• Add 20 µL sterile H2O to desired well in distribution plate.
• Incubate at room temperature for ~10 minutes.
• Transfer resuspension to microcentrifuge tube.

Materials

• Competent Cells
• DNA template
• 800 µL LB
• LB+Antibiotic Plates

Procedure

• Thaw competent cells on ice.
• Transfer 50 µL of competent cells to chilled falcon tubes.
• Add 1 µL of template to cells (2.5 µL if dilute).
• Incubate on ice for 30 minutes.
• Heat schock in 42 °C water bath for 90 seconds.
• Immediately place back onto ice for 2 minutes.
• Add 800 µL of LB to each tube.
• Incubate at 37 °C for 1 hour.
• Place 200 µL of the transformed cells on plates containing LB and the appropriate antibiotic.
• Incubate overnight at 37 °C.

Materials

• 22 µL dH20
• 1 µL BSA
• 5 µLBuffer
• 20 µL Template
• 1 µL Enzyme 1
• 1 µL Enzyme 2

Procedure

• Mix reactants, adding enzyme last.
• Place at 37 °C for 3-5 hours.
• If not purifying or running on a gel immediately, increase to 80 °C for 20 minutes to kill enzymes (some enzymes need only a 65 °C heatkill, check enzyme).
• Run on a gel and extract product.

Materials

• 10 uL Q Solution
• 5 µL 10x Buffer
• 1.25 µL 10mM dNTPs
• 1 µL Template
• 1 µL Forward Primer
• 1 µL Reverse Primer
• 0.3 µL Taq
• 0.15 µL PFU
• 30 µL dH20

Procedure

• Mix reactants into PCR tubes. A large mix can be made and then aliquoted into PCR tubes if desired.
• Run in thermal cycler.

Materials

• 10 µL Q Solution
• 5 µL 10x Buffer
• 1.25 µL 10mM dNTPs
• 2.5 µL Forward Primer
• 2.5 µL Reverse Primer
• 0.3 µL Taq
• 0.15 µL PFU
• Template based on concentrations determined by SOEing calculator: “Link”
• ±18.30 µL dH20 based on concentrations of template (above) by SOEing calculator: “Link again”

Procedure

• Mix reactants into PCR tubes.
• Run in thermal cycler.
• Continue PCR SOEing of parts until completed.

Materials

• Digested Vector
• Digested Insert
• Water
• T4 DNA Ligase
• T4 DNA Ligase Buffer

Procedure

• Mix these materials in the amounts determined by the reaction volume calculator here.

Materials

• 34 mM NaH₂PO₄
• 64 mM K₂HPO₄
• 20 mM (NH₄)₂SO₄
• 1 µM FeSO₄
• 0.1 mM MgSO₄
• 10 µM CaCl₂
• 30 mM Ethylene Glycol
• 30 mM Glycolate
• 0.5% wt/vol Casein Acid Hydrolysate
• 1.5% wt/vol Agar (if making solid media)

Procedure

• Be sure to wear a lab coat while making this media, since both glycolate and ethylene glycol are irritants.
• Mix together NaH₂PO₄, K₂HPO₄, (NH₄)₂SO₄, FeSO₄, MgSO₄, and CaCl₂.
• Titrate to pH 7 with HCl.
• Add the glycolate and casein acid hydrolysate.
• Mix in the ethylene glycol in a fume hood.
• If making solid media, also mix in agar.
• Autoclave on an appropriate cycle.

Materials

• Luria Broth Media
• Ethylene Glycol
• Tecan M200 Pro
• E. coli

Procedure

• Be sure to wear a lab coat while making dilutions and aliquots, since ethylene glycol is an irritant.
• Inoculate bacterial colony in 5 mL of luria broth (LB) media in a 15 mL falcon tube to create a starter culture.
• Incubate at 37°C overnight on a shaker at 150 rpm.
• From the incubated starter culture, take 20 µL and add it into 3 mL of LB media.
• Incubate at 37°C for 2-3 hours on a shaker at 150 rpm.
• Measure the optical density (OD) reading every thirty minutes after two hours to reach an ideal range of 0.5 to 0.7.
• Use 15 mL conical tubes to make dilutions of ethylene glycol with LB media (see diagram below).
• With a 96-well round-bottom plate, fill the outermost wells with LB media to prevent dehydration on the edges.
• Add “blank” wells of just LB media to an entire column to serve as a control for the LB.
• Include a column for 0 mM ethylene glycol and LB media as a baseline for activity without ethylene glycol.
• Aliquot the remaining dilutions as the diagram below depicts.
• Add the appropriate amount of cells to the wells for a concentration of 0.01 OD before any of the readings.
• Follow the steps of your Tecan machine appropriate to the specific bacterial strain.

Materials

• Buffer A: Minimal medium + 0.2 M Tris (pH 7.5). per liter:
  • K₂HPO₄: 10.5g
  • KH₂PO₄: 4.5g
  • (NH₄)₂SO₄: 1g
  • Sodium Citrate * 2H₂O: 0.5g
  • 1M Tris (pH 7.5): 200mL Tris

• EMS (Sigma)
• 50mL conical Corning tubes
• 15mL Falcon tubes
• LB Broth
• Ethylene Glycol Agar Plates

Procedure

• 1. Prepare a 10mL liquid culture in LB in a 50mL conical tube and grow it overnight until it reaches OD 0.2.
• Chill the cells on ice and spin down the 10mL aliquots at max speed in Eppendorf Centrifuge 5810 R for 10 minutes.
• Wash twice with 10mL Buffer A.
• Pipet up and down to mix, pellet cells, decant supernatant.
• Re-suspend the pellet in 5mL of Buffer A and transfer the medium to a Falcon tube.
• Add (35μL, 70μL, or 105μL) of EMS into each tube of re-suspended cells.
• Close tubes and parafilm the lid 2X.
• Vortex to mix and place in a secondary container (50mL conical tube).
• Transfer to mixing platform and agitate at ~30 rpm at 37°C.
• Withdraw samples at fixed time points.
• At each time point, take 1mL aliquots of the sample and place in a 15mL Falcon tube. Place in a secondary container (50mL Conical tube) and spin at max speed in Eppendorf Centrifuge 5810 R for 5 minutes.
• Discard supernatant in waste container.
• Wash twice in 5mL of Buffer A and discard supernatant in waste container.
• Re-suspend in 5mL of Buffer A and titer for viable cells.
• Add 0.5mL of mutagenized culture into 10mL of LB broth in a 50mL conical tube.
• Grow cultures overnight at 37°C and plate for mutants on EG agar plates.
• Look for viable cells

Safety: Health Hazards

• Acute toxicity (oral, dermal, inhalation), category 4
• Skin irritation, category 2
• Eye irritation, category 2
• Skin sensitization, category 1
• Specific Target Organ Toxicity – Single exposure, category 3
• LD50 – 470mg/kg

Recommended Safety & Handling

• Always work in fume hood and wear lab coat, goggles, and gloves.

Procedure

• Extract genomic DNA using any proprietary gDNA extraction kit. Once DNA is extracted, quantify your DNA and run it on an agarose gel to check DNA quality.

Procedure

• Prepare a starter culture of MG1655 E. coli with pBad regulated and his-tagged LC-Cutinase along with a negative control.
• Inoculate new 5 mL cultures from the starter cultures at an OD 600 of about 0.05.
• Let the cultures grow until and OD 600 of 0.5, then inoculate two 100 mL cultures (one for the his-tagged protein and one for the negative control).
• Once these cultures are at an OD of 0.8, separate into two 50 mL cultures. Induce one at 10 uM, leave the other uninduced.
• Take 1 mL samples at different time points.
• Centrifuge samples at 5000g for 5 minutes then take off and save media.
• Wash cells with water.
• Resuspend with 300 uL of B-PER Protein Extraction Reagent and follow the instructions provided with that kit. After this, there should be samples of culture media, and soluble and insoluble whole cell lysate.
• Take 15 uL of each sample, including controls, and run on western blot.

Materials

• pNPB buffer (should be made and used in fumehood due to acetonitriles toxicity)
  • 10mM pNPB in acetonitrile
  • 1:4:95 ratio of acetonitrile pNPB solution, 100% ethanol and 50mM Tris-HCL (pH 8)
• LB with specific resistance
• Cell Culture
• 96 well plate

Protocol

• First assign each well of the plate except for outer wells which should be filled with LB to prevent evaporation of medium.
• Fill wells with 95 uL of LB
• Fill wells with 5 uL of cell culture
• In fumehood, add 100 uL of bufer to each well
• Run in Tecan taking both ODs and absorbance 405 readings

Materials

• Pfu turbo
• 10X Pfu turbo buffer
• dNTPs (10mM)
• Forward and reverse primers (0.1ug/uL, see methods section for design tips)
• dH₂0
• Dpn1
• Competent cells

Methods

• Primer Design
• Forward primer should be between 25 and 45 bases in length and contain the desired mutation in the center with correct sequences on both sides; the reverse primer is the reverse complement of this.
• Primers should have a minimum GC content of 40% and terminate in one or more C's or G's.
• Tm should be greater or equal to 78°C and can be calculated as follows:
• Tm = 81.5 + 0.41(%GC) - 675/(length in bases) - %mismatch

• Reaction
• Template DNA: 1 µL
• 10X Buffer: 5 µL
• Forward Primer (100 ng/µL): 1µL
• Reverse Primer (100 ng/µL): 1µL
• 10mM dNTPs: 1µL
• Pfu Turbo (Stratagene): 1µL
• MilliQ H₂0: 40µL
• PCR Program
• 95°C for 1 minute
• 95°C for 50 seconds, 60°C for 50 seconds, 68°C for 1 minute/kb of plasmid length -- repeat this step 17 times, or 18 cycles total
• 68°C for 7 minutes
• 4°C hold
• Following PCR - save 4µL of PCR reaction.
• To remaining 46µLs add 1µL of Dpn1 to PCR reaction (PCR cleanup is not necessary). Incubate at 37°C for 1-2 hours to digest parental DNA. Run 5µL of the digested reaction on a gel and compare to the undigested parental plasmid - there should be some difference in band pattern. Transform into competent cells (PCR cleanup is not necessary).


• Protocol Adapted From Here