About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

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Real-time PCR is a technique that is now commonly employed in almost all molecular biology laboratories to quickly answer very specific questions. Northern and Southern blotting are now a thing of the past. No longer do we wait days to know whether a gene is expressed. We can have the answer in 45 minutes!

But with the widespread use of such a wonderful and sensitive technology comes differences in how results are reported in the literature. There are also differences between reviewers reading these papers and their understanding of the essential information required to judge the accuracy of the reported data.

To overcome this increasing problem of lack of consistency in publications, a panel of real-time PCR experts published a set of guidelines containing what they consider the minimal information required when reporting qPCR results. That paper called The MIQE Guidelines: Minimum Information for Publication of Quantitative Real-Time PCR Experiments, was published February 2009 in the Journal of Clinical Chemistry.

This is not only a great resource for authors, but it also essentially a troubleshooting guide as well. If you don’t have an answer to each of the item on the checklist, then maybe you are missing an essential piece of information in your experiment.

[PS: One of the authors of the paper, Greg Shipley is presenting live about the MIQE guidelines here on Bitesize Bio on Tuesday, click here to book your place! We first posted this article last year - so if you were wondering whether you were having a de ja vu moment, you can relax - but have re-posted it to highlight it ahead of Greg's talk]

Standardization of nomenclature:

To help standardize the lingo used in real-time PCR, the authors start off by clarifying common terminology.

• For example, the term RT-PCR is reserved for reverse transcription-PCR while the term RT-qPCR should be used for quantitative experiments. The RT in RT-PCR does not stand for “real-time” in this case. The q designates that the experiment is a real-time PCR experiment. So it is qPCR or RT-qPCR when the experiment invoves real-time methodology for DNA or RNA, respectively.
• In regards to the use of genes for normalizing qPCR experiments, the authors recommend the use of the term reference genes and not housekeeping genes.
• TaqMan probes should be called hydrolysis probes which is in contrast to hybridization probes of which FRET and Molecular Beacons are a class. TaqMan is a trade name for hydrolysis probes and is not a scientific term.
• Several terms are used to describe the point at which fluorescence becomes detectable and measurable. These include the threshold cycles (Ct), crossing point (Cp), and take-off point (TOP). The authors explain that these are marketing terms. They propose the use of the word quantification cycle (Cq) as the universal word for the cycle where fluorescence is measured.

The Checklist:

The checklist for the essential and desirable information for a publication is quite long and may be intimidating to researchers who may have not realized how much information is really critical to judging the scientific validity of their work.

But in truth, adding this information to your paper will only help make your conclusions stronger and make it more difficult for reviewers to reject your results. Topics that need to be addressed include the experimental design, the method of RNA or DNA extraction and quantification, qPCR target information, kits used, efficiency and slope information, and data analysis information including NTC information, normalization method, and repeatability.

It is truly a great checklist for making sure your paper gives a complete story.

Who should read this paper:

If you are using qPCR or RT-qPCR in your project, then read this paper. And if you are reviewing papers using qPCR, you should read this paper.

Too many critical details are being left out of papers and conclusions are being made that may not necessarily be correct. What if someone else’s paper scoops your work because it was not reviewed as critically as yours? And then the data is not even correct?

What if you are trying to repeat someone else’s work and you waste valuable hours and expensive reagents and can’t replicate the results because information was missing?

Ideally, all papers need to be reviewed for the same criteria and with the same scrutiny before being accepted for publication. This type of standardization provided by the MIQE guidelines are meant to ensure fairness during the review process and make sure that only high quality results make it to print.

What do you think about the MIQE guidelines? Do you like them? Will you follow them? Tell us your thoughts.

About the author

Jode Plank

Jode is a Postdoctoral Fellow studying DNA repair at the University of California at Davis. He received his PhD in Biochemistry from Duke University.

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Western Blotting is a long established method for which the protocol varies little from lab to lab. However, there are some new products that are available and some tweaks that can be made to the protocols that may improve your results and reduce the time it takes you to execute this popular technique.

Save time by co-incubating the primary and secondary antibodies
There are a number of kits coming out which, I believe, are based on this simple concept. Normally, we incubate the blocked membrane with the primary antibody, wash away the unbound antibody, then incubate the membrane with the secondary antibody which binds to the primary antibody, co-localizing the detection method (fluorophores, horseradish peroxidase (HRP), or alkaline phosphatase (AP)) with the protein of interest on the membrane. However, since the primary antibody binds its epitope with its variable domain and the secondary antibody with its constant domain, there is no reason that mixing the two antibodies together will interfere with the binding of the primary antibody to its epitope. Both the primary and secondary antibodies are diluted into your favorite solution (TBST + 5% milk, for example) and incubated with the blocked membrane at the same time. For the average user, this will shave over an hour off a western blot by eliminating the second antibody incubation along with the washes between the two antibody incubations. I have used this method for around five years, and have yet to have any problems with it.

Optimize the incubation times
If you find yourself doing the same western blot repeatedly (ie – same primary and secondary antibodies with similar samples), then you might be able to shave even more time off the standard protocol. A friend of mine experimented with the blocking time and incubation time with the primary/secondary antibody solution. She found that the blot was blocked after a 10 minute incubation with fresh, room temperature 5% milk, and that the western signal reached maximum after a 15 minute incubation with her antibodies (without increasing the concentration of the antibodies). She could develop a blot and be ready for exposure within 45 minutes of removing the membrane from transfer. These parameters are going to vary for each set of antibodies and sample types that one uses, so you would need to decide if it is worth you time to systematically optimize these incubation times.

Use Protein A-HRP to get clean western blots of immunoprecipitated samples
If your immunoprecipitation (IP) used an antibody from the same organism as the primary antibody that you are going to use for your western blot, then your secondary antibody will not only recognize the primary bound to the epitope, but also the antibody from the IP that was denatured and separated out on the gel, creating giant background bands around 55 and 25 kD on the blot. However, in 2005 Ashish Lal published a method to circumvent this problem: instead of using a secondary antibody to detect the location of the primary, use HRP conjugated Protein A instead. (Protein G worked well, also, but did show higher affinity to the denatured antibody.)  Unlike a secondary antibody, Protein A only recognizes correctly folded antibodies. This means that it binds tightly to the primary antibody used in the western, but largely ignores the denatured antibody stuck to the membrane. This system has (seemingly) been incorporated into several kits, but Protein A-HRP alone is also available from a number of companies.

Protein A-HRP can be used for normal western blots as well, and would replace secondary antibodies against Mouse, Rabbit, and Guinea Pig, which might be an attractive option for labs on a budget or with a fair number of undergraduates, who seem inexplicably drawn to the incorrect secondary antibody.

Multiplex Westerns
If you need to gather information about two different proteins in the same sample, you can probe the blocked membrane with an antibody against the first protein, then strip the blot and probe with an antibody against the second protein; run two identical gels and probe each with a different antibody; or if your two proteins are of very different sizes, cut one membrane in half across the lanes and probe the top and bottom sections with different antibodies. Alternatively, you can perform a multiplex western.

A multiplex western is one in which more than one primary antibody is used on the same membrane at the same time. This obviously isn’t a complicated change to the protocol – simply add the extra primary antibody (and secondary, if needed) to the appropriate solution. However, I am mentioning it here simply because I have seen many occasions when people could have used this approach to save themselves time or sample, but didn’t. If you are using chemiluminesence detection methods, then of course you need to have characterized the antibodies sufficiently to know that you are interpreting the western blots correctly. This can save you time in the day-to-day experiments, but for critical experiments (ie – paper figures) I will still blot with the individual antibodies separately, just to make sure my interpretation is correct.

The multiplex western can be taken a step further if the two primary antibodies were raised in different animals. In the high-tech version of this technique, the two secondary antibodies are labeled with two different fluorophores, and, after washing, the bands are differentially detected using a Storm imaging system or equivalent. In the low-tech version, one of the secondary antibodies is conjugated to HRP, while the other secondary is conjugated to AP. Each of these enzymes can be detected by chromogenic substrates, allowing you to develop a visible two color blot. (The caveat to the later system is that the chromogenic substrates are known to be less sensitive and reduced linearity of response compared to fluorescent or chemiluminescent methods.)

The approaches that I’ve described here can help you streamline your westerns and save you some time, but I think it is important to stress that any changes to a protocol should only be implemented with the proper testing and controls. If you are happy with your standard protocol, then you shouldn’t change it. If you are like me and have found your work day extended into a work evening on a regular basis by the standard western protocol, then the suggestions above may help you keep your work hours under control without sacrificing your productivity.

Now to hear from you: what are your best western blot tips and tricks?

About the author

Nick Oswald

Nick is a molecular biologist-turned-publisher. After a PhD in Developmental Biology and an eclectic seven years in biotech he is now Editorial Manager of Neuroendocrinology and the founder and Editor-In-Chief of Bitesize Bio. You are welcome to connect with Nick on LinkedIn

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Whatever the technique, it is essential to have the correct experimental setup to let you verify that you are measuring what you think you are measuring. And PCR is no different – you can’t conclude anything concrete from your results unless you have the correct controls in place.

Ian Kavanagh of Thermo Scientific has a very illuminating the topic of controls and standard curves in PCR lined up for today’s live seminar here on Bitesize Bio (click here to reserve your place) but in this article, I’ll give a quick run-through of the basic controls you should be including in your PCR reactions.

Negative Controls

No template controls (NTC)

Negative controls that contain everything but the template DNA are essential for detecting contamination or non-specific amplification in your reaction. Correctly used, NTCs can not only alert you to the presence of a contaminating amplicon, but can also help you identify the source of the contamination. For more details of that, see Ian’s seminar.

Please never be tempted to miss out the NTC. They are really easy to set up, but you are just asking for a beautifully amplified non-specific product to appear in the right place on your gel if you miss it out. But don’t just take my word for it – the MIQE guidelines (we have a seminar on those too!) advise that NTC’s should be considered an essential part of every PCR reaction.

No Enzyme Controls (NEC)

“No enzyme” controls are required when using PCR to quantify RNA, which involves the PCR amplification of a cDNA synthesised from the target RNA sequence using reverse transcriptase. The reverse transcriptase is omitted from the NEC (or the “no-RT” control) and this can be used to determine that any amplification that occurs in the sample is derived from the synthesised cDNA and not genomic DNA or other amplicon contamination.

If the NEC shows that contamination is present, then cleaning up the sample or using intron-targeted primers that will only bind to the RNA can solve the problem.

Positive Controls

Positive controls are needed for the verification of negative amplification results and the positive control reaction should contain the same components as the sample but include a template that is certain to amplify if the reaction goes as planned.

This could be an external positive control, which is a separate sample containing the control template. Such external control reactions can help detect when a reaction fails due to cycler or reaction component problems or when an inhibitor is suppressing the reaction.

Alternatively, you could use an internal positive control (IPC). To run a reaction with an IPC, the template and primers for the control target are included in the reaction along with those for the target of interest. The control target should of course be easily distinguished (by electrophoretic migration or Tm) from the target of interest. As well as having the advantage that of not requiring a separate reaction, IPCs are useful because they can pinpoint problems that are intrinsic to the sample reaction.

There are so many things that can make a PCR reaction go wrong. So habitually including a positive control is certain to save you a fair amount of head-scratching during the course of your career by pinpointing those times when one of the reaction components is causing you problems.

More detailed information on controls in PCR can be obtained in Ian Kavanagh’s seminar, which airs live at 9am Pacific / 12pm Eastern / 5pm BST (UK) / 6pm CET TODAY. There are still some places left and you can secure yours by clicking here.

About the author

Jode Plank

Jode is a Postdoctoral Fellow studying DNA repair at the University of California at Davis. He received his PhD in Biochemistry from Duke University.

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While almost all of you are probably familiar with the power of eBay to bring you everything from concert tickets to electronics to your very own Batmobile, you may not have realized that the world’s largest garage sale also has quite a collection of laboratory equipment. I’ve been turning to this source for equipment for several years now, and in this post I’ll fill you in on some of the ins and outs of buying lab equipment off eBay.

Where do I look?
Of course you can search for whatever you want, but if you want to surf around and see what’s available, click on the ‘Business & Industrial’ category, then go to ‘Healthcare, Lab & Life Science’, and from there you will probably want the ‘Lab Equipment’ category, but ‘Lab Supplies’ might also peak your interest. If you have ever really scoured eBay for a particular item, you probably already know that sometimes the sellers don’t list their item in the correct categories. So if there is something in particular you want, don’t just browse the category that it should be in, but search for it, and search for it in a couple different ways, as I’ve found eBay’s search engine to be a little finicky (Bio-Rad is different than BioRad, et cetera).

What is it good for?
The most obvious use for eBay is to purchase second hand equipment on the cheap. It may be that the piece of equipment you want would be nice to have, but not so essential that it warrants the bite it would take out of the lab budget, or it may be that the lab is running on fiscal fumes right now, but needs to replace a piece of equipment to maintain productivity (a scenario that is sadly all-to-common right now). Outside of research labs, eBay might also be a viable option for teaching labs, where it seems money is always an issue. For lab heads, eBay might also be a reasonable way to equip empty benches in the lab, as patience can be rewarded with some phenomenal buys.

Price of the newest version of this product - $650.00

I have found that certain items, like heat blocks and waterbaths, can be found in nice shape and at good prices, even for models that are only a couple of years old. Other items, such as microcentrifuges and electrophoresis power supplies, are more difficult to find in modern forms, and finding one at a good price is even rarer. Micropipettors are a real gamble – you can almost always find them at a third to a quarter of retail prices, but repairing a poorly maintained micropipette can easily chew up any money that you saved and then some.

The other area where eBay shines is the availability of old equipment, or parts for old equipment. In my current lab we have an older fluorimeter that is due to be replaced, but is expensive enough that it cannot be done mid-grantcycle. When the power supply went out, we discovered that the fluorimeter was no longer supported, and the company no longer stocked parts for it. However, we did find somebody on eBay selling just what we needed. Admittedly, the selection available on eBay at any given point in time isn’t so great that you can count on finding the parts that you need, but it does provide one more place to look when you are in a bind.

List price of a new digital VWR single-block heater - $662

Who to buy from?
There is inherent risk in buying used lab equipment. You can minimize the risk by employing some of the common eBay rules: don’t buy from people with low feedback scores, a low number of sales, or strict ‘as-is’ or ‘no return’ policies. Another aspect to keep in mind is the seller’s experience with lab equipment, which is usually apparent in the ad listing. If the ad for a water bath reads “the unit turns on and heats up,” then that likely isn’t as experienced a seller (with lab equipment) as one who states “the unit powers up and maintains temperature (37C) overnight.” The second unit is much less of a risk than the first. You can also gauge the experience of the seller by looking at their current listings, or by looking at their most recent sales to see how much lab equipment they deal with. Don’t be afraid to ask questions before you buy. If you do get a piece of equipment that is broken, you should contact the seller immediately, since they are likely to accept the unit back or refund your money to convince you to not leave negative feedback on their account. There are some sellers that specialize in lab equipment and have large eBay stores that you can browse – they are a very safe bet to buy from, but they are also very aware of the value of what they are selling and price their items accordingly.

How can I buy off eBay?
This is what you should ask your boss and your lab’s financial administrator. Often you can buy things yourself and the lab can reimburse you for the purchase, but different universities have different rules about this, so check first. Sometimes the lab or department will have a person who can buy the item directly from eBay. Before you do anything, though, talk to the lab head and make sure they are comfortable with purchasing used equipment and the risks that go along with that.

What are your experiences of buying equipment from eBay, or any other non-standard source for that matter?

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

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Isolating RNA is one of the more finicky protocols there is, and everyone who does it has their own personal tips and tricks to successfully isolate intact RNA from their samples with consistency. Although RNA can be somewhat unpredictable since it is so labile, there are a few common problems that occur which can be logically solved.

In my experience in helping people with RNA preps, these are the four main complaints when isolating RNA from tissues and cells. My solution to each problem is below. If you have your own tips, you’d like to share, please leave a comment.

1. Problem: Genomic DNA in the RNA
The RNA elutes with genomic DNA as evidenced by high molecular weight smearing or it appears clean on a gel but -RT controls amplify when PCR is performed.

Cause: No matter what method you use for RNA isolation, traces of DNA always carry through. This is true with TRIzol (phenol) preps and with silica spin filters. This can be caused by insufficient shearing of the genomic DNA during homogenization. If using phenol method, the pH of the phenol is key (it should be acidic) and your skill in pipetting only the aqueous phase will result in more or less DNA contamination.

Solution: The genomic DNA needs to be well homogenized so use a method that sufficiently breaks the DNA such as a high velocity bead beater or a polytron rotor stator. Samples will heat during homogenization but chilling samples in guanidine causes the salt to precipitate out. So you will need to balance the time of homogenization with time to cool down to room temperature.

The best way to remove the gDNA is with a DNase treatment. If using the spin filter RNA isolation method, most manufactures have an “on-column” DNase method that is easy and time-saving. However, if the sample is very rich in gDNA (ex. spleen tissue) it may still not be completely removed. A method that is highly recommended among the scientists I speak with is the Ambion Turbo DNA-Free kit. The DNase is inactivated without EDTA or heating and it removes the DNA with high efficiency.

2. Problem: Degraded RNA/ low integrity
The rRNA bands appear smeared on a gel or the 18s is more intense than the 28s band. On the Agilent Bioanalyzer, you see a bigger 18s peak.

Cause: Degradation occurred at some point during processing. This can be difficult to pinpoint. It could be during collection and storage or it could be during extraction. It could also occur post isolation.

Solution: If the problem occurred during storage, make sure that samples are frozen immediately after collection. Use liquid nitrogen or -80C storage. For animal tissue, use RNALater and then store at -20C.

If the problem occurred during the extraction procedure, try adding beta-mercaptoethanol (BME) to the lysis buffer. Use 10 ul of 14.3 M BME per 1 ml of lysis buffer. BME will kill the RNases and help stabilize the sample during extraction.

If the sample is coming out of a freezer to be extracted and is not in a preservative solution, do not allow it to thaw. Homogenize it quickly in the presence of BME. Make sure not to leave behind any chunks of tissue. It needs to be completely lysed.

RNase degradation can also occur after isolation. Make sure that the water used for elution or resuspension of pellets is RNase free. Many kits provide water for RNA work that is DEPC treated or purified in other ways. More on the myths behind DEPC treatment and RNases can be read here.

In addition, 10 ways to work RNase free might also be a helpful article for you.

3. Problem: Inhibitors in the RNA
The RNA has an abnormally low 260/230 reading (below 1.0) or 260/280 reading or does not work in reverse transcription.

Cause: A low 260/230 in an RNA prep is indicative of guanidine salt carry over into the sample or organic inhibitors (such as humic acids or polysaccharides if the sample is environmental). Guanidine salts are used in TRIzol and in silica preps. These salts inactivate RNases, but will also inhibit proteins such as RT enzymes if present in the final RNA.  A low 260/280 measurement indicates protein contamination.

Solution: For low 260/230 readings, the best approach is to try more washes of the RNA sample. If this is a TRIzol precipitate, try washing it with ethanol to desalt it. For silica preps, a few extra washes with 70-80% ethanol should clear the column of salts. For samples already purified that are not clean, try ethanol precipitation of the RNA to desalt.

For other inhibiting compounds, such as humic acid and polysaccharides, you may need to re-purify the sample on another column and wash it well before elution. Some of these compounds will not be removed from RNA (or DNA) with conventional methods because they are too similar to the nucleic acids. In this case, you may want to consider using inhibitor removal technology for environmental samples.

A low 260/280 reading from protein carry over most likely occurred because of using too much sample for the kit or method. The sample overwhelmed the chemistry and the proteins were not completely removed.  Try cleaning up the sample with another round of your method- either phenol:chloroform and precipitate or add the binding salts and ethanol and bind to another silica spin filter. The protein should be easily removed. Next time, try using less sample and make sure homogenization is complete.

4. Problem: Low yields of RNA
The yield of RNA is lower than expected- either based on your previous results, or, based on reported yields for a certain tissue or cell type.  RNA yields can vary greatly between different cultured cell types and in different tissues. For blood RNA, it can vary from person to person.

Cause: If the yield of RNA is lower than you expected or know it should be, and the RNA is intact, so degradation is not the cause, then the homogenization may not have been complete. To isolate RNA, a strong lysis is key. Tissues stored in RNALater will tend to be a little more difficult to homogenize. Low yields could be caused by mistakes in weighing of tissue or in the cell counts for cultured cells. You may have less cells than you think. With blood RNA, the buffy coat can vary based on your skill in collecting the white cell layer and each individual patient.

Solution:For cases where the RNA yield is low but the RNA is intact, focus on the homogenization method and make sure you are getting good shearing of the genomic DNA and release of RNA from all cells.  If you see any pieces of tissue or debris in your homogenate, that is lost RNA.

It can be difficult to weigh small pieces of tissue so make sure you are using a scale accurate to the weight you need or you may see variation from prep to prep as well as other problems with purity and clogging columns when too much is used.  For cells, it is important to have an accurate count if you want to have consistency.

If the RNA looks degraded as well as the yield being low, the problem could be the storage or the homogenization may have been too hard and the sample was heated excessively for too long. Try homogenizing in bursts of 30-45 seconds with 30 seconds of rest.  Make sure to cut your tissue section and get it into cold guanidine lysis buffer (or TRIzol) quickly to prevent RNase activity during set up.

With silica spin filters, make sure the elution from the column is in enough volume to release the RNA from the membrane. More water will elute more RNA so use the largest volume that allows the RNA to be as concentrated as you need. Don’t try to use less volume than recommended by the manufacturer or you probably are leaving your RNA on the membrane. It is better to elute with more and concentrate after using ethanol precipitation, if maximal recovery is key.

And finally…

Isolation of RNA follows a similar protocol regardless of the sample source.  For all samples, homogenization is the first step and the most important step. A good homogenization needs to break cells quickly to inactivate RNases in the lysis buffer, and needs to break genomic DNA down in size to make removal more efficient.

Let us know what you think are the biggest issues in RNA preps and your tips and tricks used to overcome it. And if you have a problem now and want some advice, just let us know!

About the author

Jode Plank

Jode is a Postdoctoral Fellow studying DNA repair at the University of California at Davis. He received his PhD in Biochemistry from Duke University.

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While there are many more methods to choose from for cleaning up your RNA or DNA than there used to be, sometimes Phenol/Chloroform extraction is still the best way to go. Here I’ll discuss some of the practical aspects of using this technique.

Nick introduced the topic of Phenol/Chloroform extraction in a previous article, touching on some of the ideas about how organic extraction will remove proteins from an aqueous solution. In brief, proteins consist of both hydrophobic and hydrophilic residues, and through protein folding, achieve a compromise with water to remain soluble. However, when they are given the opportunity to transition to an environment that can accommodate both polar and non-polar residues (ie – phenol or phenol/chloroform) with no compromise required (ie – folding), they happily move over to that phase. The more highly polar molecules, like carbohydrates and nucleic acids, are “happier” in the aqueous phase (with some exceptions noted below) and remain there. Now for the nitty-gritty.

Phenol versus Phenol/Chloroform versus Chloroform
One of the most frequent questions I’m asked when training someone is what the differences are between the different organic phases used in extraction. Here is the breakdown, as best as I understand them.

Phenol – What we are actually talking about here is buffer-saturated phenol, which consists of a solution that is actually about 72% phenol, 28% water. Since phenol is a weak acid, the solutions that we use have been equilibrated with buffer to bring the pH to a particular target – either acidic for RNA purification or slightly alkaline for DNA purification. In addition to a certain amount of water dissolving into the Phenol, there is a certain amount of phenol that dissolves into water – at equilibration the aqueous phase will contain about 7% phenol. This is thought to aid in the extraction, as this dissolved phenol helps denature proteins while they are still in the aqueous solution. Buffer saturated phenol has a density that is only slightly higher than that of water.

Phenol/Chloroform – This is a mixture of buffer-saturated phenol and chloroform, usually close to 1:1 for DNA purification with other ratios sometimes used for RNA purification. Isoamyl alcohol is sometimes included as an anti-foaming agent, but is generally thought to be an inert and optional addition. This solution is commonly used in lieu of buffer-saturated phenol for a couple of reasons. As I mentioned above, the density of buffer saturated phenol is only a little higher than water. So if your aqueous phase contains enough salt or any other solutes that would increase its density, then you could end up with phase inversion during extraction, where your aqueous phase is under the phenol, rather than on top of it. Chloroform is significantly denser than water, so adding it to the organic phase increases the overall density of that phase, helping to prevent phase inversion.

In addition, the chloroform (and some say isoamyl alcohol) help reduce the interphase – the fuzzy border between the two phases populated by molecules that can’t decide where they want to go. These can be partially denatured proteins, DNA (depending on the pH), and/or partially denatured DNA binding proteins that are still clinging to DNA, and it is a real pain in the butt. If you pipette off some of this material when removing the aqueous phase, then you decrease the purity of your sample. If you are too timid while pipetting, then you hurt your yield. If you have my luck, then whatever it was you wanted to keep most is sitting in it. Adding chloroform to the mix helps reduce this. (But I have an even better solution to this problem that I’ll tell you about below.)

Chloroform – This is normally used after phenol or phenol/chloroform extractions. While pure chloroform doesn’t work as well as the organic solutions mentioned above for protein extraction, it works well for extracting phenol from aqueous solutions. Remember when I said that the aqueous phase contained ~7% phenol after equilibration? Do you also remember when I said phenol likes to denature proteins? If you don’t get rid of (or at least severely reduce) the phenol in your now protein-free nucleic acid solution, it could come back to haunt you by partially or completely inhibiting enzymes that you treat the DNA or RNA with down the line. Presented with a nice chloroform home, however, the phenol will partition away from your nucleic acids. Chloroform itself is about 10X less soluble in water than phenol (~0.8%) and is less denaturing to proteins. I was also told long ago that it is less likely than phenol to pellet with DNA during ethanol precipitation once upon a time, but I cannot find a reference to back that point up.

Ether – This can also be used to extract phenol back out of the aqueous phase. However, because of the explosive potential of ether and the tendency of biology-types to have Bunsen burners and strikers in their labs, it has been largely replaced by chloroform.

Not so Pretty in Pink
A note of caution: don’t use your phenol or phenol/chloroform if the solution is turning pink. Oxidation of the phenol produces a pink/brown compound, and this compound will cause nicking of your DNA and degradation of your RNA. Most commercial phenol solutions contain an antioxidant to inhibit this oxidation, and phenol buffered at an acidic pH seems to be resistant to oxidation, but it isn’t a bad idea to move a portion of the buffer saturated phenol (from the brown bottle that it likely came in) to a clear bottle or tube to inspect it before you start your extraction.

pH matters – a lot
Occasionally somebody does a phenol extraction and doesn’t recover any of the DNA in the sample. If this happens to you, or somebody in your lab, your first question should be “Which phenol did you use?” Labs that do both DNA and RNA work will likely have both acidic and basic buffered phenol solutions, or somebody will buy a new bottle of phenol without paying attention to the pH. Extraction of DNA containing samples with acidic phenol results in the denaturation of the DNA, and once denatured, the DNA partitions to the organic phase. This is a key feature of many RNA purification protocols, which is one of the reasons acidic buffer-saturated phenol is used.

Now, sometimes a lab’s DNA phenol extractions start failing (no recovery of the DNA afterwards) and the pH of the phenol is called into question. If you find yourself in this spot, you can’t simply dip your pH meter into it, and you cannot use pH paper, since the pH indicator on the paper was characterized in aqueous solutions. The method that I’ve used is to dilute 1 ml of the buffer saturated phenol with 9 mls of 45% methanol, mix, and then measure the pH with a standard pH meter. The safest way of adjusting the pH is by replacing the aqueous phase on top of the phenol solution with a fresh aliquot of ~100 mM buffered water (usually Tris pH 7.9 for DNA work), mix the phases well, and then let the bottle settle until the phases are well separated again. Then pH it again.

Mixing your phases
Phenol/chloroform extractions are amazingly efficient – less than 1% of the average protein remains in the aqueous phase after the first extraction has come to equilibrium. The trick is to get the extraction to equilibrium, of course. The more surface area there is between the two phases, the faster this happens, and that surface area is greater the finer emulsion you have created. This can be achieved by vortexing the phases for a couple of minutes, as many protocols call for, but not all samples can be vortexed. If you are purifying very large DNA, like genomic DNA, then you may have to mix your sample much more gently, and therefore perform each extraction for much longer. So on this point, follow your protocol and be very cautious about trying to shave time off this step.

Effects of denaturation and digestion
Some protocols call for protein denaturation and possibly digestion with Protinase K before extraction. Both of these steps are attempts to reduce the amount of material that is trapped in the interphase and therefore improve the yield of DNA or RNA recovered. I have never seen any negative effect of denaturing the proteins with SDS before extraction. On the other hand, digestion of the protein could reduce the purity of the nucleic acid that you recover. While whole proteins are almost guaranteed to partition to the organic phase, once the protein is digested into small peptides, not all of those peptides will have the same chemical ‘character’ of the whole protein, and each will have its own partition number. It may not matter a lot if you have some peptides in your nucleic acid, depending on your downstream application, but it’s formally possible that these contaminants could effect your future quantitation of the sample. However, I discovered a better way of eliminating the dreaded interphase…

Phase Lock gel®
This is one of those things that seems to be in 50% of labs, yet less than 10% of the people I’ve talked to know what it is or how it works. I discovered this at a critical point in my research, and it saved my thesis. In short, Phase-lock gel is a gooey, Vasoline-like gel that has a density that is slightly greater than water. If you add your extraction on top of it in a centrifuge tube, and then centrifuge it, the Phase Lock gel collects between the aqueous and organic phases, separating the two and preventing the formation of the DNA/RNA hungry interphase.

A search of the internet didn’t turn up any pictures of this process that I thought were good enough, so I took some of my own. In this little demonstration, red dye is taking the place of our precious nucleic acid, and blue dye is substituting for protein.

A) The Phase Lock gel® pelleted into the bottom of a 1.5 ml Eppendorf tube.
B) After adding phenol/chloroform and the aqueous phase, complete with faux DNA (red) and faux protein (blue) in the aqueous phase.
C) After gentle shaking for 5 minutes.
D) After centrifugation. Note the gel now separates the organic phase from the aqueous phase.
E) After a second addition of phenol/chloroform and gentle shaking for 5 minutes.
F) After the second centrifugation. The faux DNA could now be extracted with chloroform (in the same tube, if space allows) to remove the residual phenol.

As you can see, the gel forms a stable partition between the two phases, and if you want to extract the sample a second time and there‘s still room in the tube, then you can do it, using the same tube two or more times with no compromise of the sample purity. You cannot vortex mix the two phases in a tube containing this reagent, but you can votex mix in a separate tube, then add the sample to the tube with the gel and centrifuge. They have this gel in two different flavors – one for regular samples (light) and another for high density samples, like solutions with a high salt or protein concentrations (heavy).

Using SDS to denature the proteins in my sample prior to extraction and then employing Phase Lock gel® to separate the phases has consistently given me DNA samples with 260/280 ratios of 1.8 and greater than 98% recovery. Seriously great stuff.

In preparing this article I came across this website, which has a lot of useful information. Visit it if you would like to learn more about phenol.

Now to hear from you: what are your tricks and tips for perfect protein extractions?

About the author

Suzanne Kennedy

Suzanne is Director of R&D at Mo Bio Laboratories in California, and the author of their blog, The Culture Dish. She has a PhD in Microbiology and Immunology from Virginia Commonwealth University.

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One of the very first things you need to do when getting set up for quantitative PCR (qPCR) is to determine the efficiency of the assay because knowing the assay efficiency is critical to accurate data interpretation. And you have to do this every time you design and purchase a new primer pair.

Ideally, the efficiency of the assay should be 100%. This means that during the logarithmic phase of the reaction, the PCR product of interest is doubling with each cycle.  Perfect PCR efficiency will demonstrate a change of 3.3 cycles between 10 fold dilutions of template.

Sometimes the efficiency is below 100% and sometimes you can get readings of PCR efficiency above 100%.  In a future article, we’ll discuss what causes high and low PCR efficiencies and how to fix it.

Today’s article is about the important considerations when setting up a PCR efficiency test for your new assay. The three key ingredients to focus on when getting started are the type of template, the primers, and the chemistry.

Template:

Your PCR template can be one of several types of nucleic acids. It can be a plasmid containing the gene of interest (in which case, linearize it), a DNA oligo (most people order a 60-70mer), genomic DNA (works best only if the target exists in multiple copies, such as actin, gapdh, or rRNA gene), and it can also be the PCR product of interest purified and quantified using a spec or picogreen.

Your template for the standard curve needs to be quantified and then diluted serially. I recommend at least 5 dilutions and 1:10 dilutions for the widest linear range. Many people use 1:4 dilutions which is fine, but limits the range in which you can be sure the primer pair is accurate.

Duplicates of each template dilution are required and triplicates will give you greater confidence in the data, especially if one data point drops out for an unknown reason.

I recommend diluting the templates so that the first sample (the most concentrated) comes up around cycle 16-18. If your first sample is coming up too early- like around cycle 8-10, you may run into problems with the subtraction of baseline fluorescence from the samples. This causes shifting curves and loss of sensitivity for detection of low copies of template.

Primers:

Every time you have a new pair of primers for qPCR, check the efficiency. Even if they are primers you know work well and you just received a new batch. You cannot assume that the primers from one lot to another work the same. Better to catch the problem of a poor synthesis early on than to perform a lot of work on irreplaceable samples only to realize later that the new primers were only working at 80% efficiency.

The concentration of primers to use depends on the enzyme chemistry. It’s a good idea to start with a concentration recommended by the kit supplier as a start and then modify if the efficiency is low.  Using too much primer can lead to dimer formation, especially at the low dilutions. This will have a greater impact on SYBR Green data but will also impact hydrolysis probe results. Too little primers and the efficiency will be negatively affected.

Sometimes no matter how much tweaking and optimizing (and enzyme kit changes) you do, you cannot get the assay efficiency to a level that allows for the sensitivity you need. In this case, re-design the primers. If you are trying to make accurate quantifications out past cycle 35, you need the most efficient assay possible. If all of your data is between cycles 18-30, maybe you can tolerate an assay that is only 80% efficient.  You can decide what works for you.

Chemistry:

The kit enzymes and buffer systems are going to play a big role in the results as well. So when buying a new lot of kit, re-check the assay efficiency as well. Chances are that the kits will not vary, but it is better to check and make sure.

When using SYBR Green assays, the PCR efficiency will be affected by the presence of primer dimers so make sure to always include the melt curve analysis. This is a great benefit of performing SYBR Green qPCR.   Primer dimers will affect a hydrolysis probe (such as TaqMan) as well. If reaction components are used up amplifying dimers instead of real product, efficiency will be low. For hydrolysis probe assays, you’ll want to save the reactions and run them on a gel if the efficiency is low so you can analyze what else may be amplifying in the reaction.

With Probe assays, there is also the consideration of the Probe design and the compatibility of three primers in the mix. The probe needs to have a higher Tm so that it lays down on the DNA first. With Probe assays, low efficiency can also be caused by the Probe design or Probe labeling. To determine if efficiency problems are the primers or the probe, run your primers with a SYBR Green kit. If they work well, then the issue is the Probe. If they do not, you know you need to optimize the primers.

Set up a qPCR experiment to test PCR efficiency:

1. Dilute the template used for the assay with clean water. Make your 10 fold dilutions and make enough for the number of reactions plus one to account for pipetting error. Use a P10 pipettor (or similar pipettor accurate to very low volumes) to achieve low standard deviations in the Cq values between replicates. 

2. To make it easy for yourself, prepare the template DNA so that the same volume is added per reaction (2 ul) in all wells.

3. In a PCR hood and using designated pipettors, prepare your mastermix (buffer, water, and enzyme) or aliquot your purchased mastermix into the wells of the plate or the special tube or cuvette you are using for your instrument. Make sure to mix the mastermix well before use, especially for commercial mastermixes that may have been sitting. In the bottle, the SYBR Green can settle out and then be distributed unequally between the wells.

4. If you are running unknowns in the same plate, make sure to have room for 10 wells (duplicates) or 15 wells (triplicates) for the standard curve and then you’ll want to leave 2 or 3 wells for negative control (water alone). If you are doing RT-qPCR, you will need to have a -RT control as well.

5. Add the template DNA to the wells in a separate location. In our lab, we use different pipettors for this.

6. Seal your plate with tape or close your cuvettes, and for plates, we centrifuge in a 96 well plate centrifuge briefly. You can also try the method described by Shoba using a Salad Spinner. We never allow the plate to touch any counter surface to protect the outside wells from attracting dust that might affect the path of light for emission or detection.

7. Put the samples or plate into the instrument and run your pre-set program.

That’s about it for getting started.

The main decisions you need to determine before getting started are the template type, primer sequences, and the chemistry (SYBR vs. probes). 

Getting good results begins with a lot of fore-thought and planning in the early steps.  The combination of primers, chemistry, and template are the three essential  ingredients for high quality data and accurate results.  Work this out systematically up front and save yourself a lot of time and pain later.

More reading on related topics:

 Ten Tips for Consistent Real-time PCR

Does Your RT-qPCR Make the Grade? New Rules for Publishing qPCR Results

About the author

Emily Crow

Emily Crow is a graduate student at Northwestern University in Chicago, Illinois. While her current work is in the field of prion biology, she is interested in a broad range of topics, including epigenetics, microbiology, and emerging diseases.

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I always keep an ear open for helpful tips in the lab – those little tricks that can make your experiments faster, easier and better. Here are a few tricks I’ve picked up for trimming down the time it takes to do your cloning:

Restriction digests

Many digests are complete within 10 minutes of digestion at room temperature – no need to be a slave to the “one hour at 37°C” rule. Check in the back of the NEB catalog for their list of high efficiency enzymes that digest quickly, or test your enzyme by yourself to minimize digestion times.

PCR amplification

There are a few ways to cur down on the time it takes to run your PCR steps…

• Trim your PCR programs to complete the run faster: check the enzyme product insert to see how short of a denaturing or annealing cycle you can get away with.
• If you’re not using the PCR product for cloning, you can eliminate the final (long) extension step.
• If you don’t need a lot of product (for instance while PCR screening), try running twenty or twenty-five cycles instead of thirty.

Ligation

Ligations can work surprisingly quickly: T4 DNA ligase can complete a reaction in just 10 minutes at room temperature. Check the product literature for your enzyme to see how short of a ligation you can get away with. Keep in mind that difficult ligations will generally be more successful if you allow the ligation to proceed longer.

Transformation

Just about every step of E. coli heatshock transformations can be shortened…

• Pre-cool the cells on ice for fifteen or twenty minutes instead of thirty
• Do a thirty second heatshock
• Plate the cells after thirty minutes recovery at 37°C, instead of one hour.
• If you are using ampicillin for selection there’s no need for a recovery step at all… more details here

All these changes will affect the efficiency of the transformation, so use them wisely. Difficult or low-frequency ligations need some coddling, so it is probably best to use the full-length steps in these cases, although you might get away with some shortcuts here if you use very highly competent cells.

Agarose gels

• Pour and run your agarose gels at 4°C. They will solidify quicker at a lower temperature, and you can run them at a higher voltage in a cool environment.
• “Pre-cast” your own agarose gels and store them in saran wrap at 4°C for one to two weeks. Then when you need to gel-purify a fragment or screen your clones, you can just pull one off the shelf and get going.

What are your favorite tips for speeding up your cloning?

About the author

Shoba Anantha

Shoba works at a biotech company in Wisconsin. She has MS from the University of North Carolina.

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PCR is highly sensitive, but the downside of that very property is that it makes the technique prone to producing false-positives. In labs where PCR is a staple, like the one I work in, any false-positives are more often than not due to amplicon contamination.

A broken capillary or a PCR plate left carelessly at the table edge is all it takes to aerosolize those amplicons. The next thing you know is it shows up in every PCR reaction that you run.  If you frequently run PCR, then you know what I am talking about.

So what do you do to decontaminate?

Well a simple, but effective way to combat an amplicon contamageddon, is to wipe down everything, equipment, workstations and pipettes, with bleach. Yes, bleach… not HCl. If you routinely use HCl for decontaminating rogue DNA then you definitely need to read on.

Sodium hypochlorite, the active ingredient in bleach, was shown in a 1992 study to effectively protect against amplicon contamination by causing extensive nicking, which prevented the 600bp fragment they tested from being amplified by PCR (see A. M. Prince, L. Andrus PCR: Biotechniques 1992 Mar;12(3):358-60).

The same study showed that even a after 5 minute exposure to 2N HCl, that same 600bp fragment could be detected by PCR. So HCl is not up to the job… its time to get the bleach out.

Which bleach should you use?

The efficacy of bleach in DNA decontamination (and as a disinfectant) depends on the amount of free and available chlorine. 0.05 – 0.5% of free and available chlorine is considered an intermediate level disinfectant and a commercial bleach (like Clorox) contains 5.84% available chlorine so a 10-100x dilution from a commercial stock will work just fine.

I tend to use a 10 dilution of Clorox and, in the above-mentioned study, Prince and Andrus used around a 20x dilution. You can make your own choice.

How and when to decontaminate

Generously spray workstations/equipment/pipettes with 10% bleach, then let it sit for 15-30 minutes (coffee break time!). Then wipe up the bleach and follow up with a water rinse and wipe — bleach is corrosive, so it will damage materials if residue is not removed by rinsing with water.

This procedure should be carried our before and after each PCR, and definitely after a spill. I religiously stick to a weekly cleaning routine of all of my PCR equipment and workspace and that has gone a long way to preventing amplicon contamination

Important points about diluting and storing bleach

Use clean water to dilute the bleach

I use regular tap water and that has worked well so far but hypochlorites are very unstable and decompose very quickly with hard water, reducing the availability of chlorine. So you may want to use a purer water, just to make sure.

Make fresh dilutions as often as possible

Potency of bleach will reduce over time so, it is essential to regularly make fresh dilutions. If you do not smell the chlorine in the bleach, then it is time to make a fresh dilution. We keep our (1:10) dilutions for about 1-2 weeks.

Store dilutions at room temperature in opaque containers

We use regular plastic spray bottles for the dilutions and store them under the sink. Temperature, light and oxygen are all catalysts for decomposition of bleach.

Make sure you use the correct PPE when handling bleach

Lab coat, gloves and safety glasses…. But you do that anyway, right?

About the author

Jode Plank

Jode is a Postdoctoral Fellow studying DNA repair at the University of California at Davis. He received his PhD in Biochemistry from Duke University.

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The release of the iPad this week may bring the long-expected replacement of the paper-bound lab notebook by electronic notebooks one step closer. But are scientists, particularly PIs, comfortable with electronic lab notebooks?

The rise of the tablets
The concept of an electronic lab notebook isn’t anything new, and even the idea of implementing it on a tablet PC has been around for a while. However, as of six months ago, the only tablet PCs widely available were specialized laptops, with a swiveling touchscreen displays. The current crop of tablets, be it Apple’s iPad, Microsoft’s Courier, or any of the other offerings, brings us a lighter, cheaper, more ‘handle-able’ computer that many of us could see sitting beside us at the bench. The power of these machines is diminished compared to the specialized laptops, generally speaking, but still more than enough for the task. So will these technical advances usher in the demise of the paper notebook? It may depend on the software, and our expectations of it.

Physical versus electronic
The advantages of an electronic lab notebook are obvious:

• You can search them. The information that you want is somewhere in the dozen lab notebooks of your predecessor – who wouldn’t want Google to help them find it?
• You can copy them with the click of a button. Creating complete back-ups of paper notebooks is time consuming and laborious, which means it often doesn’t happen. Easy (or even transparent) back-ups of electronic lab notebooks allows storage of current copies off-site in case of disaster.
• They are legible. Do not underestimate this point!
• You can put some pieces of data in them that you just can’t put into a paper notebook. Current scientific data can take the form of movies or of an interactive, multidimentional interface (think genomic and proteomic data), that just can’t be taped into a paper notebook without the “loss” of data. (The data exists, just not in the notebook.)
• Did I mention that you can search them? Just think how powerful that could be.

The advantages of a paper notebook are fewer but significant:

• The book is a universal format. You can pick up a lab notebook from 1985 and read it. Try to read a 5½ inch floppy disk from 1985. Even if you pull the data off the disk, how many files from 1985 can still be read by today’s programs?
• Lawyers love them. A prebound, handwritten notebook, properly signed, dated, and witnessed, is the gold standard for defending patents or concerns of scientific misconduct. Even though many of the high-end electronic lab notebook programs sold to pharmaceutical companies go to great lengths to ensure the integrity of the data, having cracked versions of popular mainstream programs available weeks after their release undermines confidence in these claims, at least to the non-computer science crowd.
• They are cheap. No licenses to maintain, and no worries that the notebook will stop working if the lab has to go a year without funding.

Moving towards a universal format
Some of the electronic lab notebook programs out there have recognized that proprietary file formats worry scientists, and are moving towards more universal formats. The most popular choice seems to xHTML. This seems the logical choice – it can be read by many programs including the handful of web browsers out there. It can certainly handle images, movies, graphs, and even link to particular files when proprietary programs are required to look at the data (sequencing chromatograms, for example). And while it may evolve, the world is invested enough in it to give it the best chance of survival.

How rigorous do you need to be?
This may be controversial, but in an academic lab you may not be as worried about making patent lawyers happy as in industry. I know that academic labs patent things, and that rigorous dating and witnessing of lab notebooks can be important for other reasons, but the truth is that many labs are already violating these rules with their current paper-based system. Here is the range of paper-based notebook rigorousness that I’ve seen in academic labs:

1. All lab notebooks used are prebound with numbered pages, any extraneous material (such as a picture or graph) is glued in, and all pages are signed and dated by the experimenter and a witness.
2. All lab notebooks used are prebound with numbered pages, and any extraneous material is taped into the notebook. Dates are recorded for the experiments, but the pages are not signed or witnessed.
3. Lab notebooks consist of binders, and experiments are recorded on pads of scientific paper, which are added page by page to the binder as they are used. Extraneous material may either be taped to the pages or may simply have holes punched into the edge and added directly to the binder.

Actually, I lied. Even though 1 is the most proper (rigorous) way to keep a lab notebook, I have never seen a lab that handled their notebooks this way. I’m certain there are some that do, but I haven’t seen it myself. I’ve seen some labs which function with system 2, and most that I’ve seen use system 3. System 3 works great, but only if you trust the scientist that created it. So if your lab is using a relaxed paper-based lab notebook system, do we really need to get caught up in this aspect of the security of the electronic lab notebook?

How does free sound?
With the introduction of the new tablets, developers are creating a number of programs that we haven’t really seen before: journaling programs. These programs, in broad brush-strokes, are intended for people to use to record their thoughts, insert images, and link to other files (or web pages). Do these parameters sound familiar? While they certainly don’t offer all the features of commercial electronic lab notebook programs, they do 95% of what most of us would want them to do, and they are essentially free. (If you want an example, check out this video.) Alternatively, with a little work you can use a program you’re already familiar with – Word or an equivalent – to set up your own HTML based electronic lab notebook. These programs would have the lowest form of dated security, consistent with system 3 described above, but you can’t beat the price.

I would love to hear your thoughts. Do any of you work in a lab that uses an electronic lab notebook? What system do you use, and how do you like them? How long do you think it will be before we are all using electronic lab notebooks?