TITLE: Assessing protein purity using SDS PAGE
AUTHORS: Shalo  Minette, Stephane  Fadanka, Nadine             Mowoh

[DESCRIPTION]
SDS-PAGE (Sodium Dodecyl Sulphate polyacrylamide gel electrophoresis) is an electrophoresis method that allows protein separation by mass. The gel acts as a sieve through which the proteins move in response to an electric field.
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SDS-PAGE allows an estimation of the purity of protein samples and can also be used in confirming the presence of a particular protein in a sample through its molecular weight by running it simultaneously on the gel with another protein of known molecular weight, called the standard or protein ladder. 

SDS is an anionic detergent and is used to denature the proteins. The negative charges on SDS destroy most of the secondary and tertiary structure of proteins and are strongly attracted toward the anode in an electric field. Because the charge-to-mass ratio is nearly the same among SDS-denatured proteins, the final separation of proteins is almost entirely dependent on the differences in relative molecular weight (MW) of polypeptides.



[BEFORE_START]
Ensure all materials to be used for this procedure are all ready and functional

[GUIDELINES]
This protocol can be used to assess the protein purity by assessing presence of target vs contaminating bands on SDS-PAGE.
In our lab we typically use this protocol to confirm successful protein expression in preparing cellular reagents.
In this protocol we use pre-cast TruPAGE gels

[STEPS]
SECTION: Preparing reagent working stocks
1. Running Buffer Preparation
 

For 1 L of 1x Tris-MOPS buffer

Measure out 50 mL of 20X Tris-MOPS buffer into a 1L capacity beaker and add 950 mL of distilled water.

TruPAGE Tris-MOPS SDS Express Running buffers are supplied as 20X concentrated solutions. For best results, it is recommended to prepare a fresh 1X buffer for every run.

SECTION: Preparing reagent working stocks
2. Staining reagent preparation (Coomassie Brilliant Blue G-250)

Dissolve80 mgof CBB G-250 in 1 L of bi-distilled water by stirring for 2-4 hours. 
Add3 mL of concentrated HCl to the dark blue solution with stirring for another minute and store in the dark for later use. 
NB: The solution can be stored for weeks up to several months without losing its staining efficiency.
 

SECTION: Preparing reagent working stocks
3. DOC-Lysis Buffer
For 100ml of the buffer;
 
 

 
Measure out the powders into a 250 mL capacity glass beaker as shown in the table above.
Add the buffers as indicated and finally add 96 mL of distilled water to completely dissolve the powder.
The lysis buffer can be stored at 4 °C and is stable for up to 6 months.

SECTION: Sample Preparation/Denaturation
4. Preparing whole cell samples

After protein expression with IPTG or in autoinduction media, read the optical density at 600 nm (OD600 ) of the cell culture using a photometer.

 
Calculate the amount of culture to harvest to have a cell pellet equivalent to 1mL of OD600 =1 using the formula:
Ci * Vi = Cf * Vf
 
Transfer the amount of culture as calculated above in an appropriate tube and centrifuge it at 5,000 xg for 10 minutes.
  
Discard the supernatant and resuspend the pellet in Doc lysis buffer (1mL per 0.1g of the pellet).
Vortex intensively for 2 min at room temperature.
Leave it shaking on a rotor at room temperature for 15mins.
Then pellet the cells by centrifugation at 13000 rpm for 5 mins and carefully collect the supernatant which is our cell lysate for SDS PAGE analysis.

SECTION: Sample Preparation/Denaturation
5. Preparing the protein samples

Pre-heat water bath to 95ºC. 
Determine the concentration of the protein sample via Bradford assay. 
Dilute or concentrate samples as needed to yield a final protein concentration of >0.5 mg/mL for purified proteins and <20mg/mL for cell lysates. Where this is not applicable, proceed with the next step as indicated.
Proceed to mix sample and TruPAGE LDS Sample Buffer as demonstrated in the table below. 
Samples should be prepared just prior to analysis.
 
So, dilute  your sample to an initial concentration of 25 ug/uL before preparation with a sample buffer.
buffer using this protocol.

 
    S/NReagentsAmounts (ul)Ex. 50ul reaction 1Protein Samplex3024x sample buffer12.512.53DTT (1M)3.53.54Sterilised distilled water34-x45Total volume5050
Place the samples in a water bath at 95ºC for 5 mins.
 
 
SECTION: Setting up the run
6. Setting up the apparatus

Rinse the gel wells with sterile distilled water. TruPAGE wells MUST be rinsed prior to loading samples for removal of trace storage buffer to achieve optimal band resolution and sharpness.
Remember to peel off the white tape at the bottom of the gel cassette prior to loading samples.
Place Precast gel into apparatus and secure as appropriate;
Fill the cathode chamber with 1X running buffer and ensure a proper seal prior to filling the anode chamber.

SECTION: Setting up the run
7. Loading the wells

Load the samples into the wells. Do not exceed well capacity when loading samples, 35 uL for 12 well gels and 20 uL for 10 well gels (For TruPAGE gels).
Note: Use thin gel-loading tips for better settling of the samples towards the bottom of the wells.
Load 10 µl of MW marker in the left most lane.
Load 10-30 µl of samples depending on well capacity and purity of the protein in subsequent wells. Load less for purified proteins (10 uL) and more for cell lysates (20 uL). 

SECTION: Setting up the run
8. Running the Gel

Once the samples are loaded and buffer chambers are filled, cover the chamber and firmly connect both the anode and the cathode. 
Set the voltage on the electrophoresis power supply to a constant voltage of 180 V. Turn ON the power supply.
Run the gel at constant voltage (180 V) until the blue dye front reaches the bottom of the gel cassette.

 
 

SECTION: Staining and de-staining the Gel
9. 

 

Prepare Coomassie Brilliant Blue G-250 stain solution as directed above.
After running the gel, remove the gel from the cassette. This can be tricky, use a spatula to pry open the cassette carefully not to break the gel.
Heat 100 mL of sterilized autoclaved water for 1 minute on high (600 W).
Wash the gel with warm ultrapure water for 3 to 5 minutes.
Repeat warm water wash thrice.

 

Heat 25 mL of stain solution for 20 seconds on high (600 W). Place the gel in the warm stain solution and incubate for 30 minutes with gentle agitation.  
Pour off the staining solution and add 50-100ml bi distilled water in order to de-stain the light blue background on the gel. Place on a shaker and replace water if necessary for further de-staining.
If the gel is stained for more than 30mins, warm 100 mL of de-stain solution (10% ethanol) for 30 seconds on high (600 W). Remove gel from stain solution and de-stain for 15 minutes to 16 hours. The duration of de-staining depends on the gel type used.
Document the gel (take a clear picture).