Densitometry - Science method

Hello,

I would like to perform a gradient ultracentrifugation with CsCl and then take fractions with a a given CsCl concentration. But I don't quite understand, how can I easily meassure the CsCl concentration? I know there are techniques such as refractometry or densitometry, but I somehow cannot understand the exact procedure of the meassurement. Could someone give me a protocol for this?

Thank you.

Jan

I need the density and viscosity of a molecular fluid under relatively high pressures, I expect something around 500 MPa. This kind of measurement is way outside my field, and I can find only very few groups who appear to be capable of it, all of which refused for technical reasons so far. Does anyone know a collaborator for high pressure rheology and/or densitometry?

I ran a SDS-page of a bacterial lysate and I want to quantify protein concentration in a specific band.

I was thinking of using a standards ladder or make some standards are different concentrations and compare my band to it.

2 things:

1) does anyone have a protocol they could please share with what software they use etc

2) Is it possible that this can be done thought a normal printer scanner instead of a fancy GelDoc?

Thanks.

Good day!

I need to elute proteins from polyacrylamide gel slices after BN-PAGE electrophoresis. The porblem is that the gel slices are relatively big - all the lane fragment upper from 1000 kDa region. Plus we have no equipment for electro-elution. The methods I've found are basically for elution of protein spots after 2d BN-SDS, so they can be unappliable in this case.

The subsequent use of eluted proteins would be 1D SDS-PAGE followed by immunoblotting and densitometry to analyse the difference in quantitative subunits composition of different samples.

Can you advise me on the procedure and conditions?

I plan to:

1. Disrupt the gel slice in a mortar after liquid nitrogen treatment (I wonder if it will damage proteins...?)

2. Put the powder in tubes, add elution buffer and incubate on a shaker (not sure about appropriate time and temperature).

3. Add 4 volumes of cold acetone to supernatant and incubate 1 h at -20°C - precipitate the proteins and so on.

In the attached image, it shows the cleaved PARP densitometry analysis from western blot. Cleaved PARP is commonly used for apoptosis. Treatments (A and B) were tested along with control and the two time points were used 8h and 24h.

I'm trying to run qPCR experiment looking at mRNA levels of apoptosis markers such as BCL2 and PUMA. My question is do I choose 8h for qPCR assuming that mRNA comes before protein so looking at an earlier timepoint is better? Or do I choose 24h since I see the significance in cleaved PARP at 24h in western blot?

When analyzing western blot bands by densitometry in ImageJ, I've usually been measuring the mean gray level intensity of a band inside a region of interest and then using those data.

Instead of measuring mean intensity, how could I go about asking the software to count the number of pixels above a given intensity threshold inside a ROI (if I wanted a different readout) Is this even possible with ImageJ? If so, is there a way to set my desired threshold value? I think this is different from the normal thresholding function in ImageJ that is usually used to demarcate shapes and measure area.

I am struggling to find out how to do this. Any help is greatly appreciated.

This might seem like a rather simple problem, but I am actually unable to find a helpful solution that specifically refers to this problem.

I did several Western Blots of biological replicates (n=4) of the same experimental setup of 4 samples. I quantified the band intensities of my protein of interest and normalized the densitometric values against ponceau staining of the membrane. I chose ponceau staining because the standard loading controls GAPDH and Tubulin seem to be influenced by the treatment and therefore falsify the results.

Now, since the numbers from a densitometric analysis are arbitrary values, they have to be normalized. For this I of course chose my ctrl treatment. Meaning I get values like sample 1 (ctrl): 1.0, sample 2: 4.3, sample 3: 1.3, sample 4: 3.6. I normalized each experiment individually, since they are all on seperate blots. Hence, for sample 1 (ctrl) I got the value 1.0 for each of the 4 biological replicates.

This of course creates problems for a subsequent statistical analysis. Since this ctrl value will have no standard deviation a lot of statistical analyses are technically not possible because you would violate some assumptions they make. I know that I can do a one-sample-t-test if I want to compare sample 2 with sample 1. But I want to do 3 comparisons (S1-S2, S3-S4, and S2-S4) For this I would normally choose one-way-ANOVA with an aapropriate Post-Hoc-test, but I read in several posts that having a value with no standard deviation violates the assumptions of an ANOVA.

So how can I do this analysis? How does anyone do a Western analysis? I mean, the absolute numbers from the densitometry are completely meaningless, because they depend on antibody, ECL, exposure time, and many more things.

I found in several articles where the authors use the quantitative analysis (densitometric) of the WB bands, where in most cases the significant data is not found. I think that obtaining significant densitometric data for bands is very difficult due to the difference between the intensity of the individual experimental data. At least, I have done an experiment several times, and I did not find the densitometric data differed significantly. I used LAS1000 (Fuji Film) for the analysis.

Now my question- is it really important to include this statistical analysis in the paper?

Can anyone suggest some recent papers (especially with/without the densitometric analysis of the phosphorylated protein), or give some important advice regarding this?

Thank you!

Is it reliable to use TLC densitometry for calculating kinetics of a reaction and is there any previous work that uses TLC densitometry for predicting kinetics of a reaction?

Hi,

I am trying to study the expression profile of matrix metalloproteinase (MMP-2) from human mesenchymal stem cells (hMSCs) in an osteogenic pathway. I have conditioned media (CM) collected from monolayer osteogenic differentiation of hMSCs, pass them through centrifuge filters and run them through 10% Novex Zymogram gels and incubate the gels in Coomassie stain and scan the gels. I analyse the bands using Image J software.

My technique: I convert the 600 dpi scan of gels into 8-bit images and from the "Gel" module I select the bands and use "Plot bands" to get the pixel intensity map of the bands and estimate the area under the peaks. However, the pro-MMP2 (~72kDa) and MMP2 (~62kDa) are not well resolved and the peaks overlap. I can't denconvolute the peaks and hence estimate the individual contribution of pro-MMP2 and MMP2. Can someone help me quantifying both the bands? I would really appreciate your help.

I have attached pics to give some idea about my protocol. i am following the article: "Detection of Functional Matrix Metalloproteinases by Zympgraphy" ; doi: 10.3791/2445

Dear all,

For some applications in AUC, we would like to correctly determine the partial specific volume of our biomolecules.

We currently use Sednterp as a principal approach which diplays a theoretical value.

Is someone has a nice protocole to share to calculate partial specific volume on a densitometer? We use a DMA 5000 Tm as a densitometer.

Best,

Sébastien

I am trying to understand if, two different ways of performing enzyme binding kinetics may impact the final Kd values and if yes to what extent for slow and fast reactions.

For more context :

Densitometry couple with Gel based techniques like EMSA, immunoblotting and ELISA would rely on doing these kinetic measurements by allowing varying concentrations of the reactants and observing for equilibrium state. While kinetic measurements done using advanced methods like ITC, Stopped flow or BIA core would do real time measurements for attainment of equilibrium.

Any literature with such comparison will be highly appreciated.It would also be helpful to answer this question by keeping in mind the sensitivity of the technique used here

We conducted densitometry analysis using imageJ on our Western blot (n=5) and analyzed significance using one way ANOVA. However, it always results in not significant, even when it is obvious that the results looks different. This is probably due to low n number and multiple experimental conditions tested. In this case, is it appropriate to use one way ANOVA or are there other statistical tests that can be used?

I am using the "ImageJ" software in order to quantify the protein bands on the SDS-PAGE, but I'm not sure it's the best software for this purpose. Does anyone have the experience of using other software to do this? Is there any professional software for SDS-PAGE densitometry?

I often use Chemiluminescent System for western blots. I always make sure the bands are not saturated and then perform densitometry in ImageJ. Recently, I was told that Chemiluminescent System cannot be used for western blot quantification. Is this true? Any insight? Thank you!

I am doing densitometry on bands on agarose gel, i got band area, height, width, Rf value and adjusted RF value. I don't know what is meaning of Rf ( retention factor)?

i run 2 samples in triplicate and i want to run t test on densitometry data and want to look for p value, i am confused which values should i take from densitometry?

We recently got a new BioRad digital imaging western blot detection system. All is fine except that the software (ImageLab) can export images to TIFF but then refuses to import them again for densitometry analysis. Am I doing something wrong! Surely if it can export a TIFF it should also import a TIFF?

How purity (%) of protein can be determined by densitometry of coomassie brilliant blue stained SDS-PAGE gel ?

Hello all, 

I'm very new to research, and science in general. I just started in my first lab two months ago, so I don't have a lot of experience. 

I'm currently analyzing western blot data. My PI and I are having some disagreement as to the best method to normalize the protein of interest (Neuropsin) to a total protein stain using Ponceau S.

When doing densitometric analysis with AlphaView software of the Ponceau stain, I quantify the total protein for each sample using the entire lane (meaning I make the box to be analyzed around the whole lane). My PI tells me that I should be using only a small band from each lane instead of the whole lane. But it seems to me that this defeats the purpose of quantifying the total protein for each sample because it only quantifies a portion of the total protein. If I'm analyzing the entire band of my protein of interest, but only a portion of the total protein, it seems that the data from normalization won't be reliable. 

Not much literature actually reports densitometric techniques, but in the papers that do, it appears that most researchers are using the entire lane to quantify total protein. Could anyone help clear this up for me? I want to make sure I'm using the most reliable methodologies.

Thank you all ahead of time, I appreciate any responses. 

I have been imaging western blots on a LI-COR Odyssey Fc. After looking into what the densitometry really means, I'm confused as to how only dividing the target protein's density by the same lane's loading control density is the best way to normalize the data. This imaging system uses base 10 logs to describe density, so if, for example, one lane's loading control returns a 1, then the band is 10x as dense as the surrounding background. If the target protein's band returns a value of let's say 0.1, then the band is ~1.26x the background, leading to a target protein / loading control of 0.126. If the next lane returns values "twice" as strong (i.e. target 0.2 & loading control 2), the common normalization procedure would show the same target protein / loading control as the first lane instead of 10^0.2 / 10^2 = 0.0158 != 0.126. Do the lanes actually have different target protein concentrations or am I misinterpreting the connection between density and concentration? Thanks!

What is the difference between scanning a TLC plate at 550 nm (Visible), 254 nm (UV) and 366 nm (fluorescent)?

I am working with a construct containing only the C-terminal region of my POI. It is 42 residues long, and contains no aromatic residues (but does contain a number of basic amino acids). Following SDS PAGE and Coomassie staining there is a large band visible on the gel. However on Bradford assay the concentration is determined to be very low (130 µg/ml). The concentration using BCA assay is a little higher (200 µg/ml) but I still suspect this is an underestimation. I have a number of questions. 

1) Why is there such a difference between the appearance on SDS PAGE and Bradford, since these both use Coomassie based dyes and therefore are not due to differences in amino acid composition?

2) Since I do appear to get good staining on SDS PAGE, perhaps I could estimate the concentration by running BSA standards on the gel and analysing by densitometry. However I am unsure whether it is relevant to compare intensity of staining of a small peptide (6 kDa) and BSA on SDS PAGE in this way? Is molecular weight relevant here?

3) Can anyone recommend any alternative methods?

Any advice much appreciated!

I want to quantify the change in protein expression from cell lysates by western blotting. But first, I need to find the linear range of the signal. In order to do that, I loaded increasing amounts of protein, blotted them with appropriate antibodies, measured the signal, and plotted the results in Excel as a scatter plot.

As expected, more protein gives stronger signal. With low protein loading (0-10 micrograms), there is a proportional response, but with higher protein loads (> 20 micrograms), the signal flattens out. There is a plateau because signal detection is saturated.

I want to fit the data to show the plateau more clearly. How would I do this? I also want to find the linear range of this curve. Do I need to fit a second, straight line?

If I load high amount of protein on SDS PAGE and do densitometric analysis (I am using BIO RAD's Image Lab tool) the relative intensity/amount of other bands increases to higher extent than the desired protein. However if I load a low amount, other bands almost disappear giving me higher percentage of my protein. So what is the standard amount that is loaded to do a relative quantification.

Thanks

Hello everyone, 

I'm researching leaf veination architecture, specifically leaf vein density (LVD), where I need to be able to take high resolution photos of leaves. An article posted by one of the leading authors in this field wrote a paper (Price, et al. 2014) illustrating that because your software's analysis (be it LeafGUI or ImageJ or whatever you choose) will improve with increasing photo resolution there should be a minimum requirement that needs to be met for your data to be accurate enough. They provide a formula to calculate this minimum requirement stating that,

"When a digital camera is employed to acquire images, the image size must be sufficiently large such that there are at least two pixels in the image spanning a distance equal to the magnified resolving power of the microscope."

The formula they propose is 2/(Mdmin) 

Where

M=magnification

dmin=resolving power=λ0/NA

Where 

λ0= mean wavelenth of light (assumed to be 0.53 um for white light)

NA=numerical aperature of the objective lens

My question is; is this formula translatable to a non-microscope imager such as a scanner or a DSLR camera?

Thank you for your time,

Victoria K.

To validate Methylation of the CpG site, I would like to COBRA. So basically the PCR products are digested with REs. These are then run on AGE or PAGE and do densitometry analysis to get %methylation.

I am unsure of the types of REs to be used. Should it be the common ones used in literature or as many isochizomers?

Thank you so much 

I have all my images (.TIF files), but I was wondering what would be the best route to simply determine staining versus unstained portions of my ROIs. We normally use MCID, but getting the correct density settings that work for all the images has proven troublesome. 

I began to wonder, since we have no interest in densitometry, if I even need to adjust them, or measure them for that matter. We usually do densitometry, but for this purpose we aren't really interested in that. 

Would it be as simple as just opening an image and setting the scan area to include all the stained portions of the image? 

I'm new with MCID, but I am also open to possibly learning with ImageJ, since there are so many possibilities available and it is being used almost universally at this point. 

Thanks in advance!

Any publication in which used imageJ.

What are your results? How it works and does it change the standard wb protocol?

Thanks!

I understand that the stainability of a protein with coomassie depends on the amino acid composition of the protein - how pronounced is this effect and is there a way to somehow introduce a correction factor in densitometry based on the known sequence and the amount of basic amino acids? (It seems that the arginine and lysine content is especially relevant).

Could anyone point me to some papers concerning this issue or could share experience?

Thanks in advance!

For the quantification by densitometry of supercoiled and relaxed forms of plasmid DNA stained with ethidium bromide and separated in agarose gel there is a correction factor of 1.4 because the relaxed form gives a fluorescence intensity 1.4-fold higher than the supercoiled form. Should the same factor be used to perform the quantification of plasmid DNA stained with GelRed?

I want to perform densitometric analysis of immunofluorescence in cortical plate (or developed II-III layers) and layer V in different neocortical areas at different postnatal stages and compare them using a factor to normalize, taking in account the cortical growth.

Hi, recently I saw one paper stating that, they estimated the density of the bands on SDS-PAGE using Adobe photoshop application. Anybody knows how to do that? Paper has been attached.

I need to quantify expression from SDS-PAGE-gels and I am not quite sure whether I am doing it right... (I have attached a typical gel in greyscale: lane 1 Marker, lanes 2-5 standard concentrations, all other lanes are screening results with protein only expressed in lanes 6-9). I also attached a picture of the histograms I get from the scans. The question now is: How do I quantify correctly? When I subtract the background, I get negative results from the calculatiobns, since the standard concentrations are lacking the background (they are consisting of pure standard protein)... Is there any way to circumvent that? What would be the correct way to do it?

I often use IF to develop a-tubulin as my housekeeping gene/loading control and I use ultrasensitive ECL to develop weakly expressed proteins on the same blot. Does mixing these two development techniques create any problems for data analysis?

I'm planning on carrying out westerns to assess the change in expression of certain proteins when cells are treated with or without a certain drug by quanitification. How do you ensure that the western is not saturated and is kept in the linear range. I have read that you expose blots to increasing times and draw a plot of intensity and time exposure and see when the line levels off.

Also generally for this method, how many samples would be required. Would three bioligical and three technical replicates be suitable. So for my treated and untreated that is 18 wells altogether. That would be at least 2 gels. Can you quantify lanes on different gels?

Thanks for any help!

I'm trying to get trustworthy densitometry data for some western blots. We've used SCION but I'm told using ImageJ is a better way to go. The problem is that ImageJ apparently needs each lane/box to be exactly the same for analysis and a lot of our lanes are uneven. Does anyone know of a way around this?

To set the threshold for segmenting cells by densitometry in stained immunocytochemical sections I would like to apply the following formula:

Threshold = MG – ((MG/SD) x2)

MG – Mean grey value of the section

SD – Standart deviation of grey values of the section

Please - could anybody could give me some opinion about this procedure?