Southern Blot Scientific Method Theory and Guide

Southern blot scientific method title

Southern Blots: Overview & Theory

Southern Blotting is a technique that is used to detect whether you have a specific DNA sequence available in your sample. Most commonly, you will be testing for DNA that you get from cell lysates or after creating your own plasmids. You can also do some amazing things like making transgenic mice and proving that you have selected for certain genes of interest to you! All you have to do is Southern Blot any set of DNA from the mice and you’ll have concrete proof. As you probably understand, this is one of the most common techniques that geneticists and molecular biologists know because everything they do needs to be proved by a southern blot.

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To execute a southern blot, first collect some DNA. This can be from a cell lysate or from tissue samples, etc. Next, digest the DNA using DNAse and run these fragments on an agarose gel. This will separate the DNA by size and you’ll know which well in the gel corresponds with which sample. Next, in a similar fashion to Western blots, this DNA is then transferred onto blotting paper (typically made of nitrocellulose or nylon). After the “blotting” process is complete, the DNA is then “probed” using a radioactive (or fluorescent) DNA sequence that is complementary to the sequence that you want to detect. Finally, this “probed” radioactive blot is then imaged using an autoradiograph. The steps for this process are described in the following image:
Southern Blot Scientific Method
Southern Blot Step by Step
Southern Blot Tips and Tricks
Southern Blotting Method

Nuances of Southern Blotting theory

Why is there is a second denaturing step for DNA that’s 15 KB or larger?
It’s because these large pieces of DNA don’t transfer very well to the blotting paper. As you can imagine, the larger a piece of DNA is, the slower it will migrate through an agarose gel. Even after leaving the blotting paper overnight, the transfer may not be complete!

How does the DNA actually go from the gel into the blot?
Through capillary action and wicking! Unlike a western blot where a voltage gradient is utilized to pull proteins into the blotting paper, in Southern Blots, the DNA merely moves over to the blotting paper overnight without much force at all. That’s why it is incredibly important to make sure that the blotting paper and the gel are in close contact. It’s also very important to make sure that the glass plate on top is heavy enough so that it forces the gel and the blot together.

Why is there a UV step?
By using UV after the transfer step, the DNA (which has some free aldehyde groups due to depurination) can react with the nitrocellulose/nylon membrane to form covalent bonds.

What do the NaOH and the HCl do during the denaturation step?
Essentially HCl removes some/all of the purine bases from the DNA and makes the two DNA strands less sticky to each other (because there is less hydrogen bonding). This process is called depurination. NaOH also prevents the two strands from forming hydrogen bonds due to deprotonation of all bases.

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

How does radiolabeling with P32 work?
Small amounts of DNAse introduce nicks into the single stranded probe DNA. DNA polymerase then utilizes the dATP32 from solution to repair these nicks and incorporates these radioactive phosphates into the backbone of the DNA.

Southern blot Step-By-Step Guide

Materials for Southern Blot of Mouse Tail DNA

Denhardt’s Solution 50X (5g Ficoll 70000, 5g Polyvinyl pyrrolidone, 5g BSA Fraction V in 500 ml water)
Hybridization cocktail (25 ml 50% dextran sulfate, 25 ml 20X SSC, 50 ml formamide, 1 ml Tris 1M, 2 ml Denhardt’s Solution 50X, 1ml 10% SDS) – prefilter before use
TE buffer (10 mM Tris HCl pH 7.5, 1 mM EDTA)
BamHI enzyme and buffer (#R0136S New England BioLabs)
100X BSA (B9000S New England BioLabs)
Gel loading dye (#G2526 Sigma)
TBE buffer 10X (#93290 Sigma)
SSC buffer 20X (#S6639 Sigma)
[α32P]dCTP (3000 Ci/mmol; #NHG013H250UC Perkin Elmer)
Sephadex G-50 (#G50150 Sigma)
Ready-To-Go DNA Labeling Beads (-dCTP) (#27-9240-01 GE Lifesciences)
Whatman paper (#WHA10427810 Sigma)
Salmon Sperm DNA (#15632011 ThermoFisher)

Southern Blot Protocol
DNA digest and gel electrophoresis

    1. Setup DNA digest reactions from collected tail samples as follows:

10 µl DNA (12 ug)
6 µl 10X BamHI buffer
0.6 µl 100X BSA
1 µl BamHI
42.4 µl H2O
Incubate at 37°C for at least 5 hours.

  1. Add 6 µl gel loading dye to each sample.
  2. Run samples of a 0.8% agarose gel in TBE for 16 h at 25V.
  3. Stain gel in 1 µg/ml ethidium bromide for 30 min.
  4. Take a photo of the gel.*


    1. Soak gel in 0.2 N HCl for 10 min with shaking to depurinate (remove purines from DNA). Rinse with water.
    2. Soak gel in two washes of 500 ml 0.4 M NaOH/1.5 M NaCl solution for 30 min each time.
    3. Soak gel in two washes of 500 ml 1 M Tris/1.5 M NaCl solution for 30 min each time.
    4. Setup transfer stack by placing 2 layers of Whatman paper on glass plate (wet before laying down) with soaked gel on top (flipped over). Cover with plastic wrap.
    5. Using a razor blade, cut out plastic over gel and place wetted membrane on gel. Place two layers of Whatman paper on top (wet before laying down).
    6. Place a stack of paper towels on the top and cover with a glass plate and transfer in 6X SSC overnight.
    7. Dismantle and air-dry membrane.
    8. Expose membrane to UV for 90 s (DNA side down). Bake between Whatman paper at 80°C for at least 1 h.

Labelling the probe

      1. Denature probe by adding 2 µl probe (25-50 ng) to 44 µl TE and incubating at 95°C for 4 min). Immediately place on ice.
      2. Add 46 µl denatured probe and 4 µl of [α32P]dCTP to Ready-to-Go labelling bead tube. Flick to mix.
      3. Incubate at 37°C for 15-30 min.
      4. Add 50 µl TE and run through a G50 sephadex column

Probe Hybridization and Autoradiograph

      1. Add 50 ml hybridization cocktail to the membrane.
      2. Add 300 µl of 10 mg/ml salmon sperm DNA to labelled probe and denature at 95°C for 5 min. Immediately place on ice. Add to membrane and cocktail solution.
      3. Hybridize overnight at 42°C.
      4. Wash briefly with 50 ml wash solution (0.2X SSC/0.1% SDS).
      5. Wash for 30 min with 100 ml wash solution at 50°C. Repeat.
      6. Check blot with Geiger counter to estimate cpm.
      7. Expose membrane to film and store at -70°C.
      8. Develop after 2-5 days.

Notes on this Southern Blot method

      • During DNA digest Step 5 you can note the position of the marker bands by using a fluorescent ruler or marking bands directly on the gel by punching small holes with a needle.
      • This method uses dCTP32 and not dATP32 as we talked about in the theory section
      • After undergoing HCl and NaOH treatment for the denaturation step, it is very important to neutralize the gel once again.
      • If you don’t want to buy the labelling beads, an alternative strategy is presented here at MIT Cores

Application of Southern Blots on SciGine, a Scientific Method Search Engine


ASU Southern Blots Guide
MIT Core Guide
DNA Blot from Davidson College

Southern Blot Video by Shomu’s Biology

Western Blot Theory and Method Guide

Western Blot Method Guide and Step by Step Procedure

Overview of Western Blot Method

A western blot enables sensitive detection of specific proteins from a solution containing multiple proteins. This is an essential biology technique and one of the cheapest methods that can be utilized to analyze proteins. To perform a western blot first separate proteins based on their mass and charge via gel electrophoresis, and then follow up by detecting the protein of choice with a specific antibody. Typically, researchers will use western blots to separate proteins from cell media or from cell lysates. For example, if you wanted to find out how much actin your cells are expressing, a western blot can easily compare actin amounts between different cell types. It’s also likely that you will be using western blots when producing proteins in mammalian and insect cells.

In a typical western blot procedure, cells will first be lysed and the amount of protein will be determined using a spectrophotometer. Then a gel will be made and the total protein from the cell lysate will be loaded into wells in the gel. After applying an electrical field, the proteins in the gel will begin to migrate down and separate into distinct bands based on the size and charge of the protein. After the smallest proteins reach the bottom of the gel, the electrophoresis will be stopped and all proteins on the gel will be transferred onto blotting paper so that they can easily be handled. Finally, antibodies that recognize the proteins of interest will be added and detected via chemiluminescence.

Related articles:

Here is a step by step illustration of how to perform a western blot:

Western Blot Scientific Method Guide

Western Blot Step By Step

SDS-PAGE Western Blot Step-by-Step Protocol

Western blotting can be used to examine the upregulation of RCAN1, a signaling molecule in neuronal cell types.

Materials for Western Blots:

  • Rabbit anti-RCAN1 antibody (#SAB2101967, Sigma-Aldrich)
  • SDS-PAGE gel (Criterion TGX precast Stain-free Any kD gel, #5678124, Bio-Rad)
  • TBS (20 mM TrisCl pH 7.6, 150 mM NaCl)
  • Running buffer (25 mM Tris, 192 mM glycine, 0.1% SDS)
  • Transfer buffer (25 mM Tris, 192 mM glycine, 20% methanol, 0.05% SDS)
  • 4X SDS-PAGE loading buffer (Laemmli’s sample buffer #1610747 Bio-Rad; add fresh dithiothreitol to 10 mg/ml on the day of experiment)
  • Transfer membrane (0.2 um polyvinylidene fluoride membrane, #03010040001 Roche)
  • Secondary antibody (donkey anti-rabbit horseradish peroxidase-conjugate; #711-035-152 Jackson ImmunoResearch)
  • Blocking buffer (5% skim milk powder in TBS with 0.1% Tween20)
  • Immun-Star WesternC Chemiluminescence reaction solutions (#170-5070 Bio-Rad)
  • Dual color ladder (#1610374 Bio-Rad)
  • Blotting paper (ProteanXL, #1703966 Bio-Rad)

Western Blot Experimental procedure:

  1. Unwrap precast gel and rinse wells three times with running buffer. Assemble gel in tank and fill with running buffer.*
  2. In an Eppendorf tube add protein sample (30 µg) to 10 µl 4X SDS-PAGE loading buffer and add water to a final volume of 40 µl.
  3. Heat samples to 95°C for 2 min and spin briefly to ensure contents are at the bottom of the tube
  4. Load gel with samples and include ladder in one lane.
  5. Run gel at 200V for 30 min.
  6. While gel is running, soak two pieces of blotting paper (cut to the same size as the gel) in transfer buffer (approx. 30 min). Activate transfer membrane (also cut to size) by dipping in methanol, then soak in transfer buffer for approx. 10 min.
  7. Remove gel from tank and place in transfer buffer.
  8. Assemble transfer “sandwich” by placing down soaked blotting paper, transfer membrane, gel and blotting paper onto open transfer cassette (Turbo Blot transfer unit; Bio-Rad). Use a glass rod to roll across the “sandwich” to remove any air bubbles.
  9. Close cassette and run in machine (standard minigel program for 30 min).
  10. Remove transfer membrane from cassette, taking care to snip one corner to ensure orientation.*
  11. Incubate transfer membrane in blocking buffer for 1 h at 4°C with rocking.
  12. Pour off blocking buffer and add diluted anti-RCAN1 antibody 1:200 in 5 ml TBS with 2.5% skim milk power and 0.05% Tween20. Incubate overnight at 4C with rocking.
  13. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time.
  14. Incubate with diluted secondary antibody 1:2500 in 5 ml TBS with 0.2% Tween20 at 4°C with rocking for 1 h.
  15. Wash membrane three times in TBS with 0.2% Tween20 at 4°C with rocking, for 10 min each time. Rinse membrane briefly in water.
  16. Mix 1 ml each of ECL reagents in a foil-wrapped tube and add to membrane for 5 min prior to imaging on ChemiDoc MP imager (Bio-Rad).

Procedural notes for this Western Blot Method:

  • This precast gel contains 18 lanes with a loading capacity of 10-40 ug protein in up to 30 ul per well
  • Small needle-point markings can be added to membrane in-line with color markers which reduce in intensity following subsequent incubation and washing steps.
  • To make sure you know which step you are on, cut the bottom right side of your gel after running the gel electrophoresis.
  • In this method, the protein is denatured prior to running on the gel. This is called SDS-PAGE. By denaturing, you ensure that the size and charge are all that matter, as opposed to native gel electrophoresis where the conformation of the protein also matters.
  • Blots can be regenerated (the antibodies that were used for probing can be removed) by using stripping buffer. However, blots can only be stripped a few times before they have too much background noise to be easily analyzed.

Applications of Western Blots on Scigine (Search Engine for Scientific Methods):


NIH Western Blot Reference
Western Blotting by Kurien et al

Good Video References Related to Western Blots:

Flow Cytometry and FACS: Method Guide

FACS & Flow Cytometry for Cell Population Analysis

FACS Method Overview: Using Fluorescence to Understand Cells and Cell Populations.

FACS, or Fluorescence Assisted Cell Sorting, is a type of technique that enables you to understand cells by tagging them with fluorescent markers. There are a number of things that FACS allows you to do:

  • Separate one type of cell from a mixture of cells
  • Count how many cells are in a mixture
  • Detect what kind of biomarkers a cell might have (is it cancerous? Just a regular lymphocyte? Etc.)
  • Find out if your cell is making a protein of choice (similar to biomarker detection)
  • And more.

It’s an amazing and powerful technique that you should always keep as part of your skills. Most Bio-Pharmaceutical companies have FACS experts in-house because it is so versatile!

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The principle of FACS is simple:

  1. Label cells of interest with a marker such as an antibody
  2. Pass the cells through a laser and detect which ones have the antibody
  3. Separate those cells from the rest

Here’s what this looks like in a simple picture:
FACS and Flow Cytometry Method Overall

What does the FACS instrument detect?

  • Forward Scatter Light (FSC): When you shine a laser on an object, the object blocks some of it and the rest continues to go forward towards the detector. This is called forward scattered light.
  • Side Scatter Light (SSC): An object might also bounce some of the laser in an alternate direction. There is a detector that is orthogonal to the light direction and detects this scattered light.
  • Fluorescence Emission Signal: Excite your fluorophores with a certain wavelength of light and they’ll emit a different wavelength depending on the type of fluorophore. Fluorescence detectors in the instrument will detect this emitted light.

Take a look at this illustration:
FACS Detection of Molecules and Cells
Based on the previous section, FACS provides data on forward scattered light, side scattered light, and fluorescence intensity. By graphing these different values together, you can get an idea of what your cell sample is like. Here’s the kind of data you would get:

  • Plot of FSC vs. SSC:
    • Cell debris in your solution is usually small, so it would have low FSC and SSC values.
    • Normal cells have medium sized FSC and SSC values
    • Large cells like Granulocytes have large FSC and SSC values
  • Plot of Fluorescence intensity:
    • After cleaning your cell solution and removing any non-specific fluroescence, your cells will be the only thing with large fluorescence intensity
    • A plot of intensity provides you a histogram telling you how much fluorescence is in the solution
    • You can use this to find out how many cells are there, how many antibodies bound to each cell, how much of a certain biomarker is there, etc.
    • You can compare different fluorescence histograms to compare cell types as well

Take a look at these images:
FACS Understanding Dot Plots and Histograms

Gating: How to measure cells of only one population and ignore debris

In the above images you can see that your figures may provide information on several types of particles: debris, small cells, large cells, particles with high and low fluorescence, etc. In order to narrow down your results, most instruments allow you to “Gate” your results so that the instrument provides you with a new graph within limits that you set. As an example, you may plot FSC vs. SSC and then gate only for larger particles. Then using that gate you may look at the fluorescence intensity in Red, Green, and Blue. This way you can analyze cells that have red biomarkers, or cells that have blue biomarkers specifically.

As an example, what if you wanted to only collect cells that highly express one cancer-related cellular protein (ex: folate receptor) ?

Using FACS, this would be easy to analyze! Just mark your cells with an anti-folate receptor antibody with GFP. Then, use FACS to separate out the cells that express the receptor in your sample. Take a look at this image to understand this concept.
Gating in Flow Cytometry

Note: Grammarly is a free grammar check plugin for Chrome. I used it for this article and really like it! Try it out here

Complete Step by Step Method Guide

Unfortunately, this is such a highly instrument-oriented technique that it’s difficult for me to provide details on how to operate a FACS machine. Nonetheless, take a look here to get an idea.

Tissue Culture Flask
With cells such as HT-29
Trypsin such as Tryp LE from Invitrogen
Serological pipette
Without pH indicator so that there isn’t any extraneous fluorescence


  1. Grow HT-29 to confluence in a T-75 flask in 7 ml of DMEM
  2. Use lab protocols to mark them with any antibodies that you’d like such as an anti-folate receptor antibody
  3. Aspirate the DMEM and add PBS 1x
  4. Swirl around and remove PBS
  5. Wash once again with PBS by following the previous 2 steps
  6. Add 2 ml of Trypsin (TrypLE) and wait until cells detach for approx 10 minutes
  7. Collect cells into a falcon tube by using a serological pipette
  8. Use a hemacytometer to dilute cells until they are at a concentration of 1×106 cells/ml using DMEM
  9. Load cells into the FACS using using appropriate 5 ml tubes
  10. Make sure the instrument is running well by using control samples or any extra cells that you have
  11. Replace cell collection tubes and make sure they are sterile if you plan on culturing cells that are collected
  12. Run the FACS instrument 🙂

Notes on this FACS/Flow Cytometry Methodology

  • FACS may also be referred to as Flow Cytometry on Job Postings. This is because FACS is a part of the overall group of techniques called Flow Cytometry. In biology, however, it is unlikely that you will use any other techniques besides this one.
  • The types of tubes that are necessary for loading a FACS unit vary between instruments
  • Some cells clump up and make data analysis harder. Try diluting them more or mixing them more using your serological pipette
  • It’s important to set the instrument’s gain settings to maximize/minimize signals as appropriate
  • Make sure to also set the correct excitation/emission levels for your various fluorescence markers. You will also need to set your thresholds such that your signals don’t overlap.
  • If you plan on using multiple fluorescence markers at the same time make sure you consult a fluorescence ex/em table like this fluorescence spectra table. You need to make sure that your fluorescent signals don’t overlap.

Applications of FACS on SciGine

Analysis of cell cycle and viability with a FACScan
Analysis of a GFP reporter gene using FACS
Analysis of gene disruption in diploids
Using FACS to analyze camptothecin induced apoptosis
FACS analysis of cell cycle using Propidium Iodide


Excellent Antibodies-Online FACS reference
Thermofisher FACS manual
What is FACS from UMass

HPLC: Biochemical Analysis. A Step-By-Step Method Guide

HPLC Analysis Step by Step

HPLC Method Overview

HPLC, or high performance liquid chromatography is an amazing analytical technique for chemical compounds including biopolymers, small molecules, and polymers. In this method, a sample is first dissolved to make a solution. This solution is then injected into a “column” that contains resin that will interact with the sample. This will slow down the movement of the sample through the “column” and as the sample comes out the other side of the column, it is detected. This allows you to know both the time at which the sample comes out and the intensity of the sample that was detected. Here’s an overview of this technique:

Note: If you’re writing research papers, I highly recommend Grammarly – it’s a free grammar check plugin for Chrome. Try it out here…

HPLC Bioanalytical Method Guide

So, while there is continuous flow of some buffer through the column, we also inject our sample and observe as different molecules within the sample come out at different “retention times”. The detector on the end of the column can be any kind of detector but the most common types are refractive index (RI), ultraviolet (DAD), and fluorescence (FLD). Each of these will detect different properties of the molecules that come out of the column and display a chromatogram.

HPLC Chromatogram Guide

Types of Chromatography

Different column resin compositions determine the kind of chromatography that you are running and what molecules you can separate.

  • Normal Phase: The column is filled with silica particles which are polar and the buffer running through the system is non-polar. Once you inject your sample, polar particles will stick to the silica more and have a longer retention time than non-polar molecules.
  • Reverse Phase: The column is filled with hydrophobic particles (actually they are silica particles with long hydrocarbons on the surface). The buffer that is running through the system is polar (such as acetonitrile/water or methanol/water mixtures). This means that hydrophobic molecules will stick to the resin more and be retained longer.

Complete Step by Step HPLC guide


HPLC autosampler vialsI only use autosamplers since manual injection is tedious 🙂
Centrifugal filters with 0.2 um poresTo clean up samples
Eppendorf vialsFor centrifuging
HPLC machine


In a typical HPLC procedure you can decide the following variables:

VariableWhat it does
Flow rateWith fast flow peaks come out sooner but there’s they’re harder to resolve and tend to blend together. For more resolution, run slower.
PressureAffected by flow rate and solvent
Solvent BuffersDetermines signal intensity, how quickly the peaks come out, signal fidelity
Column TypeDetermines the type of interaction with the sample
Detection ParametersIf using UV or FLD, you need to set the right excitation/emission wavelengths

Since HPLC is a very machine-variable technique, I can only provide general guidelines.

For sample preparation:

  1. Dissolve your biopolymers or small molecules in a suitable solvent such as methanol
  2. Centrifuge at 10,000 rcf in an eppendorf vial and keep the supernatant to remove any large particular matter
  3. With a centrifugal filter, add 500 ul of your sample solution onto the top
  4. Centrifuge at 10,000 rcf and collect the filtrate (the solution that successfully passes through the filter)
  5. Load this sample into an HPLC vial

For setting up the HPLC machine:

  1. Make sure you have all your buffers set up
  2. Open the purge valve and purge the system for 5 minutes.
  3. Add your samples into the autosampler tray
  4. Stop the purge
  5. Close the purge valve
  6. Run the system at a normal flow rate (1 ml/min) with your buffer to equilibrate the column for 10 minutes
  7. Make sure that your pressure is stable (ie, less than 2-3 bar of fluctuation)
  8. Set up your sequence and your method
  9. Run a standard before your actual samples or as part of the same sequence

Example buffer system to determine Fluoresceinamine levels in samples:
Sample: Add 10 ug of fluoresceinamine into 1 ml of Acetonitrile.
Buffer: Pure acetonitrile buffer on a C-18 column; this is “reverse phase”.
Flow rate: 1 ml/min.
Column: 4.6 mm x 30 cm size.
Detection: Detect via a fluorescence detector set to Excitation @ 485 nm and Emission @ 535 nm.

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Notes on HPLC methodology

  • To clean the system and equilibriate it, you need to run enough solvent. However, this amount varies column-to- column. A typical 4.6 mm x 30 cm column should be clean when you follow the procedure above.
  • Isocratic means that the solvent concentration stays constant throughout the run.
  • It is useful to run standards before your samples as well as with your samples. Standards make it easy to identify which peak pertains to your molecule of interest.
  • Always use HPLC grade solvents. This is especially true for solvents like THF which are frequently sold with inhibitors that also complicate your ability to detect your molecule of interest.

Applications of HPLC on SciGine

HPLC is such a versatile technique. Take a look at these methods on SciGine which assay different types of chemicals in various samples.


ChemGuide Summary of Technique
Method Guide from Waters
Overview on Wiki

Using PCR To Amplify DNA, A Step-By-Step Guide

PCR Method Guide on SciGine

PCR Biological Method Overview

PCR, or polymerase chain reaction, is a method to amplify a segment of DNA for analysis. Because it is such a powerful technique, there are a HUGE number of situations where PCR may be used. Some common reasons for using it are:

  1. Microbiology: You need to know if your bacteria was transformed properly with your plasmid
  2. Oncology: You need to know if a particular gene exists in your cancerous cells
  3. Forensics: You need to know if a certain criminal’s DNA was present at the crime scene

The basic steps of PCR include:

  1. Designing primers to designate a target DNA sequence to amplify
  2. Mixing together the primers with the target DNA strand, polymerase enzyme, and deoxynucleotides
  3. Running a thermocycler multiple times in “cycles” to repeatedly:
    • Separate DNA strands
    • Allow primers to anneal
    • Allow the polymerase to attach and synthesize a new DNA chain
  4. Sequencing or Electrophoresis to prove that the PCR worked

In a nutshell, here is what Step 3, above, looks like:

PCR, a biological method, amplifies DNA - SciGine

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Ingredients in a PCR mixture – What every component does

In general, a PCR tube will contain the following items:

Water: Solvent
dNTPs (or deoxynucleotide triphosphates):Single bases A, T, C, and G which are used by the polymerase while replicating the DNA. As the polymerase adds base pairs onto the new DNA strand, one base pair is used at a time.
MgCl2: An essential cofactor for the polymerase enzyme
Primers: Short segments of single-stranded DNA used to frame the DNA region that needs to be amplified. They are complementary to the template DNA strand only at defined locations around the target sequence.
Target DNA: The DNA “template” that you want to make copies of. This can be a full DNA chain or a part of a longer chain.
Taq Polymerase: An enzyme from Thermis aquaticus that uses dNTPs and replicates DNA starting from the 3′ end of a template strand towards the 5′ end. Taq  is used in PCR specifically because it is resistant to the high temperatures used for separating DNA strands during Step 3a above .

Complete Step-By-Step PCR Protocol


PCR tubes

PCR temperature Cycler


Reaction Buffer

10 mM Tris-HCl, 50 mM KCl, and 1.5 mM MgCl2, pH 8.3

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  1. For each PCR tube set up the following mixture of materials up to a 50 ul total volume on ice

    dNTP solution

    200 uM for each dNTP

    Forward Primer

    0.2 uM

    Reverse Primer

    0.2 uM

    Target DNA

    Less than 1000 ng

    Taq Polymerase

    1.25 units per tube

    Nuclease Free Water

    Q.s up to 50 ul

    Mineral Oil

    Add a little bit on top of each tube if you don’t have a heated lid on the temperature cycler

  2. Pipette each tube up and down several times, gently, so as not to add bubbles
  3. Centrifuge each tube down for 1-2 seconds at 100 rcf to bring all contents to the bottom
  4. Heat up the temperature cycler to 95oC
  5. Quickly transfer tubes from ice to the temperature cycler and begin thermocycling
  6. Typical thermocycle procedure:

    Step Name




    95 oC

    30 seconds

    Cycle Denaturation

    95 oC

    30 seconds

    Cycle Priming

    50-60 oC

    60 seconds

    Cycle Extension

    72 oC

    1 minute per kilobase of target DNA

    Final Extension

    72 oC

    5 minutes

    Hold Temperature

    4 oC


  7. Repeat the above “cycle” steps 35-45 times.
  8. Make sure to keep the samples on ice while you are not using them. The infinite hold sequence allows you the flexibility of leaving your samples in the temperature cycler while you are busy with other lab work.

Notes on this PCR Methodology

  • Note 1. It is important to design your PCR primers to be specific to only the regions flanking the target sequence. Typically, specific primers are ~30-40 bases in length.
  • Note 2. The Tm (melting temperature) of the primers affect the temperature in Step 3b and the “Cycle priming” step.
  • Note 3. For greater accuracy/fidelity while copying DNA, a Pfu polymerase (from Pyrococcus Furiosus) may also be used.
  • Note 4. With 35 cycles, the target DNA is amplified 236 times its initial concentration. For times when you have very little target DNA, you can amplify 45 times or more.
  • Note 5. For Primer design, you can use tools such as Primer 3. Tools are also available for calculating the Tm of your primers such as this Tm calculator.
  • Note 6. G-C rich sequences will need longer denaturation times (typically up to 5 minutes) because of the additional hydrogen bonding. Any template with more than 60% G-C bases is considered G-C rich.