SDS-PAGE Method

This page is part of the SDS-PAGE lab, which includes these pages:

The whole experiment will be spread over three lab days. You will also use these methods as part of the pGLO lab.

This page will show to set up and run an SDS-PAGE gel. The procedure for preparing and running the gel is the same for both of the SDS-PAGE labs you'll do this quarter, but the samples and the amounts you load on the gel will be different.

Winter 2020:

We have two different kinds of protein gels to test for this lab:

  • NuPAGE 4-12% Bis-Tris Gel. These gels are for SDS-PAGE, the most commonly used protein electrophoresis technique. In this technique, the proteins will be completely denatured, so they will be separated on the gel only on the basis of molecular mass. Fluorescent proteins such as GFP won't be fluorescent on this type of gel, because they're denatured.
  • Novex Wedge Well 10-20% Tris-Glycine. These gels are for native PAGE. This technique doesn’t use SDS or a reducing agent, so the proteins are in their native conformation (not denatured). In principle, that means that your GFP will still be fluorescent, so the fluorescence may be visible on the gel. Of course, that depends on how much GFP is present in your sample. We haven't used these gels in Bio 6B before.

I don't know which gel is going to work best, so some groups should use the native gel and others can use the denaturing gel. You should have enough of each sample for two wells, so if you work with another group you can load the samples from both groups onto one native gel and one denaturing gel.

Each gel uses specific sample and running buffers, so play close attention to which ones you use. Also, the native gels can hold a larger sample volume. Both types of gels have 10 wells.

Gel Condition  Sample buffer
Reducing Agent? Running Buffer
Sample Volume Heat Sample
Novex Wedge Well 10-20% Tris-Glycine  Native  Native Tris-Glycine Sample Buffer (2x) no Tris-Glycine Native Running Buffer (10x) 40 μl no
NuPAGE 4-12% Bis-Tris Gel Denaturing  NuPAGE LDS Sample Buffer (4x) yes MOPS SDS Running Buffer (20x) 25 μl yes

Pay attention to which gel and procedure you're using, because some parts of the procedure are different. Once you get your gel, make sure you find the appropriate sample buffer and running buffer.

First prepare your gel-ready samples.

Before you do the steps on this page, you should get your protein samples ready to run on the gel. Do that first and then set up the gel apparatus while you're waiting for the samples to denature.

Your protein samples are the lysates you prepared in the HIC lab. In that procedure, you lysed your cells using lysozyme to weaken the cell walls and freezing to burst open the cells. After centrifugation to pellet the insoluble debris, the soluble liquid is your lysate.

SDS-PAGE Sample preparation

To make your protein sample ready for SDS-PAGE, mix it as follows:

Volume Ingredient
65 μl  Protein Sample
25 μl NuPAGE LDS Sample Buffer (4x)
10 μl Sample Reducing Agent
100 μl Total

Use a 500-μl micro tube to reduce evaporation. Be sure that the ingredients are fully mixed (the purple color should be evenly distributed) and be careful not to put bubbles into it (the SDS detergent tends to get frothy). After you prepare the gel-ready samples, heat them at 70° for 10 minutes before loading on the gel. See the Electrophoresis page more detail on what these ingredients do.

These ratios will be the same for all the SDS-PAGE samples you do this quarter. The final concentration of sample buffer will be 1x, and the sample reducing agent will be 1/10 of the total. If your protein sample is less than 65 μl, use what you have and keep the proportions the same. Keep in mind that the maximum you can fit in one well of the gel is about 25 μl (or a little more). You shouldn't load less than about 8 μl, because the sample won't spread out and fill the whole lane. Highly concentrated samples should be diluted so the loading volume is between 10 μl and 25 μl.

Native PAGE Sample preparation

For non-denaturing electrophoresis, the procedure is different: a different sample bufffer, no reducing agent, an no heating.

Volume Ingredient
50 μl  Protein Sample
50 μl Native Tris-Glycine Sample Buffer (2x)
100 μl Total

The ratios of ingredients are important. If you are preparing a smaller or larger amount of sample, keep the ratios of the ingredients the same.

Assemble the gel unit

This procedure is for SDS-PAGE; if you are doing native PAGE, you'll use a different gel and running buffer.

Before you start assembling the gel unit, prepare your gel-ready samples as described above. While the samples are heating, you can get the gel unit ready.

We'll use the X-Cell Surelock Mini-cell electrophoresis system from Invitrogen. Other manufacturers make similar gel units. To set up your gel, you'll need to get all the pieces of the gel apparatus, as shown below:

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In order for electrophoresis to work, you must create a conducting path for electricity to go through the gel; the steps below are designed to accomplish that. Follow them carefully.

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We'll be using precast polyacrylamide gels (NuPAGE® Novex® 4-12% Bis-Tris gels from Invitrogen). The gel comes sealed in plastic with a little buffer solution to keep it wet. Cut open the bag and pour the extra buffer down the sink.

Rinsing the gel with deionized water

The thin polyacrylamide gel is sandwiched between two plastic plates. The whole thing is sometimes called the gel cassette. Rinse the outside of the cassette with a little deionized water.

Drying the plates

Dry the plastic plates carefully. If they're wet when you assemble the gel unit, it's likely to leak and you'll have to start over.

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Peel off the tape at the bottom of the gel cassette. If you forget to do this, your gel won't run at all! The tape covers a thin slit in the outer plastic plate. In order for your gel to run, electricity must be conducted through the gel, and this slit is the only way for the electricity to pass through.

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Carefully remove the comb. The gel is made by pouring liquid acrylamide between the plates and letting it polymerize to form polyacrylamide. The comb forms the wells into which you'll put your samples. Be careful! Between the teeth of the comb are delicate fingers of polyacrylamide to separate the wells of the gel; if you pull the comb out carelessly, you might mess up these fingers and make it impossible to get your protein samples into the wells.

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Create the inner buffer chamber by assembling the inner buffer core, the gel cassette, and the buffer dam as shown. The short plate of the gel cassette faces in toward the inner core.

The buffer dam and the plastic plates of the gel cassette must form a tight seal with the silicone gasket of the inner core; make sure all parts are clean and dry before you start.

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Line up all three pieces so they're level on the bottom.

There's a thin, delicate electrode wire on the outside bottom of the inner buffer core, and another on the inside. You're going to assemble the unit to create a conducting path for electricity from one electrode to the other, through the running buffer and the gel.

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This is how the inner buffer chamber goes into the outer buffer chamber. Note that there is a banana plug (electrical connector) sticking up on the left in this picture, and the gel cassette is facing toward you, with the writing facing out. (You could put the gel on the other side, but it would be harder to see as you load the gel.)

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Insert the assembled inner buffer chamber into the outer buffer chamber. The downward-facing banana plug on the inner buffer core must be inserted into the hole on the outer buffer chamber, as shown. Use the gel tension wedge to press the pieces of the inner buffer core together. Don't get the gel tension wedge backward, or you lid won't fit.

In this picture, the gel is on the left, away from the gel tension wedge.

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Look carefully: the gel cassette has a short plate and a tall plate. The short plate is facing in toward the inner buffer chamber, so the running buffer will be able to flow above the top of the plate and contact the gel. The buffer has to touch the gel to allow electricity to be conducted through the gel.

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Pour 1x SDS-MOPS running buffer into the inner buffer chamber. Make sure it's this exact buffer! The SDS-MOPS running buffer comes as a 20x concentrate, and you may have to dilute it to 1x before using it. If so, you can do the dilution in the 1x bottle, using the marks on the bottle to get the final volume right. Use deionized water to dilute the buffer.

Fill the inner buffer chamber all the way up so the buffer level is above the top of the short plate. This will require approximately 200 ml. You can't overfill it; any excess will just spill out the top, and that won't cause any problems. If you don't fill the chamber high enough to contact the top of the gel, your gel won't run at all. If you fill it just enough so that the buffer contacts the top of the gel only on one side, your gel will run crooked.

After you fill the inner chamber, keep an eye on the buffer level. Sometimes it leaks out.

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Fill the outer buffer chamber with running buffer, up to about 1-2 cm below the top of the gel. This large volume of running buffer acts as a heat sink, making sure your gel doesn't heat unevenly (which will cause the dye front and protein bands to curve down toward the middle of the gel). At a minimum, make sure you add enough buffer to cover the slit near the bottom of the outer gel plate. If the buffer doesn't touch the lower part of the gel, there will be no current and your gel won't run. Double check to make sure you've removed the tape at the bottom of the gel!

The gel comes packed in a storage buffer that's different from the running buffer. Rinse the wells gently as shown in this video to swish out the storage buffer and ensure that the wells are filled with running buffer. Use a P-200 pipetter for this. Don't jam the pipet tip between the two plates of the gel; just gently set the tip at the top of the short plate.

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Add 500 µl of NuPage antioxidant into the inner buffer chamber. This will help prevent disulfide bonds (previously broken by the sample reducing agent) from re-forming as you run the gel.

Don't put the antioxidant into the wells of the gel; just pipet it into the inner buffer chamber, away from the wells.

Load the gel and run it

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Load your gel-ready samples into the gel. The samples should be blue, because they have already been mixed with sample buffer and reducing agent and heated. The amount you load may be different for each experiment, but generally you should load approximately the same mass (micrograms) of protein in every lane. If you skip lanes, your samples will spread out more and you'll end up with a warped-looking gel.

The pipet tip won't fit down into the bottom of the well; hold the tip at the top of the short plate and slowly release the sample so you see it sink to the bottom of the well.

Since you have 10 lanes to load, give everyone in your group a chance to do some gel loading.

See the Lab Guide for specifics of what to put in each lane.

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Put the lid on. Make sure that the banana plugs on the unit mate with the connectors on the lid. If the lid won't go on, it might be because the gel tension wedge is in backward or because the inner gel chamber isn't inserted correctly.

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With the power supply turned off, connect the gel unit's cables to the power supply -- red to red, black to black. Turn on the power supply. Set the range switch to high and the display switch to voltage. Use the knob to adjust the voltage to approximately 200 volts.

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The power supply will maintain a constant voltage, according to how you set the knob. The current running through the gel will depend on the gel unit. Set the display switch to milliamps to see the current. At the start of your SDS-PAGE run, the current should be around 100-120 mA (milliamps); for native PAGE, it should be around 40-50 mA. If the current is much lower or higher there may be a problem with the way the gel unit is set up; see the troubleshooting section at the bottom of this page.

Also, as soon as you turn on the voltage, you should see bubbles being produced in the inner chamber due to water being split at the electrode.

 

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Once your gel is running, you should see the dye front quickly start to move down the gel. This is the blue dye from the sample buffer. Your proteins aren't visible yet, but they will be migrating down the gel more slowly than the dye front.

We're using gradient gels. The top centimeter of the gel (called the stacking gel) is made with a low percentage of polyacrylamide to allow all the proteins to start migrating into the gel. The rest of the gel ranges from 4% polyacrylamide near the top to 12% near the bottom. The lower percentage has larger pores, which are ideal for separating high-molecular-weight proteins, while the higher percentage is best for smaller proteins.

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There's no hard rule about how far to run your gel. Usually it's best to run it until the dye front gets close to the bottom of the gel; this takes about 50 minutes. If you run it too long, the dye front and the lower-molecular-weight proteins will eventually migrate out of the gel and disappear.

Wash and stain the gel

While you're waiting for the gel run to finish, fill a beaker with deionized water and warm it up in the microwave. You'll use this for washing the gel. Also, get a gel staining tray.

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When you're ready to stop, turn off the power supply and disconnect the leads, then remove the lid.

Undo the gel tension wedge and remove it. The buffer leaks out of the inner chamber.

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Remove the gel cassette from the chamber and set it on a paper towel. Use a gel knife to separate the plates as shown below:

This is a delicate operation but it takes some force to break the plastic welds and separate the plates. Start with the tall plate down and the short plate on top. Crack the plates apart all the way around, and you should be able to lift the short plate off. Make sure the gel isn't sticking to the short plate. Now you're ready to drop the gel into a gel staining tray filled with warm deionized water. The gel will tend to stick to the tall plate, because there is a "foot" of gel material sticking into the slit in the plate. Use the gel knife to gently push the foot free of the plate.

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You won't be able to see any protein bands until you stain the gel. Before you do that, you need to wash all the SDS out of the gel so the stain can bind to the proteins. Soak the gel in warm (not hot) deionized water for 3 minutes with occasional agitation.

Pour the water down the sink, holding the gel down with your fingers.

Repeat this process for a total of four 3-minute washes with warm water. Don't cut this short, or your gel won't stain well. You need to give the SDS time to diffuse out of the gel. You're using warm water to speed up diffusion.

While you're doing the washes, clean up your gel unit, put it back together, and put it away. You can throw away the comb and plates that came with gel, but don't throw away the buffer dam! That's part of the gel unit. Rinse all the parts of the gel unit briefly in deionized water, dry them carefully (don't break the wires), and put everything back together for the next lab.

After the last wash, pour off all the water and pump some SafeStain on the gel. Use enough stain to fill the bottom of your tray and cover the gel.

Cover the gel staining tray with plastic wrap and write your name and the date on the plastic. Store the gel in the refrigerator until next time. (You'll normally leave your gel in the stain from one lab period to the next.)

Destain and photograph the gel

Pour stain into waste container

If the gel has been soaking in stain for days, the whole gel will be blue. You should be able to see the protein bands, but they will be partially obscured by the background of blue stain in the gel. Destaining removes this background, leaving your gel transparent so the bands are easier to see.

When you return to lab, pour the safestain into the labeled waste container, using the large funnel. Gently hold the gel in the tray with your gloved fingers. (You'll probably have another lab activity to do on the same day, but start destaining the gel first.)

Rinse with water.

Pour some deionized water over the gel. Give it a quick rinse, pour the water down the sink, and add more water. Let the gel sit on your lab bench for a while, and you'll gradually see the background of the gel become more transparent, allowing your bands to stand out better. Leave the gel in deionized water for an hour or more, changing the water every half hour. Meanwhile, you can get some other lab work done.

Viewing the gel

Your gel should look more or less like this picture after a while -- strong blue bands against a clear background. When you're ready, carry the gel tray over to the white light transilluminator. Place a piece of plastic wrap on top of the transilluminator and write your group name and the date on the plastic wrap, leaving room in the middle for the gel.

Lift the gel out of the water using a spatula or your fingers. Slide the gel onto the plastic wrap, and you're ready to take a picture.

Camera settings Adjust the exposure

Photograph your gel. Turn on the transilluminator and the camera, and make sure the top dial is set on C mode. Raise the transparent lid on the transilluminator and put the camera on top. Make sure that the camera's hood is all the way down over the white light.

Look at the image of your gel on the camera's screen. Use the camera's zoom lever to zoom in a little, making sure that you include your group name in the photo. Use the left and right buttons to adjust the shutter speed, making the image darker or lighter until the bands show clearly. If your photo looks orange, it's because there is a UV-blocking filter on the camera, which we need for DNA gels; it will still work fine for your protein gel.

Don't try to use other picture-taking modes, like Program or Manual. The C mode includes the appropriate settings to help you get a good image. You just need to adjust the exposure.

Once you've got the right zoom and exposure, hold down the shutter release button and give the camera some time to focus. It's slow. Once it's focused, push the shutter release button all the way while holding the camera steady. The exposure may be long, and if you move the camera, you could get a blurred picture.

You can also take a picture with your own camera, but please take one with the lab camera so we'll have photos of all the gels in one place.

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Your gel photo should look something like this. The background is nearly white, the bands are clearly visible, and the name and date are easy to read. (You'll need this information to find your gel image on the website.)

When you've got your picture, put the camera into review mode (the blue arrow) and connect the usb cable to the lab's laptop. The computer should already be set up to automatically copy your image to the appropriate folder on the computer and delete it from the camera. If you took more than one image, please delete all but one.

Turn off the camera when you're done. If it's left in review mode, it won't turn off by itself, and the batteries will wear down.

When you see that you've got a good picture, throw away your gel in the biohazard trash.

Later, the instructor will upload all the images to the lab's flickr site, so you'll be able to view it and download it later. It's important to get everybody's gel picture on flickr, so you can compare your gel to others.

Alternate protocol: microwave wash and stain

In most cases, we won't have time to run the gel and then wash, stain, rinse and photograph it in the same lab period. Therefore, we'll use the long staining procedure described above. However, if you're fast, you can get it all done in one day, using the microwave to warm things up and speed up diffusion.

  1. Wash: After electrophoresis, put the gel in a tray and cover it with plenty of deionized water. Microwave on High for approximately 1 minute, until the solution almost boils. (Watch it and don't let it boil.)
  2. Let the gel sit for 2 minutes, swirling occasionally. Discard the water.
  3. Repeat steps 1 and 2 of this procedure 3 more times, for a total of 4 washes.
  4. Stain: After the last wash, add enough SafeStain to cover the gel and microwave on High for 45 seconds to 1 minute (1.5 minutes) until the solution almost boils. Watch to make sure it's not boiling.
  5. Leave the gel on your lab table for 15 minutes, swirling occasionally to speed up the diffusion of the stain. (Alternatively, you could leave the gel on an orbital shaker, such as the one we use for liquid bacterial cultures.)
  6. Wash the gel in 100 mL of warm (microwaved) deionized water for 10 minutes. Your gel is ready to photograph when the bands are visible against a fairly clear background. Take a picture as described above.

Troubleshooting the gel run

It’s possible that you could turn on your power supply and have no current running through your gel. In this situation, the voltage display could read around 200 Volts, while the milliamps reads 003. To make sense of this, you should understand how electricity works in your gel unit.

Ohm’s law and SDS-PAGE

The electrophoresis power supply provides constant voltage; you set the voltage by turning the knob. (The power supply provides direct current, in contrast to the alternating current of the electricity in the building.) The current flowing through the gel is described by Ohm’s Law:

Voltage = Current x Resistance

Current is measured in amps or milliamps, and resistance is measured in ohms. If the resistance is very large, the current will be very small. If you do not create a complete circuit allowing electricity to flow through your gel, there is no conductivity (practically infinite resistance). Even with 200 Volts applied, the current will be zero.

What breaks the circuit?

Several things could cause the circuit in your gel unit to be incomplete; if there’s no current when you turn on the power, there are several possible explanations:

  • The tape hasn't been removed from the bottom of the gel.
  • There isn’t enough running buffer in the inner buffer chamber to cover the top of the gel, or enough running buffer in the outer chamber to cover the slit in the tall plate where the tape was removed.
  • The lid is on incorrectly; it’s possible to get it on without the male and female parts of the banana plugs being connected properly.
  • A wire is broken in the buffer core or on the lid.
  • The gel unit is assembled with the tall plate facing inward toward the inner buffer chamber and the short plate facing out. If this is the case, the running buffer in the inner chamber will touch the bottom of the gel instead of the top. If you fill the outer chamber enough for the running buffer to touch the top of the gel, it will run backward! Your samples will jump out of the wells and disappear. In my years of teaching 6B, I’ve only seen one group do this.

Alternatively, if there is current, but it’s too low (milliamps below 100 at the start of the run):

  • The running buffer is too dilute. The running buffer contains salts to increase its conductivity; if it’s too dilute, the conductivity will be low, the current will be low and the gel will run slowly.
  • There might be just enough running buffer in the inner chamber to touch part of the top of the gel, but not enough to cover the gel all the way across. If this is the case, the gel will run slowly and the gel will be distorted when it runs.

Review

Terms & concepts

  • Electrodes
  • Gradient gel
  • Power supply
  • Reducing agent
  • Running buffer
  • Sample buffer
  • SDS (Sodium dodecyl sulfate; LDS, or lithium dodecyl sulfate is almost the same. The dodecyl sulfate ion is the important part.)
  • Stain for protein gels (SafeStain is what we use)

Review questions

  1. What are the functions of the sample buffer and sample reducing agent? Why do the samples need to be heated before you load them on the gel?
  2. What determines the current in your gel? What could cause the current in your gel unit to be lower than expected? Is there anything that could cause it to be higher than expected?
  3. How will you see your protein bands after running the gel?
  4. Why do you need to wash the gel before staining it? Why use warm water?
  5. Why do you destain the gel?
  6. What is a gradient gel? Why are we using them?
  7. Compare and contrast SDS-PAGE and native PAGE. Why would we want to do each of these techniques?
  8. Where are the electrodes in the protein gel unit? Which one has which charge?

References & further reading

Simply Blue SafeStain User Manual from Life Technologies. This is the protein gel stain we use in the 6B lab.

XCell SureLock Mini-Cell User Manual from Life Technologies. This is the protein gel unit we use in the 6B lab.

GFP Electrophoresis

Isoforms of green fluorescent protein differ from each other in solvent molecules 'trapped' inside this protein. On native PAGE gels with GFP (and BFP) we sometimes see two fluorescent bands. This article offers an explanation.

Native PAGE

Novex Tris-Glycine Mini Gels, WedgeWell Format quick reference.

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