SDS PAG ( Top 10 Technique Usefull for Biotech Interview )

    PROTOCOL FOR SDS PAGE.

  • SDS-PAGE, which stands for Sodium Dodecyl Sulfate-Polyacrylamide Gel Electrophoresis, is a widely used laboratory technique in biochemistry and molecular biology. It is used to separate and analyze proteins based on their molecular weight (size) in a gel matrix. SDS-PAGE is a fundamental tool for protein research, protein purification, and quality control.
  • Here’s an overview of how SDS-PAGE works:
  • Denaturation: In SDS-PAGE, proteins are first denatured by heating them in the presence of a detergent called Sodium Dodecyl Sulfate (SDS). SDS disrupts the native protein structures and coats the proteins with a negative charge, giving them a uniform negative charge-to-mass ratio.
  • Sample Preparation: The denatured protein samples are mixed with a sample buffer (commonly Laemmli buffer), which typically contains reducing agents (e.g., β-mercaptoethanol or DTT) to break disulfide bonds and a tracking dye (e.g., bromophenol blue) for visualization.
  • Gel Preparation: SDS-PAGE involves two gels stacked on top of each other: the stacking gel and the resolving gel.
  • Stacking Gel: This gel has a lower acrylamide concentration (typically 4%) and a higher pH. It helps focus the proteins into a tight band before they enter the resolving gel.
  • Resolving Gel: This gel has a higher acrylamide concentration (usually between 8% and 15%) and a lower pH. It separates proteins based on their size (molecular weight).
  • Loading and Electrophoresis: The protein samples, along with a protein size marker, are loaded into wells in the stacking gel. An electric current is then applied across the gel. The negatively charged proteins migrate towards the positive electrode, with smaller proteins moving faster through the gel matrix.
  • Separation: In the resolving gel, proteins separate based on their molecular weight. Smaller proteins travel farther through the gel, while larger proteins are impeded and move more slowly. This separation is due to the sieving effect of the polyacrylamide gel matrix and the uniform negative charge imparted by SDS.
  • Visualization: After electrophoresis, the gel is typically stained with a protein stain (e.g., Coomassie Blue or Silver Stain) to visualize the separated protein bands. The protein bands can be quantified, and their molecular weights estimated using a protein size marker.
  • SDS-PAGE is a versatile technique used for a wide range of applications, including protein characterization, purity assessment, protein quantification, and the verification of protein expression in molecular biology experiments. It is a fundamental step in many research and diagnostic processes in the life sciences

 

Materials:

 

  • Protein samples
  • Running buffer (e.g., 1x Tris-Glycine-SDS buffer)
  • SDS-PAGE gel (pre-made or self-cast)
  • Electrophoresis apparatus (e.g., mini-gel system)
  • Protein molecular weight marker
  • Loading buffer (e.g., Laemmli buffer)
  • Power supply
  • Electrophoresis tank
  • Distilled water
  • Procedure:
  •  
  • Prepare the Gel:
  • There are two kinds of gel usualy one is a
  • Assemble the gel cassette according to the manufacturer’s instructions or your self-cast gel recipe.
  • Prepare the stacking and resolving gels with appropriate acrylamide concentrations, typically between 8% and 15%.
  • Add a few drops of a polymerization initiator (e.g., TEMED) and ammonium persulfate (APS) to initiate gel polymerization. Mix well and pour the gel solution into the cassette.
  • Insert a comb to create wells for sample loading.
  • Allow the gel to polymerize for at least 30 minutes.
  •  
  • Prepare Samples:
  • Mix protein samples with an appropriate volume of loading buffer. The loading buffer should contain SDS and a reducing agent (e.g., DTT or β-mercaptoethanol).
  • Heat the sample mixture at 95°C for 5 minutes to denature the proteins.
  •  
  • Load Samples:
  • Carefully remove the comb from the gel cassette.
  • Rinse the wells with running buffer.
  • Load the protein samples and molecular weight marker into the wells.
  • Load a known volume of molecular weight marker to estimate the protein sizes.
  •  
  • Electrophoresis:
  • Place the gel cassette in the electrophoresis tank.
  • Fill the tank with running buffer until the gel is fully immersed.
  • Connect the electrodes to the power supply and set it to the desired voltage (usually 100-200 V for separating proteins).
  • Run the gel until the tracking dye reaches the bottom (typically 1-2 hours). The proteins will separate based on their size.
  •  
  • Stop Electrophoresis:
  • Turn off the power supply and disconnect the electrodes.
  • Carefully remove the gel cassette from the tank.
  •  
  • Visualize Proteins:
  • Stain the gel with a protein stain such as Coomassie Blue or Silver Stain.
  • Destain the gel if necessary to visualize protein bands clearly.
  •  
  • Analysis:
  • Measure the migration distance (Rf value) of each protein band from the wells.
  • Compare the Rf values to the molecular weight marker bands to estimate the molecular weight of your proteins.

 

  • Calculations:
  • To estimate the molecular weight of your proteins, use the following equation based on the Rf values:
  •  
  • Molecular Weight (kDa) = 10^(a + b x Rf)
  •  
  • Where:
  •  
  • a and b are constants specific to the gel and running conditions.
  • Rf is the migration distance of the protein band relative to the tracking dye.
  • Make sure to adjust the constants (a and b) based on your specific gel and running conditions.

 

 

  • Resolving Gel:
  •  
  • Determine the final volume of your resolving gel. This will depend on your gel cassette size and the thickness of the gel you want to prepare.
  •  
  • Decide on the acrylamide percentage for your resolving gel. For typical protein separation, it usually falls between 8% and 15%. Let’s assume you want to make a 12% resolving gel.
  •  
  • Calculate the volume of acrylamide and bis-acrylamide solutions needed using the desired gel volume and acrylamide percentage. The typical ratio of acrylamide to bis-acrylamide is 29:1, but this can vary depending on your specific protocol.

 

  • Volume of acrylamide solution (mL) = (desired acrylamide percentage / 100) x (final gel volume)
  •  
  • Volume of bis-acrylamide solution (mL) = (volume of acrylamide solution) / 29
  •  
  • Determine the volume of 10x resolving gel buffer to add. A common ratio is 1x resolving gel buffer, which means adding 1/10th of the final gel volume in 10x resolving gel buffer.
  •  
  • Volume of 10x resolving gel buffer (mL) = (final gel volume / 10)
  •  
  • Calculate the volume of water to make up the remaining volume of the gel. This can be done by subtracting the volumes of acrylamide solution, bis-acrylamide solution, and 10x resolving gel buffer from the final gel volume.
  •  
  • Volume of water (mL) = (final gel volume) – (volume of acrylamide solution + volume of bis-acrylamide solution + volume of 10x resolving gel buffer)

 

  • Stacking Gel:

 

  • Determine the final volume of your stacking gel. This will depend on your gel cassette size and the thickness of the gel you want to prepare. Stacking gels are usually thinner than resolving gels.
  •  
  • Decide on the acrylamide percentage for your stacking gel. Stacking gels are typically 4% acrylamide.
  •  
  • Calculate the volume of acrylamide and bis-acrylamide solutions needed using the desired gel volume and acrylamide percentage. Again, you can use the same 29:1 ratio.
  •  
  • Volume of acrylamide solution (mL) = (desired acrylamide percentage / 100) x (final gel volume)
  •  
  • Volume of bis-acrylamide solution (mL) = (volume of acrylamide solution) / 29
  •  
  • Determine the volume of 10x stacking gel buffer to add. A common ratio is 1x stacking gel buffer, which means adding 1/10th of the final gel volume in 10x stacking gel buffer.
  •  
  • Volume of 10x stacking gel buffer (mL) = (final gel volume / 10)
  •  
  • Calculate the volume of water to make up the remaining volume of the gel. This can be done by subtracting the volumes of acrylamide solution, bis-acrylamide solution, and 10x stacking gel buffer from the final gel volume.
  •  
  • Volume of water (mL) = (final gel volume) – (volume of acrylamide solution + volume of bis-acrylamide solution + volume of 10x stacking gel buffer)
  •  
  • Please note that these calculations provide a general guideline. The specific protocol and conditions may vary depending on your equipment and the manufacturer’s recommendations for the acrylamide solutions and buffers you are using. Always follow the in
  •  
  • Prepare the separation gel (10%). Mix in the following order:
· H2O· 4.1 mL
· Acrylamide/bis (30% 37.5:1; Bio-Rad)· 3.3 mL
· Tris–HCl (1.5 M, pH 8.8)· 2.5 mL
· SDS, 10%· 100 µL
· N,N,N′,N′-tetramethylethylene-diamine (TEMED) (Bio-Rad)·  10 µL
· Ammonium persulfate (APS), 10%·  32 µL
  • After adding TEMED and APS to the SDS-PAGE separation gel solution, the gel will polymerize quickly, so add these two reagents when ready to pour.
  • Pour gel, leaving ∼2 cm below the bottom of the comb for the stacking gel. Make sure to remove bubbles.
  • Layer the top of the gel with isopropanol. This will help to remove bubbles at the top of the gel and will also keep the polymerized gel from drying out.
  • In ∼30 min, the gel should be completely polymerized.
  • Remove the isopropanol and wash out the remaining traces of isopropanol with distilled water.
  • Prepare the stacking gel (4%). Mix in the following order:
· H2O· 6.1 mL
· Acrylamide/bis (30%, 37.5:1)· 1.3 mL
· Tris–HCl (0.5 M, pH 6.8)· 2.5 mL
· SDS, 10%· 100 µL
· TEMED·  10 µL
· Ammonium persulfate (APS), 10%· 100 µL
  • Pour stacking gel on top of the separation gel.
  • Add combs to make wells. In ∼30 min, the stacking gel should become completely polymerized.
  • Clamp gel into apparatus, and fill both buffer chambers with gel running buffer according to the instructions for the specific apparatus.
  • Load samples and molecular mass protein m
  •  
  • Preparing SDS Gels
  • A gel of given acrylamide concentration separates proteins effectively within a characteristic range. Very large polypeptides cannot penetrate far into a gel and thus their corresponding bands may be too compressed for resolution. Polypeptides below a particluar size are not restricted at all by the gel, and regardless of mass they all move at the same pace along with the tracking dye. Gel concentration (%T) should be selected so that the proteins of interest are resolved.
  • A typical gel of 7% acrylamide composition nicely separates polypeptides with molecular mass between 45 and 200 kDa. Polypeptides below the cutoff of around 45 kDa do not resolve. A denser gel, say 14%T, usually resolves all of the smallest polypeptides in a mix. Such a gel would be needed to resolve hemoglobin, for example. It would be useless for resolving bands much above 60 kDa, though. To analyze the entire profile of a fraction that contains heavy and light polypeptides, one should usually run two gels.
  • In the teaching lab we recommend that alternate teams prepare low or high percent gels, with each team exchanging samples with a team that prepared the other type gel. Each team, then, would load its set of samples, appropriate standards, and another team’s samples on its gel, and have its samples loaded onto another percent gel as well. In addition to expanding the range of resolution of bands, this practice allows comparison between identical fractions prepared by different teams, to control for inconsistencies in fractionation, sample preparation, etc.
  • Cassettes
  • There are many systems for setting up gel cassettes, some of which are quite expensive. A simple ‘mini-slab’ gel system can be put together for a surprisingly little amount of money and does the job quite well. Our teaching program has done well using projector slide cover glasses (Kodak cat. #140 2130) as cassette plates, with casting stand, running stands, combs and spacers supplied by Sam Lee Custom Crafting, P.O. Box 130973, Houston, Texas 77219, tel.#713-861-4636). The procedure described here employs that system.
  • We use casting stands to prepare the mini-slab gels. Two clean plates with two teflon spacers make a single cassette. We stack the cassettes upright in the stand with the bottoms of the cassettes tight to the bottom of the stand, using modeling clay to seal a thick acrylic cover in place against the last cassette to make a water-tight chamber. Using a well-former (comb) as a template, we mark a fill line about a centimeter below the bottom of the comb for the height of the first (separating) gel solution.
  • Notes on cassette preparation
  • The bevels are not essential, but they aid in the insertion of combs when the stacking solution is poured.
  • Spacers can be straighted with a thin spatula after assembly.
  • The stand must be upright, or else leaks are likely.
  • Air gaps between clay and the front cover will result in leaks.
  • Since acrylamide is toxic, the stand should be placed in a tray or on absorbent paper prior to pouring the gel mix, to confine any leaks.
  • Separating Gel Preparation
  • The total volume between the plates of our gel cassettes is ten ml, so if we prepare 10 ml separating gel mix per cassette we have more than enough. We typically prepare three cassettes per stand and use the best one of the three. From 30% acrylamide stock (see notes below) we prepare gels of composition 7 to 15% acrylamide, depending on the range of proteins that we wish to separate. Our separating gel buffer stock (4x concentrated) consists of 0.4% SDS, 1.5 M Tris-Cl, pH 8.8. Per cassette, we mix 2.5 ml buffer stock and sufficient acrylamide stock so that when the mix is brought to final volume with distilled water we have the desired percent acrylamide monomer.
  • Acrylamide polymerizes spontaneously in the absence of oxygen, so the polymerization process involves complete removal of oxygen from the solution. Polymerization is more uniform if the mix is de-gassed to remove much of the dissolved oxygen, by placing it under a vacuum for 5 minutes or so before polymerization. We initiatiate polymerization by adding freshly prepared10% ammonium persulfate (AP) to the mix followed by N, N, N’, N’-tetramethylethylenediamine (TEMED). The amounts of each depend on the quality of acrylamide used, and should be determined in advance by trial and error. We usually start with 100 µl AP and 10 µl TEMED per 10 ml gel mix, and see how it goes. Once the catalysts are added, polymerization may occur quickly, thus it is necessary to have the casting stand completely ready and to have the overlay solution ready to go (see below). After swirling to mix, we simply pour the solution into the space occupied by the cassettes. The cassettes will self-level eventually, but leveling can be hurried along by adding solution to selected cassettes with a pasteur pipet. Excess solution can be removed by tipping the apparatus and pulling off the excess with a pipet, so that the final level is at the fill mark.
  • Immediately after pouring the gel mix, it must be overlaid with water-saturated butanol to an additional height of 0.5 cm or so (butanol is the top layer in the stock container). Adding butanol to a single cassette will drive the acrylamide mix down, raising the level in the others, so care must be taken to distribute the butanol equally among the cassettes. The purpose of butanol is to produce a smooth, completely level surface on top of the separating gel, so that bands are straight and uniform. Butanol holds very little water in solution, forming a neat layer on top, which is why we use it. Water would make an effective overlay but would mix with the acrylamide solution, diluting it. In fact, the butanol we use is saturated with water so that it does not dry out the gel mix.
  • Polymerization can be confirmed by pulling some of the remaining gel mix into the pipet, allowing it to stand, and checking it after 10 min or so. When the gel mix can no longer be expelled by squeezing the bulb, the separating gel is set. It should not take more than 15 minutes for any of the gel mixes to polymerize. If it hasn’t gelled by that time, something is probably wrong. Often, first time “gel makers” are misled into thinking the gel hasn’t polymerized because the top 0.5 ml or so of the gel mix does not set (some oxygen reaches it through the overlay).
  •  
  • Stacking gel preparation
  • Ten ml of stacking gel mix is sufficient for three of our cassettes, however for the sake of accuracy it may be preferable to make 20 or 30 ml. Excess can be rinsed and tossed into a wastebasket after it polymerizes. It isn’t necessary to degas a stacking mix, because the stacker is simply designed to perform as a matrix through which samples will pass as they are caught up between moving boundaries. It is not designed for uniform separation of proteins. Our stacking gel buffer stock consists of 0.5 M Tris-Cl, pH 6.8, with 0.4% SDS. Typical stackers are 3 to 4.5% acrylamide. We use 4% in order to permit stacking of very large proteins and still retain sufficient mechanical strength to make good sample wells.
  • Before adding the final two components, which will start polymerization, the butanol should be poured off the separating gels into a sink with tap water running and excess butanol/acrylamide removed from the surfaces with a pipet. We use AP and TEMED in similar proportions as for the separating gel mix, although we sometimes increase the amount of one or both components since lower percentage acrylamide solutions tend to polymerize more slowly. After adding AP and TEMED we immediately swirl the mix and pour it into the cassettes to the tops of the plates. We insert combs one at a time, taking care not to catch bubbles under the teeth, and adjust to make them even if necessary, scraping excess stacking mix off later.
  • Notes on gel preparation
  • Acrylamide is a toxic substance so use care and wear gloves while handling solutions that contain it. Use in a well ventilated area, and report any spills. Stock solutions should be kept in a fume hood.
  • An erlenmeyer flask is good for mixing acrylamide, since the narrow neck can be stoppered to prevent toxic fumes from excaping. The wide bottom allows for a large surface area, so that oxygen can be quickly removed from the solution when it is placed under a vacuum.
  • Acrylamide gel stock is labeled according to acrylamide monomer content. Our formulation uses an acrylamide stock of 29.2% acrylamide and 0.8% bis-acrylamide, the cross-linker (cross linking gives the gel its mechanical stability). The stock solution is labeled 30% T (29.2 + 0.8 = 30), 2.5% Cbis (0.8 is 2.5% of 30).
  •  
  • Preparing SDS Gels
  • A gel of given acrylamide concentration separates proteins effectively within a characteristic range. Very large polypeptides cannot penetrate far into a gel and thus their corresponding bands may be too compressed for resolution. Polypeptides below a particluar size are not restricted at all by the gel, and regardless of mass they all move at the same pace along with the tracking dye. Gel concentration (%T) should be selected so that the proteins of interest are resolved.
  • A typical gel of 7% acrylamide composition nicely separates polypeptides with molecular mass between 45 and 200 kDa. Polypeptides below the cutoff of around 45 kDa do not resolve. A denser gel, say 14%T, usually resolves all of the smallest polypeptides in a mix. Such a gel would be needed to resolve hemoglobin, for example. It would be useless for resolving bands much above 60 kDa, though. To analyze the entire profile of a fraction that contains heavy and light polypeptides, one should usually run two gels.
  • In the teaching lab we recommend that alternate teams prepare low or high percent gels, with each team exchanging samples with a team that prepared the other type gel. Each team, then, would load its set of samples, appropriate standards, and another team’s samples on its gel, and have its samples loaded onto another percent gel as well. In addition to expanding the range of resolution of bands, this practice allows comparison between identical fractions prepared by different teams, to control for inconsistencies in fractionation, sample preparation, etc.
  • Cassettes
  • There are many systems for setting up gel cassettes, some of which are quite expensive. A simple ‘mini-slab’ gel system can be put together for a surprisingly little amount of money and does the job quite well. Our teaching program has done well using projector slide cover glasses (Kodak cat. #140 2130) as cassette plates, with casting stand, running stands, combs and spacers supplied by Sam Lee Custom Crafting, P.O. Box 130973, Houston, Texas 77219, tel.#713-861-4636). The procedure described here employs that system.
  • We use casting stands to prepare the mini-slab gels. Two clean plates with two teflon spacers make a single cassette. We stack the cassettes upright in the stand with the bottoms of the cassettes tight to the bottom of the stand, using modeling clay to seal a thick acrylic cover in place against the last cassette to make a water-tight chamber. Using a well-former (comb) as a template, we mark a fill line about a centimeter below the bottom of the comb for the height of the first (separating) gel solution.
  • Notes on cassette preparation
  • The bevels are not essential, but they aid in the insertion of combs when the stacking solution is poured.
  • Spacers can be straighted with a thin spatula after assembly.
  • The stand must be upright, or else leaks are likely.
  • Air gaps between clay and the front cover will result in leaks.
  • Since acrylamide is toxic, the stand should be placed in a tray or on absorbent paper prior to pouring the gel mix, to confine any leaks.
  • Separating Gel Preparation
  • The total volume between the plates of our gel cassettes is ten ml, so if we prepare 10 ml separating gel mix per cassette we have more than enough. We typically prepare three cassettes per stand and use the best one of the three. From 30% acrylamide stock (see notes below) we prepare gels of composition 7 to 15% acrylamide, depending on the range of proteins that we wish to separate. Our separating gel buffer stock (4x concentrated) consists of 0.4% SDS, 1.5 M Tris-Cl, pH 8.8. Per cassette, we mix 2.5 ml buffer stock and sufficient acrylamide stock so that when the mix is brought to final volume with distilled water we have the desired percent acrylamide monomer.
  • Acrylamide polymerizes spontaneously in the absence of oxygen, so the polymerization process involves complete removal of oxygen from the solution. Polymerization is more uniform if the mix is de-gassed to remove much of the dissolved oxygen, by placing it under a vacuum for 5 minutes or so before polymerization. We initiatiate polymerization by adding freshly prepared10% ammonium persulfate (AP) to the mix followed by N, N, N’, N’-tetramethylethylenediamine (TEMED). The amounts of each depend on the quality of acrylamide used, and should be determined in advance by trial and error. We usually start with 100 µl AP and 10 µl TEMED per 10 ml gel mix, and see how it goes. Once the catalysts are added, polymerization may occur quickly, thus it is necessary to have the casting stand completely ready and to have the overlay solution ready to go (see below). After swirling to mix, we simply pour the solution into the space occupied by the cassettes. The cassettes will self-level eventually, but leveling can be hurried along by adding solution to selected cassettes with a pasteur pipet. Excess solution can be removed by tipping the apparatus and pulling off the excess with a pipet, so that the final level is at the fill mark.
  • Immediately after pouring the gel mix, it must be overlaid with water-saturated butanol to an additional height of 0.5 cm or so (butanol is the top layer in the stock container). Adding butanol to a single cassette will drive the acrylamide mix down, raising the level in the others, so care must be taken to distribute the butanol equally among the cassettes. The purpose of butanol is to produce a smooth, completely level surface on top of the separating gel, so that bands are straight and uniform. Butanol holds very little water in solution, forming a neat layer on top, which is why we use it. Water would make an effective overlay but would mix with the acrylamide solution, diluting it. In fact, the butanol we use is saturated with water so that it does not dry out the gel mix.
  • Polymerization can be confirmed by pulling some of the remaining gel mix into the pipet, allowing it to stand, and checking it after 10 min or so. When the gel mix can no longer be expelled by squeezing the bulb, the separating gel is set. It should not take more than 15 minutes for any of the gel mixes to polymerize. If it hasn’t gelled by that time, something is probably wrong. Often, first time “gel makers” are misled into thinking the gel hasn’t polymerized because the top 0.5 ml or so of the gel mix does not set (some oxygen reaches it through the overlay).

 

  •  

 

 

  • Afer mking gel you have to make a protein sample to load onto the gel
  • You cant load protein directly onto the gel you have to treat  protein sample with the sample buffer the sample buffer contains the following.
  • The following calculation is for 5x
  • 10% SDS,
  • 500mM DTT,
  • 50% Glycerol
  • , 250mM Tris-HCL
  • 5% bromophenol blue dye, PH6.8.
  • So how will you make the above buffer.
  • First you decide for what volume you are going to make the above buffer
  • Suppose you are going to make above buffer for the 50 ml
  • The you will the component following order.
    10% SDS in 50ml it will come 5gm
  • 500milimolar , so molecular weight of DTT is 154.2
  • So for 50ml you will calculate following manner
  • 2* 500/1000* 50/1000
  • So why we are deviding by 1000 for milimolar and 50ml we are converting them into the molar amd liter because our definition is in molar and liter.  This
  • Same for the tris.
  • Sample preparartion.
  • For sample preparation take one  0.5 ml eppendorf tube add 50ul of protein sample and 10ul of the above sample buffer and heat for 10 minute at 95 C degree temprature.
  • After making a sample , set the gel plate in running buffer tank of the SDS PAGE,
  • Pour the required amount of running buffer in the tank , plug in the electrode and run the gel start your run at 100–120 milliamps (mA) and keep an eye on it. After reaching the dyefront to the end of gel stop the gel run.Carfully take out the gel and keep in 20ml  coomasie staining solution. Composition of staining solution includes 40% Methanol 10% glacial acetic acid , and 50% Water. gel will stain in about 20 minute after staining of the gel  , replace thg gel with destaining solution , whicn includde same content without coomasie staining poweder. Keep gel with destaining till protein bands apears.
  •  
  • How to make one liter 1x running buffer.
  • Dissolve 3.00 g of Tris base, 14.40 g of glycine, and 1.00 g of SDS in 1000 ml of H2O. The pH of the buffer should be 8.3 and no pH adjustment is required. Store the running buffer at room temperature.

 

SDS PAGE GEL BANDS

 

 

 

 

 

 

 

 

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