Experiments

Materials

1. PCR Tubes
2. Desired Plasmid/DNA
3. Desired Type IIS Restriction Enzyme
4. HI-T4 DNA Ligase
5. T4 Ligase Buffer
6. Nuclease Free H₂O

1. In a PCR tube, add the following reagants in the order listed:

   Reagent	Amount
   Nuclease Free Water	Up to 20 µL
   T4 DNA Ligase Buffer	2 µL
   DNA/Plasmid	1 µL per DNA/Plasmid
   HI-T4 DNA Ligase	1 µL
   Type IIS Restriction Enzyme	1 µL

   Note: To make the workflow more efficient, calculate the amount of water you will need to add beforehand
2. Label PCR tubes with a number and in the lab notebook, create a legend for the contents of each PCR tube
3. Complete the reaction within a thermocycler at the following conditions:
   • 42°C for 2 min → 16°C for 5 min (cycle 35X)
   • 80°C for 20 min
4. Store at -20° C in freezer

For the creation of entry vectors using the MoClo Kit the digestion and heat inactivation steps are omitted, ending the reaction on a ligation as recommended in the supplementary material¹.

1. Lee, M.E., DeLoache, W.C., Cervantes, B., and Dueber, J.E. (2015). A Highly Characterized Yeast Toolkit for Modular, Multipart Assembly. ACS Synth. Biol. 4, 975–986. https://doi.org/10.1021/sb500366v.

Materials

1. 1.5 mL Microfuge Tube
2. P10 Pipette & Tips
3. Desired DNA/Plasmid
4. Desired Restriction Enzyme
5. Restriction Enzyme Buffer
6. HPLC H₂O

1. Label 1.5 mL microfuge tubes accordingly and record in lab notebook
2. Add reagents/materials in the amount and order as listed:

   Reagent/Material	Amount
   HPLC H2O	4 µL
   Restriction Enzyme Buffer	1 µL
   DNA/Plasmid	4 µL
   Restriction Enzyme	1 µL

3. Spin down at short/pulse cycle to ensure all contents are at the bottom of the tube
4. Incubate at 37°C at one hour on a heat block
5. Heat inactivate at 80°C for 10 min
6. Store at -20°C

Materials

1. Gel Electrophoresis Cassette & Lid
2. Power Supply
3. Casting Tray & Comb
4. Red Safe
5. Agarose Powder
6. 1X TAE Buffer
7. DNA Sample
8. DNA Ladder
9. Loading Dye

1. Using a graduated cylinder, measure 30 mL of 1X TAE buffer and pour into a flask
2. Using a weighing boat, measure 0.24 g of agarose and add to the flask
3. Dissolve agarose
   • Swirl the solution gently to mix
   • Microwave in 30-second intervals, swirling between each heating, until the agarose is fully dissolved and no visible crystals remain
   • Watch closely to prevent overflow, as the solution can boil over quickly
4. Cool Solution
   • Allow the flask to cool until it is warm to the touch, but not too hot
   • If the gel solidifies prematurely, you can reheat it only if RedSafe has not been added
5. Once cooled slightly, add 0.5 µL of RedSafe and swirl gently to mix
6. Prepare casting tray
   • Use green tape to seal both open ends of the casting tray. Ensure a tight seal by pressing the edges of the cassette against the lab bench
   • Insert the comb into the cassette
7. Slowly pour the agarose solution into the cassette, avoiding bubbles
8. Allow the gel to solidify for about 30 minutes. Once fully set, remove the tape and proceed with gel electrophoresis

1. Place the gel tray into the electrophoresis chamber and fill the chamber with 1X TAE buffer until it reaches the "max" line, or the gel is fully submerged
2. Ensure the side with the comb is facing the black (negative) electrode
3. Gently pull the comb straight up to avoid damaging the wells
4. Check that the wells are intact and filled with TAE buffer
5. Gel is now ready for sample loading

1. Cut a small piece of parafilm. Dispense 2 µL of loading dye for each sample (except the ladder) onto the parafilm
2. Add 8 µL of DNA sample to each droplet of loading dye. Pipette up and down gently to mix, avoiding bubbles
   • Change tips between each sample
   • If the DNA sample being used has a high concentration, use less than 8uL, or the band will appear too bright when imaged
3. Pipette 5 µL of DNA ladder into the first well of the gel
4. Before loading, map out your sample order to keep track of what is in each well
5. Take up 10 µL of your PCR + dye mixture and carefully load it into the wells (be careful not to puncture the gel while loading)
• Change tips between each sample
6. Place the lid on the electrophoresis apparatus (black to black, red to red). Turn on the power supply and set it to 120V
   • If you see tiny bubbles forming at both ends of the chamber, the run is working
7. Wait 30 minutes, then check if the dye has migrated ⅔ of the way down the gel
   • The run typically takes ~45 minutes but check at 30 minutes to be sure
8. Once done, turn off the power supply and unplug the lid
9. Your gel is now ready to be imaged

When microwaving the agarose and TAE, the content and the flask may be hot, ensure proper protection is used such as heat-resistant gloves.

Materials

1. PCR Tubes
2. Nuclease Free H₂O
3. 5X Phusion HF Buffer
4. 10nM dNTPs
5. Forward and Reverse Primers
6. Template DNA
7. Phusion DNA Polymerase

1. Label the PCR tube accordingly and record the materials within the tube in the lab notebook
2. Add the following reagents/components into the PCR tube in the order stated:

   Component	50 µL Reaction	Final Concentrations
   H2O	Fill to Amount 50 µL
   5X Phusion HF Buffer	10 µL	1X
   10 mM dNTPs	1 µL	200 µM
   Forward and Reverse Primer	2.5 µL	0.5 µM
   Template DNA*	Variable	5 ng
   Phusion DNA Polymerase	0.5 µL	0.02 U/µL

3. In the thermocycler complete the following cycling conditions:

   Cycle Steps	Parameters	Amount of Cycles
   Initial Denaturation	98 °C for 30 sec	1
   Denature	98 °C for 10 sec	35
   Anneal + Extension	72 °C for 30 sec	35
   Final Extension	72 °C for 5 min	1
   Hold	4 °C	∞

4. For long-term storage keep PCR tubes at -20°C

1. Thermo Fisher Scientific. (2016). Phusion™ High-Fidelity PCR Kit User Guide (MAN0013363). https://assets.thermofisher.com/TFSAssets/LSG/manuals/MAN0013363_Phusion_HiFi_PCR_Kit_UG.pdf.

Materials

1. PCR Tubes
2. Nuclease Free H₂O
3. Q5 5X buffer
4. 10nM dNTPs
5. Forward and Reverse Primers
6. Template DNA

1. Select a microbial colony of interest and set plate aside for collection with pipette tip
2. Label the PCR tube accordingly and record the materials within the tube in the lab notebook
3. Add the following reagents/components into the PCR tube in the order stated:

   Component	50 µL Reaction
   DreamTaq Green PCR Master Mix 2X	25 µL
   Forward and Reverse Primers	2.0 µM
   Template DNA	1 µg
   Nuclease Free H2O	Fill to Amount 50 µL

   DNA is added by swirling some of the colony selected into PCR tube.
4. In the thermocycler complete the following cycling conditions:

   Cycle Steps	Parameters	Amount of Cycles
   Initial Denaturation	95 °C for 3 min	1
   Denature	95 °C for 30 sec	35
   Anneal	64 °C for 30 sec	35
   Extension	72 °C for 1 min	35
   Final Extension*	72 °C for 15 min	1

   Final extension time depends on length of product being amplified. For products less than 2 Kb, use a time of 1 minute. For products greater than 2 Kb, add an additional 1 minute per Kb.
5. For long-term storage keep PCR tubes at -20°C

1. Thermo Fisher Scientific. (2016). DreamTaq Green PCR Master Mix Product Information (MAN0012704). https://documents.thermofisher.com/TFS-Assets/LSG/manuals/MAN0012704_DreamTaq_Green_PCR_MasterMix_K1081_UG.pdf.

Courtesy of Dr. George van der Merwe.
(A) SDM-PCR:
Materials

1. PCR Tubes
2. Nuclease Free H₂O
3. Q5 Polymerase
4. Forward and Reverse Primers
5. Template DNA
6. dNTPs
7. Q5 buffer

1. Design SDM primers to partially overlap, with the desired mutation located in the middle of each primer
2. Label the PCR tube accordingly and record the materials within the tube in the lab notebook
3. Add the following reagents/components into the PCR tube in the order stated:

   Component	50 µL Reaction	Final Concentration
   H2O	To 50 µL
   5X Q5 Buffer	10 µL	1X
   10 mM dNTPs	1 µL	1 mM
   Template DNA	Variable	10 ng
   25 mM Forward Primer	0.5 µL	0.5 mM
   25 mM Reverse Primer	0.5 µL	0.5 mM
   2U Q5 Polymerase	1 µL	1 U

4. In the thermocycler complete the following cycling conditions:

   Cycle Steps	Parameters	Amount of Cycles
   Initial Denaturation	95 °C for 3 min	1
   Denature	95 °C for 30 sec	20
   Anneal	64 °C for 30 sec	20
   Extension	72 °C for 1 min	20
   Final Extension*	72 °C for 15 min	1

   Final extension time depends on length of product being amplified. For products less than 2 Kb, use a time of 1 minute. For products greater than 2 Kb, add an additional 1 minute per Kb.
5. For long-term storage keep PCR tubes at -20°C

1. 1.5 mL Microfuge Tube
2. Nuclease Free H₂O
3. 10X FD buffer
4. PCR product
5. Dpn1

1. Add the following reagents/ components into the microcentrifuge tube in the order stated:

Component	Volume
H2O	13 µL
10X FD Buffer	5 µL
PCR Product	10 µL
Dpn1	2 µL

2. Spin down in centrifuge on short / pulse cycle to ensure contents are in bottom of tube
3. Incubate at 37°C for a minimum of 3.5 hours
4. Remove from heat and run on agarose gel, followed by sequencing. For later use, store at –20 °C. Template DNA should be destroyed and product can now be transformed into bacteria

Materials

1. ThermoScientific Miniprep Kit
2. P1000 Pipette & Tips
3. P200 Pipette & Tips
4. 1.5 mL Microfuge Tubes

1. Lucate, J. (2018). Plasmid DNA miniprep protocol for EZ-10 Spin Column Plasmid DNA minipreps kit (BS413), 50 preps v3. Preprint, https://doi.org/10.17504/protocols.io.n8tdhwn https://doi.org/10.17504/protocols.io.n8tdhwn.

Materials

1. ThermoScientific GeneJET PCR Purification Kit
2. P1000 Pipette & Tips
3. P200 Pipette & Tips
4. 1.5 mL Microfuge Tubes
5. PCR Product

1. Thermo Fisher Scientific. GeneJET PCR Purification User Guide. (MAN0012662). https://documents.thermofisher.com/TFS-Assets/LSG/manuals/MAN0012662_GeneJET_PCR_Purification_UG.pdf.

Materials

1. BioBasic EZ-10 Spin Column DNA Cleanup Miniprep Kit
2. P1000 Pipette & Tips
3. P200 Pipette & Tips
4. 1.5 mL Microfuge Tubes
5. PCR or GGA Product

1. BioBasic. (2018). EZ-10 Spin Column DNA Cleanup Miniprep Kit Product Information. (BS367/ BS368/ BS668). https://www.biobasic.com/us/amfilerating/file/download/file_id/28865/

1. DH5α Competent Cells
2. SOC media
3. LB plate with desired antibiotic or selective media of choice
4. DNA to be transformed
5. 1.5 mL Microfuge Tubes
6. Ice Bucket
7. 42°C water bath
8. Shaking Incubator (set to 225 rpm, 37°C)
9. Sterile Spreaders (glass or disposable)

1. Thaw DH5α competent cells on ice. Chill 1.5 mL microfuge tubes on ice while competent cells thaw
2. Once thawed gently mix the cells – pipette up and down
3. Aliquot 50 μL of competent cells into chilled 1.5 mL microfuge tubes from (1)
4. Add 6 μL of DNA directly into a tube of competent cells. Mix well by gently flicking the tube several times or pipetting up and down
5. Incubate cells on ice for 30 minutes
6. Heat-shock the cells for exactly 30 seconds in a 42°C heat block
   • Do not mix or shake the tube
7. Immediately transfer back to ice and incubate on ice for 2 minutes
8. Add 250 μL of room temperature SOC recovery media to each transformation tube and mix- pipette up and down
9. Shake the tube at 225 rpm for 1 hour at 37°C. Use an empty P10 or P200 pipette tip box and lay microfuge tubes on their side with a piece of table holding all of them in place
10. Spread plate all competent cells onto LB + antibiotic plates
11. Incubate overnight at 37°C

1. Any liquids which have come into contact or was used to inoculate/grow E.coli cells must have 20% bleach added to and let sit for 30 minutes before disposal down the drain and washed with dish soap or detergent
2. Any disposable materials which into contact or was used to inoculate/grow E.coli cells can be safely disposed of in the appropriate biohazard waste

1. Thermo Fisher Scientific. (n.d.). Thermo Scientific DH5α competent cells [Product catalog page, Catalog No. EC0112]. Thermo Fisher Scientific. https://www.thermofisher.com/order/catalog/product/EC0112

1. Inoculate 5 mL YPD broth with S. cerevisiae in a sterile 16 mm test tube. One 5 mL culture makes enough cells for 10 transformations
2. Incubate overnight at 30°C, shaking at 180 rpm

1. Prewarm 50 mL YPD in a sterile 250 mL flask to 30°C
2. Dilute the overnight culture 1/10 or 1/20 in dH₂O , and of YPD media, and read OD₆₀₀. Multiply by dilution factor to determine the actual OD₆₀₀ of the overnight culture
3. Inoculate 50 mL of prewarmed YPD to a final OD₆₀₀ of 0.2. Use C₁V₁ = C₂V₂ to calculate what volume is required for inoculation
   
4. Incubate for 4-6 hours at 30°C, shaking at 180 rpm until OD₆₀₀ reaches 0.8 - 1.0

1. Transfer cells from previous step (5) to 50 mL falcon tubes
2. Centrifuge at 3000 * g, room temperature, for 5 minutes, then discard the supernatant
3. Resuspend pellet in 25 mL sterile dH₂O, centrifuge at 3000 * g, room temperature, for 5 minutes, then discard the supernatant
4. Repeat #3, for a total of 2 washes
5. Resuspend the final pellet in 1 mL sterile dH₂O

1. Boil 1 mL of single-stranded carrier DNA for 5 minutes, then immediately place on ice
2. Combine desired amount of 50% w/v PEG 3350 and 1.0 M Lithium acetate in a sterile falcon tube. On ice, add desired amount of single-stranded carrier DNA. Mix thoroughly with a pipette

   Reagent	Volume per plasmid	Final Concentration
   50% w/v PEG 3350	240 µL	33%
   1.0 M Lithium Acetate	36 µL	0.1 M
   Boiled ss-Carrier DNA (10 mg/mL)	10 µL	0.28 mg/mL
   Total	286 µL

   Note: Make enough T-mix for 1 extra plasmid to account for pipetting error.

1. Transfer resuspended cells to a 1.5 mL centrifuge tube
2. Centrifuge at 8000 * g for 30 seconds, discard the supernatant
3. Resuspend in 1 mL sterile dH₂O
4. Dispense 100 µL of the resuspended cells into separate 2 mL centrifuge tubes
5. Centrifuge at 8000 * g for 30 seconds, discard supernatant
6. Resuspend the pellet in 286 µL of T Mix
7. Add transforming DNA + sterile water to reach a final volume of 74 µL. BY- strains use 0.5 µg of DNA

1. Incubate tubes at 42°C for 20 minutes
2. Centrifuge at 1000 × g (3000 rpm) for 2 minutes
3. Discard the supernatant
4. Add 1 mL sterile dH₂O, pipette to resuspend the pellet
5. Spread 200 µL on SC-URA plates, or another selection plate for desired trait being transformed. Store any remaining yeast suspension at 4°C. Non-transformed yeast cells will lose competency and can be disposed of
6. Let liquid soak in, then incubate inverted at 30°C. Colonies should appear in 3-4 days (incubate for 7 days if none appear). Restreak onto the same selective media

1. Gietz, R. D., & Schiestl, R. H. (2007). High-efficiency yeast transformation using the LiAc/SS carrier DNA/PEG method. Nature Protocols, 2(1), Article 1. https://doi.org/10.1038/nprot.2007.13
2. Gietz, R. D., & Woods, R. A. (2002). Transformation of yeast by lithium acetate/single-stranded carrier DNA/polyethylene glycol method. Methods in Enzymology, 350, 87-96. https://doi.org/10.1016/s0076-6879(02)50957-5

Materials

1. 1L Deionized H₂O
2. 10g Tryptone
3. 5g Yeast Extract
4. 5g NaCl
5. Optional: 15g Agar or Agarose

1. Add 500 mL of dH₂O to a beaker with a stir bar and turn stir plate on. Adding reagents directly to water minimizes airborne loss of product
2. Weigh out Tryptone and slowly add to beaker
3. Weigh out Yeast extract slowly add to beaker
4. Weigh out NaCl add to bottle
5. Mix until dissolved (low amount of heat may be used to help with this process)
6. Pour mixture into a graduated cylinder and add water to 1000 mL, then transfer to bottle
7. Lightly tighten bottle lid
   • If it is too tight, it will cause an explosion inside the autoclave due to pressure buildup of gases within the bottle
8. Label with autoclave tape
9. Autoclave at L20

FOR SOLID MEDIA: Pour 20-25ml of autoclaved medium per plate

1. In an aseptic field, calculate the amount of stock antibiotics needed to achieve the appropriate working concentration using C₁V₁ = C₂V₂
2. Pipette antibiotics into the bottle of media and gently swirl to completely mix

   Antibiotic	Working Concentration
   Ampicillin	100 µg/mL
   Chloramphenicol	25 µg/mL
   Kanamycin	50 µg/mL

1. Liquid LB Medium with NO bacteria grown can safely be poured down the drain
2. Solid LB Medium with NO bacteria grown can be safely thrown away in the garbage
3. Apparatus which contains LB medium that has been INOCULATED with any form of microorganism, or which has suspected CONTAMINATION must first be autoclaved before disposal

1. Elbing, K. L., & Brent, R. (2019). Recipes and Tools for Culture of Escherichia coli. Current protocols in molecular biology, 125(1), e83. https://doi.org/10.1002/cpmb.83
2. 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

Materials

1. Cryovials
2. Sterile dH₂O
3. 100% Glycerol
4. Overnight Culture Bacteria

1. In the cryovials mix equal parts of 100% glycerol and dH₂O to make a 50% glycerol solution
   • 0.75 mL of 100% Glycerol
   • 0.75 mL of dH₂O
2. Add 500 µL of the overnight culture to cryovials from (1)
3. Ensure vials are adequately labeled and contain all necessary information
4. Store at -80°C

1. Addgene. (n.d.). Protocol - How to create a bacterial glycerol stock. Retrieved June 10, 2025, from https://www.addgene.org/protocols/create-glycerol-stock/

Materials

1. Autoclaved Culture Tubes
2. Serial pipette tips and pipette gun
3. Desired Antibiotic(s)
4. Liquid LB Media, YPD media, or SC-URA Media
5. Sterile loop
6. Petri Dishes/ Plates of E. coli for Culturing, or Petri Dishes/ Plates of S. cerevisiae for Culturing

1. Add 5 mL of appropriate media to culture tube
   • For glycerol stocks, add 6 mL
2. Add the appropriate amount of antibiotic to each culture tube to achieve the desired working concentration
   • Using the formula C₁V₁ = C₂V₂

   Antibiotic	Working Concentration
   Ampicillin	100 µg/mL
   Chloramphenicol	25 µg/mL
   Kanamycin	50 µg/mL

3. Vortex all culture tubes to thoroughly mix antibiotics and media. Avoid vortexting aggressively to prevent the media from coming in contact with the lid or spilling over
4. Using a flame-sterilized loop, pick up the colony being amplified. Tap onto the inside of the glass culture tube. Flame tip to re-sterilize
5. Vortex culture tube until the colony has been picked up by the media. Avoid vortexting aggressively to prevent the media from coming in contact with the lid or spilling over
6. Incubate at 37°C with shaking at 225 rpm for bacteria, and 30 °C with shaking at 180 rpm for yeast

1. If ANY media has come into contact with bacteria/ microorganisms, or is suspected of being contaminated, add 20% bleach and allow them to sit overnight before discarding down the drain and cleaning culture
2. If media has NOT come into contact with any bacteria/ microorganisms discard down the drain and cleaning culture tube

Materials

1. 900mL Deionized H₂O
2. 10g Yeast Extract
3. 20g Peptone
4. 2% w/v Dextrose/Glucose - prepare from 20% w/v stock solution
5. Optional: 15g Agar

1. Add 500 mL of dH₂O to a beaker with a stir bar and turn stir plate on. Adding reagents directly to water minimizes airborne loss of product
2. Weigh out Yeast Extract and slowly add to beaker
3. Weigh out Peptone and slowly add to beaker
4. Mix until dissolved
   • low amount of heat may be used to help with this process
5. Pour mixture into a graduated cylinder and add water to 900 mL, then transfer to bottle
6. Lightly tighten bottle lid
   • If it is too tight, it will cause an explosion inside the autoclave due to pressure buildup of gases within the bottle
7. Label with autoclave tape
8. Autoclave at L20
9. Add 100 mL of 20% w/v Dextrose/Glucose as media is and gently swirl to mix. Do not add Dextrose/ Glucose prior to autoclaving. Autoclaving sugars causes caramelization and the formation of toxic byproducts that can inhibit microbial growth and alter media composition

FOR SOLID MEDIA: Pour 20-25ml of autoclaved medium per plate

1. Liquid media with NO microbes grown can safely be poured down the drain
2. Solid media with NO microbes grown can be safely thrown away in the garbage
3. Apparatus which contains media that has been INOCULATED with any form of microorganism, or which is suspected of CONTAMINATION must first be autoclaved, before disposal

1. Novoprotein. (n.d.). YPD broth preparation. NovoPro Lab Tools. https://www.novoprolabs.com/tools/buffer-preparations-and-recipes/ypd-broth
2. DSMZ - German Collection of Microorganisms and Cell Cultures GmbH. (n.d.). Medium 393: YPD medium (liquid or agar) [PDF]. https://www.dsmz.de/microorganisms/medium/pdf/DSMZ_Medium393.pdf

1. 900mL Deionized H₂O
2. 6.7g Yeast Nitrogen Base without Amino Acids
3. 1.92g SC-URA
4. 2% w/v Dextrose/Glucose - prepare from 20% w/v stock solution
5. Optional: 15g Agar

1. Add 500 mL of dH₂O to a beaker with a stir bar and turn stir plate on.
   • Adding reagents directly to water minimizes airborne loss of product
2. Weigh out Yeast Nitrogen Base without Amino Acids and slowly add to beaker
3. Weigh out SC-URA and slowly add to beaker
4. Mix until dissolved
   • low amount of heat may be used to help with this process
5. Pour mixture into a graduated cylinder and add water to 900 mL, then transfer to bottle
6. Lightly tighten bottle lid
   • If it is too tight, it will cause an explosion inside the autoclave due to pressure buildup of gases within the bottle
7. Label with autoclave tape
8. Autoclave at L20
9. Add 100 mL of 20% w/v Dextrose/Glucose as media is and gently swirl to mix. Do not add Dextrose/ Glucose prior to autoclaving. Autoclaving sugars causes caramelization and the formation of toxic byproducts that can inhibit microbial growth and alter media composition

FOR SOLID MEDIA: Pour 20-25ml of autoclaved medium per plate

1. Liquid media with NO microbes grown can safely be poured down the drain
2. Solid media with NO microbes grown can be safely thrown away in the garbage
3. Apparatus which contains media that has been INOCULATED with any form of microorganism, or which is suspected of CONTAMINATION must first be autoclaved, before disposal

Materials

1. 95% Ethanol
2. MilliQ H₂O or ddH₂O
3. Graduated Cylinder
Note: When handling 95% ethanol work in the fume hood

1. Using the formula C₁V₁ = C₂V₂ determine the amount of stock solution (95% ethanol) required for the desired amount of 75% ethanol
2. Fill graduated cylinder with desired amount of MilliQ H₂O or ddH₂>O
3. In the fume hood, add ~150 mL aliquots of 95% ethanol and lightly swirl graduated cylinder to mix the solution – continue adding ethanol until the appropriate volume is achieved
4. Fill spray bottles in the fume hood with 75% ethanol, close, and remove. Close the fume hood after use and ensure 95% stock ethanol is stored in a flammable's cabinet
Consideration i) Ensure that you are wearing a lab coat and gloves, as 95% ethanol is highly flammable and can irritate the skin upon contact

Materials

1. EDTA disodium salt (MW=372.24 g/mol)
2. MilliQ H₂O
3. Sodium Hydroxide (NaOH)

1. In a 500 mL bottle dissolve 93.05 grams of EDTA disodium salt in 400 mL MilliQ H₂O
2. Add stir bar and begin mixing with low-medium heat
3. Slowly add ~2.5 grams of sodium hydroxide to EDTA solution until the mixture becomes clear (this is now roughly pH 8)
   • When adding the sodium hydroxide allow each addition of ~2.5g to fully dissolve in solution before adding the next batch
4. Once a clear solution is achieved check pH with pH stick
5. Label and store at room temperature in chemical cabinet

1. Behle, A., & Pawlowski, A. (2018, April 6). Recipe for 50x TAE buffer (Version gtvbwn6) [Protocol]. Protocols.io. https://doi.org/10.17504/protocols.io.gtvbwn6

1. Tris-base (MW = 121.14 g/mol)
2. Acetic Acid, Glacial
3. 0.5M EDTA
4. MilliQ H₂O
5. Graduated Cylinder (Glass 100 mL)
Note: This is for 500mL of buffer

1. Weigh out 121g of Tris-base and dissolve in 700 mL of MilliQ H₂O
2. Add 28.5 mL of Acetic Acid, Glacial
3. Add 50 mL of 0.5M EDTA
4. Add a stir bar and mic on low-medium heat
5. Once solution is clear, properly label and store at room temperature in chemical cabinet

1. Behle, A., & Pawlowski, A. (2018, April 6). Recipe for 50x TAE buffer (Version gtvbwn6) [Protocol]. Protocols.io. https://doi.org/10.17504/protocols.io.gtvbwn6

Materials

1. 50X TAE Buffer
2. MilliQ H₂O

1. In a 1L bottle measure and add 20mL of 50X TAE
2. Add 980 mL of MilliQ H₂O
3. Cap the bottle tightly and mix by inverting and swirling
4. Label and store in the appropriate location within the chemical cabinet

1. Behle, A., & Pawlowski, A. (2018, April 6). Recipe for 50x TAE buffer (Version gtvbwn6) [Protocol]. Protocols.io. https://doi.org/10.17504/protocols.io.gtvbwn6

Materials

1. 99% Glycerol
2. MilliQ H₂O

1. In a beaker with a stir bar, add equal parts 99% glycerol and MilliQ H₂O. Turn on stir function and mix till dissolved
2. In a sterile field, filter sterilize 50% Glycerol into sterile 50mL falcon tubes
3. Label and store in 50 mL aliquots in the appropriate location within the chemical cabinet

Materials

1. 20g D-Glucose (Dextrose)
2. 100mL MilliQ H₂O

1. Weigh out 20g D-Glucose and dissolve in 100mL MilliQ H₂O
2. In a sterile field, filter sterilize 20% D-Glucose into sterile 50mL falcon tubes
3. Label and store in 50 mL aliquots in the appropriate location within the chemical cabinet

Materials

1. 25g Polyethylene Glycol 3350
2. 50mL MilliQ H₂O

1. Weight out 25g PEG and dissolve in 50mL MilliQ H₂O
2. In a sterile field, filter sterilize 50% PEG 3350 into a sterile 50mL falcon tube
3. Label and store in the appropriate location within the chemical cabinet

Materials

1. 10.2g Lithium Acetate
2. 100mL MilliQ H₂O

1. Dissolve 10.2g Lithium Acetate in 100 mL MilliQ H₂O
2. In a sterile field, filter sterilize 1M Lithium Acetate into 2 sterile 50mL falcon tubes
3. Label and store in the appropriate location within the chemical cabinet

Courtesy of Stockinger Lab.
Materials

1. 100 mM Tris-HCl (pH 8)
2. 1 mM Dithiothreitol (DTT)
3. 20% Glycerol
4. MilliQ H₂O

1. Combine Tris-HCl, and glycerol in a breaker. Add stir bar and turn stir plate on
2. Add to graduated cylinder and top off to final volume with MilliQ H₂O
3. Pour into stock bottle, invert to mix
4. Before use, aliquot out required amount and add in DTT.
   • DTT degrades rapidly and thus should be added immediately before using a breaking buffer

1. Stebbins, J. Assay of β-Galactosidase in Yeast. Stockinger Lab. https://stockingerlab.osu.edu/sites/stockinger/files/imce/PDFs/Protocols/BetaGalAssays.pdf

Courtesy of Stockinger Lab.
Materials

1. 1.61g Na₂HPO₄ • 7H₂O
2. 0.55g NaH₂PO₂ • H₂O
3. 0.075g KCl
4. 0.0246 MgSO₄• 7H₂O
5. MilliQ H₂O to final volume of 100 mL

1. Add 50 mL of MilliQ H₂O to a beaker with a stir bar and turn on stir function
2. Weigh out reagents on a fine balance scale. Rinse out weigh boats after each use with 2 mL MilliQ H₂O and add to beaker to get any particulate matter left over. Mix till dissolved
3. Using a pH strip or pH meter to confirm pH is 7
4. Add to graduated cylinder and top off with MilliQ H₂O to 100 mL. Pour into stock bottle
5. Store at 4 °C and add DTT to 10 mM before each use.
   • DTT degrades rapidly and thus should be added immediately before using Z buffer

1. Stebbins, J. Assay of β-Galactosidase in Yeast. Stockinger Lab. https://stockingerlab.osu.edu/sites/stockinger/files/imce/PDFs/Protocols/BetaGalAssays.pdf

Courtesy of Stockinger Lab.
Materials

1. 2-Nitrophenyl-beta-D-galactopyranoside, 99% (ONPG)
2. Z buffer

1. Add 4mg ONPG per 1 mL Z buffer
2. Dissolve with a stir bar. Cover while dissolving to protect, as ONPG is photosensitive
3. Store at -20 °C

1. Stebbins, J. Assay of β-Galactosidase in Yeast. Stockinger Lab. https://stockingerlab.osu.edu/sites/stockinger/files/imce/PDFs/Protocols/BetaGalAssays.pdf

Courtesy of Stockinger Lab.
Materials

1. 26.5g Na₂CO₃ (Sodium Carbonate) • 7H₂O
2. 250 mL MilliQ H₂O

1. Add 100 mL MilliQ H₂O to a beaker with a stir bar. Turn on stir function
2. Add Na₂CO₃ and dissolve with a stir bar
3. Pour into a graduated cylinder and top off volume to 250 mL
4. Store in stock bottle at 4°C

1. Stebbins, J. Assay of β-Galactosidase in Yeast. Stockinger Lab. https://stockingerlab.osu.edu/sites/stockinger/files/imce/PDFs/Protocols/BetaGalAssays.pdf

Materials

1. 1.54mg (Dithiothreitol) DTT
2. 1 mL MilliQ H₂O

1. Read DTT SDS
2. Add 1.54mg DTT to a microfuge tube with 1 mL MilliQ H₂O. Vortex on high to dissolve
3. Store at -20 °C

CAUTION: PMSF is ANTI-PROTEASE, is a strong hydrogen-fluoride (HF) RELEASER, is highly CORROSIVE, and is ACUTELY TOXIC. Work in fume hood with appropriate PPE. Whenever possible, use a less toxic alternative.
Materials

1. 174.2mg Phenylmethylsulphonyl Fluoride (PMSF)
2. 1 mL anhydrous isopropanol

1. Read PMSF SDS
2. Add 174.2mg PMSF to a microfuge tube with 1 mL anhydrous isopropanol. Vortex on high to dissolve
3. Store at -20°C in flammables section