antibody titrations / controls
The missing sentence was: 'In addition a lot of antibodies also bind on other cells and the amount of active antibody in the bottle might vary. Therefore you should have approximately 10^3 fold excess.... Antigen concentration is indeed only of relevance with respect to depletion. Regarding the variations in the binding kinetics we have some applications where we want to stay below 5% variation. The higher the excess of antibody,the better. We use goat anti mouse*PE Fab2 at 12ug/ml and have to run at least for 30 minutes to saturate and to keep the day to day variations between the samples low. This might be due to a lower affinity or the polyclonal nature of the antibody. As most of our samples have a normal distribution of antigen load, we estimate the degree of saturation by the shape of the distribution in the fluorescence histogram.
Gerhard Nebe-v.Caron,
Unilever Research, Colworth,
I just want to add my annual reminder that the concentration of antibody to use in a staining mixture is essentially independent of the number of cells being stained, but is completely dependent on the stain volume! 1 ug of antibody is more than enough for (in general) well over 10^8 cells expressing a typical antigen--thus, it doesn't matter if you have 10^5, 10^6, or 10^7 cells in your tube! Indeed, many antibodies will stain equally well at 10^8 cells (in, for instance, 100 ul volume). On the other hand, a reagent titred to be used in a 50ul stain volume may not stain very well in 500 ul. Specifically: 1 ug of antibody (IgG) is about 3.6 x 10^12 molecules--which (in divalent fashion) binds 7.2x10^12 molecules of antigen. A typical antigen may be present in concentrations of 10^4 to 10^5 per cell, meaning that 1 ug of antibody would be enough to stain 7.2 x 10^7 to 7.2 x 10^8 cells. And, if only 10% of the cells express an antigen, you can multiply this by another factor of 10!
mario roederer, stanford
Actually, isotype controls are not a particularly good control. They are rarely matched: the F/P ratio is not the same, and how do you know if you are using them at exactly the same concentration as the reagent of choice? If you don't know that the F/P ratio is exactly the same, and if you don't know if you are using it at exactly the same concentration as your antibody reagent, then it isn't the right control. Indeed, since each one of your commercial reagents is titrated by the manufacturer to give optimal signal to background, each one is sold at a different concentration of antibody. Have you contacted the manufacturers to determine the bottled concentration of each reagent you use, so that you can use the appropriate concentration of your isotype control? And then use a different isotype concentration as the control for each reagent in your various panels? If the answer to either of these questions is "no", then how can you assert that your isotype control actually gives you the correct amount of background binding in your experiment? i.e., your "isotype control" does no more than let you that there may actually be some background binding, but doesn't give you the ability to estimate how much. In fact, I've known people to "titrate" their isotype controls to get background binding that is less than what they think their positive should be. Hmm. In another way, isotype controls are rarely used properly: most people do a single sample that has all isotype controls in all channels. This doesn't help! One must use a control for which cells are stained with all reagents EXCEPT the one of interest (and if you insist on using an isotype control for that channel, so be it). We term these controls "FMO" or "Fluorescence Minus One" controls. (For more discussion of the need of FMO controls, see my paper on Compensation in the upcoming issue of Cytometry). Staining controls are very difficult to generate. In general, the best control for antibody binding is a cell that is exactly the same as your cell of interest, but lacking the antigen of interest. Of course, this is rarely achievable. However, one will often find very similar cells that meet the bill. In immunophenotyping of peripheral blood, you can use "nonexpressing" cell types as internal controls (i.e., naive T cells can serve as a control for measuring activation markers on memory T cells). Of course, you need to be careful, because some "nonexpressing" cells actually express the marker. Isotype controls have their place. However, most people don't use them properly. In general, I counsel people NOT to use isotype controls, but rather to use their brains to come up with a set of appropriate negative controls (which MUST be included in all experiments, as others have noted). Blind reliance on isotype controls is one of the most common mistakes in publications--and leads to the erroneous placement of gates. In any case, you are correct that investigators need to be educated more. This is one of the discussions that pops up every few years on the mailing list; perhaps it's time to have a FAQ's page (no pun intended!) on the Purdue site which includes the various discussion points, and rather than coming up with a conclusion, this page can simply serve to put forth various peoples' views so that researchers can judge for themselves whether or not isotype controls are useful. mr (PS, there is no such thing as "bad data", only "bad interpretation of data.").
mario roederer, stanford
Yes... much of what you say would be true... if the use of isotype controls were scientific. But, as has been endlessly discussed on this list, they are NOT. (See also: O'Gorman MRG, Thomas JA: Isotype Controls-Time to Let Go?; Cytometry 38:78-80, 1999.) Proper science is to use a control (compared to a test sample) in which only a single variable has been changed. In the case of isotype controls, this single variable is supposedly the specificity of the isotype for the epitope. However, unless you go to great extents (and I guarantee that you have not), there are several more variables in your experiment. (1) The concentration of the isotype. If you have not matched the concentration of the isotype control to each of your antibodies (and each antibody, incidentally, is used at a different concentration, so you better be using isotypes at different concentrations for each antibody you are controlling), then the isotype control is no longer a control, but another test sample. (2) The Fluor:Protein ratio (F/P) of the isotype. If the F/P ratio is different than the test antibody (and how are you ever going to know this, unless you make both antibody and isotype control yourself?) then again, you have different variables. An isotype with a higher F/P than your test antibody, even if used at the same concentration, will give you higher "background" fluorescence. (3) Sequence-specific "nonspecific" binding. Of course the isotype has a different peptide sequence than the test antibody... are you really sure that none of these amino acid differences don't contribute to some selective binding? Until you prove that your isotype control has the same F/P ratio, and that you are using it at exactly the same concentrations as each of your test antibodies (that's a lot of isotype control stains!), then your "control" is no more than another test sample. That's the kind of scientific evidence you need to provide before you can use an isotype to determine positivity in your sample. Isotype staining certainly has its place. It can indicate IF there is an issue with background binding. It can let you know that perhaps you should be careful about interpreting your staining. And this is particularly true for myeloid cells that have high levels of FcR. But the problem is that most people go beyond this "canary in the cave" use for isotypes, and use the isotypes to set boundaries for gating and identification of positive vs. negative. And that is where the isotype ceases to be science. You are correct that we are not in the business of making things easy. And this is the insidious nature of isotype controls. They let people think that they are doing something "easily", when in fact they represent only a crutch that is being improperly used. They lull researchers into thinking that they can now identify positive vs. negative. It's so comforting to think that you have an appropriate control for your staining... whereas in fact it is much more difficult to properly control background staining. Finally, I want to address your statement: "...no scientific evidence..." When I hear of people who titrate their isotype control to give lower levels of background (to the same level as their positive antibody).... well, I don't need "scientific" evidence. Legally, this could be referred to as "prima facie" evidence of bad science. You don't need scientific evidence to prove an artefact you need scientific evidence to prove positive results. The statement that there's no evidence that the use of isotype controls has hampered any results is very much like the current "arguments" made by the religious conservatives in the US in favor of "intelligent design" (vs. evolution). There we go: experiments using isotype controls to define gating boundaries... are the "intelligent design" of experimental analyses
mario roederer, stanford
blocking
I have not had any problems (background staining) with mouse MEFs so far. For routine Fc blocking when using mouse tissue samples, I incubate the cells with 2.4G2 (anti-Fc) for 20-30 minutes on ice. Usually, I do not even wash the cells after blocking (a quick spin for removing 2.4G2 supernatant). It works really well for me. Occasionally, in my liver MNC preparations, I do get background B cell staining in spite of 2.4G2 block. I have not got around this problem yet (other than using a negative B220 gate).
I have not used the Fc block after the first blocking step. I block the cells first with the anti-Fc antibody, surface stain and directly fix and permeabilize my cells for intracellular staining. Hope this helps.
Sriram
Venkataraman Sriram, PhD
The Walther Cancer Institute
Indiana University School of Medicine
Indianapolis, IN 46202
2. In our hands we use a routine staining buffer (PBS) that includes Fc block in all staining and washing steps. I use 2.5% total protein in the block, 1% BSA,1% FBS and .5% nmIg. It can be pricey if you use at the recommended concentrations we are used to so we grow it up with a hybridoma Ig spitting cell line from ATCC ; MOPC-31C for staining mouse lymphocytes at 500ug/ml. The Bible for my generation was and is Harlow and Lane, Antibodies-A Laboratory Manual, Cold Spring Harbor Press for all this kind of stuff..
cell cycle/subg1 etc -apoptosis
Geert Martens wrote: >we have done a series of experiments with mitochondrial poisons (such as >rotenone) in rat pancreatic beta cells, and tried to determine mitochondrial membrane potential on a semi-quantitative basis with the dye JC-1 we believe our system works fine : CCCP 10 µM for 20 minutes decreases red >fluorescence en shifts all cells to green fluorescence, and we are able to produce classical JC-1 dot plots (FL2 vs FL1 ) were uncoupling shifts the cells from upper left quadrant (high red, low green) to lower right quadrant >(low red, high green fluorescence) however, we would like to know how to convert these dot plots in CORRECT >numerical data. in If a ratio of parameters is useful, the thing to do is get a value of the ratio for each cell and then plot the distribution of values of the ratio. It is often necessary to scale the ratio values so they fit on the same measurement scale as the original parameters. When taking a fluorescence ratio, you need to use linear values of the data points, not log values; to scale the values of the ratio, you must multiply the raw values by a constant. If your data are on a log scale, you can obtain the log of the ratio a/b by subtracting log b from log a; to scale this value, you add a constant rather than multiplying, because the log of a product is the sum of the logs of the multiplier and multiplicand. However, the arithmetic is only worth the effort if the two parameters used in the ratio are very well correlated with one another, i.e., if they form a "long, skinny cluster" which comes up in different regions of a 2-D measurement space (e.g., a dot plot) under different experimental circumstances. If the two parameters aren't well correlated, the distributions of the ratios won't discriminate much better between cells in different states than will the original parameter values. If the parameters are well correlated, you need to construct a calibration curve relating the scaled ratio values to what you are trying to quantify, in this case, mitochondrial membrane potential, meaning that you have to have some way of setting that to known values. A good illustration of this methodology, dealing with bacterial membrane potential measurement using DiOC2(3), appears in Novo D, Perlmutter NG, Hunt RH, Shapiro HM: Accurate flow cytometric membrane potential measurement in bacteria using diethyloxacarbocyanine and a ratiometric technique. Cytometry 35:55-63, 1999; the material is also presented on pp. 256 and 400-402 and on the back cover of the 4th Edition of Practical Flow Cytometry. Typical clusters representing JC-1 red vs. green fluorescence (presumably mitochondrial) under different experimental conditions are not nearly as well correlated as clusters representing green vs. red DiOC2(3) fluorescence in bacteria (in the 4th Edition, compare Figure 7-31, p. 399, and Figure 7-32, p.400). It thus seems doubtful to me that there is much reward to be gained from going to the trouble of calculating, scaling, and plotting fluorescence ratios. -
Howard M Shapiro
Histone H3 is phosphorylated at Ser-10 during mitosis and there is an antibody that specifically detects the phosphorylated epitope of histone H3 (e.g. provided by Sigma Chemical Co). In our hands this Ab was the most reliable marker of mitotic cells (identified from prophase to telophase) applicable to cytometry (Juan et al., Cytometry 32:71-77;1998). Phosphorylated histone H3 and other markers of mitotic cells are reviewed by Juan et al. (Methods to identify mitotic cells by flow cytometry. Meth Cell Biol, 63: 343-354, 2001)
Zbigniew Darzynkiewicz, M.D., Ph.D. Brander Cancer Research Institute
Jake Jacobberger is correct. Most likely the phenomenon reflects rapid diffusion of the dye and/or ions from the core sample stream to the sheath stream when they meet upstream in the flow channel. The diffusion leads to a decrease of dye concentration in the sample (core) stream which breaks the equilibrium between the dye and its binding sites in the cell. The changeable staining pattern is observed until new equilibrium establishes which takes some time of flow run. We observed this phenomenon using acridine orange, the dye that is extremely sensitive with respect to even minute change in its concentration or concentration of counterions such as sodium or divalent ions in the sample stream. The phenomenon is additionally exacerbated in instruments that have long sample lines such as old Ortho instruments and can be diminished by faster flow rate. We underscored this in our old papers describing the use of acridine orange (e.g. Darzynkiewicz, Z.: Simultaneous Analysis of Cellular RNA and DNA Content. In: Methods in Cell Biology, Flow Cytometry (2nd edition). Z. Darzynkiewicz, J.P. Robinson and H.A. Crissman (eds.), Academic Press, New York, N.Y. 1994, pp. 401-420, see pages 411-412.) I wish Merry Christmas, Happy Holidays, and the very best in the New Year to all FLOWERS,
Zbigniew Darzynkiewicz, M.D., Ph.D. Brander Cancer Research Institute
Hello Janet, Late apoptotic cells have many features similar to these of necrotic cells, the most apparent one is loss of plasma membrane integrity. Because they do not exclude 7-AAD, PI or DAPI the dye exclusion marker is not much help to distinguish them from necrotic cells. Microscopic examination is obviously the gold standard to distinguish apoptosis from necrosis so if you cytospin the cells from parallel sample that was measured and see only apoptotic cells then you may define that the cells in the far-left peak are indeed late apoptotic. If not, I would suggest that you use another marker, such as PARP cleavage or caspase-3 activation (each of them can be detected immunocytochemically) as a marker identifying apoptotic cells. Different strategies to distinguish apoptosis from necrosis are presented in our chapter: Darzynkiewicz Z, Bedner E, Traganos F. Difficulties and pitfalls in analysis of apoptosis. In: Methods in Cell Biology. Vol. 63, CYTOMETRY, 3rd Edition. Z.Darzynkiewicz, J.P.Robinson, and H.A.Crissman, Eds. Academic Press, San Diego, CA, 2001; 527-559
Zbigniew Darzynkiewicz, M.D., Ph.D. Brander Cancer Research Institute
The Chromatin Structure Assay is in reality analysis of susceptibility of DNA in situ to denaturation, induced by acid or heat. Extensive studies have been carried out using this assay to analyze chromatin of different cells in relation to the cycle phase, three decades ago. For example, when applied to lymphocytes this assay allows one to discriminate Go from G1 cells and G2 from mitotic cells, as well as distinguish other phases of the cell cycle. More recently this assay was mentioned, with other "historical" methods applicable to cell cycle, in the review article in Cytometry (Cytometry of the cell cycle. Cycling through history. Cytometry, 58A; 21-32, 2004). The original, earlier papers on this topic are: (1) Darzynkiewicz, Z., Traganos, F., Andreeff, M., Sharpless, T., Melamed, M.R.: Different sensitivity of chromatin to acid denaturation in quiescent and cycling cells as revealed by flow cytometry. J. Histochem. Cytochem., 27:478-485, 1979;. (2) Darzynkiewicz Z., Traganos, F., Sharpless, T., Melamed, M.R.: Cell cycle related changes in nuclear chromatin of stimulated lymphocytes as measured by flow cytometry. Cancer Res.. 37:4635-4640, 1977. The confocal analysis of DNA denaturation by this assay is described in: Dobrucki J.,Darzynkiewicz, Z. Chromatin condensation and sensitivity of DNA in situ to denaturation during cell cycle and apoptosis. A confocal microscopy study. Micron, 32: 645-652, 2001 .