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A month ago, I tried isolating cheek cells from saliva and the experiment was a success, however, I was wondering if it is possible to cultivate these cells in vitro. I'd say it's possible, but I wasn't able to find any publications about this topic online, even on PubMed (I found one, but it was outdated and the part with used media wasn't clear). Is it possible to use just DMEM media and if so, how long will the cells survive in the media, or are there any other alternatives?
Primary Cell Culture: 3 Techniques (With Diagram)
Primary culture broadly involves the culturing techniques carried following the isolation of the cells, but before the first subculture. Primary cultures are usually prepared from large tissue masses. Thus, these cultures may contain a variety of differentiated cells e.g. fibroblasts, lymphocytes, macrophages, epithelial cells.
With the experiences of the personnel working in tissue culture laboratories, the following criteria/ characteristics are considered for efficient development of primary cultures:
a. Embryonic tissues rather than adult tissues are preferred for primary cultures. This is due to the fact that the embryonic cells can be disaggregated easily and yield more viable cells, besides rapidly proliferating in vitro.
b. The quantity of cells used in the primary culture should be higher since their survival rate is substantially lower (when compared to subcultures).
c. The tissues should be processed with minimum damage to cells for use in primary culture. Further, the dead cells should be removed.
d. Selection of an appropriate medium (preferably a nutrient rich one) is advisable. For the addition of serum, fetal bovine source is preferred rather than calf or horse serum.
e. It is necessary to remove the enzymes used for disaggregation of cells by centrifugation.
Techniques for Primary Culture:
Among the various techniques devised for the primary culture of isolated tissues, three techniques are most commonly used:
1. Mechanical disaggregation.
2. Enzymatic disaggregation.
3. Primary explant technique.
An outline of these techniques is depicted in Fig. 36.1, and the procedures are briefly described:
Technique # 1. Mechanical Disaggregation:
For the disaggregation of soft tissues (e.g. spleen, brain, embryonic liver, soft tumors), mechanical technique is usually employed. This technique basically involves careful chopping or slicing of tissue into pieces and collection of spill out cells.
The cells can be collected by two ways:
i. Pressing the tissue pieces through a series of sieves with a gradual reduction in the mesh size.
ii. Forcing the tissue fragments through a syringe and needle.
Although mechanical disaggregation involves the risk of cell damage, the procedure is less expensive, quick and simple. This technique is particularly useful when the availability of the tissue is in plenty, and the efficiency of the yield is not very crucial. It must however, be noted that the viability of cells obtained from mechanical techniques is much lower than the enzymatic technique.
Technique # 2. Enzymatic Disaggregation:
Enzymatic disaggregation is mostly used when high recovery of cells is required from a tissue. Disaggregation of embryonic tissues is more efficient with higher yield of cells by use of enzymes. This is due to the presence of less fibrous connective tissue and extracellular matrix. Enzymatic disaggregation can be carried out by using trypsin, collagenase or some other enzymes.
Disaggregation by trypsin:
The term trypsinization is commonly used for disaggregation of tissues by the enzyme, trypsin.
Many workers prefer to use crude trypsin rather than pure trypsin for the following reasons:
i. The crude trypsin is more effective due to the presence of other proteases
ii. Cells can tolerate crude trypsin better.
iii. The residual activity of crude trypsin can be easily neutralized by the serum of the culture media (when serum-free media are used, a trypsin inhibitor can be used for neutralization).
Disaggregation of cells can also be carried out by using pure trypsin which is less toxic and more specific in its action. The desired tissue is chopped to 2-3 mm pieces and then subjected to disaggregation by trypsin. There are two techniques of trypsinization-warm trypsinization and cold trypsinization (Fig. 36.2).
Warm trypsinization (Fig. 36.2A):
This method is widely used for disaggregation of cells. The chopped tissue is washed with dissection basal salt solution (DBSS), and then transferred to a flask containing warm trypsin (37° C). The contents are stirred, and at an interval of every thirty minutes, the supernatant containing the dissociated cells can be collected. After removal of trypsin, the cells are dispersed in a suitable medium and preserved (by keeping the vial on ice).
The process of addition of fresh trypsin (to the tissue pieces), incubation and collection of dissociated cells (at 30 minutes intervals) is carried out for about 4 hours. The disaggregated cells are pooled, counted, appropriately diluted and then incubated.
Cold trypsinization (Fig. 36.2B):
This technique is more appropriately referred to as trypsinization with cold pre-exposure. The risk of damage to the cells by prolonged exposure to trypsin at 37°C (in warm trypsinization) can be minimized in this technique.
After chopping and washing, the tissue pieces are kept in a vial (on ice) and soaked with cold trypsin for about 6-24 hours. The trypsin is removed and discarded. However, the tissue pieces contain residual trypsin. These tissue pieces in a medium are incubated at 37°C for 20-30 minutes. The cells get dispersed by repeated pi-pettings. The dissociated cells can be counted, appropriately diluted and then used.
The cold trypsinization method usually results in a higher yield of viable cells with an improved survival of cells after 24 hours of incubation. This method does not involve stirring or centrifugation, and can be conveniently adopted in a laboratory. The major limitation of cold trypsinization is that it is not suitable for disaggregation of cells from large quantities of tissues.
Limitations of trypsin disaggregation:
Disaggregation by trypsin may damage some cells (e.g. epithelial cells) or it may be almost ineffective for certain tissues (e.g. fibrous connective tissue). Hence other enzymes are also in use for dissociation of cells.
Disaggregation by collagenase:
Collagen is the most abundant structural protein in higher animals. It is mainly present in the extracellular matrix of connective tissue and muscle. The enzyme collagenase (usually a crude one contaminated with non-specific proteases) can be effectively used for the disaggregation of several tissues (normal or malignant) that may be sensitive to trypsin.
Highly purified grades of collagenase have been tried, but they are less effective when compared to crude collagenase. The important stages in collagenase disaggregation, depicted in Fig. 36.3, are briefly described hereunder.
The desired tissue suspended in basal salt solution, containing antibiotics is chopped into pieces. These pieces are washed by settling, and then suspended in a complete medium containing collagenase. After incubating for 1-5 days, the tissue pieces are dispersed by pipetting. The clusters of cells are separated by settling. The epithelial cells and fibroblastic cells can be separated.
Collagenase disaggregation has been successfully used for human brain, lung and several other epithelial tissues, besides various human tumors, and other animal tissues. Addition of another enzyme hyaluronidase (acts on carbohydrate residues on cell surfaces) promotes disaggregation.
Collagenase in combination with hyaluronidase is found to be very effective for dissociating rat or rabbit liver. This can be done by per-fusing the whole organ in situ. Some workers use collagenase in conjunction with trypsin, a formulation developed in chick serum, for disaggregation of certain tissues.
Use of other enzymes in disaggregation:
Trypsin and collagenase are the most widely used enzymes for disaggregation. Certain bacterial proteases (e.g. pronase, dispase) have been used with limited success. Besides hyaluronidase, neuraminidase is also used in conjunction with collagenase for effective degradation of cell surface carbohydrates.
Technique # 3. Primary Explant Technique:
The primary explant technique was, in fact the original method, developed by Harrison in 1907. This technique has undergone several modifications, and is still in use. The simplified procedure adopted for primary explant culture is depicted in Fig. 36.4, and briefly described below.
The tissue in basal salt solution is finely chopped, and washed by settlings. The basal salt solution is then removed. The tissue pieces are spread evenly over the growth surface. After addition of appropriate medium, incubation is carried out for 3-5 days. Then the medium is changed at weekly intervals until a substantial outgrowth of cells is observed. Now, the explants are removed and transferred to a fresh culture vessel.
The primary explant technique is particularly useful for disaggregation of small quantities of tissues (e.g. skin biopsies). The other two techniques mechanical or enzymatic disaggregation however, are not suitable for small amounts of tissues, as there is a risk of losing the cells.
The limitation of explant technique is the poor adhesiveness of certain tissues to the growth surface, and the selection of cells in the outgrowth. It is however, observed that the primary explant technique can be used for a majority of embryonic cells e.g. fibroblasts, myoblasts, epithelial cells, glial cells.
Separation of Viable and Non-Viable Cells:
It is a common practice to remove the nonviable cells while the primary culture is prepared from the disaggregated cells. This is usually done when the first change of the medium is carried out. The very few left over non-viable cells get diluted and gradually disappear as the proliferation of viable cells commences.
Sometimes, the non-viable cells from the primary cultures may be removed by centrifugation. The cells are mixed with ficoll and sodium metrizoate, and centrifuged. The dead cells form a pellet at the bottom of the tube.
Medical Ethics and Safety Measures in Culture Techniques:
Since the culture techniques involve the use of animal or human tissues, it is absolutely necessary to follow several safety measures and medical ethics. In fact, in some countries there are established legislation/norms for selection and use of tissues in cultures. For example, in United Kingdom, Animal Experiments (Scientific Procedures) Act of 1986 is followed.
The handling of human tissues poses several problems that are not usually encountered with animal tissues. While dealing with fetal materials and human biopsies, the consent of the patient and/his or her relatives, besides the consent of local ethical committee is required. Further, taking any tissue (even in minute quantities) from human donors requires the full consent of the donor in a prescribed format.
The following issues need to be fully considered while dealing with human tissues:
1. The consent of the patient and/or relatives for using tissues for research purposes.
2. Ownership of the cell lines developed and their derivatives.
3. Consent for genetic modification of the cell lines.
6. Patent rights for any commercial use of cell lines.
In the general practice of culture techniques using human tissues, the donor and/or relatives are asked to sign a disclaimer statement (in a prescribed pro-forma) before the tissue is taken. By this approach, the legal complications are minimized.
Handling of human tissues is associated with a heavy risk of exposure for various infections. Therefore, it is absolutely necessary that the human materials are handled in a biohazard cabinet. The tissues should be screened for various infections such as hepatitis, tuberculosis, HIV, before their use. Further, the media and apparatus, after their use must be autoclaved or disinfected, so that the spread of infections is drastically reduced.
Useful Numbers for Cell Culture
There are various sizes of dishes and flasks used for cell culture. Some useful numbers such as surface area and volumes of dissociation solutions are given below for various size culture vessels.
|Catalog No.||Surface area (cm 2 )||Seeding density*||Cells at confluency*||Versene|
(mL of 0.05% EDTA). Approx. volume
(mL of 0.05% trypsin, 0.53 mM EDTA). Approx. volume
(mL). Approx. volume
|35mm||150460150318||8.8||0.3 x 10 6||1.2 x 10 6||1||1||2|
|60mm||150462150288||21.5||0.8 x 10 6||3.2 x 10 6||3||3||5|
|100mm||150464150350||56.7||2.2 x 10 6||8.8 x 10 6||5||5||12|
|150mm||150468168381||145||5.0 x 10 6||20.0 x 10 6||10||10||30|
|6-well||140675||9.6||0.3 x 10 6||1.2 x 10 6||1||1||1 to 3|
|12-well||150628||3.5||0.1 x 10 6||0.5 x 10 6||0.4 to 1||0.4 to 1||1 to 2|
|24-well||142475||1.9||0.05 x 10 6||0.24 x 10 6||0.2 to 0.3||0.2 to 0.3||0.5 to 1.0|
|48-well||150687||1.1||0.03 x 10 6||0.12 x 10 6||0.1 to 0.2||0.1 to 0.2||0.2 to 0.4|
|96-well||167008||0.32||0.01 10 6||0.04 x 10 6||0.05 to 0.1||0.05 to 0.1||0.1 to 0.2|
|T-25||156367156340||25||0.7 x 10 6||2.8 x 10 6||3||3||3–5|
|T-75||156499156472||75||2.1 x 10 6||8.4 x 10 6||5||5||8–15|
|T-175||159910159920||175||4.9 x 10 6||23.3 x 10 6||17||17||35–53|
|T-225||159934159933||225||6.3 x 10 6||30 x 10 6||22||22||45–68|
|*Seeding density is given for each culture vessel type as follows: Dishes and Flasks: Cells per vessel Culture plates: Cells per well.|
†The number of cells on a confluent plate, dish, or flask will vary with cell type. For this table, HeLa cells were used.
Because most cells are too small to be seen by the naked eye, the study of cells has depended heavily on the use of microscopes. Indeed, the very discovery of cells arose from the development of the microscope: Robert Hooke first coined the term ll” following his observations of a piece of cork with a simple light microscope in 1665 (Figure 1.23). Using a microscope that magnified objects up to about 300 times their actual size, Antony van Leeuwenhoek, in the 1670s, was able to observe a variety of different types of cells, including sperm, red blood cells, and bacteria. The proposal of the cell theory by Matthias Schleiden and Theodor Schwann in 1838 may be seen as the birth of contemporary cell biology. Microscopic studies of plant tissues by Schleiden and of animal tissues by Schwann led to the same conclusion: All organisms are composed of cells. Shortly thereafter, it was recognized that cells are not formed de novo but arise only from division of preexisting cells. Thus, the cell achieved its current recognition as the fundamental unit of all living organisms because of observations made with the light microscope.
The cellular structure of cork. A reproduction of Robert Hooke's drawing of a thin slice of cork examined with a light microscope. The lls” that Hooke observed were actually only the cell walls remaining from cells that had long since (more. )
The light microscope remains a basic tool of cell biologists, with technical improvements allowing the visualization of ever-increasing details of cell structure. Contemporary light microscopes are able to magnify objects up to about a thousand times. Since most cells are between 1 and 100 μm in diameter, they can be observed by light microscopy, as can some of the larger subcellular organelles, such as nuclei, chloroplasts, and mitochondria. However, the light microscope is not sufficiently powerful to reveal fine details of cell structure, for which resolution—the ability of a microscope to distinguish objects separated by small distances—is even more important than magnification. Images can be magnified as much as desired (for example, by projection onto a large screen), but such magnification does not increase the level of detail that can be observed.
The limit of resolution of the light microscope is approximately 0.2 μm two objects separated by less than this distance appear as a single image, rather than being distinguished from one another. This theoretical limitation of light microscopy is determined by two factors—the wavelength (λ) of visible light and the light-gathering power of the microscope lens (numerical aperture, NA)ording to the following equation:
The wavelength of visible light is 0.4 to 0.7 μm, so the value of λ is fixed at approximately 0.5 μm for the light microscope. The numerical aperture can be envisioned as the size of the cone of light that enters the microscope lens after passing through the specimen (Figure 1.24). It is given by the equation
Numerical aperture. Light is focused on the specimen by the condenser lens and then collected by the objective lens of the microscope. The numerical aperture is determined by the angle of the cone of light entering the objective lens (α) and by (more. )
The theoretical limit of resolution of the light microscope can therefore be calculated as follows:
Microscopes capable of achieving this level of resolution had been made already by the end of the nineteenth century further improvements in this aspect of light microscopy cannot be expected.
Several different types of light microscopy are routinely used to study various aspects of cell structure. The simplest is bright-field microscopy, in which light passes directly through the cell and the ability to distinguish different parts of the cell depends on contrast resulting from the absorption of visible light by cell components. In many cases, cells are stained with dyes that react with proteins or nucleic acids in order to enhance the contrast between different parts of the cell. Prior to staining, specimens are usually treated with fixatives (such as alcohol, acetic acid, or formaldehyde) to stabilize and preserve their structures. The examination of fixed and stained tissues by bright-field microscopy is the standard approach for the analysis of tissue specimens in histology laboratories (Figure 1.25). Such staining procedures kill the cells, however, and therefore are not suitable for many experiments in which the observation of living cells is desired.
Bright-field micrograph of stained tissue. Cross section of a hair follicle in human skin, stained with hematoxylin and eosin. (G. W. Willis/ Biological Photo Service.)
Without staining, the direct passage of light does not provide sufficient contrast to distinguish many parts of the cell, limiting the usefulness of bright-field microscopy. However, optical variations of the light microscope can be used to enhance the contrast between light waves passing through regions of the cell with different densities. The two most common methods for visualizing living cells are phase-contrast microscopy and differential interference-contrast microscopy (Figure 1.26). Both kinds of microscopy use optical systems that convert variations in density or thickness between different parts of the cell to differences in contrast that can be seen in the final image. In bright-field microscopy, transparent structures (such as the nucleus) have little contrast because they absorb light poorly. However, light is slowed down as it passes through these structures so that its phase is altered compared to light that has passed through the surrounding cytoplasm. Phase-contrast and differential interference-contrast microscopy convert these differences in phase to differences in contrast, thereby yielding improved images of live, unstained cells.
Microscopic observation of living cells. Photomicrographs of human cheek cells obtained with (A) bright-field, (B) phase-contrast, and (C) differential interference-contrast microscopy. (Courtesy of Mort Abramowitz, Olympus America, Inc.)
The power of the light microscope has been considerably expanded by the use of video cameras and computers for image analysis and processing. Such electronic image-processing systems can substantially enhance the contrast of images obtained with the light microscope, allowing the visualization of small objects that otherwise could not be detected. For example, video-enhanced differential interference-contrast microscopy has allowed visualization of the movement of organelles along microtubules, which are cytoskeletal protein filaments with a diameter of only 0.025 μm (Figure 1.27). However, this enhancement does not overcome the theoretical limit of resolution of the light microscope, approximately 0.2 μm. Thus, although video enhancement allows the visualization of microtubules, the microtubules appear as blurred images at least 0.2 μm in diameter and an individual microtubule cannot be distinguished from a bundle of adjacent structures.
Video-enhanced differential interference-contrast microscopy. Electronic image processing allows the visualization of single microtubules. (Courtesy of E. D. Salmon, University of North Carolina, Chapel Hill.)
Light microscopy has been brought to the level of molecular analysis by methods for labeling specific molecules so that they can be visualized within cells. Specific genes or RNA transcripts can be detected by hybridization with nucleic acid probes of complementary sequence, and proteins can be detected using appropriate antibodies (see Chapter 3). Both nucleic acid probes and antibodies can be labeled with a variety of tags that allow their visualization in the light microscope, making it possible to determine the location of specific molecules within individual cells.
Fluorescence microscopy is a widely used and very sensitive method for studying the intracellular distribution of molecules (Figure 1.28). A fluorescent dye is used to label the molecule of interest within either fixed or living cells. The fluorescent dye is a molecule that absorbs light at one wavelength and emits light at a second wavelength. This fluorescence is detected by illuminating the specimen with a wavelength of light that excites the fluorescent dye and then using appropriate filters to detect the specific wavelength of light that the dye emits. Fluorescence microscopy can be used to study a variety of molecules within cells. One frequent application is to label antibodies directed against a specific protein with fluorescent dyes, so that the intracellular distribution of the protein can be determined. Proteins in living cells can be visualized by using the green fluorescent protein (GFP) of jellyfish as a fluorescent label. GFP can be fused to a wide range of proteins using standard methods of recombinant DNA, and the GFP-tagged protein can then be introduced into cells and detected by fluorescence microscopy.
Fluorescence microscopy. (A) Light passes through an excitation filter to select light of the wavelength (e.g., blue) that excites the fluorescent dye. A dichroic mirror then deflects the excitation light down to the specimen. The fluorescent light emitted (more. )
Confocal microscopy combines fluorescence microscopy with electronic image analysis to obtain three-dimensional images. A small point of light, usually supplied by a laser, is focused on the specimen at a particular depth. The emitted fluorescent light is then collected using a detector, such as a video camera. Before the emitted light reaches the detector, however, it must pass through a pinhole aperture (called a confocal aperture) placed at precisely the point where light emitted from the chosen depth of the specimen comes to a focus (Figure 1.29). Consequently, only light emitted from the plane of focus is able to reach the detector. Scanning across the specimen generates a two-dimensional image of the plane of focus, a much sharper image than that obtained with standard fluorescence microscopy (Figure 1.30). Moreover, a series of images obtained at different depths can be used to reconstruct a three-dimensional image of the sample.
Confocal microscopy. A pinpoint of light is focused on the specimen at a particular depth, and emitted fluorescent light is collected by a detector. Before reaching the detector, the fluorescent light emitted by the specimen must pass through a confocal (more. )
Confocal micrograph of mouse embryo cells. Nuclei are stained red and actin filaments underlying the plasma membrane are stained green. (Courtesy of David Albertini, Tufts University School of Medicine.)
Two-photon excitation microscopy is an alternative to confocal microscopy that can be applied to living cells. The specimen is illuminated with a wavelength of light such that excitation of the fluorescent dye requires the simultaneous absorption of two photons (Figure 1.31). The probability of two photons simultaneously exciting the fluorescent dye is only significant at the point in the specimen upon which the input laser beam is focused, so fluorescence is only emitted from the plane of focus of the input light. This highly localized excitation automatically provides three-dimensional resolution, without the need for passing the emitted light through a pinhole aperture, as in confocal microscopy. Moreover, the localization of excitation minimizes damage to the specimen, allowing three-dimensional imaging of living cells.
Two-photon excitation microscopy. Simultaneous absorption of two photons is required to excite the fluorescent dye. This only occurs at the point in the specimen upon which the input light is focused, so fluorescent light is only emitted from the chosen (more. )
Human Cardiac Microvascular Endothelial Cells (HCMEC)
Primary Human Cardiac Microvascular Endothelial Cells isolated from heart ventricles from a single donor.
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There was no family history of breast cancer.
The cells are poorly differentiated.
The cells are negative for expression of Her2-neu and for expression of p53.
HCC1806 is positive for the epithelial cell specific marker Epithelial Glycoprotein 2 (EGP2) and for cytokeratin 19.
The cells are homozygous for deletions in the FHIT gene at 3p14.2.
- Check all containers for leakage or breakage.
- Remove the frozen cells from the dry ice packaging and immediately place the cells at a temperature below -130°C, preferably in liquid nitrogen vapor, until ready for use.
Handling procedure To insure the highest level of viability, thaw the vial and initiate the culture as soon as possible upon receipt. If upon arrival, continued storage of the frozen culture is necessary, it should be stored in liquid nitrogen vapor phase and not at -70°C. Storage at -70°C will result in loss of viability.
- Thaw the vial by gentle agitation in a 37°C water bath. To reduce the possibility of contamination, keep the O-ring and cap out of the water. Thawing should be rapid (approximately 2 minutes).
- Remove the vial from the water bath as soon as the contents are thawed, and decontaminate by dipping in or spraying with 70% ethanol. All of the operations from this point on should be carried out under strict aseptic conditions.
- Transfer the vial contents to a 75 cm 2 tissue culture flask and dilute with the recommended complete culture medium (see the specific batch information for the recommended dilution ratio). It is important to avoid excessive alkalinity of the medium during recovery of the cells. It is suggested that, prior to the addition of the vial contents, the culture vessel containing the complete growth medium be placed into the incubator for at least 15 minutes to allow the medium to reach its normal pH (7.0 to 7.6).
- Incubate the culture at 37°C in a suitable incubator. A 5% CO2 in air atmosphere is recommended if using the medium described on this product sheet.
Note: If it is desired that the cryoprotective agent be removed immediately, or that a more concentrated cell suspension be obtained, centrifuge the cell suspension at approximately 125 x g for 5 to 10 minutes. Discard the supernatant and resuspend the cells with fresh growth medium at the dilution ratio recommended in the specific batch information.
Subculturing procedure Volumes used in this protocol are for 75 cm 2 flask proportionally reduce or increase amount of dissociation medium for culture vessels of other sizes.
- Remove and discard culture medium.
- Briefly rinse the cell layer with 0.25% (w/v) Trypsin-0.53 mM EDTA solution to remove all traces of serum that contains trypsin inhibitor.
- Add 2.0 to 3.0 mL of Trypsin-EDTA solution to flask and observe cells under an inverted microscope until cell layer is dispersed (usually within 5 to 15 minutes).
Quality control specifications
The product is provided 'AS IS' and the viability of ATCC ® products is warranted for 30 days from the date of shipment, provided that the customer has stored and handled the product according to the information included on the product information sheet, website, and Certificate of Analysis. For living cultures, ATCC lists the media formulation and reagents that have been found to be effective for the product. While other unspecified media and reagents may also produce satisfactory results, a change in the ATCC and/or depositor-recommended protocols may affect the recovery, growth, and/or function of the product. If an alternative medium formulation or reagent is used, the ATCC warranty for viability is no longer valid. Except as expressly set forth herein, no other warranties of any kind are provided, express or implied, including, but not limited to, any implied warranties of merchantability, fitness for a particular purpose, manufacture according to cGMP standards, typicality, safety, accuracy, and/or noninfringement.
This product is intended for laboratory research use only. It is not intended for any animal or human therapeutic use, any human or animal consumption, or any diagnostic use. Any proposed commercial use is prohibited without a license from ATCC.
While ATCC uses reasonable efforts to include accurate and up-to-date information on this product sheet, ATCC makes no warranties or representations as to its accuracy. Citations from scientific literature and patents are provided for informational purposes only. ATCC does not warrant that such information has been confirmed to be accurate or complete and the customer bears the sole responsibility of confirming the accuracy and completeness of any such information.
This product is sent on the condition that the customer is responsible for and assumes all risk and responsibility in connection with the receipt, handling, storage, disposal, and use of the ATCC product including without limitation taking all appropriate safety and handling precautions to minimize health or environmental risk. As a condition of receiving the material, the customer agrees that any activity undertaken with the ATCC product and any progeny or modifications will be conducted in compliance with all applicable laws, regulations, and guidelines. This product is provided 'AS IS' with no representations or warranties whatsoever except as expressly set forth herein and in no event shall ATCC, its parents, subsidiaries, directors, officers, agents, employees, assigns, successors, and affiliates be liable for indirect, special, incidental, or consequential damages of any kind in connection with or arising out of the customer's use of the product. While reasonable effort is made to ensure authenticity and reliability of materials on deposit, ATCC is not liable for damages arising from the misidentification or misrepresentation of such materials.
Please see the material transfer agreement (MTA) for further details regarding the use of this product. The MTA is available at www.atcc.org.
Co-culture Experiments with Lonza's Small Airway Epithelial Cells and SAGM TM BulletKit TM Media
Data Courtesy of Furukawa, et al. Lonza's SAEC cells were co-cultured with primary normal mammary epithelial cells, breast cancer cell lines MCF-7 and MDA-MB-231 in Lonza's SAGM TM BulletKit TM Growth Media. Breast cancer cell lines showed differences in morphology and proliferation rates in the presence of small airway epithelial cells. The image above shows phase contrast imaging with a fluorescent (red) overlay of MCF-7 cell line. MCF-7 cell lines survived as tight clusters in SAGM TM Media. When co-cultured with Lonza's primary SAECs, MCF-7 cells demonstrated enhanced growth all the while staying clustered.
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The culture media used for cell cultures are generally quite complex, and culture condition widely varies for each cell type. However, media generally include amino acids, vitamins, salts (maintain osmotic pressure), glucose, a bicarbonate buffer system (maintains a pH between 7.2 and 7.4), growth factors, hormones, O2 and CO2. To obtain best growth, addition of a small amount of blood serum is usually necessary, and several antibiotics, like penicillin and streptomycin are added to prevent bacterial contamination.
Temperature varies on the type of host cell. Most mammalian cells are maintained at 37 o C for optimal growth, while cells derived from cold-blooded animals tolerate a wider temperature range (i.e. 15 o C to 26 o C). Actively growing cells of log phage should be used which divide rapidly during culture.
Process to obtain primary cell culture
Primary cell cultures are prepared from fresh tissues. Pieces of tissues from the organ are removed aseptically which are usually minced with a sharp sterile razor and dissociated by proteolytic enzymes (such as trypsin) that break apart the intercellular cement. The obtained cell suspension is then washed with a physiological buffer (to remove the proteolytic enzymes used). The cell suspension is spread out on the bottom of a flat surface, such as a bottle or a Petri dish. This thin layer of cells adhering to the glass or plastic dish is overlaid with a suitable culture medium and is incubated at a suitable temperature.
Bacterial infections, like Mycoplasma and fungal infections, commonly occur in cell culture creating a problem to identify and eliminate. Thus, all cell culture work is done in a sterile environment with proper aseptic techniques. Work should be done in laminar flow with the constant unidirectional flow of HEPA filtered air over the work area. All the material, solutions and the whole atmosphere should be of contamination-free.
If a surplus of cells is available from sub-culturing, they should be treated with the appropriate protective agent (e.g., DMSO or glycerol) and stored at temperatures below –130°C until they are needed. This stores cell stocks and prevents original cell from being lost due to unexpected equipment failure or biological contaminations. It also prevents finite cells from reaching senescense and minimizes risks of changes in long term cultures.
When thawing the cells, the frozen tube of cells is warmed quickly in warm water, rinsed with medium and serum and then added into culture containers once suspended in the appropriate media.
What Is the Function of the Cheek Cell?
A cheek cell, an epithelial cell found in the tissue on the inside lining of the mouth, continually secretes mucus to maintains a moist environment in the mouth. Together with salivary glands that secrete saliva, the cheek cells supply enough moisture in the mouth for enzymes to thrive. This moisture softens food, assists in swallowing and starts digestion.
The epithelial cells in the lining of the mouth are referred to as basal mucosa and divide roughly every 24 hours. They can easily be obtained through a simple swab or a mouth rinse. The cheek cell is very simple, but it contains the entire genetic makeup of the person's body. For this reason, cheek cells are frequently used to establish paternity and other investigations involving DNA. More recently, researchers have discovered that the cheek cell can be used to measure a person's likelihood of having high blood pressure.
A human cheek cell is thin, flat and irregularly shaped and has a large nucleus that contains the DNA. Its plasma membrane helps the cell to maintain suitable temperature while giving it its shape. Since it is selectively permeable, it only allows certain molecules into and out of the cell. Its cytoplasm contains water that dissolves nutrients and enzymes.
Basic equipment and reagents required for cell culture
To conduct research requiring cell culture work and to perform fundamental cell culture protocols, there are several key pieces of equipment and some basic reagents that are required, summarized in Tables 1 and 2.
Table 1: Basic equipment required for cell culture. 3 , 4 , 5 The images were created with BioRender.com.
Table 2: The basic reagents needed for cell culture. 2 , 5
See section below on media for your cells.
Phosphate-buffered saline (PBS) used for washing cells.
An enzyme used to detach adherent cells from culture vessels for culturing, such as trypsin.
An agent that reduces the freezing point of media and slows the cooling rate to reduce the risk of ice crystal formation which can damage cells and cause cell death. Dimethylsulfoxide (DMSO) is most commonly used.
It is also important to have access to deionized and distilled water as well as ice. The nature of the experiments to be performed will inform the reagents that will need to be acquired.
1) Tissue Procurement
Obtain nasal tissues and tracheo-bronchiolar segments from normal, CF, and disease control humans as sources of airway epithelial cells (hAE). The MLI TPC then utilizes the harvested hAE cells for culture, for ex vivo experiments, for RNA and proteins representative of in vivo conditions, and for in situ hybridization and immunohistochemistry. As a service to the greater CF research community (Function 6), the MLI TPC accepts tissues from other institutions through longstanding sources including local and distant outpatient surgical centers such as the UNC Lung Transplant Program, Carolina Donor Services, International Institute for the Advancement of Medicine (IIAM, Edison, NJ), and the National Disease Research Interchange (NDRI, Philadelphia, PA). The MLI TPC also accepts tissues from highly motivated donors nationally and internationally. Ongoing sources of CF tissues from independently recruited lung transplant programs include Loyola University, Maywood IL, Vanderbilt University, Medical College of Wisconsin, Israel, and Duke University Medical Center. Furthermore, in collaboration with the Cystic Fibrosis Foundation Therapeutics (CFFT) and NDRI, explanted CF lungs from multiple lung transplant centers are sent to UNC for processing.
2) Airway Epithelial Cell Isolation and Culture
a) Isolate epithelial cells from excised human nasal tissues, including endosinus mucosa, and cartilaginous bronchi of human lungs for distribution to investigators.
b) Prepare and maintain primary epithelial cell cultures, such as primary alveolar epithelial cells, microvascular endothelial cells and/or fibroblasts from the tissues noted above. The Core provides support for the preparation of well-differentiated airway epithelial cultures from passaged, or cryopreserved and thawed human cells, on permeable substrates.
c) Optimize and standardize epithelial culture conditions to replicate the gene expression, morphological differentiation, and physiologic functions of airways.
The MLI TPC also provides passage 1 airway epithelial cells, like primary cells, they too are suitable for producing well-differentiated cultures on porous supports. Representative sections of well-differentiated nasal and bronchial airway epithelial cell cultures are shown in Figure 1.
Figure 1. Representative photomicrographs of well-differentiated passage 1 nasal (A) and bronchial (B) human airway epithelial cells grown at an air-liquid interface for 14-21 days. OsO4 fixation, epon embedding and Richardsons stain, both panels at the same magnification, original magnification=1000x.
Routine MLI TPC procedures have significantly improved our ability to study differentiation-dependent functions (Figure 2) and have also increased the number and area of well-differentiated cultures produced from each sample. Possessing the ability to store primary cells at differing stages of maturity from the same patient sample allows repeat experiments with the same specimen. Additionally, we can perform simultaneous experiments with replicate cultures derived from multiple patients.
Figure 2. Demonstration of rotational mucus transport in well-differentiated cultures. A) 1 mm fluorescent microspheres were added to the culture surface and were propelled by the coordinated beating of cilia. A 5-second time-lapse exposure is shown. B) Linear velocities of the particles were measured and plotted as a function of distance from the center of rotation.
The in-house production of BEGM and ALI media has also resulted in significant cost savings and a deeper understanding of cell culture among staff. We are focused on producing cultures that accurately mirror in vivo gene expression, morphology, and physiology and which reproduce known differences between CF and non-CF individuals in vivo. Supplying large numbers of cultures, and large batches of media, in response to changing investigator needs is one of the major MLI TPC functions.
3) Collect Airway Surface Liquid (ASL) from in vivo and in vitro Samples
Critical testing of current advances in CF pathogenesis requires direct measurement of the chemical composition of human ASL. We have consistently collected and distributed in vivo airway mucus secretions for the study of biochemical composition, biophysical properties and as the source of the supernatant of mucopurulent material (SMM) from CF lungs. Additionally, ASL from well-differentiated cultures is used to stimulate air-liquid interface (ALI) cultures and serves as the basis for key experiments related to novel concepts in CF. The MLI TPC provides luminal contents of CF lungs explanted during transplant or removed at autopsy and harvests ASL from in vitro sources.
4) Genetic Manipulation of Cell CulturesFigure 3. Lentiviral vector genetic manipulation of hTBE cells. A) hTBE cells were sham-transduced (control) or infected with lentiviral vectors expressing a fused GFP/Blasticidin resistance protein (GFP/BSD) or red fluorescent protein linked to GFP/BSD by an internal ribosome entry site (RFP IRES), conventional epifluorescence microscopy 5 days following infection. B) RFP IRES-infected cells were trypsinized from plastic dishes, passaged to air-liquid interface cultures and allowed to differentiate for 35 days, X-Z plane confocal image. Note that the GFP/BSD fusion protein is apparently excluded from the nucleus while the RFP distributes equally. Basal cell expression is apparently lower from the CMV promoter, and alternative promoters enabling uniform expression are being tested.
Use adenovirus, adeno-associated virus, lentivirus (see Figure 3) and retrovirus vectors in combination with chemical treatment as necessary, to assist in the production of genetically manipulated, well-differentiated primary airway epithelial cell cultures.
5) Create and Characterize Novel Cell Lines
Use recent advances in cell immortalization technology to create new cell lines and characterize their biochemical properties, morphology and electrophysiologic function. The goal of the TPC is to create new and better immortalized cells retaining a greater capacity to recapitulate normal airway epithelial function. We have used retroviral transduction to serially introduce SV40 ER or HPV E6/E7 and hTERT into non-CF and CF cells. Their growth potential, morphology, biochemical features and electrophysiologic properties are studied using typical methods. Our ongoing observations have revealed that cells immortalized by SV40 ER or HPV E6/E7 and hTERT, when grown at an air-liquid-interface, are not optimal for CF-related investigation. This has led to novel studies using Bmi-1 oncogene to create superior cell lines (AJP 2009). Six novel cell lines using the Bmi-1 oncogene (3 normal and 3 deltaF508 homozygous CF) have been characterized (see Figure 4).
6) Translation of Technology and Reagents to the Greater CF Research Community
Our Core strongly observes the philosophy of developing non-proprietary technology and supporting its widespread translation. By providing material and consultative support, the MLI TPC allows others to develop or improve upon their own in-house cell culture capabilities. This involves the shipment of cells or media, hosting visitors for training, and publication of methods specifically related to improvements in human airway cell culture. In addition to hosting periodic training sessions for visitors from the US, England, Ireland, Switzerland, Canada, France, and Israel for the purpose of technology transfer, the MLI TPC consults with distant users as they encounter experimental difficulties related to their cell model work. The MLI TPC’s standing as a recharge center has allowed for a more accessible and efficient user experience. We are able to provide immortal cell lines at minimal expense, custom cell line creation for those who have sent us tissue specimens using CRC technology, and offer customized media based on manager approval.
Some of our latest updated methods have been published as a part of the Methods in Molecular Biology (MIMB) series, with the title “Epithelial Cell Culture Protocols.” The MLI TPC strives to meet the needs of academic, UNC, and commercial interests, when possible. Consultative support including advice, protocols, and more can be found on this site. If there are any requests outside of the information provided, feel free to contact the Laboratory Manager with the link provided at the top of the page.
7) New Initiatives Based on Investigator Needs
Figure 4. Robotic two plate TECC24 device that will facilitate quality control and experimental electrophysiologic testing. This device will increase our capacity to test cells from the current six, to 36 specimens per day (in quadruplicate wells). This piece of equipment is integral in improving the efficiency of our quality control testing process. The device also requires fewer cells from each specimen for analysis, which is ideal for the conservation of our resources.
Representative Ussing chamber results from UNCN1-3T (normal) and UNCCF1-3T (CF) cell lines. Note the absence of a forskolin (Fsk) response in the 4 replicate CF cell tracings, Amil = amiloride.
The TPC is responsive to the changing needs of UNC MLI investigators and provides material support for phasic motion and cyclic compressive stress studies, chambers for exposure to hypoxic conditions, mouse airway epithelial cell cultures, human lung vascular endothelial cell cultures and human alveolar epithelial cell cultures. The TPC has applied new methods enabling routine provision of well-differentiated tracheal epithelial cell cultures from normal mice as well as cultures from genetically manipulated mice, including CF mice, mice over-expressing the epithlial Na + channel, Coxsackie adenovirus receptor expressing mice, caveolin knockout mice and other genetically unique strains.