Cell culture techniques M pharmacy Sem 2

MODERN BIOANALYTICAL
TECHNIQUES
Unit 4
Contents:
1. Basic equipment used in cell culture lab.
2. Cell culture media
3. various types of cell culture
4. general procedure for cell cultures
5. isolation of cells
6. subculture
7. cryopreservation
8. characterization of cells and their applications
9. Principles and applications of cell viability assays (MTT assays)
10. Principles and applications of flow cytometry.
Prepared by:
Samruddhi P Kadam
M Pharm 2nd
semester
PIP
CELL CULTURE
TECHNIQUES

CELL CULTURE:
Cell culture refers to the technique of growing and maintaining cells
in a controlled artificial environment. This process involves isolating
cells from tissues and placing them into a suitable growth medium
that provides the essential nutrients, growth factors, hormones, and
gases (usually CO₂ and O₂), under sterile and regulated temperature
and pH conditions. It is essential for research in cell biology, drug
development, vaccine production, and cancer studies.
BASIC EQUIPMENT USED IN CELL CULTURE LAB
A cell culture laboratory requires specialized equipment to maintain a
sterile environment and provide optimal growth conditions for
cells. The following are the some of instruments used in cell culture
lab.
1. Laminar Air Flow Hood –
 It provides a sterile working area by filtering air through HEPA
filter.
 It helps in preventing contamination of cultures from airborne
particles or microorganisms.
 It is used while handling cells, media, and other materials to
protect them from microbes.
2. CO₂ Incubator –
 The CO₂ incubator is used to maintain an ideal environment for
the growth of cells.
 It controls and maintains specific conditions such as temperature
(usually 37°C), carbon dioxide concentration (typically 5%),
and high humidity.
 The incubator provides an environment that closely mimics the
conditions inside the human body, allowing cells to grow
effectively.
3. Inverted Phase Microscope -

 An inverted microscope is used to observe living cells growing
at the bottom of flasks or dishes.
 Unlike traditional microscopes, objective lenses are used
making it ideal for cell culture observation.
 It allows researchers to assess cell morphology, monitor cell
confluency (the percentage of surface covered by cells), and
detect any contamination.
4. Water Bath –
 The water bath is used primarily to thaw frozen cells quickly
and evenly at 37°C before transferring them to culture vessels.
 It is also used to pre-warm cell culture media and reagents to
physiological temperatures before adding them to cultures.
 it prevents sudden temperature changes that could shock or
damage cells.
5. Centrifuge –
 A centrifuge is used to separate cells from the culture medium
during processes such as harvesting, washing, or subculturing.
 By spinning samples at high speeds, cells are pelleted at the
bottom of the tube, allowing the supernatant to be removed.
 This step is essential for concentrating cells or preparing them
for cryopreservation, staining, or re-seeding.
6. Refrigerator and Freezer –
 A refrigerator, typically maintains temperature at 4°C and is
used to store culture media, reagents, buffers, and other heat-
sensitive solutions.
 Freezers operating at -20°C or -80°C, are used for long-term
storage of enzymes, reagents, cell lysates, and occasionally
short-term preservation of biological samples or media.
7. Autoclave –
 The autoclave is used to sterilize culture media, glassware,
instruments, and waste materials by exposing them to high-
pressure saturated steam at 121°C for about 15–20 minutes.

 This process ensures that all microbial contaminants, including
spores, are destroyed, which is crucial to prevent contamination
in cell culture work.
8. Cell Counter –
 Cell counters are used to determine the number of cells in a
suspension.
 Manual counters involve the use of a hemocytometer and
microscope, while automated counters use image analysis or
electrical impedance to count cells rapidly.
 Accurate cell counting is essential when seeding cells at specific
densities or calculating viability.
9. Micropipettes –
 Micropipettes are precision instruments used to handle very
small volumes of liquids.
 Micropipettes ensure accurate and reproducible measurements,
which is important in experiments where exact concentrations
are needed.
 They are available in fixed or adjustable volume ranges and
must be regularly calibrated.
10. Cryopreservation System –
 For long-term storage of cell lines, cryopreservation is
necessary.
 This is usually done using a deep freezer (-80°C) or a liquid
nitrogen tank (-196°C).
 Cryoprotective agents such as DMSO are added to the cell
suspension to prevent ice crystal formation during freezing,
which can damage cells.
CELL CULTURE MEDIA
Cell culture media is a nutrient-rich solution that provides all
the essential components required for the growth, survival,

and proliferation of cells in vitro. It mimics the natural
environment of cells inside the body and supports their
physiological activities.
Types of Cell Culture Media
1. Natural Media
 It is obtained from biological sources like plasma, serum, or
tissue extracts.
 These media contain a broad range of nutrients, growth factors,
and proteins that support the growth of various cell types.
 The exact composition of these media is not fully defined thus
they are often considered less reproducible than synthetic media.
 Examples: Plasma clots, embryo extracts.
2. Synthetic or Defined Media
 Defined media are chemically formulated, with known
quantities of specific components, including amino acids,
vitamins, salts, glucose, and other nutrients.
 These media are highly reproducible, making them ideal for
experimental research where precise control over growth
conditions is required.
 Examples: RPMI-1640, DMEM (Dulbecco’s Modified Eagle
Medium), MEM (Minimum Essential Medium), F-10, F-12.
3. Serum-containing Media
 Serum-containing media include animal-derived serum, usually
fetal bovine serum (FBS), which provides growth factors,
hormones, lipids, and other nutrients necessary for cell growth.
 These media are commonly used for routine cell culture due to
their ability to support the growth of a wide variety of cell types.
4. Specialized Media

 Specialized media are custom-formulated for specific cell types
or experimental needs.
 These media are optimized to provide the unique set of nutrients
and growth factors required for optimal growth and function of
certain cells.
 For example, media designed for culturing neural cells,
hepatocytes, or stem cells contain unique components to support
the specific needs of these cells.
Basic Components of Cell Culture Media
a. Amino Acids
 These are the building blocks of proteins and are necessary for
cell growth and function.
 Essential amino acids must be supplied externally as cells
cannot synthesize them.
b. Carbohydrates (Glucose)
 Acts as the primary energy source for the cells.
 Glucose is commonly added in concentrations between 1–4.5
g/L depending on the cell type.
c. Vitamins
 Act as coenzymes in metabolic reactions.
 Vitamins like B-complex and ascorbic acid are usually added.
d. Inorganic Salts
 Maintain osmotic balance and membrane potential.
 Common salts include sodium chloride, potassium chloride,
calcium chloride, and magnesium sulfate.
e. Buffering Agents (e.g., Sodium Bicarbonate)
 Maintain the pH of the medium (generally between 7.2–7.4).

 CO₂ incubators maintain a balance between CO₂ in the air and
bicarbonate in the medium.
f. Phenol Red (pH Indicator)
 Added to monitor pH changes.
 Media appears red at normal pH, turns yellow in acidic
conditions, and purple in alkaline conditions.
g. Serum (e.g., Fetal Bovine Serum – FBS)
 Contains growth factors, hormones, attachment factors, and
transport proteins.
 Typically added in 5–10% concentration depending on cell type.
 In serum-free media, these factors are added separately
VARIOUS TYPES OF CELL CULTURE
 Adherent Cells
 Suspension Cells
1. Primary Cell Culture:
 Primary cell culture is the culture initiated directly from cells
taken from tissues of an organism.
Types of cell
cultures
Primary Cell
Culture
Secondary Cell
Culture
Finite Cell Line

 These cells are isolated by enzymatic or mechanical methods
and then transferred into a suitable culture medium.
 These cells have a limited lifespan.
 They are more sensitive to conditions and contamination.
 These are further divided as
1. Adherent Cells – These cells require attachment to a solid
or semi-solid surface for their growth and proliferation in
vitro.
2. Suspension Cells – These cells grow freely floating in the
culture medium without needing to attach to a surface
2. Secondary Cell Culture:
 Secondary culture is obtained by subculturing or passaging
primary cells to a new vessel with fresh medium.
 This process is done to expand the cell population or continue
the culture.
 Cells may undergo slight changes in morphology and growth
pattern.
 Cells still have a finite life span.
3. Finite Cell Line:
 A finite cell line is a culture derived from primary or secondary
cells that can divide only for a limited number of generations
(usually 20–80 passages) before undergoing natural death.
 They have limited replicative potential.
 Commonly used in research and drug testing.
4. Continuous Cell Line:
1. A continuous or immortalized cell line is a cell line that has
acquired the ability to divide indefinitely, usually by mutation or
artificial transformation
2. Cells have an unlimited lifespan.

3. They often show altered characteristics such as rapid growth or
loss of contact inhibition.
4. Widely used in research and industrial applications.
GENERAL PROCEDURE FOR CELL CULTURES
1. Preparation of Lab and Materials
 The cell culture work is done inside a biosafety cabinet or
laminar airflow hood, which provides a sterile environment by
filtering the air.
 All glassware, media, and instruments used must be sterile to
avoid microbial contamination.
 Personal protective equipment (PPE) such as gloves, lab coats,
and masks must be worn.
 The incubator is set at the appropriate temperature (usually
37°C), with 5% CO₂ to maintain the proper pH for cell growth.
2. Preparation of Culture Media
 A suitable culture medium is selected based on the type of cells
 The medium is supplemented with fetal bovine serum (FBS),
antibiotics (penicillin-streptomycin), glucose, and amino acids
as needed.
Preparation
of Lab and
Materials
Preparation
of Culture
Medium
Thawing or
Isolating Cells
Seeding the
Cells
Monitoring
and
Maintenance
Subculturing
(Passaging)
Cells
Freezing the
Cells
(Cryopreserva
tion)
Reusing the
Cells

 The prepared medium is stored at 2–8°C until use.
3. Thawing or Isolating Cells
 If frozen cells are to be used, they are rapidly thawed in a 37°C
water bath for 1–2 minutes.
 The thawed cells are transferred into a culture flask containing
warm fresh medium.
 For isolating primary cells from tissues, enzymatic digestion or
mechanical disruption is used to separate the cells.
4. Seeding or Plating the Cells
 The cells are counted using a hemocytometer
 A suitable number of cells are seeded into a culture vessel
containing pre-warmed culture medium.
 The vessel is placed in a CO₂ incubator at 37°C for cell growth.
5. Maintenance of Cell Culture
 Cells are monitored under an inverted microscope daily to
observe cell morphology, attachment, and confluency.
 The medium is changed every 2 to 3 days to supply fresh
nutrients and remove waste products.
6. Subculturing (Passaging) Cells
 When adherent cells reach 80–90% confluence, they must be
subculture to avoid overcrowding
 The medium is removed, and the cells are washed with PBS
 An enzyme like trypsin-EDTA is added to detach the cells from
the surface.
 The cells are suspended in fresh medium and transferred to new
flasks at a lower density.
7. Cryopreservation
 Cells can be preserved for long-term use by freezing them in
liquid nitrogen (−196°C).

 Cells are suspended in a freezing medium containing
cryoprotectants such as dimethyl sulfoxide (DMSO) and FBS.
 The freezing is done gradually, typically using a controlled-rate
freezer or stepwise freezing method to prevent ice crystal
damage.
 Frozen vials are stored in liquid nitrogen tanks.
8. Revival and Reuse of Cells
 When needed, frozen cells are revived by quick thawing and
seeded into fresh culture medium for continued growth and
experimentation.
ISOLATION OF CELLS
Isolation of cells is the process of separating individual cells from a
tissue or fluid so they can be grown in vitro (outside the body)
under controlled laboratory conditions.
Sources of Cells for Isolation
1. Animal or human tissues – such as liver, skin, or kidney.
2. Body fluids – such as blood, bone marrow, or amniotic fluid.
3. Organs – used in organ or explant culture.
4. Cell lines – derived from previously established cultures.
General Methods for Cell Isolation
1. Mechanical Method
 In this method, tissues are physically broken down into smaller
pieces to release cells.
 Tissue is chopped using a sterile scalpel or scissors.
 Cells are released by shaking or pressing the pieces through a
mesh or sieve.

 The cell suspension is filtered to remove debris and large tissue
fragments.
Advantages:
 Simple and quick.
 Does not damage surface proteins of the cells.
Limitations:
 May result in lower cell yield.
 Not suitable for tissues with strong cell-to-cell adhesion.
2. Enzymatic Method
 This method uses enzymes to break down the extracellular
matrix and cell junctions to release individual cells from the
tissue.
 Tissue is cut into small pieces and incubated with the enzyme
solution at 37°C.
 The enzyme breaks down the tissue structure and releases
individual cells.
 The enzyme activity is stopped by adding culture medium with
serum.
 The cell suspension is filtered and centrifuged to collect cells.
Commonly Used Enzymes:
 Trypsin – digests proteins and detaches cells.
 Collagenase – breaks down collagen in connective tissues.
 Dispase – separates epithelial cells.
 Hyaluronidase – degrades hyaluronic acid in connective tissues.
Advantages:
 Gives a high number of viable cells.

 Gentle on cell structures when used properly.
Limitations:
 Over-digestion can damage cells.
 Some surface markers may be lost during digestion
3. Combination Method
 A combined approach uses both physical and enzymatic
treatment for better efficiency.
 Tissue is first chopped or minced mechanically.
 Then treated with enzymes like collagenase or trypsin.
 After digestion, cells are filtered, centrifuged, and suspended in
culture medium.
Advantages:
 Gives better cell yield and viability.
 Widely used in research and tissue engineering.
SUBCULTURE
Subculture is the process of transferring cells from one culture vessel
to another to provide more space and nutrients for continued growth.
It is done when the cells become too crowded (confluent) in the flask
or dish.
Purpose of Subculture:
 To prevent overcrowding of cells.
 To maintain healthy, actively growing cells.
 To increase the number of cells for experiments.
 To preserve cells at a certain passage number.

When to Subculture:
 When adherent cells cover about 80–90% of the surface area
(called confluence).
 When suspension cells reach a high density in the medium.
 Usually done every 2 to 3 days, depending on cell growth rate.
Procedure for subculture
Check Cell Confluency
Use an inverted microscope to ensure cells are 80–90% confluent
Remove Culture Medium
Aspirate the old medium carefully using a sterile pipette.
Wash Cells with PBS
Add PBS to remove residual serum and waste then discard
Add Trypsin-EDTA
Add a small amount to detach cells from the surface.
Collect and Centrifuge
Transfer to a tube and centrifuge to collect the cell pellet.
Resuspend and Replate Cells
Discard supernatant, resuspend cells in fresh medium, and transfer to a new flask
Incubate Flask at 37°C in CO₂
Place the new culture in the incubator for further growth.
Stop Trypsin Activity
Add complete growth medium (with serum) to neutralize trypsin.
Incubate at 37°C for 2–5 min
Allow cells to round up and detach from the flask.

Significance of Subculturing
 Maintains Healthy Cell Growth
 Prolongs Cell Line Life
 Increases Cell Quantity
 Reduces Toxic Waste Accumulation
 Preserves Cell Morphology and Characteristics
CRYOPRESERVATION
 Cryopreservation is based on the principle of preserving living
cells or tissues by cooling them to extremely low temperatures
(usually –196°C in liquid nitrogen), which halts all metabolic and
biochemical activities.
 This allows cells to remain viable for extended periods without
growth, degradation, or loss of function.
 At low temperatures, especially below –130°C, enzyme activity
and cellular processes completely stop. This halts the cell cycle and
prevents aging, mutation, or cell death.
 Ice crystals can physically damage cell membranes and organelles
by puncturing them. To avoid this, cryoprotective agents (CPAs)
like DMSO or glycerol are used.
Purpose of Cryopreservation:
 To store valuable or rare cell lines for long-term use.
 To maintain genetic stability and reduce the need for continuous
culturing.
 To create a cell bank for experiments, therapy, or production.

Importance of Cryopreservation:
 Ensures consistent supply of the same cell line.
 Reduces risk of contamination by minimizing handling.
 Saves time and cost by avoiding repeated subculturing.
 Maintains genetic and functional stability of cells.
 Supports research, biopharma, and clinical applications.
Steps in Cryopreservation:
Preparation of
Cells
Cryoprotective
Agent Addition
Aliquoting
Controlled
Cooling
Storage in
Liquid Nitrogen

1. Preparation of Cells:
 Healthy, actively growing cells are harvested and counted.
 Cells are centrifuged and resuspended in a
cryopreservation medium.
2. Cryoprotective Agent Addition:
 A cryoprotectant such as DMSO (Dimethyl Sulfoxide) or
glycerol is added to the cell suspension to prevent ice
crystal formation.
3. Aliquoting:
 The cell suspension is transferred into sterile cryovials and
labeled with cell type, passage number, and date.
4. Controlled Cooling:
 Cells are cooled slowly (about 1°C per minute) to avoid
cell injury. This is often done using a controlled-rate
freezer or an isopropanol freezing container.
5. Storage in Liquid Nitrogen:
 After reaching –80°C, cryovials are transferred to liquid
nitrogen tanks (–196°C) for long-term storage.
Significance of Cryopreservation
 Long-Term Storage of Biological Materials
 Preservation of Rare and Valuable Cell Lines
 Maintains Genetic Stability
 Reduces Contamination and Handling Risks
 Facilitates Cell Banking
 Enables Transportation of Cells Across Region

CHARACTERIZATION OF CELLS AND THEIR
APPLICATIONS
Cell characterization involves identifying the structural, functional,
and molecular features of cells. This is essential in fields such as
cell biology, biotechnology, pharmacology, and regenerative
medicine. The goal is to understand cell behaviour, phenotype, and
responses to various stimuli or environments.
Techniques for Cell Characterization:
1. Microscopy:
 Microscopy is a fundamental technique that allows for the
visualization of cell morphology.
 Light microscopy is used to observe basic cell structure and
count cells, whereas fluorescence microscopy employs
fluorescent dyes or proteins to highlight specific components
such as organelles or proteins.
 For higher resolution, electron microscopy is used to examine
fine details of cell ultrastructure, including membranes,
organelles, and cytoskeletal elements.
2. Flow Cytometry:
 Flow cytometry is a powerful analytical tool used to measure
physical and chemical characteristics of cells as they flow in a
fluid stream through a laser beam.
 It enables the detection of cell size, granularity, and the presence
of specific surface or intracellular markers using fluorescent-
labelled antibodies, making it invaluable for
immunophenotyping and cell sorting.
3. Cell Viability and Proliferation Assays:
 Cell viability and proliferation assays are employed to assess the
health and growth potential of cells.
 MTT and XTT assays measure metabolic activity by converting
tetrazolium salts into colored formazan products.

 The Trypan Blue Exclusion Test differentiates live cells, which
exclude the dye, from dead cells, which absorb it.
4. Molecular Techniques:
 Molecular techniques, such as PCR (Polymerase Chain
Reaction) and RT-PCR (Reverse Transcription PCR), are used to
detect and quantify gene expression at the DNA and RNA
levels.
 Western blotting identifies specific proteins using antibodies,
while ELISA (Enzyme-Linked Immunosorbent Assay)
quantifies proteins like cytokines or enzymes in cell lysates or
culture supernatants.
5. Immunocytochemistry/Immunofluorescence:
 Immunofluorescence techniques involve the use of antibodies
tagged with fluorescent dyes or enzymes to detect the presence
and distribution of specific proteins within cells.
 These techniques are crucial for studying protein localization,
signalling pathways, and cellular responses to stimuli.
Applications of Cell Characterization:
1. Drug Development and Toxicity Testing:
 Test effects of drugs on cell viability, gene expression, or
signaling pathways.
 Helps in screening and safety evaluation.
2. Cancer Research:
 Identification of tumor cell markers.
 Understanding tumor heterogeneity and progression.
3. Stem Cell Research:
 Characterize stem cells and their differentiation into
specialized cells.

 Ensures purity and potency of stem cell-based therapies.
4. Immunology and Vaccine Development:
 Characterization of immune cells (e.g., T-cells, B-cells).
 Monitor immune responses to pathogens or vaccines.
5. Regenerative Medicine and Tissue Engineering:
 Ensures the quality of cells used to repair or replace damaged
tissues.
6. Clinical Diagnostics:
 Detection of disease biomarkers (e.g., CD markers in
leukaemia).
 Prenatal and genetic disorder screening.
7. Biopharmaceutical Production:
 Characterize production cell lines (e.g., CHO cells) for
consistent therapeutic protein production.
 PRINCIPLES AND APPLICATIONS OF CELL VIABILITY
ASSAYS (MTT ASSAYS)
Cell Viability Assays
Cell viability assays are designed to determine the number of live,
healthy cells in a population. These assays measure various aspects of
cell health, such as membrane integrity, enzyme activity, or metabolic
function.
Principle of the MTT Assay:
The MTT assay works on the principle that metabolically active
(viable) cells contain NAD(P)H-dependent oxidoreductase enzymes
that can reduce the MTT compound (3-(4,5-dimethylthiazol-2-yl)-2,5-

diphenyltetrazolium bromide) into insoluble purple formazan crystals.
These crystals accumulate in live cells, while dead cells lack this
activity and do not produce formazan. The amount of formazan
formed is directly proportional to the number of viable cells. The
formazan crystals are then solubilized using a suitable solvent (such
as DMSO, isopropanol, or acidified ethanol), and the resulting
colored solution is quantified by measuring the absorbance using a
spectrophotometer or microplate reader, typically at 570 nm.
Advantages of the MTT Assay:
 Simple, rapid, and cost-effective.
 Suitable for high-throughput screening using 96-well plates.
 Provides quantitative results.
Limitations:
 MTT is not suitable for non-adherent cells unless they are
immobilized.
 The assay does not distinguish between apoptosis and necrosis.
 Only provides indirect measurement of viability (based on
metabolism).
Procedure for Conducting the MTT Assay:
1. Cell Seeding:
 Seed the cells into a 96-well plate at a desired density (typically
5,000–10,000 cells per well) in an appropriate culture medium.
 Incubate the plate for 24 hours at 37°C in a CO₂ incubator to
allow the cells to adhere and grow.
2. Treatment:
 After incubation, treat the cells with test substances (e.g., drugs,
plant extracts, nanoparticles) at various concentrations.

 Include a control group with no treatment (only culture
medium).
 Incubate the plate for an additional 24–72 hours depending on
the experimental design.
3. Addition of MTT Reagent:
 Prepare the MTT solution (usually 5 mg/mL in phosphate-
buffered saline, PBS).
 Add 10–20 µL of the MTT solution to each well containing 100
µL of medium.
 Incubate the plate for 3–4 hours at 37°C in the dark to allow
formazan crystals to form inside viable cells.
4. Solubilization of Formazan Crystals:
 After incubation, carefully remove the medium without
disturbing the crystals.
 Add 100–150 µL of solubilizing agent (commonly DMSO,
acidified isopropanol, or SDS in HCl) to each well.
 Shake the plate gently or incubate for 10–15 minutes to
completely dissolve the formazan crystals and form a uniform
colored solution.
5. Measurement:
 Measure the absorbance of each well at 570 nm using a
microplate reader.
 A reference wavelength (e.g., 630 nm) may also be used to
correct background absorbance.

Applications of the MTT Assay:
1. Cytotoxicity Testing:
 MTT assay is widely used to evaluate the cytotoxic effects of
drugs, chemicals, or nanoparticles on cultured cells.
2. Drug Screening:
 It helps in determining the effectiveness of new
pharmaceutical compounds by measuring their ability to
inhibit cell viability or proliferation.
3. Cancer Research:
 Used to assess the sensitivity of cancer cells to various
chemotherapeutic agents.
4. Cell Proliferation Studies:
 Monitors the rate of cell growth over time under different
experimental conditions.
5. Toxicological Studies:
 Evaluates the potential toxicity of environmental
pollutants or cosmetic products on living cells.
6. Biocompatibility Testing:
 Assesses whether medical devices or biomaterials are safe
for contact with human tissues based on their impact on
cell survival.

PRINCIPLES AND APPLICATIONS OF FLOW
CYTOMETRY.
Principle
 Flow cytometry is a technique used to analyze physical and
chemical characteristics of cells or particles as they flow in a
fluid stream through a beam of light, usually a laser.
 Each cell passes one by one through the laser beam, and as the
light hits the cell, it scatters.
 The amount and direction of light scatter provide information
about the cell’s size and internal complexity.
 if the cells are labelled with fluorescent dyes or antibodies
tagged with fluorescent molecules, the emitted fluorescence is
detected and measured.

Working of a Flow Cytometer:
First, a suspension of cells is prepared and stained with fluorescent
dyes or tagged antibodies, depending on what features are being
studied. This suspension is then injected into the flow cytometer.
Inside the flow cytometer, the sample passes through a narrow nozzle
where the cells are aligned into a single file using a process called
hydrodynamic focusing.
As each cell flows through a focused laser beam, it scatters light and
emits fluorescence if labeled.
There are two main types of light detected:
1. Forward Scatter (FSC): This measures the amount of light
scattered in the same direction as the laser beam. It gives
information about the size of the cell.
2. Side Scatter (SSC): This measures light scattered at a 90° angle
and provides information about the internal complexity or
granularity of the cell.
Applications of Flow Cytometry:
1. Immunophenotyping:
 Used to identify different types of immune cells based on
specific surface markers (e.g., CD4, CD8, CD19).
 Commonly used in diagnosing blood cancers like leukemia
and lymphoma.
2. Cell Counting and Sorting:
 Accurately counts thousands of cells per second.
 Can be used to sort and isolate specific cell populations
using a special form called Fluorescence-Activated Cell
Sorting (FACS).

3. Apoptosis Detection:
 Helps in identifying cells undergoing programmed cell
death using specific dyes like Annexin V and propidium
iodide.
4. Cell Cycle Analysis:
 Measures DNA content to determine the proportion of
cells in different phases (G0/G1, S, G2/M) of the cell
cycle.
5. Detection of Intracellular Proteins and Cytokines:
 After cell permeabilization, flow cytometry can be used to
detect proteins located inside the cell.
6. Transfection Efficiency:
 Evaluates how successfully foreign DNA or RNA has been
introduced into cells.
7. Stem Cell Research:
 Identifies and isolates stem cells based on specific surface
markers.
8. Microbiology:
 Used to analyze bacteria, yeast, or other microorganisms in
research and clinical samples.

Cell culture techniques M pharmacy Sem 2

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