1. The document discusses various cytological techniques used in the study and manipulation of cells including microtomy, microscopy, staining, and centrifugation. 2. Microtomy involves sectioning tissues into thin slices using a microtome for microscopic observation while staining allows cellular structures to be visualized through selective uptake of dyes. 3. Centrifugation separates particles according to properties like density, shape and size through application of centrifugal force, and is used to isolate cellular components through differential pelleting or density gradient centrifugation.

1. Lecture on
Cytological Techniques
Dr. Manikandan Kathirvel M.Sc., Ph.D., (NET)
Assistant Professor,
Department of Life Sciences,
Kristu Jayanti College (Autonomous),
(Reaccredited with "A" Grade by NAAC)
Affiliated to Bengaluru North University,
K. Narayanapura, Kothanur (PO)
Bengaluru
Email: manikandan@kristujayanti.com
ORCID ID: 0000000270066334

2. 1. Microtome
2. Microscopy techniques
3. Staining techniques
4. Centrifugation

3. Cytological Techniques
-Cytological techniques are methods used in the study or manipulation of cells.
-These include methods used in cell biology to culture, track, phenotype, sort and screen
cells in populations or tissues, and molecular methods to understand cellular function.
-Must be able to identify normal cells from abnormal cells, and inflammatory from non-
inflammatory cells.
They can be observed either directly or after preservation under the microscope.
1. For direct observation, the specimen needs sufficient contrast. Direct observation is
possible by using vital stains.
Vital stains :- Vital dyes or stains are taken up by living cells without killing them. They
selectively stain intracellular structures without affecting cellular metabolism and function.
For example, Janus green B selectively stains mitochondria, Golgi apparatus, nuclear
chromatin in a dividing cell can be stained by methylene blue; Neutral Red dye or Congo
Red dye can be used to stain yeast cells.
2. Preserved and stained tissues :- For detailed microscopic study, tissues containing cells
are passed through various stages. The stages of cell preparation on a glass slide involves
killing, fixation, dehydration, embedding, sectioning, staining and mounting.

4. PREPARATION OF BIOLOGICAL
SPECIMENS FOR LIGHT MICROSCOPY

5. Sample collection

6. Sample collection

7. Sample collection

8. EXFOLIATIVE CYTOLOGY
Sample collection

11. Methods: There are 5 methods by which the cytological specimens are prepared for
microscopical observation. They are:
1. Teasing
2. Smear preparation
3. Squash preparation
4. Whole mounting
5. Microtomy
1. Teasing:
The muscular tissue is observed by teasing. A bit of tissue is teased in a saline solution. It is
stained by safranin or eosin. The stained material is mounted on a slide and is observed
under the microscope.
2. Smear Preparation
Fluid tissues such as blood are observed by smear preparation. Smear is a thin film of
tissue on a slide.
a. A drop of blood is taken on a slide and is made into thin film with the help of edge of
another slide.
b. It is dried and stained and covered with a cover glass and observed the microscope.

12. 2. Smear Preparation: METHODS OF SMEAR PREPARATION:
i. streaking
ii. spreading
iii. pull apart
iv. touch or impression smear
STREAKING
- Used for preparing mucoid secretions, vaginal
secretions, sputum and gastric content
-use a spatula, dissecting needle or applicator
stick and streak in a zigzag fashion.
SPREADING
- used for thick mucoid secretions
- smears of fresh sputum and bronchial aspirates
PULL APART
- for serous fluids, concentrated sputum, and
enzymatic lavage form the GIT, smears of
urinary sediment, vaginal pool and breast
secretions.
TOUCH IMPRESSION
- Impression cytology being collected From a
patient , using a sterile glass slide with
polished edges.

13. 3. Squash preparation
i. Soft tissues such as testis, onion root tip, etc are observed by squash preparation.
ii. The soft tissue is placed on the slide, the cover glass is dropped over it and gentle
pressure is applied on the cover glass.
iii. The material is made into a thin layer.
- Fairly accurate,
- Simple and reliable tool for diagnosis of lesions.
Based on two essential factors:
• Availability of very small tissue fragments & good preservation of fine cellular details.
• Not effected by edema, hemorrhage, necrosis & calcification.
4. Whole mounting:
Certain objects are transparent and they are mounted entirely as such. This process is called
whole counting and the slide is called whole mount.
5. Microtomy: a technique to make a thin, transparent sections of tissues and cells of
Preserved tissues :- For detailed microscopic study, tissues containing cells are passed
through various stages. The stages of cell preparation on a glass slide involves killing,
fixation, dehydration, embedding, sectioning, staining and mounting.

14. PREPARATION OF BIOLOGICAL SPECIMENS
FOR MICROSCOPICAL OBSERVATION
Steps involved in Microtomy
Steps involved in Microtomy

15. Microtomy: is a cytological technique by which extremely thin transparent sections
of tissues and cells are prepared for microscopic observations.
For detailed microscopic study, tissues containing cells are passed through various stages.
The stages of cell preparation on a glass slide involves killing, fixation, dehydration,
embedding, sectioning, staining and mounting.
1). Killing and fixation :- FIXATION OF CYTOLOGY SPECIMENS
Fixation is the preservation of the cell structure of the material in a life like condition.
Fixation means :
- prevention of degeneration of cells and tissue
- preservation of cells as close as possible to the living state specific periods of time
changes the physical and chemical state of the cells.
-Fixation is done by chemicals or freezing. The chemical used for fixation is called Fixative.
-This process causes sudden death of cells or tissues and preserves freshly killed tissues in
as life like a condition as possible. A good fixative prevents bacterial decay and autolysis.
-It will also make cell components more visible and prepare the cell for staining.
- The commonly used fixatives are Acetic acid, formalin, picric acid, osmium tetroxide,
Formaldehyde, Bouin's solution and Carnoy's fluid.

16. 1). FIXATION OF CYTOLOGY SPECIMENS
Fixation means :
- prevention of degeneration of cells and tissue
- preservation of cells as close as possible to the living state specific periods of time
changes the physical and chemical state of the cells.
AN APPROPRIATE FIXATIVE FOR CYTODIAGNOSTIC PURPOSES SHOULD PERFORM THE
FOLLOWING FUNCTIONS
a. Penetrate cells rapidly
b. Minimize cell shrinkage
c. Maintain morphologic integrity
d. Deactivate autolytic enzymes
e. Replace cellular water
f. Facilitate diffusion of dyes across cell boundaries
g. Help cells adhere to a glass surface
h. Provide consistent results over time

17. 2). Dehydration :- is the removal of water molecules from fixed tissues. In this process
water vapour are removed from cells or tissues using chemical agents. It is done by using
ethanol and benzene.
The fixed tissue is passed through the series of increasing concentrations of alcohol such
as 30% 50% 70% 90% Absolute alcohol (95%).
After treated with 100% alcohol, the material is cleared with benzene. For electron
microscope propylene oxide is used.
Chemical removal of water and fixative from the specimen
• Replace them with dehydrating fluid - dehydrant
•Many dehydrants are alcohols. Several are hydrophilic so attract water from tissue.
•Practiced in graded series
•Progressively decreasing concentration of water
•Progressively increasing concentration of dehydrant.
•Common dehydrants are ethyl alcohol, acetone, normal butyl alcohol , tertiary
•butyl alcohol Glycerine, Dioxan etc.
•Progressively increasing concentrations – 10%, 20%, 30%, 40% …… 100%
•Time required – soft tissues ~30 minutes – Hard/ large tissue- ~6-12 hrs.
Clearing: (Dealcoholization)-Removal of alcohol from the tissues.
Clearing agents- Xylene, Toluene, Chloroform, Benzene, Petrol etc

18. 3). Embedding :- is Block making. The tissues are embedded with supporting medium such
as molten paraffin wax for sufficient hardness.
It hardens up on cooling and provides enough support to allow thin sections.
Very thin sections need to be taken for electron microscopy. Hence plastics are used for
embedding.

19. 4). Sectioning :- The embedding material is cut into thin sections of needed thickness. It is
done by using an instrument called microtome. Microtome is an instrument used to cut
sections of desired thickness.
There are three types of microtome.
They are:
a. Ordinary microtome
b. Freezing microtome
c. Ultramicrotome
For light microscope, the thickness of the section should be 6-8 microns.
For electron microscope, the thickness of the section should be 50-200 millimicrons.

20. 5). Staining :- is a process by which the sections are colored with suitable stain.
- Use of dyes to provide color to various tissue constituents
- The sections are immersed in dyes that stain structures of the cell component.
For example, cytoplasm stains pink with eosin.
Nucleus stains blue with haematoxylin or red with safranin.
6). Dehydration :- Stained sections are immersed in ethanol to remove water. The tissue
becomes more transparent. Dehydration is done gradually by using a series of increasing
concentrations of ethanol in water. Finally the section is placed in 'absolute' alcohol.
30% 50% 70% 90% Absolute alcohol (95%).
7). Mounting :- Cleaned sections are mounted on a slide using a suitable medium like canada
balsam. A drop of canada balsam is placed on the section and a cover slip is placed over it
and the medium is allowed to dry.
8). Labelling: As soon as the slide is prepared, it is marked and labelled.
canada balsam: A natural resin used as a mounting medium. Canada balsam is a commonly used mounting medium to
prepare permanent slides for microscopy. It is produced from the resin of the balsam fir tree and can be combined
with xylene-containing specimens.

21. Slides showing embedded
thin slice of the tissue
samples after microtome
Observation under light
microscope
Microtome

22. Tissue sectioning

23. Slides showing embedded thin slice of the tissue samples after microtome
Under microscope

24. 1. Microtome
2. Microscopy techniques
3. Staining techniques
4. Centrifugation

25. Centrifugation

26. Centrifugation is a technique of separating of substances which involves the application of
centrifugal force.
The particles are separated from a solution according to their size, shape, density, the
viscosity of the medium and rotor speed. The centrifuge is commonly used in laboratories for
the separation of biological molecules from a crude extract. Centrifugation is the technique
of separating components where the centrifugal force/ acceleration causes the denser
molecules to move towards the periphery while the less dense particles move to the center.
Principle:
1. The centrifuge consists of a motor and rotor.
2. The sample to be separated is placed on the rotor. The motor makes the rotor to spin.
3. The spinning produces a force called centrifugal force.
4. The force which causes a substance spinning round an axis, to move away from the
centre is called centrifugal force.
5. The movement of particles in a centrifugal field is called sedimentation.
6. The rate of movement of particles is called sedimentation rate.
7. The sedimentation rate depends on the size and density of the particles.
8. In a solution, particles whose density is higher than that of the solvent, sink (sediment),
and particles that are lighter than it floats to the top.
9. The greater the difference in density, the faster they move (sediment). This is followed by
smaller and less denser particles.

28. The sedimented particle is called pellet. The solution above the pellet is called
supernatant.
The speed of centrifuge is expressed as rpm (revolution per minute).
The greater the rpm, the greater will be the centrifugal force.
The rate of sedimentation depends on –
•The density of the particles
•The size of the particles
•The viscosity of the medium
•The gravitational pull
Revolutions Per Minute (RPM) in regards to centrifugation is simply a measurement of how fast the centrifuge rotor does a full rotation in one minute.

29. Types of Centrifuge
LOW-SPEED CENTRIFUGE
1) Low-speed centrifuge used for sedimentation of heavy particles
2) The low-speed centrifuge has a maximum speed of 4000-5000rpm
3) These instruments usually operate at room temperature.
4) Two types of rotors are used in it:
•Fixed angle- In angle type, the holders and sample tubes are kept at an angle of 30°C from
the central axis.
•Swinging bucket- The holders and sample tubes swing up and run horizontally while
spinning.
5) It is used for sedimentation of red blood cells, the particles are tightly packed into a pellet
and supernatant is separated by decantation.

30. HIGH-SPEED CENTRIFUGES
1. High-speed centrifuges are used in more sophisticated biochemical applications,
higher speeds and temperature control of the rotor chamber are essential.
2. The high-speed centrifuge has a maximum speed of 15,000 – 20,000 RPM
3. The operator of this instrument can carefully control speed and temperature which is
required for sensitive biological samples.
4. Three types of rotors are available for high-speed centrifugation-
•Fixed angle
•Swinging bucket
•Vertical rotors
ULTRACENTRIFUGES
1. It is the most sophisticated instrument.
2. Ultracentrifuge has a maximum speed of 65,000 RPM (100,000’s x g).
3. Intense heat is generated due to high speed thus the spinning chambers must be
refrigerated and kept at a high vacuum.
4. It is used for both preparative work and analytical work.

31. Types of Centrifugation
I). Differential Pelleting (differential centrifugation)- the particles are separated at
different speeds at different times. It is used in the separation of cellular components.
1. It is the most common type of centrifugation employed.
2. Tissue such as the liver is homogenized at 32 degrees in a sucrose solution that
contains buffer.
3. The homogenate is then placed in a centrifuge and spun at constant centrifugal
force (700 rpm for 10 min) at a constant temperature.
4. After some time a sediment forms at the bottom of a centrifuge called pellet I
(Nuclear fraction) and an overlying solution called supernatant I.
5. The overlying solution is then placed in another centrifuge tube which is then
rotated at higher speeds (10,000 rpm for 20 min) in progressing steps.
6. After some time a sediment forms at the bottom of a centrifuge called pellet II
(Mitochondrial fraction such as mitochondria, lysosomes, peroxisomes) and an
overlying solution called supernatant II.
7. The overlying solution is then placed in another centrifuge tube which is then
rotated at higher speeds (100,000 rpm for 1 hr).
8. After some time a sediment forms at the bottom of a centrifuge called pellet III
(Microsome fraction such as ribosomes, golgi apparatus, endoplasmic reticulum)
and an overlying solution called supernatant III (contains cytosol, Proteins, lipids,
carbohydrates).

32. Differential Pelleting
(differential centrifugation)

33. II). Density Gradient Centrifugation
1. This type of centrifugation is mainly used to purify viruses, ribosomes, membranes, etc.
2. A sucrose density gradient is created by gently overlaying lower concentrations of sucrose
on higher concentrations in centrifuge tubes
3. The particles of interest are placed on top of the gradient and centrifuge in
ultracentrifuges.
4. The particles travel through the gradient until they reach a point at which their density
matches the density of surrounding sucrose.
5. The fraction is removed and analyzed by mass spectrometry.
Density Gradient Centrifugation

34. Density Gradient Centrifugation