Questions related to Dyes
I only used 2 wells of 12-well gel for my first electrophoresis. Can I reuse the remaining wells at the end of this electrophoresis? Could the gelred dye in the gel have been affected by the previous electrophoresis? In my view, the dye molecule cannot move without binding with DNA, so I can use the free well one more time. But someone told me I should turn around the gel if I wanna use it one more time to avoid the influence on gelred dye after the first electrophoresis.
I am conducting an experiment where I am putting cultured cancer cells (B-ALL line) in whole blood and running them through a microfluidic device. I stain the cells with DAPI and DiI (cell membrane dye) and then add them to whole blood. The issue that I am facing is that clumps develop in my microfluidic device when I run the sample. I have noticed that the amount of clumps and the time it takes for them to develop are related to the concentration of cancer cells I add in the blood. (The more cells I add, the bigger the clumps and the quicker they develop.) Does anyone have experience with spiking cancer cells in whole blood and recommend any changes? Should I fix the cancer cells before spiking?
I wish to study dye degradation using these nanoparticles but I am unable to find adequate research papers on biosynthesis of iron-selenide nanoparticles.
Looking for technical guide on better dye to stain protein in starch samples for study unders confocal laser scanning microscopy.
Hi everyone! I am new in lab and I have been having problems with Western Blot, I use a Chemidoc and when I reveal I see nothing, after reincubate, or incubating with a new antibody, the signal is lost, or, is very very low, when I dye with red Ponceau, I see a lot of protein because I put 40 ug per lane, I don't have idea about what happened, someone could help me, I will be eternally grateful
HK green eyes are most suitable for the same but they're not available anywhere that's why I'm asking the question.
I have been having issues with compensation whenever I use 3 brilliant violet (BV) dyes together for flow cytometry. I heard BV buffers are the game changers but they are quite expensive. So I am looking for a substitute.
- I am using Polypyrrole Tungsten oxide nanocomposites for the Photocatalytic degradation of Methylene Blue dye(100ml).
- Kindly suggest the various scavengers which I can use and the concentration and amount of scavengers to be added for a 100ml MB dye.
I performed photocatalytic degradation experiment under direct sunlight. The original dye solution was showing lesser absorbance value than the solution kept under direct sunlight in presence of photocatalyst. How can I improve it?
I have prepared an azo dye with orange colour in polar solvents. As is seen in the absorption spectra, absorption is in the UV region and up to around 400 nm in the visible. However, an orange colour solution typically shows absorbance in the 500-600 nm range. This dye also shows a green specular reflectance in a dry solid form. Could anyone please explain the possible reasons for these two phenomena?
I tried staining beads with zombie aqua dye. but the beads did not stain at all. I use 1:400 dilution for my cells. I used the same dilution for the beads as well. do the beads do not stain at all with zombie aqua or do I need to increase the concentration?
I am currently using pHrodo Green AM as a marker for my target cells, which was used in a phagocytosis assay with my M1-like THP-1 cells and I have some questions regarding this specific dye:
(1) pHrodo Green AM was said to be a dye that is able to emit fluorescence on low pH conditions. However, when I observed the fluorescence using flow cytometry (FITC channel), I was able to see some strong fluorescence from the labeled target cells directly. Should this be happening? Considering that pHrodo dyes are supposed to only react with acidic conditions.
(2) I also perform co-culture between the target cells and the M1-like THP-1 (labeled with another dye). However, I saw that after co-culture, there seems to be a decrease in pHrodo Green fluorescence instead of an increase. Has anyone else observed this pattern?
Concerning the adsorbent dose for example iron nanoparticle to remove dye. Sometime it is written as
1- "the run was conducted by adding different dose of iron from 10 to 150 mg to 50 ml working solution of 30 mg/l of dye".
Other research paper mentioned
2- "the run was conducted by adding different dose of iron from 10 to 150 mg/l to 50 ml working solution of 30 mg/l of dye".
In case #1 it is clear that I have to add 10 to 150 mg of adsorbent to 50 ml. But I'm a bit confused in case #2. Should I have to prepare adsorbent solution with that concentration?? Or 10 to 150 mg of adsorbent will be calculated in 50 ml of adsorbate, i.e. in case of 10 mg of adsorbent dose, I have to add 0.5 mg of adsorbent to 50 ml working solution
I use a Geneious Trial Version 11.0.4 to look at my microsatellite data. As my samples run with a custom ladder I altered the ladders.txt. However, Geneious could not fit the ladder. On the support page they say that the last dye is usualy the ladder but in my case it is clearly the dye before. What I am doing wrong?
I have prepared 12% of SDS gel and a sample concentration that needs to load 24 ul in each lane to fulfill the requirement of 4uM protein concentration and the maximum volume of the lane is 20 ul. I have also prepared 2X sample loading dye. How much loading dye should I add to the sample so that it will work?
Is there a specific reason for adding EtBr to the gel instead of DNA sample/dye mixture? I use a RNA dye containing EtBr when running RNA gel so I don't add EtBr to the agarose. Just curious if I can do the same with DNA gel.
I realize that depending on the sample used in the microbiological assay, the same strain of p. aeruginosa produces a bluer or greener color. Is there any mechanism that explains this difference?
A photograph is attached.
Kindly let me know in detail. I wanna check the adsorption kinetic of my composite for dye degradation.
What is value of pseudo first order reaction. is it negative.
I am doing Photodegradation of dye. For calculation of rate constant which value should be taken Ao (zero Absorbance) -pure dye or after adsorption. Graph is plotting with help of origin.?
Has anyone used Sytox Blue, Propidium Iodide, Trypan Blue, Evans Blue, TTC, Alamar Blue, Neutral Red, and FDA dyes to determine the viability of crown and roots in perennial ryegrass and/or plant? If so, could you please share the protocol you used for this purpose?
Despite my efforts, I am finding it difficult to label overnight-grown Crypto cells grown in YPD with Aniline Blue. I have tried to alter the pH of the media I washed the cells and resuspended them in (I've tried pH 4, 6, 7, 9, and 10), the composition of the wash buffer (McIvaine's, PBS, and MES), and the concentration of Aniline Blue (0.05%, 0.1%, 0.2%). I have also suspended the stock of Aniline Blue in the variety of buffers and pHs above, to no success.
I thought my struggles might be an issue with Aniline Blue itself so I included cell wall-disrupted mutants of Crypto to see if cells with greater access to beta glucans would allow for labeling. These cells fluoresced, so Aniline Blue itself is not the issue; the issue is with YPD-grown cells.
This would indicate to me that the beta glucans are not possible to label in YPD-grown cells (for example, due to the capsule preventing labeling), but that cannot be the case since other researchers have succeeded in labeling cells with Aniline Blue with YPD-grown cells (1) or even in capsule-inducing media (2). I have been unable to replicate this success.
I am quite confused by this, as my pellets are always very blue during my wash steps. This tells me that the dye is in some way present in the cells, but that does not translate to fluorescence on the microscope.
Frustratingly, there does not seem to be an "established" means of labelling with Aniline Blue, as methods differ from publication to publication. Methods used on other fungi can vary quite a lot, in fact. Is there anyone that can offer me a protocol for this, or even a suggestion about Aniline Blue that I may be missing? Thank you.
(1) Puf4 Mediates Post-transcriptional Regulation of Cell Wall Biosynthesis and Caspofungin Resistance in Cryptococcus neoformans | mBio (asm.org)
(2) Cryptococcus neoformans Rim101 Is Associated with Cell Wall Remodeling and Evasion of the Host Immune Responses
I know that both Orange G and Bromophenol blue are negatively charged dyes and that Orange G may migrate faster in an agarose gel.
I found a resource that says Orange G can be used in Native-PAGE and in a DNA PAGE but I cannot find any resources that say whether the two dyes are interchangeable in a protein SDS-PAGE.
Any insight would be greatly appreciated.
I am doing work on plant based dyes. we do have colourimeter and UV spectrophotometer. Do any one know an equation or software to convert transmittance or absorbance value to colour coordinates
What are the Mechanisms of Adsorption of dyes from tannery effluents?
What are the detailed procedures involved in the removal of dyes from tannery effluents?
I would like to conduct dimensional gel electrophoresis (DIGE) analysis of lens tissue lysate proteins following protein solubility fractionation. I plan to label fractionated proteins with cyanine dyes for relative quantification prior to isolectric focusing (IEF), and I wanted to know if unreacted dye will affect IEF? If so, then I plan to isolate my proteins from dye using methanol/chloroform precipitation befor IEF. This is my first time trying DIGE, so any advice is most welcome! Thanks!
How long does it take to dye?
How to determine the concentration of dye?
If it is detected by ELISA, will the staining method be different?
I didn't find any publications demonstrating that the targeted cells percent in flow cytometry is only dependant to the Ab-Ag binding, and not to the dye
Can anyone help me please?
often when I document I find in the articles a ratio of absorbances of which I don't know how they determined the two values of this ratio and what is really the use of calculating it
I prepared a ssDNA sample in nuclease free water which is dye labelled and took its concentration using a nanodrop instrument. But the instrument is giving negative values for the dye and positive values for the dna concentration. I hope somebody can help me with this. Thank you
Hydrogen peroxide can be related to the generation of highly reactive hydroxyl radicals in presence of photocatalyst thereby improving its photocatalytic activity in uv light.
How can prove that the dissolution of the dye as the result of the effect of Photocatalytic Activity of nanoparticles not only the effect of peroxide alone?
Many CellROX DNA dye cell labeling methods need test ROS compound preincubation. Then when you add the CellROX dye, the dye oxidizes with the ROS and binds to the cell's DNA to fluoresce.
Is it possible to prelabel the tissue cells with the CellROX dye (non fluorescent) , wash, and then add the test ROS compound to visualize increases in fluorescence in oxidized dye binding to the DNA of the cell nucleus in a tissue.
Is there a better ROS dye labeling method to do this?
I am quite new to this technique and so would require some help. I am trying to quantify the binding of a ligand to a protein using thermal shift assay. I am using Sypro Orange Dye to bind to the protein. I am trying to optimize the protein: dye ratio and have tried the following conditions so far:
1X Dye, 2.5X Dye, 5X Dye and 10 X dye and protein concentrations ranging from 300 nM to 7 uM. The buffer I am using is Tris (pH 7.6) and 150 mM NaCl. However, in all the cases, I am getting a high background signal (buffer + dye only), which is comparable to that when low protein amount is present. What could be the possible reason for this high background fluorescence.
On another note: I am trying it in triplicates and am getting high variability in the signals (Although the trend is consistent). How can I reduce this high variability
They don't take up any dyes, and they don't activate when treated with IL-2. I've confirmed it multiple times that these cells are CD4+ T cells by immunophenotyping (FACS). So why won't these cells take up any dyes? Or lenti? Why do they look like opaque black spots under IF?
Hello the community.
I had an interesting observation recently: I trypsinized HEK293T cells in a 12-well plate and afterwards, rather than adding complete medium, I added PBS to the cells. I transferred the cells to a tube and wanted to pellet the cells down by 500g for 5mins. However, the cells formed flocculant precipitates that floating in the PBS in the end and couldn't be pelleted.
I used a P1000 to forcefully resuspend the cells and used a viability dye to see if the cells were dead (as dead cells tend to be sticky), but they were >99% viable.
Does anyone know the reasons? Thank you very much!
I've been studying oxidative stress on HL-60 and THP-1 cells. However, my cells are GFP-transfected. So, my group decided to use a probe called CellROX™ Deep Red (Thermo: Catalog number: C10422) to study it. I fallow the original protocol:
1.Treat the cells with the test compound or drug. -> I use PMA (10 ng/mL)
2.Add the CellROX® Reagent at a final concentration of 5 μM.
3.Incubate the cells for 30 minutes at 37°C.
4.Remove medium and wash the cells 3 times with PBS
I am reading it using the Flow cytometer using APC and APC-A700 (CellROX™ Deep Red -> Color: Far-Red. Excitation/Emission: 644/665).
But every time I read it, the Fluorescence Intensity is high, and all the cells (99%) are dyed. Even though when diluted it in 2.5 μM; 1.5 μM; 0.5 μM; and 0.1 μM and there is no difference of Fluorescence Intensity when compared the PMA stimulated group.
I know that both HL-60 and THP-1 cells when stimulated using PMA (10 ng/mL for 30 min, 37ºC), they activate the ROS process. I used the DHR protocol as my positive control on HL-60 WT and THP-1 WT, so it worked just fine.
That's why I know my PMA is working and my flow cytometer is working. However, I can not use the DHR protocol on my GFP-transfected cells because both emit a similar fluorescence signal, so we are trying the CellROX™ Deep Red instead.
Do you guys have any tips or similar protocols to help me to reduce the high fluorescence signal/dye pigmentation of my cells with and without PMA?
Thank you all!
Have a great day/night ahead.
I used resazurin dye on my assay for testing out the antibacterial properties of prodrugs. The wells turned pink at a higher concentration of the drug and blue at a lower concentration. I have also carried out an agar well diffusion to see if the drug works at a high concentration and it did, but I can’t think of a reason why this happened.
Does anybody know of a live membrane dye which works well with tissues and isn't internalised or pumped out of the cells in a short time frame? I'm looking to perform 1 or 2 hour imaging experiments on live vessels (ex vivo), however most of the live membrane dyes my lab has previous experience with are only really suitable for very short-term experiments. I should also add that due to the nature of the experiments I plan to carry out, using probenicid to retain the dyes for longer won't be an option.
Any advice would be greatly appreciated! Thanks!
Behavior of the adsorbent at higher or lower pH than the pHpzc in cationic and anionic dye?
I'm using the Qubit fluorometer to quantify dsDNA Broad Range. According to the manual, it should not differ if I using "1-ul sample + 199-ul Working solution (Buffer & dye)" vs. "5-ul sample + 195-ul Working solution". However, I've noticed the the latter's concentration exceeds the former by at least 5x.
I calibrate the machine with fresh standards for each run, and used the same samples on the same day in quick succession of each other. The only difference is the volume ratio of sample to working solution. I am confused about why my experiences conflict with what the user guide claims.
Good afternoon, I am trying to do calcium imaging of MDA-MB-231, U87, and CT26 cells using Cal-520 AM dye which is by all accounts nontoxic to cells, and I have used it for years with no issues...
However, recently, I've noticed that many of the cells become rounded and appear unhealthy after 1hr incubation at 37c. They do not appear to recover after the wash and RT incubation step either. This only began in recent months, but nothing obvious has changed about my protocol/cell types, and I just cant figure out what is causing it...has anyone ever seen such dyes causing cells to round up like this?
Already tried without much success:
- switching dye batches (old to new)
- Incubating in complete growth media/KRB/Hanks + 20mM HEPES
- adjusting concentration between 1, 2.5, 5uM which is the recommended range. I cant see a difference in degree of rounding, but the signal is too weak to use below 1uM...
- Different cell types
- varying cell density
- shorter incubation times. This appears to happen by about 30mins or so, but I haven't yet done a proper time course to know for sure.
Can anyone offer any suggestions?
Thank you in advance
Given that gram and acid-fast staining employ a number of dyes and a decolorizer, is it possible to substitute natural dyes with comparable properties? In acid-fast staining, mycobacteria, for example, contain long-chain fatty mycolic acids. Is it possible to substitute naturally occurring plant-based dyes with lipid-soluble properties?
In our research, leaves of Paulownia tomentosa (Paulownia) were used to extract dye sensitizers with different solvents. Thus, this study aims to identify a new natural dye from Paulownia tomentosa (Paulownia) and investigate the possibility of its potential as a natural sensitizer with simple method extraction. Paulownia leaves have been studied for their light absorbance capacity and spectroscopic analysis. Spectroscopic analysis has revealed the presence of these pigments and their specific roles in light absorbance and photoprotection. The dye extract was characterized by UV-Vis spectrophotometer (Carry 60) to observe the absorption spectra.
I am in search of a process to doubly label the proteins with different fluorophores at different positions in the same protein. Say for instance, If I conjugate dyeA at N-terminal cysteine, how can I block the other cysteine at C-terminal which is supposed to be for dye B. Any suggestions please.
Thanks in advance!
As the question already implies, i made a mistake during restriction enzyme digestion and added the 6x loading dye to the DNA insert fot checking on the gel. 10ul was added to the gel for checking but the other 60ul that was supposed to be used for ligation later on also has the 6x loading dye added to a concentration of 1x. Will this be a problem later? Will the dye be removed during the GeneJet PCR purifications later? How should i proceed? All input is welcome.
Hello, I am currently performing an experiment and need a visual aid for the media which is currently clear. The dye needs to be non-toxic and would not stain the cells in any way. Would regular food dye be safe to use with human stem cells? Should I just use phenol red or other commonly used media pH indicators?
I used the KCl extraction method to extract nitrate from soil samples which were then analysed using a skalar continuous flow analyser. I am doing this to calculate the rate of nitrification from soil samples incubated in situ for a month (amount of nitrate in month 2-amount of nitrate in month 1).
From my understanding, the analyser works by reducing nitrate to nitrite, which forms a dye, and the concentration of the dye is then measured at 540nm.
Some of the values of NO2-NO3 given (ml/L) are e.g. 6mg/L. Other values are stated as <LOD (below the limit of detection). But then I have also been given values which are negative e.g. -0.9mg/L.
I don't understand how to interpret this because it's saying there is less than 0mg/L of nitrate...which is impossible? I was assured there was no issue with the machine and that negative values are to be expected sometimes, but I don't understand why. I'm not sure I'm getting my head around it and would be grateful for an explanation, or links to useful literature that can explain.
Anyway, I don't know how to handle these values- should I replace them with 0mg/L? Should I omit them?
I want to measure reactive oxygen species in whole blood. Can anyone please suggest a protocol or method for it using 2',7'-dichlorodihydrofluorescein diacetate (DCFDA) dye ?
Hi All, I am using cell mask orange dye to label HEK 293 T cells plasma membrane. I am using 5 ug of dye as final concentration and incubating cells for 10 min at 37•C. I am facing issue with dye internalisation. I did staining before fixation as well as after fixation. In both case All dye internalize. So here is the protocol I am using:
1. Growing cells
2. wash with PBS
3. Stained with cell mask orange for 10 min
4. washing 3 time with PBS
5. Fixation with 4% PFA
6. Washing with PBS.
7. Stored in PBS,Same day Imaging.
Note: Some time I pellet down the cells after second step and then stained the cell suspension. At the end step I just mount the sample on slide.
How can I prevent dye internalization in cell? what are the possible reason of this issue? Please let me know.
I've recently started using an Edinburgh Systems FS5 fluorometer with no experience or training other than the instructional videos they provide on a memory pen.
I'd like to know what the best type of 96 well plates are to use for fluorescence emission readings, as I am currently using opaque black plates with a curved opaque bottom. I believe the Fluorometer is top-reading.
I'm using Alizarin Red S which is a dye that will become fluorescent when binding to a boronic acid and emit at 572 nm (excitation 468 nm). However the values and trends I am seeing do not seem to reflect what is expected, and this is shown even more so when I analyse completely blank well plates. I tried completely clear well plates previous to this but got similar results and also read that you should not use this as it reflects the light away from the wells.
I've also tried to use a rhodamine sample to get better results but the emission is similar to that shown for an empty well. Is there a better way to calibrate the laser rather than manually turning the mirrors for the detector etc as suggested in the instructional manual for this specific type of instrument?
In many literature they have mentioned EDTA, IPA and BQ but how much volume and concentration should be used to perform the radical scavenging tests?
Hi Research Community,
I am looking for a method to label dendritic spines in fixed hippocampal brain slices.
Due to our fixation Golgi staining seems to be off the table.
Now I found Dil labelling as an alternative, but most labs use gene guns, that I unfortunately don't have. Has anyone experience with using dye crystals added directly on the tissue or alternatives?
Who can explain the phenomenon !!! that occurs in my experiments when I do a degradation experiment of MO dye using photocatalyst ZnO:Ce where the photocatalyst works to remove the dye within the first half an hour under UV light then the dye reappears again in the solution and the same happened under sunlight the dye disappears within an hour and then reappears again as if nothing had happened at all.
i did the UV-Vis spectrometer for these experiments that approved this phenomenon.
please kindly anyone can help .
give suggestions or explanations