Science topic

Fluorescence - Science topic

The property of emitting radiation while being irradiated. The radiation emitted is usually of longer wavelength than that incident or absorbed, e.g., a substance can be irradiated with invisible radiation and emit visible light. X-ray fluorescence is used in diagnosis.
Questions related to Fluorescence
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Does anyone know some reference(s) that show/discuss the differences in ethidium bromide intercalation/fluorescence efficiency using supercoiled DNA, nicked and linear plasmids? Would nicked plasmid enhance expressively ethidium bromide staining? It is very clear from supercoiled to linear strands, but would nicked plasmid enhance expressively ethidium bromide staining?
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Supercoiled DNA has a somewhat greater affinity since EBr intercalation will relax it. This is only true at low levels or EBr. If there is a high concentration of EBr supercoiled DNA will bind less because you now induce positive supercoils. I do not think that you will see a difference in staining nicked and linear DNA, both are similar and the fact that you have an intercalation site more in the nicked substrate is not going to help you.
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Fluor-tagged protein intensities measured w/ and w/o liposomes.
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If you are looking to calculate penetration depth, add a small amount of spin-labeled or brominated lipids to the membrane. The F/Fo ratio with the membrane label at two (or more) positions will give the depth of penetration (parallax quenching).
If you are looking to calculate the quaternary structure, you can use FRET, but be warned that this is not terribly accurate
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For GFP labelling, I am using P201, VSV-G and Delta 8.9 for the transfection along with PEI in the HEK 293T cells. I used the Qiagen kit for the isolation of plasmids as per the protocol. The construct worked after a week i.e., I observed the fluorescence in the HEK293T cell line. Assuming that the virus packaging also worked, I infected my culture of interest with the supernatant (filtered). After 48hours of transduction, I was not able to see any of the cells tagged with GFP. Could you please suggest as to what should be done in order to achieve proper packaging of lentivirus or what could be done to troubleshoot the same?
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@sowmya ramesh.. 293T cells are very easily transfected and the p201 plasmid, if it is what you expect it to be, then you should be able to see GFP expression in as little as 12 hours. From your initial transfection, I would think that there is some problem with either the transfection reagent (PEI) or with the plasmid itself.
For the plasmid, you could perform a PCR for the presence of CMV-GFP and/or a diagnostic restriction digest and confirm the accuracy of the plasmid.
As for the PEI, there is a very specific way to make it and you cannot make the transfection reagent with any run of the mill PEI available. It has to be a linear unbranched polymer of at least 25kDa, which you have to dissolve in distilled or ultra pure molecular biology grade water.
The other way to confirm your transfection is to observe for syncytia in your transfected cells. VSV-G would form massive syncytia (huge cells with multiple nuclei) in most cell lines if transfected efficiently.
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Doing Stern-Volmer study on a variety of dyes in the presence of AuNPs
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Did you mean fluorescence quenching by resonance energy transfer? If you want to find the energy transfer efficiency the donor acceptor ratio should be 1:1. But for quenching study the donor dye concentration could be anything as long as the fluorescence signal is reasonable.
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Instrument: LifeSpec II(Edinburgh)
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Please answer some of my questions regarding the data fitting of anisotropy decays..
1) How do we choose the value of g factor while giving it manually not automatically?
2) can rotation time of the molecule be larger than its fluorescence lifetime?? I think rotations should be complete before the molecule comes down to the ground state??
3) is there any significance of the value of initial residual anisotropy for the lifetime values??
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My doubt concerns whether we have to prepare the reference sample (ex. Quinine Sulfate, Rhodamine, etc.) at the same con. as our compound. Also, is taking the absorption and emission spectrum for both samples and calculating the quantum efficiency by comparative method the best method or is there any other method of doing this without preparing a solution of reference sample?
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Measuring fluorescence quantum yield using a reference compound with a known fluorescence quantum yield.
Decide on a single wavelength of excitation that allows for excitation of both reference and sample.
Make sure the reference is dissolved in the solvent for which its quantum yield has been determined. Look up the refractive index for this solvent.
The sample can be dissolved in any solvent of which the refractive index is know. The fluorescence quantum yield is a property of the compound in a given solvent. Hence, the determined value is only valid for the solvent used.
Dilute sample and reference solution to obtain an optical density <0.05 at the wavelength of excitation to avoid inner filter effects. Concentration is irrelevant.
Set fluorimeter slit-widths such that both reference and sample can be measure within the detection limits of the fluorimeter.
Make sure that the emission spectra of reference and sample can be recorded in full to allow for proper calculation of the integrated emission intensity.
For compounds with very low fluorescence quantum yields Raman scattering might be visible in the spectrum. If so, it is necessary to correct the emission spectrum for the blank (solvent only) to perform a proper emission intensity integration.
Integrate emission intensity of both sample and reference.
Calculate the relative quantum yield (QY) of the sample in the solvent used according:
QY s = (O.D. ref / O.D. s) * (I s / I ref) * (RI s sol./ RI ref sol.)^2 * QY ref
Wherein s, ref, QY, O.D., I, RI, and sol. stand for sample, reference, fluorescence quantum yield, optical density at wavelength of excitation, integrated emission intensity, refractive index, and solvent, respectively.
Determine the quantum yield in triplicate and average the values.
Excellent resource:
J. R. Lakowicz, Principles of Fluorescence Spectroscopy, 3rd ed., Springer, New York, 2006.
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I want to observe the emission spectra of the flavonoid, EGCG in flurimeter.
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According to the absorption spectra that I have found it absorbs in UV, between 250 and 300 nm, and this is the region where you should excite.
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I want to quantify the difference in fluorescence intensity between two areas in a sample
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I agree with the answers above. Just a small addition to them, depending on your application, you can also normalize the signal through the desired filter setting with respect to a different emission wavelength (emission filter) for the two regions independently. This should take care of uneven staining, illumination etc and should also improve SNR.
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Looking for light sensitivity as well as fast video capture (100 fps); a rather tricky combination. It is to go on a Leica MZ10F and connect to a PC.
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From what I've seen, these cameras generally come in two categories, the $3000 arena and the $20,000. For what I'm doing the former isn't going to be enough and the latter is going to be overkill. Is there anything in between?
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I am trying to measure the fluorescence of cells at plasma membrane using ImageJ. Could anyone help me how I could do it please. I attached a sample image also.
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Hi Gulzar. What exactly do you want to analyze? The quantity of fluorescense signal at the membrane compared to cytosol? One possibility would be a 3D surface plot "interactive 3D surface plot", you get that as a plugin. Or you make a profile plot (you fin that under "analyze")
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Tryptophan fluorescence intensity is a convenient method to measure/estimate the dissociation constant of an interaction between a peptide and a partner, as it requires very low concentrations of biological material. Many papers have described the method which relies on the fluorescence intensity measured upon addition of the partner. I'm studying the interaction between peptides and real membranes extracted from bacteria using Trp fluorescence. Upon membrane addition, the maximum emission wavelength undergoes a blue shift, indicating a binding to the membrane. The fluorescence intensity overall increases with, however, an inconsistent behavior upon membrane addition (sometimes it increases, then it decreases a little bit, and increases again...). That's why I would prefer to use the maximum emission wavelength to calculate the Kd.
Can fluorescence experts recommend me an article explaining in details how to do that?
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The fluorescence of tryptophan changes because it looses energy to the solvent. The best way to measure it is not the peak intensity but the avarage distribution of the emission energy what is given by the simple calculation of the center of mass (CM) of the emission spectrum. Mathematically, the CM is the point that divides the area of the spectrum in two parts of equal area. Concerning to the emission energy, the CM is the medium point of the spectrum. It may be calculated from data in nm, and the values will be inversely proportional to the energy. Or you may convert your data to wavenumber, and then the values will be directly proportional to the energy. When you observe a blue shift of the spectra when the peptides bind to membrane, that means that the hydrophobic environment helps preserving the energy, favouring the transitions in the blue side of the spectra. The CM represents the entire shift and not only the change in intensity in one wavelength. Look for the Lakowicz book, or Gregorio weber´s papers (the pionneer), Jerson Silva´s papers, david Jameson´s papers... Those are some experts on protein fluorescence and thermodynamics.
The Stern Volmer plot could also work in what case water is the quencher and the membrane protects the peptide from being quenched. It is a little bit different from the regular quenching experiments. Anisotropy may be a problem if the membranes added sccatter too much light. Tryptophan is not really a good probe for that since the exc wavelength will be low, what increases sccattering, low signal (if you can not increase the peptide concentration) because there will be photoselection during the absorption of polarized light. The suggestion of labeling the peptide is good only if labeling does not change the membrane binding properties.
I prefer fluorescence spectroscopy in this case because it is a very sensitive method. NMR is fine but can you do it by H NMR? Or do you need to have the peptide produced with C13 or N15? In any case you will need much higher concentrations than when you use fluorescence.
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I need fluorescent markers for the liver endothelial cells. I have used anti-Pecam antibodies and they worked well, but something else that shows endothelial cell function and health would be great.
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Hello,
it is very difficult to answer your question, because except microscopic analysis (for fenestration analysis), there is no highly specif marker of this cell type. You could read the article of March et al. Hepatology 2009, it could help You, because as wrote Thomas MOhr, in in vitro study (in 2D) sinusoidal specific phenotype can be lost. This phenotype can be highly different between in vivo and in vitro conditions. For information about markers potentially expressed in these particular conditions read Am Elvevold J Physiol Gastrointest 2008
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I' believe this might be energy transfer from Trp to NAD or NADPH (excitation 340-360 emission ~460 nm).
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I wish to see an image of a ~2 keV proton beam. The beam current is ~1 microamp. I had been told that there was a simple and low cost method to do this: make a mix of doped zinc-sulphide powder and acetone (10:90 mix ratio) and "paint" the mix onto a metal surface. When the ion beam strikes the surface it fluoresces and you see the beam.
Has anyone tried this? Is there an alternative which is also simple and cheap (i.e. not employing microchannel plates)?
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We use a similar method in our tandem accelerator with 10nA of 3MeV He, only that we fix the zinc-sulphide powder on a double sided tape. It works pretty well, although the powder turns brown and loses light intensity after some time. Due to the brightness the spot may appear larger than it actually is.
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I am attaching a slide that will give you an idea of what I am talking about. The Hoe concentration is same for all cells. These are just pictures from two different wells but since these are taken at the same exposure time. I fail to understand why one is so much brighter than the other.
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Thanks. I will try it this time!
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Can anyone give me some explanations concerning aggregation induced fluorescence? I synthesized a molecule which behaves strangely and is not fluorescent in organic solvents but as solid and dissolved in water. Can anyone give me some suggestions about experiments which I can perfom?
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You should look into Tang's work about Aggregation Induced Emission Chem Commun 2009, 4332-4353.
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I don't know how to fix this bug.
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Dear Melissa,
Yes, I have the same bug with a LS-55 Perkinelmer spectrofluorometer and the software FL Winlab v4.0. The bug is due to a software problem with the file names, if you use simple short names without dots, spaces, etc, to save your spectra you will avoid this bug.
Best regards
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I am looking for a way to visualize the glycocalyx of live cells (preferably fluorescence by blue or red excitation). After googling, it appears that the standard method is to use periodic acid (which inserts between saccharides and forms a pair of aldehyde groups) and a Schiff's reagent (which binds aldehydes, which are otherwise not too common on the cell/tissue surface).
Can anyone point me to a Schiff's reagent (or a suitable alternative) that is of the fluorescent (not the typical absorptive) variety, that fluoresces either blue/green (ex/em) or red/deep red (ex/em)?
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Glycocalyx can also be stained using certain lectins. You might want to check the publications from S. Reitsma, e.g.:
The endothelial glycocalyx: composition, functions, and visualization.
Reitsma S, Slaaf DW, Vink H, van Zandvoort MA, oude Egbrink MG.
Pflugers Arch. 2007 Jun;454(3):345-59. Epub 2007 Jan 26. Review.
PMID: 17256154 [PubMed - indexed for MEDLINE]
Good luck!
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I am looking for a fourth colour for FACS analysis. I have 3 cell lines expressing DS red, GFP, CFP using lentivirus. I would like to know if there is one more colour that doesn't overlap with these three. I don't want YFP, because I'm expecting a cell fusion with DS red and GFP labelled cells leading to yellow.
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You can have as many colours as you wish the only thing is you should have. check out the web site for dyes.
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I am doing microinjections in the brain and I want to localize the location of my injection after a week or a month but avoid slicing a lot of brain and without the need to process the tissue to localize the injection. Usually it is very hard to find the location during the cryostat slicing process because there is no blood or visual lesion at the injection place.
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Hi Cristina,
I don't have an answer to your question, but you might find some hints from the link below. I could imagine not to use a dye, but some kind of colored, non-fluorescent nanopoarticles...
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I work with fluorescent compounds, particularly organic fluorescent dyes, which are doped into optical quality polymer hosts, such as PMMA. One area I'm interested in is trying to manipulate the host environment with additives to influence the resultant fluorescence by the mechanism of solvation. By including additives of high relative permittivity the bulk dielectric continuum is strengthened and this affects the emissive excited states of the dyes, redshifting them and thereby reducing overlap of absorption and emission spectra. So, I want to functionalise nanoparticles for inclusion into PMMA films with high homogeneity.
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titanium oxide compounds have strong affinity toward carboxylate-bearing molecules. You may want to look for a suitable compound (e.g. a silane) with -COOH as end-group. Titanium oxide materials are often hydroxilated on the surface, that is the trick.
I am curious about why I see a shift of the "negative" peak in my flow cytometry experiments on the benchtop Accuri C6 cytometer?
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Is it possible that scattered light from a bright sample with higher average fluorescence would produce an upshift in the designated "non-fluorescing cells" relative to a low brightness sample?
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Maybe it is the transfection procedure. After the transfection the morphology of cells may change a little bit, leading to a different backgroung fluorescence. I have seen this with electroporation of T lymphocytes and lentivirus transduction of 293T cells.
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I am recording changes in cytosolic calcium using Fura-2 AM in cells previously transfected with a GFP tagged protein. In order to obtain fluorescence background added by GFP I am trying to quench Fura-2 to obtain only the fluorescence of GFP at 510nm (emission of Fura-2) to delete that value from records and obtain only the fluorescence of Fura-2. I've tryed adding BrA23, then Digitonin and then MnCl2 2mM but did not observe quenching of Fura-2
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Mn should completely quench Fura-2 at these concentrations, so my suspicion is that you are not picking up any Fura-2 signal - it is overwhelmed by GFP. You could check this by instead using a high concentration of extracellular EGTA (mM) in the absence of calcium. This should show a reduction in signal if you are indeed picking up calcium, especially after digitonin. Are you using "classic" GFP or eGFP? The latter has a longer max excitation wavelength (488nm), and for 340/380 nm excitation would show considerably less emission (see http://www.biotek.com/resources/articles/green-fluorescent-proteins.html) . We have used this relatively successfully in the past to look at calcium in eGFP tagged cells. The alternative of course is to use a longer wavelength dye - as these are cells you could get away with a non-ratiometric dye such as calcium green.
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Do you dissect the region first, or use another method?
Array Tomography is a technique for excellent axial resolution imaging by using ultrathin sections for fluorescent microscopy. it also permits repeated staining and imaging of the same sections
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Our lab dissects the region first before embedding, then prepares the arrays. On occasion, I'll also taken semi-thin (500 nm) sections after embedding but before taking arrays and stain with toluidine blue to verify location. Then, when we image, we use DAPI to verify and locate the cell bodies.
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I am performing FRAP in live cells to check the dynamics of my interested protein tagged with GFP. Its DNA binding protein. But, when I expose the sample, the entire area bleaches even if I point a small area. I am not finding this difficulty when I do it on same cells when they are fixed. What does this tell you?
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As Radek said this probably means that either the illumination spot is too big or its duration is too long or both. Try to illuminate the sample in a smaller region for a lesser duration till you see a dark spot on a bright back ground.
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I'm reading Guide to Protein Purification (From Methods in Enzymology series) and there is:
"However, it must be pointed out that the concentration of the fluorophore cannot be calculated from the measured fluorescence emission by application of a universal constant equivalent to the molar extinction coefficient. Rather, relative changes in fluorescence emission with time are compared."
How is this so? On what does the fluorescence depend on? How can we compare relative fluorescence if it is not proportional to concentration?
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The text you've cited correctly states that luminescence intensities vary with lumophore concentration, but not always in a linear (or predictable) manner. From a practical perspective, you can certainly use emission intensities to determine analyte concentrations, but it's necessary to first construct a calibration curve (i.e. by measuring luminescence intensities from serial dilutions of an analyte solution whose concentration is known) using the same solvent, temperature, pH, etc. as the system you're hoping to study. You can then determine the lumophore concentration in your system of interest by comparing your emission measurement(s) to your calibration curve and extrapolating.
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I'm developing an adhesion assay with CFSE-labeled polyclonal T-cells. Basically, I allow the cells to adhere to a poly-l-lysine coated (0.01%) well in a 24 well plate, and after gentle washing, I read the fluorescence at an excitation of 494 and an emission of 517 nm.
Incidentally, the plate reader reports there being no signal, despite the fact that I can see cells when I look at the wells under the microscope. Should I increase the cell density? I am seeding 500,000 cells per well at this point. Or, is there something else that I am missing?
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1. Did you QC your CFSE staining? I stained with CFSE and fluorescence was very strong.
2. Are you using compatible plates? For fluorescence assay you should use black plates or the signal could be influenced from adjacent wells. Using transparent plates could give rise to high background readings caused by reflection.
3. Are you using your flourometer properly? Optimized z position etc.
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My compound shows the maximum absorbance at around 260 nm. But the maximum intensity in excitation spectrum appears at 300 nm. The excitation spectrum has been corrected. I wonder which maximum value should be involved in stokes shift calculation.
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When absorption and excitation spectra do not match, there is a severe problem. There can be a number of reasons. First: sample impurity. That has to be checked first. Absorption spectroscopy alone often is not sensive enough, depending on relative fluorescence quantum yields of sample and impurity and/or differences in absorbance at the excitation wavelength. Keep in mind that spectral calibration is not a simple issue. Also photoproducts can lead to difficulties. Check as a function of irradiation time.
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Doing fluorescence analysis, I've identified a peak related to a NADH-dehydrogenase complex, but I don't know which type of dehydrogenase is. Is there any kit I can use for this? Or any other protocol?
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Do you fractionate as a part of your protocol? If so can you see the protein(s) on a gel? Is there more than one? My immediate inclination is to MS/MS the fraction, identify proteins and look at their functional annotation to see if there is any NADH dependent anzyme present, or BLAST search with the MS/MS sequence to find an annotated homologue from another organism
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How good (or bad) was it?
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Imaging through refractive index mismatches will cause spherical and chromatic aberrations if you are out of range of the specified correction. Nevertheless, I agree that it is possible to achieve good pictures thorugh plastic. Note, that the sensetivity to aberrations is mostly depend on the NA; the lower the better the grade of freedom. I would mostly consider the chromatic aberration. Worst case I ever saw was a color shift of 50µm along the optical axes between 405nm and 561nm excitation imaging thourgh 2 mm plastic. Here, the Nikon Plan Fluor 20x/0.45 was used.
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I'm planning an assay based on the quenching / FRET with eGFP as partner. Could somebody point me to a few efficient Quenchers (or a review regarding that matter?).
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DPA (Dipicryl amine or hexanitrodiphenylamine) is a very efficient quencher of GFP/YFP. It is a hydrophobic anion that intercalates in the plasma membrane, so, if your constructs are membrane-associated, this is a very good system.
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I want to fuse a fluorescent protein to a viral protein in order to track its subcellular location and to assess colocalization with other proteins during infection. Since a red fluorescent protein is preferable for my assays, and I have a plasmid encoding RFP available in the lab, I would like to hear from you about its performance regarding to brightness and photostability when compared with other red fluorescent proteins.
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that's funny Simon, I had just the opposite case in yeast, where td-tomato caused some untypical behaviour at tagged proteins...
One point which wasn't mentioned yet, is the maturation time. mCherry is quite fast, comparable to GFP variants, but most other RFPs have at least one hour maturation half-time. So, if you are interested to see changes in localization or responses to different cues, which you expect to be fast, then use mCherry, it works even nice as threefold-tag for us but is not as bright as other RFPs. If you want I have some nice papers which list the maturation times and photostability of different fluorophores, but I'd have to search them first.
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I'm having trouble in cryostat sectioning.
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Thank you!
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I'd like to try the CLARITY method on some of my knock-in mice with fluorescent reporter molecules attached to their post synaptic proteins. It is impractical for me to set up the entire method when I don't know if it will kill the fluorescence, so it would be excellent if I could come and borrow someone's set up for a short period to test the method on my samples.
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Hi Rhiannon
Although I am not in the UK, I have recently helped to set up the CLARITY method in our lab group and have successfully used it to clear and visualize CNS tissue in GFP mutant mice. We kept our samples shielded from light throughout the protocol and did not notice any decrease in the fluorescence. If you want to try the CLARITY protocol without committing to manufacturing the whole setup, I recommend trying passive clearing instead of ETC. We have done this with great success in our lab with our tissue samples. Instead of the ETC setup, after polymerization we just keep the samples in a small amount of clearing solution in a 37C water bath and simply change the clearing solution daily. With this technique we were able to achieve completely successful clearing of our tissue in 1-2 weeks (depending on the sample size). We leave the protocol unchanged otherwise. Let me know if you have any further questions!
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I have to compare the spectra of a fluorescent dye with the producer's spectra but the FluoroMax-HORIBA that I use measures in cps, while the one from the dye producer is in relative intensity %. I checked whether I could measure mine in Relative Intensity % but I don't think I can! Is there a way to convert one to the other so that I can compare them?
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You do not really need the absolute intensities. Compare the line shapes of the two spectra (yours and the reference spectrum by the manufacturer). Is the wavelength of the fluorescence maximum similar and does the line shape match? You can measure the intensity of the maximum of your recorded spectra and divide your spectrum by this maximum. Do the same for the reference spectrum. Then, if the line-shapes of both resulting spectra superimpose without much deviation, your dye is fine.
The fluorescence intensity depends on a number of different parameters, for example the intensity of the light source, slit widths of excitation and emission monochromators etc. It is therefore not easy to compare absolute intensities recorded on different machines. A method of standardization could be to always take a spectrum of a reference substance of known concentration and use the maximum intensity as a normalization factor for all other spectra.
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I am using FITC tagged IgG to indicate its attachment to cellulose paper (Whatman #1 CHR). But the paper itself is autofluorescence, quite annoying. Does anyone know the ranges of paper's autofluorescence? What fluorescent filters can avoid the paper autoluorescence?
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You may try to photobleach it with the excitation wavelength it could get it down by half. Also, an emission wavelength over 700nm (CY5) gives you less signal.
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This is quite a specific question, I know. I'm currently determining the absorption coefficients of a range of fluorescent materials and am a bit stuck with this one because I can't find its molecular weight anywhere. From literature I know Lumogen F Red 305 has Mw = 963.956 and Lumogen F Orange 240 has Mw = 710.873. It is necessary to know these so that the molar concentration can be determined which is essential in applying the Beer-Lambert law to determine the extinction coefficient. I've emailed BASF but the lumbering giant hasn't registered my existence.
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Sadly not I'm afraid Bryce; I contacted BASF and the advisor didn't really know what this was (typical) but said they'd get a technical advisor to contact me back, which they didn't. Perhaps a follow up call with a bit more assertiveness would get me somewhere but I forgot about this along the way doing other things.
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Are there any extra tricks that I'm overlooking that are general or best-practice to get this assay to work? I am targeting a protein that should be available at the ng level and is tethered to a solid metal surface.
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Too little information on how you were trying tp target your protein
are you working with metal particles or metal plates?
Using only one antibody or a primary and secondary set ?
Cells show fluorescence under any filter. How to remove it?
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I infect my cells with lentivirus expressing GFP in serum free media. After two days, other than some cells that are really expressing GFP, almost all others show fluorescence under any filter (green and red). This is really annoying because I am trying to see how many cells are actually green and with such background it is difficult. Is it a common issue? How can I remove this background?
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Are you fixing the cells? What are the specs for the filters? What excitation are you using? To start autofluorescence is almost ubiquitous to all fluorescence imaging, you have to get as close as possible to the source and try to address the issue. Some fixation procedures induce more background (BG) than others and the same is true for mounting media. A careful selection of filters, both for your emission AND the excitation could help. There could be an excitation that although not optimal for GFP will give you lower BG and maybe better images. Also there are some computational tricks like deconvolution. Good luck
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In a project we need to stain bacteria (Gram-negative and positive ones) with a fluorescent dye for detection. It would also be ideal if the dye would absorb strongly in the UV range but would emit in the VIS range. Advantageously the dye had a high quantum yield. We tried semi-conductor particles - Quantum Dots - to stain the membranes. These would be ideal but the hydrophobic QDots are difficult to suspend in aqueous solutions (as expected…) and other types of QDots with surface coatings do not interact with the bacterial cells. Thank you for any advice!
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Do you just need to stain the membranes, or is it ok if the whole bacteria light up? If the latter, acridine orange is cheap, easy to use, and meets your other preferences.
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X rays
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I could provide more information about scintillation crystals desire sustancuas a project of fluorescent in dental organs
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I am looking for a relatively high throughput method of detecting the CMC of detergent or protein micelles. Does any one know of another high throughput method besides fluorescence or other chemical probes besides pyrene fluorescence?
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The absorbance of Coomassie Brilliant Blue R-250 is shifted upon partitioning into micelles. This method utilizes a red shift of the absorption maximum of Coomassie Brilliant Blue R-250 from 555 nm in the absence to 595 nm in the presence of detergent micelles. I used this method to determine CMC for a couple of detergents (http://shootingcupoche.com/publication/227648148_Outer_membrane_protein_A_of_E._coli_folds_into_detergent_micelles_but_not_in_the_presence_of_monomeric_detergent?ev=prf_pub). I vaguely remember that it should be the R-form of Coomassie Blue (there is also a G-form).
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Currently I am trying to make multiple immunofluorescence labelling of formalin-fixed paraffin-embedded (FFPE) liver tissue. Unfortunately, tons of red blood cells with strong autofluorescence at 488 and 546 excitation were found in tissue sections, though I have made blocking procedure using 5% BSA before antibody staining.
Does anyone have any suggestions to reduce or eliminate autofluorescence by red blood cells? May cryosections help? Thank you very much in advance!
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What species are you working with? In case you have liver samples from rats or mice you could cardially perfuse the animal with PBS to remove red blood cells and then use perfusion fixation and continue according to your standard protocol.
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We have observed that P can only interferes on W X-ray emission if they are part of the same compound. We believe there´s a non-radiative energy transfer process ocurring in our system and would like to discuss about these phenomena.
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Any standard reference: " Handbook of X-ray spectrometry" by Van Grieken & Markowicz..etc. You can look up the NIST web-site too for X-ray fundamental parameters...or X-ray "Fundamental parameters" library in google.
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Spectra calculation
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Can we compute? Well, yes and no. The fluorescence spectrum is quantum mechanically defined based on the molecule and its local environment. However, a full and accurate description might not be easily programmed into a computer. Approximations are always required and accuracy always suffers. You can almost always calculate the spectrum to a first order. Just take a look at a Jablonski diagram for your molecule. However, that's often not the interesting part of the phenomenon. It's often the second and even third order components that are the interesting or important and these are much more computationally difficult.
If you search the literature of computational chemistry, you'll see that computing every aspect of spectroscopy is a huge field. Many scientists and engineers are working to improve existing models and hardware to improve accuracy. Yet still, there's a long way to go. There are many methods and techniques. Each varies in terms of accuracy and computational expense, even in the particular field of application. One good place to get an introduction is to go to the Wikipedia website and search for "computational chemistry". There, you'll get a feel for how broad and complex the field has become.
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When analyzing free tryptophan in solution, I can see two peaks at 230nm and 280nm with the same emission ~350nm, and the peak at 230 has the highest intensity. When analyzing proteins, sometimes I see the peak at 280 higher than the one at 230nm. Why is that? Does it have to be with the location of Trp residues in the protein?
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Record an excitation spectrum with emission max at 340 nm and scan range 200-330 nm. You won't see the 230 nm peak. 230 nm band doesn't give you information about the tryptophan or tyrosine.
280 nm spectra is for the transition of diffused pi electrons of the aromatic ring of try or trp. Because the excitation of the conjugated pi electrons requires lower energy. The high energy absorption at 230 nm or below is due to the n->pi* or pi->pi* transitions of the carboxylic group or amide in protein.
Do not rely on the absorption below 250 nm if the experiment was done in aerial condition. Dissolved oxygen must be removed from the sample by passing nitrogen or inert gas through the sample and the spectra should also be recorded in continuous nitrogen flowing condition if you want to report the below 250 nm spectra. As in circular dichroism experiments where we record the differential absorption at far UV regions. Oxygen absorbs heavily at this range.
Fluorescence signal after fixation?
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I am culturing a mouse embryonic stem cell line having OCT4 fluorescently labelled. After fixation with paraformaldehyde, no fluorescence signal remains. I would like to know is there any method of fixation, upon which this inherent signal will remain intact?
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The only time I perform methanol fixation is for DAPI staining and I use it pure and ice-cold (I keep it in the -20 freezer) and fix for 15 minutes at room temperature, but I'm not sure whether this is applicable to your case. Sorry I can't help more, but I hardly ever fix my cells...
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High absorption coefficient is absolutely necessary as the amount of light absorbed by the system will have a direct effect on the efficiency of my experiment. Emission around 980 nm is another important criterion.
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Large discotic molecules like porphyrins, pthalocyanins etc can solve your purpose. You can also tune the absorption and emission of these molecules by proper substitution with different functional groups. Bodipy is another molecule which can take your absorption and emission into NIR region by proper substitution and extension of conjugation. 
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Emission spectra of those molecules must not overlap a lot.
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Quinine sulphate, if you're looking for something water soluble.
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I did the cell penetrating test for the novel peptide labelled with FITC. I incubated cell in serum free medium (DMEM) for 4 hrs. Afterward I washed with PBS and incubated cell with fetal bovine containing serum for 2 hrs.
My results were unexpected - I got the fluorescence at 488 wavelength in control plate.
Does fetal bovine serum have fluorescence at 488?
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Thank you ...
For your help...I think also. i repeated the experiment and found that it has some background..so it is mostly because of serum fluorescence...
What is the source of light used in fluorescence spectometer?
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Can anyone suggest the mechanism of fluorescence?
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just google those things!! You don't need us to answer that kind of questions!!
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I have some trouble with the flourescence staining in my whole-cell patch clamp experiments in mice, particularly with the staining of cells expressing parvalbumin. Maybe it is a problem of slice fixation.
So, does anybody know a well functioning protocol for double fluorescence staining for calcium binding proteins (parvalbumin, calretinin, calbindin)?
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There is not enough info here to get at the root of the problem. What are you using for fixation? in my case I use methyl acetic acid. It works to immediately fix the cells/tissue in addition to perforating the membrane. I then wash, add 1 antibody with blocking solution and let sit for 1hr. wash again and add the second 1 antibody for 1 hr. was and then add the 2 antibodies. Usually done in a stepwise manner, but (depending on antibody) can be grouped (e.g. all 1 antibodies at once and then all 2 antibodies at once).
J
Which phase of TiO2 is active? Is there any correlation between activity of TiO2 and fluorescence behavior?
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I have done binding interaction study between benzimidazole and all phase of TiO2 (rutile, anatase, P25, hombikat). Rutile phase gets fluorescence enhancement other gets quenching of fluorescence. Is there any correlation between activity of TiO2 and fluorescence behavior.
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P25 and hombikat are NOT phases of TiO2! They are just commercial names for nanocrystalline TiO2 . The phases for TiO2 are rutile, anatase and brookite. P25 usually contains anatase and rutile whereas hombikat is usually mostly anatase with some brookite If you do not specify activity to what, it's hard in any case that you get any meaningful answer!. For sure the various phases as well as the domain size play a role in determining the properties of the powder you work with
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I am trying to visualise the components of isolated multiprotein complexes in vitro. Can you use stoichiometric photobleaching to determine individual fluorescent proteins adhered to a surface using just an ordinary light microscope? Or do you need the sensitivity of methods such as TIRF? And are there any specific fluorophores which are best for doing this (the alexa fluor dyes i'v used seem to bleach pretty instantaneously)?
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The "ordinary" light microscope does not seem to be the limiting factor here. Provided you have an objective with a reasonable NA and good transmission, the critical factors are:
(1) quite obviously the type of fluorophore, camera and quality of your fluorescent filter
(2) whether you will be able to immobilize the proteins on the surface. If so, you could "simulate" a TIRF approach by removing labelled but unbound molecules, work with longer illumination times and, consequently, less powerful light sources.
(3) whether you protect your chemical fluorophore (most alexa dyes should definitely work here) with oxygen scavenging solutions.
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In fluorescence spectra sometimes peaks are observed at wavelengths that are multiples of the excitation wavelenght. Such peaks are not due to fluorescence, what phenomenon originates them?
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This is due to the excitation monochromator. The light of a certain wavelength is able to go through the monochromator not only when this wavelength is selected, but also to any integer multiple of it (e.g. wavelength at 250 nm is able to go through the monochromator when it is in the positions 250, 500 or 750 nm). In order to avoid that, you can put just after the sample and before the excitation monochromator an optic filter blocking the excitation light (e.g. if you are exciting at 345 nm, using a filter of 350 nm). If you do so, you will see the emision spectrum without those "disgusting peaks" from scattering.
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invivo Method for NO concentration measurement
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Hi Ata,
You can find more details regarding the chemiluminescence method in the following references:
2. The Measurement of Nitric Oxide and Its Metabolites in Biological Samples by Ozone-Based Chemiluminescence http://www.springerlink.com/content/g9t237r01636651g/#section=55157&page=1
3. Nitric oxide measurement by chemiluminescence detection http://www.sciencedirect.com/science/article/pii/105867419290045Y
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I usually take raw data and plot it using excell. I want a software where I could use these data and draw the spectra for publication
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just wanted to do some self-promotion for my optical spectroscopy software SPEKWIN32, but Pedro already did it.
However, one more hint: if your data format doesn't fit one of the formats accepted by Spekwin32, have a look at the *.fak or the *.csv format (see samples from the test spectra file in the download area). Try mimicking one of these data formats and you are done.
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I need to measure the amount of peptide (tagged with fluorophore TMR) taken up by HeLa cells. For this purpose, I plate Hela cells at an 80% confluency on a 1cm2 dish, incubate with peptide for an hour, then wash the peptides off, trypsinize the cells, remove the trypsin by centrifugation, lyse the pellet with Lysis buffer, then measure fluorescence of lysate supernatant on the fluorimeter.
The problem with my method is that even though I plate all the samples at the same 80% confluency, when I compare different peptides, I need to be sure and prove that I am comparing the same number of cells in each dish. How do I do that?
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Alright.. Will keep that in mind..
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Is there any explanation if I observe only (or mainly) the decay in fluorescence as opposed to absorbance?
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The fluorescence can be quenched by the products formed during laser irradiation. So, absorbance and emission should be independently monitored. These two parameters can be correlated to see any quenching of radiative decay, if occurs.
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This data is from a plate containing only PBS and 5% BSA excitation at 680, emission at 520, 530, 540, and 590. Seems like a lot of background fluorescence.
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I am currently performing an alpha screen assay with perkin elmer plates and beads and a biotek synergy 2 and I seem to have signal in those ranges and negative controls that go down to 4000. My excitation is at 680. Emissions at 570 and 620.
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I cannot find an article, that describes a successful use of metric other than SNR/MSE to characterize deblurring of fluorescent images. Could you please recommend me one, preferably using no reference "ground truth" image? My research shows that common metrics like SSIM, VIF and likely wavelet-based ones fail to characterize deblurring success, especially for Z-stacks.
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The only method I can think of is to use fluorescence beads with known characteristics to determine the point spread function of your imaging system. That's what I do for two-photon microscope images. (Ref: Dong,C.Y., Koenig,K. & So,P. Characterizing point spread functions of two-photon fluorescence microscopy in turbid medium. J. Biomed. Opt. 8, 450-459 (2003).)
How can I differentiate whether it is scattering or absorption?
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I did the binding between nanoparticles and imidazole derivatives. In fluorescence, spectra I got enhancement. How can I differentiate whether the enhancement is due to scattering or absorption?
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Imidazole fluoresces in the near UV and even has a tail into the blue visible region. You need an emission filter that will remove light at your exciting wavelength; this will be shorter wavelength UV light if you are pumping imidazole. All the scattered light will have essentially the same wavelength as your exciting light, so if you choose an appropriate filter you can select for only the fluorescence emission.
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I have realized that carotenoids and proteins both auto fluorescence and their excitation wavelengths are similar or close to each other.
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You could try reacting the proteins in the layer with an amine reactive fluorescent dye. The carotenoids shouldn't have amines and won't react.
What is the reason for dramatic enhancement in fluorescence spectra at the wavelength of twice of excitation?
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I recorded fluorescence spectra for imidazole derivative. I fix the excitation value at 290 nm which is predicted by absorption spectra. In 580 nm, I got dramatic enhancement having more 1000 pl intensity. What is the reason?
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Dear Karunamuoorthy, What you see at 580 nm is the scattering in the second order of the grating of the excitation monochromator. Simply insert a long pass filter blocking all radiation below about 300 to 310 nm and this spurious peak will disappear.
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I am trying to investigate the changes in motor proteins in the presence of investigational compounds that are meant to interact with tubulin polymerization.
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fuse a fluorescent protein to your protein of interest and use living cell cultures or embryos for experiments. cilia are also nice for these kind of studies, so using a ciliated cell line would be a good choice.
A nice example is shown in this publication (check the suppl. movies):
And also check the stuff from the Ephrussi group, they are doing great experiments tagging proteins and looking how they are asymmetrically localized in early drosophila embryos and eggs:
PS: fluorescent tagged tubulin proteins are around as well. so you could do a 2-color study....
Have fun! It´s a great field.
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Monomeric forms of fluorescent proteins like GFP may aggregate when they are fused to other proteins which are expressed under high levels from strong promoters. One of the solutions of this problem is to use genetically modified fluorescent proteins that are unable to aggregate. Does anyone know which amino acid substitutions in GFP or other fluorescent proteins prevent their aggregation?
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Mutations A206K, L221K, and F223R were found to essentially eliminate self-association of GFP/YFP/CFP, as reported in D.A. Zacharias et al. (2002) Science 296:913-916. A good overview of these and other studies to improve fluorescent tools for live cell imaging is found in a 2005 review by Roger Tsien (R.Y. Tsien (2005) FEBS Lett. 579:927-932.
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Option 1: I would like to label an exposed His8 tag with a compound that will stay bound to the His8 tag during denaturing SDS-PAGE resolution of protein bands. In this way, non-labeled proteins should migrate faster (i.e. lower) in the gel, whereas proteins with a successfully-labeled His8 tag should be shifted higher in the gel relative to the unlabeled counterparts (as they are running more slowly).
Option 2: I would like to label an exposed His8 tag with a fluorescent compound (with an emission profile distinct from GFP) that will stay bound to the His8 tag during SDS-PAGE. Different amounts of His8 labelling could then be compared via densitometry.
Option 3: Options 1 and 2 COMBINED (this would be an ideal scenario)
NOTE: Ideally, the probes should be commercially available.
Any ideas?
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I'm not aware of any reagents like this for His-tags, but I'm going to follow this just in case there are some!
I have used a FlAsH tag for quantifying proteins in SDS-PAGE gels. But it requires a CCPGCC sequence on your protein, not a His-tag.
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I going to work in the synthesis of fluorescent nanoparticles, but I like to compare the size and composition that are already synthesized with the nanoparticles that I like to propose!
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There are so many types of fluorocent nanoparticles with different size and composition. for example quantum dots (QDs), metal nanoparticles, lanthanides (mainly Eu,Sm,Tb,Gd) doped metal oxides/fluride bassed upconverted nanoparticles, presently, fluorescent carbon nanoparticles (FCNs) are attracting considerable attention because of its low toxicity and high chemical stebility.
except that Hybrid Architectures Involving Fluorescent Nanoprobes are also promissing, that includes Metal–Dye,Dye-doped Silica Shells,Quantum Dot-containing Microspheres.
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I am measuring miRNA levels with qPCR using taqman probe (regular, no detailed information on it). Recently I introduced some miRNA with GFP-tag to my system, and did qPCR with the taqman system.
My concern is: are there any fluorescent overlaps between GFP and taqman probe?
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Taq amn probes can be ordered with something other that FAM, like TET- 521. 536 (excitaion-emisson), JOE-520. 548 or VIC. ~555(emission). What machine are you using, atleast 3 filters should do your work. Generally ROX or TAMRA are used for internal control so your probe-primer should have a different label that these two.
Dr. Ulrike, even if GFP is tagged and its a fusion construct, then GFP along with the miRNA will be transcribed and should be a single fusion miRNA, why should it be in the protein? If its not an IRES GFP where GFP will be expressed separate as opposed to a fusion construct, is there a problem of getting a fusion RNA?
So any miRNA which is made will have a GFP tag along with the sequence of the gene/miRNA of interest.
Please correct me if I am missing something.
http://www.ncbi.nlm.nih.gov/pubmed/20554852 I am not sure if the link is very helpful.
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I am trying to find a true monomeric, stable, bright fluorescent protein to label a membrane associated protein. It seems that mcherry is currently the best... what do you think? Could you share your feelings about other fluorescent proteins ?
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Do you have any limitations for detecting the protein? Is this for an expression construct or are you labeling isolated protein? This is a good question for many looking to visualize a protein of interest but will depend largely on the laser/filter combinations available in your lab/department/core/university.
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Assuming that we have a good negative control, and the right melting temperature, what can cause very low fluorescence level of the melting curve?
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Are you running the reactions until the amplification reaches the plateau phase? You don't have to do that to get the Cq value, but generally more product will get you more signal. If you are reaching plateau and still have little fluorescence, it could be your sybr green is not fresh, as mentioned above. If you are limiting your primer concentration, you may simply run out of primer before you reach a very high fluorescence, this is also true of any other reagent like dNTPs. Of course, using a smaller reaction volume, you will have less of every reagent, and you will have less fluorescence, eg) I had less fluorescence when using 12 uL reactions than when I did 25 uL reactions.
Why do mycobacteria lose fluorescence after repeated subculturing?
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I have tagged Mycobacteria with an intrinsic RFP (red fluorescent protein), mcherry which forms bright red colonies after electroporarion but on sub-culturing in liquid medium I found out the fluorescence is decreasing significantly and after 3-4 time sub-culture it is completely disappearing.
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The same happens with GFP as well. most probably over expression of GFP/RFP cause toxicity in the liquid medium resulting in selection of GFP/RFP -ve clones.
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Please explain the basic photochemical reasons behind its fluorescent nature in comparison with adenine or guanine.
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Mr. Hepburn, thanks for the suggestion. The book is very useful.
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I have a material containing coumarin in it. When my collaborator takes fluorescence microscopic images of the material using DAPI filter, he mentioned me that the sample is photobleaching. As a result he couldn't take good fluorescence microscopic images. I didn't understand this completely.
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Just to add to Laurent's answer.
Photobleaching is a chemical reaction; typically, an excited fluorophore reacts with oxygen (but other reactions can, and do, occur). This oxidation of the fluorophore destroys its fluorescence. When using a microscope, photobleaching is observed as a continued decrease in the fluorescence intensity of a sample. If too serious, this can prevent the capture of images as the fluorophore depletes before an image can be captured.
There are three general guidelines that can aid in the reduction of photobleaching:
1) Reduce excitation intensity. This lowers the amount of fluorescence produced, meaning that a longer exposure must be used, but can reduce the amount of photobleaching during a capture through reducing the rate of fluorophore excitation in the sample. This is generally the least effective methods of controlling bleaching as lower excitation intensities result in longer exposures and (often) equivalent amount of photobleaching.
2) Use an oxygen-scavenging media. This will reduce the oxygen available for photo-oxidation reactions, thus reducing photobleaching. A number of companies sell products for this purpose, but the cheapest (and in my experience, best) method is to use 10-50mM β-Mercaptoethylamine to the media. One caveat: oxygen savaging compounds cannot generally be used with live cells, and can alter the behaviour of some fluorophores.
3) Choose better fluorophores. Older fluorophores like FITC and rhodamine are not very good dyes and photobleach readily (plus are pH sensitive and other issues). newer dyes like invitorgens Alexa series of dyes, jacksons dylight series, the ATTO dyes, etc, are generally brighter and more photostable than old-school fluorophores.
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I want to generate or purchase fluorescent live bacteria to study the uptake of fluorescent nanoparticles after biolistic transformation.
The majority of the fluorescent dyes available are used to stain mammalian cells. Please could anyone suggest a method or recommend from where I can purchase fluorescent labelled live bacteria?
Also, how can I synthesize or from where can I purchase fluorescent nanoparticles?
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I am modeling the necessary RFP(Red Fluorescent Protein) concentration in an alcoholic solution so that the solution becomes visible. To achieve this, I'm trying to relate the visual contrast (relative luminous intensity ratio) and the RFP brightness.
The problem is: this brightness doesn't seem to be related to my naive understanding of brightness.
Can someone, please, tell me how to read and use brightness values given as the product of RFP quantum yield and extinction coefficient?
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I think what is being lost here is that you are not trying to distinguish a biological structure in a microscopic field, but, instead, trying to view broad area fluoresence from a solution. In the former case, the concentration is fixed by the structure, so that the relative amount of light absorbed is related well by the extinction coefficient, and "brightness" of the observed structure makes sense.
In your case (if I understand your problem correctly), you need to understand how much light is absorbed per unit area, on average, over the entire visual field, and then convert that to how much light is emitted. If you are modeling the concentration, then Beer's Law gets you to Absorbance (if you define the path length). Then convert Absorbance to % light absorbed (1-%T). To do things right, you really need to convert this energy to photons, and then you are home using the quantum yield.
Once you have the emitted photons, you need to do two things: distribute them over the emission spectrum, and remember that fluorescence is isotropic, so you're only looking at a fraction of the emitted energy. If you are truly interested in luminous intensity, then you need to account for the photopic eye response as well.
Hope this helps.
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My protein samples are labeled and are kept in dark at -80. I am afraid they might undergo photobleaching if kept for long.
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A short-term solution is to add an enzymatic oxygen scavenging system, like glucose-oxidase/catalase. But this will only work on the minutes timescale and not at a temperature of -80 C. Otherwise, you can consider storing your samples in liquid nitrogen, but I don't think these suggestions will solve your problems, as light is required for photobleaching.
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In order to see the shape of a cell, I stained the cell with propidium iodide after fixation. After mounting the cells on coverslips, I looked at them under the fluorescence microscope, I did see the cell shape and red fluorescence when using the PI filter, but when I switch to FITC filter, I still can see the cell shape with green color fluorescence. Since FITC characteristic wavelength is 495nm/519nm and Propidium iodide is 535nm/617nm, there is no overlap area. Why is this happening?
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no worries, i woudl say that your fitc cube has a long pass filter in it. This will give a nice bright FITC signal, but will result in bleed thourhg/issues with other fluorophores. Once you know your filteres you can enter everyting into either the life tech fluoroviewer (http://probes.invitrogen.com/resources/spectraviewer/) or the BD one (http://www.bdbiosciences.com/research/multicolor/spectrum_viewer/index.jsp) and see the results of overlap, off target excitation etc.
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When I measure F content in HA, the Line of F overlaps with that of P. Because I use Rh source, I found line Fka=90.68 degree, RhLa-4=91.15, RhLa2-4=91.36, Pksat-3=90.90, Pka-3=91.69.
How can I fit the peak profile with all these five lines? For simplification, I chose only three lines for fitting, the result is good as shown. But I wonder if this is accurate enough? The intensity of RhLa-4 is small in this process. Is this result reasonable?
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First of all this broad peak doesn't seem to be an XRF peak, but a continuum. The X-axis should be in Å, (WDXRF) or keV (EDXRF), why is it in deg? Is it a diffraction peak (XRD)? From the shape of this hump (rather than peak) it's completely meaningless to analyze leave alone deconvolution of different elemental signatures. First of all please make it clear whether it's XRF or XRD data.
How can I quantify pixels with the same value (or a range of values; i.e. green pixels) on ImageJ?
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This might be useful to quantify the amount of fluorescence.
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If you go to analyze > histogram, ImageJ will produce a histogram that shows the number of pixels with each intensity value. You can use the arrow to select smaller regions, and then it will generate histograms for the selected region only. Hope this helps.
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Can I compensate with beads for my experiments. I use 4 colours in a single tube (FITC, PE, PerCP Cy5.5, APC). What does it matter if I use beads for my compensation. Why can't I just use beads to compensate? Or how would compensating with cells improve my data? (P.S. anyway compensation deals with cutting off percentages of emissions from possible overlapping fluorochromes, so it is just the property of light, how would it matter if cells autofluoresce or not?)
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Hi Sam,
Because fluorochrome spillover between channels is completely independent of the autofluorescence (AF) of your compensating particle, compensation using beads (eg BD comp beads) is much better than using cells under almost all circumstances.
The only relevant consideration with AF with respect to compensation is whether the specific "positive" and "negative" single stained control particles/cells have identical AF. This is hard to determine in a mixed biological population.
Beads have highly defined/uniform autofluorescence (AF) whereas individual cell populations can have distinct patterns of AF over a range of emission channels. This means that if using cells for single stained compensation controls, you would ideally gate your stained population initially based on scatter/AF and only then would you look for positive/negative flourochrome staining (so that the neg and pos cells have the same AF). This is not necessarily trivial, or even possible for all populations/Ab combinations. If you are interested in lymphocytes, for example, you can probably do this quite easily, but most myeloid populations (eg monocytes, neutrophils eosinophils and tissue resident macrophages) are a different matter have distinct patterns of AF.
Bottom line...use beads wherever possible!
How can I eliminate the autofluorescence of a sample of insect eye?
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The sample is without the cuticle and the controls of the immunoassay show that the tissue has autofluorescence.
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Autofluorescence can be very difficult to eliminate. I recommend that you first assess the range of emission wavelengths for your autofluorescence, using tissue that was fixed and prepared for your immunoassay, but without any antibody incubations. If you are lucky, you will be able to select a fluorophore for your assay that fluoresces in a different range from the autofluorescence. However, if the autofluorescence is broad-range (similar to chorion autofluorescence), then a spectral analysis may allow you to subtract the autofluorescent spectral signature from your immunoassay-treated spectral output.
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My solid sample shows highly intense peak at 385nm. But I couldn't the get the peak around the emission wavelength in solution. I carried out the measurement in perkinelmer LS 45. For the solution, the shape of the excitation spectrum is almost the same but shifts in the wavelength.
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I just noticed that I answered another question from you on which I did not get a reply.
Furthermore I read your other questions and conclude that you have to study (much) more before bothering other people with real questions.
For instance the question on HOMO-LUMO and proton transfer shows that you really do not know anything of the subject.
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After running qPCR plates, we would like to analyse them using the LinRegPCR software, that allows for efficiency calculation within each well. It requires fluorescence ratio before baseline correction for background fluorescence.
However, in the 7500 SDS software, we only found 'Export'>'delta Rn' to get the baseline corrected ratio. Such baseline correction is applied during analysis and can either be automatic or manual but can not be eliminated altogether.
Has anybody ever encountered (and hopefully solved) the same difficulty?
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I think you could export the component view to get the raw fluorescence values, but you would would have to do the normalization with the exported ROX values.
How to obtain the coordinates in a CIE chromaticity diagram?
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Can anyone suggest a procedure of how to obtain the coordinates in a CIE chromaticity diagram. I looked on the internet on few websites, which seems to be complicated. Can anyone give me a better way to obtain these coordinates? Here I am attaching a picture from a recently published article FYI.
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This is fairly simple and straightforward if you are familiar with the CIE system of colour measurement. If you have a spectrophotometer you may measure spectral reflectance (or in case of fluorescent specimens spectral total radiance factor). From the reflectance values (from about 380 nm to 740 nm) you may calculate XYZ tristimulus coordinates and from these the xy chromaticity coordinates. There are a number of excellent textbooks explaining the details, but if you have some more questions please contact me at robert.hirscher[at]yahoo.com
Cleavable fluorescence dye.
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Does anyone know a cleavable linker for the conjugation of an antibody with fluorescence dye (for example fluoresceine)? It is important to make the fluorescence dye cleavable. Any recommendations?
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This is similar to what Guus suggested: Take a heterobifunctinal cleavable crosslinker http://www.piercenet.com/browse.cfm?fldID=02030357 React that with NH2 groups of your protein. Then exchange the pyridylthio group with a fluorescent reagent with a thiol group, e.g. thiofluorescein.
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In setting the gates for positive events via Fluorescence Minus One (FMO) controls, do you include an isotype matched antibody in the FMO cocktail, or leave that channel empty altogether?
I.e.: To measure CXCR2 expression on Cd11b+, Gr-1+ bone marrow cells, would your FMO cocktail consist of Cd11b(apc-cy7), GR-1 (percpcy5.5) and Isotype-Pe-Cy7, or would you leave out the pe-cy7 isotype?
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Under almost all conditions it's appropriate just to leave the FMO channel empty as a measure of the population's autofluorescence.
The indiscriminate use of isotype controls is inappropriate for a number of reasons - the most important of which is simply that every monoclonal Ab has individual background binding characteristics due to its unique antigen-binding site. Many companies even specifically select their "isotype controls" based on low binding(!)
Isotype controls can have a place if you are concerned about binding of the Fc region to Fc receptors. Otherwise there are better biological background staining controls, eg:
-is there another population in your sample with similar surface characteristics to your target population that you are certain would not express your marker?
-preincubation with a molar excess of unlabelled antibody of interest (isoclonic control)
-Can you get a knockout mouse/cell line?
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I want to calculate the fluoresce quantum yield. Is there any special software to get the integrated fluorescence intensity? Can anyone give me some suggestions about how to analyse completely the fluorescence spectrum?
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Calculation of quantum yield for a a sample is little tricky. I suggest you to do it manually rather than using any software as software does approximation which will be not known to you and it may lead to wrong answer.
Read joseph lakowicz's book of fluorescence. It describes nicely quantum yield calculation and in a handy way.
Best of luck
Can anyone help me to explain the fluorescence spectra of a metal complex (see attachment)?
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Recently I synthesized a metal complex (mostly insoluble in many solutions) and tried to measure the fluorescence spectra of the compound (I am very new for this measurement). For that purpose, I measured UV for it by using reflectance mode. I observed two maxima at 275nm and 360nm (see attachment). Then I started measuring fluorescence at 360nm. But while I tried at lower wavelengths (<300), the spectra showed a broad peak (see attachment). Then I tried to measure at different wavelengths, from 300 to 400nm and started observing two more peaks at higher wavelengths along with fluorescence peak (390nm. I think so, but not very sure) by comparing all the spectra (I confirmed it by using basic understanding of fluorescence. Fluorescence must not change even we change the excited wavelength). Here my query is, as I am decreasing the excited wavelength, the other peak (other than fluorescence) started shifting towards lower wavelength and merged completely with my fluorescence peak while I excited at 300nm. I also tried to lower the wavelength, but I could see very similar pattern of broad peak from 380 to 580 nm. Is this can be considered as White Light Emission? Can I know what might be the reason for the shift in the peak to lower wavelengths upon exciting at lower wavelengths? For your information, all these fluorescence spectra were obtained by using solid compound.
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Sounds like you have Raman scattering. It is probably easier to interpret a solution spectra.
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What kind of spectroscopes do you use to get reasonable and reliable readings from the ruby fluorescence shift. I have the OceanOptics USB 4000 and I'm afraid that the resolution might be too low. Any suggestions?
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You could use a specialized spectrometer for reading ruby fluorescence shift. There are a lot of such spectrometers in a market. For example - http://www.betsa.fr/pressure-ruby-luminescence.htm
This is description from Betsa website. They are specialized in High-pressure devices.
"The [PRL] spectrometer (Pressure by Ruby Luminescence) has been designed for measuring pressures using the luminescence of a reference material (Ruby or SrB407:Sm 2+).
Its small size and weight make "on site" measurements possible.When equipped with an internal mini laser it functions as a "stand alone" instrument. Measurements are very easy and fast with custom high pressure calculation software."
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I want to conduct an experiment that determines the effect of caffeine on osteoblast uptake of Ca2+. I would like to know which Ca2+ indicator is more applicable and how to determine the appropriate concentration of the indicator to use.
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The use of a calcium indicator will depend on the nature of the calcium signal to be measured and on the equipment available.
Equipement:
Fura-2 is a ratiometric dye and to use it properly you should have an epifluorescence microscope equipped with an illumination source in the UV range to excite the probe and a filter wheel on the emission side capable of rapidly switching between 380 nm (Low Calcium) and 340 nm (High Calcium) . You would then collect emission light at around 500 nm and carry out, either online or offline, a ratiometric measurement of the signal collected at 500 nm after excitation at 340 nm/signal obtained at 500 nm after excitation at 340.
In general, ratiometric dyes enable a measurement of calibrated calcium signal and following a simple calibration protocol you can directly measure calcium concentration. Because of the ratiometric measurement and calibration you are less depend on the equipment and experimental bias. The early version of Fura-2 had rather high affinity constant but there are new version with lower ones.
Fluo-4 dyes are single wave calcium probes (excitation around 490 nm/emission around 520 nm; no ratiometric measurement) for which emission intensity will depend on the level of bound calcium i.e. the more calcium you have the brighter your signal becomes. The Fluo-X version of dyes have been developed to offer brighter signals and therefore a larger discrimination between free and bound form of the probes.
Probe to use:
Fura-2 is a so-called high affinity calcium probe (Kd: 140 nM) whereas Fluo-4 is lower (Kd: 345 nM). This difference is particularly important if you want to measure calcium variations around the basal concentration and if you are not so much interested in the maximum calcium rise. In other words, with Fura-2 you would be able to measure accurately resting calcium levels and small calcium rises but you would rapidly saturate. With Fluo-4 you would be able to measure larger calcium elevation with lower saturation but you would have difficulties to measure resting calcium levels.
The other issue concern the analysis of the calcium signals kinetics. If you want to be able to determine accurately the Calcium transient kinetics and especially decay times, you would need a calcium probe with a so-called fast k-off (a probes that would be able to rapidly release the bound calcium).
See following links on Invitrogen Molecular Probes:
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If both are the same, why should we not use absorption max wavelength for fluorescence quantum yield calculation. I can't use crystals for UV absorption measurement. Can I use a 10mm(cuvette) as the thickness of the crystals for the extinction coefficient calculation. Then how to calculate the quantum yield in the fluorescence spectrum? Is E=h* the new formula for QY calculation? Also clarify the life time calculation.
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absorption spectrum is not the same as excitation spectrum. Absorbance spectrum shows the attenuation of incident light by all interband transitions that occur at that wavelength. Not all of these transitions will yield emission at the same wavelength. The excitation spectrum shows contribution of transitions that result in emission at selected wavelength.
For UV-vis you use the length of sample, which is equal to cuvette size if it is a solution, however, you must take care to use appropriate extinction coefficients for this case. e.g. the coefficients of colloidal semiconductor nanocrystals is not same as that of the bulk material.
The easiest way to determine the QY of a sample is to compare the integrated fluorescence intensity of this sample to that of a standard fluorescence dye of know QY, while making sure that the excitation wavelength and the absorption of both samples at this wavelength are equal. In that case, QYsample=QYdye*(PLsample/PLdye).
Finally, for lifetime calculations, you need measure time-resolved fluorescence in which the decay of fluorescence after the initial excitation is shown. If you fit this curve to an exponential decay function (or multi-exponential depending on your sample), you will obtain your lifetime. Typically, I(t)=I0*exp(- t / lifetime)
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I am wondering about which wavelength is most suitable (1064 or 784 nm) to measure the Raman spectrum for a solid state sample. My organic sample is a white color and the max absorption is at 335 nm in the UV spectrum.
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You might try 785 nm or 830 nm lasers, or any IR laser. Actually you might want to check the absorption curve of your fluorescence source if you can. Sometimes your sample may be only excited by green light (for example), so you can even use red color laser or blue color laser. If you sample can only fluorescing by UV excitation, I will suggest to use shorter wavelength visible excitation (for instance green laser 532 nm), because shorter wavelength visible laser has high energy, and generally it can help you save lots of measurement time. We have an advanced Raman microscope at http://itg.beckman.illinois.edu/microscopy_suite/equipment/raman_imaging_system/. If you need any help, please let us know. Thanks.
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I am wondering about how the interaction distance between the chromophores effects the emission properties?
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We are trying to label human myoblasts and human fibroblasts while in cell culture so that after paraffin processing these can be differentiated. Does anyone know how to do this?
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If I understand your question correctly you want to stain paraffin sections of myoblast/fibroblast cultures to discern the cell types or assess their ratios?
If that is the case you can most easily stain the myoblasts directly by anti-Desmin staining. This could be combined with an anti-Vimentin staining, which stains both myo- and fibroblasts, but DAPI would suffice, if proof of proper cytoskeletal organisation is not your focus.
If you want to perform a direct double staining of both cell types a rather specific fibroblast marker in this context and in human cells/tissues is CD90 (Thy-1).
Staining of paraffin sections has its own difficulties, so please contact me, if you need more help or if I have not answered your question correctly.
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Using a 100X 1.45NA oil objective and an EM-CCD camera. In practice? In principle?
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in theory the field number (typically of the order of 22 mm divided by the transverse magnification (x100) or more if you use an additional magnification changer. In practice often smaller, for several reasons: (i) the laser beam is difficult to focus in the very small back pupil of a high-NA objective with a small pupil such as a x100, therefore the beam is smaller and hence the effective field of view; (ii) in objective-type-TIRF the field of view is often unevenly lit unless the beam is azimuthaly spun (see, e.g., van 't Hoff et al. 2008 and references therein), thus reducing the effective field of view; (iii) depending on your camera, the chip size is clipping the field of view, so to precisely answer you have to know the diagonal of the chip of your EMCCD. A reasonable value seems to be of the order of 50-80 µm field-of-view for your objective. Hope that helps.
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I want to fluorescently label tissues for intravital microscopy, in an unspecific manner. I have used Rhodamine 6G, i.v., and it worked pretty well but I need one in another wavelength.
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As a kid, I used Eosin for tissue staining. It is also fluorescent.
From the link below, you will see you can also use
Acridin Orange
Hoechst stains
Nile Red and so on...
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I have sections of frozen tissue mounted in OCT cut at 8 um. The sections were placed on a single slide, so that the 2 sections would have undergone the same processes/processing, same treatment/conditions at the same time and both are viewed under the same fluorescent microscope settings. However, as you can see (i hope) the first image (control 1) has a blue haze and there is no distinct DAPI staining whereas the second image (control 2) shows the DAPI staining.
What would cause the one section of tissue to not stain properly with DAPI, whereas the other section of tissue is fine?
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In the images, the difference between the "control" and DAPI is caused by autoexposure of the camera. If you check you would find the exposures were for each image are different (control > DAPI) due to the fact that your camera is trying to get the "same" level of staining for both- since the control has nothing to stain the software will increase exposure times to make the overall gray levels equal. if you were to adjust the overall levels of gray so that the backgrounds were equal, the haze would be gone. NOTE the background where no cells are present is "lighter" than in the DAPI section. One gets exactly the same problem in brightfield images when staining in one instance is dark and the other is weak and images are autoexposed. The problem is corrected by clampling the exposure time using a stained section, then the controls should be captured at the exact same exposure as the DAPI + section. A second feature could be the objective/filters. Fluor objectives pass UV light better improving signal to noise ratios. If your system has a UV filter, it should be taken out of the light path. As for seeing the haze with the objectives, your pupils opening more in the dark than the light (and your camera is doing the same).
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I've been trying to obtain the lifetime of some simple fluorophores that I've synthesized. The problem is that the chi-square and durbin-watson parameters of my exponential decay fit are generally far from ideal. Can I trust the results? I'd like some suggestions on how I can obtain good results in this analysis. If I extend the time of acquisition would the statistical parameters enhance? I am not sure if the problem is during data acquisition or data handling. I would appreciate some suggestions.
PS: I am using "Easy Life V" from "Optical Building Block Corporation"
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From your very brief description, it is difficult to judge whether your problem is in data acquisition or data handling.
If the chi-square or Durbin-Watson parameters are not satisfactory, it just means that the selected mathematical model does not adequately describe the experimental data, and the parameter that is obtained by fitting has no reasonable meaning. Hence, you should use another model, e.g. multi-exponential.
In fact, it is quite common that even simple organic molecules, such as e.g. fluorescein yield multi-exponential decay. In order to get a single-exponential decay, one must be quite lucky or use a standard - I use solution of 6-coumarin in ethanol.
There are many possible problems concerning the data acquisition, related to homogeneity, turbidity and stability of the sample, stability of the instrument, correct measurement of the IRF, etc. - quite difficult to advice without direct visual inspection in your lab.
Another question is what is the goal of your measurement, because it is usually very difficult to give molecular interpretation to the decay parameters obtained by fitting. If you care just for the mean fluorescence lifetime (e.g. as a parameter for FRET efficiency calculation), you can use a very simple model-free procedure described in Fiserova E., Kubala M.: J.Lumin. 132, 2059-64 (2012). If you want to estimate number of components in the decay, you can use maximum entropy method (MEM).
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Could anyone provide references or protocols to measure the binding affinity by using fluorescent ligand with protein? Ligand shows fluorescence excitation/emission: 270/340 nm.
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@rupashree how to perform experiment and how to plot scatchard plot to know binding constant and number of binding sites