cell culture: recent advance and application
TRANSCRIPT
Sharma et al. World Journal of Pharmaceutical Research
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CELL CULTURE: RECENT ADVANCE AND APPLICATION
Susrita Sharma1*, B. Ray
2 and C. K. Sahoo
3
1Final Year (B. Pharm), GCP, Sambalpur.
2Associate Prof. Dept. Pharmacology, Puri.
3Dept. Pediatric, SCBMCH, Cuttack.
ABSTRACT
Cell culture is the process by which cells are grown under controlled
conditions, generally outside their natural environment. After the cells
of interest have been isolated from living tissue, they can subsequently
be maintained under carefully controlled conditions. There are two
methods for obtaining cells: from a cell bank or by isolating cells from
donor tissue. When starting culture from cells obtained from a cell
bank, one needs to go through the procedures of "thawing," "cell
seeding" and "cell observation." culture techniques is vital tool in the
process of drug Discovery which ultimately leads to quantify the steps
of analysis of therapeutic potential of drugs. Gene therapy depends on the analysis of cell
culture to disclose the unidentified facts related to genomics. The present review Focus
different types of cell culture and related technique. It also highlights Recent progress in the
particular field.
KEYWORD:- Cell cultue, Immunolabeling, genomics, cell seeding, Biological buffer.
CELL CULTURE
INTRODUCTION
Cell culture is the process by which cells are grown under controlled conditions, generally
outside their natural environment. After the cells of interest have been isolated from living
tissue, they can subsequently be maintained under carefully controlled conditions. These
conditions vary for each cell type, but generally consist of a suitable vessel with a substrate or
medium that supplies the essential nutrients (amino acids, carbohydrates, vitamins, minerals),
growth factors, hormones, and gases (CO₂, O₂), and regulates the physio-chemical
environment (pH buffer, osmotic pressure, temperature). Most cells require a surface or an
World Journal of Pharmaceutical Research SJIF Impact Factor 8.084
Volume 10, Issue 1, 431-460. Review Article ISSN 2277– 7105
*Corresponding Author
Susrita Sharma
Final Year (B. Pharm), GCP,
Sambalpur.
Article Received on
28 October 2020,
Revised on 18 Nov. 2020,
Accepted on 08 Dec. 2020
DOI: 10.20959/wjpr20211-19435
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artificial substrate (adherent or monolayer culture) whereas others can be grown free floating
in culture medium (suspension culture). The lifespan of most cells is genetically determined,
but some cell culturing cells have been ―transformed‖ into immortal cells which will
reproduce indefinitely if the optimal conditions are provided.[1]
Cell culture systems are indispensable tools for basic research and a wide range of clinical in
vitro studies. However, conventional 2D cell cultures poorly mimic the conditions in the
living organism. This limitation may seriously compromise the reliability and significance of
data obtained from such approaches. Therefore, we present here a comparative study on
selected 3D and 2D cell cultures of U87-MG human glioblastoma cells that were processed
by means of high-pressure freezing and freeze-substitution as well as by conventional
chemical fixation and Tokuyasu cryo-section immuno-labeling. Three-dimensional cultures
comprised pseudo-vascularized cultures, fiber and bead scaffold cultures,
and spheroid cultures. Cell cultures in dishes and on coverslips were the static 2D culture
systems used as reference models. We will discuss morphological and immuno-cytochemical
observations with respect to the feasibility of the cell culture systems investigated for the
state-of-the-art electron microscopy.[2]
Various types of cell culture
1. Primary cell culture
2. Secondary cell culture
3. Suspension cell culture
4. Adherent cell culture
1. Primary cell culture: These are cells obtained directly from an organism and are directly
plated in a cell culture dish or flask. They comprise cells of a tissue or organ obtained
from an organism and immediately transferred to a suitable cell culture environment
conducive for growth. Such cells will attach to the medium, divide and grow
exponentially. They are generally termed primary cell cultures. Primary cell cultures have
a limited life span. They will only last for a short period of time (usually days to weeks).
Their only advantage is that they may exhibit some physiological behaviour similar to
that obtainable in vivo because they are freshly isolated cells. Primary cell cultures are
usually unstable and require some time to adapt to the in vitro environment they are
introduced to.
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In addition, some cells in primary cell culture may sustain injury during their isolation and
preparation, and thus eventually die in the process. In primary cell cultures, a series of
enzymatic and mechanical disruptions of the tissues or organs and selection steps are usually
employed to isolate the cells of interest from a heterogeneous population of cells. Some
examples of primary cell cultures or cell lines includes: macrophages, natural killer (NK)
cells, B and T cells, dendritic cells and cells of the spleen (splenocytes). These cells are all
cells of the immune system. They are used in primary cell cultures to decode the effects of
some certain substances (e.g. drugs) on the functions and proliferation of cells of the immune
system.
2. Secondary cell culture: These are cells taken from a primary cell culture and are
passaged (or subcultured) into a new and fresh cell culture flask/disk containing new
growth medium. Passaging which can also be referred to as sub-culturing is the
transplantation of cells from one cell culture vessel to another. Passaging gives cells the
chance to expand and increase in population. A higher cell growth is usually achieved due
to the addition of fresh growth medium and the introduction of other environmental
conditions. Normally, the number of cells obtained from a primary cell culture are may
not be enough to create sufficient cells required for a graft, and this warrant the need for
Passaging of cells obtainable in secondary cell culture.
Secondary cell cultures are transformed and immortalized cell lines with infinite growth and
proliferation capacity. They are usually derived from human carcinomas/tumours. Such cells
have been transformed in the sense that they have lost sensitivity to factors associated with
growth control and thus can grow unlimited. Secondary cell cultures are more easily cultured
than the primary cell cultures. Some sources of secondary cell cultures include: embryos and
tumours or transformed cells such as HeLa cells and Chinese hamster ovary (CHO).
Secondary cell culture has applications in a range of areas such as in vaccine production and
drug screening.
3. Suspension cell cultures: These are cells that grow freely and unattached to any surface.
Such cells are cultured in suspensions of growth medium. They are maintained in a cell
culture flask without any adherence to any surface. Examples of cells cultured in
suspension include the cells of the blood such as hematopoietic cells. Such cells are
engineered to grow in suspensions. They grow in a very much higher proportion.
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4. Adherent cell cultures: These are cells that attach or adhere to the surfaces of the cell
culture flask used for their culturing. They are referred to as anchorage-dependent cells.
These cells are cultivated in suitable growth medium that is specially suited and treated to
allow adhesion and the spreading of the cells. The cell culture flask used for adherent
cells are usually coated with materials that increase their adherence features and provide
signals needed for their growth and proliferation in the cell culture medium.[3]
General process of cell culture
There are two methods for obtaining cells: from a cell bank or by isolating cells from donor
tissue. When starting culture from cells obtained from a cell bank, one needs to go through
the procedures of "thawing," "cell seeding" and "cell observation."
When using tissue collected from a donor, unnecessary tissue are usually removed if it is
attached. There are two major methods to isolate cells from the tissue, explant culture and
enzymatic method. In enzymatic methods, isolation of cells from the tissue of interest using a
proteolytic enzyme solution. If an enzyme is used, dilute the enzyme or stop the enzyme
reaction with an enzyme reaction inhibitor, then proceed with the steps of "cell seeding" and
"cell observation" to prepare the cell culture.
Hawing
Thawing frozen cryopreserved cells to initiate a cell culture may be thought of as "waking up
the cells." A vial of frozen cells obtained from a cell bank is transferred from a liquid
nitrogen tank*1
or cryogenic deep freezer (-150 °C) to an appropriate cold storage container,
such as a liquid nitrogen container, transported to the bench, and thawed in a 37°C water bath
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or in a melting apparatus*2
. Before ice is almost melted, medium is quickly added to dilute
the cryoprotectant liquid (EX. DMSO), the cells are precipitated by centrifugation, and after
removing the supernatant, fresh medium pre-heated to 37°C is added. The cells are then
resuspended by pipetting and the number of cells/cell concentration is measured using a
microscope or cell counter.
1. *There are two ways to freeze and preserve cells in a liquid nitrogen tank: "Vapor phase"
in which liquid nitrogen's cold gas is used, and "liquid phase" in which a frozen preserved
object is directly immersed in liquid nitrogen. In the case of liquid-phase, it can be
preserved at -196 °C, which is the temperature of liquid nitrogen, but there is a high risk
of liquid nitrogen getting into the frozen vial and contaminating it with bacteria, yeast,
mycoplasma, and viruses from other vials. Therefore, storage in vapor phase is strongly
recommended.
2. *Vials that have been stored in the liquid phase could burst if they are placed in a 37°C -
water bath or a melting apparatus with liquid nitrogen remaining in a vial. The lid should
be loosened once and retightened before putting it in the 37°C -water bath.
Cell seeding
To achieve the target cell seeding density, calculate the amount of fresh medium required to
achieve the desired cell seeding density based on the measured cell numbers, and dilute the
cell suspension accordingly.
Cell observation
After seeding the cells in a new culture vessel, observe the cells in the vessel with an optical
microscope or other observation device in the following manner:
Check that there are viable cells
Check to make sure that cells are evenly distributed in the vessel
Check for the presence of foreign objects other than cells
Check the cell morphology
After confirming the above, place the cell culture vessel in a humidified CO2 incubator at
37°C and start culturing.
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From cell culture initiation to passage
Cell observation
The cells are seeded in a new culture vessel, which is placed in a CO2 incubator. Generally,
on the following day*, the below observations are performed using an optical microscope or
other observation device:
Check to make sure there are no foreign objects other than cells in the culture vessel
Determine whether the culture is proceeding normally by checking cell morphology and
condition*There are cases where they are left to stand for two days depending on culture
conditions and cell type.
Medium exchange
After having been thawed and seeded in a culture vessel, cells start to grow in the
CO2 incubator. Cells metabolize nutrients in the culture medium, therefore medium that has
been depleted of nutrients and enriched with metabolites must be replaced with fresh
medium. This process is called "medium change" or "medium replacement".
Prior to medium change, first observe the cells to verify that the culture is proceeding
normally. After removal of the old medium, quickly add new medium to prevent the cells
from drying out. Some cells may die if not immersed in medium, so there are cases where a
small amount of the old medium is left instead of being entirely discarded. Additionally, fresh
medium should be pre-warmed to 37°C so as not to expose cells to sudden changes in
temperature.
After changing the medium, check the cells for any signs of damage. Use a microscope to
acquire images of the culture in order to document that it is proceeding well, then return it to
the incubator, taking care to minimize unnecessary disturbances.
Passage
Once cells start to proliferate, divide them into new culture vessels before the current vessel
becomes full. This is called "passage." The state in which cells have grown to fill the culture
vessel is called "confluent." Generally, it is recommended that cells be passaged when the
area occupied by cells reaches approximately 70 to 80% of the vessel.
When they become confluent, cells come into contact with each other and feel that they do
not need to grow any more, a phenomenon known as "contact inhibition," and especially in
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the case of normal cells they will no longer proliferate after passage. In the case of cancer
cells, they will continue to proliferate rapidly, but they will come to lack nutrition, so it is not
good to allow them to be confluent. For this reason, it is necessary to observe and monitor
cell growth.
Passage methods differ between suspension cells and adherent cells.
Passage of suspension cells
Collect the cell culture suspension into a tube and centrifuge to collect the cells. Remove the
supernatant, leaving the cell pellet, and re-suspend in fresh medium. Take a part of the fresh
suspension, apply the vital stain trypan blue, and count the number of living cells. Calculate
the cell concentration, consider cell dilution methods, adjust the density of cells in suspension
appropriately, and place in a new container.
Passage of adherent cells
Cells that adhere to the culture vessel surface need to be somehow detached from its surface.
Generally, proteases such as collagenase, dispase, and trypsin are used. In the case of trypsin,
activity is inhibited by calcium or magnesium ions, so it is necessary to wash out medium
containing these ions prior to its application. Conversely, collagenase and dispase show
activity in the presence of calcium ions.
The general procedure is as follows
Remove the old culture medium supernatant
If necessary, wash with phosphate buffer or fresh medium
Add cell dispersion enzyme solution
Stabilize at a predetermined temperature within the active range of the enzyme for a
predetermined time to promote the enzymatic reaction
Check the degree of cell detachment under a microscope
Promote cell detachment by mild mechanical disturbance, such as tapping, etc.
If a stop solution for the enzyme(s) is available, add it to stop the reaction. If not, add
fresh medium to dilute the enzyme and decrease activity
Pipette the medium several times to dissociate cells into a single-cell suspension
Collect the medium including cells, precipitate the cells by centrifugation, and remove the
supernatant containing the enzyme and reaction stop solution.
Tap the tube to loosen cell pellet
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Add fresh medium to the tube of collected cells
Resuspend the cells by pipetting up and down
Prepare sample as a representative cell suspension to count the cell number/cell density
Count cell numbers using a microscope with hemocytometer or an automated cell counter
Determine the correct dilution for obtaining the desired cell density, add the appropriate
amount of fresh medium, and re-suspend the cells
Seed a predetermined amount of the cell suspension into a new culture vessel
Perform microscope observation
Transfer to a humidified CO2 incubator set to a temperature of 37 °C
However, some cells could get weakened if dispersing enzymes are used. In that case,
mechanically scrape and peel off cells with a scraper, or peel off cells with the flow from a
pipet to harvest cells.
Processing after cell culture (Stock preparation)
It is important to make stocks that have the same characteristics as the original cells, which
may be obtained from primary culture, purchased from a supplier, or transferred from
elsewhere. This is because cells are living materials and their characteristics may change over
time and if passage continues over a long period of time, it could cause them to differ from
the original cells.
A cell stock can be created as follows
Cell observation
Perform the following observations with an optical microscope or other observation device.
Check for foreign objects other than cells in the culture vessel
Determine if the cells are subconfluent and have not over-proliferated
Determine whether the culture is proceeding normally by checking cell morphology and
condition
Cell detachment
Harvest cells for stocks using the passage procedure described earlier.
Cell observation
Add fresh medium to resuspend the cells collected by centrifugation, but use a small amount
of medium because the density of the cell suspension needs to be higher than that for passage.
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The cells are suspended by pipetting, and the number of cells/cell density is measured using a
microscope or measuring instrument.
Dispensing
Adjust the cell suspension to the desired number of cells, add the same amount of 2x
concentrated cryopreservation solution, and re-suspend by pipetting. While pipetting or
stirring constantly, dispense a predetermined amount of solution into the cryotube.
Freeze
The cryotube should be immediately placed in a freezer container and placed in a deep
freezer (-80° C) to keep a freezing rate of -1°C per minute. Alternatively, freeze with a
programmable controlled-rate freezer. After the cells are frozen, the frozen cryotubes should
be stored in the vapor phase within a liquid nitrogen storage tank or cryogenic deep freezer (-
150 ° C). But, the procedures varies depending on the type of cryopreservation solution.
Confirmation
Thaw one or two of the frozen vials and culture them. Confirm whether the cells can grow in
the same manner and exhibit almost identical characteristics as previously. Once this is
confirmed, stop the cell culturing.
Start of experiment
Thaw prepared cell stocks as needed for research activities. After a certain period in culture,
the cells should be discarded and fresh stock thawed for use.
Regenerative medicine and stem cells
About cell culture
Cases often viewed
Accurate measurement MSC cell number and growth rate in an non-exfoliating and fast
way.
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Evaluation of cell distribution density during seeding and growth required to maintain
cell traits.
Improving work efficiency and accuracy through automatic measurement of iPS.[4]
Cell culture media
The cell culture media consists of the followings
Cell Culture Reagents
Cell Culture Supplements
Mammalian Cell Lines
Cancer Cell Lines
Primary Cells and Cell Culture
PromoCell® Human Primary Cell Culture*
Stem Cell Culture
Classical Media & Salts for Cell Culture
Sera for cell culture
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Serum Free Media
Learning Center
Specialty Media
Cellular Assays
Cell Culture Troubleshooting Guide
Biochrom Product Updates
Cell culture reagents
The Cell Culture and Insect Cell Culture Tested reagents listed in this section differ from our
research grade reagents, in that they undergo additional testing in a cell culture system. This
testing is designed to eliminate the need for screening biochemicals prior to use in a cell
culture application.
Antibiotics
Attachment and Matrix Factors
Biological Buffers
Biological Detergents
Cell Dissociation
Cell Freezing
Cell Separation
Cell Viability Kits and Reagents
Mycoplasma Kits and Reagents
Miscellaneous Reagents
Cell culture supplements
The Cell Culture and Insect Cell Culture Tested supplements listed in this section differ from
our research grade compounds, in that they undergo additional testing in a cell culture system.
This testing is designed to eliminate the need for screening these supplements prior to use in a
cell culture application.
Albumins and Transport Proteins
Amino Acids and Vitamins
Antibiotics
Cytokines and Growth Factors
Hormones
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ITS and Other Supplements
Lipids and Lipid Carriers
Primary Cells and Cell culture
Primary cells are derived directly from living tissue and therefore more closely replicate the
physiology of biological systems. They are increasingly useful in life science research,
ADME/toxicity studies and pharmaceutical development, and in diverse applications where
cell lines do not replicate the target biological system. We are pleased to offer primary cells
from Promo Cell (available in select geographies) and from Cell Applications, Inc. (CAI).
Our manufacturers have refined the isolation, purification, subculture, and growth of over
100 cell types. Cells come with the promise of purity, low passage, rigorous characterization,
and strict quality control. Harmonized media and reagents provide optimal performance,
ensuring confidence in the utility of these cells and their supporting products in your
application.
PromoCell® human primary cell culture*
Primary human cells may provide biomedical researchers with more meaningful cell models,
as their physiology more closely replicates that of the biological tissues from which they are
extracted—especially for applications where immortalized cell lines have attributes that can
confound results.
Our primary cell inventory includes cells and precisely-formulated media from industry-
leading manufacturer PromoCell. This collection offers the broadest range of validated,
ethically-sourced human primary and stem cells, plus human blood cells—along with culture
media optimized for these phenotypes.
Primary cell culture protocols, plus advanced media and reagents for 3D culture from
patient cancers
Unique formulations for groundbreaking cancer studies include a xeno-free (XF) medium for
cultivating cancer lines in 3D. The Primary Cancer Culture System is a complete solution for
the selective culture of malignant cells extracted from primary tumors or patient xenografts.
Stem cells have the unique ability to self-renew or to differentiate into various cell types in
response to appropriate signals. These properties provide stem cells with unique capabilities
for tissue repair, replacement, and regeneration. Accordingly, human stem cells are of special
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interest in medical research. Embryonic stem cells have the ability to differentiate into more
cell types than adult stem cells. Differentiation is triggered by various factors in vivo, some
of which can be replicated in in vitro stem cell cultures. The nature of stem cells necessitates
the use of special stem cell culture media and reagents. Since suboptimal media may change
the differentiation potential of stem cells, it is vital to choose the correct stem cell validated
media and reagents at the start of your research process. Use the selection guide provided
below during the planning phase of your research project. With the correct reagents, stem
cells can be cultured like any other cell lines. They are no more or less finicky. Some stem
cell lines are immortal and can be cultured indefinitely in the lab, but many are not. Be sure
to choose the right cell line for your purpose!
Recently, there have been advances in the realm of the stem cell research due to the advent
of CRISPR genome editing technology and more advanced 3D cell culture techniques. These
cell culture protocols, such as organoid cell cultures, have provided more predictive in
vitro cellular ―Disease-in-a-Dish‖ models. Combining our CRISPR, ZFN gene editing, and
stem cell expertise, We now offer novel stem cell lines, optimized media, and innovative kits
for all areas of stem cell biology, including induced pluripotent stem cells (iPSCs), neural,
mesenchymal and hematopoietic stem cell culture. In addition to our expansive portfolio of
assay ready stem cells, serum-free cell culture media, and 3D culture solutions, we offer
custom engineered stem cell lines through our easy-to-use Cell Design Studio. Choose your
favorite host or iPSC line and watch our dedicated team of scientists knock-out, modify, or
knock-in your gene of interest.
Classical Media & Salts for cell culture
It is a testament to Dulbecco, Eagle, Ringer and other pioneering physicians and scientists
that the media formulations they developed are still widely in use today.
Media variations have been refined in response to the need for physiologically-relevant
environments for diverse mammalian cell cultures. Whether you're growing adherent
suspension phenotypes, with or without FBS, need high- or low-glutamine, ready-to-use
liquid or easy-to-store powder – you'll find just what you need for cell culture here. These
media and salts, along with their components, have been qualified for cell culture
applications, and are manufactured in our state-of-the art facilities.
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Some examples are
Basal Salts for Cell Culture
DMEM
DMEM/F12 Media
Ham's Nutrient Mixtures
Medium 199 (M199)
MEM (Minimum Essential Medium Eagle)
RPMI-1640 Media
Isolation of cells
It is the process of separating individual cells from a solid block of tissue or cell suspension.
This may be performed by using enzymes to digest the proteins that binds these cells together
within the extracellular matrix.
There are multiple methods that can be used when performing cell isolation.
Cell separation methods
Immunomagnetic Cell Separation
Fluorescence-activated Cell Sorting
Density Gradient Centrifugation
Sedimentation
Adhesion
Microfluidic Cell Separation
The cell separation method you choose typically depends on what you intend to use the
isolated cells for, and the choice may involve a trade-off. For example, if you need very pure
cells, you will likely choose a method with high purity but that may result in lower yield.
Immunomagnetic cell separation
Immunomagnetic cell separation is a technique whereby magnetic particles are used to isolate
target cells from heterogeneous mixtures. To accomplish this, the magnetic particles are
bound to specific cell surface proteins on the target cells via antibodies, enzymes, lectins, or
streptavidin. The sample is then placed in an electromagnetic field that pulls on the magnetic
particles, bringing the labeled cells with them. The unlabeled cells remain in the supernatant,
thus creating a physical separation between target and non-target cells within the sample.
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Due to its speed and simplicity, immunomagnetic cell separation is one of the most
commonly used methods by which scientists isolate highly purified populations of specific
cell subsets. Immunomagnetic cell separation has several advantages, including:
High purity
Fast protocols
Ease of use
Low equipment cost
Many cells can be isolated at once
Potential for automation
High cell viability
Fluorescence-activated cell sorting
Fluorescence-activated cell sorting (FACS) is a method that uses flow cytometry and
fluorescent probes to sort heterogeneous mixtures of cells. Fluorophore-tagged antibodies
bind to epitopes on specific antigens on the target cells within a single-cell suspension. After
tagging, the flow cytometer focuses the cell suspension into a uniform stream of single cells.
This stream is then passed through a set of lasers that excites the cell-bound fluorophores,
causing light scattering and fluorescent emissions. Based on the wavelengths produced by the
laser excitation, the resulting photon signals are converted into a proportional number of
electronic pulses that assign a charge to the droplet that is formed around the cell. As each
droplet falls between the deflection plates, its charge causes the droplet to either be deflected
into collection tubes or fall into the waste chamber.
Immunomagnetic cell separation is a much faster and simpler procedure than FACS, and is
often the preferred cell isolation method for common cell types. FACS has several
advantages over immunomagnetic cell separation including the ability to:
Sort single cells
Isolate cells based on intracellular markers (e.g. GFP)
Isolate cells based on surface marker expression levels
Sort complex cell types with multiple markers at higher purity
Pre-enrich samples prior to FACS
Isolating rare cell types by FACS can be time consuming, expensive and can result in low
cell recovery. Researchers can pre-enrich their samples for target cells using
immunomagnetic cell separation to reduce the sort time and improve purity and recovery.
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Density gradient centrifugation
Density gradient centrifugation relies on the varying densities of cells within a heterogeneous
sample. The sample is layered on top of a density gradient medium before being centrifuged.
During centrifugation, each cell type will sediment to its isopycnic point, which is the place
in the medium gradient where the density of the cells and medium are equal.
Common applications include the fractionation of peripheral blood mononuclear cells,
exclusion of dead cells from a cell culture, and separation of plasma from blood cells.
There are several types of density gradient media, each with unique properties that render
them ideal for different purposes. The following are examples of the most well-known types:
Lymphoprep™, Lympholyte®, and Ficoll-Paque
® are similar media that consist of
saccharides and sodium diatrizoate; they have a density of 1.077 g/mL. These media are
commonly used to isolate mononuclear cells from peripheral blood, cord blood, and bone
marrow. See our comparison data >
Percoll® (density: 1.131 g/mL) consists of colloidal silica particles coated with
polyvinylpyrrolidone (PVP) and is widely used to separate cells, organelles, viruses, and
other subcellular particles.
OptiPrep™ is a medium consisting of iodixanol in water that is used to isolate viruses,
organelles, macromolecules, and cells.
Density gradient centrifugation is an inexpensive cell separation technique but has limited
specificity, low purity, and low throughput. In addition, even though it is a common
laboratory technique, density gradient centrifugation can be a slow and laborious process that
is difficult to master. Scientists typically need to carefully layer their sample over the density
gradient medium, centrifuge for 30 minutes without brakes, then carefully harvest and wash
the appropriate layer of cells. Technologies like SepMate™ make this method easier and
faster. SepMate ™ is a specialized tube that allows users to quickly layer blood over the
density gradient medium, prevents the layers from mixing and facilitates fast and easy
harvesting of the target cells. With SepMate™, cells can be obtained in as little as 15 minutes.
Immunodensity cell separation
Immunodensity cell separation, also referred to as erythrocyte rosetting, is a negative
selection method that uses a combination of antibody-based labeling and density gradient
centrifugation. With this method, antibodies are added to a whole blood sample, labeling the
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unwanted cells and cross-linking them to red blood cells. This results in the formation of
complexes called immunorosettes that are much denser than the mononuclear cells being
isolated. During centrifugation, the unwanted cells pellet with the red blood cells, leaving the
target cells in a layer above the density medium.
Immunodensity cell separation doesn’t require any specialized equipment beyond a
centrifuge, can be easily incorporated into established density gradient centrifugation
protocols, and can be used to isolate specific cell subsets directly from whole blood. However,
the technique is limited to negative selection, relies on the operator’s blood sample layering
technique, and requires a high concentration of red blood cells in the starting sample.
RosetteSep™ is an example of a commercially available immunodensity cell separation
reagent (Figure 1). RosetteSep™ can be combined with SepMate™ PBMC isolation tubes for
even faster and easier immunodensity cell separation.
Sedimentation
Sedimentation works on the basis that gravity will cause larger and denser components to
sediment faster than materials that are smaller and less dense. The largest and densest
components in a sample can be pelleted through an initial low-force centrifugation due to
their high rate of sedimentation. The supernatant can then be spun again. Through successive
centrifugations, components with an increasingly lower rate of sedimentation can be isolated.
Leukocytes are commonly separated from erythrocytes through dextran
sedimentation. HetaSep™ is an example of an erythrocyte aggregation agent that is used to
separate nucleated cells from red blood cells (RBCs) in whole blood.
Sedimentation is inexpensive but generally results in lower purity than other methods.
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View protocols for cell isolation by sedimentation >
Adhesion
The unique adhesion profiles of different cell types can be used to separate target cells from
heterogeneous populations. By choosing suitable growth factors and cell culture plates to
selectively favor or inhibit adhesion, adherent cells can be separated from cells in suspension.
For example, macrophages are inherently adherent and they are often isolated from peripheral
blood and bone marrow by adhesion. Mononuclear cells can be cultured with serum and a
differentiation cocktail, promoting the formation of an adherent monolayer of macrophages.
After removing the supernatant containing unwanted cells, the macrophages can be isolated.
Alternatively, cells that naturally grow in suspension or have lost anchorage dependency can
be isolated by culturing the heterogeneous cell population in plates designed for ultra-low
attachment. Without a surface to adhere to, adherent cells will fail to survive and the target
cells will remain in suspension.[1]
Microfluidic cell separation
Microfluidics is an umbrella category of cell separation methods.[2]
Designed to manipulate
fluids on a microscopic level in order to facilitate single-cell isolation, microfluidic
technologies are frequently built onto microchips and are also commonly known as ―lab-on-
a-chip" devices. These devices have several advantages, including the smaller volumes of
samples and reagents required for use. Lab-on-a-chip devices are also portable, allowing
them be used virtually anywhere, making them particularly useful as field-based diagnostic
tools.
Microfluidic methods can be divided into active and passive systems. Active microfluidic
systems involve external forces, whereas passive microfluidics make use of the cell’s density
and mass in combination with gravity. These methods can also be classified by the presence
or absence of cell labeling; although some methods involve labeling cells with antibodies,
most methods are known for being label-free. There are several different microfluidic
methods used for cell isolation, including:
Acoustophoresis
Aqueous two phase systems
Biomimetic microfluidics
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Cell affinity chromatography
Deterministic lateral displacement
Electrophoretic sorting
Field flow fractionation
Gravity and sedimentation
Magnetophoresis
Microfiltration
Optical sorting
There are some other techniques of isolation.
Other cell separation techniques
This section summarizes other, less commonly used, cell separation methods.
Aptamer technology
Aptamers are single-stranded RNA or DNA oligonucleotides that form structures that can
bind to highly specific targets. Through systematic evolution of ligands by exponential
enrichment (SELEX) technology, aptamers can be screened and synthesized to target any cell
type. These aptamers have high affinity and specificity toward their targets, and can be
labeled with fluorochromes or magnetic particles to facilitate cell separation. The main
advantage of aptamers is that they lack immunogenicity.
Fluorophore-labeled aptamers have been used to sort mesenchymal stem cells[3]
from bone
marrow and RNA aptamers have been used to isolate mouse embryonic stem cells.[2]
Buoyancy-activated cell sorting
Buoyancy-activated cell sorting is a cell separation technique that utilises glass microbubbles
labeled with antibodies specific to the target cells. When mixed into the sample, the
microbubbles bind to the target cells. Due to the augmented buoyancy force, the
microbubbles float to the surface, separating the target cells.
Complement depletion
The complement depletion method takes advantage of the proteolytic cascade initiated by the
complement system of the immune system. The complement system consists of plasma
proteins that can be activated by pathogens or antibodies. Once activated, the plasma proteins
induce the formation of a membrane-attack complex on a cell, resulting in cell lysis. With
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specific monoclonal antibodies, any cell population can be targeted and lysed through the
complement cascade.
Laser capture microdissection
Laser capture microdissection (LCM) is a technique that uses a narrow laser beam to cleave
target cells or areas from mostly solid tissue samples. Through microscopic visualization,
LCM can isolate cell populations from heterogeneous mixtures using cell morphology or
specific histological and immunological staining. LCM is particularly useful when working
with small sample sizes.
Immunoguided laser capture microdissection
Immunoguided laser capture microdissection combines immunostaining with laser capture
microdissection (see above). This allows immunophenotypes to be used, in addition to
morphology and tissue location, to identify and isolate target cells from the tissue sample.
This technique employs immunohistochemistry or immunofluorescence to guide the
dissection process for isolating cells expressing a specific molecular marker, and is
particularly useful when histological stains do not recognize certain cell populations.
Limiting dilution
Limiting dilution involves isolating single cells through the dilution of a cell suspension. This
technique can be carried out with standard pipetting tools and is commonly used to produce
monoclonal cell cultures and single cell cultures for single-cell analysis.[4]
Micromanipulation
Micromanipulation, a form of manual cell picking, is a cell isolation technique involving the
use of an inverted microscope and ultra-thin glass capillaries connected to an aspiration and
release unit. The system moves through motorized mechanical stages, allowing the operator
to carefully select a specific cell and apply suction via micropipette to aspirate and isolate the
cell.[5]
Cryopreservation
Cryo-preservation or cryo-conservation is a process where organelles, cells, tissues,
extracellular matrix, organs, or any other biological constructs susceptible to damage caused
by unregulated chemical kinetics are preserved by cooling to very low temperatures (typically
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−80 °C using solid carbon dioxide or −196 °C using liquid nitrogen). At low enough
temperatures, any enzymatic or chemical act.
Cryopreservation, the preservation of cells and tissue by freezing. Cryopreservation is
based on the ability of certain small molecules to enter cells and prevent dehydration
and ice crystal formation, which would otherwise destroy cells during the freezing
process
Cryopreservation is based on the ability of certain small molecules to enter cells and
prevent dehydration and formation of intracellular ice crystals, which can cause cell death
and destruction of cell organelles during the freezing process. Two common cryoprotective
agents are dimethyl sulfoxide (DMSO) and glycerol. Glycerol is used primarily for
cryoprotection of red blood cells, and DMSO is used for protection of most other cells and
tissues. A sugar called trehalose, which occurs in organisms capable of surviving extreme
dehydration, is used for freeze-drying methods of cryopreservation. Trehalose stabilizes cell
membranes, and it is particularly useful for the preservation of sperm, stem cells,
and blood cells.
Most systems of cellular cryopreservation use a controlled-rate freezer. This freezing system
delivers liquid nitrogen into a closed chamber into which the cell suspension is placed.
Careful monitoring of the rate of freezing helps to prevent rapid cellular dehydration and ice-
crystal formation. In general, the cells are taken from room temperature to approximately
−90 °C (−130 °F) in a controlled-rate freezer. The frozen cell suspension is then transferred
into a liquid-nitrogen freezer maintained at extremely cold temperatures with nitrogen in
either the vapour or the liquid phase. Cryopreservation based on freeze-drying does not
require use of liquid-nitrogen freezers.
An important application of cryopreservation is in the freezing and storage of hematopoietic
stem cells, which are found in the bone marrow and peripheral blood. In autologous bone-
marrow rescue, hematopoietic stem cells are collected from a patient’s bone marrow prior to
treatment with high-dose chemotherapy. Following treatment, the patient’s cryopreserved
cells are thawed and infused back into the body. This procedure is necessary, since high-dose
chemotherapy is extremely toxic to the bone marrow. The ability to cryopreserve
hematopoietic stem cells has greatly enhanced the outcome for the treatment of
certain lymphomas and solid tumour malignancies. In the case of patients with leukemia,
their blood cells are cancerous and cannot be used for autologous bone-marrow rescue. As a
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result, these patients rely on cryopreserved blood collected from the umbilical cords of
newborn infants or on cryopreserved hematopoietic stem cells obtained from donors. Since
the late 1990s it has been recognized that hematopoietic stem cells and mesenchymal stem
cells (derived from embryonic connective tissue) are capable of differentiating into skeletal
and cardiac muscle tissues, nerve tissue, and bone. Today there is intense interest in the
growth of these cells in tissue culture systems, as well as in the cryopreservation of these
cells for future therapy for a wide variety of disorders, including disorders of the nervous and
muscle systems and diseases of the liver and heart.
Cryopreservation is also used to freeze and store human embryos and sperm. It is especially
valuable for the freezing of extra embryos that are generated by in vitro fertilization (IVF). A
couple can choose to use cyropreserved embryos for later pregnancies or in the event that
IVF fails with fresh embryos. In the process of frozen embryo transfer, the embryos are
thawed and implanted into the woman’s uterus. Frozen embryo transfer is associated with a
small but significant increase in the risk of childhood cancer among children born from such
embryos.[6]
Characterisation of cells and their application
cell culture is the process in which cell growth in a suitable medium under controlled
condition for desire purpose or product. but during this process cells undergoes cross-
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contamination or miss-identification so detect to this require characterization of cell lied. it
also requires to confirm that cell lines derived from its tissue of origin. so characterization
requires to what cell got during culture.
Characterization of cultured cells or cell lines is important for dissemination of cell lines
through cell banks, and to establish contacts between research laboratories and commercial
companies.
Characterization of cell lines with special reference to the following aspects is generally
done
1. Morphology of cells
2. Species of origin.
3. Tissue of origin.
4. Whether cell line is transformed or not.
5. Identification of specific cell lines.
Morphology of cells
A simple and direct identification of the cultured cells can be done by observing their
morphological characteristics. However, the morphology has to be viewed with caution since
it is largely dependent on the culture environment. For instance, the epithelial cells growing
at the center (of the culture) are regular polygonal with clearly defined edges, while those
growing at the periphery are irregular and distended (swollen).
The composition of the culture medium and the alterations in the substrate also influence the
cellular morphology. In a tissue culture laboratory, the terms fibroblastic and epithelial are
commonly used to describe the appearance of the cells rather than their origin.
Fibroblastic cells
For these cells, the length is usually more than twice of their width. Fibroblastic cells are
bipolar or multipolar in nature.
Epithelial cells
These cells are polygonal in nature with regular dimensions and usually grow in monolayers.
The terms fibroblastoid (fibroblast-like) and epitheloid (epithelial-like) are in use for the cells
that do not possess specific characters to identify as fibroblastic or epithelial cells.
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Species of origin of cells
The identification of the species of cell lines can be done by
a. Chromosomal analysis.
b. Electrophoresis of isoenzymes.
c. A combination of both these methods.
In recent years, chromosomal identification is being done by employing molecular probes.
Identification of tissue of origin
The identification of cell lines with regard to tissue of origin is carried out with
reference to the following two characteristics
1. The lineage to which the cells belong.
2. The status of the cells i.e. stems cells, precursor cells.
Tissue markers for cell line identification
Some of the important tissue or lineage markers for cell line identification are briefly
described.
Differentiated products as cell markers
The cultured cells, on complete expression, are capable of producing differentiation markers,
which serve as cell markers for identification.
Some examples are given below
a. Albumin for hepatocytes.
b. Melanin for melanocytes
c. Hemoglobin for erythroid cells
d. Myosin (or tropomyosin) for muscle cells.
Enzymes as tissue markers
The identification of enzymes in culture cells can be made with reference to the
following characters
a. Constitutive enzymes.
b. Inducible enzymes.
c. Isoenzymes.
The commonly used enzyme markers for cell line identification are given in Table 35.1.
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Tyrosine aminotransferase is specific for hepatocytes, while tyrosinase is for melanocytes.
Creatine kinase (MM) in serum serves as a marker for muscle cells, while creatine kinase
(BB) is used for the detection of neurons and neuroendocrine cells.
Filament proteins as tissue markers
The intermediate filament proteins are very widely used as tissue or lineage markers.
For example
a. Astrocytes can be detected by glial fibrillary acidic protein (GFAP).
b. Muscle cells can be identified by desmin.
c. Epithelial and mesothelial cells by cytokeratin.
Cell surface antigens as tissue markers
The antigens of the cultured cells are useful for the detection of tissue or cells of origin. In
fact, many antibodies have been developed (commercial kits are available) for the
identification cell lines (Table. 35.2). These antibodies are raised against cell surface antigens
or other proteins.
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The antibodies raised against secreted antigen a-fetoprotein serves as a marker for the
identification of fetal hepatocytes. Antibodies of cell surface antigens namely integrin’s can
be used for the general detection of cell lines.
Transformed cells
Transformation is the phenomenon of the change in phenotype due to the acquirement of new
genetic material. Transformation is associated with promotion of genetic instability.
The transformed and cultured cells exhibit alterations in many characters with
reference to
a. Growth rate
b. Mode of growth
c. Longevity
d. Tumorigenicity
e. Specialized product formation.
While characterizing the cell lines, it is necessary to consider the above characters to
determine whether the cell line has originated from tumor cells or has undergone
transformation in culture.
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Identification of specific cell lines
There are many approaches in a culture laboratory to identify specific cell lines
1. Chromosome analysis
2. DNA detection
3. RNA and protein analysis
4. Enzyme activities
5. Antigenic markers.[7]
Application
Disease diagnosis: Cell culture techniques are applied in clinical medicine for the
diagnosis of infectious diseases especially diseases caused by pathogenic viruses. Cell
culture techniques aid in rapid viral detection from clinical samples. It also aids in the
early treatment of viral infections once the causative viral agent have been detected. Over
the years, viral disease diagnosis has traditionally relied on the isolation of viral
pathogens in cell cultures which some perceive as being slow and requires special
technical expertise. However, advances in cell culture-based viral diagnostic products and
techniques including but not limited to cryopreserved cell cultures, centrifugation-
enhanced inoculation, precytopathogenic effect detection, co-cultivated cell cultures, and
transgenic cell lines have made cell culture to be useful for the diagnosis of viral diseases.
Biomedical research: In biomedical research, cell culture techniques are most preferable
than the use of animals for research. Since the use of animals such as monkeys and
chimpanzees for research could lead to the extinction of these animals, cell culture
techniques is a good alternative and replacement to prevent the extinction of some
wildlife. Cell culture techniques can be applied in biomedical research especially in the
area of studying some molecular disease processes, and finding out ways via which these
diseases of non-microbial origin could be better treated. With the application of cell
culture techniques in biomedical research, improved and prompt ways of detecting
disease causative agents could be developed. Cell culture techniques could also be used as
model system to study basic cell biology, metabolism and the physiology of living
systems.
Virology: In the field of virology, animal cell culture techniques can be used to replicate
the viruses used for vaccine production instead of using animals for this purpose. Cell
culture techniques can also be used to detect and isolate pathogenic viruses from clinical
samples. It can also be used to study the growth and development cycle of viruses. Cell
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culture techniques can also be used in virology to study the mode of infection of viral
disease agents.
Genetic engineering: In genetic engineering, cultured animal cells can be used to
introduce new genetic material like DNA or RNA into another cell. Such exchange of
genetic information amongst cells or organisms can be used to study the expression of
new genes and its effect on the health of the recipient host cell. The recipient host cell
starts expressing novel proteins that could be of immense industrial and medical
importance. Animal cell cultures are used to produce commercially important genetically
engineered proteins or immunobiologicals such as monoclonal antibodies, polyclonal
antibodies, insulin, anticancer agents and hormones.
Model systems: Cell culture techniques are used in model systems to study the effect of
drugs in human or animal host. It can also be used to study the process of aging in
humans. In model systems, cell culture techniques are used to study the major triggers for
ageing in man. It can also be used to study how host cell and disease causing agents like
bacteria, fungi and viruses interact in vivo.
Cancer research: Cell culture techniques is used in cancer research to study the basic
difference between normal cells and cancer cells since both cells can be cultured in
vitro in the laboratory. Normal cells can be converted into cancer cells by using radiation,
chemicals and viruses. This allows the mechanism and cause of cancer to be studied in
vitro using cell culture techniques. Cell culture techniques can also be used to determine
the effective chemotherapeutic drugs that can selectively destroy only cancer cells
without harming the host cells since most cancer drugs have several untoward effects on
the host.
Toxicity testing of novel drugs: Cell culture techniques can be used to study the effects
of novel drugs, cosmetics and other chemical agents in order to determine not just their
efficacy but also the level of their toxicity (i.e. cytotoxicity). The toxicity of the newly
developed drugs to vital organs of the body such as the liver and kidney (that are involved
in drug metabolism) is also evaluated using cell culture techniques. Drug dosages for
novel drugs can also be determined using cell culture techniques.
Gene therapy: Gene therapy is an experimental technique that uses genes to treat or
prevent disease especially molecular or non-infectious diseases such as cancer. It allows
clinicians to treat a genetic disorder by inserting a functional gene (to replace a
dysfunctional gene) into a patient’s cells instead of using the conventional treatment
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methods such as the use of drugs, chemotherapy or surgery. In gene therapy techniques, a
dysfunctional gene is replaced with a functional gene. Through cell culture techniques,
cultured animal cells are genetically altered and made functional so that they can be used
in gene therapy techniques. Briefly, cells are removed from the patient lacking a
functional gene or missing a functional gene; and the extracted cells are cultured in
vitro through cell culture techniques. These dysfunctional genes are replaced by
functional genes. Gene therapy uses a vector, typically a virus, to deliver a gene to the
cells where it is needed. Once inside the host cell, the host cell’s gene-reading machinery
uses the information in the introduced functional gene to build ribonucleic acid (RNA)
and protein molecules which will now replace the lost activities of the replaced
dysfunctional gene.
Vaccine development: Cell culture techniques can be used in vaccine development since
they help to culture animal cells in vitro. Cultured animal cells are in turn used in the
production or propagation of viruses that are used to produce vaccines. These vaccines
are used clinically for the prevention of communicable diseases caused by pathogenic
viruses including measles, polio, rabies, hepatitis and chicken pox and there preventable
diseases.[8]
CONCLUSION
Cell culture techniques is vital tool in the process of drug Discovery which ultimately leads to
quantify the steps of analysis of therapeutic potential of drugs. Gene therapy depends on the
analysis of cell culture to disclose the unidentified facts related to genomics. Some parts of
vaccines development and designing also take note from cell culture report. In the field of
communicable disease the fate of cellular behaviour helps a lot. Furturresearch must be focus
and emphasise in this field.
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