Questions related to DNA Quantification
I am culturing MSCs on 3D-printed hydroxyapatite scaffolds. We need to detach cells from the 3D to analyze/quantify overall DNA content using Quant-iT PicoGreen dsDNA Reagents and Kit. However, these protocols have not been tested or adapted for complex 3D cultures. We aren't sure what the best method would be to detach & cells from the 3D scaffold and lyse the cells. We also need a technique to verify that our adapted method is effective. I'm interested in hearing what techniques/protocols others are using or any recommendations. Thanks!
Our currently drafted protocol, which is subject to change, involves the following steps:
1. Get DNA standard lysates using PureLink Genomic DNA Kit of cells prior to seeding.
2. After culturing cells seeded on 3D scaffolds for _____ days, at different timepoints, transfer the scaffolds to new wells in 24-well plates so that cells adhered to the wells are excluded.
3. Add TrypLE to the scaffold wells and incubate them, on an oscillating shaker to promote detachment, for 20 minutes.
4. Collect the trypsinized cells and transfer to centrifuge tubes.
5. Add TrypLE to the scaffold wells again and incubate for 10 minutes. Then, repeat collection & transfer of trypsinized cells.
6. Do a 2x rinse using trypLE to try to "knock off" remaining cells and collect as many cells as as possible from the matrix. Repeat until the TrypLE collected is clear, not turbid, hinting that there are little cells remaining in trypsinized suspension.
7. Centrifuge the trypsinized cells to isolate the cell pellet.
8. Resuspend cells in PBS.
9. Follow the protocol in the PureLink Genomic DNA kit to prepare unknown content of DNA in the cell lysates.
10. Follow the Quant-iT PicoGreen Kit protocol to complete the reactions & quantify dsDNA in the samples.
We are considering purchasing a Bioanalyser to increase the accuracy of DNA quantification during library preparation for next generation sequencing. However I have heard that some laboratories now use a Tapestation instead.
Given that Tapestations are more expensive, I wondered if anybody had any advice regarding either of these machines and could offer their own recommendations.
Hi folks - I am buying a nanodrop (or equivalent) for our lab, and Polygen has a version which is less than half price than other instruments on the market. It is the EzDrop 1000. Do any of you have experience with it? Is it OK or not worth saving the money?
I'm processing some leaf samples collected in different rivers for shotgun metagenomic sequencing (Illumina NovaSeq 6000).
The company that will sequence my DNA samples (Novogene in UK) requires a 260/280 ratio =1.8-2.0 (no degradation or RNA contamination).
But I've sent samples in the past (to the same company but for amplicon sequencing) with 260/280 = 1.7-1.8 and they passed the quality control and went full analysis.
Regarding the 260/230 ratio, they do not refer any requirement but my 260/230 ratios are even lower (the lowest is 0.6).
I'm using the DNeasy PowerSoil kit (Quiagen).
So my question is, how low can these ratios be for shotgun metagenomic sequencing?
What's the limit?
I know that I will probably have to clean some samples but I just want to have an idea to help me select the ones I definitely have to clean.
Thanks a lot!!!
I am using the Qubit dsDNA HS Assay Kit with the Qubit 4.
However, I am noticing that I am getting different readings from my standards between batches.
As per the instructions, I make new standards with every batch. I've noticed Standard 1 is relatively consistent at a lower concentration standard ~30 RFU (raw fluorescence units). However, Standard 2, the higher concentration standard, varies significantly and impacts the readings I get between batches.
For instance, in a few batches, Standard 2 was about ~11,000-16,000 RFU, while in other batches, it was 24,000-29000 RFU).
I've had instances where I've double-checked the same sample, and when standard 2 was around 11,000 RFU, the readings were too high- out of range, but when Standard 2 was at a higher level ~24,000-29,000 RFU, the sample concentration was around 44ug/ul.
I have contacted the company, who said it should be working as long as Standard 2 is 50 times greater than Standard 1. But I am noticing differences between batches.
I don't know which RFU is correct for standard 2. I was wondering if anyone could share their typical readings for Standard 2? or any potential trouble-shooting suggestions?
I really appreciate any help you can provide.
I purified about 5 uL of amplicon from about 1 month ago, but after running the quantititation gel, I don't have enough remaining to ideally perform my downstream applications (Sanger sequencing). I still have remaining amplicon which I have now purified once again using EXOSAP-IT. Should I run quantititation gels on this product as well, to confirm the concentrations? Ideally, I would like to simply rely on the results of my first quantititation gel from this amplicon.
The amplicon has been sitting in a +4 fridge for about a month, undisturbed. Please let me know if there's any additional information or details I can provide to help inform an answer - thank you in advance!
You may know we can do a back calculation to get a rough measurement of the concentration of the PCR product we amplified in comparison of both ladder and PCR product intensities. According to the manual.
If we load 4 µL of both ladder and the PCR sample; think roughly our PCR band locates close to 400 bp fragment size. Intensity of our PCR product greater than 2 folds than that of the 400bp ladder fragment, therefore we can roughly say quantity of our PCR product is about 80 ng (According to the manual I have provided).
However my question is can't we do this quantification using Nanodrop Spectrophotometer?
If can what is the blanking solution we have to consider? (If we use Go Taq ® Green master mixture for the above amplification)
I am working on MSCs culture and to normalise the data, I am doing Picogreen DNA quantification. However, the total amount of DNA is downgraded by 30% when I repeated the quantification using stored cell lysate after a week time (-80 degree) and after 2 weeks, further decline was observed. So, is this trend is normal or quantification on the fresh cell lysate is always better to avoid this kind of change in data. Please share your opinions. Thanks
I am looking for a method to normalize alcian blue OD in micromasses, with the amount of cells. Normally cells are fixed, AB is extracted with GuHcl 8M, then OD is measured. At this point is there an easy way to evaluate cells number? I have tried SyBR green quantification, but together with GuHcl I have seen a 6 fold increase in the blank signal when I mix the two of them alone. Problem is that GuHcl is required. I have seen that neutralin red can be used on the same extracts (ref. http://ajpendo.physiology.org/content/299/2/E325.long#ref-29), but I could not find any specific protocol nor explanation. Also, I read that it's used with viable cells, but mine are fixed, so I was wondering whether someone could confirm or explain how it can be used as in the paper I linked.
I know that it's possible to extract GAGs in a cell Lysate, and from there quantify DNA as well, but it is more complex and longer as a protocol.
Thanks for any help!
Basically i need to remove some monocyte macrophages from a 96 well culture plate to store them for later use, which may well be cellular gene expression analysis or more simple viral DNA quantification using PCR. Any help is most appreciated, particularly with regard to the removal of cells from tiny wells. If anybody has protocols to share :) thanks
I have frozen cells (around 20.000-100.000) per well in cell media (280ul total) (without phenol red or any other reactants, just base medium and nutrients).
I need to quantify these cells somehow to have a relative quantification between my samples.
First I thought I could use BCA to quantify, but as I have frozen with medium the concentration (proteins outside in the media due to freeze-unfreeze cycle) is low. I do not know how to concentrate the samples so I could have ALL the protein in a small volume (e.g, 20ul RIPA or Triton X-100).Maybe a freeze dryer would be useful? (not sure if the one in our lab works, though :S)
Second idea would be to measure the DNA content of my samples. My idea would be to use a comercial DNA kit (Blood & Cell Culture DNA Mini Kit we have from Quiagen) and use Qbit and Nanodrop to measure.
Would this give me a relative "quantification" of DNA content between my samples?
I have like 40 wells to do this with and cannot repeat the original experiments, so I will try first with "control" wells I prepared for this purpuse the other day, but any help to design this better would be appreciated.
Thank you! :)
For my internship, I was wondering whether it is possible to perform a DNA quantification measurement without the DNA-extraction from blood. So, you have a pure blood sample and without putting any chemicals in the blood sample you can perform a quantification.
Does anybody know any current researches on this subject?
We have used standard UV-spec and a Nanodrop 1000 to quantify DNA in the past. Can anyone recommend the fastest, most accurate way to quantify DNA? The Nanodrop was, quite frankly, a very slow process and students with poor pipetting skills often missed the pedestal. However, standard specs needed large volumes to be read. Has anyone found a system that you are satisfied with?
I would like to quantify nuclear DNA content in cultured cells for the purpose of identifying copy number changes over time. I don’t need to know specifics of which chromosomes are duplicated or deleted; that will come later if I see any differences in this initial study. I cannot do flow cytometry as I will only be dealing with a maximum of 20-50 cells. The cells can be grown and fixed on a microscope slide. Any recommendations or comments are appreciated!
We are currently running multiple analyses on our culture media (to test what the cells were expressing , DNA quantification, etc..). We have aliquotted several for each type of sample and stored them in -20 degrees, however, we are running out of these ones. I have saved the thawed out aliquots in 4 degrees C for about a month now after taking what I need from them for other analysis. Would it be okay still use these ones?
I quantified the extracted DNA on Multiskan Go. The provider of the instrument directed me to take the absorbance at 260 and multiply it by 1000. According to him this would give the concentration of DNA in ug/ml. I read somewhere else that it is actually the difference in A260 and A320 which is then multiplied by dilution factor and in turn multiplied by 50 which will give concentration. Could someone tell me what is the correct way and if later is correct then what dilution factor is he referring to?
regarding DNA concentration determination by nanodrop , i just read that ((For reliable spectrophotometric DNA quantification A260 readings should lie between 0.1 and 1.0. ))
does that mean that any reading above this range is not acceptable ?
because I have high DNA concentration samples i.e 300ng/ul and I have the A260 is 3 and 4
so can i accept this reading ?
I am currently trying to set up at a new lab in order to begin experiments on osteoclasts. Before starting anything, I wanted to make sure that BMMs have indeed differentiated into osteoclasts by using TRAP staining and PCR.
In our lab, we harvest bone marrow-derived macrophages (BMM) from the tibia of 5 week-old female ICR mice for osteoclast differentiation. For BMM differentiation, I seeded 2 x 10^5 cells per well with full alpha MEM and M-CSF (30ng/ml). I used 3 6-well plates in order to retrieve cells on Day0, Day2, and Day4 of differentiation. On the next day, I retrieved the Day0 plate using 1ml of Tri-RNA reagent and changed the media (containing M-CSF and RANKLE (1:1000)) for the other two plates. I retrieved the rest of the plates on appropriate days.
After retrieving all cells, I performed RNA isolation followed by RNA quantification (ND-1000), reverse transcription, PCR, and gel electrophoresis. My problem here is that I'm getting nothing on gel for Day0 with actin, GAPDH, and HPRT primers. I triple-checked all my steps for gel, PCR, and reverse transcription using other cells and the technique does not seem to be the problem. I performed RNA and DNA quantification using Nd-1000 (I know they are not super accurate) and I've attached the results as image files.
Please help me figure out what made the Day0 bands disappear! Thank you in advance:)
I am currently doing research on Drosophila suzukii (pest of fresh thin-skinned fruit) and my experiments do not seem to be working.
1. I extracted DNA from single fly and from a pool of 5 flies to compare which one works best. I tested the quality using Nanodrop and got the following:
5flies gDNA concentration is 824.1ng/ul, A260/280 is 2.14, and A260/230 is 2.06 (I did not use RNase in the extraction protocol)
1fly gNDA concentration is 241.8ng/ul, A260/280 is 2.11, and A260/230 is 1.90 (I did not use RNase in the extraction protocol).
2. I ran PCR at 45, 50, 60 degrees. Then tested in the gel for amplification. However, I got smears each time. Next, I tested House Keeping gene (HK gene) in Drosophila melanogaster to see if something was wrong with the chemistry. I confirmed that reagents and chemistry was not the issue because I got bands for HK gene at 60degreesC. We also know that HK gene works at 60C only.
3. Next, I ran PCR on single fly and a pool of 5 flies using 1ul DNA template and 2ul DNA template to see which works best. I got smears again.
4. I have ruled out TAE buffer and the gel process as not the problem because I do get bands for the 100bp ladder that I am using. I have ruled out reagents and chemistry as not the problem because they work with HK gene in melanogaster. I have ruled out that it doesn't matter if I use 1 fly or 5 flies because the A260/280 and A260/230 is similar in range for 1 fly and pool of 5 flies.
5. Non-specific binding is maybe ruled out because the primers that I use are universal identification markers for suzukii. I am using CO1 and ND4.
5. Could it be leftover ethanol? or DNA shearing? or non-specific binding? I wash my DNA pellet with 50ul 70% ethanol twice and then leave it to air for 40min in inverted position and 20mins right side up. Last step is to add 25ul of deionized water and let it soak...followed by storing at -20C.
I have a casework trace/contact DNA swabbed samples been extracted using Chelex method. 5uL of the extracted sample was transferred to a clean 2mL microtube and 2uL was used for quantification using Quantifiler Human via ABI QuantStudio5. It gives a reading of 0.98ng/uL. Amplification was proceed and unfortunately no STR profile was developed. Re-quantification of the sample was performed by taking 2ul of the extracted sample directly from original extraction tube and come out with concentration 0.005ng/uL.
All negative control, positive control, reagent blank of the extraction give expected results.
Slope and y-intercept of the quantification are within the range.
Can anyone explain what cause this variation?
In our experiment we are trying to assess the effect of certain drugs on ATP output. We are using an ATP Luciferase kit as a functional assay for ATP production, however the process involves lysing our cell cultures. It would really help sharpen up our measurements if I could normalise the ATP output with a reasonably accurate estimation of cell number from this lysate, either from protein, DNA or another type of quantification method, has anyone done this before?
Does anyone know the DeNovix equipment? Could anyone tell me if it is better than the NanoDrop for microvolume spectrophotometry (DNA, RNA and protein)?
I am trying to quantifying DNA after propidium monoazide (PMA) treatment. I found PMA impacted DNA quantification by either PicoGreen or Nanodrop. For Nanodrop, I got strong background noise from Nanodrop; For PicoGreen, the measured DNA concentration was much lower than the amount I added (I assume PMA reduced the binding of PicoGreen to DNA). I wonder if there is other ways to measure the DNA concentration in the presence of PMA. Thanks a lot.
I want to measure cell viability in a 3D culture. Usually I would use the ratio between prestoblue and Hoechst to quantify the relative viability per cell. Presto blue quantifies the reducing environment due to cell metabolism, Hoechst quantifies DNA - their ratio is "metabolism per cell".
However, my 3D culture matrix is black and the emitted signal is therefore weak (which effects the DNA quantification).
Are there any recommended methods for quantifying the amount of DNA in a cell culture without using fluorescence.
I plan on conducting GBS or RAD sequencing on several hundred tissue samples of Macroalgae (Cystoseira) that were collected previous 1 year. Extract DNA for Macroalgae it's not easy because there are a lot of inhibitors in these tissues. I try to extract DNA with Qiagen power plant pro kit with 24h of tissues incubation (Wilson et al., 2016) and I have these results: 570 ng of DNA measured to Qubit but the purity it to low: 1,58 (260/280) and 0,86 (260/230). With a step of purification (Qiagen power clean kit) the purification ratio is better: 2.06 for 260/280; 0.7 (260/230) but the DNA quantity is divided par two: 250 ng.
Do you think it's feasible to conduct Radseq or GBS for these samples?
I am conducting a study in which I am growing cells underneath a hydrogel for a period of 7 days. The cells are grown in a 24 well plate, and there are 3 different groups. After the 7 day period, the media is removed, the hydrogel is dissociated, and the cells are lysed. Is there a commonly used DNA quantification kit that could be used for a portion of the cell lysate sample? I am aiming to determine the relative amount of cells in the different groups of the study.
I am trying to build libraries starting from RNA, doing first RT with Superscript III and then second strand synthesis with Klenow. I start from the maximum input in RT (good quality RNA, considering both Bioanalyzer RIN number and Nanodrop purity measures) which is 5 ug and according to the dilutions in the different steps I would expect something like 100 ng/ul of ds cDNA. Nevertheless I quantified my ds cDNA after second strand synthesis and I got around 10 ng/ul. I can imagine that the efficiency of the process is not 1:1 (1 molecule of cDNA generate from 1 molecule of RNA input), but 1:10 of the expected is really not good. If I quantify my ds cDNA with Bioanalyzer and Qubit which only measure ds DNA I get such values (10 ng/ul) but if I quantify with Nanodrop which measures any nucleic acid present I get the expected 100 ng/ul. So maybe the RNA is not efficently converted in cDNA or cDNA is not efficiently converted in ds cDNA, and the templates are still there. Has anybody ever tried to measure the efficiency of such a process? Are the concentrations that I get OK or is there something wrong in my workflow?
Thanks so much in advance for any help on this
Why isn't everyone using Sybr Green I instead of Picogreen(Qubit) to quantify low amounts of DNA? It seems to be roughly 10x cheaper and without significant drawbacks.
I am hestitating to switch because it seems systematically under-utilised in the community. Please help me see the missing point (if there is any).
Are there any well established fluorescent dyes (alternatively to PicoGreen) that can differentiate between supercoiled and relaxed DNA, so that they are able to measure the difference in supercoiling of plasmid?
I am trying to get pure cytoplasmic and nuclear fractions from mammalian cells after infection, for further viral DNA quantification. After fractionation and DNA extraction, I quantified mitochondrial DNA in both fractions ( as what I have are DNA extracts) to validate my fractionation, but I got mitochondrial DNA in the nuclear fraction. Perhaps some cytoplasmic structures are pelleted with nuclei. Does anyone have a protocol to get pure cytoplasmic and nuclear fractions, and how to validate it on the DNA level (as I am looking for DNA extraction, not protein, and if there is a better control other than mitochondrial DNA)? Thanks
Is anybody aware if there is a relevant differential dye intercalation between supercoiled plasmid and linear marker during Electrophoresis run? does this difference can affect DNA quantification?
Is there any reliable reference?
Thanks for any advice!
I am looking for a citation to show that even after glutaraldehyde cross linking DNA should be detectable and quantifiable by a nano drop?
I performed DNA extraction from fecal fish contents using the Qiagen fast stool mini kit.
I have the following results in the nanodrop:
A260 / A230 - 2.607
A260 / A280 - 2.074
-Do you think the sample is good for performing qPCR (SYBR green method) for quantification of bacteria?
-Is RNase treatment required?
I'm currently trying to run MNase digestion assays using zebrafish embryos. I have all my protocols down and working well, only problem is I don't seem to be able to quantify the nuclei/DNA so that I can normalise across tubes. Most papers/protocols seem to state that they quantified using A260. I've tried taking sub-samples of my resuspended nuclei and quantifying at A260 on a nanodrop, but don't get any consistency. There is also a lot of interference from other components. I also tried lysing nuclei in the sub-sample to make the DNA more accessible, but still problems. Any help or advice would be much appreciated.
I am using serial samples from a pdx mouse model and so can only take low volumes of blood, but need enough ctDNA to perform ddPCR reactions. I am currently using the Invitrogen MagMax cell-free DNA isolation kit, and i am getting very low yields <1ng.
Hi, I am doing DNA extraction of Mycobacteria like M.bovis or M.avium. I have to analyse the effectiveness of my extraction method. Therefore I have to calculate, how much DNA (weight) can be estimated in the sample I have used.
In real, a had extracted of 1 mL sample a DNA content of 0,1 µg/mL.
1 mL sample used (M.intracellulare cultivated in 7 ml Kirchner Media)
OD at 600 nm of the sample = 0,53 AU
Genom of a Mycobacterium = 4.4 * 106 bp
Weight of a Genom = 650 Da
I need a formula or idea to caluclate the estimated DNA content in my sample. Thx for the help.
I had already quantified my DNA using a Spectrophotometer, and most of the samples were around the 200-300 ng/ul. When I quantify using Picogreen assay, my standards range from 10-1000 ng/ml. I know the conversion is 1000ng/ml= 1ng/ul... So, if one of my samples concentration with the picogreen is 600ng/ml that means I only have .6ng/ul??
I have been trying to find out about this, maybe I'm not doing the conversion right? Please, someone help.
We ideally need to start from quite some gDNA (>20ug total, >300ng/uL).
Most kits, looked at so far, deliver at most 5ug from 1.5ml culture and we found from 4 prep experiments that a second clear band on agarose gel (<500bps) participates to most of the OD (Nanodrop) and that the Qbit measure is 50x less (and far under specs).
A 2-fold RNAse-A treatment did not remove that band!
Any kit that does the job (from larger culture if necessary)?
Thanks in advance
hello i am currently working on bio-materials i am using a protocol to extract Extracellular Matrix from porcine adipose tissue. so far i have done H & E staining, DNA quantification and collagen content.
how to take SEM images of my sample? and what are other test can be done for the justification of appropriate ECM?
i am beginner in this field.
I am preparing some high quality genomic DNA for NGS and some of my sample have high RNA concentrations. I would like to treat them with RNAse, but I've had some previous problems of DNA degradation due to RNAse. So I am trying to figure what will be the best option: leave it with the RNA or add RNAse. Additionally, I've purchased a RNAse that will not be inactivated by heating (only spin column and phenol/chlorophorm), so I was wondering if the left over RNAse will affect my DNA quantification using the pico-green method.
I got a really high value for the 260/280 ratio on a nanodrop. I am not sure if I am just interpreting my data wrong, but my results were:
Nucleic acid concentration: 1.7 ng/uL
Sample type: DNA
The doctoral student that was helping me run the nanodrop said my sample was fine and fairly pure, but I am not sure how he came to that conclusion given the 260/280 ratio.
I used Qiagen mini kit for isolation of RNA from mouse bone marrow cells and spleen tissue. The samples were in stored 4 days before at -80C. The 260/280 ratio in 4 different samples is 1.532, 1.836, 1.991, 0.963 and the 260/230 ratio is 1.156, 1.815, 2.053, 3.519 respectively. The conc. (ug/ml) of my RNA is 77.41, 183.8, 253.5, and 7.451. I have used thermo fisher plate reader with plate compatible to RNA measurement (1ul RNA sample used). I want to use it for RT-PCR gene expression study.
Please suggest me.
I am purifying dsRNA that I intend to inject into insects. I need the dsRNA resuspended in 0.9% PBS.
Usually, I precipitate the dsRNA, resuspend in Nuclease-free water, Qubit, speedvac, and resuspend to the desired concentration in 0.9% PBS.
I would like to avoid the speed vac step and resuspend directly in 0.9% PBS, but I am not sure I will get an accurate reading.
As far as for the qubit manual, the salts concentrations in 0.9% PBS would be very little to interfere with the reading but I would like to know if anyone's tried it before.
DNA quantification using micro spectrometer like nanodrop and gel electrophoresis shows difference in their results. why its higher with spectrometer? Can anyone please share their experience and if possible explanation for this.
We are trying to quantify dna content by Hoechst 33342 staining after seahorse experiment. One of our cell conditions consistently yield higher hoechst staining readouts that we are wondering if it is true or it is because of artifact caused by lipid droplets in the cell. Will lipid content in the cells interfere with Hoechst staining? Are there other suggested methods for DNA content quantification after seahorse experiment?
Thanks for your help
I have 3 animal /group for control (C) and treatment (T) and used housekeeping gene (IC) for each. Now after running gel, Image j densiometric results - lets say are C - 11.9, 13.3, 4.4 ; T- 14.5, 11.4, 5.2 and IC for c -9.2,5.3,5.5 ; IC for t- 5.1,7.6,10.1 respectively. How to calculate 1. Expression level of T compared to C. 2. How to calculate fold change 3. Which type of t-test to verify significance.
I am looking to do hMSCs DNA quantification assay by picogreen assay. does this assay need DNA extraction first as it does not mention that in the protocol? if yes what is the best method for doing DNA extraction ? is it possible to use DNA extraction kit from company other than the invitrogen? or i can use the conventional way for DNA extraction .
I have also run the internal control gene. How to normalise, plot graph and in which units to express the data.
Hi, I have been utilizing the Picogreen DNA quantification assay as a method of approximating cell number increase over a 7 day period in 6 well plates. My question is whether flow cytometry or BrdU incorporation assays would be better options as I see them being more frequently used in papers looking at cellular proliferation and cell cycle.
I have been a tad apprehensive because of the typical use of BrdU assays in much smaller plates making a 7 day assay more difficult, which is necessary as we seem to see the effects of the conditions showing from Day 4-7. Below I have a brief description of the protocol.
-Plate cells and allow them to adhere
-Move cells to different conditions
-Take out plates at days 1, 3, 5 and 7 image
-Lyse with buffer (20mM Tris-HCl, 2mM EDTA, 1% Tween 20 and pH 6.9)
-Scrape plate rubber policeman
-Spin down lysate at 21,000g and pipette out supernatant
-Run picogreen assay in black bottom 96 well plate with negative and positive controls along with standards to make up a 10 pt curve
-Incubate at RT in dark for 4 min then read on PheraStar plate reader
Thanks for any replies.
I am quantifying the DNA on nanodrop 1000. when I dilute the DNA and quantify, it gives more concentration than the diluted concentration. How can I overcome this problem?
I am looking for any kit or method to quantify cellular DNA. We have already done the ploidy analysis successfully by flow but we need to cross check our result. We tried the Pico green kit but it is not working probably due to buffer incompatibility. I will try the quantification by gel this week. I will highly appreciate if anybody can help.
I need to quantify DNA from human pancreatic islets. I plan to use PicoGreen for quantification but dont know what is the best way to isolate DNA.
Does anyone have any experience with isolation of DNA from pancreatic islets? We will have quite a hugh number of samples so a simple method would be ideal. Do you recommend kits or the traditional isolation methods?
I also found this in a paper by Suszynski et al. (2008): 'To measure DNA content, the same samples, which had been analyzed using the CellTiter-Glo and ApoGlow kits for ATP and ADP/ATP measurement were diluted an additional 10-fold in an aqueous solution of 1 mol/L ammonium hydroxide (Mallenckrodt, St. Louis, Mo) and 3.4 mmol/L Triton X-100 (Sigma-Aldrich). DNA content was then determined using the Quant-iT PicoGreen dsDNA assay kit per manufacturer’s instructions.'
Do you have any opinion on this very simple protocol and whether it will be sufficient for the PicoGreen quantification? The link for the paper is here:
Thank you very much for your help.
Standard graph for known bacterial samples ca be prepared with different CFU/ml corresponding to different DNA concentration. But when we want to quantify different groups of bacteria present in fecal sample and we don,t have a pure culture of representative group, how to represent data or with respect to which gene should we normalize the data.
My query is regarding the amount of DNA in my sample using Picogreen assay kit.
I normally use a total reaction volume of 100 microlitre out of which 50 microlitre is 1:200 fold diluted Picogreen (as per manufacturer recommendation) and 50 microlitre is standard DNA or my sample.
For the standard, the final concentration expressed in ng/ml is as per the manufacturer after diluting with TE buffer.
I use a 5 microlitre of a sample (which has been pre-diluted already 10 times before being used) and obtain a DNA concentration of
e.g 800 ng/ml, the actual concentration of my sample would be
= (800 ng/1000 ml)*20*10 = 160 ng/ ml (5 microlitre of sample in 100 microlitre of reaction volume and 10 fold prior dilution of sample).
Please suggest as I got faint band for normal tissues following agarose (1.5%) gel electrophoresis using TBE buffer.
I have some issues regarding the direct SYBR gold assay for DNA quantification. My values are 2- 3 fold off from the NanoDrop value (NanoDrop2000c). Is there anyone familiar with this situation ?
Hello friends I am getting actin band of 150 bp in negative control while in other samples there is no bands
I am not understanding this trouble shoot while I am using the same reagents in all the samples except negative where I am adding nuclease free water instead of cDNA as per rule.
If there is any reagent contamination, then it should show the 150 bp band in positive control while there is no band.
I am attaching the pic of run gel with detail:-
Lane 1 50bp ladder
Lane 2 & 3 - Positive control of actin
Lane 4,5, 6 & 7 - Tested primers for ATPase gene
Lane 8 - negative control showing actin band 150
Kindly reply to this query???
Does anyone have # of adherent cells to seed in 48 well for 7-day DNA quantification assay to study cell proliferation? I would imaging the seeding # will be slightly different from absorbance-based proliferation assays?
Thanks in advance,
The methods involved in qPCR for fungi community analysis. It would be very helpful if anyone can share the information. The quantification of fungal DNA is necessary for MiSeq sequence preparation.
I am aware of both the ∆∆Ct and Pfaffl methods for analyzing qPCR results, the issue is both of these methods are based on a 2 gene (target and reference), 2 sample (treated or test sample and control) model and that is not what I'm working with. Essentially, my experiment is rather unusual as I did not start with RNA. I isolated mtDNA from mussels (who inherit sequentially distinct male and female-type mtDNA types or "mitotypes" from both parents) and performed qPCR on that. So on each sample I ran: a male-type mtDNA-specific primer, a female-type mtDNA specific primer, and a primer that would amplify both types (this being my "endogenous control"). This is supposed to be a sex determination assay (the ratio of male-type to female-type mtDNA differing in males and females). My primer efficiencies are not similar enough to do direct quantification or use the ∆∆Ct method. As a stated previously, the Pfaffl method is designed for comparing Ct number yielded using target gene primer to that yielded using a reference gene primer in 2 samples, treated and control. I know the background info. on my experiment is unusual since this isn't an assay that's looking at gene expression, but there must be something I can do? Can't I just use my known efficiencies to correct my Ct numbers to be what they would be if efficiency were 100% and go from there? I going to assume the answer is no since I've not read of anyone doing this:) I would really appreciate any suggestions regarding how I could normalize my data. Thanks!
Etarget[∆Ct-target (Calibrator - Test)]
RQ = _________________________________________
EReference [∆Ct-reference (Calibrator - Test)]
What my data looks like:
for each sample I need to take into consideration-
Male-specific primer Ct / Male-specific primer efficiency
Female-specific primer Ct / Female-specific primer efficiency
Control primer (amplifies both types) / Control primer efficiency