Purpose

Prepare 500 mL worth of LB + 20 μg/mL CM and 500 mL worth of PYE agar plates for use in future cloning experiments.

Materials & Methods

Chloramphenicol stock (20mg/mL)

For 1 mL recipe:

1. Weigh out 20 mg of chloramphenicol (Sigma, Cat. C0378-100G) in a sterile 1.5 mL eppendorf tube.
2. Add 1 mL 95% ethanol.
3. Mix well by shaking the tube and vortexing.
4. Filter solution using syringe. into a new tube.
5. Store at -20°C.

PYE agar solution (500 mL)

See PYE Medium Recipe for PYE media protocol.

1. Create the PYE base by mixing in a bottle filled with some distilled water (~300 mL):

   Chemical	Amount (for 500 mL)
   Peptone	1.5 g
   Yeast extract	1 g
   $CaCl_2$	109.54 mg
   $MgSO_4$	184.85 mg
   Agar	6 g

2. Top up to 500 mL with distilled/Milli-Q water.

LB agar solution (500 mL)

3. Mix in a bottle filled with some distilled water (~300 mL):

   Chemical	Amount (for 500 mL)
   Luria Broth base (Invitrogen, Cat. 12795084)	12.5 g
   Agar	6 g

4. Top up to 500 mL with distilled/Milli-Q water.

Preparing plates

*CM = chloramphenicol

1. Swirl the bottle(s) to mix and dissolve all solutes.
2. Sterilize the media by autoclaving, ensuring that the cap is loose.
3. Cool to room temperature/50 - 60°C before using.
4. If needed, add the appropriate antibiotic. For chloramphenicol plates, add 20 mg/mL CM stock to the agar solution:
   1. 1000x dilution of stock for 20 µg/mL (500 uL in 500 mL media) for E. coli
   2. 10,000x dilution of stock for 2 µg/mL (50 uL in 500 mL media) for CB2A Vortex the antibiotic to mix the solution immediately before adding to the media.
5. Using the Bunsen burner or BSC, pour 20 - 25 mL of media per plate.
6. When the agar has solidified, store the plates in plastic sleeves in 4C room.

Results

Stored in antibiotic box in -20°C freezer:

• 1.5 mL tube of 20 mg/mL CM stock (less then 500 µL remaining)

Stored in 4ºC room in iGEM box:

• 17 PYE-20 µg/mL CM plates (off to the side of the box because may not be able to use them)
• 20 12 LB plates

Notes for protocol optimization:

When creating media, mix everything in a large beaker first then transfer to media bottles.

While waiting for media to cool, bring plates to BSC to pre-label.

Before pouring plates, swirl the bottle to ensure agar hasn’t settled onto the bottom.

• Some of the LB plates did not solidify at all (or would have taken to long so were discarded).

Summary

LB and PYE plates were prepared and 20 ug/mL chloramphenicol stock was made. 500 µL of chloramphenicol antibiotic was added to PYE plates instead of LB, so PYE plates cannot be used for CB2A cloning due to the incorrect concentration (20 µg/mL is too high).

Hand-off

Prepare more plates and broth:

• LB + 20 µg/mL CM plates
• PYE + 2 µg/mL CM plates
• PYE plates
• 500 mL of PYE broth
• 500 mL of LB broth Need to buy more petri dishes.

Purpose

Aliquot and prepare BG-11 stock components, obtain UTEX starter culture, and receive guidance from Nannaphat (Patrik) Sukkasam. Prepare 900 mL of BG-11 medium and 500 mL worth of BG-11 agar plates.

Materials & Methods

The following recipe for BG-11 medium is based on the recipe from UTEX (the cell bank, not the strain): . Adjustments were made for UTEX by Patrik. It will be written up in the protocol page: BG-11 Medium Recipe.

BG-11 liquid medium

Working/final concentrations

• 17.6 mM $NaNO_3$
• 0.22 mM $K_2HPO_4$
• 0.3 mM $MgSO_4$
• 0.24 mM $CaCl_2$
• 0.012 mM citric acid
• 0.02 mM ferric ammonium citrate
• 0.002 mM $Na_2EDTA$
• 0.18 mM $Na_2CO_3$
• BG-11 trace metals solution (see link above)
• AGAR ONLY: 1 mM $Na_2S_2O_3$ (sodium thiosulfate)

Stock concentrations and recipe

Component	Relative stock concentration	mL stock per 1L medium
$MgSO_4$	1000x	1
$K_2HPO_4$	1000x	1
$CaCl_2$	1000x	1
$Na_2CO_3$	1000x	1
ferric ammonium citrate + citric acid	1000x	1
$Na_2EDTA$	1000x	1
BG-11 trace metals	1000x	1
$NaNO_3$	100x	10
AGAR ONLY as a final step	not added in liquid medium
$Na_2S_2O_3$ (sodium thiosulfate)	1000x	1

BG-11 agar medium

For 1L worth of plates:

2x medium solution

1. Add the amounts listed in the above table to 400 mL MilliQ water.

2. Add MilliQ water to bring final volume to 500 mL and mix well.

Agar solution

3. Add 15g phytagel to 400 mL MilliQ water.

4. Add water to bring final volume to 500 mL and mix well.

Final preparation

5. Autoclave both solutions then cool to 45-50C, or use paper towels

6. Under a flame or in a BSC, add 1 mL 1000x sterile $Na_2S_2O_3$ to the agar solution.

7. Combine both solutions and mix well.

8. Refrigerate or pour 30 mL per plate using a sterile 50mL falcon tube.

9. Seal and store in 4ºC room.

Notes

• Use MilliQ water. If not available, use $dH_2O$ (white taps by sinks).
• $Na_2S_2O_3$ slows the hardening of agar
• Phytagel is used in place of agar-agar because it is more transparent, allowing cyanobacteria to receive light.
• Often the antibiotic concentration needs to be gradually increased with each restreak. In these cases, it is more practical to make medium without AB, then add the desired amount to the bottom of a plate (by gently scooping out medium) instead. For this reason, a relatively precise volume of agar poured is required.

Results

The UTEX starter culture (small flask, green tape) is stored in the light incubator to the right of the clean bench, in the green room.

11 aliquots of 1000x stock components have been received and are placed on the bench in a green tube rack.

900 mL of sterile BG-11 medium is made and placed in bay C of the bench.

24 BG-11 agar plates were made and stored in the 4ºC room. The remaining BG11 agar is kept on bay C of the bench.

Summary

BG11 culture medium and agar plates were prepared and aliquots of 1000x medium stock was received with the help of Nannaphat (Patrik) Sukkasam.

Hand off

• Materials generated in this session are logged in our inventory page.
• Check the starter culture on for the color or cell density.
• Purchase a large 50mL tube rack.

Purpose

Continue from previous experiment to create solid media to use in future cloning experiments. For LB and PYE media, prepare 1000 mL of agar solution. 250 mL worth of each media will be used to make CM antibiotic plates.

Materials & Methods

PYE agar solution (1000 mL)

See PYE Medium Recipe for PYE media protocol.

1. Create the PYE base by mixing in a bottle filled with some distilled water (~800 mL):

   Chemical	Amount (for 1000 mL)
   Peptone	3 g
   Yeast extract	2 g
   $CaCl_2$	219.08 mg
   $MgSO_4$	369.71 mg
   Agar	12 g

   Note: You may want to prepare the solution in a larger beaker first for easier mixing of chemicals and then transfer over to a bottle.

2. Top up to 1000 mL with distilled/Milli-Q water.

LB agar solution (1000 mL)

3. Mix in a bottle filled with some distilled water (~800 mL):

   Chemical	Amount (for 500 mL)
   Luria Broth base (Invitrogen, Cat. 12795084)	25 g
   Agar	12 g

   Note: You may want to prepare the solution in a larger beaker first for easier mixing of chemicals and then transfer over to a bottle.

4. Top up to 1000 mL with distilled/Milli-Q water.

Preparing plates

*CM = chloramphenicol

1. Swirl the bottle(s) to mix and dissolve all solutes.

2. Sterilize the media by autoclaving, ensuring that the cap is loose.

3. Cool to room temperature/50 - 60ºC before using.

4. If needed, add the appropriate antibiotic. For chloramphenicol plates, add 20 mg/mL CM stock to the agar solution:

   1. 1000x dilution of stock for 20 μg/mL (500 μL in 500 mL media) for E. coli
   2. 10,000x dilution of stock for 2 μg/mL (50 μL in 500 mL media) for CB2A Vortex the antibiotic to mix the solution immediately before adding to the media.

5. Using the Bunsen burner or BSC, pour 20 - 25 mL of media per plate.

6. When the agar has solidified, store the plates in plastic sleeves in 4ºC room.

Workflow

7. Prepare and sterilise LB and PYE agar solutions. Split each into two bottles, one with 750 mL and the other with the remaining 250 mL.

8. To the 250 mL bottles, add the appropriate amount of CM stock.

• LB: Add 250 μg of 20mg/mL CM to 250 mL media to get 20 μg/mL final concentration.
• PYE: Add 25 μL of 20mg/mL CM to 250 mL media to get 2 μg/mL final concentration.

1. Pour and label ~10 plates for each media type. When solidified, store the plates in the plastic sleeves for petri dishes.
2. Store broth at room temp on the bench and plates in 4ºC room.
3. Make 5 mL of CM stock.

Calculations:

C1V1 = C2V2

For LB:

C1 = 20 mg/mL → 20,000μg/mL

V1 = x

C2 = 20 μg/mL

V2 = 250 mL

V1 = C2V2/C1 = (20 μg/mL 250 mL) / 20,000 μg/mL) = 0.25 mL

For PYE:

C2 = 2 μg/mL

V1 = (2 μg/mL * 250 mL) / (20,000 μg/mL) = 0.025 mL

Results

For each media type (LB or PYE), made 1000 mL of agar solution. 250 mL of each media was portioned out into 500 mL flasks and appropriate amount of CM was added to create plates. The remaining solution was stored away for future use.

Amount of plates prepared:

• 10 PYE-2 μg/mL CM
• 10 LB-20 μg/mL CM (not enough media for 10 plates so the last plate is very thin layer of agar, avoid using)

Plates were stored in iGEM box in 4ºC room and agar solutions were stored on the bench shelf:

LB and PYE agar solutions

Making plates in BSC

Prepared LB and PYE antibiotic selection plates stored in 4ºC room

2 of the LB plates were wrapped in parafilm and passed off to Beth Davenport, who will streak out pCB2A_Disp plasmid and pCB2A_Sec plasmid vectors in DH5α (E. coli) the next day for CB2A (Caulobacter) display, secretion and intracellular vector preparation in a future experiment.

Summary

LB and PYE antibiotic selection plates were prepared for future cloning experiments with E. coli and Caulobacter, respectively.

250 mL of media makes around 9 - 10 plates. To create exactly 10 plates, slowly pour the media to evenly distribute it across all the plates.

Hand off

• The prepared plates and agar solution were stored away in 4ºC room and room temperature on bench shelf to be used by members in future experiments.
• Need to prepare LB and PYE broth to use for liquid culture.
• Need to prepare more chloramphenicol antibiotic stock.

Purpose

1. Make LB medium
2. Prepare LB-Agar plates with KanR and AmpR for future E. coli transformation and cloning experiments.

Materials & Methods

• 2x 500 mL media bottles

• 1x 1000 mL media bottle

• 20x sterile Petri dishes

• 1x 500 mL graduated cylinder

• 1x clean beaker

• 250 µL Kanamycin (Kan) (50 mg/mL)

• 250 µL Ampicillin (Amp) (100 mg/mL)

• LB powder

• Agar powder

• Plenty of MilliQ water

• Aluminum foil for weighing

• Parafilm + scissors

• 1x 1000 µL pipette

• Tips for pipette

• Stir plate and bar

• Ethanol for sanitation

• Autoclave + safety equipment

Protocol - Preparation of LB broth

• Referencing a blank binder containing a list of protocols named “iGEM 2021 Protocols”. Recipes for LB media and LB-Agar media were sourced from here.

• LB Media

1. Recipe:

• 12.5 g LB powder
• 500 mL MilliQ water
• Liquid 20 Autoclave

1. The recipe creates 500 mL of media, so it was scaled up to 1 L.
2. 25 g of LB powder was measured using a top-loading scale and transferred into a smaller beaker alongside some MilliQ water. This was mixed into the liquid for easier pouring.
3. The 1 L flask was topped up using more MilliQ water measured in the graduated cylinder. As the increments went up to only 400 mL, the last 100 mL was measured in the difference in the graduated cylinder.
4. The flask was then tightly sealed, gently inverted ~20 times, then prepared for autoclave.
5. Flask was autoclaved under setting “liquid 20” for about approximately 50 minutes, or until completion.
6. Flask was tightened again and after cooling, stored.

• LB-Agar Media

1. Recipe:

• 12.5 g LB powder
• 7.5 g Agar powder
• Liquid 20 Autoclave
• 500 mL MilliQ water
• 500 µL Antibiotic

1. This recipe was used as is to produce 500 mL total of LB-Agar medium. Note it was split into two portions.
2. Two clean 500 mL bottles were prepared, one was set aside for later. 12.5 g of LB powder and 7.5 g of Agar powder were measured using a top-loading balance and poured directly into one of the flasks.
3. The flask was topped up using MilliQ water through the graduated cylinder. The final 100 mL was added once the volume in the flask reached 400 mL and was measured via difference in the graduated cylinder.
4. A stir bar was inserted and the media was stirred until it was mostly homogeneous (about 30 minutes of on and off stirring, 3/5s power of the stir plate).
5. The bottle was prepared for “liquid 20” and subsequently autoclaved using that setting for approximately 50 min, or until completion. The bottle saved earlier was also autoclaved under the same settings for sterility.
6. The bottles were sealed tightly after returning from autoclave.

• Pouring LB-Agar medium plates

1. 20 sterile plates (Petri dishes) were prepared. 10 were used with Kan and 10 were used with Amp.
2. After cooling the bottles, Petri dishes, pipette & tips and Kan and Amp stocks were moved into the BSC.
3. The LB-Agar medium was split into both bottles (250 mL in each)
4. 250 µL Kanamycin (Kan) (50 mg/mL) was added to one, 250 µL Ampicillin (Amp) (100 mg/mL) was added to the other using the pipette.
5. The Kan Petri dishes were poured first, in batches of 5, and placed aside. The Amp Petri dishes were then completed in batches of 5 and placed aside.
6. The pipette, tips, and bottles were all removed from the BSC. The dishes were left to cool, with Kan on the left and Amp on the right. This was to ensure correct labelling after solidifying.
7. After solidifying, a pen, tape, scissors, and parafilm were placed into the BSC. The Petri dishes were individually labelled with iGEM, antibiotic type, date and initials. They were then individually sealed using parafilm (cut using scissors in ~1.5cm strips). Wrap parafilm around entire plates, ensuring contact with both halves of the plate.
8. After completion, the Kan plates were packed into the bottom of the original Petri dish bag. The Amp dishes were arranged on top. In total all 20 Petri dishes were packed into the original bag. The bag was sealed using tape and labelled with iGEM, date, Kan + Amp dishes, and initials. It was then moved to storage in the 4ºC .

Summary

1 L of LB media was prepared (25 g LB to 1L H₂O). 10 plates of Kan LB-Agar were prepared.

10 plates of Amp LB-Agar were prepared.

Hand off

• The prepared plates and LB media were stored away in 4ºC room and room temp on bench shelf to be used by members in future experiments.

Purpose

• Make chemically competent DH5α E. coli cells

Materials & Methods

Subculturing & Making Competent Cells

• Mother culture of DH5α and BL21
• Two 125mL erlenmeyer flask
• LB Broth
• 0.1M of CaCl2
• 30% glycerol
• Sterile Oakridge Tube
• Ice bucket and Ice
• Water
• Spectrophotometer
• Curvette Subculturing Procedure:

1. Next morning, subculture 1 mL of mother culture into 99 mL of fresh LB without antibiotics

2. Shake in incubator at 37ºC and 200 rpm for 3-4 hours

3. Take 1 mL and read in OD600 in spectrophotometer to confirm OD is approximately 0.4 Calcium Chloride Wash Procedure:

4. Make sure all reagents are 4ºC

5. Separate culture into multiple Oakridge tubes

6. Place on ice for 20 minutes

7. Centrifuge at 4ºC at 4000 rpm for 10 minutes

8. Discard all media by tipping tubes and aspirate any remaining media

9. Resuspend pellet with 20 mL ice-cold 0.1 M calcium chloride

10. Incubate on ice for 30 minutes

11. Centrifuge at 4ºC at 4000 rpm for 10 minutes

12. Discard supernatant and combine pellets by resuspending in 5 mL ice-cold 0.1 M calcium chloride with 15% glycerol

13. Store in -80ºC freezer or use immediately for transformation (Chang et al., 2017)

• Added 49 mL of LB Broth into a 125mL erlenmeyer flask, with 1 mL of DH5α and BL21 into each respective flask for subculturing.
  • Jade mentioned that it will likely grow fast, so be there within 1-2 hour time frame.
  • It was incubated from 11:01 AM→12:47PM.
• After centrifuging, cells were white/yellow-ish and clumped onto the side of the oakridge tube.
• I waited ~10 minutes before resuspending the pellet in 0.1M CaCl2 with 15% glycerol after centrifuge.
• Made five Eppendorf tubes with an aliquot of 750 µL each
  • Placed in -70ºC freezer

Notes

• 1 loopful of glycerol-contained lab stock E. coli was inoculated into 50 mL of LB overnight to make the mother culture

• Used Av-Gay centrifuge to pellet culture

• 0.1 CaCl2 + 15% glycerol was used accidentally in 2/4 falcon tubes due to human error therefore the competency may have been affected

Results

• Mother culture was made and is stored in the 4ºC fridge

• Subcultured

• At the end, Pattarin 21 1 mL aliquots of chemically competent E. coli DH5a is stored in the iGEM 2025 stocks box in the top shelf of the -70°C freezer

Summary

• DH5a E. coli were cultured and made chemically competent and stored in the -70°C freezer in 1 mL aliquots in 0.1 CaCL2 + 15% glycerol

Hand off

• Caulocoli & UTEX working groups can use these cells for transformation purposes

Purpose

Continuing from the previous experiment to create liquid PYE broth for future culturing experiments.

Materials & Methods

PYE Broth (500 mL or 1000 mL)

See PYE Medium Recipe for PYE media protocol.

1. Create the PYE base by mixing in a bottle filled with some distilled water (~400 mL for 500 mL total, or ~800 mL for 1000 mL total):

Chemical	Amount (for 500 mL)	Amount (for 1000 mL)
Peptone	1.5 g	3 g
Yeast extract	1 g	2 g
CaCl2	109.54 mg	219.08 mg
MgSO4	184.86 mg	369.71 mg

1. Top up to 500 mL or 1000 mL with distilled/Milli-Q water.

Procedure

2. Swirl the bottle(s) to mix and dissolve all solutes.

3. Sterilize the media by autoclaving, ensuring that the cap is loose.

4. Cool to room temperature/50ºC - 60ºC before using.

Workflow

5. Prepare 500 mL or 1000 mL of PYE broth solution.

6. Sterilize the PYE broth solution by autoclaving.

7. Store the PYE broth at room temperature on the bench once cooled.

Results

PYE broth was prepared in 500 mL or 1000 mL volumes and autoclaved for sterilization.

Summary

1000 mL PYE broth was prepared.

Hand off

PYE broth will be used in future cloning experiments.

Purpose

The starter culture from 2025.05.19 UTEX Medium and Agar Prep, Starter Culture is starting to show green, indicating the starter culture is alive and can be plated for maintenance. Two plates will be streaked.

Materials & Methods

• 2x BG-11 agar plates
• 20uL pipette and tips
• BSC
• Parafilm

1. Two plates are taken out and left ajar (agar side still facing up) in the BSC to dry out and warm to room temperature for 30 mins.
2. 20 µL of the starter culture is added to each plate and streaked with the pipette tip

Results

Two plates labelled iGEM UTEX WT 21-5-25 PB are kept in the top shelf of the lighting chamber

Notes

• According to Nannaphat (Patrik) Sukkasam, cyanobacteria like UTEX tend to form large colonies, therefore it requires careful attention when streaking for selection to get individual colonies.

Summary

See above.

Hand off

• Because the culture was very dilute, the plates should be ready to be restreaked on .

Purpose

Beth Davenport streaked DH5α E. coli harboring pCB2A_Disp and pCB2A_Sec on PYE (+ 2 µg/mL chloramphenicol) plates (prepared the previous day) on May 21st. These plates were incubated in the 37C room at 3:15 pm and were checked at 9:15 pm the following day (30 hrs post-inoculation).

Materials & Methods

• Plates streaked with DH5α E. coli harboring harboring pCB2A_Disp and pCB2A_Sec plasmid
• 37ºC room
• 4ºC room

Results

A line of small, white colonies was observed on both plates. To avoid overgrowth plates were stored in 4C room.

pCB2A_Disp and pCB2A_Sec plasmid harboring plates (agar side down), PYE + 2 µg/mL CM

pCB2A_Disp vector plate (agar side up), PYE + 2 µg/mL CM

pCB2A_Sec vector plate (agar side up), PYE + 2 µg/mL CM

Summary

Plates growing DH5α harbouring pCB2A_Disp and pCB2A_Sec plasmid vectors yielded sufficient colonies for plasmid extraction and maintenance. Note that plates should be stored agar side up in the future.

Hand off

Colonies will be picked to start a culture for plasmid prep to extract pCB2A_Disp and pCB2A_Sec.

Purpose

Beth Davenport inoculated 5 mL PYE from CB2A glycerol stock on culture wheel on May 23 @ 12:37 pm. The overnight culture has been incubating for over 48 hours so glycerol stocks will be made to keep a stock of wild type CB2A cells for future use.

Materials & Methods

CB2A Glycerol stocks

• 5 mL CB2A culture in 30C room (‘for iGEM, CB2A’)

Making glycerol stocks:

800 µL culture + 200 µL 50% glycerol to get 10% glycerol final conc.

5 mL / 800 µL = 6.25 tubes

So make 6 glycerol stocks with 200 µL leftover to start new culture

Maintenance CB2A culture:

200 µL preexisting culture + 50 mL PYE (250x dilution)

Incubated in 30C room shaker at 6:51 pm

10 mL of 20 mg/mL chloramphenicol:

For 10 mL recipe:

1. Weigh out 200 mg of chloramphenicol (Sigma, Cat. C0378-100G) in a sterile 15 mL tube.
2. Add 10 mL 95% ethanol.
3. Mix well by shaking the tube and vortexing.
4. Filter solution using syringe into a new tube.
5. Store at -20C.

Results

CB2A maintenance culture

Glycerol stocks in -70C freezer box

Chloramphenicol stocks in -20C freezer box

Summary

• CB2A culture can achieve OD600 > 1 overnight if at least 200 µL starting culture is used to inoculate a 50 mL culture.
• Glycerol stocks were made with overnight culture.
• Made 20 mg/uL chloramphenicol stocks

Hand off

Glycerol stocks stored in -70°C will be used to inoculate future CB2A cultures

Chloramphenicol stocks stored in Antibiotics and Restriction Enzymes box

Purpose

Extract and purify small amounts of Caulobacter backbone plasmid DNA from bacterial cells.

Materials & Methods

Plasmid Mini-Preparation

Prepare miniculture

• pick two colonies from pCB2A_Disp plasmid plate, designate as #1 and #2
• pick two colonies from pCB2A_Sec plasmid plate, designate as #1 and #2
• Each tube has 5 mL LB + 5 uL 20 mg/mL CM (final conc is 20 µg/mL CM)

Marina - pCB2A_Disp plasmid #1

Talha - pCB2A_Disp plasmid #2

Arda - pCB2A_Sec plasmid #1

Jessie - pCB2A_Sec plasmid #2

Results

Sample	Concentration (ug/uL)	A260/280	A260/230
CB2A Display Parent plasmid #1	53.5	2.00	1.62
CB2A Display Parent plasmid #2	89.2	1.93	1.87
CB2A Secretion Parent plasmid #1	-	-	-
CB2A Secretion Parent plasmid #2	-	-	-

Summary

Plasmids were successfully purified and can be used to assemble the backbone plasmids pCaD, pCaS and pCaN.

Hand off

Stored plasmids in iGEM 2025 box in -20°C freezer.

Purpose

UTEX colonies appear on agar plates from 2025.05.21 Streaking UTEX on BG-11 plates, but there is contamination, so it needs to be carefully picked and restreaked. LB+Cm agar plates will also be prepared in advance to amplify distribution kit parts for Transformation and Golden Gate Assembly Training. As can be seen, the majority of colonies are not green but white.

Materials & Methods

Restreaking

• 2x BG-11 plates
• sterile loop
• bunsen burner

1. Warm agar plates to room temperature
2. Pick colonies and streak for isolation

Subculture

• 250 mL flask
• BG-11 medium
• sterile tips

1. Under a flame, add 50 mL BG-11 medium to the 250 mL flask
2. Take 10 µL from the starter culture and add to the new flask

Cm LB plates

• 10 petri dishes
• ~250 mL LB agar
• 20 mg/mL Cm stock

1. Melt LB agar in microwave in 30 s intervals until boiling. Keep the cap loosely screwed on.
2. Under a flame, pour ~250 mL into a 500 mL flask, wait to cool slightly.
3. Add 250 µL Cm stock into agar solution
4. Pour into petri dishes under a flame
5. Once plates solidify, wrap, put in bag, and label bag, store in 4°C room.

Results

A colony from each plate was streaked onto a new plate, stored in the light incubator.

UTEX was subcultured to a new flask, but the starter culture will be kept until the new flask turns green. Contamination on the plates may have been from the pipette tips in the BSC, or the starter culture itself had a contaminating species. Therefore, using plate colonies of UTEX for experiments ensure not only monoclonality but reduced contamination.

7 Cm LB plates were prepared stored in the 4°C room.

Summary

See results.

Hand off

Check back by to see if green patches appear on the new plates.

Purpose

To prepare a 50 mL preculture in advance for preparing electrocompetent CB2A cells on . Additionally, create a preliminary growth curve for CB2A to better estimate the time required for culturing CB2A for future protocols.

Materials & Methods

Preculture for electrocompetent cells

Materials

• Bunsen burner
• CB2A #1 glycerol stock
• 250-mL flask
• PYE broth medium
• Sterile pipette tips (200 µL or 20 µL size works)
• Ice Procedure

1. Prepare all other materials, then take out glycerol stock and sit it on ice to prevent thawing it out completely.
2. Under a flame, add 50 mL PYE broth to a sterile 250-mL flask.
3. Using a pipette tip, gently scrape the top of the glycerol stock and dip the tip into the broth. Swirl the tip around, then discard.
4. Secure the flask to a shaker in the 30°C room.
5. Shake for 48 hours. Notes

Start incubating at 7:00 pm.

CB2A growth curve

Materials

• Bunsen burner
• Preculture from other day (O.D. 1.142)
• PYE broth medium
• Sterile pipette tips
• 2x disposable cuvettes
• Spectrophotometer
• Parafilm squares Procedure

Same as the preculture but use ~200 µL preculture to inoculate 50 mL of PYE media.

Results

Time	OD600
7:30 pm	~0.1
8:00 pm	-0.003 (too low to read)
9:20 pm	0.018

CB2A culture after 2 hours:

CB2A preculture for making competent CB2A cells after >48 hrs

Summary

CB2A has a longer doubling time so it was not possible to complete the growth curve within a few hours. We will try again on a different day, starting in the morning. The CB2A preculture appears to have passed or is currently in log growth phase and has enough cells to scale up for competent cells.

Hand off

CB2A preculture is kept shaking at 30°C to make competent cells on .

Purpose

To practice a full cloning cycle using parts from the distribution kit. Operators will learn how to resuspend oligos from iGEM distribution kits, chemically transform cells, purify plasmids, and perform Golden Gate assembly.

Materials & Methods

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 42ºC

• Bunsen burner

• Timer

• Ice bucket

Procedure

Retrieving Parts from Distribution Kits

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	BBa_J435300	2	G7	Amp	a

Day 1 from Transformation and Golden Gate Assembly Training

Results

None of the plates yielded colonies the next day.

Summary

The distribution plates were switched (e.g., plate 1 was used instead of plate 2) so no colonies observed from transformations.

Purpose

Preparing plain PYE plates for future cell cultures.

Materials & Methods

• PYE agar media made from 2025.05.20 LB and PYE Media Prep, part 2
• Microwave
• Ethanol
• 10x Petri dishes
• Parafilm
• Masking tape

Procedure

1. Bottle of PYE agar media was melted using microwave in 60 second intervals.
2. Media bottle was brought to the BSC, along with 10 petri dishes.
3. Petri dishes were labelled inside the BSC.
4. PYE agar media was poured for a total of 10 plates.
5. Let the plates sit in the BSC for 30 minutes to solidify.
6. Wrap plates using parafilm, stack and store in plastic sleeve.
7. Seal and label the stack using masking tape, store in 4ºC room.

Results

• 10 prepared plates of plain PYE agar media were created. Plates were stored inside iGEM box in 4ºC room.
• The remaining PYE media solution was stored back on the bench shelf for future use.

Fig 1. Poured and labelled PYE plain plates

Fig 2. Prepared plates stored inside iGEM box in 4ºC room

Summary

10 plates of plain PYE agar media were prepared for use in future experiments with Caulobacter.

Hand off

• 10 prepared plates of plain PYE agar media were stored inside iGEM box in 4ºC room.
• Remaining solution was stored back on the bench shelf.

Purpose

1. Perform E. coli transformation training once again, following protocol from 2025.05.28 Transformation Training, Day 1.
2. Restreak new plates of UTEX Goal is to prepare to finish transformation training and to train on UTEX?

Materials & Methods

-Transformation training-

iGEM distribution kit plate 1 & 2

(maybe write in the specific plasmids extracted)

Molecular H2O aliquot

DH5_alpha E. coli competent cells

LB broth

Water bath and water + beaker (water + beaker to adjust if needed).

Ice

8x CM LB-agar plates

2x Kan LB-agar plates

2x Amp LB-agar plates

-UTEX restreaking-

Innoculating loop

Ethanol spray

UTEX plate

2x BG-11 AGAR plates.

-Both-

Bunsen burner

Pipettes and tips

Purpose

To pick recombinant colonies that were successfully transformed following Day 1 of Transformation and Golden Gate Assembly Training to culture transformed colonies that express the desired DNA fragments to be used later on in Golden Gate Assembly and final transformation of E. coli to produce purple colonies.

Materials & Methods

Antibiotics used per plate and label (label in LABEL column is used to identify plates).

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	BBa_J435300	2	G7	Amp	a

Other materials

E. coli plate cultures

LB broth

CM, Kan and Amp antibiotic stock.

Pipettes and tips

Serological pipette and tips

Culturing tubes

Bunsen burner

Procedure:

Protocol from Day 2 of Transformation and Golden Gate Assembly Training

Results

Shown: green fluorescent protein (GFP) expression in the “0” plate. This plate contains an E. coli with the plasmid from well “A10” of iGEM distribution kit 1. This plasmid serves a purpose for the golden gate assembly at the end, but also has GFP as marker.

The expression of GFP means that the cells used were indeed competent, and that the transformation for “0” was successful. While it does not guarantee that the transformations for plates “1”, “2”, “3”, “4” and “a” were successful, it may suggest that these plates were also able to take up their plasmids.

checking the plates again showed one or two colonies forming, which made sense given using Cm as a marker tends to take longer than Kan or Amp. Examining the culture tubes from the morning, only tube 0 was cloudy and thus successfully picked. Unfortunately, no colonies for the “a” plate formed, even though it should have formed fastest with Amp.

Colonies for part plates 1,2,3, and 4 were picked, and liquid culture tubes were placed in the 37°C room for overnight culture.

Summary

5 / 6 of the plates had colonies picked and moved to liquid cultures. “a”, the pDest plasmid, did not have visible colonies. “a” uses Amp, which means it should have shown colonies the fastest. Since it did not, this means something went wrong during the transformation. The Amp stock used may have been too high of a concentration, or the cells were damaged somewhere else along the way.

Hand off

Liquid cultures generated today will be used the following day to miniprep plasmids and perform golden gate assembly.

Purpose

Make LB CM stock and plates for E. coli transformation. Done due to potential issues with the previous inventory.

Pictured above: plating experiments done previously. The plates used must be replenished. Cm stock must also be replenished.

A lot of CM plates (8 total) were used in a plating experiment. Due to using the wrong distribution kit, these will not grow, and we will have to make more CM stock and agar plates.

Pictured: side of distribution kit. For future reference: check the side of the plates for the numbering.

Additionally, informed us that the CM powder used was a different color to the CM powder used in this session. As such, there is concern as to if the CM used for the previous experiment was not the correct powder. It was taken from the general lab stockroom, and was noted to be pink. Based on online search results, Chloramphenicol appears as a white-yellow solid.

Materials & Methods

Chloramphenicol (CM) - dry form

Ethanol (70%, we used the spray bottle)

LB agar aliquot.

Plates

Parafilm

5 sterile Eppendorf tubes

1 non sterile falcon tube

1 sterile falcon tube.

1 small beaker

1 medium Erlenmeyer flask

1 syringe and sterilization filter

Pipette and tips.

Weight measurement equipment.

Microwave + oven mitts

Bunsen burner and lighter

Vortex machine

Results

Protocol: making 5x 1 mL CM stock aliquots. Concentration of 20 mg/mL.

1. Get CM powder and measure on an ANALYTICAL BALANCE to 0.100 or 0.1000g → please check over this to ensure the precision is necessary. As of writing, there is aluminum foil (weigh boat) and disposable spatulas for use in the Hallam lab.

Note: in this session, an toploading balance was used that measures to 0.10g, and it is uncertain if this is accurate enough for the CM stock. Regardless, we mo ved forward.

1. Spray ethanol into a small beaker, ~ 5-6 mL. Pour this ethanol into the NON STERILE falcon tube until it hits the 5mL graduation. The antibiotic is more soluble in this than in plain water.

2. Seal the falcon tube tightly and vortex ~ 10 s to mix. Prepare 2nd falcon tube in holder, and 5 Eppendorf tubes in holder. Set up and light the bunsen burner as well for sterile flame.

3. [flame] Take your two falcon tubes near sterile flame. Keep the CM stock, and all other sterile materials near flame for as long as its being worked on and unsealed. Grab the syringe and attach the filter.

4. [flame] Remove the plunger from the syringe if possible, and pour the CM stock into the syringe from the back. Otherwise find a way to get the stock into the syringe then attach the filter. Open the 2nd falcon tube and place the syringe atop it, and replace the plunger. Press until as much stock as possible has been transferred into the sterile tube through the filter. The filter should remove microbes.

5. [flame] take the 1000 µL pipette and sterile tips, and then aliquot 1mL of the 5mL of stock into each Eppendorf tube. Ensure they are sealed upon completion.

This method produces 5x 1mL aliquots of 20 mg/mL CM stock. A mistake in lab resulted in the contaminiation of one, but 4 full aliquots were good to go.

Protocol: making CM agar plates.

1. Reheat LB agar in microwave until at least 250 mL of it is liquid. Do this in 30s intervals, or 1 min intervals in the microwave on standard settings. To check, gently tip to the side to see how much is liquid. When ~ 250 mL is liquid, remove with oven mitts to cool.

2. [flame] light the burner and flame all glassware after opening for the 1st time. The LB-Agar should be autoclaved and sterile. Pour off a 250 mL aliquot into the Erlemeyer flask, then cover and wait for it to cool. If the agar is too hot, use oven mitts to handle.

3. [flame] into the LB-agar aliquot, deposit 250 µL of the CM stock solution to make an ~ x1000 dilution. . Preferably, swirl gently in Erlenmeyer to mix. This was not done in this session, but the result should function just fine. → PLEASE CHECK OVER THIS ADDITION TO THE PROTOCOL.

4. [flame] pour the 250 mL of agar into ~ 10 plates. in session 9 were managed, but this should make 10. Once done, the flame may be turned off.

5. Let the agar set. In session, we waited about 2 hours. Once done, seal individually with parafilm then label with iGEM, date, contents and initials.

This should produce 10x LB-Agar CM plates for use with E. coli.

Results: 9x plates were made, with not enough to fill up the 10th. 5x aliquots of Cm stock were made with 1 being contaminated and discarded, resulting in 4x aliquots in the end.

Summary

CM stock was made and LB - CM plates were made. A total of 9x were made, and 4x 1 mL aliquots of CM were made, though the protocol, if followed, makes 5 aliquots and 10 plates.

Hand off

These materials will likely be used for the same transformation practice as 2025.05.28 Transformation Training, Day 1.

Purpose

To assemble the fragments of DNA isolated and quantified after Miniprep and Nanodrop through Golden Gate Assembly to transform E. coli into a recombinant strain that forms purple colonies

Materials & Methods

Golden Gate Assembly

• 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

Transformation

• LB medium

• Competent cell stock (start thawing in the middle of the GGA incubation cycle)

• Foam tube rack

• Water bath at 42ºC

• Glass beads

• Sterile microcentrifuge tube

• 4 Kan LB agar plates + each additional transformation replicate Procedure:

Protocol from Day 4 of Transformation and Golden Gate Assembly Training

Modification to Step 2: total volume was calculated to be 20 µl instead of 15 µl as written on the original procedure phase to increase the amount of DNA to maximise the chances of successful Golden Gate Assembly

Component	Volume (µL)
10x T4 Ligase Buffer	2
T4 DNA Ligase	0.5
DNA fragments (Promoter - BBa_J23100, RBS - BBa_B0034_m1, tsPurple - BBa_K1033906, terminator - BBa_B0015, pJUMP28-1A - BBa_J428353, pDEST)	3 µL per fragment = 15
BsaI-HFv2	1
Water	1.5
Total	20

Controls for transformation (4 controls):

This is to verify that Golden Gate Assembly was successful and the transformation took place as expected.

Label	Condition	Expected Results
C	Cell + Construct (experimental)	Purple Colonies
0	Cell + Backbone
(positive control)	Green/fluorescent colonies
B	Cell Only (negative control 1)	No colony formation
Media Only	Medium (negative control 2)	No colony formation

Results

Figure 1: The transformed E. coli colonies on Kan plates following Golden Gate Assembly and transformation. Top left: negative control - cell only. Top right: negative control - media only. Bottom left: construct colony. Bottom right: positive control - backbone only

An overnight incubation period was enough for significant colony growth, as marked by the numerous colony formations in the backbone plate. Additionally, no colony formation was seen in the negative controls. However, while the construct colonies should have been purple, they were observed to be green/yellow, identical to the backbone only colonies. This suggests that the assembly was unsuccessful while the transformation was successful. It is unclear why the assembly failed.

Figure 2. Cell + Construct (experimental) plate. Top Left: overview of plate, Top Right: zoomed in photo with purple colony labelled, Bottom Row: 3 Purple colonies found and an example of untransformed colony.

upon inspecting the experimental plate again, 3 small purple colonies were spotted. This indicated successful golden gate assembly, even with the expired reagents, although the efficiency is very low. The recipe should be adjusted to increase enzyme concentration relative to DNA fragments, and reduce backbone concentration relative to other fragments. The colony will be restreaked and picked the following day.

restreaking from the first plate yielded mostly white colonies with a few being purple. The colonies started out small grew slowly, suggesting it is possibly not E. coli. There was one liquid culture tube that was cloudy with a dark streak on the top. Upon spinning it down there there was a clear purple pellet. The plate was restreaked again and picked for miniprep. The pellets are kept at -20°C for future viewing and will be disposed of later.

Figure 3. Left side: Restreak of purple colonies, Top Right: Pelleted mini culture, Bottom Right: Miniculture of purple colony

Summary

To summarize, Golden Gate Assembly and transformation was carried out to yield two colonies - a positive control consisting of only the backbone and the construct. However, the construct colonies and the backbone-only colonies were both green/yellow, indicating that the assembly failed. We are going to redo the assembly with different sets of conditions (e.g different volume/different BsaI/use of master mix vs one-by-one addition of ligase/BsaI) to figure out why the assembly failed and culture the desired recombinant strain of E. coli that produces purple colonies.

Hand off

• pick colonies and restreak
• check plates from 2025.06.03 GG constructs and AB_T7_lacO Transformation for purple colonies

Purpose

To purify the plasmids in E. coli cultures generated in 2025.05.31 Transformation Training Day 2 Colony Picking and nanodrop to quantify the plasmids for Golden Gate Assembly Training.

Materials & Methods

From Transformation and Golden Gate Assembly Training:

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) Procedure: Day 3 Miniprep protocol from Transformation and Golden Gate Assembly Training

5 samples - 0, 1, 2, 3, 4 (samples from Transformation and Golden Gate Assembly Training Day 1 (also see lab notebook entry 2025.05.30 Transformation Training Day 1 Redo + UTEX restreak)

• The backbone (0) was eluted once and stored in an Eppendorf tube and it was also eluted twice to give sample 0.1 to compare the effectiveness of single and double elution. The quantity of DNA measured through Nanodrop significantly decreased in sample 0.1 compared to 0.

Results

Nanodrop measurement results showing DNA concentration and total quantified DNA in constructs 0-4

Construct Name	Concentration (ng/µL)	A260/A280	Total (ng) (~40 µL)
0	28.2	1.77	1132
0.1	3.4	4.92	136
1	23.8	1.93	952
2	27.7	1.79	1108
3	23.7	1.87	948
4	20.8	1.95	832

Summary

Miniprep and Nanodrop quantification was performed to quantify the isolate and quantify the DNA from the transformed competent E. coli cells. Consistent amounts of DNA were isolated from all samples except sample 0.1 (eluted twice) - shows multiple elutions decrease the concentration of DNA in samples. Using the isolated DNA from this experiment, we’re going to perform Golden Gate Assembly and Transformation.

Purpose

Results from 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation showed successful transformation but unsuccessful assembly of our construct. Among possible causes, such as incorrect proportions of enzymes to fragments, nonfunctional expired enzymes and/or reagents, and incorrect fragments created, we will address the first two causes. Two replicates using the same recipe but with corrected proportions (0.05 pmol per fragment, not 5 pmol) and different reagent vials, as well as another recipe (using the BsaI GG master mix).

This experiment will be done in hopes of identifying the cause of the failed transformation, and to address potential causes in the hopes of creating a successful construct.

Due to a shortage of LB-agar Kan plates, some of those were also created during this session.

Materials & Methods

Materials:

Golden Gate Assembly

• 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

Revised reaction recipe

Component	Volume (µL)
10x T4 Ligase Buffer	1
T4 DNA Ligase	1
DNA fragments (Promoter - BBa_J23100, RBS - BBa_B0034_m1, tsPurple - BBa_K1033906, terminator - BBa_B0015, pJUMP28-1A - BBa_J428353, pDEST)	1 µL per fragment = 5 µL
BsaI-HFv2	1
water	7
Total	15

Recipe using GG master mix

Component	Volume (uL)
10x T4 Ligase Buffer	2
DNA fragments	1 uL each = 5 uL
BsaI GG master mix	2
water	11
Total	20

GG protocol used was the same as in 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation, which was Transformation and Golden Gate Assembly Training, day 4.

LB agar stock is made from the black binder with plate recipes at the iGEM desk.

Results

An error was made and a P20 was used instead of a P2, wasting most of the DNA ligase and some of the BsaI and fragments.

Transformed cells were plated onto antibiotic plates however no purple colonies were observed.

Summary

Transformation was successful however it appears that the assembly efficiency and transformation efficiency is low for cultivation of purple colonies.

Hand off

Will observe plates over several days for purple colony growth on LB/Kan plates.

Purpose

We created 1 L of plain PYE media to be used to create electrocompetent cells.

Materials & Methods

1. To a 1 L autoclavable flask, add:

• 2 g of peptone
• 1 g of yeast extract
• 0.2 g of magnesium sulfate
• 0.1 g $CaCl_2$ dihydrate

1. Top up to 1 L with dH20
2. Loosely tighten the cap and add autoclave tape
3. Autoclaved on L20 setting for 50 mins

Results

Summary

Sterile media was prepared and will be used for 2025.06.03 experiment to make electrocompetent Caulobacter.

Hand off

The PYE media was handed off to Ada to create a larger bacterial culture from our starter culture.

Purpose

To perform a transformation on constructs from Golden Gate Assembly training and AB_T7_lacO constructs, validate transformation efficiency with appropriate controls, and miniprep plasmid DNA from successfully transformed colonies.

Materials & Methods

• Constructs tested:
  • AB_T7_lacO: iGEM Distrbution Plate 2 - O15 (AMP)
  • Golden Gate Training Construct: A1, A2, Z1, Z2, P1
• Controls:
  • Positive: cells + backbone (pJUMP28-1A, AmpR)
  • Negative 1: cells only
  • Negative 2: medium only

1. labeled and pre-chilled tubes on ice
2. aliquoted 50 µL competent E. coli cells per condition
3. added 2 µL GG construct DNA (or 1 µL backbone control)
4. incubated on ice for 20 min
5. heat shocked at 42 °C for 45 sec, returned to ice for 3 min
6. added 200 µL LB (no antibiotic) to each tube
7. recovered at 37 °C for 45 min with shaking
8. plated 50 µL of each transformation on LB + Amp (100 µg/mL) plates. Spread with sterile glass beads
9. incubated plates overnight at 37 °C

Purpose

To make competent CB2A for subsequent electrotransformation experiments where plasmids can be introduced into our Caulobacter cells.

Materials & Methods

Reference protocol: Electrocompetent CB2A Cell Preparation

Set up the preculture Sunday

1. Measuring OD600 on Tuesday:

• Scale up to 500 mL from 50 mL (1.1232)
• 10:00 am - 0.087 (wasn’t actually 0.1 because there wasn’t enough preculture)
• 11:30 am - 0.12 (beth)
• 2:10 pm - 0.2 (beth)
• 4:30 pm - 0.205 (wendy)
• 6:20 pm - 0.621 (cell growth program)
• 6:26 pm - 0.623

1. Set up prechilled water: Milli-Q in beaker placed in -20C
2. Using the centrifuge:
   1. Set to 3700g (21°C) → start
3. For first wash, didn’t top up to the top with water → repeated step 5
4. Followed the remaining protocol
5. Starting at step 1 of glycerol steps, all tubes were kept on ice
6. Aliquoted cells under the flame.

Calculations to make 10% glycerol:

C1V1 = C2V2

(50%)V1 = (50ml)(10%)

V1 = 10ml of 50% glycerol in 40ml sterile water

Results

Figure 1. 50 µl aliquot of electrocompetent CB2A in 10% glycerol.

Figure 2. -70°C iGEM freezer box containing the electrocompetent CB2A cells.

Summary

Electrocompetent cells will be used in subsequent electroporation experiments. For future reference, make more than 50 ml of 10% glycerol as some may evaporate during autoclaving.

Hand off

~45 x 50µl aliquots of electrocompetent CB2A were stored in the iGEM freezer box (-70˚C), labelled as “EC CB2A”.

Purpose

This experiment was done to make colourful bacteria to use in our HP workshops.

Materials & Methods

pDest Miniprep

Plasmid Mini-Preparation

Results

DNA (plasmid)	Plate Name	Concentration	260/280	260/230
mRFP1	R	87.4	1.90	1.54
tannenRFP	O	54.7	1.97	2.22
sarcGFP	G	46.9	1.91	2.08

Purpose

Electrocompetent CB2A cells created in 2025.06.03 Making Electrocompetent CB2A cells (Training) were validated by transforming the cells with pCB2A_Disp #2 extracted in 2025.05.26 Miniprep training with CB2A parent. This backbone plasmid confers chloramphenicol resistance, so we expect to see colonies on chloramphenicol PYE plates within a few days if the cells are competent and successfully take up the plasmid. Through this experiment, we will be testing out a preliminary electroporation protocol.

Materials & Methods

Materials

• PYE liquid medium

• Ice

• PYE-CM (2ug/mL) Agar plates

• Electroporation cuvettes, 0.1 cm interelectrode gap

• Disposable 10-mL round bottom culture tubes with snap-cap

• Sterile Eppendorf tubes

• Electrocompetent CB2A cells (labelled “EC CB2A”), pre-aliquoted into 50 uL portions (in Eppendorf tubes)

• pCB2A_Disp plasmid #2 miniprepped plasmid DNA (75.0ng/uL)

• Plating tools (glass spreader)

Equipment

• Shaking incubator at 30C

• Gene Pulser Xcell Microbial System #1652662 (Av-Gay machine)

• MicroPulser Electroporator #1652100 (Haney Lab machine)

1. Plasmid DNA preparation

We initially planned to transform CB2A with plasmids from the 2024 iGEM distribution plates, but these plasmids needed to be prepared in advance via transformation in E. coli followed by plasmid mini-preparation.

The following wells were resuspended, following Retrieving Parts from Distribution Kits. Since we did not end up using the DNA, the distribution kits were returned to -20C.

Part	Plate	Well	Resistance	Ori
pJUMP25-1A(sfGFP)	1	G10	Kanamycin	pBBR1 (high copy number broad host)
pJUMP23-1A(sfGFP)	2	J21	Kanamycin	RSF1010 (same as pCB2A_Disp ori)

The concentration of the pCB2A_Disp plasmid #2 miniprepped plasmid was remeasured as the concentration obtained in the miniprep entry was inconclusive.

Concentration	A260/280	A260/230
78.0 ng/uL	1.91	2.30

The concentration was 78.0ng/uL. The 260/280 ratio 1.91 and 260/230 ratio 2.30 indicates that this DNA product is relatively pure.

2. Electrotransformation

Electrotransformation was conducted by following the reference protocol CB2A Electroporation. As we intended to try out both machines available in the lab, we transformed two tubes of CB2A with pCB2A_Disp #2 (one for each machine).

2.A Calculating amount of DNA required for transformation:

The reference protocol calls for 1uL of 100-1000ng/uL undiluted DNA (i.e., 100-1000ng of DNA added to cells). We did not know the transformation efficiency of the cells, so we decided to add a volume of DNA that was around the middle of this range (500 ng) to promote a higher chance of successful transformation.

1 uL of 500ng/uL = 500 ng of DNA

500ng / (78.0ng/uL) = 6.41 uL (we rounded up to 6.5 uL)

2.B Preparation

1. Thaw two 50 uL aliquot tubes of electrocompetent Caulobacter cells on ice (one tube per condition).
2. Put two electroporation cuvettes on ice to chill.
   1. The cuvettes were cleaned prior to this step (re-used from previous years).
3. Put 6.5 uL of 78.0ng/uL undiluted pCB2A_Disp #2 DNA in the new Eppendorf tubes (labelled) on ice.
4. Pre-warm 950 uL of PYE medium in culture tubes. (950 uL of PYE was added to culture tubes under the flame, then transferred to 37C room to warm up).

2.C Pre-electroporation steps

1. Add 50 uL of cells to each of the tubes with 6.5 uL of DNA on ice.
2. Let the tubes sit for 2 minutes.
3. Transfer 50 uL DNA/cell mix to cuvette, and pulse.

2.D Electroporation

As we did not see the full demonstration of the machines, we followed the written protocol and modified the steps for each machine.

2.E Incubation and plating steps

1. Incubate at 30C for 2 hours.
   1. The tubes were placing in the culture wheel in the 30C room.
2. Warm up two PYE-CM (2ug/mL) plates.
3. Plate on PYE-CM and incubate 30 degrees for 2-3 days.

2.E.1 Making dilutions:

After shaking, serial dilutions of the transformation cultures were carried out under the flame. The dilutions were prepared in the following manner:

1. Tube A (10x): 20 uL from each replicate sample added to 180 uL of PYE liquid media in an Eppendorf tube and flicked gently to mix. 20 uL was immediately removed for the next dilution.
2. Tube B (100x): 20 uL from tube A was added to 180 uL of PYE → mix → immediately remove 20 uL.
3. Tube C (1000x): 20 uL from tube B was added to 180 uL of PYE.

In the end 6 dilution tubes were prepared (3 per replicate), labelled with sample replicate number and the dilution letter. Along with the original transformation cultures, 8 samples/tubes were plated.

2.E.2 Plating

Each plate was split into quadrants and labelled accordingly (Q = quadrant).

Example of plating scheme for Rep 1 plate (the same thing was done for Rep 2 but with Rep 2 set of samples):

Quadrant	Label (dilution)	Sample
Q1	ORIGINAL (undiluted)	Replicate 1 transformation tube
Q2	1/10	Tube A-1
Q3	1/100	Tube B-1
Q4	1/1000	Tube C-1

50 uL plated for each sample.

For each plate (all steps done under the flame):

1. 50 uL of sample was dispensed into the centre of the quadrant. This was repeated for all 4 quadrants. The same pipette tip was used, starting from the LOWEST dilution up to the undiluted sample.
   1. If you do not start at the lowest dilution, change tips to avoid accidently increasing the concentration of the lower dilutions.
2. Using a sterilised glass spreader, start spreading from Q4 (lowest dilution) and proceed with the same spreader to the other quadrants, moving up the dilutions in a sequential manner (Q3 → Q2 → Q1).
3. Let the plates dry for 1 - 2 mins. Flip the plates over (agar side facing up) and
4. Wrap the plates in parafilm.

The plates were incubated in the 30C room at 10:30 pm.

Figure 3. Plates with electrotransformed Caulobacter bacteria

Results

Checked plates on (43.5 hrs incubation, ~ 2 days):

Both plates had plenty of small white colonies. Rep 1 appeared to have more colonies but colony enumeration was not conducted.

Figure 4. Plates with electrotransformed Caulobacter bacteria

Summary

Transformation of CB2A with pCB2A_Disp plasmid yielded colonies and what we assume are chloramphenicol-resistant transformants appeared in a little less than 2 days. This is somewhat expected as according to the reference protocol, C. crescentus cells have electroporation efficiencies comparable to E. coli cells and require only a small amount of DNA, yielding thousands of colonies within 2 - 3 days. However, according to Beth it may take transformed colonies may take 4-6 days to appear due to the antibiotic in the plates, so our transformants formed quicker in this case, and may possibly not be Caulobacter cells.

Additionally, there were steps during the protocol that we would like to standardize for future electrotransformation experiments, so more testing should be done to optimize the transformation protocol for recombinant plasmids and calculate the transformation efficiency of CB2A. Another electrotransformation experiment should be carried out following more training from Hallam lab members.

Things to standardize:

• Amount of DNA added to cells (6.5 uL might be too much)
• Procedure for using electroporation machines
• Incubation before plating (see whether shaker or culture wheel makes a difference)
• Plating procedure

Hand off

The plates were stored in 4C room to preserve colonies for further analysis. To double-check that the cells took up pCB2A_Disp plasmid #2, a restriction digestion experiment will be carried out.

Purpose

Colonies expressing the correctly-assembled tsPurple construct from 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation are picked for overnight culture for agar art. A miniprep, glycerol stock, and agar plate, will be made the following day.

Materials & Methods

• LB medium
• Kan stock
• Sterile tips
• Sterile capped culture tubes

Colony picking

1. For each colony, prepare a culture tube with 4 mL LB medium with 4 uL Kan stock
2. Use a p20 tip to touch a desired colony and drop the tip into the culture tube # Results

3 culture tubes of picked colonies were made.

unfortunately the culture tubes still remain clear.

In hopes of reviving the purple colony, the remains of it on the previous plate was restreaked onto a fresh LB/Kan plate.

Summary

Colony picking into liquid culture appears unsuccessful as of , the remains of the purple colony was restreaked.

Hand off

Check in the coming days to see if purple colonies form on the new plate.

Purpose

To prepare for future experiments, we need to check if the UTEX strains are naturally resistant to any of the marker antibiotics we plan on using. Kanamycin (Kan), Ampicillin (Amp), and Chloramphenicol (Cm) will be tested, alongside a non-antibiotic control plate. In theory, it should be resistant to none of these.

In addition to this, our previous experiments, such as 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation produced a lot of liquid culture waste. We planned on eliminating 1x UTEX culture and 2x E. coli cultures in this session. The biohazard bin was also getting full, so we intended on emptying it.

Finally, we discovered a purple colony in one of the plates and a culture. We are unsure if it is E. coli, but to see if the plasmid replicates we have restreaked it in an LB-Kan plate as well as made a liquid culture.

Materials & Methods

Materials

• Petri dishes
• BG-11 agar
• Antibiotic stocks
• Sterile loop
• Disposable culture tube

Procedure

Remaining BG-11 agar from 2025.05.19 UTEX Medium and Agar Prep, Starter Culture was used to pour plates according to BG-11 Medium Recipe. Antibiotic was added after 30 mL of agar medium was poured into a 50 mL tube. A new 50 mL tube was used for each antibiotic, starting with the lower concentration.

Amp: stock is 100 mg/mL

• high 100 µg/mL ---> 30 µL stock
• low: 33 µg/mL ---> 10 µL stock

Kan: stock is 50 mg/mL

• high 50 µg/mL ---> 30 µL stock
• low: 20 µg/mL --->12 µL stock

Cm: stock 30 mg/mL

• high 30 µg/mL ---> 30 µL stock
• low 10 µg/mL ---> 10 µL stock

New plates were streaked from the plate generated in 2025.05.30 Transformation Training Day 1 Redo + UTEX restreak, using the same tools and protocol, but more plates.

Culture disposal was done using 10% bleach, 1M NaOH, and ethanol to wipe.

Purple colony restreaking was done with the same method as the UTEX, replacing BG-11 plates with LB-kan plates.

Results

Six (6) total plates were streaked from the successful plate from 2025.05.30 Transformation Training Day 1 Redo + UTEX restreak.

2x Kan (1x high concentration, 1x low) BG-11 plates

2x Amp (1x high concentration, 1x low) BG-11 plates

1x Cm (low concentration) BG-11 plate

1x No AB BG-11 plate

Note that we intended on having both a high concentration CM and a low concentration CM plate. However, the BG-11 medium solidified before the high concentration plate could be poured. We do not currently intend on using CM, so we have less of a need to thoroughly test the CM plates. A non-AB plate is also there to test if the plating was successful.

Results: After ~2 days, the BG-11 (UTEX) plates showed no sign of green colonies.

After ~6 days, the BG-11 CONTROL plates showed very small signs of colonies, whereas none of the other plates did. Although it was slow to grow, this shows that our strain has no natural resistance to Kan, Amp or Cm antibiotics.

One (1) plate was streaked from the purple bacterium grown from the results of 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation in LB-Kan agar.

Pictured: After waiting ~2 days, the restreaked colony has successfully grown into a purple plate. Note that the date on the originating plate is not 01.06.2025, this appears to have been a more recent plate from the same colony as the linked notebook entry.

One (1) liquid culture was made from another purple colony (from the same plate as the above restreaking) in LB-Kan medium.

There is possible contamination in the old LB medium bottle so 10% bleach was added to it in a 1:1 ratio, and disposed of.

Note that the contaminant is potentially a filter from a serological pipette, as it does not appear to be a living colony. However, the broth pictured is from last year, and with the added contamination it is likely better to use another batch entirely.

A glass culture tube shattered, likely due to improper handling after flaming, near the top. As such it was disposed of in the sharps waste. It was also slightly cracked before it shattered.

Summary

A set of UTEX plates were streaked with varying antibiotics to test for innate resistance.

After ~6 days, only the control plate grew colonies, indicating no natural AB resistance.

A plate and culture were made from purple colonies to test for plasmid viability. Essentially, we are testing for the success of 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation. The plate was successful and displayed purple colonies.

Liquid waste was disposed of. Vessels were cleaned. Biohazardous waste bin was emptied and re-bagged.

Hand off

Plates were moved to their respective incubation chambers.

Purpose

Following 2025.06.04 Electrotransformation Training with pCB2A_Disp #2, a miniculture will be created for each replicate to miniprep the following day.

To prepare for upcoming experiments, a starter/mother WT CB2A culture will be made from glycerol stock to inoculate a new culture for growth curve analysis. CB2A in liquid culture takes a day less time to grow than colonies so we expect the culture to reach sufficient O.D. (> 1) in 1-2 days. Additionally, maintenance plate of WT CB2A colonies will be made, which can be used to easily create more liquid cultures in the future.

Lastly, a series of plates will be created to evaluate the antibiotic resistance of WT CB2A along with plating colonies from 2025.06.04 Electrotransformation Training with pCB2A_Disp #2 onto a higher concentration PYE/CM plate as an additional test to confirm that the colonies only grow at the expected antibiotic concentration.

Materials & Methods

Materials & Equipment

• Sterile tips (P200)
• Sterile 250-mL flask
• PYE liquid medium
• CB2A #1 glycerol stock (WT CB2A)
• Ice
• Agar plates:
  • 1x PYE
  • 1x PYE-CM (2ug/mL) - LOW concentration
  • 2x PYE-CM (20ug/mL) - HIGH concentration
• Cm antibiotic (20 mg/mL)
• Disposable 10-mL round bottom culture tubes with snap-cap
• Inoculation loops
• Shaker & incubator at 30C

Plates were taken out of 4C room, labelled and placed in 30C room for 10 minutes to warm up.

The glycerol stock was taken out of -70C freezer and placed on ice after all materials were prepped to prevent it from fully thawing.

The existing Cm antibiotic (AB) stock was a bit too high to easily use for creating minicultures, so a lower concentration stock was created:

2 mg/mL Cm AB stock

For 100 uL of stock (because we didn’t need that much):

Calculation C1V1 = C2V2

C1 = 20 mg/mL CM stock

V1 = x

C2 = 2mg/mL CM stock

V2 = 0.1 mL (100 uL)

x = 2 * 0.1 / 20 = 0.01 mL (10 uL)

Procedure

1. Add 10 uL 20 mg/mL CM stock to 90 uL 70% ethanol in a sterile 1.5 mL Eppendorf tube under the flame.
2. Vortex to mix, then store at -20C when done using. The stock tube labelled “Cm 2mg/mL” was stored in the “Antibiotics & Enzymes” freezer box in -20C.

pCB2A_Disp #2 Miniculture

Under the flame:

1. Add 4 mL PYE to each culture tube.
2. Add 4 uL of a 2mg/mL to achieve a final concentration of 2ug/mL (1000x dilution).
3. Use a clean pipette tip to touch the top of the colony and drop it into the tube.
4. Shake at (~200 rpm) at 30C and incubate for 14 - 24 hours. Note: This is the first time creating minicultures with Caulobacter cells so they may required longer to grow than E. coli cells (12 - 18 hrs), this can be tested in future experiments.

Figure 2. Inoculated minicultures, labelled “transformed CB2A rep 1 #1” and “transformed CB2A rep 2 #1” was set to shake at 30C at 11:00 pm.

Figure 3. Colonies picked from pCB2A_Disp #2 transformant plates (the chosen colony was circled and labelled #1 on each plate).

Cm Antibiotic Resistance Test Plates and CB2A Maintenance Plate

Under the flame:

1. Use a clean pipette tip to scratch the top of the glycerol stock.
2. OR pick a colony from the transformant plate using an inoculation loop.
   1. For the transformant plates (Rep 1 and Rep 2), one colony was picked from each one (same colony used to create minicultures) and streaked across half of an agar plate.
3. Touch the tip/loop near the edge of the plate to transfer majority of the cells onto the agar (if using glycerol stock, the cells will start thawing at the end of the tip and form a droplet). Then gently streak the cells down the plate in a zig-zag manner.
4. Flip the plates agar side up to let them dry for 1 min then wrap them in parafilm.
5. Incubate at 30C for 48 hours.

Plates created:

Name (source of bacteria - condition)	Shorthand	Medium type	Cells streaked	Expected result after 2 - 3 days
WT CB2A maintenance plate	maintenance	PYE	WT CB2A	Colonies
WT CB2A Neg Ctrl - Low [Cm]	WT-Low [Cm]	PYE-Cm (2ug/mL)	WT CB2A	No colonies, WT CB2A shouldn’t have Cm resistance
WT CB2A Neg Ctrl - High [Cm]	WT-High [Cm]	PYE-Cm (20ug/mL)	WT CB2A	No colonies, WT CB2A shouldn’t have Cm resistance
pCB2A_Disp #2 transformed CB2A Neg Ctrl - High [Cm]	pCB2A_Disp #2-High [Cm]	PYE-Cm (20ug/mL)	Colony #1 from Rep 1 and Colony #1 from Rep 2 transformant plates	No colonies, transformed CB2A shouldn’t confer resistance to 20ug/mL Cm (only lower concentration of 2 ug/mL)

Note: The glycerol stock started thawing after plating the high Cm concentration plate, so instead of scratching the surface, ~2 uL of stock was dispensed onto the remaining plates.

The plates were incubated at 30C at 11:00 pm.

Figure 4. Cm Antibiotic Resistance Test Plates and CB2A Maintenance Plates created (incubated at 30C at 11:00 pm). Top left is CB2A maintenance plate, top right is WT-High [Cm], bottom left is WT-Low [Cm] and bottom right is pCB2A_Disp #2-High [Cm].

Mother CB2A culture

Under the flame:

1. Pour ~50 mL of PYE into a sterile 250-mL flask.
2. Using a clean pipette tip (attach to P200 pipette), lightly scratch the top of the glycerol stock until you get a small amount of cells at the end of the tip (about half of a sesame seed).
3. Swirling the tip in the flask and gently pipette up and down to inoculate the PYE medium.
4. Shake (~200 rpm) at 30C and incubate for 1 - 2 days.

Figure 5. Mother culture labelled “CB2A mother culture” set to shake at 30C at 11:00 pm.

Results

:

Since the colonies from the transformant plates were confirmed to not be Caulobacter in 2025.06.11 Troubleshooting CB2A Electrotransformation Remaking PYE Medium and PYE-Cm Agar Plates, the minicultures were discarded.

Measuring O.D.600 of mother culture:

The O.D.600 measurement of the CB2A mother culture was 0.161 (post-23 hrs shaking at 30C).

Checking the plates:

Upon checking on the antibiotic and maintenance plates, no colony growth was observed on any of the Cm plates, and the maintenance plate had a small lawn of growth at the one edge of the plate.

The lawn of growth is unexpected as we did not expect WT CB2A to form colonies so fast (this was only 1 day instead of the typical 2 - 3 days under optimal laboratory conditions).

Figure 6. Results of CM Antibiotic Resistance Test Plates and CB2A Maintenance Plates after 23 hours incubation at 30C.

Figure 7. CB2A maintenance plate after 23 hours incubation at 30C.

:

Measuring O.D.600 of mother culture:

The culture O.D.600 was 1.21 (post-44.25 hrs shaking at 30C). The culture will be used to inoculate a new culture to start a growth curve experiment.

Sample time	Culture time	OD600
10:00pm, Jun 11	23 hrs	0.161
7:45pm, June 12	44.25 hrs	1.21

Table 1. Summary of OD600 measurements of mother culture

Checking the plates:

No colony growth was observed in the WT-High [Cm] plate, but a few colonies was observed in the WT-Low [Cm] plate. This means that the WT CB2A may have resistance to 2ug/mL Cm which is unexpected, as this strain is typically selected with this level of Cm concentration. We suspect the plate may be contaminated.

For the pCB2A_Disp #2-High [Cm] plate, colonies formed along the streak lines for the rep #1 sample, indicating that these colonies conferred resistance to high concentration level of Cm. However, in light of learning that our transformants were not Caulobacter in the previous day, these result is no longer relevant.

The maintenance plate formed more colonies on the streak lines.

Checking the plates:

Still no growth observed in the WT-High [Cm] negative control plate, and a few more colonies formed on the WT-Low [Cm] plate.

For the #2-High [Cm] plate, colonies formed for rep #2 as well as rep #1.

The maintenance plate had lots of tiny colonies speckled all over the plate, making it difficult to isolate individual colonies. We will try this experiment again.

The plates were discarded.

Summary

1 eppendorf tube of 2 mg/mL Cm stock was prepared and stored in the “Antibiotics & Enzymes” freezer box in -20C.

The CB2A mother culture reached 1.21 OD600 after 44 hrs (almost 2 days).

The CB2A maintenance plate yielded colonies, but the pattern of growth was not suitable for colony picking and due to the fast appearance of the lawn of growth after 1 day of incubation, we suspect that the colonies may not be Caulobacter. In our next attempt, we will use less glycerol stock and streak more lines along the plate to cover a wider surface area and spread out the colonies.

The appearance of colonies on the pCB2A_Disp #2-High [Cm] plate indicates that whatever bacteria was growing in the transformant plates were indeed resistant to Cm, but since the original colonies from the transformant plates were proven to no be Caulobacter, these negative control plates were not further analysed.

The WT CB2A Cm antibiotic test plates gave unexpected results. The Low-[Cm] plate yielded a few colonies after 2 days of incubation, indicating that our WT CB2A has some resistance to chloramphenicol. This does not align with literature, so it is possible that the colonies we observed was not Caulobacter. No colonies was observed in the High-[Cm] plate, which was made using the same Cm stock (from the pink-coloured powder) as the Low-[Cm] plates, so this at least confirms that the Low-[Cm] plates were effective and that those few colonies that formed were resistant to chloramphenicol.

The WT CB2A maintenance plate and Cm antibiotic test plates will be attempted again, and we will recruit the help of Beth to examine the resulting colonies under light microscopy to confirm that they are Caulobacter.

Hand off

The CB2A mother culture will be used to inoculate a new culture for growth curve analysis.

Purpose

To confirm that we extracted the correct plasmids in 2025.05.26 Miniprep training with CB2A parent, a restriction digest will be carried out, followed by gel electrophoresis analysis. Since these plasmids both be cut by the restriction enzymes BsaI and EcoRI, a double digest will be conducted using these enzymes.

• pCB2A_Disp can be digested by any of the following unique site cutting enzymes we have available in the lab: BsaI, EcoRI, PstI
• pCB2A_Sec can ve digested by the following enzymes: BsaI, EcoRI, BamHI
• We chose to digest both plasmids with BsaI and EcoRI to simplify the experiment

Materials & Methods

Before setting up the restriction digest reaction, the concentration of the plasmids were remeasured (as the Nanodrop results were inconclusive when measurements were taken on May 26th).

Remeasuring Nanodrop DNA

NanoDrop Spectrophotometer

Blank = Elution buffer

Sample	Concentration [ng/uL]	A260/280	A260/230	Volume needed to get 1 ug (1000 ng) for digest
pCB2A_Disp #1	87.2 ng/uL	1.96	2.17	11.5 uL
pCB2A_Disp #2	78.0 ng/uL	1.91	2.30	12.8 uL
pCB2A_Se #1	27.2 ng/uL	2.19	1.78	36.8
pCB2A_Sec #2	22.6 ng/uL	2.39	3.45	44.2 (rounded 43 as 44.2 would take up too much volume in 50 uL rxn)

Restriction Digestion reaction

Initially we planned to do two sets of digestions (a double digestion with both BsaI and EcoRI + single digestion with just EcoRI). However, we realized there wasn’t enough DNA so only the double digestion was carried out.

Protocol: Restriction Digest with NEB Enzymes

Enzymes and buffer:

• BsaI-HFv2 (Exp 02/24)
• EcoRI-HF (Exp 9/20)
• 10X rCutSmart Buffer

Component		per tube
10X rCutSmart Buffer	20 uL	5
BsaI-HFv2	4 uL	1
EcoRI-HF	4 uL	1
TOTAL	28 uL	7 uL

Table 2. Master Mix 1 (BsaI and EcoRI) (tube is labelled ”+“)

Component	Tube 2	Tube 3	Tube 4	Tube 5	Tube 6	Tube 7	Tube 8	Tube 9
DNA	11.5 uL pCB2A_Disp #1	12.8 uL pCB2A_Disp #2	37 uL pCB2A_Sec #1	43 uL pCB2A_Sec #2	11.5 uL pCB2A_Disp #1	12.8 uL pCB2A_Disp #2	37 uL pCB2A_Sec #1	43 uL pCB2A_Sec #2
Master Mix	7 (+)	7 (+)	7 (+)	7 (+)	6 (-)	6 (-)	6 (-)	6 (-)
Water	31.5 uL	30.2 uL	6.2 uL	0 uL	32.5 uL	31.2 uL	7.2 uL	1 uL
TOTAL volume (uL)	50	50	50	50	50	50	50	50

Table 3. Recipe scheme for each reaction tube. tubes were labelled according to the lane they would be loaded in the 10-lane agarose gel.

*Master mix was created and aliquoted for tubes 6 - 9 but there wasn’t enough DNA so those tubes were discarded.

Results

The reaction tubes (labelled 2 - 5, with “CC 6.11” written in the PCR strip tab) were stored at -20C in the iGEM PCR freezer box.

In the future a protocol should be written out (and order of reaction components should be added as water → buffer → DNA → restriction enzymes to prevent premature digestion). Additionally, a negative control without template DNA should be prepared. Since this digestion experiment was intended for plasmid confirmation and the fragments would not be gel extracted, the reaction was not heat inactivated, but this should be done if another step in the workflow follows without DNA purification.

Hand off

The samples were stored at -20C to be run in agarose gel .

Purpose

For further training and fun, we plan on making E. coli colony art. To do this, we will need a few more Cm plates to grow the cultures needed. As we already have the materials to do so, we have decided to do this early. We aim to make 6x plates.

Materials & Methods

LB-Agar

Cm stock from 2025.05.29 Preparing LB CM plates and stock

Plates (Petri dishes)

250 mL Erlenmeyer flask

Parafilm

→ making about 6 plates.

Bunsen burner

Microwave and mitts

Pipette & tips

Ethanol

Protocol

Protocol from 2025.05.29 Preparing LB CM plates and stock.

Making 6 plates, using about 25 mL of agar each. Need 150 mL of agar. Use 150 µL of Cm for a x1000 dilution.

Results

Figure 1: Label and bag of the produced plates.

There was more than enough agar to cover each plate’s bottom. The remainder was distributed unevenly, eyeballed to be about the same for each plate.

No bag was available, so saran wrap was used. All plates are individually wrapped with parafilm regardless.

Summary

6x LB-Cm plates made and placed in 4°C room, iGEM box.

Hand off

These will be used for future experiments with E. coli, specifically the culture art.

Purpose

Miniprep liquid culture containing the tsPurple synthetic construct for later use. Because we are unsure if the organism carrying the plasmid is E. coli, the liquid culture will not be made into a glycerol stock.

Materials & Methods

Plasmid Mini-Preparation

• GeneJET plasmid miniprep kit
• x2 microcentrifuge tubes
• x1 collection tube NanoDrop Spectrophotometer

Results

Initial centrifugation showed a purple cell pellet, indicating the culture contains the synthetic construct generated in 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation.

The restreaked and repicked purple construct liquid culture from 2025.06.01 Transformation Training Day 3 Golden Gate Assembly and Transformation was successfully miniprepped and the concentration is reported below. It is placed in the iGEM 2025 -20 box.

Construct Name	Concentration (ng/ul)	A260/A280	A260/A230
purple construct	37.0	1.83	0.91

Eppendorf tubes with 500 uL, 850 uL, and 1500 uL water were also made and labelled for balancing the centrifuge. They are kept on the pink rack on the right side of the bench shelves.

Summary

The purple construct was miniprepped and resuspended to a concentration of 37.0 ng/uL.

Purpose

When discussing the 2025.06.04 Electrotransformation Training with pCB2A_Disp #2 results with Beth, it was revealed that the colonies from the transformant plates were not Caulobacter but a contaminating species.

Initially we believed that the colonies were Caulobacter based on morphological observations. Caulobacter colonies appear translucent or opaque (in various shapes of yellow/gold/orange/re-orange depending on the species), with margins that are entire, circular outline and convex elevation; the texture varies between soft and watery to cohesive and smooth ([1]). The colonies we had were tiny (1 mm), round, translucent, white/cream coloured, had raised elevation and a mucoid consistency.

However the colony yield was very high (we were able to form a lawn of colonies in the undiluted and 1/10 dilution quadrants), which is not typical for transformation. Based on this factor alone, that would indicate our cells were exceptionally electrocompetent OR that they were not transformed but happened to tolerate the PYE-CM plate. Along with the fact that the colonies formed much quicker than expected for transformed Caulobacter, which usually takes 4 - 6 days in Beth’s experience, we began to suspect that the bacteria was a different species (that either took up the plasmid or was already tolerant of the low Cm concentration of the plate).

Figure 1. Rep 1 transformation plate from Jun 4th experiment, lawn of growth/colonies is outlined in red.

Beth looked at the colonies from one of transformant plates under a light microscope and confirmed that the colonies were NOT Caulobacter. She informed us that it was a rod shape bacterium, similar to E. coli, which explains why the colonies grew so fast. She suggested checking whether the PYE medium used to recover the bacteria was contaminated or whether the electroporation was accidently done with E. coli instead of Caulobacter (as we also had stocks of competent DH5alpha right beside the competent CB2A). The medium that was used didn’t appear cloudy upon inspection, and we confirmed that the correct cells were used based on our lab notebook documentation.

Then Beth looked at one of our electrocompetent CB2A stocks under the microscope and verified that they were pure Caulobacter, so our competent cells were ruled out as a source of contamination. The cells had a distinctive crescent shape and were wriggling across the field of view. Many of the cells were in pre-divisional stage, characterized by a swarmer mother cell with an attached stalked cell (appearing as two crescent-shaped bacterium stuck together at their poles). An image from literature is provided below as reference.

Figure 2. Snippet of figure 2 from Hillson et al., 2007 depicting Caulobacter under phase-contrast microscopy.

Thus the contamination likely occurred during the electrotransformation procedure. Since the transformant plates yielded many colonies which formed fairly quickly, we hypothesize that the source of contamination would be from a reagent that could support something would have been growing for a while before we unsuspectingly used it for transformation. This could be either the PYE medium used to recover the bacteria or the PYE-CM agar plates used to plate the transformants (Beth said that plate contamination was less likely in her experience). We also suspect the electrotransformation cuvettes we reused could be another source of contamination.

So to pinpoint the true source(s) of contamination, we planned to make/order new materials in preparation for our next electrotransformation attempt.

Materials & Methods

We inspected the PYE broth medium used in the previous electrotransformation experiment and it didn’t appear to be contaminated (the solution looked clear with no signs of anything growing). However it is possible that the contaminating bacteria is hard to see so we made new medium.

As a test, we put the bottle in the 30C room and will check again in 1 - 2 days to see if anything grows.

Since we were concerned that the PYE agar used to make the previous plates could be another source of contamination, we also made new agar in this lab session.

Making new PYE medium (500 mL) + PYE-CM agar plates

Following the Version 1 recipe from PYE Medium Recipe.

Calculations for PYE-CM plates:

Stock Cm = 20mg/mL

We intend to make 2 plates so only need 50 mL (25 mL per plate).

C1V1 = C2V2

C1 = 20mg/mL = 20,000 ug/mL

V1 = x

C2 = 2ug/mL

V2 = 50 mL

x = 2 * 50 / 20,000 = 0.005 mL = 5 uL

Notes:

We noticed that the volume of the agar solution was ~50 mLs above 500 mL despite following the recipe. We may have to add less Milli-Q water next time.

To make the plates:

1. The PYE agar solution was prepared and 50 mL was removed into a sterile 100-mL beaker in the BSC.
2. 5 uL of 20mg/mL Cm stock was added to the 50 mL and swirled the beaker to mix.
3. Two plates were poured, let to solidify for 30 mins then stored in 4C (sealed with parafilm and wrapped in saran wrap).

Purpose

Start a subculture of WT UTEX and measure OD750 at hourly intervals to get a growth curve of UTEX, particularly its exponential phase. Data during the exponential phase will be used to determine the doubling time and growth rate of WT UTEX.

Materials & Methods

Materials

• Cuvettes
• Culture flask

Procedure

Experiment is conducted following this reference protocol: Generating a Growth Curve, except BG-11 medium is used as the diluent instead of PBS, therefore will blanked with it.

BG-11 blank is 0.001

OD750 of mother culture already 0.302 at 1x

Start culture with 125 mL medium, aim for OD 0.05

(125*0.05)/(0.302-0.05) = inoculate with 24.8 mL, then inoculate new flask with 20 µL culture in 100 mL medium and discard rest

Results

Note: UTEX colony was removed at 5:20pm and not placed back in the incubator until around 5:50pm. this means it was outside for ~30min, interrupting the incubation conditions. May have negatively impacted the growth curve.

after some reflection, several factors could have contributed to slow growth:

• UTEX appears to have a particularly long lag time - should this experiment be repeated, the culture should have been inocculated overnight.

• There could be strong substrate limitation (mainly CO2), since even though there is a lot of headspace and interfacial area in the flask, it is sealed off. Growth rate might be higher if cultured in a well plate with even more area per volume, or with CO2 sparging. In air, CO2 is 0.04 mol%, but [1] achieved the 2h doubling time at 3% CO2

• [1] used $OD_{730}$ instead of 750, it is worth seeing if there is a significant difference in OD.

• There could be insufficient light, and the temperature could be slightly higher, to achieve the theoretical doubling time of ~2h. Nannaphat (Patrik) Sukkasam had comments about growth curves and transformation

• He usually starts at OD 0.1, 0.1-0.5 are acceptable. This was the original plan, so now we have another reason not to start lower

• For transformation, cells should be taken in the middle of the exponential phase, and definitely not from the stationary phase - this is when a lot of the population is “young” and will be able to recover from electroporation.

• The reason growth curves are important is for the above, so transformation experiments also need to be timed accordingly.

• Kalen Dofher would be the contact for electroporation, since strains Patrik works with are naturally competent.

Growth Condition

This matrix will help dry lab as starting point for their optimization, and as their negative control.

Parameter	Value	Unit	Remarks
Temperature	29	ºC
% CO2	0.04	%	1
Media	BG-11	-
Media pH	N/A	-
Shaking rotation		rpm	2
Lighting	White light - 70	µmol photons·m−2·s−1	-
Culture Vessel Type	Erlenmeyer flask	-
Culture Vessel Volume	250	mL	3
Actual Culture Volume	125	mL	4

1: If unenriched, the atmosphere is 0.04% $CO_2$.

2: If not rotating, enter 0 rpm.

3: e.g. a 500 mL Erlenmeyer flask

4: e.g. 200 mL actual volume in a bigger Erlenmeyer flask

Growth curve

Graph 1: UTEX growth curve detailing the growth of culture over 125 hours measured at $OD_{750}$

Growth rate and doubling time

Doubling time of UTEX is ~24 h (similar to a mammalian cell). This is much higher than the expected ~2h doubling time for UTEX 2973.

Summary

Growth rate of UTEX 2973 is significantly slower than the expected growth rate of the strain reported in literature. This may be due to a number of culturing conditions including CO2 concentration, light concentration and temperature conditions. UTEX also appears to have a significantly low lag time so future growth curve experiments should start at a culture OD between 0.1-0.5.

Hand off

Send results to dry lab for optimising culture growth conditions. Repeat growth curve experiments starting at a higher OD and improved culture conditions to lower doubling time.

Purpose

Many previous attempts at restreaking UTEX have failed, despite following otherwise functional protocols. Using the same method, the purple colony streaked in 2025.06.09 Checking UTEX AB resistance + Culture Disposal Training + Restreaking & Culturing Purple Colony was very successful. Currently, only Wendy’s initial streaking attempt worked, alongside Pattarin’s. As such, we will attempt to determine the best method for streaking using a control plate and BG-11.

This experiment is to determine the optimal streaking method for UTEX.

Pictured: purple colony by date 11.06.2025 (day, month, year). As shown, it has shown to be healthy, despite being restreaked with the same method as the UTEX colonies that were unsuccessful. The current theory is that the streaking method is compatible with the purple colony (theorised to be E. coli), but not UTEX. Since the plating tool is hot after being flamed, cooling time will be tested as a factor.

Test various plating methods based on cooling time before plating.

Materials & Methods

Materials UTEX colony

BG-11 plate

Flame

Sterile loop

P200/20 sterile tip

Parafilm

Follow general plating protocol of 2025.06.09 Checking UTEX AB resistance + Culture Disposal Training + Restreaking & Culturing Purple Colony, with some changes listed below.

Results

Test	Procedure
1: Loop 0s	Cool by pressing hot loop against control agar. Immediately streak afterwards.
2: Loop 15s	Set loop aside and wait 15s before streaking.
3: Loop 30s	Set loop aside and wait 30+s (waited 35 in experiment) before streaking.
4: Pipette Tip	Take a sterile P200 tip and streak as you would with the loop.
5: Cooled, 15s	Cool by pressing hot loop against control agar. Wait 15s, then streak.
6: Control	No streaking, test for contamination.

Pictured: the initial plate (above) and the plate it was restreaked onto (below). Note below has the various tests written on it.

The original plate by Wendy was returned to the 4°C room, and the new plate was placed in the incubator. Note that a potential source of error is the original plate, as it was placed in the 4°C room. We have no experience with storing UTEX or cyanobacteria in this condition, and we know that UTEX does not fare well in glycerol stocks.

After 2 days:

No visible change

After 4 days:

Pictured: the aforementioned plate with large colonies seen on the “0s”, “30+s”, and “pipette tip” segments.

Test	Results after 4 days
1: Loop 0s	Large colonies
2: Loop 15s	Small, isolated colonies
3: Loop 30s	Thin colonies
4: Pipette Tip	Moderate colonies
5: Cooled, 15s	Small, isolated colonies
6: Control	Small or no colonies, appears to be none.

As seen, this is inconclusive about the plating method theory. The most effective seen is the “0s” plating method. This was the one believed to be used for all previous UTEX streaking. However, it proved to be completely fine in this experiment. The other methods are lesser or similar in effectiveness. Notably, the pipette tip method has been seemingly validated as one of the more effective methods.

Despite this, these results should be taken skeptically, as there are many factors potentially influencing these results. Poor streaking technique (as in the spreading part), which colony was taken, and the density of said colony may all impact streaking results. As such, we cannot conclusively say which streaking method is the best.

On the other hand, we can confirm that storing UTEX in the 4°C room is likely safe. As this plate was restreaked using the UTEX plate made by Wendy, and that was placed into the 4°C room, we can confirm that UTEX remains alive and viable during and after its placement into the room.

If this were to be redone, it would be preferable to have an E. coli control to see if the differences between species.

Summary

Results of the plating experiment are inconclusive, but we have a small confirmation that using the 4°C room for UTEX is effective and does not hamper restreaking attempts.

Hand off

The results of this experiment may be useful for continued work on UTEX (4°C viability).

Purpose

Run the digestion products from 2025.06.11 CB2A parent plasmids Restriction Digestion on a 1% agarose gel to visualize the digested plasmids.

Materials & Methods

Agarose Gel Electrophoresis

Making 1X TAE Buffer (500 mL recipe)

Component	Amount
50X TAE Buffer (on shelf G)	10 mL
Distilled or Milli-Q water (from the	490 mL

1. Measure out 10 mL mL of 50X TAE Buffer into a 500-mL glass bottle.
2. Top up to 500 mL with distilled/Milli-Q water to dilute the buffer to 1X concentration.
3. Mix by inverting the bottle 3 - 6 times.

Preparing gel

• Made 40 mL, 1% gel

• Use masking tape on gel casting mold

Preparing samples

• Digestion reaction products (tube # 2 - 5)

• Keep everything on ice

Lane	1	2	3	4	5	6	7	8	9	10
Sample tube ID		1	2	3	4	5	6	7	8	Blank
DNA	-	10 uL (tube 2)	10 uL (tube 3)	10 uL (tube 4)	10 uL (tube 5)	20 (tube 2)	20 (tube 3)	20 (tube 4)	20 (tube 5)	-
6x dye	-	2	2	2	2	4	4	4	4	4
Ladder	5	-	-	-	-	-	-	-	-	-
TOTAL		12	12	12	12	24	24	24	24	4

The samples were loaded in the lane order assigned in the table above.

Two sets of samples were created, one where 10 uL of reaction was used to run on agarose gel, and the other using 20 uL of reaction in case 10 uL did not yield enough DNA to be visualized on the gel.

• Since each digestion reaction was a final volume of 50 uL containing 1000 ng of DNA the concentration of the reaction product would be ~ 20 ng/uL.

• 10 uL * 20 ng/uL is 200 ng (which is more than enough to see on the gel)

Running gel

• Removing tape from mold and orientate it the correct way in running tray

• Pour 1X TAE buffer until the gel is just submerged (only need ~0.5 cm of buffer above the gel)

• Ran at 120V for ~45 mins.

Visualized on UV imager.

Results

Expected fragment sizes:

• pCB2A_Disp: 5846 bp
  • After digestion with BsaI and EcoRI: 3292 bp and 2554 bp
• pCB2A_Sec: 3970 bp
  • After digestion with BsaI and EcoRI: 2886 bp and 1084 bp

Snapgene simulation:

Summary

Plasmids were correct. May need to digest longer (maybe 30 mins), use less DNA and run the gel at a slower setting.

Hand off

The remaining digestion reaction (< 20 uL) was stored in the PCR tube box in -20C.

Purpose

To create a growth curve and determine the growth rate of WT CB2A in typical bench growing conditions.

Materials & Methods

Experiment was conducted following this reference protocol LP-20 Generating a Growth Curve, except PYE broth is used as the diluent and OD600 reference/standardization medium instead of PBS.

Setup Notes

June 12,2025 9:00PM

Based on the results of 2025.06.11 UTEX Growth Curve, and previous growth data from 2025.06.03 Making Electrocompetent CB2A cells (Training) and other experiments, we hypothesized that the CB2A culture would still be in lag phase for several hours before reaching exponential phase. Thus we decided to inoculate the culture the evening before the day of continuous measurements to let the culture run through lag phase overnight.

Using the preculture prepared in 2025.06.10 Testing antibiotic resistance of WT CB2A, we made 100ml of CB2A at 0.05A in a sterile 500mL flask (1st passage).

The OD600 reading for the preculture was 1.21 (measured by blanking the spectrophotometer with PYE).

• 1.21A * x = (0.05)(100ml)
• x = 4.1ml
  • volume of starter culture: 4.1ml
  • volume of PYE media: 95.9ml

The OD600 of the new culture was measured and it came out to be 0.07 (close enough to 0.05).

The culture was set to shake at 30°C overnight.

The preculture was stored at 4°C (can use as waste flask the next day).

June 13,2025 6:27 AM

Following the reference protocol, the initial measurement of PYE media was 0.002 and the overnight culture was 0.821. The culture is already in exponential phase at this point, so a new culture will be created using this culture (2nd passage).

For a 500mL flask we don’t want to exceed 25% of the flask’s volume (i.e., no more than 125 mL of culture).

Thus add 100 mL media + x amount of overnight culture for a final OD600 of 0.05 to start the growth curve.

$$OD_{inoc}V_{add}=OD_{start}(100 + V_{add})$$ $$V_{add}=\frac{100\text{ mL}\cdot OD_{start}}{OD_{inoc}-OD_{start}}$$

V_add = x

OD_inoc = 0.821

OD_start = 0.05

x = (100 * 0.05) / (0.821 - 0.05) = 6.485 mL

6.485 mL of the overnight culture (using P1000 pipette) was added to 100 mL of PYE in a sterile 500mL flask under the flame.

Overnight culture was left on bench for the whole day and measured OD at 9:55 pm:

0.915

June 13, 2025 6:55 AM

The machine was turned off, so reblanked with water and measured OD600 of PYE again.

OD_PYE = 0.03

OD_inoc = 0.793 (wanted to see if the machine would give a similar reading after being turned off, this reading is lower than the initial measurement but that could be due to cells settling at bottom of cuvette)

OD_start = 0.064, 0.063, 0.063 (took 3 replicate measurements to confirm machine precision)

Workflow

1. Take the culture flask out of shaker in 30ºC room.
2. Remove 1 mL from the culture into a clean cuvette under the flame.
   1. Shake/swirl the flask to resuspend the cells.
   2. Use P1000 to take a sample, then pipette up-and-down 2-3 times before dispensing into cuvette.
3. Measure OD600 on spectrophotometer (machine should be set to 600 nm).
   1. Wipe cuvette body with kimwipe.
   2. The program should be set to growth curve program, and blanked with water. Press “measure sample”.
   3. Record the number in the “C” column of the program in the “Measured OD” column of the spreadsheet (also include the time the measurement was taken, the culture time since the first measurement, and your initials).
4. Return the culture flask to the shaker (if you need to switch off the machine to put the flask back, remember to turn it back ON).
5. Dump the contents of the cuvette into the WASTE flask and rinse the cuvette with water at least 3x. Shake out as much water as you can.
6. Set the cuvette upside down on a 70% EtOH soaked paper towel to let it dry.
7. Repeat steps 1-6 for every measurement (approximately every hour).

Started first measurement at 8:15 am, and took measurements in 1 hr increments afterwards (9:15, 10:15, 11:15, 12:15, 1:15 …).

Once the OD is around 0.8 or above, you’ll likely need to dilute the sample with PYE media for the next measurement .

If you get to the machine and the growth curve program is gone you can go back by pressing this sequence of options:

Bio Test → use arrow keys to go down to Growth Curve → press enter → press the button corresponding to Run Test → you will need to reblank with dH2O then measure the PYE blank again (both of these are in parafilm sealed cuvettes at our bench, these can be reused so don’t dump out the contents and wash them).

Demos

1. Taking the flask out of shaker in 30°C

211d65dd-82be-80f4-b719-e5fae14033c7.mp4

2a. Taking a sample for OD600 measurement, part 1 (this step is split up because of filming purposes ).

211d65dd-82be-8065-8e36-f05af72f8064.mp4

2b. Taking a sample for OD600 measurement, part 2

211d65dd-82be-80a2-8649-cbabf38a1737.mp4

3. Reading OD600 of sample

211d65dd-82be-803d-980a-d309de5817a4.mp4

4. Returning flask to shaker

211d65dd-82be-8091-bf31-c8bf41fa7dda.mp4

5. Cleaning cuvettes

211d65dd-82be-80b2-8f11-fbbdd5fc0727.mp4

Results

Spreadsheet

%% START :: skip %%

%% END :: skip %%

Growth Condition

This matrix will help dry lab as starting point for their optimization, and as their negative control.

Parameter	Value	Unit	Remarks
Temperature		ºC
% CO2		%	1
Media	PYE	-
Media pH		-
Shaking rotation		rpm	2
Culture Vessel Type		-
Culture Vessel Volume		mL	3
Actual Culture Volume		mL	4

1: If unenriched, the atmosphere contains 0.04% CO2.

2: If not rotating, enter 0 rpm.

3: e.g. a 500 mL Erlenmeyer flask

4: e.g. 200 mL actual volume in a bigger Erlenmeyer flask

Graph

<Create a true OD vs. time AND log graph>

Growth rate and doubling time

(report the numbers)

Type here…

Purpose

A maintenance plate of WT CB2A colonies will be made, which can be used to easily create more liquid cultures in the future. This is the second attempt of this experiment, following the results of 2025.06.10 Testing antibiotic resistance of WT CB2A in which the first attempt produced a plate with too many colonies. To overcome this, we will use less glycerol stock when streaking with the inoculation loop.

Materials & Methods

• Sterile tips (P200)
• CB2A #2 glycerol stock (WT CB2A)
• Ice
• Agar plates:
  • 1x PYE
• Incubator at 30C

Under the flame:

1. Use a clean pipette tip to scratch the top of the glycerol stock.
2. Touch the tip near the edge of the plate to transfer majority of the cells onto the agar (if using glycerol stock, the cells will start thawing at the end of the tip and form a droplet). Then gently streak the cells down the plate in a zig-zag manner.
3. Flip the plates agar side up to let them dry for 1 min then wrap them in parafilm.
4. Incubate at 30C for 48 hours.

Results

Plate streaked and incubated at 9:45 pm.

Fig 1. CB2A maintenance plate before incubating at 30C on June 13, 2025.

There was a prominent growth of colonies along the streak line and many tiny colonies covered the plate. This is similar to the undesirable result of the first streaking attempt linked above, so we will need to reattempt this experiment.

Fig 2. CB2A maintenance plate after 36 hours of incubation at 30C.

Upon discussion with our instructor Antonio, we will implement these adjustments to the protocol:

• Use loops instead of pipette tips (either the reusable metal or plastic disposable ones)
  • And instead of scratching the top as we would typically do to inoculate liquid culture, we will try just touching/jabbing the top of the glycerol stock as very little bacteria is needed to streak plates for isolated colonies
• Serially dilute the sample by following the streaking pattern described in https://www.addgene.org/protocols/streak-plate/
• Alternatively, we can try spreading the sample all over the plate like in this tutorial video: https://www.youtube.com/watch?v=_r79BHaFf-c&ab_channel=HoweLab

Summary

This second attempt in streaking WT CB2A from glycerol stock to obtain isolated colonies was not successful as too many colonies grew on the plate, making it difficult to use for picking colonies to inoculate culture. This experiment will be reattempted with changes to the protocol.

Hand off

This plate was stored in 4C room for further analysis.

Purpose

Transform the following parts from the distribution kit to be used for agar art or to check transformation in CB2A. Unless otherwise stated, the parts will be resuspended from the 2025 distribution kit.

Materials & Methods

Heat Shock Transformation with a Thermocycler

SOC Medium Recipe

• PCR tubes

• SOC medium from -20°C

• 2025 Distribution kit plates 1 and 2, 2024 Distribution plate 1

  Name	Shorthand	Kit Plate	Well	Antibiotic
  mRFP1	R	1	O3	Cm
  tannenRFP	O	1	I3	Cm
  sarcGFP	G	2	I23	Amp
  aeBlue	B	1	O1	Cm
  pJUMP25-1A (2024)	25	1	G10	Kan
  pJUMP24-1A	24	1	E10	Kan

Results

The above parts were resuspended, transformed, and plated onto their corresponding agar plates with antibiotic.

100 mL SOC medium was prepared and autoclaved. The recipe is written up in SOC Medium Recipe.

@ June 16, 2025 Upon inspection, plates appear contaminated, having widespread growth instead of individual white colonies.

Hand off

• Come in evening or morning to pick colonies.
  • not done due to contamination
• Redo, see 2025.06.16 Transforming Additional Distribution Kit Parts Redo + SOC Medium Aliquots

Purpose

• Create a WT CB2A maintenance plate from glycerol stock with changes to the streaking protocol as described in 2025.06.13 CB2A maintenance plate attempt 2.
• Redo test of Cm antibiotic susceptibility of the WT CB2A and competent CB2A cells on PYE-Cm plates.

Based on the results of 2025.06.10 Testing antibiotic resistance of WT CB2A, the WT CB2A seems resistant to low concentration (2 ug/mL) of chloramphenicol (Cm) which is not expected. Since the previous plates were discarded, the experiment will be redone so we can analyse the colonies under light microscopy and determine whether the colonies observed were Caulobacter.

Additionally, while we confirmed that the competent CB2A cells we created were not a source of contamination in our first electrotransformation attempt (see 2025.06.11 Troubleshooting CB2A Electrotransformation Remaking PYE Medium and PYE-Cm Agar Plates), we would like to test the Cm susceptibility of these cells to ensure that they would not grow on PYE-Cm plates used for colony selection. If they do grow on Cm-plates, this would lead to false positives in our transformation experiments and thus the stock would have to be discarded.

Materials & Methods

CB2A maintenance plate + PYE-Cm test plates

Materials

• EC CB2A (electrocompetent CB2A cells) from -80C
• CB2A #2 glycerol stock (WT CB2A) from -80C
• Ice
• Agar plates:
  • 2x PYE
  • 2x PYE-Cm (2ug/mL)
  • 2x PYE-Cm (20ug/mL)
• Incubator at 30C
• Inoculation loops
• Bunsen burner

Procedure

Under the flame:

1. Use a sterile loop to touch/jab at the top of the glycerol stock. OR Dip the loop into the thawed competent cells.
2. Touch the tip near the edge of the plate then start to drag the loop along the surface of the agar in a zig zag motion to spread the bacteria in about a third of the plate.
3. Using a freshly sterilized loop, drag through streak #1 and spread the bacteria over the 2nd un streak portion of the plate (leaving enough room for a final streak).
4. Using another freshly sterilized loop, drag through streak #2 to create streak #3.
5. Flip the plates agar side up to let them dry for 1 min then wrap them in parafilm.
6. Incubate at 30C for 48 hours.

Streaking pattern:

Plates created:

Name (source of bacteria - condition)	Shorthand	Medium type	Cells streaked	Expected result after 2 - 3 days
WT CB2A maintenance plate (attempt 3)	maintenance	PYE (no antibiotic)	WT CB2A	Many isolated colonies

*This plate also acts as the positive control for the WT CB2A plates, equivalent to “Electrocompetent CB2A Pos Ctrl - No Cm” | WT CB2A Neg Ctrl - Low [Cm] | WT-Low [Cm] | PYE-Cm (2ug/mL) | WT CB2A | No colonies, WT CB2A shouldn’t have Cm resistance | WT CB2A Neg Ctrl - High [Cm] | WT-High [Cm] | PYE-Cm (20ug/mL) | WT CB2A | No colonies, WT CB2A shouldn’t have Cm resistance | Electrocompetent CB2A Pos Ctrl - No Cm | EC-No Cm | PYE (no antibiotic) | EC CB2A | Many colonies (not sure if they’ll be isolated because the inoculant wasn’t spread as typically would for transformation) | competent CB2A Neg Ctrl - Low [Cm] | EC-Low [Cm] | PYE-Cm (2ug/mL) | EC CB2A | No colonies, EC CB2A shouldn’t have Cm resistance | competent CB2A Neg Ctrl - High [Cm] | EC-High [Cm] | PYE-Cm (20ug/mL) | EC CB2A | No colonies, EC CB2A shouldn’t have Cm resistance

The plates were placed in 30C room at 11:00 pm.

Fig 3. CB2A maintenance plate and the PYE/PYE-Cm test plates were incubated at 30C.

Contaminated PYE

While conducting this experiment, we noticed that the PYE created on 2025.06.11 Troubleshooting CB2A Electrotransformation Remaking PYE Medium and PYE-Cm Agar Plates was slightly cloudy, so it was likely contaminated during the 2025.06.13 CB2A Growth Curve experiment (probably during dilution steps). This bottle will be treated with 10% bleach the following day.

In the future, 50 mL aliquots of PYE can be prepared for experiment that require frequent use of the media and the stock bottle can be stored in 4C.

Results

Check plates

Both EC-No cm and EC-Low [Cm] had some growth along the first streak line after 23 hrs of incubation at 30C. This is a bit faster than expected as colonies should appear within 2 - 3 days, and the presence of colonies on the Low [Cm] plate indicates that what we presume to be electrocompetent CB2A colonies are resistant to Cm.

There was no growth on any other plates. In terms of the CB2A maintenance plate, this result is different from the previous two attempts (where a lawn of growth appeared within a day) and we expect to see colonies on this plate in the next 1 - 2 days. However, we also observed a light dusting of white dots along the plate (doesn’t seem to be colonies but not all of the plates have this).

All the plates were returned to the 30C room.

CB2A maintenance plate and the PYE/PYE-Cm test plates after 23 hrs incubation at 30C.

Close up of EC-No Cm plate after 23 hrs incubation at 30C.

Close up of EC-Low [Cm] plate after 23 hrs incubation at 30C.

July 1st:

• Maintenance plate had growth along the side of the plate → plate is contaminated

• EC-High [Cm] → not contaminated Beth to look at some plates under the microscope to confirm whether they are Caulobacter species.

• WT-Low [Cm] → contaminated with rod-like species

• EC-Low [Cm] → contaminated with rod-like species

• EC-No Cm → mainly CB2A but also some rod-like species

Hand off

• Remake PYE plates

Purpose

Redo 2025.06.13 Transforming Additional Distribution Kit Parts since the plates appear contaminated. Instead of using the thermocycler, out of concern that the cell volume is too small, the water bath will instead be used for transformation, where a larger competent cell volume can be used. The purple construct from 2025.06.11 Miniprepping purple construct will also be transformed to be sure it is in fact E. coli carrying the plasmid.

Name	Shorthand	Kit Plate	Well	Antibiotic
mRFP1	R	1	O3	Cm
tannenRFP	O	1	I3	Cm
sarcGFP	G	2	I23	Amp
aeBlue	B	1	O1	Cm
pJUMP25-1A (2024)	25	1	G10	Kan
pJUMP24-1A	24	1	E10	Kan

Materials & Methods

Heat Shock Transformation with a Water Bath

• Added 250 uL SOC medium instead of 950.

• 6+10 sterile microcentrifuge tubes

• competent cells

• 3 petri dishes

• x2 LB/Kan plates

• x1 LB/Amp plate

Make new Cm agar plates for today since the previous ones appear contaminated (cloudy) and would be difficult to pick colonies from.

• LB agar
• Cm stock
• 3 petri dishes

Results

10x 1 mL aliquots of SOC medium were made and placed in the -20C freezer box.

Transformations were redone, in addition with the purple construct, totalling 7 plates.

only 2 plates, R and purple, showed a distinct colony, however the purple one is not purple. Will check again in the evening.

Both 24 and 25 had a small colony, R had no new colonies and there were no colonies on the other plates. Two new colonies formed on the purple plate, both of which had a faint purple colour visible by eye.

Summary

See above

Hand off

Check for colonies and pick if transformations are successful.

Purpose

To test what temperature an eppendorf tube is able to withstand in an oven. The subsequent dry cell weight experiment for CB2A will be altered based on these results, where the oven will be used to dry out the pellets in microcentrifuge tubes.

Materials & Methods

• 6 Eppendorf tubes
• Tinfoil
• Oven

Generating a Dry Cell Weight Curve

• Label the eppendorf tubes (1,2,3…).
• Weigh 6 eppendorf tubes and record their initial masses.
• Fold 3 pieces of tinfoil into “weigh boats”.
• Place 3 eppendorfs (tube #1,2,3,) into the oven rack and 3 eppendorf tubes (#4,5,6) into the tinfoil boats.
• Set the oven to 90˚C and heat the eppendorf tubes for 2 hours.
• Let the eppendorf cool, until cool to touch.
• Weigh the 6 eppendorf tubes and record their final masses.

Results

Table 1. Initial weight of eppendorf.

Tube #	Replicate 1	Replicate 2	Replicate 3
1	1.0519g	1.0520g	1.0519g
2	1.0818g	1.0819g	1.0820g
3	1.0371g	1.0375g	1.0376g
4	1.0574g	1.0578g	1.0576g
5	1.0547g	1.0549g	1.0549g
6	1.0588g	1.0587g	1.0587g

Table 2. Final weight of eppendorf.

Tube #	Replicate 1	Replicate 2	Replicate 3
1	1.0514g	1.0517g	1.0516g
2	1.0819g	1.0814g	1.0814g
3	1.0371g	1.0372g	1.0373g
4	1.0574g	1.0575g	1.0579g
5	1.0545g	1.0544g	1.0545g
6	1.0585g	1.0584g	1.0585g

Table 3. Change in weight of eppendorfs.

Tube #	Avg Initial Weight	Avg Final Weight	Change in weight
1	1.0519	1.0517	-0.0002
2	1.0819	1.0816	-0.0003
3	1.0374	1.0372	-0.0002
4	1.0576	1.0576	-0.0000
5	1.0548	1.0545	-0.0003
6	1.0587	1.0585	-0.0002

Summary

There was negligible loss of mass. However, we should conduct this experiment for the required 24 hour timespan to test for further melting. Additionally, it was observed that the eppendorf tubes were still cool to the touch after 1.5hrs of heating, so there is a chance that the oven is not well insulated.

Purpose

Extracting plasmid pCB2A_Disp plasmid and pCB2A_Sec from dH5α for digest reaction and agarose gel.

Materials & Methods

Minicultures (June 16th)

Materials

• pCB2A_Disp DH5α stock plate (stored in 4C)
• pCB2A_Sec DH5α stock plate (stored in 4C)
• LB broth
• Sterile pipette tips
• Sterile glass culture tubes
• Cm (20 mg/mL) stock (stored in -20C)
• Bunsen burner
• Shaker at 37C

Procedure

• Both Culture plates and four sterile culture tubes were labeled for picking and containing proper colonies.
• Into each labeled tube, 4 mL of LB broth was added.
• 4 µL of Chloramphenicol antibiotic were added to each of the 4 mL LB broth in tubes to selectively grow plasmid-containing bacteria. Before adding Cm to the culture, the eppendorf tube was vortexed to ensure uniform distribution of Cm concentration.
• Colonies were picked from previously prepared LB plates using a sterile pipette tip.
• The picked colonies were then inoculated into the prepared 4 mL LB broth solutions with Cm.
• The tubes were incubated at 37°C with shaking overnight to allow for bacterial growth and plasmid amplification.
• The resulting minicultures will be used for subsequent miniprep procedures to extract the pCB2A_Disp and pCB2A_Sec plasmids.

Notes:

Since we confirmed that pCB2A_Disp and pCB2A_Sec colonies contained the correct plasmid in our digestion experiment, we could’ve picked from them again but since the plates were not incubated in 37C to regenerate the colonies in advance there wasn’t much of those colonies left to pick. Instead we selected two new colonies, labelled #3 and #4 to pick.

For pCB2A_Sec, we confirmed the original #1 and #2 colonies contained the right plasmid in the same digestion experiment but our original miniprep yield was low so we will pick new colonies (also labelled #3 and #4) as well.

All tubes were placed on a shaker in the 37C room at 7:00 pm.

Note: p4a #3 tube was spilled so the culture was remade in a new tube and set to shake at 11:00 pm

And the stock plates were incubated at 37C to generate more colonies (they will be moved back to 4C the next day).

Miniprep (Talha and Yuki)

Plasmid Mini-Preparation

• GeneJET plasmid miniprep kit
• x2 microcentrifuge tubes
• x1collection tube NanoDrop Spectrophotometer

Tube	Construct (Backbone plasmid)	Colony #
1	pCB2A_Disp	3
2	pCB2A_Disp	4
3	pCB2A_Sec	3
4	pCB2A_Sec	4

Results

Construct Name	Concentration (ng/ul)	colony	A260/A280	A260/A230
pCB2A_Disp	12.7	3	1.92	2.12
pCB2A_Disp	197.7	4	1.89	2.29
pCB2A_Sec	18.9	3	1.73	1.91
pCB2A_Sec	36.0	4	1.83	2.42

pCB2A_Disp #3 was cultured late and pCB2A_Sec #3 300μl was discarded, which may have affected final results.

DNA stored in mini fridge inside white iGEM 2025 box.

Purpose

Plates from 2025.06.16 Transforming Additional Distribution Kit Parts Redo + SOC Medium Aliquots were slow to show colonies, but by had grown enough to be picked, although a few look like possible contamination. The plates supposedly with JUMP plasmids had several colonies but were not fluorescent.

Materials & Methods

• 6 glass culture tubes
• LB medium
• Cm and Kan stock
• Sterile pipette tips

Results

Colonies were picked and culture tubes were placed in the 37C room shaker.

The Cm plates poured in 2025.06.16 Transforming Additional Distribution Kit Parts Redo + SOC Medium Aliquots had a lot of particulates which made it hard to see colones. In the future poorly melted agar like that should not be used.

The Amp plate did not have any bacterial colonies but instead a mold colony. It was discarded.

Summary

See above.

Hand off

Return to examine morning, especially if the JUMP plasmid colonies have a yellow-ish colour or are fluorescent. If all are sufficiently cloudy, either place them in the 4C room or miniprep.

Purpose

Set up CB2A culture for dry weight cell experiment.

Materials & Methods

• Sterile tips (P200)
• Sterile 250-mL flask
• PYE liquid medium
• CB2A #2 glycerol stock (WT CB2A)
• Ice
• Shaker & incubator at 30C

Under the flame:

1. Pour ~50 mL of PYE into a sterile 250-mL flask.
2. Using a clean pipette tip (attach to P200 pipette), lightly scratch the top of the glycerol stock until you get a small amount of cells at the end of the tip (about half of a sesame seed).
3. Swirling the tip in the flask and gently pipette up and down to inoculate the PYE medium.
4. Shake (~200 rpm) at 30C and incubate for 1 - 2 days.

Results

Incubated the culture at 30C around .

Summary

See above.

Hand off

Use the culture for the dry cell weight curve experiment starting on evening.

Didn’t do curve so discarded culture.

Purpose

To assemble the fragments of DNA isolated and quantified after Miniprep and Nanodrop through Golden Gate Assembly to transform BL21 E. coli.

Materials & Methods

Parts from plate map:

• BL21-D_LacI

• BL21-E_T7pro_MCS_His Backbone plasmid, either:

• ‘pdest’ = undiluted

• ‘pdes 100’ = diluted to 100ng/µL

Resuspension guidelines from twist:

Materials & Methods

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
• Pipette tips

Resuspension of DNA from Twist Plate

Materials

• IDTE pH 7.5 (1X TE solution) → make aliquot
• Twist Plate
• Ice
• Centrifuge with plate adaptors
• PCR tubes

Procedure

1. For resuspension, briefly centrifuge the tube or plate before opening and resuspend in nuclease free Tris-EDTA (TE) buffer, pH 8.0 or 10 mM Tris-HCl, pH 8.0 to the desired concentration.
2. A concentration of at least 10 ng/μL is recommended for the stock dilution, but the optimal concentration will need to be determined based on your desired application.
3. Prepare aliquots of the stock dilution and separate working aliquots to limit chances of contamination and to reduce the number of freeze/thaw cycles. Use working aliquots as soon as possible after preparation and minimize exposure to high temperatures.

Each well contains 1000 ng.

To make a final concentration of 10 ng/µL, resuspend each well with 100 µL TE buffer.

• 12 wells * 100 µL = 1200 µL → take out 1200 µL TE to use instead of directly from stock (have a little excess) Then split each resuspension into 25 µL aliquots, so 4 PCR tubes per well.

12 *4 = 48 PCR tubes

Well Location	DNA part name	PCR tube label	Final concentration
A1	SazCA_co_BL21	CC1	10ng/µl
B1	SazCA_co_CB2A	CC2	10ng/µl
C1	BtCAII_co_CB2A	CC3	10ng/µl
D1	HpCA_co_BL21	CC4	10ng/µl
E1	BL21-C_T7term_CmR	CC5	10ng/µl
F1	BhCA_co_CB2A	CC6	10ng/µl
G1	BL21-D_LacI	CC7	10ng/µl
H1	BtCAII_co_BL21	CC8	10ng/µl
A2	HpCA_co_CB2A	CC9	10ng/µl
B2	BL21-E_T7pro_MCS_His	CC10	10ng/µl
C2	BL21-B_pelB_INPN-Re	CC11	10ng/µl
D2	BhCA_co_BL21	CC12	10ng/µl

Golden Gate of pBL21-Int (pLiN)

Golden Gate Assembly with NEBridge Ligase Master Mix

Parts to use:

Name	Length (bp)	Mass (ng) for 0.05 pmol	Concentration (ng/µl)	Volume (µL)
pDest 100	2723	83.86	103.8	0.81
BL21-E_T7pro_MCS_His	594	18.30	10 (1000ng/100ul)	1.83
BL21-D_LacI	1255	38.65	10 (1000ng/100ul)	3.87

Reaction mix:

Component	Volume (µL)
10x T4 Ligase Buffer (03/23)	5
T4 DNA Ligase (02/25)	0.5
pDest 100	0.81 (round to 0.8)
CC7	3.87 (round up to 3.9)
CC10	1.83 (round down to 1.8)
BsaI-HFv2 (03/23 exp)	01
water	1.99 (round up to 2 uL)
Total	15

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 # Results

Resuspended parts stored in PCR box in -20C freezer.

The pLiN assembly did not include fragment BL21-C_T7term_CmR so the construct is incorrect.

Purpose

To resuspend and prepare DNA oligos from Twist, GenScript, and IDT plates/tubes for downstream cloning and transformation experiments.

Materials & Methods

Resuspending dried oligos

Resuspending Dried DNA

Twist UTEX plate

• each well contained 1000 ng DNA
• resuspended with 20 µL TE to 50 ng/µL
• transferred to PCR tubes

Name	Well Location	Volume (uL)
1_NS1_US_700_gg	A1	20
1_VCBS_US-SpecR_TU_gg	B1	20
4_KanR_TU_gg	C1	20
5_NS1_DS_700-bom_gg	D1	20
5_VCBS_DS_768-bom_gg	E1	20
SapI-tag-VCBS_C-term_gg	F1	20
sfGFP-term_gg	G1	20
BhCA_UTEX_gg	H1	20
BtCAII_UTEX_gg	A2	20
HpCA_UTEX_gg	B2	20
SazCA_UTEX_gg	C2	20

GenScript plate

• each well contained 500 ng DNA
• resuspended with 10 µL TE to 50 ng/µL
• transferred to PCR tubes

Name	Well Location	Volume (uL)
0_pANS_RFP_1	A1	10
0_pANS_RFP_2	B1	10
0_pANS_RFP_3	C1	10
0_pANS_RFP_4	D1	10
0_pBR322_ori+sepT2_gg	E1	10
3_VCBS_Myc_sfGFP_gg	F1	10

IDT plate

• each well contained 5 nmol (per primer in pair)
• resuspended with 50 µL TE to 100 µM per primer (200 µM per pair)
• diluted aliquots prepared at 10 µM per primer (20 µM per pair)
  • generics tube strip
  • VCBS cPCR strip
  • NS1 cPCR strip
  • UTEX misc strip

Pair name	Well	PCR tube strip name	Label	Volume (μl)
twist	A1	generics	twist	50
twist	B1	generics	twist	50
genscript	C1	generics	GS	50
M13	E1	generics	M13	50
M13	F1	generics	M13	50
M13	G1	generics	M13	50
VCBS locus	A3	VCBS cPCR	locus	50
VCBS HR US	B3	VCBS cPCR	HR US	50
VCBS HR DS	C3	VCBS cPCR	HR DS	50
VCBS ori US	D3	VCBS cPCR	ori US	50
VCBS ori DS	E3	VCBS cPCR	ori DS	50
NS1 locus	A4	NS1 cPCR	locus	50
NS1 HR US	B4	NS1 cPCR	HR US	50
NS1 HR DS	C4	NS1 cPCR	HR DS	50
NS1 ori US	D4	NS1 cPCR	ori US	50
NS1 ori DS	E4	NS1 cPCR	ori DS	50
BhCA UTEX	A6	UTEX misc	BhCA	50
BtCAII UTEX	B6	UTEX misc	BtCAII	50
HpCA UTEX	C6	UTEX misc	HpCA	50
SazCA UTEX	D6	UTEX misc	SazCA	50
pANS 1	H3	UTEX misc	pANS 1	50
pANS 2	H4	UTEX misc	pANS 2	50
pANS 3	H5	UTEX misc	pANS 3	50
pANS 4	H6	UTEX misc	pANS 4	50

IDT tubes

Name	Label	Amount (ng)	Volume	Storage concentration (ng/uL)
2_J23107-RBS_gg	UTEX prom LO	250	10	25
2_J23119-RBS_gg	UTEX prom Med	250	10	25
2_Pcpc560-RBS_gg		500	10	25
1_VCBS_US_short_gg		250	10	25
BL21-A_T7pro_lacO	BL21-A	250	10	25
CB2A_MCS_v2	MCS_v2	250	10	25
CB2A_prsaA	prsaA	250	10	25

Results

mRFP1 CDS plasmid miniprep

• Yield: 8.2 ng/µL

• 260/280 ratio: 1.75

• 260/230 ratio: 0.85

• likely due to new resuspension buffer lacking RNase A resulting in low yield and contamination

• Sample discarded. Will repeat with corrected buffer

Oligo resuspension

• All Twist, GenScript, and IDT plates (@ June 20, 2025) successfully resuspended.

• Stocks stored at 50 ng/µL (plasmids) and 100 µM (primers).

• Working aliquots prepared at 10 µM (primers).

Purpose

To test our newly received DNA oligos, we will be checking their plasmid maps to find two enzymatic restriction sites. These, once used to cut the DNA, will result in distinct parts of known length. This will go into a gel electrophoresis, which will allow us to check if the experimental fragment lengths are the correct ones.

We are validating our DNA via double digestion into gel electrophoresis. Success will be confirmed by observing the expected banding pattern at the predicted b

The plasmids we will be digesting today:

AF_lacZ_pDest

CB2A Display Parent plasmid

CB2A Secretion Parent plasmid

Materials & Methods

Lane	Plasmid	Enzyme(s)
MW	Ladder	N/A
1	pDest - digested	BsaI
2	CB2A Secretion Parent plasmid - digested	BsaI+EcoRI
3	CB2A Secretion Parent plasmid - digested	BsaI+EcoRI
4	CB2A Display Parent plasmid - digested	BsaI+EcoRI
5	CB2A Display Parent plasmid - digested	BsaI+EcoRI
6	CB2A Display Parent plasmid (+)	N/A
7	CB2A Secretion Parent plasmid (+)	n/a
8	CB2A Secretion Parent plasmid (+)	n/a

Pictured: simulated gel scheme with numbering. We should see something similar to this.

From protocol LP-30 Restriction Digest with NEB Enzymes, we plan on using ~ 1 µg of DNA for each. However, due to low concentration, we aim for 500 ng, which should still be visible on gel.

DNA	Conc	Amount we’re digesting	Mass	Using?
pDest	103.8 ng/µL	4.82 µL	500 ng	Y
CB2A Display Parent plasmid #3	12.7 ng/µL	39.37 µL	500 ng	N, conc. low
CB2A Display Parent plasmid #4	- 18.6ng/µl (10x diluted)	26.88 µL	500 ng	Y
CB2A Secretion Parent plasmid #3	18.9 ng/µL	26.46 µL	500 ng	Y
CB2A Secretion Parent plasmid #4	36.0 ng/µL	13.89 µL	500 ng	Y

Required volumes - total should be ~50 µL for each.

Component	Tube 1 pDest	Tube 2 #4 CB2A Display Parent plasmid	Tube 3 #3 CB2A Secretion Parent plasmid	Tube 4 #4 CB2A Secretion Parent plasmid	Tube 5 pDest (+)	Tube 6 #4 CB2A Display Parent plasmid (+)	Tube 7 #4 CB2A Secretion Parent plasmid (+)	Tube 8 (- ctrl)
Water	39.2 µL	17.1 µl	16.5 µL	29.1 µL	40.2 µL	18.1 µl	31.1 µL	43 µL
DNA	4.8 µL	26.9 µL	26.5 µL	13.9 µL	4.8 µL	26.9 µL	13.9 µL	-
Buffer	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart	5 µL rCutSmart
BsaI	1 µL	1 µL	1 µL	1 µL	-	-	-	1 µL
EcorI	-	1 µL	1 µL	1 µL	-	-	-	1 µL
TOTAL	50 µl	50 µl	50 µL	50 µL	50 µL	50 µL	50 µL	50 µL

Add largest first!

Note: Tube 2 has a very low concentration, we likely won’t be using this recipe. Use a negative control instead, replacing tube 2. This will check for contamination of DNA.

Tube 8 (-) control	Vol
Water	43 µL
DNA	-
Buffer	5 µL
EcorI	1 µL
BsaI	1 µL

Set up thermocycler:

• 37°C - infinite (will take out reaction then switch to next one)
  • ran reaction for 15 mins
• 80°C - 20 mins
  • only deactivated for 2 mins

Component	Tube 1 pDest	Tube 2 #4 CB2A Display Parent plasmid	Tube 3 #3 CB2A Secretion Parent plasmid	Tube 4 #4 CB2A Secretion Parent plasmid	Tube 5 pDest (+)	Tube 6 #4 CB2A Display Parent plasmid (+)	Tube 7 #4 CB2A Secretion Parent plasmid (+)	Tube 8 (- ctrl)
Rxn	10 µl	10 µl	10 µl	10 µl	10 µl	10 µl	10 µl	10 µl
6X dye	2 µl	2 µl	2 µl	2 µl	2 µl	2 µl	2 µl	2 µl

Notes for gel: Wells 7 and 8 were redone. Should have a minimal impact on seeing the dyes.

Well 2 was redone in well 10. The expected signature for well 2 should be observed in well 10 instead.

Results

Prep: Plasmid CB2A Display Parent plasmid from colony #4 was quantified by nanodrop, as its concentration was not known at the start of this experiment. We found that its concentration was 18.6 ng/µL after a 10x dilution.

Image here:

Lane	Description	Volume loaded (not including dye)	Digested with	Expected bands
1	DNA Ladder	5 µL	N/A	-
2	pDest (100 diluted)	10 µL	BsaI	2106 and 617 bp
3	CB2A Display Parent plasmid #4	10 µL	BsaI + EcoRI	3292 and 2554 bp
4	CB2A Secretion Parent plasmid #3	10 µL	BsaI + EcoRI	2886 and 1084 bp
5	CB2A Secretion Parent plasmid #4	10 µL	BsaI + EcoRI	2886 and 1084 bp
6	pDest	10 µL	None, positive ctrl	2723 bp SUPERCOILED
7	CB2A Display Parent plasmid #4	10 µL	None, positive ctrl	5846 SUPERCOILED
8	CB2A Secretion Parent plasmid #4	10 µL	None, positive ctrl	3970 SUPERCOILED
9	None	10 µL	BsaI + EcoRi, negative ctrl	None
10	pDest	10 µL	BsaI	(Redo of Lane 1 because the sample wasn’t loaded very well)

Lanes 3, 4, 5 and 8 yielded expected results (the additional bands that differ from the simulated gel correspond to the expected size of undigested plasmid DNA). All the lanes containing pDest (lanes 2 and 10) did not show any bands. Given that the other lanes showed bands, we can confirm the UV spectrophotometer is working so we suspect the following reasons for the negative result:

• No DNA was loaded (error)
• Nuclease contamination
• ‘pDest 100’ tube did not contain the expected concentration of DNA written on the tube

Summary

Double digestion and gel electrophoresis of CB2A Secretion Parent plasmid and CB2A Display Parent plasmid succesfully corresponded with expected DNA lengths, thus we validated the DNA of these plasmids for downstream assemblies. However, there were no bands in pDest lanes which could have resulted from an error in DNA loading, nuclease contamination and difference between the reported concentration of pDest DNA on the tube compared to the actual concentration of pDest.

Hand off

We will use the backbones for future assemblies of CA constructs. Repeat double digestion & gel electrophoresis for pDest.

Purpose

To provide hands-on practice for team members with essential molecular biology techniques, including: Golden Gate Assembly and Bacterial Transformation. As well as to provide our Human Practice team with “Colourful colonies” to introduce students to the basics of synthetic biology by showing how DNA can be inserted into bacteria.

Additionally, attempt Golden gate assembly for pBL21-Int (pLiN).

Materials & Methods

Procedure Heat Shock Transformation with a Thermocycler.

Plasmid DNA from 2025 distribution kits for creating colourful colonies:

Name	Shorthand	Kit Plate	Well	Antibiotic
mRFP1	R	1	O3	Cm
tannenRFP	O	1	G3	Cm
sarcGFP	G	2	I23	Amp
aeBlue	B	1	O1	Cm

DNA (plasmid)	Transformation tube name	Media - Antibiotic	Plate Name
pBL21-Int GG rxn	pBL21-Intra	LB-Cm (20 ug/mL) and if there’s enough plates, do a LB-Amp (100 ug/mL) one because this construct is based off of pDest plasmid which confers Amp resistance
BL21-Intra
mRFP1	mRFP1	LB-Cm (20 ug/mL)	R
tannenRFP	tannenRFP	LB-Cm (20 ug/mL)	O
sarcGFP	sarcGFP	LB-Amp (100 ug/mL)	G
aeBlue	aeBlue	LB-Cm (20 ug/mL)	B

Notes:

• Used 2025 distribution plates (the DNA was already resuspended from last time), let it sit on the bench to thaw
• Since there was a mistake with tannenRFP from last transformation (I3 instead of G3), resuspended G3 with 10 uL nuclease free water (waiting 2 mins after adding water and pipetting up and down)
• 2 uL of DNA to 9uL of cells for every tube (even the distribution kit ones, although igem recommends 1 uL usually)
• BL21 rxn was from 2025.06.18 Resuspension of Twist fragments + Golden Gate for pLiN, did 5 mins of 60C on thermocycler since these rxn product was stored for a while (and also probably thawed during the freezer failure) as recommended by this:
• Only heat shock at 42C for 30sec (mistake, should be 45sec)
• Added 100uL soc (the tube is in the container labelled “iGEM media & solutions” in 4C room)

Results

Plate Name	July 22, 1:20 PM (16 hrs)
BL21-Intra (Cm)	No colonies
BL21-Intra (Amp)	No colonies
R	4 colonies
O	2 colonies
G	1 colony
B	No colonies

Checked (~16 hrs incubation):

• All the distribution part plates yielded colonies (R, O, G), except for B
• Colonies formed were white, opaque circular and looked like E. coli

Figure 1. tannenRFP LB-Cm (20 ug/mL) plate with colonies labelled.

Colonies were picked and inoculated into 3 mL of TB broth with respective antibiotics

R/O/G tubes were cloudy, purple tube was not- cloudy tubes stored at 4C

• tubes were then miniprepped (following Plasmid Mini-Preparation )

DNA (plasmid)	Plate Name	Concentration	260/280	260/230
mRFP1	R	87.4	1.90	1.54
tannenRFP	O	54.7	1.97	2.22
sarcGFP	G	46.9	1.91	2.08

Purpose

We have just received our UTEX shuttle vector backbone, pANS, as a set of fragments. We must PCR the fragments to get more of them, then we will be using Gibson Assembly to create the complete backbone plasmid.

Materials & Methods

PCR amplification of fragments

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

Depending on the NEB polymerase/kit you are using, gather the following and keep on ice:

If using Q5 High-Fidelity 2X Master Mix (NEB M0492S or M0492L):

• Deoxynucleotide (dNTP) mix
• Q5 High-Fidelity 2X Master Mix

Table 1: Fragments used in the experiment

Name	Primer name	Annealing Temp (°C)	PCR strip	Label
pANS_1	pANS_1	61	Genscript	A1
pANS_2	pANS_2	60	Genscript	B1
pANS_3	pANS_3	62	Genscript	C1
pANS_4	pANS_4	61	Genscript	D1

Using protocol from Polymerase Chain Reaction (PCR) with NEB Q5 High-Fidelity 2X Master Mix

1. 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. See LP-24 Hallam Lab Thermocycler for setting up the machine. 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	10 seconds
   	Annealing	72	30 seconds
   	Extension	72	20 seconds
   Final Extension		72	2 minutes
   Hold		4	Infinity

2. Calculate how much of each fragment should be added such that there is 10 ng of template DNA in the reaction. Genscript fragments are resuspended to 50 ng/µL per 2025.06.20 Resuspending plates → need 0.2 µL per fragment.

3. Make the following tubes for a PCR reaction:

   Component	pANS 1	pANS 2	pANS 3	pANS 4
   Q5 2x master mix (µL)	12.5	12.5	12.5	12.5
   10 uM primer pair mix (µL)	2.5	2.5	2.5	2.5
   DNA fragment (µL)	0.2	0.2	0.2	0.2
   Nuclease-free water (µL)	9.8	9.8	9.8	9.8
   Total (µL)	25	25	25	25

4. Place tubes into thermocycler and run cycle.

From https://www.neb.com/en-ca/protocols/2014/11/26/nebuilder-hifi-dna-assembly-reaction-protocol?srsltid=AfmBOoqotwpUWHXDP6y56bD8gkQFkWW-ko2IX2vPX8lN43LIHs4NhmNB

Gibson assembly protocol:

Components	4—6 Fragment Assembly
Total Amount of Fragments	1 µL each = 4 µL
NEBuilder HiFi DNA Assembly Master Mix	10 μL
Nuclease- free Water	6 μL
Total Volume
20 μL

First, we must NanoDrop to get the concentrations of each PCR product (see above).

Sample	pANS_1	pANS_2	pANS_3	pANS_4
ng/uL	526.6	495.8	455.5	407.2

Because all are of similar size and concentration, we will be using 1 µL for each fragment.

This was calculated using the NEB protocol, which states an upper limit of 0.5 pmol. This is approximately 610 ng of DNA. Each of our PCR products has a concentration of 400-500, so it will likely sit in the 0.3-0.5 pmol range.

Gel Extraction

Agarose Gel Electrophoresis

• NEB Quickload 1kb plus ladder
• NEB 6x purple loading dye

Gel Extraction using GeneJET Kit

Results

PCR was performed using a vial of Q5 2X master mix from NEB having expired on 10/24.

Nanodrop of PCR products

• pANS 1: 526.6 ng/µL
• pANS 2: 495.8 ng/µL
• pANS 3: 455.5 ng/µL
• pANS 4: 407.2 ng/µL 260/280 and 260/230 ratios were not recorded, but each PCR product was roughly ~1.75 and 0.8 respectively.

Gel electrophoresis of Gibson Assembly Product

The last remaining 2X HiFi master mix was used for assembly. Gel electrophoresis was run at 120 V.

Figure 1: Image of the gel from the gel electrophoresis experiment

Figure 2: Simulated gel electrophoresis on Snapgene

The lane contents are as follows:

1. Ladder
2. Gibson product (pANS_RFP)
3. pANS_1
4. pANS_2
5. pANS_3
6. pANS_4 The Gibson product would have been expected to be high up in the ladder, although no band was seen. Nothing is seen for pANS 1, suggesting failed amplification.

Because no band was seen for the Gibson product, gel extraction was not performed.

Another thing to note is that 5 µL of PCR product was added for the fragment-only lanes, whereas in the Gibson assembly reaction only 1 µL of each fragment was added. There are very faint bands aligning with pANS 2,3, and 4 in lane 2, but nothing for pANS 1 in either lanes 2 or 3. A post-electrophoresis stain may help make it more visible.

A PCR cleanup should also be considered so that more DNA can be added in the reaction (recommended for PCR product volume to not exceed 20% of reaction volume), and that there are less interfering species in the reaction mix.

Not all of the gel loading samples were used even though the intention was to use all of them. They are kept in the -20 °C freezer.

Summary

Gibson assembly of pANS_RFP failed, at the moment we think the likely cause is fragment 1 being poorly amplified. Possible troubleshooting steps are listed in the hand-off section below.

Hand off

• Perform nanodrop on the storage and working stocks the pANS_1 primer pair to check if it was correctly resuspended.
• Verify the ordered primers bind at the desired sites on the pANS_1 fragment on SnapGene to rule out an incorrect order. Retrieve the sequence from the order portals.
  • Ordered pANS_1 sequence verified to match design
  • Ordered primer sequences bind at desired locations
• If new HiFi assembly kits do not arrive soon, ask around to use one Gibson reaction worth of master mix.
• Consider running the PCR products through PCR cleanup.
• Perform a post-electrophoresis stain with SYBR safe according to
• Consider using the remains of the gel loading samples to rerun a gel experiment.

Purpose

To validate that pDest plasmid is the correct plasmid through a restriction digest.

Materials & Methods

Restriction Digest with NEB Enzymes

Remeasured pdest and pdest dil, similar concentrations as before (at most 1 ng/uL difference)

263.5 ng/uL

103.8 ng/uL pDest

Digest using 500 ng

Component	Tube 1 pDest digest	Tube 2 pDest dil digest	Tube 3 pDest (+)	Tube 4 pDest dil (+)	Tube 5 Neg Ctrl	Tube 6 pCB2A_Disp #4 10x dil
Water	42.1	39.2	43.1	40.2	46	16.2
DNA	1.9	4.8	1.9	4.8	0	28.8
Buffer 10x rCutSmart	5	5	5	5	5	5
BsaI	1	1	0	0	1	0
TOTAL	50	50	50	50	50	50

Running the gel:

Lane 1: 1 kb neb ladder

Lane 2: Patt’s pcr tube 1

Lane 3: Patt’s pcr tube 2

Lane 4: Tube 1

Lane 5: Tube 2

Lane 6: Tube 3

Lane 7: Tube 4

Lane 8: Tube 5

Lane 9: Tube 6

Lane 10: empty

Results

No bands appeared for Lane 4 - 8, and bands for Lane 10 is incorrect.

Purpose

Amplify the following fragments for future assemblies and transformation experiments:

	Vendor	Primer	Label	Concentration
2_J23107+RBS_gg	IDT	Twist	prom L
2_J23119+RBS_gg	IDT	Twist	prom M
2_Pcpc560+RBS_gg	IDT	Twist	prom H
1_VCBS_US_short_gg	IDT	Twist	VCBS US S
0_pBR322_ori+sepT2_gg	genscript	genscript	ori CS
3_VCBS_Myc_flexible_gg	Genscript	Genscript	* (star)
4_KanR_TU_gg	Twist	Twist	KanR
SazCA_UTEX_gg	Twist	Twist	SazCA	30.1
BhCA_UTEX_gg	Twist	Twist	BhCA	29.4
BtCAII_UTEX_gg	Twist	Twist	BtCAII	41.7
HpCA_UTEX_gg	Twist	Twist	HpCA	19.1

Assemble the following:

• pRepv2
• pNS1v2-L
• pNS1v2-M
• pNS1v2-H
• pKOv2
• pANS_RFP

Materials & Methods

Component	Volume (µL)
Q5 2x master mix	12.5
10 uM primer pair mix	2.5
DNA fragment	0.2
Nuclease-free water	9.8
Total	25

Golden Gate Assembly

Golden Gate Assembly with NEBridge Ligase Master Mix

pRepv2

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (µL)
Ligase master mix					5
1_VCBS_US_short_gg		241	7.424	29.7	0.25
3_VCBS_Myc_flexible_gg		2182	67.20	80.4	0.84
4_KanR_TU_gg		1032	31.78	79.9	0.4
5_VCBS_DS_768+bom_gg		981	30.21	50	0.6
0_pBR322_ori+sepT2_CS_gg		1541	47.46	99.8	0.5
BsaI-HF-v2					1
Water					6.4
Total					15

pNS1v2-L, pNS1v2-M

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (µL)
Ligase master mix					5
1_NS1_US_700_gg		776	23.19	50	0.5
2_J23107+RBS_gg		133	4.098	L: 13.4
M: 5.0	L: 0.31

M: 0.82 | 3_VCBS_Myc_flexible_gg | | 2182 | 67.20 | 80.4 | 0.84 | 4_KanR_TU_gg | | 1032 | 31.78 | 79.9 | 0.4 | 5_NS1_DS_700+bom_gg | | 917 | 28.24 | 50 | 0.6 | 0_pBR322_ori+sepT2_CS_gg | | 1541 | 47.46 | 99.8 | 0.5 | BsaI-HF-v2 | | | | | 1 | Water | | | | | L: 5.85 M: 5.34 | Total | | | | | 15

pNS1v2-H

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (µL)
Ligase master mix					5
1_NS1_US_700_gg		776	23.19	50	0.48
2_Pcpc560+RBS_gg		659	20.30	37.8	0.54
3_VCBS_Myc_flexible_gg		2182	67.20	80.4	0.84
4_KanR_TU_gg		1032	31.78	79.9	0.4
5_NS1_DS_700+bom_gg		917	28.24	50	0.56
0_pBR322_ori+sepT2_CS_gg		1541	47.46	99.8	0.5
BsaI-HF-v2					1
Water					5.68
Total					15

pKOv2

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (µL)
Ligase master mix					5
1_VCBS_US+SpecR_TU		1809	55.71	50	1.1
5_VCBS_DS_768+bom_gg		981	30.21	50	0.6
0_pBR322_ori+sepT2_CS_gg		1541	47.46	99.8	0.5
BsaI-HF-v2					1
Water					6.8
Total					15

Color constructs

• using preps from 2025.06.21 Transformation of pLiN + distribution parts for Colourful colonies

DNA (plasmid)	Plate Name	Concentration	260/280	260/230
mRFP1	R	87.4	1.90	1.54
tannenRFP	O	54.7	1.97	2.22
sarcGFP	G	46.9	1.91	2.08

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (µL)
Ligase master mix					5	20
pJUMP28	BBa_J428353	3359	103.4	28.2	1	4
Promoter	BBa_J23100	2424	74.65	23.8	2	8
RBS	BBa_B0034_m1 (BBa_J428038)	2411	74.25	27.7	2	8
CDS (R, O, G)		2740	84.39	23.7	2
Terminator	BBa_B0015	2520	77.61	20.8	2	8
BsaI-HF-v2					1	4
Water					0
Total					15	13

Gibson Assembly

Component	Concentration (ng/µL)	Fragment length	mass for 0.05 pmol (ng)	Volume (µL)
2X HiFi master mix				10
pANS_RFP_1	50	1989	61.26	1.2
pANS_RFP_2	50	1311	40.38	0.8
pANS_RFP_3	50	1301	40.07	0.8
pANS_RFP_4	50	2200	67.76	1.4
Water				5.8
Total				20

Results

pRepv2, pNS1v2-L, pNS1v2-M, pNS1v2-H, pKOv2, and pANS_RFP are assembled and placed in the -20 freezer for subsequent transformation .

Constructs for mRFP1, tannenRFP, and GFP have also been assembled.

We performed a gel electrophoresis.

Summary

See above.

Hand off

The assembled plasmids will be used for subsequent transformation experiments.

Purpose

Prepare BG-11 medium for UTEX 2973 for optimised growth and use in future experiments.

Test if using different concentration of Na2CO3 increases the growth rate of UTEX. Concentrations tested are 10x, 20x and 50x.

Materials & Methods

Na2CO3

• MW = 106 g/L
• BG-11 working conc: 0.02 g/L

For 100 mL:

• 10x: 0.02 g/L * 10 * 0.1 mL = 0.02 g
• 20x = 0.04 g
• 50x = 0.1 g

BG-11 Medium Recipe

Results

There was no significant difference in the growth rate of UTEX 2973 between the different concentrations of Na2CO3 used. There was a biomass increase (marked by increased OD) in the media containing higher concentrations of Na2CO3.

Summary

Prepared BG-11 media containing different concentrations of Na2CO3 to test if increased Na2CO3 concentration contributes to increased growth rate. We will continue to use 10X stock of BG-11 media for future experiments.

Hand off

Use prepared 10x BG-11 medium for future UTEX experiments.

Purpose

To create microscope slides from CB2A liquid culture

Materials & Methods

Materials:

Coverslip

Microscope slides

CB2A liquid culture

CB2A agar plate (pure culture Wendy prep)

Methods:

Everything done under Bunsen burner

CB2A liquid culture slide:

5 microliters of liquid culture onto microscope slide

CB2A culture slide:

5 microliters of dH2o+ colony picked from plate

Cover with coverslip at an angle

Results

2 microscope slides

Labeled cb2a liquid culture and cb2a culture, currently left on lab bench.

Purpose

Type here…

Materials & Methods

Miniprep

LP-9 Plasmid Mini-Preparation (Miniprep) Protocol

LP-10 Nanodrop DNA/RNA

Golden Gate Assembly with SapI

LP-17 Golden Gate Assembly with NEBridge Ligase Master Mix

Concentrations for inserts from 2025.06.27 UTEX Assembly 1 + Amplifying Fragments

Name	Length (bp)	Mass (ng) for 0.05 pmol	Concentration (ng/μl)	Volume (μL)
pRepv2	5677	87.42	25.5	3.43
pNS1v2-H	6723	103.5	22.7	4.56
BtCAII_UTEX_gg	854	26.30	41.7	0.63
BhCA_UTEX_gg	908	27.97	29.4	0.95
HpCA_UTEX_gg	764	23.53	19.1	2.46
SazCA_UTEX_gg	836	25.75	30.1	0.86

Reaction compositions (volumes in μL)

Component	pRepv2-BtCAI	pRepv2-BhCA	pRepv2-HpCA	pRepv2-SazCA
NEBridge Ligase Master Mix	5	5	5	5
pRepv2	3.43	3.43	3.43	3.43
DNA fragments	0.63	0.95	1.23	0.86
SapI	1	1	1	1
Molecular water	4.94	4.62	4.34	4.71
Total	15	15	15	15

Component	pNS1v2-H-BtCAI	pNS1v2-H-BhCA	pNS1v2-H-HpCA	pNS1v2-H-SazCA
NEBridge Ligase Master Mix	5	5	5	5
pNS1v2	4.56	4.56	4.56	4.56
DNA fragments	0.63	0.95	1.23	0.86
SapI	1	1	1	1
Molecular water	3.81	3.49	3.21	3.58
Total	15	15	15	15

Results

Miniprep and Nanodrop

Name	Concentration (ng/ul)	A260/A280	A260/A230	QBIT
pANS_RFP 1	23.4	1.84	1.75	7.92
pANS_RFP 2	19.32	1.70	1.50	8.38
pANS_RFP 3	17.7	1.95	1.56	10.3
pANS_RFP 4	18.8	1.78	1.5	10.8
pANS_RFP 5	24.6	1.81	1.70	9.32
pANS_RFP 6	17.8	1.76	1.36	9.3
pRepv2	25.5	1.94	1.66
pNS1v2-M	5.0	1.64	0.79
pNS1v2-H	22.7	1.95	1.76

Summary

• Preformed a plasmid miniprep achieving normal concentrations and lower purity except for pNS1v2-M which was unsuccessfully miniprepped.
• Preformed a transformation with new and old competent cells
• Preformed a Golden Gate Assembly and subsequent plating overnight.

Hand off

N/A

Purpose

Due to the results from 2025.07.06 Colony PCR and Restriction Digest Redo, pNS1v2-L, pNS1v2-H Retransformation, Miniprep of Minicultures in TB being inconclusive, we will be redoing the gels to see if better technique can give us more useful results.

Notable issues: 1. Both gel 1 and 2 displayed strangely under UV. This may be attributed to poor formation during pour as a result of waiting too long.

1. Gel 1 “no PCR” lane 3 did not show. The amount of DNA added was not very high (6 uL) for its concentration. However, lane 2 seemed to appear faintly about where it was expected.
2. Gel 1 “digestion” lanes did not show either. Potentially due to low concentration.
3. Gel 1 “colony PCR” lanes showed, indicating good concentration and volume used, but the location was much further (therefore size smaller) than initially expected. They were found much further than lane 2, which was the non-amplified DNA, which indicates that the DNA in the PCR amplfied wells is shorter than expected.
4. Gel 2 did not show anything for any lane except the ladder. Initial guess seems to be no DNA in the wells.

Non-issues:

1. The DNA is likely not decayed due to lane 2 (no pcr) showing at approximately the correct location.
2. Primer design is likely to be correct due to previous experiments.
3. Gel concentration is probably fine, as the ladder was intact for each.
4. Gel itself is likely not to blame for gel 2 not displaying anything.

Changes: → Add more DNA into each gel well wherever possible.

→ Do PCR cleanup on the redone PCR products, see if that fixes plasmid size.

→ Redo Gel 2 entirely, with increased DNA per well, using a mix of 4 uL DNA + 1uL water + 1uL dye to 6 uL total.

→ Pour gel at a higher temperature to ensure correct formation.

→ Better technique may avoid contamination of DNA/RNAses.

→ Use midiprep instead of miniprep DNA to ensure more concentration and to have more stock. (This stock of midiprepped DNA in theory has more DNA).

Additionally, also do colony PCR for SazCA colonies to check for expression, will also do PCR cleanup on these.

Materials & Methods

Plan:

1. start colony PCR again (30-40min), then set in the thermocycler for 1:30 (hour, minute).
2. No need to do restriction digest, just finish protocol and set up cleanup kit, make PYE.
3. At around one hour in the thermocycler, Set up gels, put in dye once its warm to the touch. May be done during next step. Once dye in, pour immediately then cover.
4. Once thermocycle done, pcr clean up then assemble non pcr, and get all other dnas for gel loading

Making TAE:

2025.06.12 Agarose Gel for CB2A parent plasmid digest

For 500mL bottle of 1x TAE

10mL 50x TAE + 490mL milliQ water → 500mL total.

This is for gels, no need to do under sterile.

Colony PCR

Polymerase Chain Reaction (PCR) with NEB Q5 High-Fidelity 2X Master Mix

Protocol adapted from 2025.07.05 Colony PCR and Restriction Digest of pRepv2 and pNS1v2-H.

PCR on miniprepped plasmids using M13 primers. Total 20 uL.

Note that the volumes were reduced from the original to save on resources.

Performed on pRepv2 and pNS1v2-H minipreps from 2025.07.04 Miniprep of pANS_RFP, pRepv2, pNS1v2-M, pNS1v2-H + Assembly of CAs to pRepv2, pNS1v2-H

Component	Volume (uL)
2X Q5 master mix	10
10 uM M13 primers	1
Plasmid	1
Water	8
Total	20

STEP		TEMP (°C)	TIME
Initial Denaturation		98	30 seconds
25 Cycles	Denaturation	98	10 seconds
	Annealing	57	20 seconds
	Extension	72	100 seconds
Final Extension		72	2 minutes
Hold		4	Infinity

Times are preserved from 2025.07.06 Colony PCR and Restriction Digest Redo, pNS1v2-L, pNS1v2-H Retransformation, Miniprep of Minicultures in TB.

Note that denaturation was changed to 10 seconds to see if it had improvements on PCR.

Another colony PCR was done on SazCA colonies. This was placed in the thermocycler with the rest of the PCR tubes.

Component	Volume (uL)
2X Q5 master mix	10
SazCA primers	1
Add small amount of
colony via p2 tip	n/a
Water	9
Total	20

PCR DNA Cleanup By Centrifugation

Base protocol on the NEB #T1130S/L Spin PCR & DNA Cleanup Kit online protocol, using the Monarch kit in lab. There is also a sheet that comes with the box that has the protocol.

Steps are similar to miniprep.

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

1. (if kit is unopened, prepare buffers by following instructions on website).

BIND

1. In the Adjust PCR results to 20uL if needed. Mix 5 to 1 parts of 5 Monarch buffer BZ to 1 DNA PCR results. Mix by flicking or pipetting, DO NOT VORTEX.

Name	Vol uL
DNA (PCR’d)	20
BZ buffer	100
total	120

1. 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

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

1. Transfer the column (top part) onto a clean 1.5mL microfuge (Eppendorf) tube.
2. Add 5-20uL of EY elution buffer to the center of the matrix. 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. Collect flow-through for further use.

Note: 5 uL didn’t come through. Use more (15-20 uL) or centrifuge for 2-2.5 minutes instead of 1.

Gel electrophoresis

1% gel 30 mL → 0.30g agar

1/10 000 volume of SYBR safe dye → 3 uL

Do 2 rows.

Gel 1: colony pcr and restr. dig. + old gel 2

1st row digest and pcrs

#	name	DNA:H2O:dye uL
1	Ladder	n/a
2	pRep No PCR	4:1:1 6 total
3	pNS1 No PCR	4:1:1 6 total
4	new pRep PCR	20:0:4 use 12
5	new pNS1 PCR	20:0:4 use 12
6	pRep Digest	14 total
7	pNS1 Digest	14 total
8	old pRep PCR	14 total
9	old pNS1 PCR	14 total

2nd row full construct w/ CAs

#	name	DNA:H2O:dye + uL
1	Ladder	n/a
2	pRep BtIICA	4:1:1
3	pRep BhCA	4:1:1
4	pRep HpCA	4:1:1
5	pRep SazCA	4:1:1
6	pNS1 BtIICA	4:1:1
7	pNS1 BhCA	4:1:1
8	pNS1 HpCA	4:1:1
9	pNS1 SazCA	4:1:1

100V for 30-40min

Note: We won’t be doing the 2nd row anymore.

Gel 2:

1.2% of 30mL agar→ 0.36g powder

#	name	DNA:H2O:dye + uL
1	Colony 1	5:0:1 aliquot
2	Colony 2	5:0:1 aliquot
3	Colony 3	5:0:1 aliquot
4	Colony 4	5:0:1 aliquot
5	Colony 5	5:0:1 aliquot
6	Colony 6	5:0:1 aliquot
7	Ladder	N/A

100V for 30-40min

Results

Gel 1:

Note: it appears that lane 6-7 (digests of pRep and pNS1) were empty, despite adding more DNA. this indicates that there was NO DNA resulting from the digest experiment.

Lanes 4-5 and 8-9 show digests (note that 8-9 are from 2025.07.06 Colony PCR and Restriction Digest Redo, pNS1v2-L, pNS1v2-H Retransformation, Miniprep of Minicultures in TB, the previous day’s PCR experiment). Curiously, they appear much lower than expected. We think this maybe a result of mispriming, and therefore the extension being incomplete. A potential cause for this may be.

Note that 2025.07.08 Qubit Measurement of Prepped UTEX Vectors indicates a different, and much lower, quantity of DNA from our mini/midipreps than the nanodrop. Qubit is a more intensive, but supposedly accurate method of measuring DNA, so this may indicate that our DNA concentrations across the board are too low.

Gel 2:

We didn’t see any bands for the colonies picked, despite doing colony PCR.

These were done on the SazCA colonies, we believe it to be contamination, Pattarin mentions accidentally slipping the cover open (for this plate) outside of flame, which is a potential cause. Additionally, we may also just be not seeing successful transformation.

Note: try doing a touchdown pcr to optimize annealing temps instead of relying on lit notes.

Summary

Gel 1: Digestion likely unsuccessful (hardly any DNA seen), retry digestion with much higher quantities.

PCR of pRep and pNS1 backbones is yielding unexpected results, potentially do touchdown PCR (varying annealing temp) to optimize PCR for the future.

“no PCR” lanes show approximately the expected size. We think that these are correct, something is likely off with the PCR.

PCR cleanup seems to have eliminated the upper (larger) band of DNA for lanes 4-5. It is not known whether or not this means it was successful in increasing the purity (neither band were of expected size).

Gel 2:

We did not see anything, the SazCA colonies are potentially contamination. All lanes were PCR, so quantity of DNA should not be an issue.

It is unknown if PCR cleanup was successful for this batch.

Hand off

→ design future experiments based on PCR touchdown

→ review digestion of pRep and pNS1, maybe redo

• Consider modifying M13 primers to overcome high Tm difference (56 and 62 C on NEB Tm calculator for Q5 polymerase)

Purpose

To prepare wash buffer (2 mM Tricine, 2 mM EDTA) and DTN buffer for future use according to [LP-40 DTN Medium Recipe]. These buffers are required for downstream experiments for recovery of cyanobacteria after electroporation.

Materials & Methods

For wash buffer: 2mM Tricine and 2mM EDTA for total of 1L with Milli-Q water.

For DTN buffer: Followed exactly LP-40 protocol

Results

Buffers successfully prepared and DTN buffer was autoclaved. Wash buffer clear, colourless, no solubility issues observed.

Summary

Buffers are ready for use in recovery of cyanobacteria after electroporation experiments.

Hand off

Buffers should be stable. Prepare fresh if contamination or precipitation occurs.

Purpose

To verify the colonies yielded from 2025.07.11 Transformation of pCaS and pCaD + CA assemblies, we will perform colony PCR to check for successful clones. In addition we will perform Colony PCR on some colonies from the pCaS and pCaN plates from 2025.07.05 Transforming pCaS, pCaN, pLiD, pLiN to check if previous transformations yielded any successful clones as well (initially we suspected the entire plate was contaminated).

Materials & Methods

Colony PCR

TUBE	1	2	3	4	5	6	7	8	9	Final
DNA template	pCaD-BhCA	pCaD-BtCAII	pCaD-HpCA	pCaD-SazCA	pCaD-BhCA	pCaD-BtCAII	pCaD-HpCA	pCaD-SazCA	pCaD
Rxn for amplifying:	linearize the plasmid	linearize the plasmid	linearize the plasmid	linearize the plasmid	amplify the inserts	amplify the inserts	amplify the inserts	amplify the inserts	amplify the inserts/MCS
Amplicon size (bp):	6625	6580	6490	6484	881	836	746	740	86
Ideal annealing temp (C)	68	68	68	68	61	61	61	61	61
COMPONENTS (uL)
Water	8.75	8.75	8.75	8.75	8.75	8.75	8.75	8.75	7.5	-
2X Q5 master mix	10	10	10	10	10	10	10	10	10	1X
10 uM Primer pair	1.25 (C8)	1.25 (C8)	1.25 (C8)	1.25 (C8)	1.25 (A8)	1.25 (A8)	1.25 (A8)	1.25 (A8)	1.25 (A8)	0.5 uM
DNA template	Picked colony	Picked colony	Picked colony	Picked colony	Picked colony	Picked colony	Picked colony	Picked colony	1.25 uL	~2 ng
Total	20	20	20	20	20	20	20	20	20

C8 master mix (C8 MM)

Water = 8.75 * 5 = 43.75

2X Q5 Master mix = 10 * 5 = 50

C8 primer = 1.25 * 5 = 6.25

Total = 100 µL

Aliquot 20 into each tube.

A8 master mix (C8 MM)

Water = 8.75 * 5 = 43.75

2X Q5 Master mix = 10 * 5 = 50

A8 primer = 1.25 * 5 = 6.25

Total = 100 µL

Aliquot 20 into each tube.

C8 master mix 2 (C8 MM)

Water = 8.75 * 4.4 = 38.5

2X Q5 Master mix = 10 * 4.4 = 44

C8 primer = 1.25 * 4.4 = 5.5

Total = 88µl

Aliquot 20 into each tube.

A8 master mix 2 (C8 MM)

Water = 8.75 * 4.4 = 38.5

2X Q5 Master mix = 10 * 4.4 = 44

A8 primer = 1.25 * 4.4 = 5.5

Total = 88 µL

Aliquot 20 into each tube.

pCaD miniprep product (labelled “CB2A display”) used to assemble the pCaD-CA constructs is 172 ng/µL → too concentrated so will dilute then amplify the CA insert region via PCR

• 100x dilution: 5 µL of DNA in 495 uL of water
• Then use 1.25 µL of that for PCR

Thermocycler for C8

20 sec/kb

6.6 * 20 sec = 132 sec

STEP		TEMP (°C)	TIME
Initial Denaturation		98	10 min
25 Cycles	Denaturation	98	10 seconds
	Annealing	68	30 seconds
	Extension	72	132 sec (2:12)
Final Extension		72	2 minutes
Hold		4	Infinity

Thermocycler for A8

20 sec/kb

0.8 * 20 sec = 16 sec

STEP		TEMP (°C)	TIME
Initial Denaturation		98	10 min
25 Cycles	Denaturation	98	10 seconds
	Annealing	61	30 seconds
	Extension	72	16 seconds
Final Extension		72	2 minutes
Hold		4	Infinity

Results

Used too much primer for ‘s PCRs, should be 1 µL in 20 µL reaction.

Purpose

To verify if the new batch of electrocompetent CB2A cells from 2024.07.08 Making electrocompetent CB2A cells can indeed take up the plasmids.

Materials & Methods

Based on the following protocol as a reference, but with changes:

CB2A Electroporation

Transform with pCB2A_Sec plasmid, pCB2A_Disp plasmid, no DNA but transformed with EC cells (negative), 4th plate is just empty - nothing plated (negative)

• CB2A plasmids confer Cm resistance
• Used the miniprep products with the highest concentration

Set-Up/Preparation

1. Put 3 new cuvettes on ice for at least 20 minutes to chill (or pre-chill by putting them in the fridge).
2. Thaw 3 electrocompetent Caulobacter cells on ice (one tube per plasmid + one for negative control). Do not handle the cells vigorously, as this will reduce transformation efficiency.
3. Put undiluted DNA in new eppendorf tubes (labelled) on ice.
   1. Concentration of pCB2A_Disp plasmid is 69.2ng/µl (potentially 89.2, writing font hard to tell), use 1µl

• Concentration of pCB2A_Sec plasmid #4 is 36ng/µl, use 1.5µl

1. Pre-warm 3 tubes of PYE medium 950µl in sterile aeration tubes in 30C room, along with 4 PYE + CM plates.

Electroporator Settings ( Eppendorf)

• Set to 2500V

From Beth about the previous protocol:

The ratio is quite inportant here I beleive so keeping the DNA to 1µL would be the way to go. Whenever I miniprep the pCB2A_Disp plasmid I get >100ng/µL, so I just put 1µL of that directly into the electroporation mix. The more volume of DNA you put, the more salt you put, which affects the electricity efficiency. I know you only got 78ng/µL, so maybe popping 1.5µL in the electroporation mix would be more appropriate, or even just trying with 1µL.

Procedure: Electrotransformation

1. Transfer 50µl electrocompetent CB2A cells to each tube containing DNA
2. Wait 1-2 minutes.

• In this time, dry cuvette thoroughly with kipwipes and place in pulser. Uptake 950µl PYE medium from aeration tube into p1000 pippette.

1. Transfer the 50µl DNA/cell mix to the cuvette (careful to place in 1mm slit), and pulse.
2. Resuspend cells gently but immediately with 950µl PYE medium with p1000 pipette.
3. Transfer cells to aeration tube.
4. Incubate in 30°C room at 200rpm shaking for 2 hours
5. Plated 100 µL of each transformation onto PYE-Cm plates. Put in 30°C at .

Shorthand	Plate Label	Expected Result
pCB2A_Disp/CB2A-EC	CB2A-EC pCB2A_Disp plasmid ctrl 7/18/2025 AS iGEM	Expect colonies in 2-3 days?
pCB2A_Sec/CB2A-EC	CB2A-EC pCB2A_Sec plasmid ctrl 7/18/2025 AS iGEM	Expect colonies in 2-3 days?
CB2A-EC -ve ctrl	CB2A-EC -ve ctrl 7/18/2025 AS iGEM	No colonies
No trans -ve ctrl	unlabelled, only has the original plate label “PYE+Cm 2025-06-27”	No colonies

Results

:

No colonies on any plates, as expected:

:

• 48 hours incubation

• A lawn of colonies on pCB2A_Disp/CB2A-EC plate

• pCB2A_Sec/CB2A-EC and CB2A-EC -ve ctrl plates have speckles that appear to be indents from vigorous plating with glass beads

• No trans -ve ctrl plate is completely clear

• pCB2A_Disp plasmid/CB2A-EC looks the same

• Lawn of growth developed on pCB2A_Sec plasmid/CB2A-EC and CB2A-EC -ve ctrl

• 3 colonies on No trans -ve ctrl

Get Wendy to check pCB2A_Disp plasmid plate under the microscope to verify that colonies observed are Caulobacter.

Checked a colony from pCB2A_Disp/CB2A-EC and pCB2A_Sec plasmid/CB2A-EC under the microscope, Wendy suspects they’re not Caulobacter cells.

Troubleshooting

These mini-experiments were preformed prior to the electrotransformation experiment described above.

• Plated with beads but they were recently sterilised (and didn’t have any issues using those beads with E. coli)
• The batch of plates used in this experiment were checked earlier in the month and didn’t grow anything on their own so likely not a source of contamination (+ the :
  • : No colonies after 2 days of incubating at 30°C
• PYE media used was a fresh batch.
• The competent cells were made in full sterile conditions (using the BSC), and streaked out on a PYE/Cm plate to check for Cm resistance:
  • : There was growth on the streak lines but only after a few days (CM could have possibly degraded)
• DNA used was extracted a while ago, did have some issues with contamination in E. coli transformations

Summary

Contamination of plates again so troubleshoot.

Hand off

• Check CB2A-EC -ve ctrl under the microscope?
  • Or just look at a sample of the EC cell stock
• For next trial:
  • Use new miniprepped DNA
  • New batch of plates
  • New media
  • Parafilm plates after plating
  • Use disposable inoculation loops
• Ask Wendy/Beth for advice on electroporation protocol

Purpose

Golden Gate Assembly of pCaD-CAs, pCaS-CAs, pCaN-CAs.

Materials & Methods

Golden Gate Assembly with NEBridge Ligase Master Mix

CA insert sizes:

pCaD size: 5830 bp

pCaS size: 3974 bp

pCaN size: 2915 bp

Assembly of pCaD + CAs

Reaction mix

D = pCaD

S = pCaS

N = pCaN

Bh = BhCA

Bt = BtCA2

Hp = HpCA

Saz = SazCA

pCaD-CAs

Component	D-Bh	D-Bt	D-Hp	D-Saz
NEBridge Ligase Master Mix, 3X	5	5	5	5
pCaD (backbone)	1.04	1.04	1.04	1.04
CA	2.7	2.7	2.3	2.3
SapI	1	1	1	1
Water	5.3	5.3	5.6	5.6
TOTAL	15	15	15	15

pCaS-CAs

Component	S-Bh	S-Bt	S-Hp	S-Saz
NEBridge Ligase Master Mix, 3X	5	5	5	5
pCaS (backbone)	3.6	3.6	3.6	3.6
CA	2.7	2.7	2.3 (2.7)	2.3 (2.7)
SapI	1	1	1	1
Water	2.7	2.7	3.0	3.0
TOTAL	15	15	15	15

pCaN-CAs

Component	N-Bh	N-Bt	N-Hp	N-Saz
NEBridge Ligase Master Mix, 3X	5	5	5	5
pCaN (backbone)	2.3	2.3	2.3	2.3
CA	2.7	2.7	2.3	2.3
SapI	1	1	1	1
Water	4.0	4.0	4.4	4.4
TOTAL	15	15	15	15

Thermocycler (program for 2 fragments, SapI)

STEP		TEMP (°C)	TIME
1 Cycle	Digestion	37	15 min
Deactivation		60	5 minutes
Hold		4	Infinity

Results

All products stored in -20°C.

Purpose

To perform miniprep on:

• pCaD-BtCAII
• pCaN-BtCAII
• pCaS-BtCAII from the minicultures started from the July 22 transformation plates.

Materials & Methods

Plasmid Mini-Preparation

Procedure was slightly modified to include an extra ethanol step to improve efficiency.

Results

Nanodrop results:

All samples had good curves.

Sample	ng/μL	260/280	280/260
pCaD-BtCAII	654.7	1.95	2.15
pCaN-BtCAII	553.88	1.98	2.30
pCaN-BtCAII	736.28	2.04	2.45

Summary

Miniprepped pCaD-BtCAII, pCaS-BtCAII, and pCaN-BtCAII, stored final miniprep products in -20°C.

Hand off

Further validation to be performed through PCR and agarose gel electrophoresis to confirm transformations.

Purpose

Redo transformations for BL21 backbones (pLiD and pLiN) and CB2A CA constructs. In 2025.07.22 Transformation of pCaD-CAs, pCaS-CAs, pCaN-CAs, we used 45 seconds instead of the recommended 30 seconds for heat shock, in this attempt we will revert back to the shorter heat shock duration to improve transformation efficiency.

Materials & Methods

Heat Shock Transformation with a Thermocycler

Materials

• Comp cells: 10-beta NEB

Thermocycler

STEP	TEMP (°C)	TIME	Sample volume
Ice incubation	4	20 min	52 µL
Heat shock	42	30
DNA uptake	4	5 min
-	4	Infinity
Recovery	37	Infinity (timer set for 1 hr)

Samples transformed:

• pLiD
• pLiN
• pCaD-BhCA/HpCA/SazCA
• pCaS-BhCA/HpCA/SazCA
• pCaN-BhCA/HpCA/SazCA

Plated 50 µl of each transformation incubated at .

Results

• All plates yield colonies except for -ve ctrl, pLiD and pLiN

• 1 colony will be picked from each pCaD/pCaS/pCaN-CA plate for gel analysis validation (PCR + digest) Fig 1. Example of plate with colony marked for picking, labelled with the current date.

• pLiD and pLiN plates returned to 37°C, check again

• pLiD and pLiN plates have colonies

• 1 colony will be picked from each plate for gel analysis

Summary

Switching to 10-beta cells improved transformation efficiency, but pLiD and pLiN are still slow to yield colonies.

Hand off

• Perform colony PCR and gel analysis to validate clones
• Validate that the competent cells were able to take up DNA plasmid prepped using the modified miniprep protocol
  • Pick 1 colony from +ve ctrl to to test whether the clones have successfully taken up CB2A Secretion Display plasmid plasmid

Purpose

Colony PCR of pCaD-BhCA, pCaS-BhCA/HpCA/SazCA, and pCaN-BhCA.

Materials & Methods

Master mix

9 tubes 110%→ 9.9

• water 9.9*4.5= 44.55( round to 45)
• 2X Q5 MM: 9.9 * 5 = 49.5 (round to 50)
• A8 primer pair: 9.9 * 0.5 = 4.95 (round to 5)

1. Aliquot 10 uL of master mix per tube, remaining can be used as negative ctrl
2. Instead of DNA miniprep product, pick a colony with sterile p200 or p10 tip
   1. Then dip and swirl into PCR reaction mix
3. Spin down

TUBE label
DNA template name (miniprep products)	D-BhCA	S-BhCA x2	S-SazCAx2	S-HpCA x2	N-BhCA
COMPONENTS (uL)
Water (UPH2O)	4.5	4.5	4.5	4.5	4.5
2X Q5 master mix (10/24)	5	5	5	5	5
10 uM Primer pair (A8)	0.5	0.5	0.5	0.5	0.5
DNA template	colony	colony	colony	colony	colony
Total volume	10	10	10	10	10

STEP		TEMP (°C)	TIME (MM:SS)	Sample volume (uL)
Initial Denaturation		98	00:30(supposed to be 5 min)	10
25 Cycles	Denaturation	98	00:10
	Annealing	61	00:30
	Extension	72	00:25
Final Extension		72	02:00
Hold		4	Infinity
Total duration			53:45

Results

PCR products stored in green tip rack in -20°C freezer.

Summary

See above. There was a mistake in the thermocycler protocol used in this experiment, the initial denaturation time should be 5 minutes. However, 30 seconds may be sufficient for lysing the cells --- if so, adjust protocol to use 30 seconds for initial denaturation.

Hand off

Run on 1% agarose gel (use the gel already made 2 days ago)

Purpose

Previous transformation attempts of pNS1v2-L, pKOv2, pDisp-L and pDisp-H have not been successful and their assembly products are running low so they will be reassembled.

Also assemble pExp-L and pExp-H

Transform the products and the following backbones to check for reporter functionality, for a total of 13:

• Materials & Methods

Recipes from 2025.06.27 UTEX Assembly 1 + Amplifying Fragments

LP-17 Golden Gate Assembly with NEBridge Ligase Master Mix Run the cycle for 3-6 reactions

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

pNS1v2-L

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
NS1_US_700_gg	twist A1	776	23.19	50	0.5
J23107+RBS_gg	UTEX Prom L	133	4.098	13.4	0.31
VCBS_Myc_flexible_gg	star	2182	67.20	80.4	0.84
KanR_TU_gg		1032	31.78	79.9	0.4
NS1_DS_700+bom_gg	twist D1	917	28.24	50	0.6
pBR322_ori+sepT2_CS_gg	ori+CS	1541	47.46	99.8	0.5
BsaI-HF-v2					1
Water					5.85
Total					15

pKOv2

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
VCBS US homology arm+SpecR TU: VCBS_US+SpecR_TU	twist B1	1809	55.71	50	1.1
VCBS DS homology arm+pBR322 bom: VCBS_DS_768+bom_gg	twist E1	981	30.21	50	0.6
pBR322_ori+sepT2_CS_gg	ori+CS	1541	47.46	99.8	0.5
BsaI-HF-v2					1
Water					6.8
Total					15

pDisp-L

Recipe from 2025.07.15 UTEX Assembly 3

skip this for now, managed to get colony of this

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
J23107+RBS_gg	UTEX prom L	133	4.098		0.31
VCBS_Myc_flexible_gg	star	2182	67.20	80.4	0.84
pANS_RFP	pANS 2	6651	204.8	122	0.6
BsaI-HF-v2					1
Water					7.25
Total					15

pDisp-H

Recipe from 2025.07.15 UTEX Assembly 3

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
Pcpc560+RBS_gg	UTEX prom H	659	20.30	37.8	0.54
VCBS_Myc_flexible_gg	star	2182	67.20	80.4	0.84
pANS_RFP	pANS 2 7/16	6651	204.8	122	0.6
BsaI-HF-v2					1
Water					7.02
Total					15

pExp-L

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
J23107+RBS_gg	UTEX Prom L	133	4.098	13.4	0.31
sfGFP+term_gg	twist G1	917	28.24	50	0.56
pANS_RFP	pANS 2 7/16	6651	204.8	122	0.6
BsaI-HF-v2					1
Water					7.56
Total					15

pExp-H

Component	Label	Length with adapters	Mass for 0.05 pmol (ng)	Concentration	Volume (μL)
Ligase master mix					5
Pcpc560+RBS_gg	UTEX prom H	659	20.30	37.8	0.54
sfGFP+term_gg	twist G1	917	28.24	50	0.56
pANS_RFP	pANS 2 7/16				0.6
BsaI-HF-v2					1
Water					7.3
Total					15

Results

13 transformations were plated. Since these are all backbones, a GFP fluorescence signal is expected for all.

The pRepv2 (R2) ethanol prep (labelled mira) clearly has many more fluorescent colonies than the old midiprep, suggesting a much higher true concentration.

Like was seen on the gel in 2025.07.30 Restriction Digest and Gel Electrophoresis of UTEX Vectors, there appeared to be nothing in the pNS1v2-M (N2M) and pNS1v2-H (N2H) ethanol prep (no colonies). Looks like there is also a very low concentration in the N2H midiprep.

Out of the 5 transformations of assembly products from this experiment, only pExp-H and pExp-L were successful. pDisp-H had no fluorescent colonies, and pNS1v2-L and pKOv2 were blank.

pKOv2 had been blank several times now on LB/Kan/Spec plates. This could indicate a problem with the synthesized gene fragment. The ordered sequenced matched the design sequence, so it could be a problem with the design itself. The SpecR cassette shares the same promoter, RBS, and terminator with the KanR cassette.

Summary

• Reassembly and transformation of UTEX constructs yielded success for pExp-L and pExp-H only.
• pNS1v2-L, pKOv2, and pDisp-H failed, producing no colonies. pKOv2 in particular has repeatedly failed despite correct fragment sequencing, suggesting a design issue with the SpecR cassette.
• Backbone quality appears to strongly influence outcomes (R2 ethanol prep effective; N2M/N2H essentially empty)

Hand off

check plates for green fluorescent colonies and pick the new assemblies