the machine is not properly calibrated or the sample concentration is too high (need to be diluted).
14. store final product in -20°C

Nanodrop

See NanoDrop Spectrophotometer

Objective

Author(s) : Ada Jiang

Preparation of DNA samples for Nanopore sequencing.

Submitting samples

To receive results the next business day, complete these steps by 2:00 PM (LSI pickup time).

Prepare the samples

1. Place an order and print out the first page of the order confirmation.
   1. Draw dinosaur at the back of the page if you’d like!
2. If necessary, resuspend your DNA in nuclease-free water or elution buffer (10 mM Tris, pH 8.5).
   1. Avoid using buffer containing DMSO if possible.
   2. Do not include any primers with or mixed into your samples.
3. Measure your sample using the Nanodrop and verify that:
   1. The concentration is within the concentration range specified in your order submission.
   2. The quality of your sample is good - aim for intact circular double-stranded plasmids (usually extracted via plasmid prep).
   3. The purity of your sample is sufficient. The recommended ratios of absorbance are A260/280 > 1.8 and A260/230 between 2.0-2.2.
      1. If using Qiagen miniprep kit, use the optional PB buffer wash step to increase purity.
      2. If purity is poor, try performing gel extraction or reaction purification as an additional purification step.
4. If possible, use a high-precision instrument like a Qubit fluorometer to measure concentration. Nanodrop is not as precise and may lead to an inappropriate amount being submitted.
5. Aliquot enough volume of each sample into a strip of transparent, 200 uL PCR tubes in a contiguous manner (fill the the strip sequentially, don’t skip tubes).
   1. Ensure the caps are firmly closed.
   2. Wrap the tubes in a light layer of parafilm to keep the caps closed during shipping.

Package the samples

1. Label the tubes:

• Write the first 2 characters of your order ID on the 1st tube of each strip. (Can be found on the printed order confirmation).
• Write the respective sample number on each tube of the strip.

1. Place the tubes in a small protective vessel (like a falcon tube, small box or bubble wrap) to protect them during shipping.
2. Place the samples into a small bag with your order confirmation sheet (folded or cut out so that the QR code is visible).

For 96-well plate submission, see the detailed instructions for packaging on the plasmidsaurus site.

Deliver the samples

1. Go to Level B2 of LSI (take the East elevators).
2. Once you exit the elevator, turn around the corner, then walk straight until you see a table with boxes on it (next to LSI stores).
3. Find the dropbox for Plasmidsaurus orders and drop off our package.

Objective

Author(s): Ada Jiang

Amplify specific DNA sequences using PCR for downstream analysis or cloning.

Materials

Gather the following:

• Ice OR Pre-chilled PCR tube rack

• PCR tubes

• Pipettes and tips

• Labelling tools Keep the reaction components on ice:

• Primer(s)

• Template DNA sample

• Molecular grade/Nuclease-free water

• Q5 High-Fidelity 2X Master Mix Precipitates in the Q5 Master Mix might appear in the first few freeze/thaw cycles. To ensure optimal performance, allow the reagent to fully thaw and resuspend it prior to use.

Equipment

• Thermocycler

Procedure

1. Use the NEB Tm Calculator to calculate the required annealing temperature for your PCR program. Follow the instructions at the top of the page.

2. Turn on and set up the thermocycler program to allow the machine enough time to reach the desired temperature of the first step prior to adding your reaction samples. Set up the program according to the NEB recommended protocol for your reaction reagent/kit. The duration for the Cycles steps depends on several factors, including the size of your DNA template.

   STEP		TEMP (°C)	TIME
   Initial Denaturation		98	30 seconds
   25 - 35 Cycles	Denaturation	98	5 - 10 seconds
   	Annealing	50-72	10-30 seconds
   	Extension	72	20 - 30 seconds/kb
   Final Extension		72	2 minutes
   Hold		4-10	Infinity

3. Store completed samples on ice or at -20

Objective

Author(s): Jessie Luo, Ada Jiang

Quantify amount of extracted and purified plasmid DNA from bacterial cells using Qubit fluorometer and Invitrogen DNA assay kits.

Materials

• Qubit assay tubes
• Qubit dsDNA BR / HS buffer (stored at room temperature)
• Standard #1 (stored in 4˚C), keep on ice
• Standard #2 (stored in 4˚C), keep on ice
• Dye: Qubit dsDNA BR / HS reagent, 200x (stored in 4˚C)
• DNA samples

Procedure

Prepare solutions:

1. Choose the correct assay kit. Select HS (High Sensitivity) or BR (Broad Range) depending on expected DNA concentration. Retrieve the appropriate standards from 4°C.

2. Label Qubit assay tube lids for the standards and samples.

3. In an eppendorf tube, prepare the working solution:

   1. Mix buffer with dye so that when the working solution is added to the standards and samples, the final ratio of solution:dye is 199:1.
   2. Prepare enough for all samples and standards. Each Qubit sample should be a final volume of 200 µL.
   3. You may need to prepare a second working solution at a different dilution for the samples ,depending on how much you want to dilute the DNA.

4. Prepare the standards: 10 µL of each standard + 190 µL of working solution

5. Prepare the samples: Add up to 1-10 µL of DNA sample in each tube. Bring the final volume to 200 µL. with working solution.

6. Mix by gentle pipetting, avoiding bubbles. Spin down.

7. Incubate tubes at room temperature for 10 mins.

Measurment

1. On the Qubit fluorometer, select: DNA → dsDNA → (HS or BR depending on kit)
2. Insert Standard 1 and Standard 2 as prompted by the machine for calibration.
3. Measure each sample in its assay tube.
4. For each sample, input the volume of DNA used.
5. Record results.

Objective

Author(s): Ada Jiang

Cut DNA fragments at specific restriction sites in preparation for analysis or downstream cloning. This protocol is based off of the NEB Restriction Digest Protocol.

Materials

• Digestion enzymes
• 10X rCutSmart Buffer (pr r1.1, r2.1, or r3.1 buffer for optimal enzyme activity)
• Molecular grade water
• Ice
• PCR tubes and rack
• Thermocycler

Procedure

1. Find your protocol by inputting the desired enzymes into NEBcloner 1.16.3 Restriction Enzyme Single/Double Digestion calculator. This will generate a custom protocol for a 50 µL reaction. All components can be reduced for a 25 µL reaction. Generally, digests require the following components:

   Component	50 uL Reaction	Final Conc/Amount
   DNA	200-1 ug	n/a
   10x rCutSmart Buffer	5 µL	1X
   Enzyme 1	1 µL	20 units
   Enzyme 2	1 µL	20 units
   Nuclease-free Water	to 50 µl	n/a

   Table 1. General double digest protocol from NEB *5 - 10 units of enzyme per 1000ng of DNA is recommended (most enzymes are 20 units per 1 uL)

2. Preheat thermocycler to 37°C.

3. Set up the reaction (keeping everything on ice).

4. Mix components by pipetting the reaction mixture up and down, or by “flicking” the reaction tube.

5. Quick (“touch”) spin-down in a microcentrifuge. Do not vortex the reaction.

6. Incubate at 37°C in the thermocycler for at least 15 minutes (for time-saver enzymes), up to 1 hr if needed.

7. If appropriate for your enzyme, deactivate the reaction by incubating at 60°C or 80°C for 20 minutes.

8. Transfer the tubes to ice. Store at -20°C if agarose gel/downstream analysis will be run in a separate lab session.

Objective

Author(s): Pattarin Blanchard

Resuspend DNA oligos and fragments for use in downstream cloning experiments.

Resources

• IDT:

• Twist:

• GenScript:

Materials

• RNase-free 1x TE buffer

• Pipette tips

• Centrifuge

• PCR or microcentrifuge tubes

• Molecular water

Procedure

Resuspension into storage stocks

1. Briefly centrifuge plates at 2000 RPM for 5 seconds (setting is not too important) to make sure pellets are at the bottom of wells.
2. Calculate how much TE buffer should be added to each well to properly resuspend plates. Add TE buffer accordingly and pipette up and down to resuspend.
   1. Primers (if in pairs, this applies for each individual one) should be resuspended to 100 uM for a storage/master stock.
   2. Fragments, amount depends on yield/manufacturer
      1. Twist normalizes yield to 1000 ng, resuspend in 20 uL TE buffer for a storage concentration of 50 ng/uL
      2. Genscript normalizes yield to 500 ng, resuspend in 10 uL TE buffer for a storage concentration of 50 ng/uL.
      3. IDT depends on fragment length
         1. For fragments up to 750 bp, resuspend in 10 uL TE buffer for a storage concentration of 25-50 ng/uL
         2. For fragments above 750 bp, resuspend in 20 uL TE buffer for a storage concentration of 50 ng/uL
   3. Note: a relatively high stock concentration is desired to minimize freeze-thaw cycles on the storage stocks.

Making working stocks

1. For each fragment desired, calculate how much storage stock and water are needed to make a diluted aliquot (working stock) such that it is a convenient volume to work with.
   1. Both Golden Gate and Gibson Assemblies typically require 0.05 pmol per fragment, choose a concentration that allows 0.5-1 uL of working stock to be used each time. Keep in mind a typical reaction volume is 15 uL for GGA, 20 for Gibson.
   2. Aim for a volume such that the working stock does not go through more than 10 freeze-thaw cycles to avoid DNA degradation.
2. In PCR tubes or microcentrifuge tubes, use molecular water to make aliquots, and label appropriately.

Objective

Author(s): Pattarin Blanchard

Resuspend and amplify parts from iGEM distribution kits. Adapted from the various distribution handbooks published by iGEM. Note that each part comes as a plasmid, not linear fragments.

A. Locate desired parts on a plate

1. Use the following platemaps to identify which plate and well contains a desired part.

Year	Links
2025	All parts: 2025 platemap

Cloning vectors only: 2025 cloning vectors | 2024 | All parts: 2024 platemap Cloning vectors only: 2024 cloning vectors | 2023 | All parts: 2023 platemap | 2022 | plate 1: https://parts.igem.org/assembly/plates.cgi?id=6437 plate 2: | 2021 | https://parts.igem.org/Help:2021_DNA_Distribution | 2019 | https://parts.igem.org/Help:2019_DNA_Distribution

1. Make a note of the following properties. Note you should already know what enzymes and fusion sites are used with each part.

• Resistance marker
• Kit plate (if applicable)
• Well

1. For good practice, also save the full plasmid sequences of each part to be retrieved on SnapGene to simulate assembly.

B. Gather materials

Materials

• Competent E. coli cells, kept on ice

• Molecular water

• Two LB agar plates with appropriate antibiotic from step A2 per part retrieved

• Sterile spreader or glass beads

• LB or SOC medium

• Floating foam tube rack

• Microcentrifuge tubes

• Desired distribution kit plate

• One culture tube or flask per part retrieved

Equipment

• Water bath at 42C

• Microcentrifuge

• Bunsen burner

• Timer

• Ice bucket

C. Resuspend DNA

If this is the first time a part is being retrieved, follow these steps. Otherwise, simply thaw the distribution kit and take 1 uL of resuspension.

1. Mark a desired well on the distribution kit with a permanent marker. A plate is oriented like so:

2. Use a pipette tip to carefully pierce the well and discard the tip. Do not peel off the foil cover.

3. Pipette 10 uL of molecular-grade water into a well and pipette up and down several times to resuspend, then let it sit for ~5 minutes. The resuspension should look red because it has cresol red dye.

4. Place the plate(s) in the -20C freezer with the lid closed. There is no need to wrap the plates. The resulting resuspension has a concentration of 100-200 pg/uL DNA.

D. Transform into cells

1. Label and pre-chill centrifuge tubes on ice. Ensure the water bath is ready at 42C.
2. Pipette 50 uL of competent cells per tube and keep on ice.
3. Pipette 1 uL of reuspended DNA into its corresponding centrifuge tube with comp cells. Close tubes and flick gently to mix, but do not vortex.
4. Incubate tubes on ice for 30 mins.
5. Using a floating tube rack, heat shock tubes in the 42C water bath for 1 min. Rest tubes on ice for 5 mins.
6. Pipette 200 uL LB or SOC medium (no AB needed) into each tube.
7. Incubate tubes in the 37C room for 2 hours, preferably on a shaker or rotor.
8. Pipette 20 uL onto a corresponding plate and spread immediately with a sterile loop or glass beads. Label the plate with appropriate information, including the inocculation volume.
9. Repeat step D8 on a second plate, this time with 200 uL.
10. Incubate plates at 37C overnight and continue the following day with liquid culture, to prep DNA or make glycerol stocks.

Objective

Author(s): Pattarin Blanchard

Prepare nutrient-rich Super Optimal Broth (SOB) medium for culture of E. coli. This protocol is adapted from NovoPro Bioscience Inc’s SOB recipe.

This recipe has the same components as SOC Medium Recipe, but without glucose.

Procedure

1. Follow the same procedure as LB Medium Recipe, using the following reagents instead:

   Component	Amount (g)	Final Conc (g/L)
   Tryptone	20	40
   Yeast extract	5	10
   NaCl (Sodium chloride anhydrous)	0.5	1
   KCl (Potassium chloride anhydrous)	0.186	0.372
   MgSO₄ (Magnesium sulfate anhydrous)	1.2	2.4

   *Note that MgCl₂ has been substituted with MgSO₄ in this version to avoid having to add MgCl₂ solution after autoclaving

2. Top up with Milli-Q water and mix.

3. Optionally adjust pH to 7 using NaOH pellets or solution.

4. Sterilize by autoclaving.

Objective

Author(s): Pattarin Blanchard, Matthew Chan

Prepare a nutrient-rich Super Optimal broth with Catabolite repression (SOC) medium for improved recovery rate in bacterial cell transformation procedures.

Protocol

Follow the same procedure as LB Medium Recipe, using the following reagents instead:

For 100 mL SOC medium:

Component	Amount	Notes	Final Conc.
Tryptone	2 g	On our bench	20g/L
Yeast extract	0.5 g	On our bench	5g/L
Sodium chloride anhydrous	0.05 g	Hallam Chemical Cabinet	0.5g/L
Potassium chloride anhydrous	0.018 g	Hallam Chemical Cabinet	0.18g/L
D-Glucose	4 g	Should be 3.6 g, but accounting for degradation under heat

Hallam Chemical Cabinet | 40g/L | Magnesium sulfate anhydrous | 0.24 g | Hallam Chemical Cabinet | 2.4g/L | Milli-Q water | Fill to 100 mL | Start with some water first to dissolve components then top up | N/A

For 500 mL SOC medium:

Component	Amount	Notes	Final Conc.
Tryptone	10 g	On our bench	20g/L
Yeast extract	2.5 g	On our bench	5g/L
Sodium chloride anhydrous	0.25 g	Hallam Chemical Cabinet	0.5g/L
Potassium chloride anhydrous	0.09 g	Hallam Chemical Cabinet	0.18g/L
D-Glucose	20 g	Should be 3.6 g, but accounting for degradation under heat

Hallam Chemical Cabinet | 40g/L | Magnesium sulfate anhydrous | 1.2 g | Hallam Chemical Cabinet | 2.4g/L | Milli-Q water | Fill to 500 mL | Start with some water first to dissolve components then top up | N/A

After mixing and ensuring powders have fully dissolved, autoclave the SOC medium (LIQUID20)

• Do not fully close the lid Store at 4°C for long term storage

Objective

Author(s): Matthew Chan

To quantify the number of viable bacterial cells in a sample by performing serial dilutions, plating them on agar media, and using a spectrophotometer to estimate bacterial concentration based on optical density.

Materials

For 1 Bacterial Strain:

• Flask of UTEX or CB2A culture
• 7 sterile tubes w/ 9 ml of media each
• 14 LB agar plates
• Micropipettes and sterile tips
• Ice
• Glass spreader, Petri dish, ethanol
• Spectrophotometer
• Disposable plastic cuvettes

Procedure

Making Serial Dilution Standards

1. label six sterile tubes of media with 1/10, 1/100, 1/1000, etc., as per Table 1.

2. vortex the culture and transfer 1 ml of the undiluted sample to tube 1.

3. using a sterile pipette tip, transfer 1 ml from tube 1 to tube 2, and repeat for all tubes.

4. Vortex-ing each tube before transferring

5. store the tubes on ice to prevent growth Table 1 — Decimal Dilution Series:

   Tube Label	Starting Volume (ml)	Volume from Previous Tube (ml)	Total Dilution
   1	9	1	1/10
   2	9	1	1/100
   3	9	1	1/1,000
   4	9	1	1/10,000
   5	9	1	1/100,000
   6	9	1	1/1,000,000
   7	9	1	1/10,000,000

Plating Cells

1. pipette 100 ul from each dilution tube onto an agar plate.
2. sterilize a glass spreader with ethanol and flame before use or use plastic spreader
3. spread the sample evenly across the agar surface, then allow it to dry.
4. repeat for all dilutions in duplicate, then incubate the plates upside-down

Optical Density Measurement

1. measure OD of the undiluted, 1/10, 1/100, and 1/1000 dilution tubes (OD600 - CauloColi + OD750 - UTEX)
2. vortex then transfer 2 mL to cuvette
3. set the wavelength to the desired 600 nm
4. blank the machine with a media
5. record the data

Objective

Author(s): Pattarin Blanchard

Produce a recombinant strain of E. coli that appears as purple colonies. Do so by using 2024 Distribution Kit parts and Golden Gate Assembly to produce a construct. This multi-day experiment involves transformation, colony picking, plasmid purification, stock preparation, and assembly.

Construct Design

|GGAG| A A[Promoter: BBa_J23100] --> |TACT|B B[RBS: BBa_B0034_m1] --> |AATG|C C[CDS: tsPurple] --> |GCTT|D D[Terminator: BBa_B0015] --> |CGCT|Y -->

Backbone: pJUMP28-1A

If you are in the UTEX group, hopefully some of these parts ring a bell to you. We use the BBa_B0034 RBS and BBa_B0015 terminator in our constructs. BBa_J23100 is a strong promoter from the same family of Anderson promoters we use.

Why the BBa_B0034_m1 RBS? The m1 suffix indicates this variant of B0034 has a short spacer after the RBS sequence. According to the registry, generally RBSs require 6-7 base pairs for optimal translation efficiency, so the m1 variant is used over the minimal B0034 part, also found in the kit. Use this as a rule of thumb unless explicitly indicated in a part’s documentation.

Since these are from the distribution kit, they use the golden gate affixes prescribed in iGEM’s Type IIS standard, which allows them to be assembled in the desired promoter-RBS-CDS-terminator order. Parts in the distribution kit come in vectors so a minimal amount is shipped to us, meaning we are responsible for amplifying the parts prior to use.

TsPurple (short for tinselPurple) is a chromoprotein, meaning unlike fluorescent proteins like GFP, RFP, etc., it is visible without needing UV light. This makes chromoproteins See this page for more commonly-used chromoproteins and reporters in iGEM: .

Day 0: Simulating the assembly

pJUMP28-1A(sfGFP) can be downloaded from the Oligonucleotide/Gene Database.

For the rest, obtain plasmid sequences from the 2024 kit platemap: https://airtable.com/appgWgf6EPX5gpnNU/shrb0c8oYTgpZDRgH/tblNqHsHbNNQP2HCX

• Each of these should automatically be annotated, but for tsPurple you may want to manually label the translated feature yourself. Simulate Golden Gate Assembly using BsaI. The construct should look like this:

Day 1: Resuspension and transformation

Materials

• Competent E. coli cells, kept on ice

• Molecular water

• 8x Cm LB agar plates

• 2x Kan LB agar plates

• Sterile loop

• Floating foam tube rack

• 5x sterile microcentrifuge tubes

• LB medium, 1.5 mL aliquot

Equipment

• Water bath at 42C

• Bunsen burner

• Timer

• Ice bucket

The parts/plasmids can be retrieved from the following wells from the 2024 kit.

Name	ID	Plate	Well	Ab resistance	LABEL
Promoter	BBa_J23100	1	A5	Cm	1
RBS	BBa_B0034_m1 (BBa_J428038)	1	I19	Cm	2
tsPurple	BBa_K1033906	1	G1	Cm	3
terminator	BBa_B0015	1	C1	Cm	4
pJUMP28-1A	BBa_J428353	1	A10	Kan	0
pDest		2	G7	Amp	a

Preparation

The following take some time so they should be the first things you do, in addition to setting up your bench space.

1. Place agar plates in the BSC with the agar side on top and slightly off to dry and warm to room temperature
2. Place competent cells and empty microcentrifuge tubes on ice
3. Turn on the water bath and set the temperature to 42C.

Resuspending DNA

If this is the first time a part is being retrieved, follow these steps. Otherwise, simply thaw the distribution kit and take 1 uL of resuspension.

Reference protocol: Retrieving Parts from Distribution Kits

Transformation

Follow Heat Shock Transformation with a Water Bath.

For step 3, use 1 uL of resuspended DNA into its corresponding centrifuge tube with comp cells.

For step 6, use 200 uL LB medium (no AB needed) into each tube.

For step 7, incubate tubes in the 37°C room for 1 hour.

Note: The iGEM protocol for distribution kit parts says 2 hours incubation, but the time tends to vary between protocols. According to addgene, this step is to allow bacteria to produce Ab resistance proteins in advance.

For step 9, pipette 20 uL onto a corresponding plate and spread immediately with a sterile loop or glass beads. Label the plate with appropriate information, including the inocculation volume.

Note: The recommended amount varies by protocol and person. This is from the iGEM protocol for distribution kit parts.

Repeat step 9 on a second plate, this time with 200 uL. Having one plate with little and another with a lot maximizes the chance of getting a colony.

Incubate plates at 37C overnight.

*In a typical transformation experiment you would have four or more plates for positive and negative controls e.g. cell only, cell+backbone, no cell, cell+construct, but we will only do it for Day 3 because we already know these vectors in Day 1 should work.

Day 2: Colony picking

Materials

• 5x sterile glass or disposable culture tubes
• LB medium
• 1000x Cm and Kan stock
• Sterile 20/200uL pipette tips
• 25 mL serological pipette

Procedure

1. Thaw Cm and Kan stock on ice. The following steps should be done aseptically:

2. Add 4mL LB medium to each culture tube. Label as appropriate.

3. Once thawed, add 4 uL Ab stock to the corresponding tubes. From the table above, 1-4 should get Cm, 0 should get Kan.

4. For each plate, use a marker to circle the colony to be picked. > Note: Ampicillin (Amp) when used as a resistance marker tends to form satellite colonies that appear to have resistance but are just piggybacking off of a resistant colony, therefore when using Amp, pick the large colony in the middle instead of small ones around a larger one. Cm, Kan, and Spec do not exhibit this behavior so we do not need to worry about it.

5. For each plate, use a sterile pipette tip to gently touch the desired colony, and drop that pipette tip into the culture tube. Do not poke into the agar.

6. Bring the culture tubes over to the 37C room in a tube rack and place on a shaker for overnight culture. Continue the following day. # Day 3: Plasmid purification

Materials

• GeneJET miniprep kit
  • Resuspension Solution (Buffer P1)
  • Lysis Solution (Buffer P2)
  • Neutralization Solution (Buffer N3)
  • Wash Solution (Buffer PB)
  • RNase A
  • Elution Buffer (Buffer EB; 10 mM Tris-HCL, pH 8.5)
• 10x microcentrifuge tubes
• 5x collection tubes+spin columns (close the bag tightly afterward)

Miniprep & Quantification

1. Pre-label tubes. Each culture to be miniprepped gets two microcentrifuge tubes and a collection tube + spin column. Arrange them on a tube rack like so: ------microcentrifuge tubes (culture)------ ------empty------ ------collection tubes+spin columns------ ------empty------ ------microcentrifuge tubes (product)------

2. Proceed with Plasmid Mini-Preparation

3. Measure DNA concentration using NanoDrop Spectrophotometer.

Day 4: Golden Gate Assembly and Transformation

Golden Gate Assembly

Materials

• NEBridge ligase master mix
• DNA fragments (miniprepped DNA from the day prior) stored on ice
• Molecular-grade water
• BsaI stock (BsaI-HFv2), thaw on ice
• 1 PCR tube per replicate
• ice bucket (multiple if many people doing it)
• Thermocycler

Procedure

Reference protocol: Golden Gate Assembly with NEBridge Ligase Master Mix

1. Calculate how much each fragment is required to get 0.05 pmol, the sequence length of each fragment is tabulated below:

   Name	Label	Length (bp)
   Promoter	1	2424
   RBS	2	2411
   tsPurple	3	2736
   terminator	4	2520
   pJUMP28-1A	0	3359

2. Add the components according to the reference protocol.

3. For a 5-fragment assembly (like in this training session, including the backbone), follow the reference protocol for 3-6 fragment assembly.

4. Chill on ice once reaction is complete. Proceed to transformation.

Transformation

Materials

• LB medium
• Competent cell stock (start thawing in the middle of the GGA incubation cycle)
• Foam tube rack
• Water bath at 42C
• Glass beads
• Sterile microcentrifuge tube
• 4 Kan LB agar plates + each additional transformation replicate

Procedure

This time we will make proper controls for our transformation experiment, so there will be 4 plates per replicate:

cell+construct | cell+backbone

(positive control)
cell only
(negative control 1)	medium

(negative control 2)

1. Label and pre-chill centrifuge tube(s) on ice. Ensure the water bath is ready at 42C.
2. Make 1 mL aliquot of LB medium in a sterile microcentrifuge tube.
3. Pipette 50 uL of competent cells per tube and keep on ice.
4. Pipette 2 uL of GGA construct (or 1 uL pJUMP28-1A, part 0) into a centrifuge tube with comp cells. Close tubes and flick gently to mix, but do not vortex.
5. Incubate tubes on ice for 20 mins.
6. Using a floating tube rack, heat shock tubes in the 42C water bath for 45s, then rest tubes on ice for 3 mins.
7. Pipette 200 uL LB medium (no AB needed) into each tube.
8. Incubate tubes in the 37C room for 45 mins, preferably on a shaker or rotor.
9. Pipette 50 uL onto a plate and spread with 7-10 glass beads by moving the plate around horizontally (try not to let the beads touch the lid), then carefully deposit the beads in an ethanol-filled beaker (dirty beads beaker). Label the plate with appropriate information.
10. Stack the plates and tape, bring to 37C room to incubate plates overnight.
11. Close the microcentrifuge tubes well and dispose of them in the biohazard bin.

Day 5: done!

Upload pics of plates here so we have a running record.

Objective

Author(s): Alya Kaynak

Introduce plasmid DNA into UTEX via triparental conjugation. These protocols are adapted from Yu et al., 2015 and iGEM team Toulouse_INSA-UPS 2021.

Protocol 1 (Yu et al., 2015)

1. Transform shuttle/suicide vectors carrying the CA of interest into competent DH5α E. coli cells that already contained the pRL623 helper plasmid to form cargo strains.
2. Wash 100 μl overnight cultures of the conjugal strain (pRL443) and 100 μl overnight cultures of the cargo strain with distilled water and mix with pre washed UTEX 2973 cells (200 μl at OD730 ~0.4-0.6).
3. Plate mixed cells onto BG11+5% LB (v/v) agar plates containing selective antibiotics.
   1. Place plates under bright continuous illumination (200 μmole photons·m−2·s−1). Mutant colonies were usually apparent within 3—4 days

Protocol 2 (iGEM Toulouse)

1. Growth of the cyanobacterial culture (day 1)

1. Set up a fresh culture of S. elongatus UTEX 2973 by inoculating a 100 mL conical flask of fresh BG11 medium (50 mL) with cells sourced from an axenic BG11 agar plate. Grow S. elongatus UTEX 2973 at 40°C, 300 μmol photons m2 s1 at 100 rpm. Grow cultures until OD750nm = 0.5−1.5 (typically 1−2 days). NOTE: S. elongatus UTEX 2973 cultures can be grown at 40°C in high light intensities (e.g., 2000 μmol photons m-2s-1)

2. Growth of helper and cargo E. coli strains (day 2)

1. Inoculate LB medium containing ampicillin (final concentration 100 μg/mL) and chloramphenicol (final concentration 25 μg/mL) with a DH5α E. coli strain containing vectors pRL443 (conjugal strain) and pRL623 (the helper strain) and grow at 37°C overnight at 225 rpm in a shaking incubator. Grow up a sufficient volume of helper strain culture, assuming 1 mL of culture is required per conjugation.

2. Inoculate LB medium (5 mL) containing appropriate antibiotics with the E. coli culture carrying the cargo vector. Grow the culture at 37°C overnight at 225 rpm in a shaking incubator.

3. Conjugal transfer (triparental mating) (day 3)

3. Prepare the E. coli helper and cargo strains. Centrifuge the helper and the cargo E. coli overnight cultures at 3,000 g for 10 min at room temperature. Discard the supernatant without disturbing the cell pellet.

4. Wash the pellet by adding fresh LB medium without antibiotics. Use the same volume as the initial culture. Resuspend the pellet by gently pipetting up and down. Do not vortex the culture. Repeat this step 3x to remove residual antibiotics from the overnight culture.

5. Centrifuge the resuspended culture (as in step 3.3.1), discard the supernatant and resuspend in half the volume of LB medium of the initial culture volume (e.g., 2.5 mL if the overnight culture was 5 mL). Combine 450 μL of the helper strain with 450 μL of the cargo strain in a 2 mL tube and set aside (leave at RT) until step 3.3.6.

6. Prepare the cyanobacterial culture. For each conjugation reaction, use 1 mL of cyanobacterial culture (OD750nm = 0.5−1.5).

7. Centrifuge the required total volume of cyanobacterial culture at 1,500 g for 10 min at RT, then discard the supernatant carefully without disturbing the cell pellet. Wash the pellet by adding fresh BG11 medium of the same initial volume. Resuspend the pellet by gently pipetting up and down, do not vortex the culture. Repeat this step 3x and set the washed culture aside.

8. Add an aliquot of washed cyanobacterial culture (900 μL) to the combined E. coli strains (helper and cargo) (900 μL) in a 2 mL tube. Mix the cultures by gently pipetting up and down. Do not vortex. Incubate the mixture at RT 2 hr for S. elongatus UTEX 2973.

9. Centrifuge the mixture at 1,500 g for 10 min at RT. Remove 1.6 mL of the supernatant. Resuspend the pellet in the remaining ~200 μL of supernatant. Place one 0.45 μm membrane filter on an LB-BG11 agar plate lacking antibiotics (section 8). Carefully spread 200 μL of the E. coli cyanobacterial culture mix on the membrane with a sterile spreader or a sterile bended tip and seal the plate with paraffin film

10. Incubate the LB-BG11 plate with the membrane for 24 hr. Maintain membranes with S. elongatus UTEX 2973 cultures at 40°C in 150 μmol photons m2 s1.

Objective

Author(s): Wendy Wu

Electroporate UTEX cells to introduce plasmid DNA into cells. This protocol is adapted from iGEM team HK_SSC 2021.

Materials

• Sterile Milli-Q water
• UTEX wash buffer: 2mM Tricine 2mM EDTA in Milli-Q water, autoclaved
• 1mm electroporation cuvette
• Electroporator
• Culture tube, 1x per electroporation
• Recovery medium
• Sterile tips

Procedure

1. Cells were grown and normalize to OD750 of 0.5. Spin the cells down
   1. volume: 500uL
   2. speed: 3000 rpm
   3. duration: 20 min.
2. Resuspend ONCE with UTEX wash buffer (see above). Spin down after washing using settings in step 1.
3. Wash twice with distilled water
4. Resuspend in distilled water to 10X concentration ( 1/10 of the original volume)
5. Mix 40µl aliquot of the suspension with 4 µg of plasmid DNA
6. Chill on ice for 2 minutes
7. Electroporated with a single square pulse with strength 8kV/cm-1 for 5 ms
8. Immediately after pulsation, cells were transferred to fresh DTN medium to recover for 24 hours prior to antibiotic challenge in liquid culture

Objective

Author(s): Pattarin Blanchard

Electroporate UTEX cells to introduce plasmid DNA into cells.

Notes:

The wash steps are meant to remove salt and weaken the cell wall of UTEX. Dissolved salts make the solution electrically conductive and could cause arcing at lower voltages than otherwise.

Materials

• 4 ug DNA per transformation
• UTEX wash buffer; 2 mM tricine, 2 mM EDTA, autoclaved
• BG-11 medium supplemented with 12 mM NaHCO3
• Sterile Milli-Q water
• Sterile tips
• 1x 1mm electroporation cuvette, pre-chilled, per transformation
• 1x culture tube per transformation
• 1x BG-11 plate with antibiotic per transformation
• UTEX liquid culture in mid-exponential phase (36-48h)
• Spectrophotometer and cuvette
• Centrifuge
• Sterile (micro)centrifuge tubes - pre-chill some on ice

Procedure

1. Retrieve liquid UTEX culture in mid-exponential phase. Measure OD750 and calculate how much culture is required for the number of transformations. Account for a negative control as well.

   $$V_{culture}=\frac{\text{\# of transformations}\cdot400 \text{ uL}\cdot 0.5}{OD_{culture}}$$

2. Pellet cells by centrifuging at 3000 g for 20 minutes. Decant supernatant and resuspend in n*400 uL wash buffer by pipetting up and down. Do not vortex.

3. Pellet cells by centrifuging at 3000 g for 10 minutes. Resuspend in the same volume sterile water by pipetting up and down.

4. Repeat step 3 once.

5. Pellet cells by centrifuging at 3000 g for 10 minutes. Resuspend with sterile 10% glycerol to 10x concentration (1/10th volume; n*40 uL).

6. Make 40 uL aliquots of concentrated cell suspension in sterile microcentrifuge tubes on ice.

7. Add (4) ug worth of DNA to each aliquot for transformation. Incubate on ice for 2 minutes. While waiting, set the electroporator to 800 V.

8. For each electroporation, pre-label and add 900 uL of supplemented BG-11 medium to a sterile 1.5 mL tube for recovery. Perform the next steps quickly and next to a flame.

9. Add the mixture to a chilled cuvette, making sure it is in the gap. Ensure there are no bubbles - pipette slowly and use it to pop any visible bubbles.

10. Place the cuvette in the holder of the electroporator with the cap off, slide the holder into the slot, and press run. If successful, it should report the actual voltage and duration.

11. Remove the cuvette, and add 900 uL of supplemented BG-11 to it. Pipette gently 2-4 times to mix the electroporated cells.

    1. Alternatively use a gel loading tip to retrieve the electroporated cells and add to a prealiquoted BG-11 tube.

12. Increase the pipette volume to ~900 uL and collect as much of the contents as possible, then transfer it to a culture tube.

13. Place the culture tube on a shaker in the 37C for (24h) recovery.

14. Once the recovery phase is done, plate onto _ ug/mL antibiotic BG-11 plates using beads or a spreader.