Unit V

Tools and
Techniques:
Chapter 12
Tools and Techniques Basic Concepts
Considering the fact that biotechnology is
an experimental science and involves a lot of
experimentations; therefore, research in this field
depends highly on sophisticated laboratory methods.
Advances in biotechnology were closely followed by
the development of newer tools and techniques in
biological sciences. These new methods opened new
avenues for research and investigation in the field
of biotechnology. It is, thus, important to appreciate
the experimental tools available to biotechnologists
in order to understand the progress and future
directions of this rapidly moving area of science.
Some of the important experimental methods
including methods of cell and molecular biology
will be discussed in this unit.

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 Frederick Sanger
(1918-2013)

Frederick Sanger (1918–2013) was

a British biochemist and molecular
biologist who had two Nobel Prizes
in Chemistry to his credit. He was
awarded the first Nobel Prize in
1958 for the discovery of structure
of insulin molecule, and second
Nobel Prize in 1980 for his work (in
collaboration with Paul Berg and
Walter Gilbert) on the determination
of base sequences of nucleic acids.
He is, by far, the most influential
biochemist in history. His technique
of deciphering DNA sequences
was based on ‘read-off’ methods
using acrylamide gel. In 1977,
Sanger sequenced the genome of
bacteriophage ΦX174, the first
genome to be completely sequenced.
Most of his later contributions laid
the foundation of molecular biology
and are being utilised in every
biotechnology application.

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 Tools and
Chapter 12 Techniques

12.1 Microscopy 12.1 Microscopy

12.2 Centrifugation Biological studies and explorations cannot be imagined
12.3 Electrophoresis without a microscope as it enables us to see something
12.4 Enzyme-linked which is beyond the scope of our eyes. Today, the technique
Immunosorbent of microscopy has become so much advanced that a
Assay (ELISA) researcher can not only see a highly magnified image of
a very minute structure but also can visualise the three
12.5 Chromatography
dimensional structure of such objects. Using powerful
12.6 Spectroscopy electron microscopic techniques, even the DNA molecule
12.7 Mass Spectrometry of bacteria and viruses have been visualised.
The use of first microscope dates back to 1665 when
12.8 Fluorescence in situ
the British physicist Robert Hooke designed a simple
hybridisation (FISH)
microscope using combination of magnifying lenses (Fig.
12.9 DNA Sequencing 12.1) and observed the slices of cork, and coined the term
12.10 DNA Microarray ‘Cellulae’ or ‘cell’ to that honeycomb like structure. You
12.11 Flow Cytometry are aware that Matthias Jacob Schleiden and Theodor
Schwann proposed cell theroy in 1838 on the basis of
observation of cells in plants and animals.

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Eyepiece

Body tube
Coarse adjustment
knob
Revolving nose
piece
Fine adjustment
knob

Arm Objective

Stage

Condenser
Inclination joint
Sub stage

Mirror

Base

Fig. 12.1: Microscope

12.1.1 Magnification and Resolution

Let us now focus on the principle on which the technique of
microscopy is based. Two optical properties are extremely
important for an optical instrument like microscope. One is
the power to magnify and the other is the ability to resolve.
Magnification or magnifying power of a microscope is
the ability by which the retinal image size can be increased.
Thus in simple terms magnification is—

Size of retinal image with the help of microscope

__________________________________________________
Size of the retinal image without using microscope

You may have studied in physics that magnification (M) of

a lens is measured as per the following formula (in which
f is focal length of the lens and d is the distance of object
from the lens).
f
M=
f -d

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Normally, the microscope which is used in the laboratory

is a compound microscope in which two sets of lenses are
there. One is called objective lens, which remains close to
the object to be seen, and the other is eyepiece through
which the observer sees. It is needless to mention that the
object, objective lens, eyepiece, and the eye of the observer
have to be in the same line for the passage of light to
see the magnified image of the object. In simple words,
magnification of a microscope is product of the magnifying
power of the objective lens and the magnifying power of
the eyepiece (Mo × Me).
Resolving power is another important property of a
microscope, which is the ability to form separate images
of the two objects situated very close to each other. It can
be measured by the smallest distance between two points.

12.1.2 Functioning of a light microscope

You have already studied about the structure of a compound
microscope in previous class, yet to recapitulate, as you
can see in the Fig. 12.1, a compound microscope consists
of a base on which a stage is fitted with a central hole.
Attached to the base is an arm to which a body tube is fitted
in such a way that it aligns with the hole of the stage. At the
lower end of the body tube, a nose piece is fitted on which
two to four objective lenses may be present. By rotating the
nose piece, one of the objective lenses can be placed above
the hole present on the stage where object to be seen is
placed on a glass slide. At the upper end of the body tube,
an eye piece is fitted through which an observer can see
under the microscope. There are adjustment screws (coarse
and fine) on the arm which facilitate in adjusting the
distance of objective lens from the object present on the
stage. Below the stage, there is a source of light (which may
be a reflective mirror or a bulb to illuminate the object and
facilitate the formation of image through objective lens and
eyepiece) (Fig. 12.2). In addition, there is a condenser
present between the light source and the stage, which is
important for focusing light on the object. You might have
observed that both objective lenses and eye pieces are of
different magnifying power. In a student microscope the
eyepiece has the magnifying power of 10× or 15× and that
of the different objective lenses fitted on the nose piece are
of 4×, 10×, 40/45× and 100×. The technique of microscopy

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just discussed is also called bright field microscopy as

light is used to illuminate the object to be seen. Therefore,
in order to distinguish different regions of the object, the
same is stained with specific dyes or stain. Carmine, eosin,
safranin, methylene blue, giemsa, etc., are few such stains
commonly used for light microscopy.
Eye
Magnified
Image

Eyepiece

Projector
Lens

Objective

Specimen

Condenser Lens

Light Source

Fig. 12.2: Pathway of light in a light microscope

12.1.3 Different forms of microscopy

Studying minute details of internal organisation of tissues/
cells is so diverse that it cannot be achieved by light
microscopy only. Therefore, by making one or the other
kind of maneuvering, quite diverse forms of microscopy
are used. In one such maneuvering, light falling on the
object from the central condenser is blocked by a disc
and illumination of the object is done by an oblique light
beam, which is reflected off from the slide and the image

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is illuminated against the dark background. Therefore,

such a microscopy is known as Dark Field Microscopy.
Mitochondria, nuclei, vacuoles, etc., can be easily detected
using this. Similarly, in a different form called Phase
Contrast Microscopy, wave amplitude and phase of light
passing through the transparent object is changed. This
change depends on the density of the part of the object or
specimen. Such a change is more in the area where density
is comparatively high and as a result of which, varied
contrast of different regions of the object can be seen.
This is especially helpful in the study of cell organelles
and chromosomes. Staining of the object or specimen
with some specific dye is routinely done. There are some
special types of dyes e.g., acridine orange, bisbenzimide,
merocyanine (also called fluorophores). These dyes are
capable of emitting light of longer wavelength after being
illuminated, a property called fluorescence. As a result of
this, the fluorophore stained object looks more illuminated
and of different colour depending on the dye used. In
fluorescence microscopy, the same principle is applied.
Object to be seen is stained with fluorophore to study a
specific part of organelle or molecule. After illuminating
the object under fluorescence microscope, the specifically
stained region is easily seen or observed. This is helpful
in identifying bacteria or viruses to know the cause of
infection and immunodiagnosis.
Electron microscopy is a highly sophisticated
technique in which the object to be studied is bombarded
with electron beam which is approximately 1,00,000 times
shorter in wavelength than visible light. The electron
beam in an electron microscope magnifies the image with
the help of electromagnetic lenses. The entire passage of
electron is in vacuum and the generated image is viewed
on a fluorescent screen and not through eyepiece. Owing
to the very shorter wavelength of the electron, the image
produced by an electron microscope is of very high
resolution. Two types of electron microscopy are used;
transmission electron microscopy and scanning electron
microscopy. In Transmission electron microscopy, the
ultra-thin heavy metal salt (of lead, tungsten, etc.) coated
section of the object or specimen is placed in such a way
that the electron beam passes through it to create the
image. In the other technique of electron microscopy,

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reflected electron beam from the gold or platinum coated

surface of the object creates the image. In this technique,
highly magnified and resolved image of the object surface
is generated, therefore, it is called Scanning electron
microscopy.
In last few decades, yet another more
sophisticated microscopic imaging technique has been
developed and used called the confocal microscopy.
Confocal microscopy is useful in resolving detailed
structures within fixed cells/tissues and gives sharp
images of the objects. To examine an object using confocal
microscopy, it is first fluorescently labelled and then
analysed under a confocal microscope in high resolution.

12.2 Centrifugation
You have studied about various biomolecules like proteins,
nucleic acids, etc., present in cells of all living organisms.
To study these biomolecules, you need to isolate them by
using one or the other separation techniques. Centrifugation
is one such technique in which particles or molecules are
separated based on their densities under the influence of
gravitational force (g), by spinning them in a solution
around an axis at high speed using centrifugal force. The
equipment used is called centrifuge (Fig. 12.3), which is of
different types depending on its use. It consists of a base,
a rotating container (spinning vessel/rotor) and a lid. The

Latch
Lid
Chamber

Control

Rotor

Motor

Sample

Fig. 12.3: Basic structure of centrifuge

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spinning vessel contains several centrifuge tubes. The cell

extract or mixture is taken in centrifuge tubes and allowed
to spin at a desired speed (revolution per minute; rpm) for
a specific period of time leading to settlement of particulate
material at the bottom of the centrifuge tubes.

12.2.1 Types of Centrifuge

Based on the principle and application, following types of
centrifugations are performed—
Differential Centrifugation —It is based upon the
differences in the sedimentation rate (centrifugal force)
of particles of different size and density. It is used to
separate large cellular structures, the nuclear fraction,
mitochondria, chloroplasts, or large protein.
Density-gradient Centrifugation — In order to separate
biological particles of similar size but differing in densities,
one can use density gradient centrifugation. In this type
of centrifugation, a density gradient is developed in
centrifuge tubes. Depending upon their densities, different
molecules get sedimented at different levels. The heavier
molecules move outward and lighter ones remain in inner
part in the centrifuge tubes. The greater the difference in
density, the faster they move.
Ultracentrifugation— When centrifugation is carried out
at very high speed, i.e., 100,000 x/g or more to separate
the molecules, it is called ultracentrifugation. Commonly
used centrifuges are:
• Table top/clinical centrifuge or microfuge
• High-speed centrifuge
• Ultracentrifuge

12.3 Electrophoresis
Electrophoresis is a method of separation on the basis
of charge to mass ratio of macromolecules under the
influence of an electric field. Electrophoresis is a Greek
word meaning ‘to bear electrons’. The prefix electro refers
to electricity which is required to migrate molecules and
the suffix phoresis means ‘migration’ or ‘movement’. It was
observed for the first time in 1807 by Russian professors

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Peter Ivanovich Strakhov and Ferdinand Frederic Reuss.

They noticed the migration of the clay particles dispersed
in water in the presence of constant electric field.

Principle
Many important biological molecules such as nucleotides,
DNA, RNA, peptides and proteins bear ionisable groups,
and therefore at any given pH exist in solution as
electrically charged species either as cations or anions.
Under the influence of an electric field, these particles will
migrate either towards cathode or anode, depending on
their net charge.
The mobility of a molecule is inversely proportional to
its size and directly proportional to its charge, allowing
them to be separated from one another.

12.3.1 Agarose Gel Electrophoresis

In this type of electrophoresis, the gel is a matrix of agarose
molecules that are held together by hydrogen bonds and
form tiny pores. Gels for DNA separation are often made
out of a polysaccharide called agarose, which comes as
dry and powdered flakes. When the agarose is heated in a
buffer and allowed to cool, it will form a solid and slightly
squishy gel.
Gel is a slab of jelly-like material, placed in a gel box. One
end of the box is connected to a positive electrode, and
another end with a negative electrode. The gel box is filled
with a salt-containing buffer solution that can conduct
current. The end of the gel with the wells is positioned
towards the negative electrode. The other end of the gel is
positioned towards the positive electrode to which the DNA
fragments will migrate (Fig. 12.4).
DNA molecules are negatively charged.
Gel electrophoresis of DNA fragments
separates them on the basis of size only.
Using electrophoresis, we can check the
different DNA fragments present in a
sample and determine their absolute size
with the help of DNA ladder made up of
DNA fragments of known sizes.
When the power is turned on, the
Fig 12.4: Agarose gel electrophoresis unit to current begins to flow through the gel.
separate nucleic acid

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DNA molecules have a negative charge due to the presence

of phosphate groups in their sugar-phosphate backbone;
therefore, they move through the matrix of the gel towards
the positive electrode (anode).
The voltage for running an agarose DNA gel lies in
the range of 80–120 V. As the electric current is applied,
shorter pieces of DNA travel through the pores of the gel
matrix faster than longer ones. Thus the longest pieces
of DNA remain near the wells while the shortest pieces of
DNA are close to the positive end of the gel.

12.3.2 Visualising the DNA fragments

The equipment used for visualisation of target
DNA is ultra-violet (UV) trans-illuminator.
Ethidium bromide (EtBr) is most commonly
used for staining DNA. This stain can be
mixed in the gel mixture, in the electrophoresis
buffer or the gel is stained after it is run.
Molecules of the EtBr intercalates within DNA
bases and fluoresce under UV light. Despite its
advantages, ethidium bromide is a potential
carcinogen, so it must be handled with great Fig. 12.5: Visulalisation of DNA bands
care (Fig. 12.5). under UV light

12.3.3 Polyacrylamide Gel Electrophoresis (PAGE)

PAGE is an analytical method used to separate components
of a protein mixture based on their size. To provide uniform
charge to the protein molecules, an anionic detergent
called sodium dodecyl sulfate (SDS) is used to bind the
proteins and give them negative charge. Proteins are then
separated electrophoretically according to their size using
a gel matrix made of polyacrylamide in an electric field.
Polyacrylamide is produced as a result of the
polymerisation reaction between acrylamide and N,N'-
methylene-bis-acrylamide (BIS) using a catalyst. The
degree of polymerisation or cross-linking can be controlled
by adjusting the concentration of acrylamide and BIS.
The more the cross-linking, the harder the gel. Hardness
of the gel, in turn, modulates the friction experienced by
macromolecules when they travel through the gel during
PAGE, thus affecting the resolution of separation. Loose gels
(4–8% acrylamide) allow higher molecular weight molecules

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to migrate faster through the gel while hard gels (12–20%

acrylamide) restrict the migration of large molecules and
selectively allow small ones to move through the gel.

12.3.4 Tracking dye

As DNA, RNA, and proteins are mostly colourless, their
progress through the gel during electrophoresis cannot be
easily followed. Anionic dyes of a known electrophoretic
mobility are therefore, usually included in the sample
buffer. A very common tracking dye is bromophenol blue.
This dye is coloured at alkali and neutral pH, and is a
small negatively charged molecule that moves towards
the anode and it is coloured at alkali and neutral pH.
Being a highly mobile molecule it moves ahead of most
proteins and nucleic acids. As it reaches the anodic end
of the electrophoresis medium, electrophoresis is stopped.
Other common tracking dyes are xylene cyanol, which has
lower mobility, and Orange G, which has a higher mobility.

Visualisation of protein on gel — Coomassie Brilliant

Blue R-250 is the most popular protein stain. It is an
anionic dye, which non-specifically binds to proteins.
Box 1
Surprise Discovery of Jelly property
A Japanese Emperor and his Royal party were lost in the mountains during a snow
storm and arrived at a small inn; they were treated by the innkeeper with a seaweed jelly
dish with their dinner. Maybe the innkeeper prepared too much jelly or the taste was not
so palatable but some jelly was thrown away, freezing during the night and crumbling
afterwards by thawing and draining, leaving a cracked substance of low density. The
innkeeper took the residue and, to his surprise, found that by boiling it up with more
water, the jelly could be remade.
Agar
One of the scientists named Koch used to culture bacteria on the sterile surfaces of
boiled cut potatoes. This was unsatisfactory because bacteria would not always grow
well on potatoes. He then tried to solidify regular liquid media by adding gelatin but it
was digested by many bacteria and melted when the temperature rose above 28°C. A
better alternative was provided by Fanny Eilshemius Hesse, the wife of Walther Hesse,
one of Koch’s assistants. She suggested the use of agar as a solidifying agent—she had
been using it successfully to make jellies for some time. Agar was not attacked by most
bacteria and did not melt until reaching a temperature of more than 42°C. Agarose is
one of the two principal components of agar, and is purified from agar by removing
agar's other component, agaropectin.

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Proteins in the gel are fixed by methanol and acetic acid

and simultaneously stained. The excess dye incorporated
into the gel can be removed by destaining with the same
solution without the dye. The proteins are detected as blue
bands on a clear background. After the visualisation by a
staining (protein-specific) technique, the size of a protein
can be calculated by comparing its migration distance
with that of a known molecular weight ladder.

12.4 Enzyme-linked Immunosorbent Assay (ELISA)

Enzyme-linked immunosorbent assay (ELISA) was
invented by two Swedish scientists, Eva Engvall and Peter
Perlman in 1971. ELISA is a quantitative method used for
the measurement of antigen and antibody concentration
in a given sample. This is done by monitoring the antigen-
antibody interaction with the help of an enzyme catalysed
reaction. This detection system (an enzyme-conjugate) is
covalently linked to a specific antibody that recognises
a target antigen. The intensity of the color produced is
detected by ELISA reader or spectrophotometer. ELISA is
a safer and less costly assay as compared to many other
immunological assays.
A number of modifications of ELISA have been developed,
allowing qualitative detection or quantitative measurement
of either antigen or antibody. These different types of
ELISA can be employed qualitatively to detect the presence
of antibody or antigen. Using the known concentrations
of antibody or antigen, a standard curve is prepared to
determine the unknown concentration of a sample.
Direct ELISA — In direct ELISA, the antigen or sample
is coated on the microtiter plate wells and the enzyme-
conjugated antibody ‘directly’ binds to the antigen. The
enzyme linked to the antibody reacts with its substrate
to produce a colourful product that can be measured
using spectrophotometer/ELISA reader. The direct ELISA
is faster and less error prone and does not require a
secondary antibody. Disadvantage of direct ELISA is non-
specific binding of antibody due to cross-reactivity.
Indirect ELISA — The indirect ELISA detects the presence
of antibody in a sample in two stages (Fig. 12.6). Firstly,

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E
S

E
S

E
Antigen-coated well Add specic antibody Add enzyme-conjugated Add substrate(s)
to be measured secondary antibody and measure colour

Fig. 12.6: Indirect ELISA

E E E E S

Antibody Add antigen Add enzyme- Add substrate and

coated well to be measured conjugated secondary measure color
antibody
Fig. 12.7: Sandwich ELISA

E
E

Incubate antibody with Add Ag.Ab mixture to Add enzyme conjugated Add substrate and
antigen to be measured antigen-coated well secondary antibody measure color

Fig. 12.8: Competitive ELISA

an unlabelled primary antibody (Ab1) is applied to antigen
coated microtiter wells. The unbound extra primary
antibody (Ab1) is then washed off. Next, an enzyme-
conjugated secondary antibody (Ab2) specific for primary
antibody (Ab1) is added. Any unbounded secondary
antibody (Ab2) is washed off and the substrate is added.
The amount of coloured product formed can be measured
using spectrophotometer/ELISA reader. Indirect ELISA
has increased sensitivity since more than one labelled
antibody is bound per primary antibody.
Sandwich ELISA — In sandwich ELISA, the antibody
is coated on the microtiter plate and is referred to as
captured antibody. Using sandwich ELISA technique, the

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antigen can be detected or measured (Fig. 12.7). After

the captured antibody is coated on the plate, antigen is
added and allowed to react with the captured antibody.
Excess antigen is washed off and then a second enzyme-
conjugated antibody specific for a different epitope (the
part of an antigen molecule to which an antibody attaches)
on the antigen is loaded. The excess unbound enzyme-
conjugated antibody is washed off and substrate is added
and the coloured product produced is then measured using
spectrophotometer/ELISA reader. High specificity and no
requirement of antigen purification or limiting antigen are
the main advantages of sandwich ELISA.
Competitive ELISA — The competitive ELISA helps in
measuring amounts of antigen (Fig. 12.8). In this method,
the antibody and antigen are first incubated in solution. In
this step, the antibody is present in excess to the antigen;
it will bind to its antibody in a concentration dependent
manner leaving unbound antibody, accordingly. This
antigen-antibody complex is then added to an antigen
coated microtiter well. Thus, if more antigens are present
in the sample, less free antibody will be available to bind
to the antigen-coated well and vice versa. Finally, the
enzyme-conjugated secondary antibody (Ab2), specific
for the primary antibody is added to the plate and these
bind to the primary antibody bound to the antigen on
the plate. As in the case of indirect ELISA, addition of
an enzyme-conjugated secondary antibody (Ab2) specific
for the primary antibody helps in determining the
amount of primary antibody bound to the well. Thus, in
the competitive ELISA, the higher the concentration of
antigen in the sample; the lower will be the free antibody
and accordingly, the intensity of the developed colour will
be less.
Application of ELISA
1. Presence of antigen or antibody in a sample can be
estimated.
2. Concentration determination of serum antibody in a
virus test can be carried out.
3. Detection of potential food allergens in food industry.
4. Disease outbreaks can be tracked down e.g., HIV, bird
flu, cholera, etc.

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12.5 Chromatography

Principle
Chromatography is a separation procedure for isolating
various components of a mixture. The whole operation is
time dependent, and involves a mobile phase and a
stationary phase. The mobile phase may be a solution
containing solutes to be separated and the eluent (liquid)
that carries the solution through the stationary phase.
Stationary phase may be adsorbent, ion-exchange resin,
porous solid, or gel. The basis of chromatography is
different migration properties of the solute molecules
during passage through the stationary phase. Each solute
in the original solution moves at the rate proportional to
its relative affinity for the stationary phase. The stationary
phase is usually packed in a cylindrical column. Fig. 12.9
reveals an outline description of the separation of three
solutes from a mixture through chromatography.
The various components of a mixture, i.e., the solute
molecules travel with the eluent molecules at different speeds

Fig. 12.9: Chromatography — a schematic description

depending upon their relative affinities for the resin particles.

As a result, they separate and appear for collection at the
other end of the column at different time intervals. The
pattern of solute peaks emerging from a chromatography
column is called a chromatogram. A typical HPLC
chromatogram is shown in Fig. 12.10.
Liquid chromatography is used both as a laboratory
method for sample analysis and as a preparative
technique for large-scale purification of biomolecules.

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In recent year there have been rapid developments in the

technology of liquid chromatography aimed at the isolation
of recombinant products from genetically engineered
organisms. Chromatography is a high resolution technique
and therefore suitable for the recovery of high-purity
therapeutics and pharmaceuticals. Chromatographic
y

0. 5

0. 4
Intensity

0. 3

0. 2

0. 1

x
Time

Fig. 12.10: HPLC chromatogram

methods are available for purification of proteins, peptides,

amino acids, nucleic acids, alkaloids, vitamins, steroids
and many other biological materials. These methods differ
in the principles by which molecules are separated in the
chromatography column.
Adsorption Chromatography (ADC): This
chromatography is based on the adsorption of solute
molecules onto polar adsorbents such as silica gel,
alumina, diatomaceous earth and charcoal. Because the
mobile phase is in competition with solute for adsorption
sites, solvent properties are also important. Polarity scales
for solvents are available to aid mobile-phase selection.
Liquid-liquid Partition Chromatography (LLC)—Partition
chromatography relies on different partition coefficients
(solubility) of solute molecules between two immiscible
solvents. This is achieved by fixing one solvent as the
stationary phase and passing the other solvent containing
solute over it, a mobile phase. The solvents make intimate
contact allowing multiple extractions of solute to occur.

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Ion-exchange chromatography (IEC)— The basis of

separation in this procedure is the adsorption of solute
ions on ion exchange resin particles on the column packing
by electrostatic forces. Ion exchange chromatography may
provide high resolution of macromolecules. Commercially,
it is used for fractionation of antibiotics and proteins.
Column packings include silica, glass and polystyrene;
carboxymethyl and diethylaminoethyl groups attached to
cellulose, agarose or dextran provide suitable resins for
protein chromatography. Solutes are eluted by changing
the pH or ionic strength of the liquid phase.

Gel Filtration Chromatography —This technique is

also known as molecular sieve chromatography,
size exclusion chromatography and gel-permeation
chromatography. This is based on the size of
molecules to be separated. Solute molecules present
in the solution are separated in a column packed with
gel particles of distinct porosity. Mostly, the cross-linked
dextrants, agaroses and polyacrylamide gels are used
for the packing of column. The penetration of the solute
molecules through the column depends on the shape
of solute molecules and their effective molecular size.
Gel filtration can be used for separation of proteins and
lipophilic compounds. Large-scale gel filtration columns
are operated with upward-flow elution.

Affinity Chromatography (AFC): This separation

technique exploits the binding specificity of
biomolecules. Enzymes, hormones, receptors, antibodies,
antigens, binding proteins, lectins, nucleic acids, vitamins,
whole cells and other components capable of specific and
reversible binding are amenable to highly selective affinity
purification. Column packing is prepared by linking a
binding molecule called a ligand to column, only solutes with
appreciable affinity for the ligand are retained.
High-pressure /performance Liquid Chromatography
(HPLC): It is based on general principles of chromatography.
With the development of HPLC, the particle size of stationary
phase used has become progressively smaller. Small size
of these particles leads to a considerable resistance to

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solvent flow. The mobile phase has to be pumped through

the column under high pressure.
Gas Chromatography (GC): The gas chromatography
(GC) is widely used for separation and purification of
volatile components such as alcohols, ketones, aldehydes
and many other organic and inorganic compounds.
In gas chromatography, the mobile phase is gas.
Gas chromatography is used widely, however, liquid
chromatography is of great relevance to bioprocessing.

12.6 Spectroscopy
Spectroscopy is a technique used for the study of
interaction of electromagnetic radiation with matter. It is
used for the identification of substances through the
spectrum emitted from or absorbed by them. It is used for
the estimation of concentration of coloured compounds,
analysis of chemical structure of molecules, types of
molecular changes occurring during various enzymatic
reactions; and in the study of intermolecular bonding. List
of various spectroscopic techniques is given in Table 12.1.
Spectrophotometric techniques offer a very rapid and
convenient means of qualitative and quantitative
estimation of biomolecules. Spectrophotometer consists of
the following parts: light source, which provide the desired
wavelength of light, collimator, which transmits a straight
beam of light, monochromator, which splits the light into
its component wavelength, and wavelength selector, that
transmits only the desired wavelength (Fig. 12.11).
Sample

Monochromator
Detector

Incident light Monochromator

light
Light source

IO d I

Fig. 12.11: Diagrammatic representation of instrumentation of spectroscopy

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Box 2

The Beer-Lambert Law

The Beer-Lambert law states that the absorbance of a solution is directly
proportional to the concentration of the absorbing substance in the solution and
the path length. For this reason, Beer-Lambert law can only be applied when there
is a linear relationship. Beer’s law is written as:
A= εlc

Where A is the measure of absorbance (no units),

ε is the molar extinction coefficient or molar absorptivity (or absorption coefficient),
l is the path length and
c is the concentration.

Transmittance is the relationship between the amount of light that is transmitted to

the detector once it has passed through the sample (I) and the original amount of
light (I0). This is expressed in the following formula.

T = I / I0

Where T is the transmittance,

I is the intensity of the light coming out of the sample, and
I0 is the intensity of the incident light beam.
Absorbance equals the negative log of transmittance and this relationship between
transmittance (T) and absorbance (A) can be expressed as:
A = - log (T)
A = - log (I/I0)
A = log (I0/ I)
Therefore,
A= εlc = log (I0/ I)

If you know the absorption coefficient for a given wavelength, and the thickness
of the path length for light transmitted through the solution, you can calculate
concentration.

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Table 12.1 List of various spectroscopic techniques

Types of Region of
Spectroscopic
Energy Electromagnetic Application
Technique
Transfer Spectrum

Detection of functional groups, extent

UV/Visible
UV/Visible of conjugation, and determines the
spectroscopy
configurations of geometrical isomers

Atomic absorption Determines the amount of various levels of

UV/Visible
spectroscopy metals and other electrolytes within samples

Infrared
Infrared Determines the functional groups
Absorption spectroscopy

Raman Contaminant identification, gemstone and

Infrared
spectroscopy mineral identification

Nuclear magnetic
Provides information about the structure and
resonance Radio wave
chemical environment of atoms
spectroscopy

X-ray absorption Determines the elemental composition and

X-ray
spectroscopy chemical bonding of molecules

Atomic emission Detection of trace metals, minerals, sodium,

UV/Visible
spectroscopy potassium and lithium
Emission
Analysis of proteins, peptides, checking water
Mass spectrometer
quality and food contamination

Fluorescence Detection of many of organic compounds,

UV/Visible
Photolumi- spectroscopy numerous aromatic active substances in drug
nescence
Phosphorescence
UV/Visible -
spectroscopy

12.6.1 Colorimetry
Colorimetry technique utilises the interaction of light
energy with coloured solutions. This instrument is used
to measure transmittance and absorbance of light passing
through liquid sample. It measures the intensity of the
color that develops upon adding a specific reagent into
a solution. The intensity of color is directly proportional
to the concentration of the compound being measured.
Wavelength is selected using coloured filters which
absorb all but a certain limited range of wavelength. This
limited range is known as bandwidth of the filter. The
three main components of colorimeter are light source

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(tungsten-filament lamp), filter, cuvette containing

sample and a photocell for detecting the transmitted
light (light passed through the solution) (Fig. 12.12).
The principle of colorimeter is based on Beer-Lambert’s
law (Box 2) which is a combination of two laws, each
dealing separately with absorption of light related to the
concentration of absorber and the path length or thickness
of the absorbing medium. Colorimeter is inexpensive,
easily transportable and used for quantitative analysis of
colored compounds.

Polychromatic
Light
Monochromatic
Light

Light
source
Output

Slit Lens Filter Cuvette Photo cell

Fig. 12.12: Components of colorimeter

12.6.2 UV-visible Spectrophotometry

This instrument is used to measure the amount of light
absorbed at each wavelength of UV and visible regions of
electromagnetic spectrum. A spectrophotometer is a
sophisticated type of a colorimeter where monochromatic
light is provided by grating of a prism. In a colorimeter,
filters are used which allow a broad range of wavelengths to
pass through, whereas in the spectrophotometer a prism
(or) grating is used to split the incident beam into different
wavelengths. A ‘photometer’ is a device for measuring light,
Slit
Collimator (Wavelength
selector) Detector
(Photocell) 0.45

lo l
Light
Source Monochromator Cuvette Digital
(Prism of Grating) (Sample solution) Display

Fig. 12.13: UV-visible spectrophotometer

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and ‘spectro’ means the whole range of continuous

wavelength that the light source is capable of producing.
Spectrophotometer is made up of different components,
such as light sources (UV and visible), wavelength selector
(monochromator), sample containers (cuvette), detector,
signal processor etc. (Fig. 12.13).

12.7 Mass Spectrometry

Mass spectrometry is used to identify unknown
compounds, to quantify unknown materials, and to
elucidate structure and chemical properties of molecules.
The complete process involves the conversion of the
sample into gaseous ions by electron ionisation with or
without fragmentation, which are then characterised
by their mass-to-charge ratios (m/z) and relative
abundances. The unit of measurement of mass is Dalton
(Da for short form). One Dalton is equal to 1/12th of the
mass of single atom of the isotope of carbon-12. Three
major components of mass spectrometry are: ion
source for producing gaseous ions from the substance
being studied, analyser for resolving the ions into their
characteristic mass components according to their mass-
to-charge ratio, detector system for detecting the ions,
and recording the relative abundance of each of the
resolved ionic species (Fig. 12.14).

Gas Phase Ion Ion

lons Sorting Detection
Mass Spectrum

Ion Data
Inlet Source Analyser System
Detector

Sample
Introduction Vacuum Pumps Data Output

Fig. 12.14: Mass spectrometer

12.8 Fluorescence In Situ Hybridisation (FISH)

Fluorescence in situ hybridisation (FISH) is a cytogenetic
(study of chromosomes number and structure) technique
that uses fluorescent molecule that binds to highly
complementary region of chromosome. FISH is useful to
identify where a particular gene falls within an
individual’s chromosomes. Fluorescence microscopy is

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used to find the location of the fluorescent molecule in

the chromosome. It is an important tool to understand
chromosomal abnormalities.

Target

Probe Labelling b

Indirectly labelled Directly labelled

probe probe

Denaturation of target DNA c

and probes

Combining denatured probe

and target DNA allows d
annealing of complementary
DNA sequences

Since indirect labelling in e

non-uorescent detection
Hapten Fluorophore
systems is based on enzymatic
and immunological.

Fig. 12.15: Basic steps of FISH

The main components of FISH are:

(i) A fluorescent DNA molecule (probe), and
(ii) The chromosome (target sequence)

Here is how FISH works

1. Design a molecule (probe) complementary to the known
sequence. The probe is labelled with a fluorescent
molecule e.g. fluorescein (Fig. 12.15).
2. Put the chromosomes on a microscope slide and
denature them.
3. Denature the probe and add it to the microscope slide.
4. The probe hybridises to its complementary site.

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5. The excess probe is washed off and the chromosome

is observed under a fluorescent microscope. The
probe will show as one or more fluorescent signals in
the microscope, depending on how many sites it can
hybridize to.

Application of FISH
There are various applications of FISH including
chromosome painting. For chromosome
painting, multifluor FISH probe can be used to
generate a karyotype in which each chromosome
appears to be painted with a different colour
(Fig. 12.16).
First, a collection of DNA sequences
are prepared for using as probe for each
chromosome. Then these DNA sequences are
labelled with combinations of fluorochromes
that produce a unique color. The fluorescent Fig. 12.16: Chromosome painting
DNA probes and metaphase chromosomes are
mixed together; then the hybrids are visualised
under fluorescent microscope.

12.9 DNA Sequencing

As you are aware, DNA is made up of four types of
nucleotides (A,T,G,C), which are linked together through
phosphodiester linkages, and is the carrier of genetic
information. DNA sequencing refers to finding the order of
nucleotides (ATGC) on a piece of DNA. The order of these
four bases is key to the unique feature of the functional unit
of a given DNA (e.g., a gene). In other words, the sequence
of DNA comprises the heritable genetic information that
forms the basis for the developmental programs of all
living organisms. The advent of DNA sequencing has
significantly accelerated biological research and discovery.
The unique order of these bases greatly influences
health, e.g., what disease a person is prone to and how the
person will react to different medications. Understanding
a particular DNA sequence can shed light on a genetic
condition (e.g., disease) and offer hope for the eventual
development of treatment. Thus, an alteration in a DNA
sequence can lead to an altered or non functional protein,
and hence to a harmful effect. Also, in order to understand
the structure, function and evolutionary history of a cloned

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DNA, its primary structure, i.e., the nucleotide sequence

is required. DNA sequencing technology is also extended
to environmental, agricultural and forensic applications.
Thus, determining the DNA sequence is useful in basic
research studying fundamental biological processes, as
well as in applied fields such as diagnostic or forensic
research. The rapid speed of sequencing attained with
modern DNA sequencing technology has been helpful in
the sequencing of the complete DNA sequences of many
animal, plant, and microbial genomes including that of
the human.

12.9.1 DNA sequencing methods

Historically there are two main methods of DNA
sequencing, namely, (1) Enzymatic method (Sanger’s
method, dideoxynucleotides chain termination method)
and (2) Chemical degradation method (Maxam and Gilbert
method).
The two methods are described in detail in the following
sections:
(a) Sanger’s method — Sanger’s method works on the
principle that dideoxynucleotides (dideoxyadenine,

O O O
Base
O P O P O P O CH2 O (A,T,C or G)

O O O
Deoxynucleotide
triphosphate
H H H
(dNTP)

OH H

O O O
Base
O P O P O P O CH2 O (A,T,C or G)

O O O
Dideoxynucleotide
triphosphate H H H
(ddNTP)

OH missing H H

Fig. 12.17: S
 tructure of normal deoxynucleotide (top)
dideoxynucleotide (bottom)

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dideoxyguanine, etc., which resemble normal nucleotides

but lack the normal-OH group at 3' position) get
incorporated, instead of the normal deoxynucleotide,
into the newly synthesised chain (daughter chain)
which leads to termination of synthesis of the new
strand at that point (and hence it is called as chain
termination method) (Fig. 12.17). Sanger’s method is
considered as a gold standard for DNA sequencing. It
is used even today for routine sequencing applications
and also for validation of Next Generation Sequencing
(read in following sections of this chapter) data.
In Sanger’s method of DNA sequencing, the template
DNA (the sequence of which has to be determined) is mixed
with a primer (a small piece of chemically synthesised DNA

3'

5'

Fig. 12.18: Sanger's method of DNA sequencing

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of defined sequence that can pair with the template DNA to

act as a starting point for DNA synthesis) complementary
to the template DNA and the four normal dNTPs, one
of which is labelled radioactively or fluorescently. This
mixture is then split into four different tubes that are
labelled A, C, G, and T. Each tube is then ‘spiked’ with
a different ddNTP (ddATP for tube A, ddCTP for tube C,
ddGTP for tube G, or ddTTP for tube T). Following this,
DNA polymerase is added, and using the DNA template
and its complementary primer, the synthesis of new
strands of DNA complementary to the template begins.
Occasionally, a dideoxynucleotide is added instead of the
normal deoxynucleotide and synthesis of that strand is
terminated at that point. Thus, all fragments in lane A
will end in an A, fragments in lane C will all end in a C,
fragments in lane G will all end in a G, and fragments in
lane T will all end in a T. After carrying out the reaction for a
fixed time the newly synthesised DNA strands (fragments)
are separated through high resolution polyacrylamide
gel electrophoresis (PAGE) and the DNA fragments are
visualised by exposing the gel to X-ray film for nucleotides
labelled radioactively. Finally, the sequence of the DNA is
read from the gel by starting from the bottom and reading
upward on X-ray film (Fig. 12.18).

(b) Chemical Degradation Method (Maxam and Gilbert

Method) — In the chemical degradation method of DNA
sequencing, the DNA fragment, whose sequence is to
be determined is cleaved in a base specific manner
using chemicals as shown in Fig.12.19. Before the
chemical mediated degradation of the DNA, the DNA is
labelled at the 5' end using enzyme polynucleotidyl

Heat
*DNA + Dimethylsulphate (DMS) Specific for G
Acid
*DNA + Dimethylsulphate (DMS) Specific for A
High NaCl
*DNA + Hydrazine Specific for C

*DNA + Hydrazine Specific for C+T

Fig. 12.19: Base specific cleavage reactions used in Maxam and Gilbert method

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kinase and gamma-32P labelled nucleotide. The

fragments generated are subsequently separated using
high resolution PAGE to resolve the sequence order.
These gels are placed under X-ray film, which then
yields a series of dark bands which show the location of
radiolabeled DNA fragments. The fragments are ordered
by size and therefore, we can deduce the sequence of
the DNA molecule.

12.9.2 Automated DNA sequencing

Majority of DNA sequencing is now done through automated
method. These automated sequencers primarily work
on Sanger method of DNA sequencing. In automated
sequencing, fluorescent labelled dideoxynucleotides
are used which has eliminated the need for radioactive
isotopes. The slab gel has been replaced by polymer filled
capillary tubes in automated equipment. Automation of
DNA sequencing has made the method much quicker and
more reliable. For example, in one year, by using manual
sequencing, on an average, a person can sequence
20,000 to 50,000 bases while automated sequencer can
sequence that long in just a few hours. Furthermore,
total cost of material for one gel using the automated A
method is approximately half, compared to that of the T
G
manual method. In automated DNA sequencing, all four C
T
dideoxy reactions are carried out in a single tube, which T
is possible because each dideoxynucleotide is labelled C
G
with a different fluorescent dye such as Rhodamine 110 G
C
(RHO)-A (which gives green fluorescence), Rhodamine A
A
6G (REG)-C (which gives blue fluorescence), Tetramethyl G
A
Rhodamine (TAMRA)-G (which gives black fluorescence) G
and X-Rhodamine (ROX)-T (which gives red fluorescence). A
C
The contents of the single tube reaction are loaded onto T
C
a single lane of a gel and electrophoresis is done. The A
A
sequence is determined by the order of the dyes coming off A
A
the gel. A fluorimeter and computer are hooked up to the A
gel and they detect and record the fluorescent dye attached A
T
to the fragments as they come off the gel. All the DNA A

fragments labelled with fluorescent dideoxynucleotide are

‘read’ by a laser and the fluorescence intensity translated Fig. 12.20: Chromatogram
into a data ‘peak’ (Fig. 12.20). of automated
sequencing

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Box 3

Next-generation Sequencing (NGS)

NGS methods have facilitated the sequencing of very large DNA such as whole genome
as the above mentioned methods work optimally for short sequence analysis up to
few Kb size DNA and it is difficult to analyse the whole genome as it will take too
much time and cost. The evolution of DNA sequencing from the historical methods
of Sanger (Sanger sequencing) and Maxam & Gilbert (Maxam-Gilbert sequencing ) to
today’s high-throughput technologies has occurred at a breathtaking speed. In the
past 30 years or so, these high-throughput technologies have given rise to super-
exponential growth in sequence data generation and the resultant data have led to
transformative applications ranging from basic biology to criminal investigation and
prenatal diagnostics.
NGS allows massively parallel sequencing reactions and therefore, they are
capable of analysing millions or even billions of sequencing reactions at the same
time. The widely used NGS platforms are Roche/454 sequencing, Solexa/Illumina
and SOLiD platforms.

12.10 DNA Microarray

DNA microarray technology is a high throughput
hybridisation-based technique that is used to analyse a
large number of DNA fragments in parallel for quantification
of the expression of large number of genes. It uses the
property of two DNA strands to pair with each other by

Fig. 12.21: DNA Microarray chip

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forming hydrogen bonds between complementary

nucleotide bases. Hence, the principle of DNA microarray
technology is that complementary DNA sequences can be
used to hybridise to immobilised DNA fragments on the
chip and individual hybridisation events can be recorded.
Thousands of single stranded DNA (ssDNA) segments
corresponding to the gene transcripts (mRNA) or other
genomic regions, are immobilised on a small solid surface
and are referred as microarray chips (Fig. 12.21). These
chips are usually made of either glass or nylon, and are
coated with a special surface coating that allows spotting
of DNA on the chip. ssDNA segments immobilised on the
chips are called as probes and are arranged in rows and
columns on the chip. This arrangement of the probes
helps to find the location of any specific fragment on the

Fig. 12.22: An example of a cDNA microarray experiment

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chip. Usually probes are either cDNAs, PCR amplicons

or oligonucleotides that correspond to the mRNAs and
are referred as cDNA or oligonucleotide probes.
Oligonucleotide probe-based arrays are very popular.
These probes are short sequences that are complementary
to the known/predicted transcripts from a single species,
and allows to analyse the expression of thousands of
genes in parallel.
A typical microarray experiment includes the
following steps (Fig. 12.22).
1. Extraction of mRNA
2. Probe labelling
3. Hybridisation and washing
4. Scanning and data analysis

Extraction of mRNA
In a cell, transcription from a gene produces a mRNA,
which makes proteins by translation. Depending upon the
requirement, many RNA copies of the same gene are formed.
Therefore, activity of a particular gene can be quantified
by quantification of mRNA. Because mRNA is degraded
very easily, it is converted into complimentary DNA (cDNA)
through reverse transcription, which is more stable and
represents actively transcribing genes in the cell.

Probe Labelling
cDNA fragments are digested with restriction endonucleases
and resulting fragments are attached with fluorochrome
dyes. Cy3 and Cy5 are most commonly used fluorescent
dyes to this purpose. Therefore, probes are made of
thousands of labelled nucleic acid fragments.

Hybridisation and Washing

Labelled DNA fragments are hybridised with the microarray
chip. For hybridisation, DNA chips (having thousands
of single stranded probes) are exposed and allowed to
react with single stranded fluorochrome labelled DNA
fragments. These fragments bind to their complementary
single stranded probes on the chip and form duplexes. The
number of duplexes formed is the reflection of the number
of its DNA segments complementary to its probe. Other
DNA fragments, which do not find their complementary

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probe on the chip are washed away.

Scanning and Data Analysis

Finally, microarray chip with hybridised labelled DNA
fragments are scanned using a highly sophisticated
scanner. Image analysis is performed using sophisticated
software program that helps to determine how much
labelled cDNA is bound to target probes on the chip.
Unique addresses of these target probes and their
association with specific genes is used to interpret and
quantify data. Microarray data analysis software uses
different colours to represent the expression level of genes
in one condition with respect to other condition. Genes
whose expression increases in one condition with respect
to an other condition are called as upregulated genes,
and other whose expression decreases are called as
downregulated genes. Classically, green colour is used
to represent upregulated genes and red colour is used to
represent downregulated genes.

12.11 Flow Cytometry

There are diverse varieties of cells in organisms, which
perform one or the other function. Understanding
features of a specific cell, whether physical or chemical
may provide valuable information from different
perspectives. Such an understanding of cells based on
either of the parameters mentioned earlier may be used
for qualitative or quantitative measurement of cells.
History of quantifying cells based on physical or chemical
properties can be traced back to the Coulter counter
used during late 1950s , which was invented to quantify
particle in suspension based on the principle of change
in impedance proportional to the particle volume. Such
impedance can be detected under electric field when
the particle passes through an area separated by two
chambers having electrolyte solution (Fig. 12.23).
Present day’s flow cytometry is also based on the same
principle of impedance due to particle passing along
with flowing channel of fluid. In order to achieve this,
cells (sample) are passed through the flow cytometer in
such a way that cells flow one by one in the fluid stream.

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Laser beams fixed in the passage detects cells one by one

based on its properties or the dye used to label the same. The
deflected light is then detected by a sensor, which based on
the intensity of light source and the deflected light provides
information about particle or cell. These detectors may
either be in the line of the light beam to detect the surface
property or volume, or it may be perpendicular, which
can detect the internal properties. Signals are received
by sensor and ultimately an image of the object emerges
comprehensively (Fig. 12.23). Similarly, cells present in
the immune system may have thousands of antigens on
their surface. In order to locate any antigen or protein in a
cell, specific antibodies are used quite often. Fluorescence
dye is commonly used to label antibodies for the purpose
of easy identification and localisation. Different cells of a

Sample

Red detector

Flow Chamber
Q1 Q2
Q3 Q4

Blue detector

Sensor
Laser

Waste

Fig. 12.23: A diagrammatic representation of flow cytometry

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mixture activated with different fluorescent dye can also

be mechanically separated by this technique called Flow
Cytometry.
Sometimes, cells labeled with different fluorescent
dyes in a mixture is detected on the basis of fluorescence
present on it. For the purpose of separation of cells, first
the mixture of fluoresce labeled cells are loaded on a
charging electrode followed by its release drop-wise. Laser
based sensor present in the path of dropped cells detect
these based on their fluorescent label and the information
is recorded in a computer, which can be seen as the plot
shown in Fig.12.23.
The development of modern, rapid and sophisticated
biological tools and techniques has made biological studies
more accurate, fast, quantitative and reproducible.

Summary
• Simple light microscopy enables us to see things that are
otherwise too small to be observed by our naked eyes.
Studying minute details of internal organisation of cells
is so very diverse that it requires more refined microscopy
like contrast microscopy or electron microscopy.
• Different types of centrifuges like differential centrifuge,
high speed centrifuge, density gradient centrifuge
and ultracentrifuge helps in the separation of various
biomolecules present within cells based on their densities
under the influence of gravitational force and spinning
them in a solution around an axis at high speed using
centrifugal force.
• By means of electrophoresis, many important biological
molecules like DNA, RNA and proteins can be separated
and studied on the basis of charge to mass ratio of
macromolecules under the influence of an electric field.
• Enzyme-linked immunosorbent assay (ELISA) is a
highly sensitive and quantitative immunological assay
for measurement of antigen and antibody concentration
in given sample. Different types of ELISA like direct,
indirect, sandwich and competitive are used in diagnosis
and scientific studies.
• Various different chromatography methods: Adsorption
chromatography, Ion-exchange chromatography, affinity
chromatography and gas chromatography are exploited
for purification of proteins, peptides, amino acids,
sugars, nucleic acids, alkaloids, vitamins and steroids.

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• Similarly, to elucidate the chemical structure of molecules,

spectroscopy techniques are used. Colorimetry technique
measures the transmittance and absorption of light passing
through liquid sample and measures the concentration of
the sample.
• Fluorescence In Situ Hybridisation (FISH) technique uses
fluorescent molecules binding to highly complementary
regions of chromosome and facilitates in the identification of
a particular gene in an individual chromosome and hence,
play crucial role to understand chromosomal abnormalities.
• Sequence of DNA comprises the heritable genetic information
that forms the basis for the developmental programs of
all living organisms. The advent of DNA sequencing has
significantly accelerated biological research and discovery.
Sanger method of DNA sequencing are developed about four
decades earlier is even used today for routine sequencing
applications. Many other sequencing methods called Next
Generation Sequencing are available these days such as;
Roche/454, Solexa/Illumina and SOLiD plateforms.
• DNA microarray analysis assist in analysing expression
levels of large number of genes.
• In flow cytometry, cells pass through a laser beam, allowing
their physical and chemical properties to be analysed.

Exercises
1. The function of ethidium bromide in electrophoresis is to
(a) track the progression of electrophoresis
(b) visualise the DNA molecules
(c) separate the DNA molecules
(d) provide charge to DNA molecules
2. Match the following

Column I Column II
(a) Separation of ionic Affinity chromatography (AFC)
solutes
(b) Separation of Gas chromatography (GC)
biomolecules with
different binding
specificities
(c) Separation of volatile Ion-exchange chromatography (IEC)
components

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3. Mass spectrometry is used to

(a) identify unknown compounds
(b) elucidate the structure of molecules
(c) quantify compounds
(d) All of the above
4. Match the following table with reference to Antigen

ANTIGEN ANTIBODY Procedure

(i) Free Bound to surface Direct ELISA

(ii) Bound Only one labeled primary Indirect ELISA

antibody used

(iii) Bound Labeled secondary Sandwich ELISA

antibody used

5. In DNA gel electrophoresis,

I. Longer DNA fragments remain close to the well.
II. Longer DNA fragments move towards the positive end
of gel.
III. Smaller DNA fragments move close to the positive end
of gel.
IV. Smaller DNA fragments remain close to the well.
Which of the above options are correct
(a) I and III
(b) II and IV
(c) Only II
(d) None of the above
6. For a resolved image of the surface of an object, which of
the following microscopes would you prefer
(a) Transmission electron microscope
(b) Scanning electron microscope
(c) Phase contrast microscope
(d) Fluorescence microscope
7. Match the following:

Column I Column II
(a) Engvall and Perlman Microscopy
(c) Robert Hooke DNA sequencing
(c) Sanger ELISA

8. Which of the following techniques is feasible to quantify

the expression of a large number of genes
(a) Mass spectrometry
(b) Microarray
(c) FISH
(d) Agarose gel electrophoresis

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9. Differentiate between the following types of microscopy

techniques
(a) Scanning electron microscopy (SEM) and transmission
electron microscopy (TEM)
(b) Dark field microscopy and bright field microscopy
(c) Phase contrast microscopy and confocal microscopy
10. Discuss the principle of agarose gel electrophoresis.
11. Name a tracking dye which is used to track DNA as well
as proteins during electrophoresis. What will happen
if you forget to add tracking dye to your sample during
electrophoresis?
12. Two polyacrylamide gels A and B were prepared. Gel A had
4% acrylamide whereas Gel B had 12% acrylamide. Based
on the given information answer the following
(a) Which gel is harder: A or B?
(b) Which gel offers greater friction to the proteins: A or B?
(c) Which gel (A or B) will be used to separate a mixture
containing low molecular weight proteins?
(d) Which gel (A or B) will be used to separate a mixture
containing both low and high molecular weight
proteins?
13. What is a chromatogram? Draw a well labeled diagram of
a chromatogram of a mixture containing three different
solutes.
14. Explain the principle of FISH. How is FISH technique
applied in chromosome painting? What are the advantages
of chromosome painting?
15. Mention the various applications of spectroscopy
techniques.
16. What are major components of UV-visible
spectrophotometer? Explain each in brief.
17. Write the major differences between the Sanger’s method
and Maxam and Gilbert’s method of DNA sequencing.
18. Write the principle of flow cytometry.

Reprint 2025-26

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