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In vitro Cell Migration, Wound Healing Full...
Continue ReadingWhat is Cell Migration?
Cell migration, movement of cells from one area to another generally in response to a chemical stimulus.
Migration is a central to achieving functions such as wound repair, cell differentiation and embryonic development, immune response, and disease processes such as cancer metastasis and inflammation.
Methods to examine cell migration are very useful and important for a wide range of biomedical research such as cancer biology, immunology, vascular biology, cell biology and developmental biology
Principle of Cell Migration
Migration assay is commonly used test to study the migratory response of endothelial cells or tumor cells to tumor promoters or inhibitors.
In migration assay the cells are seeded and when it reaches a confluency of 90% a scratch is made to mimic a wound.
This wounded cells are exposed to different concentration of test compounds to check its effect on the migratory property of cells.
Cell Migration Assay Requirements
1. Cancer cell line
2. Complete growth media
3. 1X PBS (pH-7.2)
4. 6-well plate
5. Micro-tips
6. Pipettes
7. Light Microscope
8. CO2 incubator
9. Biosafety cabinet level II
Procedure of Cell Migration Assay
1. Seed appropriate number of cells (e.g. for MDA-MB-231 seed approx. 2.5×10^5 cells per well) in 6 well plates.
Make duplicates for each treatment and control group. Incubate in 5% CO2 incubator for 24-36 hours until they reach ̴ 90% confluence.
2. When the cells are ̴ 90% confluence, make a scratch with 200 μL tip in the middle of each well and flush the scratch area with 1 ml pipette gently, so as any of adhered cells from the scratch area get detached.
3. Discard the media wash with 1 ml PBS and add freshly prepared treatment media.
4. Take the images at 0, 12, 24 and 48h, as per the requirement of your experiment (Observe the cells before taking images each time, sometimes some percentage of cells at higher doses get died or undergo apoptosis after 12h or 24h.
These cells come out in the scratch area and we get blurred image of the scratch area with undefined boundaries.
In such case, first collect the treatment media in separate tubes, wash with PBS and keep PBS in wells while taking images.
After taking images, replace PBS with treatment media).
5. Arrange the representative images for each treatment group and perform ImageJ analysis to measure the scratch area and represent your results as percent migration area or wound closure area.
Cell Migration Assay Precautions
Do not use non sterile tips to make the scratch.
Make the scratch straight with steady hands otherwise the wound will not be clear and the cells will be disturbed which will result in insignificant data.
Always take the image of area which shows least dead or floating cells to produce significant and publishable data.
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Introduction to Cell Culture: SubCulture, Media, Protocol
Continue ReadingWhat is Cell Culture?
Cell culture refers to the removal of cells from an animal or plant and their subsequent growth in a favourable artificial environment.
The cells may be removed from the tissue directly and disaggregated by enzymatic or mechanical means before cultivation, or they may be derived from a cell line or cell strain that has already been established.
Mammalian cell culture is used widely in academic, medical and industrial settings.
It allows the study of physiology and biochemistry of the cell and its development.
It has also widen the scope and its application in the field of cell and molecular biology where the use of reproducible model systems is attained by cultured cell lines.
For medical use, cell culture provides test systems to assess the efficacy and toxicology of potential new drugs.
Large- scale mammalian cell culture has allowed production of biologically active proteins, initially production of vaccines and then recombinant proteins and monoclonal antibodies; recent innovative uses of cell culture include tissue engineering to generate tissue substitutes.
Basic Requirements for Successful Cell Culture
The first necessity is a well-established and properly equipped cell culture facility.
The level of bio containment required (Levels 1–4) is dependent on the type of cells cultured and the risk that these cells might contain, and transmit, infectious agents.
All facilities should be equipped with the following: a certified biological safety cabinet a centrifuge, preferably capable of refrigeration and equipped with appropriate containment holders that is dedicated for cell culture use; a microscope for examination of cell cultures and for counting cells; and a humidified incubator set at 37°C with 5% CO2 in air.
The second requirement for successful cell culture is the practice of sterile technique prior to beginning any work, the biological safety cabinet should be turned on and allowed to run for at least 15 minute to purge the contaminated air.
All work surfaces within the cabinet should be decontaminated with an appropriate solution; 70% ethanol or isopropanol are routinely used for this purpose.
A third necessity for successful cell culture is appropriate, quality controlled sterile reagents, culture media and required sterile plastic wares.
Adopted from BioRender
Types of Cell Culture
There are two types of cell culture system, Primary and continuous culture.
1. Primary Culture: Primary cultures are derived directly from excised, normal animal tissue and cultures either as an explant culture or following dissociation into a single cell suspension by enzyme digestion.
The preparation of primary cultures is labour intensive and they can be maintained in vitro only for a limited period of time.
During their relatively limited lifespan primary cells usually retain many of the differentiated characteristics of the cell in vivo.
2. Continuous Culture: Continuous cultures are comprised of a single cell type that can be serially propagated in culture either for a limited number of cell divisions (approximately thirty) or otherwise indefinitely.
Cell lines of a finite life are usually diploid and maintain some degree of differentiation.
The fact that such cell lines senesce after approximately thirty cycles of division means it is essential to establish a system of Master and Working banks in order to maintain such lines for long periods.
Continuous cell lines that can be propagated indefinitely generally have this ability because they have been transformed into tumour cells.
Tumour cell lines are often derived from actual clinical tumours, but transformation may also be induced using viral oncogenes or by chemical treatments.
Transformed cell lines present the advantage of almost limitless availability, but the disadvantage of having retained very little of the original in vivo characteristics.
Cell Culture Morphology
In terms of growth mode cell cultures take one of two forms, growing either in suspension (as single cells or small free floating clumps) or as a monolayer that is attached to the tissue culture flask.
The form taken by a cell line reflects the tissue from which it was derived e.g. cell lines derived from blood (leukaemia, lymphoma) tend to grow in suspension whereas cells derived from solid tissue (lungs, kidney) tend to grow as monolayers.
Attached cell lines can be classified as endothelial, epithelial, neuronal or fibroblasts and their morphology reflects the area within the tissue of origin.
There are some instances when cell cultures may grow as semi-adherent cells, ( e.g. marmoset B-lymphoblastoid cell line), where there appears to be a mixed population of attached and suspension cells.
For these cell lines it is essential that both cell types are subcultured to maintain the heterogeneous nature of the culture.
In vitro age of cell culture Two terms are predominantly used to define the age of a cell culture:
1. Passage number indicates the number of times the cell line has been sub-cultured.
2. The population doubling (pd) number indicates the number of cell generations the cell line has undergone i.e. the number of times the cell population has doubled.
Cell Culture Maintenance
In culturing mammalian cells in vitro, one attempts to reproduce in a culture vessel the physiological environment and characteristic responses of individual cell types.
At a minimum, the fluid medium in which cells are cultured must provide for their nutritional requirements, provide an energy source, maintain pH, and provide a level of osmolarity compatible with cell viability.
Culture media commonly used today consist of two parts: a basal nutrient medium and supplements.
The basal nutrient medium, such as Dulbecco’s modified Eagle’s medium (DMEM; also known as Dulbecco’s minimal Eagle’s medium), RPMI 1640, or Ham’s F-12, is a buffered aqueous solution of inorganic salts, vitamins, amino acids and other anabolic precursors, energy sources such as glucose and glutamine, and trace elements.
Supplements are either undefined, such as fetal bovine serum (FBS), tissue extracts, and conditioned medium, or defined, such as hormones and growth factors, transport proteins, and attachment factors.
The compositions of basal nutrient media and medium supplements may vary considerably; however, both components of the complete medium are necessary for support of cell viability and proliferation.
There are two formats of media available i.e. dehydrated and liquid media.
Preparation of Cell Culture Media
Requirements:
1. Dehydrated medium (i.e. RPMI or DMEM)
2. Double autoclaved water
3. Autoclaved bottles for storing medium
4. 0.22 micron filtration unit (capacity 1 litre)
5. Antibiotics (Penicillin, Streptomycin, Amphotericin B)
6. Additives for the medium as required (like sodium bicarbonate, sodium pyruvate, glutamate etc.)
7. 1N NaOH and HEPES
8. Conical Flask (1L or 2L for dissolving the dehydrated medium)
9. Measuring Cylinder
10. Discard beaker
Cell Culture Procedure
1. In a sterile biosafety cabinet dissolve powdered medium with constant stirring in a 0.8× to 0.9 × volume of water. (If a commercially prepared liquid medium is being used, add penicillin and streptomycin from commercial stock solutions and proceed to step 8)
2. Add an amount of HEPES that yields a concentration of 15 mM in the final volume of medium. (Omit this step if the powdered medium is formulated with HEPES.)
3. Add the amount of sodium bicarbonate recommended by the medium supplier for use in a CO2-controlled atmosphere (e.g., 14 to 36 mM in 5% CO2 atmosphere). (Omit this step if the powdered medium contains sodium bicarbonate.)
4. Add glutamine to give a final concentration of 2 mM and pyruvic acid to give a final concentration of 0.01% (w/v). (Omit this step if the powdered medium contains glutamine & pyruvic acid.)
5. Add penicillin G to give a final concentration of 100 IU/ml and streptomycin to give a final concentration of 50 μg/ml.
6. (Other antibacterial agents or antifungal agents should not be routinely included in culture medium. Gentamicin at a final concentration of 50 μg/ml or kanamycin at 100 μg/ml may be useful in eliminating gram-positive and gram-negative bacteria from primary cultures or from irreplaceable cultures, but it is best to discard any cultures that are contaminated with bacteria, yeast, or fungi.)
7. Adjust the pH of the medium to 7.4 with 1 N NaOH, and add water to achieve the final (1×) volume. Readjust the pH of the medium to 7.4, if necessary.
8. Sterilize the medium by filtration through a 0.2-μm filter. Store the medium at 4°C in the dark. Vacuum-operated filtering units or bottle-top filters are useful for small volumes of medium (0.1 to 2 liters), whereas filter capsules (2 to 5 liters) or filter stands (>10 liters) that are used under positive pressure are more suitable for larger volumes.
9. Make an aliquote of prepared media and keep at 37°C to check for any contamination.
10. Add serum (5-20%) to the desired final concentration at the time of use.
Cell Culture Precautions
Basal nutrient medium and the serum supplement should be stored individually at 4°C, and the complete medium should be made up at the time of use and only in the volume necessary.
Working volumes of serum should be stored at 4°C and used within several weeks.
Serum should not be subjected to repeated freezing and thawing, but it can be stored for at least 2 years at −20°C with little deterioration in growth-promoting activity.
In this way, medium components are not wasted, and the chances of detecting, isolating and eliminating contamination with minimal losses are increased.
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What is Immunocytochemistry? Immunocytochemistry Methods, Techniques
Continue ReadingAbout Immunocytochemistry
Immunocytochemistry (ICC) is a common laboratory technique that uses antibodies that target specific peptides or protein antigens in the cell via specific epitopes.
The antibodies bound on diffirent epitotoes can then be detected using several different methods.
Immunocytochemistry enables researchers to evaluate whether or not cells in a particular sample express the antigen in question.
In cases where an immunopositive signal is found, ICC also allows researchers to determine which sub-cellular compartments are expressing the antigen.
In the present experiment we are identifying the neurons by the presence of neuron-specific marker Neurofilament-H in the primary cotical neuronal culture.
Adopted from BioRender
Immunocytochemistry Requirements
1. 4% Para formaldehyde in PBS
2. Triton X-100, Milli Q water
3. Bovine serum albumin (BSA)
4. 5% w/v BSA in Phosphate buffered saline (PBS)
5. 0.1% w/v BSA in PBS
6. Primary antibody produced in rabbit
7. Goat-Anti rabbit secondary antibodies.
8. Inverted florescent microscope
9. TMB substrate for localization(20X)
Procedure of Immunocytochemistry
1. Wash the cultures two times with PBS and then fix for 30 minutes with 4% paraformaldehyde – PBS at room temperature followed by permeabilisation with 0.2% Triton X-100 for 5 minutes.
2. Block the non-specific binding by incubating the cells with PBS containing 5% BSA w/v for 1 h at room temperature.
3. After blocking, wash once with PBS and incubate with primary antibody overnight at 4°C In the present experiment we are using specific antibody as neuronal marker.
The antibody should be diluted with PBS with 0.1% BSA (w/v) according to manufacterur’s instructions.
In the present case we are diluting 1part of antibody with 399 parts of 0.1% BSA-PBS.
4. Once the incubation time is over, remove the primary antibody and wash three times with 0.1% BSA-PBS
5. After washing incubate the cultures with secondary antibodies which was goat anti rabbit IgG conjugated with HRP (1:3000) in 0.1% BSA-PBS for 1 hour at room temperature.
6. Following the incubation, remove the antibody solution and wash three more times with 0.1% BSA-PBS.
7. Add 200μl of TMB substrate to each of the well and incubate in dark for few minutes.
Remove the substrate as soon as color starts appearing and observe under the microscope
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Cell Viability Assays: Overview of Cell Viability...
Continue ReadingAbout Calcein AM / PI Cell Viability Assays
The Calcein –AM/PI assay is a two-colour fluorescence cell death assay that is based on the simultaneous determination of live and dead cells with two different probes.
It measure recognized parameters of cell viability by evaluating intracellular esterase activity and plasma membrane integrity : calcein-AM and Propidium iodide (PI) .
The assay principles are general and applicable to most eukaryotic cell types, including adherent cells, suspension cells, and certain tissues, but not to bacteria or yeast.
The viable cells are distinguished by the presence of ubiquitous intracellular esterase activity, determined by the enzymatic conversion of the virtually nonfluorescent cell- permeant calcein AM to the intensely fluorescent calcein.
The calcein, a polyanionic dye is well retained within viable cells, producing an intense uniform green fluorescence in viable cells (ex/em ~495 nm/~515 nm).
Propidium iodide enters cells with damaged membranes in dead cells and undergoes a 40-fold enhancement of fluorescence upon binding to nucleic acids (DNA, RNA) thereby producing a bright red fluorescence in dead cells (ex/em ~535 nm/~617 nm).
Adopted from from BioRender
Cell Viability Assays Principle
PI is excluded by the intact plasma membrane of live cells.
The cell viability determination depends on these physical and biochemical properties of cells.
Cytotoxic events that do not affect these cell properties may not be accurately assessed using this method.
Background fluorescence levels are inherently low with this assay technique because the dyes are virtually non- fluorescent before interacting with cells.
This assay is suitable for use with a wide variety of techniques, including microplate assays immunocytochemistry, flow cytometry, and in vivo cell tracing
Cell Viability Assays Requirements
1. Calcein-AM fluorescent dye
2. Propidium iodide fluorescent dye
3. Phosphate Buffer Saline (PBS) (i.e. 137mM NaCl, 2.7mM KCl, 10mM Na2HPO4.2H2O and 2 mM KH2PO4, pH 7.4).
4. Fluorescence microscope with blue (FITC) and green (Rhodamine) filters 5. 37oC water bath
Cell Viability Assays Procedure
1. Prepare the stock of Calcein-AM dye in DMSO at a concentration of 1mg/ml. Similarly prepare PI stock in sterile water at a concentration of 10mg/ml
NOTE: These dyes are highly light sensitive and should be stored in smaller aliquots frozen in -20`C till use; PI is a suspected mutagen and hence should be handled with great care
2. Adherent cells may be cultured on sterile glass coverslips as either confluent or subconfluent monolayers (e.g., fibroblasts are typically grown on the coverslip for 2–3 days until acceptable cell densities are obtained).
The cells may be then cultured inside 35 mm disposable petri dishes or other suitable containers; non-adherent cells may also be used.
NOTE: If inverted fluorescence microscope is used, then the cells can be grown in the culture plates and observed directly in the microscope If a normal (upright) fluorescence microscope is available, cells can be grown on coverslips, loaded with dye and carefully mounted on glass slides, and observed under microscope.
3. Remove the medium from the cells in the plates (for cells in suspension, centrifuge and remove the medium; for adherent cells medium can be directly removed).
4. Wash twice with PBS buffer. It is important to wash twice as traces of serum in the medium might interfere with the loading of the dyes.
5. Add fresh buffer and add Calcein-AM dye so that the final concentration of the dye is 1μg/ml.
6. Incubate at 37`C in dark for 15 minutes
7. Now add PI dye so that the final concentration is 10μg/ml. Incubate in dark at 37`C for a further 5 minute duration
8. Now remove the dye containing buffer and add fresh buffer.
9. Observe under the microscope FITC filter can be used for viewing the calcein stained live cells and Rhodamine filter can be used for PI stained dead cells
10. Count the number of calcein positive cells and PI positive cells also the total number of cells in each field of view. Count atleast five fields for each plate.
11. The cell viability can be expressed as (the number of calcein positive cells/ total number of cells) x 100
Features of About Calcein AM / PI Cell Viability Assays
1. Suitable for both proliferating and non-proliferating cells
2. Ideal for both suspension and adherent cells
3. Rapid (no solubilization step)
4. Ideal for high-throughput assays
5. Better retention and brightness compared to other fluorescent compounds (i.e. fluorescein)
6. Useful in a variety of studies, including cell adhesion, chemotaxis, multi-drug resistance
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Electroporation – an Overview, An Efficient Electroporation...
Continue ReadingWhat is Electroporation?
The process of introducing nucleic acids (DNA or RNA) into eukaryotic cells by nonviral methods is defined as transfection.
Using various chemical, phospho-lipid or physical methods, this gene transfer technology is a powerful tool to study gene function and protein expression.
Transfection is a method that neutralizes or obviates the issue of introducing negatively charged molecules (e.g., neagtively charged phosphate backbones of DNA and RNA) into cells with a negatively charged membrane.
Various chemicals such calcium phosphate and DEAE-dextran or cationic lipid-based reagents coat the DNA, neutralizing or even creating an overall positive charge to the molecule and thus enable sucessful delivery inside the cells.
This makes it easier for the DNA or RNA:transfection reagent complex to cross the membrane, especially for phospho-lipids that have a “fusogenic” component, which enhances fusion with the lipid bilayer.
Physical methods like microinjection or electroporation simply punch through the membrane and introduce DNA directly into the cytoplasm. .
Electroporation Principle
In electroporation a high-intensity electrical field transiently permeabilizes the cell membrane, enabling uptake of exogeneous molecules from the surroundings.
This technique has been used to introduce nucleotides, DNA, RNA, proteins, carbohydrates, dyes and virus particles into prokaryotic and eukaryotic cells.
It provides a valuable and effective alternative to other physical and chemical methods for transfection.
Adopted from from BioRender
Requirements for Electroporation
Sterile:
1. Electroporation cuvettes
2. 6 well plates
3. Microtips
4. Growth medium
5. Cells (A 549 Cell line)
6. Trypsin EDTA
7. Phosphate Buffer Saline (PBS) (i.e. 137mM NaCl, 2.7mM KCl, 10mM Na2HPO4.2H2O and 2 mM KH2PO4, pH 7.4).
8. DNA
Non sterile:
9. 70% IPA
10. Cotton balls
11. Electroporator
Cell Preparation
1. One – two days prior to electroporation, transfer the cells into 25cm2 flasks with fresh growth medium so that they will be 50–70% confluent on the day of the experiment.
For most cell lines the cell density will be 2–10 x 10^6 cells/ flask; about 1–20 x 105 cells are needed per electroporation.
2. Trypsinize the cells as described in previous experiment and scrap the cells if required with scrapper and transfer the cells along with the medium in to 15 ml centrifuge tube, and centrifuge the cells at 1300 rpm for 7 min at 40 C.
3. Discard the supernatant and re-suspend the cells in the required quantity of PBS (Phosphate buffer saline)
4. Count the cells using Neubar’s chamber and calculate the volume of the cell suspension to be used in the experiment (2 x 10^6 in 200 μl)
Electroporation Procedure
1. Set the parameter required for electroporation likes voltage, time pulse, no. of pulse (V 75 to 200, Pulse length 15 to 30 mili seconds)
Note: This is variable from instrument to instrument and cell type
2. Add plasmid DNA to the cuvettes (5 to 15 μg)
3. Then add 200 μl of cell suspension (containing 2 x 10^6 cells) to the 0.2 cm cuvette and tap the side of the cuvette to mix.
4. Place the cuvette in the shockPod. Push down the lid to close. Pulse once
5. Immediately after the pulse, transfer the cells to a 6 well plate and add 1.8 ml fresh media
6. Rock the plates gently to assure even distribution of the cells over the surface of the plate. Incubate the plates at 37 oC in 5% CO2 in a humidified incubator.
7. Add suitable concentration of antibiotics after 24- 48 hrs.
8. Do not keep the cells out for longer time after electric shock.
9. Do not rock the plates forcefully.
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Apoptosis Assays: A Step by Step Complete...
Continue ReadingAbout Apoptosis
Annexin V is a phospholipid binding protein with high affinity for binding to phosphotidyl serine (PS), which is a phospholipid present exclusively in the inner leaflet of the cell membrane.
Annexin V does not bind to normal cells because of its inability to penetrate the lipid bilayer.
During apoptosis, due to changes in membrane permeability, PS is translocated to the outer surface of the membrane with which Annexin V binds with high affinity in the presence of Ca .
Detection of cell surface PS with Annexin V thus serves as a marker for apoptotic cells. Similarly, propidium iodide (PI), which specifically stains the dead cells, is used as a marker for necrosis.
Adopted from from BioRender
Apoptosis Principle Assay
In this assay, we are using fluorescently tagged Annexin V and PI as markers for apoptotic and necrotic cells respectively.
The number of annexin positive cells and PI positive cells are counted under the fluorescence microscope to estimate the extent of apoptosis and necrosis in response to specific drug treatment.
Materials Required for Apoptosis Assays
Non Sterile:
1. Phosphate Buffer Saline (PBS) (i.e. 137mM NaCl, 2.7mM KCl, 10mM Na2HPO4.2H2O and 2 mM KH2PO4, pH 7.4).
2. Annexin binding buffer (10mM HEPES (pH 7.5), 140mM NaCl, 2.5mM CaCl2)
3. Annexin V FITC (1mg/ml in PBS)
4. Propidium iodide (1mg/ml in PBS)
5. Fluorescence microscope with green and blue filters
Apoptosis Assay Procedure
1. Remove the medium and wash the cells with PBS thrice and once with annexin binding buffer
2. Add 100μl of annexin binding buffer and subsequently add annexin V-FITC (final concentration should be between 2 μg/ml)
3. Incubate the cells in dark at 37oC for 30 minutes
4. Add PI (Final concentration 1 μg/ml) and keep at RT for 10 minutes
5. Wash the cells twice with annexin binding buffer and observe under the fluorescence microscope
6. Capture atleast 5 fields for each treatment and don’t forget to take phase contrast images of all the fields.
Apoptosis Assay Calculation
Percentage of apoptotic cells = [no. of annexin positive cells/total no. of cells] x 100 Percentage of necrotic cells = [no. of PI positive cells/total no. of cells] x 100
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Cell Viability Assay: Neutral Red Uptake Assay...
Continue ReadingAbout Neutral Red Cell Viability Assay
The NRU cytotoxicity assay or Neutral red uptake assay procedure is a cell survival or cell viability chemosensitivity assay based on the ability of live cells to incorporate and bind neutral red (NR), a supravital dye.
Principle of Neutral Red Cell Viability Assay
NR is a weak cationic dye that rapidly enters into cells via cell plasma membranes by non-ionic diffusion and accumulates intracellularly in lysosomes.
Cell surface alterations or the sensitive lysosomal membrane lead to lysosomal fragility and other modifications that gradually become irreversible.
These changes brought about by the action of any toxic compound result in a decreased uptake and binding of NR.
It is thus possible to distinguish between dead cell, viable cells, damaged cells, which is the basis of the NRU cytotoxicity assay or Neutral red uptake assay.
Healthy and viable mammalian cells, when maintained in culture conditions, continuously proliferate and divide over time.
Any chemical compound that will interfere with this process and result in a reduction of the growth rate due to toxic nature of compound as reflected by cell number or cell death.
Cytotoxicity is expressed as a concentration dependent reduction of the uptake of the NR after chemical exposure thus providing a sensitive, integrated signal of both cell integrity and growth inhibition.
Created with BioRender
Neutral Red Cell Viability Assay Requirement
Sterile:
1. Secondary cell line
2. Growth medium with 10% FCS; Growth medium with 5% FCS
3. Trypsin (0.25%) + EDTA, (1 mM) in PBS
4. Multiwell plates (96 well), Reservoir for multichannel pipette.
5. Pipettor tips, in an autoclavable tip box
6. Universal containers or tubes
7. Cytotoxic drug, 70% IPA
8. Neutral red stock solution (3.3 mg/ml in water)
9. Neutral red medium (1:100 dilution of neutral red stock in medium)
10. Neutral red desorbing medium(50% ethanol, 1% glacial acetic acid)
Non-sterile:
11. Plastic box (clear polystyrene, to hold plates)
12. Multichannel pipettor, neuber’s chamber
13. Dimethyl sulfoxide (DMSO)
14. ELISA plate reader
Neutral Red Cell Viability Assay Procedure
Cells are plated in 96 well plate at a concentration of 1×104 cells / well.
The medium used is RPMI-1640 medium supplemented with10% FBS (Fetal Bovine Serum).
The plate is incubated for 24 hr at 37 C in CO2 incubator with 5% CO2.
The dilutions of the drugs are prepared in RPMI-1640 medium supplemented with 5% FBS.
After 24 hr of seeding, medium is removed and cells are treated with different concentrations of drug.
Treatment is done in 96 well plates and is same as given for MTT assay.
150 μl of drug containing medium is added per well and incubated for 72 hr at 37oC in CO2 incubator.
Control wells are treated with drug free medium. DMSO control wells are treated with medium containing DSMO (as calculated by the dilutions used for the assay)
After the incubation period of 72 hr, the medium is removed.
150 μl of Neutral red medium is added to each well and the plate is incubated at 37C in CO2 incubator.
After the incubation period of 3 hr, medium is removed and cells are washed with 1X PBS and 100 μl of neutral red desorbing solution is added.
Record optical density (O.D.) at 540/570 nm using ELISA plate reader.
Cell viability is calculated as a ratio of control and plotted against log concentration of drug to calculate the IC50.
Analysis of Neutral Red Cell Viability Assay
Plot a graph of the absorbance (y-axis), considering cells without drug as 100% against the concentration of drug (x-axis).
Calculate the LC50 as the drug concentration that is required to reduce the absorbance to half that of the control.
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Breast Cancer : Types of Treatment Options...
Continue Reading"Main challenge with the surgery is the recurrence of the tumor from the cells which are left during surgery there should not be a single tumor cells left during surgery"
Therapies for breast cancer
Surgery: Surgical removal of visible cancer cells and tissues by experts to avoid spread of the tumor to other parts of the body. Amount of tissues removed depends upon the size of the tumor.
There are many types of surgeries needle biopsy, lumpectomy, mastectomy, modified mastectomy are common types of surgeries in breast cancer.
During needle biopsy minute amount of cells or tissue is removed with the help of needle under the guidance of imaging techniques.
Needle biopsy is done for the diagnostic purpose. In lumpectomy a small lump of the tumor tissue is removed for the diagnosis purpose.
"Usually surgery follows chemotherapy or adjuvant chemotherapies to further eradicate the remaining breast cancer cells and prevent the recurrence of the disease"
In mastectomy a large portion of the breast removed along with normal surrounding tissue while in modified mastectomy tumor tissue, surrounding normal tissue along with armpit lymph node are removed are removed.
Main challenge with the surgery is the recurrence of the tumor from the cells which are left during surgery there should not be a single tumor cells left during surgery.
Surgery is also associated with inflammatory complications. Usually surgery follows chemotherapy or adjuvant chemotherapies to further eradicate the remaining cells and prevent the recurrence of the disease.
Sometime radiation therapy is recommended after mastectomy to reduce the recurrence of the tumor.
"When breast cancer cells exposed to high energy radiation such as X-rays cancer cells are not able to repair the DNA damage"
Radiotherapy: It involves the administration of ionizing radiation, X-ray or gamma rays to the tumor site.
Normal cells suspend cell division when their DNA get damage by halting their DNA replication does not proceed until get repaired when the DNA damage is irreparable cells undergoes to programmed cell death.
Cancer cells are genetically unstable cells which has lost the power repairing the DNA damage for their own benefits.
When cancer cells exposed to high energy radiation such as X-rays cancer cells are not able to repair the DNA damage induced by high energy radiations and finally get died while normal cells repairing mechanism is intact and are able to repair the damage induce by these radiations.
So these radiations specifically targets cancer cells.
Gibhardt CS et. al, 2015, showed that challenging cells with ≥1 Gy X-rays or with UV-A laser micro-irradiation causes a rapid rise of H2O2 in the nucleus and in the cytosol which result in the production and glutathione-buffering, is sufficient for triggering a signaling cascade that involves an elevation of cytosolic Ca2+ and eventually an activation of hIK channels.
A course of radiation therapy is preceded by a simulation session in which low-energy beam are used to produce radiograghic images that indicate the exact beam location.
Radiation therapy is usually delivered in fractionated doses such as 180 to 300 cGy per day, five times a week for a total course of 5-8 weeks.
Success of radiotherapy depends in the difference in the radio sensitivity between the tumor and normal tissue.
Radiation therapy either used individually or in combination with chemotherapy.
Although radiation therapy is effective against many types of cancers this therapy is also associated with side effects, Fatigue and serious skin problems (dryness, itching, blistering, or peeling), GI toxicity, oropharyngeal mucositis& xerostomia, myelosuppression.
There is also possibility of developing a secondary cancer after radiation therapy.
"Radiation therapy either used individually or in combination with chemotherapy to treat breast cancer"
Chemotherapy: Chemotherapy is a treatment method for cancer with one or more chemically synthesized anti-cancer drugs which mainly targets rapidly dividing cancer cells and normal cells (epithelial cells of mouth, intestine lining).
Chemotherapy either given intravenously or orally in cycles.
Usually chemotherapy given after surgery or mastectomy to kill cancer cells which are left during surgery to reduce recurrence of tumor.
Alkylating agents, antimetabolites, antitumor antibiotic, plant alkaloids, hormonal agent, immunotherapy are the important agents of chemotherapy.
The most common chemo drugs used for early breast cancer include the anthracyclines such as doxorubicin and the taxanes ( paclitaxel and docetaxel).
These may be used in combination with certain other drugs, like fluorouracil (5-FU), cyclophosphamide, and carboplatin while for advance breast cancer docetaxel, paclitaxel, cisplatin, carboplatin, liposomal doxorubicin, mitoxantrone, vinorelbine etc.
Chemotherapy is associated with loss of appetite, hair loss, mouth sores, nausea and vomiting, low blood cell counts increases the chance of getting infection, irregular menstrual cycle and fatigue.
E.G, Myelosuppression, nausea &vomiting, Stomatitis and alopecia.
"Chemotherapy is associated with loss of appetite, hair loss, mouth sores, nausea and vomiting, low blood cell counts increases the chance of getting infection, irregular menstrual cycle and fatigue"
Hormone therapy or endocrine therapy: A gold standard therapy for ER positive breast cancers.
The main purpose of hormone therapy is to reduce the level of steroids (estrogens, androgens and progesterone) in the body or blocking the actions.
But hormone therapy does not work in ER negative breast cancer. Aromatase inhibitors, selective estrogen receptor modulators (SERMs) and selective estrogen receptor down regulators (SERDs) mainly use in hormone therapy.
"Hormone therapy or endocrine therapy is a gold standard therapy for ER positive breast cancers"
In alternate chemotherapy fallopian tube, ovary and adrenal glands surgically removed to reduce the hormone level in the body.
Tiredness, weight gain, hot flush, menopausal symtoms, digestive problems, headache, hair thinning, sweating, breast tenderness, memory problems and mood swing are associated side effects of the endocrine therapies.
Biological therapy or Targeted therapy: Targeted therapy is the use of drugs which specifically target the cancer by interfering the molecular targets of the cancer cells which have important role in growth and progression of cancer cells.
Most of the targeted drugs are cytostatic which block the cell proliferation.
Molecular targets are decided by comparing the molecular profiles of the normal and cancer cells and those proteins which are over expressed in cancer cells are selected as molecular target such as HER2/neu over expressed in ER negative breast cancer cases.
Trastuzumab, a monoclonal antibody against HER2/neu protein used to target the over expressed HER2/neu, effective therapy for the ER negative HER2/neu over expressing breast cancer cases.
"Most of the targeted drugs are cytostatic which block the breast cancer cell proliferation"
Another approach is to analyse the protein for the mutation which only found in cancer cells and specifically target the mutated proteins.
Target therapy includes hormone therapies, signal trasducer inhibitors (gefitinib), gene expression modulators, apoptosis inducer, angiogenesis inhibitors (bevacizumab), cancer vaccine, gene therapy and immunotherapies.
Toremifene, Trastuzumab, fulvestrant, exemestane, lapatinib , letrozole, pertuzumab, ado-trastuzumab emtansine, palbociclib, Everolimus, tamoxifen, anastrozole are the targeted drugs approved for the treatment of breast cancer.
Skin problems, high blood pressure, problem with blood clotting and wound healing are the associated side effects of the targeted therapies.
Bone Directed therapy: When breast cancer cells metastases to bone serious complications arises. Bisphosphonates (pamidronate, zoledronic acid) and denosumab are mainly used in such conditions. Biphosphonates mainly increases the bone strength and reduces bone thinning.
Osteonecrosis is serious side effect with the use of biphosphonates.
Denosumab another drugs for the bone directed therapy it is more effective than biphosphonates. Low blood calcium and phosphates are the side effects associated with Denosumab.
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Electrophoresis: an Overview, How to Run Gel...
Continue ReadingGel Electrophoresis Objective
To perform SDS-PAGE Gel Electrophoresis for separation of proteins.
About Gel Electrophoresis
Almost most of the analytical electrophoresis of proteins is carried out in polyacrylamide gels under conditions (generally SDS-PAGE) that ensure dissociation of the proteins into their individual polypeptide subunits.
Most common anionic detergent SDS (Sodium Dodecyl Sulfate) is used in combination with a reducing agent such as 2-mercaptoethanol or dithiothreitol (DTT) followed by heating to dissociate or break ionic and disulphide bonds of the proteins before they are loaded on the gel for Gel Electrophoresis.
The denatured proteins or polypeptide binds SDS and become negatively charged.
Because the amount of SDS bound is proportional to the molecular weight of the polypeptide or protein and is independent of its sequence, SDS-polypeptide complexes migrate through polyacrylamide gels in accordance with the size of polypeptide.
Using various markers of known molecular weight, it is therefore possible to estimate the molecular weight of the polypeptide chain(s).
Hence, SDS-PAGE Gel Electrophoresis is the most widely used method for qualitatively analyzing any protein mixtures.
The method is based on the separation of proteins according to their size and then locating by binding them to a dye.
Gel Electrophoresis is particularly used for monitoring protein purity and to determine their relative mass.
Adopted from BioRender
In, most cases SDS-PAGE Gel Electrophoresis is carried out in a discontinuous buffer system in which buffer in the reservoirs is of a various pH gradient and ionic strength from the buffer used to cast the polyacrylamide gel.
The Sodium Dodecyl Sulfate (SDS)-polypeptide complexes in the protein sample that is applied to the gel are swept along by a moving fence created when an electric current is passed between the electrodes.
After moving through a stacking gel of high porosity, the protein complexes are stacked on the surface of the resolving gel.
The ability of discontinuous buffer systems is to concentrate all of the complexes into a very small volume greatly increases the resolution of Sodium Dodecyl Sulfate (SDS)-polyacrylamide gels.
Mechanism of Gel Polymerization in Electrophoresis
The sample and the stacking gel contain Tris-Cl (pH 6.8), and upper top and lower buffer tank contain Tris-glycine (pH 8.3), and the resolving gel contains Tris-Cl (pH 8.8).
All components of the system contain 0.1% SDS.
The gels are composed of chains of polymerized acrylamide that are cross-linked by a bifunctional agent such as N,N’ -methylene bisacrylamide. The effective range of separation of Sodium Dodecyl Sulfate (SDS)-polyacrylamide gels (SDS-PAGE ) depends on the concentration of polyacrylamide used to prepare the gel and on the amount of cross-linking.
Polymerization of acrylamide in absence of cross-linking agents generates viscous solutions that are of no practical use.
Cross-liks formed by bisacrylamide add rigidity and tensile strength to the gel and form pores through which the SDS-polypeptide complexes must pass.
The size of these pores decreases with the increase of bisacrylamide:acrylamide ratio, reaching a minimum when the ratio is approximately 1:20.
Most gels are cast with molar ratios of bisacrylamide:acrylamide of 1:29.
The sieving properties of the gel are evaluated by the size of the pore which is determined by the absolute concentrations of acrylamide and bisacrylamide.
The following table shows the linear range of separation obtained with gel cast with concentrations of acrylamide that range from 5% to 15%:
Acrylamide Concentration and Protein Size in Electrophoresis
Acrylamide concentration (%) Linear range of separation (kD) 15 12-43 10 16-68 7.5 36-94 5.0 57-212 Component of Gel Electrophoresis
Separating / resolving gel:
The lower (separating, running or resolving) gel is prepared using 5-15% acrylamide which is more than that used in the stacking gel (the amount of acrylamide used depends upon the molecular weight of the macromolecule under separation).
Hence the pores are numerous and of a smaller diameter imparting the molecular sieving property to this gel.
It is in this gel that the macromolecules subsequently separate.
The separating gel constitutes about two-thirds of the length of the gel plates.
The buffer used in the running gel is Tris.Cl at pH 8.8.
Stacking gel:
The stacking gel is layered on top of the separating gel after it has polymerized completely.
It is prepared using 2-5% of acrylamide and is consequently highly porous and devoid of any molecular sieving action.
The stacking gel constitutes one-third of the length of the gel plates in which the comb is placed in order to form wells for sample loading.
The buffer used in preparation of stacking gel is Tris.Cl. at pH 6.8.
Gel Electrophoresis Process
Glycine in the upper buffer reservoir exists in two forms; as a zwitterion which does not have a net charge, and as glycinate ion which is negatively charged.
When the power is switched on, chloride, protein and glycinate anions begin to migrate towards the anode.
Upon entering the stacking gel, the glycinate ions encounter a condition of low pH (pH of stacking gel buffer is about 2 pH units lower than that of the upper reservoir) which shifts the pH towards formation of zwitter ion.
Since zwitter ions are devoid of charge, they are immobile.
This immobility of glycine zwitterions to migrate into stacking gel coupled with high mobility of the chloride ions creates high localized voltage gradient between the leading chloride ion and the trailing glycinate ions.
Since protiens have their mobility intermediate between the trailing and the leading ions, the proteins carry the current in this region and migrate rapidly in this high local electric field.
However, they cannot overtake the chloride ions since the strong local fields exist only between the chloride and the glycinate ions.
As a consequence the proteins migrate quickly until they reach the chloride rich region and then drastically slow down.
This two speed movement of the proteins results in a piling up of protein sample in a tight sharp disc between the glycinate and the chloride ions.
It is in this form that the protein enters the resolving gel.
The small pore size of the gel retards the protein band for a while which allows the glycinate ions to catch up.
As soon as the glycinate ions enter the resolving gel they encounter a higher pH range and thus resume their full charge upon diasappearance of the localized high voltage gradient.
Now the separation of the proteins takes place according to their molecular weight.
Note: The buffer in the upper and the lower reservoir is Tris-glycine at pH 8.3, whereas the buffer used for preparing the stacking gel is Tris-Cl at pH 6.8.
Gel Electrophoresis Requirements
1. Acrylamide and N,N’ -methylene bisacrylamide: Prepare a stock solution containing 29% (w/v) acrylamide and 1% (w/v) bisacrylamide in deionized warm, water (to assist the dissolution of bisacrylamide).
Note: Acrylamide and bisacrylamide are slowly converted to acrylic and bisacrylic acid upon storage.
The reaction is catalyzed by light and alkali.
Hence always check that the pH of the solution is 7.0 or less, and store it in dark bottles at room temperature.
The fresh solutions should be prepared every few months.
Caution: Both are neurotoxins.
Polyacrylamide is considered to be non-toxic but care is to be taken while handling it as it might contain some amount of unpolymerized material.
2. Sodium dodecyl sulphate (SDS): Prepare a 10% (w/v) stock solution in Deionized water and store at room temperature.
3. Tris buffers for the preparation of resolving and stacking gels: Prepare 1.5 M Tris Cl. (pH 8.8) for resolving gel and 1.0 M Tris.Cl (pH 6.8) for stacking gel.
4. TEMED (N,N,N’N’-tetramethylenediamine): TEMED accelerates the polymerization of acrylamide and bisacrylamide by catalyzing the formation of free redicals from ammonium persulfate.
5. Ammonium persulfate (APS): It provides free radicals for polymerization of acrylamide and bisacrylamide.
Ammonium persulfate decomposes slowly and fresh solution is to be prepared weekly.
Prepare a 10% stock solution in Deionized water and store at 4C.
6. Tris-glycine electrophoresis buffer: 25 mM Tris base, 250 mM glycine and 0.1% SDS.
Adjust pH 8.3.
Prepare a 5X stock by dissolving 15.1 g of Tris base and 94 g of glycine in 900 ml water.
Adjust pH 8.3.
Add 50 ml of 10% (w/v) stock solution of SDS.
Make up the volume to 1000 ml with water.
7. 2X SDS gel-loading buffer: 100 mM Tris Cl (pH 6.8), 10% β-mercaptoethanol, 4% SDS, 0.2% bromophenol blue, 20% glycerol.
Gel Electrophoresis Procedure
1. Clean the whole Gel Electrophoresis apparatus with methanol so as to ensure them to be clean and detergent free.
Assemble the glass plates.
Clamp the assembly to the gel casting apparatus.
Ensure the Gel Electrophoresis assembly is leak proof by filling methanol between the plates where the gel is to be casted.
2. Determine the volume of the gel mold.
3. Prepare the resolving gel as shown below:
For preparing 15 ml, 12% gel
Solution components Volume in ml Water 4.9 30% acrylamide mix 6.0 1.5M Tris.Cl (pH 8.8) 3.8 10% SDS 0.15 10% APS 0.15 TEMED .006 Add APS and TEMED in the last because polymerization will begin as soon as TEMED has been added.
4. Pour 13 ml of acrylamide solution in the gap between the glass plates (leave sufficient space for the stacking gel- the length of the teeth of the comb plus 1 cm).
Carefully overlay the gel with isobutanol to prevent oxygen from diffusing into the gel (oxygen inhibits polymerization).
5. After polymerization is complete (around 30-40 min), pour off the isobutanol and wash the top of the gel several times with water so as to remove the traces of unpolymerized material.
Remove the remaining water using a filter paper towel.
6. Prepare the stacking gel as follows:
Solution components Volume in ml Water 5.5 30% acrylamide mix 1.3 1.0M Tris.Cl (pH 6.8) 1.0 10% SDS 0.08 10% APS 0.08 TEMED .008 8. Pour the stacking gel solution directly on to the surface of the polymerized resolving gel.
Immediately insert a clean Teflon comb (cleaned with water and dried with ethanol).
Avoid trapping air bubbles.
Add more stacking gel solution from the sides of the comb to fill the spaces completely.
9. After polymerization is complete (30-40 min), remove the Teflon comb, clean the wells with water to remove any unpolymerized material.
Mount the gel in the electrophoretic apparatus.
For ease mark the wells.
Add tris-glycine electrophoretic buffer to the top and the bottom reservoir.
First add the buffer in the upper reservoir and check for leak.
10. Load upto 20 ul of each sample in a pre-determined order into the bottom of the wells with a Hamilton microliter syringe.
Load the samples in an order so that the gel can be cut in two exact replicas after run so as to use one part for staining and other for blotting.
11. Attach the electrophoretic apparatus to electric power supply.
Positive electrode should be connected to the lower reservoir.
Apply voltage of 8V/cm to the gel.
When the dye front has moved into the resolving gel increase the voltage to 15V/cm and run the gel until the dye reaches the bottom of the gel.
12. Turn off the power supply. Remove the gel plates from the apparatus.
Carefully remove the gel with the help of spacers.
Cut it into two and place one part in a reservoir to stain with coomassie brilliant blue so as to visualize the protein bands, and use the other part for western blotting.
Store the part to be used for western blotting in transfer buffer (39 mM glycine, 48 mM Tris base, 0.037% SDS and 20% methanol, pH 8.3.
Make up the volume with water) at 4C.
Gel Staining
Polypeptides separated by SDS-PAGE Gel Electrophoresis can be simultaneously fixed with methanol:glacial acetic acid and stained with coomassie brilliant blue R250.
The gel is immersed for several hours in a concentrated methanol and acetic acid solution of the dye and the excess dye is then allowed to diffuse from the gel during a prolonged period of destaining.
1. Dissolve 0.25 g of coomassie brilliant blue R250 (COBBR-250) in 90 ml of methanol: H2O (1:1 v/v) and 10ml of glacial acetic acid.
Filter the solution through a whatmann No. 1 filter to remove any particulate matter.
2. Immerse the gel in atleast 5 volumes of staining solution and place on a slowly rocking platform for a minimum of 4 h at room temperature.
3. After that, remove the stain and save it for future use.
Destain the gel by soaking it in methanol/acetic acid solution (step 1) without the dye on a slowly rocking platform for 4-8 h, changing the destaining solution three or 4 times.
4. The more thoroughly the gel is destained, the smaller the amount of protein that can be detected by staining with COBBR250.
5. After destaining the gels can be stored indefinitely in water containing 20% glycerol in a sealed plastic bag.
Gel Electrophoresis Citations:
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Thawing of Frozen Cell Lines: Cell Revival...
Continue ReadingAbout Thawing of Frozen Cell
Cells that are cryopreserved can be used again whenever required.
The procedure of bringing frozen cells to room temperature from a metabolically inactive state to an active state without causing much damage is called as thawing.
Principle of Thawing or Cell Revival
When cryopreserved cells are needed for study, they should be thawed rapidly and plated at high density to optimize recovery.
The ampoule should be thawed as rapidly as possible, to minimize intracellular ice crystal growth during the warming process.
This can be done in warm water, in a bucket or water bath.
The cell suspension should be diluted slowly after thawing as rapid dilution reduces viability.
Some mammalian cells (often suspension-growing cells) are more sensitive to cryoprotectants, particularly DMSO, and must be centrifuged after thawing but still need to be diluted slowly in medium first.
Thawing Requirements
Sterile:
1. Culture flask
2. Centrifuge tube (if centrifugation is required)
3. Growth medium
4. Pipettes, 1 ml, 10 ml
Non-sterile:
5. Protective gloves and face mask
6. Sterile water at 37°C, 10 cm deep in a clean, alcohol-swabbed bucket with lid
7. 70% IPA, cotton swabs.
Thawing Procedure
1. Bring the cryovial out of freezer in ice. Thaw.
2. Incubate in water bath at 37o C for 1-2 minutes.
3. Double-check the label to confirm the identity of the cells; then swab the vial thoroughly with 70% alcohol, and open it in a laminar-flow hood.
4. Transfer the contents of the vial to a 2ml microfuge tube and seal with parafilm.
5. Centrifuge at 1300 rpm for 10 minutes at room temperature.
6. Resuspend the pellet in 2 ml medium.
7. Recentrifuge at 1300 rpm for 10 minutes at room temperature to remove traces of DMSO. Pellet should be resuspended in 1 ml of medium.
8. Transfer the suspension to 25 cm2 flask. Add medium slowly to the cell suspension (5 ml). For cells in suspension culture also require centrifugation to remove the cryoprotectant.
9. Incubate cells in a humidified incubator with 5% CO2, at 37oC.
10. Check after 24 h.
Post Thawing Observations
For attached monolayer cells, confirm attachment and try to estimate percentage survival based on photographs of cells at the expected density (cells/cm2 ) with full survival.
For cells growing in suspension culture, check appearance (clear cytoplasm, lack of granularity and dilute to regular seeding concentration.
This can be made more precise if the cells are counted and an estimate of viability is made, in which case the cells can be diluted to the regular seeding concentration of viable cells.
Thawing Precaution
Dilution should be slow in case of DMSO because sudden dilutions can cause severe osmotic change and reduce the viability.
On removal from storage, extreme caution must be exercised to prevent explosion of the cryo vial because of sudden expansion of the trapped nitrogen.
To retain maximum viability during cryopreservation, cells must be cooled at a constant slow rate, -1 to -5 C/min.
To minimize contamination risk, all components should be pretested.
Resuspending cells for freezing in precooled freeze medium, 0-4 C, may improve their survival prior to freezing.
When this approach is used, it is important to maintain the cells at a constant temperature during all subsequent handling by placing ampoules in ice until they are frozen.
After thawing cells it is necessary to slowly dilute the cryoprotectant to prevent osmotic shock.
If the cryovial is stored in the liquid N2 it may expand and on thawing it may crack.
Thawing Citations:
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