Cell culture and cell analysis (2024)

Introduction

In vitro cell culture is a method used for studying the behavior of animal cells in a controlled environment, free of systemic variations. Currently, different types of cell cultures have been adapted and developed. Animal cell cultures have been applied for studying basic cell biology, interactions of drugs and other chemicals with cells, production of vaccines and proteins, etc. This chapter covers a brief summary of the main features, types, requirements and applications of cell culture methodology. Also, we describe applications and principles for cell separation techniques, ranging from basic to more advanced applications.

Development of cell culture

Cell culture was developed in the early twentieth century as a method to study the behavior of animal cells in an environment free of the systemic changes that can be found in an animal during the normal homeostasis and stress of an experiment. The first model chosen for cell culture were amphibian cells, presumably for being exothermic animals and given that their cells would not require successive incubations. Later, medical science breakthroughs led to an interest in endothermic animals, where the normal and pathological development is similar to that of humans. The advent of mouse strains genetically pure brought mammals to the research laboratories. While the embryos provided a wide range of cell types in primary culture, mouse models had the advantage of continuous cell lines and a substantial tumor cell repertoire.

The progression of cell culture as a modern and sophisticated technique is based mostly on the needs of two major medical branches: virology and oncology. Cell culture has also been welcomed in many medicine and manufacturing routine applications. Cytogenetically analysis of amniocentesis derived-cells has the ability to disclose genetic disorders in the fetus. Likewise, viral infections can be evaluated quantitatively and qualitatively in host-cultured cells. Toxic effects of potential pharmaceutical compounds and contaminants can be evaluated also by using cell cultures.

Cell culture applications

The main uses of cell culture systems include:

Experimental model systems in basic and medical sciences. Cell culture offers certain advantages over the environmental and biological variability of other models. In addition, the use of genetically defined and characterized cell lines can simplify the analysis of experimental data. On the other hand, results obtained with specific cellular systems may not be representative of a wide range of other types of cells.

Study of physiological requirements for certain cell types. These include studies on positive effects of growth factors, growth-promoting substances, negative effects of cytotoxic compounds or xenobiotics and events related to programmed cell death (apoptosis), as well as cell proliferation, cell activation, cell signaling or any other cellular process.

Studies of cell development and differentiation. these include aspects of cell cycle and gene expression.

Pathological studies. these are enclosed in the characterization of cells using karyotyping to determine their genetic status.

Genetic Manipulation. cell culture techniques have played an essential role in the development of molecular biology, through the development of methods such as transfection.

Biotechnology. based on the manufacturing and industrial production of therapeutic proteins, vaccines and monoclonal antibodies.

Animal cell culture systems

Cellular systems can be established from whole organisms (e.g., chicken embryo), discrete organs (e.g., mouse liver), or blood cells (e.g., lymphocytes). While, in theory, it is possible to grow nucleated cells from any source, in practice, the highest probability of success is achieved with active young cells. The main considerations for the development of cell cultures are:

Biosafety. It is important to be aware of the potential infection risks when culturing animal cells. Although animal cells have a reduced risk of disease transmission compared to human cells, cultures should always be handled as a potential source of pathogenic microorganisms.

Using primary culture or continuous cell lines. freshly isolated cells more easily reflect the biochemical dynamics of the cells in vivo, although having a limited life span. Continuous cell lines are easy to use and offer the advantage of a priori knowledge of their specific growth requirements.

Culture media requirements. These include the provision of inorganic ions (such as a balanced salt solution), a carbon source, organic nutrients and other supplements that include antimicrobial agents to counteract the risk of contamination. For growth support, usually a basal medium supplemented with serum (e.g., fetal bovine serum) or a hom*ologous serum supplement with a set mixture composition of proteins, polypeptides, hormones, lipids, and trace elements. Levels of CO₂ and O₂ should be considered; many cultures are buffered with bicarbonate and should be kept in a CO₂ rich atmosphere.

Advantages and disadvantages of cell culture

Clearly, the study of cellular activity in vitro has several advantages and disadvantages. The main advantage is the consistency and reproducibility of results that can be obtained from using a batch of clonal cells. Cell cultures have a highly control of the physicochemical environment (i.e., pH, temperature, osmotic pressure, oxygen, and carbon dioxide tension) which can be controlled very accurately, and the control of physiological conditions, which can be constantly examined.

The disadvantages of cell culture are: highly skilled personnel, techniques must be performed using strict asepsis techniques because animal cells grow slower than many of the common contaminants (e.g., bacteria, viruses and fungi). Additionally, animal cells may not survive when isolated and therefore are not capable of an independent sustainable existence without providing a complex environment. One of the main limitations of cell culture is the expense and effort that has to be applied to obtain a relatively low amount of cells.

In addition, tissue composition is variable and heterogeneous. Replicas from the same sample have various constituents. To replicate an experimental result, cell lines must be manipulated many times in serial passages. For instance, every culture is going to be different from the original and less uniform in its constitution. In order to resolve this issue, the replicas are randomly mixed in each passage and the selective pressure of growing conditions tends to produce an optimal prevalent phenotype.

Main types of cell culture

Cell culture is a collection of techniques and resources in which cells that were part of an organism are growth in an artificial controlled environment. Usually, tissue must be previously treated to disrupt it by mechanical or enzymatic processes depending on the origin of the tissue and the purpose of the cell culture. However, some cells can be culture without this treatment such as in the case of liquid samples.

During tissue culture, tissue characteristics and architecture must be retained, at least partially. For instance, growth is slow and restricted to the use of embryonic tissue or 3D cell culture procedures. The 3D cell culture mimics the microenvironment, architectural design and functionality of a normal tissue under control conditions. In some cases, cell migration is observed through the solid phase when a piece of tissue is placed on a solid liquid-interface. Thus generating primary explants that grow outside the original tissue

The Primary Culture is the first culture that grows successfully after the cell isolation from a tissue. As mentioned above, cell should be subculture in a series of passages in order to keep the best condition for cell growth. As result, these cell lines go into senescence after the thirtieth division cycle. For this reason, cells are storage in a cell bank system to maintain them for long periods of time. In some cases, the cells can be immortalized, e.g., B cell lymphocyte can be immortalized with the Epstein- Barr virus to confer them the ability to proliferate indefinitely. These transformed cells have the advantage of unlimited availability, but they have the disadvantage of losing its initial characteristics. Figure 1 shows the main ways to start cell cultures (for further information please see recommended readings).

Figure 1

Main type of sources for cell cultures. Adapted from (1).

Preliminary preparation and sampling

Cell samples are obtained and prepared from organs, tissues and biological fluids depending on the target cell type, type of genetic material and type of analysis to be performed (Table 1). In all cases one must follow processes to ensure the quality of the sample. Generally, the preparation is performed after obtaining the sample, however, if this is not possible, conditions should be provided to ensure that the sample does not undergo degradation.

Table 1

Some types of cell cultures and their applications.

The success of each technique depends on the proper acquisition, processing and preservation of the sample.

Considerations for blood samples

Blood-drawing is a critical step for the experimental design. There are many factors that influence the conditions of blood components such as nutritional status, hormone levels and the circadian rhythms. If an experiment or project depends on the analysis and comparison of multiple parameters in blood samples collected from different individuals, it must be ensured that blood drawing should be carried out under the same conditions. For example, nutritional state and circulating hormones and cytokines may have important effect on the performance and behavior of the cells. Thus, in fasting state, the leptin levels in the blood are diminished and as a result, there are differences in T cell processes such as differentiation and increasing IFNγ secretion and decrease IL-4 production. In addition, differences in leptin levels have been shown to interfere with the T-regulatory cells proliferation.

Cell separation methods

To obtain tissue samples suitable for laboratory analysis, several procedures like separation, fractionation and characterization are usually performed and they are applied based on the properties of each cell type in the sample. The choice of each protocol depends on the desired degree of separation, preservation of viability, and technical analyses that would be studied.

Cell separation techniques have the advantage to allow high yield and recovery in a shorter time. Some examples are separation by density sedimentation and/or flow cytometry. In general, high performance techniques used for cell separations rely on differences like: 1. Cell size, 2. Cell density (specific gravity), 3. Cell load, 4. Cell surface chemistry, 5. cellular complexity and 6. fluorescence emission of two or more cellular constituents or adsorbed antibody.

Cell separation by sedimentation and centrifugation Methods

Centrifugation depends not only on the centrifugal force but also on other factors which modify sedimentation and are dependent on the cell characteristics.

Differential centrifugation

This process is normally the most simple in practice, given that it only can separate cells showing large differences in size (at least 10 times) (Figure 2a). In the case of blood with anticoagulant, at 200 g (g = gravities) erythrocytes sediment in the lower zone, leukocytes appear at the interface and at the supernatant there will be a platelet rich plasma phase. It is possible to obtain plasma with less platelets using higher speed (3.000g) (Figure 2b).

Figure 2

Differential centrifugation. Separation is done according to size and the sedimentation coefficient, which is dependent on the mass. A. Peripheral blood separation. B. Differential centrifugation by size.

Density gradient centrifugation

Barrier methods (centrifugation through a continuous centrifugation media). To achieve more effective separations each sample is centrifuged on a bed of density intermediate between the two cell types that are to be separated.

Zonal centrifugation or sedimentation rate. This method is used to separate cell types whose sedimentation coefficient differs. To be able to perform this separation it is necessary to form a density gradient, which favors the concentration of each cell type in a narrow band or zone. The density gradient is generated before adding the sample, using several different solutions with suitable concentration of a compound (Ficoll, sucrose, albumin, fetal bovine serum).

After the gradient is formed, a small volume of sample is deposited upon it and centrifuged for a short time. Typically, for this process a gradient less than the maximum density of cells is used and therefore it may not reach the sedimentation equilibrium. Always shorter times are used, if cells are centrifuged for longer times they will leak and end up in the background, this is how the result is dependent on the time of centrifugation. This technique can separate all blood cell types, viable sperm, viable and non-viable cells from disaggregated tissues and suspension cell samples (Figure 3a).

Figure 3

A. Zonal centrifugation, separation depending on the coefficient of sedimentation. Centrifugation stops before the sedimentation equilibrium is reached. B. Centrifugation on a gradient of constant centrifugation media, separation media have constant density (more...)

Peripheral mononuclear blood cells separation. The peripheral blood mononuclear cells (PBMCs) include lymphocytes, monocytes and macrophages. They share different characteristics as the presence of a circular nucleus and their density. As a consequence of these characteristics they can be isolated by different techniques (see above-Cell separation methods) and also by the use of special type of tubes as the BDVacutainer® CPT™ which is a vacuum-driven drawing tube containing anti-coagulant and a cell separation medium. The procedure to isolate the cells may also affect them, as for the use of different components to the separation and the serial washes to clean them.

Using Ficoll-Hypaque, which has a density of 1.077 g/ mL, density identical to that of lymphocytes and monocytes, it is possible to recover PBMCs. The Ficoll-Hypaque is a combination of a polymer of high molecular weight sucrose (Ficoll) and an organic compound (sodium diatrizoate: 3–5 bis acetylamino-2, 4, 6 tri-iodobenzoic acid). Granulocytes and erythrocytes have a higher density and when peripheral blood is centrifuged in a Ficoll-Hypaque gradient, it passes through a package formed on the bottom of the tube. Platelets have a lesser density and remain in the plasma and the mononuclear fraction located at the interface (Figure 3b).

There are other solutions for the separation of mononuclear cells using density gradients, the most effective one is the use of Percoll. To separate mononuclear, the commercial solutions most commonly used is Lymphoprep, Hystopaque and Lymphopure. Several media solutions are available with suitable densities for the separation of other specific cell types: Nycoprep -1.077 for mononuclear cells, Nycoprep -1.068 for monocytes, Polymorphoprep for polymorphonuclear cells and Nycoprep -1.063 for platelets.

Isopycnic centrifugation or sedimentation equilibrium. This is a method in which cells are separated solely according to its density using another centrifugation variant, called isopycnic (der. Greek. Similar density). It is also performed on a density gradient, but in this case the centrifugation time is sufficiently long to reach the sedimentation equilibrium. To achieve the sedimentation equilibrium continuous gradients are used to cover the entire range of cell densities: at the bottom of the tube has to have the greater density than the denser cells. Thus, independent of the time of centrifugation, the cells will never sediment at the bottom but instead will reach a stable intermediate position in the gradient.

Centrifugal elutriation. This process involves the separation of cells according to their sedimentation rate. The original process consists on performing successive cycles of sedimentation and decanting provided by the incorporation into a system of liquid flow suspension. Finally, when it is combined with the effect of the centrifugal force it results in centrifugal elutriation. Here the cells are exposed to two opposing forces; firstly by centrifugal force and the drag force by the continuous flow of the medium counter force ().

Figure 4

Elutriation. A. Elutriation separation chamber. B. Elutriation process. 1. Introduction of the cell suspension. 2. The centrifugal force pushes the cells to the bottom of the chamber, continuously introducing a liquid medium, which opposes the centrifugal (more...)

The above method yields a versatile, rapid and effective form of separation for cell subpopulations according to their size from a mixture. Moreover this process, admits high cell concentrations and even allows for a greater recovery rate of viable cells compared to the original cell.

Blood cell storage: cryopreservation and thawing process

Sometimes cells are not necessarily cultured immediately after the blood drawing, thus it is necessary to keep and storage them for future experiments. Freezing is the best way to storage cells. However, freezing affects the proliferation and cytokine secretion as well as the protein production and mRNA expression. Other consequences of the freeze-thaw process are mechanical injuries produced by crystal formation, alteration of physical properties and shape of cellular structures. That is why protocols for freezing cells include cryoprotectant substances in the freezing media that prevent crystal formation and avoid osmotic injury. Some examples are the dymethylsulphoxide (DMSO), glycerol, ethylene glycol or hydroxyethyl.

Hence, it is necessary to take into account many considerations when freeze-thawing cells. Firstly, it is extremely important to process and store the cells within a period between 8 to 24 hours after samples are collected and they should be conserved in especial freezing media. Secondly, it could be helpful to include in the freezing media caspase inhibitors to avoid the apoptosis of the cells as a consequence of the stress. And finally, the rate by which the temperature will decrease should be slow enough to avoid the rupture of the cells. This is done using special containers that have alcohols (i.e Isopropanol) which surround the sample tubes. Thus, the rate of temperature decreasing is gradually, and close to 1°C/min to -70°C. After this process frozen samples should be transferred to liquid nitrogen promptly within the next 24 – 72 hours.

As for the thawing process it is recommended to thaw the cells rapidly by transferring the cryovials directly from liquid nitrogen to 37°C. Immediately after the samples are thawed, they should be diluted and washed to eliminate the cryopreservant that could be toxic for the cells. The rapid change of temperature and media diminish the osmotic variation and protects the integrity of the cells. However, despite all these considerations there may be some problems, for example, cell clumping is frequent following the thawing process as a consequence of the release of DNA of dead cells. In that case, DNase can be used to avoid the aggregation of cells. Finally, it is recommended giving the cells a resting period before the experiment so they can eliminate the components that they were producing before the freezing process. This resting period allows them also to get used to the new conditions and favors to normalize the conditions for future comparisons. General recommendations about the treatment of blood samples for cell culture are summarized in table 2.

Table 2

General guidelines for sample blood collection and cell processing.

Characterization and separation of cells by cellular markers

Flow cytometry

Today, flow cytometry is an important method in biomedical research and clinical laboratories, especially for its ability to analyze automatically different cell suspensions. Flow cytometry is based on the transportation of a cell suspension (e.g., blood cells, bone marrow aspiration and dissociated tissues) driven by the flow of an isotonic solution to the measuring point or flow chamber. Flow cytometry uses include analysis (biomarker detection) and separation (sorting) of cells previously labeled with fluorochromes (Figure 5). Some flow cytometers only perform the first, while others carry out both.

Figure 5

Flow cytometry. Principles of operation of the flow cytometer (see text). Adapted from (1).

Characterization of a cell population. Briefly, the cell suspension arrives in laminar flow conditions, forming a very fine line containing individual cells in succession. These cells pass one by one through a laser whose wavelength allows excitation of previously incorporated fluorescent markers. The light emerging from each cell is analyzed for scattering and fluorescence intensity.

Cell characterization is accomplished by measuring multiple physical characteristics of the cells, as they flow to a beam of light. The properties measured include the relative size, relative granularity or internal complexity, and relative fluorescence intensity given by the use of fluorocromes. In order to make these measurements, the cytometer has three main systems: fluidics, optics, and electronics. The first one transports the cells to the interrogation point where the laser bean pass through. The second one consists of the different lasers that illuminate the cells in the interrogation point, and directs the light to the filters and detectors. And finally the third one is the electronic system that helps to convert the changes in light signals from the detectors to values that can be interpreted by the computer.

This technique has multiple functional applications. study different cell surface markers and intracellular signaling by using monoclonal antibodies, assessment of DNA and RNA content of the cell and the determination of its shape and size.

Characterization of a cell population by flow cytometry

The first characteristics than can be determined by the simplest cytometer are the shape and the internal complexity of the cells. These characteristics are measured by the changes in the light scattering. It occurs when a cell deflects incident laser light. Hence there are many cellular factors that affect light scattering such as cell’s membrane, nucleus, and any granular material inside the cell. Also, the cell shape and surface topography can contribute to the total light scatter.

There are two ways to measure the scattered light. First, the forward-scattered light (FSC) is proportional to cell-surface area or size. As a consequence, FSC can be interpreted as the shadow projected by the cell, finally the detection of FSC is done parallel to the lasers. On the other hand, the side-scattered light (SSC) is proportional to cell granularity or internal complexity. For instance, SSC is the measurement of mostly refracted and reflected light that occurs at any interface within the cell, it means that SSC indicated how much the light is diverted from the original source as a consequence of the content of the cells. As results, SSC is collected at approximately 90 degrees to the laser beam (figure 6).

Figure 6

A. Schematic representation of side and forward scatter light (SSC and FSC). B. Results of FSC vs SSC analysis of leucocytes from whole blood.

In addition, superficial and internal cell markers can be detected by flow cytometry and they allow a better characterization of the cell population within a sample. In order to do these analyses it is necessary to use antibodies that bind to the markers of a specific cell population, but there is not enough binding of the antibodies to identify the cell. As a result, the antibodies should be labeled with different flurochromes. As an example, a cytometer with three lasers can detect 8 colors. It means that it can be used to find 8 different cell markers plus SSC and FSC.

Fluorochromes

Typically, a fluorescent compound absorbs light energy over a range of wavelengths that is characteristic for that compound. The absorbed light causes excitement of electrons in the compound raising them to a higher energy level. Finally, when the source energy finishes the excited electrons quickly decay to their basal state, emitting the excess energy as photon light. This process is called fluorescence.The range over which a fluorescent compound is excited is known absorption spectrum. On the other hand, the range of emitted wavelengths for a particular compound is known as emission spectrum.

Fluorochromes are fluorescent components that are excited by different wavelength, and they emit light in specific wavelengths in the visual spectra that can be detected by an instrument (Figure 7). Hence, this property is used for labeling antibodies; commonly, a fluorescent dye is conjugated to a monoclonal antibody, then it can be used to identify a particular cell type based on the characteristic cell markers (Figure 8).

Figure 7

Fluorochrome characteristics. A. Specific wavelength values of excitation and emission. (Colors represented the fluorescence emission visible color for each fluorochrome). B. Relative brightness scale of common fluorochromes. It is recommended to use (more...)

Figure 8

Antibodies labeled with fluorochromes against specific cell markers allow the characterization of a cell population within a sample.

The correct detection and characterization of cell populations depends on the right choice of fluorochromes and the correct calibration of the Flow cytometry. In the first case, it is important to know which cells are going to be analyzed and how is the expression pattern of the marker that is going to be used for their characterization. For example, a marker which has a low expression in the cell surface should be labeled with a strong fluorochrome in order to intensify the signal. On the contrary, a marker highly expressed can be labeled with a fluorochrome with mild intensity (Figure 7).

Secondly, the cytometer should be calibrated in order to provide good quality results. One of these calibration processes is called compensation and it should be done every time that an experiment is run. Compensation is the correction for the spectral overlap during multicolor flow cytometry experiments. The goal of color compensation is to correctly quantify each dye with which a particular cell is labeled. This is done by subtracting the portion of the signal overlapping between fluorochromes (Figure 9. For further information please go to the recommended lectures).

Figure 9

Compensation. In an experiment with more than two fluorochromes there can be spillover. Spillover takes place when the presence of the other fluorescent reagent can contribute significantly to optical background in proportion to the brightness of a specific (more...)

All this together, allows the characterization of a mixed population of cells by using different fluorochromes. This method leads to distinguishing and characterizing the subpopulations within the sample in combination with FCS and SSC. The combination of different markers, FCS or SSC enables the definition and sub-analysis of populations known as “gating’. (Figure 10)

Figure 10

Gating approach to analyze peripheral blood mononuclear cells (PBMC). Left panel shows the results for the stained control and right panel shows the results for the unstained/negative control, which includes all the events registered in panel A. Panel (more...)

Cell separation by flow cytometry or fluorescence activated cell sorting (FACS)

This process allows the separation of cell samples fractions according to their morphologic and fluorescent characteristics. A regular flow cytometry analysis must be done before to define which populations will be sorted. Then, cells are separated in different fractions when the flow passing through the detection point is transformed into small droplets. These droplets contain a single cell (as a result of an ultrasonic vibration). Finally, a voltage pulse which provides an electrical charge is applied according with the cell characteristics. This pulse allows the cells separation when the droplet passes through an electrical field. Then, cells are deflected according to their charge, thus falling into different sample collection tubes (Figure 5).

Affinity separation by magnetic particles

This process is of great application. Currently in use are commercially available microspheres (beads) in several sizes (0.5 - 10 microns) for different applications and formed by a super-paramagnetic material (iron oxide) coated with a thin layer of plastic polymer which allows the absorption and/or covalently binding of different molecules. Often an antibody is bound to magnetic or immunomagnetic microspheres antibodies (Figure 11).

Figure 11

Affinity separation using magnetic beads. A. Direct labeling: microsphere covalently linked to the Fc domain specific antibody for the target cells. B. Indirect labeling: uses two types of reagents, an unmodified primary antibody covalently linked to (more...)

Count and cell viability

The accurate determination of the amount and viability of the cells is very important for correct standardization of reagents and conditions for cell culture experiments. Counting is performed using a hemacytometer counting chamber consisting of a central chamber (double counting chamber) which is divided into two parts by a transverse slit of 1mm. Each chamber consists of a silver film etched in a grid of 3×3 mm. Each rack is divided into nine side frames, each of 1×1 mm. Boxes in the corners are divided into 16 squares and center box in 25. The hemacytometer is accompanied by a thin cover glass slide with a weight that determines the exact depth when placed on the chamber (Figure 12). Alternatively, cells can be counted by using flow cytometry according to the number of events registered in the analysis or the use of special tubes which allows the determination of absolute cell counts (Further information can be found in recommended readings or in the Becton Dickinson website http://www.bdbiosciences.com/research/).

Figure 12

Hemocytometer. The coverslip is 0.1 mm above the grid, and the lines etched on the grid are at preset dimensions. The four outer squares, marked 1–4, each cover a volume of 10–4 mL. The inner square, marked as 5, also covers a volume of (more...)

Cell viability refers to the ability of a cell to perform its biochemical and physiological processes, particularly in regards to its metabolism and ability to divide. However, in practice the term is relative as it is used with different criteria; viability is commonly spoken of when referring to cell integrity or metabolic activity or their proliferative capacity. In fact, with the use of a vital dye exclusion is possible to determine cell viability base on the integrity of the membrane from living cells. This membrane excludes certain dyes such as trypan blue, eosin, 7-AAD or propidium iodide, whilst dead cells allow their passage into the cytoplasm (Figure 13). For this method, the cell suspension is mixed with a known volume of dye and examined visually to determine the dead and live cells. In the case of trypan blue, cytoplasm is seen in refractive (clear) when the cell is alive while the cytoplasm of dead cells is seen blue. Propidium iodide stains dead cells red when observed in the fluorescence microscope. Alternative, flow cytometry can be used instead of the microscope with the colorants 7-AAD or propidium iodide (Figure 14).

Figure 13

Cell viability - Exclusion assays These tests determine the number of viable cells present in a cell suspension. Live cells possess intact cell membranes that exclude certain dyes like trypan blue. If the cell membrane is damaged (dead cells) trypan blue (more...)

Figure 14

Cell viability - Exclusion assays in flow cytometry. The 7-AminoActinomycin D (7-AAD) has the ability of inserting itself between the tops of successive CG bases of the DNA double strand. This occurs when the interior of the cell and the nuclear chromatin (more...)