Purpose:

Prepare kanamycin LB agar plate and LB broth for clone selection

Methods:

Prepare 9 kanamycin LB agar plate and 500mL LB broth for Cloning experiments

Materials:

• LB powder
• Agar powder
• Kanamycin powder
• dd water

Note from day of experiment:

2024-05-04

1. 250ul of kanamycin (50mg/ml) is added once the mixture is cooled to ~40 degree Celcius
   1. Kan concentration of 50ug/ml
2. 250 mL of mixture is used to prepare 9 agar plates
3. Plates are left inside BSC to cool down

2024-05-05

1. Plates are checked and no sign of bacteria growth
2. Plates are sealed with parafilm and stored upside down in 4C cool room

Results:

Plates and LB broth were successfully prepared.

Purpose:

To create Kanamycin Stock solution preparation

Materials:

• Distilled water
• Kanamycin sulphate to solutioN

Methods:

1. Prepare 0.8mL of distilled water in suitable container.
2. Add 0.05g of Kanamycin sulphate to solution.
3. Add distilled water until 1mL.
4. Keep stock at -20C.

Results:

Successful preparation

Purpose:

Remove C-terminus 6x His-tag and linearize the pET-28b(+) vector

Materials:

• Forward primer
• Reverse primer
• pET-28b(+) vector
• 2x Q5 Mastermix
• Water

Methods:

See Polymerase Chain Reaction (PCR) Protocol.

Note From Day of Experiment:

• Since we are doing inverse PCR of pET-28b(+) vector (~5500bp), it requires a longer denaturation and extension time.

• dilute pET-28b(+) to 1ng/ul, the original concentration is 0.5ug/ul
  • take 1ul from the plasmid tube and dilution with 500ul water
• dilute P5 and P6 primer to 10uM
  • P5: take 10ul primer and add 90ul water
  • P6: take 10ul primer and add 90ul water
• Concentration of each primer
  • P5: tm = 63.7C; 100uM in 1187ul
  • P6: tm = 63.4C; 100uM in 747ul

Results:

Further results of this experiment can be seen at 2024.06.08 - Gel Electrophoresis for Vector Confirmation.

Purpose:

• PCR to add Gibson assembly overlap region to ThTdT from IDT

Materials:

• Fwd + Reverse Primer
• 2x Q5 Mastermix
• thermostable TdT sequence from IDT
• Water

Procedure:

Polymerase Chain Reaction (PCR) Protocol

Concentration of primer

• P9: tm=66.4C
• P10: tm = 63.6C

gBlock

• 1000ng, dry
• dilute to 10ng/ul, add 100ul water

Results

See 2024.06.08 - Gel Electrophoresis for Insert Confirmation.

Purpose:

Make Kanamycin agar plate for future cloning

Materials:

• Agarose
• Agar
• Water
• Kanamycin (30ug/ml)

Methods:

Refer to protocol: Antibiotic Resistant Agar Plate and Broth

• Dilute Kanamycin stock (50mg/ml) to 30 ug/ml
  • 150ul in 250ml molten agar
• 8 plates are made, sealed using parafilm.
• Plates are put into a plastic bag, stored in -4 degree cold room.

Results:

Successful plates

Purpose:

• Confirm whether the PCR amplification was successful.

Materials:

• DNA gel loading dye (6x) - Thermofisher
• 1Kb Plus DNA Ladder - FroggaBio
• SYBR Safe
• 1x TAE buffer

Methods:

Refer to protocol: Agarose Gel Electrophoresis

1. Made 65ml of 1% agarose gel in 1x TAE
   1. Add 6ul of SYBR safe in the liquid when its lukewarm
2. Run at 130V for 25min .
3. Gel visualized under UV light

Results:

6th + 7th lanes

Purpose:

• Confirm whether the PCR amplification worked.

Materials:

• DNA gel loading dye (6x) - Thermofisher
• 1Kb Plus DNA Ladder - FroggaBio
• SYBR Safe
• 1x TAE buffer

Methods:

Refer to protocol: Agarose Gel Electrophoresis

1. Make 1% Agarose Gel
   1. 65ml 1x TAE buffer + 0.6g Agarose
   2. Add 6ul of SYBR safe in the liquid when its lukewarm
2. Run at 130V for 25min
3. Gel visualized under UV light

Results:

Important Notes:

• Vectors can be visualized in lanes 3 and 4 shown above. The approximate length is around 5000 — 6000bp.

Purpose:

Isolate PCR products from agarose gel

Materials:

• GeneJET Gel Extraction Kit from Thermo Scientific

Methods:

Refer to protocol: GeneJet Gel Purification

Results:

Important Notes:

• The DNA purified is relatively pure indicated by a distinctive peak. 260/280 ratio is slightly higher than ideal (1.8). 260/230 ratio is lower than ideal (2-2.2). There may be unwanted organic materials.
• The yield is not very high.

Purpose

solate PCR products from agarose gel

Materials:

• GeneJET Gel Extraction Kit from Thermo Scientific

Methods:

Refer to protocol: GeneJet Gel Purification

Results:

After the extraction of the Insert PCR product concentration was determined to be 48.6ng/ul

Purpose:

Ligate together the insert and vector

Materials:

• Gibson assembly master mix from NEB
• ThTdT insert
• pET-28b(+) vector
• water

Methods:

Refer to Gibson Assembly Protocol

Vector: 38.6ng/ul → Need 1.3ul

Insert: 48.6ng → Need 3.1ul

Water: 0.6ul

• No positive or negative control

Results:

Transformation is performed directly after Gibson assembly, for results please see: 2024.06.09 - Transformation

Purpose:

• Have lab members practice the transformation protocol and aseptic technique.

Materials:

• 2 Kanamycin-containing agar plates
• Plasmid DNA
• Chemically-Competent Cells
• Sterile Centrifuge Tubes
• Thermocycler
• Ice Bucket
• Sterile LB Broth
• Glass Beads
• Plate shaker
• Incubator
• p2 + tips
• p200 + tips
• p1000 + tips

Procedure:

Refer to Transformation (General protocol)

1. Mix 1.5 µL of plasmid DNA (~5 ng) into 50 µL of competent cells, with the reaction being on ice the entire time, pipetted up and down, in a 1.5 mL centrifuge tube.
2. Incubated the mixture on ice for 20-30 minutes.
3. Heat-shocked the culture for 45 secs at 42ºC using thermocycler.
4. Placed the tube back on ice for 2 minutes.
5. Added 950 µL of LB broth to the culture.
6. Taped the centrifuge tube against the wall of a falcon tube and placed the tube on a shaker for 60 minutes at 250 rpm.
7. Pipetted an aliquot of 300 µL of the transformed culture onto a Kanamycin-containing agar plate, repeated for a 30 µL aliquot.
8. Plated the bacteria using glass beads and shaking the plate until the surface was dry, then inverted and placed in 37ºC incubator.

Results

Transformation was successful.

Purpose:

Intake of Gibson assembly product (assembled plasmid) into BL21 (DE3) E.coli

Materials:

• BL21 (DE3) E.coli Competent cell
• Gibson Assembly product from the same date:

2024.06.09 - Gibson Assembly for Thermostable TdT

• LB Broth

Methods:

Refer to Transformation (General protocol)

• Plated 30ul, 300ul, and a 1 in 2 dilution to 1/10 on 30ug/ml Kanamycin LB agar plate
• Plated 30ul, 300ul, on 50ug/ml Kanamycin LB agar plate

Important notes:

• No colony can be seen growing on the plate indicating that the transformation has failed

We suspect the following:

1. Heat shock too long
   1. do 10s next time
2. BL21 DE3 is hard to transform
   1. try comp cell from training
3. Gibson Assembly efficiency was low
   1. Use agarose gel to check and see whether the insert and vector as ligated
   2. Try 1hr incubation instead of 30min

• We will begin with using agarose gel to check whether insert and vector as ligated or not

Purpose:

Confirm whether Gibson assembly from 2024.06.09 worked

Materials:

• 2024.06.09 gibson assembly mix
• pET-28b(+) Vector
• Mutant TdT Insert
• Purple loading buffer

Methods:

Agarose Gel Electrophoresis

• 45min at 12V, 40ml 1% agarose gel

Results:

Important notes:

• Looking at Gibson Assembly product, no band longer than vector can be seen, indicating that the vector and insert didn’t assemble together, so the assembly failed.
• Additional lane of around 4000bp is also seen, which may be due to the inserts anneal with each other.

Purpose:

Confirm whether Gibson assembly worked or not

Materials:

• 2024.06.15 gibson assembly mix
• pET-28b(+) Vector
• Mutant TdT Insert
• Purple loading buffer

Methods:

Agarose Gel Electrophoresis

• 0.7% Agarose instead of 1%
• Make 30ml agarose gel for analytical gel
  • 0.21g of Agarose
  • 3ul Sybr Safe
• Run for 1hr at 120V

Results:

• The Gibson assembly product only contains two bands same length of the vector and insert, no longer band was seen, so no sign of ligation after running agarose gel to confirm.
• Gel contaminated, no picture available.

Purpose:

Improve efficiency of Gibson Assembly to troubleshoot 2024.06.09 gibson assembly

Materials:

Gibson Assembly

• Gibson assembly master mix from NEB
• mutant TdT insert
• pET-28b(+) vector
• water

Methods:

1. Add everything according to above
2. 1hr at 50C

Results:

2024.06.15 - Agarose Gel

Purpose:

Isolate and prepare TDT insert PCR products from agarose gel

Materials:

• ABM Gel Purification Kit from ABM

Methods:

Followed procotol from Gene Jet protocol

ABM Column-Pure Gel and PCR Clean-Up Kit

• Used 1:1 Binding buffer ratio instead
• Eluted with 30ul water instead of elution buffer

Results:

NanoDrop results from gel isolation of QingRu’s PCR product in 2024.06.22 - Gel electrophoresis for gel purification.

Purpose:

To confirm whether the PCR amplification worked.

Materials:

Agarose Gel Electrophoresis

• DNA gel loading dye (6x) - Thermofisher
• 1Kb Plus DNA Ladder - FroggaBio
• Sybr Safe
• 1x TAE buffer
• PCR product

Methods:

1. Made 60ml of 1% agarose gel in 1x TAE
   1. Add 6ul of SYBR safe in the liquid when its lukewarm
2. Run at 120V for 30min
3. Gel visualized under UV light

Results:

All except Diego’s PCR products were successful.

Purpose:

To improve the Gibson Assembly, possible reasons were identified:

• Primers with longer overhangs were ordered to allow more efficient assembly. Inverse PCR is performed with longer overhang primer.
• This trial will test the old insert (with 15bp overhang primer: P9+P10) and new insert (with 30bp overhang primer:P3+P4) to see whether Gibson assembly can proceed successfully. New incubation condition will also be tried out to try to improve efficiency.

1. 15bp overhang primer (P9+P10) insert for 45C for 1hr
2. 30bp overhang primer (P3+P4) insert for 50C for 1hr

Materials:

• Gibson assembly master mix from NEB
• Old mutant TdT insert from 2024.06.06 - PCR for ThTDT to Add Gibson Assembly Overlap Region
• New mutant TdT insert from 2024.06.06 - PCR for ThTDT to Add Gibson Assembly Overlap Region
• pET-28b(+) vector
• Sterilized Water

Methods:

Results:

See 2024.06.22 - Transformation attempt 2

Purpose:

To add 30 bp overlap to TdT gene insert to enable Gibson Assembly.

Materials:

• Fwd and rev primers (30 bp overlap: P3+P4)
• 2x Q5 Master Mix
• Thermostable TdT gene sequence from IDT
• Distilled water

Methods:

PCR Protocol

• Add water to dilute the concentration of the primer to 100uM for storage
• Dilute forward and reverse primer to 10uM for reaction
  • Forward, P3: take 10ul primer and add 90ul water
  • Reverse, P4: take 10ul primer and add 90ul water

Results:

• PCR was successful. See 2024.06.22 - Gel electrophoresis for gel purification for agaorse gel results.

Purpose:

Transformation with DH5α strain of E. coli with various gibson assembly products

Materials:

• E. coli DH5α cells

Methods:

Transformation Protocol

• New New = 30bp overhang gibson assembly product
• New old = 15bp overhang, 45C for 1 hr gibson assembly product
• old old = 15bp overhang 50C for 1 hr gibson assembly product
• - CNTL = no insert, only vector 50C for 1 hr gibson assembly product

Results:

• new old = gibson assembly product from 2024.06.15 - Gibson Assembly Attempt 2
  • 250 = 250ul of Transformation product plated
• old old = gibson assembly product from 2024.06.09 - Gibson Assembly for Thermostable TdT
  • all = 300ul of Transformation product plated
• new = gibson assembly product from 2024.06.22 - Gibson Assembly Attempt 3
  • 45 = 45ul of Transformation product plated
  • 250 = 250ul of Transformation product plated
• CNTL = suppose to be linearized vector 2024.06.08 - GeneJet Gel Purification; but might be the original non-linearized vector
  • all = 300ul of Transformation product plated
  • Error observed with -CNTL, likely accidentally used original, non-linearized vector as -CNTL instead of linearized vector made 2024.06.08 - GeneJet Gel Purification, hence the growth of colonies on the -CNTL plate
• All plates had colony growth

Purpose:

Pick colonies from agar plate that has intake TdT plasmid and developed resistance to Kanamycin

Materials:

• LB kanamycin agar plate with transformed bacteria
• LB broth
• Kanamycin

Methods:

Colony Picking

• Pick 8 colonies, put them in 4ml
• Make kanamycin LB media (30ug/mL):
  • 50mg/mLv = 30ug/mL8*4mL
  • v = 19.2ul
  • Add 19.2ul of kanamycin stock to 32mL LB broth
• Pick 8 colonies into kanamycin LB broth
  • 3 colony from new new
  • 3 from new old
  • 2 from -CNTL

Results:

NA

Purpose:

Extract plasmid from dh5α E. coli cells and quantify the plasmid concentration

Materials:

• 3 colony from 30bp overhang gibson
• 3 colony from 15bp overhang 45C 1hr gibson
• 2 colony from -CNTL
  • Overnight incubation for all colonies in LB + Kanamycin broth

Methods:

Miniprep

Results:

Nanodrop results from elute collected through Miniprep

Below are plasmid concentrations from colonies with 30bp overhang gibson:

Below are plasmid concentration from colonies with 15 bp overhang 45C gibson:

Below are plasmid concentrations from the control colonies with just the vector:

Purpose:

• Eliminate false positive plasmid from miniprep

Materials:

• Q5 polymerase
• miniprep product
• 15bp Insert forward primer (P9): “TDT clone Forward 10uM”
• 15bp Insert reverse primer (P10): “TDT clone Reverse 10uM”
• water

Methods:

• 8 samples from 2024.06.24 - Miniprep and Plasmid Quantification
  • 3 new new
  • 3 new old
  • 2 -CNTL

Polymerase Chain Reaction (PCR) Protocol

Results:

Sample labels:

• C1 = -CNTL1
• C2 = -CNTL2 old1, old2, old3 correspond with miniprep names new1, new2, new3 correspond with miniprep names

For -C1, originally added less than 2.5ul P10 (there was a bubble in the primer soon) and added a small amount later to make it about ~2.5ul

Put PCR samples in -20C fridge, will run electrophoresis next day

See 2024.06.27 - Gel Electrophoresis

Purpose:

• To confirm if the insert was ligated into the vector.

Materials:

• Used PCR products from 2024.06.26 - PCR Insert Confirmation of miniprep

Methods:

Agarose Gel Electrophoresis

1. Made 30ml of 1% agarose gel in 1x TAE
   1. Add 3ul of SYBR safe in the liquid when its lukewarm
2. Fill the running tray with 1x TAE
3. Add loading buffer into the
4. Run at 120V, stop when the bottom line of the loading buffer reaches midway through the gel ~30-1hr
5. Gel visualized under UV light
   1. make sure to place the orange light filter on top of the gel before opening the UV light
   2. NEVER look directly into the blue light!!!!!!

Notes during experiment:

• Had to remove the masking tape after loading the wells, so maybe some samples escaped
• Loaded about 5ul samples in each well (5ul sample + 1ul loading dye)

Results:

Lane 5 (Old1) has a band between 1,000 bp and 3,000 bp, so the PCR amplification product likely contains the correct insert.

Legend:

• -Ctrl1-2: No ThTdT gene insert, only vector 50C for 1 hr gibson assembly product
• Old1-3: 15bp overhang 45C for 1 hr gibson assembly product
• New1-3: 30bp overhang 50C for 1 hr gibson assembly product IMG_1995 (original picture of electrophoresis results)

Purpose:

Make Kanamycin agar plate for future transformation and CODE agar art project

Materials:

• Agar
• LB
• Water
• Kanamycin (30ug/ml)

Methods:

Refer to protocol:

• Dilute Kanamycin stock (50mg/ml) to 30 ug/ml
  • 300ul in 500ml molten LB agar mixture
• 24 plates are made, sealed using plastic wrap.
• Plates are put into a plastic bag, stored in -4 degree cold room the second day.

Results:

Plates were successfully prepared.

Purpose:

1. Amplify “old 1” plasmid (generated on June 24th) to make sure we prepare enough TDT plasmid for later use.
2. Pick other colonies from the “new old 15” plate to extract DNA to send for sanger sequencing

Materials:

• dh5 alpha e.coli competent cell
• extracted plasmid DNA “old 1”
• LB Broth
• Kanamycin (50mg/mL)
• “new old 15” plate

Methods

Transformation - Amplify old 1 plasmid

• Mix 2.5 µL of old 1 plasmid miniprep product(~50 ng) into 50 µL of dh5 alpha E.coli
• Mixed 2.5ul of linearized vector product from 2024.06.08 - GeneJet Gel Purification into 50ul of dh5 alpha E.coli as negative control
• Transfer the resulting bacterial suspension into 4mL of LB + kanamycin
  • 50mg/mLV = 30ug/mL5mL*2
  • V = 6ul in 8mL of LB broth

Colony picking

1. Pick 5 colonies from “new old 15” plate, put them in 5ml of LB + kanamycin
2. Make kanamycin LB media (30ug/mL):
   • 50mg/mLv = 30ug/mL4mL
   • v = 2.4ul

Results

Successful inoculation of colonies. Further experiments and results following this experiment can be observed at 2024.07.01 - Miniprep and Plasmid Quantification

Purpose:

Extract plasmid from da5 alpha e.coli and quantify the plasmid concentration

Materials:

• 5 colony from 15bp overhang 45C 1hr gibson
• 1 transformed “old 1” from 06.30
• 1 control vector only

Methods:

Miniprep

Results:

Raw results below:

Nanodrop results from elute collected through Miniprep Below is the plasmid concentration from the supposedly control (vector only) transformed bacteria

• Below is the plasmid concentration from bacteria transformed with “old 1” purified plasmid DNA

• Below are plasmid concentrations from newly picked colonies with 15bp overhang 45C 1hr gibson

Summary:

• The plasmid DNAs are extracted and are ready for sequencing. If transformation was successful, these plasmids DNA should contain sequences that produce TDT.

Purpose:

To verify extracted plasmid DNA from transformed da5 alpha E. coli

Materials:

• Purified plasmid DNA
• P14 - T7 Promoter
• P15 - T7 Terminator

Methods:

1. Used Genewitz to send plasmid (low concentration) for sequencing. Attachments below:

Results:

• Unfortunately, no amplification from the primers we sent.
• There might be a slight amplification in old1 with P15.
• Requested for a free repeat and placed order with Azenta universal T7 Terminator primer.

Purpose:

• Run gel electrophoresis with control and sample from 2024.07.09 - PCR for Plasmid Verification.
• If the sample shows a band of expected size (~1400bp?) then we have verified that the ThTdT insert has been added to the pET+28 vector?.

Materials:

• DNA gel loading dye (6x) - Thermofisher
• 1Kb Plus DNA Ladder - FroggaBio
• SYBR Safe
• 1x TAE buffer
• “Old1” PCR product
• Control PCR product

Methods:

1. Make 30ml of 1% agarose gel in 1x TAE
   1. Add 3ul of SYBR safe in the liquid when its lukewarm
2. Fill the running tray with 1x TAE —> ran out of 1x TAE while filling the electrophoresis tank so we added a bit of old 1x TAE from the electrophoresis station
3. Add loading buffer into the DNA sample(s) (6x loading buffer)
   • For the sample and the control, we took out 5ul from each PCR tube and put it into a new tube, then added 1ul loading buffer to the new tube.
     • Loading buffer was accidentally added to the Old1 PCR tube.
   • We loaded 5ul ladder each time.
   • While loading the first ladder, we lifted the gel to see the wells better and some of the ladder spilled out.
4. Run at 120V, stop when the bottom line of the loading buffer reaches midway through the gel —> it took 40 mins
5. Gel visualized under UV light
   1. make sure to place the orange light filter on top of the gel before opening the UV light
   2. NEVER look directly into the blue light!!!!!!

Results:

Original image of gel

Figure 1. Gel electrophoresis result. Lane 5 to 8 were used for loading ladder and samples, every other well was empty. Lane 6 is the control. Lane 7 is the sample. Lane 8 is the DNA ladder. Visualized using a Dual LED Blue/White Light Transilluminator.

Summary:

• While loading the ladder in Lane 5, the gel was lifted out of the electrophoresis tank and some of the ladder spilled out of the well. This is likely why Lane 1 to 4 shows diffused bands.
• Additionally, the higher bp bands of the ladder have not resolved very well. This could be a result of not running the gel long enough.
• Lane 7 shows a band around 1500bp which is the expect size of the plasmid containing the ThTdT insert.
• We will use the “Old1” miniprep product to transform the plasmid into Top10 E. coli for mass plasmid production and BL21 (DE3) E. coli for protein production.

Purpose:

• Today’s experiment will confirm whether or not the “Old1” plasmid from the MiniPrep 2024.06.24 - Miniprep and Plasmid Quantification contained the insert.
• This insert was used to transform the competent E. coli 2024.06.30 - Transformation + Colony Picking.
• This will be confirmed by conducting PCR of the plasmid and gel electrophoresis.

Materials:

• Q5 polymerase (Master Mix)
• Old1 MiniPrep Sample
• 15bp Insert forward primer, P9: “TDT clone Forward 10uM”
• 15bp Insert reverse primer, P10: “TDT clone Reverse 10uM”
• Nuclease-Free Water

Methods:

Part I - PCR of Old1 Sample

• Old1 Sample from 2024.06.24 - Miniprep and Plasmid Quantification
  • 1x “Old 1”
  • 1x control (nuclease-free water)
• See PCR Protocol for further information: Polymerase Chain Reaction (PCR) Protocol
• Set up program on Thermocycler according to the chart below and waited until lid temperature reached 98C.

Results:

• Prior results were from 2024.06.27 - Gel Electrophoresis.
• See results in 2024.07.09 - Gel Electrophoresis

Purpose:

• To prepare phe-comp E. coli and kanamycin resistant agar plates for C.O.D.E. agar art activity.

Materials & Methods:

• Phe-comp cells (-80 fridge, first shelf, rightmost column all the way to the back)
• LB agar (autoclaved, on bench)
• Kanamycin resistant agar plates (4C room)
• 37ºC incubator room

Results (where to find things):

• Took out 10 agar plates from 4C room (originally took out 12 but put 2 back) and put them on top shelf of second trolley in 37C room
  • labelled “iGEM 06.30.2024 kan 30 ug/ul”
  • some of the plates had condensation on the lid

• E. coli: took out 1 eppendorf tube from phe-comp box and added 1ml LB broth to the E. coli on the bench, then transferred that solution to a loose capped tube (note. some spilled on the lab bench)
  • closed the cap loosely so bacteria don’t die
  • labelled the tube “iGEM phe-comp cells C.O.D.E. 7.10.2024”
  • put on green rack shaker (190 rpm)

Purpose:

• After confirming that the old1 plasmid has the TdT Thermostable part, in this experiment, we are transforming it into BL21 and Top10 E.coli strains, which are ideal for protein expression and high-efficiency cloning respectively.

Materials & Methods:

Refer to Transformation Protocol

1. Mixed 1.5 µL of old1 plasmid DNA (~5 ng) into 50 µL of competent cells BL21 and in another tube TOP10, with the reaction being on ice the entire time, pipetted up and down, in a 1.5 mL centrifuge tube. The negative control for this reaction was 50 µL of competent cells Top10 cells.
2. Incubated the mixture on ice for 30 minutes.
3. Transferred each mixture to a PCR tube.
4. Heat-shocked the culture for 45 secs at 42ºC using a thermocycler.
5. Placed the tube back on ice for 2 minutes.
6. Transferred back to 1.5 mL centrifuge tube.
7. Added 950 µL of LB broth to the culture.
8. Taped the centrifuge tube against the wall of a falcon tube and placed the tube on a shaker for 60 minutes at 220 rpm.
9. Pipetted an aliquot of 30 µL of each of the transformed cultures onto a Kanamycin-containing agar plate.
10. Plated the bacteria using glass beads and shaking the plate until the surface was dry, then inverted and placed in a 37ºC incubator.

Results:

No growth due to the use of wrong old1 plasmid. Results can be seen in the next experiment 2024.07.16 - Plating PHE competent cells after no growth for colony picking. Retrial of this experiment was done on 2024.07.17 - 2nd transformation attempt of old1 plasmid into BL21 and Top10 E.coli strain

Purpose:

• To plate PHE competent cells in a kanamycin agar plate to check if it can grow on its own.
• Original plan was to pick colonies from the transformation experiment performed on 2024.07.15.
• These cells are another chassis that can be used to express the TdT vector.

Materials & Methods:

1. Materials (with measurements etc.)

• 25 uL PHE competent cells
• x1 30ug/mL kanamycin agar plate
• P200 pipette and tip
• 9 glass beads
• parafilm

1. Procedure
   1. Defrost a PHE comp cell tube in ice. Let agar plate warm up to room temperature in the meantime.
   2. Once defrosted, add 25 uL PHE competent cells to the plate.
   3. Add glass beads and shake to spread bacteria.
   4. Wrap plate in parafilm and place in 37C room.
   5. Place cell tube back in -80C fridge. Transformed BL21, top10, and the negative control plates were wrapped and placed in the 4C room after inspection.

Results:

Kept in 37ºC room for further analysis.

Purpose:

• To re-transform old1 plasmid into BL21 and Top10 E.coli strain.
• We discovered that old1 PCR product was used for transformation instead of old1 miniprep DNA product in 2024.07.15 - Transformation of old1 plasmid into BL21 and Top10 E.coli strain.

Materials & Methods:

Refer to Transformation Protocol and Transformation Top 10 e.coli

1. Mixed 1.5 µL of old1 plasmid DNA (~5 ng) into 50 µL of competent cells BL21 and in another tube TOP10, with the reaction being on ice the entire time, pipetted up and down, in a PCR tube. The negative control for this reaction was 50 µL of competent cells Top10 cells.
2. Incubated the mixture on ice for 30 minutes.
3. Heat-shocked the culture for 45 secs at 42ºC using a thermocycler.
4. Placed the tube back on ice for 2 minutes.
5. Transferred samples to 1.5 mL centrifuge tubes.
6. Added 950 µL of LB broth to the BL21 culture, and 250 µL to the TOP10 culture and the negative control.
7. Taped the centrifuge tube against the wall of a falcon tube and placed the tube on a shaker for 60 minutes at 220 rpm.
   1. At some point the rack on the shaker fell off so the samples weren’t shaking for the full 60 minutes. We put the samples back on the shaker for another 15 min.
8. Pipetted an aliquot of 200 µL of each of the transformed cultures onto a Kanamycin-containing agar plate.
9. Plated the bacteria using glass beads and shaking the plate until the surface was dry, then inverted and placed in a 37ºC incubator.
   1. Note: The plates had bumps and there was some liquid on the agar which we assumed was from condensation. Three plates (200ul of each transformation product plated):

• negative control (Top10 E. coli with no plasmid added)
• Top10 E. coli
• BL21 E. coli

Results:

• No growth observed on any of the plates.

• All three plates kept in 37C room for further analysis.
• We will send the linearized vector from 2024.06.06 - Inverse PCR for Vector and “old1” plasmid from 2024.06.24 - Miniprep and Plasmid Quantification for sequencing. On July 24th (7 days after the initial transformation), the plates were checked again. Irregular and sparse colony growth was observed on the negative control Top10 E. coli plate.

Purpose:

• To prepare the linearized pET-28b(+) vector product for whole plasmid sequencing to verify vector sequence.

Materials:

• Linearized vector from 2024.06.06 - Inverse PCR for pET-28b(+) Vector

Methods:

1. Use Plasmidsaurus and followed instructions provided, attachments below:

Vector order sheet.pdf

Screenshot from order page on Plasmidsaurus. The row highligted in green is the one we follow to prepare the vector for sequencing.

1. Measured concentration of vector and Old1 plasmid using Qubit fluorometer.
   1. Old1 plasmid concentration was too low so only prepared vector for sequencing.
   2. Quality of vector sample:
      1. A260/A280 (Nanodrop): 1.98
      2. A230/A280 (Nanodrop): 1.38
      3. Concentration (Qubit): 40.4 ng/ul

Vector

Old1 plasmid

1. Secured samples using bubble wraps in the basement next to the drop off area for packaging
2. Printed order sheet and taped to packaging
3. After prepping the sample, drop it off in B1 of LSI, once you exit the elevator walk straight and you’ll see a table with boxes on it, find the one that says plasmidsaurus and just leave it in there

Results:

Purpose:

• making the EDTA reagent that will be included in the Urea PAGE for visualization of ssDNA formed by the TdT enzyme

Materials & Methods:

• from the following protocol: TBE Urea Gel **500mM EDTA (pH 7.4), 100 mL **

1. Weigh out 18.61 g EDTA disodium salt, dihydrate and add to a 100 mL Duran bottle.
2. Measure out 30 mL distilled water and add to the Duran bottle.
3. Add a stir bar and place on a magnetic stirring plate to mix the solution. The EDTA salt will not go into solution until the pH reaches 8.0.
4. Add a pH meter into the solution to observe the pH.
5. Add enough 1 M NaOH to correct pH using 10 mL serological pipet
   1. Add a drop a time, wait for the pH to stabilize before another drop
6. Once fully dissolved (this will take some time so be patient), top up the solution to 100 mL using distilled water, if necessary.
7. To sterilize, autoclave the solution on a liquid cycle (at least 30 min) 1 aliquot is made from the big bottle and stored in 50ml falcon tube

Results:

• the first reagent that was made ⇒ the volume was overshot
• 1 M NaOH was used but it took more volume than Vt = 100 mL to reach pH 7.4
• note that original volume so the amount of water added to the EDTA powder should be no more than 20 mL
• there was solute at the bottom of the bottle despite pH 7.4 and stirring it
• though the 500 mM EDTA in 100 mL with pH 7.4 being made, the pH 8 one beside it wasn’t made unfortunately due to time constraints.

Figure 1. 500 mM EDTA in 100 mL of deionized water and 1 M NaOH with pH 7.4.

Summary:

• Troubleshooted the initial volume of dH2O
• Solute remained undissolved in the bottle after pH 7.4
• Need to remove the magnetic stir rod using a anti-paramagnetic wand
• In Figure 1., the right flask is unfinished and may need to be remade as there is more than 30 mL of deionized water in there initially and a stronger NaOH solution may be required if this one is to be salvaged.

Purpose:

• This was not an initially scheduled experiment. It was discovered prior to entering the lab that the Gibson assembly product previously prepared was discarded, thus this is a remedial experiment to prepare more for our sequential transformations.
• By the end of the day we should have produced a ligation for both the insert and the Tdt + I1. This Gibson assembly will provide us with vector-bound inserts for future transformation.

Materials:

• 1.5 mL Eppendorf tubes which were labelled as follows:
  • Vector
  • Insert
  • Tdt + I 1
• Distilled Water
• NEBuilder HiFi DNA Assembly Master Mix (stored in the -80ºC freezer door)
  • HiFi master mix
  • Assembly positive control
• PCR strip tubes
• Thermo-cycler
• Micropipette and tips (P2, P20)
• Ice bucket and ice

Methods:

(ADAPTED FROM NEBuilder HiFi DNA Assembly Master Mix) Remove the DNA fragments and the NEBuilder Kit from the -20ºC and -80ºC freezers, respectively, and allow them to thaw on ice. Experimental Trials

1. In a 1:3 ratio of vector:insert, add 50 ng of vector and 150 ng of insert to a PCR strip tube (see Appendix Below)
2. Add 10 uL of master mix to each tube
3. Add enough distilled water so that there is a total volume of 20 uL in each tube following the addition of the DNA fragments and master mix
4. Repeat twice for each insert enough times to produce duplicates of: There should be 4 trials total: 2 x [Vector + Insert], 2x [Vector + (TdT + I 1)] Negative Control
5. Add 50 ng of vector to a PCR strip tube
6. Add 10 uL of master mix
7. Add enough water to achieve a 20uL solution Positive Control
8. Add 10 uL of master mix to a PCR strip tube
9. Add 10 uL of positive control master mix Following preparation, incubate in the thermo cycler for 30 minutes at 50ºC then either store on ice if being used immediately, or store at -20ºC for future use.

Results:

The success of this assembly will be confirmed through the transformation experiments that follow it and plasmid sequencing if deemed necessary based on the transformation results.

Summary:

During the assembly, 6 tubes were prepared and stored in the -20º C freezer for future transformation use. The PCR tubes are labelled as follows: A1: Vector + insert, replicate 1 A2: Vector + insert, replicate 2 B1: Vector + (TdT + I1), replicate 1 B2: Vector + (TdT + I1), replicate 2 (-) : negative control [no insert] (+) : positive control

Appendix:

RAW Calculations for reagent volumes

Transcribed; unrounded values

Purpose:

• The purpose of this procedure is to transform both Top10 and dh5alpha E. coli with the Gibson assembly products prepared on July 23, 2024. Successful completion of this assembly will allow us to move forward with our experimental plan.
• Due to lack of plates, an experimental sample from each Gibson assembly group (A1 from A group, B1 from B group) was transformed with each strain of E. coli, so that we can compare transformation efficiency and results between Top10 and Dh5alpha. This will help troubleshoot previous transformation attempts.
• Positive control gibson assembly sample was transformed into both Top10 and Dh5alpha.
• Negative control gibson assembly sample was transformed into only Dh5alpha, due to lack of plates. Dh5alpha was chosen over Top10 E. coli because we have not tested negative controls on Dh5alpha before. Expected results:
• At least some colony growth among the 4 experimental group plates (if gibson assembly was succesful)
• Colony growth on both positive control plates (Top10 and Dh5alpha)
• No colony growth on negative control Dh5alpha plate Note: if there is no growth in the experimental group, we could try transforming A2 and B2 (but need to make more plates)

Materials:

• Competent *E. coli: *Top10 and Dh5alpha
• Plasmid DNA from 2024.07.23 - Gibson Assembly
  • A1: Vector + Insert
  • B1: Vector + (TdT + I 1)
  • Gibson positive control
  • Gibson negative control
• LB Broth

Procedure:

Adapted from ThermoFisher Routine Cloning Using Top10 Competent Cells and ThermoFisher Dh5alpha Competent Cells Guide Day 1:

1. Thaw competent cells on ice.
2. Mix 1.5 µL of plasmid DNA (~5 ng) into 50 µL of competent cells (reaction has to be on ice all the time) in PCR tubes.
   • Mix by tapping gently/flicking the tube several times. Do not mix by pipetting up and down or vortexing.
3. Incubate mixture on ice for 30 min.
4. Heat shock for 30 secs at 42ºC (using thermocycler set at 42ºC for infinte amount of time and set for 52ul volume).
5. Put tubes back on ice for 2 min.
6. Add 250 µL LB media to samples and transfer mixture into eppendorf tubes.
7. Shake for 60 min. at ~225 rpm, and warm agar plate (with kanamycin antibiotic) at 37C.
   • Taped eppendorf tubes into 15-ml falcon tubes.
   • The samples were put in a shaker in the 37C room that was already running (speed dial was set in the middle of “slow” and “fast”)
8. Plate the bacteria and incubate in 37C room:
   • Plated 100ul of each transformation product and used 4 - 5 glass beads to spread sample on the plates
     • Each sample had ~300ul total volume of tranformation product, so we plated 100ul which is about half of that volume and 100ul is a volume between the 20−200 µL range described the Dh5alpha transformation protocol
   • Plates were flipped upside down, wrapped in parafilm
   • Made two stacks of plates, each stack wrapped in saran wrap
     • 3 Top10 plates
     • 4 Dh5alpha plates Day 2:
9. Check and see whether there is any colony growth on Kanamycin-containing agar plate.
10. If there is growth, this means that the plasmid was successfully incorporated into the bacteria and is expressing the Kanamycin-resistance gene.

Results:

Completed transformation at 9:15 pm, will check the next day (~13 hrs) for results.

• A1 - Top10: no colonies
• B1 - Top10: no colonies
• (+) - Top10: no colonies
• A1 - Dh5alpha: no colonies
• B1 - Dh5alpha: no colonies
• (+) - Dh5alpha: no colonies
• (-) - Dh5alpha: no colonies

Purpose:

• PCR to add Restriction enzyme (XhoI, NcoI) Gibson assembly overlap region to mutant TdT from IDT

Materials:

• P12 + P13 primers
• 2x Q5 Mastermix
• thermostable TdT sequence from IDT
• Water

Procedure:

Polymerase Chain Reaction (PCR) Protocol gBlock

• 1000ng, dry
• dilute to 10ng/ul, add 100ul water

Results

• Please see 2024.07.26 - Gel Electrophoresis & Purification

Purpose:

• To visualize the PCR product from restriction digest and purify the product from the gel

Materials & Methods:

• make 1.5% agarose using 60mL TAE and 0.9g agarose Samples loaded:
• 5ul ladder (2)
• PCR (2) in 10x loading buffer
• Original insert in 10x loading buffer Gel is visualized using gel imager Target bands containing PCR products are cut out and purified according to purification protocol
• Since the cut gel is too big to fit into 1 tube, 2 tubes were used
• 2 purification column are used for the insert
• Gel purification is also done on the digested plasmid DNA concentration is measured using NanoDrop on July 26 DNA concentration of P and I1 are re-measured using Qubit on July 28

Results:

Higher resolution image to be loaded from the LSI computer — will update

• Lane 1: 1kb DNA ladder
• Lane 3: original mutant TdT DNA insert
• Lane 5&6: PCR product
• Lane 9: 1kb DNA ladder

• Looking at lane 5 and 6, thick bands at between 1000 and 1500 bp. PCR is successful.
• Lane 3 has a faint band that is not very visible by eye on this image. The length is between 2000 and 2500 bp.
• Lane 7 seems to have some spill over from lane 6. Nanodrop:
• Digested plasmid (labelled P): 9.6 ng/ul
• Digested insert 1 (labelled I1): 33.6 ng/ul
• Digested insert 2 (labelled I2): 6.9 ng/ul The purity of all the samples do not seem great, no peak visible. Qubit:
• Digested plasmid (P): 16.1 ng/ul
• Digested insert 1 (I1): 31.0 ng/ul

Summary:

• The concentration for the digested plasmid seems a bit low, so restriction digest may need to be redone
• Gibson assembly using digested plasmid and insert are still performed
• Remeasured DNA concentration is used for later Gibson assembly

Purpose:

• The previous transformations have been unsuccessful, so we redid the Gibson assembly to create new plasmids. The product consists of the vector and the Thermostale Tdt insert. This Gibson assembly will provide us with vector-bound inserts for future transformation.

Materials:

• Distilled Water
• NEBuilder HiFi DNA Assembly Master Mix (stored in the -80ºC freezer door)
  • HiFi master mix
• PCR strip tubes which were labelled as follows:
  • Ex1
  • Ex2
  • ctrl
• Thermo-cycler
• Micropipette and tips (P20)
• Ice bucket and ice

Methods:

(ADAPTED FROM NEBuilder HiFi DNA Assembly Master Mix) Remove the DNA fragments and the NEBuilder Kit from the -20ºC and -80ºC freezers, respectively, and allow them to thaw on ice. Experimental Trials

1. In a 1:2 ratio of vector:insert, add 50 ng of vector and 100 ng of insert to a PCR strip tube (see Appendix Below)
2. Add 10 uL of master mix to each tube
3. Add enough distilled water so that there is a total volume of 20 uL in each tube following the addition of the DNA fragments and master mix
4. Repeat twice for each insert enough times to produce duplicates of: Following preparation, incubate in the thermocycler for 30 minutes at 50ºC then either store on ice if being used immediately.

Results:

• The plasmids were transformed 2024.07.26 - Transformation of Gibson Assembly Product into Dh5a and Top10 E. coli strain and the plates should determine if it was successful..

Summary:

• During the assembly, 3 tubes were prepared Ex1 (replicate 1) Ex2 (replicate 2), ctrl (negative control [no insert]). Since we transformed right away we didn’t store them. This was a mistake, which later on affected our experiments, since we wanted to transform the product again later.

Purpose:

Making the 20% 29:1 (bis)acrylamide urea, 7M Urea and loading dye that will be included in the Urea PAGE to visualize ssDNA formed by TdT enzyme.

Materials:

• 3 Duran bottles
• 1 falcon tube to store loading dye Liquid Reagents
• 0.5M EDTA solution (pH 8.0)
• 500mM EDTA (pH 7.4)
• double-distilled H2O
• 40% 29:1 acrylamide:bisacrylamide (ordered)
• Formamide (Hallam in the flammable cabinet) Dry Reagents
• Boric acid
• Tris base
• Urea (Hallam)
• Bromophenol Blue (borrowed)

Methods:

TBE Urea Gel 10X TBE, recipe of 1 L

• 108 g tris base
• 55 g boric acid
• 900 ml double-distilled H2O
• 40 ml 0.5 M EDTA solution (pH 8.0) Adjust volume to 1 L 20% 29:1 (Bis)acrylamide Urea, recipe of 1 L
• 420 g Urea
• 500 mL 40% 29:1 acrylamide:bisacrylamide
• 100 mL 10X TBE
• 90 mL H2O Store in 4 ºC 7 M Urea, recipe of 1 L
• 420 g Urea
• 590 mL H2O
• 100 mL 10X TBE

1. Mix the materials in a 1L bottle
2. Add a stir bar, stir the solution on a hot plate, keep heating (50C +/- 10C) while stirring the solution
   1. You can leave it stirring overnight
3. Once done, cool to RT, store in 4 ºC within 24 hours. Loading Dye (25 mM EDTA/ Formamide/ Bromophenol Blue), recipe of 10 mL

• 500 µL EDTA, 500 mM, pH 7.4
  • prepared from EDTA powder
• 9.5 mL Formamide
• 0.5 mg Bromophenol Blue (add up to 1 mg where appropriate) Store in RT. If precipitate forms, use the supernatant.

Results:

• Solutions have been made and are in the inventory.

Purpose:

Transform Top10 and dh5alpha E. coli with 2024.07.26 - Transformation of Gibson Assembly Product into Dh5a and Top10 E. coli strain. Successful completion of this assembly will allow us to move forward with our experimental plan. Expected results:

• At least some colony growth among the 4 experimental group plates (if gibson assembly was succesful)
• Colony growth on both positive control plates (Top10 and Dh5alpha)
• No colony growth on negative control Dh5alpha plate

Materials:

• Competent *E. coli: *Top10 and Dh5alpha
• Plasmid DNA from 2024.07.23 - Gibson Assembly
  • Vector+insert replicate 1
  • Vector+insert replicate 2
  • Gibson negative control
• LB Broth

Procedure:

Adapted from ThermoFisher Routine Cloning Using Top10 Competent Cells and ThermoFisher Dh5alpha Competent Cells Guide Friday, 26th July

1. Thaw competent cells on ice.
2. Mix 1.5 µL of plasmid DNA (~5 ng) into 50 µL of competent cells (reaction has to be on ice all the time) in PCR tubes.
   • Mix by tapping gently/flicking the tube several times.
3. Incubate mixture on ice for 30 min.
4. Heat shock for 30 secs at 42ºC (using thermocycler set at 42ºC for infinite amount of time and set for 52ul volume).
5. Put tubes back on ice for 2 min.
6. Add 250 µL LB media to samples and transfer mixture into eppendorf tubes.
7. Shaked at 200 rpm overnight.
   • Taped eppendorf tubes into 15-ml falcon tubes.
   • The samples were put in a shaker in the 37C room that was already running (speed dial was set in the middle of “slow” and “fast”)

Saturday, 27th July

1. Plate the bacteria and incubate in 37ºC room:

Sunday 28th July:

1. Check and see whether there is any colony growth on agar plate.

Results:

Plates will confirm if transformation it was successful.

Summary:

• A positive control was also required (unlinearized pET-28b vector), yet the members working on this experiment, only did a negative control.
• Additionally, after the transformation, the bacteria were left overnight in the mixer at 37ºC. This happened due to the fact that the members had been working in the lab since 3 pm and at 9:30 pm when the transformed bacteria were finally placed in the mixer, the members went home instead of waiting an hour and then plating.
• Unfortunately, the members doing the experiments did not had the foresight to consider that the bacteria might loose the plasmid, since they were grown in LB without antibiotic resistance. This oversight might result in the experiment to be inconclusive.

Purpose:

• To transform Top10 and Dh5a strain with Gibson assembly products
• To test the growth of wt bacterial strains

Materials & Methods:

Protocol from Top10 and dh5a website

Top10 competent cells

DH5Alpha Competent Cells

• 2ul of DNA is added to each sample.
• 100ul of transformed bacteria are plated

Conditions:

Results:

Growth

July 28th: Checking plates (after ~17 hrs?)

Monday, 29th July:

1. Check again and see whether there is any colony growth on agar plate.

Summary:

• The 4 plates (kanamycin) with the gibson assembly A2, B2 for both dh5alpha and top10, no growth
• The top10 and DH5alpha plates (2) without kanamycin grew
• Plates (2) without kanamycin, with cells dh5alpha and top10 having the B2 assembly mix, grew
• The plates (kanamycin, 2) with the positive control having the original plasmid Pet28b, no growth
• The plate (kanamycin) with the negative control having water in DH5alpa, no growth

Purpose:

Plate transformed Top10 and DH5a E. coli cells from 2024.07.26 transformation experiment. Expecting out of 7 plates:

• growth for transformed DH5a + insert 1, DH5a + insert 2, Top10 + insert 2 cells in kanamycin (3)
• no growth for the negative DH5a and Top10 controls in kanamycin (2)
• growth for the negative DH5a and Top10 controls in no-kanamycin (2) Getting results as expected would allow us to proceed to colony picking to select the transformed cells

Materials & Methods:

Materials (with measurements etc.)

• x5 kanamycin agar plates
• x2 regular agar plates
• glass beads
• Transformed cells from 2024.07.26

Procedure (should be VERY detailed, enough to be replicated)

1. Place agar plates in 37C room to warm up
2. Once warmed, label each plate for which cells will be plated
3. Pipette 100uL cell mix into each corresponding plate, pipetting up and down to mix before adding the mix
4. Add ~9 glass beads and shake horizontally to spread the cells. Once done, carefully remove the beads over a waste container into the waste tube.

Results:

The transformed cells from 2024.07.27 were plated and labelled accordingly. They are put in the 37C room, first rack to the right, on the second highest shelf. They are grouped as follows:

• Transformed DH5a and Top10 (3)
• Negative control on kan+ (2)
• Negative control on kan- (2)

Sunday, 28th July: Checking the plates (after ~19 hrs?) After checking the plates, they were returned to 37C room to incubate longer.

Monday, 29th July:

1. Check again and see whether there is any colony growth on agar plate.

Summary:

• The positive control (plates with no antibody) both Top10 and DH5alpha grew
  • DH5a and Top10 controls in no-kanamycin grew colonies as expected, which indicates that the DH5a and Top10 E. coli are viable on their own.
• The negative controls (plates with antibody) neither Top10 or DH5alpha no growth
  • This indicates that the plates with antibiotic are effective in preventing bacteria without antibiotic resistance from growing.
• the 3 plates containing the gibson assembly products from 26.07.2024, no growth
• From this we assume that the cells nor the plates are the problem.
• Thus the problem is either the gibson assembly, the transformation protocol, or the antibiotic.

Purpose:

• After assessing the results of the transformed plates from 2024.07.27 - Transformation of Gibson Assembly Product into Dh5a and Top10 E. coli strain, continued and 2024.07.27 - Transformation of Gibson Assembly products into Dh5a and Top10 E. coli strain, replicate we adjusted the Gibson assembly and transformation protocols to troubleshoot issues with transforming Top10 and Dh5a strain with Gibson assembly products.

Materials & Methods:

Gibson assembly:

protocol source: Experimental Trials

1. In a 1:2 ratio of vector:insert, add 50 ng of vector and 100 ng of insert to a PCR strip tube (see Appendix Below)
2. Add 10 uL of master mix to each tube.
3. Add enough distilled water so that there is a total volume of 20 uL in each tube following the addition of the DNA fragments and master mix
4. Incubated samples at 50C for 30 mins

Kanamycin stock:

Labeled as Kanamycin stock 50 mg/ml and is in the -20 fridge.

Ampicillin Stock

protocol link:

Labeled as Amp 100mg/ml iGEM (on the side the date says 7/28/2024) and is in -20 box in -20 fridge

Transforming and plating

• 2ul of DNA is added to each sample of E. coli
• 100ul of transformed bacteria are plated
• plates were spread with silica beads then incubated in 37C room at around 4:30pm **Top10 **

1. Thaw, on ice, one 50 µl vial of One Shot cells for each ligation/transformation.
2. Pipet 2 µl of each ligation reaction directly into the vial of competent cells and mix by tapping gently. Do not mix by pipetting up and down. The remaining ligation mixture(s) can be stored at -20°C.
3. Incubate the vial(s) on ice for 30 minutes.
4. Incubate for exactly 30 seconds in the 42°C water bath. Do not mix or shake.
5. Remove vial(s) from the 42°C bath and place them on ice.
6. Add 250 µl of pre-warmed S.O.C medium to each vial. S.O.C is a rich medium; sterile technique must be practiced to avoid contamination.
7. Place the vial(s) in a microcentrifuge rack on its side and secure with tape to avoid loss of the vial(s). Shake the vial(s) at 37°C for exactly 1 hour at 225 rpm in a shaking incubator.
8. Spread 20 µl to 200 µl from each transformation vial on separate, labeled LB agar plates. The remaining transformation mix may be stored at +4°C and plated out the next day, if desired.
9. Invert the plate(s) and incubate at 37°C overnight.
10. Select colonies and analyze by plasmid isolation, PCR, or sequencing. Source from: https://www.thermofisher.com/ca/en/home/references/protocols/cloning/competent-cells-protocol/routine-cloning-using-top10-competent-cells.html DH5alpha protocol Note: for DH5a we made to to incubate on ice for exactly 2 minutes.
11. Thaw competent cells on wet ice. Place the required number of 1.5-mL polypropylene microcentrifuge tubes on wet ice.
12. Gently mix the cells, then make 50 µL aliquots of competent cells in the chilled 1.5-mL microcentrifuge tubes.
13. Add 1−5 µL of sample DNA directly into a tube of competent cells. Mix well by gently flick the tube several times.
14. Incubate the cells on ice for 30 minutes.
15. Heat-shock the cells for exactly 30 seconds in a 42°C water bath. Do not mix or shake the tube.
16. Incubate the cells on ice for 2 minutes.
17. Add 250 µL of room-temperature S.O.C. Medium.
18. Place the tube on its side in a shaking incubator. Use tape to secure the tube in place.
19. Shake the tube at 225 rpm for 1 hour at 37°C.
20. If necessary, dilute the cells 1:10 with S.O.C. Medium.
21. Spread at least two differet volumes (20−200 µL) of cells from each transformation reaction on separate LB plates containing the appropriate selective antibiotic. Label the plates with the plating volume.
22. Invert the plates and incubate overnight at 37°C. Source from: https://www.thermofisher.com/document-connect/document-connect.html?url=https://assets.thermofisher.com/TFS-Assets%2FLSG%2Fmanuals%2FMAN0018520_DH5alpha_competent_cells_UG.pdf Preparing Amp plate for positive control:
    • prepared working solution of 2mg/ml Amp from 100mg/ml Amp
      • using a sterilized glass spreader, spread 150 ul of 2mg/ml Amp on LB (no antibiotic) plate in presence of bunsen burner followed protocol from:

• originally wetlab lead suggested 50mg/ml Amp to spread over the plate (based on the ideal antibiotic conentration for the plasmid from the gibson assembly positive control solution) but WL members missed the message
  • thus the positive control from this experiment may be unreliable

Results:

Summary:

1. Interpret the results that you attained during this lab session.
   1. What does the data mean?
   2. How do the results relate to the iGEM project aims as a whole?
2. Explain why the results can be trusted. Do you have controls that show expected results which inform the validity of experimental results?
3. Are the results similar to the literature or do they diverge from expectations?
4. Discuss any issues you may have experienced during this session. Any possible sources of error? How are you going to troubleshoot them?
5. What are you gonna be doing moving forward?

Appendix

raw calculations for Gibson Assembly Bottom half is raw calculations for 1.5ml of kanamycin stock (but we ended up making 10ml so these calculations were not used)

Purpose:

• The original purpose was to check the plates from 2024.07.28 - Gibson assembly with R.E. Vector and Insert, transformation, Kan and Amp stock and older plates for colonies. One plate of transformed cells from 2024.07.27 - Transformation of Gibson Assembly Product into Dh5a and Top10 E. coli strain, continued showed two colonies, so the experiments proceeded as follows:

1. Culture the colonies in LB broth + 30ug/ml kanamycin to check if they contain plasmid
   1. 4 mL per tube with 2.4 uL Kanamycin
2. Run 1% agarose gel electrophoresis of restriction enzyme product 7.28 C1 (plasmid with insert), pET-28b (positive control), and P (restriction digest product) to check if the plasmid from 202
3. Make 1.5 mL aliquots of 50mg/mL kanamycin stock from the 15 mL falcon tube for easier use.

Materials & Methods:

• x2 4mL LB media
• 50mg/mL kanamycin stock
  • x2 2.4uL for colony picking
  • x2 1.5 mL for smaller aliquots
• x2 culture tubes
• x4 Eppendorf tubes
• P1000 tips
• P20 tips
• P2 tips
• x2 10mL stripettes
• 2 mL stripette See below protocols for detailed procedures: 2024.06.23 - Colony picking 2024.06.27 - Gel Electrophoresis 2024.05.04 - Kanamycin Agar Plate and LB Broth Preparation Alterations to above procedures include:
• Aseptic technique used for colony picking and used an inoculation loop for one colony
• Below are the volumes used for each well:

Figure 1. Contents of each lane and the volume of sample and loading buffer added.

Results:

Figure 2. Plates from 2024.07.28 - Gibson assembly with R.E. Vector and Insert, transformation, Kan and Amp stock with no colonies on any plates. Many bubbles and droplets observed.

• The transformations seem to have failed as no colonies were produced even in the positive control condition.
• Refer to figure 4, as the lanes with the DNA samples had no bands other than one band for the linearized plasmid lane at the right.

Figure 3. Top10 E. coli Transformants observed on plate with kanamycin. Two colonies circled.

• The two colonies were picked into two culture tubes. One was picked with a P2 tip and the other was picked with an inoculation loop. The tubes were labeled accordingly.

Figure 4. Gel electrophoresis product of DNA samples visualized by gel imager. Refer to figure 1 for the lanes and their corresponding contents. Left to right in figure 1 is the same as left to right in the gel image above.

Summary:

• To our surprise, we found two colonies exhibiting E. coli colony phenotype on a plate from 2024.07.27 - Transformation of Gibson Assembly Product into Dh5a and Top10 E. coli strain, continued

• This plate contains Kanamycin as written on the lid and says that it has the insert in it so we are hopeful that the colonies we have picked have taken up the recombinant plasmid and are resistant to Kanamycin.

• We expect the turbulence of the culture tubes to increase telling us that the cells are Kanamycin resistant and are expressing the TdT enzyme, with the assumption that they do not have innate antibiotic resistance which will be further tested via DNA extraction and gel electrophoresis at a later date.

• It seems that the transformations conducted during 2024.07.28 - Gibson assembly with R.E. Vector and Insert, transformation, Kan and Amp stock failed as no colonies were produced.

Calculations:

1. volumes needed for kanamycin stock
   1. Colony picking c1 = 50mg/mL c2 = 30ug/mL v2 = 8ug/mL v1 = c2*v2/c1 = 4.8 uL (2.4 uL kan stock per 4 mL LB)
   2. LB agar media prep 500 uL per 500 mL, so 1 mL kan stock for 1L LB agar media

Purpose:

• We hope to observe high bacterial density in the culture tubes that contain the colonies picked from the LB Kanamycin plate containing two colonies of Top10 *E. coli *assumed to contain the plasmid and TdT insert.
• By running colony PCR and gel electrophoresis of the subsequent PCR product, we hope to observe that the plasmid with the TdT insert have been successfully transformed into the bacteria.
• We hope to make progress on our project and prepare for optimization by preparing 3 bottles of LB broth and making 10% APS and sample loading buffer for UREA PAGE.

Materials & Methods:

2024.06.26 - PCR Insert Confirmation of miniprep

2024.06.27 - Gel Electrophoresis 2024.07.29 - Transformation Plate Review, Colony Picking, Gel Electrophoresis Samples to test:

• cells in LB Broth with kan *2 (1 and 2) July 29
  • restriction digest primer, P12+P13
• Dh5a Insert 2 July 27
  • restriction digest primer, P12+P13
• Dh5a B2 July 27
  • 30bp primer, P3+P4
• Top10 A2 July 27
  • 15bp primer, P9+P10
• Dh5a Control July 27
  • restriction digest primer, P12+P13
• Positive control Ammonium Persulfate, APS, 10% w/v, recipe of 10 mL
• Dissolve 1 g APS in 10 mL H 2 O Store in 4ºC, away from light. Loading Dye (25 mM EDTA/ Formamide/ Bromophenol Blue), recipe of 10 mL
• 500 µL EDTA, 500 mM, pH 7.4
  • prepared from EDTA powder
• 9.5 mL Formamide
• 0.5 mg Bromophenol Blue (add up to 1 mg where appropriate) Store in RT. If precipitate forms, use the supernatant.

Results:

• the figure below can be followed in order to understand what was in each of the lanes; the top of the note is from right to left which is downwards the note

Summary:

• Transformation was unsuccessful.

Purpose:

• After verifying that the previous Gibson Assembly product does not contain the expected plasmid, this experiment aims to redo Gibson Assembly using restriction digest products.
• The products will be analyzed using gel electrophoresis, and if expected products are observed, transformation will be done

Materials & Methods:

Gibson Assembly mix:

Transformation condition:

Results:

Gel electrophoresis:

• Lane 2 & 6: 1000kb DNA ladder
• Lane 3: Gibson Assembly products
• Lane 4: digested vector P
• Lane 5: Gibson Assembly positive control
• 4 bands visible in lane 3. We suspect the one band slightly higher than Lane 4 is our expected Gibson assembly product, which should have the digested vector integrated with insert.

Summary:

• This gel indicates that the Gibson assembly is most likely successful. Transformation is then proceeded.

Purpose:

Given the recent failures regarding the transformations of our construct into e.coli, we decided to confirm that the gibson assembly had worked and that we had correctly purified the inserts.

Materials & Methods:

Preparing Media for LB agar plates

Following the instructions from Agar plate and broth preparation and based concentration of antibiotics from:

Materials:

• LB powder (labelled “iGEM”, at the weighing station)
• Agar powder (labelled “iGEM”, at the weighing station) 1L LB agar for kanamycin plates
• The protocol x4 in two 1L bottles, each having 500mL.
• 300 uL of 50mg/ml Kanamycin was input per 500 mL bottles of LB agar. 500mL LB agar for plates with no antibiotic
• The protocol x2 in one 1L bottle. 250mL LB agar for ampicillin plates
• Exactly the same protocol in one 500 mL bottle.
• 250 uL of 100mg/ml Ampicillin was input into the LB agar bottle. The plates were finalized and placed in the -4ºC.

Rerunning PCR analysis

Following protocol: Polymerase Chain Reaction (PCR) Protocol

Materials

• Master Mix (2x Q5) (-20 iGEM box)
• Molecular grade water (small bottle at lab bench)
• Broth 1 (from 37C room, incoulated with colony from
• Broth 2 (from 37C room, incoculated with colony from
• Insert 1 “i1” (restriction digest insert (-20 iGEM box)
• “7.28 C1” (gibson assembly with restriction digest Vector + Insert)
• Restriction digest forward primer, P12 (-20 iGEM box)
• Restriction digest reverse primer, P13 (-20 iGEM box) After checking the condition of Broth 1 and 2, both 1 appeared more cloudy (indicating more colony growth) so less broth was used for PCR.

Gel Electrophoresis

Following protocol from: Agarose Gel Electrophoresis

Materials

• Agarose powder (at the weighing station)
• 1x TAE (by the sink next to eletrophoresis station)
• 6x NEB Purple Gel Loading Dye (at gel eletrophoresis station)
• 1kb DNA Ladder Froggabio (-20 iGEM box)
• SYBRSafe (from pink rack on iGEM bench)

1. First run: (5 ul sample + 1 ul 6x NEB Purple Loading dye), 120V for about 40 mins
   • Lane 1, 2, & 7: 5ul Ladder
   • Lane 3: 5ul Broth1 PCR
   • Lane 4: 5ul Broth2 PCR
   • Lane 5: 5ul Insert1 PCR
   • Lane 6: 5ul 7.28 C1 gibson product PCR
2. Visualized the gel once the red/pink part of the loading dye travelled halway down the gel
3. Second run: Using the remainng 3 wells of the gel
   • Lane 7: 3ul 7.28 gibson product (just the product itself, no PCR)
   • Lane 8: 5ul Insert1 (just the product itself, no PCR)
   • Lane 9: Ladder

Results:

Fig 1. Gel eletrophoresis results from initial run.

• If colonies in the broth contain a plasmid with ThTdT insert we expect to see a band at ~1500bp:
  • Lane 3 and 4: Both broth PCR products do not show a 1500bp band, this indicates they did not have the desired plasmid with ThTdT insert but somehow grew on kanamycin resistance plates
• Lane 5: After amplifying the insert using the P12+P13 used to added restriction enzyme overlap regions in 2024.07.25 - PCR for Restriction Digest Insert, Insert 1 PCR does not show a band at 1500bp which is unexpected
• Lane 6: No band at 1500bp, but this lane was not supposed to be run (see results for 2nd gel eletrophoresis run below)

Fig 2. Gel eletrophoresis results from second run of the same gel.

• Lane 7: Expect to see a few bands, but no visible bands at all, indicating that the C1 Gibson Assembly product does not contain the expected plasmid
• Lane 8: Insert 1 is a bit above 1500bp as expected, indicating that “i1” did contain the purified ThTdT with added restriction enzyme overhang region
  • Comparing with Lane 5, this verifies that the PCR amplification of insert 1 DIDN’T fail because the wrong template was used

Summary:

Prepared LB agar plates for future experiments:

• 8 Ampicillin resistant plates, labelled “iGEM 100 ??”
• ?? Kanamycin resistant plates, labelled “iGEM 30 ??”
• ?? LB plates (no antibiotic), labelled “iGEM ??” After running PCR amplification and gel electrophoresis analysis, we confirmed that:
• broth 1 and broth 2 did not contain the desired plasmid
• the latest gibson assembly product did not contain the expected plasmid, thus why 2024.07.28 - Gibson assembly with R.E. Vector and Insert, transformation, Kan and Amp stock 8353e5d46a324f948c950569ab7ffbe5) was unsuccesful
• insert 1 was purified correctly

Purpose:

• Redo transformation using competent cells

Materials & Methods:

Gibson assembly:

• 1 for restriction digest products
• 1 for products from inverse pcr using 30bp primers

• run at 50c for 30min

Transformation:

Results:

See plate results in

2024.08.02 - Colony PCR + Colony picking

Purpose:

Transformants observed so we’ll be conducting a colony PCR followed by gel electrophoresis of the PCR product confirm plasmid incorporation.

Materials & Methods:

2024.07.09 - PCR for Plasmid Verification

• The PCR products were placed into the gel in the following order from LEFT to RIGHT:

  • Lane 1 = DNA Ladder
  • Lane 2 = Positive Control colony PCR product
  • Lane 3 = Negative Control (no bacteria)
  • Lane 4 = 100 ul bp30
  • Lane 5 = 50 ul bp30
  • Lane 6 = DNA Ladder 2024.06.23 - Colony picking

• (4 mL of LB broth + 2.4 uL of diluted Kanamycin) x4 in culture tubes

• 1 colony per tube (2 tubes for each dilution: 100 µL and 50 µL of Bp 30 TdT Transformed Competent DH5α cells)

Results:

• no growth observed on the following plates Fig 1. No colonies observed on plates.
• more colonies observed on the following plates when I returned to the lab Fig 2. Chemically competent DH5α successfully transformed with plasmid containing TdT insert.
• Insert observed in 50 µL Bp 30 PCR product in the gel electrophoresis product shown below:
  • Lane 1 (left) = DNA Ladder
  • Lane 2 = Positive Control colony PCR product
  • Lane 3 = Negative Control (no bacteria)
  • Lane 4 = 100 ul bp30
  • Lane 5 = 50 ul bp30 (has TdT insert)
  • Lane 6 (right) = DNA Ladder

Fig 3. Gel electrophoresis of colony PCR products

Summary:

• We have a transformant that has the TdT insert after weeks of trying.
• This confirms that the cells were not competent previously and by using competent cells we have E. coli with the plasmid.
• It is strange to not see the insert in Lane 4 as it was the same treatment just with a higher volume plated.

Purpose:

• Perform colony PCR to amplify the insert region for the 4 samples from 2024.08.02 - Colony PCR + Colony picking

• Verify if the colonies contain the insert/desired plasmid

Materials & Methods:

• Master mix of 6 samples are made
• 24ul of master mix is added to each pcr tube, 1ul of each sample is then added

The resulting solutions are ran on TAE gel and are sent for sequencing to verify the sequence.

Results:

• All the colonies have the desired insert at the correct band size.

Plasmidsaurus sequencing result: Plasmid 1 and 2 has the correct transformation product:

• Plasmid 1 - 1 mutation
• Plasmid 2 - 2 mutations

Summary:

• Plasmid 3 and 4 failed to amplify

Purpose:

• Extract plasmid from dh5a e.coli and quantify the plasmid concentration

Materials:

• 4 colonies transformed with 30bp overhang TDT insert, incubated overnigh
  • 1: colony that has been verified with insert with colony PCR
  • 2: “50ul plated colony”
  • 3: “100ul plated colony #1”
  • 4: “100ul plated colony #2”
• GeneJet Plasmid Miniprep Kit

Methods:

Miniprep

Results:

Raw data below:

Summary

• The miniprep successfully extracted plasmid DNA from the overnight bacteria samples Plasmid 1 and plasmid 2 had the correct sequences

Plasmid 1:

4VP7YG_1_1_-_50ul.fastq

• 1 silent mutation

Plasmid 2:

4VP7YG_2_2_-_50ul.fastq

• 2 silent mutation

Purpose:

• prepare Urea gel for WT testing

Materials & Methods:

TEMED 10% solution:

• Measured 0.6g of TEMED powder in fume hood
• Add 600ul water Remake APS 10% as we cannot find the old one:
• weigh 1g of APS powder, add 10ml water
• wrap in Alum foil to prevent exposure from light Both TEMED and APS are stored in 4c cold room

Purpose:

• To visualize WT TdT DNA addition on Urea-PAGE gel.

Materials & Methods:

• WT TdT from NEB
• Primer P8
• 10x TdT reaction buffer
• dNTPs (ATCG)
• molecular grade water

For materials needed for Urea PAGE and protocol, see TBE Urea Gel

Design:

1. Dilute 100mM dNTP to 1mM dNTP
   1. 1 in 100 — 1/100 dilution
2. Dilute TdT
   1. 1 in 9 — 1/10 dilution 5ul loading dye of each ul of sample

• 50ul for 10ul sample
• Both primer and negative control are added with loading dye Samples loaded:
• Start with lane 3
• Lane 1 and 2 are not washed with TBE, so samples in those 2 lanes are reloaded
  • Lane 3: A
  • Lane 4: T
  • Lane 5: C
  • Lane 6: G
  • Lane 7: negative control
  • Lane 8: primer/ladder
• The cast runs within ice to cool down the cast with high voltage (400V)
• During imaging, we realize that the primer we used does not have a fluorescence tag, so the bands cannot be visualized. The gel is incubated in SYBRSafe for 15min before imaging again using green laser Cy3 Gel staining protoco

Results:

No bands visible

Summary:

• Next trial should use fluorochrome-tagged primers so no extra step of staining is needed
• 20% gel should be used for visualization of better separation

Purpose:

• Try transforming Miniprep generated TDT containing plasmid into BL21 for protein mass production. Also, to generate more plasmid in dh5a.

Materials & Methods:

1. Transformation of the following cells:

Results:

• Tube of Dh5α is used for plasmid extraction.
• Plates checked on August 5th, 2024.
• Wrap with parafilm and stored in 4c for further use.

Purpose

• Extract plasmids from transformed cells

Materials & Methods:

Miniprep

https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0013117_GeneJET_Plasmid_Miniprep_UG.pdf

1. Spin down 25ml of transformed dh5a from
   1. 4000rpm for 10min

2024.08.04 - Transformation of TdT-containing Plasmid to BL21

1. Resuspend in 500ul of resuspension solution
2. perform first step in 2 eppendorf tubes

Results:

Plasmid concentration: 90.1 ng/ul

• purity relatively ok

Purpose

• The purpose of this experiment is to quantify the concentration of the plasmids miniprepped on 2024.08.03 - Miniprep for picked colonies from 08.02, which we expect to have our construct in its plasmid. We had previously measured its concentration using a less accurate method, Nanodrop.
• Lastly the 10 uL of the plasmids were sent for sequencing to plasmidsaurus.

Materials & Methods:

Dna Quibit

• Qubit® dsDNA HS Reagent (Component A) —Stored next to the qubit machine at room Temp, which is in one of the benches next to the computer.
• Qubit® dsDNA HS Buffer (Component B) —Same as component A
• Qubit® dsDNA HS Standard #1 (Component C) —Stored in 4ºC in one of the boxes on the right as you walk in, on the floor underneath the rack (Always keep on ice).
• Qubit® dsDNA HS Standard #2 (Component D)—Stored in 4ºC (Always keep on ice)
• Special qubit tubes next to the Qubit machine
• Plasmids
  • 1: colony that has been verified with insert with colony PCR
  • 2: 50ul plated colony
  • 3: 100ul plated colony #1
  • 4: 100ul plated colony #2

Results:

Plasmid 1: (Incorrect)

Plasmid 2: (Incorrect)

Plasmid 3: (Incorrect)

Plasmid 4: (Incorrect)

Summary:

• Sadly, the results are way to high and most likely incorrect. This might have been due to some mistake when creating the working solution or perhaps a reagent gone bad, yet unlikely.
• The experiment was going to be repeated, but there were not enough qubit tubes available, thus we had to send the plasmids for sequencing knowing only the NanoDrop concentration.

Purpose:

• Transformation of BL21 with plasmid #1 + #2

Materials & Methods:

Refer to protocol: https://www.neb.com/en/protocols/0001/01/01/transformation-protocol-for-bl21-de3-competent-cells-c2527

• Both BL21 samples are transformed onto LB-Kanamycin (LB-Kan) plates
• Both DH5α samples are directly transformed and amplified in LB-Kan broth
• Make LB-Kan broth: 25ml (30ug/ml) x 3 tubes
• For each tube: 50mg/ml * v = 25ml * 30ug/ml
• v = 15ul in 25mL LB media
• Leftover transformed BL21 are combined and add to another LB-Kan broth
• All the broth is 25ml with the addition of 15 µl Kan stock (conc. = 50mg/ml)

Results:

• Observed colonies on both plates, growth in all broths.
• Broth pictures are not available

Summary:

• Successful transformation with plasmid #1 and #2

Purpose:

• To store BL21 bacteria containing plasmid #1 for long-term use.

Materials & Methods:

1. Make 50% glycerol by mixing 4mL of pure glycerol with 4mL of dH2O
2. Add 500ul of overnight BL21 bacteria TDT plasmid #1 culture into 500ul of 50% glycerol
3. Freeze in -80 freezer. Directly thaw for future use.
   1. A total of 15 vials are generated

Results:

• All cryo vials are stored in the -80C freezer in IGEM box. Ready for future use.

Purpose:

• To extract plasmids from transformed bacteria

Materials & Methods:

Miniprep

Using plates generated from

2024.08.08 - Transformation of TDT-containing Plasmid to BL21 (with plasmid #1)

Results:

Purpose:

• To visualize WT TdT DNA addition on Urea-PAGE gel.

Materials & Methods:

• WT TdT from NEB
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• molecular grade water

For materials needed for Urea page and protocol, see TBE Urea Gel

Design: Ladder Negative control Reaction 1 Reaction 2 Reaction 3 Reaction 4 Master mix of 5 reaction Primer 0.5pmol 5ul of 100mM 0.5pmol 0.5pmol 5ul of 100mM 0.5pmol 5ul of 100mM 0.5pmol 5ul of 100mM 0.5pmol 5ul of 100mM 25ul dNTP / G A 0.1ul of 10mM 1ul of 1mM T 1ul of 1mM C 1ul of 1mM G 1ul of 1mM TdT buffer / 0.5ul 0.5ul 0.5ul 0.5ul 0.5ul 2.5ul TdT / / 0.1ul of original 1ul for 1/10 dilution 1ul for 1/10 dilution 1ul for 1/10 dilution 1ul for 1/10 dilution Water / 3.5ul 2.5ul 2.5ul 2.5ul 2.5ul 12.5ul Total 5ul 10ul 10ul 10ul 10ul 10ul 40ul each 8ul Reaction time / 5 min 5 min 30 min 5 min 30min 5 min 30min 5 min 30 min Load all - 5ul 5ul 5ul 5ul 5ul 5ul

After 5 min, take out 5ul of each tube, add to 25ul loading dye Leave the rest 5ul for another 25min reaction Notes: there are bubbles present in the gel made. When loading samples, 2 gels are used, lanes are selected to load to exclude any with bubbles.

Results:

Lanes from left to right: Primer only, negative control, A, T, C, G

Lanes from left to right: A, T, C, G, Primer only

We notice that the lanes shifted outwards, possibly due to no samples were loaded in the center. Lanes at the edges ran out of the frame. The pink lane shows saturation, indicating the primer concentration might be too high.

Summary:

• Despite seeing over saturated bands, no bands with longer length were visible, indicating the TdT did not add nucleotides to the primer. Follow-up research shows that even WT TdT needs to function at 37 C, while we incubated it at RT, so this maybe why. Future trial will try incubating TdT at 37 C.
• Future trial will use a lower concentration of primer to minimize saturation.

Purpose:

Note. This experiment was performed in the Perin Lab with Graduate Advisor Antonio Wang

• To visualize WT TdT DNA addition on Urea-PAGE gel.

Materials & Methods:

• WT TdT from NEB
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• molecular grade water

For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

Design: Stock 0 1 2 3 4 5 6 7 8 9 10 11 12 Final Conc ALL Bfr 10 X 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1 X ALL TdT 20 U/µL 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 250.0E-9 0.5 U/µL ALL 5’-Cy5 Primer 1.0E-6 M 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 1.0E-6 100.0E-9 M MASTER 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 2.3E-6 1 dCTP 1.0E-6 M 3.0E-6 300.0E-9 M 2 dCTP 10.0E-6 M 3.0E-6 3.0E-6 M 3 dCTP 50.0E-6 M 4.0E-6 20.0E-6 M 4 dATP 1.0E-6 M 3.0E-6 300.0E-9 M 5 dATP 10.0E-6 M 3.0E-6 3.0E-6 M 6 dATP 50.0E-6 M 4.0E-6 20.0E-6 M 7 dTTP 1.0E-6 M 3.0E-6 300.0E-9 M 8 dTTP 10.0E-6 M 3.0E-6 3.0E-6 M 9 dTTP 50.0E-6 M 4.0E-6 20.0E-6 M 10 dGTP 1.0E-6 M 3.0E-6 300.0E-9 M 11 dGTP 10.0E-6 M 3.0E-6 3.0E-6 M 12 dGTP 50.0E-6 M 4.0E-6 20.0E-6 M W 7.8E-6 4.8E-6 4.8E-6 3.8E-6 4.8E-6 4.8E-6 3.8E-6 4.8E-6 4.8E-6 3.8E-6 4.8E-6 4.8E-6 3.8E-6 0 1 2 3 4 5 6 7 8 9 10 11 12

Results:

Summary:

1. Successful DNA addition is observed in all four dNTPs. However, dCTP seems not to work as efficiently as the rest. Further research into dCTP may be required.
2. Other than dCTP, this can be a great reference and baseline for our thermostable mutant TdT.

Purpose:

• Induce BL21 E.coli transformed with plasmid 1 (mutant TdT) with IPTG for mass mutant TdT production

Materials & Methods:

Protocol Page Link: Shake in 50ml

1. Shook 3ml of LB broth with 5ul of Plasmid 1 transformed BL21 glycerol stock for 1hr
2. Divide up 3ml of LB broth into two 1.5ml broth and put into 50ml falcon tube and top up to 25ml LB broth, shake for 2.5hrs
3. Add 500uM IPTG (25ul of 1M stock concentration) and shook after night
4. Measure OD

Results:

• For unpurified BL21 protein lysate please see 2024.08.19 - SDS-PAGE Confirmation for mutant TdT
• OD for uninduced BL21 - 1.483
• OD for IPTG Induced BL21 - 2.578 B - Blank (LB Broth only) NI - No IPTG Induction I - IPTG Induction

Summary:

• The IPTG Induced E. coli grew better than no IPTG Induction E. coli, overall the growth was significant and we will proceed to *E. coli *Lysis step.

Purpose:

• Follow protocol provided by Beth Davenport from UBC Hallam lab to lyse IPTG induced E.coli from 2024.08.16 - BL21 Protein Expression. The unpurified protein will then by proceed to extract mutant TdT.

Materials & Methods:

Protocol:

• Matrix A from MP Biomedical
• Homogenizer
• E.coli

Procedure:

• 1. Resuspend pellets
  • resuspend cell pellets (from overnights, for example) with binding buffer on ice
  • add to purple lidded tube with sonication beads
  • keep on ice
• 2. Lysis with sonicator
  • load tubes into white tube holder boxes - the 2 must balance
  • tighten boxes until they dont move
  • settings: 30seconds, 20Hz frequency
  • Run for 30s, then rest tubes on ice 2 minutes (sonication heats up samples and don’t want proteins to denature)
  • repeat this three times (should get cloudier each time, cell debris should rest on top of beads)
• 3. Centrifuge
  • take tubes on ice and tabletop centrifuge to cold room
  • centrifuge 30 minutes, max speed (14,300 rpm)
  • supernatent is the lysate - take this and use it in SDS Page etc.
  • can store purple lidded tubes with rest of cell debris and lysate (supernatent) in -20 freezer

Results:

See 2024.08.17 - Mutant TdT Extraction

Purpose:

• Extract mutant TdT from BL21 whole E.coli lysis using anti-His Magnetic beads from NEB

Materials & Methods:

Protocol:

1. Unpurified protein lysate from 2024.08.17 - E. coli Lysis was run through the magnetic beads following the NEB protocol
2. Extracted 200ul of mutant TdT in total

Results:

See 2024.08.19 - SDS-PAGE Confirmation for mutant TdT

Purpose:

• Run SDS-PAGE of mutant TdT to check purity and molecular size

Materials & Methods:

• SDS-PAGE gel pre-made
• Pre-Made SDS-PAGE running buffer
• Comassie stain

Protocol for SDS-PAGE

1. Use pre made SDS-PAGE and running buffer
2. Run at 80V for the stacking gel and 120V for the resolving gel

Protocol for Coomassie stain

1. After SDS-PAGE, put membrane in container and rinse with tap water (drain out tap water from container after rinse)
2. Pour coomassie stain over the membrane until the membrane is fully covered
3. Shake for 1 hour (slow speed)
4. Remove coomassie stain (can pour it back to stock solution)
5. Pour de-stain solution until membrane is fully covered
6. Place scrunched up kimwipes in the container around the edges of the membrane to help soak up excess dye
7. Shake for overnight

Results:

• WT TdT molecular weight ~ 58kDa

Summary:

• The mutant TdT has been successfully induced in BL21 and successfully extracted using the anti-His magnetic beads from NEB

Purpose:

To prepare glass microscopic slides with functional groups, SH and NH, which is required for the oligo to stick to the glass microscopic slide. The glass should have exposed OH groups if the procedure works correctly. This is the beginning of the preparation for SPS.

Materials & Methods:

Materials Used

• Sulfuric acid (98%)
• Hydrogen peroxide (30%)
• Sodium bicarbonate (500 g powder, ACS grade)
• pH papers
• APTES (3-Aminopropyl)triethoxysilane (Ambeed, ACS grade) (Antonio will bring)
• Acetone (ACS grade)

• MPTS
• EtOH [Anhydrous]
• Acetic Acid [1.7485 M]

Protocols:

• Glass Slide Preparation biotin
• Glass Slide Preparation thiol

Procedure

Preparation of Piranha solution

All of this work occurs in the fumehood.

1. A solution of 60 g sodium bicarbonate was prepared with 300 mL. The solution does not need to be dissolved.
2. 30 mL of sulfuric acid was measured into glass graduated cylinder poured into a beaker equipped with stir bar on stirrer. The graduated cylinder was rinsed with the sodium bicarbonate solution made in step 1.
3. 10 mL of hydrogen peroxide was measured into a glass graduated cylinder, then added drop-wise so the beaker with sulfuric acid, taking care to adjust the rate based on the bubbling seen.
4. The solution is then mixed with the stir bar for [??].
5. The resulting mixture is very hot, and chilled on an ice bath.
6. After the mixture is cooled, pour onto a wide glass plate or beaker, and place glass microscopic slides, making sure the slides are one layer.
7. Leave the slides in the solution for overnight.

Preparation of 1% MPTS, 95% EtOH and 16mM acetic acid

1. Into a 50 mL falcon tube, was measured, with the 1000 ul micropipette, 0.5 mL MPTS, 47.5 EtOH and fill to the 50 mL line with 16mM acetic acid*.*

Preparation of Vectabond/acetone (1% v/v)

1. Into a 50 mL falcon tube, was measured, with the 1000 ul micropipette, 0.5 mL APTES and 50 mL acetone*.*

Preparation of Acetic Acid [16 mM]

1. Into a 50 mL falcon tube, was measured, with the 1000 ul micropipette, 457 ul acetic acid [1.7485 M] and declorinated (instead of deionized) water*.*

Controls

Will be in next notebookwebp entry.

Purpose:

• To determine if the extracted mutant thermostable TDT is able to functionally add dNTP to the primer by running Urea PAGE Gel.

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• molecular grade water

For materials needed for see TBE Urea Gel

Made 2 Urea PAGE Gel according to the following: TBE Urea gel

Sample preparation according to the following excel sheet:

Results:

Gel 1:

First 4 lanes using WT TdT:

• Old sample from

2024.08.14 - LPS with WT TdT (Trial 3)

• Sample 0 (primer)
• Sample 3: dATP (0.3uM)
• Sample 4: dATP (3uM)
• Sample 4: dATP (20uM)

Rest: Mutant TdT

• Sample 13 - 16

Gel 2: Mutant TdT

• Sample 1 - 12

• All conditions of mutant TDT saw no nucleotide addition.

Summary:

• The mutant TDT is either not functioning or other reagents interfere with mutant TDT function.

Purpose:

• To determine if the extracted mutant thermostable TDT is able to functionally add dNTP to the primer by running Urea PAGE Gel.

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

WT

MT1 (one brand of dNTPs)

WT2 (different brand of dNTP)

Master mix of mutant TdT are made together

Results

Lane loaded (left to right in order):

• Primer only (0)
• Old sample from sample 9: dTTP (20mM)
• sample 1 - 12
• primer (0)

Summary:

• Sample 2 - 5 use WT TdT from NEB. Since addition is observed, this indicates that the other components of the reaction are good.
• No addition is observed for all the reactions involving mutant TdT, showing that it is the mutant that is not working. Further investigation is needed.
  • Since we used Imidazole for purification of our mutant TdT, we suspect there may be a possibility that the imidazole will affect mutant TdT’s function. For next trial, purification column will be first applied to the mutant TdT extract before proceeding to downstream DNA addition reaction.

Purpose:

• Prepare 95% EtOH and 16mM acetic solution for rinsing glass slides
• Rinse one thiol glass slide

Materials & Methods:

Using protocol from Glass Slide Preparation thiol Modifications:

• Using parafilm instead of gloves Procedure outline:

1. Prepared 40mL of 95% EtOH and 16 mM acetic acid (see calculations below)
2. Rinsed glass slide under DI water for 10 mins
3. Rinsed each side of the glass side once using ethanol usin P1000 micropipette
4. Submerged glass slide functionalized side up in 1% MPTS 95% EtOH 16mM acetic for 30 mins
5. Placed functionalized side up inside a beaker, wrapped in 2 layers of parafilm.
6. Pierced hole with glass pipette for venting, then pierced another hole through the parafilm to displace air with argon from filled balloon.
7. Wrapped 2 more layers of parafilm to seal holes and keep argon inside beaker.

Results:

Summary:

• the slides have been made and stored

Purpose:

• To further purify the extracted protein.

Materials & Methods:

1. 14,000 rpm 10 minutes spin for step 7
2. After first spin, flow-through is re-added to the column for another spin
3. 100 µL of TdT buffer is added, followed by another spin
4. The overall flow-through + eluted protein is combined and rerun once again later
5. Protein concentration is measured using NanoDrop

Results

• Protein concentration: 0.19mg/ml

Summary

• Protein was purified.

Purpose:

• To generate enough LB broth for future bioreactor experiments

Materials & Methods:

• LB broth powder
• Water

1. Mix 12.5g of LB broth powder in 500mL of water.
   1. 4 bottles
2. Autoclave LIQUID20

Results:

• The 4 LB broth bottles are stored on the bench.

Summary:

• Enough broth has been made for future experiments.
• More will have to be made if contaminated.

Purpose:

• To resuspend purified His protein 2+3 in water to remove the imidazole that might interfere with the mutant thermostable TDT function.

Materials & Methods:

Refer to the above Sigma protocol; additionally:

1. Used 100ul of Purified his protein 2+3
2. Spinned at 14000g for 20 minutes
3. Exchanged buffer by adding 400ul of RNase free water
4. Spinned at 14000g for 15 minutes
5. Reverse spin at 1000g for 2 min (2 times) to collect the concentrated protein in water

Results:

• Collected 0.4mg/ml 20ul of purified proteins.

Summary:

• The purified proteins are ready to be use for liquid phase.
• This may have resolved our issue with MT TdT unable to add nucleotides onto the primer.

Purpose:

• To test effectiveness of the bioreactor against traditional bench-top methods for bulk production of liquid bacterial culture. Using our transformed BL21 E. coli, a large amount of culture can be generated and used for future experiments.
• For the first trial, we want to compare the yield of BL21 E. coli cultivated by the Mk. 1 bioreactor with the traditional method of shaking a culture flask at 37°C. for approximately 10 hours.
  • The timeframe of 10 hours was determined based on general protocols for generating E. coli growth curves (Aryal, 2023) and due to time limitations of manual OD measurement readings.
  • Measurement timepoints were determined based on the fact that the average doubling time for E. coli is 20 minute (Tuttle, 2021).

Materials:

• Disposable cuvettes (minimum 10 needed)
• 1x Spectrophotometer
• 1x 1-litre Erlenmeyer flask
• 1x BL21 #1 glycerol stock vial (transformed BL21 E. coli with ThTdT-containing plasmid #1)
• 400 mL of LB Broth media
• 50 mg/mL Kanamycin stock
• Mk. 1 bioreactor

Methods:

LB Broth stock was prepared on 08.31.2024.

1. Thaw a vial of BL21 #1 glycerol stock made on 08.09.2024.
2. Take 200 mL of LB Broth for each condition (200 for control condition shaking at 37°C, 200 for bioreactor).
3. For each condition, add Kanamycin to the removed media (not the LB Broth stock) to a final concentration of 30 µg/mL.
4. For each condition, pipette 5 µL of BL21 #1 glycerol stock into the media.
5. For the bioreactor, place a large magnetic stir bar into the container and place it on a heating plate (set to 37°C).
6. Take 1 mL of each condition at designated times for OD measurement.
   • Always use Kanamycin LB broth as control.
   • Start with timepoint 0 hours.

Results:

Observations during experiment

After testing Mk. 1 bioreactor with water, the fan wasn’t working so it was sent back for repairs. Carried on experiment with control condition.

• To sterilise bioreactor container, remove the filter then flush the tubing with isopropoanol. Avoid letting fluid flow through the filter.

To prepare the control condition, an autoclaved 1-litre Erlenmeyer flask containing Kanamycin LB Broth was prepared as described above.

• A clean 50-mL falcon tube was used to measure out 200 mL of LB Broth.
• Loosely wrapped the top of flask with aluminum foil.
• Masking tape was used to secure flask in place in the 37°C incubation room. The shaker speed was set to approximately 210 rpm.
• Culture flask was left shaking overnight after 4 hours of OD measurements.

Kanamycin antibiotic calculations:

• v = volume needed from stock Kanamycin

For control condition:

• 50 mg/mL * v = 200 mL * 30 µg/ml
• v = 120 µl in 200mL LB Broth

For blanking spectrophotometer:

• 50 mg/mL * v = 10 mL * 30 µg/mL
• v = 6 µL in 10 mL LB Broth

O.D. Measurements

Relevant Figures

Figure 1. Flask was secured to shaker. The culture looked clear. It was left to shake overnight at 7:40 pm.

Figure 2. Flask was removed form shaker at 2:30 pm (almost 24 hours of incubation at 37°C). The culture looked very cloudy. A foam of bubbles was formed due to vigorous shaking.

Figure 3. Image of spectrophotometer reading at 24 hour timepoint. The machine was blanked with 1 mL of LB media containing 30 µg/mL of Kanamycin antibiotic.

Figure 4. Image of cuvettes for 24 hour measurement. Blank sample is on the left, 24 hour sample is on the right.

Summary:

This experiment demonstrated that for the control condition, a larger amount of BL21 #1 glycerol stock would have to be used to start the maxiculture to generate enough reliable OD measurements values within a 10-hour timeframe for comparison with bioreactor condition for future bioreactor testing.

References:

Aryal, S., & Y, P. (2023, August 3). Bacterial growth curve protocol. Microbe Notes. https://microbenotes.com/bacterial-growth-curve-protocol/

Tuttle, A. R., Trahan, N. D., & Son, M. S. (2021). Growth and Maintenance of Escherichia coli Laboratory Strains. Current protocols, 1(1), e20. https://doi.org/10.1002/cpz1.20

Purpose:

• The BL21 E. coli containing plasmid #1 maxiculture created from 2024.09.01 Mk. 1 Bioreactor Trial 1 experiment will be used to produce more ThTdT via IPTG induction.

Materials:

• 1x 200-mL Erlenmeyer flask
• 200 mL maxiculture from 2024.09.01 Mk. 1 Bioreactor Trial 1 experiment
• 100 mL of LB Broth media
• 1M IPTG stock reagent (keep on ice)
• 50 mg/mL Kanamycin stock

Methods:

Modified from BL21 IPTG induction protocol.

1. Add 100 mL of LB Broth into 200mL Erlenmeyer flask
   • Add Kanamycin to a final concentration of 30 µg/ml.
2. Add 4 mL of the maxiculture into the flask, then loosely wrap the opening of the flask with aluminum foil (not airtight), and shake at 37°C, 220 rpm.
3. After 2.5 hours of incubation or when the culture reaches 0.50-0.8O OD (whichever occurs first), add IPTG.
   • If the culture doesn’t reach 0.5-0.8 OD after 2.5 hours, just add IPTG.
   • Add IPTG to a final concentration of 500 µM.
4. Shake at 37°C, 220 rpm overnight (16 - 18 hours).

Results:

Observations during experiment

1. Started shaking the incoulated 100 mL culture at 4:02pm, with shaker set at 196 RPM (not 220 rpm because the shaker was shared with other lab personnel).

2. Periodically removed flask from shaker to check OD (each measurement resulting in the culture not shaking for 5 minutes):

3. Put culture back in 37C room to shake overnight
   1. Secured with paper towels
4. Disposed of E. coli BL21 plasmid 1 starting culture by pouring in bleach, swirling and dumping down the sink with tap water running to dilute bleach solution

Kanamycin antibiotic calculations:

• v = volume needed from stock Kanamycin
• 50 mg/mL * v = 100 mL * 30 µg/ml
• v = 60 µL in 100 mL LB Broth

IPTG calculations:

• v = volume needed from stock IPTG
• 1M * v = 97 mL * 500 µM (or 0.0005M)
• v = 0.0485 mL or 48.5 µL 1M IPTG in 97 ml culture

Figure 1. 1 mL samples were taken from the BL21 culture to check OD via spectrophotometry. From left to right the cuvettes correspond to: Blank (Kanamyci LB Broth), sample after 35 minutes incubation, 1 hour incubation, and 2.5 hours incubation.

Figure 2. BL21 culture flask after 2.5 hours of shaking at 37°C. A tube of Kanamycin LB broth is placed on the left for comparison of media appearance. The culture media was cloudy but would still let through the blue colour from a glove passing behind the flask.

Figure 3. BL21 culture was left shaking at 37°C overnight at 196 RPM.

Summary:

BL21 E. coli containing plasmid #1 was induced with IPTG and left to grow for protein production. We will proceed with subsequent ThTdT extraction and purification steps.

Purpose:

• Prepare suitable buffer to store purified ThTdT.

Materials & Methods:

Prepare 10mL of 0.5M KPO4

• Weigh 114.249mg of Potassium Phosphate dibasic
• Weigh 46.83mg of Potassium Phosphate Monobasic
• Add 10mL of dH2O Prepare 10mL of 1M NaCl
• Weigh 0.5844g of NaCl
• Add 10mL of dH2O Make: Storage buffer
• 50 mM KPO4
• 100 mM NaCl
• 1.43 mM β-ME
• 50% glycerol
• 0.1% Triton X-100
• pH 7.3 @ 25C KPO4: (10mL)(50mM) = (0.5M)(X) X = 1mL NaCl: (10mL)(100mM) = (1M)(X) X = 1mL b-ME (10mL)(1.43mM) = (14.3M)(X) X = 1ul 50%glycerol X = 5mL 0.1% Triton X (10mL)(0.1%) = (10%)(X) X = 1mL Rest dH2O = 10mL - 5mL - 1mL -1mL - 1mL = 2mL

1. Mix KPO4, NaCl and b-ME first and filter through 0.22uM filter
2. Add glycerol and triton X after

Results:

See 2024.09.03 - Protein purification using centrifugal filter

Summary:

• The new buffer is around pH 7.0 and is ready to use.

Purpose:

• To determine if the extracted mutant thermostable TDT (ThTdT) is able to functionally add dNTP to the primer by running Urea PAGE Gel.

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

Design:

• Purified ThTdT Condition:
• concentration gradient
  • 0.5, 3 ul
• Temperature gradient
  • 37, 47
• With/without CoCl2
  • 0, 1ul
• Samples are prepared in 10ul, divide 5ul to a new tube.
• Incubate one at 37ºC for 30min.
• Incubate the other one at 47C for 30min.

• Master Mix 1:

• Master Mix 2:

• Lysate:

Samples are subjected to 4 different conditions:

1. 37ºC for 1 min.
2. 37ºC for 30 min.
3. 47ºC for 1 min.
4. 47ºC for 30 min.

Results:

• Purified ThTdT Protein:

• Unpurified Lysate

Summary:

• DNA addition is observed for purified protein** at all conditions**
  • without CoCl2, the protein does not perform as good, only single addition is observed for 0.1ug/ul protein condition.
  • Smear is seen in 0.6ug/ul protein condition.
  • with CoCl2, longer bands are produced. Therefore, it seems that CoCl2 facilitates the DNA addition to a greater extent.
  • Our enzyme can function at both 37ºC and 47ºC with comparable activity.
• No DNA addition is observed in lysate reactions
• Two bands are observed in primer only lane, possibly due to primer degrade overtime.

Purpose:

• To purify mutant TdT proteins by exchanging its buffer to the new storage buffer that is used.2024.09.02 - Protein Purification Storage Buffer Preparation

Materials & Methods:

https://www.sigmaaldrich.com/deepweb/assets/sigmaaldrich/product/documents/105/563/pr05485w-ug-ms.pdf

Refer to the above sigma protocol; additionally;

1. Used 200ul of Purified his proteins and top up to 500ul using the storage buffer
   1. 495ul storage buffer + 5ul 10mM 2,2’-bipyridyl
2. Spun at 14000g for 10 minutes
3. Exchanged buffer by topping up to 500ul of storage buffer
4. Spun at 14000g for 10 minutes
5. Reverse spin at 1000g for 2 min (2 times) to collect the concentrated protein in water

Results:

Summary:

Purpose:

• To test our mutant TdT to see if it can allow DNA addition in certain conditions
  • Here we are testing the condition without CoCl2, and at** higher **dNTP concentrations

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

Design:

• 10ul reaction is divided to 5, incubated at 37ºC and 47ºC separately for 30 minutes

Results:

Summary:

• Single nucleotide addition is observed for 20uM dCTP condition with CoCl2 addition, under both 37ºC and 47ºC.
• We can conclude that cobalt chloride is necessary for dCTP addition using ThTdT.
• No dATP, dTTP or dGTP addition was observed in this experiment.
• The other conditions without cobalt chloride do not show obvious DNA addition.

Purpose:

• To create Urea gel mixture for making Urea PAGE gels in future optimization experiments.

Materials:

For materials needed for see TBE Urea gel.

• Urea (solid form, white colour)
• 40% 29:1 acrylamide:bis-acrylamide stock (kept in 4ºC room)
• 10X TBE stock
• ddH2O

Methods:

20% 29:1 (Bis)acrylamide Urea, recipe of 1 L

• 420 g Urea
• 500 mL 40% 29:1 acrylamide:bis-acrylamide
• 100 mL 10X TBE
• 90 mL ddH2O Store in 4 ºC

20% 29:1 (Bis)acrylamide Urea, recipe for 200 mL

• 84g Urea
• 100 mL 40% 29:1 acrylamide:bis-acrylamide
• 20 mL 10X TBE
• 18 mL ddH2O Store in 4 ºC

Results:

• Measured out urea powder into beaker
• Added ddH2O, 40% 29:1 acrylamide:bis-acrylamide and 10X TBE (measured with graduated cylinder)
• Mixed with stir bar until urea appeared fully dissolved in solution (~30 min)
• Poured solution into 250 mL bottle
• Stored in 4ºC room

Figure 1. Image of finalized urea gel mixture solution.

Summary:

Prepared 20% 29:1 (Bis)acrylamide Urea solution, ready to use to create Urea PAGE gels for visualizing oligonucleotides.

Purpose:

• To characterize ThTdT using temperature gradient, test the temperature range where ThTdT can function

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

Design:

Make master mix, then divide 3ul to one tube for a total of 18 tubes for each condition.

Primer only lanes:

Reaction condition:

The temperature gradient is achieved using the gradient pcr protocol on a thermocycler.

Primer only conditions were incubated at 37 degree for 30 minutes.

Results:

Figure 1. Urea PAGE showing dCTP and dTTP addition by ThTdT after 5 minutes.

Figure 2. Urea PAGE showing dCTP and dTTP addition by ThTdT after 15 minutes.

Figure 3. Urea PAGE showing dCTP and dTTP addition by ThTdT after 30 minutes.

Summary:

ThTdT adds dCTP and dTTP effectively across 37C-55C with at least 5 minutes of reaction time. However, the primer concentration is either too high or the primer is not being utilized effectively by ThTdT.

Purpose:

• To validate P2 primer can be extended by WT TdT with full primer utilization, i.e. primer is consumed quantitatively

Materials & Methods:

• WT TdT from NEB
• Primer P2
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

Set up reactions tubes 0-8 according to the following. The first three rows (Bfr for Thermopol Buffer, WT TdT and fluorescent primer P2 are premixed as master (see right most column).

For example, tube 2 received 2.3 µL master mix, 1.0 µL dCTP, and 6.8 µL H_2O.

At time = 3, 10, and 15 min, 2 µL samples were removed to be suspended in 18 µL loading dye. 5 µL of each quenched sample was loaded onto the gel, which is equivalent to 50 fmol.

Results:

Processed file

Figure 1. 3’-extension by wild type terminal transferase (WT TdT) at 37ºC, using [Primer P2] = 100 nM. 20% D-PAGE, 40 min, 400 V. Cy3 mode detecting fluorescein is overlaid onto Cy5 mode detecting bromophenol blue. The overlaid images demonstrated non-uniform migration front.

Raw file (for traceablilty purposes only)

Figure 2. Raw image to produce Figure 1.

Hand off:

All samples generated in this experiment were handed off to @Diego for further uses in LSI.

Summary:

1. Complete primer utilization is observed.

2. CoCl_2 improves WT TdT activity.

3. 3 min reaction is sufficient for all nucleotides when CoCl_2 is used.

Purpose:

• Perform ThTdT vs. WT TdT liquid phase synthesis experiments at 37ºC, 47ºC, and 55ºC to demonstrate ThTdT function at higher temperatures and loss-of-function of WT TdT at 47ºC and 55ºC

• Observe addition of all dNTPs to the primer

• Culture 4x100 mL from transformed *E. coli *BL21 ThTdT stock and perform IPTG induction to produce more ThTdT.

Materials & Methods:

• ThTdT
• Primer P1
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

• The following table are compositions for the reaction tubes using WT TdT and ThTdT respectively. A master mix was made with WT TdT and ThTdT separately.

The following are the reaction conditions tested

24 reaction tubes (4 bases x 3 temps x 2 proteins) + 2 negative controls (WT, ThTdT with primer only at 37C)

1.2 uL dNTPs were added to each PCR tube first before 1.8 uL master mix.

1.2 uL water was added to primer only/negative control (0) in place of dNTP, then add 1.8ul of master mix

When preparing the master mix the primer was put last

Results:

Figure 1. WT liquid phase elongation of all dNTPs (C,A,T,G) at different temperatures. From the left, the first band shows the negative control of the primer without dNTP. A blank space. The following 3 bands are all elongations using dCTP at 37ºC, 47ºC, and 55ºC, respectively. The next 3 bands constitute elongations with dATP at the same temperatures in the same order. Then follows dTTP and lastly dGTP.

Figure 2. Mutant liquid phase elongation of all dNTPs (C,A,T,G) at different temperatures. From the left, the first band shows the negative control of the primer without dNTP. A blank space. The following 3 bands are all elongations using dCTP at 37ºC, 47ºC, and 55ºC, respectively. The next 3 bands constitute elongations with dATP at the same temperatures in the same order. Then follows dTTP and lastly dGTP. 

• The urea PAGE for ThTdT was done with only one gel loaded, with the other side using a buffer gate. Voltages and durations were otherwise the same.

• Another 20% 29:1(Bis)acrylamide Urea was made since the last one (9.5.2024) got contaminated. The following entry was used to make the new one 2024.09.05 - Making 20% 29:1 (Bis)acrylamide Urea mixture.

• Roughly 300 mL of 1x TBE was prepared to top up the existing bottle.

Summary:

• Liquid phase synthesis was performed to compare elongation ability of all dNTPs at a range of temperatures between WT TdT and ThTdT. Urea PAGE was used to show elongation.

• WT TdT shows elongation with all bases at 37ºC. Elongation rate drops as temperature rises but WT TdT remains functional. WT TdT elongates dGTP at a far slower rate than other dNTPs.

• Mutant TdT (ThTdT) results are inconclusive. It will need to be redone since the small elongation is inconsistent with results from 2024.09.07 - LPS with ThTdT (Trial 5) which tested elongation over a similar range of temperatures of ThTdT on dCTP and dTTP.

• Some possible errors on the ThTdT gel may have been not thawing the reagents enough for liquid phase synthesis, incorrect amounts in the master mix,

Purpose:

• Prepare the mother culture to be used for the bioreactor growth rate measurement project

• Reactivate the mother culture and start bioreactor growth rate measurements

Materials & Methods:

1. Aliquot 10 µL of E. coli BL21 ThTdT-positive 15% glycerol stock into 200 mL of LB-Kanamycin media.

2. Place in 37ºC incubator and leave to shake.

3. Leave overnight for 16 hours.

Refer to the following protocol for next steps:

Bioreactor Efficacy Evaluation

Results:

• The mother culture was made and the growth curve analysis took place. Please refer to the following notebook entry to observe the experiment.

2024.09.09 - Mk. 1 Bioreactor Growth Rate Measurements Preparation

Summary:

• The mother culture was made.

Refer to the following notebook entry:

2024.09.09 - Mk. 1 Bioreactor Growth Rate Measurements Preparation

Purpose:

• Based on inconclusive results for ThTdT from 2024.09.08 - LPS with ThTdT (Trial 6), perform mutant vs. WT TdT liquid phase synthesis experiments at 37ºC, 47ºC, and 55ºC to show mutant TdT function at higher temperatures and LOF of WT TdT at 47ºC and 55ºC.

• Observe addition of all nucleotides CGAT to the primer

Materials:

• Terminal Transferase (WT TdT)

• Terminal Transferase Buffer (Bfr)

• CoCl2

• Diluted 5’-Cy5 primer (1 uM) (Primer P1)

• Diluted dNTPs: dCTP (1 mM), dATP (1 mM), dTTP (1 mM), dGTP (1 mM)

• Purified ThTdT aliquot (PCR tube labelled “P”)

• Urea PAGE reagents

• Quenching reagents

Equipment:

• Thermocycler(s) set to correct temperatures

• Gel cassette, electrode chamber, electrophoresis tank, power source

• Amersham TYPHOON biomolecular imager

• Timer

Methods:

• The experiment followed the protocol described in TdT reaction + TBE Urea gel (modified)

• The following table are compositions for the reaction tubes using WT TdT and ThTdT respectively. A master mix was made with WT TdT and ThTdT separately.

• The following are the reaction conditions tested:

September 9th: Sample preparation

• 24 reaction tubes (4 bases x 3 temps x 2 proteins) + 2 negative controls (WT, ThTdT with primer only at 37C).

  • mixed by pipetting and light tapping (not vortex)

• Order of reagents added to mastermix: Buffer, Water, CoCl2, TdT, primer

• 1.2 uL dNTPs were added to each PCR tube first before 1.8 uL master mix for a total reaction volume of 3ul per sample.

• 1.2 uL water was added to primer only/negative control (0) in place of dNTP, then add 1.8ul of master mix.

• Samples were incubated in thermocyclers:

  • Machine 1 set to 37-55C gradient (37C in the front row, 55C in the back row) for infinite time

  • Machine 2 set to 47C (entire block) for infinite time

• Samples were quenched, put in an empty pipette tip box and left in -20C fridge for continuation of experiment the next day.

• Urea PAGE gels were wrapped in water-soacked paper towels, covered in saran wrap and left in -4C room to use for next day

September 10th: Running Urea PAGE and imaging

• Gels from previous day had bubbles, remade gels using the same protocol

• Thawed samples from previous day (on ice):

• Loaded gels according to the following diagram:

  • 47C condition for dTTP was skipped so added at the end

  • Ran gels for 40 minutes

Results:

Figure 1. Wild type (WT) TdT reaction with different dNTPs, over a range of temperatures. The first lane is the negative control, excluding dNTPs from the reaction mixture. To the control’s right are three lanes showing the incorporation of dCTP nucleotides at three different temperatures: 37°C, 47°C, and 55°C. Next is dATP incorporation using the same order of temperature conditions, followed by dTTP and lastly dGTP.

**Figure 2. **ThTdT reaction with different dNTPs, over a range of temperatures. The first lane is the negative control, excluding dNTPs from the reaction mixture. The next lane is a gap that separates the control from the rest of the samples. To the right of the gap are three lanes that show the incorporation of dCTP nucleotides at three different temperatures: 37°C, 47°C, and 55°C. Next is dATP incorporation using the same order of temperature conditions, followed by dTTP and lastly dGTP.

• See appendix for alternate versions of the figures.

Summary:

• This liquid phase synthesis experiment was conducted to compare the incorporation of dNTPs across a range of temperatures between wild type TdT and ThTdT.

• Wild type TdT exhibits highest efficiency at 37°C across all dNTPs and while still functional, its activity drops as temperature rises. In figure 1, each dNTP shows the same pattern, the bands above the primer band appear lower as temperature point increases.

• Wild type TdT is least efficient at incorporating dGTPs. In figure 1, dGTP lanes show lower bands above the primer band compared to other dNTP lanes.

• ThTdT has very low efficiency at 55°C, which contradicts the results from 2024.09.07 - LPS with ThTdT (Trial 5) which demonstrated that ThTdT could add dCTP and dTTP with comparable number of dNTP additions to lower temperature points. Between 37°C and 47°C, results are inconclusive as there is very little difference between the bands in those temperature conditions for most dNTPs.

• ThTdT appears to favour pyrimidine addition, as seen in figure 2 where dCTP and dTTP lanes show the highest bands above the primer band.

• Overall, the elongated bands for both WT TdT and ThTdT are fairly faint compared to the primer band. This could be an indication that the reaction was very weak or concentrations of reagents were incorrect. More testing will be required to identify the issue.

Appendix:

more versions of the gel image: Drive

Purpose:

• Prepare the mother culture

• Make more LB Broth

• Prepare the bioreactor and conventional control

Materials & Methods:

Refer to Bioreactor Efficacy Evaluation

Refer to Agar plate and broth preparation

Results:

• Prepared the mother culture and stored it in 4ºC fridge and Burak will come tomorrow at 3 PM to run this experiment.

• Bioreactor Mrk1 is ready.

• Refer to Bioreactor Efficacy Evaluation

• Mother culture has been activated and then chilled at 4C to prevent overgrowth.

Summary:

• First trial of the bioreactor will occur tomorrow.

• The mother culture has been prepared and activated.

• We will split it in two and then place it in the bioreactor and conventional liquid culturing flask and run it for 4 hours, 20 min. interval measurements.

Purpose:

• Extract mutant TdT for LPS, SPS, and microfluidics experiment

Materials & Methods:

Protocol:

Purpose:

Culture BL21 E.coli and induce with IPTG for ThTdT expression

Materials:

• 1M IPTG
• Transformed BL21 E.coli
• LB Broth

Procedure:

Protocol Page Link: Protein Expression

1. Pick a single colony from transformed BL21 plate and shake in 3ml of LB broth with 30ug/ml Kanamycin for 1hr
   1. Stock concentration 50mg/ml
      1. 0.6ul/ml Kanamycin → 1.8ul of 50mg/ml in 3ml LB broth
2. Add the 3ml of E.coli culture into 55ml of LB broth with Kanamycin and shake for 2.5hr
3. Add 500uM of IPTG to the solution, shake overnight
   1. Stock concentration 1M IPTG
      1. 27.5ul of 1M IPTG in 55ml of LB

References:

Protein expresion

1. Resuspend pellets - resuspend cell pellets (from overnights, for example) with binding buffer on ice - add to purple lidded tube with sonication beads - keep on ice 2. Lysis with sonicator - load tubes into white tube holder boxes - the 2 must balance - tighten boxes until they dont move - settings: 30seconds, 20Hz frequency - Run for 30s, then rest tubes on ice 2 minutes (sonication heats up samples and don’t want proteins to denature) - repeat this three times (should get cloudier each time, cell debris should rest on top of beads)

• 3. Centrifuge
  • take tubes on ice and tabletop centrifuge to cold room
  • centrifuge 30 minutes, max speed (14,300 rpm)
  • supernatent is the lysate - take this and use it in SDS Page etc.
  • can store purple lidded tubes with rest of cell debris and lysate (supernatent) in -20 freezer

References:

Beth from Hallam Lab

Magnetic beads typical reaction protoco

• Shook 400ml of LB, However only extracted Mutant TdT from 300ml LB
  • 400ml of LB was split into 4 flasks of 100ml
  • Flask 2 was added 3x the IPTG, instead of 500uM, used 1500uM - didn’t use this flask for mutant TdT extraction

Purification refer to 2024.09.03 - Protein Purification using Amicon® Ultra-0.5 Centrifugal Filter

Purpose:

• Reactivate overnight culture in LB+Kan and seed the bioreactor and conventional flask

• Take spectrophotometry measurements in triplicate every 20 minutes to generate a growth curve and compare

Materials & Methods:

Refer to Bioreactor Efficacy Evaluation

Results:

• 2 Eppendorf tubes with 1 mL each of Kanamycin stock solution (50 mg/mL) was made.

Common subculture OD at start: 0.033 A

• 8:45 PM the trial began

• Blanked with LB+Kan

Figure 1. OD Measurements in 20-minute intervals using spectrophotometer at 600 nm for bioreactor & conventional culturing flask method.

Figure 2. Growth curves of the bioreactor and control culture flask generated. Using spectrophotometer and collecting OD at 600 nm in 20-minute intervals for 2 hours.

Statistics:

Null hypothesis of no difference

Critical value = 0.05

P-value = 0.73847536

Significance level = 2.447

• P-value > significance level, therefore we reject the null hypothesis of no difference and support the alternative hypothesis that there is a difference in the amount of bacteria both methods when compared to each other with the bioreactor performing better and culturing more bacteria in the same amount of time and from the same volume and same mother culture.

  • Only difference is the air added directly into the culture in the bioreactor and the magnetic stir bar for bioreactor Mk. 1, as you can observe below:

Summary:

• Prior to data analysis, it seems that there is an increased OD for the bioreactor compared to the flask culturing method.

• This supports our hypothesis that the bioreactor would have a higher OD than the culturing flask.

• Statistically significant results.

• Increased OD change for bioreactor compared to the conventional method.

• Newer bioreactors will be built with additional features.

Purpose:

Testing the ThTdT enzyme with new primers.

Materials & Methods:

• ThTdT
• Primer P2
• 10x TdT reaction buffer
• dNTPs (ATCG)
• CoCl2
• molecular grade water

For materials needed for see TBE Urea Gel

Protocol: TdT reaction + TBE Urea gel (modified)

• The following table are compositions for the reaction tubes using WT TdT/ThTdT.

Incubation at 37C for 15min

Results:

• No addition occurred in either WT TDT or mutant TTDT.

Summary:

• The experiment needs to be repeated. It is hypothesized that adding 0.3ul for the volume might be too little. The concentration is also significantly lower than what we have done before, so it needs to be validated if this concentration works.

Purpose:

• Rerun liquid-phase synthesis and urea PAGE using Antonio’s fluorescent P2 primer at 37ºC, based on original experiment from 2024.09.11 - LPS with ThTdT Trial 8 using Different Primers.

• Looking for extension of all bases for both WT and ThTdT. Meant to see if there is improved extension with a different base other than G terminating the primer for TdT-mediated nucleotide addition.

Materials & Methods:

Materials:

• pipette tips, ~60

• PCR tubes

• Urea gel components

• liquid-phase synthesis components (see below)

• Loading dye mix

Methods:

• The experiment followed the protocol described in Bioreactor Efficacy Evaluation

• The following table are compositions for the reaction tubes using WT TdT/ThTdT, with the rightmost column being the master mix composition, made separately for WT TdT and ThTdT. The primer was the last component added to the master mix.

• Cy3-fluorescent P2 primer provided by Antonio were used in place of the P1 primer

Table 1: Reaction tube and master mix compositions

• Once made, each reaction tube was divided into 2x5uL tubes so that one may be incubated for 15 minutes and the other for 30 minutes at 37C.

Results:

• Following the use of the new P2 Primer during LPS, UREA gels were ran and imaged (figure 1).

• Incorrect fluorescence wavelengths were used when imaging leading to bands being not visible. Imaging was repeated the following day using the correct fluorescence wavelength (see Figure 1 below).

• Due to the contents of the gel degrading slightly overnight, the experiment was performed again. The notebook entry can be found at 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8).

Left gel: LPS with WT TdT, in order from left to right:

• 15 mins

• primer only

  • dATP
  • dTTP
  • dCTP
  • dGTP

• 30 mins

  • same order

Right gel: LPS with ThTdT, same order as WT TdT

• Lack of a ‘row’ of primers on the image, as seen in previous experiments using the P1 primer, indicate better utilization by both versions of TdT. However, the Cy3 fluorophore is seen to be generally less fluorescent than Cy5.

Summary:

• Liquid-phase synthesis using the P2 primer showed that it is better utilized by TdT than the P1 primer. Gel degradation, low fluorescence, and debris in the image make it hard to qualitatively interpret the addition rates for each dNTP. The experiment was repeated for a third time in 2024.09.11 - LPS with ThTdT Trial 8 using Different Primers to obtain a cleaner image that can be more easily interpreted.

Purpose:

• The goal of this experiment was to attempt immobilization of the DNA primer on a glass solid support and to extend DNA strands using an adapted protocol from liquid-phase synthesis (LPS), particularly the reaction composition used earlier in the day in 2024.09.11 - LPS with ThTdT Trial 8 using Different Primers. The previous lessons learned from LPS should inform the protocol to apply to the solid phase. Expected results should show a glass slide with the immobilized fluorescent primer and a gel with the results of immobilization.

Materials & Methods:

• 4x 10ul 100nM biotinylated Cy5-fluorescent DNA primer P1

• 4x 10ul 1M NaOH

• 5uM dTTP

• WT TdT

• Quenching solution (see TdT reaction + TBE Urea gel (modified))

• 4x 10ul 0.1% biotin-PEG-SVA and 100% PEG-SVA in 0.1M NaHCO3

• Reagents for TdT extension reaction

• 4x 10ul 2mg/ml Neutravidin

• Nitrogen gas

• Microcentrifuge tubes

• PCR tubes

Protocol page link: Glass Slide Preparation biotin

Urea PAGE protocol: TdT reaction + TBE Urea gel (modified)

1. On the opposite face of the glass slide to be functionalized, 4 dots were made with sharpie, denoting C,1,2,tdtRXN

2. 10ul 0.1% biotin-PEG-SVA and 100% PEG-SVA in 0.1M NaHCO3 was added to each dot for 3hs

3. The glass slide was rinsed off with water

4. 10ul 2mg/ml Neutravidin was added to each dot for 10min. The avidin solution was collected back to the eppendorf tube and the glass slide was washed with water

5. 10ul 100nM biotinylated DNA primer was added to each dot for 10min then washed with water

6. To the tdt dot was added the necessary reagents for tdt synthesis as carried out by 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8)

7. The glass slide was placed in the 37 degree room for 10min

   1. **not preheated prior to adding TdT, therefore true reaction time likely was lower

8. The glass slide was rinsed with water then imaged

9. To each of the dots was added a NaOH solution for 10min

   1. 2: 2ul 1M NaOH

   2. 1: 10ul 0.1M NaOH

   3. C,tdt: 10ul 1M NaOH

10. The NaOH solution was pipetted out and mixed with the appropriate dye and loading buffer to run in urea PAGE. It should be noted that dot C, tdt and 1 had 20ul loaded into the gel while dot 2 had 10ul loaded into the gel.

Results:

• Annotated spots were marked on the bottom of the glass slide for where reagents are to be added:

(need a regular pic of glass slide for reference)

• The glass slide was imaged for Cy5 fluorescence. The dark large circles on spots 1, 2, C, and tdt indicate successful immobilization of the fluorescent primers, where a 10uL drop of 100nM primer solution was each dispensed. No primer solution was dispensed to spot 3, which makes sense that it does not have a large circle.

• After cleaving the immobilized primers with varying concentration/volume of NaOH, the glass slide was reimaged. The large dark circles no longer seen where the smaller spots are indicate successful cleavage (this is particularly visible on C).

• After cleavage, the droplets were carefully pipetted up and dispensed into PCR tubes. Quenching solution was added to each sample in a 3:7 ratio as was done for LPS, and incubated at 95ºC for 5 minutes. The use of 10uL 1M NaOH used to cleave the TdT spot (which received TdT and dTTP to facilitate nucleotide addition) was noted since the NaOH concentration in the final sample was far higher than in LPS, which could affect the gel electrophoresis. 20% Urea PAGE was run at 250V for 30 minutes.

Fig. 4: Urea gel image of cleaved samples from the glass slide. From right to left:

1. primer solution with loading dye, no incubation

2. primer solution with loading dye, incubated at 95C for 5 mins

3. Spot 1, cleaved with 10 uL 0.1 M NaOH

4. Spot 2, cleaved with 3 uL 1 M NaOH

5. Spot C, cleaved with 10 uL 1 M NaOH

6. Spot TdT, cleaved with 10 uL 1M NaOH

• The cleavage step is expected to result in an oligo shorter than the original primer. The gel image confirms successful cleavage, as the band on the second lane from the left higher up than the four lanes on the right indicates the cleaved primers are indeed shorter. Lane 3 suggests a lower concentration of NaOH (0.1 M) is sufficient for cleavage.

• However, the lack of another band or smudge above the last lane on the right suggests either no or too little nucleotide addition.

Summary:

• Cy5-fluorescent primers were successfully immobilized onto a biotin-functionalized glass slide, confirmed with fluorescence imaging. Cleavage with concentrated NaOH was successful and confirmed with imaging the glass slide and urea PAGE. TdT-mediated nucleotide addition appeared unsuccessful.

• To try next:

  1. Urea PAGE with standard curve of primer concentrations to approximate amount of primers immobilized and cleaved on glass slide.

  2. Cleavage using lower concentrations of NaOH and loading dye solution (~ 4mM NaOH)

  3. Nucleotide addition using ThTdT and other dNTPs, over a range of reaction conditions (dNTP concentration, time)

Purpose:

• To test ThTdT vs WT TdT nucleotide addition using new primers.

Materials & Methods:

• The experiment followed the protocol described in TdT reaction + TBE Urea gel (modified)

• The following table are compositions for the reaction tubes using WT TdT/ThTdT.

**** Use Antonio’s primer please P2

Add dNTP and water separately

Make master mix, adding buffer, TdT and CoCl2

Mix well, then aliquot to each tube

Repeat the reaction for WT

A total of 10 reactions

• WT * 5

• Mutant *5

Incubation at 37°C for 15min

Quenching:

• Took 3ul from each sample added dye + NaOH solution, then incubate at 95°C for 5 min

• For the remaining 7ul in each sample:

  • Incubate at 70°C for 10 minutes or by adding 10 µl of 0.2 M EDTA (pH 8.0)

Results:

will annotated soon, but the order of lanes (ignoring the empty lane) is:

Lane 4: WT, primer only

Lane 5 - 8: WT; A, T, C, G

Lane 9: mutant, primer only

Lane 10 - 13: mutant; A, T, C, G

Raw file saved as “20240914 wt vs mutant new primer no plastic-[Cy3].webp” on imaging computer in LSI-5.435

Summary:

Purpose:

The purpose of this experiment is to attempt SPS using primer P7, a non-fluorophore primer. As it is hypothesized that the cy5 tag may sterically hinder the TdT enzyme from extending DNA effectively, seen in 2024.09.14 - Immobilization and SPS test 1 , primer P7 (lacks any fluorescent tag) will instead be used.

Additionally, for cleavage off of the solid phase, we will attempt to cleave with 10ul of loading buffer containing 0.1M NaOH. The high dilution of the previous SPS test may have resulted in poor gel contrast, this approach may alleviate this.

Downstream imaging will necessarily require SYBR Green as an imaging reagent due to the lack of any fluorophore on the primer.

Materials & Methods:

SPS for 10uL rxn volume:

• 1 uL TdT buffer

• 0.25 uL purified ThTdT stock

• 2 uL 5uM dTTP

• 1 uL CoCl2

• 5.75 uL water

For 50 uL quench solution with 0.1M NaOH for cleavage:

• 10 uL water

• 5 uL 1M NaOH

• 35 uL loading dye mix

Procedure

Protocol based on 2024.09.14 - Immobilization and SPS test 1

Rather than using NaOH solution to cleave immobilized oligos off the glass slide, a quench solution was prepared with 0.1 M NaOH so that the contents are less dilute for urea PAGE.

4 trials will be conducted and labelled as follows:

• control: no avidin

• 1: no TdT extension

• 2: no TdT extension replicate

• 3: ThTdT extension

Reactions were collected and frozen to be imaged later.

Results:

See 2024.09.18 - SPS test 2 UREA Gel Electrophoresis for imaging results

Purpose:

• To create UREA gel mixture for making UREA gels in future optimization experiments.

Materials:

• Urea (solid form, white colour)

  • Kept in a plastic bag on the bench

• 40% 29:1 acrylamide:bisacrylamide stock

  • Brown bottle, ept in ºC room

• 10X TBE stock

  • Kept on lab bench

• ddH2O (Av-Gay lab has a dispenser)

Methods:

20% 29:1 (Bis)acrylamide Urea, recipe for 200ml

• 84g Urea
• 100ml 40% 29:1 acrylamide:bisacrylamide
• 20ml 10X TBE
• 18ml ddH2O

Store in 4 ºC

Results:

• Measured out Urea into beaker
• Added ddH2O, 40% 29:1 acrylamide:bisacrylamide and 10X TBE (measured with graduated cylinder)
• Mixed with stir bar until urea appreaed fully dissolved in solution (~30 min)
• Poured solution into 250ml bottle
• Stored in 4ºC room
• There are blue and black precipitate in the bottle, doesn’t affect gel composition, use the supernatant when making gel

Summary:

• Prepared 20% 29:1 (Bis)acrylamide Urea

Purpose:

Run urea PAGE on reaction products of 2024.09.17 - Immobilization and SPS test 2 with three primer standards to approximate concentration or amount of primers immobilized and collected, as well as standards for the three primers (P1, P2, P7) identify a qualitive relationship between primer concentration and fluorescence strength.

Primer standards should approximate 1000fmol, 100fmol, and 10fmol of primer molecules in 10uL, as that is the volume normally loaded into gels.

Materials & Methods

• PCR tubes
• Urea PAGE components
• 502uL Quench solution (500uL loading dye mix + 2uL 1M NaOH)
• pipette tips
• water
• 1uM P1
• 10 uM P2
• 1uM P7
• SYBR safe
• reaction products from 2024.09.17 - Immobilization and SPS test 2

Primer standards were prepared with serial dilutions as follows:

1. P1

2. 3.33uL 1uM P1 + 6.67uL water → 10 uL 333 nM P1 2. 1uL 333 nM P1 + 9uL water → 10uL 33 nM P1 3. 1uL 33nM P1 + 9uL water → 10uL 3 nM P1 4. 1uL 3nM P1 + 9uL water → 10uL 0.3 nM P1, mix and remove 1uL 5. All tubes should now have 9uL, add 21uL quench solution to each for 30uL final volume. Final concentrations should be 100nM, 10nM, 1nM, and 0.1nM repsectively.

3. P2: only 10uM stock available

   1. 0.33uL 10uM P2 + 9.67 uL water → 10 uL 333 nM P2
   2. Repeat steps a.2 - a.5

4. P7:

   1. Steps a.1 - a.4 were repeated, without removing 1uL for 0.3 nM
   2. 1uL 0.3nM P7 + 9uL water → 10uL 0.03 nM P7, mix and remove 1uL
   3. ll tubes should now have 9uL, add 21uL quench solution to each for 30uL final volume. Final concentrations should be 100nM, 10nM, 1nM, 0.1nM, and 0.01 nM repsectively.

100nM ~ 1000fmol

10nM ~ 100fmol

1nM ~ 10fmol

100pM ~ 1fmol

10pM ~ 100amol

2 20% urea PAGE gels were prepared according to TdT reaction + TBE Urea gel (modified) and run at 250V for ~40min, imaged using Cy5, Cy3, and UV wavelengths.

The two gels were stained in SYBR safe solution for 10 minutes according to Sheet, which will make any oligos in the gel Cy3-fluorescent.

Results:

SYBR safe staining appears successful given the visible bands of 100nM and 10nM P7 primers on the rightmost side of the right gel under Cy3 fluoresence. However, it appears 1nM and below is too dilute to be visible.

This would explain the lack of visible bands in the left gel with dilute primer standards although not having any bands in any wells in the left gel also suggest either unsuccessful cleavage with the 0.1 NaOH quench solution, poor staining, or no immobilized P7 primer in the first place.

@Qingru @Tina do you have the UV prints

Summary:

SPS test 2 appears unsuccessful, with either failed immobilization or failed cleavage. More experiments will have to be done to successfully perform solid-phase synthesis. The primer standards that are Cy3-fluorescent will also have to be made with higher concentrations.

Purpose:

Rerunning a D-PAGE gel using remaining reaction products created in 2024.09.14 - Immobilization and SPS test 1 and serial dilutions of the primer standard.

Materials & Methods:

TdT reaction + TBE Urea gel (modified)

• P1 primer (100nM biotinylated Cy5-fluorescent DNA primer)
• SPS Test 1 reaction products (labelled “1”, “C”, “TdT” in -20C mini fridge)
• Urea PAGE reagents and materials
• Imaging room

Note: steps 2-4 will be done while waiting for the gel to solidify

1. Create 20% (Bis)acrylamide Urea PAGE gel

2. Make dilutions of P1 primer

   1. Make diluted primer solutions such that when loading in the gel, the final mols of primer are 10fmol, 1 fmol, 100attomol

3. Create primer standards:

   1. Add loading dye/quenching solution + NaOH such that each standard ends up with 0.1M NaOH

   2. Heat denaturation: 95C for 5 min

4. Thaw SPS Test 1 reactions

5. Run the gel, loading the lanes in the following order (whichever lane is loaded first is considered lane 1):

   Lane 1: primer standard, 10fmol

   Lane 2: primer standard, 1 fmol

   Lane 3: primer standard, 100 attomol

   Lane 4: Sample “1”

   Lane 5: Sample “C”

   Lane 6: Sample “TdT”

   Note: 10ul total volume will be loaded for each lane.

   250v for 30 mins.

6. Image gel using Cy5 setting

Primer 1 dilution

nano (n) = 10e-9

pico (p) = 10e-12

femto (f) = 10e-15

atto (a) = 10e-18

Starting stock of P1 was 10 nM. In PCR tubes, created 10ul of each cncentration listed below:

• 3.33nM: 3.33ul 10nM + 6.67 ul water
• 0.333 nM: 1ul 3.33nM + 9 ul water
• 33.3 pM: 1ul 0.333nM + 9 ul water

Formula for calculation: x = 3e-6 L * conc.

Making quenching solution

• First made a 4x master mix

[NaOH] in quenching solution: 4ul 1M NaOH/28ul total volume = 0.14285714285 M

[NaOH] in primer standard: 7 ul quenching/10ul total volume * 0.14285714285 M = 0.1 M

Creating primer standards

Added the following to PCR tubes:

Results:

Summary:

There seems to be no nucleotide addition in the TdT condition as the primer band in the 6th lane is similar in size to the primer band in the 5th lane (control).

Need to troubleshoot the gel sample preparation as the samples ran in a curved manner. It is suspected that the different in concentration of NaOH between the standard and reaction samples affects the migration of bands.

Purpose:

Following the results from the previous sps test, the issue of dilution affecting downstream imaging and validation is a problem. So in this set of experiments, we will attempt to rectify this problem by cleaving the primer of two replicate trials with the same NaOH cleavage solution, thereby hopefully effectively doubling the signal from downstream assays.

This approach will be tested using both P1 and P7

• dot 1: P1
• dot 2: P1
• dot 3: P7
• dot 4: P7

Materials & Methods:

• 2x 10ul 100nM biotinylated Cy5-fluorescent DNA primer P1

• 2x 10ul 100nM biotinylated non-fluorescent DNA primer P7

• 5uM dTTP

• Thermostable TdT

• Loading buffer in 0.1M NaO

  • 10 uL water
  • 5 uL 1M NaOH
  • 35 uL loading dye mix

• 4x 10ul 0.1% biotin-PEG-SVA and 10% PEG-SVA in 0.1M NaHCO3

• Reagents for TdT extension reaction

• 4x 10ul 2mg/ml Neutravidin

• Nitrogen gas

• Microcentrifuge tubes

• PCR tubes

Protocol page link: Glass Slide Preparation biotin

Urea PAGE protocol: TdT reaction + TBE Urea gel (modified)

1. On the opposite face of the glass slide to be functionalized, 4 dots were made with sharpie, denoting 1, 2, 3, 4. The 1/2 trials denoted the use of primer P1, the 3/4 trials denoted the use of primer P7.

2. 10ul 0.1% biotin-PEG-SVA and 10% PEG-SVA in 0.1M NaHCO3 was added to each dot for 3hs

3. The glass slide was rinsed off with water

4. 10ul 2mg/ml Neutravidin was added to each dot for 10min. The avidin solution was collected back to the eppendorf tube and the glass slide was washed with water

5. 10ul 100nM biotinylated DNA primer was added to each dot for 10min then washed with water and imaged

6. To the tdt dot was added the necessary reagents for tdt synthesis as carried out by 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8)

7. The glass slide was placed in the 37 degree room for 10min

   1. **not preheated prior to adding TdT, therefore true reaction time likely was lower

8. The glass slide was rinsed with water

9. To 1 and 3 was each added 10uL loading buffer in 0.1 M NaOH solution for 5min

10. The 10uL loading buffer from 1 was pipetted to 2. The 10uL loading buffer was pipetted from 3 to 4. This solution was allowed to sit for 5 minutes.

Results:

Fig. 1: The glass slide was imaged following immobilization

• Immobilization of the DNA primer appeared to have been unsuccessful. No signal is detected around the loci of primer addition. The glass may have been incorrectly functionalized due to poor exposure to the silane reagents. As such, this experiment will have to be repeated.

Purpose:

To compare efficacy of Mk. 2 Bioreactor to conventional flask shaking method of culturing E. coli:

• Reactivate overnight culture in LB+Kan and seed the bioreactor and conventional flask

• Take spectrophotometry measurements in triplicate every 30 minutes to generate a growth curve and compare

Materials:

• 250ml and 1L erleynmeyer flask

• LB Broth (at least 500ml)

Methods:

Bioreactor Efficacy Evaluation

1. Subculture 1 mL of old mother culture to create 100 mL of new mother culture (grow for 3-4 hours until 0.4-0.5 OD)

   1. In a 250 mL erlenymeyer flask, prepare fresh media by adding the appropriate amount of kanamycin antibiotic to 100 ml of LB Broth.

   2. Shake the old culture such that bacterial cells are evenly dispersed, then take 1 ml of that culture and add that to the fresh media.

   3. Securely attach the culture flask to a shaker in the 37ºC room, and measure OD after 3- 4 hours of incubation.

2. Set up two 200ml cultures (one for control and one for bioreactor):

   1. For each condition, use 20ml of the new mother culture to inoculate 180 ml LB media (with added antibiotic). Then start measuring OD for each condition for 6 hours

Calculation of Kanamycin AB Addition:

For mother culture:

C1V1 = C2V2

C1 (stock conc.)= 50mg/ml = 50000ug/ml

V1 (volume of stock needed)= x

C2 (final conc.)= 30ug/ml

V2 (total volume of culture) = 100ml

x = (30ug/ml * 100ml) / 50000ug/ml

x = 0.06ml = 60 µL of 50 mg/mL Kanamycin Stock

For individual broth:

C1V1 = C2V2

C1 (stock conc.)= 50mg/ml = 50000ug/ml

V1 (volume of stock needed)= x

C2 (final conc.)= 30ug/ml

V2 (total volume of culture) = 200ml

x = (30ug/ml * 200ml) / 50000ug/ml

x = 0.12ml = **120 µL **of 50 mg/mL Kanamycin Stock

Results:

• Mother culture looks opaque therefore growth is present and high OD too which is what we want.

• Each condition will have an equal amount of the mother culture added to it.

• The control flask has been seeded with the mother culture, LB+Kan, however as the bioreactor was not ready, I placed it in the 4ºC freezer in the meantime at 5:45 PM.

• No results as the bioreactor was not ready.

Summary:

• This experiment has been delayed to September 23rd.

Purpose:

Troubleshooting immobilization and gel visualization of SPS.

Materials & Methods:

Immobilization was carried out based on previous results in 2024.09.19 - SPS Test 1 Urea PAGE with modifications.

Preparing 100nM P1 master mix (40ul total):

C1V1 = C2V2

C1 = 1 uM

V1 = x

C2 = 100 nM

V2 = 40ul

x = 100*40/1000 = 4ul

Preparing 100nM P7 master mix (40 ul total):

C1V1 = C2V2

C1 = 25 uM

V1 = x

C2 = 1 uM

V2 = 10ul

x = 1*30/25 = 1.2 ul

+28.8 ul water

C1V1 = C2V2

C1 = 1 uM

V1 = x

C2 = 100 nM

V2 = 40ul

x = 100*40/1000 = 4ul

For 50 uL quench solution with 0.1M NaOH for cleavage:

• 10 uL water
• 5 uL 1M NaOH
• 35 uL loading dye mix

Results:

Immobilization of the DNA primer appeared to have been unsuccessful. No signal is detected around the loci of primer addition. The glass may have been incorrectly functionalized due to poor exposure to the silane reagents. As such, this experiment will have to be repeated.

Purpose:

• To subculture the mother culture for Mk.2 bioreactor

• Conducting Trial 1 growth curve generation of Mk.2 bioreactor

Materials & Methods:

Refer to Bioreactor Efficacy Evaluation

For subculturing:

1. Thaw out 50 mg/mL of Kanamycin sulphate solution.

2. Take 60 µL of 50 mg/mL of Kanamycin sulphate solution and add to 99.94 mL of LB Broth under flame.

3. Add 1 mL of mother culture containing *E. coli *B21 ThTdT-positive culture.

4. Incubate in 37ºC room while shaking overnight for 16 hours.

Results:

• The subculture was made and is in the incubator room.

Summary:

• The subculture turned into an overnight culture and will be placed in the 4ºC fridge tomorrow morning to be subcultured once more on Saturday morning.

Purpose:

• To prepare E. coli BL21 ThTdT-positive for bioreactor growth rate assessments

• Make LB Broth & Kanamycin Stock Solution

Materials & Methods:

1. Bioreactor Experimentation: Bioreactor Efficacy Evaluation

   1. subcultured 1 mL of overnight culture in 99 mL of LB-Kan liquid culture for 1 hour

   2. sterilized bioreactors Mk.2 and Mk.3 with 70% EtOH

   3. all bioreactors and control were propagated at the same time in the same environment

   4. measurements taken every 30 minutes for 6 hours

   5. 160 mL of culture in control, Mk.2 and Mk.3

      1. 144 mL LB broth

      2. 96 uL of Kanamycin Stock (50mg/mL)

      3. 16 mL of subculture E. coli BL21 ThTdT-positive

2. LB Broth Preparation: 2024.08.31 - LB Broth Preparation for Bioreactor

3. 2024.05.04 - Kanamycin Agar Plate and LB Broth Preparation

Results:

Subculture OD was 0.251

Spreadsheet for OD measurements:

OD measurement spreadsheet

Purpose:

Troubleshooting immobilization and gel visualization of SPS.

Materials & Methods:

two spots, 1 and 2

Added 10ul of 2mg/ml to each dot, let it sit for 10 mins

Rinsed off with diH2O

Added 10ul 100nM P1 to 1 and 10ul 100nM P7 to 2, let it sit for 10 mins

Rinsed off with diH2O

Preparing reaction mixture

Added 10ul of reaction mixture to each spot, then added water drops around the slide to prevent dehydration.

Incubate for 25mins.

Cleave off primer.

For 50 uL quench solution with 0.1M NaOH for cleavage:

• 10 uL water
• 5 uL 1M NaOH
• 35 uL loading dye mix

Results:

Immobilization of the DNA primer appeared to have been unsuccessful. No signal is detected around the loci of primer addition. The glass may have been incorrectly functionalized due to poor exposure to the silane reagents. As such, this experiment will have to be repeated.

Purpose:

• The goal of this experiment was to attempt immobilization of the DNA primer on a glass solid support and to extend DNA strands using an adapted protocol from liquid-phase synthesis (LPS). The previous lessons learned from SPS suggests concentrating the reaction mixture prior to loading into SDS-PAGE would allow for better visualization. Expected results should show a gel with the results of immobilization.

Materials & Methods:

• 3x 10ul 100nM biotinylated Cy5-fluorescent DNA primer P1

• 3x 10ul 100nM biotinylated non-fluorescent DNA primer P7

• 5uM dTTP

• Thermostable TdT

• Loading buffer in 0.1M NaO

  • 10 uL water
  • 5 uL 1M NaOH
  • 35 uL loading dye mix

• 6x 10ul 0.1% biotin-PEG-SVA and 10% PEG-SVA in 0.1M NaHCO3

• Reagents for TdT extension reaction

• 6x 10ul 2mg/ml Neutravidin

• Nitrogen gas

• Microcentrifuge tubes

• PCR tubes

Protocol page link: Glass Slide Preparation biotin

Urea PAGE protocol: TdT reaction + TBE Urea gel (modified)

1. On the opposite face of the glass slide to be functionalized, 6 dots were made with sharpie, denoting A1, A2, A3, B1, B2, B3. The A trials denoted the use of primer P1, the B trials denoted the use of primer P7.

2. 10ul 0.1% biotin-PEG-SVA and 10% PEG-SVA in 0.1M NaHCO3 was added to each dot for 3hs

3. The glass slide was rinsed off with water

4. 10ul 2mg/ml Neutravidin was added to each dot for 10min. The avidin solution was collected back to the eppendorf tube and the glass slide was washed with water

5. 10ul 100nM biotinylated DNA primer was added to each dot for 10min then washed with water

6. To the tdt dot was added the necessary reagents for tdt synthesis as carried out by 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8)

7. The glass slide was placed in the 37 degree room for 10min

   1. **not preheated prior to adding TdT, therefore true reaction time likely was lower

8. The glass slide was rinsed with water

9. To A1 and B1 was each added 10uL loading buffer in 0.1 M NaOH solution for 5min

10. The 10uL loading buffer from A1 was pipetted to A2. The 10uL loading buffer was pipetted from B1 to B2. This solution was allowed to sit for 5 minutes.

11. The 10uL loading buffer from A2 was pipetted to A3. The 10uL loading buffer was pipetted from B2 to B3. This solution was allowed to sit for 5 minutes.

12. The 0.1M NaOH loading buffer solution was pipetted out and run in urea PAGE.

Results:

Fig. 1: Urea gel image of cleaved samples from the glass slide.

• Bands corresponding to the primer can be seen in lane 4, indicating successful immobilization. No bands can be seen from the P7 lanes, indicating that the SYBR staining and visualisation was unsuccessful.

Summary:

• Cy5-fluorescent primers were successfully immobilized onto a biotin-functionalized glass slide and successfully cleaved from the support, confirmed by the presence of the primer from gel imaging. However the band is extremely faint, preventing detection of any extension product. Thus it is impossible to validate any successful TdT primer extension.

• To try next:

  1. Use a higher concentration of primer for immobilization to increase loading density. A higher concentration of biotin-PEG-SVA may also yield similar results.

  2. Use a higher concentration of DNTP’s during the extension reaction to increase primer extension.

Materials & Methods:

Making solutions:

Preparing reaction mixture

Added 10ul of reaction mixture to each spot, then added water drops around the slide to prevent dehydration.

Incubate for 25mins.

Cleave off primer.

For 50 uL quench solution with 0.1M NaOH for cleavage:

• 10 uL water
• 5 uL 1M NaOH
• 35 uL loading dye mix

Purpose

Test LPS on microfluidic chip.

Materials:

• SAR Microfluidic Chip
• Microfluidic Chip system
• CoCl2
• Thermostable Terminase Buffer
• P2 Primer
• ThTdT
• ddH2O
• dCTPs

Methods:

Set up microfluidic chip system.

Prepare reaction mixture:

Prepare a 180ul reaction mixture with the same concentration as a typical LPS reaction, using the optimized reagent ratios based on data from previous LPS experiments.

20uL of 100nM P2 Primer was prepared. 18ul was used for standard asssay (same concentrations as typical LPS reaction) with teh reamining 2 uL used to diluted the primer 10x.

• 179 uL will be used for the SAR microfluidic chip condition (the entire system can take 1ml max but the total volume that fills up the channels of the chip is 180 uL).

• 1ul of reaction will be removed prior to adding enzyme to act as a control.

• Concentrations of LPS reagents will be based on reaction conditions in 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8).

Results:

Summary

Need to troubleshoot fluorescence intensity. We suspect the oligonucleotides could be binding to the chip.

Purpose:

1. To verify whether primers were irreversibly bound to the microfluidic chip during the reaction interval
2. To compare the performance of liquid phase synthesis (LPS) in microfluidic chip against a PCR tube

Materials:

• Primer P2 as defined in Oligonucleotides, /56FAM/AGCCTGTTGTGAGCCTCCTAAC, 1.0 µM
• 10X Terminal transferase reaction buffer (NEB)
• dTTP, 10.0 µM
• $CoCl_2$, 2.5 mM
• ThTdT, 40X relative to in-house final standard assay concentration
• pET-28b, 33.6 ng/µL
• Water
• 20% denaturing PAG

Methods: part 1 - addressing non-specific binding

1. pET-28b (33.6 ng/µL, 180 µL) was introduced to a split end recombination chip for blocking for 1 hr in RT
2. The solution was discarded.
3. A mock reaction mixture (1X Terminal transferase reaction buffer, 100 nM primer P2, 180 µL) was introduced the chip for 15 min in RT.
4. The solution was recovered.
5. 5 µL of the mock reaction mixture before and after manupulations in the chip were applied to DNA PAGE for analysis.

Methods: part 2 - comparing LPS performance between microfluidic chip and thermocycler

1. The following reaction master mixture was assembled:

   Item	Stock		Volume (L)	Final Conc
   Terminal transferase reaction buffer	10	X	33.0E-6	1	X
   ThTdT	40	X	8.3E-6	1	X
   Fluorescent Primer	1.0E-6	M	33.0E-6	100.0E-9	M
   dTTP	10.0E-6	M	33.0E-6	1.0E-6	M
   CoCl2	2.5E-3	M	33.0E-6	250.0E-6	M
   Water			264.0E-6

   The contents of this tube is labeled AB

2. An aliquot from AB (30 µL) was diluted to 300 µL with water. In other words, this reaction was diluted 10 fold (0.1X).

3. An aliquot from AB (50 µL) was transferred to a PCR tube and reacted in a thermocycler programed to heat at 37ºC for 15 min. This served as a standard assay control completed in a thermocycler, known as reaction condition A.

4. The remaining aliquot (250 µL) was introduced to the microfluidic chip. This served as a standard assay condition completed in a microfluidic chip at 37ºC, known as reaction condition B.

5. An aliquot from CD (50 µL) was transferred to a PCR tube and reacted in a thermocycler programed to heat at 37ºC for 15 min. This served as a 0.1X standard assay control completed in a thermocycler, known as reaction condition C.

6. The remaining aliquot (250 µL) was introduced to the microfluidic chip. This served as a 0.1X standard assay condition completed in a microfluidic chip at 37ºC, known as reaction condition D.

7. 5 µL was removed from each reaction condition for analysis in 20% denaturing PAGE.

Results:

Figure 1. LPS using ThTdT with dTTP and primer P2. Reaction manifolds: - not used; T thermocycler; M: microfluidic chip.

Raw data for traceability purposes:

Summary:

1. Blocking the microfluidic chip with plasmids appears successful compared with results in 2024.09.29 - Microfluidic Chip LPS Experiments.
2. Regretfully, ThTdT activity was not observed across all reaction manifolds. The efficacy of ThTdT in microfluidic chips remains inconclusive in this entry, and warrants future studies.

DBTL

Run 1 Learn (from the previous cycle)

The recovery of the fluorescent oligonucleotide was poor. This was evidenced by the faint band signal intensity in the DNA gel, suggesting that the amount recovered was lower than the Typhoon biomolecular imager’s detection limit.

Of the faint bands observed, the evidence of primer elongation by ThTdT in LPS was inconclusive. Both research questions require further investigation in the next cycle.

Run 2 Design

To address the non-specific binding hypothesis, and experiment with a comparator (control) and intervention arm is devised. The intervention, using plasmids as blocking agents, is inspired by Enzyme-Linked ImmunoSorbent Assay (ELISA). In this assay, analytes were absorbed onto a solid support non-covalent. To ensure specificity, a relative low cost protein was sacrificially applied to block the surfaces that were not supposed to react. Common examples include fetal bovine serum (FBS), bovine serum albumin (BSA), and fat-free milk power. While it would be easy to use common nucleic acids carriers, such as tRNA and salmon sperm DNA, for the purpose of blocking, the presence of free 3’-ends will compete resources over nucleotides during the elongation reaction. Therefore, plasmids were chosen since they are DNA will no free 3’-termini.

Run 2 Build

A two part experiment was proposed to study the effect of non-specific binding and reaction manifolds. In part 1, the effect (with vs. without use) of the microfluidic chip was studied, with the following experimental plan:

Figure 2. Experimental plan to test the effect of non-specific binding of primer P2 /56FAM/AGCCTGTTGTGAGCCTCCTAAC by blocking the microfluidic chip with plasmid pET-28b.

In anticipation for this test, the chip was blocked by plasmid pET-28b prior to microfluidic LPS.

Run 2 Test

The following experimental plan tested the efficacy of LPS on the chip, such that a three-arm study is devised. The negative control was no manipulation (stored over ice, expecting no elongation activity); the positive control was 37ºC heating in a thermocycler; and the experimental intervention was 37ºC heating in a microfluidic chip.

Figure 3. Experimental plan to study the effect of reaction manifold (thermocycler, no manipulation, and microfluidic chip) on ThTdT 3’-elongation activity with Primer P2 /56FAM/AGCCTGTTGTGAGCCTCCTAAC.

Run 2 Learn

Figure 4. LPS using ThTdT with dTTP and primer P2. Reaction manifolds: - not used; T thermocycler; M: microfluidic chip.

We learned that blocking the microfluidic chip with plasmids appears successful compared with the unblocked results in 2024.09.29 - Microfluidic Chip LPS Experiments.

Regretfully, ThTdT activity was not observed across all reaction manifolds. The efficacy of ThTdT in microfluidic chips remains inconclusive in this entry, and warrants future studies.

Purpose:

Compared to previous experiments, this trial will attempt to increase loading density and reaction power of ThermoTdT. To increase the loading density, a higher concentration of Biotin-PEG-SVA will be used (1% instead of 0.1%). To increase the reaction power of TdT, a 10x concentration of DNTP will be used, and the reaction will be allowed to progress for 30 minutes.

Materials & Methods:

• 3x 10ul 300nM biotinylated P1
• 3x 10ul 300nM biotinylated P7
• 4x 10ul 1M NaOH
• 10uM dTTP
• ThermoTdT
• Quenching solution (see TdT reaction + TBE Urea gel (modified))
• 4x 10ul 1% biotin-PEG-SVA and 5% PEG-SVA in 0.1M NaHCO3
• Reagents for TdT extension reaction
• 4x 10ul 2mg/ml Neutravidin
• Nitrogen gas
• Microcentrifuge tubes
• PCR tubes

Protocol page link: Glass Slide Preparation [biotin]

Urea PAGE protocol: TdT reaction + TBE Urea gel (modified)

1. On the opposite face of the glass slide to be functionalized, 6 dots were made with sharpie, denoting A1, A2, A3, B1, B2, B3
2. 10ul 1% biotin-PEG-SVA and 5% PEG-SVA in 0.1M NaHCO3 was added to each dot for 3hs
3. The glass slide was rinsed off with water
4. 10ul 2mg/ml Neutravidin was added to each dot for 10min. The avidin solution was collected back to the eppendorf tube and the glass slide was washed with water
5. 10ul 300nM biotinylated DNA primer was added to each dot for 10min then washed with water
6. The glass slide was imaged for cy5
7. To the tdt dot was added the necessary reagents for tdt synthesis as carried out by 2024.09.14 - LPS with ThTdT Trial 10 using Different Primer (3rd attempt at Trial 8)
8. The glass slide was placed in the 37 degree Celsius room for 30min
9. The glass slide was rinsed with water then imaged for cy5
10. To A1 and B1 was each added 10uL loading buffer in 0.1 M NaOH solution for 5min
11. The 10uL loading buffer from A1 was pipetted to A2. The 10uL loading buffer was pipetted from B1 to B2. This solution was allowed to sit for 5 minutes.
12. The 10uL loading buffer from A2 was pipetted to A3. The 10uL loading buffer was pipetted from B2 to B3. This solution was allowed to sit for 5 minutes.
13. The 0.1M NaOH loading buffer solution was pipetted out and run in urea PAGE.
14. The glass slide was imaged for Cy5
15. The gel was stained with SYBR Safe and imaged for Cy5 and the stain.

Results:

• Annotated spots were marked on the bottom of the glass slide for where reagents are to be added:

• The glass slide was imaged for Cy5 fluorescence three times.

• The dark spots indicate successful immobilization of the fluorescent primers, where a 10uL drop of 300nM primer solution was each dispensed.

Figure 1. Cy5 fluorescence image of P1 primers immobilized onto a biotinylated glass slide prior to enzymatic DNA synthesis. The dark spots indicate strong successful binding of the Cy5 labelled primers.

• After TdT-mediated extension, the glass slide was rinsed with water to quench the reaction and remove reagents. The glass slide was then imaged under Cy5 parameters. We see retention of primer on the glass slide following TdT extension

Figure 2. Cy5 fluorescence image of P1 primers immobilized onto a biotinylated glass slide following enzymatic DNA synthesis. The dark spots indicate successful retention of Cy5 labelled primers.

• After cleavage, the glass slide was imaged one final time. The results show a stark loss of signal, indicating successful cleavage of the product.

Figure 3. Cy5 fluorescence image of the biotinylated glass slide following cleavage of the previously immobilized P1 primers. The lack of dark spots indicate no P1 primers remain on the glass slide and thus successful cleavage.

• Droplets were carefully pipetted up and loaded into SDS PAGE (20% Urea PAGE was run at 250V for 30 minutes). In addition to the SPS samples, a varying concentrations of primer standard were loaded to benchmark the results. The result is shown below. The SPS lane shows numerous bands and an intense signal.

Figure 4. 3’-extension by ThTdT on using primer P1 /5Biosg/ATTCGrATCA/iCy5/CTAGCATACTATCATTCGGGG immobilized on microscope glass slide at 37ºC for 30 min. Lanes 1, 2, and 3 contains 10 fmol, 1 fmol, and 100 amol P1. Lane 4 contains reaction crude from SPS, reacted in [CoCl₂] = 250 µM, [dTTP] = 10 µM. 20% D-PAGE, 30 min, 400V. Imaged using Cy5 setting.

Summary:

• Cy5-fluorescent primers were successfully immobilized onto a biotin-functionalized glass slide, confirmed with fluorescence imaging. Cleavage was successful and confirmed with imaging the glass slide and urea PAGE. TdT-mediated nucleotide was successful, resulting in many bands of higher molecular weight.
• To try next:
  1. Optimize the results of SPS to achieve near quantitative conversion of the starting material to a singly extended product with few by-products.
  2. Apply the established methodology to a microfluidic platform

Purpose:

• to prepare agar plates for agar art to be conducted by students
• expose students to plating techniques and bacteria colony growing

Materials:

• 70% ethanol
• dh5a - bacteria
• Kanamycin resistant agar plates
• 37C incubator
• Lab Coat
• Mask
• Gloves
• Swab

Procedure:

1. Take agar plates out from 4ºC fridge.
2. Put it in 37ºC incubator for 1 hour to warm it up.
3. Take plates to Michael Smith.
4. Participants wash hands for 20s with soap, wear lab coat, gloves, masks.
5. Mark the agar plate with your name, iGEM, and the date.
6. Spray your hands (wearing glove) with 70% ethanol.
7. Gently dip the toothpick/swab into the tube containing dh5a E.coli.
8. Open the lid, draw freely on agar plate.
9. Close the lid.
10. Spray your hands (wearing glove) with 70% ethanol.
11. Wash you hands with soap for 20s.
12. Coordinators will take away the plates.

Note. MAKE SURE THE KIDS DON’T TOUCH ANY PERSONAL BELONGINGS OR THEMSELVES DURING THE EXERCISE