Posted :10/7/2008 7:24:00 PM
Hi Ansh,
Your reasoning is exactly right. Migration and invasion assays do measure different activities. The typical migration assay, which measures the number of cells traversing a porous membrane, can be used, for instance, to evaluate chemotaxis. Chemotaxis can occur entirely in the fluid phase without need for degradation of basement membrane or extracellular matrices. Invasion assays, on the other hand, are used to investigate cellular properties involved in such phenomenon as tumor progression and metastasis. For these activities, extracellular proteases are frequently required to degrade protein-containing barriers to migration.
A good description can be found here:
link
Posted :10/7/2008 7:34:00 PM
It sounds like the new prep may have some contaminant that is inhibiting the PCR reaction. You might want to try cleaning up the DNA with a phenol/chloroform extraction or a clean-up kit of some sort. It would also be good to sequence the band you get just to make sure it's what you think it is. Finally, is there some housekeeping gene you could amplify to check things out? For RT-PCR of mammalian mRNA, I routinely run a PCR with GAPDH primers to see if the cDNA reaction worked. If I get no band, I reprecipitate the RNA and try again. If still no band, I toss it and start over.
Have fun.
mike
Posted :10/7/2008 7:44:00 PM
Protease activity in the tissue prep could explain both the contamination problem and the cleaved Lamin A problem. Are you using a protease inhibitor cocktail?
Posted :10/7/2008 7:52:00 PM
The reason ELISA kits mention azide is that it's an irreversible inhibitor of HRP. It doesn't matter if the azide is in the Ab solution since it will be washed away well before the HRP step. However, it should not be in the wash solutions. Same goes for immunoblots with HRP-conjugated secondary Abs. As far as azide inhibiting Ab-antigen interaction, sure it's possible. However, it would probably have to be a very specific issue with a particular Ab-antigen pair. Otherwise, azide would never be recommended as a preservative for Abs.
Posted :10/8/2008 8:43:00 PM
It sounds like you have something in your set of reagents that the others in the lab don't. You could run someone else's RNA and primers (that work) using your reagents; and have someone in the lab whose reactions work run their primers on your RNA using their reagents. Basically, you want to find out if the problem is in your RNA or your reagents.
Posted :10/8/2008 8:51:00 PM
How dilute are you making the cDNA? It could be that you are losing most of it to the sides of the tube. Although plasmids can usually stand to be stored at relatively dilute concentrations, cDNA (and protein) are different. I would suggest only diluting enough cDNA for the current qPCR run. Toss the remainder of the diluted sample and store the concentrated sample. I have frozen (-20C) and thawed relatively concentrated cDNA multiple times over many months and have had no problem.
Posted :10/8/2008 9:05:00 PM
Hi,
I have done some proliferation assay using MTT kit. I use siRNA to knockdown a particular protein in glioma cell lines and then checked the effect on cell proliferation. I found a decrease in cell proliferation after siRNA knockdown. Now I want to see if this decrease in proliferation is due to increase in number of dead cells (because of apoptosis??) or due to decrease in the growth of cells ie cells now divide much slower after knockdown.
Can someone share some protocol to do it?
any help of suggestion would be appreciated.
Cheers!
The nuclear antigen, Ki67, is frequently used as a comparative measure of proliferation. It is more abundant in cycling cells. The paper referenced here
link describes an ELISA to quantify this protein. You could also look at the level of apoptosis in the cells. There are numerous ELISAs designed to measure various stages of apoptosis. (I'm assuming you've ruled out any effect of the procedure itself on the decrease in cell number. I.E. did you check the transfection method alone and use a scrambled siRNA as negative controls?)
Posted :10/10/2008 4:15:00 PM
Hi everyone,
Can anyone tell me about the colony forming assay? What does it tell us and which kind of agar do we use for this assay.
i am working on some glioma and melanoma cell lines and checking the effect of knockdown of a particular protein in these cell lines. will this assay tell me something useful.
AA
This assay is most frequently used to evaluate how "transformed" or neoplastic a cell is; i.e. how many features of a malignant cell does it have. Compared to non-transformed cells, neoplastic cells are much less contact-inhibited, exhibit anchorage-independent growth, and can proliferate in the absence of exogenous growth factors. In the assay, a single cell suspension of cells is mixed with molten soft agar at 37C and then poured over a base layer of agar. The cells are then allowed to grow in a conventional CO2 incubator. Depending on how rapidly the cells grow, it may take 2-4 weeks to get decent-sized colonies to count. The presence of colonies, as opposed to single cells or tiny colonies consisting of just a few cells, indicates that the cells at least have some of the characteristics of transformed cells. The literature is loaded with protocols for this assay. Agarose as well as different types of agar can be used for the assay. The following was taken from Oncogene. 1999 Nov 11;18(47):6469-76:
"For anchorage-independent growth in soft agar, 3´104 cells were resuspended in a final volume of 2 ml containing 0.3% agar Noble agar (DIFCO, Detroit, MI, USA) in melanocyte growth medium. Cells were added to 6-well plates (Falcon; Becton Dickinson Labware, Lincoln Park, NJ, USA) containing 1 ml/well of 0.5% agar in melanocyte growth medium. After 5 days, new growth medium in agar was added and 5 days later, colonies of four or more cells/colony were counted in five random fields."
Posted :10/13/2008 8:25:00 PM
Has anyone had a bad experience with buying precasted gels and using them in another companies gel tank? I'm looking at a agarose system for DNA analysis but I don't want to get stuck with something that limits the brand of precast gel available. Does anyone know which companies offer the most compatible gel box or gels for that matter?
If you really don't want to be limited, pour your own. That way you can choose the agarose, the %, the dye, the size, and the well number. Pre-cast SDS-acrylamide gels for proteins are much more commonly used than pre-cast agarose gels because pouring your own SDS-PAGE gels is a major pain. Also, unpolymerized acrylamide is toxic and it's very difficult to pour gradient gels. On the other hand, agarose gels are extremely easy to pour and the components are not toxic.
Posted :10/15/2008 2:55:00 PM
hi, all,
I am looking for the technology, which help me to solve a trouble technology.
I am want to use in situ hybridization method to detect the co-localization of two mRNA in tissue section.
these two mRNA are splitted from one DNA sequence, and differs only for one exon.
does anyone know any sensitive and successful method to solve the problem??
thanks a lot........................
my email: roy_liu@genetimes.com.hk
Roy,
You might want to add some details to your question. For example, will you be using frozen sections, formalin-fixed paraffin-embedded sections, or cells? The protocols differ somewhat for each of these. Also, are you asking about probe design? When using a sensitive detection method, such as enzymatic signal amplification, ISH probes can be as small as 16 bases or as large as 500 bases. With direct fluorescent labeling of probe molecules, which would easily allow colocalization experiments through the use of different fluorophores, the probes are frequently fairly large in order to increase the signal intensity. There is also the question of probe chemistry; i.e. DNA, RNA, or PNA.
Basically, you will most have to employ 2 probes, one that hybridizes to the exon, and the other that spans the junction created when the exon is absent. Fluorescently labeled probes would be the most direct but may not be sensitive enough to detect your message. Otherwise, you will need to probe serial sections with each probe, use some sort of an antibody- or biotin-based detection system, and overlay the images to obtain colocalization information. The internet is replete with in situ hybridization protocols and most are pretty straightforward.
Posted :10/15/2008 8:46:00 PM
Hi,
I recently started working with 3T3 cells and noticed that when I plate them in wells containing coverslips, most of the cells grow (and crowd) on the plastic well surface, and very few on the coverslip. These are high quality glass coveslips (EMS) that we also use for primary cultures, so I assume they should not be the problem. The coverslips are extensively washed in water and sterilized with 70 % EtOH before plating, but have not been coated in any way. I've been using them to grow a variety of other cell lines (HeLa, Cos7 etc.), and never seen anything similar.
Do 3T3 cells need a coated surface?
Thanks!
Phyll
They may. Whether a cell line adheres to an uncoated coverslip or not depends entirely on the cell line. I've used 2 derivatives of 3T3 cells, as well as a lung adenocarcinoma line and a large-cell lung cancer line, and all grew very well on the coverslips without coating. It could also be partially dependent on the brand of coverslip. I have used VWR #48366227 with good results. You might want to try a different brand, or just try coating your coverslips with poly-L-lysine (Sigma P4707, 0.01% (w/v) solution). After sterilizing the coverslips with EtOH, cover with the poly-L-lysine and let sit for 1 h at room temp. Then, aspirate and do 3 x 5 min washes with sterile water. Best of luck!
mike
Posted :10/20/2008 3:34:00 PM
Hello all,
I need some help with understanding immunoprecipitation. For instance I IP'ed protein A, did the entire protocol, ran a SDS-PAGE, transfer, then i blocked.....and finally i probed with proteing B because I wanted to see if it bound with protein A. When I expose am I suppose to see just protein A bands, only protein B bands, or both protein A and B bands?? Please help
Both proteins, A & B, should be on the blot. Which one you see depends on what antibody you use to probe the blot. If you IP with an anti-A antibody, it should bring down both protein A and whatever binds to it, which you hope is protein B. When you run the IP on the gel and blot it, you should have protein A and whatever other proteins bind to it. So, if you probe the blot with an anti-A antibody, you see protein A. The same goes for probing with an anti-B antibody; you should see B.
Some caveats: If the antibody you use for doing the IP is from the same species as the antibody you use to probe the blot, you have to be careful how you perform the experiment in order to avoid huge contaminating bands of IgG on the immunoblot. For example, if you IP with a mouse antibody and bring down the antibody-antigen complex with protein A-beads, you are also bringing down the mouse IgG, which will end up on the immunoblot and be recognized by the anti-mouse secondary antibody. There are a couple ways around this: One is to use antibodies from different species to do the IP and the immunoblot. Another way is to covalently attach the IP antibody to beads so it doesn't end up in the mix of what gets loaded onto the gel. Some companies also carry secondary antibodies that only recognize native IgG, thus making the denatured and reduced IgG on the blot invisible. Pierce sells one (cat.no. 21230).
Another general caveat is that even though proteins A & B may interact in vivo, they may not in vitro if the IP antibody interferes with the interaction. Finally, you will need to perform solid negative control IPs to rule out the presence of protein B in the protein A IP being due to non-specific binding. Immunoblots are very sensitive and can detect proteins that are just along for the ride. One common control is to denature the proteins in the lysate prior to adding the IP antibody. This should disrupt the A-B interaction but not the antibody-protein A interaction, provided the antibody works with denatured protein A.
Posted :10/21/2008 2:07:00 PM
Does anyone know why cells clump in the middle of the well and how this can be prevented? I am trying to transfect primary lung fibroblasts in a 24-well plate and I believe that the over-confluency in the center of the well is inhibiting the transfection. I've noticed that as I get higher in passage number, the cells take longer to trypsinize - is it possible that the longer trypinisation time could lead to clumping? Is there a trick to stopping this?
Thanks...
The most frequent cause of cells clumping in the middle of the well is swirling the plate after the addition of the cells. This causes all the cells to go to the middle of the well. It's better to gently shake the plate from side to side, in both directions, to distribute them evenly. It also helps to have sufficient medium in the wells as the surface tension can also cause uneven distribution.
How many passages are you carrying out with the cells? Primary lung fibroblasts are guaranteed by Clonetics for only 15 doublings, which is only 3-4 passages. After that, they begin to either die or senesce. Whatever they do, they are no longer classical primary lung fibroblasts.
Posted :10/22/2008 1:57:00 PM
We are currently using caco2 cells that were cryopreserved 10yrs ago, at passage 70-80. We would like to know if this may be problematic in terms of protein expression etc. given the high passage number and age of the cell line. We are contemplating purchasing new cryopreserved cells (of a lower pasage number) and would like to know if anyone can suggest a good suppplier for the UK.
The effect of passage number on the characteristics of the cells is entirely dependent on the particular cell line in question. There's no hard and fast rule concerning what passage number is "too high." That being said, it's good that you are thinking about this since passage 70-80 may indeed have resulted in changes in the caco2 cells. ATCC carries these cells and they can be obtained in the UK from:
GC Standards
Queens Road, Teddington
Middlesex TW11 0LY
United Kingdom
Tel: 44 (0)20 8943 8489
Fax: 44 (0)20 8943 8405
Email: atcc@lgcstandards.com
Technical Inquiries Email: atcc-tech@lgcstandards.com
Website: www.lgcstandards-atcc.com
Posted :10/22/2008 2:24:00 PM
I was wondering if there was any type of DNA in which two pyrimidines bonded with one another?
Thanks.
Not that I know of. The pairing of purines with pyrimidines ensures that the DNA helix will be of uniform diameter. If 2 pyrimidines bonded with each other, there would be a bulge in the helix and probably quite a bit of strain. You might be able to force a pyrimidine-pyrimidine pairing in vitro but it would most likely get repaired in vivo.
Posted :10/23/2008 2:17:00 PM
Please can I ask why TEERs of 900 isn't good enough for transport studies ? We are using cells with lower TEER values than this, with cut-off of 500 after experiments. Is this too low ?
This article contains an excellent explanation of the entire procedure: Nature Protocols 2, - 2111 - 2119 (2007)
Published online: 23 August 2007 | doi:10.1038/nprot.2007.303
Determination of drug permeability and prediction of drug absorption in Caco-2 monolayers
Ina Hubatsch1, Eva G E Ragnarsson1 & Per Artursson1
This is from that article: TER values of the Caco-2 monolayers might depend on the Caco-2 clone used, on the number of days after seeding and on the passage number and culture conditions (including the material the filter support is made of). In our laboratory, standard TER values at 37 °C are 260 +/- 65 ohm cm2. Cell monolayers with TER values below 165 ohm cm2 are discarded.
Hope this helps.
Posted :10/23/2008 2:21:00 PM
We know that plates should not be seeded until the 2nd tripsinisation, but can anyone explain why we can't do it on the 1st ?
Please see:
Nature Protocols 2, - 2111 - 2119 (2007)
Published online: 23 August 2007 | doi:10.1038/nprot.2007.303
Determination of drug permeability and prediction of drug absorption in Caco-2 monolayers
Ina Hubatsch1, Eva G E Ragnarsson1 & Per Artursson1
Here's a quote:
Critical Use only about ten passages (i.e., cells within a defined interval of passages). We use Caco-2 cells of a high passage number (95–105), but it is possible to use cells of lower passage numbers (e.g., 40–50). The phenotypes of Caco-2 cells taken from high and low passage intervals will differ.
Critical After thawing a vial of Caco-2 cells from the stock,
cultivate the cells for two passages before seeding cells on the filter supports to stabilize the cell phenotype.
Posted :10/23/2008 2:23:00 PM
how i can get research articles from website?
It depends mostly on which journals your institution subscribes to. Many journals offer abstracts for free but you either have to pay for each article, be a subscriber, or be a member of an institution that subscribes. Then you can download the pdf.
Posted :10/27/2008 7:19:00 PM
Dear Sir, I am growing Mesenchymal stem cells on scaffold and now I want to do confocal microscopy to see the cell attachment and morphology.can you suggest me which antibody or kit or fluorochrome should I use to do the same.
please help?
If you just want to see morphology, you could use Differential Interference Contrast (DIC). No stain is used. Using an antibody will limit you to visualizing a single protein and wouldn't be very helpful for looking at morphology. If you had a protein in mind that was involved in the cell attachment, you could try an antibody against that but the DIC would still probably be more informative.
Posted :10/31/2008 6:18:00 PM
Does anyone know the best place to look for the following equipment:
Microscopes (plus counting slide with the criss-cross boxes) suitable for looking a Caco2/HEK293 cells
Vaccum pumps plus galssware attachments (for media removal using glass pippetes)
Voltohmeters with the 'chopstick' attachment that can fit 12-well plates
A general lab supplier like VWR
link will have everything with the possible exception of the voltohmeter. They have international offices as well. Millipore has a voltohmeter
link, also World Precision Instruments has a couple
link