Science method

Fluorescence Spectroscopy - Science method

Fluorescence spectroscopy is a type of electromagnetic spectroscopy which analyzes the emission of light by a substance that has absorbed light or other electromagnetic radiation.
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I have an oxindole derived compound. To study its photophysics the compd was dissolved in DCM with varying concentrations but when emission was recorded it was observed that at low concentration there is an emission at around 420 nm, however on increasing concentration a low broad peak at near 600 nm starts to appear. Moreover, in low concentration solution where initially on peak was shown at 420 nm, a newer starts to appear as time passes. What could be the reason possibly. Any answers from experts, if anyone who is reading this are acknowledged.
- Bidyut
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It sounds like excimer (excited dimer) formation, meaning that the compound is self-associating. The classic example of this is pyrene.
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Is it possible to get the reverse TRES spectra in TRIS buffer?
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Dear Anjali,
I think (but I'm not sure) in your case the protein is already in excited state (unstable state). So when you give absorbance of particular wavelength it excites to higher state. During emission spectra, it might give lesser wavelength. Please refer following attached case:
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If a compound is dissolved in DMSO and the resultant solution if freezed to solid form (as DMSO freezes at nearly 10 ⁰C) what can be the possible implications on the photophysiccal characteristics of the compound or how the characteristics may be expected to be affected by the freezed DMSO compared to liquid DMSO?
Anyone with their valuable suggestions is gratified.
- Bidyut
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Edward Sanders thank you.
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I am trying to study the interaction of my protein with a small molecule ligand by trp fluorescence spectroscopy. My protein has molecular weight of 16 kDa and has 2 tryptophan residues. I am keeping protein concentration 2uM and gradually increasing the ligand concentration (0-10uM). I am setting up separate binding reactions for each concentration of ligand. Excitation wavelength is 295 nm and emission spectra recorded from 310 to 400 nm. Excitation and emission slits both are 10 nm. Scan speed is 100nm/sec. The buffer used in the study has the following composition: 25mM HEPES-NaOH (pH 7.4), 150 mM NaCl and 10% (v/v) glycerol. The ligand is soluble in DMSO and I am keeping a final 2% (v/v) DMSO in all the reactions. I wish to find the binding affinity and stoichiometry from this study.
The problem is that I am not getting any definite pattern in the emission spectra of the protein upon increasing the ligand concentration; I am getting fluorescence enhancement for some ligand concentrations, for others I am getting fluorescence quenching. Please suggest a solution to this problem. (The protein and ligand interacts, that has been checked by other methods).
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Hello.
You need to take the absorption and fluorescence spectra of each concentration of only ligand and subtract them from the corresponding concentration of ligand-protein. If your protein and ligand both absorb at the same wavelengths, UV and fluorescence are not very good methods for the study. You need to correct the absorbance for the inner filter effect as well as the fluorescence intensity by easily available mathematical equations.
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Protein of interest has been purified using denaturing conditions from inclusion bodies and refolded by removal of denaturants and adding refolding enhancing additives. Upon refolding the it delivers catalytic activity. We tried to heat denature this protein and at the same time monitored its intrinsic fluorescence (Trp at Ex.280).Analyzing the 3D structure revealed that most of the Trp residues were not exposed. We got no Unfolding transition, whatsoever.
Also tried to probe nile red along with it at 1:1 ration of protein, but unable to fetch any transition. We also checked the post heated sample for activity, didnt got any. Hence the protein is transitioning from catalytically active state to inactive state.
Protein concentration : 1uM to 4uM
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Hi Diptesh,
You mention that there is some aggregation following thermal denaturation, which suggests that the high temperature states undergo unfolding/aggregation/precipitation. Do you also observe aggregation at low temperatures right after the refolding procedure is finished?
I believe you mentioned that your protein solution is clear in the pre-melting state. In this case DLS could be useful to determine whether you have soluble aggregates present in the solution, which could be indicative of a misfolding pathway towards inactive species. The latter could still exhibit some secondary structure and no detectable tertiary structure, explaining why you did not see a transition using Trp fluorescence. If this is the case CD would be the ideal method to use for the thermal denaturation studies, as you could would be able to monitor both, unfolding and aggregation.
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I am using fluorescence spectroscopy to determine stability parameters for human acidic fibroblast growth factor 1. (FGF1). The fraction unfolded for FGF1 is determined from the ratio of emission at 350 nm to 308 nm corresponding to tryptophan and tyrosine emissions respectively. So, what I need is the fraction unfolded, do I still need to subtract blank from the 350 nm and 308 nm fluorescence reading separately at each denaturant concentration?
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That's something. Thank you.
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1. Can you please suggest to me how to select a reference standard for quantum yield calculation and what are the reference standards available?
2. If at one excitation two emission bands show in emission spectra then how to calculate the quantum yield for it.
3. While taking the absorption and emission spectra of reference standard what parameter need to be considered for e.g concentration of reference standard should be the same as our sample or the absorption value should be the same for reference standard and sample.
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I mostly agree with Prayasee. However I would like to point out the following since these measurements are very difficult to carry out with a reasonable precisoin (estimated experimental uncdrtainty is usually considered to be a minimum of 10%, considering all factors):
- Always consider the integrated spectra (excitation and emission) and not simply the intensity at maximum (especially in your case when you have two emission peaks) since the definition of QY involves the total number of photons absorbed/emitted and absorption/emission bands may not have the same shape.
- You have to be careful with concentrations since too concentrated solutions may lead to re-absorption (inner-filter effect) or to the formtion of exciplexes. Moreover, theory shows that in emission the intensity is proportional to the concentration only up to an absorbance of 0.05 at excitation wavelength! Usually you have a linear dependence upt to A = 0.05-0.1 then a levelling off and then the intensity decreases with increasing concentration.
- If for some reasons (particularly to avoid dissociation of complexes) you have to have solutions with A > 0.05 then you have to correct and replace A with (1-10^-A)
- It is best to have the same excitation wavelength both both the sample and the standard and, also, to have the same absorbance for both solutions; this avoid many errors. In this case, solutions may have A up to 0.1.
- Note that emission spectra (standard and sample) have to be corrected for the instrumental function
- A good way to test if your solutions are "ideal" is to prepare several solutions of the sample (typically with A = 0.01, 0.02, 0.03, 0.04, 0.05, 0.1) and to plot the integrated emission intensity versus A. If everything is OK (no exciplex formation, no dissociation of complex, no inner-filter effect), then you should get a straight line the slope of which is the quantum yield.
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Can we predict/explain fluorescence behavior (conventional steady-state fluorescence spectra) of a material like quantum dot, perovskite, etc. from its pXRD data? Whether the lattice plane arrangement have a major role in fluorescence behavior of these type of materials?
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Dear Dr Saptarshi Mandal . See the following useful RG link:
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Is a binding site value of 0.8 a good value in fluorescence spectroscopy analysis?
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Hello.
By binding site value, are you referring to the Hill coefficient, n? If yes, 0.8 can be approximately considered equivalent to 1, implying a mono-site binding. In moet cases, the values obtained by fitting are a little higher than 1 but 0.8 should work.
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Dear colleagues
I would like to know if there are some methods to quantify nitrogen
(from urea) in water and/or soil matrix, using UV - Vis spectroscopy or
fluorescence spectroscopy techniques.
Thank you so much for your help.
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There are colorimetric methods for urea determination in the biochem literature, and the indophenol reaction of chloramines with phenol or salicylate is also well documented in compilations of 'standard' methods, e.g., APHA. Nitrate too is readily determined and there are several routes. There is a direct UV measurement, but that is susceptible to organic interferences. There is an ion-selective electrode but it is susceptible to interference from chloride. If you have access to the technology you can reduce the nitrate to nitrite and form and measure azodyes. If you can't proceed that way there are older methods by which nitrate is reduced to ammonia, after which the indophenol reaction can be applied.
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#research #biotechnology #spectroscopy
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Molecular fluorescence spectroscopy is one of the most sensitive and highly selective spectroscopic methods, which can detect extremely low amounts of chemical substances. In the milk industry, it is used for the determination of vitamins, fatty acids, residual amounts of antibiotics, and the identification of different milk species in dairy products.
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I'm conducting research on an anion receptor. I would like to ask, if the PET mechanism can be used to describe fluorescence study or can it also be used for UV-visible studies? Because most of the papers that I'd read discussing about PET mechanism use fluorescence spectroscopy.
Thank you
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i m work on thisd field
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how to calculate stokes shift of a molecule which is having absorbance at 307 nm and emission at 469 nm?
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  • Stokes shift usually calculates in wavenumber (unit = cm-1).
  • Convert to wavelength (nm) to wavenumber (cm-1) [for that, abs 307 nm = 107/307 cm-1 = 325732.9 cm-1 , emi 469 nm = 107/469 cm-1 = 213219.6 cm-1 ,]
  • Stokes shift = Absorption (wavenumber) - Emission(wavenumber)
  • Stokes shift = 325732.9 cm-1 - 213219.6 cm-1 = 112513.3cm-1
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In fluorescence spectroscopy (in the region 200-800nm), generally we get emission at longer wavelength with positive stoke shift, but what will be the reason to get anti-stoke emission of dyes in particular solvent ?
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Anti Stokes emission means emitted photon has more energy than absorbed photon. The origin of this extra energy can be : (i) two photon absorption (where simmultenously two photons are absorbed to excite a molecule and then it jumps back to ground state as one photon having energy higher than both the protons). (ii)hot band absorption (If absorption occurs from higher vibrational states (hot bands) of ground electronic state and emission is common fluorescence, i.e, from excited state to ground vibtrational level of ground electronic state; then emissive energy is higher. (iii) Upconversion process ( There are many mechanisms of this process. Simple one is absorbed photon receives extra energy from some sensitizer to move to higher excited state and from their emission occurs). There are many interesting notes on anti Stokes shift emission. For organic dyes oit could be hot absorption process or some sort of upconversion process.
This is a beautiful paper regarding this
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I am trying to develop a method for quantification of cinnarizine (1-trans-Cinnamyl-4-diphenylmethylpiperazine) in various pharmaceutical products using the fluorescence spectroscopy since UV/Vis method I developed isn't sensitive enough for that purpose. Cinnarizine is cinnamic acid derivative and it is already known that cinnamic acid is susceptible to photoisomerization (from trans to cis form).
I prepared sample of cinnarizine (0,5 ug/mL) using 70% methanol : 30% water solvent and set excitation wavelength at 256 nm (Perking Elmer LS55 instrument). After irradiating the same sample every ten minutes at mentioned wavelength, intensity of the fluorescence emission (309 nm) decreased continuously decreased. I tried using other wavelengths for excitation and other solvents for preparation of the samples but this "problem" remains. I assume that photoisomerization from trans to cis form is going on (or maybe photodegradation)? There are several reported methods for quantification of cinnarizine using the fluorescence spectroscopy (or using the HPLC with fluorescence detector) but none mentioned this problem. Any advices for solving this problem?
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I think looking at the overlay spectrum photodegradation is taking place rather than photoisomerization. You may check the fluorescence behavior of cis and trans isomers in isolation if available with you by following their kinetics.
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I don't know how to fix this bug.
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Dear Irene,
I know this question has been posted along time ago, but I would like to add an answer to this question as other researcher may find it useful.
I had the same problem, I figured out that it was because of the occurrence of high concentration of the molecule I am investigating.
you can solve such a problem by diluting your sample, reading the absorbency of the diluted sample, then multiply the readings to the dilution factor. for example, if you make the 10 fold dilution once, you have to multiply the result by 10.
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Does anyone have experience running any Shimadzu XRF instrument, in particular the EDX8000? We are looking for application notes or advice for analysing geological samples. We appear to be having problems with the software. If you can help, please contact Nathan Halcovitch email: n.r.halcovitch@lancaster.ac.uk
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I think the best advice would be to contact the manufacturer. They should be able to send you the documentation you need, or if they're good, even diagnose what's going on remotely, or help you navigate through the software.
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Dear all,
I am interested to know the methods and detailed analysis of X-ray fluorescence spectroscopy. In general, from the XRF spectra, one can evaluate the elemental and chemical analysis of metals, glass, chemical composition, etc.
I am eager to understand that from the XRF spectra do we can do any other analysis like thickness, refractive index, etc.
Thanking you
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XRF can give you qualitative data, i.e. identifying which elements are present in a sample by looking at their X-ray lines in the spectrum, and quantitative data (either coating thickness or elemental composition).
Quantitative data is obtained through measuring samples against a calibration. Calibrations can be empirical (known reference materials are measured and their elements' x-ray line intensities are plotted against their known concentration/thickness, establishing a relationship between both)), or they can be based on x-ray physics (fundamental parameters method). Depending on the analyzer and the type of sample, you can determine low ppm levels from very high %s, and measure the thickness of multi-layers from nm to several um.
Do not hesitate to contact me if you want more information.
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Photoluminescence quantum yields (PLQY) represent one of, if not the, most important characteristic of a fluorophore. I would say there are two different wide-spread versions to record them:
  1. Relative PLQY: the emission spectrum of a fluorophore is compared to a different flurophore with known PLQY (so-called standard).
  2. Absolute PLQY: by using an integrating sphere, a fluorophore's PLQY can be determined without any further reference measurement.
PLQY can either be indicated as decimals, with values between 0 and 1, or as a percentage, with values between 0 % and 100%. When you read about the accuracy in the determination of PLQY, you will find statements such as:
"It should be noted that the accuracy of the determination of fluorescence quantum yields cannot be better than 5–10%, due to the small additive errors relevant to the absorbances at the excitation wavelength, the correction factors of the detection system and the quantum yield of the standard." ( Valeur, B.; Berberan-Santos, M.N. Molecular Fluorescence: Principles and Applications; Second.; Wiley-VCH Verlag GmbH & Co. KGaA: Weinheim, Germany, 2012; page 161)
However, what does this 5-10 % mean? Is this indicating a relative or an absolute error of PLQY determination? Or, in different words, how accurately can you assign a PLQY? To illustrate, two examples.
I determine the PLQY of a fluorophore as 0.80. If I take the 5 % as relative error, I could not write it more accurately than "0.80 ± 0.04", but as an absolute error, I could never describe it more accurately than "0.80 ± 0.05".
For a different fluorophore, I find a PLQY of 0.20. With relative error, this would be "0.20 ± 0.01", with absolute error, "0.20 ± 0.05"
I find the interpretation as a relevant error counter intuitive, as this means that weak fluorophores have a much smaller (absolute) error than strong fluorophores. In the interpretation as an absolute error, small fluorophores have a much higher relative error, which I find intuitive. However, I feel like a large part of the chemistry community has no problem with publishing a "PLQY = 0.005 ± 0.0003", saying they measured it three times and the relative error is in the order of 5-10 % (let's ignore significant digits here for a second). However, this would be two orders of magnitude more accurate than a determined PLQY of 0.50!
Am I completely on the wrong track here and do I have a completely wrong understanding of accuracy? it only adds to the confusion that a lot of the literature reports PLQY in percentage.
Mind you, this discussion only holds for relative PLQY measurements. For absolute PLQY, the error is 0.02 in the absence of systematic errors (dx.doi.org/10.1038/s41598-019-51718-4). This only increase my doubts on the above interpretation as a relative error, as this would mean that absolute PLQY measurements are only more accurate than relative ones for fluorophores with PLQY > 0.40.
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There are a lot of parameters impacting accuracy and precision of relative quantum yield determination, but you should read the given accuracy values as relative error. We are routinely able to determine quantum yields in the 0.01-0.001 range by using spectrometer equipment that is sensitive enough. Which wouldn't make any sense with an achievable absolute accuracy of 5-10%. Of course, in a statement like "0.80 ± 0.04", the 0.04 value is an absolute number.
Unfortunately, even current textbooks still replicate accuracy statements from the 1960-1970s science, where the total uncertainty was strongly influenced by the achievable measurement precision of that times. With today's equipment, we can achieve PLQY accuracies of 0.02 with moderate effort and even better with some more effort. The uncertainty in the reference QY then becomes the limiting parameter...
I did quite a bit of analysis in that area during my PhD, if you dare to read a German PhD thesis, you can find it in my profile:-)
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I have seen papers where they use small and large unilameller liposomes (size upto few hundred nanometers). But can we measure fluidity of giant unilameller liposomes (size in few micrometers) using the same dye in fluorescent spectroscopy?
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Fluidity refers the "fluid-likeness" of lateral diffusion, i.e. inverse of viscosity. DPH anisotropy reports better on acyl-chain order/disorder that most of the time correlates fairly well with the fluidity, but not always, as exemplified by the cholesterol-induced liquid-ordered phase where the lipids diffuse as in (2D) fluid but are ordered.
If you are putting your GUVs in a cuvette and use fluorometer, then DPH is fine for chain order. If you are using a fluorescence microscope to do anisotropy imaging, you must note that the anisotropy will be strongly dependent of the projection of the membrane. E.g., if you are using DPH-PC (and your should use it) that stays fairly well along the acyl chains and the acyl chains are in a gel phase or otherwise very ordered, then at the top and bottom of the GUV the excitation and emission transition dipole moments will lie in the direction of your lightpath and you will get hardly any signal, and even if you get enough signal, the changes in orientation will all be symmetrical for both excitation polarizations, and your anisotropy is expected to be zero. On the other hands, at the sides of the GUV where the lipid chains and DPH are oriented perpendicular to you light path, you will tend to excite them very well with the excitation light that has polarization along the excitation transition dipole moment, and if the chains are fairly immobile, you will tend to get values close to 1 unlike in cuvette in solution where you would get values close to the maximum value of 0.4. Of course, if the chains were very ordered, and you integrated/averaged the anisotropy signal over the whole GUV surface area, then you would end up close to the value 0.4.
If you are genuinely interested in local variations in fluidity along your GUV bilayer and want to image it, choose a phosphatidylcholine with pyrene attached to the sn-2 acyl chain (often abbreviated PPDPC). Then you can image the excimer/monomer intensity ratio. Here is still the possible problem that it also reports on the local clustering (i.e. signal increases both with increasing rate of diffusion and increasing local mole ferction in the membrane), but this you can see if you are imaging the vesicle.
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I have cells expressing mvenus and mcherry and I need an intrument to quantify the fluorescence of the cells (not a flow cytometer). Can anyone please tell me if there is a fluorescence plate reader at the uni-Düsseldorf?
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Du findest mich unter www.foranthrop.com
Beste Grüße Wolfgang
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Hi, I have a single crystal .cif file and I want to see the electronic distributions under 1GPa pressure and the corresponding structure. How can I get the optimized structure? Can Gaussian be used for this conditions?
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Dear Banchhanidhi,
Please note that neither pressure nor temperature can affect your geometry within optimization. In other words, you cannot expect that your geometry is affected by a change in pressure or temperature.
These parameters (P or T) can only affect the absolute values of thermochemical functions obtained via a frequency calculation. In fact, and depending on the phase of your system, translational as well as vibrational partition functions can be affected by P and T. Consequently, a change in the contribution of corresponding thermochemical values is expected due to change in P or T.
Regards,
Saeed
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The relative light unit measures luciferase activity. I am curious to what the unit is relative to.
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RLU do not have any physical meaning and are often not comparable between different instruments. The output is affected by the PMT mode, the voltage applied, the efficiency of the PMT and other parameters, that make it really difficult to relate the output to any physical magnitude depending on the quantity of light emitted by the sample.  As a consequence, some instruments can give values between 0 and 10 million, others between 0.0001 and 100, others something completely different. Hence, light units are kept relative to other measurements taken in the same instrument.
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I noticed the fluorescence emission of one of my big bio-polymer, which has 2-4% protein is quenching when the another big bio-polymer, which does not show fluorescence emission and does not have protein, is being added. As I am not specialist in fluorescence spectroscopy,
I have three questions here:
1- Is static quenching formula applicable in this case?
2- Is there any way we can conclude the quenching is static or dynamic, without changing the temperature?
3- Is it possible to get the binding site number of 0.5? what does it mean then?
Thank you,
Best regards,
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Static quenching can be distinguished from dynamic quenching by measuring the effect of the quencher on the fluorescence lifetime, which requires special equipment, namely a lifetime fluorometer.
A binding stoichiometry of 0.5 B:A could indicate that it requires 2 moles of A to bind 1 mole of B. This could occur if B binds at the interface of 2 A molecules. It could also result from inaccuracies in the concentrations of A and/or B.
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I have done the equilibrium unfolding of a protein in presence of chemical denaturant (urea) with CD and tryptophan emission spectroscopy. I am getting different delt G (H2O) from both the experiments at pH 7.0. Since the protein is multi-domain having three domains, and TRP fluorescence gives tertiary structure information as this protein has six tryptophan. While in CD (190-240nm) it only give information about secondary structure elements. Please comment on the situations where equilibrium experiments with CD and fluorescence give different results.
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Your curves show that the protein unfolds with intermediate state, since the CD and fluo unfolding curve is not overlapping. Though it is appearing biphasic. To confirm this, u should perform CD and fluorescence studies at different protein concentrations (may be in log ratio, as 1, 10, 100 concentrations). The curves now obtained for either CD/fluo will not overlap i.e. the CD curves will have different Cm values. This will confirm the folding to be non-co-operative. Additional experiments can be done, u can discuss with me if needed or even rajanish may help u out.
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Hi, I am running excitation-emission fluorescence spectra (on algal samples) and I want to use the EEM R package to visualize and analyze the data. However, this package only supports data files coming from the following instruments:
JASCO FP-8500
Hitachi Hi-tech F-7000
Shimadzu RF-6000
Horiba Aqualog
I am using TECAN.
Can anyone send me a data file = .csv, .txt or .xlsx output of a 3D excitation-emission spectrum measurement from any of the above instruments so, that I can adjust my dataset to one of the supported formats? I tried to do it following the EEM manual (https://github.com/chengvt/EEM/blob/master/vignettes/file-io.md#intro) but I can' t make it work. It needs to have certain keywords in certain positions of the file as well as the correct position of the excitation and emission information.
Thanks
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Thanks for this helpful info!
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I am working on analysis of some phenolic compounds via TLC scanning in fluorescence mode.
I noticed a decrease in peak areas upon increasing concentation in all used concentration ranges starting from nanogram/ml range to zeptogram/ml (10 -21) range in one of my compounds. Even in zepto range there are still peaks with large areas and decreasing areas upon increasing concentrations. I can’t establish linearity.
The other compound showing fluctuating readings up and down upon decreasing concentrations.
How to fix this problem and can TLC scanner in fluorescence mode reach single molecule detection or what is wrong?
I am using Camag TLC scanner
  • Mercury lamp
  • Excitation wavelength 230 nm
  • Silica gel 60 as stationary phase
  • K 320 as a filter
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yes I am using Camag TLC scanner 4 ,working on natural phenolic compounds and how degradation can affect readings if I am preparing fresh stock solutions every time and measuring readings instantly after developing plates and air drying the developing system (I also tried oven drying of eluent with the same problem).
I compared it with Agilent Cary eclipse spectroflorimeter, unfortunately it couldn't even detect concentrations in low nanogram range( TLC scanner is far more sensitive)
yes absorption mode gave the same behavior as florescence going up and down in the same way. how to avoid this problem???
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Shift of ThT excitation maximum (from 385 nm to 450 nm) and its emission maximum (from 445 nm to 482 nm) occurs only upon binding to amyloid fibrils ( doi:10.1016/j.bbapap.2010.04.001). And we do not see the formation of amyloid fibirls in the lag phase, right? Then why does the shift occurs at such early "lag phase" stage?
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Devanshu Mehta - Amyloid fibers are likely bound to the surface of the cuvette from a previous experiment. SDS is often not sufficient to remove them. Try washing in aqua regia
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How would the dissociation constant vary if I am (1) Titrating 2uM of protein with increasing concentrations of ligand or (2) Titrating 5uM of protein with increasing concentrations of ligand. Isn't the dissociation constant intrinsic property of ligand/binding molecule or it varies with concentration of protein/substrate in the cuvette.
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Adrian Velazquez-Campoy Can you please share link/paper for correct estimation of Kd(app).
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I have a problem with a spectrofluorometer LS55 PerkinElmer. 
The SCAN FL WINLAB application program does not run. And an error message is displayed: DDE Setup: Client failed to connect to server Lamp September. FL = On.
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I am going to perform ligand and heme-protein interaction studies using Time resolved Fluorescence spectro. I want to know what would be the concentration used in it, or we can use the concentration of normal fluoroscence spectroscopy. As We incubate overnight, the samples to observe the changes between ligand and the protein in UV visible, fluoro, CD spectroscopy and other techniques......My second question is that should i incubate the samples overnight or there is no need to incubate whilst using time dependent fluorescence spectroscopy
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Nilimesh Das if it so brother, then what is the use of Time in this technique,,,,,already we have given a maximum time for the changes to occur.
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The protein HSF1 (6X-His tagged at the N-terminus) was overexpressed in BL21(DE3) strain of E.coli and subsequently purified by Ni-NTA affinity chromatography. 300mM imidazole was used to elute the protein from Ni-NTA agarose beads. This excess imidazole was later removed by dialysis of the eluted protein sample against a dialysis buffer having no imidazole. When freshly purified protein sample was used for tryptophan fluorescence study, good results were obtained. But protein sample stored in -80 freezer in absence of glycerol didn't yield expected result for the same study. What percentage of glycerol is ideal for storing proteins to be used for biophysical studies (Fluorescence spectroscopy, CD spectroscopy)? Can glycerol interfere with such studies? Should glycerol be removed by dialysis from the protein sample before doing the experiments?
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In my experience, glycerol is not essential for storage of most proteins at -80o. What is important is that the protein be frozen very rapidly to prevent separation of solutes from the ice. This means preparing small aliquots and using liquid nitrogen, or powdered dry ice, or a dry ice-ethanol or dry ice-acetone bath to freeze them.
Having said that, it is useful to include 10% glycerol in the samples for storage because it makes it quicker to thaw them out, and may also act as a stabilizer in some cases. I don't think it causes a problem for fluorescence spectroscopy as long as the batch of glycerol is not itself fluorescent due to contamination.
The increased refractive index and viscosity of the solution containing glycerol should be factored into any calculations that require these numbers, such as for dynamic light scattering or fluorescence polarization measurements of molecular weight.
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I am trying to evaluate potential electrochemical halogenation of unsaturated sterols to exploit their properties using fluorescence spectroscopy. I am interested to know if this has been attempted and can this be applied to conjugated aliphatic unsaturated sterols.
Thank you for suggestions.
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Dear Jean,
Thank you for this paper. I also looked searched this topic on the Internet however I could not find any fluorescence evaluation of the produced halogenated derivatives. I would therefore appreciate if anyone could point me in the right direction.
Regards,
Cezary
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Hello,
What are the useful tools (experimental devices) to determine the interaction site (cavity) of a protein with a specific ligand. In my country I do not have access to ITC device (Isothermal titration calorimetry). I have also used fluorescence spectroscopy but I need accurate tests to determine the site of interactions.
Thanks
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X-ray crystallography is the best method. Perhaps you can find a collaborator to help with this.
For large protein complexes, cryoelectron microscopy can also be used. Obviously, you would have to collaborate with an expert in the field.
Multidimensional protein NMR can be used to identify sites of interaction of ligands with a protein if the resonances are assigned to specific residues. This is a very complicated and specialized study, so you would need to collaborate with an expert.
I don't know whether this applies to your situation, but it sometimes happens that if you select for mutant cells resistant to a drug, the mutations occur in the target protein in and around the drug binding site, but not always.
If you create a modified version of the ligand that has a mildly reactive chemical moiety, or a photoaffinity analog, you may be able to identify residues in or near the ligand binding site using proteolytic digestion and mass spectrometry by comparison to the unlabeled protein.
If you have some idea of residues that might be involved in binding, then site-directed mutagenesis can be used to test hypotheses. However, these results are not definitive without structural information.
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Can anyone please suggest how is it possible to calculate EEM fluorescence intensities from the EEM graphs?
Should first and second order Raleigh scattering regions (FORS/SORS) be taken into consideration when calculating the intensities?
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Dear Dimas,
Thank you very much for the valuable hints. The paper is in fact discussing what I needed to know. Greatly appreciate your help!
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Hi everyone,
I am reading an article which has measured "extinction spectra" for a dispersion of MoS2, but hasn't explained the details. I wish to measure the same spectrum for a dispersion of graphene flakes obtained by aid of a surfactant. Actually, I don't know much about the difference between "extinction spectra" and "absorption spectra".
I would be deeply appreciated if someone could explain the concepts and the applicable way to measure an extinction spectra.
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For definitions check https://goldbook.iupac.org/
extinction This term, sometimes used as equivalent to absorbance, is no longer recommended. (https://goldbook.iupac.org/html/E/E02293.html)
absorption spectrum The wavelength dependence of the absorption cross-section (or absorption coefficient); usually represented as a plot of absorption cross-section versus wavelength of the light (https://goldbook.iupac.org/html/A/A00043.html)
Sea also for "extinction coefficient", "attenuation coefficient," "molar absorption coefficient"
Read textbooks for "the concepts and the applicable way to measure an extinction spectra. "
extinction coefficient attenuation coefficient
Analogous to absorption coefficient but taking into account also the effects due to scattering and luminescence. It was formerly called extinction coefficient.
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I have a chemosensor which after binding analyte shows white emission. Now how can I plot CIE diagram?
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Hello subhash,
Just go for google search, type Gocie then you will get direct link. From there you can download the software easily.
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Is there any book to study of fluorescence spectroscopy at reasonable price
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Principles of Fluorescence Spectroscopy - Joseph .R.Lackowicz is a great book for learning Fluorescence Spectroscopy
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Zinc-tetrakis(4-N-methyl pyridyl)porphyrin has an absorption soret band at λ=437 nm whereas λexcitation=458 nm. ( λemission=640 nm).
How can we explain this difference??
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I would like to revert to the definition of the terms. I think when one defines precisely the terms there will be smaller difference between the people.
If one define the photo excitation of the material as: In order to excite a materials by photons, the photon energy needs only to be just equal to its energy gap. In order to absorb a photon its energy needs to larger or equal to the energy gap. Look at the absorption curves of the material as a function of lambda.
So absorption wavelength may be smaller equal to than the excitation wavelength
Best wishes
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Hi There,
I am trying to use the bear Lambert law for fluorescence spec of my sample. in the equation I is the intensity of excitation light, I do not know how to find the value of I from my data. Anybody have idea?
Thanks,
Sima
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Beer Lambert Law is not for fluorescence intensity measurement, Fluorescence intensity usually defined in Arbitrary Unit (A.U.) a Procedure Defined Unit (PDU). The general equation for the measurement of intensity could be like this:
Intensity (Em) = Constant * ((Em + noise (Em)) /(Ex + noise (Ex))
Constant : a constant defined by the maker of spectrometer or by anyone making the spectrometer
Em : Emission intensity measured by PMT (or other sensor)
Ex : Excitation intensity measured by photodiode (or other sensor)
of course Em and Ex will be affected by noise
to obtain more global unit, one of the way is by using intensity calibration such as conversion to Raman Unit (R.U.) as explained in the following paper:
Hope this can help.
Sincerely.
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I would like to determine emission time decay and excited state lifetime of QD'S but we only have Perkin Elmer LS 55 Fluorescence Spectrometer can it be use for such measurements?
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yes you can.
by a manual method
mesure the emission spectrat at diffrent time and plot the intensity (maxcimum = f(t))
and you will get your decay courv
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Hi,
I have a protein which is a hexamer (360kDa) and I want to check whether it binds to its ligand (17kDa only) . Will titration of increasing ligand to fixed conc of protein help in getting this information? If yes, How??
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Another possibly useful approach to measure the affinity is to attach a fluorescent dye (e.g. fluorescein) to the ligand and measure the increase in fluorescence polarization of the dye when the large protein is titrated to a fixed concentration of the ligand. There are several other methods, which have been discussed previously on RG (e.g. surface plasmon resonance, analytical ultracentrifugation, microscale thermophoresis, isothermal titration calorimetry). The method you choose depends upon what equipment and expertise is available to you, and how much of the proteins you have.
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Hello. In my laboratory we need to perform the quantification of curcumin in some mice organs after subcutaneous administration of that molecule (“biodistribution” bioanalysis). Our experimental groups are treated and untreated mice, in two different ways (schemes) and we will take samples at different times (days) to check the clearance of the molecule. What we have observed a priori is that curcumin is retained at the site of the injection. We want to use a simple method, such as absorption or fluorescence spectroscopy, and not necessarily HPLC because we don’t have that equipment and we also think it’s not that necessary. Please, I need publications about this approach since I did not find any; most of the determinations are by HPLC. Do you have any suggestions about the procedure or the idea? Do you think it’s a correct approach? Methodologically, what we will do first is a liquid-liquid extraction of curcumin using ethyl acetate, then evaporate the solvent and then reconstitute with a suitable organic solvent for measurement (methanol? ethanol?). Additionally, we want to calculate limit of detection, matrix effects (a priori there should not be such effects) and recovery percentage of the sample preparation process. Do you have any information about this? I would appreciate any protocol or method or tips. Thank you very much.
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Commercially available curcumin, a bright orange-yellow color pigment of turmeric, consists of a mixture of three curcuminoids, namely, curcumin, demethoxycurcumin, and bisdemethoxycurcumin. These were isolated by column chromatography and identified by spectroscopic studies. The purity of the curcuminoids was analyzed by an improved HPLC method. HPLC separation was performed on a C18 column using three solvents, methanol, 2% AcOH, and acetonitrile, with detection at 425 nm. Four different commercially available varieties of turmeric, namely, Salem, Erode, Balasore, and local market samples, were analyzed to detect the percentage of these three curcuminoids. The percentages of curcumin, demethoxycurcumin, and bisdemethoxycurcumin as estimated using their calibration curves were found to be 1.06 ± 0.061 to 5.65 ± 0.040, 0.83 ± 0.047 to 3.36 ± 0.040, and 0.42 ± 0.036 to 2.16 ± 0.06, respectively, in four different samples. The total percentages of curcuminoids are 2.34 ± 0.171 to 9.18 ± 0.232%.
Improved HPLC Method for the Determination of Curcumin, Demethoxycurcumin, and Bisdemethoxycurcumin
Guddadarangavvanahally K. Jayaprakasha, Lingamullu Jagan Mohan Rao, and Kunnumpurath K. Sakariah*Central Food Technological Research Institute, Mysore 570 013, India
J. Agric. Food Chem., 2002, 50 (13), pp 3668–3672
DOI: 10.1021/jf025506a
Publication Date (Web): May 18, 2002
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I do not own a qPCR machine. If a fluorimeter is available, would it be possible to generate a large amount of amplicons from PCR, where there after SYBR Green 1 is added to the PCR reaction vial in the double stranded state, fluorescence is then evaluated to see if the SYBR Green mounts itself in the double strands of the targeted sex genes, and deduce the presence or absence of the male sex chromosome?
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1) Unless you are working with high resolution microscopy and look for the seq chromosome, staining would not help.
2) For your yes/no analysis, you don't need qPCR.
3) For the sex chromosome, there must be some marker gene and specific primers could easily be designed and simple PCR could do the job, of course followed by gel electrophoresis. (there are some spectroscopic protocols available which dont require gel electrophoresis for the yes/no analysis)
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As you know, BSA is commercially available as 96% and 98% pure. It contains also fatty acids. Therefore, i'm wondering how to get BSA purified. 
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If you are trying to polish purified BSA by removing fatty acids, you can run it over a cationic resin.
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I am trying to denature a protein which is known for high stability even at high concentration of salts(Urea and GnCl) . I am currently trying denature it by EDTA; and as per fluorescence spectroscopy results, the apex peak after treatment of EDTA up to 100mM EDTA is 333nm . But after increasing the concentration of EDTA beyond 100mM , there is a peak shift in the sample which is bi-directional- one apex peak is at 313 nm and other at 344 nm. Also there is a gradual decrease in intensity of emission spectra after 20mM of EDTA.
Does this indicates denaturation of the protein?
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Unfolded proteins tend to aggregate sometimes. Aggregate formation can cause some of the chromophores to be buried and other ones to be exposed to the solvent. This explains the bidirectional shift and the occurrence of peaks at 313 nm and 344 nm in the emission spectra.
Tryptophans show red shifted emission as polarity of the solvent increases i.e., on moving from a hydrophobic environment to hydrophilic environment.
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Am working on development of the semiconductor materials for the Photocatalysis processes, and i will need to study the rate of production of hydroxyl radicals. My questions are;
1/ why some people use HPLC and others just fluorescence spectroscopy?
2/ how can I quantify the rate of *OH knowing that we intends to use the terephthalate acid (TPA)?
Thank you
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I would suggest to work with spin traps like DMPO or BMPO
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While performing synchronous mode of Tyrosine amino acid residues in protein at Δλ 15 nm, it shows one single peak and one hump (another peak), what is the possible explanation.
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Raman scattering by solvent?
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I am conjugating a drug on gold nanoparticles. The unbound drug absorbs well between 350-400 nm. The naked gold nanoparticle absorbs between 250-500 nm. After conjugation of the drug to the nanoparticle, the UV features of the drug are skewed and blue shifted. I thought of creating a standard curve of the plain drug and calculate the amount of drug on the nanoparticle by correlation. However, because of this aforementioned problem, I do not know if it is a good idea to use the same technique.
The drug also shows fluorescence. If I take an excitation scan of the nanoparticle keeping fluorescence at 500 nm constant, the excitation spectra matches to that of unbound drug. Also exciting at 350 nm, the fluorescence spectra of the nanoparticles matches with that of the drug. Gold nanoparticles has absolutely no contribution in the fluorescence spectra. I proceeded to make a standard curve of fluorescence with different concentrations of the drug and calculate the amount of drug on the nanoparticle by correlation of fluorescence intensities. Is fluorescence a correct technique to use for quantification of the amount of drug? Any help would be appreciated.
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It may be a good approach to verify that there is drug on the nanoparticles. However, for quantification, you need to assume that the quantum yield in DMSO is the same as the quantum yield in water/nanoparticle conjugate, which is a huge leap of faith.
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I am looking for research papers/data regarding above mentioned tests for YAG and reason or mechanism particular for these peaks.
UV VIS result shows single peak at 205 nm.
Fluorescence spectroscopy with excitation wavelength = 205 nm shows two peaks at 308 nm and 412 nm. Thnakyou.
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Thankyou Verymuch.I will follow your lead.
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Catalase has intrinsic fluorescence due to its tryptophan and tyrosine residues, but I wonder if it can fluoresce at a longer wavelength, due to other residues (such as NADPH). I haven't seen anything reported other than the peak of tryptophan. I analyzed pure catalase in the lab and I saw a weak peak around Ex/Em ~ 310/420nm, but I can't find anything related to that in the literature.
Thank you
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Catalase contains heme. Perhaps that is the source of the additional spectral feature. It could also be a minor contaminant. It looks like a very minor feature compared to the aromatic amino acid fluorescence. NADPH fluorescence excitation and emission peaks are at 340/460 nm.
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I am writing a script for Processing fluorescence spectra. In Order to make it useful for a large audience, I am collecting Sample Data of Export Files from fluorescence spectrometers of different manufacturers (Perkin Elmer, Varian, Thermo Fisher, etc.). Anyone who would like to contribute may record a fluorescence Emission spectrum (excitation 280 nm) plus an excitation spectrum (Emission 350 nm) of water with an available spectrometer and post the Output Text File Here with all necessary information on the Device used for recording (model, manufacturer,etc). If multiple Output File Formats are selectable, pleased provide a Sample of each.
Thank you all for your help! :)
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There is nothing wrong in creating tools for research, this happens all the time. Unfortunately, researchers investing their time into such efforts, are rarely credited for their work, as there is (as we see above) little understanding among researchers about this necessity. However, one these tools are ready, everyone likes to use them:-) How should anyone do research without proper tools?
Apart from this, the GRAMS is becoming really outdated, there was no substantial improvement or updates since nearly ten years, while the prices are staying huge... I can fully understand when people are looking for alternatives.
Speaking about alternatives: the _free_ optical spectroscopy software Spectragryph (http://Spectragryph.com) recognizes 60+ file formats from UV-VIS, NIR, FTIR, Raman, LIBS, XRF and fluorescence spectrometers and has a lot of powerful spectra processing functions. Just that you know.
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Is this data(emission and excitation wavelength ) available from literature ?If Yes please provide me.Thankyou.
(My aim is to get emission and excitation spectrum for YAG
)
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Dear Rishabh,
What do you mean by YAG (Y3Al5O12 doped with Ce3+) or simply host YAG material. For Ce3+ doped YAG which is used in commercially available white LEDs have excitation in the UV-Blue part of the spectrum and the emission in yellow region. Different rare earth doping have different emission and excitation wavelength.
Reference paper is attached herewith.
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Hi,
I wanted to study the conformation changes in BSA using Three dimensional fluorescence spectroscopic analysis in a spectrofluorometer. I am using BSA (10 uM) for this analysis and my parameters for scan are emission wavelength scanning range is from 200 to 600 nm and the excitation wavelength scan range is recorded from 200 to 400 nm at 5 nm increments. 
I am not getting the 4 characteristic peaks that are Peak 1, the Rayleigh scattering peak, Peaks 2 and 3 which are the two typical fluorescence peaks of BSA and peak 4 which is the second order scattering peak. Instead I always end up getting multiple peaks of equal heights at random wavelengths.  What could be the reason for this? I feel my protocol may not be correct or some scan settings need to be changed.
Thanks in advance :)
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You may be seeing water Raman scattering peaks, although they will not be at random wavelengths. They will be at a fixed offset from the excitation wavelength (about 3500 cm-1) when measured in wavenumbers, not in nm. As a result, the offset in nm changes with the excitation wavelength.
You should not allow the the emission and excitation wavelengths to be the same, as this allows a lot of light to enter the detector, potentially damaging it. The emission wavelength should always be longer than the excitation wavelength.
Tryptophan excitation maximum is at about 280 nm. The emission maximum is between 330 and 355 nm, and the emission peak wavelength does not change with the excitation wavelength (the intensity does). Therefore, it should only require 2 scans over a narrow range of wavelengths to capture the pertinent information: one excitation scan at a fixed emission wavelength, and one emission scan at a fixed excitation wavelength. The approach you are using seems unnecessarily elaborate. Also, there is no useful information about the protein in the Raman scattering peak or the 2nd harmonic peak, so there is no point in scanning over such a broad range.
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How to draw Scatchard plot, using data of fluorescence intensity and concentrations of ligands and protein only. Please help..
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To use a Scatchard plot, you have to convert the fluorescence intensity measurements into concentrations of bound and free ligand. The Scatchard plot is a linearization. To avoid having to prepare this secondary plot, you can simply plot fluorescence versus concentration of the titrant and fit the data to the binding isotherm equation or Hill equation using nonlinear regression. This will give you the Kd of the binding pair and the Hill coefficient. (If you are working in the tight binding region, you will have to use the Morrison equation instead).
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Is there any reference about the difference in kd value of protein and ssDNA interaction, although isothermal calorimetry and fluorescence spectroscopy are two different technique.
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Something that can throw off the measurement of a Kd is if the concentration of the receptor (non-titrated partner) is not well below the Kd of the ligand-receptor complex. Since fluorescence and ITC may require different concentrations of the components to get a sufficient signal, you may inadvertently get into this "tight-binding" regime in one method but not the other, resulting in the different Kd, if you are not aware of the problem.
Fluorescence measurements can be perturbed in several ways. Be careful about possible artifacts, which depend on the details of the experiment.
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What are the methods involved? What information can be obtained from it?
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Dear Sir. Concerning your issue about the study of domestic wastewater by UV - VIS/Fluorescence spectroscopy. Estimation of wastewater composition (Chemical Organic Demand – COD and nitrogen-related species such as ammonia and organic nitrogen) for on-line control of treatment plants based on Principal Components Regression (PCR) and Partial Least Squares (PLS) applied to spectrometric data (turbidity, UV-visible and synchronous fluorescence spectra). Satisfactory models were obtained based on the total UV-visible spectra. The synchronous fluorescence spectra should be divided into regions of interest that could be related to urine and humic and fulvic acids. I think the following below links may help you in your analysis:
Thanks 
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Can you help me please,
How can I make sure about the formation J-aggregation of (PIC) dye on solid substrate by Langmuir-Blodgett technique?
Is there any other way except uv-visible absorption spectroscopy?
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Mrs/Miss Asgari,
The uploaded spectra are IR. These are completely different spectroscopies. There was question about your Fs spectra. Can you upload them?
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Is it possible to measure it with the help of basic fluorescence spectroscopy?
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Dear Michal Vik,
Thank you so much for your answer. Preparing some basic equipment with some modification might help me to solve this problem. Could you please explain in detail this metal strip which i can prepare?
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I am curious how it could be determined by using conventional spectroscopy techniques.
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Define conventional? The presence of an unpaired electron and its chemical properties in relation to a SOMO model can be clearly obtained with EPR. If really only a check of the presence of a unpaired electron is desired, a SQUID could suffice. If you want to use an optical technique, and assuming you are dealing with a transition metal complex, you'd have to find a method to take advantage of spin-orbit coupling effects, but that would depend on the particular electronic structure of what you are trying to study.
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I am trying to compare compound fluorescent activity in different solvent. first i observed Absorption spectra of compound, then found max absorption wavelength/ excitation wavelength. on this excitation wavelength excited to observe emission of the compound in different solvent. and also run the blank (solvents) on same excitation  wavelength. now blank showing the peak (solvents) on this excitation wavelength. now i want to know that, should i subtract the solvents emission spectra from compound emission spectra. And solvents emit spectra or do not?. or only compound can emit, solvent not.
please, please suggest me, my work is stopped at this stage. 
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The apparent fluorescence you observe for the solvent is probably Raman scatter, or a contaminant. Subtract the solvent spectrum from the compound-in-solvent spectrum to obtain the compound spectrum in that solvent. If the compound spectrum is very weak compared to the solvent spectrum, this subtraction may be inaccurate. To improve the result, use a high concentration of the compound and minimize the dimensions of the cuvette. By minimizing the dimensions of the cuvette, you reduce the amount of solvent through which the light passes, thus reducing the solvent background. The compound concentration can be increased up to a point, but once the concentration gets too high, the absorption of light by the compound will begin to affect the spectrum (inner filter effect). Measure the compound spectrum at a range of compound concentrations. You should see that the fluorescence intensity increases in direct proportion to the compound concentration until the inner filter effect starts to become significant, at which point the increase will be sublinear.
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Which emission is more likely to occur or will have more intensity then? How can we justify this with selection rules? Any references and comments will be highly appreciated. 
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There is the famous Kasha's rule, even in Wikipedia
Exception is azulene
As for the simplest answer: when you excite a molecule/entity in the first excited electronic state it will emit from there
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Hi, 
I am trying to determine critical micelle concentration using nile red and I am using Agilent Cary Eclipse Fluorescence Spectrophotometer. I don't know why I am I always getting values under the base line (negative values). I see positive values at nile red emission wavelength but I don't have a good baseline. Does anybody know what it is the underlying reason for this?
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Dear Colleague,
If you really can measure negative fluorescence, then you should go to Stockholm to accept a new Nobel Price for the discovery of "Antiphotons", may be an analogon to ANTI-Materia.....Until you get the invitation from Stockholm follow your way by asking the question: WHY do I see a funny phenomenon? Many people can help you to find the bug. e.g.
Often you can make an error with the baseline or reference fluorescence. Even in full darkness any fluorimeter can indicate a positive or negative signal due to a non-precise setting of the instrument signal for complete darkness = zero light = no photons, but not necessarily NO CURRENT.
If the reference signal refers not to zero photons, but to a reference light beam, then you may be trapped by quenching-phenomena and you see less fluorescence.
An other trivial situation can be due to the setting of the zero light (or reference light) from where ever it can come. An other case is possible that your "lower signal" is not due to a different intensity of photons but due to any kind of electric offset due to a slightly wrong setting of the dark current in real darkness.
You will find the reason, which will be easily corrected. The most important thing is, that you DID OBSERVE THE PHENOMENON AS AN ARTEFACT and that you where taking actions to get rid of it. BRAVO. Good luck for your safe measurements leading to interesting scientific RESULTS. Any time you can contact me for funny questions about Bioenergetics, Photobiology and Spectroscopy:
Bioenergetics Laboratory
University of Geneva (Switzerland)
or at my home:
Hofgutweg 51B
CH 3400 BURGDORF
Switzerland
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For a fluorescent probe which is sensitive to hydrophobic environment, what factors impact on the intensity and what factors impact the wavelength shifting? In our probe, we observed increase in intensity and blue shift in maximum emission wavelength in more hydrophobic milieu. Despite of similarity in pattern between changes in intensity and shifting, in condition 1 we found higher increase in intensity and lower Blueshift, while in condition 2 we found lower increase in intensity and higher Blueshift. My expectation was that increase in hydrophobicity should increase the intensity and Blueshifting in similar manner. How can we interpret this at the molecular level?
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There are different reasons for blue shift and for enhancement of fluorescence. These are two different phenomena. Restriction of intramolecular motion usually causes enhanced fluorescence while the change in environment of fluorophore or restriction of TICT causes the shifting of spectra. When the aggregates are formed the solvent moves out of the aggregates so the fluorophore is now facing different environment. If the polarity of environment is lesser in aggregates then we observe blue shift. If somehow your fluorophore has TICT state, then aggregation can cause restriction of TICT state and leads to enhanced fluorescence and blue shift.
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Hi, I intend to purchase a Fluorescence Spectrophotometer. I am thinking for two models: (i) Shimadzu Model RF-6000 Spectrofluorophotometer  and (ii) Model: F-7100, Hitachi High Technologies Corporations.
Please suggest me which one might be better for research purpose.
Thanking you, Basith
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Dear Dr. Benat,
Thank you very much for your comment and information. Useful indeed.
Best regards, Basith
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I want measuring the   PL decay time  profile of a sample by LS50-B FLUORESCENCE SPECTROMETER(Perkin elmer ) . but I do not know how to measure it.Has anyone done this kind of experiment? Please suggest what I have to do.
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I have never used this model but I have used Cary Eclipse fluorimeter where it is possible to measure long lifetime decay (phosphorescence, rare earth metal) in single shot. If your sample gives normal fluorescence you have to use TCSPC like system (already mentioned by Jose Hodak). What I saw online is that you can measure phosphorescence in LS-50B. If you are able to vary the delay time in phosphorescence mode then you can measure the decay from steady state intensity. The delay time vs integrated intensity plot will be the decay profile. Please see the attached decay of Tb3+ I have measured in Cary Eclipse using this method
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Hello everyone,
I am currently running a lot of protein expression experiments with a plate reader, using GFP as a reporter protein.
I need to try varying different parameters and assess the parameter's influence over the global protein production, a.k.a the fluorescence intensity. However, as the measurements are given in Relative Fluorescence Units, I can make Intra-experiment comparisons but not Inter-experiment comparisons.
I am therefore looking for something that could be use as a standard, allowing me to have a control with a constant fluorescence no matter what. I have looked at the Spectral Fluorescence Standard Kit (Sigma 69336), but its purpose seems to be more machine calibration for experiment reproducibility than providing an experment standard.
Thank you in advance for your help !
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Although it is true that many fluorescence dyes are pH sensitive it is not always a problem. Most such dyes have specific pH sensitivity range and some, such as hydrophilic Quantum dots should have good constant fluorescence. Alternatively, if your protein concentration is high enough and if you can excite in the UV range you can use Tryptophan fluorescence (Max Ex. ~280nm, Max Em. ~360nm)  and it is quite stable between pH 4 and 10.
For Tryptophan fluorescence 
For hydrophilic QD
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In Synchronous spectrum, both Exc and Fl monochromators are scanned simultaneously. MY questions are;
1) How to determine delta wavelength
2) if delta wavelength is 20nm, does it mean the data point at 400nm corresponds to excitation at 380nm? and point at 480nm corresponds to excitation at 460nm.
3) which synchronous technique ( wavelength or energy) is more preferred and informative.
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It depends on what you are looking for. The Excitation Emission Matrix is generally more informative than the Synchronous scan...
The issue is it takes a long time to perform on a system with 2 monochromators so a synchronous scan can sometimes be used in rare cases.
Attached is a schematic of an EEM with an excitation spectrum, emission spectrum and a synchronous scan (all plotted using wavelength).
Attached also is an EEM of water with and without the Rayleigh lines masked. Note the Raman shift in wavenumbers is constant for water... The gap in energy difference may be useful if one wants to look exclusively at the Raman line... 
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I found in literature that authors plot x-axis as emission wavelength and sometimes as excitation wavelength. Is it dependent on spectrophotometer used?
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It doesn't really matter if the difference between the monochromators for the synchronous scan is 30 nm then one can simply add/subtract the 30 nm if they want to change the axis from excitation to emission or vice versa.
If you are unsure how your system works then plot the output from the reference photodiode R1 as well as the signal S1. You know the peak of the xenon lamp is 467 nm. If the plot says its 497 nm then you know your system has plotted x as the emission axis. If the plot says its 467 then you know your system has plotted x as the emission axis...
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 Hello! I am just starting to explore self-labeling tags technologies in the course of my internship in MPIMF Heidelberg, and am playing around with the machines that are available. One is Tecan Spark 20m, and I'm trying to make sense/optimize the usage of it for tag binding efficiency evaluation and potentially other related processes.
I was wondering, what exactly do Gain and RFU parameters do with the signal? I understand that Gain is essentially an amplification of the signal once it's been transformed from light into electricity. But what is the point of RFU? Isn't it somewhat just the opposite of Gain (well, in different units), so what is the actual point of having those two parameters next to each other? Wouldn't it be sufficient to have just one Gain for example? And how do you work with it in your lab? 
Any input is highly appreciated, thank you!
Artur
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Hi Artur, I spend some years of my life  as a system engineer developing the Tecan Spark series.  Pulling nights and weekends, Donating blood, sweat and tears (well, not tears really), creating inventions for 6 new patent applications, and finally get laid off after the product being launched. Tecan readers might be great products, but as an employer, the company utterly sucks...
About your question:
1) it is explained in your manual, just read it!
2) Tecan support has to answer all user questions. Give them a call!
3) it means that you can set the auto-gain to meet a percentage of the intensity scale (RFU), so that if the signal increases later on, you will not run into saturation. This is good for time kinetic cycles, when you expect the signal to increase over time, but still need to decide for a gain value at the beginning. You don't want to have the gain adapted during the kinetics measurement, because there is a non-linear relationship between gain and fluorescence signal.
Now I've done the job for Tecan again, and don't even get paid for it...
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I want to study the fluorescence of a peptide having tryptophan and tyrosine. As I want to excite tryptophan only, I am planning to excite at 295 nm. Its reported that at 295 nm, tyrosine will not be excited.
For this, if I select excitation slit width as 1 nm, what range of wavelength will pass through the excitation slit. Is there any relation between the slit width selected and the range of wavelength that it allows to pass through it?
Intrinsic tryptophan fluorescence of protein. Available from: https://www.researchgate.net/post/Intrinsic_tryptophan_fluorescence_of_protein#5961f4dbf7b67e927e52f094 [accessed Jul 9, 2017].
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The narrower the excitation slit width, the narrower the range of wavelengths (centered on the selected wavelength of the monochromator) that will pass through it. Choosing a narrow slit width like 1 nm gives better wavelength selectivity than a wider one, at the cost of less intense illumination, hence less sensitivity of fluorescence detection. I believe the slit width number refers to the width of the spectrum that passes through it at 1/2 the maximal intensity (full width at half-maximum):
If the signal intensity is too low with 1 nm slits, you can probably get away with 2 nm slits, and/or you can increase the sample concentration.
  • asked a question related to Fluorescence Spectroscopy
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So I have the PL spectra and lifetime transients for a fluorophore in solution (Perylene based polymer in Chloroform) and so by fitting the transients I am able to extract 2 lifetimes one attributed to the perylene emission and one to the excimer that is formed in the solution. I also have the quantum yield for it. The question is how can I calculate the decay rates for each individual species since I have 2 lifetimes and I can't use the simple Q=tau*K equation. I also have the lifetime and rates for the perylene monomer in solution, will this be of any help ?
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1. There must be a difference in decay rates as a function of wavelength when this decay is that of monomer and dimer
2. The dimer emission will grow in with the decay time of the monomer: there cannot be a simple bi-exponential decay?
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I have a doubt and difficulty interpreting my tryptophan fluorescence graph since the concentrations do not seem to be creating a pattern in the curves.
It would be expected that in larger concentrations, such as 1000nM, would have a higher quenching when compared to 800nm and 300nm for example (Figure B). But by looking at the graph we see that the opposite occurs (Figure A). The fluorescence was done in triplicate and similar results were observed.
Anyone have any idea what might be generating these curves without patterns?
Protein: DHPS
Ligand: Pteroic Acid
The data was collected as follow:
Signature=ISS_Experiment_Ver_ 1_0
Product=Vinci2
SoftwareVersion=BETA.2.1
BuildNumber=322
AcquisitionType=Photon Counting
AcquisitionFormat=L
AcquisitionSide=Right
Measurable=Intensity
MaxIterations=10
RightEmissionWavelengthBandwid th=8
ExcitationWavelengthBandwidth= 8
Visualization=Y:Intensity,X: EmissionWavelength,PlotType:2D
EmissionWavelength=type: numeric,unit:nm,from:300.0,to: 420.0,step:1.0
ExcitationWavelength=type: numeric,unit:nm,fixed:295.0
Space=EmissionWavelength
Columns=EmissionWavelength, Intensity,IntensityStdError