Transcription, epigenetic, epigenomic techniques - BioWiki Knowledge Base

Sonication Vs MNase for ChIP assay

Fri, 26 Aug 2011 00:20:34 +0000

I am sure that this debate has been covered before, but I just wanted to share my own personal experiences with it. I am working with a TF and fairly recently switched to MNase. I initially tried MNase digestion out of pure curiousity, but I have to say that I (and my experiments) have not looked back since. I was deeply unhappy with sonication as it just didn't feel right and my experiments were not reproducible to the level that I would have liked.

I have found that MNase fragmentation is superior to sonication in EVERY way. I am talking better %input, better specificity, lower background and much, much, much higher reproducibility. I have even compared the two in ChIP-Seq experiments and there is a night-and-day difference between the two (tighter peaks, much less noise/bias). The only downside is that a lot of optimization is required but, once you are over the initial hurdle, it is so much easier in my hands.

I am sure that most of you will respond with the usual "yeh but you should use sonication for TF, but MNase for histones etc". But I think this dogma is just that, dogma. If you get the MNase shearing right (i.e. not down to single nucleosomes), you will be amazed at the results. Seriously, try it!

Just wanted to see how many others are starting to use MNase for TF ChIP?

chabraha
Posted 15 April 2011 - 07:30 AM
Dukey,

I am glad to hear that someone is really willing to vouch for MNase. I really want to switch to it before i start ChIP-chip or ChIP-seq, whichever one I can justify doing. Can you elaborate on the optimization step and explain why it is important not to have single nucleosomes? Thanks for the post.........personally, the variability of sonication makes me sick.

Dukey
Posted 18 April 2011 - 12:12 PM
Sure!

Once you get down to single nucleosomes (<150 bp), you really start to see dramatic drops in % input values. I think that this is mostly because you are working blind in the primer design stage (at least I was) and so you may not be right on top of the binding site and you really have no flanking region to "buffer" the PCR. So your chances of capturing the binding event dimish, unless of course your primers are right on top of the site. If you go for slightly larger range of fragments (i.e. one to six nucleosomes) you see a huge jump in % input and I think it is related to the primer design point above. Note that there is really no parallel increase in background with less fragmentation.

I am convinced that this over-digestion problem is why MNase digestion has gotten a bad rap with TF ChIP. Most of the objections to it are also theoretical in nature and there is really little data to suggest it is inferior to sonication. In fact, all of my data suggests the complete opposite, as does a whole bunch of data from some vendors who are pushing it (i.e. Cell Signaling Technology). Once optimized, it is also very reproducible, assuming experimental conditions are kept consistent.

In terms of optimization, it is quite simple. You just have to titrate very carefully the MNase and be very careful to keep cell number consistent. In my hands MNase was very potent in chopping up my DNA and I ended up using a very small quantity in my reactions. One big simple optimization experiment should take care of it. I would be more than happy to give you some more details should you decide to switch.

I am submitting my ChIP-Seq data for publication this week and so we will see how it copes in peer review. I am not anticipating any major issues with the ChIP methodology.

chabraha

Posted 18 April 2011 - 07:57 PM
Good to know about the vendors pushing it....I will buy from one like cell signaling I'm sure. Right now I am waiting on a cell line and have to do a few tests on it before I can do my ChIPs, but if everything goes well I will for sure be contacting you. Best of luck on the manuscript!

roelq
Posted 19 April 2011 - 02:21 AM
Hey Dukey,

I was wondering which kit you are using for the MNase. We are currently trying the SimpleChIP kit (Cell Signaling), but we are having problems with the sonication step (too bad it still needs sonication).

How many cells do you normally use, how much MNase, how much lysis buffer, which type of sonicator?

Hope you can help, because we are really struggling right now.

Cheers,
Roel

Dukey
Posted 20 April 2011 - 09:20 AM
OK, so here is my brief protocol. I use the SimpleChIP enzymatic magnetic kit from CST (#9003).

- Harvest 5 x 10(7) cells per experiment. For me that is about 4 x 10cm dishes of cells.
- Follow protocol exactly (except for 30 min protein lysis on ice), up until the addition of MNase
- I add just 0.3 ul of MNase/5 x 10(7) cells in 1 ml of buffer B and leave for 20 min at 37 degrees
- The protocol states that 5 ul of MNase is what CST used for HeLa cells. When I tried this amount, I had nothing but single nucleosomes.
- I then resuspend the nuclear pellet in 1 ml ChIP buffer as per the protocol and split into two 500 ul aliquots
- I then sonicate very mildy just to break open the nuclei. For this I use a Branson Sonifier on 20% and do 2 x 10 second pulses for each aliquot.
- 100 ul of the clarified chromatin is then used in each IP reaction (final volume 500 ul). Therefore, I use about 5 x 10(6) cells per IP reaction. This is pretty much per the protocol
- IPs are left O/N and then I follow the protocol exactly from there on in.
- For ChIP-Seq, I will usually do 4 - 5 individual ChIPs for my TF and then combine at the DNA purification step to get enough material for quantification and library prep.

Hope this helps a little.

Mighty Mouse
Posted 20 April 2011 - 03:47 PM
Interesting post. I always shied away from the MNase because I heard from others who tried it that it simply resulted in too small of fragments, as you suggested. I wonder if it would work as consistently in tissue samples in which it is much more difficult to determine and control the number of cells harvested and thus to obtain consistent optimization.

Interesting discussion.

MM

roelq
Posted 21 April 2011 - 05:31 AM
Thanks a lot.

I am wondering if you ever tried to use lower cell number. We are currently trying with 4 x 10^5 cells because there is no way we would ever get 10^7 cells for our purpose.

One of our problems is that we seem to lose our DNA during column purification. So now that you say that you use only like 0.3 µl MNase for 120x the amount of cells that we have, I guess we are just overdigesting our DNA and it is just too small to stick to the columns. Would you agree with that?

Cheers,
Roel

Dukey
Posted 12 May 2011 - 12:44 PM
I have not tried lower cell numbers but I guess it could work, as long as you titrate the MNase very carefully. You may need to dilute it to make it easier.

chabraha
Posted 13 May 2011 - 02:17 PM
Dukey,

In a not so related question......What about MNase digestion on viral chromatin, which has a low nucleosomal content compared to cellular chromatin? Any down (or up)-side to using MNase on irregularly chromatinized with low nucleosome content?

nanook
Posted 06 July 2011 - 08:47 AM
my major concern regarding MNase-ChIP is the chromatin release. are you sure you get a representative fraction of chromatin released from the nucleus? did you sequence your input chromatin, how evenly do the reads distribute?

according to my expts a huge part of chromatin remains in the nuclei after digestion with the released one being mainly chopped down to mono/di-nucleosomes.

i would be really interested in trying the MNase approach but without a proper release this technique is useless.

Dukey
Posted 03 August 2011 - 01:01 PM

nanook, on 06 July 2011 - 08:47 AM, said:

my major concern regarding MNase-ChIP is the chromatin release. are you sure you get a representative fraction of chromatin released from the nucleus? did you sequence your input chromatin, how evenly do the reads distribute?

according to my expts a huge part of chromatin remains in the nuclei after digestion with the released one being mainly chopped down to mono/di-nucleosomes.

i would be really interested in trying the MNase approach but without a proper release this technique is useless.

Useless????? Wow.

The answer to your question is YES I did sequence the input chromatin and the reads distribute pretty nicely. In fact, the results looked a whole lot better than with sonication. That is for certain. Actually when you compare my data to other published studies, my results are VERY consistent. Actually the paper is in press now so obviously the reviewers didn't have a problem with my technique.

As I stated in my original post, the reason that MNase has gotten less attention is because of people like you following the crowd. If you digest down to mono-nucleosomes, then that is what you will get. However, if you do the proper experiments before hand and ensure that this doesn't happen, I personally believe that you get a good chromatin prep - or "proper release" as you call it.

How do you know you get "proper release" when you blast the crap out of your DNA-protein complexes with a sonicator? How do you know you don't destroy epitopes with sonication?

wangjing
Posted 03 August 2011 - 01:28 PM

Dukey, on 20 April 2011 - 09:20 AM, said:

OK, so here is my brief protocol. I use the SimpleChIP enzymatic magnetic kit from CST (#9003).

- Harvest 5 x 10(7) cells per experiment. For me that is about 4 x 10cm dishes of cells.
- Follow protocol exactly (except for 30 min protein lysis on ice), up until the addition of MNase
- I add just 0.3 ul of MNase/5 x 10(7) cells in 1 ml of buffer B and leave for 20 min at 37 degrees
- The protocol states that 5 ul of MNase is what CST used for HeLa cells. When I tried this amount, I had nothing but single nucleosomes.
- I then resuspend the nuclear pellet in 1 ml ChIP buffer as per the protocol and split into two 500 ul aliquots
- I then sonicate very mildy just to break open the nuclei. For this I use a Branson Sonifier on 20% and do 2 x 10 second pulses for each aliquot.
- 100 ul of the clarified chromatin is then used in each IP reaction (final volume 500 ul). Therefore, I use about 5 x 10(6) cells per IP reaction. This is pretty much per the protocol
- IPs are left O/N and then I follow the protocol exactly from there on in.
- For ChIP-Seq, I will usually do 4 - 5 individual ChIPs for my TF and then combine at the DNA purification step to get enough material for quantification and library prep.

Hope this helps a little.

hi, What MNase are you using? Cat. No? and what buffer condition? I really want to switch to MNase soon! thanks!

Dukey
Posted 04 August 2011 - 11:06 AM
I am using the SimpleChIP Enzymatic Chromatin IP Kit #9003 (Cell Signaling Tech). Everything you need is in there and, apart from the modifications I have already mentioned, I pretty much follow the protocol.

Edited by Dukey, 04 August 2011 - 11:06 AM.

angelawu
Posted 22 August 2011 - 06:54 PM
Hi Dukey (and others),

Thanks for this post. I've been working with MNase for years now, but not been able to optimize it well enough for a good prep - caveat though, is that I'm trying to do it for thousands of cells instead of millions of cells, in which case the enzyme kinetics might be a lot different. My question for you is, do you know what buffer is used for lysing the cell prior to MNase treatment? The protocol from cell signalling doesn't give the specific buffer conditions. I'm just curious as to what buffer would be compatible with both sufficient cell lysis to release the chromatin for digestion, but also allow the enzyme to still function... For me, I think my buffer does the job for both, but badly, which is why my chromatin prep isn't so great.

Theoretically speaking, overdigestion with MNase of the chromatin into mono-nucleosomal fragments would give you bad qPCR results because you may not capture the protein binding site with your primer since the fragment length is smaller than your amplicon. However, this should be a non-issue if you are sequencing. So my question is, if you are just sequencing your chromatin-IP and not doing qPCR, does it even matter if the digestion has gone to completion and all the chromatin is in mono-nucleosomal fragments? Surely, for histone analysis this is even beneficial because it would give a higher resolution of the histone binding site? Anyone care to comment on this, or maybe have any experience with it?

Thanks in advance.

Edited by angelawu, 22 August 2011 - 07:00 PM.

Dukey
Posted Today, 04:25 PM
angelawu,

You ask two very good questions. My impression from the buffer in the Cell Signaling kit is that it is a pretty standard lysis buffer with a detergent, most likely SDS (judging on the consistency and appearance of the buffer). Beyond that of course, I don't know the exact details of the solution. However, note that this protocol uses a second nuclear lysis/wash. The initial cell lysis buffer is spun out and the nuclei are lysed and washed in buffer B (+ DTT) and the enzyme digestion is then performed in this same buffer B. So the composition of buffer B is the most important for successful digestion with MNase. I would guess that buffer B is a standard MNase reaction buffer.

Now to your second point. I actually agree with you 100% in that it might be beneficial in some cases to allow the digestion to go all the way to single nucleosomes, at least theoretically in ChIP-Seq. But you answered the question really because the issue with that is that it becomes impossible to validate your library prior to running the sample on the machine. It also becomes more complicated when you want to compare ChIP signals to ChIP-Seq signals because for standard ChIP there is no way you should digest all the way. I know from experience that an even slightly over-digested chromatin will give dramatically different results to a well prepared sample. So you would have to have two different protocols and two different sizes of chromatin for ChIP and ChIP-Seq and I'm not sure that is ideal.

Hope this helps a little.

How to avoid foaming during sonication in ChIP...

Sat, 07 Aug 2010 06:20:07 +0000

ChIP - Foaming in sonication and other woes - (Jan/27/2005 )Hi all! I was wondering if anyone out there has got any trick to avoid foaming during sonication in the SDS lysis buffer for ChIP assays. Supossedly, placing the tip of the sonicator probe 13 mm below the surface of the liquid should avoid it, but is just not working on my hands. After 10 sec on the first sonication pulse a get lots of foaming, but I like to do longer pulses (20 sec) and foaming is supposed to denature the sample. For how long do you sonicate, and how many pulses? I'm using a Branson 250 sonicator.

Also, which type of vessel do you use for sonication? I use 2 ml eppendorf tubes, but the instruction manual of my sonicator recomends metal or glass vessels, better than plastic ones, because of heating considerations.

And last but not least, have you noticed differences in the sonication quality when using different cell lines? For some reason, I get really small fragments with 3T3 cells but not with 10T/2, and both of them are fibroblast cell lines. I don't change the conditions and sometimes I perform the experiments in parallel

Any thoughts or suggestions will be very welcome. Thanx!

-badcell-

It is tricky to avoid foaming during sonication. I have tried both 1.5 ml (conival bottom) and 2 ml (round bottom) tubes and found 1.5 ml tubes are less likely to produce foam. In addition, according to this protocol, conical bottom improves sonication efficiency.

To avoid foaming, dip the probe all the way to the bottom of the tube.
To avoid heating, hold the tube in ice water during sonication.

Hope that is useful.

-pcrman-

Thanx alot for your answer, pcrman. I'm surprised to see that in the protocol you attach the sonication step is performed in RIPA buffer instead of the more astringent 1%SDS buffer. I've tried before to sonicate the sample in RIPA and the results were dismal, 10 kb DNA fragments. Maybe is because I used larger volumes (2-3 ml) corresponding to the final volume of the IP sample? I see that in this protocol the use smaller volumes and then dilute after sonication. I'll try that. Thanx again!

-badcell-

If you want to avoid foaming during sonication, cup horns are the way to go. They utilize indirect probe sonication, and are able to sonicate sealed vessels. Another advantage is that you can do multiple samples at once. The following link gives a better description:

http://www.misonix.com/Products/index.cfm?...6&div=11&cat=21

Contact Misonix and they will be able to provide lab procedures for DNA shearing using the cup horn.

-sonix82-

QUOTE (badcell @ Jan 27 2005, 12:12 PM)After 10 sec on the first sonication pulse a get lots of foaming, but I like to do longer pulses (20 sec) and foaming is supposed to denature the sample. For how long do you sonicate, and how many pulses? I'm using a Branson 250 sonicator.
Hi Badcell,

We use a Branson Sonifier B-30. I had real problems with foaming initially and I came to realise that the chromatin is not shearing properly when it foams up. We were using maximum settings on the sonifier with no luck.

So I tried lowering the settings with the same pulse times. The settings I used and were sucessful are a duty cycle of 30% and power setting to 3. I performed 15 continuous pulses for ten seconds each in 1.5 eppy tubes set in an ice bath to reduce the heat from sonication. We resuspend and sonicate our chromatin in RIPA buffer also.

I would suggest lowering the settings of the sonicator, that certainly stopped the foaming issue and still sheared the chromatin quite nicely.

The cells we work with are CHO hybrids but I am sure different cell lines behave differently. You could try optimising formaldehyde fixation times as I would think cell lines would behave differently to the fixation. (for our cells, 1% formaldehyde in media for 10 minutes exactly is suffice)

hope this helps!

Nick

-methylnick-

Thanks all of you for your suggestions, they really help!

Nick, I recently tried something similar to what you said: I fixed the cells using a lower concentration of formaldehyde than usual (0.5% for 10 min) and the shearing seemed to improve. I'll try your suggestion to decrease the power of the sonicator and increase the number of pulses.

Sonix, I thought water bath sonicators are not fit for shearing the DNA for ChIP assays, 'coz they're not powerful enough. I checked the webpage you indicated and it says that this cup horns are high intensity sonicators, but will the sample not heat up a lot during sonication? Heating of the sample during sonication, I heard, is supposed to denature the proteins, even break the formaldehyde bonds.

Thanx again both of you for your help!

-badcell-

QUOTE Nick: I would suggest lowering the settings of the sonicator, that certainly stopped the foaming issue and still sheared the chromatin quite nicely.

Woe, this tip works and solved my foaming problem too! Previously I used a power setting of 15% (setting 3) and got foaming easily. After I lowered the setting to 10% (setting 2), I hardly got any foaming. Thank you Nick!

Here is my result with high and low power (volume 400 ul)

-pcrman-

no problemo pcrman,

I always thought that you needed the high setting to ensure proper chromatin shearing. but you don't

N

-methylnick-

I'm using sonication to lyse E. COLI in a protein purification procedure. I'm trying to explain some strange results I observed.

I used a 25ml sample in a sonication test on duty-60%, intensity=6 for 5 minutes, taking samples every minute. Running the crude extract and the insoluble suspension on an SDS-PAGE gel for each time point showed very little difference in sonicating one minute and sonicating five minutes.

In addition, the insoluble and soluble bands were identical, which makes the presence of inclusion bodies less likely.

My lab is relatively new to sonication, so the hope is that the error is ours and not in the experiment. If sonication produced foaming, therefore denaturing the proteins, would this explain the results - at least in part?

-mwwall-

Hi mwwall,

Here we are talking about sonicating DNA. It seems that you are sonicating protein, right?

Once foaming occurs, sonicating efficiency goes almost to zero. I found your power setting is very high, have you experienced foaming? If yes, decrease your setting. Your are using 25 ml sample for sonication, that sounds to much to me. For ChIP sonication, the max volume I tried is 400 ul. Certainly the bigger the volume, the lower the efficiency.

-pcrman-

I just noticed the photo of you gel pcrman,

there have been some changes to the website, and the inclusion of photos with the messages is just awesome!!!

great to see lowering the settings worked for ya! seems so simple a change doesn't it?

Nick

-methylnick-

I have been using the Upstate ChIP kit quite a while. Here are some tips which I think are important but not documentated in the protocol.

1) When taking the agarose beads using a yellow tip, cut off its end otherwise you will end up with dry beads only left (because the diameter of the beads is roughly the same of lumen of tip end.

2) After reverse crosslinking at 65C for 4 hr and one hour of proteinase K digestion at 45C, before going to phenol/chloroform extraction, you have to transfer the samples into a new tube because the cap no longer fit tightly on the tube after intensive heating. If you don't, your sample will leak when you vortex.

-pcrman-

Thanx for all, although I haven't take ChIP in practice....
There are many nice man tell me the tricky point in experiment...
These help me to improve my exp.
Thanks very much, Everybody...

-Ming-

Posted 03 March 2009 - 03:51 AM There are several reasons for foaming, but there are some main:
wrong vessel (wrong turbulence while sonication), too high power setting of the ultrasonic device, bad vibration behaviour of the system especially used microtips, wrong proceedure and system handling due to bad support by sales.
But foaming is not the only problem to have best results!
If you have any specific questions, do not hesitate to ask. I am very experienced in this by serving ultrasonic users in bio sciences since more than 45 years and Germany, but also worldwide.
We usually propose to use a special set up based on a BRANSON SONIFIER 450 (and OEM units, not other manufacturers, because their over all performance and quality is poorer!) using a special cup horn including a cooling device for full temperature control.
It is no problem to achieve highest ultrasonic power (is not combined with WATT of the power supply....!!!!) if necessary with full temperature control at around 2 - 4 �C. We are able to handle 7 test tubes at a time! Sample volumes smaller 100 �l is no problem!
We know all systems on the market, so we can give recommendations for use of other units to recieve the best goals!
Ask for more details
Andreas

Posted 03 March 2009 - 03:59 AM The use of the right vessel is always a question of sample volume:
The conical tubes (1.5 ml) are good for sample volumes starting around 250 �l,
when using the round bottom one (2 ml) the minimum volume for successful sonication (without foaming - foam is killing hte performance down to approx 30 % left power only!) is around 1 - 1.3 ml.
The recommendation regaring glass or metal vessel is for heat transfer reason only - and usually given by people, who never worked with a system or may not even has seen one except in a brochure!
Count on people who are really experienced, but also know the technology background of the ultrasonic technology!
Ask me for more details in ultrasonic treatment!
Andreas

Posted 11 March 2009 - 11:02 AM
Hi Andreas I need help with a sonication that is not working.. but I'm using a Misonix 4000 sonicator with a microplate cup horn and I need to shear the chromatin of mammalian cells (Saos2 and MCF-7).
No results till now.
I have been using a bioruptor in the past with NO problem.. but this Misonix is killing me!
Help!

Posted 01 May 2009 - 06:52 AM

I use the Branson 450, which is similar to your 250. I also had problems with foaming initially but reducing the power output and increasing the number of pulses solved the problem.
Don't hold the tip too close to the bottom of the tube because you don't evenly shear your chromatin if you do.

Different cell lines certainly can behave differently in sonication based on their size and number. If your cells are small then you get more of them in a dish. The more cells you have the longer you need to sonicate for to fragment more chromatin. Seeing differences in the sonication efficieny between cell lines is certainly not unusual as a result.

Be careful with water bath sonicators because not all of them are powerful enough to shear DNA, and for some the power distribution around the horn isn't equal. Thus, seek some serious advise before getting one - and I don't mean advise from the company who will tell you anything to get you to buy their machine even if it's not suitable. From what I've heard, bioruptor's are supposed to be the best for ChIP, why the switch for whoever it was using the Misonix? First rule of science (and the first 10 rules for ChIP) - if it's working, don't change it!

How to amplify long fragment from bisulfite tre...

Sat, 07 Aug 2010 01:05:26 +0000

Quote

I am able to very successfully get small (150-200 bp) products using a nested PCR strategy on bisulfite modified DNA; I do not see products from the larger primers but they are certainly there since the nest works so well. I'd love if I could sequence those larger products (400-500 bp). Does anyone have any hints to get a visible PCR product? Can I just ramp up the cycles? (I do 35 cycles and an MgCl2 concentration of 3 mM) Before you tell me this is just a normal PCR problem, you should know that most papers specify that a nest is necessary, one should not expect to see the larger product, and larger products after modification are fragile and difficult to get.

Thanks!!

labtechie

Hey Labtechie,

I have been able to successfully amplify and sequence 1kb from bisulfite treated DNA.

A nested PCR strategy is the best option. I typically design a hemi-nested primer set that is, I pick a primer pair (A and that amplifies around 900-1.2kb in length and then a third primer � within the pair.

So this would require two rounds of PCR amplification. In the first round of PCR I would use Primer A and B, take 1uL of this into a second round of PCR with primer C and either A or B depending on how you have designed things.

If this fails a magnesium titration will do the trick. Again with the hemi nested PCR I would titrate MgCl2 at 0.5mM final concentration intervals.

so I typically do 0.5, 1, 1.5, 2, 2.5 and 3mM.

good luck techie!

the cycle number I use is 30 cycles for each round of PCR.

Here are my cycling conditions:

95C 4 minutes denaturation followed by 5 cycles of :

95C 30sec (denaturation)
annealing for 90sec (annealing temperature set at 2 degrees below calculated Tm)
72C for 120sec extension.

and then 25 cycles of:

95C 30sec (denaturation)
annealing for 90sec (annealing temperature set at 2 degrees below calculated Tm)
72C for 90sec extension.

Final extension period at 72C for 4 minutes and then a hold at 4C.

This is repeated for the second round of PCR.

I am unsure how you get the degrees symbol so I have omitted it.

The bisulfite treatment was overnight at 50C in the dark, and desalted with Wizard Columns.....

I have also used Human Genetic Signatures' MethylEasy Kit with the same success. The kit is certainly much easier than the conventional method.

I have used AmpliTaq from Applied biosystems and PCR mastermix from Promega and they both perform very well with bisulfite PCR in my hands.

Nick

Hi, I think the primer design ought to be the critical step when you do the PCR. You need to avoid use too many degenerate base for the primer. I used up to 4 sites for one of my primer and it work pretty good.
I usually use two pairs of primers to do the nesting PCR. The first round PCR should be in a higher strigent condition, i.e. use higher annealing temp and ramp at -.1 or -.2 degree per cycle. I run it about 30-35 cycles. The second round can be less strigent.
I usually did many different PCR in one block. There's some variations of Tm in each primer pairs but PCRs worked really good. Only very few primer pairs did not work. So I guess you ought to work more on primer design rather than titrate the MgCl2.
BTW, I usually amplify 400-700bp. I did succeed a 1.2kb products. But larger ones are indeed harder and sometimes not very necessary.

Differences between MSP and BSP methods

Sat, 07 Aug 2010 00:50:58 +0000

Quote

Hello everyone I'm new here and I was wondering if I could get some assistance.
In my research of the literature I thought the differences between the two methods were based on the proximity of the primers to the CpG sites and that BSP primers were designed without including CpG dinucleotides.
I was wondering if someone could clear it up for me and what are the pros and cons of each method. Thank you very much in advance

Hi Gungrave,

I assume by BSP you mean bisulfite PCR and sequencing. If so, the advantages of this over MSP are:

A greater number of CpG residues analyzed for methylation when compared with MSP (MSP is only one and that is identified with a methylation specific primer). Bisulfite sequencing looks at every CpG residue across the whole amplicon.

I would also say that primer optimization is crucial for an MSP assay to work, it is not so crucial for BSP because you will be sequencing through the amplicon anyway.

MSP has it advantages in that it is far more quicker than BSP in terms of hands-on lab time.

hope this helps. I would be interested to see what other people think about this also.

Cheers

nick