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There have been 6 items by B.B. (Search limited from 18-April 20)


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#128584 Can you freeze-thaw DMEM and it still work?

Posted by B.B. on 07 February 2012 - 09:33 AM in Tissue and Cell Culture

Hey guys,

many thanks for your comments.

Perhaps I didn't make it clear but I would essentially be making a large volume of the peptide/media mix; aliquoting this mixture; freezing these down and then thawing out the required aliquot when needed - so just freezing and thawing once as 'almost a doctor' suggests :)

So the "one round" of freeze-thaw of the 'peptide/media mixed aliquot' should be fine.

Just to be on the safe side I will research a little more into the solubility of my peptides and see if I can find a different solvent to dissolve them in to.
:)



#128570 Can you freeze-thaw DMEM and it still work?

Posted by B.B. on 07 February 2012 - 08:28 AM in Tissue and Cell Culture

Hi,

Odd question - but hear me out! Posted Image
I'm currently testing the effect of some peptides on the migration of 3T3 fibroblast cells over a scratch wound.
I need to incubate my cells in DMEM media containing the peptide (which is purchased as a powder) for a couple of hours before assaying my cells.

The concentrations of the peptide in the media is really quite small (i.e. 100uM) and so I'm measuring out miniscule amounts of the peptide powder to dissolve into the media - which is leading me to believe that I'm losing some accuracy along the way.

An obvious solution would be to dissolve more powder into a larger aliquot of DMEM media = less room for error.
But I won't be using the peptide/media combination that often and I don't want to just throw it away when my media naturally degrades as the peptide is very expensive.

So I would naturally want to freeze my DMEM/Peptagon combination (in aliquots to thaw out when and as needed).

BUT - Can you freeze-thaw DMEM and it still work optimally?
I can't find anything to say you can't - so I'm guessing it's OK....



#113790 Housekeeping protein issues

Posted by B.B. on 29 June 2011 - 05:04 AM in SDS-PAGE and Western Blotting

Hello,
this is an unusual problem, but there may be someone out there who has been in a similar situation.

I have an experiment where I induce hypoxia on cell samples, collecting them at specific time points over 24 hours, and measuring a particular connexin level in them.
I measure the protein concentration of the samples using a Bradford Assay, to ensure that I have the same level of protein in all my lanes.

In my control samples (no hypoxia), my housekeeping antibody (alpha tubulin) levels are pretty similar over the 24 hour period, reassuring me that I do have similar levels of proteins in my lanes.

However, in my hypoxic samples, this is not the case. The levels of alpha tubulin rise progressively over the 24 hour time period, looking like I have not considered protein concentration over time.

Is there a slim possibilty that my house keeping antibody is being affected by the hypoxic nature of my experiment? I have heard that alpha tubulin is a pretty reliable house keeping antibody, so I don't know if this will be the case.
I have triple checked my calculations of my protein concentration (and will be doing another Bradford assay in the near future), but I really don't think that I have made a mistake here.

If anyone else has come across their housekeeping antibody being affected by their experiment, please could you throw some advice this way!
I guess one obvious option that I have is to change my housekeeping protein - but I don't want to run into the same problem.

Thanks in advance
Bev



#113783 Help! Half membrane exposed perfectly. Half membrane is cloudy

Posted by B.B. on 29 June 2011 - 04:40 AM in SDS-PAGE and Western Blotting

Hi,
thanks for the reply.
It turns out that my secondary antibody for my housekeeping protein was dud!
I had been given an aliquot of it from another post-doc, which I stored in the freezer. Turns out it can only be stored at 4*C! Whoops! So when I defrosted it to use on another Western Blot, my blots were fuzzy and messy.
Never mind - lesson learnt. And at least it was only a small aliquot :)
Bev



#111698 Oxygen Probe

Posted by B.B. on 03 June 2011 - 12:14 AM in Biochemistry

Good Morning,

I'm measuring the oxygen concentration of samples using a FOXY probe, and have been instructed through the manual to use sodium hydrosulfite (also known as sodium dithionite) to create a 0% standard, so I can create a calibration curve.

My problem is, that I don't know how much to use. From what I have read, this white crystaline powder is a strong reducing agent and removes oxygen from solution. Made fresh is can last up to 24 hours. From the research I've done online, I can't seem to find a standard weight to use.

One paper I came across quoted that they used 1M (so with MW of 174.11, and volume required 20ml, that's 3.48g).
Whereas another paper stated that they used a 2-5% solution (again, in 20ml, that's 0.4-1g - nearly x9 difference!)
Again, another paper uses just 0.2g in 20ml.

My ultimate question: can you use too much sodium hydrosulfite? I guess the most you could do is 'overstaurate' the solution, and you can't have a negative oxygen level.

Has anyone else used this chemical to create an 0% O2 standard before? What values did you use?

Thanks!
Bev



#110344 Help! Half membrane exposed perfectly. Half membrane is cloudy

Posted by B.B. on 20 May 2011 - 04:33 AM in SDS-PAGE and Western Blotting

Hello,

I'm new to this forum and also to Western Blots.
I've been following a protocol given to me by a post doc and am having a little trouble with exposure.
Here is the protocol in breif:

Cells are grown and harvested with lysis buffer. The sample is homegenised, spun down and collected.
The samples are run on a lab-made 10% gel and transfer is done on ice at 100V.
I check for protein transfer for Ponceau Red (which was positive) and I cut my membrane according to MW (referring to the ladder) so that I am able to stain for two proteins (housekeeping proteins and protein of interest).
The membrane (nitrocellulose) is blocked in 5% milk for 1 hour at room temp.
The first antobody (made in 5% milk) is incubated on a rocker at 4*C overnight.
The second antibody (also made in 5% milk) is incubated on a rocker at room temperature for 1 hour.
All washes are done with TBST. I aim for at least 5 washes for 5 minutes for each membrane.
I use West Pico Pierce Chemiluminiscent for developing.

The problem I am experiencing is that one half of the membrane (with the protein of interest on it) is exposing perfectly. The other half (with the housekeeping antibodies) is messy, unspecific and prdocuing a black outline on the membrane.

As I treat these two half of the membranes exactly the same, I can only think it's the antibodies I use for the housekeeping protein.

The concentrations I ues are:
HOUSEKEEPING PRIMARY 1:500, SECONDARY 1:1000
PROTEIN OF INTEREST PRIMARY 1:4000 SECONDARY 1:1000

Has anyone else had this very unusual problem? Or thinks they may be able to shed some light on it!

Bev




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