Shreffler: SDS-PAGE

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Current revision (16:00, 22 March 2012) (view source)
(Running Gel)
 
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===Running Gel===
===Running Gel===
-
#Make 1X  
+
#Make MES SDS Running Buffer 1X using 20X stock (~one liter should be more than enough).
 +
#Set aside 200 ml of Running Buffer and add 0.5 ml Antioxidant.
#Remove precast 4-12% Bis-Tris gels from packages, carefully and over a sink (pouch contains liquid). Pull off white stripe at bottom and carefully remove plastic comb from gel. Gently wash cassette and (esp. wells) under DI water (can apply gentle centrifugal force to eject residual liquid from wells so that they can be washed).
#Remove precast 4-12% Bis-Tris gels from packages, carefully and over a sink (pouch contains liquid). Pull off white stripe at bottom and carefully remove plastic comb from gel. Gently wash cassette and (esp. wells) under DI water (can apply gentle centrifugal force to eject residual liquid from wells so that they can be washed).
 +
#Place gel cassette on tube rack (stabilize vertically) and fill wells with standard (5 µl)  and samples of interest - can run varying volumes of loading solution to vary protein quantity (load 2 µl for 1 µg protein, 10 µl for 5 µg protein).
#Lower buffer core into cell and insert gel cassettes on both sides, such that the shorter well side faces inward (if only running one gel, use a dummy cassette). Insert tension wedge and lock in place.
#Lower buffer core into cell and insert gel cassettes on both sides, such that the shorter well side faces inward (if only running one gel, use a dummy cassette). Insert tension wedge and lock in place.
-
#
+
#Fill buffer core with the 200 ml of Running Buffer + Antioxidant. Pour slowly and make sure that core does NOT leak into outer cell.
-
 
+
#Fill outer space (between cell and core) with Running Buffer (no Antioxidant) until level is sufficient (~600 ml)
-
Fill the buffer core with 200 mL 1X RB. Make sure this section does NOT leak before adding liquid to the outside. Carefully remove the comb and pipette out any air bubbles. Then, fill outer part of case with 600 mL of RB.
+
#Match electrode ends and turn voltage source on. Set to 100V to begin with, but can increase to 150V when gel is halfway done running (gel has density gradient, so bands will run slower the further down they are).
-
Fill the wells with samples:
+
#Stop running the gel when blue indicator bands are almost at bottom.
-
Add 10 μL MW markers to well #1.
+
#Remove gel cassette from gel box, and carefully crack edges open using metal spatula (knife works too). Start at corners, inserting tool and twisting until separation occurs, then work way down sides. Do not insert tool deep enough to touch gel, or else gel will rip when separating cassette sides.
-
Fill remaining 11 wells with 10 μL of prepared protein sample solution.
+
#Fill deep tray with DI water and open gel cassette while submerged under water. Gel should be stuck on one side of the cassette. Carefully trim off bottom blue bands, and the wells on top. Then, carefully remove gel.  
-
Fill each well slowly. Try to avoid bubbles in wells that may cause overflow into other wells
+
#Place gel in pipette box cover with DI water and let shake at very low speed. Replace DI water every five minutes for fifteen minutes of washing.
-
When working with 2 gel plates, make sure to label the outside of the container.
+
#Pour off DI water completely and stain gel with 15 ml of SimplyBlue SafeStain. Set on shaker (very low speed) for one hour.
-
Match the + and -electrode ends (same colors)
+
#Pour off blue stain completely and wash gel with DI water, shaking at very low speed for two hours. Replace DI water every 20 minutes or so.
-
Turn on (be sure Range Select is OFF). Set Range to 200V/400mA, then hit Start.
+
#Carefully place gel between transparent folder sheet. Place against white paper backdrop and scan or photograph as desired.
-
Run for 35 min. When done, then turn off.
+
-
Gel stain:
+
-
Take gel cassette out of Invitrogen gel box.
+
-
Fill microwave safe glass container with MilliQ water enough as to immerse gel.
+
-
Crack open gel cassette and place gel in microwave safe glass container.
+
-
Microwave gel for two minutes or until water boils.
+
-
Place container on shaker until cool. Then decant the water. Rinse one more time with MilliQ water.
+
-
Add Denville Blue Protein Stain to gel container and immerse gel.  
+
-
Microwave gel for two minutes or until protein stain solution boils and place on shaker until cool.
+
-
Decant gel stain into waste container, and rinse gel with MilliQ water.
+
-
Add MilliQ water to immerse gel and microwave for two minutes and then shake until cool.
+
-
Carefully place gel into plastic gel box filled with water.
+
==Discussion==
==Discussion==

Current revision

Contents

Overview

From wikipedia: "SDS-PAGE, sodium dodecyl sulfate polyacrylamide gel electrophoresis, describes a technique widely used in biochemistry, forensics, genetics and molecular biology to separate proteins according to their electrophoretic mobility (a function of the length of a polypeptide chain and its charge). In most proteins, the binding of SDS to the polypeptide chain imparts an even distribution of charge per unit mass, thereby resulting in a fractionation by approximate size during electrophoresis."

We use the Invitrogen NuPAGE system.

Materials

  • Appropriate gelbox/voltage source
  • NuPAGE 4-12% BIS-TRIS Gel (Invitrogen NP0322BOX)
  • NuPAGE MES SDS Running Buffer (20x) (Invitrogen NP0002)
  • NuPAGE SimplyBlue SafeStain (Invitrogen LC6060)
  • NuPAGE LDS Sample Buffer (4X) (Invitrogen NP0007)
  • NuPAGE Antioxidant (Invitrogen NP0005)
  • NuPAGE Reducing Agent (Invitrogen NP0004)
  • Novex Sharp Pre-Stained Protein Standard (Invitrogen 57318)

Procedure

Preparing Samples

  1. Make a 20 µl loading solution for each protein consisting of:
    • 5 µL LDS Sample Buffer (4X)
    • 2 µL Reducing Agent
    • Volume containing 10 µg protein (e.g. if protein stock solution is 1 µg/µl, add 10 µl)
    • Bring up to 20 µl total with PBS.
  2. Heat samples on heating block (or water bath) 70°C for 10 mins (protein denaturing and SDS binding).

Running Gel

  1. Make MES SDS Running Buffer 1X using 20X stock (~one liter should be more than enough).
  2. Set aside 200 ml of Running Buffer and add 0.5 ml Antioxidant.
  3. Remove precast 4-12% Bis-Tris gels from packages, carefully and over a sink (pouch contains liquid). Pull off white stripe at bottom and carefully remove plastic comb from gel. Gently wash cassette and (esp. wells) under DI water (can apply gentle centrifugal force to eject residual liquid from wells so that they can be washed).
  4. Place gel cassette on tube rack (stabilize vertically) and fill wells with standard (5 µl) and samples of interest - can run varying volumes of loading solution to vary protein quantity (load 2 µl for 1 µg protein, 10 µl for 5 µg protein).
  5. Lower buffer core into cell and insert gel cassettes on both sides, such that the shorter well side faces inward (if only running one gel, use a dummy cassette). Insert tension wedge and lock in place.
  6. Fill buffer core with the 200 ml of Running Buffer + Antioxidant. Pour slowly and make sure that core does NOT leak into outer cell.
  7. Fill outer space (between cell and core) with Running Buffer (no Antioxidant) until level is sufficient (~600 ml)
  8. Match electrode ends and turn voltage source on. Set to 100V to begin with, but can increase to 150V when gel is halfway done running (gel has density gradient, so bands will run slower the further down they are).
  9. Stop running the gel when blue indicator bands are almost at bottom.
  10. Remove gel cassette from gel box, and carefully crack edges open using metal spatula (knife works too). Start at corners, inserting tool and twisting until separation occurs, then work way down sides. Do not insert tool deep enough to touch gel, or else gel will rip when separating cassette sides.
  11. Fill deep tray with DI water and open gel cassette while submerged under water. Gel should be stuck on one side of the cassette. Carefully trim off bottom blue bands, and the wells on top. Then, carefully remove gel.
  12. Place gel in pipette box cover with DI water and let shake at very low speed. Replace DI water every five minutes for fifteen minutes of washing.
  13. Pour off DI water completely and stain gel with 15 ml of SimplyBlue SafeStain. Set on shaker (very low speed) for one hour.
  14. Pour off blue stain completely and wash gel with DI water, shaking at very low speed for two hours. Replace DI water every 20 minutes or so.
  15. Carefully place gel between transparent folder sheet. Place against white paper backdrop and scan or photograph as desired.

Discussion

discuss this protocol

Contact

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