Introduction

We will cover only the very basics of nucleic acid structure here, just enough to understand how to isolate them and work with them.

We will go through several procedures for isolating DNA and RNA and then how to check the amount and purity of the samples you have isolated. These procedures are quite standard in many labs. Although you may find some of the details a little different in a lab you work in (depending on the organisms you are studying), the basic principles are pretty much the same and understanding these principles will give you an edge in your future lab work.

A. DNA and RNA structure:

Here is a link to a video that nicely explains the basics of DNA and RNA structure. It covers the subject at the right level for our course.

Nucleic acids are polymers of nucleotides and nucleotides have 3 parts: a pentose sugar, a phosphate group, and a nitrogenous base. You don’t need to be able to draw every detail of the structure of a nucleotide but you should be able to:

• sketch the pentose sugar (a stick diagram- no details)
• identify the carbons, 1′ to 5′
• indicate where the base and phosphates are attached to the sugar
• indicate how the ribose sugar differs from the deoxyribose sugar and which bases have a single ring and which have a double ring.

I remember the purines are the larger bases but have the shorter name and vice versa. It is useful to have prompts like this for remembering terminology and other basic information. That is why I suggest using your transcript – a copy of your grades – as a prompt for distinguishing between transcription and translation.

You also need to know the complementary base pairing, A with T and G with C. Although it is not mentioned in the video, the A – T pair is bonded more weakly than the G – C pair. A and T have two hydrogen bonds between them and G and C have three hydrogen bonds between them, giving the G – C pair a stronger attachment. This will be relevant later in the course.

And finally and maybe the MOST important concept to know is the polarity of the RNA and DNA molecules (5′ to 3′) and the fact that complementary base pairing is antiparallel – the molecules line up with opposite polarity. The complementary base pairing won’t work if the two molecules are lined up with the same polarity; the shapes of the bases don’t match and cannot form hydrogen bonds in that arrangement. How primers recognize their targets, how PCR products are made and how pieces of DNA can be attached to each other by several different methods all rely on complementary base pairing. So it is important to understand.

B. Principles of DNA isolation:

Here we will cover several common methods of DNA isolation. In this section the main focus is on the components of the buffers and other solutions that are used during the preps, and the purposes of each step. No details will be given except where relevant (such as the pH of a solution). There is no need to memorize exactly how much salt is in a buffer because if you are working in a lab you will probably have made the buffer yourself and will know very well how much salt is in it. But what you should know is WHY the salt is in the buffer. What is it doing? How does it affect the DNA or the cells that are being lysed?

There are three important aims in making a DNA prep. We want the prep to be pure DNA and free from other components such as RNA and most importantly, proteins. We want the DNA to be isolated in large pieces. Handling (pipetting mainly) will cause some breakage of the DNA strands, but so long as the pieces we’ve isolated are larger than the piece we want to clone, then they are large enough. We also want the DNA to be concentrated enough to work with. There must be enough of it to see on a gel for instance, which requires a fairly high concentration. And we want to perform downstream reactions like PCR and restriction digests or sequencing in fairly small volumes. I’m usually pretty happy if my prep is between 100 and 1000 ng/μl. Lower concentrations can work too, depending on what we plan to do with the DNA.

For each procedure there are three main steps:

1. The tissue or cells are mechanically broken up if needed (some cells don’t require this)
2. A crude lysate is produced using a buffer that both disrupts cellular structure and protects DNA from degradation
3. The unwanted materials are removed from the solution, leaving behind just the DNA, purifying and concentrating it. The details of this last step are most variable between the methods.

B-1. Prepare the material (all methods)

The beginning of any procedure for DNA isolation is to obtain the material from which you will isolate the DNA and to break open the cells to release the contents, including the DNA. The way the cells are disrupted can differ greatly depending on your starting material.

Cheek swab cells or blood samples require only an extraction buffer to be added. The buffer causes the cells to break open, and the contents are released. Bacterial preps need only to be spun in the centrifuge to pellet the cells. The media the cells have grown in is discarded and the cells can be directly resuspended in the extraction buffer.

Small samples of tissues may be mashed in a mortar or crushed with a small homogenizer. Harder tissues such as bone or tree bark, or other plant materials are frequently ground in a mortar with liquid nitrogen. The ultra cold temperature is advantageous both in making the tissues very brittle and thus easier to crush, and in keeping the material so cold that enzymes such as nucleases are essentially inactive. Some materials- especially very tough substances like pieces of bone or some types of tree bark- are put into a mechanical mill and are pounded with steel balls. If you go online, you can see many copyrighted images of different types of equipment used to disrupt tissues in preparation for DNA (or RNA) isolation.

If your material doesn’t have the extraction buffer in it at this point, is important to move quickly to the next step and get the prepared material into that buffer as quickly as possible. Nucleases are everywhere, including the tissue that you’ve just prepared!

B-2. Prepare a crude lysate (all methods)

Extraction buffer recipes might vary somewhat from lab to lab but most will contain the following ingredients:

Tris, pH 8.0: Tris is a buffer which will keep the solution slightly basic, the pH at which DNA is most stable

EDTA: this chelating agent will chelate Mg⁺² ions. These are a needed cofactor for DNases, enzymes that degrade DNA. Without the cofactor, it is much harder for the DNases to degrade the DNA in your prep.

A strong detergent such as SDS (sodium dodecyl sulfate) to disrupt the cell membranes, and the nuclear envelope and to disrupt hydrogen bonds that keep proteins folded correctly. A strong detergent will denature proteins, including DNases.

NaCl helps solubilize the DNA in this buffer and will help precipitate it in a later step.

Some buffers also contain a guanidinium salt which disrupts hydrogen bonds and promotes the sticking of DNA to silica beads. This is the case for the column chromatography and solid phase methods, listed below.

Proteinase K, freshly made, is added to the buffer after the buffer is added to the cells/tissue. The tubes are then incubated at 37-65^oC. The enzyme works faster at higher temperatures, and in the lab we usually use 65^oC though other labs have different protocols. In any case the protease degrades proteins, including DNases. Degradation is not the same as denaturation.  Denatured proteins are unfolded but not destroyed; in theory if they could re-fold correctly, they could still function. Degraded proteins have been broken into pieces and are thus destroyed.

Incubation in this buffer is usually followed by centrifugation to remove cellular debris. The lysate – the liquid – is removed to a new tube, and the step-wise removal of unwanted components of the lysate begins.

B-3. Remove the non-DNA components of the lysate

There are several methods that can be used for the next step.  These are the most common.

B3-i. Phenol-Chloroform method

The crude lysate is moved into a new tube and 1V of phenol ph 8.0 is added. Phenol is a caustic agent that can cause burns, so care is used in pipetting it. You must wear gloves when handling it and we keep the phenol in the fume hood and do all the pipetting of the phenol in the hood as well. Phenol, being an acid has a naturally low pH and so must be buffered to maintain the pH at 8.0. If you have worked in a lab or if you do in the future you will have to take care when pipetting the phenol that you do not pipette instead the buffer that is above the phenol in the bottle. This is an easy mistake to make when you are starting out. The buffer is an aqueous solution so if you spin your tubes and there is no obvious organic and aqueous separation you have most likely pipetted in the buffer rather than the phenol.

The tube is tightly capped and shaken to ensure that the phenol contacts all of the solution. Phenol, being hydrophobic, will attract the hydrophobic components of the cell lysate: lipids and proteins. The tube is then spun in the centrifuge and the phenol, along with the hydrophobic components of the lysate, settles to the bottom of the tube. In this organic layer or phase, some lipids will be found, along with proteins that are predominantly hydrophobic. The aqueous layer lies above the organic layer and contains the nucleic acids along with other molecules from the cells. Depending on the tissue used, there may be a thick layer of material between the organic (lower) and aqueous (upper) layers. In this interphase, most proteins and some types of lipids, that have both hydrophobic and hydrophilic regions will be found. Because they can interact with both the organic and the aqueous phases, they end up lying between the two. The aqueous phase is pipetted into a new tube, taking care to avoid touching the interphase with the pipette tip – the proteins there are sticky and many could be accidentally transferred into the tube. It is better to be careful and leave a little of the aqueous layer behind and have a clean prep, rather than trying to get every microlitre of aqueous and risk transferring nucleases into the new tube.

If we want extremely clean DNA – some procedures require this – we can follow this step with several extractions of phenol:chloroform:isoamyl alcohol, in this proportion: 25:24:1. Chloroform is also an organic solvent and in this step it enhances the separation between the phases while the phenol is still attracting hydrophobic components of the cell lysate. The alcohol is added to prevent the solution from getting frothy or foamy – which interferes with good separation. These extractions ensure that the vast majority of proteins and lipids are removed from the prep. The final extraction is 1V chloroform:isoamyl alcohol (24:1). This removes any tiny droplets of phenol that may be left behind in the aqueous solution. Phenol will greatly inhibit any downstream enzymatic reactions so we need to remove it from the solution.

Whether two extractions (a phenol followed by a chloroform) or several (including one or more phenol:chloroform) are done, after the final extraction, the aqueous is moved into a new tube and in some protocols 0.1 V 3M sodium acetate, ph 5.2 is added. This is needed if the original extraction buffer did not contain enough salt to promote the precipitation of the DNA. Then 2.5 V ice cold ethanol (95% or 100%) or 0.6 to 0.7 V isopropyl alcohol is added to the tube. If there is sufficient DNA in the solution, inverting the capped tube a few times should cause the precipitating DNA to become visible as slimy threads or a small “knot” in the tube. In a very small prep you may not see this. Procedures may vary at this point. Some protocols suggest keeping the tubes at -80C or in the regular freezer (-20^oC) overnight. Some suggest an hour or two on ice or even at room temperature. Some go directly to the next step- it depends on how efficient you need to be. A long incubation at low temperature will increase the yield of DNA.

The precipitated DNA is spun in the centrifuge to bring the DNA to the bottom of the tube in a pellet. Generally the pellet will be whitish in appearance. The fluid in the tube is removed carefully, without disturbing the pellet. The pellet has quite a bit of salt in it, which helped the DNA come out of solution but next we want to get rid of some of that salt. Otherwise it might interfere with downstream procedures. So we do one or several “wash” steps, also called “desalting” steps. We add a large amount of 70% ethanol, and flick or invert the tubes multiple times to ensure the solution interacts well with the pellet. The DNA will stay precipitated but some of the salt will go into solution in the lower percent alcohol. The tube is spun again in the centrifuge. The less salty pellet is often clearer and harder to see. The alcohol solution is removed and the pellet is allowed to air dry for just a few minutes. Then TE ph8.0 is added. TE buffer is Tris (10mM) plus EDTA (1mM). You should be able to explain why these components of the buffer promote DNA stability.

B3-ii. Protein precipitation method

In the protein precipitation method, the proteins are not removed by organic solvent extraction but instead by using high salt and low temperature conditions to preferentially precipitate proteins while leaving the DNA in solution. We start with the crude lysate, that has been treated with proteinase K, but add more salt to the solution and incubate the tubes on ice for an hour or more. This causes the protein fragments resulting from cleavage by the protease to come out of solution. Carbohydrates will also precipitate out of solution. The tubes are then spun in the centrifuge and the resulting supernatant is removed to a new tube for precipitation of the nucleic acids with ethanol or isopropyl alcohol (as described for the previous method.

The advantage of this method – which we usually use in the DNA isolation lab – is to reduce the use of organic solvents. Phenol can cause burns. It is also – unfortunately – a topical anaesthetic so that you cannot feel the burn on your skin right away. Chloroform is a cumulative carcinogen – this means that small exposures over time can build up to increase the chances of a cancer developing. I will post the MSDS (the Material Safety Data Sheets) on these two substances in the Lab Archive notebook. You should familiarize yourself with the hazards and what to do if a spill or exposure occurs. Our general approach in the lab is to minimize the usage of any hazardous chemicals in the interests of protecting students.

This method works fairly well in our lab, but we do not have enough time to incubate the solution long enough for an efficient protein precipitation. So we do follow with a chloroform extraction to ensure the DNA is protected from nucleases. We use the fume hood and exercise care when working with even the small amount of chloroform in this procedure. When pipetting chloroform you need to work quite quickly. Chloroform has almost no surface tension so it drips out of the tip almost immediately. It takes a little practice to pipette it accurately.

The video below is the second portion of the DNA isolation lecture, focusing on the main points of the above two methods (Click here to download the powerpoint slides presented in the recording):

B3-iii. Column purification

Column purification is usually done for preps where there is a lot of the same DNA molecule- billions of copies of a PCR product after a reaction, or a large prep of bacteria, with hundreds of copies of a plasmid per cell. The kits are more expensive than the home-made procedures already described, and in the lab, we find that the yields are not huge – at least for some of the kits we’ve used. But the DNA that is isolated is usually very clean and we get sufficient yields for our purposes.

This procedure makes use of silica columns and high salt concentration, especially guanidinium salts, that promote DNA sticking to the silica beads in the column. The lysate is added to the column and metabolites, proteins, and other molecules flow through the column when it is centrifuged. When the lysate is added to the column, it is important to apply it carefully- you want to pipette the lysate slowly to the centre of the column, avoiding dripping it down the sides. When this happens, the solution mostly bypasses the beads and little or no DNA can bind to them. You also avoid touching or disrupting the beads of the column as this can also reduce yield. Several wash steps are performed with wash solutions containing ethanol, which keeps the DNA out of solution and stuck to the beads while washing away other, unbound substances. Finally the DNA is eluted in a low salt solution which dislodges it from the beads and collects it in a small volume in the collection tube. There is no use of organic solvents, so the method is relatively safe and there is no alcohol precipitation step. Unfortunately it is possible to accidentally “lose the pellet” with alcohol precipitation, especially if the pellet was small and fairly clear – difficult to see – to begin with. You can accidentally pipette it out of the tube and discard it.

In this procedure, each kit has slightly different instructions and these must be followed carefully, especially in the speeds at which you spin. There are given in g (rcf) because the users will be using a variety of different centrifuges.  The relationship between the rotations per minute (RPM) of the centrifuge and the force relative to g (gravity) depends on the radius of the rotor. Therefore you would need to check the table that comes with the centrifuge you have which shows the g values and the rpms that correspond to those values. These will be available in whatever lab you are working in. Here is the table that corresponds to the centrifuges we generally use when the lab is in person. Our centrifuges have a radius of 73 mm.

RPM to RCF conversion table for 73 mm rotor

RPM                RCF (g-force)
1000                82
2000                325
3000                735
4000                1310
5000                2040
6000                2940
7000                4000
8000                5220
9000                6610
10000              8160
11000              9880
12000              11750
13000              13800
14000              16000

B3-iv. Solid phase isolation (using magnetic beads)

In the solid phase isolation method very tiny beads (nanoparticles) of iron oxide, 20- 30 nm in diameter are used. These are manufactured for different purposes: to isolate nucleic acids from a crude lysate, or to isolate proteins, mRNA, mitochondrial DNA, and other components of a lysate. What determines the specificity of isolation is the coating on the beads and the components of the buffer used. For the isolation of nucleic acids, the beads are silica coated. In this respect they are much like the beads of the column purification method described above. Guanidinium is used in the buffer as well, to promote the binding of DNA to the beads.

Although they are called magnetic beads, the beads are not magnetic unless a magnetic field is applied. So when the beads are added to the lysate, they don’t clump together and DNA can interact with the silica coating. When a magnetic field is applied, the beads stick together, bringing the bound DNA with them. The other components of the lysate are removed and a series of wash steps ensures that only the beads and the bound DNA remain in the tube or microtitre plate. Finally the DNA is eluted from the beads using a low salt buffer, commonly TE.

The video below briefly describes the column purification method and the magnetic bead method. They are more expensive than the first two “home made” methods. They also share in common the affinity between DNA and a silica matrix under the right conditions. In the video, the magnetic bead method is shown using a single eppendorf tube. This is mainly because I cannot draw well in powerpoint. This method is frequently performed on large scale for multiple preps at once, using microtitre plates with 24, 48 or 96 wells.

(Click here to download the powerpoint slides presented in the video below)

B-4. Use of RNase

It is not possible to isolate only DNA (and no RNA) by these methods. So there is some RNase A (the most frequently used RNase) added to either the extraction buffer at the beginning of the procedure or the TE or other buffer that the DNA is dissolved in at the end of the procedure. The presence of contaminating RNA in some of the downstream treatments – such as PCR, sequencing, even restriction digest – can interfere with the procedures. It can also make the quantification of the prep inaccurate, which will be discussed in section D.

Using RNase in DNA preparation means having to carefully protect the RNA preps you may be making in the same lab. RNases are found in all cell types you are likely to work with and are on your body as well. In humans RNases are found in tears, mucous membranes and even in shed skin. It is a first line of protection against RNA viruses. This is why you must be careful in handling RNA preps.

RNases are surprisingly robust enzymes that are not easily inactivated. For example unlike most proteins, you can boil or autoclave RNases and when they return to normal temperature they regain their activity. Many labs set aside a special area for isolating RNA where no other work is done and conditions are kept extremely clean. The benches are wiped frequently and products manufactured to inactivate these enzymes may be sprayed on the bench. The RNA preps themselves may also contain RNase inhibitors to promote stability of the RNA.

Reagents used in the process of RNA isolation are generally made using DEPC treated water. Diethylpyrucarbonate (DEPC) is a carcinogen and very hazardous to work with. It is added to distilled, deionized water (ddH₂O) and it icleaves any proteins that might be found in the water. It reacts with amine, hydroxy and thiol groups of proteins. Although RNase can be boiled and will still be able to refold and regain its activity, if it is cut into pieces it cannot do so. DEPC is added to the water, given time to cleave any and all proteins in the sample and then it is autoclaved. When it is autoclaved, it degrades to water and CO₂ and thus is rendered harmless. Note that the DEPC is now gone from the water so careful handling is required when using it to make solutions to avoid introducing RNase into the solution. We used to do DEPC treatments ourselves but currently we purchase nuclease free water from chemical supply companies. This reduces the chance of our technicians being exposed to hazardous substances.

C. Principles of RNA isolation:

C-1. RNA Isolation Introduction

RNA isolation requires special care to avoid RNases in the prep as described above.

RNA is very much like DNA chemically, so we have to exploit the small differences in chemistry in order to isolate just RNA. We cannot use the nuclease method of removing unwanted DNA from a prep due to the expense. DNase is not efficient enough to remove all DNA from a prep of mixed RNA and DNA. There are two methods we can use that take advantage of chemical differences between RNA and DNA. The first is that RNA is most stable at a much lower pH than is DNA, and the other is its ability to interact with lithium ions.

C-2. TRIzol Method and Lithium Chloride Precipitation

In the first case, the TRIzol method, a modification of the phenol method already described except that we perform the extraction with acidified phenol (at pH about 4.5). Recall that when isolating DNA we keep the pH at 8.0 for good stability. At this low pH DNA will precipitate out of solution while RNA will remain soluble. We can exploit this difference to preferentially isolate RNA.

There is another RNA isolation method which uses lithium chloride in the isolation. Lithium cations will form an insoluble salt with RNA but not DNA. So in this case we precipitate RNA but not DNA from a cell lysate and then resuspend the RNA in nuclease free water. We don’t need to use TE. RNases do not use divalent cations as a cofactor and RNA does not particularly require a high pH.

This video briefly describes the two methods (Click here to download the powerpoint slides presented in the recording):

There are also kits that can be purchased for RNA isolation – these use column chromatography (or magnetic beads) and some are used for total RNA isolation while others allow the preferential isolation of mRNA. These are easy to use and follow a protocol fairly similar to the DNA isolation. But they are expensive. I mention expense frequently because it can often be an important consideration. If you are lucky enough to work in a very well funded lab you may be able to buy all the most expensive reagents, but many of us work in labs that have relatively small funding and so we always need to keep an eye on the price tag.

D. Quality and quantity of your sample:

Once we have isolated our nucleic acid samples, we need to know if we’ve succeeded in getting a clean prep with a good yield of DNA or RNA. There are two main ways we can determine both the quantity and quality of our DNA or RNA: spectrophotometry (we use the nanodrop machine in our lab), and gel electrophoresis. Ideally both should be used.

D-1. Spectrophotometer

The spectrophotometer beams light of a mixture of wavelengths through a sample and measures the difference between the light beamed at the sample and the light detected after it has passed through the sample. This measures how much light and of which wavelengths was absorbed by the sample. Different substances absorb light of different wavelengths. DNA and RNA absorb maximally at 260 nm wavelength. Proteins absorb maximally at 280 nm. So does phenol. Carbohydrates and guanidinium absorb maximally at 230nm. In assessing the purity of your RNA or DNA sample you need to know the absorbance properties of these, the most likely contaminants.

The amount of light absorbed at 260 nm, called A₂₆₀, is proportional to the amount of nucleic acid in the sample and the A₂₈₀ is proportional to the amount of protein in the sample and so on. We can use the A₂₆₀ itself to determine the concentration of nucleic acids in our sample and the ratio between the A₂₆₀ and A ₂₈₀ to estimate the purity of the sample. Recall that we don’t want a lot of protein in our sample so we’re hoping the ratio will be high (lots of nucleic acid and less protein).

Here is how the A₂₆₀ relates to nucleic acid concentration:

• For double stranded DNA, 1 absorbance unit corresponds to a concentration of 50 ng/μl
• For RNA, 1 absorbance unit corresponds to a concentration of 40 ng/μl
• For single stranded DNA, 1 absorbance unit corresponds to a concentration of 33 ng/μl.

In our lab we use a nanodrop spec which does all the work and calculations for you. It also uses just a tiny amount of the sample (0.5 to 2.0 μl). You first make a blank reading by pipetting 2 μl (we usually use 2 μl) of the buffer your sample is in, TE usually for a DNA prep and nuclease free water for an RNA prep. If you have used a column purification then you use elution buffer. The machine measures the absorbance of this buffer and then subtracts that from the absorbance of your sample. You ONLY want to measure the absorbance of the nucleic acids and proteins etc. in your sample and not that of the buffer. You blot away the liquid on the stage and then pipette 2μl of your sample onto it, and press measure – the machine reads the absorbance and tells you the concentration of your sample.

The nanodrop also reads the A₂₆₀, A₂₈₀, and A₂₃₀ all at the same time and displays the ratios: A₂₆₀/A₂₈₀ and A₂₆₀/A₂₃₀. Your job is just to interpret the ratios to determine whether your prep is clean enough to use.

Pure ds DNA in a buffer has A₂₆₀/A₂₈₀ of 1.85–1.88 (we generally say that 1.8 is ideal– there are many publications with slightly contradictory information). If the ratio is higher, it can indicate that there is RNA in the prep, while a lower ratio indicates protein or phenol contamination; if it is phenol you will be able to smell it in the prep when the tube is open. In determining the likely contaminant in your prep ensure that it was part of your preparation mechanism. For instance, if you used a column prep, with no phenol  extraction step, phenol will not be the reason for a ratio that is too low. Even if your A₂₆₀/A₂₈₀  ratio is not perfect, you may still be able to use your nucleic acid prep. In practice, we’ve found that a sample with a ratio of 1.6 and higher is pure enough for the procedures we will do in our lab.

Pure RNA has a ratio of around 2.1. (2.0 to 2.2 are considered “ideal”). A lower value will usually mean protein contamination, and thus potentially nucleases. A ratio of 1.8 and higher is generally acceptable.

The contaminants that absorb at 230 nm are numerous. They include chaotropic salts such as guanidine thiocyanate (GTC) and guanidine hydrochloride (GuHCl), EDTA, non-ionic detergents. Substances like polysaccharides also absorb at this wavelength and are most likely to be a problem in preps from plants. You can guess at the contaminant if you understand the buffers and solutions used in your prep. If you did not do an organic extraction, chloroform will not be a contaminant, for example.

Pure ds DNA has an A₂₆₀/A₂₃₀ of 2.3–2.4 and RNA has a ratio of 2.1–2.3. In our lab, we’ve seen great variation in the A₂₆₀/A₂₃₀ ratios and sometimes they are bizarrely high when we do a column plasmid prep. We’ve often just gone ahead and used the DNA and the procedures usually work out. If the ratios are much less than 1.8 indicating, for instance, guanidinium contamination, we might do an additional chloroform extraction to clean up the DNA or RNA a little more before using it. To do that, we would add more buffer to the prep to increase the volume to 500 μl, do a 1V chloroform extraction, add 0.1 V 3M sodium acetate to the sample, and then do an ethanol precipitation. We would resuspend the pellet in quite a small volume of nuclease free water (for RNA) or TE (for DNA) because sometimes you lose some of the nucleic acids when you re-extract and re-precipitate. Ratios around 2.0 are ideal, but 1.8 and above is usable.

For our purposes we consider that for DNA the ideal A260/A280 ratio is 1.8 and the ideal A260/A230 is 2.0 (a little higher is not a problem). And we will consider that for RNA the ideal for both ratios is 2.0. Different sources list the ideals slightly differently, but we will use these in our lab.

In a DNA prep, if we have RNA also present, it will be measured by the spectrophotometer along with the DNA. This is because both nucleic acids absorb maximally at 260nm. This can lead to a huge overestimation of the DNA concentration, perhaps up to a 10-fold overestimation because you will be measuring all the DNA and RNA in the sample. This can be problematic in downstream applications, as you will put too little DNA into reactions.  The use of RNAse should deal with this problem. RNase does not work instantly. In the lab, your DNA prep has RNase added to the TE buffer but some RNA will be left in the tube when you measure the concentration. By the next week, the RNA should be degraded.

(Click here to download the powerpoint slides presented in the video below)

D-2. Gel electrophoresis

Our spec readings might indicate that the DNA is pure and a good yield. But these are only two of the three criteria listed way at the beginning of the chapter. We also want the DNA to be relatively intact (unbroken). The spec cannot give us this information. We need to run a gel. Gel electrophoresis separates DNA pieces by size (when we talk about the size of a band we are talking about the length in base pairs). Sometimes gels are used to purify DNA- we may discuss this a bit later. For now, we will talk about its use for confirming that our DNA is relatively undamaged. Agarose gel electrophoresis uses an electrical field to move DNA through an agarose gel. The agarose is melted in a buffer, such as TAE (tris-acetate-EDTA) or TBE (tris-borate-EDTA). You know two of the components of those buffers and what they are for. The acetic acid or boric acid help with the buffering – to keep the pH in range.

An indicator such as ethidium bromide, gel red or sybr safe is added to the agarose once it is melted. These substances interact with DNA or RNA and when subjected to light of a particular wavelength they fluoresce. This will allows us to see where the DNA is in the gel after we’ve run it. Ethidium bromide is a known mutagen and we don’t use it as often as we used to. Gel red is a safer alternative although in the lab we handle it with caution to be on the safe side. Both ethidium bromide and gel red fluoresce when exposed to UV light.

The gel is cast in a tray and wells for the DNA are made in one end. The agarose forms a kind of mesh or matrix and when the apparatus is turned on, the DNA moves from the wells at the negative end, through the pores in the gel, toward the positive pole. Shorter DNA fragments migrate through the gel more quickly than longer ones. Thus, you can determine the approximate length of a DNA fragment based on the distance it travels through the gel. You run a ladder alongside the DNA. This commercially prepared ladder has DNA in known sizes so that you can compare your bands to the ladder to determine the approximate size of the fragments.

This link shows the basic procedure of making and running an agarose gel:

https://www.addgene.org/protocols/gel-electrophoresis/

If the case of a genomic DNA prep, we would run a small amount of the prep in the gel and we would hope to see one quite large band. Good quality genomic DNA will run as if it is about 20–40 kb in length. DNA that is damaged will have some DNA up near the top of the gel- the large pieces of 20–40 kb, with a smear down the lane that gets fainter the further you go. This is what DNA looks like when it has been randomly broken and there are pieces of all different sizes in the prep.

RNA is single stranded but still undergoes complementary base pairing. As a result it can form secondary structure. That means that the molecule can fold back on itself and base pair in some locations, forming many loops and stems. A molecule that has folded up this way won’t run through a gel based on length- if it is tightly compacted, it may run much faster than a linear molecule of the same length. And if there are many big loops, it may have more difficulty navigating through the agarose and move more slowly than a linear molecule of the same length. So when we run RNA on a gel, we run a denaturing gel. It is run under conditions that interfere with the hydrogen bonds of base pairing and keep the molecule linear. RNA gels usually contain formaldehyde and formamide. They smell bad and the fumes that are produced while the gel is running are not good for you. We run these gels in the fume hood.

It is also impossible to see all of the RNA in your prep by viewing the gel. That is because the RNA molecules are all different sizes- even transcripts from the same gene will vary a little in size because the poly A tail won’t be exactly the same length for every RNA. Also, mRNA is a very small proportion of the total RNA that is isolated. The majority is ribosomal RNA. A mammalian cell can have up to 10,000,000 ribosomes in it. That is a huge amount of rRNA! mRNA is about 1-5% of the total RNA isolated, and again, those RNAs are all different sizes. So when we are assessing the quality of the prep, we can only see the ribosomal RNAs. We use them as an indicator of the quality of all the RNA. If the rRNAs are intact, we can assume the mRNAs you are interested in are also intact.

The video in the link below briefly describes the gel procedures for RNA and DNA and has examples from the Mattsson lab that show good quality DNA and RNA (Click here to download the powerpoint slides presented in the recording).

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