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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If you have ever attempted to load a SDS-PAGE gel only to miss the well, stab the divider, and then watch helplessly as your sample squirts off into the wrong well, then this tip is for you.

The fortunate among us are able to use pre-cast gels with the wells outlined on the gel plate, but home-made gels don’t have this feature. I’ve seen labs with various loading guides printed off on acetate that they stick to the front plate of the gel before loading, which is better than nothing, but only shows you where the well is supposed to be, not where it is. It is true that after loading enough gels you start to develop an eye for finding the wells, but there is an easier way.

The trick is as simple as this: add a bit of bromophenol blue to your stacking gel. If you remember, bromophenol blue is already in your loading dye, so you aren’t adding anything new to the SDS-PAGE equation. I find that adding the dye to 0.003% is enough to color the stacker, but you can adjust the concentration to your liking. To make things even easier, I simply add the dye to my 4X Stacking Gel Buffer (0.5M Tris-HCl pH 6.8, in my case) to a final concentration of 0.012%. Now you don’t even have to add a new line to your recipe. You can see the results below.

Before loading the samples (Yes, I know these are ugly wells.)

Samples Loaded...

...a third of the way into the run...

...and close to the end of the run.

As you can see, the wells are easy to visualize. Once the voltage is applied, all the dye in the gel collapses down into the dye front and the gel runs normally. If the color of the stacking gel is too similar to the color of your protein samples, then you can simply add more bromophenol blue to your concentrated loading buffer (you’ll find that recipes vary on this point anyway). For the sake of the photographs, I ran this gel without the cover – this is potentially dangerous, so don’t try this at home…urr… I mean, your lab.

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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I recently visited a lab that had a salad spinner on their lab bench and at first I wondered if they were putting together a salad lunch there but when I took a peek I got a nice surprise. It turns out that the salad spinner was actually a bench top, “minifuge” version of a plate centrifuge.

What a great idea I thought. A cheap, quick-to-build plate centrifuge that also worked pretty well for a quick spin just before PCR. So, we tried to built one in my lab and we loved it so much that we now have one sitting near almost EVERY PCR plate instrument, and have even gifted a couple to others!

Building one for your lab is simple, here is how…

You will need

1. A salad spinner – We use the Zyliss brand pull-cord salad spinner.
2. Multi-purpose cable ties found at any hardware store.
3. 96-well plate inserts – We use the ones from ABI

Gathering the components is as complicated as it gets! All you need to do now is use those cable ties to secure the 96-well plate inserts to the inner bowl of the spinner as shown below. Then start using the new mini-plate-fuge!

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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We received the following question from Bitesize Bio reader, Beheroze Sattha. It relates to a problem with absolute quantification using plasmids for standard curves. Since many people use this technique it is an interesting one question for us to explore, and it also gives us a great opportunity to cover some important tips for performing qPCR with a new template for standard curves.

Question:

I would like your help to assign copy numbers to plasmid standard dilutions. I cloned a portion of the mitochondrial D-loop gene and after plasmid prep of a single clone made serial dilutions (1:10). In my lab previously they would do a lightcycler run on those serial dilutions using SYBR Green and then assign copy numbers based on the crossing points. The theory used was that a crossing point (CP) of 32.7 would be equal to 10 copies. So the dilution closest to CP of 32.7 would be 10 copies with increasing 10 fold copies for the earlier dilutions (because I had made 1:10 serial dilutions: 1:10, 1:100, 1:1000, etc.

Then I learned from a co-worker that it is better to use the following website:
http://www.uri.edu/research/gsc/resources/cndna.html

So I entered the nanodrop reading of the undiluted plasmid standard and the length of template (TOPO vector and length of my insert) and got the copy number.

The problem is that there is a 10 fold difference between these two methods. 1:1000 dilution of the plasmid standard gives me 8.44E7 copies using the website and 9.99E6 using lightcycler CP.

******

Thanks for your question!

There are a couple things to keep in mind here and I think you’ll be able to solve this problem.

1- The standard curve results and crossing point or Cq numbers are not going to be identical every time. You didn’t mention whether the original standard curve data was performed with the cloned gene or with gDNA or with the PCR product for the gene itself.  If the original standard curves were determined with one type of template and now you have a plasmid template, there will be a difference. Even a change of 1 cycle can have a big effect in absolute quantification.

2- Have you checked the efficiency of the plasmid template? How does it compare to the efficiency of the template used to generate the previous standard curve? If they are different, the quantification will be different.  Ideally you want the efficiency to be above 90%. If the efficiency is below 80%, you won’t want to use this data and you may need to redesign the primers or optimize the chemistry or running times.

2- Every time you order new primers or a new enzyme kit (of a different lot#), you will want to repeat the standard curve results because the numbers can shift. As long as PCR efficiency is high, the data will be accurate, but the Cq may very well be different with new reagents.

3- If you used a plasmid for a standard curve, did you linearize it for qPCR? Many people report that using supercoiled plasmid for standards can cause some variance in results. Try linearizing it first. Here is a recent publication on the subject.

4- When calculating the copy numbers, you may use the length of the PCR amplicon or the entire plasmid. When using the website above, if you use the size of just the amplicon as your input, make sure to adjust the amount of DNA going in to reflect the proportion of the plasmid (see the example in the comments).  If you do not, your reading will be 10 fold off.

5- Make sure your negative controls are negative. Working with plasmids can be tricky because they can easily contaminate solutions. Make sure you have negative controls that are not amplifying because this will boost the real samples and result in inaccurate quantification.

6- Always do the melt curve analysis when using SYBR green and make sure you amplified a single product. Amplification from dimers will add fluorescence and result in an artificially low Cq. You can remedy this by performing an extra data acquisition step at a temperature above where the dimers melt and below where the real product melts. Alternatively, you may want to redesign the primers if dimer formation occurs even in samples with the highest amount of plasmid.

7- With plasmids, it is easy to overload the reaction and have Cq values so early that the detection won’t be accurate. Some instruments have a pre-set baseline setting where they subtract any fluorescent signals generated too early, assuming it is background noise. If you have too much signal in cycles 1-10, this can happen. You don’t want to have samples coming up early so dilute until the first sample has a Cq of 15-18. The subtraction of strong fluorescence in the early cycles will cause all of the data from the more dilute samples to shift right, causing later Cq values than what they are.

Summary:

When using a new plasmid as a standard in qPCR, do an efficiency check  first and compare to the efficiency of the previous assay. For best results, an assay needs to have >90% efficiency, although there are formulas you can use that normalize for differences in efficiency. It is not uncommon that the Ct values are not exactly the same from user to user and from assay to assay for the same gene but with different primers. Just make sure that when calculating copy numbers, you are using the length of the template being amplified and not the entire plasmid which is probably 30 fold bigger than the template and could be the cause of the 10 fold difference in copy number results between the two methods you are comparing. Finally, it’s always good to re-check your standard curve with each new purchase of reagents to make sure no new variables are introduced that could throw off the quantification and make months of work unusable.

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 every lab has a small toolbox with some screwdrivers, pliers, and such, here are some tools that may not have obvious utility at the bench, but could make your life easier.

A Butane Torch
If your OCD is as bad as mine, then watching a bubble flow out of the flask onto the Petri plate you’re pouring bothers you… a lot. A short (~6 inch) butane torch comes in quite handy for flaming bubbles off the top of freshly poured plates. I also use this to pop bubbles that form while pouring agarose gels, but you have to be particularly careful while doing this not to damage the casting tray or any other part of the gel rig. In addition, these guys come in real handy as a portable Bunsen burner when you need to work away from a gas outlet.

What to look for: Almost all of them can be refilled with the generic cans of butane, but double-check to be sure the one your looking at doesn’t require some proprietary refill system, which will almost certainly be more expensive. In addition, make sure that it has an auto-start feature. If you think you might use it as a Bunsen burner, make sure that it has a large, stable base and a button to lock the trigger ‘on’.

What I use: Ronson Tech Torch

An Infrared Thermometer
Ideally, you would cool a flask of autoclaved agar in a waterbath, but that isn’t always possible. And once you leave it cooling on the counter, the gamble begins. Pull it too early, and you risk inactivating your antibiotic. Pull it too late, and you’ll end up with lumpy plates. (Did I mention my OCD?) One solution to use an infrared thermometer to monitor the flask.

After acquiring mine, I set up a test where I outfitted a flask of boiling water with a stir bar and a traditional thermometer, and compared the readings of the IR thermometer (aimed at the outside of the flask, below the fluid level) and traditional thermometer (in the fluid) as the water cooled on a stirplate. The two readings didn’t differ by more than a degree or so the entire time the flask cooled. This is particularly valuable if you have inexperienced people (ie – undergrads) making the plates in your lab. In addition, these things are great for getting an immediate temperature on anything without having to wait for a traditional thermometer to equilibrate. Once you have one of these, you will use it more than you think.

What to look for: Make sure that the thermometer you’re looking at can be switched to output Celsius (if you’re in the US or Canada). Also, you might want to get one that has “adjustable emissivity”. Emissivity is the ability of a material to radiate energy, and in practical term this means that glass at a particular temperature will emit a different amount of infrared radiation than aluminum at that same temperature. If you find that you want high sensitivity for one particular application, then you can adjust this parameter to fine tune the thermometer. (A quick Google search for “emissivity coefficient” will turn up tables of emissivity settings for common materials.)

What I use: Advanced Tool Design Deluxe Infrared Thermometer

Strap Wrenches
As a group, we scientists aren’t known for our intimidating physiques, and this is usually revealed in the lab when the liquid nitrogen knob, or the top of a centrifuge canister or bottle gets stuck. You then have to go find the largest scientist you can think of to help you out of the jam, which for guys is “The Walk of Shame”. Strap wrenches will save your pride or even save your experiment if nobody is around to help. These tools consist of a flexible strap that wraps around and grips the object, with a straight handle that allows you to get some leverage on the beast. (If you’ve ever watched a mechanic change your oil, he likely used a specialized version of this tool to remove your oil filter.)

What to look for: Choose ones that have a urethane coated nylon strap to make sure that it doesn’t damage anything you might use it on. They are adjustable, so choose ones with a large capacity (~5 inch diameter), as they will also likely adjust down to less than 2 inches for smaller tops. You will want two, since you may have to wrap one around the bottle and a second (in the opposite direction) around the lid. Of course, be careful if you are removing the stuck top off a glass bottle.

What I use: Klein 12-Inch Strap Wrench

Are there any tools that I missed?

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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Recently we received a question from Bitesize Bio reader Sonia after our article How to: Get Better Plasmid Midiprep Yields. She asked: “What could be the problem when one sample gives a good yield while the other plasmid gives poor a one, when both the samples were processed simultaneously, and in the same way.”

This is a good question because many things can cause differences in yields between plasmid preps.  Let’s resolve this mystery one point at a time and go over some reasons why you might get low yields when you prep plasmids.

1. Plasmid backbone

You prepped two plasmids simultaneously using the exact same protocol and had different yields. If the plasmids have the same backbone (that is, they are both pUC, or pBluescript, etc.) then the reason leans towards the insert playing a role. Some inserts can be problematic for bacteria. It might be that a protein is made that makes the bacteria sick (for example, DNase) or it could be that the insert is unstable (for example, repetitive sequences). To overcome the problem, try using a specialized competent cell line. For unstable inserts, try the STBL2 cells from Life Technologies and for growing clones with toxic proteins, try the T7 Express LysY/Iq Competent cells from NEB.

Another important point is how the insert size changes the copy number of the plasmid. Large inserts will reduce the number of copies of the plasmid.

2. Copy number

If the genes are cloned into different vectors then the issue could be that the plasmids are replicating at different rates. One may be high copy and one may be a medium or even low copy plasmid. Some examples of  low copy plasmids are ones using the backbone pBR322 and pACYC, which are older and not used often in cloning work today. Many vectors used for protein expression are medium copy. This is desireable because when producing proteins, sometimes if growth is too fast, it enhances the chance of the protein becoming insoluble or forming inclusion bodies.

3.  Culture issues and antibiotic

Nick went over the technique for growing bacteria to obtain a healthy culture in late log phase where you can get the most plasmid in our original midiprep how-to article.  This technique works well as does using a 1:1000 dilution of starter culture into a large scale culture (so 100 ul into 100 ml) if you want to grow the bacteria overnight for 12-16 hrs.  One important consideration is the antibiotic. The bacteria are going to break down the antibiotic while they are growing in the culture. If not enough antibiotic is added or if the stock is old and not at the correct strength, the antibiotic selection pressure may not last very long and you could end up having a culture that was antibiotic-less for most of the culture time. Plasmid yields will go down without the selective pressure to keep it.

As a reminder, the state of the culture is critical for high yields of plasmid. For maximal yields, the culture should be in late log or early stationary phase. If the culture overgrows, you will be harvesting more dead bacteria than live cells and this also leads to genomic DNA contamination in the prep. If the culture is undergrown, then of course, yields are lower than expected. You can mistakenly undergrow a culture by using old colonies from plates or starting direct from a frozen stock and not from a colony. The lag time for the bacteria to ramp up is much longer when you use either of these approaches.

Streak fresh colonies:

One more point that people forget when setting up their starter cultures or overnight cultures is the age of the plate you are using to pick your colony. If your plate is old, you may have picked a nice big colony but it will not be all living cells. And if there were satellite colonies sitting around the original colony where the antibiotic no longer exists, those will not have plasmid and will be introduced into your culture. So streak a fresh plate before starting to ensure the best result.

So let’s assume that Sonia’s plasmids were the same vector, same antibiotic, grown exactly the same, and each have different inserts that are not too large or unstable or toxic. What else can it be?

4. Processing steps:

If the culture conditions are not the problem, then we have to look at something with the downstream steps. Since we are talking about Midipreps, let’s discuss what can go wrong with the anion-exchange procedure next.

Nick covered alkaline lysis in great detail and typically these reagents in the kits are stable and fine. Solution 2 (the one containing NaOH and SDS) can break down over time with exposure to air, but in general, they work for lysing bacteria for the life of the kit.

The other area of plasmid preparation where DNA can be lost are the final steps after anion-exchange which is the final precipitation step in isopropanol and finding the DNA pellet.

Isopropanol quality:

Many labs have isopropanol in large containers that have been opened and closed over the course of a year. For the best result in the precipitation step, make sure the isopropanol used is not the old bottom-of-the-barrel stuff. Use some isopropanol from a new bottle or a smaller bottle that is not who-knows how many years old. This makes a huge difference in the size of the DNA pellet you obtain after centrifugation.

Don’t lose the pellet!

Isopropanol pellets are glassy and clear and difficult to see. The best practice is to mark the side of the tube where you expect the pellet to form after centrifugation in a fixed angle rotor so when you decant the isopropanol, you know where to look for it. Keep an eye on the spot and look for the glassy material. Sometimes this is difficult because many people use the oakridge plastic tubes which are opaque. If you have glass corex tubes, this is a nice alternative and they can be baked to make them pyrogen free.

Sometimes, if you have concerns about losing the pellet, it is good practice to pour the isopropanol supernatant into a 15 ml tube to save it, just in case the pellet slipped off the wall. But this does not normally happen as long as you do not let the sample sit for long after the centrifuge stops. Once it is done, be right there to decant the sample. The only times I have seen a pellet come loose from the wall is when I was late getting to the centrifuge and it sat still for a few minutes.

Whether you use Oakridge tubes or glass, just note where that pellet should be. Once you wash with 70% ethanol, the pellet becomes visible.  When you are ready to resuspend your pellet, you’ll know exactly where to find it because you marked the tube.

Caution! It is not always a pellet!

Sometimes with fixed angle rotors, the DNA may not always form a nice tight pellet at the side wall. It can sometimes smear down the side. For this reason, I always use my resuspension buffer to wash down the side of the wall above my pellet to make sure I solubilize every molecule of plasmid that may be present even though I can’t see it.

It is a shame to do all that work and then lose the DNA pellet right at the end! More tips on perfect DNA pellet recovery is found here too.

We had a nice discussion about DNA precipitation in a previous article and it was the consensus that the most important factor in obtaining high yields is centrifugation speed and time. Don’t cut the centrifuge time short unless you can turn up the speed.

I thought it would be good to mention that many plasmid kit manufacturers have recognized that the pelleting step is problematic for some users so have developed kits that desalt the DNA using “precipitators” or silica disc filters. These are a fast alternative to centrifugation. However, you still need good isopropanol for these to work so always use fresh.

Summary:

Large scale plasmid DNA preps have a lot of steps where things can go wrong but in my experience, the problem is usually either the culture or the DNA precipitation.  To check if your culture is healthy, just take 1-2 mls out of your 100 ml flask and then do a quick miniprep on it to see how much plasmid/ml is there. That will give you a good idea of what you will get from the rest of the sample.

So remember to get great plasmid yields, do a little background first on your vector and insert to make sure there is no reason for the DNA itself to be a problem and then start with a fresh colony and a starter culture and fresh antibiotic. And at the end, fresh isopropanol will be key to a thorough precipitation of all the plasmid DNA.

That about covers it folks. If you have any more questions, problems, or concerns, send them our way and we will put our heads together to figure out how to help!

About the author

John Mackay

John is interested in using molecular diagnostics to solve industry issues - working on samples as diverse as wine, truffles, grapevine and human. He also consults on PCR and real-time PCR assay development as technical director for the company dnature.

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If you’re performing DNA/RNA precipitations, you will have read Suzanne’s excellent article on which alcohol to use for precipitating your precious samples (check out some useful info in the comments for that article as well).

Its publication prompted the recall of a useful tip I learned from a post-doc many years ago,  one of those ‘magic-hands’ scientists where everything seemed to work first time. This tip of his is on the removal of the 70% ethanol in the final step of the precipitation process.

The usual wisdom is to carefully drain off the ethanol and leave it to either air-dry or place in a speed-vac. The problem is that air-drying can be too slow, while the speed-vac can be too fast meaning that you risk over-drying the pellet making solubilisation more difficult.

So here’s the tip: After draining or pipetting off the 70% ethanol, simply:

• recap the tube
• pulse the centrifuge to bring down the remaining ethanol
• remove this liquid with a pipette and 200ul tip – you can get right alongside the pellet if visible. Leave the tube open as you move to the next sample.

By the time you’ve finished your series of tubes (even if n= 3), the pellets are dry always enough to add diluent immediately. Job done. This saves time and give you samples are just the right degree of “dry”, every time.

Any other tips for ethanol (or even isopropanol!) precipitation would be great to hear. Meanwhile – thanks Michael.

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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There’s something in the water, and it would love to go after your experiments.

Straight out of the tap, water contains microorganisms, endotoxins, DNase and RNase, salts and other impurities that could gobble up your experiment in one bite. Of course we avoid this drama completely by using purified water from which these nasties have been removed. But how is this purification done?

Well in practice a number of techniques are used, each of which can remove a different set of impurities.

So here are the techniques:

1. Distillation. A technology as old as the hills (or at least as the stills that were hidden in the hills). Water is heated to its boiling point then condensed back to liquid. This will remove many impurities but impurities with a boiling point equal to or less than that of water will also be carried over in to the distillate.

2. Microfiltration. In this technique, pressure is used to force the water through a filter with pore sizes of 1 to 0.1 micron in order to remove particulate matter. Filter diameters lower than 0.2 micron removes bacteria – so-called cold sterilisation.

3. Ultrafiltration uses even smaller pore sizes (down to 0.003 micron). These are essentially molecular sieves, which remove molecules with a diameter larger than the pore size. It can be used to remove viruses, endotoxins, RNase and DNase

4. Reverse osmosis. If you thought that ultrafiltration used impressively small pore sizes, you’ll be even more impressed by reverse osmosis . Reverse osmosis filters have pore sizes of less than 0.001 microns, which allows them to sieve ions depending on their diameter. This is used for desalting the water.

5. Filtration through a bed of activated carbon is useful for removing things like chloride ions and organic compounds, which are adsorbed onto the surface of the carbon.

6. UV radiation. We all know what UV radiation, at specific wavelengths, can do to DNA and microorganisms. So UV is an obvious way to remove microorganisms from the water. It can also clean up the water by breaking down certain organic compounds into less harmful products.

7. Deionization/ Ion exchange. This technique removes ions from the water by passing it through a resin bed containing a mixture of cationic and anionic resins. Positive ions in the water are attracted to the anionic resin particles and negative ions are (yes, you’ve guessed it) attracted to the cationic resins. The result is that nicely deionised water comes out of the other end of the resin bed.

Commercially available water, or water purification systems will typically use a combination of these. The higher the water purity grade, the more techniques used.

Any questions or comments? Just jump in and join the discussion in the comments section below. The water’s lovely.

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.

To enable tagging you will need to register on Bitesize Bio. We're sorry for the inconvenience, but it's free, only takes a few seconds, and it will enable you to view our seminars for free, ask questions from the professional community, and take part in the lively community of Bitesize Bio

Since our most popular article of all time (“The Basics: How Ethanol Precipitation of DNA and RNA Works”) was published, many of our readers have asked us to further explain the difference between precipitating DNA with ethanol vs. isopropanol and which is the better choice. So today, I’ll meet the challenge and discuss the pros and cons of ethanol vs. isopropanol.

First, let’s review what we know about what is needed for precipitation of DNA or RNA with ethanol:

1. Salt to neutralize the charge on the nucleic acid backbone, causing the DNA to become less hydrophilic and fall out of solution.

2. Ice to chill the sample. Lower temperatures promote the flocculation of the nucleic acids so they form a larger complex that readily pellets under the centrifugal forces of a microcentrifuge.

3.  A nucleic acid concentration high enough to force the DNA out of solution (if the conc is not high enough, you can add a carrier nucleic acid or glycogen to enhance the recovery).

4. Centrifugation to pellet the sample

The difference between isopropanol and ethanol is the solubility of DNA in each solvent.

DNA is less soluble in isopropanol so it will fall out of solution faster and at a lower concentration, but the downside is that the salt will too. With ethanol, the DNA needs to be at a higher concentration to flocculate but the salt tends to stay soluble, even at cold temperatures.

DNA falls out of solution in 35% isopropanol and 0.5M salt. Using ethanol, the final concentration needs to be around 75% with 0.5M salt. So for the typical precipitation protocol, isopropanol is added from between 0.7-1 volumes of sample and ethanol is added at 2-2.5 volumes of sample.

The choice of which solvent to use depends largely on the volume of sample you need to precipitate.

If you are precipitating small volumes of DNA, and you can fit the required amount of solvent into the sample tube, then ice cold ethanol is the preferred choice. You can chill it (some people use liquid nitrogen or -80C to accelerate the precipitation) and precipitate more DNA without the salt contamination that would occur from chilling isopropanol. Afterwards you need to wash the pellet with 70% ethanol to remove salt.

Isopropanol use useful for precipitations where you have a large sample volume  (e.g. the eluate you get after using a Qiagen plasmid Maxi Kit) because less solvent is needed, so you can fit the whole lot in the (15ml) tube. But because salts are generally less soluble in isopropanol than in ethanol, they have more of a tendancy to co-precipitate with the DNA. So to lessen the chances of salt precipitation, isopropanol precipitations are carried our at room temperature with minimal incubation times.   Once the DNA or RNA pellet is recovered from the isopropanol, you’ll want to wash it with cold 70% ethanol to remove excess salt and to exchange the isopropanol with the more volatile ethanol. It is ok to chill the isopropanol precipitated sample, if you are sure that it is not excessively salty.

Because DNA is less soluble in isopropanol, isopropanol allows precipitation of larger species and lower concentrations of nucleic acids than ethanol, especially if you incubate it cold and long. If you do this, just remember to wash the pellet several times in  70% ethanol after pelleting, to reduce the amount of salt you carry over.

So how do you choose when to use isopropanol and when to use ethanol?

Use ethanol if:

1. You have room to fit two volumes of ethanol to sample in your tube
2. The sample needs to be stored for a long period of time and will be chilled
3. You need to precipitate very small pieces of DNA or you have a very low concentration of sample so you want to chill it longer and colder.

Use Isopropanol if:

1. You have limited in space in your tube and can fit only 1 volume of sample
2. You need large molecular weight species because incubation at room temperature for short periods of time will not be conducive to precipitating small species of nucleic acid
3. You are in a hurry and want to accelerate the precipitation of nucleic acids at room temperature

What do I prefer? I use ethanol over isopropanol for most cases, but will use isopropanol if I need to make everything fit in one tube. My preferred protocol is 2 volumes of ethanol and freeze at -20C for at least an hour or overnight for best results. I centrifuge the sample at full speed for 20 minutes to make sure I get everything down. I always wash with 70% ethanol and then centrifuge for 10-15 minutes and keep my eye on the pellet when I decant everything. You need to note or mark the side of the tube where the pellet is expected to be and don’t let it out of your sight when decanting the ethanol!

If I use isopropanol, I avoid cold temperatures because of the excess salt that usually comes down with it. If I want to increase the yields precipitated, I prefer to leave it incubating at room temperature longer vs. chilling the sample. When the DNA is pelleted, the pellet is sometimes more difficult to see compared to the ethanol pellet. It can be clear and glassy. Make sure, again, to note the side of the tube where the pellet should be. Look for it before decanting the isopropanol and 70% ethanol wash. After washing with ethanol, the pellet becomes visible and white. I always make sure it doesn’t slip off the side of the tube wall before decanting the supernatant. Allow the tube to drain upside down for a few minutes and then air dry or speed vac dry (5 minutes is enough) and then resuspend in buffer.

Finally, for dry DNA pellets, heating the sample in buffer at 50-60C will help the DNA dissolve faster and won’t damage the DNA. For RNA, heating can be used too (in water) at temps around 42C.  Overdried DNA and RNA will take longer to dissolve so make sure not to speed vac for too long.

So now you know the difference between ethanol and isopropanol and when to use which. If you have any questions, or anything to add, please drop a comment in below.

About the author

Eric J. Perkins

Eric is a Senior Scientist at Addgene, the non-profit plasmid repository. He also consults for LabLife.org.

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Working at a plasmid repository, I get a lot of feedback from scientists who are relieved we exist simply because that means they don’t have to request a plasmid directly from another academic lab. Either they’ve had a bad experience making requests in the past, or they really don’t know how to go about doing it in a professional manner.

Nothing you can do will guarantee that a plasmid request will be fulfilled, but here are a few rules of etiquette to consider. Note that I’m talking about plasmids, but most of these “rules” can be applied to nearly any biological reagent:

Asking for the Plasmid

1. Introduce yourself and explain why you want the plasmid

Remember that you are essentially asking your colleague for a favor. Yes, if a scientist publishes a plasmid,   there is an obligation to distribute it, but remember who put in the work to make the reagent. Explain who you are, where you found out about the plasmid, and if possible, explain how you might be using it. An ethical scientist should provide you with the plasmid regardless of how  you’re using it, but if  you can demonstrate that you’re not a direct competitor, that might move things along for you.

2. Pay for shipping

This one might seem like a no-brainer, but I’ve talked to scientists who had essentially stopped honoring plasmid requests because they were spending a small fortune on shipping. Asking a scientist to send a reagent and pay for the shipping takes some hutzpah. Cough up your FedEx number and you’ll likely get your plasmid a lot faster.

3. Cooperate with the Material Transfer Agreement (MTA) process

The vast majority of plasmid sharing between academic labs is probably done outside the MTA process. That might be the easiest way of doing things for the parties involved, but it’s not necessarily the best (or legal) way.  Many institutions, especially HHMI, are quite rigid about their MTA guidelines and they require researchers to fill them out for every reagent that’s sent out. Don’t begrudge scientists for making you fill out this paperwork–they’re just following the rules. That plasmid you’re requesting is technically “owned” by the institution, not the lab.

After Receiving the Plasmid

1. Show some gratitude

Congratulations! You received your plasmid. Be polite and say thank you. You never know when you might run into this person at a meeting or want to propose a collaboration.

2. Make sure you received what you asked for

Before you go transfecting cells all willy-nilly, you better make sure this plasmid is what you think it is. At the very least, you should do a diagnostic digest. But to really be on the safe side, sequence it. Whether the lab published on this plasmid 1 month ago or 10 years ago, mistakes can be made. Play it safe and do some testing before you commit to an important experiment. If you do find that the plasmid has a problem, let the person who gave it to you know, diplomatically.

3. Acknowledge

If you publish anything that uses these requested reagents, give credit where credit is due. Not only is it the right thing to do, but but it will send people to the right lab should they read your paper and want to request the plasmid in question. Unless you modified the plasmid in some way, you should not be redistributing it without the original lab’s permission. A proper acknowledgment should prevent the issue from arising.

And finally, if you’ve ever used Addgene, you know that we take care of a lot of these steps for you. If you see a published plasmid that you want and it’s not already in the Addgene repository, e-mail help@addgene.org and let us know. We’ll request it for you (and take care of some of the quality control, AND take are of the MTAs).

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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TAE or TBE, which is best? Well, of course, it depends on what you want to do. Here are the pros and cons of both:

• TBE (Tris-borate-EDTA) is a better conductive medium than TAE (Tris-acetate EDTA) so is less prone to overheating so use TBE for long runs
• Borate is an enzyme inhibitor so TBE is not a good buffer to use if you will be isolating the DNA for downstream enzymatic steps. For example, borate carry-over could affect your ligations, so use TAE instead.
• Acetate gives improved separation of large DNA fragments
• On the other hand, borate resolves <2kb fragments better, so use TBE for smaller fragments
• And finally, TBE is reputed to give nicer bands than TAE (although I’m not sure why – does anyone know?)

But you might be better of using neither of these buffers. Despite the fact that they have been firmly established as the most popular buffers for DNA electrophoresis since their emergence in the early 1970s, TBE and TBE are not really optimised for the job and have a lot of disadvantages.

Borate and acetete are simply carry-overs from earlier work on  protein electrophoresis buffers upon which the technology of DNA electrophoresis was built.  And while the reasons that Tris was chosen as the primary cation by early pioneers of DNA electrophoresis are not clear, the draw-backs certainly are. As well as being expensive, Tris-based buffers are not ideal for DNA electrophoresis because, even under optimal conditions, they are prone to a temperature-current feedback loop that causes the solution, and therefore your gel, to overheat. And finally, EDTA is a relic from RNA electrophoresis, which has been unecessarily retained in DNA electrophoresis buffers.

In an earlier article here on Bitesize Bio, Bala described new and improved DNA electrophoresis buffers developed by scientists at John’s Hopkins University. These are optimised for the job and are faster, cheaper and less prone to overheating than TAE and TBE.

For more information on this, check out this article, which details the history of DNA electrophoresis buffers and the development of the new buffers. The authors of this paper (and the inventors of the new buffers) have started a company called Faster, Better Media to manufacture and distribute the buffers.

So what do you normally use; TBE, TAE, or something else?