unit 12 tools and techniques in Biology 2.pdf

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Histochemical and Immuno techniques
Pab and Mab
 In general , Antibodies are normally produced by B cells, which are part of the immune
system, in response to the introduction of foreign substances ,such as infectious agents
into the animals body.
 The antibodies bind to the antigens that cause destruction, thus helping to fight infection.
 Pab - Polyclonal Antibody’s ,(react with multiple epitope) Mab - Monoclonal
Antibody’s.(react with one epitope) .
 Mab also called MCA (Mono Clonal Antibodies)
 Pab - Poly clone: Several type of antibodies that target specific epitope of antigen.
 It is naturally occurring in blood serum.
 Mab: Single clone : Single type of Antibody that target specific antigen protein, it is
prepared in laboratory.
 Mab in laboratory , single B cell produce identical antibodies with similar paratope can
recognize specific epitope of an antigen , specific one antigen and one epitope.
 Pab involves different B.cells  produce different antibody and each with specific
paratope , each antibody recognize specific epitope which present on same antigen.
 Pab – in the serum , in presence of different multiple epitopes on antigen many clones of
B-cells produce antibody of a mixture called polyclonal antibodies.(heterogenous
Antibodies)
 Pab – A heterogenous mixture of antibodies is known as polyclonal antibodies.
 For example: Antibody’s derived from immunized man or animals are heterogenous.
 Immunization with serum albumin, a multi determinant globular protein antigen , usually
results in the production of antibody with different specificity.
 Vaccination (or) immunization with antigen , result B cell response is production of Pab.
 Pab’s are produced against the Antigen entering into the body.

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 The antigen’s will have several epitopes. each can induce specific Antibody production.
 Pab are made using several different immune cells. They have affinity for the same
antigen but have different epitopes.
 Different multiple epitopes on the same antigen (or) protein.
 Pab maybe generated in a variety of species including rabbit ,goat , sheep and donkey.
Conclusion :
 The antibodies produced against different antigenic determinants of an antigen are called
polyclonal antibodies. Each bind specific epitope . They bind with different determinant
sites on the antigen.

epitope
Different

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Pab : Among several , each Antibody bind with specific epitope so Pab bind to different
areas of the target molecule (antigen).
Mab: Single type of antibody bind with single specific epitope of an antigen.
Mab (also called Rocket Antibodies , it is specific for one antigen and one epitope)
 An antibody that is specific for one antigen and is produced by a single B-cell clone is
called Mab.
 A single type of antibodies having the same type of paratope produced against a single
epitope by a single hybridoma clone, is called monoclonal antibody.
 Monoclonal antibodies are pure antibodies with a single specificity to a given antigen.
they are monospecific.
 The two main sources of monoclonal antibodies are myelomas and B lymphocytes.
 Presence of a single type of paratope is the useful feature of the monoclonal antibodies.
Mab’s bind with only one or same type of epitope which is on the same antigens or
different antigen.
 Monoclonal Antibodies were first named by Kohler and Milstein et al in 1973.
 The B cells one get activated will produce Antibody with unique specificity against one
epitope called Mab.
 Single epitope specific B cells can be separated and made immortal after fusion with
tumour cell (Myeloma cell) in a lab to secrete large quantities of Mab artificially in
laboratory.
 Mab produced by Kohler and Milstein used in the diagnosis and treatment of several
diseases. they are also used in screening specific proteins.
 Mab also abbervates as MCA.
 When compare to Pab , the homogeneity of monoclonal antibodies (Mab) is very high.
 Mab can be grown in cell cultures and collected as hybridoma supernatants.
 Specimen used to making Mab is mice or rat.

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Mab : Rocket Antibody

All Mab’s 1,2,3 are similar type

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Pab Mab

Same Antigen with different epitope different antigen with same epitope

Different types of antibody Only one type of antibody identical antibody
(ideal antibody)
Can find different epitope of Can find same epitope of different antigen.
the same antigen.
Used for the diagnosis of variety of diseases.
Among several types each bind with
Different Specific epitope of an Antigen
 Mab’s are monospecific antibody that are same identical because they are made by
identical immune cells that are all clone of a unique parent cells.

 Mab are monovalent affinity , in that they bind same epitope.

Pab : Polyvalent nature

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Difference between Pab and Mab
Pab Mab
Derived from different lymphocyte cell
lines (polyclone)
Derived from a single B cell clone
(monoclone)
Mixed population of Antibody’s produced
(non-identical to each other)
Single antibody specific (identical to each
other ) all antibody are identical
Recognizes multiple epitopes (polyvalent)
on anyone antigen each recognize
specific epitope
Recognize only one epitope
(monovalent)of different antigen
Will have affinity with different epitope of
same antigen (same antigen with different
epitope)
Will have affinity with same epitope of
different antigen (different antigen and
same epitope)
Antibody structurally differ from each
other
they are identical to each other
Multivalent / polyvalent Monovalent
Produce large amount of non-specific
antibody (different antibody)
Can produce large amount of too specific
antibody.
Antibody produced several type Single type (identical)
Inexpensive to produce (cheap) Expensive to produce
No technology is require , naturally
occurring
Hybridoma technology used in laboratory
Short time duration Long time duration
Not powerful tool for clinical diagnosis test Powerful tool for several clinical diagnosis
test , research , therapeutic purpose
Tolerent of small changes in protein
(Antigen) structure
May only recognize a particular protein
(Antigen ) structure
Recognize several epitope (each single
epitope)
Single and same epitope , same
antibodies same epitope

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Several B cells involved Single B cell involved
Synthesis naturally inside the body Artificially in laboratory
Precipitate formed (cross linking formed) No precipitate formed because if an
antigen contain only a single copy of
each epitope (No cross linking)
It shows variability in antibody Once hybridoma is made it is a constant
and renewable source and all antibody
are identical.
B cells and myeloma cells involved
Different parental B cells involved Unique parental B cell
Pab’s are multi specific Mab’s are mono specific
Take participate in precipitation test Don’t take part in precipitation test

Mab
 In general , an antigen typically has many epitopes, several different clones of plasma cells
produce different antibody against antigen called Pab.
 A single type of antibody having the same type of paratope produced against a single
epitope by a single hybridoma clone is called Mab.
 If a single plasma cell isolated and induced to proliferate into a clone of identical plasma
cells, then a large quantity of identical antibody are produced.
 Mab can be raised by fusion of B lymphocytes with immortal cell cultures to produce
hybridomas.
 Hybridomas will produce many copies of the exact same antibodies.
 Monoclonal antibodies react with one epitope on the antigen.(Pab antibodies often
recognize multiple epitopes)
 The hybridoma is made by fusion of a lymphocyte (B cell) with a myeloma cell (Tumour
cell)

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 Presence of single type of paratope is the important feature and useful feature of the mab.
 Mab can bind with the only one type of epitope of different antigen.
 Hybridoma technique developed by Jerne Kohler and Milstein in 1973 (They got NP in 1984)
 Kohler and Milstein , Mab in the diagnosis and treatment of several diseases.
 Used for the screening specific proteins
 used for the diagnosis of pregnancy,allergy and diseases such as hepatitis , rabies and
some STD’s.
 Mab used to study metastasis - Cancer (spread by blood and lymph body fluid)
 Mab used to prepare vaccines to counteract the rejection associate with transplants ,
auto immune diseases and to treat cancer.

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How Mab produced?

Have ability to produce specific antibodies in a large amount
to make cell immoral
antigen antigen
Mab: identical antibodies can
recognize specific epitope which
present on different Antigen

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Steps:
 Isolation of B lymphocytes
 Isolation of myeloma cells
 Somatic cell fusion
 Selection of hybrids
 Screening of the hybridoma’s
1. Isolation of B lymphocytes
 2 – 4 week old mice are immunized with same antigen consist single epitope.
 A mouse is killed 3 – 4 days of immunized and its spleen is taken and fragmented.
 A small fragment of spleen is spliced and sterilized
 Spliced fragments treated with enzymatic treatment for the isolation of spleenocytes.
 Spleenocytes are antibody producing B lymphocytes (B-cells).
 Result : B cells are isolated by this method from the immunized mouse.
 B cells are grown in the fresh medium for cell fusion.
2. Isolation of myeloma cells
 Myeloma cells are bone marrow tumour cells
 They are fast growing large cells of haemopoietic portion of bone marrow.
 Myeloma cells is taken from a bone affected by tumour.
 The myeloma cells have the ability to produce a specific antibody in a large amount
 In general – tumour cells lack the enzyme HGPRT (hypoxanthine Guanine Phospho
Ribosyl Transferase).
 In this enzyme added in the medium it can synthesis purine nucleotide.
3. Somatic cell fusion
 B cell and myeloma cells are mixed together.
 B cell  from spleen
 Myeloma cell  From bone marrow
 at the ratio 2 – 5 : 1 with PEG.

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 Other fusogens are PVA, Sandai virus .
 The fusogen brings the two cells get together and induce cell fusion.
 Result : Spleenocyte – myeloma hybrids called hybridoma’s are formed.
4. Selection of hybrids :
 Hybridoma cells are cultured in the medium containing HAT.
 The cell suspension is a diluted with a serum free medium containing HAT.
 HAT – Hypoxanthine Aminopterin Thymidine.
 Selection breeds are kept in incubated 25 – 29°C for 2 – 3 weeks.
 HAT medium helps to fuse B and Myeloma cells hybrids and prevent fusion of B-B cell
hybrids and myeloma – myeloma hybrid.
5. Screening of hybridomas:
 Hybridoma cells is transferred into subcultured.(secondary culture)
 The hybridoma clone, producing Mab is higher quantities (large amount of identical
Mab’s)
 All Mab’s are identical and with single identical paratope.
 In the Mab’s are used for
o Analysis of viral antigen
o Analysis of surface marker (CD’s)
o Identify human lymphocytes and sub population of lymphocytes
o Diagnosis of HLA
o Study to autoimmune diseases.
o Study of cardiovascular diseases.
o Diagnosis of cancer and also in transplanted rejection.

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Applications of monoclonal antibodies:
1. Research field:
 In genetic engineering, monoclonal antibody is used to screen recombinants.

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 In immunological studies, Mab’ s are used to identify various cell types involving in
immune responses and to detect interactions among them. For this purpose . Mab’s are
raised against the cell surface antigens and used as markers to identify the cells.
 The rejection of transplanted organs, especially kidney transplants, is associated with
OKT-3 antigen present on the surface of T cells.
 Monoclonal antibody against OKT -3 antigen is made and injected into the patient to
suppress the transplant rejection. The Mab named orthoclone OKT -3 is mainly used for
this purpose.
 Monoclonal antibodies are used to determine the structure of cell membranes.
 They are employed in serological classification of closely related bacteria, viruses and
protozoans.
 They are used in radioimmunoassay , ELISA and immune fluorescence assay in research
to identify and detect some target products.
 Mab’s are being used to detect and classify enzymes.

2. Treatment of diseases: (Therapeutic purpose)
 Monoclonal antibodies used against some surface antigens of cancer cells.
 The cancers of lungs, breast, pancreas,etc. are cured by using monoclonal
antibodies.
 Mab’s have prepared against the surface antigens of some parasites.
 These Mab’s are used to control the spread of Troponema pallidium, mycobacterium
leprae, Haemophilus influenza etc.

Clinical diagnosis:
 Typing tissue and blood
 Identifying infectious agents
 Identifying and quantifying hormones
 Measuring protein and drug levels in serum
 Identifying tumour antigens and auto-antibodies
 Identifying specific cells involved in immune response
 Identifying CD molecules for classification and follow-up therapy of leukemias and
lymphomas.

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Pab : different B cells used for their production , Pab are heterogenous , mixed antibodies comprising a
mixture of antibodies , each specific one Epitope.
Mab : single B cells used , Mab derived from specific clone , specific for a single epitope , Mab used for
research , therapeutic , diagnosis of diseases.
Pab and Mab classified based on epitope specificity.
1. Pab : React with various epitopes on a given antigen.
2. Mab : React with specific epitope on a given antigen.

Cross linking
and
precipitation
occur
No cross linking
and No
precipitation
occur.

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Mab : Used as Marker antibodies, for the
diagnosis of varieties of diseases

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List of Monoclonal antibodies for research applications

Types of Mab: (developed in laboratory and injected into patients)
-Ximab – chimeric antibody
-Umab - Human antibody
-Zumab - Humanized antibody
-Omab – Murine antibody (1
st
prepared antibodies by Hybridoma technic by Jerne Kohler
and Milstein).
Chimeric antibody (-Ximab)
 Origin of 2 different sources in Man and Mice.
 Light chain derived from mice OR VH derived from mice and CH1,CH2,CH3
derived from Human. Example: Ritu-ximab.
Human antibody (-Umab)
 Also called mixed Mab , created sources of two different individuals which
belongs to same species.
 VH of one individuals and CH1,CH2,CH3 of another individual.
 It is mixed antibody of Ximab and Umab.
Humanized antibody (-Zumab) :
 VH is synthetic sources and CH1,CH2, CH3 are human sources.
Murine antibody (-Omab)
 First developed Mab , created in mice.
 VH from one mice and CH1, CH2, CH3 from other mice.
 It is highly purified Mab because of single sources.
 It is short life span antibody
 It is obtained by hybridoma technic by Jerne Kohler and Milstein.
------------------------------------

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ELISA ( Enzyme-Linked Immunosorbent Assay ) TEST
o It is very sensitive immunological and serological test.
o ELISA – Technique to identify AIDS Patient.
o When HIV is present in a man, his blood will contain Ab’s (especially Ig G , Ig M), against
HIV.
o ELISA – Identify the specific antigen and Ab’s.
o It is used to detect unknown Ag with the help of a known Ab’s
o Here Ab is labeled with specific enzyme.
o The enzyme act on substrate and produces a colour in a positive test.
o The intensity of the colour can be read by ELISA Reader or Spectrophotometer
o A positive direct ELISA to detect presence of HIV Ag’s
o A positive indirect ELISA to detect Presence of Ab’s against HIV.
o Substrate Enzyme widely used ELISA is “Alkaline Phosphatase”.

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Detection OF Molecules using ELISA
Principle:
ELISA (Enzyme-Linked Immunosorbent Assay) is a highly sensitive technique used to
detect and quantify specific antigens (proteins, peptides, hormones) or antibodies in a
sample. It uses an enzyme-labeled antigen or antibody and a color change reaction for
detection.

An antigen-antibody reaction is used for specific binding. An enzyme (e.g., horseradish
peroxidase, alkaline phosphatase) linked to the antibody or antigen produces a color
change when a suitable substrate (like TMB or OPD) is added. The intensity of the color
(measured using a spectrophotometer) is directly proportional to the amount of the target
molecule.
Steps in ELISA for Detection:
● Coating: The wells of a microtiter plate are coated with either antigen (to detect
antibodies) or antibody (to detect antigens).
● Blocking: Unoccupied sites in the well are blocked with a non-specific protein (like BSA or
skimmed milk).
● Sample Addition: The sample containing the target molecule (antigen or antibody) is
added and allowed to bind to the coated well.
● Enzyme-linked Antibody Addition: A secondary antibody conjugated with an enzyme is
added to bind to the antigen-antibody complex.
● Substrate Addition: A chromogenic substrate (like TMB) is added. The enzyme converts it
into a colored product.
● Detection: The color intensity is measured using an ELISA reader at a specific wavelength,
indicating the amount of target molecule present.
Types of ELISA:
● Direct ELISA: Uses a single enzyme-linked antibody.
● Indirect ELISA: Uses a primary antibody and an enzyme-linked secondary antibody.
● Sandwich ELISA: Uses two antibodies (capture and detection antibody) – highly specific.
● Competitive ELISA: Involves competition between labeled and unlabeled antigens for
antibody binding.

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Enzymes Used in ELISA
In ELISA (Enzyme-Linked Immunosorbent Assay), enzymes are conjugated to antibodies or
antigens. These enzymes react with specific substrates to produce a detectable signal,
usually a color change, which indicates the presence of the target molecule.
Common Enzymes Used in ELISA:
● 1. **Horseradish Peroxidase (HRP)**
- Substrate: TMB (3,3′,5,5′-Tetramethylbenzidine) or OPD (o-phenylenediamine).
- Produces a blue or yellow color depending on substrate and reaction stop solution.
● 2. **Alkaline Phosphatase (AP)**
- Substrate: PNPP (p-nitrophenyl phosphate).
- Produces a yellow color.
● 3. **β-galactosidase**
- Substrate: ONPG (o-nitrophenyl-β-D-galactopyranoside) or CPRG (chlorophenol red-β-
D-galactopyranoside).
- Produces a color change (yellow or red).
● 4. **Glucose Oxidase**
- Substrate: Typically used with peroxidase for color development.
● 5. **Acetylcholinesterase (AChE)** (less common)
- Substrate: Specific acetylcholine derivatives, used in specialized ELISA formats.
Enzyme-Substrate-Color Table:
Enzyme Substrate Color Produced
Horseradish Peroxidase
(HRP)
TMB or OPD Blue or Yellow
Alkaline Phosphatase
(AP)
PNPP Yellow
β-galactosidase ONPG or CPRG Yellow or Red
Glucose Oxidase Coupled with
Peroxidase
Color varies
Acetylcholinesterase
(AChE)
Acetylcholine
derivatives
Color varies

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Detection of Molecules using RIA (Radioimmunoassay)
Principle:
Radioimmunoassay (RIA) is a highly sensitive technique used to detect and quantify
small amounts of biological molecules (e.g., hormones, drugs, peptides) in a sample. It is
based on the principle of antigen-antibody binding, where a radioactively labeled antigen
competes with the unlabeled antigen in the sample for binding to a specific antibody.

The amount of radioactivity measured is inversely proportional to the concentration of
the target molecule.
Steps in RIA for Detection:
● Preparation: A known amount of radioactively labeled antigen is prepared.
● Binding: A specific antibody is added to the mixture of labeled and unlabeled antigens.
● Competition: The labeled and unlabeled antigens compete for binding sites on the
antibody.
● Separation: The bound antigen-antibody complex is separated from the free antigen.
● Detection: The radioactivity of the bound complex is measured using a gamma counter.
● Quantification: The concentration of the target molecule is calculated using a standard
curve.
Applications of RIA:
● Measurement of hormones (e.g., insulin, thyroid hormones, cortisol).
● Detection of drugs and toxic substances.
● Diagnosis of infectious diseases through antibody/antigen detection.
● Detection of tumor markers.
● Monitoring therapeutic drug levels in patients.

Detection of Molecules using Immunoprecipitation
Principle:
Immunoprecipitation (IP) is a widely used technique to isolate and detect specific
proteins or molecules from a complex mixture (e.g., cell lysates) using an antibody that
specifically binds to the target protein. The antigen-antibody complex is precipitated and
then analyzed, usually by SDS-PAGE and Western blotting.
Steps in Immunoprecipitation for Detection:
● Sample Preparation: Cell lysate or sample containing the target protein is prepared.
● Antibody Binding: A specific antibody against the target protein is added to the sample.
● Immune Complex Formation: The antibody binds to the target molecule forming an
immune complex.

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● Capture of Immune Complex: Protein A/G-coated beads (agarose or magnetic) are
added to bind the Fc region of the antibody, allowing the complex to be pulled down
(precipitated).
● Washing: The beads are washed to remove non-specific proteins.
● Elution: The target protein is eluted from the beads.
● Detection: The isolated protein is detected using techniques like SDS-PAGE, Western
blotting, or mass spectrometry.
Applications of Immunoprecipitation:
● Detection and analysis of protein-protein interactions (Co-immunoprecipitation).
● Purification of a specific protein from complex mixtures.
● Identifying post-translational modifications (e.g., phosphorylation, ubiquitination).
● Verifying the presence of specific antigens in a sample.
● Studying protein expression and regulation in cells.

Detection of Molecules using Flow Cytometry
Principle:
Flow cytometry is a powerful analytical technique used to detect and quantify
physical and chemical properties of cells or particles as they pass in a fluid stream through a
beam of light (usually a laser). Fluorescently labeled antibodies or dyes bind to specific
molecules, and the emitted fluorescence is measured to determine the presence and
quantity of the target molecules.

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Steps in Flow Cytometry for Detection:
● Sample Preparation: Cells or particles are suspended in a fluid medium.
● Staining: The sample is treated with fluorescently labeled antibodies or dyes that
specifically bind to target molecules.
● Flow Through Laser: The sample passes through a narrow nozzle, and individual cells flow
one by one through a laser beam.
● Light Scattering: Forward scatter (size) and side scatter (granularity) of light are
measured.
● Fluorescence Detection: Fluorescent signals emitted by the labeled molecules are
detected by photodetectors.
● Data Analysis: The detected signals are analyzed using computer software to quantify
the molecules or cell populations.
Applications of Flow Cytometry:
● Detection and quantification of cell surface and intracellular molecules.
● Cell counting and sorting (FACS – Fluorescence Activated Cell Sorting).
● Immunophenotyping of cells (e.g., T-cells, B-cells).
● Measurement of DNA content and cell cycle analysis.
● Detection of apoptosis, viability, and functional assays in research and clinical
diagnostics.

Detection of Molecules using Immunofluorescence Microscopy
Principle:
Immunofluorescence microscopy is a technique used to detect and visualize specific
molecules (usually proteins) within cells or tissues using antibodies conjugated with
fluorescent dyes.
When illuminated with specific wavelengths of light, these dyes emit fluorescence,
allowing the target molecules to be visualized under a fluorescence microscope.

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Types of Immunofluorescence:
1. **Direct Immunofluorescence:** Uses a primary antibody directly conjugated to a
fluorescent dye.
2. **Indirect Immunofluorescence:** Uses an unlabeled primary antibody and a
fluorescently labeled secondary antibody that binds to the primary antibody.
Steps in Immunofluorescence Microscopy for Detection:
● Sample Fixation: Cells or tissues are fixed (e.g., with formaldehyde) to preserve structure
and immobilize molecules.
● Permeabilization (if needed): Detergents are used to allow antibodies to enter the cell
(for intracellular targets).
● Blocking: Non-specific binding sites are blocked using BSA or serum.
● Primary Antibody Incubation: The sample is incubated with a specific primary antibody
against the target molecule.
● Secondary Antibody Incubation (for indirect method): A fluorescently labeled secondary
antibody is added.
● Washing: Excess antibodies are washed off.
● Visualization: The sample is observed under a fluorescence microscope, and the emitted
fluorescence indicates the presence and location of the target molecule.

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Applications of Immunofluorescence Microscopy:
● Localization of specific proteins within cells or tissues.
● Detection of antigens in medical diagnostics (e.g., autoimmune diseases).
● Studying cell structure and function.
● Visualizing protein-protein interactions and cellular pathways.
● Research on infectious diseases by detecting pathogen antigens in samples.

Detection of Molecules in Living Cells
Introduction:
Detection of molecules in living cells involves techniques that allow visualization,
quantification, or tracking of specific biomolecules (proteins, nucleic acids, ions) in real time
without disrupting the normal cellular processes. These techniques are crucial for studying
dynamic biological processes.
Key Techniques for Detection in Living Cells:
● Fluorescent Proteins (e.g., GFP, RFP): Genes encoding fluorescent proteins are fused to
the target protein to visualize its localization and dynamics in living cells.
● Fluorescent Dyes: Small fluorescent molecules (e.g., DAPI, Calcein-AM) are used to label
specific cellular structures or molecules.
● FRET (Fluorescence Resonance Energy Transfer): Used to study molecular interactions and
conformational changes in real time.
● Live-Cell Imaging: Time-lapse fluorescence microscopy is used to monitor cellular events.
● Biosensors: Genetically encoded sensors detect specific ions or molecules (e.g., Ca²⁺
sensors like GCaMP).
● Quantum Dots: Nanoparticles with fluorescent properties are used for tracking molecules
over long periods.
● Confocal and Two-Photon Microscopy: High-resolution imaging to detect and quantify
molecules in living cells.
Steps (General Workflow):
● Labeling of molecules using fluorescent proteins or dyes.
● Transfection or delivery of biosensors or labeled molecules into cells.
● Maintaining living cells under physiological conditions during imaging (using live-cell
imaging chambers).
● Visualizing the labeled molecules using advanced microscopy techniques (e.g., confocal
microscopy).
● Analyzing the fluorescence signals to determine localization, interaction, or quantity of
the molecules.

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Applications:
● Studying real-time cellular processes such as protein trafficking and signal transduction.
● Monitoring ion fluxes (Ca²⁺, Na⁺, H⁺) in live cells.
● Drug screening and understanding drug-target interactions in living cells.
● Tracking infection pathways of pathogens in host cells.
● Understanding molecular dynamics during cell division, apoptosis, and differentiation.

In Situ Localization by FISH (Fluorescence In Situ Hybridization)
Principle:
Fluorescence In Situ Hybridization (FISH) is a molecular technique used for the detection and
localization of specific nucleic acid sequences (DNA or RNA) within cells or tissues. It utilizes
fluorescently labeled DNA or RNA probes that hybridize to complementary sequences in the
sample, allowing visualization under a fluorescence microscope.
Steps in FISH Technique:
● Sample Preparation: Cells or tissue sections are fixed on a microscope slide to preserve
morphology.
● Denaturation: The DNA or RNA in the sample is denatured (strands separated) to allow
probe binding.
● Probe Hybridization: A fluorescently labeled DNA/RNA probe is applied to the sample
and hybridizes with its complementary sequence.
● Washing: Excess unbound probe is washed off to reduce background fluorescence.
● Visualization: The sample is observed under a fluorescence microscope, and the
fluorescent signals indicate the location of the target sequence.
Applications of FISH:
● Detection of chromosomal abnormalities (e.g., deletions, translocations).
● Gene mapping and karyotyping.
● Detection of specific RNA transcripts within cells.
● Cancer diagnostics (e.g., HER2 gene amplification).
● Identification of microorganisms in clinical samples or environmental samples.
Advantages:
● Highly specific and sensitive for detecting genetic sequences.
● Can be performed on non-dividing cells and tissue sections.
● Allows simultaneous detection of multiple targets using probes with different fluorophores.

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In Situ Localization by GISH (Genomic In Situ Hybridization)
Principle:
Genomic In Situ Hybridization (GISH) is a cytogenetic technique used to localize and
differentiate entire genomes or large chromosomal regions within a sample. It is based on
the hybridization of labeled total genomic DNA from one species to chromosome spreads,
with unlabeled DNA from a closely related species used as a blocking agent to reduce non-
specific binding. The hybridization is visualized using fluorescence microscopy.
Steps in GISH Technique:
● Sample Preparation: Chromosome spreads are prepared on microscope slides.
● Probe Preparation: Total genomic DNA from the species of interest is labeled with a
fluorescent dye.
● Denaturation: Both the chromosome DNA on the slide and the labeled probe DNA are
denatured to form single strands.
● Hybridization: The labeled genomic DNA is hybridized to the chromosome spreads,
binding to complementary sequences.
● Blocking: Unlabeled genomic DNA from a related species is used as a blocking agent to
minimize cross-hybridization.
● Washing: Excess probes are washed off to reduce background fluorescence.
● Visualization: The hybridized chromosomes are visualized under a fluorescence
microscope.
Applications of GISH:
● Identification of parental genomes in hybrids and polyploids.
● Detection of introgression or alien chromosomal segments in breeding programs.
● Comparative genome analysis between related species.
● Karyotyping and studying genome organization.
● Analyzing chromosome evolution in plants and animals.
Advantages:
● Allows the discrimination of genomes from different species in hybrids.
● Useful for studying interspecific hybridization and genome evolution.
● Applicable in both plants and animals.
Limitations:
● Requires high-quality chromosome preparations.
● Cross-hybridization can occur if blocking DNA is not optimized
Specialized

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BIOPHYSCIAL METHODS
Molecular Analysis using UV/Visible Spectroscopy
Principle:
UV/Visible spectroscopy is based on the principle that molecules absorb light in the
ultraviolet (200–400 nm) and visible (400–700 nm) regions of the electromagnetic spectrum.
The amount of light absorbed at a specific wavelength is related to the concentration of the
absorbing molecules (as per Beer-Lambert’s law). This technique is widely used for qualitative
and quantitative molecular analysis.
Beer-Lambert’s Law:
A = ε × c × l
Where:
- A = Absorbance (no units)
- ε = Molar absorptivity (L·mol⁻¹·cm⁻¹)
- c = Concentration of the sample (mol/L)
- l = Path length of the cuvette (cm)
Steps in UV/Visible Spectroscopy:
● Sample Preparation: The sample is prepared in a suitable solvent.
● Blank Measurement: A blank (solvent without analyte) is placed in the
spectrophotometer to set a baseline.
● Wavelength Selection: A specific wavelength where the molecule shows maximum
absorption (λmax) is selected.
● Sample Measurement: The sample is placed in the cuvette, and the absorbance is
recorded.
● Data Analysis: The concentration of the sample is determined using a calibration curve or
Beer-Lambert's law.
Applications of UV/Visible Spectroscopy:
● Determination of nucleic acid (DNA, RNA) and protein concentrations (e.g., A260/A280
ratio).
● Quantitative analysis of drugs and chemicals in pharmaceutical industries.
● Monitoring enzyme kinetics by measuring changes in absorbance over time.
● Characterizing chromophores and conjugated compounds.
● Environmental testing (e.g., water quality analysis).
Advantages:
● Simple, fast, and non-destructive method.
● Requires small sample volume.
● Can be used for both qualitative and quantitative analysis.

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Molecular Analysis using Fluorescence Spectroscopy
Principle:
Fluorescence spectroscopy is based on the principle that certain molecules (fluorophores)
absorb light at a specific excitation wavelength and then emit light at a longer emission
wavelength. The intensity of emitted fluorescence is proportional to the concentration of the
fluorophore, allowing both qualitative and quantitative molecular analysis.
Steps in Fluorescence Spectroscopy:
● Sample Preparation: The sample is dissolved in a suitable solvent and placed in a
cuvette.
● Excitation: The sample is illuminated with light of a specific wavelength (excitation
wavelength).
● Emission Measurement: The emitted light (fluorescence) at a longer wavelength is
measured by the spectrofluorometer.
● Data Analysis: Fluorescence intensity is analyzed to determine the presence and
concentration of target molecules.
Key Features:
● Fluorescence is more sensitive than absorbance-based methods (like UV/Vis).
● Emission wavelength is always longer (lower energy) than excitation wavelength due to
energy loss (Stokes shift).
● Fluorophores can be natural (e.g., aromatic amino acids, chlorophyll) or synthetic dyes.
Applications of Fluorescence Spectroscopy:
● Quantification of biomolecules such as DNA, RNA, and proteins.
● Detection of enzyme activities and kinetics using fluorogenic substrates.
● Studying molecular interactions (e.g., protein-ligand binding, FRET analysis).
● Medical diagnostics (e.g., detection of biomarkers, fluorescence immunoassays).
● Environmental analysis (e.g., detecting pollutants like PAHs).
Advantages:
● Extremely high sensitivity (detects molecules in nanomolar concentrations).
● Can provide information about the environment of the fluorophore (e.g., polarity, pH).
● Suitable for real-time analysis of dynamic processes.
Limitations:
● Not all molecules are fluorescent (requires labeling with fluorophores).
● Fluorescence quenching due to environmental factors can affect accuracy.
● Requires specialized and expensive equipment.

32

Molecular Analysis using Circular Dichroism (CD) Spectroscopy
Principle:
Circular Dichroism (CD) spectroscopy is a technique used to study the chiral properties of
molecules. It measures the difference in absorption of left-handed and right-handed
circularly polarized light by chiral molecules. CD is particularly useful in analyzing the
secondary and tertiary structures of biomolecules such as proteins and nucleic acids.
Steps in Circular Dichroism Spectroscopy:
● Sample Preparation: A solution of the chiral molecule (e.g., protein or DNA) is prepared in
a suitable buffer.
● Light Source: Circularly polarized light (both left and right-handed) is passed through the
sample.
● Measurement: The difference in absorption between left and right circularly polarized
light is measured.
● Data Analysis: The resulting CD spectrum provides information about the structure and
conformational changes of the molecule.
Key Features:
● CD signals are sensitive to the secondary structures of proteins (α-helices, β-sheets,
random coils).
● Far-UV CD (190–250 nm) is used for studying protein secondary structure.
● Near-UV CD (250–320 nm) provides information on tertiary structure and environment of
aromatic residues.
Applications of CD Spectroscopy:
● Determination of secondary structures of proteins (e.g., α-helix content).
● Studying protein folding, unfolding, and stability.
● Analyzing conformational changes in nucleic acids and biomolecules.
● Monitoring ligand binding and structural modifications of macromolecules.
● Comparative analysis of wild-type and mutant proteins.
Advantages:
● Non-destructive method requiring small sample quantities.
● Provides structural information in solution state (close to native environment).
● Fast and relatively easy to perform.
Limitations:
● Cannot provide detailed atomic-level structure like X-ray crystallography or NMR.
● Sensitive to buffer components that absorb in UV range.
● Requires relatively pure samples for accurate analysis.

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Molecular Analysis using NMR and ESR Spectroscopy

1. Nuclear Magnetic Resonance (NMR) Spectroscopy
Principle:
NMR spectroscopy is based on the absorption of radiofrequency radiation by nuclei in a
strong magnetic field. Certain atomic nuclei (such as ¹H, ¹³C) possess a property called spin.
When placed in a magnetic field, these spins align with or against the field. Irradiation with
radiofrequency energy causes transitions between spin states, which are detected and
translated into a spectrum that provides structural information about the molecule.

Steps in NMR Spectroscopy:
● Sample Preparation: The sample is dissolved in a deuterated solvent (e.g., D₂O or CDCl₃).
● Magnetic Field Exposure: The sample is placed in a strong magnetic field.
● Radiofrequency Pulse: A radiofrequency pulse is applied to excite the nuclear spins.
● Signal Detection: The nuclei relax and emit signals that are detected as free induction
decay (FID).
● Data Analysis: Fourier transformation is used to convert FID into an NMR spectrum,
providing structural data.
Applications of NMR:
● Determination of molecular structure (including stereochemistry).
● Identification of functional groups and connectivity of atoms.
● Study of molecular dynamics and conformational changes.
● Quantitative analysis of compounds in mixtures.
● Characterization of complex biomolecules like proteins and nucleic acids.

34

2. Electron Spin Resonance (ESR) Spectroscopy
Principle:
ESR spectroscopy (also called Electron Paramagnetic Resonance, EPR) detects
species with unpaired electrons. When placed in a magnetic field, unpaired electron spins
can transition between energy states when exposed to microwave radiation. This transition is
detected and analyzed to provide information about the electronic environment.
Steps in ESR Spectroscopy:
● Sample Preparation: The sample containing unpaired electrons (e.g., free radicals or
transition metal complexes) is prepared.
● Magnetic Field Exposure: The sample is subjected to a variable magnetic field.
● Microwave Radiation: Microwaves are applied to induce electron spin transitions.
● Signal Detection: The resonance absorption is detected and recorded as an ESR
spectrum.
● Data Analysis: The spectrum is analyzed to determine the structure and environment of
paramagnetic species.

Applications of ESR:
● Detection and characterization of free radicals and reactive intermediates.
● Study of metal complexes and transition state chemistry.
● Investigation of oxidative stress in biological samples.
● Analysis of radiation-induced damage in materials.
● Study of enzyme reaction mechanisms involving radical intermediates.

Molecular Structure Determination using X-ray Diffraction and NMR

1. X-ray Diffraction (XRD)
Principle:
X-ray diffraction is based on the scattering of X-rays by the electron clouds of atoms
in a crystal lattice. When a crystalline sample is irradiated with X-rays, the rays are diffracted
in specific directions. According to Bragg's Law (nλ = 2d sinθ), the diffraction pattern is used
to determine the spacing between atomic planes, and thereby the three-dimensional
arrangement of atoms in the molecule.
Steps in X-ray Diffraction:
● Crystal Preparation: A pure crystal of the compound is obtained.
● Irradiation: The crystal is exposed to a beam of monochromatic X-rays.
● Diffraction Pattern Recording: The scattered X-rays are recorded on a detector.
● Data Processing: Bragg's law is applied to calculate interatomic distances.

35

● Structure Determination: A three-dimensional electron density map is constructed to
model the molecular structure.
Applications of X-ray Diffraction:
● Determination of 3D molecular structures of small molecules and macromolecules
(proteins, nucleic acids).
● Identification of crystal structures and polymorphs.
● Drug design and structural biology.
● Analysis of lattice parameters and defects in solid materials.

2. Nuclear Magnetic Resonance (NMR) Spectroscopy
Principle:
NMR spectroscopy is based on the absorption of radiofrequency radiation by nuclei with a
magnetic moment (e.g., ¹H, ¹³C) when placed in a strong magnetic field. The resulting
resonance signals provide information about the local chemical environment of nuclei,
allowing for detailed structural determination.
Steps in NMR Structure Determination:
● Sample Preparation: The sample is dissolved in a deuterated solvent.
● Magnetic Field Exposure: The nuclei align with or against the applied magnetic field.
● Excitation: Radiofrequency pulses cause transitions between nuclear spin states.
● Signal Detection: Relaxation signals are detected as free induction decay (FID).
● Spectral Analysis: The data is processed (via Fourier transformation) to generate an NMR
spectrum, providing chemical shifts, coupling constants, and structural connectivity.
Applications of NMR in Structure Determination:
● Determination of 3D structures of proteins and nucleic acids in solution.
● Identification of functional groups and molecular connectivity.
● Study of molecular dynamics, folding, and conformational changes.
● Analysis of small molecules, natural products, and complex mixtures.

Molecular Analysis using Light Scattering
Principle:
Light scattering techniques are based on the principle that particles or molecules
scatter incident light. The intensity and angle of the scattered light provide information
about the size, shape, molar mass, and interactions of the molecules. Two commonly used
techniques are Static Light Scattering (SLS) and Dynamic Light Scattering (DLS).

36

Types of Light Scattering:
1. **Static Light Scattering (SLS):** Measures the average intensity of scattered light to
determine molecular weight and radius of gyration.
2. **Dynamic Light Scattering (DLS):** Measures fluctuations in scattered light
intensity due to Brownian motion, allowing determination of hydrodynamic size and size
distribution.
Steps in Light Scattering Analysis:
● Sample Preparation: The sample is filtered or centrifuged to remove dust and
aggregates.
● Illumination: The sample is irradiated with a monochromatic laser beam.
● Scattering Measurement: The scattered light is measured at one or more angles relative
to the incident beam.
● Data Collection: Fluctuations in the scattered light intensity are recorded over time (for
DLS).
● Data Analysis: Using appropriate models (e.g., Stokes-Einstein equation), molecular size
and other parameters are calculated.
Applications of Light Scattering:
● Determination of molecular weight and size of polymers and proteins.
● Characterization of nanoparticles and colloids.
● Studying aggregation and stability of biopharmaceuticals.
● Monitoring conformational changes in macromolecules.
● Analyzing complex fluids and suspensions.
Advantages:
● Non-invasive and requires minimal sample preparation.
● Provides rapid and accurate size measurements.
● Can analyze a wide range of particle sizes (from nanometers to micrometers).

Molecular Analysis using Different Types of Mass Spectrometry

Principle:
Mass spectrometry (MS) is an analytical technique used to determine the molecular mass,
structure, and composition of molecules. It works by ionizing chemical compounds to
generate charged particles (ions), measuring their mass-to-charge ratio (m/z), and
detecting them to produce a mass spectrum.

37

General Steps in Mass Spectrometry:
● Ionization: The sample is ionized to produce charged particles (ions).
● Acceleration: The ions are accelerated into a mass analyzer using an electric or
magnetic field.
● Separation: Ions are separated based on their mass-to-charge ratio (m/z).
● Detection: The ions are detected, and their abundance is measured.
● Data Analysis: The resulting mass spectrum is analyzed to determine molecular mass,
structure, or composition.
Types of Mass Spectrometry:
1. Electron Ionization (EI):
In EI, a high-energy electron beam is used to knock electrons off the sample molecules,
producing positively charged ions. It is commonly used for small organic molecules.
2. Electrospray Ionization (ESI):
ESI involves spraying a solution of the sample through a fine needle under high voltage,
producing ions from large biomolecules like proteins, nucleic acids, and peptides.
3. Matrix-Assisted Laser Desorption/Ionization (MALDI):
MALDI uses a laser and a matrix compound to ionize large biomolecules with minimal
fragmentation. It is widely used for analyzing proteins and polymers.

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4. Time-of-Flight (TOF):
TOF mass spectrometers measure the time it takes for ions to travel a fixed distance. The time
is proportional to the mass-to-charge ratio, allowing for precise mass determination.
5. Quadrupole Mass Analyzer:
A quadrupole uses oscillating electric fields to filter ions based on their mass-to-charge ratio.
It is commonly used in tandem MS (MS/MS) for targeted analysis.
6. Orbitrap and Fourier Transform Ion Cyclotron Resonance (FT-ICR):
These high-resolution techniques measure ion oscillation frequencies or cyclotron motion in a
magnetic field to determine exact masses with very high accuracy.
Applications of Mass Spectrometry:
● Determination of molecular weight and structure of organic and inorganic compounds.
● Protein identification and characterization (proteomics).
● Detection of metabolites, drugs, and toxins.
● Isotopic analysis and elemental composition determination.
● Studying post-translational modifications in biomolecules.
Advantages:
● High sensitivity and accuracy in molecular mass determination.
● Can analyze complex mixtures and trace components.
● Applicable to a wide range of molecules, from small compounds to large biomolecules.

Surface Plasmon Resonance (SPR)
What is SPR?
Surface Plasmon Resonance (SPR) is an optical technique used to study biomolecular
interactions in real-time without the need for labeling. It measures changes in the refractive
index near a metal surface (usually gold) when molecules (e.g., proteins, antibodies) bind to
it.

Surface plasmon Resonance (SPR)

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Principle of SPR:
When polarized light hits a thin metal film (gold) at a specific angle, surface plasmons
(electron oscillations) are generated. When a molecule binds to the surface, the refractive
index changes, causing a shift in the resonance angle of reflected light. This change is
detected and plotted as a sensorgram, which shows binding and dissociation events in real-
time.
Key Components:
● Sensor Chip: Coated with a thin gold layer where biomolecules (ligands) are immobilized.
● Light Source: A monochromatic, polarized light beam.
● Detector: Measures changes in reflected light angle.
● Microfluidics: Allows sample flow across the sensor surface.
Applications of SPR:
● Drug Discovery: Studying drug-target interactions.
● Kinetics & Affinity: Determining binding rates (association/dissociation constants).
● Protein-Protein and Protein-DNA interactions.
● Epitope mapping in antibody characterization.
● Detection of biomolecules (biosensors).

Advantages:
● Label-free detection (no need for fluorescent or radioactive labels).
● Real-time analysis of binding interactions.
● Provides kinetic (ka, kd) and equilibrium (Kd) data.
Limitations:
● Requires high-quality sensor surfaces.
● Sensitive to bulk refractive index changes (e.g., temperature, buffer variations).
● Equipment is costly and requires expertise.

Molecular Analysis using Surface Plasmon Resonance (SPR) Methods
Principle:
Surface Plasmon Resonance (SPR) is an optical technique used for real-time, label-free
analysis of molecular interactions. It measures changes in the refractive index near a sensor
surface when biomolecules (ligands and analytes) interact. SPR occurs when polarized light
hits a thin metal film (usually gold), causing collective electron oscillations (plasmons) at a
specific angle. Binding of molecules to the sensor surface changes this angle, which is
detected and used to study interactions.

40

Steps in SPR Analysis:
● Sensor Chip Preparation: A metal-coated chip (gold) is functionalized with a ligand (e.g.,
antibody, protein).
● Baseline Measurement: A flow of buffer solution establishes a baseline signal.
● Sample Injection: Analyte molecules are introduced to bind with the immobilized ligand
on the sensor surface.
● Resonance Signal Detection: The change in refractive index due to binding is detected
as a shift in resonance angle.
● Data Analysis: Binding kinetics (association and dissociation rates) and affinity constants
(Kd) are determined from sensorgrams.
Key Features of SPR:
● Real-time monitoring of biomolecular interactions.
● Label-free detection (no need for fluorescent or radioactive tags).
● Quantitative analysis of binding affinity and kinetics.
Applications of SPR:
● Drug discovery: Screening of drug-target interactions.
● Studying protein-protein, protein-DNA, and protein-ligand interactions.
● Antibody characterization (e.g., epitope mapping, affinity measurement).
● Biosensor development for detecting pathogens, toxins, or biomarkers.
● Analyzing conformational changes in biomolecules upon binding.
Advantages:
● Real-time and highly sensitive measurement of molecular interactions.
● No labeling required for detection.
● Provides both kinetic and equilibrium binding data.
Limitations:
● Requires high-quality sensor surfaces and proper immobilization of ligands.
● Sensitive to changes in bulk refractive index (e.g., temperature, buffer composition).
Instrumentation is expensive and requires expertise for data interpretation.

Radio labeling Techniques

Detection and Measurement of Different Types of Radioisotopes Used in Biology
Introduction:
Radioisotopes are unstable isotopes that emit radiation and are widely used in biology for
labeling, tracing, and quantifying biomolecules or biological processes. The detection and

41

measurement of these radioisotopes rely on their radiation type (alpha, beta, or gamma)
and require specialized detectors.
Commonly Used Radioisotopes in Biology:
● ¹⁴C (Carbon-14): Used in metabolic studies, radiocarbon dating, and tracer experiments.
● ³H (Tritium): Used for labeling nucleic acids, proteins, and drug compounds.
● ³²P (Phosphorus-32): Used in DNA and RNA labeling (e.g., radioactive probes).
● ³⁵S (Sulfur-35): Used to label sulfur-containing proteins and amino acids.
● ¹²⁵I (Iodine-125): Used in immunoassays and labeling of proteins (e.g., radioimmunoassay).
● ⁹⁹mTc (Technetium-99m): Used in imaging and diagnostic studies.
Detection and Measurement Techniques:
● **Geiger-Müller (GM) Counter:** Detects beta and gamma radiation by ionization of gas
in the detector tube.
● **Scintillation Counters:** Measure beta and gamma emissions by converting radiation
into light signals, detected by photomultiplier tubes.
● **Autoradiography:** Detects radioactively labeled molecules on X-ray films, commonly
used with ³²P and ³⁵S.
● **Gamma Spectroscopy:** Measures gamma radiation energy spectra for isotopes like
¹²⁵I and ⁹⁹mTc.
● **Liquid Scintillation Counting:** Used for detecting weak beta emitters like ³H and ¹⁴C in
liquid samples.
● **Cherenkov Counting:** Detects beta particles (e.g., from ³²P) by their interaction with
water.
Applications of Radioisotopes in Biology:
● Tracing metabolic pathways and biochemical reactions.
● DNA sequencing and mapping (using ³²P-labeled nucleotides).
● Protein labeling and turnover studies.
● Measuring enzyme activity and receptor-ligand interactions.
● Medical imaging and diagnostics (e.g., PET scans using positron emitters).
Safety Precautions:
● Use of shielding (lead or acrylic barriers) to reduce exposure.
● Proper waste disposal of radioactive materials.
● Regular monitoring of radiation exposure using dosimeters.
● Use of fume hoods and protective gear when handling radioisotopes.

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Incorporation of Radioisotopes in Biological Tissues and Cells
Introduction:
Radioisotopes are incorporated into biological tissues and cells to trace and study
biochemical processes, molecular pathways, and cell functions. The radioactive labels allow
detection of molecules with high sensitivity. This method is widely used in research,
diagnostics, and metabolic studies.
Methods of Incorporation:
● **1. Direct Incorporation into Biomolecules:** Radioisotopes such as ³²P, ³⁵S, ³H, or ¹⁴C are
incorporated into biomolecules (e.g., nucleotides, amino acids) which are then taken up
by cells and tissues during normal metabolic processes.

● **2. Use of Radioactively Labeled Precursors:** Cells or organisms are exposed to
radiolabeled precursors (e.g., ³²P-labeled phosphate), which become integrated into
nucleic acids or phosphoproteins.
● **3. In Vivo Labeling:** Radioisotopes are injected into an organism, where they are
distributed and incorporated into tissues through normal physiological processes.
● **4. In Vitro Labeling:** Cells or tissue cultures are incubated with radioactive compounds
to allow uptake and incorporation.
● **5. Covalent Labeling:** Radioisotopes are chemically attached to biomolecules (e.g.,
proteins, antibodies) to track their localization and interaction.
Common Radioisotopes Used:
● ³²P – for DNA/RNA labeling and phosphate metabolism studies.
● ³⁵S – for labeling sulfur-containing amino acids (methionine, cysteine).
● ³H (Tritium) – for labeling nucleotides, steroids, and drugs.
● ¹⁴C – for tracing metabolic pathways and organic compounds.
● ¹²⁵I – for labeling proteins and hormones.
Applications:
● Tracing metabolic pathways (e.g., glycolysis, Krebs cycle).
● Studying DNA/RNA synthesis using radiolabeled nucleotides.
● Protein labeling and turnover studies.
● Receptor-ligand binding studies using radiolabeled hormones or drugs.
● Medical diagnostics and autoradiography for imaging tissue distribution of molecules.
Detection Methods:
● Autoradiography – visualizing radioactive molecules in tissues using X-ray film.
● Liquid Scintillation Counting – for β-emitters like ³H and ¹⁴C.
● Gamma Counters – for γ-emitters like ¹²⁵I.
● Geiger-Müller Counters – for general radiation detection.

43

Safety Precautions:
● Use of shielding and protective clothing to minimize radiation exposure.
● Proper disposal of radioactive waste.
● Work in designated areas with radiation monitoring devices.
● Avoid direct contact and inhalation of radioactive materials.

Molecular Imaging of Radioactive Material
Introduction:
Molecular imaging of radioactive material involves using radiotracers to visualize, track, and
quantify biological processes in living organisms at the molecular and cellular levels. By
combining radioactive markers with imaging techniques, researchers and clinicians can
obtain both functional and anatomical information non-invasively.
Techniques for Molecular Imaging of Radioactive Material:
● 1. **Positron Emission Tomography (PET):** Uses positron-emitting isotopes like ¹⁸F or ¹¹C to
visualize metabolic activity.
● 2. **Single Photon Emission Computed Tomography (SPECT):** Uses gamma -emitting
isotopes (e.g., ⁹⁹mTc, ¹²³I) to obtain 3D images of tissue function.
● 3. **Gamma Camera Imaging:** Detects gamma rays emitted from radiotracers for
planar imaging.
● 4. **Autoradiography:** Uses radiolabeled compounds and X-ray film to visualize
radioactive distribution in tissues.
● 5. **Hybrid Imaging:** PET/CT or SPECT/CT combines functional and structural imaging for
precise localization.
Applications of Molecular Imaging:
● Studying drug distribution, metabolism, and pharmacokinetics.
● Detecting tumors and monitoring cancer progression.
● Analyzing brain activity and neurological disorders (e.g., using ¹⁸F-FDG PET).
● Assessing cardiac function and perfusion.
● Tracking biological processes like glucose metabolism or receptor-ligand interactions.
Common Radioactive Tracers Used:
● ¹⁸F-fluorodeoxyglucose (¹⁸F-FDG) – for cancer and metabolic imaging.
● ¹¹C-labeled compounds – for neurotransmitter studies.
● ⁹⁹mTc-labeled compounds – for SPECT imaging.
● ¹²³I – used in thyroid imaging and brain studies.
● ⁶⁸Ga – used in PET imaging of neuroendocrine tumors.

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Advantages:
● Provides functional and molecular information in addition to anatomical details.
● High sensitivity, allowing detection of minute amounts of tracer.
● Non-invasive and suitable for real-time imaging.
Safety Guidelines for Handling Radioactive Materials
Introduction:
Handling radioactive materials requires strict safety protocols to minimize exposure to
harmful ionizing radiation. Radiation safety guidelines focus on controlling exposure, proper
waste management, and emergency preparedness.
Basic Safety Principles (ALARA):
The ALARA principle (As Low As Reasonably Achievable) is followed to minimize radiation
exposure by optimizing three main factors:
● 1. **Time:** Minimize the time spent near radioactive sources.
● 2. **Distance:** Maximize the distance from radioactive materials (exposure decreases
with distance).
● 3. **Shielding:** Use appropriate shielding (e.g., lead, acrylic) to block or reduce
radiation.
Personal Protective Measures:
● Wear lab coats, gloves, and safety goggles.
● Use dosimeters or film badges to monitor personal exposure.
● Wash hands thoroughly after handling radioactive materials.
● Avoid eating, drinking, or applying cosmetics in radiation work areas.
Handling and Storage:
● Work in designated radiation areas with clear warning signs.
● Use tongs or remote handling tools for strong radioactive sources.
● Store radioactive materials in shielded and labeled containers.
● Regularly check for contamination using radiation survey meters.
Radioactive Waste Disposal:
● Separate solid, liquid, and sharps waste.
● Use properly labeled containers with radiation symbols.
● Follow institutional and government guidelines for waste storage and disposal.
● Never dispose of radioactive waste in regular trash or drains.
Emergency Guidelines:
● In case of a spill, evacuate the area and notify radiation safety officers.
● Use spill kits and follow decontamination procedures.
● Report any accidental exposure immediately.
● Conduct periodic radiation safety training.

45

MICROSCOPIC TECHNIQUES
 Microscopy is the technical field that uses microscopes to observe samples which are
not in the resolution range of the normal-unaided eye.
 Microscope is a scientific-instrument consisting of magnifying lens that enables an
observer to view the minute features distinctly.

THE HISTORY OF MICROSCOPES

 Most read with multiple lenses placed in a tube, observed greatly enlarged objects.
1609 Galileo Galilei developed a compound microscope with a convex and
concave lens.
 Most read 1590-Zaccharias Janssen and Hans Janssen (Dutch eye glass makers)
experimented 1625 Giovanni Johannes Faber coined the term microscope.
 1660s- Extensive use of microscopes in research (Italy, Holland and England). 1665
Robert Hooke looked at a silver of cork through microscope lens & noticed "cells".
 1670 Antonie Van Leeuewenhoek (Father of microscopy) made the single lens
Microscope & developed magnifying lens (~300X).
 17th century Christiaan Huygens, developed a simple 2 lens ocular system 1893
August Kohler developed a key technique for sample illumination.
 1903 Richard Zsigmondy developed ultramicroscope (Nobel Prize in Chemistry, 1925).
 1931 Ernst Ruska co & Max Knoll invented the electron microscope.
 1932 Fritz Zernike invented the phase-contrast microscope that enabled the study of
colourless and transparent biological materials (Nobel Prize in physics, 1953).
 1981- Gerd Binnig & Heinrich Rohrer invented Scanning tunnelling microscope (Nobel
Prize, 1986

USE OF MICROSCOPES IN CYTOLOGY
 Life-scientists use the invaluable tool in the field of medicinal diagnosis and research.
 To visualize the crystalline and molecular structures of cells.
 To conduct cytological screening for blood disorders and other diseases
 To study microorganisms, this allows scientists to develop the vaccines. Being able to
identify the infecting agent is the basis for effective treatment.
 To map the fine details of the spatial distribution of macromolecules within cells.
 To measure the biochemical events in the living tissues.
 To interpret the function of proteins within cells by labeling the proteins with a tag.
 To review chromosomal structure particularly in chromosome abnormalities by staining
techniques.
 To Examine Forensic Evidence.

46

 To study the failures in immune function and molecular studies
 To obtain Digital imaging for storing images and in obtaining second opinions or
returning results to remote locations.
 To monitor the health of a particular ecosystem.
 To diagnose and get symptoms details in the veterinary clinic.
 NOTE: The word "lens" comes from the lentil because the shape of a convex lens is
similar to that of a lentil.

MICROSCOPE Two types :
OPTICAL MICROSCOPE
 Simple Microscope
 Compound Microscope
o Stereozoom Microscope
o Phase Contrast Microscope
o Fluorescent Microscope
Electron Microscope
o Transmission Electron Microscope
o Scanning Electron Microscope

PRINCIPLE: MAGNIFICATION AND RESOLVING POWER MAGNIFICATON
 Magnification is defined as "The degree of enlargement of an object provided by the
microscope for detailed analysis of sample".
 The magnification by microscope is the product of individual magnifying powers of
ocular lens (eye piece) and objective lens.
Magnification = Magnifying Power of ocular lens X Magnifying Power of objective Lens
For example:
If ocular lens is 10X and objective is 40X. Then,
Magnification = Magnifying Power of ocular lens X Magnifying Power of objective lens =10
X 40
= 400X
 The Magnifying Power of Microscope is defined as "The ratio of the final image observed
through the microscope to the size of sample observed via naked eye".

Magnifying Power = The ratio of the final image observed through the microscope

The size of sample observed via naked eye

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 Magnification has no limit, but beyond certain point the view becomes blur or
unclear. This is termed as EMPTY MAGNIFICATION.
 herefore, magnification alone does not provide quality information of the sample.
Thus, Resolution plays a crucial role.

RESOLVING POWER
 Resolving Power is defined as "the performance capacity or ability of the microscope
to distinguish between two very closely associated particles".
 For example;
 Human eye has resolving power of 0.25nm.
 Resolving Power of the microscope is the reciprocal of limit of resolution. Limit of
Resolution is the shortest distance between the two objects when they can be
distinguished as two separate entities.
 Limit of Resolution (d) = 0.61 x λ / n Sin 
 Where, λ - wavelength of light ; n - refractive index of the medium between
specimen and objective.  → half angle formed between the specimen and lens.
 As, Resolving Power of the microscope = 1 / Limit of Resolution
 Therefore, Resolving Power of the microscope = n Sine  / 0.61 Χ λ
 Where, n refractive index of the medium between specimen and objective.  
half angle formed between the specimen and lens. Λ  wavelength of light.

 Since, Resolving Power of the microscope X n Sin  / λ

 Resolving power can be increased by following 3 steps:

o 1. by increasing Refractive Index; n immersion oil = 1.5 n air = 1
o 2. by increasing Sin 
o 3. by decreasing wavelength of light;
 λ blue light = 400nm
 λ red light = 600nm

 NOTE: Numerical Aperture (NA) of the objective is defined as the property of lens that
decides the quantity of light that enters into objective.
Numerical Aperture = n Sin 

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GENERIC CONSTRUCTION OF MICROSCOPE
Any microscope is constructed based on mechanical system and optical
adjustments. Henceforth, could be separated into;
1. MECHANICAL PARTS provides physical support to the optical parts and help in focusing the
sample.
2. OPTICAL PARTS confers required adjustments for magnification and optical pathway.
LIST OF MECHANICAL PARTS
1. Base / Metal stand / Foot
2. Pillar
3. Inclination joint
4. Curved arm
5. Stage
6. Stage knobs
7. Stage clips
8. Revolving nosepiece
9. Coarse adjustment
10. Fine adjustment
11. Body tube
12. Draw tube
LIST OF OPTICAL PARTS
1. Light source / illuminator
2. Diaphragm
3. Sub stage Condenser
4. Objective lens
5. Ocular lens / eye piece
NOTE: Optical pathway is the light path from source of illumination passing through the
optical parts (condenser, specimen, objective lens, ocular lens) and finally creates a
magnified virtual image in eyes of an observer).

SIMPLE MICROSCOPE
A simple microscope works on the principle that when a tiny object is placed within
its focus, a virtual, erect and magnified image of the object is formed at the least distance of
distinct vision from the eye held close to the lens.

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Working Principle:
Light from a light source (mirror) passes through a glass stage with slide containing a
thin transparent specimen. A biconvex ocular lens based on its capacity magnifies the size
of the object, resulting in an enlarged virtual image.

APPLICATIONS OF SIMPLE MICROSCOPE
1. Simple microscope is used to obtain small magnifications such as morphology.
2. Simple microscope is usually used for study of microscopic algae, fungi and biological
specimen.
3. Simple microscope is used by skin specialists to scan for various skin disorders.
4. Simple microscope is used to see the magnified view of different particles present in
diverse soil forms.

COMPOUND MICROSCOPE
 A compound microscope is an optical instrument used to observe the magnified
images of small objects on a glass slide.
 Compound microscopes are so called because they are designed with a
compound lens system.
 The objective lens provides the primary magnification which is compounded
(multiplied) by the ocular lens (eyepiece).
 It provides higher magnification and overcomes the limited clarity of image
observed by stereo or other low power microscopes and reduces chromatic
aberration.
 It facilitates detailed study of specimen in a two-dimensional spatial lane. High-
quality Compound Microscopes are available in Monocular, Binocular, and
Trinocular configurations.

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It has a series of two lenses;
 (i) the objective lens ((4x, 10x, or 100x)close to the object to be observed and
 (ii) the ocular lens or eyepiece (5x-30x), through which the image is viewed by
eye.

Structural parts of a microscope and their functions
There are three structural parts of the microscope i.e. head, arm, and base.
1. Head – The head is a cylindrical metallic tube that holds the eyepiece lens at one end and
connects to the nose piece at other end. It is also called a body tube or eyepiece tube. It
connects the eyepiece lens to the objective lens. The light coming from objectives will
bend inside this tube. In binocular microscopes, they are adjustable so that the viewer can
adjust the eyepiece for maximum visualization.
2. Arm – This is the part connecting the base to the head and the eyepiece tube to the base
of the microscope. It supports the head of the microscope and is also used when carrying
the microscope. Some high-quality microscopes have an articulated arm with more than
one joint, allowing more movement of the microscopic head for better viewing.
3. Base – The base is the lowermost part of the microscope that supports the entire
microscope structure. It provides stability for the microscope. Illuminators, light switches,
and electrical wiring systems are fitted in the base.

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Parts of a microscope.
The optical parts of the microscope are used to view, magnify, and produce an image from
a specimen placed on a slide. These parts include:
1. Eyepiece – The eyepiece (ocular Lens) is closest to the viewer’s eye. They are located at
the top of the microscope. This part is used to look at the specimen. These lenses come in
different magnification powers from 5X to 30X, but the most common ocular lenses are of
10X or 15X magnification. They magnify the image for the second time.
2. Eyepiece tube – It’s the eyepiece holder. It carries the eyepiece just above the objective
lens. In some microscopes, such as the binoculars, the eyepiece tube is flexible and can be
rotated for maximum visualization for variance in distance. For monocular microscopes,
they are none flexible.

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3. Diopter Adjustment – Diopter Adjustment is a control knob present only in the binocular
microscope that is used to change focus on one eyepiece. It is used to correct any
difference in vision and compensate for the differences in vision between the viewer’s two
eyes.
4. Nose piece – A nose piece is a movable circular structure that houses all the objective
lenses. It is also called the revolving turret. It is connected to the body tube and lies just
above the stage. It can be rotated clockwise or counterclockwise to increase or decrease
the magnification. The change in magnification results due to a change in the objective
lens.
5. Objective lenses – The objective lens is the lens that is closest to the specimen. They are
fitted on the nosepiece. A standard microscope has 3 to 4 objective lenses of different
magnifying powers: 4X, 10X, 40X, and 100X. The objective lenses first receive the light
transmitted from the specimen and magnify the image for the first time. Objective lenses
are color-coded and are of different sizes. Size and color depend on the power of the lens.
The smallest lens is of the lowest power, and gradually, the longest will be of the highest
power. The high power lenses i.e. 40X and 100X, are retractable, i.e., their end can be
pushed inward. In most optical microscopes, objective lenses with 100X or more
magnification are of oil immersion type.
6. The Adjustment knobs – Adjustment Knobs are the control knobs used to focus the
microscope on the specimen. These knobs are of two types;
a. Fine Adjustment Knob: Fine Adjustment Knob is used for fine adjustment. It is a smaller
knob and is used to move the stage up or down very slowly. The stage covers a very small
distance on each rotation of the fine adjustment knob. It is used to sharpen the image. It is
mostly used while viewing under high power.
b. Coarse Adjustment Knob: Coarse Adjustment Knob is used for focusing the image under
low power magnification. It is a larger knob and is used to move the stage up or down very
rapidly. The stage is raised or lowered rapidly with the help of a coarse adjustment knob.

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7. Stage – This is the section in which the specimen is placed for viewing. They have stage
clips that hold the specimen slides in place. The most common stage is the mechanical
stage, which allows the control of the slides by moving the slides using the mechanical
knobs on the stage instead of moving them manually.
8. Stage Control Knobs – Stage Control Knobs are the control knobs used to move the stage
mechanically. There are two knobs; one for moving left and right and the other for moving
forward and backward. This will move the slide in the field of vision.
9. Aperture – This is a hole in the microscope stage through which the transmitted light from
the source reaches the stage.
10. Microscopic illuminator – A microscopic illuminator is a light source. In some compound
microscopes, a mirror, which reflects the light from an external source to the sample, is
used. In other optical microscopes, different electric bulbs of low voltages are used as a
constant light source. Commonly used illuminators are tungsten-halogen lamps, 75-150W
Xenon lamps, tin-halide lamps, mercury vapor lamps, etc. The selection of types of bulbs is
based on the requirement of intensity and wavelength for illumination.
11. Condenser – These are lenses that are used to collect and focus light from the illuminator
into the specimen. They are found under the stage next to the diaphragm of the
microscope. They play a major role in ensuring clear, sharp images are produced with a
high magnification of 400X and above. The higher the magnification of the condenser, the
clearer the image. More sophisticated microscopes come with an Abbe condenser that
has a high magnification of about 1000X.
12. Diaphragm – It’s also known as the iris. It is found under the stage of the microscope, and
its primary role is to control the amount of light that reaches the specimen. It’s an
adjustable apparatus, hence controlling the light intensity and the size of the beam of light
that gets to the specimen. For high-quality microscopes, the diaphragm comes attached
with an Abbe condenser, and combined, they are able to control the light focus and light
intensity that reaches the specimen.

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13. Condenser focus knob – This is a knob that moves the condenser up or down, thus
controlling the focus of light on the specimen.
14. Abbe Condenser – This condenser specially designed for high-quality microscopes makes
the condenser movable and allows very high magnification above 400X. High-quality
microscopes normally have a higher numerical aperture than objective lenses.
15. The rack stop – It controls how far the stages should go, preventing the objective lens from
getting too close to the specimen slide, which may damage the specimen. It is responsible
for preventing the specimen slide from coming too far up and hitting the objective lens.
16. Light Switch – Light Switch is an electrical control device. Light switches are used to on and
off the illuminator.
17. Brightness Adjustment – The brightness adjustment system controls the voltage supplied to
the light bulb, controlling the intensity (brightness) of the light bulb.

Compound microscopy classified based on the field observed;
1. Bright-field microscopes
2. Dark-field microscopes

1. BRIGHTFIELD MICROSCOPES
 The bright-field microscope is the simplest optical microscope and is popularly
employed.
 The object to be inspected is normally placed on a clear glass slide, and light is
transmitted though the object.
 This makes the object appear against a bright background, hence the term Bright-
field.

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WORKING PRINCIPLE
 Light from the illumination (light) source from the base of the Microscope stand is
aimed at sub-stage condenser lens.
 The sub-stage condenser lens focuses light through slit in the stage onto the sample.
 The sample absorbs some amount of light based on stain, pigmentation or thickness.
 The projected light from the sample is collected by objective lens and is magnified
according to its capacity, creating a primary image.
 The primary image is magnified by ocular lens (eye piece), which also act as
magnifying glass by allowing the observer to view virtual and magnified image of the
sample.
APPLICATIONS
 Widely used for stained or naturally pigmented or highly contrasted specimens
mounted on a glass microscope slide.
 Used in biology classrooms (mitosis & meiosis, etc.) and clinical laboratories. Used in
pathology to view fixed tissue sections or cell smears / smears.

2. Dark field Microscopes
 Used to observe unstained - transparent specimens. Samples having very close
refractive indices value as that of surroundings are difficult to observe with

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conventional bright-field microscopes, such samples are ideal for observation with
dark background.
 Example: small aquatic organisms, oocytes and other thin-transparent materials with
Refractive Index from 1.2 to 1.4

WORKING PRINCIPLE:
 Light from the illumination (light) source from the base of the Microscope stand is
aimed at dark-field ring.
 Dark-field ring is an opaque disk blocks the central rays of the light.
 The marginal/peripheral light rays are directed to sub-stage dark-field condenser
lens.
 The specimen on the stage is illuminated only with the peripheral oblique rays.
 As a result of this, the field appears dark.
 The scattered ray from bright specimen is collected by objective lens and is
magnified according to its capacity, creating a primary image.
 The primary image is magnified by ocular lens (eye piece), which also act as
magnifying glass by allowing the observer to view virtual and magnified image of
the sample.

APPLICATIONS
 Used for examination of live sample.
 Unstained or lightly stained specimen or fluids could also be observed.

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 Useful for diagnosis of disease.
 The bacterial motility can be studied. Precious stones are viewed.

STEREO MICROSCOPE
 The stereo microscope, also called a Dissecting microscope, as it allows the operator to
manipulate/dissect the specimen while it is being observed through the microscope.
 It provides relatively lower magnification usually below 100x. They provide a close-up, 3-
Dimensional view of objects surface textures.
 Stereo microscopes are used for large biological samples (insects, leaf, tissues...) and
medical science applications as well as in the electronics industry, such as by those who
make circuit boards or watches.
WORKING PRINCIPLE
 The Optical binocular stereo microscope consists of two objective lens and two ocular
lens. Two spatially separated optical path focuses sample on the same point from
slightly different angles. The laterally correct, upright-erect image is obtained.

ADVANTAGES
 They can have a single fixed magnification, several discrete magnifications, or a
zoom magnification system.
 Many stereo microscopes are modular in design. It does not require a slide
preparation.
 It enables to switch from bright-field to dark-field and vice-versa.

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PHASE-CONTRAST MICROSCOPE
 The first phase contrast microscope was developed by FRITZ ZERNIKE (FREDRICK
ZERNIKE) in 1933, hence also referred as ZERNIKE MICROSCOPE.
 The phase contrast microscope enables to differentiate transparent, unstained, living
(without killing or altering the living component) structures.
 Phase contrast is an illumination technique provides greater degree differentiation
inside the cells by phase contrast optics.
WORKING PRINCIPLE
 Light from the illumination (light) source from the base of the Microscope stand is
aimed at annular diaphragm stop.
 The annular-diaphragm-stop allows only the hollow cone of light rays to pass through
sub-stage condenser lens.
 The sub-stage condenser lens focuses light through slit in the stage onto the sample.
 The projected light from the sample is collected by special set of objective lens with
phase plate and phase rings which are placed in the back/rare focal plane of the
objective.
 The direct rays (unaltered amplitude and phase, but retarded by ½ wavelength) from
transparent sample converge on the phase ring within the objective and produce
phase shift.
 The most diffracted rays (altered rays due to difference in density) pass through plate
plate by missing phase ring.
 The convergence of diffracted and direct rays on the image plane results in image.

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ADVANTAGES
 Specimens which have a refractive index similar to their surroundings can be invisible
in Brightfield, but are well defined in Phase Contrast.
 Phase Contrast is normally used to examine unstained biological specimens. Living
microorganisms and their minute details such as Cilia, flagella can be observed.

FLUORESCENT MICROSCOPE
 When a substance absorbs light, the electrons present at the outermost orbit absorbs
energy and get excited; on the way back to the ground state, it emits a part of the
energy absorbed.
 This phenomenon is termed as fluorescence. Fluorescent microscope involves staining of
specimens with special fluorescent dyes (fluorescein, acridine orange, etc). When a
fluorescent dye is applied to a substance, it absorbs a wavelength of light (excitation
wavelength) and emits light of different wavelength (emission wavelength).

WORKING PRINCIPLE
 Illumination (light) is provided by a bright mercury vapor lamp (very expensive
harmful), produces light range of 200-400nm and generates considerable amount of
heat.
 The heat filter absorbs heat, allows UV rays and visible rays by blocking infrared rays.
The exciter filter ensures high energy short wavelength monochromatic light towards
dichroic mirror.
 Dichroic mirror (beam splitter) eliminates visible light and reflects excited UV light to
the dark-field condenser, which provides high contrast for fluorescence and also
deflects majority of UV light.
 The excitation light is focused on to the fluorochrome specimen. The fluorescent
labeled specimen absorbs light and emits excitatory light along with florescent light,
which reaches objective lens.
 As per the capacity of objective lens, the specimen would be magnified and are
directed towards barrier filter.
 The additional barrier filter permits only the fluorescent wavelength and rejects
excitation light.
 The fluorescent light passing through ocular lens creates the magnified image, which
can also be detected by detector to give a photographic image.

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APPLICATIONS
 Imaging the genetic material (DNA & RNA) and other structural components.
 Monitoring the environment for microbial contamination.
 Certain micro-organism can be detected and identified only by this microscopy.

ELECTRON MICROSCOPE

 Electron beam is the source of illumination.
 Image is produced by magnetic field.
 Contrasting features between light microscope and electron microscope are
construction, working principle, specimen preparation, cost-expenses and designed
room (vacuum chamber).
ELECTRON
 Electrons are sub-atomic particles around the nucleus with negative charge. Electrons
have high velocity and shorter wavelength about 0.05 A° [105K times shorter than
wavelength of visible light- 5500 A
 Shorter the wavelength, higher is the resolution.
 Electrons are sensitive to magnetic field.
 In 1924, BROGLIE proposed Dual nature of electrons (wave and particular)
PRINCIPLE OF ELECTRON MICROSCOPE

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 A vacuum chamber with heating metal filament such as tungsten [at about ~6000volts]
generates electron rays.
 Multiple electro-magnetic lenses i.e., the copper wires coiled around hallow cylindrical
tube induces electromagnetic field during current flow and converts electron rays into
electron beam.
 Electron beam is similar to light rays, but have shorter wavelength. Electron beam on
interaction with atoms of the biological sample produces image and is displayed on
fluorescent screen.
 Faster the electron moves, shorter the wavelength and greater is the image quality.
TYPES OF ELECTRON MICROSCOPE
 1.Transmission Electron Microscope [TEM]
 2. Scanning Electron Microscope [SEM]

1. Transmission Electron Microscope (TEM)

 The transmission electron microscope is used to view thin specimens through which
electrons can pass generating a projection image.
 The TEM is analogous in many ways to the conventional (compound) light microscope.
 TEM is used, among other things, to image the interior of cells (in thin sections), the structure
of protein molecules (contrasted by metal shadowing), the organization of molecules in
viruses and cytoskeletal filaments (prepared by the negative staining technique), and the
arrangement of protein molecules in cell membranes (by freeze-fracture).

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Scanning Electron Microscope (SEM)
Transmission Electron Microscopy (TEM)

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 Conventional scanning electron microscopy depends on the emission of secondary
electrons from the surface of a specimen.
 Because of its great depth of focus, a scanning electron microscope is the EM analog of a
stereo light microscope.
 It provides detailed images of the surfaces of cells and whole organisms that are not
possible by TEM. It can also be used for particle counting and size determination, and for
process control.
 It is termed a scanning electron microscope because the image is formed by scanning a
focused electron beam onto the surface of the specimen in a raster pattern.

Scanning Electron Microscopy (SEM)

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Parts of Electron Microscope

Electron Microscope is in the form of a tall vacuum column that is vertically mounted. It
has the following components:
1. Electron gun
 The electron gun is a heated tungsten filament, which generates electrons.
2. Electromagnetic lenses
 The condenser lens focuses the electron beam on the specimen. A second condenser lens
forms the electrons into a thin tight beam.
 The electron beam coming out of the specimen passes down the second of magnetic
coils called the objective lens, which has high power and forms the intermediate
magnified image.
 The third set of magnetic lenses called projector (ocular) lenses produce the final further
magnified image.
 Each of these lenses acts as an image magnifier all the while maintaining an incredible
level of detail and resolution.
3. Specimen Holder
 The specimen holder is an extremely thin film of carbon or collodion held by a metal grid.
4. Image viewing and Recording System
 The final image is projected on a fluorescent screen.
 Below the fluorescent screen is a camera for recording the image.

Application of Electron microscope

 Electron microscopes are used to investigate the ultrastructure of a wide range of
biological and inorganic specimens including microorganisms, cells, large molecules,
biopsy samples, metals, and crystals.
 Industrially, electron microscopes are often used for quality control and failure analysis.
 Modern electron microscopes produce electron micrographs using specialized digital
cameras and frame grabbers to capture the images.
 The science of microbiology owes its development to the electron microscope. The study
of microorganisms like bacteria, virus, and other pathogens have made the treatment of
diseases very effective.

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Advantages of Electron Microscope
 Very high magnification
 Incredibly high resolution
 Material rarely distorted by preparation
 It is possible to investigate a greater depth of field
 Diverse applications

Limitations of Electron Microscope
 The live specimen cannot be observed.
 As the penetration power of the electron beam is very low, the object should be ultra-
thin. For this, the specimen is dried and cut into ultra-thin sections before observation.
 As the EM works in a vacuum, the specimen should be completely dry.
 Expensive to build and maintain
 Requiring researcher training
 Image artifacts resulting from specimen preparation.
 This type of microscope is large, cumbersome extremely sensitive to vibration and
external magnetic fields.

TRANSMISSION ELECTRON MICROSCOPE [TEM]
 The Transmission Electron Microscope [TEM] was first type of Electron Microscope. TEM
was developed by MAX RUSKA in 1931 and was awarded Nobel Prize for Physics in
1986.

WORKING PRINCIPLE
 Electron generator is the source of illumination with a tungsten filament. When heated
by electric current, it emits a stream of electrons.
 The stream of electrons is directed through anode aperture into a condenser lens
system.
 The condenser lens system (1st electromagnetic coils) adjusts the beam and guides the
beam towards the specimen.
 As the electron beam passes through the specimen placed below the condenser,
electron beam is scattered depending on the varying refractive index of the specimen.
 From the specimen, the beam of electrons passes through objective/intermediary lens
(2nd set of electromagnetic coils) forming an intermediary image. The projection lens
(3rd set of electromagnetic coils) produces final image and is projected on a
fluorescent screen/photographic plate.

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PREPARATION OF THE SPECIMEN FOR TEM
1. DEHYDRATION Specimen is dehydrated i.e., water molecules are removed, in order to
avoid shrinkage of specimen under high temperature and preserve the structural
integrity.
2. FIXATION The specimen is mounted in proper orientation and fixed in a required
angle. This minimizes any disturbance in the specimen observation. Cryo-fixation could
also be used.
3. ULTRA-SECTIONING Very thin section Specimen is necessary to visualize their internal
structures. Ultra- sectioning is done with the help of ultra-microtome, which uses a
mechanical instrument to move specimen (embedded in renin) slowly across a knife
surface (made up of glass/diamond) to create thin slices.
4. STAINING Staining is used to improve the contrast between the specimen and the
background. The stains used in TEM contain electron dense heavy metal salts.
There are two types of staining;
1. Positive staining and
2. negative staining.
 In Positive staining, the cell components are combined with metals of high atomic
weight (lead-Pb207, U238) and the specimen appears dark in light background.
 In Negative staining, electron opaque materials (phospho-tungsic acid) are deposited
which does not combine with cell components but make background appear dark
and specimen appears light.

Tem Advantages
 TEM provides most powerful magnification.
 TEM offers detailed and high quality image. T
 They are easy to operate with proper training.

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 TEM is ideal for a number of different fields such as life-sciences, nanotechnology,
medical, biological and material research, forensic analysis, gemology and metallurgy.
 TEM provides topographical, morphological, compositional and crystalline information.
TEM DISADVANTAGES
 TEMs are large and very expensive.
 Dehydration may alter morphological features dealing to mis-interpretation. Requires
large, special housing and maintenance.
 They are expensive and as laborious sample preparation Images are black and
white. Operation and analysis requires special training.

SCANNING ELECTRON MICROSCOPE [SEM]
 Scanning Electron Microscope [SEM] was developed by DENNIS MC MULLAN (PhD
student - England) and CHARLES OUTLAY (Engineer) in 1948.
 SEM generates an image by scanning the specimens with a beam of electrons and
enables topographical study of the specimen surface.
 NOTE: The path of the electron beam within SEM differs from that of the TEM.

WORKING PRINCIPLE
 Electron gun is the source of illumination in a vacuum chamber that produces a
stream of electrons and is directed into a condenser lens, thus generating the narrow
electron beam.
 Rapidly moving electron beam passes through the beam deflector, enters the
objective lens and primary electron beam is created.
 The primary electron beam strikes the specimen, the surface atoms discharge shower
of second electrons and are called as Secondary electrons.
 The secondary electrons are collected by a Scintillator detector (composed of
scintillator and photomultiplier) which generates an electronic signal.
 These signals help in the formation of the final image on a CRT/Video screen. The
secondary electrons emitted from each point on the specimen are characteristic of
the surface.
 The image on the screen thus reflects the composition and topography of the
specimen surface. This image gives a three-dimensional appearance.

PREPARATION OF THE SPECIMEN FOR SEM
1. DEHYDRATION
 SEM allows observing the surface topography. So, dehydration is achieved by critical
point drying which minimizes artifact formation (disturbance in surface configuration).

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 In critical point drying, at a particular temperature and pressure the liquid changes to
gas without any surface tension damage to the specimen.
 The specimen is first immersed in ethanol or acetone to remove water and then in
pressurized liquid of CO2.
 Simultaneously, rising the temperature above 32°C (the critical point of CO2).
 At this temperature, the liquid vaporizes without surface tension leaving the specimen.

SHADOW CASTING
 In this technique, the specimen is coated with an extremely thin layer of gold, gold
palladium or platinum at an oblique angle, so that the specimen produces a shadow
on the uncoated side.
 The shadow casting technique results in three dimensional topographic image of the
specimen. Coating is done with a device called sputter coater.
SURFACE REPLICA
 In this technique a thin layer of a coherent material is coated on to the specimen
evenly.
 The coated specimen is then floated on to a water surface, from where it is transferred
to a strong acid or alkali.
 This dissolves the specimen without damaging the replica.
 This replica is then dried and kept on the mental grid for viewing.

SEM ADVANTAGES
 SEM provides detailed three-dimensional and topographical imaging. Easy to operate
with proper training, associated with user-friendly software.
 SEM is used as research tool and as got various application in the industrial fields. SEM
samples require relatively minimal preparation than TEM.
SEM DISADVANTAGES
 SEM is expensive and occupies large space. Special training is mandatory. Additional
cooling and system maintenance is required.
 SEMs are limited to solid, inorganic samples. Sample size must be small enough to fit
inside the chamber.

Different fixation and staining techniques for electron microscope
1.The coagulant fixatives
1. Ethanol (ethyl alcohol)
2. Picric acid (Trinitrophenol)

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3. Mercuric chloride and
4. Chromium trioxide (Chromic acid)
1.Ethanol
 Ethanol penetrates the tissues quickly but causes shrinkage and hardening.
 Mitochondria are destroyed, lipids may be dissolved. It does not affect
subsequent staining greatly.
2.Picric acid
 Picric acid is maintained under water and dispensed as a saturated solution.
 It is explosive when dry. It may also be used as a dye.
 Picric acid penetrates rather slowly with almost as much shrinkage as ethanol , but
leaves tissues soft and less liable to hardening by ethanol.
 Chromosomes are well preserved , lipids are not dissolved in this acid.
3.Mercuric chloride
 Mercuric chloride penetrates moderately quickly with slight initial shrinkage but
gives about 70% shrinkage on embedding in paraffin.
 It preserves mitochondria , nuclear membrane, nucleolus and the external shape
of the cell particularly well and is often regarded as a good mucopolysaccharide
fixative.
 It produces black pepper grain crystals in the cytoplasm, which are removable by
treatment with iodine solution.
 It is a pre-eminent fixative for producing brilliant staining effects.
4.Chromium – trioxide
 Chromium trioxide penetrates slowly and gives moderate shrinkage and moderate
hardening.
 It converts polysaccharides to aldehydes and provides good fixation of
chromosomes.
 It is a good neutral fixative in the presence of isotonic sodium chloride.

2. Non-coagulant Fixatives
 Formalin (Formalin is a 40% solution of formaldehyde gas in water.
 Osmium tetroxide (osmic acid)
 Potassium dichromate and
 Acetic acid
Formalin
 Formalin penetrates moderately quickly , producing some initial swelling, often with
considerable attendant shrinkage on paraffin embedding.

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 It is a good preservative of lipids.
Osmium tetroxide
 Osmium tetroxide penetrates slowly with little shrinkage and no hardening.
 Mitochondria are exceptionally well preserved, lipids are rendered less soluble in
certain solvents by the use of this fixative.
 Osmium tetroxide preserves the structure of the cell better than any other fixative
which is why it is used in electron microscopy.
Potassium dichromate
 Potassium dichromate penetrates quickly but without much fixing action and causes
no hardening.
 It is very sensitive to pH in its staining reaction.
Acetic acid
 Acetic acid penetrates quickly with great attendant swelling and almost no
hardening.
 It precipitates nucleic acids and mitochondria are not seen in paraffin work.

Plant fixatives
Formalin – alcohol
This is very good for woody twigs, and material may be left in the fluid indefinitely
without harm.
Formalin –acetic alcohol (F.A.A or AFA)
 This is very good for stems, leaves and roots.
 It may be left in the fluid indefinitely without harm.
Animal fixatives
 Formal saline
 Zenker’s fluid
o This is an excellent general fixative giving a good picture with many stains.
 Helly’s fixative
o This has essentially the same formula as Zenker’s differing in only one
respect, and is recommended for use with tissue containing large quantities
of red blood cells, e.g. liver
 Bouin’s fluid
o Bouin’s fluid gives excellent results with a wide variety of tissues and
organisms.
o Fixation for 12 hours to several days is recommended.

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Staining categories
1. Basic dyes
2. Acid dyes and
3. Amphoteric dyes
Basic dyes:
o These are so called because they are essentially alkaline.
o Positively charged and are therefore attracted to and stain acidic tissue
constituents such as the chromatin in nuclei.
o Basic dyes are positively charged (cationic) dyes that readily bind to
negatively charged cell components like nucleic acid.
o There are commonly used in staining because most bacterial cell surfaces
carry a negative charge.
 Typical basic dye will be positive at all pHs.
 Basic dyes are also classified as cationic dyes since they are positively charged and
will move toward a negative charge.
 Examples for basic dyes:
Crystal violet
Methylene blue
Malachite Green
Basic Fuchsin
Safranin
Toluidine blue
Neutral red
Acid dyes:
 These are so called because they are essentially acidic.
 The dye ion are negatively charged and are therefore attracted to and stain
alkaline tissue constituents such as cytoplasm generally.
 Acid dyes are also classified as anionic dyes since they are negatively charged
and will move towards a positive charge.
 The classic amphoteric dyes are carmine and orcein.
 Acidic dyes are negatively charged (anionic) dyes that bind to positively
charged components of cells like proteins and cytoplasm.
 Examples for Acidic dyes
o Eosin
o Acid Fuchsin
o Nigrosin
o Congo Red

72

o Rose Bengal
o India ink
o Picric acid
Amphoteric Dyes
These dyes are capable of both acid and basic reactions, because the negativity or
positivity of their charges changes with pH and they will stain different tissue elements in
different pH solutions.
 The classic amphoteric dyes are carmine and orcein.
 Amphoteric dyes also called Zwitterionic dyes.
 It can act as either basic (cationic) or acidic (anionic) depending on the pH of
the solution.
 They have both positive and negative ionisable groups.
 Examples
o Congorubine
o Aician blue
o Amido Black 10B
o Janus Green B
o Aniline Blue

Types of staining Methods
Single staining : When only one stain is used to give a single colour.
Double staining : Where two stains are used , on the same tissue , each staining a
specific area or structure.
Multiple staining : When more than two stains are used on the same tissue, each
being used for a specific area or structure.

Mordanting
 Mordanting is effectively the conditioning of tissues so that they will accept basic dyes
more readily than they would otherwise.
 Mordants are usually metal salts , most often alums . e.g.Potash alum (which is
aluminium potassium sulphate) , Potassium permanganate , ammonium chromate
,ammonium dichromate.
o The salts are used sometimes in advance of the stain and sometimes with the stain.

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Difference Between Freeze Fracture and Freeze Etching
 The key difference between freeze fracture and freeze etching is that
 freeze fracture is the breaking of a frozen specimen to reveal internal structures,
 while freeze etching is the vacuum-drying of an unfixed, frozen and freeze-fractured
biological sample.
 Freeze fracture and freeze etching are two procedures that are helpful to study details
of three-dimensional structures of biological samples.
 Freeze fracturing is always followed by freeze etching. Freeze fracturing involves the
breaking the frozen biological sample, while freeze etching involves making a platinum-
carbon replica of the fracture face through frozen cells.
What is Freeze Fracture?
 Freeze fracture is a technique of fracturing a frozen biological sample.
 This procedure helps to understand the structure of different cell structures with a
detailed analysis of their functions. In freeze fracturing, the frozen sample is cracked
using a microtome.
 A microtome is a knife-like instrument that cuts thin tissue slices. Freezing of the sample
is done using liquid nitrogen.
What is Freeze Etching?
o Freeze etching is a technique of vacuum drying the frozen fractured biological
sample.
 It involves making a platinum-carbon replica of the fracture face through frozen cells.
Generally, freeze etching is done after freeze fracturing the biological sample.
 Once the replica is made, it is examined under electron microscopy. The freeze etching
process is similar to the normal freeze-drying process of fruits and vegetables selling at
grocery shops.
 Fixation and dehydration are not involved in freeze etching.
What are the Similarities Between Freeze Fracture and Freeze Etching?
 Freeze fracture is followed by freeze etching.
 They are done to biological samples.
 Moreover, the sample needs to be frozen prior to fracturing and etching.
 They are electron microscopy sample preparation procedures.

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What is the Difference Between Freeze Fracture and Freeze Etching?
Freeze fracture is the procedure for breaking apart a frozen biological sample. Freeze
etching is the making of a platinum-carbon replica of the fracture face through frozen cells.
So, this is the key difference between freeze fracture and freeze etching.
Freeze Fracture Vs Freeze Etching
Freeze Fracture Freeze Etching
Definition Freeze fracture is the
procedure that involves
rapid freezing of a
biological sample and
cracking it
Freeze etching is a
vacuum procedure that
makes a platinum carbon
replica of the fractured
surface
Instrument involved Microtome No instrument is used
Major event Cracking of the sample Sublimation of surface ice
under vacuum
Followed by Rapid freezing is followed
by freeze fracture
Freeze fracture is followed
by freeze etching

*****************
Image processing methods in Microscopy
 Microscopy image processing involves various techniques to enhance, analyze, and
extract information from microscopic images.
 These techniques can be broadly categorized into 1.preprocessing, 2.segmentation,
3.feature extraction, and 4.analysis.
 Common methods include noise reduction, contrast enhancement, ima ge
restoration, and object detection, often utilizing both spatial and frequency domain
approaches.
Preprocessing:
 Noise reduction:
Methods like averaging, median filtering, and Wiener filtering are used to reduce noise
and improve image clarity.
 Contrast enhancement:
Techniques such as histogram equalization and contrast stretching are applied to
improve the visibility of features.

75

 Image restoration:
Algorithms like deconvolution and Richardson-Lucy deconvolution can be used to
compensate for imperfections in the imaging system and improve image resolution.
Segmentation:
 Thresholding:
Simple thresholding separates objects from the background based on intensity
values.
 Edge detection:
Algorithms like Canny or Sobel edge detection help identify object boundaries.
 Watershed segmentation:
This technique is useful for separating touching or overlapping objects.
 Clustering:
Algorithms like k-means can group pixels based on similarity, aiding in object identification.
Feature extraction:
 Shape-based features:
Measurements like area, perimeter, and circularity can be extracted from segmented
objects.
 Texture-based features:
Texture analysis methods can be used to quantify surface properties of cells or tissues.
 Intensity-based features:
Measurements of average intensity, maximum intensity, or intensity distributions can be
informative.
Analysis:
 Object counting: Counting the number of objects in an image is a common task.
 Object tracking: Tracking the movement of objects over time can reveal dynamic
processes.

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 Pattern recognition: Identifying patterns or structures in the images can provide insights into
biological processes.
 3D reconstruction: Techniques like confocal microscopy and light sheet microscopy
generate 3D images that can be processed using specialized algorithms.
 Image stitching: Combining multiple image tiles to create a larger field of view is crucial for
analyzing large samples.
 Deep learning: Convolutional neural networks (CNNs) are increasingly used for tasks like
image segmentation, object detection, and image restoration.
Specific applications:
 Confocal microscopy: Quantitative analysis of fluorescent intensity, particle locations, and
dynamics.
 Single-molecule localization microscopy (SMLM): Deep learning-based methods for
improving spatial resolution and processing speed.
 Fluorescence microscopy: Analysis of cellular structures and protein interactions.
 Multispectral imaging: Analyzing images across multiple wavelengths to identify different
components.
 Time-lapse imaging: Studying dynamic processes in living cells.

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