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Basic step by step methods for PCR & Gel electrophoresis class

Basic step by step methods for PCR & Gel electrophoresis class


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I'm teaching a class how to do PCR & gel electrophoresis soon and would like it if you could check through my basic step by step instructions - it's a while since I've done one. Is there anything there that is wrong, missing, or should be excluded in the following?

A) Collect all ingredients from the refrigerator and keep cool (on ice). Leave the TAQ in the freezer until required.

B) Make master mix by adding all ingredients (ddH20, MgCl2, PCR Buffer, F Primer, R Primer, dNTP) (excl. TAQ)

W) Take the gel to the UV camera and take a picture. Large DNA molecules move through the gel slowest so will be nearest to the wells.

(The students will use primers they designed to sex 6 DNA samples from 3 species of birds, males should produce 1 band & females 2 bands in the gel)


for the full version see the answer below


I've run a test following this protocol to the letter…

PCR step by step:

  1. Collect all ingredients (excluding TAQ) from the refrigerator and keep cool (on ice). Leave the TAQ in the freezer until required.
  2. Make master mix by adding all ingredients (excluding TAQ) using fresh pipette tips for each ingredient and using the volume specified in the PCR reaction mix recipe.
  3. Vortex & lightly spin down on a centrifuge.
  4. Add the required amount of TAQ to master mix (insert in to the liquid) and mix using the pipette to draw some liquid up and down.
  5. Add master mixes to labeled PCR tubes in the required volumes as specified by the protocol.
  6. Add DNA to PCR tubes (using fresh pipette tips for each DNA sample) again using the volume specified in the protocol. Label your tubes carefully (and on both the lid and tube).
  7. Lightly spin down the PCR tubes on a centrifuge.
  8. Run PCR in the thermocycler according to your protocol.
  9. Collect samples from thermocycler and place in fridge until needed.

Gel Electrophoresis step by step:

  1. Make 1xTAE (enough to make the agarose gel and to place in the electrophoresis apparatus).
  2. Make an agarose gel (about 5 mm thick) by melting agarose and 1xTAE in the microwave; allow the liquid to cool a little before adding it to the mold.
  3. Once the gel is cool, place it in the electrophoresis apparatus, cover it with 1xTAE (just covering the gel) and then remove the comb.
  4. Collect the PCR products from the fridge/thermocycler.
  5. Make spots of loading dye on parafilm. The loading dye is premixed with GelRed so your samples show under UV light.
  6. Add one sample to a spot, mix it by drawing the liquid in and out of the pipette tip, and then carefully add to a well in the agarose gel. Take care not to puncture the gel or spill any of the liquid out of its well. Discard the pipette tip.
  7. Repeat the previous step for each sample and for a negative control (master mix and milli-q water instead of DNA). Use a fresh tip for every sample.
  8. Load PCR Ladder, also mixed with loading dye, in to a well.
  9. Connect the electrophoresis apparatus to a power supply, with negative electrode (black) nearest to the wells containing the samples.
  10. Set the power to run 80-120 volts.
  11. Wait until about a finger-width gap appears between the blue and purple bands (approximately 20-30 minutes).
  12. Disconnect the power and collect the gel carefully from the liquid.
  13. Take the gel to the UV camera and take a picture. Large DNA molecules move through the gel slowest so will be nearest to the wells.

And got a perfect result…

The order is Control, female, male, female, male, female, male, ladder. Males have one band, females have two. The final pair didn't show too well on the picture but were clear enough on the screen in the camera room. Thanks for the tips everyone! Sorry the picture is low quality.

Feel free to recreate these guidelines but remember to credit the Biology Stackexchange Community!


  • The first step to study or work with nucleic acids includes the isolation or extraction of DNA or RNA from cells.
  • Gel electrophoresis depends on the negatively-charged ions present on nucleic acids at neutral or basic pH to separate molecules on the basis of size.
  • Specific regions of DNA can be amplified through the use of polymerase chain reaction for further analysis.
  • Southern blotting involves the transfer of DNA to a nylon membrane, while northern blotting is the transfer of RNA to a nylon membrane these techniques allow samples to be probed for the presence of certain sequences.
  • denaturation: the change of folding structure of a protein (and thus of physical properties) caused by heating, changes in pH, or exposure to certain chemicals
  • electrophoresis: a method for the separation and analysis of large molecules, such as proteins or nucleic acids, by migrating a colloidal solution of them through a gel under the influence of an electric field
  • polymerase chain reaction: a technique in molecular biology for creating multiple copies of DNA from a sample

The broad steps involved in a common DNA gel electrophoresis protocol:

1. Preparing the samples for running

2. An agarose TAE gel solution is prepared

TAE buffer provides a source of ions for setting up the electric field during electrophoresis. The weight-to-volume concentration of agarose in TAE buffer is used to prepare the solution. For example, if a 1% agarose gel is required, 1g of agarose is added to 100mL of TAE. The agarose percentage used is determined by how big or small the DNA is expected to be. If one is looking at separating a pool of smaller size DNA bands (<500bp), a higher percentage agarose gel (>1%) is prepared. The higher percentage of agarose creates a denser sieve to increase the separation of small DNA length differences. The agarose-TAE solution is heated to dissolve the agarose.

3. Casting the gel

The agarose TAE solution is poured into a casting tray that, once the gel solution has cooled down and solidified, creates a gel slab with a row of wells at the top.

4. Setting up the electrophoresis chamber

The solid gel is placed into a chamber filled with TAE buffer. The gel is positioned so that the chamber wells are closest to the negative electrode of the chamber.

5. Loading the gel

The gel chamber wells are loaded with the DNA samples and usually, a DNA ladder is also loaded as reference for sizes.

6. Electrophoresis

The negative and positive leads are connected to the chamber and to a power supply where the voltage is set. Turning on the power supply sets up the electric field and the negatively charged DNA samples will start to migrate through the gel and away from the negative electrode towards the positive.

7. Stopping electrophoresis and visualizing the DNA

Once the blue dye in the DNA samples has migrated through the gel far enough, the power supply is turned off and the gel is removed and placed into an ethidium bromide solution. Ethidium bromide intercalates between DNA and is visible in UV light. Sometimes ethidium bromide is added directly to the agarose gel solution in step 2. The ethidium bromide stained gel is then exposed to UV light and a picture is taken. DNA bands are visualized in from each lane corresponding to a chamber well. The DNA ladder that was loaded is also visualized and the length of the DNA bands can be estimated. An example is given in the figure below.


Touchdown PCR

Another approach to promoting specificity is to modify the PCR cycling parameters. In touchdown PCR, the annealing temperature of the first few cycles is set to be a few degrees higher than the highest melting temperature (Tm) of the primers [1,2]. Higher temperatures help destabilize the formation of primer-dimers and nonspecific primer-template complexes, thus minimizing undesirable amplification. As such, higher annealing temperatures reduce nonspecific PCR products and promote specific amplification at the start of PCR (learn more about PCR annealing step).

While preventing primer-dimers and nonspecific primer binding, the higher annealing temperatures may result in lower PCR yield due to increased dissociation of primers from their intended target. To overcome this challenge, the annealing temperature is often decreased 1°C at every cycle of the initial few cycles to produce a sufficient yield of the desired amplicon. Once the annealing temperature reaches, or “touches down”, at the optimal temperature (usually 3–5°C lower than the lowest primer Tm), it is maintained throughout the remaining cycles for primer annealing. In this manner, desired PCR products are selectively increased with little or no amplification of nonspecific targets over the course of PCR (Figure 2).

Figure 2. Touchdown PCR. The method promotes specificity (yellow curve) by starting with a higher-than-optimal annealing temperature, which is then gradually lowered (black line) as cycling continues until the optimal annealing temperature is reached. The yield of the intended amplicon (green curve) accumulates considerably with the optimized annealing temperature.


GEL ELECTROPHORESIS TECHNIQUE

The term “electrophoresis” refers to the movement of a solid particle (e.g. nucleic acids) through a polymer matrix or gel under the influence of electric field. Electrophoresis is a molecular biology technique that is used to separate nucleic acid molecules and other macromolecules mainly on the basis of their charge to mass ratio as they migrate through a gel in an electric field. Gel electrophoresis technique is a molecular biology technique that is used to separate nucleic acid molecules (DNA and RNA) according to their sizes and conformation or charges. It is generally used in the molecular biology laboratory for the separation and purification of nucleic acid fragments. The process occurs in an electrophoretic tank or chamber laden with specialized type of gel (e.g. agarose) through which controlled electric charge is allowed to pass through (Figure 1).

Figure 1. Agarose gel electrophoresis apparatus (arrowhead) for the separation of DNA fragments and protein molecules. DNA is negatively charged, and when an electric current is applied during electrophoresis, the DNA molecules will move from the cathode (coloured black) end towards the anode (coloured red) end of the electrophoresis tank.Photo courtesy: https://www.microbiologyclass.com

A dye (e.g. ethidium bromide) is added to the gel so that the nucleic acid fragments can be visualized under ultraviolet (UV) light. The size range of nucleic acid fragments that can be separated using agarose gel is usually in the range of 0.2 kb to 20 kb. Gel electrophoresis is an important technique that is used to analyze the products of a PCR reaction. It allows separated fragments of nucleic acid molecules or protein molecule from a given organism or cell to form characteristic pattern of bands known as fingerprints when they are passed through a gel under the influence of electric charges. Other examples of gel used in gel electrophoresis technique apart from agarose include polyacrylamide or acrylamide. Agarose gel is used for the separation of DNA fragments while acrylamide or polyacrylamide is the gel matrix used for the separation of protein molecules. While agarose is used in horizontal gel apparatus as shown in Figure 1, polyacrylamide is mainly used in vertical gel apparatus applied in advanced separation techniques such as blotting. Thegel electrophoresis apparatus is used for separating nucleic acids based on their mobility under the influence of an electric field in an electrophoresis tank.

Gel electrophoresis is the technique of separating charged molecules such as DNA in an electric field. Fragments of separated nucleic acid molecules move through gel in electric fields according to their different sizes. This serves as the basis for the utilization of electrophoresis to identify the individual fragments of a particular DNA. It is worthy of note that after isolating a piece of DNA from an organism, and cutting same into different fragments using restriction enzymes there is need to study the individual fragments. This can only be made possible through electrophoresis which gives a detailed analysis of each fragment of the nucleic acid. Deoxyribonucleic acid (DNA) is a negatively charged nucleic acid molecule because of its phosphate groups.

When the separated DNA fragments is placed in a gel and allowed to move through an electric field (with positive and negative ends), the DNA molecule tends to move towards the positive terminus (i.e. the anode) of the electric charge than the negative terminus (or cathode) because of its notable negative charge. Smaller molecules of DNA migrate through the gel faster than the larger molecules because of the sieving nature of the gel used for gel electrophoresis technique. The separated DNA fragment is allowed to run for a specific amount of time, and the DNA fragments are visualized under UV light after the addition of ethidium bromide (EtBr) which makes the bands visible. DNA is a colourless macromolecule, and EtBr is used in gel electrophoresis to make the different bands of the nucleic acid (DNA) visible. EtBr is mutagenic and can cause cancer. Thus it should always be handled with utmost care and always with gloved hands.

The EtBr intercalate between the nitrogenous bases of the double stranded DNA molecule, and this causes the DNA molecule to fluoresce or produce an orange colour when the gel carrying the DNA fragments is photographed or illuminated with UV light. Electrophoretic technique is the most versatile method of analyzing, identifying and purifying the fragments of nucleic acid molecules (DNA and RNA) and proteins. It is unique because it separates macromolecules according to their sizes and charges. Agarose and polyacrylamide are the two notable gels used in electrophoresis experiment. While agarose gel is used in most simple electrophoresis techniques (e.g. separation of nucleic acid molecules), polyacrylamide gel is mainly used in advanced electrophoresis such as those that has to do with protein separation. Several electrophoresis techniques are available and they include agarose gel electrophoresis, pulse field gel electrophoresis (PFGE) and polyacrylamide gel electrophoresis (PAGE).

STEPS OF PERFORMING GEL ELECTROPHORESIS

Gel electrophoresis is one type of electrophoresis technique, and its procedure shall be highlighted in this unit. The following materials and steps are employed in gel electrophoresis technique:

  • Agarose gel is used for performing gel electrophoresis in the microbiology or molecular biology laboratory. It is noteworthy that the agar powder used for gel electrophoresis is different from the powdered agar used for the preparation of routine culture media plates for microbial cultivation. In gel electrophoresis, agarosegel powder is used to prepare the gel. The agarose gel is prepared by mixing a particular amount of agarose powder (e.g. 1.5 %) in a deionized water or buffer solution such as tris boric acid EDTA (TBE) buffer. Agarose gel could be made with varying concentrations of agarose ranging between 0.6 % – 3 %. This usually depends on the size of the nucleic acid fragments the researcher wishes to resolve or separate. Larger fragments of nucleic acids are separated or resolved better in a gel with a lower percentage of agarose while smaller nucleic acid fragments are separated better in a gel with a higher percentage of agarose.

To prepare 1.5 % agarose gel for example, measure out 3 g of agarose powder and dissolve same in 250 ml buffer or deionized water in a conical flask as aforesaid. Stir the mixture properly to break up all clumps. Heat the mixture by boiling at a particular temperature and time in a microwave oven until the solution becomes clear as water. A Bunsen burner flame could also be improvised for heating the agarose solution in places where microwave oven is unavailable. The agarose gel solution should be heated until a homogenous solution is formed. After heating, the homogenate gel should be allowed to cool to about 60 o C before pouring gel onto the gel casting apparatus or slab. Agarose, a white powder and the buffer solution are the two basic components of an agarose gel and both needed to be heated sufficiently to make the gel required to run the gel electrophoresis technique. After cooling, the molten gel should be poured into the gel casting chamber with the toothed comb in place.

  • A toothed comb is used to form wells known as sample wells in the agarose gel (Figure 2). Samples for gel electrophoresis analysis are individually inoculated or dispensed into each of the toothed wells using micropipette (Figure 3). Multiple pipette tips (Figure 4) also exist for multiple analyses during molecular biology experimentation.
Figure 2. Illustration of toothed combs of various sizes used for making sample wells in agarose gel. The toothed combs exist in different sizes, and the type used is mainly dependent on the number of samples the researcher wishes to run. The toothed comb is significant in gel electrophoresis because it creates cavities generally known as wells in the agarose gel and it is in these wells that the nucleic acid samples are pipetted into. It is noteworthy that the toothed comb is inserted into the electrophoretic gel chamber before the molten agarose gel is poured and allowed to gel. Photo courtesy: https://www.microbiologyclass.com Figure 3. Single tip micropipettes. Photo courtesy: https://www.microbiologyclass.com

The toothed comb should be placed into the gel casting apparatus or tray prior to pouring of the gel so that the wells will be formed appropriately. The toothed comb is removed prior to the insertion of the DNA samples into the wells. The number of wells or holes formed is usually dependent on the number of samples or organisms to be analyzed. And the type of toothed comb used in agarose gel experimentation varies. The comb should be removed or pulled out in an upward manner from the casted gel after its solidification.

Figure 4. Multiple tips micropipette with tip container for pipette tips (arrowhead). Photo courtesy: https://www.microbiologyclass.com
  • Dispense the cooled homogenous solution into the gel casting apparatus or tray (Figure 5). The pouring should be done slowly, and all air bubbles formed during the pouring should be removed using a disposable pipette. Air bubbles could generally affect the shape of the wells if allowed to settle around the comb. The air bubbles could also prevent the free flow of the electric current through the gel.
Figure 5. Gel casting apparatus. The agarose gel slab is formed in the gel casting apparatus or tray, and then transferred to the electrophoresis tank for further analysis. It is critical to ensure that the gel casting apparatus or tray is placed on a horizontal surface before pouring the gel so that the gel formed will be uniform. The toothed comb should be inserted before pouring the molten agarose gel into the gel casting apparatus. Photo courtesy: https://www.microbiologyclass.com
  • The poured gel is allowed in the gel casting apparatus for some minutes (e.g. 20 min) so that it will set or gel to form agarose gel slab. The gel casting apparatus gives the poured gel its characteristic horizontal shape required for agarose gel electrophoresis technique. Once cooled and gelled, the gel is now ready for agarose gel electrophoresis experimentation. It is then inserted into the electrophoretic matrix or chamber in which a buffered solution is also added to. In practice, the agarose gel slab is submerged in the buffered solution in the electrophoretic tank. The buffer solution can also be poured in the electrophoretic chamber after adding the gel slab
  • The individual DNA samples are pipetted into the sample wells created in the agarose gel by the comb (Figure 6). Ensure that the pipette tip is changed for each sample to be pipetted. A DNA fragment or ladder (with known or standard size) is added in one of the wells (usually the first well) and the ladder is used to compare the separated DNA fragments (with unknown sizes).
Figure 6. Illustration of loading samples onto wells created on an agarose gel. When loading the samples, the pipette should be held at an angle in order to avoid puncturing the wells. Punctured wells will cause the DNA samples to leak out into the buffer and/or electrophoresis tank and this will cause contamination of the electrophoresis process. https://www.microbiologyclass.com

In some agarose gel experimentation, ethidium bromide (EtBr) solution is added alongside the DNA solution to be analyzed. However, the EtBr is usually added to the prepared gel after cooling and before pouring onto the gel electrophoresis tank.

EtBr act as a chemical staining agent which helps to visualize the DNA bands or fragments after the electrophoresis experimentation. (EtBr is a dye that binds to DNA and clearly marks the position of the individual DNA fragments). In some agarose gel experimentation, the staining dye (in this case EtBr) is not added alongside the DNA solution to be electrophoresed. But it is added prior to or after the electrophoresis analysis since its main function is to aid the visualization of the DNA fragments. Note: EtBr is mutagenic or carcinogenic in nature, and thus should be handled with care. SYBR Green, a nucleic acid gel stain is another staining agent that could be used in gel electrophoresis technique to visualize separated nucleic acid fragments. However, EtBr solution is the most commonly used dye in gel electrophoresis experimentations. It is critical that the researcher wears gloves when handling EtBr since the dye is a mutagen and could easily be absorbed by the skin to cause health problems in the individual.

  • An electric current (e.g. 100 volts) is passed through the gel and the process is allowed to run for the appropriate time limit. DNA, a negatively charged molecule moves from the negatively charged electrode (cathode) towards the anode (positive electrode). The DNA moves through the gel matrix. Smaller DNA molecules move faster than the larger DNA molecules. The electric charge or current is switched off once the electrophoresis process is completed.
  • Separated DNA fragments is visualized under UV light and photographed after soaking the gel slab in EtBr or any other staining dye as may be available.

Further reading

Cooper G.M and Hausman R.E (2004). The cell: A Molecular Approach. Third edition. ASM Press.

Das H.K (2010). Textbook of Biotechnology. Fourth edition. Wiley edition. Wiley India Pvt, Ltd, New Delhi, India.

Davis J.M (2002). Basic Cell Culture, A Practical Approach. Oxford University Press, Oxford, UK.

Mather J and Barnes D (1998). Animal cell culture methods, Methods in cell biology. 2 rd eds, Academic press, San Diego.

Noguchi P (2003). Risks and benefits of gene therapy. N Engl J Med, 348:193-194.

Sambrook, J., Russell, D.W. (2001). Molecular Cloning: a Laboratory Manual, 3rd edn. Cold Spring Harbor Laboratory Press, New York.

Tamarin Robert H (2002). Principles of Genetics. Seventh edition. Tata McGraw-Hill Publishing Co Ltd, Delhi.


Destroying Membranes Within the Cell

The cell's plasma membrane is made of phospholipid bilayers they are made of fat. To disrupt them, that mesh of fat molecules is broken up with soap. The structure of soap is very similar to that of fat and grease.

A soap molecule has two parts: a head and a tail. The head is polar and is attracted to water while the tail is non-polar and is attracted to oil and fat. When soap molecules are in water, they group themselves into micelles — a roughly spherical structure in which all the polar heads point outwards (in contact with water) and all the non-polar tails point inwards at the center of the sphere (away from the water). They can effectively trap the fat molecule inside the micelle and dissolve the cell membranes. How does this micelle break down the phospholipid bilayer? The molecules in the phospholipid bilayer (Figure 5) also contain molecules that are made up of a hydrophobic head and a hydrophilic tail. The soap molecules orient themselves so that their head associates with the tail of the phospholipid bilayer. In this way, the soap is able to break up the bilayer molecule by molecule.


Extracting DNA from Cells

To perform DNA fingerprinting, you must first have a DNA sample! In order to procure this, a sample containing genetic material must be treated with different chemicals. Common sample types used today include blood and cheek swabs.

These samples must be treated with a series of chemicals to break open cell membranes, expose the DNA sample, and remove unwanted components – such as lipids and proteins – until relatively pure DNA emerges.

PCR Amplification (Optional)

If the amount of DNA in a sample is small, scientists may wish to perform PCR – Polymerase Chain Reaction – amplification of the sample.

PCR is an ingenious technology which essentially mimics the process of DNA replication carried out by cells. Nucleotides and DNA polymerase enzymes are added, along with “primer” pieces of DNA which will bind to the sample DNA and give the polymerases a starting point.

PCR “cycles” can be repeated until the sample DNA has been copied many times in the lab if necessary.

Treatment with Restriction Enzymes

The best markers for use in quick and easy DNA profiling are those which can be reliably identified using common restriction enzymes, but which vary greatly between individuals.

For this purpose, scientists use repeat sequences – portions of DNA that have the same sequence so they can be identified by the same restriction enzymes, but which repeat a different number of times in different people. Types of repeats used in DNA profiling include Variable Number Tandem Repeats (VNTRs), especially short tandem repeats (STRs), which are also referred to by scientists as “microsatellites” or “minisatellites.”

Once sufficient DNA has been isolated and amplified, if necessary, it must be cut with restriction enzymes to isolate the VNTRs. Restriction enzymes are enzymes that attach to specific DNA sequences and create breaks in the DNA strands.

In genetic engineering, DNA is cut up with restriction enzymes and then “sewn” back together by ligases to create new, recombinant DNA sequences. In DNA profiling, however, only the cutting part is needed. Once the DNA has been cut to isolate the VNTRs, it’s time to run the resulting DNA fragments on a gel to see how long they are!

Gel Electrophoresis

Gel electrophoresis is a brilliant technology that separates molecules by size. The “gel” in question is a material that molecules can pass through, but only at a slow speed.

Just as air resistance slows a big truck more than it does a motorcycle, the resistance offered by the electrophoresis gel slows large molecules down more than small ones. The effect of the gel is so precise that scientists can tell exactly how big a molecule is by seeing how far it moves within a given gel in a set amount of time.

In this case, measuring the size of the DNA fragments from the sample that has been treated with a restriction enzyme will tell scientists how many copies of each VTNR repeat the sample DNA contains.

It’s called “electrophoresis” because, to make the molecules move through the gel, an electrical current is applied. Because the sugar-phosphate backbone of the DNA has a negative electrical charge, the electrical current tugs the DNA along with it through the gel.

By looking at how many DNA fragments the restriction enzymes produced and the sizes of these fragments, the scientists can “fingerprint” the DNA donor.

Transfer onto Southern Blot

Now that the DNA fragments have been separated by size, they must be transferred to a medium where scientists can “read” and record the results of the electrophoresis.

To do this, scientists treat the gel with a weak acid, which breaks up the DNA fragments into individual nucleic acids that will more easily rub off onto paper. They then “blot” the DNA fragments onto nitrocellulose paper, which fixes them in place.

Treatment with Radioactive Probe

Now that the DNA is fixed onto the blotting paper, it is treated with a special probe chemical that sticks to the desired DNA fragments. This chemical is radioactive, which means that it will create a visible record when exposed to X-ray paper.

This method of blotting DNA fragments onto nitrocellulose paper and then treating it with a radioactive probe was discovered by a scientist name Ed Southern – hence the name “Southern blot.”

Amusingly, the fact that the Southern blot is named after a scientist and not the direction “south” did not stop scientists from naming similar methods “northern” and “western” blots in honor of the Southern blot.

X-Ray Film Exposure

The last step of the process is to turn the information from the DNA fragments into a visible record. This is done by exposing the blotting paper, with its radioactive DNA bands, to X-ray film.

X-ray film is “developed” by radiation, just like camera film is developed by visible light, resulting in a visual record of the pattern produced by the person’s DNA “fingerprint.”

To ensure a clear imprint, scientists often leave the X-ray film exposed to the weakly radioactive Southern blot paper for a day or more.

Once the image has been developed and fixed to prevent further light exposure from changing the image, this “fingerprint” can be used to determine if two DNA samples are the same or similar!


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Steps of p olymerase chain reaction-PCR

To perform PCR, the extracted sample (which contains target DNA template) is added to a tube containing primers, free nucleotides (dNTPs), and Taq polymerase. The PCR mixture is placed in a PCR machine. PCR machine increases and decreases the temperature of the PCR mixture in automatic, programmed steps which generates copies of the target sequence exponentially.

Polymerase Chain Reaction (PCR) has three major steps.

  1. Denaturation (strand separation): The separation of the two hydrogen-bonded complementary chains of DNA into a pair of single-stranded polynucleotide molecules by a process of heating (94°C to 96°C)
  2. Annealing (primer binding): The temperature is lowered (45-60 °C) so the primers can attach themselves to the single-stranded DNA strands.
  3. Extension (synthesis of new DNA): It starts at the annealed primer and works its way along the DNA strand (72°C).

Once the first round is completed, the process is repeated by cycling back to the first reaction temperature and the next round of denaturation, annealing, and extension is started (an automatic process in thermocycler). This 3 steps temperature cycle is repeated approximately 30 times which results in exponential amplification of the target gene sequence.

Animation

Detection of PCR products

  1. It allows visualization of the PCR product
  2. It provides specificity by ensuring that the amplicon is the target sequence of interest and not the result of non-specific amplification.

Apart from DNA based hybridization method, sometimes a simple gel electrophoresis method is sufficient to confirm the presence of specific amplicons.


PCR: The Basic Theory

The capacity of the technique called the polymerase chain reaction (PCR) to amplify many million-fold any known DNA fragment from a complex mixture in a short time has revolutionized all areas of the life sciences, making it one of the most widely used molecular techniques in use today.

Quite simply, PCR is a laboratory technique used to makes a huge number of copies of a piece of DNA. In other words it is a way of amplifying DNA or increasing the amount of a specific piece of DNA. The core principle of PCR is the use of an enzyme called DNA polymerase to make a copy of a DNA strand. Normally DNA exists as a double strand, but the enzyme can only work on a single strand. Therefore it is first necessary to separate the strands of DNA. This is done by applying heat.

Applying heat to DNA denatures the double strand to single strands. In the first PCR step the double strand melts to single stranded DNA, a step known as denaturation. The enzyme can work on the single strand to make a copy. However, the enzyme needs a small region of double strand to get started. So it is necessary to add a short piece of single strand DNA, called the primer , that binds specifically to a particular place on the single strand molecule. This binding, or annealing, is achieved by cooling the PCR mixture again. This step in the PCR process is called annealing.

Then the DNA polymerase enzyme gets to work and copies the single strand molecule, starting at the bound primer region. This final step, extension, is so-called because the DNA extends the DNA from the annealed primer and makes a complementary copy of the single strand. In practice, two different primers are used. A forward primer that binds to one strand and a reverse primer that binds to the opposite strand. The choice of primers is important as each primer binds the DNA in a specific place. The only DNA that is copied is the region between the forward primer binding location and the reverse primer binding location.

Both single strands of the original double-stranded DNA molecule are copied. Therefore the result is two double stranded molecules each identical to the original double-stranded DNA fragment.

The three PCR steps are repeated for around 30 or 40 cycles. Each cycle doubles the number of double stranded DNA molecules.

Normally PCR is performed on a machine called a PCR thermal cycler or PCR machine. The PCR thermal cycler rapidly heats and cools the PCR reaction mixture thus allowing the denaturation, annealing and extension to occur.


Basic step by step methods for PCR & Gel electrophoresis class - Biology

The Polymerase chain reaction (PCR) is a ubiquitous technique utilized extensively for diagnostic purposes and molecular biology research. PCR is the in vitro amplification of specific nucleic acid (NA) sequences by a DNA Polymerase enzyme. The PCR technique was transformed by Kary Mullis in 1983 (who died in August 2019 at the age of 74), when he expanded the use of a heat-stable polymerase with temperature cycling [2-4]. The universal utility of PCR is that it amplifies small quantities of target nucleic acid sequences, yielding an amount of product that is detectable by downstream methods, such as visualization of NA on an agarose gel. This is due to the exponential amplification of the sequence and the resulting millions of copies of the original template.

PCR reactions amplify target nucleic acid sequences via the use of a DNA Polymerase, primers, and nucleotides. The template for a PCR reaction may be any nucleic acid sequence of interest, and the NA source may be DNA, RNA, or cDNA. Primers are short sequences of nucleotides synthesized in vitro. They are designed to anneal to opposite strands of a specific NA template target and usually are between 15-40 bases long. Primers ideally lack secondary structure and are not complementary to each other, to prevent primer-dimer formation. A variety of DNA Polymerases have been utilized for PCR, but the thermostable Taq DNA Polymerase is probably the most widely used. This enzyme adds the deoxyribonucleoside triphosphates (dNTPs or nucleotides) onto the ends of the primers to extend the NA based on the template NA sequence.

The PCR reaction mixture is temperature cycled, typically 20-40 times. Denaturation of the NA template sequence is achieved at 95ºC. The temperature is cooled to 37-60ºC to anneal primers to the target sequence. Extension of the primers with nucleotides by the DNA Polymerase is achieved at temperatures ranging from 60-72ºC. Conventional cycling conditions are 95ºC for 5 minutes initially to denature all template NA, followed by 2-40 repeats of 95ºC for 30 sec, 60ºC for 30s and 72ºC for 1 minute. The time spent at each temperature can be optimized for specific assays. For instance, the amplification of very short target sequences requires much shorter incubations at each temperature than that of very large target sequences. Each round of temperature cycling results in two times more target sequence than the prior round. This leads to the exponential amplification of the original template, often resulting in millions or billions of copies of the original NA target.

The basic PCR reaction occurs in three phases. The exponential phase is the period in which exact doubling of a nucleic acid product occurs every cycle. Real-time PCR detection is carried out during this exponential phase. The linear phase occurs as the reaction is slowing due to the consumption of the reagents and the degradation of the products. The final stage is the plateau phase, which occurs when the reaction has stopped, and no additional amplicon is being generated. This is the point at which the PCR reaction product is analyzed via gel electrophoresis for conventional PCR reactions.

Downstream detection of the PCR product is done in many ways. A common method of visualization is via agarose gel electrophoresis. This involves separating NA fragments via electrophoresis and staining the NA with an intercalating dye such as ethidium bromide or SybrSafe and subsequent detection using a UV light source and imaging system (Figure 2).

Real-time PCR uses specialized thermocyclers that detect the fluorescent signal in each well. This signal is indicative of the amount of double-stranded NA within the reaction tube or well. This signal, in relative fluorescent units, is plotted by the thermocycler software versus cycle number (Figure 3).

Multitudes of research, clinical and forensic applications of PCR exist. In molecular biology research, PCR is often used for genetic engineering, DNA sequencing, and gene expression analyses. In clinical laboratories, PCR is crucial in the detection of infectious disease. Forensic applications of PCR include paternity testing and DNA fingerprinting. These tests benefit from the exquisite sensitivity of PCR because target sequences can be detected from a single human hair. Table 1 lists the major applications and references to manuscripts describing them.

applicationsreferences
detection of foodborne pathogens [5, 6]
diagnosis of infectious disease [7]
genotyping [8]
human genetic mutation detection [9, 10]
expression profiling of microRNA [11, 12]
gene expression analysis [13, 14]
DNA sequencing [15, 16]
forensics - genetic fingerprinting [17-19]
forensics - parental testing [20]
genetic engineering [21, 22]

A wide variety of PCR methods exist, and each has advantages and limitations. Standard or conventional PCR is the most basic type of PCR reaction. It gives qualitative results and requires a post-PCR step for detection or visualization of the DNA. A major advantage to conventional PCR is the ready access to conventional thermocyclers that almost all research facilities have, and the fairly low cost. Often, conventional PCR reactions are loaded onto an agarose gel and are resolved by size via electrophoresis. The DNA is visualized by using an intercalating dye such as ethidium bromide or Sybr Safe, and a UV light source. The specificity of the PCR reaction is confirmed by size as compared to a DNA ladder, which is a mixture of known sized fragments of DNA. The bands may be isolated from the gel, and the DNA can be purified and sequenced, which is a more dependable means of specificity determination than sizing via comparison to a DNA ladder. Figure 2 gives an example of an agarose gel with a DNA ladder on the left and several PCR reactions to its right. Limitations of this type of end-point PCR include low sensitivity and non-quantitative results.

experimental needsPCR methodadvantageslimitations
determine if a target NA sequence is present or absent in a few samplesstandard or conventionaleasy access to equipment
minimal cost
qualitative results only
post reaction handling
determine quantities of target NA in many samplesreal-timecan generate quantitative results
can be sequence specific
more expensive than conventional
PCR speed is mid-level
determine quantities of many target NA for a few samplesPCR arraysup to 88 genes can be measured per sample at a timeone array is needed per sample
costly for many samples
detect target NA in the fieldmicrofluidic chipfast results
small size
portable
specialized equipment
costly
detect very low abundance NA targets or need extremely accurate quantitationdigital and drop digitalvery precise absolute quantitationspecialized equipment
costly

Real-time PCR, also called qPCR (quantitative PCR), is a more recent but already extremely common method of PCR that offers several advantages over conventional PCR. First, the PCR product can be detected in real time, so the need for an agarose gel to visualize the DNA post-PCR is unnecessary. Further, real-time PCR can be both quantitative and specific. Starting quantities of a sequence of interest can be determined by comparison of samples to a standard curve of known quantities of DNA. The increased specificity is achieved through the use of specific NA probes and/or a melt curve analysis that follows the PCR reaction. See Figure 4 for an example of a melt curve of a single PCR product. The presence of a single peak indicates the presence of a single amplicon at the end of the PCR reaction. If multiple peaks were detected, it would indicate that multiple PCR products were produced and the PCR assay would require further optimization and development. Comparison of mRNA expression levels between species or other circumstances, by qPCR, may be achieved through the addition of a synthetic RNA to correct for differences in RNA isolation and reverse transcription. The limitations of Real-time PCR are the increased costs, the specialized thermocyclers, and the limitation of precision in quantifying the starting quantity of target NA sequences. RT-PCR can be accomplished through commercial kits such as Cells-to-CT 1-Step TaqMan Kit from Thermo Fisher [23], Qiagen OneStep Ahead RT–PCR [24] or Integrated DNA Technologies PrimeTime qPCR assay kits [25]. Human ACTB or mouse Actb gene is often as the control [25, 26]. Zeng Q et al, for example, normalized RT-PCR results to the housekeeping genes ACTB or GAPDH for human cells and Rpl19 or Gapdh for mouse cells [27]. Corman VM et al quickly established a real-time PCR assay for detecting 2019-nCoV [28].

PCR arrays utilize Real-time PCR thermocyclers are based on the Real-time PCR SYBR green assay. PCR arrays have multiple primer sets within a 96-well plate to measure the expression of up to 88 genes and eight normalization or control reactions for a given sample. PCR arrays measure a single gene-specific product within each well of the plate. These arrays normally include control reactions and allow quantitative gene expression results. Arrays are often generated to analyze the expression of a group of genes involved with a specific biological pathway, such as DNA repair, cell cycle, or oxidative stress. The advantage to PCR arrays is that expression of many different genes can be obtained at once for a sample. The limitation is that each sample must be processed separately, so it is time-consuming and also costly. De Cecco M et al employed the Qiagen RT2 Profiler Human Type I Interferon Response PCR Array (PAHS-016ZE-4) to study the involvement of L1 retrotransposon during cellular senescence [29]. D Thomas et al examined human ECM and adhesion molecules through RT2 profiler PCR array (PAHS-013Z) from QIAGEN / SABiosciences [30].

Microfluidic chip PCR utilizes microfabrication and microfluidics to amplify DNA much more quickly than conventional or Real-time PCR. Besides, the small size and integration with detection components offer improved portability and field accessibility to PCR. Digital and drop digital PCR partitions NA molecules and quantifies end-point PCR product without the need for a standard curve. This allows for very precise copy number determination and provides detection for very low copy number NA sequence targets. Although Chip PCR is robust and precise, its accessibility to many researchers is fairly low. This is due to the need for microfabrication equipment, or purchasing premade chips, which can be quite costly. However, Chip PCR is evolving rapidly, and the costs will likely be lowered significantly soon.

Many other PCR methods are well utilized in biological research. Colony PCR can be used to screen for the presence of a specific genomic insert from bacterial colonies without the need for culturing or plasmid purification [31]. Genotyping often uses allele-specific PCR [32]. Epigenetic research is heavily reliant upon methylation-specific PCR. This technique detects the methylation of CpG islands in genomic DNA [33]. Touchdown PCR reduces non-specific amplification by systematically lowering the annealing temperature as a reaction progresses. This technique can improve the yield of a specific target sequence [34]. Nested PCR can increase the sensitivity of detection. For example, de Goffau MC et al used a nested PCR-qPCR approach to detect the Streptococcus agalactiae sip gene in human placental samples [35].

Although PCR is a ubiquitous method, opportunities to reduce the length of time required to obtain results and to reduce the reaction volumes are abundant. The speed of a reaction is dependent on the types of heating and cycling mechanisms as well as reaction volume, and thus these are prime targets for assay enhancement. Metal-block-based PCR systems, which use indirect conductive heating, are widely utilized, yet the block has a high thermal mass and requires relatively large reaction volumes, making the rates of thermal change fairly slow (3ºC/s) [36]. The Roche Lightcycler uses convective heating and requires just several microliters of volume, so the speed of temperature change is much quicker (10ºC/s) [37]. Microchip PCR devices offer even greater speed and volume reduction. These are most frequently made from silicon or glass, which are thermally conductive. These chips are heated by thermoelectric heating, convection-based rotary platforms, or embedded resistive heaters [38-40] and require just a few microliters of volume. Direct heating from infrared radiation via a tungsten filament lamp has been used in microchip PCR [41]. IR mediated direct heating can also be carried out by lasers [42, 43]. Both give PCR results in minutes and require just picoliters of volume. See Labome review on PCR machines for a comprehensive review.

False positive and negative PCR results are detrimental to the usefulness of a PCR assay, and so optimization to prevent these is imperative. False positive results occur when DNA amplification is detected even though no starting temple NA was added to the reaction. This occurs when contamination is present. Clean laboratory practices are necessary to avoid aerosolization of DNA so that it isn’t unintentionally introduced into reactions. False negative results occur when no NA amplification is detected, yet the target NA is present. This can be caused by poor primers, suboptimal thermocycling parameters, or quantities of product amplicon that are below the limit of detection of a system. To reduce the occurrence of false negatives, the optimization of PCR assay design and cycling conditions is necessary. This includes the selection of quality primers [44-46], optimization of temperatures during cycling, and the use of quality template NA.

Real-time PCR offers significant advantages over conventional endpoint PCR. Real-time PCR utilizes detection systems that measure fluorescence indicative of DNA amplification within a closed tube, normally in a 96 well plate format. This negates the need for post-PCR manipulations or detection via agarose gel electrophoresis, and thus significantly reduces the time required to obtain results. The detection of the PCR product occurs after every cycle, and thus allows for the measurement of reaction kinetics. The fluorescent detection of PCR product is also much more sensitive than detection of DNA via agarose gel electrophoresis, and so this technology can identify as little as a two-fold difference in DNA quantities, something that is not possible with conventional PCR and agarose gels. Furthermore, when utilizing probe based Real-time assays, detection of the PCR amplicon is sequence specific.

DNA binding dyes: Real-time PCR employs fluorescent dyes or probes that interact with the PCR products. The two primary types of fluorescent detection are DNA binding dyes, such as SYBR Green, or fluorescently tagged sequence-specific probes, such as TaqMan or Molecular Beacon probes. When the DNA binding dyes attach to any double-stranded DNA segment, they emit a fluorescent signal [47]. When in the presence of single-stranded nucleic acids, these dyes do not attach to the NA and emit only low levels of fluorescence. Although SYBR Green is commonly used, several other DNA binding dyes are also utilized. These include SYTO 9 [48], SYTO-13, SYTO-82, [49] and EvaGreen [50]. Since detection of a fluorescent signal from these dyes is not sequence-specific, melting temperature analysis must be performed to ensure the production of a single PCR product [51]. See Figure 4 for an example of a single PCR product in a melt curve plot. If a melt curve had multiple peaks, it would indicate the presence of multiple PCR products, and the PCR assay would require further optimization. Although this may be time-consuming, SYBR Green and other DNA binding dyes remain popular because they are significantly less expensive than most alternative probe-based assays.

Nuclease dependent probes: Sequence-specific fluorescently labeled probes are the second main type of detection chemistry utilized in Real-time PCR. These probe systems can be nuclease dependent or simply hybridization probes. The nuclease cleaved probes include TaqMan, HybProbe (two oligonucleotides), minor groove binding (MGB), and locked nucleic acid (LNA) probes. These probes are complementary to a target nucleic acid sequence within the PCR amplicon, and they have both a reporter and a quencher fluorophore covalently attached to opposing ends. These systems utilize Florescent Resonance Energy Transfer (FRET) technology. When the dyes remain near one another, the fluorescent signal of the reporter dye is quenched, preventing any detectable signal. When the dyes are separated, the reporter dye’s fluorescence is unquenched and thus detectable [52]. A probe anneals to a sequence internal to the PCR primers’ binding sites, and as the Taq DNA Polymerase enzyme extends the primers to produce the PCR product, its 5’ exonuclease activity cleaves the end of the probe. The cleavage removes the quencher dye and allows excitation of the reporter dye, resulting in a fluorescent signal. Cycling probe technology (CPT) probes differ from the TaqMan type probes in that they include an RNA nucleotide. These probes form an RNA-DNA duplex upon hybridization to the target sequence. Then RNase H enzyme is used to cleave the quencher dye from the probe [53].

Hybridization probes: Although hybridization probes are also sequence-specific, they do not require the exonuclease cleavage of a dye from the probe. These include Molecular Beacons, which have a loop region between two inverted repeats, creating a hairpin structure. When the probe is denatured and anneals to a target sequence, the hairpin is released, and the fluorophores are separated from one another enough that the reporter dye generates increased fluorescence [54]. Other detection probe technologies include the Scorpion (probe and one PCR primer are combined in one molecule [55] ), and LUX (Light Upon eXtension) assays [56]. The Lux primer probe has a dye near its 3’ end that is quenched by the hairpin structure of the primer. Once it binds to a target sequence and DNA Polymerase extends the sequence, the dye’s signal increases [56].

Real-time PCR thermocycler systems detect fluorescence, which is proportional to the accumulation of PCR amplicon. However, the analysis is performed only with the data from the exponential phase of the reaction, because this is the optimal point for precise quantitation. The threshold cycle (CT) is the cycle number at which the fluorescence within a reaction crosses the threshold, a level of signal set above the baseline but within the exponential phase of the amplification. Two general methods of quantification determination are used for Real-time PCR. Relative quantification uses a relative expression ratio of the amount of target sequence from control versus experimental treatments, as normalized to the expression of a control gene such as Gapdh, Hsp90ab1, and Gusb [29] or cyclophilin A [57]. It is critical to use a control gene that has uniform expression across treatments. For instance, GAPDH served as the normalization control in experiments with human cells and the mean of Gapdh, Hsp90ab1, and Gusb served as a normalization control of RT–qPCR experiments with mouse tissues (with the exception of liver) [29]. The primary advantage to relative quantification is the reduced assay development time because the creation of a calibration curve is not required. Both a relative standard curve method and the comparative CT method (ΔΔCT) are commonly used. A mathematical model to calculate the later was described by Pfaffl [58].

Absolute quantification measures the actual amount of starting target sequence in a sample by comparing the sample amplification signal to that of a standard curve of target DNA. Calibration curves may be based on dilutions of nucleic acid molecules such as recombinant plasmid DNA containing a subcloned amplicon, Real-time PCR product, or large synthesized oligonucleotides. Plasmid DNA is widely used because it has higher stability than PCR products or oligos. However, the shorter templates are less time consuming to prepare. A plasmid standard curve is generated by performing Real-time PCR on known concentrations of the plasmid (with amplicon insert) and graphing the CT values against the log of the initial target copy number (Figure 5). The CT value is inversely proportional to the log of the initial copy number [59]. Initial copy numbers of target sequences for experimental samples are determined by linear regression against the standard curve [60].

Since the 1990s, innovative miniaturization of PCR reactions and thermocyclers has been well pursued in the form of microfluidic chip PCR. For instance, in 1998, Northrup et al reported their construction of a miniature analytical thermal cycling instrument (MATCI) that was silicon-based and contained chambers with integrated heaters [61]. The device performed fluorescence measures in real-time as PCR cycling occurred, and was as portable as a briefcase. Since then, the field of microfluidic chip PCR has exploded, and advancement of the technology has included multiple fabrication materials and architectures [62].

A majority of PCR chip reaction wells are silicon-based. Silicon has excellent thermal conductivity, which allows for quick temperature changes during thermocycling. However, chips have also been fabricated from glass, polymers including polycarbonate and polydimethylsiloxane, ceramic, and 317 stainless steel. Chip architectures are either stationary chamber based or dynamic continuous flow based. The stationary chamber chips have a PCR reaction solution that is held stationary while the temperature of the chamber is cycled. These chips may be single or multi-chambered. The dynamic continuous flow based chips have amplification while the sample is pumped through a microfluidic channel during temperature cycling. With these chips, the flow rate of the sample determines the time for temperature transitions. Microchip PCR offers the benefit of short assay times, low consumption of reagents, rapid rates of heating and cooling, and reduced power consumption. Microchip PCR may require downstream handling for DNA detection, such as agarose gel or capillary electrophoresis. However, capillary electrophoresis, as well as fluorescence detection or DNA hybridization methods of DNA detection, may be included on the chip [38, 63]. Microchip PCR continues to be developed with advanced integration of on-chip detection systems and optimization of both fabrication materials and temperature cycling methodologies.

The microfluidic chamber based digital PCR (cdPCR) method is capable of absolute quantification of nucleic acids without the use of standard curves [64]. With digital cdPCR, the sample is partitioned into discrete water-in-oil droplets that may have no target NA or may have one or more copies. The presence of the target sequence is measured at the end of PCR amplification, and the concentration is calculated from the fraction of positive partitions versus the total number of partitions. The “Drop” digital PCR (ddPCR) is a cutting-edge version of digital PCR. It has twenty-five times more partitioned droplets, which allows for very accurate copy number estimates [65]. M Aubert et al quantified the presence of latent herpes simplex virus genpme in mouse cells with ddPCR [66]. de Morree A et al compared the expression of Pax3 RNA isoforms between diaphragm and limb muscle stem cells through digital PCR [67]. Lee J et al measured allele-specific expression of LMNA wild-type and mutant allele in iPSC-derived cardiomyocytes with ddPCR [68]. Nam AS et al detected CALR somatic mutations from various subsets of CD34+ blood cells with ddPCR by using the QX200 Droplet Digital PCR System from Bio-Rad [69]. Researchers have directly compared ddPCR and real-time PCR platforms for the detection of low abundance DNA and found that ddPCR was superior over real-time PCR [70]. In February 2019, Bio-Rad released the first FDA-cleared droplet digital PCR system and BCR-ABL fusion test for monitoring chronic myeloid leukemia treatment response.


Watch the video: Native gel electrophoresis (June 2022).