A detail note on Plasmid

DNA isolation methods
Many different methods and technologies are available for the isolation of genomic DNA.
In general, all methods involve disruption and lysis of the starting material followed by
the removal of proteins and other contaminants and finally recovery of the DNA.
Removal of proteins is typically achieved by digestion with proteinase K, followed by
salting-out, organic extraction, or binding of the DNA to a solid-phase support (either
anion-exchange or silica technology). DNA is usually recovered by precipitation using
ethanol or isopropanol. The choice of a method depends on many factors: the required
quantity and molecular weight of the DNA, the purity required for downstream
applications, and the time and expense. Home-made methods often work well for
researchers who have developed and regularly use them. However, they usually lack
standardization and therefore yields and quality are not always reproducible.
Reproducibility is also affected when the method is used by different researchers, or with
different sample types. The separation of DNA from cellular components can be divided
into four stages:
1. Disruption
2. Lysis
3. Removal of proteins and contaminants
4. Recovery of DNA
In some methods, stages 1 and 2 are combined.
DNA isolation from Bacteria
Plasmid Preparation
A plasmid preparation is a method used to extract and purify plasmid DNA. Many
methods have been developed to purify plasmid DNA from bacteria. These methods
invariably involve three steps:
• Growth of the bacterial culture
• Harvesting and lysis of the bacteria
• Purification of plasmid DNA
Growth of the bacterial culture
Plasmids are almost always purified from liquid bacteria cultures, usually E. coli, which
have been transformed and isolated. Virtually all plasmid vectors in common use encode
one or more antibiotic resistance genes as a selectable marker (Ex :kanamycin,
Ampicillin), which allows bacteria that have been successfully transformed to multiply
uninhibited. Bacteria are grown under favorable conditions.

Harvesting and lysis of the bacteria
When bacteria are lysed under alkaline conditions both DNA and proteins are
precipitated. Some scientists reduce the concentration of NaOH used to 0.1M in order to
reduce the occurrence of ssDNA. After the addition of acetate-containing neutralization
buffer the large and less supercoiled chromosomal DNA and proteins precipitate, but the
small bacterial DNA plasmids can renature and stay in solution.

Preparations by size
Kits are available from varying manufacturers to purify plasmid DNA, which are named
by size of bacterial culture and corresponding plasmid yield. In increasing order, these
are the miniprep, midiprep, maxiprep, megaprep, and gigaprep. The plasmid DNA yield
will vary depending on the plasmid copy number, type and size, the bacterial strain, the
growth conditions, and the kit.
Mini preparation
Minipreparation of plasmid DNA is a rapid, small-scale isolation of plasmid DNA from
bacteria. It is based on the alkaline lysis method invented by the researchers Birnboim
and Doly in 1979. The extracted plasmid DNA resulting from performing a miniprep is
itself often called a “miniprep”. Minipreps are used in the process of molecular cloning to
analyze bacterial clones. A typical plasmid DNA yield of a miniprep is 20 to 30 µg
depending on the cell strain.
Miniprep Protocols
http://www.protocol-online.org/prot/Molecular_Biology/Plasmid/Miniprep/
Midipreparation
The starting E. coli culture volume is 15-25 ml of LB broth and the expected DNA yield
is 100-350 µg.
Maxipreparation
The starting E. coli culture volume is 100-200 ml of LB broth and the expected DNA
yield is 500-850 µg.
Megapreparation
The starting E. coli culture volume is 500 ml – 2.5 L of LB broth and the expected DNA
yield is 1.5-2.5 mg.
Gigapreparation
The starting E. coli culture volume is 2.5-5 L of LB broth and the expected DNA yield is
7.5–10 mg.
Purification of plasmid DNA
Addition of phenol/chloroform can dissolve and denature proteins, like DNase. This is
especially important if the plasmids are to be used for enzyme digestion. Otherwise,
smearing may occur in enzyme restricted form of plasmid DNA.

Alkaline lysis
Alkaline lysis was first described by Birnboim and Doly in 1979 (Nucleic Acids Res. 7,
1513-1523) and has, with a few modifications, been the preferred method for plasmid
DNA extraction from bacteria ever since. The easiest way to describe how alkaline lysis
works is to go through the procedure and explain each step, so here goes.
1. Cell Growth and Harvesting
The procedure starts with the growth of the bacterial cell culture harboring your plasmid.
When sufficient growth has been achieved, the cells are pelleted by centrifugation to
remove them from the growth medium.
2. Re-suspension
The pellet is then re-suspended in a solution (normally called solution I, or similar in the
kits) containing Tris, EDTA, glucose and RNase A. Divalent cations (Mg2+, Ca2+) are
essential for DNase activity and the integrity of the bacterial cell wall. EDTA chelates
divalent cations in the solution preventing DNases from damaging the plasmid and also
helps by destabilizing the cell wall. Glucose maintains the osmotic pressure so the cells
don’t burst and RNase A is included to degrade cellular RNA when the cells are lysed.
3. Lysis
The lysis buffer (aka solution 2) contains sodium hydroxide (NaOH) and the detergent
Sodium Dodecyl (lauryl) Sulfate (SDS). SDS is there to solubilize the cell membrane.
NaOH helps to break down the cell wall, but more importantly it disrupts the hydrogen
bonding between the DNA bases, converting the double-stranded DNA (dsDNA) in the
cell, including the genomic DNA (gDNA) and your plasmid, to single stranded DNA
(ssDNA). This process is called denaturation and is central part of the procedure, which is
why it’s called alkaline lysis. SDS also denatures most of the proteins in the cells, which
helps with the separation of the proteins from the plasmid later in the process.
It is important during this step to make sure that the re-suspension and lysis buffers are
well mixed, although not too vigorously (see below). Check out my article on 5 tips on
vector preparation for gene cloning for more information and tips. Also remember that
SDS and NaOH are pretty nasty so it’s advisable to wear gloves and eye protection when
performing alkaline lysis.
4. Neutralization
Addition of potassium acetate (solution 3) returns decreases the alkalinity of the mixture.
Under these conditions the hydrogen bonding between the bases of the single stranded
DNA can be re-established, so the ssDNA can re-nature to dsDNA. This is the selective
part. While it is easy for the the small circular plasmid DNA to re-nature it is impossible
to properly anneal those huge gDNA stretches. This is why it’s important to be gentle
during the lysis step because vigorous mixing or vortexing will shear the gDNA
producing shorter stretches that can re-anneal and contaminate your plasmid prep.
While the double-stranded plasmid can dissolve easily in solution, the single stranded
genomic DNA, the SDS and the denatured cellular proteins stick together through
hydrophobic interactions to form a white precipitate. The precipitate can easily be
separated from the plasmid DNA solution by centrifugation.
5. Cleaning and concentration
Now your plasmid DNA has been separated from the majority of the cell debris but is in
a solution containing lots of salt, EDTA, RNase and residual cellular proteins and debris,

so it’s not much use for downstream applications. The next step is to clean up the
solution and concentrate the plasmid DNA.
There are several ways to do this including phenol/chloroform extraction followed by
ethanol precipitation and affinity chromotography-based methods using a support that
preferentially binds to the plasmid DNA under certain conditions of salt or pH, but
releases it under other conditions. The most common methods are detailed in my article
on 5 ways to clean up a DNA sample
Differences between plasmid and Genomic DNA
If you want to isolate plasmid DNA, you crack your cells open and carry out a miniprep,
trying very hard not to get any contaminating genomic DNA in your sample. If you want
genomic DNA, you crack your cells open in a different way and try to isolate as much of
the stuff as possible.
So what’s the difference?
In this article, I’ll explain how both plasmid and genomic DNA preps work and how they
are different.
Genomic DNA Extraction:
1. Lysis: Just Crack Them Open
Genomic DNA extraction is the simpler of the two procedures because all that is needed
is a good strong lysis to release the genomic DNA into solution. For yeast, plant cells and
bacteria, this involves breaking down the strong, rigid cell wall before mechanically
disrupting the membrane. The cell wall can normally be broken down using enzymes
such as lysozyme, which catalyses the hydrolysis of the cell wall peptidoglycans and the
serine protease, proteinase K (and for gram+ species, lysostaphin will help). For more
exotic species with different cell wall compositions, different enzymes may be required.
A more universal method of lysis for genomic DNA extraction involves mechanically
breaking the cell wall. One method for this is bead beating, which can be easily
performed on a vortex using 0.1 mm glass beads or 0.15 mm fine garnet beads. Special
vortex adapters help with performing multiple extractions at the same time with equal
efficiency. Bead beating is faster than enzymatic lysis and generally more thorough.
2. …and purify
Once the sample has been lysed so bringing the genomic DNA into solution, all that is
needed is to purify the sample. This can be achieved using either phenol-chloroform or a
spin filter membranes by adding guanidine salts that promote binding to silica.
3. Some words of advice
The chromosome is going to break during purification because it is much too big to stay
in one piece. But for most applications this is not a problem and for PCR, the breakage
will be an advantage because it allows better melting the DNA and result in a more
efficient reaction.
The E.coli chromosome is 4,638, 858 bp long and this comes to roughly .005 picograms
per cell. In a typical overnight culture started from a single colony, the bacteria number
around 1-2×109 bacteria/ml. That means that 1 ml of culture should yield about 5 µg of
genomic DNA per 109 bacteria.
Plasmid DNA Extraction

Plasmid DNA extraction is a bit more complicated because it involves separating the
plasmid from the genomic DNA. The separation of the two forms of DNA is based on
size…
…and the trick is in the lysis method.
1. Alkaline Lysis
For plasmid DNA extraction, the lysis has to be a lot more subtle than simply chewing up
the cell wall with enzyme or bashing it with glass beads. The (virtually) universal method
for plasmid DNA extraction was invented by Birnboim and Doly in 1979 (and was
explained by Bitesize Bio in 2008!)
The lysis buffer contains sodium hydroxide and SDS, the purpose of which is to
completely denature of the plasmid and genomic DNA (i.e. separate the DNA into single
strands). It is critical that this step is performed quickly because too long in the
denaturing conditions of this solution may result in irreversibly denatured plasmid at the
end.
Next the sample is neutralized in a potassium acetate solution to renature the plasmid.
And this is the key to the separation of the plasmid and genomic DNA.
Because plasmid is small, it can easily re-anneal. But the genomic DNA is too long to reanneal properly and instead it becomes tangled so the complimentary strands stay
separated.
When the sample is centrifuged, the genomic DNA is still bound to protein and gets
pulled down while plasmid DNA is soluble and free. It is key at this step not to vortex or
mix the sample vigorously because the genomic DNA is easy to break, and broken
genomic DNA can be small enough to re-anneal and go into solution with the plasmid.
2. Purification
The plasmid DNA is recovered in the supernatant and can now be ethanol precipitated for
a crude prep or cleaned up using phenol-chloroform or a spin filter based prep. If you are
using a spin filter prep, the neutralization buffer will already contain guanidine salts so
the lysate can be bound directly onto silica for further washing and elution. The pure
DNA is fine for most everything from cloning to sequencing. If the plasmid is to be used
for transfection, anion-exchange purification is a better choice to remove the endotoxin,
although endotoxin removal is available using faster silica based purification also.
The method for purifying plasmids can also be used for mammalian plasmids transfected
in eukaryotic cells or for any other small extra-chromosomal DNA. The difference for
mammalian cells or chloroplast/mitochondrial DNA is that the copy numbers are much
smaller compared to the high copies of plasmid that can be obtained. So expect a lower
yield if you try the plasmid method on another type of DNA isolation or scale up your
buffer accordingly if you decide to start with more sample.
3. …and some words of advice.
Plasmid DNA is typically 3-5 kb and then the size is increased based on the insert. The
type of origin of replication will affect how high the copy number will be per cell. A
typical high copy number plasmid such as pUC or pBluescript should yield between 4-5
µg of DNA per ml of LB culture.
To isolate high yields of plasmid DNA, the culture should be in late log phase or early
stationary phase. Prepare cultures using fresh single colonies from plates and make sure
the antibiotic is fresh and the correct strength to maintain the plasmid during growth. It is

important not to overgrow the culture or it may result in genomic DNA contamination in
the plasmid prep.

Nucleic Acids Res. 1979 Nov 24;7(6):1513-23.
A rapid alkaline extraction procedure for screening recombinant plasmid DNA.
Birnboim HC, Doly J.
Abstract
A procedure for extracting plasmid DNA from bacterial cells is described. The method is
simple enough to permit the analysis by gel electrophoresis of 100 or more clones per day
yet yields plasmid DNA which is pure enough to be digestible by restriction enzymes.
The principle of the method is selective alkaline denaturation of high molecular weight
chromosomal DNA while covalently closed circular DNA remains double-stranded.
Adequate pH control is accomplished without using a pH meter. Upon neutralization,
chromosomal DNA renatures to form an insoluble clot, leaving plasmid DNA in the
supernatant. Large and small plasmid DNAs have been extracted by this method.

Plasmid DNA Isolation
Isolation of plasmid DNA from E. coli is a common routine in research laboratories. You
will perform a widely-practiced procedure that involves alkaline lysis of cells. This
protocol, often referred to as a plasmid “mini-prep,” yields fairly clean DNA quickly and
easily.

Procedure
1. Fill a microcentrifuge tube with saturated bacterial culture grown in LB broth +
antibiotic. Spin tube in microcentrifuge for 1 minute, and make sure tubes are
balanced in microcentrifuge. Dump supernatant and drain tube briefly on paper
towel.
2. Repeat step 1 in the same tube, filling the tube again with more bacterial culture.
The purpose of this step is to increase the starting volume of cells so that more
plasmid DNA can be isolated per prep. Spin tube in microcentrifuge for 1 minute.
Pour off supernatant and drain tube on paper towel.
3. Add 0.2 ml ice-cold Solution 1 to cell pellet and resuspend cells as much as
possible using disposable transfer pipet.
o Solution 1 contains glucose, Tris, and EDTA. Glucose is added to increase
the osmotic pressure outside the cells. Tris is a buffering agent used to
maintain a constant pH (= 8.0). EDTA protects the DNA from degradative
enzymes (called DNAses); EDTA binds divalent cations that are necessary
for DNAse activity.
4. Add 0.4 ml Solution 2, cap tubes and invert five times gently. Let tubes sit at
room temperature for 5 minutes.
o Solution 2 contains NaOH and SDS (a detergent). The alkaline mixture
ruptures the cells, and the detergent breaks apart the lipid membrane and

solubilizes cellular proteins. NaOH also denatures the DNA into single
strands.
5. Add 0.3 ml ice-cold Solution 3, cap tubes and invert five times gently. Incubate
tubes on ice for 10 minutes.
o Solution 3 contains a mixture of acetic acid and potassium acetate. The
acetic acid neutralizes the pH, allowing the DNA strands to renature. The
potassium acetate also precipitates the SDS from solution, along with the
cellular debris. The E. coli chromosomal DNA, a partially renatured tangle
at this step, is also trapped in the precipitate. The plasmid DNA remains in
solution.
6. Centrifuge tubes for 5 minutes. Transfer supernatant to fresh microcentrifuge tube
using clean disposable transfer pipet. Try to avoid taking any white precipitate
during the transfer. It is okay to leave a little supernatant behind to avoid
accidentally taking the precipitate.
o This fractionation step separates the plasmid DNA from the cellular debris
and chromosomal DNA in the pellet.
7. Fill remainder of centrifuge tube with isopropanol. Let tube sit at room
temperature for 2 minutes.
o Isopropanol effectively precipitates nucleic acids, but is much less
effective with proteins. A quick precipitation can therefore purify DNA
from protein contaminants.
8. Centrifuge tubes for 5 minutes. A milky pellet should be at the bottom of the tube.
Pour off supernatant without dumping out the pellet. Drain tube on paper towel.
o This fractionation step further purifies the plasmid DNA from
contaminants.
9. Add 1 ml of ice-cold 70% ethanol. Cap tube and mix by inverting several times.
Spin tubes for 1 minute. Pour off supernatant (be careful not to dump out pellet)
and drain tube on paper towel.
o Ethanol helps to remove the remaining salts and SDS from the preparation.
10. Allow tube to dry for ~5 minutes. Add 50 ul TE to tube. If needed, centrifuge tube
briefly to pool TE at bottom of tube. DNA is ready for use and can be stored
indefinitely in the freezer.
Solutions:
Solution 1:
per 500 ml:
50 mM glucose
9 ml 50% glucose
25 mM Tris-HCl pH 8.0 12.5 ml 1 M Tris-HCl pH 8.0
10 mM EDTA pH 8.0 10 ml 0.5 M EDTA pH 8.0
Add H2O to 500 ml.
Solution 2: per 500 ml:
1% SDS
50 ml 10% SDS
0.2 N NaOH 100 ml 1 N NaOH

Add H2O to 500 ml.
Solution 3: per 500 ml:
3 M K+
300 ml 5 M Potassium Acetate
5 M Acetate 57.5 ml glacial acetic acid

Add H2O to 500 ml.
TE
per 100 ml:
10 mM Tris-HCl pH 8.0 1 ml 1 M Tris-HCl pH 8.0
1 mM EDTA
0.5 ml 0.5 M EDTA pH 8.0
Add H2O to 100 ml.
Optional: RNAse can be added to TE at final concentration of 20 ug/ml.

Preparation of crude lysates
An easy technique for isolation of genomic DNA is to incubate cell lysates at high
temperatures (e.g., 90°C for 20 minutes), or to perform a proteinase K digestion, and then
use the lysates directly in downstream applications. Considered “quick-and-dirty”
techniques, these methods are only appropriate for a limited range of applications. The
treated lysate usually contains enzyme-inhibiting contaminants, such as salts, and DNA is
often not at optimal pH. Furthermore, incomplete inactivation of proteinase K can result
in false negative results and high failure rates. It is not recommended to store DNA
prepared using this method, as the high levels of contamination often result in DNA
degradation.
Salting-out methods
Starting with a crude lysate, ”salting-out” is another conventional technique where
proteins and other contaminants are precipitated from the cell lysate using high
concentrations of salt such as potassium acetate or ammonium acetate. The precipitates
are removed by centrifugation, and the DNA is recovered by alcohol precipitation.
Removal of proteins and other contaminants using this method may be inefficient, and
RNase treatment, dialysis, and/or repeated alcohol precipitation are often necessary
before the DNA can be used in downstream applications. DNA yield and purity are
highly variable using this method.

Organic extraction methods

Organic extraction is a conventional technique that uses organic solvents to extract
contaminants from cell lysates. The cells are lysed using a detergent, and then mixed with
phenol, chloroform, and isoamyl alcohol. The correct salt concentration and pH must be
used during extraction to ensure that contaminants are separated into the organic phase
and that DNA remains in the aqueous phase. DNA is usually recovered from the aqueous
phase by alcohol precipitation. This is a time-consuming and cumbersome technique.
Furthermore, the Methodology uses toxic compounds and may not give reproducible
yields. DNA isolated using this method may contain residual phenol and/or chloroform,
which can inhibit enzyme reactions in downstream applications, and therefore may not be
sufficiently pure for sensitive downstream applications such as PCR. The process also
generates toxic waste that must be disposed of with care and in accordance with
hazardous waste guidelines. In addition, this technique is almost impossible to automate,
making it unsuitable for high-throughput applications.
For all different sources to extract genomic and plasmid DNA extraction, the ready-touse kits are available on this site of Promega.
http://www.promega.com/a/dnaextraction/?gclid=CP2j1KDRpqsCFYUntAodcCoBzw

Isolation of DNA from Plant Tissue
Aim
To isolate DNA from plant tissue and to determine its purity
Principle
The cell wall must be broken or digested to release the cellular components. This is done
by grinding the tissue in liquid nitrogen with a pestle and mortar. Usually phenol
extraction yields a fairly pure DNA sample. This treatment however may not be sufficient
to give pure DNA if the cells also contain significant quantities of other biochemical like
the plant tissue which often contain large amounts of carbohydrates that are removed by
phenol extraction. DNA from the plant tissue is therefore usually obtained by using a
detergent either SDS or CTAB (cetyltrimethylammonium bromide). CTAB forms
insoluble complex with nucleic acids (Nucleic acid – CTAB complex) leaving behind
carbohydrates, protein and other contaminants in the supernatant. The insoluble nucleic
acid-CTAB complex precipitate is then collected by centrifugation and re-suspended.
EDTA in the lysis buffer inactivates endogenous nucleases by chelating Mg2+ ions. The
tissue – buffer mixture is emulsified with chloroform – phenol to denature and separate
remaining proteins from the DNA. The genomic DNA can be precipitated by using
absolute ethyl alcohol.
Materials
Microfuge tubes, centrifuge, oven, EDTA, Tris, CTAB or SDS, Chloroform, Ethanol,
isoamyl alcohol, isopropanol, sodium acetate buffer.
Methodology

Around 0.5 g of fresh leaf tissue or cauliflower was ground into powder in a pre-cooled
pestle and mortar. Exactly 9 ml of pre-heated CTAB extraction buffer was poured on to
the leaf powder in a polypropylene centrifuge tube. The mixture was gently inserted
several times and incubated for 60 minutes in an oven at 65°C. The tubes were gently
agitated during the incubation time at regular intervals. The tubes were cooled after
incubation period and 5 ml of chloroform – isoamyl alcohol (24:1) was added and mixed
gently for 5-10 min. The tubes were spun at 3000 rpm for 10 minutes. The layers were
clearly seen after centrifugation. The top aqueous phase was poured off into new tube and
re-extracted with chloroform-isoamyl alcohol.
To the aqueous phase 6 ml of cold isopropanol was added and inverted gently to
precipitate the nucleic acid.
The precipitate was dissolved with 1 ml of TE buffer and re-extracted using phenol –
chloroform (1:1) at 3000rpm for 10 minutes. The aqueous phase was transferred to a new
tube. About 1 ml of chloroform: isoamyl alcohol was added to the aqueous phase and
spun at 3000 rpm to separate the phase. To the aqueous phase 50 ml of 3M sodium
acetate and 2.5 ml of absolute ethanol was added to precipitate the DNA. The tube was
gently inverted for several times and incubated at 20°C for 20 minutes. The pellet in the
microfuge after centrifugation was collected with 70% ethanol. Ethanol was discarded
and the pellet was air-dried. Dry powdered form of DNA was collected as pellet was then
dissolved with 150 µl of TE buffer.
Estimation of the concentration of DNA
DNA has a maximum absorbance at 260nm. Based on the extinction co-efficient, an
optical density of 1.0 at 260 nm corresponds to 50µg/ml of double stranded DNA. The
ratio of O.D 260/280 provides the estimate of purity. A typically pure DNA has a ratio of
approximately 1.8. Ratio less than 1.8 indicates the probable presence of proteins or other
UV absorbers. Ratio higher than 2.0 indicates that the sample may be contaminated with
chloroform or phenol and should be reprecipitated with ethanol.
Methodology
To measure the concentration of DNA, the sample was diluted with TE buffer. The
spectrophotometer was “blanked” using TE buffer.
Absorbance of the sample at 260nm was recorded and the DNA concentration was
calculated from the following relation. Conc. of double stranded DNA = O.D (260) X 50
(µg/ml) X Dilution factor / 1 O.D (260)
Result:
White threads of DNA was precipitated and stored in TE buffer. The DNA was then
subjected to agarose gel electrophoresis and DNA band was visualized using ethidium
bromide.
Discussion
Organic extraction is a conventional technique that uses organic solvents to extract
contaminants from cell lysates. The cells are lysed using a detergent, and then mixed with
phenol, chloroform, and isoamyl alcohol. The correct salt concentration and pH must be

used during extraction to ensure that contaminants are separated into the organic phase
and that DNA remains in the aqueous phase. DNA is usually recovered from the aqueous
phase by alcohol precipitation. This is a time-consuming and cumbersome technique.
Furthermore, the Methodology uses toxic compounds and may not give reproducible
yields. DNA isolated using this method may contain residual phenol and/or chloroform,
which can inhibit enzyme reactions in downstream applications, and therefore may not be
sufficiently pure for sensitive downstream applications such as PCR.

Lab: DNA Extraction from Human Cheek Cells
Introduction
DNA…you hear about it all the time. DNA is used every day by scientists and lawyers to
help in criminal investigation, paternity suits, cloning, etc. Your DNA is your “genetic
fingerprint”—this means that your DNA is like no one else’s in the world! The procedure
that we will use to see your DNA includes the same basic processes that researchers use
to isolate, analyze, and manipulate DNA in a laboratory setting (although the DNA
isolated here is not nearly as “pure” as the research lab version).
If you remember back to Chapter 2, DNA is a nucleic acid, made of carbon, hydrogen,
oxygen, nitrogen, and phosphorous. DNA can be considered the hereditary “code of life”
because it possesses the information that determines an organism’s characteristic and is
transmitted from one generation to the next. You receive half of your genes from your
mother and half from your father. Day to day, DNA’s job is to direct the functioning
within the cells of your body.
DNA is in the nucleus of almost every cell in your body. The length of DNA per cell is
about 100,000 times as long as the cell itself. However, DNA only takes up about 10% of
the cell’s volume. This is because DNA is specially packaged through a series of events
to fit easily in the cell’s nucleus. The structure of DNA, the double helix, is wrapped
around proteins, folded back onto itself, and coiled into a compact chromosome.
Individual chromosomes can be studied using microscopes, but the double helix of a
chromosome is so thin that it only be detected through innovative, high-tech procedures.
Chromosomal DNA from a single cell is not visible to the naked eye. However, when
chromosomal DNA is extracted from multiple cells, the amassed quantity can easily be
seen and looks like strands of mucous-like, translucent cotton.
We will first collect cheek cells by swishing a sports drink in our mouths and using our
teeth to scrape cells off our cheeks. (The more vigorous and the longer that you swish, the
more cells are removed, and the more materials you’ll have from which to extract DNA.)
Then, we will lyse the cell membranes by adding a detergent based cell lysis solution,
which allows the DNA to be freed. DNA is soluble in water, but much less soluble in
alcohol. Thus, alcohol will be slowly added, and DNA will precipitate to the sports
drink/alcohol interface, and you will be able to see your own DNA! The white, stringy
material is thousands of DNA molecules stuck together (with some proteins too).

Materials and Methods
1. Label your 15mL test tube with a piece of tape and your initials
2. Obtain a small cup of sports drink and swish it around in your mouth for 1 full
minute. As you swish, gently and continuously scrape the sides of your cheeks
with your teeth to help release your cheek cells.
3. Spit the drink (with your collected cheek cells) back into the small cup.
4. Pour the contents of the cup into your labeled test tube (discard the cup).
5. Holding the test tube at an angle, use the provided plastic pipet to add 2mL of cell
lysis solution to your collected cheek cells.
6. Cap your test tube, and invert it 5-8 times. (This mixes the lysis solution with the
cheek cells.)
7. Allow this to stand for 2 minutes.
8. Using the provided pipet, add the cold alcohol by letting it run gently run down
the side of the test tube (hold the test tube at an angle). Add the alcohol until your
total volume reaches 12-13mL. You should have 2 distinct layers. DO NOT mix
the cheek cell solution with the alcohol!!!
9. Watch as wispy strands of translucent DNA begin to clump together where the
alcohol layer meets the cheek cell solution. (It kind of looks like cobwebs
extending upward.)
10. Place your 15mL test tube in a test tube rack and let it stand undisturbed for 15
minutes. During this time the DNA will continue to precipitate out.
11. Optional—use a plastic pipet to transfer your DNA into a smaller test tube. To do
so, place the pipet near the DNA and draw the DNA into the pipet (along with
some alcohol). Do not move your pipet up and down into the bottom layer.

Summary
In order to understand what you are doing in this activity, it is
important that you know the “big picture” behind the methods we will be
using:
•

Cells may be physically and chemically treated to break open the outer cell
membrane and inner nuclear membrane.

•

The portion of the cell mixture containing DNA (the watery portion) will be
separated from the cell membranes and organelles (the gloppy portion).

•

The solution containing dissolved DNA will be chemically altered so that the
DNA can precipitate out of the solution in its solid, string-like state.

Additional Notes
• The recipe for the Salt/detergent mixture is: 2 L distilled water, 100 mL detergent
(we use Palmolive dishwashing detergent), 15 g salt.
•

The ethanol needs to be ice cold–keep in freezer until the time it is needed.

•

There is about a 15-minute wait time–plan something DNAish during this time.

Anal Biochem. 1987 Apr;162(1):156-9.
Single-step method of RNA isolation by acid guanidinium thiocyanate-phenolchloroform extraction.
Chomczynski P, Sacchi N.
Abstract
A new method of total RNA isolation by a single extraction with an acid guanidinium
thiocyanate-phenol-chloroform mixture is described. The method provides a pure
preparation of undegraded RNA in high yield and can be completed within 4 h. It is
particularly useful for processing large numbers of samples and for isolation of RNA
from minute quantities of cells or tissue samples
Since its introduction, the ‘single-step’ method has become widely used for isolating total
RNA from biological samples of different sources. The principle at the basis of the
method is that RNA is separated from DNA after extraction with an acidic solution
containing guanidinium thiocyanate, sodium acetate, phenol and chloroform, followed by
centrifugation. Under acidic conditions, total RNA remains in the upper aqueous phase,
while most of DNA and proteins remain either in the interphase or in the lower organic
phase. Total RNA is then recovered by precipitation with isopropanol and can be used for
several applications. The original protocol, enabling the isolation of RNA from cells and
tissues in less than 4 hours, greatly advanced the analysis of gene expression in plant and
animal models as well as in pathological samples, as demonstrated by the overwhelming
number of citations the paper gained over 20 years.

Phenol–chloroform extraction (abbreviated PC or PCIA, see reagents below) is a
liquid–liquid extraction technique in biochemistry. It is widely used in molecular biology
for isolating DNA, RNA and protein. Equal volumes of a phenol:chloroform mixture and
an aqueous sample are mixed, forming a biphasic mixture.
It was originally devised by Piotr Chomczynski and Nicoletta Sacchi and published in
1987 (referred to as Guanidinium thiocyanate-phenol-chloroform extraction).[1][2] The
reagent used specifically for RNA extraction is sold by Sigma-Aldrich by the name TRI
Reagent, by Invitrogen under the name TRIzol, and by Bioline as Trisure.
Guanidinium thiocyanate denatures proteins, including RNases, and separates rRNA
from ribosomes, while phenol, isopropanol and water are solvents with poor solubility. In
the presence of chloroform or BCP (bromochloropropane), these solvents separate
entirely into two phases that are recognized by their color: a clear, upper aqueous phase
(containing the nucleic acids) and a bright pink lower phase (containing the proteins
dissolved in phenol and the lipids dissolved in chloroform). Other denaturing chemicals
such as 2-mercaptoethanol and sarcosine may also be used. The major downside is that
phenol and chloroform are both hazardous and inconvenient materials, and the extraction
is often laborious, so in recent years many companies now offer alternative ways to
isolate DNA

•

•

•

Phenol: The phenol used for biochemistry comes as a water-saturated solution
with Tris buffer, as a Tris-buffered 50% phenol, 50% chloroform solution, or as a
Tris-buffered 50% phenol, 48% chloroform, 2% isoamyl alcohol solution
(sometimes called “25:24:1”). Phenol is naturally somewhat water-soluble, and
gives a fuzzy interface, which is sharpened by the presence of chloroform, and the
isoamyl alcohol reduces foaming. Most solutions also have an antioxidant, as
oxidized phenol damages the nucleic acids. For RNA purification, the pH is kept
around pH 4, which retains RNA in the aqueous phase preferentially. For DNA
purification, the pH is usually near 7, at which point all nucleic acids are found in
the aqueous phase.
Chloroform: Chloroform is stabilized with small quantities of amylene or ethanol,
because exposure of pure chloroform to oxygen and ultraviolet light produces
phosgene gas. Some chloroform solutions come as pre-made a 96% chloroform,
4% isoamyl alcohol mixtures that can be mixed with an equal volume of phenol to
obtain the 25:24:1 solution.
Isoamyl alcohol: Isoamyl alcohol may reduce foaming and ensure deactivation of
RNase.

Diethyl pyrocarbonate (DEPC)
Diethylpyrocarbonate (DEPC), is used in the laboratory to inactivate the RNase
enzymes from water and other laboratory utensils. It inactivates the RNases by the
covalent modifications of the histidine residues. DEPC is unstable in water and
susceptible to hydrolysis to carbon dioxide and ethanol, especially in the presence of a
nucleophile. For this reason DEPC cannot be used with Tris buffer or HEPES. In contrast
it can be used with PBS or MOPS. A handy rule is that enzymes or chemicals which have
active -O:, -N: or -S: cannot be treated with DEPC to become RNase-free as DEPC reacts
with these species. Furthermore DEPC degradation products can inhibit in vitro
transcription.

1. Autoclaving is not effective at eliminating RNase in solution because the RNases
simply renature as the solution cools.
FALSE, but… Autoclaving alone does indeed inactivate a substantial amount of RNase A.
2. Autoclaving inactivates DEPC.
TRUE. Autoclaving does inactivate DEPC by causing hydrolysis of
diethylpyrocarbonate. CO2 and EtOH are released as reaction by-products. DEPC has a
half-life of approximately 30 minutes in water, and at a DEPC concentration of 0.1%,
solutions autoclaved for 15 minutes/liter can be assumed to be DEPC-free.
3. Solutions containing Tris cannot be treated with DEPC.
TRUE. Tris contains an amino group which “sops up” DEPC and makes it unavailable to
inactivate RNase
4. 0.1% DEPC is sufficient to inhibit any amount of RNase in a solution.
FALSE, but… The amount of DEPC required to inactivate RNase increases as the
amount of contaminating RNase in a solution increases
5. If 0.1% DEPC works well to inhibit RNase, 1% should work even better.
TRUE, but… Increasing DEPC concentrations inactivate increasing amounts of RNase A
contamination (Figure 3). However, it is also true that high levels of residual DEPC or
DEPC by-products in a solution can inhibit some enzymatic reactions or chemically alter
(carboxymethylate) RNA

The RNeasy principle and procedure
The RNeasy procedure represents a novel technology for RNA isolation. This technology
combines the selective binding properties of a silica-gel-based membrane with the speed
of microspin technology. A specialized high-salt buffer system allows up to 100 µg of
RNA longer than 200 bases to bind to the RNeasy silica-gel membrane. Biological
samples are first lysed and homogenized in the presence of a highly denaturing guanidine
isothiocyanate (GITC)-containing buffer, which immediately inactivates RNases to
ensure isolation of intact RNA. Ethanol is added to provide appropriate binding
conditions, and the sample is then applied to an RNeasy mini column where the total
RNA binds to the membrane and contaminants are efficiently washed away. High-quality
RNA is then eluted in 30 µl, or more, of water.
With the RNeasy procedure, all RNA molecules longer than 200 nucleotides are isolated.
The procedure provides an enrichment for mRNA since most RNAs primer extension, RNase and S1 nuclease protection, cDNA synthesis, differential
display, expression-array and expression-chip analysis.
RNA stabilization in tissue
RNA stabilization is an absolute prerequisite for reliable gene-expression analysis.
Immediate stabilization of RNA in biological materials is necessary because, directly
after harvesting the biological sample, changes in the gene-expression pattern occur due
to specific and nonspecific RNA degradation as well as to transcriptional induction. Such
changes in gene-expression pattern need to be avoided for all reliable quantitative geneexpression analyses, such as biochip and array analyses, quantitative RT-PCR, or other
nucleic acid-based technologies.
The RNeasy principle and procedure
The RNeasy procedure represents a novel technology for RNA isolation. This technology
combines the selective binding properties of a silica-gel-based membrane with the speed
of microspin technology. A specialized high-salt buffer system allows up to 100 µg of
RNA longer than 200 bases to bind to the RNeasy silica-gel membrane. Biological
samples are first lysed and homogenized in the presence of a highly denaturing guanidine
isothiocyanate (GITC)-containing buffer, which immediately inactivates RNases to
ensure isolation of intact RNA. Ethanol is added to provide appropriate binding
conditions, and the sample is then applied to an RNeasy mini column where the total
RNA binds to the membrane and contaminants are efficiently washed away. High-quality
RNA is then eluted in 30 µl, or more, of water. With the RNeasy procedure, all RNA
molecules longer than 200 nucleotides are isolated. The procedure provides an
enrichment for mRNA since most RNAs cleared lysate, creating conditions which promote selective binding of RNA to the
RNeasy silica-gel membrane. The sample is then applied to the RNeasy mini column.
Total RNA binds to the membrane, contaminants are efficiently washed away, and highquality RNA is eluted in water. RNeasy Plant Mini Kits can also be used for total RNA
minipreparation from animal cells and tissues, bacteria, and yeast.
RNA Cleanup
The RNeasy Plant Mini Kit can be used to purify RNA from enzymatic reactions (e.g.,
DNase digestion, RNA labeling) or for desalting RNA samples (maximum 100 µg RNA).
GITC-containing lysis buffer and ethanol are added to the sample to create conditions
that promote selective binding of RNA to the RNeasy membrane. The sample is then
applied to the RNeasy mini column. RNA binds to the RNeasy silica-gel membrane,
contaminants are efficiently washed away, and high quality RNA is eluted in water.
Disruption and homogenization of starting materials
Efficient disruption and homogenization of the starting material is an absolute
requirement for all total RNA isolation procedures. Disruption and homogenization are
two distinct steps.
Disruption: Complete disruption of cells walls and plasma membranes of cells and
organelles is absolutely required to release all the RNA contained in the sample. Different
samples require different methods to achieve complete disruption. Incomplete disruption
results in significantly reduced yields.
For disruption using a mortar and pestle, freeze the sample immediately in liquid nitrogen
and grind to a fine powder under liquid nitrogen. Transfer the suspension (tissue powder
and liquid nitrogen) into a liquid-nitrogen-cooled, appropriately sized tube and allow the
liquid nitrogen to evaporate without allowing the sample to thaw. Add lysis buffer and
continue as quickly as possible with the homogenization (below).
Homogenization: Homogenization is necessary to reduce the viscosity of the cell lysates
produced by disruption. Homogenization shears the high-molecular weight genomic
DNA and other high-molecular-weight cellular components to create a homogeneous
lysate. Incomplete homogenization results in inefficient binding of RNA to the RNeasy
membrane and therefore significantly reduced yields.
Use of QIAshredder modules is a fast and efficient way to homogenize cell and tissue
lysates without cross contamination of the samples. The lysate (maximum volume 700
µl) is loaded onto the QIAshredder spin column sitting in a 2 ml collection tube, spun for
2 min at maximum speed in a microfuge and the homogenized lysate collected.
QIAshredder spin columns are supplied in the RNeasy Plant Mini Kits.
Determining the amount of the starting material
It is essential to begin with the correct amount of plant material in order to obtain optimal
RNA yield and purity with RNeasy columns. A maximum of 100 mg plant material or 1
x 107 cells can generally be processed with RNeasy mini columns. For most plant
material, the binding capacity of the column (100 µg RNA) and the lysing capacity of
Buffer RLT will not be exceeded by these amounts.

If you have no information about the nature of your starting material, we recommend
starting with no more than 50 mg of plant material or 3–4 x 106 cells. Depending on the
yield and purity obtained, it may be possible to increase the amount of plant material to
100 mg or to increase the cell number to 1 x 107 in subsequent preparations. Do not
overload the column. Overloading will significantly reduce yield and quality.
Important notes before starting
Fresh or frozen tissue can be used. To freeze tissue for long-term storage, flash-freeze in
liquid nitrogen, and immediately transfer to –70°C. Tissue can be stored for several
months at –70°C. To process, do not allow tissue to thaw during weighing or handling
prior to disruption in Buffer RLT. Homogenized lysates (in Buffer RLT, step 4) can also
be stored at –70°C for several months. To process frozen lysates, thaw samples and
incubate for 15–20 min at 37°C in a water bath to dissolve salts. Continue with step 5.
The RNeasy Plant Mini Kit provides two different lysis buffers, Buffer RLT and Buffer
RLC, which contain guanidine isothiocyanate (GITC) and guanidine hydrochloride,
respectively. In most cases, Buffer RLT is the lysis buffer of choice due to the greater
cell disruption and denaturation properties of GITC. However, depending on the amount
and type of secondary metabolites in some tissues (such as milky endosperm of maize or
mycelia of filamentous fungi), GITC can cause solidification of the sample, making
extraction of RNA impossible. In these cases, Buffer RLC should be used.
β-Mercaptoethanol (β-ME) must be added to Buffer RLT or Buffer RLC before use. β-ME
is toxic; dispense in a fume hood and wear appropriate protective clothing. Add 10 µl βME per 1 ml Buffer RLT. Buffer RLT is stable for 1 month after addition of β-ME. Buffer
RPE is supplied as a concentrate. Before using for the first time, add 4 volumes of
ethanol (96–100%), as indicated on the bottle, to obtain a working solution.
Generally, DNase digestion is not required since the RNeasy silica-membrane technology
efficiently removes most of the DNA without DNase treatment. However, further DNA
removal may be necessary for certain RNA applications that are sensitive to very small
amounts of DNA (e.g., TaqMan RT-PCR analysis with a low-abundant target). In these
cases, the small residual amounts of DNA remaining can be removed using the RNaseFree DNase Set for the optional on-column DNase digestion or by a DNase digestion
after RNA isolation.
Buffer RLT may form a precipitate upon storage. If necessary, redissolve by warming,
and then place at room temperature.
Buffer RLT, Buffer RLC, and Buffer RW1 contain a guanidine salt and are therefore not
compatible with disinfecting reagents containing bleach. Guanidine is an irritant. Take
appropriate safety measures and wear gloves when handling.
All steps of the RNeasy protocol should be performed at room temperature. During the
procedure, work quickly.
All centrifugation steps are performed at 20–25°C in a standard microcentrifuge. Ensure
that the centrifuge does not cool below 20°C.

Flowchart of RNA Extraction using RNeasy Kit

NucleoSpin® technology and products
These days, most labs use commercial kits, which employ spin columns, for the isolation
of nucleic acids. The spin columns contain a silica resin that selectively binds DNA/RNA,
depending on the salt conditions and other factors influenced by the extraction method.
These kits make the whole process much easier and faster than the methods of old, when
things are going well, but the downside of using a kit is that we don’t always know what
is in the mysterious and proprietary set of solutions that each company uses in its kit,
which makes troubleshooting more difficult.
Lysis
The lysis formulas may vary based on the whether you want DNA or RNA, but the
common denominator is a lysis buffer containing a high concentration of chaotropic salt.
Chaotropes destabilize hydrogen bonds, van der Waals forces, and hydrophobic
interactions. Proteins are destabilized, including nucleases, and the association of nucleic
acids with water is disrupted setting up the conditions for the transfer to silica.
Chaotropic salts include guanidine HCL, guanidine thiocyanate, urea, and lithium
perchlorate.
Besides the chaotropes, there is usually some detergents involved, to help with protein
solubilization and lysis. There can also be enzymes used for lysis depending on the
samples type. Proteinase K is one of these, and actually works very well in these
denaturing buffers; the more denatured the protein, the better Proteinase K works.

Binding:
The chaotropic salts are critical for lysis, but also for binding, as we discussed.
Additionally, to enhance and influence the binding of nucleic acids to silica, alcohol is
also added. Most of the time this is ethanol but sometimes it may be isopropanol. The
percent ethanol and the volume have big effects. Too much and you’ll bring in a lot
of degraded nucleic acids and small species that will influence UV260 readings and
throw off some of your yields. Too little, and it may become difficult to wash away all of
the salt from the membrane.
The important point here is that the ethanol influences binding and the amount added is
optimized for whatever kit you are using. Modifying that step can help change what you
recover so if you are having problems and want to troubleshoot recovery that can be a
step to evaluate further.
Another way to diagnose problems is to save the flow-through after binding and
precipitate it to see if you can find the nucleic acids you are searching for. If you used an
SDS-containing detergent in lysis, try using NaCl as a precipitant to avoid contamination
of the DNA or RNA with detergent.

Fig. Binding and elution mechanism of Nucleic acids

Washing Steps:
Your lysate was centrifuged through the silica membrane and now your DNA or RNA
should be bound to the column and the impurities, protein and polysaccharides, should
have passed through. But, the membrane is still dirty with residual proteins and salt. If
the sample was from plants, there will still be polysaccharides, maybe some pigments too,
left on the membrane, or if the sample was blood, the membrane might be tinted brown or
yellow.
The wash steps serve to remove these impurities. There are typically two washes,
although this can vary depending on the sample type. The first wash will often have a low
amount of chaotropic salt to remove the protein and colored contaminants. This is always
followed with an ethanol wash to remove the salts. If the prep is something that didn’t
have a lot of protein to start, such as plasmid preps or PCR clean up, then only an ethanol
wash is needed.
Removal of the chaotropic salts is crucial to getting high yields and purity DNA or RNA.
Some kits will even wash the column with ethanol twice. If salt remains behind, the
elution of nucleic acid is going to be poor, and the A230 reading will be high, resulting in
low 260/230 ratios.
Dry Spin:
After the ethanol wash, most protocols have a centrifugation step to dry the column. This
is to remove the ethanol and is essential for a clean eluant. When 10 mM Tris buffer or
water is applied to the membrane for elution, the nucleic acids can become hydrated and
will release from the membrane. If the column still has ethanol on it, then the nucleic
acids cannot be fully rehydrated.
Elution:
The final step is the release of pure DNA or RNA from the silica. For DNA preps, 10
mM Tris at a pH between 8-9 is typically used. DNA is more stable at a slightly basic pH
and will dissolve faster in a buffer. This is true even for DNA pellets. Water tends to have
a low pH, as low as 4-5 and high molecular weight DNA may not completely rehydrate
in the short time used for elution. Elution of DNA can be maximized by allowing the
buffer to sit in the membrane for a few minutes before centrifugation.
RNA, on the other hand, is fine at a slightly acidic pH and so water is the preferred
diluent. RNA dissolves readily in water.
What other things can go wrong:
Low yields: If you experience yields lower than you expected for a sample, there are
many factors to think about. Usually it is a lysis problem. Incomplete lysis is a major
cause of low yields. It could also be caused by incorrect binding conditions. Make sure
to use fresh high quality ethanol (100% 200 proof) to dilute buffers or for adding to the
binding step. Low quality ethanol or old stocks may have taken on water and not be the
correct concentration. If the wash buffer is not made correctly, you may be washing off
your DNA or RNA.

Low Purity: If the sample is contaminated with protein (low 260/280) then maybe you
started with too much sample and the protein was not completely removed or dissolved.
Degradation: This is more of a concern for RNA preps. Mainly with RNA, degradation
occurs from inproper storage of the sample or an inefficient lysis, assuming of course that
you eluted with RNase-free water. For DNA preps, degradation is not a huge problem
because for PCR, the DNA can be sheared and it works fine. But if you were hoping to
not have so much sheared DNA, then you may have used too strong a lysis method.
The term “chaotropic” means chaos-forming, a term which in biochemistry usually refers
to a compound’s ability to disrupt the regular hydrogen bond structures in water.
Hydrogen bonding profoundly affects the secondary structure of polymers such as DNA,
RNA, and proteins, as well as how water-soluble a molecule is. Under native conditions,
nucleic acids are covered by a hydrate shell consisting of water molecules that maintain
the solubility of DNA in aqueous solutions. With the addition of chaotropic ions to the
nucleic acid, this relatively ordered structure of water molecules of the hydrate shell is
destroyed. The chaotropic salts create a hydrophobic environment. Under these
hydrophobic conditions, the silica membrane of the NucleoSpin columns is the most
suitable binding partner for the nucleic acids. Proteins, metabolites, and other
contaminants do not bind to the membrane and therefore are washed away during the
subsequent washing steps. As a further feature of the chaotropic salts, the respective
cations saturate the silica membrane with positive charges, which still improves the
binding of nucleic acids under hydrophobic conditions. Chaotropic salts increase the
solubility of nonpolar substances in water. They denature proteins because they have the
ability to disrupt hydrophobic interactions. They do not denature DNA or RNA. Their
function in the NucleoSpin Extraction Kit is to denature cellular proteins (such as DNase
and RNase). The high concentration of salt also facilitates binding of the nucleic acids
DNA and RNA to the silica membrane in the column.

Effect of pH on nueleic acids
The ribose 2′-OH group of RNA is absent in DNA. Consequently, the ubiquitous 3′-O of
polynucleotide backbones lacks a vicinal hydroxyl neighbor in DNA. This difference
leads to a greater resistance of DNA to alkaline hydrolysis. To view it another way, RNA
is less stable than DNA because its vicinal 2′-OH group makes the 3′-phosphodiester
bond susceptible to nucleophilic cleavage (Figure 1). For just this reason, it is selectively
advantageous for the heritable form of genetic information to be DNA rather than RNA.
Hydrolysis of Nucleic Acids
Most reactions of nucleic acid hydrolysis break bonds in the polynucleotide backbone.
Such reactions are important because they can be used to manipulate these polymeric
molecules. For example, hydrolysis of polynucleotides generates smaller fragments
whose nucleotide sequence can be more easily determined.
Hydrolysis by Acid or Base

RNA is relatively resistant to the effects of dilute acid, but gentle treatment of DNA with
1 mM HCl leads to hydrolysis of purine glycosidic bonds and the loss of purine bases
from the DNA. The glycosidic bonds between pyrimidine bases and 2′-deoxyribose are
not affected, and, in this case, the polynucleotide’s sugar-phosphate backbone remains
intact. The purine-free polynucleotide product is called apurinic acid.
Depurination is a DNA alteration in which the hydrolysis of a purine base (Adenine or
Guanine) from the deoxyribose-phosphate backbone occurs. After a depurination, the
sugar phosphate backbone remains and the sugar ring has a hydroxyl (-OH) group in the
place of the Adenine or Guanine. Around 1,000 Purines are lost this way each day in a
typical mammalian cell. One of the main causes of depurination is the presence of
endogenous metabolites in cell undergoing chemical reactions. This breaks the bond
linking the purine with the pentose sugar.

DNA is not susceptible to alkaline hydrolysis. On the other hand, RNA is alkali labile
and is readily hydrolyzed by dilute sodium hydroxide. Cleavage is random in RNA, and
the ultimate products are a mixture of nucleoside 2′- and 3′-monophosphates. These
products provide a clue to the reaction mechanism (Figure 1). Abstraction of the 2′-OH
hydrogen by hydroxyl anion leaves a 2′-O2 that carries out a nucleophilic attack on the δ+
phosphorus atom of the phosphate moiety, resulting in cleavage of the 5′-phosphodiester
bond and formation of a cyclic 2′,3′-phosphate. This cyclic 2′,3′-phosphodiester is
unstable and decomposes randomly to either a 2′- or 3′-phosphate ester. DNA has no 2’OH; therefore DNA is alkali stable.

Figure 1.The vicinal OH groups of RNA are susceptible to nucleophilic attack leading to
hydrolysis of the phosphodiester bond and fracture of the polynucleotide chain; DNA
lacks a 2′-OH vicinal to its 3′-O-phosphodiester backbone. Alkaline hydrolysis of RNA
results in the formation of a mixture of 2′- and 3′-nucleoside monophosphates.

Helping lecture tips
A. Laminar flow hoods. There are two types of laminar flow hoods, vertical and
horizontal. The vertical hood, also known as a biology safety cabinet, is best for working
with hazardous organisms since the aerosols that are generated in the hood are filtered out
before they are released into the surrounding environment. Horizontal hoods are designed
such that the air flows directly at the operator hence they are not useful for working with
hazardous organisms but are the best protection for your cultures. Both types of hoods
have continuous displacement of air that passes through a HEPA (High Efficiency
Particle Air) filter that removes particulates from the air. In a vertical hood, the filtered
air blows down from the top of the cabinet; in a horizontal hood, the filtered air blows out
at the operator in a horizontal fashion. NOTE: these are not fume hoods and should not
be used for volatile or explosive chemicals. They should also never be used for bacterial
or fungal work. The hoods are equipped with a short-wave UV light that can be turned on
for a few minutes to sterilize the surfaces of the hood, but be aware that only exposed
surfaces will be accessible to the UV light. Do not put your hands or face near the hood
when the UV light is on as the short wave light can cause skin and eye damage. The
hoods should be turned on about 10-20 minutes before being used. Wipe down all
surfaces with ethanol before and after each use. Keep the hood as free of clutter as
possible because this will interfere with the laminar flow air pattern.
➢ EDTA is a chelating agent and has great affinity with metal ions and Mg-ion present
in DNase as a cofactor and responsible for DNase action that degrade the DNA, here
EDTA bind with Mg-ion and nullify the action of DNase.
➢ Focusing material is DNA, to extract the DNA from Human, BLOOD must be in
preserved form and must not be coagulated so as to perform subsequent extraction
steps.
So
to
prevent
Blood
Clotting
EDTA
is
used.
It behaves as chelating agent for divalent ions (Ca++, Mg++).
The question is that, why these ions are needed to be captured. Simple is that Ca++
ions are initiator of Blood Clotting and behave as cofactor of clotting enzymes, so it
must
be
captured
by
EDTA.
Hence EDTA prevents blood Coagulation (Clotting) by chelating these ions.
Divalent ions are also co-factors of DNAase enzymes (that degrade DNA).
So
EDTA
also
prevents
DNAase
activity
by
chelating
ions.
➢ Chelation is the formation or presence of two or more separate coordinate bonds
between a polydentate (multiple bonded) ligand and a single central atom.[1] Usually
these ligands are organic compounds, and are called chelants, chelators, chelating
agents, or sequestering agents.

➢ SDS is an anionic detergent which disrupts cell membrane and destabilizes all
hydrophobic interactions holding macromolecules in their native form. It lyses the
cell membrane and nucleus to make the extraction possible.
➢ Sucrose provides Osmotic Shock to the Blood cells (leukocytes). By adding sucrose
to the solution, it absorbs (osmosis) glucose from the cell; hence cell is shrinked and
ruptured.
So
nucleus
is
available
for
extraction.
➢ Lysozyme is an enzyme that is used during DNA extraction to degrade the cell wall.
➢ NaCl stabilizes the double helical structure of DNA. It also provides Na+ that
neutralizes the negative charge present on DNA. Due to the negative charge the DNA
molecules repel each other. When the charge is neutralized the DNA molecule come
together and could be precipitated.
➢ TAE (Tris-acetate-EDTA) buffer is used as both a running buffer and in agarose
gel.[3] Its use in denaturing gradient gel electrophoresis methods for broad-range
mutation analysis has also been described.[4] TAE has been used at various
concentrations to study the mobility of DNA in solution with and without sodium
chloride.[5] However, high concentration of sodium chloride (and many other salts) in
a DNA sample retards its mobility. This may lead to incorrect interpretations of the
resulting DNA banding pattern.
➢ Compared with TBE buffer, TAE buffer offers advantages in subsequent enzymatic
applications for the DNA sample. For example, if a DNA sample is going to be used
in a cloning experiment, the step that follows its running on an agarose gel is to ligate
(covalently link) to a cloning vector (most likely a plasmid). DNA sample from TAE
Buffer is suitable for this purpose, while DNA from TBE buffer is not. Borate in the
TBE buffer is a strong inhibitor for many enzymes. This enzyme inhibiting property
made TBE buffer very popular in its realm for two reasons. First, a DNA sample run
in a TBE buffer can better keep its integrity. The other main reason is that the purpose
for many agarose gels electrophoreses is to analyze the size of DNA molecules.
➢ Extraction with phenol and phenol/chloroform mixtures is a universal method for
purification of DNA and RNA. Proteins and restriction enzymes are removed by
phenol and chloroform in disrupting protein secondary structure causing proteins to
denature and precipitate from solution. Although each of these solvents is capable of
performing this function alone, the two materials together remove proteins from
solution much more effectively. Nucleic acids are recovered in the liquid phase.
➢ During phenol extraction, the precipitated proteins collect at an interphase between
the aqueous and organic layers. It is very important to understand the effect of pH on
the
performance
of
phenol.
For the purification of DNA, the use of phenol at pH above 7.0 is desirable in order to
collect DNA in the upper aqueous layer of a phenol extraction (phenol saturated,
pH8). At lower pH (4.5), DNA and proteins are collected at the interphase although
RNA can be collected in the aqueous phase (phenol saturated, pH4.5).

➢ Phenol – helps in removing protein impurity from the dna, so we can get pure dna.
➢ Chloroform – prevent shearing of dna during isolation.
➢ Ethanol is a dehydrating agent. In DNA it is used for the precipitation of DNA
molecule. When a molecule is to be precipitated it should be dehydrated as the water
molecules forming a film around it prevent their interaction. In DNA isolation the
DNA molecules are dehydrated by ethanol.
➢ The lysozyme is used to degrade the cellular cell wall of the bacteria.
➢ C-TAB is a detergent that helps lyse the cell membrane, however it is pretty poor
with denaturing proteins so something with a longer tail is usually used for extraction.
➢ Sodium ions neutralize the negative charge of the DNA backbone (phosphates)
making the DNA less hydrophilic (less likely to be solubilized) in the wash solution.
➢ Glucose is added to increase the osmotic pressure outside the cells. glucose should
also be added to maintain osmolarity and prevent the buffer from bursting the cells.
➢ Glacial acetic acid is used to neutralize the alkaline pH of the buffers.
➢ In plasmid isolation RNA behaves as an unwanted material so to separate it out
RNAase is required which breaks down the RNA. This is done to get pure quality of
the product.
➢ For RNA purification, the pH is kept around pH 4, which retains RNA in the aqueous
phase preferentially. For DNA purification, the pH is usually near 7, at which point
all nucleic acids are found in the aqueous phase.
➢ Isoamyl alcohol: Isoamyl alcohol may reduce foaming and ensure deactivation of
Rnase
➢ Lysis, or breaking open the cells, is the first step of DNA extraction. This is
accomplished by a buffer containing tris and EDTA (ethylene diamine tetraacetic
acid). EDTA binds divalent cations such as calcium and magnesium. Since these ions
help maintain the integrity of the cell membrane, eliminating them with EDTA
destabilizes the membrane. Tris is the main buffering component; its chief role is to
maintain the pH of the buffer at a stable point, usually 8.0. Additionally, tris likely
interacts with the LPS (lipopolysaccharide) in the membrane, serving to destabilize
the membrane further.
➢ DNA is a polar molecule due to the negative charges on it’s phosphate backbone, so it
is
very
soluble
in
water.
➢ Lysogeny broth (LB), a nutritionally rich medium, is primarily used for the growth
of bacteria. The acronym has been incorrectly interpreted as Luria broth, Lennox
broth, or Luria-Bertani medium; according to its creator Giuseppe Bertani, the
abbreviation LB was actually intended to stand for lysogeny broth. Measure out the
following:
➢ 10 g tryptone
➢ 5 g yeast extract
➢ 10 g NaCl
➢ Suspend the solids in ~800 ml of distilled or deionized water.
➢ Add further distilled or deionized water, in a measuring cylinder to ensure accuracy,
to make a total of 1 litre.

➢ Autoclave at 121 °C.
➢ After cooling, swirl the flask to ensure mixing, and the LB is ready for use.

PCR Primer Design Guidelines
PCR (Polymerase Chain Reaction)

Polymerase Chain Reaction is widely held as one of the most important inventions of the
20th century in molecular biology. Small amounts of the genetic material can now be
amplified to be able to a identify, manipulate DNA, detect infectious organisms,
including the viruses that cause AIDS, hepatitis, tuberculosis, detect genetic variations,
including mutations, in human genes and numerous other tasks.

PCR involves the following three steps: Denaturation, Annealing and Extension. First, the genetic

material is denatured, converting the double stranded DNA molecules to single strands.
The primers are then annealed to the complementary regions of the single stranded
molecules. In the third step, they are extended by the action of the DNA polymerase. All
these steps are temperature sensitive and the common choice of temperatures is 94oC,
60oC and 70oC respectively. Good primer design is essential for successful reactions. The
important design considerations described below are a key to specific amplification with
high yield.

1. Primer Length: It is generally accepted that the optimal length of PCR primers is 1835 bp. This length is long enough for adequate specificity and short enough for primers to
bind easily to the template at the annealing temperature.

2. Primer Melting Temperature: Primer Melting Temperature (Tm) by definition is the
temperature at which one half of the DNA duplex will dissociate to become single
stranded and indicates the duplex stability. Primers with melting temperatures in the
range of 55-58 oC generally produce the best results. Primers with melting temperatures
above 65oC have a tendency for secondary annealing. The GC content of the sequence
gives a fair indication of the primer Tm. For primer melting temperature, the neighbor
thermodynamic theory, accepted as a much superior method for estimation, which is
considered the most recent and best available.

Formula for primer Tm calculation:
Melting Temperature Tm(oK)={ΔH/ ΔS + R ln(C)}, Or Melting Temperature Tm(oC) =
{ΔH/ ΔS + R ln(C)} – 273.15 where
ΔH (kcal/mole) : H is the Enthalpy. Enthalpy is the amount of heat energy
possessed by substances. ΔH is the change in Enthalpy. In the above

formula the ΔH is obtained by adding up all the di-nucleotide pairs
enthalpy values of each nearest neighbor base pair.
ΔS (kcal/mole) : S is the amount of disorder a system exhibits is called
entropy. ΔS is change in Entropy. Here it is obtained by adding up all the
di-nucleotide pairs entropy values of each nearest neighbor base pair. An
additional salt correction is added as the Nearest Neighbor parameters were
obtained from DNA melting studies conducted in 1M Na+ buffer and this
is the default condition used for all calculations.
ΔS (salt correction) = ΔS (1M NaCl )+ 0.368 x N x ln([Na+])
Where
N is the number of nucleotide pairs in the primer ( primer length -1).
[Na+] is salt equivalent in mM.
[Na+] calculation:
[Na+] = Monovalent ion concentration +4 x free Mg2+.

3. Primer annealing temperature: The primer melting temperature is the estimate of the
DNA-DNA hybrid stability and critical in determining the annealing temperature. Too
high Ta will produce insufficient primer-template hybridization resulting in low PCR
product yield. Too low Ta may possibly lead to non-specific products caused by a high
number of base pair mismatches,. Mismatch tolerance is found to have the strongest
influence on PCR specificity.
Ta = 0.3 x Tm(primer) + 0.7 Tm (product) – 14.9
where,
Tm(primer) = Melting Temperature of the primers
Tm(product) = Melting temperature of the product

4. GC Content: The GC content (the number of G’s and C’s in the primer as a percentage
of the total bases) of primer should be 40-60%.

5. GC Clamp: The presence of G or C bases within the last five bases from the 3′ end of
primers (GC clamp) helps promote specific binding at the 3′ end due to the stronger

bonding of G and C bases. More than 3 G’s or C’s should be avoided in the last 5 bases at
the 3′ end of the primer.

6. Primer Secondary Structures: Presence of the primer secondary structures produced
by intermolecular or intramolecular interactions can lead to poor or no yield of the
product. They adversely affect primer template annealing and thus the amplification.
They greatly reduce the availability of primers to the reaction.

i) Hairpins : It is formed by intramolecular interaction within the primer and should be
avoided.

ii) Self Dimer: A primer self-dimer is formed by intermolecular interactions between the
two (same sense) primers, where the primer is homologous to itself. Generally a large
amount of primers are used in PCR compared to the amount of target gene. When primers
form intermolecular dimers much more readily than hybridizing to target DNA, they
reduce the product yield.
iii) Cross Dimer : Primer cross dimers are formed by intermolecular interaction between
sense and antisense primers, where they are homologous.

7. Repeats: A repeat is a di-nucleotide occurring many times consecutively and should
be avoided because they can misprime. For example: ATATATAT. A maximum number
of di-nucleotide repeats acceptable in an oligo is 4 di-nucleotides.

8. Runs: Primers with long runs of a single base should generally be avoided as they can
misprime. For example, AGCGGGGGATGGGG has runs of base ‘G’ of value 5 and 4. A
maximum number of runs accepted is 4bp.

9. 3′ End Stability: It is the maximum ΔG value of the five bases from the 3′ end. An
unstable 3′ end (less negative ΔG) will result in less false priming.

10. Avoid Template secondary structure: A single stranded Nucleic acid sequences is
highly unstable and fold into conformations (secondary structures). The stability of these
template secondary structures depends largely on their free energy and melting
temperatures(Tm). Consideration of template secondary structures is important in
designing primers. If primers are designed on a secondary structures which is stable even
above the annealing temperatures, the primers are unable to bind to the template and the
yield of PCR product is significantly affected. Hence, it is important to design primers in
the regions of the templates that do not form stable secondary structures during the PCR
reaction.

11. Avoid Cross homology: To improve specificity of the primers it is necessary to
avoid regions of homology. Primers designed for a sequence must not amplify other
genes in the mixture. Commonly, primers are designed and then BLASTed to test the
specificity.

Primer Design Using Software
A number of primer design tools are available that can assist in PCR primer design for
new and experienced users alike. These tools may reduce the cost and time involved in
experimentation by lowering the chances of failed experimentation.
Primer Premier follows all the guidelines specified for PCR primer design. Primer
Premier can be used to design primers for single templates, alignments, degenerate
primer design, restriction enzyme analysis. contig analysis and design of sequencing
primers.
The guidelines for qPCR primer design vary slightly. Software such as AlleleID and
Beacon Designer can design primers and oligonucleotide probes for complex detection
assays such as multiplex assays, cross species primer design, species specific primer
design and primer design to reduce the cost of experimentation.
PrimerPlex is the software that can design ASPE (Allele specific Primer Extension)
primers.

Primer3 is the most commonly used online server for the designing of primers to be used
for the detection or diagnosis of genes.

DESIGNING DEGENERATE PRIMERS
• Degenerate primers may be used to amplify DNA in situations where only the protein
sequence of a gene is known, or where the aim is to isolate similar genes from a variety
of species.
• A six or seven residue peptide sequence should be selected, corresponding to an oligo
of about 20 nucleotides.
• If the oligo is designed to amplify several similar protein sequences, then the most
conserved regions of the proteins need to be selected. If some of the residues are not
completely conserved, then the oligo sequence will need to accommodate all possible
codons of all amino acid residues at that site (e.g. if one protein has an E at a particular
site, while all the others have a D, then the corresponding oligo sequence will be GAN –
see table overleaf).
• The peptide sequence should avoid amino acids that have a lot of codons, such as
leucine (L), arginine (R), and serine (S). Instead, aim for regions that are rich in amino
acids that have only one or two possible codons (i.e. M, W, C, D, E, F, H, K, N, Q, Y –
see table overleaf).
Primer sequence:
1. Avoid degeneracy in the 3 nucleotides at the 3´end.
2. If possible, use Met or Trp-encoding triplets at the 3´end.
3. To increase primer-template binding efficiency, reduce degeneracy by allowing some
mis-matches between the primer and template, especially towards the 5´end (but not
the 3´end).
4. Try to design primers with less than 4-fold degeneracy at any given position.
Primer concentration:
1. Begin PCR with a primer concentration of 0.2 micromolar.
2. In case of poor PCR efficiency, increase primer concentration in increments of 0.25
micromolar until satisfactory results to be obtained.
Try and avoid degeneracy at the 3’ end of the oligo (note that it is not necessary to have
whole codons), and especially avoid ending in inosine.

• The table overleaf may be used to help with primer design:
Amino acid

Symbol

Nucleotide sequence Complement (for

methionine
tryptophan
cysteine
aspartic acid
glutamic acid
phenylalanine
histidine
lysine
asparagine
glutamine
tyrosine
isoleucine
alanine
glycine
proline
threonine
valine
leucine
arginine
serine

M
W
C
D
E
F
H
K
N
Q
Y
I
A
G
P
T
V
L
R
S

(With degeneracy)

designing
primers)

ATG
TGG
TGY
GAY
GAR
TTY
CAY
AAR
AAY
CAR
TAY
ATH
GCN
GGN
CCN
ACN
GTN
YTN
MGN
WSN

TAC
ACC
ACR
CTR
CTY
AAR
GTR
TTY
TTR
GTY
ATR
TAD
CGN
CCN
GGN
TGN
CAN
RAN
KCN
WSN

reverse

Key to symbols:
R=A+GY=C+T
M=A+CK=G+T
S=G+C
W=A+T
H=A+T+CD=G+A+T
B=G+T+CV=G+A+C
N=A+T+G+C
PCR WITH DEGENERATE PRIMERS
The PCR is pretty standard, with the main difference being that a higher concentration of
primer is used. The concentration of MgCl2 and the annealing temperature may be varied
to optimise the reaction. The extension time may need to be increased for larger (>1 kb
fragments). It also sometimes helps to do a hot-start reaction (reduces non-specific
annealing of primers) – i.e. the Taq is not added until the tubes have been heated to 94oC
(pause the run during the 94oC for 2 minutes step, add the Taq, then proceed). Note that
1.25 µl DMSO or 5 µl of 5 M betaine should be added if the reaction is carried out on
maize.
1. Set up PCR, on ice:
genomic DNA
50-100 ng
10 x PCR buffer
2.5 µl

MgCl2 (25 mM)
dNTPs (10 mM)
each primer (100 µM)
dH2O to
Taq

1-2.5 µl
0.5 µl
0.5 µl
24.5 µl
0.25 µl

2. Run PCR:
94 °C
then 35 rounds of:
94 °C
45-55 °C
72 °C
followed by:
72 °C

2 minutes (add Taq at this stage, for hot-start)
30 secs
30 secs
1 minute
10 minutes

Ammonium sulfate precipitation is a method used to purify proteins by altering their
solubility. It is a specific case of a more general technique known as salting out.
Ammonium sulfate is commonly used as its solubility is so high that salt solutions with
high ionic strength are allowed.

The solubility of proteins varies according to the ionic strength of the solution, and hence
according to the salt concentration. Two distinct effects are observed: at low salt
concentrations, the solubility of the protein increases with increasing salt concentration
(i.e. increasing ionic strength), an effect termed salting in. As the salt concentration (ionic
strength) is increased further, the solubility of the protein begins to decrease. At
sufficiently high ionic strength, the protein will be almost completely precipitated from
the solution (salting out).
Since proteins differ markedly in their solubilities at high ionic strength, salting-out is a
very useful procedure to assist in the purification of a given protein. The commonly used
salt is ammonium sulfate, as it is very water soluble, forms two ions high in the
Hofmeister series, and has no adverse effects upon enzyme activity. It is generally used
as a saturated aqueous solution which is diluted to the required concentration, expressed
as a percentage concentration of the saturated solution (a 100% solution).

The Hofmeister series or lyotropic series is a classification of ions in order of their
ability to salt out or salt in proteins. The effects of these changes were first worked out by
Franz Hofmeister, who studied the effects of cations and anions on the solubility of
proteins.
Hofmeister discovered a series of salts that have consistent effects on the solubility of
proteins and (it was discovered later) on the stability of their secondary and tertiary
structure. Anions appear to have a larger effect than cations, and are usually ordered

This is a partial listing; many more salts have been studied. The order of cations is
usually given as

The mechanism of the Hofmeister series is not entirely clear, but does not seem to result
from changes in general water structure, instead more specific interactions between ions
and proteins and ions and the water molecules directly contacting the proteins may be
more important. The salting out effect is commonly exploited in protein purification
through the use of ammonium sulfate precipitation.

In the preliminary test, the ammonium sulfate concentration is increased stepwise, and
the precipitated protein is recovered at each stage. This is usually done by adding solid
ammonium sulfate, but calculating how much ammonium sulfate to add to a solution at
one concentration to achieve a desired higher concentration is tricky, since addition of
ammonium sulfate significantly increases the volume of the solution. The amount to add
can be determined either from published nomograms or by using an online calculator.
Each protein precipitate is dissolved individually in fresh buffer and assayed for total
protein content and amount of desired protein. The aim is to find the ammonium sulfate
concentration which will precipitate the maximum proportion of undesired protein, whilst
leaving most of the desired protein still in solution or vice versa.
The precipitated protein is then removed by centrifugation and then the ammonium
sulfate concentration is increased to a value that will precipitate most of the protein of
interest whilst leaving the maximum amount of protein contaminants still in solution. The
precipitated protein of interest is recovered by centrifugation and dissolved in fresh buffer
for the next stage of purification.
This technique is useful to quickly remove large amounts of contaminant proteins, as a
first step in many purification schemes. It is also often employed during the later stages
of purification to concentrate protein from dilute solution following procedures such as
gel filtration.

Online calculator
http://www.encorbio.com/protocols/AM-SO4.htm

Restriction Endonucleases
A restriction enzyme (or restriction endonuclease) is an enzyme that cuts DNA at specific
recognition nucleotide sequences known as restriction sites. Restriction enzymes are
commonly classified into three types, which differ in their structure and whether they cut
their DNA substrate at their recognition site, or if the recognition and cleavage sites are
separate from one another. To cut DNA, all restriction enzymes make two incisions, once
through each sugar-phosphate backbone (i.e. each strand) of the DNA double helix.
These enzymes are found in bacteria and archaea and probably evolved to provide a
defense mechanism against invading viruses. Inside a bacterium, the restriction enzymes
selectively cut up foreign DNA in a process called restriction; while host DNA is
protected by a modification enzyme (a methylase) that modifies the bacterial DNA and
blocks cleavage. Together, these two processes form the restriction modification system.
Over 3000 restriction enzymes have been studied in detail, and more than 600 of these
are available commercially. These enzymes are routinely used for DNA modification and
manipulation in laboratories, and are a vital tool in molecular cloning.

History
The term restriction enzyme originated from the studies of phage λ and the phenomenon
of host-controlled restriction and modification of a bacterial virus. The phenomenon was
first identified in work done in the laboratories of Salvador Luria and Giuseppe Bertani in
early 1950s. It was found that a bacteriophage λ that can grow well in one strain
of Escherichia coli, for example E. coli C, when grown in another strain, for example E.
coli K, its yields can drop significantly, by as much as 3-5 orders of magnitude. The E.
coli K host cell, known as the restricting host, appears to have the ability to reduce the
biological activity of the phage λ. If a phage becomes established in one strain, the ability
of that phage to grow also becomes restricted in other strains. In the 1960s, it was shown
in work done in the laboratories of Werner Arber and Matthew Meselson that the
restriction is caused by an enzymatic cleavage of the phage DNA, and the enzyme
involved was therefore termed a restriction enzyme.
The restriction enzymes studied by Arber and Meselson were type I restriction enzymes
which cleave DNA randomly away from the recognition site. In 1970, Hamilton O.
Smith, Thomas Kelly and Kent Welcox isolated and characterized the first type II
restriction enzyme, HindII, from the bacterium Haemophilus influenzae. This type of
restriction enzymes is more useful for laboratory use as they cleave DNA at the site of
their recognition sequence. It was later shown by Daniel Nathans and Kathleen Danna
that cleavage of simian virus 40 (SV40) DNA by restriction enzymes yielded specific
fragments which can be separated using polyacrylamide gel electrophoresis, thus
showing that restriction enzymes can be used for mapping of the DNA. For their work in
the discovery and characterization of restriction enzymes, the 1978 Nobel Prize for
Physiology or Medicine was awarded to Werner Arber, Daniel Nathans, and Hamilton O.
Smith. Their discovery led to the development of recombinant DNA technology that
allowed, for example, the large scale production of human insulin for diabetics using E.
coli bacteria.

Recognition site
Restriction enzymes recognize a specific sequence of nucleotides and produce a doublestranded cut in the DNA. The recognition sequences usually vary between 4 and 8
nucleotides, and many of them are palindromic, meaning the base sequence reads the
same backwards and forwards. In theory, there are two types of palindromic sequences
that can be possible in DNA. The mirror-like palindrome is similar to those found in
ordinary text, in which a sequence reads the same forward and backwards on a single
strand of DNA strand, as in GTAATG. The inverted repeat palindrome is also a sequence
that reads the same forward and backwards, but the forward and backward sequences are
found in complementary DNA strands (i.e., of double-stranded DNA), as in GTATAC
(GTATAC being complementary to CATATG). Inverted repeat palindromes are more
common and have greater biological importance than mirror-like palindromes. For
example, the common restriction enzyme EcoRI recognizes the palindromic sequence
GAATTC and cuts between the G and the A on both the top and bottom strands, leaving
an overhang (an end-portion of a DNA strand with no attached complement) known as
a sticky end on each end, of AATT. This overhang can then be used to ligate in
(see DNA ligase) a piece of DNA with a complementary overhang (another EcoRI-cut

piece, for example). Some restriction enzymes cut DNA at a restriction site in a manner
which leaves no overhang, a blunt end.
EcoRI digestion produces “sticky” ends,

Whereas SmaI restriction enzyme cleavage produces “blunt” ends:

Recognition sequences in DNA differ for each restriction enzyme, producing differences
in the length, sequence and strand orientation (5′ end or the 3′ end) of a stickyend “overhang” of an enzyme restriction.
Different restriction enzymes that recognize the same sequence are known
as neoschizomers. These often cleave in different locales of the sequence. Different
enzymes that recognize and cleave in the same location are known as isoschizomers.
Palindromic sequences play an important role in molecular biology. Because a DNA
sequence is double stranded, we read the base pairs, not just the bases on one strand to
determine a palindrome. Many restriction endonucleases (restriction enzymes) recognize
specific palindromic sequences and cut them.
Enzyme Source

Recognition Sequence Cut

EcoR1

Escherichia coli

5’GAATTC
3’CTTAAG

5′—G AATTC—3′
3′—CTTAA G—5′

BamH1 Bacillus amyloliquefaciens

5’GGATCC
3’CCTAGG

5′—G GATCC—3′
3′—CCTAG G—5′

Taq1

Thermus aquaticus

5’TCGA
3’AGCT

5′—T CGA—3′
3′—AGC T—5′

Alu1*

Arthrobacter luteus

5’AGCT
3’TCGA

5′—AG CT—3′
3′—TC GA—5′

* = blunt ends
Palindromic sequences may also be methylation sites.

Types

Naturally occurring restriction endonucleases are categorized into four groups (Types I, II
III, and IV) based on their composition and enzyme cofactor requirements, the nature of
their target sequence, and the position of their DNA cleavage site relative to the target
sequence. All types of enzymes recognize specific short DNA sequences and carry out
the endonucleolytic cleavage of DNA to give specific fragments with terminal 5’phosphates. They differ in their recognition sequence, subunit composition, cleavage
position, and cofactor requirements, as summarized below:
• Type I enzymes (EC 3.1.21.3) cleave at sites remote from recognition site; require
both ATP and S-adenosyl-L-methionine to function; multifunctional protein with
both restriction and methylase (EC 2.1.1.72) activities.
• Type II enzymes (EC 3.1.21.4) cleave within or at short specific distances from
recognition site; most require magnesium; single function (restriction) enzymes
independent of methylase.
• Type III enzymes (EC 3.1.21.5) cleave at sites a short distance from recognition
site; require ATP (but do not hydrolyse it); S-adenosyl-L-methionine stimulates
reaction but is not required; exist as part of a complex with a modification
methylase (EC 2.1.1.72).
• Type IV enzymes target modified DNA, e.g. methylated, hydroxymethylated and
glucosyl-hydroxymethylated DNA

Nomenclature
Since their discovery in the 1970s, more than 100 different restriction enzymes have been
identified in different bacteria. Each enzyme is named after the bacterium from which it
was isolated using a naming system based on bacterial genus, species and strain. For
example, the name of the EcoRI restriction enzyme was derived as shown in the box.
Derivation of the EcoRI name
Abbreviation

Meaning

Description

E

Escherichia

genus

co

coli

species

R

RY13

strain

I

First identified

order of identification
in the bacterium

PLASMIDS:
Plasmids are extrachromosomal elements found inside a bacterium. These are not
essential for the survival of the bacterium but they confer certain extra advantages to the
cell.
Number and size: A bacterium can have no plasmids at all or have many plasmids (2030) or multiple copies of a plasmid. Usually they are closed circular molecules; however
they occur as linear molecule in Borrelia burgdorferi. Their size can vary from 1 Kb to
400 Kb.
Multiplication: Plasmids multiply independently of the chromosome and are inherited
regularly by the daughter cells.
Types of plasmids: R factor, Col factor, RTF and F factor.
F factor: This is also known as fertility factor or sex factor. Most plasmids are unable to
mediate their own transfer to other cells. Vertical (inheritance) or horizontal (transfer)
transmissions maintain plasmids. F factor is a plasmid that codes for sex pili and its
transfer to other cells. Those bacteria that possess transfer factor are called F+, such
bacteria have sex pili on their surface. Those cells lacking this factor are designated F-.
The F factor plasmid is transferred to other cells through conjugation. An F- cell will
become F+ when it receives the fertility factor from another F+ cell.
R factor: Those plasmids that code for the transmissible drug resistance are called R
factor. These plasmids contain genes that code for resistance to many antibiotics. R
factors may be transferred by conjugation and its transfer to other bacteria is independent
of the F factor. Bacteria possessing such plasmids are resistant to many antibiotics and
this drug resistance is transferred to closely related species. R factors may simultaneously
confer resistance to five antibiotics. They are usually transferred to related species along
with RTF.
Significance of plasmids:
1. Codes for resistance to several antibiotics. Gram-negative bacteria carry plasmids that
give resistance to antibiotics such as neomycin, kanamycin, streptomycin,
chloramphenicol, tetracycline, penicillins and sulfonamides.
2. Codes for the production of bacteriocines.
3. Codes for the production of toxins (such as Enterotoxins by Escherichia coli, Vibrio
cholerae, exfoliative toxin by Staphylococcus aureus and neurotoxin of Clostridium
tetani).
4. Codes for resistance to heavy metals (such as Hg, Ag, Cd, Pb etc.).
5. Plasmids carry virulence determinant genes. Eg, the plasmid Col V of Escherichia coli
contains genes for iron sequestering compounds.
6. Codes resistance to uv light (DNA repair enzymes are coded in the plasmid).
7. Codes for colonization factors that is necessary for their attachment. Eg, as produced
by the plasmids of Yersinia enterocolitica, Shigella flexneri, Enteroinvasive Escherichia
coli.
8. Contains genes coding for enzymes that allow bacteria unique or unusual materials for
carbon or energy sources. Some strains are used for clearing oil spillage.

Application of plasmids:
1. Used in genetic engineering as vectors.
2. Plasmid profiling is a useful genotyping method.
Episomes: Jacob and Wollman coined the term episome. Previously, it was considered
synonymous with plasmids. F factors are those plasmids that can code for self transfer to
other bacteria.
Occasionally such plasmids get spontaneously integrated into
chromosome. Plasmids with this capability are called episomes and such bacterial cells
are called Hfr cells i.e. high frequency of recombination.

Cloning
Cloning is a procedure which generates a large number of copies of a single sequence of
DNA. These procedures depend upon the ability of vectors to continue their life cycles in
bacterial or yeast cells in spite of having foreign DNA inserted into them. Because the
foreign DNA is carried into the bacterial or yeast cell by these molecules, they are called
cloning vectors.
Cloning Vectors
The molecular analysis of DNA has been made possible by the cloning of DNA. The two
molecules that are required for cloning are the DNA to be cloned and a cloning vector.
Cloning vector – a DNA molecule that carries foreign DNA into a host cell, replicates
inside a bacterial (or yeast) cell and produces many copies of itself and the foreign DNA
Three features of all cloning vectors
1. sequences that permit the propagation of itself in bacteria (or in yeast for YACs)
2. a cloning site to insert foreign DNA; the most versatile vectors contain a site that
can be cut by many restriction enzymes
3. a method of selecting for bacteria (or yeast for YACs) containing a vector with
foreign DNA; usually accomplished by selectable markers for drug resistance
Types of Cloning Vectors
•

•

•

Plasmid – an extrachromosomal circular DNA molecule that autonomously
replicates inside the bacterial cell; cloning limit: 100 to 10,000 base pairs or 0.110 kilobases (kb)
Phage – derivatives of bacteriophage lambda; linear DNA molecules, whose
region can be replaced with foreign DNA without disrupting its life cycle; cloning
limit: 8-20 kb
Cosmids – an extrachromosomal circular DNA molecule that combines features
of plasmids and phage; cloning limit – 35-50 kb. Cosmids are plasmid vectors that
contain cos sites. The cos site is the only requirement for DNA to be packaged

•
•

into a phage particle. Cosmids were developed in light of this observation. (One
of the cohesive, single-stranded extensions present at the ends of the DNA
molecules of certain strains of λ phage.)
Bacterial Artificial Chromosomes (BAC) – based on bacterial mini-F plasmids.
cloning limit: 75-300 kb
Yeast Artificial Chromosomes (YAC) – an artificial chromosome that contains
telomeres, origin of replication, a yeast centromere, and a selectable marker for
identification in yeast cells; cloning limit: 100-1000 kb

Insert Size of Various DNA Cloning Vectors
Vector

Insert size (kb)

Plasmid

Foreign DNA is inserted into a plasmid (or any cloning vector) by ligating the DNA into
a complementary site in the plasmid. These sites are generated by digesting the DNA and
vector with the same restriction enzyme. (The site for the restriction enzyme that is
chosen should only be represented once in the plasmid. Thus, when the plasmid is
digested, a single, linear molecule would be generated.) The foreign DNA is then inserted
into the plasmid by the action of the enzym…