Why does my gel have such poor resolution?

Why does my gel have such poor resolution?

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Brand new user to bio This Site here. I ran this gel (0.9% agarose, run starting at 60V then to 105V over ~30-45 min) with DNA samples containing 20uL of DNA and 1X tracking dye in each of the wells. I do not know why the resolution of this gel is so poor. I'm assuming it's from DNAse activity, as this is what my lab tech said it probably was. I'm pretty sure I did everything right regarding the procedure. I'm not sure what other info I could provide, other than the samples were prepared and stored at -20C a week prior to the gel being run. Here's a picture of the gel below.

Although your tech is referring to your DNA samples, your ladder also indicates there is some room for improvement of your running technique. -20C for a week should not have impacted the quality of the DNA assuming that it was good quality going into the freezer. You can mitigate DNAase activity by adding EDTA to your storage buffer, which will pull metal ions out of solution. Just make sure to take that into account if you're doing a reaction later that requires metal ions (like PCR).


Focusing first on the DNA samples. There are a few things to check.

How much did you load? I see you stated 20uL, but what was the concentration? It it more important to know the amount of ng being used as the sample. Various stains have different concentrations they can detect (EtBr, SYBR, etc.) I would check the specifics of the stain you're using to make sure you're loading an appropriate amount of DNA.

Are these PCR products? If so, how many cycles was your PCR run? I have seen similar results from PCR if the samples are run for too many cycles. Non-specific products begin to be produced in later cycles, and will cause degradation of your specific products. I would also re-check your primers, the extension time, and the polymerase you're using. It is more difficult to do extensions of the 5.8kb and 6.8kb sizes you mentioned, but not impossible.

Are these restriction digests? Do you have a control well showing the undigested plasmid, or other source? Your source should have a clean band first to expect a clean band in the results. If your source is clean and bright, and the digest looks like the samples above, check your digest buffer and enzymes.


There are several things which can improve the quality of an agarose gel. Thermofisher provides a guide for their tips and tricks. Below are things that I have found useful in my own experience.

  1. The time between casting the gel and running it. A common practice is to make gels and store them in TBE/TAE buffer until they're ready to be used. In my experience, the quality of the gel will degrade over time, yielding diffuse bands. The best results are achieved with a freshly poured and cast gel.

  2. Loading technique. Carefully loading the samples into the wells so that they settle as closely to the bottom without mixing with the surrounding buffer is key. The best results are achieved by having your pipet tip touching the bottom of the well, and slowly pulling it out as you release your sample so the sample does not have to settle down into the well on its own.

  3. Run immediately after loading, and image immediately after running. Diffusion will cause nice tight bands to become poor the longer the samples sit in the well.

  4. Voltage, timing, and agarose concentration. Getting these right takes practice. You'll want to try to run your gel as quickly as possible. The more time it sits in the gel, the more diffusion will occur. However, have the voltage too high, and you risk melting your gel. Vary the concentration of agarose to target the size of DNA you're after. I have previously used anywhere between 0.5% and 1.5%.

  5. Prestaining the gel or poststaining? I prefer prestaining for the stains that work well, as it reduces the time the samples will sit in the gel allowing diffusion to occur.

  6. I am looking at your ladder, and the link to the provided standard, and it is difficult to tell which bands are which. The 3.0kb fragment should stand out as the brightest, however the bottom band in your gel is the brightest. Is this the 3.0kb band? Maybe. It's difficult to count the number of bands above it to confirm. If it's not, which might be indicated by the fact that none of your samples seem to have run to the end of the gel, then this would be the 0.5kb band. If it is the 0.5kb band, the ladder intensity doesn't match up with your expected intensity. Double check your ladder so you can know what sizes you're looking at.

Blurry bands . - (Jul/07/2012 )

I've been running SDS Pages for a while . lately I was doing some DNA affinity chromatography experiments and was running the elutions on a 16% gel. My particular interest is the identification of smaller proteins (around 10 - 20 kDa). I don't know why, but apparently the smaller proteins on my gel, appear as blurry bands (see attached file) thus giving really troubles for further MALDI analysis.

Does anyone know how I can compress the lower bands? Would it make sense to run a 10% gel and just stopping the run earlier? Why does this actually happen?

Thanks in advance,
Cheers, Daniel
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I would check the salt content of the elution buffer. You could try dialysing against a lower salt buffer to bring the salt down a bit.

yeah .. I thought about this issue already before . the thing is, with a DNA affinity chromatography, you have to increase the salt concentration in the elution buffer . dialysis would be an option .. but would prefer an easier way .

as for the attached image (from left to right) from 100 mM NaCl to 1 M NaCl.
As you can see, the bands I'm interested in, are from 100 mM to 500 mM NaCl (with the strongest band around 350, 500 mM) .. actually I was assuming that this salt concentrations would not interfere with the run .

. would you have any suggestions on how to concentrate the elution? In a sense, how would you do a dialysis when you only have about 40-50 ul of elution .

I suggest you might need to look into your buffer's voltage and temperature especially after a log run. Also, perhaps you might need to change by using new APS and B-mercaptoethanol. I used to have blurry bands when such things were not right.

Adrian K on Sun Jul 8 05:24:08 2012 said:

I suggest you might need to look into your buffer's voltage and temperature especially after a log run. Also, perhaps you might need to change by using new APS and B-mercaptoethanol. I used to have blurry bands when such things were not right.

I like to use 3kDa amicon filters when I run serum samples, those would probably work well with your application. I have used those filters on samples anywhere from 50 uL to 400 uL.

Also, I can't really tell from the picture you posted, but are you using a stacking gel? That can definitely help to increase resolution.

proteaMatt on Mon Jul 9 12:41:36 2012 said:

I like to use 3kDa amicon filters when I run serum samples, those would probably work well with your application. I have used those filters on samples anywhere from 50 uL to 400 uL.

Also, I can't really tell from the picture you posted, but are you using a stacking gel? That can definitely help to increase resolution.

Hi, Thanks for your reply! The idea with the filter sounds great .. hope they are not too expensive. . And yeah, I do have a stacking gel, but cut it away before I started with the staining .

you can desalt by drop dialysis

resolution of low mw proteins and peptides is often poor, even with a high acrylamide percentage, with tris-glycine-sds gels. you can get exceptional resolution of your proteins of interest with a 10% tris-tricine-sds gel (shagger and von jagow).

mdfenko on Mon Jul 9 16:30:18 2012 said:

you can desalt by drop dialysis

resolution of low mw proteins and peptides is often poor, even with a high acrylamide percentage, with tris-glycine-sds gels. you can get exceptional resolution of your proteins of interest with a 10% tris-tricine-sds gel (shagger and von jagow).

Hi, Thanks for your answer .. I might give this a shot tomorrow .. is there any chance you have a protocol available for preparing the gel as well as the buffer? Can I run those gels with any system? Since I found this statement:

Tris-Tricine-SDS (TTS) running buffer is the cathode (upper reservoir) buffer

I'm a little bit confused .. what does it mean with the upper reservoir? Do I need a special system for that .. do I also need an anode buffer?

EDIT: So I guess the upper reservoir is the part of the assembly between the glass plates and the lower reservoir is whole tank?

Microscopes magnify and show more detail

When we talk about how microscopes work, we often say that they make things look bigger – that is, they magnify them. We describe what we see down the microscope in the same way, for example, we might say that the dead fly we’re looking at has been magnified 200 times. This helps us to make sense of what we’re seeing. It also helps others who are looking at our photographs or drawings to understand what they’re looking at. This is why all micrographs published in scientific journals must indicate the extent of magnification.

However, making things bigger is only part of the story. If microscopes did nothing but make what we can already see bigger, they wouldn’t be much use! Instead, microscopes increase the amount of detail that we can see. Another word for the level of detail we can see is ‘resolution

To understand the difference between magnifying something and increasing the detail that’s visible, have a look at this digital photo of harakeke.

PCR product: clone, or directly sequence?

7-8 clones to ensure capturing that information with cloned product with direct sequencing, such heterozygosity will usually be visible as two overlapping peaks that are roughly ½ as high as surrounding peaks. Nevertheless, sometimes the data cannot be obtained without cloning. See "Other information 'gold mines' on this website? for further information.

a) Single-product: Try direct sequencing of purified PCR product (by commercial columns, standard ethanol precipitation, or ExoSap) when the sample runs as a clean, single band in an agarose gel (typically 0.8 - 3.0%, depending on fragment size).

b) Multiple bands: Try direct sequencing of gel-purified PCR product when the sample runs as a clean band amongst other bands in an agarose gel.

c) 'Single-base' regions: If you know that your sequence contains a pure poly-'singlebase' region (>8-10 bases), either clone the product or try direct sequencing in both directions. (Due to strand-slippage during the original PCR, sequence data becomes unreadable after the pure-base region.)

d) Go with Plasmids: With purified plasmid DNA, stretches of pure poly-“single-base” regions(e.g., if you can get a clean read in both the forward and reverse directions because you can create a contig from both reads to recreate the full PCR product.

    Primer-Dimers: Typically, primer-dimers generate

70-bp into the base-called read and may obscure the real peaks. In cases where longer reads are involved, samples with extremely high levels of UDTs (post-cleanup) will have additional UDT peaks later in the sequence (

First, although the majority of same-sized fragments basically migrate together as a band in a gel, fragments of all sizes in the sample are actually present throughout the gel lane. Thus, even under the best of circumstances, gel-purified products will not be entirely pure (even though the impurities won’t actually matter in those ‘best’ cases). Here, though, the lack of good band separation makes the situation much worse and there are likely to be high levels of non-target templates even in a gel-purified sample. leading to poor quality reads.

PCR Primer Titrations: A very simple approach is run a set of test reactions using serially-diluted primers. This primer-titration approach is likely to eliminate primer-dimers (by minimizing the probability of primers coming into contact with each other) unless the primers are simply a very poor design that promotes dimerization. It will also minimize or eliminate non-target priming, unless the primers simply match the non-targets as well as they do the targets. Thus, as primer concentrations fall, although the target band will become ever weaker, it will do so less rapidly than will occur for primer-dimers or non-specific products.

Ideally, at some point, the target band (using

3-5 ul in the gel well) will be distinctly visible (but not very bright) and there will not be any visible primer-dimers or other non-specific products. In that case, purify those PCR products and then use the equivalent of 3-5 ul of the original PCR products to perform DNA sequencing.

  • Residual primers: Even cleaned PCR products may contain residual primers from the original reaction to help offset residual primers, use 1 ul of sequencing primer at 10-20 uM (vs. 2-5 uM).
  • BigDye: Normal levels of BigDye (e.g., 0.5 ul in a 10 ul reaction) can cause two problems with very short templates.
    >> First, very short templates don’t consume as much of the dye terminators as do long products. Thus, when cleaning the completed reactions, it may be difficult to remove enough of the UDTs simply because so many remain.
    >> Second, if the target is very short, the standard amount of BigDye can create excessive signal intensity because the polymerase can quickly move to additional templates. Excessive signal can cause problems with basecalling.
    >> Frankly, even as little as 0.1 ul of BigDye can generate acceptable signal intensities for products shorter than

Can I directly sequence genomic DNA?

4-kb vector template, the minimum input that will reliably generate an acceptable, albeit low, signal is

20-25 ng. and stronger signal requires at least 50-100 ng of DNA. Thus, if the organism's genome were merely 25X as large (i.e., 100 kb), you would need at least 500 ng of DNA to have sufficient copies for sequencing. However, even the genomes of Mycoplasma exceed 1000 kb, which would require on the order of 5 ug of DNA input. It is highly unlikely that the sequencing reaction would function with 5 ug of DNA present. let alone with the milligrams of DNA input that would be needed for many other organisms.

How to achieve better cloning results?

Both 'sample-to-sample' and 'experiment-to-experiment' consistency are improved if you:

a) Minimize variation: keep bacterial growth periods and processed volumes consistent.
b) Minimize growth time: yields better quality DNA and better sequencing results.

Overnight growth at 37 o C (i.e.,

14 hours ) can produce cultures with so many cells that they 'overwhelm' the capacity of commercial purification columns. Further, it results in cells that are in late log phase/early stationary phase in which not all of the genomic DNA is intact and conjugated to cell wall.

By contrast, overnight growth at 30 o C or limited growth at 37 o C (i.e., 5 - 8 hours) tends to produce better quality DNA from minipreps than does standard overnight growth at 37 o C. Finally, sequencing results are often better if purified by 'midi' preps rather than by miniprep-size columns.

How to avoid multiple signals?

Most commonly, multiple signals in your sequence data will arise from the existence of multiple templates in the sequencing reaction (although occasionally the sequencing primer will find more than one suitable location on the template). See also What causes a low-level signal after the PCR-stop? for another possibility.

a) Cloned DNA: Either there are multiple copies of a vector (containing different inserts) within the same cell, or you failed to pick a clean, single colony. When the inserts are of different sizes, both problems can be be avoided by testing cloned DNA with standard PCR.

Your chances of a clean colony pick are enhanced by picking colonies when they are just barely big enough to be visible, such that they are still well separated. Further, examining the colonies under a low-power scope will reveal cases in which one apparent colony was actually generated by two adjacent cells (colony shape will be 'dumb-bell' like, rather than circular). Picking colonies early also ensures that there has not been sufficient time for the satellite colonies (i.e., those lacking the antibiotic-resistance gene from the insert) to grow next to the 'real' colonies. Typically, the antibiotic simply prevents growth by the bacteria thus, once antibiotic-resistant colonies begin to exude (on plates or in a broth) compounds that neutralize the antibiotic, the bacteria from satellite colonies will begin to grow &mdash as such, their inserts will also be harvested when the DNA is extracted. If sequencing is done with vector primers, there will be a double-signal if done with insert primers, the signal might be weaker than expected.

b) PCR products: Typically, the existence of multiple PCR products in a reaction are apparent from a simple agarose gel experiment in that case, you need to at least gel-purify the DNA. However, sometimes even a single, clean band is actually composed of multiple PCR products in that case (which you will discover through sequencing), you need to clone the DNA first.

c) Primer issue: If your primer has degraded on the 5'-end or was manufactured with "n-1" (or more) nts (which would omit nts on the 5'-end of the primer due to the way primers are synthesized), it will generate templates which will be 'frame-shifted' on the DNA sequencer.

Is it essential to clean PCR products for sequencing?

Aside from removing potential contaminants that might shut down a sequencing reaction, the purpose of cleaning the PCR product is to remove the original primers and other PCR reagents. Ideally, cleaned templates should be resuspended in a low TE buffer (e.g., TVLE, 10 mM Tris, 0.05 mM EDTA pH 8) alternatively, they can be resuspended in nuclease-free water &mdash however, the issues noted in Choice of primer resuspension buffer? apply to templates as well.

Nevertheless, while purification of the PCR products is highly recommended, it might not be essential. For instance, if the PCR product is sufficiently diluted, sequencing can be successful without first cleaning the PCR product. It is particularly important that the original PCR primers are diluted to the point that they will not generate noticeable levels of sequenced product – otherwise, the original primers will introduce noise into the signal. Determining the required level of dilution is an empirical process. In addition to diluting the PCR product, you can use kits (e.g., ExoSAP-IT&trade) to eliminate the original PCR reagents. Both approaches have been used successfully with samples submitted to the Genomics Facility however, they should be used with appropriate caution.

How to clean DNA templates for sequencing?

a) PCR products: Assuming a robust reaction, a standard ethanol precipitation (inexpensive method) will often do an excellent job of eliminating residual primers and primer-dimers however, do not use sodium acetate. as it is likely to co-precipitate the un-consumed primers at high enough concentrations to generate multiple signals in your sequencing reactions (due to the presence of both forward and reverse primers). A better option is to use 70-150 mM EDTA (available from the Genomics Facility) as described in Templates_EtOH-EDTA_Preciptation.docx (for 96-well plates) or Sequencing-Templates EtOH-EDTA Preciptation.docx (for 1.5-ml tubes).

However, please note that some primers are particularly resistant to removal by the standard EtOH-EDTA protocol for such primers, the Genomics Facility has developed a 'proprietary' modified EtOH-EDTA protocol that effectively removes primers and primer-dimers to levels that are of no appreciable consequence to the sequencing reaction. To avail yourself of this 'proprietary' method, please make the appropriate online submission with the Genomics Facility.

b) Plasmids: Commercial columns give the best template for sequencing. If you use a homemade technique, please consult the “Plasmid Prep” document on the website.

c) DNA Clean & Concentrator-5 Kit (distributed by Genesee Scientific [Zymo Research product]): For those who prefer column technologies, this is an excellent product that both does a good job of cleaning DNA samples and allows for elution in very small volumes (>6µl). As usual, we recommend eluting in the TVLE (see Choice of primer resuspension buffer?) to avoid any problems with pH issues or minor DNAse contamination, while not interfering with downstream applications. Zymo states that recovery is 70-95% for DNA ranging from 50 bp to 10 kb as such, the column ought to remove the majority of primers (typically

18-25 bp oligos) &mdash however, in some cases, you might need to use the Select-a-Size Columns (D4080) instead to achieve sufficient primer removal. especially if primer-dimers are an issue. You can either purchase columns for your lab or have samples processed by the Genomics Facility.

d) Notes regarding Commercial columns: First, if using a column to remove primers and primer-dimers, please read the product specifications very carefully. some kits retain sufficient amounts of primer to create problems for sequencing. Second, many protocols finish with a 5-min spin after adding the wash ethanol however, the ethanol vapor pressure under the column prevents some ethanol from spinning out of the filter. Thus, spin out the wash ethanol (1 min), dump the ethanol and blot collection tubes on a Kimwipe, and finish with a final spin (5 min).

How much DNA to use in a sequencing reaction?

The most common sequencing error is to use too much DNA (either volume or quantity).

(1) Volume: higher volumes increase the potential for including ‘reaction-killing’ contaminants in the mix.

(2) Quantity: excessive initial template has two synergistic effects, which can result in dramatically reduced sequence read length: (a) BigDye reagents are exhausted by making very small fragments and, (b) excessive quantities of small fragments clog the capillary on the sequencer, preventing the injection of longer fragments.

Most commonly, researchers will use a standard spectrophotometer to determine the concentration of DNA samples. If they are really concientious, they will use a specialized spectrophotometer such as the Nanodrop. However, it is critical to recognize that these instruments have significant limitations for such work (see Why are Spectrophotometers "Bad", and what to do instead?), and that there are better alternatives (see below).

In any case, unless your protocols consistently generate appropriate concentrations of DNA for sequencing, you will save time and money by taking steps to properly quantitate your DNA templates prior to sequencing them. Ultimately, you will use the DNA concentration estimates to determine how manyµl of template to add to the sequencing reaction in order to use the optimimum mass of DNA.

  • It is advisable to run at least some of the plasmid samples on an agarose gel, as discussed in Why are Spectrophotometers "Bad", and what to do instead?.
  • Otherwise, it is usually sufficient to take A260/A280 readings on a random sample of purified templates, and dilute the DNA accordingly. Nevertheless, if readings are not relatively consistent, process all templates then, for future samples, see How to achieve better cloning results? to minimize the occurrence of variant yields.
  • Finally, unless your samples are highly concentrated, you will obtain more reliable results from a low-volume spectrophotometer. Nanodrops can be especially useful in this regard and will also provide some clues as to the presence of contaminating salts (in the absorbance graph, as discussed in Nanodrop tips.pdf) see also Nanodrop Nucleic Acid Guide.pdf.

i) Option A . inexpensive: Purify enough PCR product for a valid A260 reading (subtracting your false reading), and make a serial dilution reference photo down to 1 ng in a clean 2% gel. With ethidium bromide, the 1 ng lane should not show a visible band however, an extremely faint band should appear with 3-5 ng of DNA. Compare products to reference photo to estimate needed dilutions.

ii) Option B. expensive: Quantitate your cleaned PCR product with an instrument that relies on a dye that intercalates in the dsDNA. For instance, you could use the Agilent Bioanalyzer (concentration, size distribution & sample integrity) or the Qubit (concentration only). Both platforms have several different kits for different applications further, they both have their pros & cons. Unfortunately, both options are expensive if you need to accurately quantitate numerous samples.

iii) Option C. inexpensive: Run the equivalent of 1 µl & 3 µl of samples (purified or raw) in a clean 2% gel (stained with ethidium bromide) e.g., mix 5 µl of each template with 20 μl of loading dye (diluted to 20% with TVLE) and then electrophorese 5 vs. 15 µl in adjacent wells for each sample. If running raw (i.e. unpurified) PCR products, use the results to determine volumes for eluting your DNA from a column (or for resuspension, if doing an ethanol precipitation).

Criteria: If the 1-µl band is barely visible and 3-µl band is faint (but distinct), use 1-3 μl of DNA in the sequencing reaction. By contrast, use >3 µl template if only the 3 µl band is visible or use a dilution of your template if the 1 µl band is bright.

In this Weak-Strong PCR products.jpg example, all of the templates might be suitable for use with 1-3 µl in a sequencing reaction. However, ‘A’ is verging on becoming too bright ‘B’ is perfect ‘C’ is getting somewhat weak ‘D’ is extremely weak (especially for a 1-kb product, which should be

2X as bright as a 500-bp product for the equivalent number of copies) and at least 6 ul should be used and, even the

450-bp product for 'E’ would likely do much better if

6 μl of template were used. In the photo, bands sizes are referenced to the Biorad EZ Load 100 bp Molecular Ruler (#1708352).

By contrast, in this Strong PCR products.jpg example, sequencing reactions using 1-3 µl of DNA would be appropriate for all of the samples. Still, while samples such as ‘A’ are perfect for using 1-3µl, samples like ‘B’ are verging on being too bright and you might want to consider using

1 µl rather than 3 µl for such samples. In the photo, bands sizes are referenced to the Biorad EZ Load 100 bp Molecular Ruler (#1708352).

Why are Spectrophotometers "Bad", and what to do instead?

Okay. so "Bad" is a dramatic overstatement. However, with respect to DNA, standard spectrophotometers have some significant limitations:

  1. PCR products & Plasmids: It is crucial that the sample not be overloaded on the gel otherwise, the presence of multiple products (of nearly the same size) can be masked by the excessively broad and bright bands.
  2. PCR products: can be run directly in gels of the appropriate concentration (based on fragment size).
  3. Plasmids: ideally, these should be linearized first to eliminate the coiled and supercoiled forms.
    • Restriction enzyme(s) can be used to cut the vector either once (generating a single, long fragment) or twice (to release the insert from the vector) both methods can have their advantages.
    • Linearizing a plasmid can also sometimes be helpful for getting better sequencing results of course, the restriction site cannot be within the desired read or between the desired read and the primer site!

How to interpret 260/230 & 260/280 OD ratios?

Should I use a Nanodrop to quantitate DNA?

220-300 nM, the Nanodrop can provide useful information for essentially no-cost. However, one should be aware of the various factors that can badly skew readings from a Nanodrop for additional details, please see Nanodrop tips.pdf and Nanodrop Nucleic Acid Guide.pdf.

Nanodrop Guide for Nucleic Acids?

The Nanodrop Nucleic Acid Guide.pdf provides nucleic acid measurement support information relevant to Thermo Scientific NanoDrop 2000/2000c, 8000 and 1000 spectrophotometers. Please refer to the model-specific user manual for more detailed instrument and software feature-related information.

The patented NanoDrop™ sample retention system employs surface tension to hold 0.5 μL to 2 μL samples in place between two optical fibers. Using this technology, NanoDrop spectrophotometers have the capability to measure samples between 50 and 200 times more concentrated than samples measured using a standard 1 cm cuvette.

Nanodrop Guide for Protein?

The Nanodrop Protein Guide.pdf is meant to provide some basic protein measurement support information for direct A280 methods relevant to Thermo Scientific NanoDrop 2000/2000c, 8000 and 1000 spectrophotometers. Please refer to the model-specific user manual for more detailed instrument and software feature related information.

The patented NanoDrop™ sample retention system employs surface tension to hold 0.5 μL to 2 μL samples in place between two optical fibers. Using this technology, NanoDrop spectrophotometers have the capability to measure samples between 50 and 200 times more concentrated than samples measured using a standard 1 cm cuvette. The Protein A280 method is applicable to purified proteins that contain Trp, Tyr residues or Cys-Cys disulphide bonds and exhibit absorbance at 280 nm. This method does not require generation of a standard curve and is ready for protein sample quantitation at software startup. Colorimetric assays such as BCA, Pierce 660 nm, Bradford, and Lowry require standard curves and are more commonly used for uncharacterized protein solutions and cell lysates.

Image 3

The DNA in wells 50, 51, 52, 53, 54 and 55 are degraded. The concentration of the DNA is very high in the well 59 hence it can not come out of the well. The comb is not removed properly from wells 56, 59,60, 61 and 62. The DNA samples are highly contaminated with proteins as well as RNAs (59 to 62).


2-D electrophoresis begins with electrophoresis in the first dimension and then separates the molecules perpendicularly from the first to create an electropherogram in the second dimension. In electrophoresis in the first dimension, molecules are separated linearly according to their isoelectric point. In the second dimension, the molecules are then separated at 90 degrees from the first electropherogram according to molecular mass. Since it is unlikely that two molecules will be similar in two distinct properties, molecules are more effectively separated in 2-D electrophoresis than in 1-D electrophoresis.

The two dimensions that proteins are separated into using this technique can be isoelectric point, protein complex mass in the native state, or protein mass.

Separation of the proteins by isoelectric point is called isoelectric focusing (IEF). Thereby, a pH gradient is applied to a gel and an electric potential is applied across the gel, making one end more positive than the other. At all pH values other than their isoelectric point, proteins will be charged. If they are positively charged, they will be pulled towards the more negative end of the gel and if they are negatively charged they will be pulled to the more positive end of the gel. The proteins applied in the first dimension will move along the gel and will accumulate at their isoelectric point that is, the point at which the overall charge on the protein is 0 (a neutral charge).

For the analysis of the functioning of proteins in a cell, the knowledge of their cooperation is essential. Most often proteins act together in complexes to be fully functional. The analysis of this sub organelle organisation of the cell requires techniques conserving the native state of the protein complexes. In native polyacrylamide gel electrophoresis (native PAGE), proteins remain in their native state and are separated in the electric field following their mass and the mass of their complexes respectively. To obtain a separation by size and not by net charge, as in IEF, an additional charge is transferred to the proteins by the use of Coomassie Brilliant Blue or lithium dodecyl sulfate. After completion of the first dimension the complexes are destroyed by applying the denaturing SDS-PAGE in the second dimension, where the proteins of which the complexes are composed of are separated by their mass.

Before separating the proteins by mass, they are treated with sodium dodecyl sulfate (SDS) along with other reagents (SDS-PAGE in 1-D). This denatures the proteins (that is, it unfolds them into long, straight molecules) and binds a number of SDS molecules roughly proportional to the protein's length. Because a protein's length (when unfolded) is roughly proportional to its mass, this is equivalent to saying that it attaches a number of SDS molecules roughly proportional to the protein's mass. Since the SDS molecules are negatively charged, the result of this is that all of the proteins will have approximately the same mass-to-charge ratio as each other. In addition, proteins will not migrate when they have no charge (a result of the isoelectric focusing step) therefore the coating of the protein in SDS (negatively charged) allows migration of the proteins in the second dimension (SDS-PAGE, it is not compatible for use in the first dimension as it is charged and a nonionic or zwitterionic detergent needs to be used). In the second dimension, an electric potential is again applied, but at a 90 degree angle from the first field. The proteins will be attracted to the more positive side of the gel (because SDS is negatively charged) proportionally to their mass-to-charge ratio. As previously explained, this ratio will be nearly the same for all proteins. The proteins' progress will be slowed by frictional forces. The gel therefore acts like a molecular sieve when the current is applied, separating the proteins on the basis of their molecular weight with larger proteins being retained higher in the gel and smaller proteins being able to pass through the sieve and reach lower regions of the gel.

The result of this is a gel with proteins spread out on its surface. These proteins can then be detected by a variety of means, but the most commonly used stains are silver and Coomassie Brilliant Blue staining. In the former case, a silver colloid is applied to the gel. The silver binds to cysteine groups within the protein. The silver is darkened by exposure to ultra-violet light. The amount of silver can be related to the darkness, and therefore the amount of protein at a given location on the gel. This measurement can only give approximate amounts, but is adequate for most purposes. Silver staining is 100x more sensitive than Coomassie Brilliant Blue with a 40-fold range of linearity. [3]

Molecules other than proteins can be separated by 2D electrophoresis. In supercoiling assays, coiled DNA is separated in the first dimension and denatured by a DNA intercalator (such as ethidium bromide or the less carcinogenic chloroquine) in the second. This is comparable to the combination of native PAGE /SDS-PAGE in protein separation.


A common technique is to use an Immobilized pH gradient (IPG) in the first dimension. This technique is referred to as IPG-DALT. The sample is first separated onto IPG gel (which is commercially available) then the gel is cut into slices for each sample which is then equilibrated in SDS-mercaptoethanol and applied to an SDS-PAGE gel for resolution in the second dimension. Typically IPG-DALT is not used for quantification of proteins due to the loss of low molecular weight components during the transfer to the SDS-PAGE gel. [4]

  • Magnification is the ability to make small objects seem larger, such as making a microscopic organism visible.
  • Resolution is the ability to distinguish two objects from each other.
  • Light microscopy has limits to both its resolution and its magnification.
  • airy disks: In optics, the Airy disk (or Airy disc) and Airy pattern are descriptions of the best-focused spot of light that a perfect lens with a circular aperture can make, limited by the diffraction of light.
  • diffraction: the breaking up of an electromagnetic wave as it passes a geometric structure (e.g., a slit), followed by reconstruction of the wave by interference

Magnification is the process of enlarging something only in appearance, not in physical size. This enlargement is quantified by a calculated number also called &ldquomagnification. &rdquo The term magnification is often confused with the term &ldquoresolution,&rdquo which describes the ability of an imaging system to show detail in the object that is being imaged. While high magnification without high resolution may make very small microbes visible, it will not allow the observer to distinguishbetween microbes or sub-cellular parts of a microbe. In reality, therefore, microbiologists depend more on resolution, as they want to be able to determine differences between microbes or parts of microbes. However, to be able to distinguish between two objects under a microscope, a viewer must first magnify to a point at which resolution becomes relevant.

Resolution depends on the distance between two distinguishable radiating points. A microscopic imaging system may have many individual components, including a lens and recording and display components. Each of these contributes to the optical resolution of the system, as will the environment in which the imaging is performed. Real optical systems are complex, and practical difficulties often increase the distance between distinguishable point sources.

At very high magnifications with transmitted light, point objects are seen as fuzzy discs surrounded by diffraction rings. These are called Airy disks. The resolving power of a microscope is taken as the ability to distinguish between two closely spaced Airy disks (or, in other words, the ability of the microscope to distinctly reveal adjacent structural detail). It is this effect of diffraction that limits a microscope&rsquos ability to resolve fine details. The extent and magnitude of the diffraction patterns are affected by the wavelength of light (&lambda), the refractive materials used to manufacture the objective lens, and the numerical aperture (NA) of the objective lens. There is therefore a finite limit beyond which it is impossible to resolve separate points in the objective field. This is known as the diffraction limit.

Estimate apparent molecular mass for unknowns

Relative mobility should be calculated for each band of interest and the standard curve used to estimate apparent molecular mass. Because the relationship between mass and Rf is logarithmic, one should interpolate the standard curve data rather than use a trendline that may miss some of the data points. It is especially important to avoid extrapolating the standard curve, since even the logarithmic relationship begins to break down in the top 20% or so of a gel. One can report the mass of an unknown to exceed that of the highest mass standard, but cannot estimate a molecular mass for an unknown with Rf smaller than that of any of the standards. For example, if the Rf for the myosin standard (205 kDa) was 0.18 and the Rf for unknown 1 was 0.15, one reports that unknown 1 had apparent molecular mass > 205 kDa.

Consider resolution in the appropriate region of a gel and thickness of the band of interest when determining significant figures with which to report a mass estimate. For example, suppose the distance between 97,000 and 116,000 kDa standards is 0.5 cm and a band between them is 1 mm thick. You have resolution to the nearest 4,000 Daltons. An estimate of, say, 110 kDa should probably be written as 110 ± 4 kDa.

STING condensates on ER limit IFN response

STING is a key player in the IFN response to cytosolic DNA, and its multimerization is commonly associated with activation of the pathway. A new study now shows that STING forms ‘puzzle’-like condensates to limit the IFN response and constrain antiviral immune activation.

Formation of higher-order assemblies including liquid and gel condensates has recently emerged as a major mechanism of signal transduction in innate immunity 1 . Indeed, activation of pathways through multimerization of sensor and adaptor proteins is known for Toll-like receptor signalling 2 , inflammasome activation 3 , retinoic acid-inducible gene I (RIG-I) 4 and cyclic GMP–AMP synthase (cGAS) 5 nucleic acid detection, as well as many others. However, the question as to whether and how a multimerization principle may also be involved to inhibit immune pathways has not been fully addressed. In this issue of Nature Cell Biology, Yu et al. report that excess 2′,3′-cyclic GMP–AMP (2′,3′-cGAMP) induces STING to phase separate on the endoplasmic reticulum (ER) in a puzzle-like structure, thereby preventing STING overactivation 6 .

Restriction Digestion/Gel Electrophoresis Assignment 1

What we're going to do now is give you some experimental results and let you interpret them, so let's jump right in. You have performed Restriction Digestion and Agarose Gel Electrophoresis on a plasmid you purified, using 3 different Restriction Enzymes, and the gel is shown below. Unfortunately, you forgot to label your tubes or keep good records, and the only things you can remember about the experiment are that your standards are in Lane 5 and your uncut control is in Lane 1, and that you loaded roughly the same amount of total DNA in your sample lanes (1-4). Hey, at least you remembered that much!

2. How many times did the enzyme used in Lane 2 digest the plasmid? Does the data seem reasonable? What is the likely number of base pairs this enzyme recognizes?

3. When DNA appears as a messy, continuous band as it does at the bottom of Lane 3, rather than independent, discreet bands, the effect is known as smearing. What are some likely explanations for the smearing detected in Lane 3? You should be able to come up with at least two.

4. How many times did the enzyme used in Lane 4 digest the plasmid? Does the data seem reasonable?

Answers to Questions

2. How many times did the enzyme used in Lane 2 digest the plasmid? Does the data seem reasonable? What is the likely number of base pairs this enzyme recognizes?    The enzyme digests the plasmid in two places. It is important to think about the state of the DNA before digestion. The DNA used in this experiment was a plasmid, and plasmids are circular. If you cut a circle once, you get one linear fragment. You must cut it a second time to get 2 linear fragments like in Lane 2. The data does seem reasonable because if you add up the approximate sizes of the resulting fragments (roughly 4 kb and 2.5 kb), you get the original size of 6.5 kb. Lastly, it is likely that the enzyme used recognizes a sequence of 6 bases. 6-cutters, if you'll recall, cut an average of once every 4,096 bases. This is just an average, however, so in this case where we have a piece of DNA 6,500 bp long, cutting twice is very reasonable.

3. What are some likely explanations for the smearing detected in Lane 3?    In general terms, smearing is when you have many bands together close enough in size that you cannot distinguish between adjacent bands (i.e., no resolution). With beginning molecular biologists, the most likely reason for the smearing is contamination by some stray nuclease that degraded the DNA into dozens, hundreds, or even thousands of little pieces. Another beginning mistake is to use the wrong buffer, wrong temperature, or wrong conditions. Any or all of these could make the enzyme behave badly, including cutting away at your DNA at multiple, random sites. However, as you do more and more experiments like this, personal error becomes less of a concern and you need to start thinking in terms of the science. If this experiment was performed without significant error, the likely explanation is that a 4-base cutter was used. Cutting an average of once every 256 bases in a 6.5 kb plasmid yields roughly 25 fragments, all smaller than the original. It is unlikely that one could see 25 individual fragments of such a small size, and the smearing pattern is probably what would be detected.

4. How many times did the enzyme used in Lane 4 digest the plasmid? Does the data seem reasonable?    If you said twice, you are correct, but let's see if you were correct for the right reasons. In question 2, it was pointed out that to get two fragments from a circular piece of DNA, you need two cuts. So far so good. It was also mentioned that the total size of the resulting DNA fragments must add up to the original size. Uh oh--they don't, do they? Looking at the gel you see one band approximately 6.5 kb and one large band at roughly 3 kb. Does 6.5 + 3 = 6.5? Not in this class.

Science doesn't lie, it's just sometimes hard to interpret. So break it down. Is there anything significant about 6.5 kb? Yes, it's the size of the original plasmid. This, plus the fact that there is a band in the uncut control (Lane 1) which migrates to the same position, should suggest to you that not all of your DNA was digested (a common occurrence). This leaves the band around 3 kb. Could that band be 3.2 or 3.3 kb instead of 3.0? Easily. Unless we plot a standard curve, we're just approximating anyway. Is there anything significant about 3.3 kb? Yes, it's about half of our original sample. If the enzyme cut the plasmid into two roughly equal sized pieces, those pieces would run about the same, and would likely be indistinguishable on a gel. This is further supported by the information about this experiment which states that roughly equal amounts of DNA were loaded into Lanes 1-4. Notice how much darker the 3 kb band in Lane 4 is than the bands in Lane 2. There is twice as much DNA in that band than there is in either of the bands in Lane 2, and the data supports this conclusion.