Trypsinization or Scraping for adherent cells in ChIP assay? - (Sep/19/2010 )

hi!

we're working on h9c2 rat cardiac myocytes(adherent),trying to standardize X-chip by Ez-ChIP protocol,though we've not bought the kit.


which one is the best?
a)i trypsinize my cells first and then add formaldehyde?
b)add formaldehyde first and then go for trypsiniztion?
c)add formaldehyde and then scrape cells off?

the b part is a bit tricky since the cells just don't come off after trypsinization.probably they bind to the poly lysine bottom or something else.

thanks for any help that you may offer!

cheers
sameer

The standard protocol is to add formaldehyde, then glycine for neutralization of the formaldehyde and then scrape the cells in PBS with protease inhibitors.

yup..the problem is we're low on resources.so would better try to trypsinize and use the flask again rather than scrape and throw away the flask.

i did find some answers here:http://www.protocol-online.org/forums/topic/9525-getting-cells-from-flasks/
but..
still my saving on the flasks question remains.

sameerbau on Sun Sep 19 17:43:03 2010 said:

Most do not use trypsin as there is a potetntial risk that it may alter protein-DNA interactions. For the price of a flask (I mean, really?) it is not worth it in my opinion. What if you complete your experiment only to find out a year later that your trypsinization introduced some bias into your whole study? At the very least, you should compare both methods and if there is no difference, then you can re-use as many flasks as you like.

You really should not be re-using flasks anyway. It is not good cell culture practice. If you can't afford the flask, you probably shouldn't be doing any culture at all.

Do follow the classic protocol by scraping. Once your sample is safely on the ChIP track, you can try to save your plates: 4hrs at 65°C to decrosslink the remaining cell debris from the plate and then trypsin as you would do.

thank you all!

we did agree on crosslinking-quenching-scraping in PBS for our cell extraction.

1.i'm concerned with the low cell yield..i mean..everyone says they start off with atleast 10 million or so cells.the problem is we hardly reach 2.5million in out T75s.
would scaling up to t150 be an option?like would it really increase the cell number to atleast 10 million.i say..for a low budget lab like ours..thats a task.

secondly,about the phenol chloroform.

we use around 2.5 million cells,add 500ul of sds lysis buffer,followed by 500ul of phenol choloroform(250ul each).Take the around 500ul aqueous layer,add 500+500ul 100%ethanol and 150ul NaCl,spin down,wash with 70%ethanol (1000ul),spin down.dissolve in 20ul TE.

the problems:
1.we use 2ml eppys.till the time we're using 500ul of SDS lyis,things are ok.if we go for 1ml SDS lysis buffer,things have to be aliquot-ted (1ml SDS lysis+ 2volumes of 100% ethanol=3ml)
is there a way around that?

2.as sambrook recommends,after the last 70% wash,we run the 20ul TE up and down the tube for dissolving the DNA..which seems a bit unscientific.is it really necessary?is there an alternative?

thanks for your inputs
sameer

i would really appreciate some advice!

Usually I trypsinize the cells and do the cross-linking in suspension, just because I get better yelds at qPCR and the results seem to be more consistent this way.
(a long time a ago I did some side-by-side comparisons and this was what worked better in my hands.

The SDS volume can be changed, you just have to reoptimize the sonication conditions (this way you can keep the volumes within a manageable level) or you can split the sample in more eppendorfs.

The TE step is important, some times you even have to warm up the sample, when the DNA pellets get to dry it is close to impossible to re-dissolve them (I just changed to mini-elute columns, it was more expensive to re-run the ChIP because of a bad PC extraction than to just buy the columns and get it right the first time)


Hope this helps