Conversation with Gemini

Cell culture: Basic techniques of cell culture.

Cell culture is a fundamental technique in biology where cells are grown and maintained in an
arti cial environment outside of their natural habitat. It's an invaluable tool for studying cell
physiology, biochemistry, disease, and for producing biological compounds like vaccines and
therapeutic proteins.

Here are the basic techniques involved in cell culture:

1. Aseptic Technique: This is paramount in cell culture to prevent contamination by

microorganisms (bacteria, fungi, mycoplasma) or other cell types. It involves:

• Working in a Laminar Flow Hood (Biosafety Cabinet): This provides a sterile workspace
with ltered air.
• Sterile Equipment and Reagents: All instruments, media, and solutions must be sterile.
• Proper Personal Protective Equipment (PPE): Lab coats, gloves, and eye protection are
essential.
• Minimizing Air Exposure: Keeping culture vessels open for the shortest possible time.
• Disinfection: Regularly cleaning work surfaces with ethanol or other disinfectants.
2. Cell Types and Culture Systems:

• Primary Cell Culture: Cells are directly isolated from tissues (e.g., biopsy) and grown.
These have a limited lifespan in culture.
• Cell Lines: Primary cultures that have been subcultured (passaged) become cell lines.
◦ Finite Cell Lines: Have a limited number of divisions before senescence (aging).
◦ Continuous (Immortalized) Cell Lines: Through genetic changes (spontaneous or
induced), they can divide inde nitely.
• Adherent Culture: Most common for cells derived from vertebrates. Cells attach and grow
on a solid surface (e.g., tissue culture-treated asks) as a monolayer.
• Suspension Culture: Cells grow free- oating in the culture medium, suitable for some cell
types (e.g., hematopoietic cells) or cells adapted for suspension.
3. Culture Medium: This is the "food" for the cells, providing essential nutrients and maintaining a
suitable environment. It's a complex mixture of:

• Salts: For osmotic balance.

• Carbohydrates (e.g., glucose): Energy source.
• Amino Acids: Building blocks for proteins.
• Vitamins: Co-factors for metabolic reactions.
• Minerals: Essential ions.
• Growth Factors and Hormones: To stimulate cell proliferation and differentiation.
• Buffering System (e.g., bicarbonate buffer with CO2): To maintain stable pH (often
indicated by phenol red).
• Antibiotics/Antimycotics (optional and generally avoided if possible): To prevent
bacterial/fungal contamination, but can mask underlying issues.
• Serum (e.g., Fetal Bovine Serum - FBS): Provides a rich source of growth factors and
other components, though serum-free media are increasingly used.
4. Incubation Conditions: Cells require speci c environmental conditions to thrive:
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 • Temperature: Typically 37°C for mammalian cells.
• Humidity: To prevent evaporation of the medium.
• CO2 Concentration: Usually 5% CO2, essential for maintaining pH with bicarbonate-
buffered media.
5. Thawing Cells: Frozen cells (cryopreserved) need to be thawed correctly:

• Rapid Thawing: Vials are quickly transferred from liquid nitrogen (-196°C or -150°C) to a
37°C water bath.
• Dilution of Cryoprotectant: Immediately after thawing, the cryoprotectant (e.g., DMSO) is
diluted with pre-warmed medium.
• Centrifugation (optional): To remove the cryoprotectant and concentrate cells.
• Resuspension and Seeding: Cells are resuspended in fresh medium and seeded into a new
culture vessel.
6. Cell Seeding and Observation:

• Cell Counting: Before seeding, cells are counted (e.g., using a hemocytometer or automated
cell counter) to ensure the desired density.
• Seeding Density: Varies by cell type and experimental needs.
• Observation: Regularly check cells under an optical microscope for:
◦ Viability and morphology: Healthy cells have a characteristic appearance.
◦ Even distribution.
◦ Absence of contamination.
◦ Con uency: The percentage of the culture vessel surface covered by adherent cells.
7. Medium Exchange (Feeding Cells): As cells grow, they consume nutrients and release
metabolic waste products.

• Frequency: Depends on cell type and growth rate, often every 2-3 days.
• Procedure: Carefully aspirate old medium and replace with fresh, pre-warmed medium,
ensuring cells don't dry out.
8. Subculturing (Passaging Cells): When adherent cells reach a certain con uency (e.g., 80-90%)
or suspension cells reach a high density, they need to be "split" into new vessels to provide more
space and fresh nutrients.

• For Adherent Cells:

◦ Washing: Rinse with a balanced salt solution (e.g., PBS) to remove residual medium
and serum.
◦ Dissociation: Use an enzymatic solution (e.g., trypsin-EDTA) or mechanical
scraping to detach cells from the surface.
◦ Inactivation (for enzymes): Add fresh medium containing serum to inhibit the
enzyme.
◦ Centrifugation: To pellet cells and remove the enzyme/inhibitor.
◦ Resuspension and Dilution: Resuspend cells in fresh medium and dilute to the
desired split ratio for new asks.
• For Suspension Cells:
◦ Simply dilute the existing culture into new asks with fresh medium, or centrifuge
and resuspend in fresh medium at a lower density.
• Passage Number: Keep track of the passage number, as cell lines can change characteristics
with increasing passages.
9. Cryopreservation (Freezing Cells): To preserve cells for long-term storage and prevent genetic
drift or contamination:
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 • Preparation: Harvest cells and resuspend them in a freezing medium containing a
cryoprotectant (e.g., DMSO) and serum.
• Slow Freezing: Gradual cooling (e.g., using a controlled-rate freezer or a Mr. Frosty
container) prevents ice crystal formation that can damage cells.
• Storage: Store in liquid nitrogen (vapor phase is recommended) at -150°C or -196°C.
Essential Equipment:

• Laminar Flow Hood/Biosafety Cabinet: For sterile work.

• CO2 Incubator: To maintain temperature, humidity, and CO2 levels.
• Microscope (Inverted): For observing cells in culture vessels.
• Centrifuge: For pelleting cells.
• Water Bath: For warming media and reagents.
• Aspirator/Vacuum Pump: For removing media.
• Sterile Culture Vessels: Flasks, dishes, plates.
• Sterile Pipettes and Tips.
• Liquid Nitrogen Storage Tank: For cryopreservation.
By mastering these basic techniques, researchers can successfully grow and maintain cells in vitro
for a wide range of scienti c applications.

development of primary cell culture

The development of primary cell culture is a meticulous process that begins with the isolation of
cells directly from living tissues. Unlike established cell lines, primary cells retain many of the
characteristics of their tissue of origin, making them highly valuable for research that aims to mimic
in vivo conditions more closely.

Here's a breakdown of the key steps and considerations in the development of primary cell culture:

1. Tissue Collection:

• Aseptic Retrieval: This is the absolute rst and most critical step. The tissue must be
collected from the organism (human or animal) under strict aseptic conditions to prevent
microbial contamination. This often involves surgical excision or biopsy.

• Prompt Transport: The collected tissue should be transported to the laboratory

immediately in a sterile, physiological solution (e.g., balanced salt solution or transport
medium) containing antibiotics to minimize cell damage and bacterial growth. Keeping the
tissue cold (on ice) during transport is often bene cial.
2. Tissue Processing and Disaggregation: The goal here is to break down the complex tissue
structure into a single-cell suspension while maintaining cell viability. This can be achieved
through:

• Mechanical Disaggregation:
◦ Mincing/Chopping: The tissue is nely minced with sterile scissors or scalpels into
small pieces (e.g., 1-2 mm). This increases the surface area for subsequent enzymatic
digestion.
◦ Pressing/Filtering: For softer tissues, pieces can be pressed through sieves of
decreasing mesh size or forced through a syringe and needle to release cells.
◦ Scraping/Teasing: Some tissues can be gently scraped or teased apart to release
cells.
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 • Enzymatic Digestion: This is the most common method for most solid tissues. Enzymes
break down the extracellular matrix (ECM) that holds cells together.
◦ Common Enzymes:
▪ Trypsin: A widely used protease that cleaves peptide bonds. Often used with
EDTA (ethylenediaminetetraacetic acid) to chelate calcium ions, which are
important for cell-cell adhesion.Can be used in "warm trypsinization" (37°C
with stirring) or "cold trypsinization" (extended incubation at 4°C, often
yielding higher viability).

▪
Collagenase: Breaks down collagen bers, which are abundant in connective
tissues. Different types of collagenase (e.g., Type I, Type II) are available,
chosen based on the speci c tissue.
▪ Dispase: A neutral protease that can gently separate epithelial cells from
underlying connective tissue.
▪ Hyaluronidase: Breaks down hyaluronic acid, a component of the ECM.
◦ Optimization: The type and concentration of enzymes, incubation time, and
temperature are critical and need to be optimized for each tissue type to ensure
maximal cell yield and viability while minimizing cell damage. Over-digestion can
severely harm cells.
3. Cell Isolation and Puri cation (Optional but Recommended): After disaggregation, the cell
suspension is often heterogeneous, containing various cell types, tissue debris, and dead cells.
Puri cation steps can enhance the quality of the primary culture:

• Filtration: Passing the suspension through cell strainers with progressively smaller pore
sizes to remove undigested tissue fragments and clumps.
• Centrifugation: To pellet the cells and separate them from enzymes, debris, and dead cells.
Repeated washing with a balanced salt solution helps remove residual enzymes.
• Density Gradient Centrifugation (e.g., Ficoll-Paque): Used to separate different cell types
based on their density (e.g., isolating peripheral blood mononuclear cells from whole blood).
• Selective Adhesion: Some cell types adhere to plastic surfaces faster than others. This
property can be used to enrich for speci c cell populations (e.g., broblasts often adhere
quickly).
• Immunomagnetic Cell Separation (MACS) or Fluorescence-Activated Cell Sorting
(FACS): Highly speci c methods using antibodies conjugated to magnetic beads or
uorescent markers to isolate target cell populations.These are typically used when a highly
pure population of a speci c primary cell type is required.
4. Cell Culture Setup (Seeding):

• Resuspension: The isolated cells are resuspended in a complete growth medium.

• Cell Counting and Viability Assessment: Use a hemocytometer and trypan blue exclusion
assay or an automated cell counter to determine the total cell number and viability.
• Seeding Density: Cells are seeded into sterile tissue culture vessels ( asks, dishes, plates) at
an optimized density. Too few cells may struggle to establish, while too many can lead to
rapid overgrowth and senescence.
• Culture Medium: A specialized medium is crucial. Primary cells often have more stringent
nutritional requirements than immortalized cell lines. The medium typically contains:
◦ Basal medium (e.g., DMEM, RPMI-1640)
◦ Fetal Bovine Serum (FBS) or other animal serum (e.g., horse serum) as a source of
growth factors, hormones, and adhesion factors (though serum-free alternatives are
increasingly available for speci c primary cells).
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 ◦
Growth factors and supplements speci c to the cell type (e.g., EGF for epithelial
cells, FGF for broblasts, insulin, hydrocortisone).
◦ Antibiotics (e.g., penicillin, streptomycin) and antimycotics (e.g., amphotericin B)
are often included during the initial phase to prevent contamination from the tissue,
but their long-term use is generally discouraged.
◦ A buffering system (e.g., bicarbonate buffer with 5% CO2 in the incubator) to
maintain physiological pH.
• Coating (Optional but often recommended): For many adherent primary cells, coating the
culture surface with extracellular matrix proteins (e.g., collagen, bronectin, laminin, poly-
L-lysine) can signi cantly enhance cell attachment, spreading, and survival, especially for
sensitive cells.
5. Incubation and Maintenance:

•
Incubation Conditions: Typically 37°C, 5% CO2, and high humidity for mammalian cells.
•
Initial Observation: After seeding, observe cells daily under an inverted microscope for
attachment, morphology, and signs of contamination.
• Medium Change (Feeding): Regularly replace the culture medium to provide fresh
nutrients and remove metabolic waste products. The frequency depends on the cell type and
growth rate.
6. Monitoring and Characterization:

•
Morphology: Observe the characteristic shape and appearance of the desired primary cells.
•
Growth Rate: Monitor cell proliferation. Primary cells have a nite lifespan and will
eventually undergo senescence.
• Purity: Assess the purity of the desired cell population. Fibroblast contamination is a
common challenge in many primary cultures derived from solid tissues, as they tend to grow
rapidly. Techniques like differential trypsinization or speci c medium formulations can help
control broblast overgrowth.
• Speci c Markers: Use immunocytochemistry, ow cytometry, or PCR to con rm the
identity and purity of the primary cells by checking for speci c cell surface markers or gene
expression.
Challenges in Primary Cell Culture Development:

•Low Cell Yield: Obtaining suf cient numbers of viable cells from some tissues can be
challenging.
• Contamination: High risk of microbial contamination from the source tissue.
• Heterogeneity: Primary cultures are often a mix of different cell types, making it dif cult to
study a speci c population.
• Limited Lifespan: Primary cells undergo senescence after a limited number of divisions
(Hay ick limit), which restricts long-term experiments.
• Donor Variability: Cells from different donors or even different sites within the same donor
can exhibit signi cant biological variability.
• Senescence and Differentiation: Cells may differentiate or undergo senescence in culture,
losing their original characteristics over time.
Despite these challenges, primary cell cultures remain an indispensable tool in research, providing a
more physiologically relevant model for studying cellular processes, disease mechanisms, and drug
responses compared to immortalized cell lines.
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 separation in cell culture

In cell culture, "separation" refers to the process of isolating speci c cell types from a
heterogeneous mixture. This is crucial for many research applications, as working with a pure
population of cells ensures more accurate and reliable experimental results. Cell separation
techniques exploit various physical, chemical, or biological properties of cells to achieve this
isolation.

Here are the main methods of cell separation in cell culture:

1. Based on Physical Properties:

• Density Gradient Centrifugation:

◦ Principle: Cells are separated based on their buoyant density. A density gradient is
created using solutions like Ficoll, Percoll, or sucrose. When the cell mixture is
centrifuged, cells migrate to the layer in the gradient where their density matches the
density of the medium.

◦ Application: Widely used for isolating peripheral blood mononuclear cells (PBMCs)
from whole blood using Ficoll-Paque, where lymphocytes and monocytes form a
distinct band above the red blood cells and granulocytes.
◦ Advantages: Relatively simple, cost-effective, can handle large sample volumes.
◦ Limitations: May not provide very high purity for closely dense cell populations,
can sometimes activate cells.
• Differential Centrifugation (Sedimentation):

◦ Principle: Cells are separated based on their size and sedimentation rate. Larger and
denser cells pellet faster at lower centrifugation speeds, while smaller and less dense
cells remain in the supernatant and can be pelleted at higher speeds.

◦ Application: Often used as a preliminary step to remove larger debris or red blood
cells from tissue dissociates, or to concentrate cells.
◦ Advantages: Simple, fast, inexpensive.
◦ Limitations: Low purity, often not suf cient for isolating speci c cell types from
complex mixtures.
• Filtration/Sieving:

◦ Principle: Cells are separated based on their size by passing the suspension through
lters or sieves with de ned pore sizes.
◦ Application: Used to remove large aggregates, undigested tissue fragments, or
clumps of cells after tissue dissociation, ensuring a single-cell suspension.
◦ Advantages: Simple, quick, gentle.
◦ Limitations: Primarily for removing larger particles, not for ne separation of
similarly sized cells.
• Differential Adhesion (Plastic Adherence):

◦ Principle: Different cell types adhere to tissue culture plastic surfaces with varying
af nities and rates.
◦ Application: Commonly used in primary cultures. For example, broblasts often
adhere more quickly and strongly to plastic than epithelial cells or hematopoietic
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 cells. By performing sequential washes or timed aspirations, non-adherent cells can
be removed, or the adherent cells can be further cultured.
◦ Advantages: Simple, inexpensive, no special equipment needed.
◦ Limitations: Not highly speci c, purity can be low, relies on differences in cell
behavior.
2. Based on Biological Properties (Immunological Methods):

These methods typically involve using antibodies that speci cally recognize surface markers
(antigens) expressed on the target cells.

• Magnetic-Activated Cell Sorting (MACS):

◦ Principle: Cells expressing speci c surface antigens are labeled with antibodies
conjugated to magnetic beads. The cell suspension is then passed through a column
placed in a strong magnetic eld. Labeled (target) cells are retained in the column,
while unlabeled (unwanted) cells pass through. The magnetic eld is then removed,
and the labeled cells are eluted.

◦ Positive Selection: Directly isolates the target cells.

◦ Negative Selection: Removes unwanted cells, leaving the untouched target cells
behind. This is preferred when the target cells are sensitive to antibody binding or
bead presence.
◦ Application: Highly versatile for isolating various cell types (e.g., T cells, B cells,
stem cells, endothelial cells) from blood, bone marrow, or dissociated tissues.
◦ Advantages: Relatively fast, high purity (often >90%), gentle on cells, does not
require expensive equipment like FACS. Can be automated.
◦ Limitations: Requires speci c antibodies for the target or unwanted cells, magnetic
beads might affect cell function in some applications.
• Fluorescence-Activated Cell Sorting (FACS) / Flow Cytometry:

◦ Principle: This is a highly sophisticated method that uses uorescently labeled

antibodies to identify and sort cells based on their surface (or intracellular) markers,
as well as their physical properties (size and granularity). A cell suspension ows in
a single le through a laser beam. The scattered light and emitted uorescence from
each cell are detected. Based on the signal, an electric charge is applied to droplets
containing speci c cells, which are then de ected by an electric eld into collection
tubes.

◦ Application: Unparalleled for isolating highly pure populations, even rare cells, and
for simultaneously analyzing multiple cell parameters. Used in immunology, stem
cell research, cancer biology, and diagnostics.
◦ Advantages: Very high purity (>99%), can sort multiple cell populations
simultaneously, allows for complex phenotyping (identifying cells based on multiple
markers), can sort single cells.
◦ Limitations: Expensive equipment, requires highly trained operators, can be slow
for large numbers of cells, cells may experience stress from the sorting process,
requires viable single-cell suspension.
• Immunodensity Cell Separation (e.g., RosetteSep™):

◦ Principle: Combines antibody labeling with density gradient centrifugation.

Antibodies speci cally label unwanted cells and link them to red blood cells,
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 forming denser aggregates (rosettes). During density gradient centrifugation, these
heavy aggregates pellet, leaving the untouched target cells in a less dense layer.
◦ Application: Primarily for negative selection of speci c cell types directly from
whole blood (e.g., isolating PBMCs while depleting red blood cells and
granulocytes).
◦ Advantages: Simple, no specialized equipment beyond a centrifuge, maintains high
cell viability.
◦ Limitations: Limited to negative selection, relies on effective rosetting.
3. Other Methods:

• Laser Capture Microdissection (LCM):

◦ Principle: Uses a laser beam under a microscope to precisely cut out and isolate
speci c cells or small regions of interest from a tissue section on a slide.
◦ Application: Ideal for isolating cells from heterogeneous tissues for molecular
analysis (e.g., genomics, proteomics), particularly for rare cells or cells in speci c
histological contexts.
◦ Advantages: Extremely precise, preserves spatial information.
◦ Limitations: Low throughput, typically for xed cells, not for live cell culture.
• Micro uidics:

◦ Principle: Utilizes micro-scale channels and chambers to manipulate and separate

cells based on various physical properties (size, shape, deformability, electrical
properties) or by incorporating speci c antibodies for af nity-based separation.
◦ Application: Emerging eld for single-cell analysis, rare cell isolation (e.g.,
circulating tumor cells), and high-throughput screening.
◦ Advantages: High precision, low sample volume, potential for automation and
integration, gentle on cells.
◦ Limitations: Still under development for many applications, speci c chip design
needed for different applications.
The choice of cell separation method depends on several factors:

• Starting sample type and volume.

• Desired cell type and its characteristics.
• Required purity and viability of the isolated cells.
• Downstream application (e.g., culture, genomic analysis, functional assays).
• Available equipment and budget.
Often, a combination of these techniques is used for optimal cell separation (e.g., density gradient
centrifugation followed by MACS or FACS).

harvesting and maintance of cell line

Harvesting and maintaining cell lines are fundamental practices in cell culture, ensuring the
continuous propagation of cells for research, development, and industrial applications. The speci c
techniques vary slightly depending on whether the cell line is adherent (grows attached to a surface)
or in suspension (grows free- oating in the medium).

I. Maintenance of Cell Lines

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 Routine maintenance is crucial for the health and optimal growth of cell lines. It involves:

1. Aseptic Technique (Paramount):

• Sterile Environment: All procedures must be performed in a laminar ow hood (biosafety

cabinet) with sterilized equipment and reagents.
• Personal Protective Equipment (PPE): Wear lab coats, gloves, and eye protection.
• Disinfection: Regularly wipe down surfaces and equipment with 70% ethanol.
• Minimize Exposure: Keep culture vessels open for the shortest possible time.
2. Optimal Growth Conditions:

• Temperature: Most mammalian cell lines grow optimally at 37°C. Insect cells, however,
require 27-30°C.
• CO2 Concentration: Typically 5% CO2 for bicarbonate-buffered media, which is essential
for maintaining the physiological pH (7.2-7.4). Some media use HEPES buffer and may
require less CO2 control.
• Humidity: High humidity in the incubator (usually >90%) is necessary to prevent
evaporation of the culture medium.
• Culture Medium:
◦ Selection: Choose a basal medium (e.g., DMEM, RPMI-1640, MEM, F-12)
speci cally formulated for your cell line. Refer to the cell line's supplier information.
◦ Supplements: Most media require supplementation with:
▪ Serum (e.g., Fetal Bovine Serum - FBS): Provides growth factors,
hormones, attachment factors, and essential nutrients. Concentration varies
(e.g., 5-20%). Serum-free media are also available for speci c applications.
▪ L-Glutamine: An essential amino acid for cell growth, often added freshly
due to instability.
▪ Antibiotics/Antimycotics: (Optional, use with caution) Penicillin/
Streptomycin (Pen/Strep) and Amphotericin B are sometimes used to prevent
bacterial and fungal contamination, but can mask underlying issues or
promote resistance. Avoid long-term use if possible.
▪ Other Supplements: Some cell lines require additional growth factors,
hormones (e.g., insulin), or non-essential amino acids.
• Cell Density:
◦ Adherent Cells: Monitor con uency (percentage of the ask surface covered by
cells). Subculture when cells reach 70-90% con uency to prevent overcrowding,
nutrient depletion, and senescence.
◦ Suspension Cells: Monitor cell density by counting. Subculture when cell density
reaches a certain threshold (e.g., 1x10^6 cells/mL for some lines) to ensure
logarithmic growth.
3. Routine Monitoring:

• Daily Visual Inspection: Use an inverted microscope to check:

◦ Cell Morphology: Healthy cells exhibit characteristic shapes (e.g., epithelial,
broblast-like, round for suspension) and appear bright and refractile. Changes can
indicate stress, differentiation, or contamination.
◦ Medium Color: Phenol red, a pH indicator in most media, changes color. Yellow
indicates acidity (cells producing lactic acid, overgrowth, or bacterial
contamination). Purple indicates alkalinity (under-gassing CO2, fungal
contamination, or low cell density).
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 ◦
Turbidity: Clear medium is desired for adherent cultures. Turbidity indicates
contamination (bacterial or yeast) or excessive cell debris. Suspension cultures are
inherently turbid due to cells.
◦ Contamination: Look for characteristic signs of bacterial (small moving particles),
fungal ( laments, budding yeast), or mycoplasma (invisible to naked eye, causes
"fuzzy" appearance on high magni cation) contamination.
4. Medium Exchange (Feeding):

• Frequency: Every 2-3 days, or when the medium color changes signi cantly, depending on
cell growth rate and density.
• Procedure: Aseptically aspirate the old, spent medium and replace it with fresh, pre-
warmed complete medium.
II. Harvesting of Cell Lines

Harvesting refers to the process of collecting cells from the culture vessel. The method depends on
the cell's growth characteristics.

A. Harvesting Adherent Cell Lines (for subculturing or experiments):

Adherent cells are attached to the plastic surface and need to be detached.

1. Aspirate Spent Medium: Carefully remove the old medium using a sterile aspirator.
2. Wash with Balanced Salt Solution (BSS): Add a sterile, warmed BSS without Ca2+ and
Mg2+ (e.g., PBS without Ca/Mg) to rinse the cell monolayer. This removes residual serum
(which can inhibit trypsin) and metabolic waste products. Gently rock the ask.
3. Aspirate Wash Solution: Remove the BSS.
4. Add Dissociation Reagent: Add a small volume of enzymatic dissociation solution (most
common) or use a non-enzymatic method:
◦ Trypsin-EDTA:
▪ Trypsin is a protease that cleaves cell adhesion proteins. EDTA chelates Ca2+
and Mg2+, which are crucial for cell-cell and cell-surface adhesion,
enhancing trypsin's activity.
▪ Add enough trypsin-EDTA to just cover the cell monolayer. Gently rock to
ensure even coverage.
▪ Incubate at 37°C (or room temperature for sensitive cells) for a short period
(1-5 minutes typically).
▪ Monitor under Microscope: Periodically check until cells begin to round up
and detach. The ask can be gently tapped to aid detachment. Do not over-
trypsinize, as this can damage cells and reduce viability.
▪ Inactivate Trypsin: Once cells are detached, immediately add fresh
complete growth medium (containing serum, if used) to dilute and inactivate
the trypsin. If using serum-free medium, a speci c trypsin inhibitor (e.g.,
soybean trypsin inhibitor) must be added.
◦ Non-Enzymatic Cell Dissociation Solutions: Gentler alternatives (e.g., Accutase,
Versene/EDTA alone) for sensitive cell lines or when surface protein integrity is
crucial.
◦ Cell Scrapers: For very robustly adherent cells or when enzymatic treatment is
undesirable. Physically scrape cells from the surface into the medium. Can cause cell
damage.
5. Collect Cell Suspension: Pipette the cell suspension up and down several times to break up
clumps and create a single-cell suspension. Transfer the suspension to a sterile centrifuge
tube (e.g., 15 mL or 50 mL conical tube).
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 6. Centrifugation: Centrifuge the cell suspension (e.g., 200-300 x g for 5 minutes) to pellet
the cells. This separates cells from the old medium, dissociation reagent, and debris.
7. Aspirate Supernatant: Carefully remove the supernatant without disturbing the cell pellet.
8. Resuspend Cells: Gently resuspend the cell pellet in a desired volume of fresh, pre-warmed
complete growth medium. Pipette up and down carefully to ensure a homogeneous
suspension.
9. Cell Counting and Viability: Take an aliquot to count cells (e.g., with a hemocytometer or
automated cell counter) and assess viability (e.g., using Trypan Blue exclusion).
10. Seeding (for subculturing): Dilute the cell suspension to the appropriate seeding density
and transfer to new, pre-labeled culture vessels with fresh medium.
B. Harvesting Suspension Cell Lines (for subculturing or experiments):

Suspension cells do not adhere to the ask, making harvesting simpler.

1. Collect Cell Suspension: Directly transfer the desired volume of cell suspension from the
culture ask to a sterile centrifuge tube. If cells have settled, gently swirl the ask to
resuspend them before transferring.
2. Centrifugation: Centrifuge the cell suspension (e.g., 100-200 x g for 5 minutes) to pellet
the cells. Use lower speeds than for adherent cells, as suspension cells can be more fragile.
3. Aspirate Supernatant: Carefully remove the old medium.
4. Resuspend Cells: Gently resuspend the cell pellet in fresh, pre-warmed complete growth
medium.
5. Cell Counting and Viability: Take an aliquot for cell counting and viability assessment.
6. Seeding (for subculturing): Dilute the cell suspension to the desired seeding density and
transfer to new, pre-labeled culture vessels.
III. Subculturing (Passaging) Cell Lines

Subculturing is the process of transferring cells to a new culture vessel to allow for continued
growth when the current vessel becomes too crowded.

• Adherent Cells: The full harvesting procedure (trypsinization, centrifugation, resuspension,

counting) described above is typically performed before subculturing. Cells are then diluted
to a speci c "split ratio" (e.g., 1:3, 1:5, 1:10) into new asks.
• Suspension Cells: This is often simpler. Once the cell density reaches the desired level, a
portion of the culture is removed, and the remaining cells are simply diluted with fresh
medium in the same ask, or transferred to new asks at a lower density. Centrifugation is
usually only performed if a complete medium change is needed or if cells need to be
concentrated or frozen.
•
IV. Cryopreservation (Freezing) of Cell Lines

For long-term storage and to maintain a consistent cell stock, cell lines are cryopreserved.

1. Harvest Cells: Follow the appropriate harvesting procedure (as above) to obtain a single-
cell suspension.
2. Count Cells and Assess Viability: Ensure cells are healthy and in exponential growth
phase.
3. Prepare Freezing Medium: This typically consists of complete growth medium (often
with extra serum) plus a cryoprotectant like dimethyl sulfoxide (DMSO) at 5-10%. DMSO
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 protects cells from ice crystal formation during freezing. Keep the freezing medium ice-
cold.

4. Resuspend Cells in Freezing Medium: Resuspend the cell pellet in the ice-cold freezing
medium at a high concentration (e.g., 1x10^6 to 1x10^7 cells/mL, depending on the cell
line). Minimize exposure time to DMSO at room temperature.
5. Aliquot into Cryovials: Dispense 1 mL aliquots into labeled cryogenic vials.
6. Controlled Freezing: Slow freezing is critical to prevent intracellular ice crystal formation.
◦ Mr. Frosty Container: Place cryovials in an insulated Mr. Frosty (or similar device)
containing isopropanol, which provides a ~1°C/minute cooling rate in a -80°C
freezer overnight.
◦ Programmable Freezer: Use a controlled-rate freezer for more precise temperature
control.
7. Transfer to Liquid Nitrogen: After overnight freezing at -80°C, transfer the vials to a
liquid nitrogen storage tank (-150°C or -196°C) for long-term storage.
8. Record Keeping: Log the cell line name, passage number, date, and location in the liquid
nitrogen tank.
Proper harvesting and maintenance are essential for successful and reproducible cell culture
experiments. Neglecting these basic techniques can lead to contamination, cell stress, altered cell
characteristics, and ultimately, unreliable results.

types of cell culture

Cell culture is a broad eld with various classi cations based on the origin of the cells, their growth
characteristics, and the dimensionality of the culture system. Here's a breakdown of the main types
of cell culture:

I. Based on Origin and Growth Potential:

This is the most common classi cation, focusing on how cells are initially isolated and their
capacity for proliferation in vitro.

1. Primary Cell Culture:

◦ De nition: Cells directly isolated from tissues (e.g., biopsy, organ explant) of an
organism (human, animal, or plant) and grown in vitro.
◦ Characteristics:
▪ Finite Lifespan: Most primary cells have a limited number of divisions
(Hay ick limit) before undergoing senescence (aging) and eventually dying.
▪ Retention of In Vivo Characteristics: They tend to maintain many of the
physiological and genetic characteristics of the original tissue, making them
more physiologically relevant for studying normal biological processes and
disease.
▪ Heterogeneity: Initially, primary cultures can be heterogeneous, containing
various cell types from the tissue. Over time, faster-growing cells (often
broblasts) may dominate the culture.
▪ Labor Intensive: Isolation and optimization of culture conditions can be
complex and time-consuming.
▪ Variability: There can be signi cant donor-to-donor variation.
◦ Examples: Human dermal broblasts, bovine aortic endothelial cells, primary
hepatocytes, neurons.
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 2. Cell Lines:
◦ De nition: A cell line is established when a primary culture is subcultured
(passaged) at least once. Cells in a cell line have adapted to growth in vitro.
◦ Sub-types of Cell Lines:
▪ Finite Cell Lines (Diploid Cell Strains):
▪ De nition: These are cell lines with a limited number of population
doublings before they enter senescence. They usually retain a normal
diploid chromosomal number.
▪ Characteristics: Similar to primary cells in retaining some
differentiated features, but they have undergone a few passages.
▪ Examples: WI-38 (human lung broblasts), MRC-5 (human lung
broblasts).
▪ Continuous Cell Lines (Immortalized Cell Lines):
▪ De nition: These cell lines have undergone a transformation process
(spontaneously, virally, or chemically induced) that allows them to
proliferate inde nitely in vitro, bypassing senescence.They often
exhibit aneuploidy (abnormal chromosome number).
▪ Characteristics:
▪ Unlimited Lifespan: Can be passaged inde nitely, providing
an endless supply of cells.
▪ Ease of Culture: Generally more robust, easier to grow, and
have faster growth rates compared to primary cells.
▪ Homogeneity: Tend to be more homogeneous due to selective
pressures during immortalization.
▪ Loss of In Vivo Characteristics: May lose some
differentiated functions or physiological relevance due to
genetic and phenotypic changes during transformation.
▪ Examples: HeLa (human cervical carcinoma), HEK293 (human
embryonic kidney), CHO (Chinese Hamster Ovary), NIH/3T3 (mouse
broblasts).
II. Based on Growth Characteristics/Adherence:

This classi cation describes how cells grow within the culture vessel.

1. Adherent Cell Culture (Monolayer Culture):

◦ De nition: Cells that require a solid surface (e.g., tissue culture-treated plastic,
glass) to attach and spread for growth. They form a single layer (monolayer) of cells.
◦ Characteristics: Most mammalian cells, particularly those of epithelial and
broblastic origin, are anchorage-dependent.
◦ Applications: Widely used for studying cell morphology, migration, cell-cell
interactions, drug screening, and virus propagation.
◦ Examples: Fibroblasts, epithelial cells, endothelial cells.
2. Suspension Cell Culture:

◦ De nition: Cells that grow freely oating in the culture medium without attaching to
a surface.
◦ Characteristics: Typically derived from hematopoietic tissues (blood, bone marrow)
or some transformed cell lines that have lost anchorage dependence. Requires
agitation (shaking asks, bioreactors) for oxygen and nutrient distribution.
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 ◦ Applications: Ideal for large-scale production of biological products (antibodies,
vaccines, proteins), studies requiring high cell yields, or when cell harvesting needs
to be quick and gentle.
◦ Examples: Lymphocytes, hybridomas, some leukemia cell lines (e.g., Jurkat, K562).
III. Based on Dimensionality:

This classi cation relates to the complexity of the culture environment.

1. 2D Cell Culture (Conventional Cell Culture):

◦ De nition: Cells are grown on a at, two-dimensional surface (e.g., Petri dishes,
asks, well plates).
◦ Characteristics:
▪ Simplicity: Easy to set up, observe, and manipulate.
▪ Cost-effective: Relatively inexpensive.
▪ Standardized: Well-established protocols and equipment.
▪ Limitations: Does not accurately mimic the in vivo environment where cells
are surrounded by a 3D extracellular matrix and interact with other cells in all
dimensions. This can lead to altered cell morphology, gene expression, and
function compared to in vivo.
◦ Applications: Routine cell expansion, basic drug toxicity screening, gene expression
studies.
2. 3D Cell Culture:

◦ De nition: Cells are grown in a three-dimensional environment that better mimics

the in vivo tissue architecture.
◦ Characteristics: Allows for more realistic cell-cell and cell-matrix interactions,
formation of complex structures, and establishment of gradients (nutrients, oxygen).
◦ Advantages: Provides a more physiologically relevant model, better prediction of in
vivo responses (e.g., drug ef cacy and toxicity), and can model tissue development
and disease progression more accurately.
◦ Sub-types/Approaches:
▪ Spheroids/Organoids: Self-assembling aggregates of cells that form sphere-
like structures. Organoids are more complex, multicellular structures that
mimic the architecture and function of speci c organs.

▪
Scaffold-Based Systems: Cells are grown within or on a porous scaffold
(e.g., hydrogels, porous polymers) that provides structural support and
mimics the extracellular matrix.
▪ Micro uidic Devices (Organs-on-a-Chip): Micro-engineered devices with
channels and chambers that allow precise control over the cellular
microenvironment, including uid ow, gradients, and mechanical forces, to
simulate speci c organ functions.
▪ Bioprinting: Using 3D printing technologies to precisely deposit cells and
biomaterials layer-by-layer to create complex tissue constructs.
◦ Applications: Drug discovery and screening, regenerative medicine, disease
modeling, stem cell research, tissue engineering.
IV. Other Specialized Types:
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 • Co-culture: Growing two or more different cell types together in the same culture system to
study their interactions (e.g., epithelial cells with broblasts, immune cells with cancer
cells).
• Feeder Layer Culture: A layer of supporting cells (feeder cells, often inactivated
broblasts) is cultured to provide essential growth factors and an optimal microenvironment
for the growth of more fastidious cells (e.g., stem cells, primary neurons).
• Organ Culture: Involves culturing intact pieces of organs or tissues (rather than dissociated
single cells) to study their integrated functions, preserving the complex tissue architecture.
• Explant Culture: A type of primary culture where small pieces of tissue (explants) are
placed directly onto a culture surface, and cells migrate out from the tissue onto the surface
to proliferate.
The selection of a cell culture type depends entirely on the research question, the desired level of
physiological relevance, and the resources available. While 2D cell lines remain a staple for many
basic studies due to their simplicity and cost-effectiveness, the increasing demand for more accurate
in vivo models is driving the adoption and innovation in 3D and organoid cultures.

cryopreservation cell line

Cryopreservation is the process of preserving cells or tissues by cooling them to very low
temperatures, typically $-80^\circ C$ or in liquid nitrogen ($-196^\circ C$). At these ultra-low
temperatures, all biological activity, including metabolic processes, is effectively halted, allowing
cells to be stored for extended periods, theoretically inde nitely, without deterioration. This is a
crucial technique in cell culture for maintaining valuable cell lines, preventing genetic drift,
reducing the risk of contamination, and ensuring a readily available supply of cells for future
experiments.

Principles of Cryopreservation

The primary challenge in cryopreservation is preventing damage to cells during the freezing and
thawing processes, mainly due to ice crystal formation. The key principles involve:

1. Controlled Cooling Rate (Slow Freezing): Rapid freezing can lead to the formation of
large, damaging intracellular ice crystals. Slow freezing (typically around $-1^\circ C$ per
minute) allows water to move out of the cells, concentrating intracellular solutes and
minimizing the formation of lethal ice crystals inside the cells.

2. Cryoprotective Agents (CPAs): These chemical compounds are added to the freezing
medium to protect cells from freezing-induced damage. They work by:
◦ Lowering the freezing point: Reduces the amount of ice formed at a given
temperature.
◦ Increasing viscosity: Slows down ice crystal growth.
◦ Protecting cell membranes: Interacting with cell membranes to stabilize them
during dehydration and rehydration.
◦ Common CPAs:
▪ Dimethyl Sulfoxide (DMSO): The most commonly used CPA for
mammalian cells. It's a permeating agent, meaning it can enter cells.
Typically used at 5-10% concentration.

▪ Glycerol: Another permeating CPA, often used for cells sensitive to DMSO
or for speci c applications.
▪ Non-permeating CPAs: Sugars like Trehalose or Sucrose, or polymers like
Polyethylene Glycol (PEG) or Hydroxyethyl Starch (HES), are sometimes
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 used in combination with permeating CPAs. They primarily act
extracellularly to dehydrate cells.
3. Optimum Cell Concentration: Freezing cells at too low a concentration can reduce
viability upon thawing. A recommended density is usually $1 \times 10^6$ to $1 \times
10^7$ cells/mL, though optimization might be needed for speci c cell types.

4. Ultra-Low Temperature Storage: Once frozen, cells must be stored at temperatures below
$-130^\circ C$ (the glass transition temperature of water) to ensure that all metabolic
activity is suspended.
Steps for Cryopreservation of Cell Lines

Aseptic technique is critical throughout the entire process.

1. Preparation of Cells:
◦ Select Healthy, Log-Phase Cells: Cells should be actively growing (in the
exponential/logarithmic growth phase) and healthy, with high viability (e.g., >90%).
Avoid freezing cells that are overcon uent or stressed.

◦ Check for Contamination: Ensure the cell culture is free from bacterial, fungal, and
especially mycoplasma contamination before freezing, as contaminants will also be
preserved.
◦ Harvest Cells:
▪ Adherent Cells: Gently detach cells from the culture vessel using an
appropriate dissociation agent (e.g., trypsin-EDTA, Accutase). Neutralize the
dissociation agent (e.g., by adding serum-containing medium) and gently
centrifuge to obtain a cell pellet. Wash with a balanced salt solution (e.g.,
PBS) to remove residual dissociation agent if necessary.
▪ Suspension Cells: Directly transfer the cell suspension to a sterile centrifuge
tube and gently centrifuge to pellet the cells.
◦ Count Cells and Assess Viability: Resuspend the cell pellet in a small volume of
complete growth medium.Count cells using a hemocytometer or automated cell
counter and perform a viability test (e.g., Trypan Blue exclusion assay). Adjust the
cell concentration to the desired density for freezing.

2. Preparation of Freezing Medium:

◦ Prepare the freezing medium by mixing the basal culture medium (often with
increased serum content, e.g., 20% FBS) and the chosen cryoprotectant (e.g., 5-10%
DMSO).
◦ Crucially, keep the freezing medium on ice to minimize the time cells are exposed
to the CPA at warmer temperatures, as CPAs can be toxic.
3. Mixing Cells with Freezing Medium:

◦ Carefully resuspend the cell pellet in the cold freezing medium to achieve the desired
cell concentration (e.g., $1 \times 10^6$ to $1 \times 10^7$ cells/mL).
◦ Pipette gently to ensure a homogeneous single-cell suspension and minimize
clumping.
◦ Minimize the time cells spend in the freezing medium at room temperature/warm
temperatures before cooling.
4. Aliquot into Cryovials:
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 ◦ Dispense uniform aliquots (e.g., 0.5-1.5 mL) of the cell suspension into pre-labeled,
sterile, cryogenic vials.Use vials speci cally designed for cryopreservation, which
are robust and have leak-proof caps.
◦ Labeling: Clearly label each vial with essential information: cell line name, date of
freezing, passage number, cell concentration, and your initials.
5. Controlled Freezing (Slow Cooling):

◦ This is a critical step for maximizing post-thaw viability. The goal is to achieve a
cooling rate of approximately $-1^\circ C$ per minute until the vials reach around
$-80^\circ C$.
◦ Methods for Slow Freezing:
▪ Programmable Freezing Unit: The most ideal method, providing precise
control over the cooling rate. The vials are placed in the unit, and a pre-
programmed cooling pro le is executed.
▪ Mr. Frosty Freezing Container (or similar): A widely used and effective
passive device. Vials are placed inside an insulated container lled with
isopropanol (which acts as a thermal buffer) and then placed in a $-80^\circ
C$ freezer overnight. The isopropanol helps achieve the desired cooling rate.
▪ Styrofoam Box: A less precise but sometimes used method. Vials are placed
in a Styrofoam box (thick walls provide insulation) and placed in a $-80^\circ
C$ freezer overnight. This method is less recommended for valuable or
irreplaceable cultures due to less uniform and reproducible cooling.
6. Transfer to Long-Term Storage:

◦ After the initial slow freezing (usually overnight at $-80^\circ C$), the vials must be
transferred rapidly to a liquid nitrogen storage tank for long-term preservation.
◦ Storage Temperature: Below $-130^\circ C$, ideally in liquid nitrogen vapor phase
($-150^\circ C$ to $-190^\circ C$) or directly submerged in liquid nitrogen
($-196^\circ C$). Vapor phase storage is generally preferred to minimize the risk of
cross-contamination if a vial leaks, or explosion during thawing due to liquid
nitrogen entering the vial.
◦ Record Keeping: Crucially, record the exact location (rack, box, position) of each
vial in the liquid nitrogen tank for easy retrieval.
Thawing Cryopreserved Cells

Rapid thawing is essential to minimize the formation of damaging ice crystals during the warming
process.

1. Retrieve Vial: Locate the desired vial from the liquid nitrogen tank using appropriate
cryogenic gloves and face shield.
2. Rapid Thawing: Immediately immerse the cryovial in a $37^\circ C$ water bath, gently
swirling it until only a small ice crystal remains (usually 1-2 minutes).
3. Disinfect Vial: Once thawed, wipe the outside of the vial with 70% ethanol before opening
it in a laminar ow hood.
4. Transfer Cells: Transfer the cell suspension from the vial to a sterile centrifuge tube
containing a large volume of pre-warmed complete growth medium (e.g., 5-10 mL). The
large volume rapidly dilutes the cryoprotectant (like DMSO), which can be toxic to cells at
warmer temperatures.
5. Centrifugation (Optional but Recommended): Gently centrifuge the cell suspension (e.g.,
100-300 x g for 5 minutes) to pellet the cells. This step removes the cryoprotectant and dead
cells/debris.
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 6. Resuspend and Seed: Aspirate the supernatant and gently resuspend the cell pellet in fresh,
pre-warmed complete growth medium. Count cells and assess viability. Seed cells into a
fresh culture vessel at an appropriate density.
7. Initial Culture: Place the culture in the incubator. For adherent cells, check for attachment
after a few hours. For both adherent and suspension cells, perform a medium change after
12-24 hours to completely remove any residual cryoprotectant and metabolic waste
products.
Key Considerations for Successful Cryopreservation

• Cell Health: Only freeze healthy, contamination-free cells in their logarithmic growth
phase.
• Asepsis: Strict adherence to aseptic technique is paramount to prevent contamination.
• Cryoprotectant Choice and Concentration: Optimize for your speci c cell line.
• Cooling Rate: The $-1^\circ C$/minute rule is a general guideline; some cells might require
different rates.
• Storage Temperature: Maintain storage below $-130^\circ C$ at all times.
• Thawing Rate: Always thaw rapidly.
• Record Keeping: Meticulous records of cell line details, passage number, freezing date,
viability, and storage location are critical for proper cell line management.
By following these principles and steps, you can effectively cryopreserve your cell lines, ensuring
their long-term viability and maintaining the integrity of your research materials.
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