Team:CMUQ/Dspb Lab

Dspb Lab

DspB Lab

B1: Agarose gel

Materials

  • Agarose
  • dH2O
  • Ethedium Bromide
  • 10X TBE
  • Gel box + power
  • DNA ladder (check size)
  • DNA loading dye

Procedure

Making the gel:

  1. Weigh appropriate mass of Agarose into an Erlenmeyer flask
  2. Add appropriate volume of dH2O and 10% TBE, For example, to make 1% mini gel: 0.5 g agarose + 45 mL dH2O + 5 mL 10x TBE
  3. Microwave for 2 min, take out every 30 sec and swirl to dissolve
  4. Using hot hand protectors, take to bench and let cool
  5. Add appropriate volume of Ethidium Bromide and swirl to mix
  6. Pour onto previously setup gel box with comb
  7. Allow to solidify for ≈20 min
  8. Add 1% TBE
  9. Run for 30 - 45 min at 100 Volts
  10. Image and discard gel

B2: LB Media Preparation

Materials

  • Tryptone
  • NaCl
  • Yeast extract
  • dH2O
  • Autoclave
  • 1N NaOH

Procedure

  1. Dissolve 10 g tryptone, 5 g yeast extract, and 10 g NaCl in 950 mL deionized water.
  2. Adjust the pH of the medium to 7.0 using 1N NaOH and bring volume up to 1 liter.
  3. Autoclave on liquid cycle for 20 min at 15 psi. Allow solution to cool to 60°C.
  4. Store in the fridge.

B3: SOC Media Preparation

Introduction

Protocol adapted from: Here

  • Tryptone
  • Yeast Extract
  • NaCl
  • 250 mM KCL
  • 1N NaOH
  • 1M glucose
  • 2M MgCl2
  • dH2O

B4: LB Chl Plates Prep

Introduction

Adapted from: Here

Genetics lab recipe book in 2033

Materials

  • Antibiotic: Chloramphenicol
  • De-ionized Water
  • LB-Agar
  • Plates with lids
  • Erlenmeyer flasks
  • 60˚C water bath
  • Autoclave
  • Flame

Procedure

Preparing 250 mL LB-Agar

  1. Add the following into a 500 mL Erlenmeyer flask:
    2. PCR reagents
    Reagent Mass/Volume
    Tryptone 2.5 g
    Yeast Extract 1.25 g
    Sodium Chloride 2.5 g
    Agar 3.75 g
    Thymine (2mg/mL) 1.75 mL
    1N NaOH 0.5 mL
    De-Ionized Water 250 mL
  2. Swirl flask and cover the opening with aluminum foil and tape the bottle with autoclave tape.
  3. Label the bottle with your initials, the date, and the bottle contents.
  4. Place the gel mix in the autoclave and run on a setting that gets the sample to at least 121 ℃ under 20 psi for at least 30 min. The high pressure will prevent your gel mix from boiling over at high temperature.
  5. Turn the water bath and heat to 60˚C
  6. After 1 - 1.5 hours the LB agar autoclave was complete, the flask was taken using rubber hot hand protector and placed in a 60˚C water bath

    7. Preparing Chloramphenicol antibiotic

  7. Brenadette weighed 12.5 mg chroamphinicol in powder form (box in the 4˚C fridge) into a 2.5 mL microcentrifuge tube
  8. Dissolve in 500 µL ethanol to make 25mg/mL (I used 70% Ethanol)
  9. Vortex to mix and leave on a rack untill 10 min has elapsed on the LB agar solution in water bath and the pouring station is prepared

    10. Pour Plating

  10. Use the hood next to the 4˚C fridges in 2033
  11. Spray down the bench with bleach and 70% ethanol solution and wipe down with a paper towel.
  12. Obtain a stack (20) small plates, and stack them in the hood.
  13. Open the plates and place the lids on the side
  14. Obtain LB Agar flask from water bath, add 250 µL of 25 mg/mL Chloramphenicol and swirl to mix
  15. Pour directly from the Erlenmeyer flask, a little more than half of the plate.
  16. Once all the plates are poured, ignite the flame and quickly run over the plates to avoid contamination.
  17. Leave your plates to solidify. it takes ≈ 20 min to solidify at room temperature.
  18. Close the plates with the lids. and stack on top of each other.
  19. Label the plates withblue and orange markers coding for Chloramphenicol LB
  20. Label the plates with the date and initials
  21. The plates are then placed in a box with iGEM label and stored in 4 ℃ fridge.

B5: Competent Cell Test

Introduction

The kit includes three vials of purified plasmid DNA from BBa_J04450 (RFP construct) in plasmid backbone pSB1C3. Each vial contains 50 µL of DNA at a different concentration: 100 pg/µL, 50 pg/µL, 10 pg/µL. Perform transformations with each of these to determine how efficient your competent cells are.

Protocol from:

Materials

  • 70% ethanol
  • Paper towels
  • Lab marker / Sharpie
  • 1.5 mL microcentrifuge tubes
  • Container for ice
  • Ice
  • Competent cell aliquot(s)
  • Competent Cell Test Kit
  • Agar plates with chloramphenicol
  • 42°C Waterbath (or hot water source and thermometer)
  • 37°C Incubators (oven and shaker)
  • SOC media
  • Sterile glass beads or sterile cell spreader
  • Pipettor
  • Pipette tips

Procedure

Estimated time: 30 minutes active, 1.5 hours incubation

  1. Clean your working area by wiping down with 70% ethanol.
  2. Thaw competent cells on ice. Label one 1.5 mL microcentrifuge tubes for each transformation and then pre-chill by placing the tubes on ice.

    3. Do triplicates (3 each) of each concentration if possible, so you can calculate an average colony yield.

  3. Spin down the DNA tubes from the Competent Cell Test Kit/Transformation Efficiency Kit to collect all of the DNA into the bottom of each tube prior to use. A quick spin of 20-30 seconds at 8,000-10,000 rpm will be sufficient. Note: There should be 50 µL of DNA in each tube sent in the Kit.
  4. Pipet 1 µL of DNA into each microcentrifuge tube.
  5. TPipet 50 µL of competent cells into each tube. Flick the tube gently with your finger to mix.
  6. Incubate on ice for 30 minutes.

    7. Pre-heat waterbath now to 42°C. Otherwise, hot water and an accurate thermometer works, too!

  7. Heat-shock the cells by placing into the waterbath for 45 seconds (no longer than 1 min). Be careful to keep the lids of the tubes above the water level, and keep the ice close by.
  8. Immediately transfer the tubes back to ice, and incubate on ice for 5 minutes.
  9. Add 950 µL of SOC media per tube, and incubate at 37°C for 1 hour shaking at 200-300rpm.

    10. Prepare the agar plates during this time: label them, and add sterile glass beads if using beads to spread the mixture.

    Next Day

  10. Count the number of colonies on a light field or a dark background, such as a lab bench. Use the following equation to calculate your competent cell efficiency. If you've done triplicates of each sample, use the average cell colony count in the calculation.

    11. Efficiency (in cfu/µg) = [colonies on plate (cfu) / Amount of DNA plated (ng)] x 1000 (ng/µg)

    *Note: The measurement "Amount of DNA plated" refers to how much DNA was plated onto each agar plate, not the total amount of DNA used per transformation. You can calculate this number using the following equation:

    Amount of DNA plated (ng) = Volume DNA added (1 µL) x concentration of DNA (refer to vial, convert to ng/µL) x [volume plated (100 µL) / total reaction volume (1000 µL)]

    Results:

  11. Competent cells should have an efficiency of 1.5x108 to 6x108 cfu/µg DNA, where "cfu" means "colony-forming unit" and is a measurement of cells.
  12. Here are some sample results:

    13. DNA concentration 10 pg/µL 50 pg/µL 100 pg/µL

    of colonies 280 - 360 500 - 1000+ 1000+

B6: Transformation into ultra-competent DH5α

Introduction

Protocol adapted from:

Materials

  • Resuspended DNA to be transformed
  • 10pg/µl Positive transformation control DNA (e.g. pSB1C3 w/ BBa_J04450, RFP on high-copy chloramphenicol resistant plasmid. Located in the Competent Cell Test Kit.)
  • Competent Cells (50µl per sample)
  • 1.5mL Microtubes
  • SOC Media (950µL per sample)
  • Petri plates w/ LB agar and antibiotic (2 per sample)
  • Floating Foam Tube Rack
  • Ice & ice bucket
  • Lab Timer
  • 42°C water bath
  • 37°C incubator
  • Sterile spreader
  • Pipettes and Tips (10µl, 20µl, 200µl recommended)
  • Microcentrifuge

Procedure

Estimated bench time: 1 hour. Estimated total time: 2 hours (plus 14-18 hour incubation)

  1. Resuspend DNA in selected wells in the Distribution Kit with 10µl dH20. Pipet up and down several times, let sit for a few minutes. Resuspension will be red from cresol red dye.
  2. Label 1.5ml tubes with part name or well location. Fill lab ice bucket with ice, and pre-chill 1.5ml tubes (one tube for each transformation, including your control) in a floating foam tube rack.
  3. Thaw competent cells on ice: This may take 10-15min for a 260µl stock. Dispose of unused competent cells. Do not refreeze unused thawed cells, as it will drastically reduce transformation efficiency.
  4. Pipette 50µl of competent cells into 1.5ml tube: 50µl in a 1.5ml tube per transformation. Tubes should be labeled, pre-chilled, and in a floating tube rack for support. Keep all tubes on ice. Don’t forget a 1.5ml tube for your control.
  5. Pipette 1µl of resuspended DNA into 1.5ml tube: Pipette from well into appropriately labeled tube. Gently pipette up and down a few times. Keep all tubes on ice.
  6. Pipette 1µl of control DNA into 2ml tube: Pipette 1µl of 10pg/µl control into your control transformation. Gently pipette up and down a few times. Keep all tubes on ice.
  7. Close 1.5ml tubes, incubate on ice for 30min: Tubes may be gently agitated/flicked to mix solution, but return to ice immediately.
  8. Heat shock tubes at 42°C for 45 sec: 1.5ml tubes should be in a floating foam tube rack. Place in water bath to ensure the bottoms of the tubes are submerged. Timing is critical.
  9. Incubate on ice for 5min: Return transformation tubes to ice bucket.
  10. Pipette 100µL of each transformation onto petri plates Spread with sterilized spreader or glass beads immediately. This helps ensure that you will be able to pick out a single colony.

    11. If higher concentration needed: spin down cells at 6800g for 3mins and discard 800µL of the supernatant. Resuspend the cells in the remaining 100µL, and pipette each transformation onto petri plates Spread with sterilized spreader or glass beads immediately. This increases the chance of getting colonies from lower concentration DNA samples.

  11. Incubate transformations overnight (14-18hr) at 37°C: Incubate the plates upside down (agar side up). If incubated for too long, colonies may overgrow and the antibiotics may start to break down; un-transformed cells will begin to grow.
  12. Pick single colonies: Pick single colonies from transformations: do a colony PCR to verify part size, make glycerol stocks, grow up cell cultures and miniprep.
  13. Count colonies for control transformation: Count colonies on the 100μl control plate and calculate your competent cell efficiency. Competent cells should have an efficiency of 1.5x10^8 to 6x10^8 cfu/µg DNA.

B7: Plasmid extraction (mini prep)

Introduction

Protocol adapted from:

Protocol fromcQIAprepⓇ Spin Miniprep Kit (50)

Materials

  • QIAprep Spin Miniprep Kit (50)

Procedure

Overnight Culture

  1. Pick a single colony and inoculate in 5 mL SOC media (might want to pick two colonies to have duplicates)
  2. Leave overnight shaking at 37˚C

Next Day: Mini-prep

  1. Pellet 1–5 ml bacterial overnight culture by centrifugation at >8000 rpm (6800 x g) for 3 min at room temperature (15–25°C).
  2. Resuspend pelleted bacterial cells in 250 μl Buffer P1 and transfer to a microcentrifuge tube.
  3. Add 250 μl Buffer P2 and mix thoroughly by inverting the tube 4–6 times until the solution becomes clear. Do not allow the lysis reaction to proceed for more than 5 min. If using LyseBlue reagent, the solution will turn blue.
  4. Add 350 μl Buffer N3 and mix immediately and thoroughly by inverting the tube 4–6 times. If using LyseBlue reagent, the solution will turn colorless.
  5. Centrifuge for 10 min at 13,000 rpm (~17,900 x g) in a table-top microcentrifuge.
  6. Apply 800 μl supernatant from step 5 to the QIAprep 2.0 spin column by pipetting.
  7. Centrifuge for 30–60 s and discard the flow-through
  8. Recommended: Wash the QIAprep 2.0 spin column by adding 0.5 ml Buffer PB. Centrifuge for 30–60 s and discard the flow-through. Note: This step is only required when using endA+ strains or other bacteria strains with high nuclease activity or carbohydrate content.
  9. Centrifuge for 1 min to remove residual wash buffer.
  10. Place the QIAprep 2.0 column in a clean 1.5 ml microcentrifuge tube. To elute DNA, add 50 μl Buffer EB (10 mM TrisCl, pH 8.5) or water to the center of the QIAprep 2.0 spin column, let stand for 1 min, and centrifuge for 1 min. If the extracted DNA is to be analyzed on a gel, add 1 volume of Loading Dye to 5 volumes of purified DNA. Mix the solution by pipetting up and down before loading the gel.

B8: PCR DspB DspA

Introduction

Will do total 6 PCR tubes + 1 negative control

3 samples of DspB DspA, from parts: 211 and 201

2 annealing temperatures: 53˚C and 58˚C for each sample

Materials

  • Master Mix
  • DNA
  • Nuclease free water
  • Primers

Procedure

  1. Prepare PCR tubes:




B9: Gel Extraction

Introduction

The goal of this experiment is to extract a plasmid DNA from an agarose gel This protocol is for cleanup of DNA fragments of 40 bp to 50 kb. The yellow color of Buffer QX1 indicates a pH ≤7.5. Add ethanol (96–100%) to Buffer PE concentrate before use (see bottle label for volume). All centrifugation steps are carried out at 17,900 x g (~13,000 rpm) in a conventional tabletop microcentrifuge at room temperature (15–25°C).

Materials

  • Heating block or water bath at 50°C

Procedure

  1. Excise the DNA band from the agarose gel with a clean, sharp scalpel. Use a 1.5 ml microfuge tube for processing up to 250 mg agarose per tube.
  2. Weigh the gel slice in a colorless tube. Add Buffer QX1 according to DNA fragment size: 6 volumes for <100 bp; 3 volumes for 100 bp – 4 kb; 3 volumes with 2 volumes of water for >4 kb. Add 6 volumes of Buffer QX1 when using >2% or Metaphor agarose gels.
  3. Resuspend QIAEX II by vortexing for 30 s. Add QIAEX II to the sample and mix: Use 10 μl QIAEXII for ≤2 μg DNA; 30 μl for 2–10 μg DNA; and an additional 30 μl for each additional 10 μg DNA.

B10: Ligation

Introduction

Adapted from: Here

Procedure

  1. Set up the following reaction in a microcentrifuge tube on ice.

    2. T4 DNA Ligase should be added last. Note that the table shows a ligation using a molar ratio of 1:3 vector to insert for the indicated DNA sizes. Use the calculator from:

    The T4 DNA Ligase Buffer should be thawed and resuspended at room temperature.

  2. Gently mix the reaction by pipetting up and down and microfuge briefly.
  3. For cohesive (sticky) ends, incubate at 16°C overnight or room temperature for 10 minutes.
  4. For blunt ends or single base overhangs, incubate at 16°C overnight or room temperature for 2 hours (alternatively, high concentration T4 DNA Ligase can be used in a 10 minute ligation).
  5. Heat inactivate at 65°C for 10 minutes.
  6. Chill on ice and transform 1-5 μl of the reaction into 50 μl competent cells.

B11: IPTG induction

Introduction

Protocol adapted from: Here Induction in bacteria can be performed using one of two basic methods. Fast induction will not work for all proteins and may give you suboptimal yields. Slow induction may enhance the solubility of some proteins. The best method for your research will depend on your particular protein and the application. If you want optimal solubility both should be tested before scaling up. This protocol is generalized and optimal conditions may vary based on factors such as the bacterial strain, recombinant protein, and parent plasmid.

Procedure

Fast induction

  1. From a relatively fresh plate, pick a colony and grow overnight at 30°C (or 37°C) in 1-2 ml LB+Antibiotic in a 15 ml tube on a rotator or shaker.
  2. Dilute 1:50 (1:100 if 37°C overnight) in 2 ml LB+Antibiotic and grow 3-4 hours at 37°C in 15 ml tube in a rotator.
  3. Prepare 1 ml LB+Antibiotic+1mM IPTG (GoldBio Catalog # I2481) in a 15 ml conical and prewarm to 37°C about 10 minutes before use.
  4. After 3-4 hours remove 1 ml from tubes at 37°C and place in labeled 1.5 ml tubes. Centrifuge at maximum speed for 30 seconds at RT and remove supernatant. Freeze pellet at -20°C until
  5. needed. THIS IS THE UNINDUCED CONTROL.
  6. Add 1 ml prewarmed (37°C) LB+Antibiotic+1mM IPTG to 15ml tube and return to 37°C for 3-4 hours. This will get the final volume back to 2 ml and the final concentration of IPTG to 0.5mM.
  7. After 3-4 hours, transfer 1 ml from the induced sample to labeled 1.5 ml tubes and centrifuge at maximum speed for 30 seconds at RT and remove supernatant. Freeze pellet at -20°C until needed. THIS IS THE INDUCED SAMPLE.
  8. Sample preparation for SDS-PAGE: Add 100 μl of 1X Loading Buffer (see Solutions) with 1% BME to uninduced and induced samples. Vortex the samples for 10 seconds to 1 minute or until there are no clumps of bacteria. Boil the samples for 3 to 5 minutes. Centrifuge at maximum speed for 30 seconds at RT and load 5-25 μl (usually 10 μl) depending on gel (amount of protein, size of pellet, Western, etc.).

Slow induction

    For slow induction of protein follow fast induction protocol with the following changes:

  1. Add 1 ml LB+Antibiotic+1mM IPTG (prewarmed to 20°C) to 15 ml tube and incubate rotating or shaking at 20°C for 12-16 hours. This will get the final volume back to 2 ml and the final concentration of IPTG to 0.5mM.
  2. After 12-16 hours, transfer 1 ml from induced sample to labeled 1.5 ml tubes and centrifuge at maximum speed for 30 seconds at RT and remove supernatant. Freeze pellet at -20°C until needed. THIS IS THE INDUCED SAMPLE.

    3. *NOTES for induction:

     Fast Induction times may vary from 2-5 hours.

     IPTG can be varied from 0.1-1.0M.

    If you boil your sample too long they will become viscous from total release of cellular DNA. You can still use them if you can find an area of low viscosity. However, it is usually better just to repeat the experiment.

    *Solutions

    Loading Buffer - 4X Stock (to make total volume 40 ml)

    50mM Tris-HCl; pH 6.8 (Tris-HCl, GoldBio Catalog # T-095) (2.0 ml 1M Tris-HCl; pH 6.8)

     2% SDS (0.8 g SDS)

     10% Glycerol (4.0 ml 100% Glycerol)

     12.5mM EDTA (EDTA Disodium, GoldBio Catalog # E-210) (1.0 ml 0.5M EDTA)

    0.02 % Bromophenol Blue (Bromophenol Blue, GoldBio Catalog # B-092) (8 mg Bromophenol blue)

    Fill to volume with dH2O (~33 ml dH2O)

    Add fresh β-mercaptoethanol (BME) to 1% before using.