Cell Lysis Protocol

Share protocols and ask for sample preparation advice.
Firelark
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Cell Lysis Protocol

Postby Firelark » Fri Jul 15, 2011 9:43 pm

I am looking for a good method to lyse cells for subsequent proteomic analysis. Can anybody recommend a method? I am currently working with human cancer cell lines. Thanks.

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Postby David » Mon Jul 18, 2011 4:06 am

Well, I am sure there are going to be multiple opinions and responses for this, but here is our experience:

Our lab has multiple projects concerning quite varied human cell lines which are both cancerous and "normal" (A-431, RKO, hFF, etc). Though we initially were trying home-made lysis buffers, we eventually found that some commercial surfactants worked pretty well. We tried both Rapigest and ProteaseMAX, and found that ProteaseMAX works pretty well, and in fact better than several other more complicated buffers which we made up and tried out.

Lysis Buffer*:

ProteaseMAX (from Promega: http://www.promega.com/resources/protocols/technical-bulletins/101/proteasemax-surfactant-trypsin-enhancer-protocol/) 1%
Ammonium Bicarbonate (50mM) 90%
ACN 10%
Total Volume 1 ml

*We have found that since we do this fairly quickly, we do not need to add extra protease inhibitors. If you would like, you may of course also add protease inhibitors to the lysis buffer; I would recommend the Roche mini-Complete inhibitors and Roche PhosSTOP if you are interested in phosphoproteomics.


Lysis Protocol (for 10cm^2 flask):

1) Remove old media.
2) Rinse with PBS.
3) Add trypsin (0.5 mL). Incubate at 37C for ~3-5 min.
4) Add 3 mL of PBS to the flask then take up all solution containing cells to tube.
5) Centrifuge at 950 rpm (~1000 rcf?) for 5 minutes (3 times), remove supernatant.
6) Add cold lysis buffer to the pellet (on ice); 3:1 lysis buffer to cell pellet. Resuspend.
7) Remove to Eppendorf tube and spin.
8) Heat at 40 ̊C for 20 minutes.
9) 3x Probe sonication - 60 sec, pause 90 sec, Amplitude 30%. (Alternatively, can sonicate in water bath for ~30 min.)
10) Vortex, then centrifuge at ~13,000 rcf for 20 minutes.
11) Remove the supernatant to new tube.

I hope this helps, and will be interested to see what varied methods get posted here.

Good luck!

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Doug
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Postby Doug » Mon Jul 18, 2011 8:54 pm

This is the lysis protocol we have used for human embryonic stem cells but it seems like it should work for cancer cell lines as well.

LYSIS BUFFER
8M Urea
40 mMNaCl
50 mM tris
2 mM mgCl2
50 mMNaF,
50 mM b-glyceradelhyde phosphate
1 mM sodium orthovanadate
10 mM sodium pyrophosphate
1 (or part of one depending on volume) mini EDTA-free protease inhibitor (Roche Diagnostics)
1 (or part of one depending on volume) phosSTOP phosphatase inhibitor (Roche Diagnostics).

1) Add this to pelleted cells in a ration of ~3:1 (buffer to pellet)
-We often add a little benzonase too to digest the DNA and decrease the viscosity of the solution.
2)Vortex (on ice) 30 seconds on, 60 seconds off (repeat 3 times total)
3) Spin max speed on benchtop centrifuge to pellet debris
4) Keep supernatant (determine protein concentration)

If digesting with trypsin

5) Add LysC in a ratio of 1:100 (enzyme : protein) (~2 hours at 37 degrees C)
6) Dilute with 50 mM tris pH 8.0 until urea concentration is 1.5 M
7) Add trypsin in a ration 1:50 (enzyme : protein) (overnight at 37 degrees C)

I believe a similar method was used for this paper.

Swaney DL, Wenger CD, Thomson JA, Coon JJ. Human embryonic stem cell
phosphoproteome revealed by electron transfer dissociation tandem mass
spectrometry. Proc Natl Acad Sci U S A. 2009 Jan 27;106(4):995-1000. Epub 2009
Jan 14. PubMed PMID: 19144917; PubMed Central PMCID: PMC2633571.

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Postby domthe » Sat Jul 23, 2011 2:33 pm

Hi David,

Could you comment on downstream sample processing (in-solution digestion, labelling etc) following this method of sample prep? Is this method compatible with iTRAQ labelling of the peptides for instance?

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Postby RDUnwin » Sun Jul 24, 2011 11:49 pm

Domthe,

For iTRAQ labelling we just use 1M TEAB with 0.1% SDS (+Benzonase to digest DNA) for cell lysis. This seems to solubilise most of the proteins (for example we get histones and some membrane proteins (e.g. GF receptors) coming up in our subsequent analysis. We just lyse in this buffer (+ phosphatase inhibitors, if we are doing subsequent phosphopeptide enrichment) for 30 mins on ice with plenty of vortexing/manual pipetting to break up the cells. All you then need to do is collect the s/n, protein assay, and when you reduce/alkylate and digest ensure that you have feectively diluted the SDS out to 0.05% (anything about 0.1% and trypsin will start to work less well, so we stay on the safe side. For a detailed protocol, see Unwin et al in Nature Protocols.

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Postby Doug » Mon Jul 25, 2011 10:49 pm

David, I am also interested in how this works with downstream steps. The website says that the surfactant is designed to "degrade over the course of a digestion reaction" and that reactions (digestion) only takes one hour. This sounds truly amazing. And much easier that the buffer exchange required in FASP protocols. So can you just to Solid Phase Extraction and forget about the surfactant? Is it compatible with isobaric labeling? And do you see any extra peak in your mass spectra from these reagents?

It sounds very promising. Does anyone else have experience with ProteaseMAX?

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Postby David » Tue Aug 02, 2011 5:51 am

Yes, ProteaseMAX is designed to degrade over the course of digestion and should speed up digestion as well. We have used ProteaseMAX for both in-solution and in-gel digestions. However, with that being said, we do not change our digestion protocols, and I still typically set up digestions at the end of the day and let them go overnight. We have not played around with any conditions to see the effects of "speeding up" digestions. I must say, though, that we get very few missed cleavages in our final results and our digestions are very "complete", resulting in mostly 2+ and 3+ precursors over the course of the entire gradient. Also, we are actually using a robotic system for many of our digestions to aid in keeping high robustness between replicates, so again we just set this up near the end of the day and let it do its thing over the course of the night.

As for labeling compatability, we have personally not tried to do any labeling lately (we are a label-free lab). So, I called Promega and spoke to them about this. Here is what they had to say:
"ProteasMax is compatible with iTRAQ triethyl ammonium bicarbonate buffer, but not with the iTRAQ Reagent. ProteasMAX generates primary amines that compete with the iTRAQ amine labeling reagent.

The iTraq literature it is clear that primary amines are an interfering substance that need to be avoided. At this point the only recommendation that we could offer is that you could use solid phase extraction (either reverse-phase or SCX) to remove the interfering material before doing the iTraq labeling chemistry. My guess is this would not be an attractive alternative, however.

Just to follow up on why, iTRAQ reagents are amine labeling reagents that target N-terminal amino and epsilon amines of lysine. When proteasemax degrades one of the products is a primary amine that would compete with the peptide amines that the iTRAQ reagent is meant to label. iTRAQ instructions warn against using any buffers or components (in this case the degradation product of proteasemax) that would compete in this fashion."


However, we simply do either ZipTip, Spin-Trap, or SepPak clean up after digestion, and I believe that should be enough to remove these primary amines. All of these techniques have proven to work quite well for us, and we vary them only due to sample amount (SepPak for large sample quantity (>50-100 ug), Spin-Trap for "medium" (~20ug), and ZipTip for single gel pieces (at most ~2-4 ug). In regards to seeing any left over peaks in our mass spectra, I must say that none of us has specifically looked for any weird features. However, nothing atypical has stood out in the 100s of runs which we have done using protocols similar to the one I provided above. So, I am fairly confident in saying that nothing is left over which would hinder our LC-MS/MS.

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Postby LEE » Wed Sep 14, 2011 12:18 pm

Hi,
I have a question. How do people extract (mouse fibroblast) cells off of plates? I have of course tried SDS or Triton-X in my buffer, but I really don't want to use these considering those require additional clean-up later down the road. I have also tried a buffer containing MOPS and Urea (plus all of the protease inhibitors, etc.). I had to scrape the cells off the plate and it was not easy. And after performing the BCA assay, the protein concentration using the non-detergent buffer was a lot lower than using one with a detergent. Has anyone tried anything else to extract cells... like using RapiGest directly on a plate (instead of using it later on after the cells have been pelleted).

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Postby justin » Thu Sep 15, 2011 4:04 am

There are a couple of ways that you can remove cells before lysis. We prefer TrypLE from Invitrogen. The key is that you want to move quickly so that you minimize any changes that occur when removing the cells. Here is a step-by-step protocol:

-pre-warm TrypLE to 37 degrees C
-remove media from cells
-apply enough TrypLE to cover the culture surface (i.e., 2 ml for a 10 cm2 dish)
-wait approximately 5 minutes for cells to lift from the culture surface. If after 5 minutes the cells are still firmly attached, wait longer, checking every minute or so until you can see cells floating.
-add an equal volume (relative to the TrypLE) of ice-cold PBS.
-centrifuge at 1000 X g for 4 minutes
-wash by resuspending the cell pellet in 10 ml of PBS
-centrifuge at 1000 X g for 4 minutes
-wash again by resuspending the cell pellet in 10 ml of PBS
-centrifuge at 1000 X g for 4 minutes
-You can then either proceed with lysis or freeze the cells.

The volume of PBS that I listed is for 10^8 cells. You can probably go down if you have fewer cells. I've also used Trypsin from Invitrogen and Accutase. These work fine as well. A lot of people quench the reaction using 10% FBS but we avoid this because it triggers all kinds of signaling events. If you wash well with an excess of really cold PBS, you'll stop and then remove the trypsin.
Hope this helps.

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8M uea lysis buffer on ice?

Postby Duncan » Fri Jan 20, 2012 5:57 am

Hello Doug

I'm interested in this protocol where you effectively lyse cells in 8M urea and incubate on ice (I had avoided this due to solubility concerns)

Does urea fall out of solution when its stored on ice? I would be surprised if it all stayed in solution at this temperature!

I presume this doesnt cause an issue? So are you working on the principle that you have a urea saturated solution whilst on ice?

Cheers

Duncan

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Doug
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Postby Doug » Fri Jan 20, 2012 8:16 am

Hi Duncan,

That is a good question. Once in solution we don't have much of a problem of urea precipitation. I couldn't tell you why but these are my experiences.

It's definitely a bit challenging to get the urea into solution. This cannot be done on ice (or not easily). Room temp is fine but warming to 37 degrees can make it faster. Once in solution the urea is fine though. Aside from freezing the excess lysates we don't get them that cold. We sonicate on ice and leave on ice while determining concentration (1-3 hours on ice). But then reduction, alklyation, and Lys-C digestion are all done at room temp or higher. And after a few hours of Lys-C digestion we dilute down to 1.5-2 M urea.

After lysis we commonly store the lyse cells at -80 (in 8M lysis buffer) with no problems. Once thawed there is no noticeable precipitate (as far as I can remember, it's been a year or so). And samples that were processed and analyzed prior to and after a freeze that don't show any major discrepancy between number of IDs.

Quenching tryptic digestions (and preparing samples for SPE) with acid definitely causes precipitation of urea. And I always give the samples a hard spin here prior to SPE. I don't know if there is anyway around that.

Anyway, I know others on the site use that protocol and I am sure they have used it more recently than I have. Does anyone else have problems with urea precipitation?

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Postby Duncan » Fri Jan 20, 2012 9:23 am

Hello Doug

Thanks for the information. Its good to know You can keep 8M in solution on ice (despite my instinct).

Do you guys that perform urea extractions/urea buffer exchange followed by LysC and trypsin digestion have information as to the extent of lysine carbamylation induced by your procedure? Obviously, extensive lysine carbamylation is going to be particularly problematic with your proteolytic enzymes of choice. Do you perform LysC digestion at room temp in 8M urea in an attempt to limit this modification or is there another reason? If you perform trypsin digests at 37 degrees C in the presence of 1.5-2M urea, doesn't this induce lysine carbamylation?

Any comments on the extent of lysine modification, it's impact on these analyses and ways to limit its impact would be really useful.

Cheers

Duncan

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Postby Doug » Fri Jan 20, 2012 10:53 am

Sorry. That sentence was a little misleading. We perform LysC digestion and tryptic digestion at 37 overnight (alkylation is at room temp).

I can't speak to how much carbamylation we see. I am really a protocol user, not a protocol developer. I have been told that carbamylation is not that bad if you keep the temperatures below 60 degrees. I can't remember right now (and i just noticed its missing from the protocol) but I believe the reduction is done at 37 degrees to minimize carbamylation.

Again, perhaps other SharedProteomics members who use this protocol could provide more information.

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Postby mvlee » Fri Feb 03, 2012 10:19 pm

My experience has been there are a few things that you can do keep carbamylation to a minimum:

1. Always make the urea lysis buffer fresh immediately before lysing cells
2. As Doug said, don't go avoub 60C.
3. I do not allow my digestions to incubate longer than O/N

In regards to percipitation:
To minimize ppt when quenching a digest, I ususally add the acid incrimentallly rather than just one shot x% of acid of choice. By adding small volumnes of acid, you can acidify your digest just enough to quench which minimizing percipitation.

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Postby KLee » Tue Oct 02, 2012 2:31 pm

David wrote:Well, I am sure there are going to be multiple opinions and responses for this, but here is our experience:

Our lab has multiple projects concerning quite varied human cell lines which are both cancerous and "normal" (A-431, RKO, hFF, etc). Though we initially were trying home-made lysis buffers, we eventually found that some commercial surfactants worked pretty well. We tried both Rapigest and ProteaseMAX, and found that ProteaseMAX works pretty well, and in fact better than several other more complicated buffers which we made up and tried out.

Lysis Buffer*:

ProteaseMAX (from Promega: http://www.promega.com/resources/protocols/technical-bulletins/101/proteasemax-surfactant-trypsin-enhancer-protocol/) 1%
Ammonium Bicarbonate (50mM) 90%
ACN 10%
Total Volume 1 ml

*We have found that since we do this fairly quickly, we do not need to add extra protease inhibitors. If you would like, you may of course also add protease inhibitors to the lysis buffer; I would recommend the Roche mini-Complete inhibitors and Roche PhosSTOP if you are interested in phosphoproteomics.


Lysis Protocol (for 10cm^2 flask):

1) Remove old media.
2) Rinse with PBS.
3) Add trypsin (0.5 mL). Incubate at 37C for ~3-5 min.
4) Add 3 mL of PBS to the flask then take up all solution containing cells to tube.
5) Centrifuge at 950 rpm (~1000 rcf?) for 5 minutes (3 times), remove supernatant.
6) Add cold lysis buffer to the pellet (on ice); 3:1 lysis buffer to cell pellet. Resuspend.
7) Remove to Eppendorf tube and spin.
8) Heat at 40 ̊C for 20 minutes.
9) 3x Probe sonication - 60 sec, pause 90 sec, Amplitude 30%. (Alternatively, can sonicate in water bath for ~30 min.)
10) Vortex, then centrifuge at ~13,000 rcf for 20 minutes.
11) Remove the supernatant to new tube.

I hope this helps, and will be interested to see what varied methods get posted here.

Good luck!


You use 1% ProteaseMAX final conc? This seems pretty high compared to their protocols? Do you dilute your lysis buffer?
Cheers
Kate

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Postby isobaric » Sat Oct 06, 2012 9:10 am

I also want to bring up discussion about C18 Sep-Pak(waters). Hope someone can make some comments
Our lab has both Sep-Pak light and Pep-Pak plus cartridges, which works for different amount of starting material. I was told that Sep-Pak light works for < 1 mg of starting material whereas Sep-Pak plus works for > 1mg of starting material(could be wrong but I never test it). Can anyone make a comment on how to choose between Sep-Pak light and Sep-Pak plus?
In my experience, Sep-Pak can lose tons of sample, at least 50% based on BCA assay at protein level and at peptide level after Sep-Pak. It could be something wrong with protein(peptide) estimation or I did Sep-Pak in a wrong way. Anyway, do someone also have similar sample loss issue?
I almost gave up SepPak-based protocol for quite a while and switched to FASP method because FASP method requires much less material and works quite well for my sample. However, I am going to use antibody to enrich acetylated peptides, FASP method seems not quite convenient because in FASP method, I cannot start too much material(100 ug protein at most) because the filter definitely will get clogged(I am working with really dirty samples). I saw in open literature people generally would start with 10mg of protein for enriching acetylated peptides later which seems impossible with FASP method in my case. I will have to pick up Sep-Pak again. Can anyone having experiences with antibody enrichment for acetylated peptides make a comment on how efficient the enrichment is and what the recovery rate is?
Thanks

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Fasp

Postby proteovations » Sun Oct 07, 2012 5:29 am

Did you feel that FASP method gave you more IDs? Or work better? Have you tried comparing it? Looking for an independent assessment of the method.


At a conference, I recently saw people suggesting that using lots of cheap trypsin (maybe 1:1 or 1:2) instead of the 1:20 of sequencing grade is better for identification and quantitation. Thought I would mention it even though I have not tested it yet.
Benjamin J Cargile
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Proteomics Services | Protein Identification

isobaric
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Postby isobaric » Tue Oct 09, 2012 8:33 am

I did get better result with FASP method in my sample, though I did not systematically compare those methods.

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Postby bphinney » Tue Nov 20, 2012 3:31 pm

Just wanted to follow up on this. is the 1% correct? Did you find it works better than 0.01%? thanks!!

Brett

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Postby karthikuttan » Sun Dec 02, 2012 11:58 pm

RDUnwin wrote:Domthe,

For iTRAQ labelling we just use 1M TEAB with 0.1% SDS (+Benzonase to digest DNA) for cell lysis. This seems to solubilise most of the proteins (for example we get histones and some membrane proteins (e.g. GF receptors) coming up in our subsequent analysis. We just lyse in this buffer (+ phosphatase inhibitors, if we are doing subsequent phosphopeptide enrichment) for 30 mins on ice with plenty of vortexing/manual pipetting to break up the cells. All you then need to do is collect the s/n, protein assay, and when you reduce/alkylate and digest ensure that you have feectively diluted the SDS out to 0.05% (anything about 0.1% and trypsin will start to work less well, so we stay on the safe side. For a detailed protocol, see Unwin et al in Nature Protocols.


Dear Unwin and other members,

I am wondering whether I need to add protease inhibitors along with 1 M TEAB with 0.1% (wt/vol) SDS (I am not interested in phophoproteins). Also in Unwin et al article in Nature Protocols, I don't think I have seen Benzonase addition. So, I just wanted to clarify these points (and more below!) since I am also planning to do iTRAQ soon.

Also, it would be helpful if somebody could share some alternative iTRAQ-tested buffers for cell lysis (for human cell lines). I have seen people using RIPA buffer and then do acetone precipitation. How good is this method?

Thanks a lot for your help,

Kart

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Postby Doug » Mon Dec 03, 2012 7:56 am

You definitely want to add protease inhibitors but you probably don't need to add the phosphatase inhibitors. You don't need the benzonase but I would highly recommend it (and it is very easy to add). It makes the lysates less viscous and much easier to work with. I haven't done comparisons with and without it to know if it results in more chromatin protein IDs but there is some logic to suggest that it might.

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Postby domthe » Tue Dec 04, 2012 1:43 am

Doug wrote:You definitely want to add protease inhibitors but you probably don't need to add the phosphatase inhibitors. You don't need the benzonase but I would highly recommend it (and it is very easy to add). It makes the lysates less viscous and much easier to work with. I haven't done comparisons with and without it to know if it results in more chromatin protein IDs but there is some logic to suggest that it might.


But trypsin will not work with protease inhibitors! Most inhibitor cocktails has components inhibiting tryspin?! How do you deal this problem?

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Postby Doug » Tue Dec 04, 2012 8:32 am

I have used the roche mini protease inhibitor tablet as well as other protease inhibitors, followed by digestion with Lys-C, trypsin, and AspN, and other enzymes with no problem. The inhibitors are all still in solution during digestion. I haven't thought about why this works but it does. Perhaps, the excess of enzyme added. It is a good question I would be interested to know the answer to.


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