Experiments

Purpose:

Determine Size of PCR product

Materials:

• Agarose
• 1x TAE buffer
• DNA loading buffer
• DNA ladder
• SYBR safe

Procedure:

1. For gel purification - if we need the product after running a gel
   • Small gel: 60ml TAE buffer : 1% agarose (0.6g)
   • Large gel: 110ml
2. For gel analysis - if the gel is only for confirmation that we have the correct size
   • Small gel: 30-40ml TAE buffer : 1% agarose
3. Microwave for 1:30-2:00 min
   • Check every 30s after the initial 1 min, to make sure no over spill due to bubbling
   • Stop when the liquid has no or minimal visible translucent pellets
4. Add SYBR safe into agarose when the liquid is comfortable to the touch
   • 10,000 dilution (60ml + 6ul SYBR safe)
   • Swirl to mix well
5. Pour into the cast, insert size comb, wait until the agarose harden (~15-20min)
   • Use comb to remove bubble
   • For gel purification, use large comb
6. Add 1x loading buffer into sample
   • 6x loading buffer (45ul sample, 9ul loading buffer)
7. Put the gel with the cast into the electrophoresis box, then top up with 1X TAE buffer
8. Load 5-10ul ruler, add DNA sample
   • Gel purification: maximum amount of DNA that the well can hold (~50ul)
   • Gel analysis: 4-6ul DNA
9. Run the gel at 110-160V until desired distance
10. Check the band of the sample using gel imager, make sure it’s the correct size.
    1. Or take a picture at UV light station if imager is not working
11. Use gel imaging device to visualize the bands and print a copy of the gel.

Purpose:

Prepare antibiotic resistant LB agar plate and LB broth for Clone selection

Materials:

• LB powder
• Agar powder
• Antibiotic
• dd water
• molecular grade water

Procedure:

For 250mL of Kanamycin Agar Plate (~9-12 plates)

1. Find a clean 500mL bottle
2. Weigh 5g (20g/L) of LB broth powder and 3.75g (15g/L) of agar powder
3. Pour 250mL of ddH2O
4. Stir until it looks dissolved (there will be some powder in the middle)
5. Autoclave in liquid cycle 20
   1. Make sure to put autoclave tape, screw the cap so it doesn’t fall off but don’t go all the way, and put aluminum foil on top
6. Cool it down to around 55 degrees (just feel it with hand and you can comfortably touch it)
7. Add antibiotics according to their recommended concentration (usually 1:1000, ex. 250mL of liquid with 250uL of antibiotic)
   1. if antibiotic is in powder form, dissolve in molecular grade water to desire concentration
8. Pour the plates in the BSC so it is just enough to cover the bottom
9. You should make about ~9-12 plates
10. Let it dry on the bench and then once it cools, put it in 4c
11. Can put 1 plate in 37degree overnight to check if the antibiotic works (if it works, nothing should grow)

For 500mL LB Broth

1. Find a clean 1000mL bottle
2. Weigh 12.5g of LB broth powder
3. Pour 500mL of ddH2O
4. Autoclave in lquid cycle 20
   1. Make sure to put autoclave tape, screw the cap so it doesn’t fall off but don’t go all the way, and put aluminum foil on top
5. Once cooled, store at room temperature on the shelf

Purpose:

• to determine if the bioreactor is able to culture more bacteria in a shorter amount of time compared to the control condition which involves the conventional culturing method of using a flask

Materials:

• Glycerol stock BL21 #1 MT E. coli
• LB broth
• Incubation room, shaker and magnetic stir bar
• Hemocytometer
• Brightfield microscope
• Erlenmeyer Flask
• Bioreactor Mrk1

Procedure:

Trial 1

Based on:

TN52236-differences-bacterial-optical-density-uv-vis.pdf

1. Make the mother culture by taking the glycerol stock and make a mother LB liquid culture with Kanamycin (719.5 mL LB broth, 432 uL Kan, 80 mL E. coli)
2. Incubate it at 37ºC overnight
3. Aliquot 16 mL of overnight culture and inoculate 800 mL of pre-warmed 37ºC LB+Kan (with 50 mg/ml Kanamycin sulphate stock solution) media
4. Take OD
5. Split in half into the bioreactor and the conventional flask
6. Keep both in the 37ºC incubation room for 4 hours while mixing.
7. Blank spectrophotometer with LB Broth
8. Take a 1-3 mL aliquot from the control flask and bioreactor every 30 minutes.
9. Pipet into cuvette.
10. Take the OD measurement at 600 nm using the spectrophotometer.

To make the mother culture:

1. 432 µL Kan + 719.5 mL of LB Broth
2. 360 mL of LB+Kan in each
3. 40 mL of mother culture in each

Trial 2

1. Subculture 1ml of old mother culture to create 100ml of new mother culture (grow for 3-4 hours until 0.4-0.5 OD)
2. Set up two 300ml cultures (one for control and one for bioreactor):
   1. For each condition, use 30ml of the new mother culture to inoculate 270ml LB media (with added antibiotic). Then start measuring OD for each condition for 10-12 hours

• 100 mL
• 1 mL
• 60 ul Kan + 99.94 mL of LB for subculture
• 1 hour wait until 0.4-0.5 OD

And for the control, there’s two ways we could go about it:

• use a large 1000ml flask, which would be the typical method for a 100-200ml culture (technically 300ml is pushing it, but a 2000ml flask may be too big for the shakers in Hallam lab), this would be realistic to what someone would do in this situation so if the bioreactor is better in comparison then it would make sense for someone to use the bioreactor over conventional method
• OR use a flask that is closest in total volume capacity to the bioreactor (probably 500ml) so that it’s comparable to using the bioreactor in terms of vessel volume (in theory the bioreactor would be better because it’s continuously aerate the culture whereas the control flask will likely run out of oxygen even with shaking)

Safety, containment and contamination control:

1. H2SO4 + H2O2. This combination is known as the Piranha solution. This is no joke. Any organic matter vaporizes to CO2 upon contact. You don’t want this on your skin, or your clothes. If this spilled onto your lab coat, remove immediately. Contamination control: secondary containment and have sodium bicarbonate on stand by. These are very dangerous reagents to work with. MUST ADD H2O2 TO H2SO4 SLOWLY.

https://www.youtube.com/watch?v=cLpSapjKcxM

The rest are relatively a cake walk, but use caution and common chemical lab sense.

1. APTES and MPTS. These silanols are category 1 skin sensitizers. Not great on your skin. Degrades on contact with water. Contamination control, rinse affected area with water, copiously.
2. Acetone, ethanol, acetic acid. Flammable solvent. Contamination control, rinse affected area with water, copiously.

Materials:

1. Sulfuric acid (98%)
2. Hydrogen peroxide (30%)
3. Sodium bicarbonate (500 g powder, ACS grade)
4. NaOH pellets
5. pH papers
6. APTES (3-Aminopropyl)triethoxysilane (Ambeed, ACS grade) (Antonio will bring)
7. Acetone (ACS grade)
8. Poly(ethylene glycol) succinimidyl valerate, MW 5000 (mPEG-SVA)
9. Biotin-PEG-SVA at a ratio of 99:1 (w/w) (Laysan Bio
10. Sodium bicarbonate (0.1 M, can be made from line 3, no need to be sterile)

Procedure:

Coverslips were soaked in piranha solution (25% H2O2 and 75% concentrated H2SO4) and left overnight, followed by multiple rinses in water (Thermo Fisher Scientific, molecular-biology grade) and acetone (Thermo Fisher Scientific, HPLC grade). Dry and clean coverslips were then treated with Vectabond/acetone (1% v/v) (Marketed by Vector Labs, is actually APTES (3-Aminopropyl)triethoxysilane, purchased from Ambeed) solution for 5 min and then rinsed with water and left in a dried state until used. In order to prevent non-specific adsorption of biomolecules onto the glass surface, coverslips were functionalized prior to use with a mixture of poly(ethylene glycol) succinimidyl valerate, MW 5000 (mPEG-SVA) and biotin-PEG-SVA at a ratio of 99:1 (w/w) (Laysan Bio) in 0.1 M sodium bicarbonate (Thermo Fisher Scientific) for 3 h. Excess PEG was rinsed with water, and the coverslips were dried under a N2 stream. The surface was incubated with 7 μL of a 2 mg/ml neutravidin solution (Thermo Fisher Scientific). Excess neutravidin was then washed off with 100 μL of 1X PBS buffer.

The biotinylated primer is added to the glass slide in a 10uL volume on the functionalized loactions at a concentration of 300uM and incubated for 10 minutes. The slide is washed with water then the reaction mixture was added, consisting of:

1. 1x TdT buffer
2. 0.25uL TdT enzyme
3. 250uM CoCl2
4. 10uM DNTP

The slide was incubated at 37 degrees Celsius for 30 minutes at which point the reaction mixture was washed off with water. Loading dye in 0.1M NaOH was added to the reaction centre for 10 minutes to cleave the primer from the solid phase. The droplet was collected and analysed through gel electrophoresis.

References:

Improved immunoassay sensitivity and specificity using single-molecule colocalization

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:

https://qb3.berkeley.edu/education/lab-fundamentals-bootcamp/manual/proteins/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:

https://qb3.berkeley.edu/education/lab-fundamentals-bootcamp/manual/proteins/protein-expression/

Purpose:

To create buffer with neutral pH that is suitable for the storage of ThTDT and prepapre 10mM 2,2’-bipyridyl to stabilize ThTDT

Materials:

For storage buffer

• KH₂PO₄ solid
• K₂HPO₄ solid
• NaCl solid
• 14.3M β-ME
• Glycerol
• Triton® X-100
• Water

For 10mM 2,2’-bipyridyl,

• Solid 2,2’-bipyridyl
• Water

Procedure:

Storage buffer recipe:

• 50 mM KPO₄
• 100 mM NaCl
• 1.43 mM β-ME
• 50% Glycerol
• 0.1% Triton® X-100

Make 10mL 0.5M KPO₄ stock buffer:

1. Prepare 8mL of DI water.
2. Measure 571.24mg of K₂HPO₄ solid and 234.15mg of KH₂PO₄ solid, add into the water and mix well.
3. Add water till the volume of 10mL.

Make 1M NaCl stock solution:

1. Add 584.4mg NaCl in 10mL of water, mix well

2. Mix stock KPO₄, NaCl and β-ME to achieve a final concentration of 50 mM KPO₄, 100 mM NaCl. 1.43 mM β-ME

3. Use a 0.22uM filter, sterile filter the mixed solution

4. Add the same volume of Glycerol to achieve 50%

5. Add 0.1% Triton® X-100

6. Measure the pH of the buffer using a pH meter according to the pH Meter Use protocol

7. If the pH is not 7.3, adjust the pH to 7.3

Materials:

• p20 pipette tip

Procedure:

1. Using a p20 pipette tip, gently touch ONE SINGLE colony and transfer this to a tube filled with 4-10 mL of LB broth with 30 µg/mL Kanamycin.
   1. 4 mL for miniprep
   2. 10 mL for IPTG induction
2. Transfer as many colonies as needed to guarantee one of them has the correct plasmid integrated.

See BL21 IPTG induction for inducing protein expression once colony has been expanded.

Purpose:

To confirm whether purified protein is correct size. Coomassie blue dye stains proteins in protein gels, making visible bands that can be compared to a protein ladder in the same gel to identify size of sample proteins.

Materials:

• Container to hold gel (be sure the gel can move freely in water or stain to allow diffusion of solution)
  • The base of the container should be at least the size of the lid of a pipette box for standard protein gel
• Orbital Shaker
• See below for materials needed for coomassie stain and de-stain
  • Note: Both solutions have a strong smell due to acetic acid and methanol and coomassie dye can stain clothing, advised to wear appropriate PPE.

Coomassie Stain, recipe of 1L

0.1% Commassie R250, 10% acetic acid, 40% methanol

Reagents needed:

• 1g Comassie R250
• 100 ml glacial acetic acid
• 400 ml methanol (note: there are recipes with ethanol as an alternative)
• 500 ml ddH2O

Directions:

1. In a 1L beaker/duran bottle, add a stir bar and place on a magnetic stirring plate to mix the solution.
2. Add 100 ml of glacial acetic acid to 500 ml of ddH2O.
3. Add 400 ml of methanol and mix.
4. Add 1g of Coomassie R250 dye and mix. The solution should turn a deep blue.
5. Filter to remove particulates (a coffee filter works).
6. Store at room temperature in a sealable container/duran bottle.
   1. Optional: Cover bottle in alumnimum foil.

De-stain for Coomassie, recipe for 1L

20% methanol, 10% acetic acid

Reagents needed:

• 200 ml methanol (note: there are recipes with ethanol as an alternative)
• 100 ml glacial acetic acid
• 700 ml ddH2O

Directions:

1. In a 1L beaker/duran bottle, add a stir bar and place on a magnetic stirring plate to mix the solution.
2. Add 100 ml of glacial acetic acid to 700 ml of ddH2O.
3. Add 200 ml of methanol and mix.
4. Store at room temperature in a sealable container/duran bottle.

Procedure:

Coomassie R250 Staining Protocol (without microwave)

(This protocol may take longer but is safer. The sources below provide a quicker protocol using a microwave.)

1. After electrophoresis of protein gel, carefully disassemble the electrophoresis casing and release the gel from the glass sandwich using a plastic wedge.
   • At this point, you may cut off the top and bottom sides of the gel to help fit into the container, but be careful not to cut off any portion of the gel containing protein. It may help to leave some of the top of the wells remaining for easier identification of individual lanes later on.
2. Transfer it to a container and rinse the gel with tap water (fill up container with water, shake the gel in a side-to-side motion then carefully decant water from the container without losing the gel).
3. Pour Coomassie stain over the gel until the gel is fully covered.
   • 100 - 200 ml stain for a container close to the size of the gel
4. Shake (slow speed) the container for at least 2 hours on orbital shaker.
   • 1 hour may be sufficient but this depends on the amount of protein loaded
5. Decant the coomassie stain (can pour it back into stock solution or dispose in appropriate liquid waste).
6. Pour de-stain solution over the gel until the gel is fully covered
7. (Optional) Place scrunched-up Kimwipes in the container around the edges of the membrane to help soak up excess dye.
8. Shake (fast speed) for at least 2-3 hours, overnight if needed until the desired background is achieved.
   • Looking at the gel in the container, you should see that some or all of the lanes contain blue bands (unless experiment was not successful).
9. Take a picture of the gel.
   • Lay the gel on a clean surface such as a glass lid.
   • Put a piece of masking tape below the gel and label it with the date, sample in each lane and any additional information about the gel.
   • If the gel is left out for too long it will deform and not be flat anymore. Quickly put the gel back into de-stain solution. The gel can be kept on the bench for as long as desired, however the bands will fade with time.
10. To dispose of the gel:
    • Decant the de-stain solution (can pour it back into stock solution or dispose in appropriate liquid waste)
    • Throw the gel into the appropriate waste bin (generally protein gels are not biohazardous so they can go into normal waste)
    • Clean the container

References:

Modified protocol from:

• https://www.thermofisher.com/ca/en/home/references/protocols/proteins-expression-isolation-and-analysis/staining-protein-gels/coomassie-staining.html
• https://www.fredhutch.org/content/dam/stripe/hahn/methods/biochem/Coomassie Stain of Protein Gel.pdf

Materials:

• Competent DH5α E. coli (ThermoFisher)
• Ice bucket with ice
• Thermocycler or water bath set to 42°C
• LB broth or SOC media
• 37°C shaking incubator (37°C room)
• 10 cm diameter LB agar plates with appropriate antibiotic
• Timer
• Plasmid DNA

Procedure:

Modified protcol from: https://www.thermofisher.com/document-connect/document-connect.html?url=https://assets.thermofisher.com/TFS-Assets/LSG/manuals/MAN0018520_DH5alpha_competent_cells_UG.pdf

Note: Skip step 1 and 2 if correct volume aliquots have already been made.

1. Thaw competent cells on wet ice. Place the required number of 1.5-mL polypropylene microcentrifuge tubes on wet ice.
2. Gently mix the cells, then make 50 µL aliquots of competent cells in the chilled 1.5-mL microcentrifuge tubes.
3. Add 1−5 µL of sample DNA directly into a tube of competent cells. Mix well by gently flicking the tube several times.
   • For a control condition, replace DNA with dH2O.
   • The remaining ligation/transformation mixture(s) can be stored at -20°C.
4. Incubate the cells on ice for 30 minutes.
5. Heat shock: Incubate tube(s) for exactly 30 seconds in the 42°C water bath. Do not mix or shake.
6. Cool down: Remove tube(s) from the 42°C bath and place them on ice (keep on ice for at least 2 mins).
7. Add 250 µL of room-temperature S.O.C. Medium or LB Broth to each tube.
   • S.O.C is a rich medium; sterile technique must be practiced to avoid contamination.
8. Place the tube on its side in a shaking incubator. Use tape to secure the tube in place.
9. Shake the tube at 225 rpm for 1 hour at 37°C.
10. If necessary, dilute the cells 1:10 with S.O.C. Medium/LB Broth.
11. Spread at least two different volumes (20−200 µL) of cells from each transformation reaction on separate LB plates containing the appropriate selective antibiotic.
    • Label the plates with “iGEM”, the date and plating volume so that the amount providing the best colony density can be identified.
12. Invert the plates and incubate overnight at 37°C.
13. Check plates the next day.

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. Lyse with sonicator
  • load tubes into white tube holder boxes - the 2 must balance
  • tighten boxes until they don’t 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)
  • supernatant is the lysate - take this and use it in SDS PAGE etc.
  • can store purple lidded tubes with rest of cell debris and lysate (supernatant) in -20 freezer

References:

Beth from Hallam Lab

Adapted from GeneJet Gel Extraction Manual from Thermofisher

Materials:

GeneJET Gel Extraction Kit from Thermo Scientific (NOT the green boxes)

Procedure:

1. Pre-weigh 1.5mL tubes and record the weight on the tubes.
2. Excise gel slice with desired DNA fragment using clean razor blade, place into pre-weighed 1.5mL tube and measure the weight of the gel.
3. Add 1:1 volume of Binding Buffer to gel slice
   1. ex. 100ul Binding Buffer to 100mg of agarose gel
4. Incubate gel mixture at 50-60C for 10min until gel slice is completely dissolved. Mix tube by inversion every few minutes.
   1. The solution should be yellow in colour.
5. Set up and label columns accordingly (one column per sample). Vortex the gel mixture briefly before loading on GeneJET Purification column.
6. Transfer up to 800uL of solubilized gel solution to GeneJET purification column. Centrifuge for 1min at 12000rpm. Discard flow-through and place column back into same collection tube.
   1. If the total volume of a given sample exceeds 800ul, repeat this process again with left over gel mixture solution
7. Add 700ul of Wash Buffer to the GeneJET purification column, centrifuge for 30sec at 12000rpm. Discard flow-through and place column back in same collection tube
8. Repeat step 6 but centrifuge for 1 min at 12000rpm.
9. Centrifuge empty GeneJET column for 1min at 12000rpm to remove residual wash buffer
10. Transfer column to a clean 1.5mL tube, add 50ul of Elution Buffer (or distilled water) to center of purification column member (make sure the liquid covers the membrane completely). Incubate at room temperature for 1min. Centrifuge for 1min at 15000rpm.
    1. The amount of elution buffer used will depend on amount of DNA recovered. Use a lower volume (as low as 20ul) to elute if needed.
    2. If DNA fragment is >10 kb, prewarm Elution Buffer to 65°C before applying to column.
11. Store the purified DNA at -20°C.

Materials:

• NEB Gibson Assembly Master Mix
• Insert (thermostable TdT)
• Vector (pET-28b(+))
• Water

Procedure:

• 2 conditions
  • With insert
  • Without insert (negative control, only vector)
• 10ul reaction
  • 40-50ng vector
  • Insert 3: vector 1
  • 5ul master mix
  • Water

1. Set up the experiment according to the above table, keep everything on ICE
2. Incubate in Thermocycler for 30min at 50C
3. Store product at -20C

Materials:

• Refer to Thermo Scientific GeneJET Plasmid Miniprep Kit

Procedure:

• Refer to Thermo Scientific GeneJET Plasmid Miniprep Kit

Purpose:

Use pH meter to adjust the pH of media to the desired pH

Materials:

• pH meter

• pH4, 7 and 10 calibration buffer (if you want to calibrate)

• dH2O

• NaOH

• HCl

• paper towel

Procedure:

Everytime when switching to the testing probe to a different solution, rinse with dH2O thoroughly, and dip on the paper towel to remove excess water

• Turn on the pH meter

• Test pH in pH 7 buffer

  • take out the testing probe

  • rinse with dH2O

  • Dip the probe gently on a piece of paper towel to remove excess water

  • Put the probe in pH 7 buffer (yellow)

  • press Read

  • Wait for the auto sign to not flash — this means the reading is stabilized

  • If the reading is off, go forward with calibration

• Calibration:

  • Start with pH 4 buffer, same as above

  • After reading is stabilized, press calibrate

  • Switch to pH 10 buffer, repeat

  • After finishing both, test again with pH 7 buffer

• If the reading is correct, go directly measure the pH of the buffer you are preparing for

  • If the pH is lower than expected, add NaOH

  • If the pH if higher than expected, add HCl

  • Add each dropwise, and keep track of the pH

  • When adjusting the pH, you may need to stir the solution using the stirring bar to consistently mix the solution

• When the pH is adjusted, take out probe, rinse thoroughly with dH2O, dip on paper towel, put it back into its buffer to prevent drying out the probe.

• Turn off the meter

Purpose:

• Add or remove DNA sequences from template

Materials:

• DNA
• Forward + Reverse Primers
• Q5 High-Fidelity 2X Master Mix

Procedure:

1. Keep everything on ice and mix the reagents together using the concentration and volume indicated below
   1. Can make either 25ul or 50ul PCR reaction

1. Step up the thermocycler, use the table below as an example ONLY
   1. adjust temperature and time according to each reaction
      1. dependent on DNA length
   2. Put your sample in once the thermocycler is heated up
   3. The 4C hold at the end is so that you can pick up your sample anytime after the cycles has ended, samples can be left overnight in 4C hold if needed

Purpose:

• The 5’ modified disulfide primer will be immobilized on an activated glass slide via disulfide exchange

Materials:

• 5’-disulfide oligonucleotides
• NaHCO3/NaH2CO3 buffer (pH 9.0) or 0.1 M citric acid buffer (pH 4.5)
• ETNTw (10 mM Tris–HCl, pH 7.5, 150 mM NaCl, and 0.05% Tween 20).

Procedure:

The 5’-disulfide oligonucleotides were diluted to a concentration of 20 micromolar in 500 mM NaHCO3/NaH2CO3 buffer (pH 9.0) or 0.1 M citric acid buffer (pH 4.5) and arrayed onto the mercaptosilanecoated glass slide surface. Arrayed primers were incubated in a humid chamber for 5 min to overnight, followed by three washes in TNTw (10 mM Tris–HCl, pH 7.5, 150 mM NaCl, and 0.05% Tween 20).

Rogers, Y.-H.; Jiang-Baucom, P.; Huang, Z.-J.; Bogdanov, V.; Anderson, S.; Boyce-Jacino, M. T. Immobilization of Oligonucleotides onto a Glass Support via Disulfide Bonds: A Method for Preparation of DNA Microarrays. Analytical Biochemistry 1999, 266 (1), 23–30. DOI:10.1006/abio.1998.2857.

Purpose:

To quantify the concentration of DNA within a sample.

Materials:

• 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

Procedure:

1. Prepare working solution in a falcon tube. For every standard and sample you need to multiply the amount of needed working solution. So that is at least x3. (Standard 1, Standard 2, and your sample)
   1. Working solution — 200:1, buffer:dye. Add 200 uL of Buffer (Component B) and 1 uL of Reagent (Component A) per sample and standard to measure.
2. For each standard (two in total), in a Qubit tube pipette 190 uL of the working solution and 10 uL of one standard. Do the same for the other standard.
3. For each sample, in a qubit tube pipette 199 uL of the working solution and uL of your sample.
4. Allow all tubes to incubate at room temperature around 2 minutes.
5. Turn on the qubit. On the Home screen of the Qubit® 3.0 Fluorometer, press DNA, then select dsDNA High Sensitivity as the assay type. The “Read standards” screen is displayed. Press Read Standards to proceed.
6. Insert the tube containing Standard #1 into the sample chamber, close the lid, then press Read standard. When the reading is complete (~3 seconds), remove Standard #1.
7. Insert the tube containing Standard #2 into the sample chamber, close the lid, then press Read standard. When the reading is complete, remove Standard #2.
8. On the assay screen, select the sample volume and units: Press the + or – buttons on the wheel to select the sample volume added to the assay tube (from 1–20 μL).
9. Insert a sample tube into the sample chamber, close the lid, then press Read tube. When the reading is complete (~3 seconds), remove the sample tube.
   1. Note that the initial concentration will be very high, press the button above Sample to fix. The button might be called “dilution concentration”.
10. Repeat until all samples have been read.

References:

Protocol based on the following link and help from Lucas from the Hallam lab. https://tools.thermofisher.com/content/sfs/manuals/Qubit_dsDNA_HS_Assay_UG.pdf

Purpose:

To end TdT addition reaction prior to loading into the gel if not directly proceed with Urea Page

Procedure:

• 0.2M EDTA (pH 8.0)

1. Heat the sample at 70 °C for 10 min, or add 10ul of 0.2M EDTA (pH 8.0)

2. After quenching, the sample can be stored in -20, ready for future Urea Page visualization

Purpose:

To visualize ssDNA oligo produced by TdT

Materials:

• Urea (Hallam)

• 40% 29:1 acrylamide:bisacrylamide (ordered)

• 1x TBE (Hallam)

• Tris base

• Boric acid

• Ammonium Persulfate (Hallam)

• EDTA disodium salt (Hallam)

• Formamide (Hallam in the flammable cabinet)

• Bromophenol Blue (borrowed)

• TEMED (Hallam)

• 1M NaOH

• Water

• DNA sample

Procedure:

Preparation of solutions

**500mM EDTA (pH 8), 100 mL **

1. Weigh out 18.61 g EDTA disodium salt, dihydrate and add to a 100 mL Duran bottle.

2. Measure out 80 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 NaOH to correct pH

   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)

500mM EDTA (pH 7.4), 100mL

Same as above, adjust pH to 7.4 instead

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.

Ammonium Persulfate, APS, 10% w/v, recipe of 10 mL

• Dissolve 1 g APS in 10 mL H 2 O

Store in 4 ºC, 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.

Pouring a Polyacrylamide Gel

1. Clean your glass plate with soap water, EtOH and finally glass surface cleaner.

2. Wipe them dry with KimWipe (Caution: do not scratch nor drop.)

3. Select a comb with the same thickness as the spacer. Clean the comb as outlines in Steps 1-2.

4. Depending on polyacrylamide gel percentage, prepare an unpolymerized gel mixture in a plastic Falcon Tube. total to 5 mL:

5. Add 5 μL TEMED (N′,N′,N′,N′-Tetramethylethane-1,2-diamine) to the mixture under the water level to prevent rapid evaporation. (Caution: TEMED can cause irritation. It has a pungent fishy odor.)

6. Do not forget to return the 20% 29:1 (Bis)acrylaimide Urea and 7 M Urea to the 4ºC refrigerator.

7. Clip the glass plates and the spacers together, face the |¯______/¯| side on the top (not the |¯¯¯¯¯¯¯¯¯| side)

8. Add 50 μL 10% Ammonium Sulfate (Caution: forms radicals. Store away from light.).

9. Pour the mixture into the glass sandwich. Avoid any air bubbles. Consult an experienced colleague before you start if this is your first attempt.

10. Allow the gel to polymerize overnight in room temperature. Do not disturb.

PAGE Preparation

1. Connect the red (+) wire to lower reservoir (female) and to red input (male) to power supply.

2. Connect the black (-) wire to higher reservoir (female) and to black input (male) to power supply.

3. Install the gel. (The |¯______/¯| side, in contrary to the |¯¯¯¯¯¯¯¯¯| side, should face the inner reservoir.)

4. Fill the upper reservoir, and then lower reservoir, with 1x TBE.

5. Use a 19-gauge syringe to retrieve some 1x TBE from the pool and flush out excess urea in the PAG well. Notice the urea leaving gel.

6. Set Voltage . (Caution: High voltage.)

   1. 400 voltes connecting 2 machines

7. Allow the system to run for 10 mins. Colorless gas bubbles, due to electrophoresis of water, will form on both exposed Platinum wires.

Sample Preparation

1. Prepare the following solution “Dye solution” (only mix this prior to usage. Over-incubation of NaOH with formamide causes undesired gel electrophoresis results):

2. Add 5 μL of the dye solution per µL sample to be electrophoresed (e.g. terminal transferase product). (Caution: For radioactive samples, work behind shielding.)

3. Resuspend by slight vortexing. Heat at 95ºC for 5 min (Allow denaturing by NaOH, allow chelation by EDTA).

PAG Loading

1. Set up the cassette.

2. At 45º relative to the water level at upper reservoir, add the sample at a slow constant rate in order to allow the sample to fall into the well by gravity (formamide is denser than water).

3. Allow some time between pushing the first pipette stop and the second pipette stop.

4. Push at most one small gas bubble in order to avoid turbulence.

PAG Electrophoresis

1. Set Voltage to 400V. (Caution: High voltage.)

2. In case of overheating, reduce the voltage to 250V.

3. Stop the gel when the bromophenol blue migrated to an appropriate length.

Methods:

Pouring a 20% Polyacrylamide Gel

1. Take the 1.5mm glass with its cover glass

2. Take 2 1.5mm 15-well combs

3. Assemble the gel cassette using the glasses

   1. https://www.youtube.com/watch?v=EDi_n_0NiF4 (here is a video you can watch on how to assemble gel cassettes, notice in the video that it only pours the solution half way not full, DON’T DO THIS, pour it all the way in for Urea PAGE and insert comb right away)

4. Add water in between the slides to test if there is leakage, if no leakage is observed overtime, proceed to make the gel

   1. Make sure to remove the water as much as possible (can use paper towel to do this)

5. Add the following solutions in order in a 50mL falcon tube (this recipe is for 1 gel)

   1. 10mL 20% 29:1 (Bis)acrylamide Urea (In cold room iGEM box)

   2. 10ul TEMED (On biomod’s bench)

   3. 100ul 10% APS (In cold room iGEM box (wrapped with Aluminum foil) - once you add both TEMED and APS in to the mixture, make sure to act FAST and pour the solution into the glass cassette and insert the comb

      • Typically we need to make 2 gels for one set of experiment. In this case, you can double the volumes up

      • Even if making only one gel, use a 50 mL falcon tube, as a 15mL tube is too skinny for the gel to solidify

6. Once mixed, vortex slightly, and quickly transfer the solution into the cassette, pour all the way to the top. Insert the comb quickly before the gel solidifies, when inserting the comb keep your eyes away from the glass, the liquid tends to splash out from the cassette towards your face!!!!!

7. Wait for around 10-15 minutes for the gel to solidify

   1. If it does not solidify within 30min, then something is wrong, go figure out the issue and redo from the top.

   2. Keep the leftover solution in the falcon tube to check solidification

8. If you are going to use it the same day, leave it in the rack. If you want to use it overnight, wrap in wet paper towel put it in a plastic bag and store in 4C cold room

Pre-run (PAGE preparation) (you can choose to do this after the DNA addition reaction is done during quenching, or after the samples are ready)

1. Insert the gels into the reservoir. Fill the upper reservoir, and then the lower reservoir, with 1x TBE. This means pour into the middle and let it overflow to the sides.

   1. We have a bottle of used 1X TBE you can use

   2. If there is gel buildup on the OUTSIDE of the cassettes, try to wipe or rinse them off. They will detach during pre-run and cause the machine to stop.

   3. If only running one gel, use a buffer dam to take the place of another glass cassette. It is plastic and is the same dimensions.

   4. Be sure to pull up on the cassettes while holding it in the electrode casing to create a seal. This is easier when the glass and your gloves are dry.

2. Set Voltage to 400V.

3. Plug in the electrodes corresponding to colors (black in black, red in red)

4. Allow the system to run for 10 mins. Colorless gas bubbles, due to electrophoresis of water, will form on both exposed Platinum wires.

   1. This means you should see bubbles coming up from the bottom of the reservoir

   2. If you don’t see a continuous flow of bubbles up, it means that your circuit is not complete. Stop the electricity and check your setup.

Sample preparation

All the reagents except the protein are stored in -20 freezer in a separate box named iGEM TdT with pink tape

• Before the reaction, grab a bucket of ice, thaw the reagents you need in ice

Proteins are stored in -80, top shelf, left corner, within the box storing BL21 bacteria stock. You can find tubes in the right side of the box. Those are our proteins.

• Proteins are already aliquoted in 10ul each in PCR tube

• Grab one tube, and return the rest

  • You may need to use scissors to cut out 1

• Thaw the proteins are ice

• Keep protein on ice ALL TIME

PLEASE KEEP EACH REAGNT CLEAN AND NON-CONTAMINATED AT ALL TIME. IF ANY GET CONTAMINATED, THE REACTION WON’T HAPPEN

• If you don’t feel comfortable using the stock, aliquot into smaller tubes

• Always change tips

1. Prepare the correct dilutions of each reagent if necessary

2. Preheat the thermocycler at certain temperature

   1. new protocol

   2. select the first column, set temperature, turn the time to infinite

   3. run (change the reaction volume)

   4. Leave it there

3. Take out new clean pcr tubes, correctly label for different reactions

   1. I usually mark 0,1,2, etc

4. Aliquot a small volume of molecular water from the bottle

5. Add the correct amount of water to each tube

   1. You can reuse the tip

6. Add CoCl2 if required

7. Add dNTPs to each tube

   1. DO NOT REUSE TIPS

8. Prepare the master mix by adding TdT buffer, TdT and primer, add correct amount of master mix to each tube

   1. Add drops at the side

   2. Once finishing adding all the tubes, tap the strip to mix

   3. **Use new tip each time please **

9. Quick spin to spin down all the residues

10. If you are doing different temperatures, you may need to divide your samples into multiple tubes

    1. In this case, label your tubes well

11. Incubate the tubes in a thermocycler

    1. Make sure you set the right temperature

    2. Time your reaction

While incubating:

1. Prepare the dye solution

   1. Dye (25mM EDTA/ Formamide/ Bromophenol Blue)              250 μL

      1. On bench, blue solution in 15ml tube

   2. 1M NaOH (NaOH/ Water)                                                                1 μL

      1. Cold room, middle shelf on your right

      2. Andrea’s box

      3. small, colorless tube

You cannot reuse this for multiple days, always make new

Once finished preparing the solution, return the NaOH immediately

1. Mix the solution

Quenching (after incubation is done):

1. Take new PCR tubes, label them well

2. Add 7ul of dye solution to new pcr tubes

3. Pipette up and down of the sample using a multichannel pipette, add 3ul of sample to the 7ul dye solution in the new pcr tubes. Slightly pipette up and down to mix

4. Quick spin to spin down all the residues

5. Incubate the tubes at 95c in a thermocycler for 5 min

Run gel

1. Before adding the sample, flush the wells with 1XTBE in the reservoir

   1. Use a p1000 pipette, suck up 1xTBE from a side reservoir, pipette the TBE into each well to remove excess urea. Rinse every single well

2. Load samples into the well

   1. Can use tape to mark individual well with their separate reaction

   2. DO NOT PICK UP AND ROTATE THE TANK

3. Run at 250V for around 30-40 min, stop once the bands migrate close to the bottom (ideally around 2/3 the way)

Disassembly

1. Pick up the casing and pour the TBE in the middle reservoir into the tank. Release the cassettes from the electrode.

2. Using a plastic wedge, metal spatula, or metal scoopula, gently wedge open the glass cassette.

   1. To minimize chance of breaking the glass, try to insert the wedge in as straight as you can before applying a lever force (do not try to just insert a corner on the side, it puts more stress on the glass).

   2. Remember the orientation of each gel.

3. Transfer the gel onto a transparent plastic film. Blot dry excess liquid using a piece of KimWipe where appropriate.

4. Stick the tape with the well labels onto the correct side of the gel.

5. Recycle 1xTBE - pour it back into our bottle.

Imaging

Machine usage rights courtesy to Dr. Eric Jan

1. On the 5 Floor of LSI, locate Room 5.435. Locate the GE Amersham Typhoon Scanner.

2. Open the door. Retrieve the Fluor Stage AmTyphoon screen (usually the middle one).

3. Gently slide the stage through the door. Put in the gel at the bottom left corner Insert the stage until it stops at the point indicated by the arrows. Make note of the grid regions then close the door.

   1. To get a better image (like one for putting on the wiki):

      1. Use ethanol or isopropanol solution to wipe clean the stage

      2. Pour a small amount of water onto the stage (a 2cmx2cm blob is sufficient)

      3. Carefully place your gel WITHOUT THE PLASTIC onto the water blob. This will allow you to move it freely.

4. Open Desktop on the associated computer. Open Typhoon Amersham software. Select Fluorescence mode. Customize acquisition parameters

   1. **Select “Cy5” **If you have other fluorophores, browse this list of possibilities.

   2. **Select “200 μm” **(lower number means higher resolution but it will also be slower)

   3. Double check file size < 40 MB, scan time < 20 min

   4. Select “Fluor” Stage/Area

   5. **Select Imaging Area **Drag corresponding area, you may have more than one region.

   6. **Define file path for data **C:\Typhoon images\igem\2024(create folder for what you’re imaging)

   7. **Define dataset file name **The File name is mandatory, while the Notes field is optional.

5. Select “Scan” to acquire

6. After completion, analyze using ImageJ.

NOTE: Check if your image matches the migration pattern of bromophenol blue by comparing with your original gel. If not, the acquisition is invalid. Double check your imaging parameters and your stage.

Please read this entire document through before attempting any of the wet lab work. There is information about safety, disposal and handling of various chemicals. At the end of the document are some schematics that may be of help to your understanding of the general workflow.

Required materials

The required chemicals and materials are listed below. Please ensure that you have access to them prior to initiating work.

Literature procedure

Commercially available glass slides were immersed in the 25% ammonium hydroxide solution overnight. They were then rinsed with running DI water for 10 minutes followed by a brief rinse in anhydrous EtOH. The slides were immersed in a mixture of 1% MPTS, 95% EtOH and 16mM acetic acid for 30 minutes. The slides were rinsed with 95% EtOH/16 mM acetic acid (pH 4.5) once and then cured either under dry nitrogen for overnight at room temperature or in a vacuum oven for 2 h at 150°C.

Rogers, Y.-H.; Jiang-Baucom, P.; Huang, Z.-J.; Bogdanov, V.; Anderson, S.; Boyce-Jacino, M. T. Immobilization of Oligonucleotides onto a Glass Support via Disulfide Bonds: A Method for Preparation of DNA Microarrays. Analytical Biochemistry 1999, 266 (1), 23–30. DOI:10.1006/abio.1998.2857.

Literature procedure – What you need to know

Notes

Before doing anything, notice that the glass in the petri dish is immersed in ammonium hydroxide. Notice that the glass slide has a side that is facing up towards the lid and a side that is facing down. We need to treat the side facing up with extreme delicacy. Do not touch this face!! You can handle the glass slide from the edges or the bottom, but we need to keep the top face relatively sterile. The following work will be conducted in the fumehood. Ammonium hydroxide will burn your nose, so be sure to open the sealed petri dish in the fumehood only.

Procedure

You will start by carefully peeling away the plastic parafilm wrapping on the petri dish, being careful not to jostle the materials inside. You will then remove the microscope slide inside and take it from the fumehood to the sink, leaving the petri dish filled with ammonium hydroxide in the fumehood. This petri dish should be diluted with water then quickly dumped down the sink with flowing water. Rinse the petri dish a couple times with tap water then it can go in the waste.

You will rinse the microscope slide with DI water for 10 min. When done, use a micropipette (the largest one) and rinse the slide once with EtOH (rinse each face of the slide once). Collect the EtOH filtrate in a waste beaker and dispose of the liquid down the sink with running water. Then grab a new petri dish from a bag that is already opened and place the slide in the petri dish, with the correct face up. Immerse the slide in the 1% MPTS, 95% EtOH and 16mM acetic acid for 30 minutes, ensuring that the top face is completely submerged. Place the lid on the petri dish to make sure nothing evaporates too much.

After 30 min, remove the glass slide. The remaining 1% MPTS, 95% EtOH and 16mM acetic acid solution in the petri dish can be disposed in the non-halogenated waste jerry can in the fumehood (should be located on the right or left inside the fumehood). Rinse the petri dish with water and dispose of it in the waste (same place as before). Rinse the glass slide with the 95% EtOH/16 mM acetic acid using a large micropipette. Just like the EtOH rinse from before, collecting the liquid into a waste beaker (this filtrate can be disposed in the sink with running water). Place the glass slide in a beaker with the correct face up. Ensure that it leans against the wall. Then cover the lid of the beaker with parafilm to create a seal over the beaker. Then select a balloon filled with Ar (g) and insert the needle into the glove, ensuring that the air from the balloon is released into the beaker as shown below. You can now place this beaker containing the slide on the igem bench.

Materials:

• Competent Top10 E. coli (ThermoFisher)
• Ice bucket with ice
• Thermocycler or water bath set to 42°C
• LB broth or SOC media
• 37°C shaking incubator (37C room)
• 10 cm diameter LB agar plates with appropriate antibiotic
• Timer
• Plasmid DNA

Procedure:

Modified protocol from: https://www.thermofisher.com/ca/en/home/references/protocols/cloning/competent-cells-protocol/routine-cloning-using-top10-competent-cells.html

1. Centrifuge the vial(s) containing the ligation reaction(s) briefly and place on ice - skip this step if transforming Gibson assembly product or miniprep plasmid DNA
2. Thaw, on ice, one 50 µL vial of One Shot cells for each ligation/transformation.
3. Pipet 2 µL of each ligation reaction directly into the vial of competent cells and mix gently with pipette tip (or finger tap the tube after adding DNA). Do not mix by pipetting up and down.
   • For a control condition, replace DNA with dH2O.
   • For transformation reactions, 1-5 uL of sample DNA is sufficient
   • The remaining ligation/transformation mixture(s) can be stored at -20°C.
4. Incubate the vial(s) on ice for 30 minutes.
5. Heat shock: Incubate vial(s) for exactly 30 seconds in the 42°C water bath. Do not mix or shake.
6. Cool down: Remove vial(s) from the 42°C bath and place them on ice (keep on ice for at least 2 mins).
7. Add 250 µl of pre-warmed S.O.C medium or LB Broth to each vial.
   • S.O.C is a rich medium; sterile technique must be practiced to avoid contamination.
8. 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.
9. Spread 20 µl to 200 µl from each transformation vial on separate LB agar plates containing the appropriate selective antibiotic.
   • Label the plates with “iGEM”, the date and plating volume so that the amount providing the best colony density can be identified.
   • The remaining transformation mix may be stored at +4°C and plated out the next day, if desired.
10. Invert the plate(s) and incubate at 37°C overnight.
11. Check plates the next day.

Materials:

• Competent E. coli
• Plasmid DNA
• LB Broth

Procedure:

Day 1:

1. Mix 1.5 µL of plasmid DNA (~5 ng) into 50 µL of competent cells (reaction has to be on ice all the time)
   • thaw competent cells on ice
2. Don’t vortex when mixing DNA with competent cell
   • pipette up and down
3. Incubate mixture on ice for 20-30 min. Pipette into PCR tube
4. Heat shock for 30-60 secs at 42ºC
5. Put tubes back on ice for 2 min.
6. Add 950 µL LB or SOC media
7. Shake for 60 min. at 250rpm, warm agar plate (with antibiotic)
8. Plate the bacteria:
   • option 1 — make one plate by adding 300 µL of culture and 30 µL of culture onto the second plate
   • option 2 — complete a three-factor (10-1, 10–2, 10-3) serial dilution using 100 µ , 1/3 cells + 2/3 LB or PBS), plate 5 µL

Day 2:

1. Check and see whether there is any colony growth on Kanamycin-containing agar plate.
2. If there is growth, this means that the plasmid was successfully incorporated into the bacteria and is expressing the Kanamycin-resistance gene.

Purpose:

To test our primers, see if wt TdT can successfully perform nucleotide addition on our primers, and if these additions can be visualized.

Materials:

• WT TdT from NEB

• Primer

  • Biotin linked, fluorochrome tagged

  • (Thiol modified)

• dNTPs

• TdT buffer

• CoCl2

Procedure:

1. For 50ul reaction, mix:

   1. 5.0 μl (10X) TdT Buffer

   2. 5.0 μl (2.5 mM) CoCl2 solution

   3. 5.0 pmols DNA primers

   4. 0.5 μl 10 mM dNTP

   5. 0.5 μl Terminal Transferase (20 units/μl)

   6. deionized water to a final volume of 50 μl.

2. For 10ul reaction, each will have:

   1. 1.0ul (10x) TdT Buffer

   2. 1.0 ul (2.5 mM) CoCl2 solution

   3. 1.0 pmols DNA primers

   4. 0.1 ul 10mM dNTP == 1 ul 1mM dNTP

      1. dilute dNTPs before preparing the mix

   5. 0.1 ul TdT (20 units/ul)

   6. deionized water for a final volume of 10 ul

To test different factors:

1. Prepare a master mix containing:

   1. 16ul (10x) TdT Buffer

   2. 16 pmols DNA primer

   3. 1.6ul TdT

   4. Deionized water (volume depend on dNTP and DNA primer concentration)

2. Divide the master into half:

   1. Half add 8ul (2.5 mM) CoCl2 solution

   2. Leave the other half without CoCl2

3. Further divide each half into 4, add 1nmol of individual dNTPs

4. 37 C incubation, for 5 min, take up half, add loading dye to stop the reaction

5. Leave the other half for another 25 min (total of 30min), then add loading dye to stop the reaction

6. Load the sample to Urea gel for visualization