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 coat coverslips before seeding cells?

Mon, 22 Aug 2011 20:24:36 +0000

jennycharlies Posted 18 August 2011 - 06:55 AM
Hi,

I am going to seed my cells onto coverslips inside the wells of a 12 well plate and was wondering what is the best way to coat the coverslips?

I have 18mm coverslips and need to coat with poly-D-lysine. But presumably I need to sterilise the coverslips first.. What is the best way to go about this? I assume I need to coat the slides while in the wells of the plate?

Thanks for your advice!!

Chakchel Posted 18 August 2011 - 11:10 PM

Hi!

Yes, you first should sterilize your coverslips. You can do this either by putting them into the autoclave, e.g. wrap several in a little "bag" of aluminium foil, or by dipping them in Ethanol and shortly flame them (too long flaming will let them burst).

Then you can directly cover them in your well. Put your poly-D-lysine in the well, so that the coverslip is covered, but not too much, then let it dry in the hood for several hours. If then there is still some liquid, you can suck it of.
Finally wash the wells once or twice with PBS.

Done, and ready to use.

Edited by Chakchel, 18 August 2011 - 11:11 PM.

fysio lab Posted 19 August 2011 - 02:25 AM
Hello,

we put non-sterile coverslips in a glass petridish, wrap that in alu-foil, autoclave it and before use we put the CS in the wells with sterile forceps. After that we put our coating solution on the CS (in our case 2% gelatin), we leave it 30' in the incubator. Then we remove the gelatin and let it dry for a few minutes in the flow, lid open... meanwhile we split and count our cells. All together this doesn't take much time.
Good luck

DNA migrate differently in agarose vs PAGE gel

Tue, 16 Aug 2011 07:28:19 +0000

Quote

i am currently working a gene in leukemia. The mutant allele has been sequenced to contain 527 base pair. when i run an agarose gel to confirm, it did appear in the position approximately equal to 527 base pair according to the DNA ladder. HOWEVER, when i run a PAGE, it shows up at a different poistion!!(it is at about 600 base pair in PAGE!!!!). Yet the conditions were the same when i run the PAGE and agarose gel (150v, 30 mins)

Many things can affect DNA migration in gel. Check this page "Top 10 Fun Facts for DNA Electrophoresis" to see if there is anything that might have caused the abnormal migration. For example "On a polyacrylamide gel, DNA fragments having AT-rich regions migrate slower than other DNA fragments of the same size."

According to Fermentas troubleshooting, here is a list of things that may affect migration of DNA in gel:

Atypical migration due to different DNA sequence or structure. During high resolution electrophoresis DNA fragments of equal size can migrate differently due to differences
in DNA sequences. AT rich DNA may migrate slower than an equivalent size GC rich DNA fragment. DNA structures such as nicked, supercoiled or dimeric molecules will always show different mobility on gels compared to an equivalent DNA size standard.

Gel shift effect. The presence of DNA binding proteins in the sample, such as ligases, phosphatases or restriction enzymes may alter DNA migration in the gel or cause the DNA to remain in the gel wells. High salt concentration in the sample may also cause gel shift effects.

Tips on freezing and thawing of cells

Sun, 15 Aug 2010 01:50:14 +0000

I am facing problems in thawing of cells. The procedure which we follow is- we take stock from liq. N2 then though it in water bath at 37 degree. Then we take medium in falcon (5ml) then add thawed cells dropwise into it. We then centrifuge it at 1500rpm for 5 mins.
but the problem is that after thawing i am unable to get viable cells.
Almost all cells are dead. So what all modification shall i make ? Its really hard to revive cell line from frozen stock?

The general rules for freezing and thawing cells is: freeze cells slowly but thaw cells quickly. It seems to me that you have not frozen your cells in a gradual manner. There are many ways for achieving gradual decrease in temperature. Check the section for cell preservation on this site and you will find protocols of different flavors for this purpose.
http://www.protocol-...tion/index.html

One thing you could possibly try is not to spin your cells straight away, just add the warm medium, and let them sit over a few hours to overnight, then wash out DMSOcontaining medium, and feed with normal medium plus serum (increase serum conc to 10 or even 20% for a little while). Sometimes brittle cells get sheared while centrifuging, so that's why the modified protocol. DMSO is toxic though, so change medium as soon as you see under the microscope that your cells have settled.
If that's the problem you could instead decrease your spin time- even 1min at 1500 should pellet enough cells.
Also, I add medium to my cells, no vice versa, I'm not sure if that would make any difference at all.
As for freezing cells- I do it your way, it seems fine, but your cells could be very sensitive, so could be the problem.

Though my experience of handling the cells is less than 2years. I would like to suggest you some things. Even though if you leave DMSO in the media after thawing the cells the dmso in low cocentration is not toxic 2 cells more over once you dilute the small volume of cell suspension in 5-10 ml of media, it further dilutes the dmso. It looks to me like the protocol you are following may be causing the cells to die, I think you recognise the fact that centrifuging at high speed can lead to cell necrosis. One method is to lower the speed of centrifuge maybe 900-1000 rpm for 4-5 min should be adiquate.
Alternatively you should also consider the method you are using for preserving the cells. If you freeze the cells directly in liquid nitrogen this can also lead to cold shock induced cell lysis or damage the membrane, one way of preventing this is gradually lowering the temperature such as storing 4 degrees for few hours then transfering to -20 degree for overnight and then storing in -70 degrees for an couple of days then transfering to liquid nitrogen tanks, by following this method u r not only making the cells to adjust to the lower temp you are also ensuring that they donot undergo cold induced necrosis leading to cell membrane damage and lysis.

This is how to freeze and thaw cells:

Always check the cell viability before freezing. They should be highly viable: about 95%. keep your freezing media ( 10% DMSO in FBS) cold. Centrifuge cells for 1000 RMP for 5 minutes. Prepare labeled cryogenic vials. Cell concentration should be about for e.g. collected from a 50 ml flask, about 50 million cells. You can have 10 vials and about 0.5 milion cells per vial. After you centrifuge, get rid of media completely, gently tap the pellet to make it loose. Add 5 mls of cold freezing media resuspend with a pipet and transfer 0.5ml to each vial. Close the cap tight and place cryogenic tubes in special cryogenic container that has alcohol at the bottom and cool down gradually, they usually hold up to 20 vials. Close the top and transfer the container to -70 freezer. Wait 24 hours and no longer than 15 days before transferring them out of the container into the liquid nitrogen boxes.

To thaw the cells, be very quick, take the viall out still with some liquid nitrogen, walk to the water bath. Take the vial, make sure the cap is very tight, sometimes it becomes loose. thaw the vial holding the opening upward so the water from the water bath does not contaminate cells or if there is some detegent in the water it doesn't become in contact with the inside of the vial . When frozen cell media is almost half thawed (about few minutes), take it under the hood and add 0.5 warm media to the cells and immediately transfer to a flask with about 10 to 15ml warm media in it. DMSO will be diluted and you can check the viability.

Handling RNA - How to avoid RNase contamination?

Sat, 07 Aug 2010 20:55:25 +0000

Hi Susan,

I would second what tea-test has pointed out. Any tissue you process is going to be loaded with RNases (cells contain RNases). Any residual RNase present on your homogenizer is going to be very minor compared to what is already present in your cells. All you need to do is wash your homogenizer with soap and water and give it a rinse with clean MilliQ or DEPC water.

In my experience working with RNA, the majority of the degradation happens at the initial stages of the isolation. You need to work quickly to get the cells lysed and homogenized within whatever lysis buffer you are using. Sometimes RNA degradation is a problem for very inexperienced users who are very slopping (touching the insides of tubes/lids with fingers even with gloves on, leaving lids open so that bacteria (that contains RNases) can float in, taking way too much of the top layer and getting the white goop (protein) when using Trizol etc.). That not withstanding, your biggest challenge will be to get the tissue homogenized quickly.

A note on RNases: Autoclaving is not effective at eliminating RNases. Check:
http://www.ambion.co.../tb/tb_178.html

Also, even if autoclaving destroyed RNases, it would not be necessary to do. As tea-test pointed out, there should be very little RNases on your homogenizer, compared to what is present in your cells so autoclaving is not helpful here. Don't worry about using RNase Zap or any of that stuff on your homogenizer.

Another tip: When I do RNA work, I make sure I start with sterile/RNase free tubes. I take a whole bag and dump it out in the flow hood (not fume hood). Put on a clean pair of gloves. Cap all the tubes and store them capped in a jar. Now you have a clean supply of tubes ready to be used for any application, including RNA work. If you don't do this you may find yourself touching the inside of the caps when you close them when you go to label them. Your gloves will pick up RNases from the environment so don't consider them RNase free when you touch things.

How to avoid contamination in PCR?

Sat, 07 Aug 2010 20:51:01 +0000

PCR contaminations are like ghosts... you know they are there but you dont know WHERE??

I have followed the following with good results...

Prepare your master mix in a separate room from your current location, somewhere PCR is not the main practice... Be sure to have a separate lab coat, gloves, tubes, pipette tips tobe used only in that clean room.
Use a separate aliquot of DEPC water stock for each round of PCR
Prepare your mix in a hood with laminar flow. Decontaminate it with bleach, alcohol, RNAse, DNase, etc... Be sure to UV-irradiate pipettes, pipette tips, tubes, racks, gloves, and also your aliquots of water and PCR buffer... before the procedure.
Use a different pipette tip when pipetting all your reagents, even the same master mix to each tube
Keep your tubes closed during the procedure, even your master mix tube. Be sure that your tubes are closed when discarding the pipette tip!!! Aerosols are dangerous!!! Open the tubes only when necessary.
More important... schedule your PCR when not handling plasmids!!!

Hope this help...
Good luck

How to get rid of PCR primer dimer

Sat, 07 Aug 2010 20:48:39 +0000

1) Try the DMSO up to 5%. Some polymerases are ok with DMSO and others aren't. You never know until you try.

2) Try a two step PCR. There is a paper in Biotechniques about a two step PCR reaction but I don't have the reference with me since I'm at home and it's in lab. Its worth looking up. Basically you've got template plus forward primer in one tube and template plus reverse primer in another tube and then do between 1-15 rounds of PCR. Then combine the contents of the two tubes and do your normal 25 rounds of PCR. Sometimes this allows the primer to bind to the template effectively and allow you to get some initial transcripts which will then bind to each other when you combine the contents of the two tubes.

3) If you're not using a polymerase for high GC content, then get one. It will save you a lot of hassle. It won't solve your problem but it will help you narrow down things when troubleshooting. KOD XL polymerase (novagen) isn't sensitive to DMSO up to 5% and it helped me get all of my mutants but not without a lot of tries just because the GC content of primer and template were over 60%. Some people like the quik change kit by stratagene but I never got it to work, which is really expensive when its $600 a kit for 30 rxn or something like that. To each his own.

4) I wouldn't go below 55 deg for an annealing temp. 50 might be fine but below that you're pushing your luck with trying to have a high enough temp to resolve any hairpin structures which are common in high GC rich primers not to mention possibly decreasing the specificity of matching primer and template.

good luck! it takes a lot of troubleshooting and its hard to get consistent results.

Normalization for RNA or cDNA during two step R...

Sat, 07 Aug 2010 20:44:51 +0000

Curtis 20 April 2009 - 03:13 AM
Hello all,

I have a two-step SYBR Green RT-PCR kit for RotorGene. After I extract RNA from my samples, I proceed with cDNA synthesis right away without measuring the RNA concentration or without normalizing equal amount of RNA for all my samples. It is only after making cDNA that I measure the concentration and add around 100 ng in each PCR tube.

my results are a bit funny now and I get my first signal after 35 cycles and surprisingly the samples that should give signal sooner are always the last ones. my friends are suggesting to measure my RNA before proceeding with cDNA synthesis. They say maybe too much RNA in the sample does not let cDNA to be properly synthesized !?

is this comment true? Reply Edit
public/style_images/mobile/profile/default_thumb.png littleaxt 20 April 2009 - 05:42 AM
Hi

How do you measure cDNA concentration? Do you purify it before measureing it? Because I was told that if I measure with the Nanodrop I would also measure the unused dNTPs and therefore I could also start making it up :-)...

public/style_images/mobile/snapback.pngCurtis, on Apr 20 2009, 12:13 PM, said:

Hello all,

I have a two-step SYBR Green RT-PCR kit for RotorGene. After I extract RNA from my samples, I proceed with cDNA synthesis right away without measuring the RNA concentration or without normalizing equal amount of RNA for all my samples. It is only after making cDNA that I measure the concentration and add around 100 ng in each PCR tube.

my results are a bit funny now and I get my first signal after 35 cycles and surprisingly the samples that should give signal sooner are always the last ones. my friends are suggesting to measure my RNA before proceeding with cDNA synthesis. They say maybe too much RNA in the sample does not let cDNA to be properly synthesized !?

is this comment true? Reply Edit Delete
uploads/profile/photo-thumb-6817.jpg Curtis 20 April 2009 - 06:45 AM
good point, but if I purify the cDNa I'm gonna lose a lot of it during the purification steps. after all we are not talking about a PCR product that has been amplified. it's just cDNA from our RNA which is also not a lot. Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png gfischer 20 April 2009 - 07:45 AM
I always normalize the amount of RNA I use in my RT reaction. This way, you know that the only source of variation in the amount of signal in the quantitative step is from a higher level of relative expression of target RNA. Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png littleaxt 20 April 2009 - 08:48 AM
Oh, it seems possible, see http://www.biomedcen.../1471-2199/9/18 they also did it. Seems like cDNA clean-up also enhances the PCR.

But I never tried it myself, I normalize against total RNA prior reverse transcription and use relative quantification.

All the best

Jan

public/style_images/mobile/snapback.pngCurtis, on Apr 20 2009, 03:45 PM, said:

good point, but if I purify the cDNa I'm gonna lose a lot of it during the purification steps. after all we are not talking about a PCR product that has been amplified. it's just cDNA from our RNA which is also not a lot. Reply Edit Delete
uploads/profile/photo-thumb-6817.jpg Curtis 20 April 2009 - 09:07 AM
littleaxt: yeah, relative quantification. that's what I'm doing now. the link you gave doesn't open for me.

gfischer: that would be for one-step RT-PCR? or applies to two-step as well? how do you make sure you won't have more than 100 ng of cDNA in your sample? because if you have more than 100 ng the amplification might not work properly. Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png littleaxt 21 April 2009 - 12:04 AM
ok, this might help:

Libus & �torchov� 2006 �Quantification of cDNA generated by reverse transcription of total RNA provides a simple alternative tool for quantitative RT-PCR normalization� BioTechniques 41, 156-164

Liss 2002 �Improved quantitative real-time RT-PCR for expression profiling of individual cells� Nucleic Acids Research 30 (17): e89

Hibbeler et al. 2008 �Housekeeping genes for quantitative expression studies in the three-spined stickleback Gasterosteus aculeatus� BMC Molecular Biology 8:18

First is another idea you could try, second is a precipitation protocol for cDNA (never tried it myself) third is another paper where they made a cDNA clean-up prior PCR. Hope it helps, good luck!

public/style_images/mobile/snapback.pngCurtis, on Apr 20 2009, 06:07 PM, said:

littleaxt: yeah, relative quantification. that's what I'm doing now. the link you gave doesn't open for me.

gfischer: that would be for one-step RT-PCR? or applies to two-step as well? how do you make sure you won't have more than 100 ng of cDNA in your sample? because if you have more than 100 ng the amplification might not work properly. Reply Edit Delete
uploads/profile/photo-thumb-6817.jpg Curtis 21 April 2009 - 07:08 AM
ok thank you, I'll have a look now Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png Trof 22 April 2009 - 04:33 AM
We do two-step RT real-time PCR and always measure the amount of RNA only. Usually we put 1-3 ug of RNA to the RT reaction, depending how much cDNA we need. We don't measure nor clean-up the cDNA, use it directly to the real-time PCR.
We can dilute the cDNA if we need more volume for more samples (like 2 times or 3 times), but usually only when using more than 1 ug. Never exceed the 10 % of cDNA in a reaction mix. That makes from 1 ug in 20ul RT mix, when using maximum 2ul to a 20ul PCR reaction just about 100 ng.

I think we would see it on the aplification curve if there is any problem with high amount of cDNA, we never encountered such thing. Reply Edit Delete
uploads/profile/photo-thumb-6817.jpg Curtis 23 April 2009 - 10:18 PM
public/style_images/mobile/snapback.pngTrof, on Apr 22 2009, 03:33 AM, said:

We do two-step RT real-time PCR and always measure the amount of RNA only. Usually we put 1-3 ug of RNA to the RT reaction, depending how much cDNA we need. We don't measure nor clean-up the cDNA, use it directly to the real-time PCR.
We can dilute the cDNA if we need more volume for more samples (like 2 times or 3 times), but usually only when using more than 1 ug. Never exceed the 10 % of cDNA in a reaction mix. That makes from 1 ug in 20ul RT mix, when using maximum 2ul to a 20ul PCR reaction just about 100 ng.

I think we would see it on the aplification curve if there is any problem with high amount of cDNA, we never encountered such thing.

Thank you Trof,
your explanation was very helpful. Reply Edit Delete
uploads/profile/photo-thumb-10294.jpg Chimp 16 June 2009 - 12:54 AM
Checking [RNA] is an important step. You could also run a denaturing RNA gel to check RNA integrity, you should get clear rRNA bands Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png Freddynb 02 September 2009 - 05:43 AM
I have developed a method to amplify multiple random genes in a single real-time RT-PCR reaction to assess quality of cDNA between samples. It can be used to normalize gene expression between samples. Please see
http://www.springerl...28327382188710/ Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png sssbio 29 May 2010 - 03:32 PM
public/style_images/mobile/snapback.pngFreddynb, on Sep 2 2009, 01:43 PM, said:

I have developed a method to amplify multiple random genes in a single real-time RT-PCR reaction to assess quality of cDNA between samples. It can be used to normalize gene expression between samples. Please see
http://www.springerl...28327382188710/

Hi,

I need your help regarding quantification of mRNA levels of trageted genes in control and treated samples. I have no choice of reference housekeeping genes to be used for relative quantification as these genes are showing significant down regulation in treated samples. So what are other methods that I can use to quantify mRNA levels?

I will be very happy if you guide me for this.

Thanks Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png gradechips 02 June 2010 - 11:05 PM
Please help me, I have been doing RT-PCR for the past six months and i still have problems. During this period i was only able to get decent Cts for the house keeping gene (18) twice but all the other times the Cts for both GAPDH/beta actin have been really high 25 or inconsistent within triplicate samples (20, 24, 29). Obviously i cant use these data to analyze gene expression and am getting frustrated. After extracting RNA using Trizol i quantify it by ODing and i usually get ~2.500ng/ul and i use 2ug for the cDNA synthesis and i normally dilute the cDNA 1:50. My lab mate normally gets low and consistent Ct for the house keeping gene using the same reagents. I have been trouble shooting for the past 6 months with his help but am not getting it.
Is it because vortexing shear my samples? Please someone help me.
Thanks in advance Reply Edit Delete
public/style_images/mobile/profile/default_thumb.png Nataliya 24 June 2010 - 01:19 AM
public/style_images/mobile/snapback.pnggradechips, on Jun 2 2010, 11:05 PM, said:

Please help me, I have been doing RT-PCR for the past six months and i still have problems. During this period i was only able to get decent Cts for the house keeping gene (18) twice but all the other times the Cts for both GAPDH/beta actin have been really high 25 or inconsistent within triplicate samples (20, 24, 29). Obviously i cant use these data to analyze gene expression and am getting frustrated. After extracting RNA using Trizol i quantify it by ODing and i usually get ~2.500ng/ul and i use 2ug for the cDNA synthesis and i normally dilute the cDNA 1:50. My lab mate normally gets low and consistent Ct for the house keeping gene using the same reagents. I have been trouble shooting for the past 6 months with his help but am not getting it.
Is it because vortexing shear my samples? Please someone help me.
Thanks in advance

I usually use RNeasy Mini Kit (50) and RNase-Free DNase Set (50) from Qiagen. Once I have got RNA extracted I use it for cDNA synthesis or keep at -70 C. For cDNA synthesis I used SuperScriptTM III First-Strand Synthesis System for RT-PCR (Invitrogen, Cat. No: 18080-051). For RT-PCR I diluted cDNA 1:5. Sometimes I measured cDNA by nanodrop and the concentration of the diluted cDNA samples was around 200 ng/ul I have never faced the problem with low values for house-keeping genes. I normally get 13-14 Ct for GAPDH. Probably, you don't need to dilute samples up to 50 ul/ml.

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!

Useful Dilution Techniques & Calculations

Sat, 07 Aug 2010 05:55:42 +0000

Today was a very bad day at the university... beside that some students answered in the quiz question that if you get blue color in Gram staining that means your bacteria is Gram negative I noticed most of the students including myself had trouble with the dilution process. So I thought maybe I'd make a quick explanation for what is dilution and how it is done and all the ratios and other confusing stuff. I tried my best to simplify it for people who find trouble in doing their homeworks, labwork, or other calculation problems regarding dilution questions and solutions.

So let's start out with definitions:

Dilution: is the mixing of a small accurately measured sample with a large volume of sterile water or normal saline called (diluents or dilution blank)

Laws:

Dilution = V of Sample / Total V of (sample + diluent)

Dilution Factor = Total V of (sample + diluent) / V of sample
** or we can simply say the reciprocal of Dilution

Starting off with this simple example to understand how these laws are applied.

It has been known that if we use a larger volume we obtain a more accurate dilution.
So for better results, we use 1:1000 dilution. And that is by adding 1ml of sample to 999 ml of diluent. But practically we cannot use 999 ml of diluent. So we do what is called a serial dilution.

Serial Dilution: is a dilution made of a series of smaller dilution, and the total dilution is the product of each dilution in the series.

To understand this more, let's see this example.

Here's an example combining all of the above.

I hope this brief explanation proves helpful to you...let me know if i've made any mistakes...

-Property of Yulia-
thank u for your explanation..

what makes me always confused is the last volume used for inoculation (0.1ml in your example)..

-strawberry-

Yeah that's why i almost jumped with 'HAVE' there :blush: sorry about that
i hope my mini-tutorial and examples helped you through it strawberry...

-Property of Yulia-

I'm very thankful to you to provide the easier explaination...it's true that many undergrad students have problem in dilution..and the worst thing is if the lecturers/tutors assume that the students are familiar with the concept. it is true that they have learned it in high school, but most of the time they will forget..

Regarding the example 2 given, you dilute a sample (10 ml the final volume) 10x10x10=10 to the power of 3. and u inoculate 0.1 ml in your medium.so supposed the final volume of your medium is 1ml right?

-kent19-

How to convert g into rpm?

Sat, 07 Aug 2010 05:54:42 +0000

The relationship between revolutions per minute (RPM) and relative centrifugal force (xg) is:
g = (1.118 � 10^-5) R S²
where g is the relative centrifugal force, R is the radius of the rotor in centimeters, and S is the speed of the centrifuge in
revolutions per minute.

You can use this for any centrifuge, just measure the radius of the rotor from the center to outer edge.

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