The steps in your sucrose step gradient dissipate mainly due to diffusion, not centrifugal force. This would be true also for PEG and glycerol gradients. For self-forming gradients, given your time and CF, you would need to use a non-uniform gradient-forming material like Percoll, or else dense compounds like sodium iodide or cesium chloride (the latter two probably not appropriate, depending on what you actually want to do with your membranes). As a rule of thumb you put your sample in the bottom fraction if what you want to collect ends up closer to the top, and in the top fraction if what you want to collect ends up closer to the bottom. Theoretically it should make no difference, since the membranes should find their buoyant density either way. However, membranes tend to be sticky and will form micro-aggregates so that you will have contamination of light membranes with heavy membranes floating at intermediate density, for example. An important process that is often overlooked in gradient fractionation is fluid drag. This results from the movement of the membrane through the viscous solution to find its buoyant density. The farther the membranes travel, the more opportunity the fluid drag will have to pull apart adherent membranes of different origin. Therefore, if you follow the rule of thumb above you will get purer fractions. Sharp gradient interfaces can be obtained by centrifuging for shorter times (less time for diffusion). Sharper gradients will give discreet sharp bands of membrane at the interfaces. Collecting these is easier with just a pipettor and your sample will be much more concentrated, which may be useful. Some workers insist that sharper membrane bands are not as pure, and they prefer to let the gradient diffuse and then collect the broader band and recentrifuge it to obtain a pellet. My own experience with plasma membrane fractionation from CHO cells and primary human fibroblasts is this: You can get the same purity fraction in a 2 h spin (plus no-brake spin-down) as in a 24 h spin. The sharp interface means you can collect fractions that are concentrated and useable immediately for assays without spending more time recentrifuging (and having to deal with resuspending dense pellets). The result for us was that we were able to demonstrate enzyme activities and get undegraded bands in Western blots because our total prep time was only a few hours. In the 24 hour method we had significant loss of activity and some degradation of protein bands was evident. For viable biological activity, time is usually the most important factor. A further note: although we started with a published protocol, we tested many different times/methods and characterized fractions with TEM, marker enzyme assays and marker protein ELISA to completely characterize our fractions before going on to do more meaningful experiments. Hope this helps.

I think you've done well. It is natural that step gradient is dissapear after centrifugation. Your gradient will smoothly formed from 80% to 25%. So, you cannot distinguish any border. But, gradients do exist! Don't worry about that.

Hey, thank you for the quick answer! That's great news! Good then that I didn't trash the samples ... ;-)

The interface of gradients usually mix-up during spinning or if you keep overnight undisturbed (due to minor background vibrations). Its better to mark the level of gradients before spinning so that the approximate levels of gradients can be recognized. Mark around the tube because the friction due to overnight spin may erase the markings on some sides (dont use stickers which will lead to tube breakage). Mark when the tube is not cold. I usually calculate how much volume I will be loading per % gradient and I use similar volume of water to mark the level for each step. Then discard water and layer the gradients+samples.

Thanks for your feedback, Goodwin. Next time, I will mark my tubes before the run ...

It is natural that step gradient is dissapear after centrifugation. Your gradient will smoothly form from 80% to 25%. So, you cannot distinguish any border. But, gradients do exist! Don't worry about that.

Many thanks for your feedback, Golam!

As stated above, your various steps used to form the gradient behaved as expected for the formation of a linear gradient.

Recommended modification:

rather than separating your membranes by loading at the bottom of the tube and then allowing the various fractions to separate by floating up, you could load your membrane fraction in the lowest percentage sucrose and layer it on the top before centrifugation.

Thanks for your suggestion, Fedora. I had been wondering about this before I started my experiment. Suspending my sample in a solution as viscous as 80% sucrose was not easy. I would much rather prefer to get my sample into 25% sucrose and centrifuge it down into the gradient. Do you by any chance know why some people choose to load their sample at the top, whereas others prefer to place their sample at the bottom of the gradient? Is this merely a matter of personal preference or has it something to do with the nature of the sample or the type of gradient?

Sample loading from the top has several advantages.

1. There is less contamination of your high density material with low density fractions as occurs when loading from the bottom.

2. Depending on your sample, it is much easier to predict and visualize the final locations of the fractionated components of the sample.

3. Having sample components moving against the gradient as it is forming can be disruptive to obtaining the desired outcome.

4. A significant advantage to loading your sample from the top is that you can preform your gradient and then load your sample.

Thanks so much! This is very helpful information!

There is usually a distinct advantage to the float up method rather than the precipitation method. The float up is occuring against the centrifugation force and so a purer preperation is achived whereas in the precipitation method, everything heavier/ denser that the sucrose environment it is in will precipitate and the preperation will have contaminations.

I have prepared membrane fractions several times and have a simpler method. I resuspended the Tx-100 insuluble fraction in 2.5M sucrose to achieve 1.4M, then layer it with 1.2M, followed by 0.25M sucrose. Spin in TLS 55 if volumes can be kept within 3 ml or SW41 or SW28 again depending on volumes for 100,000g for 2 hours. Membrane fractions float up to the 1.2M-0.25M interface. Usually this is enriched in almost all organellar membrane but not 100% of the plasma membrane as that tends to be heavier. You may check out my publications or email me for details. Good Luck with your preparations and yes remember to mark your tubes the next time.

The method described above does work.

However layering from the top is not a precipitation method. It is a fractionation method. Components fractionate according to density. Certainly if you have components denser than the sucrose environment they will pellet. Obviously, if you did not want to achieve that result you would generate a more appropriate gradient.

Thanks Aparna and Fedora for your continued input to the discussion. In the literature you typically find people only describing the concrete conditions they chose for their experiment, but rarely why they chose them. This is extremely helpful for me for planning my next experiment!

The steps in your sucrose step gradient dissipate mainly due to diffusion, not centrifugal force. This would be true also for PEG and glycerol gradients. For self-forming gradients, given your time and CF, you would need to use a non-uniform gradient-forming material like Percoll, or else dense compounds like sodium iodide or cesium chloride (the latter two probably not appropriate, depending on what you actually want to do with your membranes). As a rule of thumb you put your sample in the bottom fraction if what you want to collect ends up closer to the top, and in the top fraction if what you want to collect ends up closer to the bottom. Theoretically it should make no difference, since the membranes should find their buoyant density either way. However, membranes tend to be sticky and will form micro-aggregates so that you will have contamination of light membranes with heavy membranes floating at intermediate density, for example. An important process that is often overlooked in gradient fractionation is fluid drag. This results from the movement of the membrane through the viscous solution to find its buoyant density. The farther the membranes travel, the more opportunity the fluid drag will have to pull apart adherent membranes of different origin. Therefore, if you follow the rule of thumb above you will get purer fractions. Sharp gradient interfaces can be obtained by centrifuging for shorter times (less time for diffusion). Sharper gradients will give discreet sharp bands of membrane at the interfaces. Collecting these is easier with just a pipettor and your sample will be much more concentrated, which may be useful. Some workers insist that sharper membrane bands are not as pure, and they prefer to let the gradient diffuse and then collect the broader band and recentrifuge it to obtain a pellet. My own experience with plasma membrane fractionation from CHO cells and primary human fibroblasts is this: You can get the same purity fraction in a 2 h spin (plus no-brake spin-down) as in a 24 h spin. The sharp interface means you can collect fractions that are concentrated and useable immediately for assays without spending more time recentrifuging (and having to deal with resuspending dense pellets). The result for us was that we were able to demonstrate enzyme activities and get undegraded bands in Western blots because our total prep time was only a few hours. In the 24 hour method we had significant loss of activity and some degradation of protein bands was evident. For viable biological activity, time is usually the most important factor. A further note: although we started with a published protocol, we tested many different times/methods and characterized fractions with TEM, marker enzyme assays and marker protein ELISA to completely characterize our fractions before going on to do more meaningful experiments. Hope this helps.

Wow, thanks Norbert for this very detailed information!

There is no one-answer-fits-all to the question of should your sample of interest sink down or float up. If you are going for the light fraction, contamination with denser fractions will be less if you float up. If you are going for the dense fraction you should layer at the top and let the dense fractions sink. If you want both or some thing in between, you might consider a self-forming gradient, as described in a previous comment. It all depends on your goals. You might have to try several methods to see which gives you the most satisfactory result, which means you need to select reliable markers for your fraction of interest and the fractions you most want to avoid.

Good luck!

Thanks Lawrence! All this information is definitely going to help me plan a better experiment next time!

If you are trying to separate only TX-insoluble from soluble fractions I would suggest you use a different type of step gradient. The setup from bottom to top is : 80% sucrose 1 ml, sample (I used 0,5% TX-100 to solubilize my samples not more) adjusted to 40% sucrose 1ml, 30% sucrose 7-8 ml, 5% sucrose 1 ml. The TX-insoluble protein-lipid complexes float to the interface of 5-30% sucrose and can be isolated easily without mass contamination. I strongly advise using a peristaltic pump for sucking the fractions slowly. More details with some controls at http://www.ncbi.nlm.nih.gov/pubmed/19022244

Good luck!

Your membrane fraction will be cleaner if you do layer them at the bottom and let them float than if you do load them at the top...... In addition you have to harvest the gradient as soon as possible (you mentioned on reply to one of the first comments that you keept the gradients), otherwise there will be diffusion

I think your gradient is working nicely. The interface normally disappear during time even you did not move it. If you really need some better sharp interface, you could try other gradient medium such as Optiprep.

I used to make up samples in 40% sucrose and layer steps of 5 or 10% decreases above that and then spin. The I would elute by pushing the gradient up with 80% sucrose solution, we had a tool for this. After running the gradient, I would use my blank for determining the density of my fractions. There is always some mixing and each step would be different by a few percent. May sure you use ice-cold sucrose solutions and get the gradients into the centrifuge as soon as possible, and slow acceleration and zero break. Are you using a swing bucket, fixed-angle, or vertical rotor for your spins? I used to use fixed-angle for mine. Good luck.

Thanks guys for your continued input. It is amazing how many responses I am getting to my problem. I will incorporate your suggestions in the planning of my next experiment.

From the http://en.wikipedia.org/wiki/Coriolis_effect

The Coriolis force is proportional to the rotation rate and the centrifugal force is proportional to its square. The Coriolis force acts in a direction perpendicular to the rotation axis and to the velocity of the body in the rotating frame and is proportional to the object's speed in the rotating frame. The centrifugal force acts outwards in the radial direction and is proportional to the distance of the body from the axis of the rotating frame. These additional forces are termed inertial forces, fictitious forces or pseudo forces.[1] They allow the application of Newton's laws to a rotating system. They are correction factors that do not exist in a non-accelerating or inertial reference frame. Sometimes at less than 100gs, the rotors decelerate too quickly. Sucrose also rapidly diffuses. Anyway, a linear gradient is better, since it maximizes the volume of gradient.

Robert C. Leif, Ph.D.

Thanks Robert for your feedback!

Usually gradients are made the day before and let at 4°C to equilibrate. The day after, you can not see the different interfaces. When you are running your centrifuge, gravity transform your initial gradient in a linear gradient. So, if you have light fractions and heavy fractions, it is normal. So to summurize, you have a mix of chemistry effect (sucrose dissolution) and gravity effect.

I think that the gradient is working great, but there is diffusion after the ultra centrifugation. You can test the density with a refractometer to test the gradient in differents parts of the tube. You need make first a calibration with and without Triton for see if there are differences due to Triton. Other possibility is make two ultra centrifugations, in the first refloat your interesting protein and pellet it in the second.

Thanks Stephane and Roberto for your kind responses!

Most of the prior responses contain useful information for using density gradients to purify membrane fractions. You can use either linear or nonlinear gradients depending on the density of the membranes of interest and the density of the most troublesome contaminants. Obviously, as mentioned by someone earlier, the longer the path through the gradient taken by the membranes of interest and the greater the distance between the membranes of interest and the predominant contaminants on the gradient, the easier it will be to obtain relatively pure samples. Jason Blum's tips are particularly practical. I too, used high density sucrose or Percol to displace the gradient from the top of the tube into fractions, using a simple tool we constructed in the lab. (Description available on request.). If your membranes are available in sufficient quantity, you can pellet them and reisolate from a second gradient for added purification. It may be useful to place the sample at the bottom or the top or alternate between the two in successive isolations, to optomize the isolation of the target from the contaminates. These choices will depend on the design of the gradients. Good luck

Nonlinear sucrose gradient is a great way to purify membranes.

One of the ways to make linear sucrose gradient is to use multiple fractions at evenly distributed concentrations that are spaced fairly closely, and let them diffuse through long-time spin or waiting. You probably want to avoid that.

For my non-linear / discontinuous gradients, I tend to use a few concentrations that are not evenly spaced, the volumes of fractions tend to have some differences too. The choice of sucrose concentrations depends on what membrane fractions you are trying to isolate, and what membranes you are trying to separate it from. Different membranes have different sucrose density equivalence. You might want to find that out and design your gradient accordingly. For example, if Golgi membranes has an equivalent density as 34% sucrose (I just made it up, don't remember exactly the numbers anymore), you want to use, let's say, 25% and 45% to sandwich the Golgi fraction. The volume of each fraction depends a lot on how tall your tube is and how much sample you are loading.

This brings up my second point, you don't want to overload your gradient. Each gradient and each fraction of it has its own capacity for samples, in terms of both volume and protein amount.

Also, I've used about 100,000 g for only two hr to separate Golgi, ER, plasma membrane and Mitochondria. You can still use brake, just use the lower (lowest?) setting.

Good luck!

Thanks guys, this is really helpful!

Steps in a gradient will always flatten during centrifugation, due to diffusion and unavoidable mechanical shaking during acceleration and deceleration of the rotor. Indeed, one method to make linear gradients is to make a step gradient and leave o/n in the cold room before loading the sample the next morning. Fractioning a gradient leads to some further mixing.

You may want to rethink switching the brake of, especially if you are using a swinging bucket or NVT rotor. Each rotor has a velocity where it is in the process of reorientation, during this phase the rotor runs less stably. Brakes make sure that this phase is crossed as fast as possible.

How do you balance the sample tubes, with or without buckets? The correct procedure is without, because slight differences in bucket weight are used to balance the rotor. This is why each rotor should always be used with "its" buckets, and the buckets always be put in the correct place on the rotor (numbers on rotor and bucket should agree). If you add balanced tubes (to within 1 mg) to a balanced rotor/bucket system, precession will be minimal. This will not only improve your samples, but also reduce wear and tear of the centrifuge drive. 1 mg may sound little, but at 260,000*g the centrifugal force will be as high as the gravitational force is on a mass of 260 gram.

Other than that, Methods in Enzymology are an invaluable source of protocols.

Thanks for your feedback Engelbert. I balanced the tubes without the buckets within 1 mg as you had suggested.

Hey, thanks David. This is very interesting! In my next run then, I will use the fastest acceleration and deceleration.