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Protein Purification

Transformation of plasmid DNA into competent E. coli cells

Material and Reagents

  • LB

  • 1.5 mL microfuge tubes

  • 42° C dry-bath

  • Ice

  • 37° C shaker

 

Protocol

 

    1. Thaw competent cells on ice. 20–200µL per tube

    2. Add max.4µL of pure plasmid DNA or of Infusion ligation reaction

    3. Mix very gently (tap on tube walls)!

    4. Incubate the tubes on ice for 30 min

    5. Heat shock the cells for 90 sec at 42°C

    6. Place the tubes immediately on ice for at least 2 min

    7. Add 900µL of LB medium to each tube

    8. Incubate for 1 hour at 37°C and shake vigorously (225 RPM)

    9. Spin down briefly (4100 rpm for 10 minutes) and remove most (~850uL) of the supernatant

    10. Resuspend cell pellet with the rest LB medium in the tube by pipetting up and down

    11. Streak the suspension on a LB agar plate (containing the appropriate antibiotic as the attached picture shows.

    12. After streaking, incubate the plates overnight at 37°C

Protein expression & purification (general) 

Protein Purification (introduction)

Strategies of Protein Purification

Principle of affinity chromatography

Principle of ion-exchange chromatography

Principle of gel filtration chromatography

Principle of

hydrophobic interaction chromatography

'How-to' videos from GE Healthcare

Solution (or 'buffer') preparation from solid

Solution (or 'buffer') preparation from stock

Phosphate buffer recipe

Cleaning Ni resin

Regeneration by stripping and recharging:


Stripping and recharging of Ni-NTA Superflow Cartridges is usually not
necessary. If an increase in back pressure or significant contamination of the
resin is observed, a cleaning-in-place procedure usually restores performance.
However, if performance is still not satisfactory, the Ni-NTA resin in the
cartridge can be stripped and recharged using the protocol below.


Procedure
1. Strip the resin by washing with 10 column volumes of stripping
buffer (50 mM Na phosphate; 300 mM NaCl; 100 mM EDTA;
pH 8.0).
2. Wash the resin with 20 column volumes of deionized water.
3. Recharge the water-washed cartridge by loading 2 column volumes
of 100 mM NiSO4 (in deionized water). Salts of other metals
(chlorides or sulfates) may also be used.
4. Wash with 10 column volumes of deionized water and re-equilibrate
with 10 column volumes of Buffer NPI-10.
The cartridge is now ready for use. Store cartridge in 20–30% ethanol or
10–100 mM NaOH.

SDS-PAGE introduction

SDS-PAGE Gel Recipe

Bradford assay in 97 wells

The bradford dye-binding assay is a colorimetric assay for measuring total protein concentration. It involves the binding of Coomassie Brilliant blue to protein.

 

There is no interference from cations nor from carbohydrates such as sucrose. However, detergents such as sodium dodecyl sulfate and triton x-100 can interfere with the assay, as well as strongly alkaline solutions.

 

General Overview of Procedure:

  1. Dilute 5x Bradford Reagent (BioRad) 4:1 with water and run through a gravity filter.

  2. Get a 96-well ELISA plate (Limbro microtitration plate).

  3. Turn on machine to allow bulb to warm up (approx. 10 min. before use).

  4. Prepare standards.

  5. Plate standards in triplicate (10 uL per well). Leave column 1 blank, begin plating in column 2.

  6. Plate samples in triplicate (10 uL per well). Since your standard curve is from 0-1 mg/mL (for BSA), your samples should be in this range. Taking an O. D. reading on your sample and generalizing that 1 O. D.=1 mg/mL is a good way to get a ballpark value. Dilute your samples appropriately, and if you don't know the concentration of protein make several samples of various dilutions.

  7. Add 200 uL of diluted Bradford reagent to each well, and let stand 5 minutes.

  8. Insert the 595 nm. filter into the machine. Set the plate on the reader. Press "blank" and after the machine gives a printout of the blank reading, press "start". The reading is complete when the plate slides back out from underneath the reader.

  9. Use the results to graph the standard curve (Axes are commonly labeled as y=A, 600 nm and x=mg/mL). Use the curve and data from bradford to determine unknown protein concentration.

 

Preparation of Standard

A set of standards is created from a stock of protein whose concentration is known. The Bradford values obtained for the standard are then used to construct a standard curve to which the unknown values obtained can be compared to determine their concentration. Use a protein as your standard that most closely resembles the protein you are assaying. BSA and IgG are typical standards used to construct the curve. For BSA, use 0-1 mg/mL as your standard curve concentration; for IgG, use 0-1.6 mg/mL.

Example of typical standards created from BSA stock (1 mg/mL), to give a standard curve from 0-1 mg/mL:

Sample #1 (0.0 mg/mL): 0 uL BSA + 30 uL buffer.

Sample #2 (0.2 mg/mL): 6 uL BSA + 24 uL buffer.

Sample #3 (0.4 mg/mL): 12 uL BSA + 18 uL buffer.

Sample #4 (0.6 mg/mL): 18 uL BSA + 12 uL buffer.

Sample #5 (0.8 mg/mL): 24 uL BSA + 6 uL buffer.

Sample #6 (1.0 mg/mL): 30 uL BSA + 0 uL buffeer.

*Note: The ubiquitously used BSA is a very poor standard for the bradford assay. To adjust for this, multiply the results you get for your protein concentration by 2.1 to get a closer approximation of your protein's concentration. Lysozyme, ovalbumin and catalase make much better standards, and no adjustment is necessary for these standards.

Bradford Protein Concentration Assay (in 1ml volume)

Abbreviations:

 

  • mcg = micrograms

  • mcL = microliters

  • BSA = bovine serum albumin

  • O.D. = optical density

  • dI = deionized

 

Background

 

The Bradford protein assay (1) is one of several simple methods commonly used to determine the total protein concentration of a sample. The method is based on the proportional binding of the dye Coomassie to proteins. Within the linear range of the assay (~5-25 mcg/mL), the more protein present, the more Coomassie binds. Furthermore, the assay is colorimetric; as the protein concentration increases, the color of the test sample becomes darker. Coomassie absorbs at 595 nm. The protein concentration of a test sample is determined by comparison to that of a series of protein standards known to reproducibly exhibit a linear absorbance profile in this assay. Although different protein standards can be used, we have chosen the most widely used protein as our standard - Bovine Serum Albumin (BSA).

 

Procedure

 

  • Prepare a 4-fold dilution of a 2 mg/mL BSA sample by adding 50 mcL of 2 mg/mL BSA to 150 mcL of dI water to make 200 mcL of 0.5 mg/mL BSA.

  • Generate test samples for the reference cell, blank, BSA standards and the protein sample to be tested according to Table 1 in disposable cuvettes.

  • Note that the "reference cell" and "blank" are identical. A reference cell test sample is only required when using a double-beam UV-visible spectrophotometer for absorbance measurements.

  • Note that a dilution of the protein sample may be required for the resulting absorbance to fall within the linear range of the assay.

  • Allow each sample to incubate at room temperature for 10-30 minutes. (Record the actual incubation time in your notebook.)

  • Measure the absorbance of each sample at 595 nm using a UV-visible spectrophotometer. Be sure to allow the instrument to warm up for at least 15 minutes prior to use.

  • Plot the absorbance of each BSA standard as a function of its theoretical concentration. The plot should be linear.

  • Determine the best fit of the data to a straight line in the form of the equation "y = mx + b" where y = absorbance at 595 nm and x = protein concentration.

  • Use this equation to calculate the concentration of the protein sample based on the measured absorbance. Note: If the absorbance of the test sample is outside of the absorbance range for the standards, then the assay must be repeated with a more appropriate dilution, if any. The linear range for the assay (and for most spectrophotometers is 0.2 - 0.8 O.D. units.

 

Table 1. Preparation of test samples for the Bradford protein assay.

​

Making a long term stock of bacteria

Pick a single colony of the clone off a selective plate and grow an overnight culture in the appropriate selectable liquid medium (3-5ml).

  1. Enter all the relevant information (e.g. vector, strain, date, researcher, etc.) in the online Parashar lab inventory inventory at Quartzy.com. This will also be a good time to record the strain information and record the location. Label strain numbers as VPXXXa and VPXXXb where XXX are laboratory's strain numbers.

  2. Take two cryogenic vials and label them with the XXXa and XXXb internal strain numbers.

  3. Add 333 micro liter of 60% glycerol in H2O to each cryogenic vial.

  4. To each cryogenic vial, add 666 micro liter sample from the culture of bacteria to be stored.

  5. Gently vortex the cryogenic vial to ensure the culture and glycerol is well-mixed.

  6. Put in a box corresponding to sdata entry in inventory  and store it at -80C freezer.

More info at:

http://openwetware.org/wiki/Making_a_long_term_stock_of_bacteria

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