Introduction to animal tissue culture science

Animal tissue culture technology is now becoming a significant model for many scientists in various fields of biology and medicine. Despite the various developments in animal cell and tissue culture since the late 1800s, until the early 1950s progress in animal tissue culture was stalled due to the non-availability of a suitable cell line. In the early 1950s, for the first time, successful growth of cells derived from the cervical cancer of Mrs Henrietta Lacks was demonstrated. This breakthrough using Mrs Henrietta Lacks's cells in culture successfully transformed medical and biological research, allowing numerous cellular, molecular and therapeutic discoveries, including the breakthrough of the first effective polio vaccine [1, 2]. This culture is now called HeLa, on which there were more than 60 000 publications by 2017, and which has been involved in numerous Nobel prize-winning innovations [2–4].

Animal cell culture is a significant tool for biological research. The importance of cell culture technology in biological science was realized a long time ago. Earlier dedifferentiation based experiments of cells due to selective overgrowth of fibroblasts resulted in the enhancement of culture techniques. Animal cell culture involves isolation of cells from a tissue before establishing a culture in a suitable artificial environment. Initial isolation of the cells from the tissues can be achieved by disaggregation using enzymatic or mechanical methods. The source of the isolated cells is usually an in vivo environment, but sometimes cells are also derived from an existing cell line or cell strain. Animal cell culture offers suitable model systems for investigating the following factors:

• Drug screening and development.
• Mutagenesis and carcinogenesis.
• Normal physiology and biochemistry of cells.
• Potential effects of drugs and toxic compounds on the cells.

In addition, it also permits reliable and reproducible results, and is thus considered as a significant model system in cellular and molecular biology. Mammalian cell culture requires an optimal environment for growth. Environmental conditions are divided into nutritional requirements and physicochemical requirements. Nutritional requirements include a substrate or medium that provides support and essential nutrients such as amino acids, carbohydrates, vitamins, minerals, growth factors, hormones and gases (O₂, CO₂). All these factors control physical and chemical factors such as pH, osmotic pressure and temperature. In animal tissue culture the majority of cells are anchorage-dependent and therefore require a solid or semi-solid support in the form of a substrate (adherent or monolayer culture), whereas others can be cultured in the culture medium, called a suspension culture. Cell culture technologies have emerged as a tool to assess the efficacy and toxicity of new drugs, vaccines and biopharmaceuticals, and also play a major role in assisted reproductive technology. Animal cell culture is one of the more important and diverse techniques in current research streams. Animal, plant and microbial cells are always cultured in predetermined culture medium under controlled laboratory conditions. Animal cells are more complex than micro-organisms. Due to their genetic complexity it is difficult to determine the optimum nutrient requirements of animal cells cultured under in vitro conditions. Animal cells require additional nutrients compared to micro-organisms, and they usually grow only when attached to specially coated surfaces. Despite these challenges, different types of animal cells, including both undifferentiated and differentiated ones, can be cultured successfully.

Tissue culture involves the in vitro maintenance and propagation of cells in optimal conditions. Culturing animal cells, tissue or organs in a controlled artificial environment is called animal tissue culture. The importance of animal tissue culture was initially realized during the development of the polio vaccine using primary monkey kidney cells (the polio vaccine was the first commercial product generated using mammalian cell cultures). These primary monkey kidney cells were associated with many disadvantages [5–8] such as:

• Chances of contamination with adventitious agents (risk of contamination by various monkey viruses is high).
• Most of the cells are anchorage-dependent and can be cultured efficiently only when they are attached to a solid or semi-solid substrate (obligatorily adherent cell growth).
• The cells are not well characterized for virus production.
• A scarcity of donor animals as they are on the verge of extinction.

The foundation of animal tissue culture can be considered to have occurred in 1880, when Arnold showed that leukocytes can divide outside the body [9]. Then, in the beginning of the 19th century, Jolly investigated the behavior of animal cells in serum lymph [9]. The development of animal tissue culture commenced after the breakthrough frog tissue culture technique, which was discovered by Harrison in 1907. Due to this effort Harrison is considered as the father of tissue culture. In his experiment he introduced tissue from frog embryos into frog lymph clots and showed that not only did the tissue survive, but nerve fibers grew out from the cells. During the mid-20th century, human diploid fibroblast cells were established by Hayflick and Moorhead [10]. They named this cell line MRC-5 (a cell line of fibroblasts derived from lung tissue). Later, Wiktor et al (1964) explored the utilization of this cell line in the production of rabies virus for vaccine production [11]. After a couple of years they suggested a large-scale production protocol along with a method for the assessment of purified rabies vaccine immunogenicity. During the same time, BHK-21 (C13) cells (baby hamster kidney cells) were established. These cells are susceptible to human adenovirus D, reovirus 3 and vesicular stomatitis virus. The commercial production of inactivated foot and mouth disease (a viral disease that causes sores in the mouth and a rash on the hands and feet of children) vaccine began using a suspension process [12]. Back in 1914, Losee and Ebeling [13] cultured the first cancer cells and after a few decades the first continuous rodent cell line was established by Earle (1943) [14]. In 1951, Gay established that human tumor cells can give rise to continuous cell lines. The cell line considered as the first human continuous cell line was derived from a cancer patient, Henrietta Lacks, as mentioned above, and HeLa cells are still used very widely. Continuous cell lines derived from human cancers are the most extensively used resource in the modern laboratory. The HeLa discovery was followed by FDA approval for the production of interferon from HeLa cell lines [15]. In addition to the progress in the field of cell culture, different media have been explored, which are typically based on specific cell nutritional requirements, such as serum-free media, starting with Ham's fully defined medium in 1965. In the 1970s, serum-free media were optimized by the addition of hormones and growth factors. Currently, thousands of cell lines are available and for the establishment and maintenance of these cell lines many media are available.

Broadly, animal tissue culture can be divided into two categories:

• Cultures that allow cell–cell interactions and encourage communication or signaling between cells.
• Cultures in which cell–cell communication or interactions are lost or the signaling between them is missing.

The first category includes three different types of culture systems: organ cultures, histotypic cultures and organotypic cultures. The second category includes cultures in monolayers or as suspensions. Organ culture is a culture of native tissue that retains most of the in vivo histological characteristics, whereas culturing cells for their re-aggregation to yield tissue-like structure is known as histotypic culture. In histotypic cultures, individual cell lineages are initially derived from an organ and then cultured separately to high density in a 3D matrix to study interactions and signaling between homologous cells. In organ cultures, whole embryonic organs or small tissue fragments are cultured in vitro in such a manner that they retain their tissue architecture, i.e. the characteristic distribution of various cell types in the given organ.

In an organotypic culture, cells from different origins are mixed together in specific proportions and spatial relationships so as to re-form a component of an organ, i.e. the recombination of different cell types to yield a more defined tissue or organ. Some terms frequently used in animal tissue culture are as follows.

Cell culture. Cell culture is the process of removing cells from an animal or plant and their subsequent growth in an artificially controlled environment.

Primary cell culture. This is the first culture (a freshly isolated cell culture) or a culture which is directly obtained from animal or human tissue by enzymatic or mechanical methods. These cells are typically slow growing, heterogeneous and carry all the features of the tissue of their origin. The primary objective of this culture is to maintain the growth of cells on an appropriate substrate, available in the form of glass or plastic containers, under controlled environmental conditions. Since they are directly obtained from original tissue they have the same karyotype (number and appearance of chromosomes in the nucleus of a eukaryotic cell) as the original tissue. Once subcultured, primary cell cultures can gives rise to cell lines, which may either die after several subcultures (such cell lines are known as finite cell lines) or may continue to grow indefinitely (these are called continuous cell lines). Usually, normal tissues give rise to finite cell lines, whereas cancerous cells/tissue (typically aneuploid) give rise to continuous cell lines. Nevertheless, there are some exceptional examples of continuous cell lines which are derived from normal tissues and are themselves non-tumorigenic, e.g. MDCK dog kidney, fibroblast 3T3, etc. The evolution of continuous cell lines from primary cultures is assumed to involve mutation, which alters their properties compared to those of finite lines. Serial subculturing of cell lines over time can increase the chances of genotypic and phenotypic variation. Bioinformatic studies based on proteomic phenotypes discovered that the Hepa1–6 cell lines lacked mitochondria, reflecting a rearrangement of metabolic pathways in contrast to primary hepatocytes. With the emergence of newer technologies such as 3D culture, the use of primary cells is becoming increasingly prevalent and achieving improved results. Primary cells which are directly obtained from human or animal tissue using enzymatic or mechanical procedures can be classified into two types:

• Anchorage-dependent or adherent cells. Adherent cells are those cells which require attachment for growth and are also called anchorage-dependent cells. In other words, these cells are capable of attaching on the surface of the culture vessel. These types of cells are often derived from the tissues of organs, for example from the kidney, where the cells are immobile and embedded in connective tissue.
• Anchorage-independent or suspension cells. Suspension cells do not require attachment or any support for their growth and are also called anchorage-independent cells. All suspension cells are isolated from the blood system, for example white blood cell lymphocytes, and are suspended in plasma.

For several reasons cells obtained from primary cultures have a limited life span, i.e. the cells cannot be maintained indefinitely. An increase in cell numbers in a primary culture results in exhaustion of the substrate and nutrients, which can influence cellular activity and lead to the accumulation of high levels of toxic metabolites in the culture. This may ultimately result in the inhibition of cell growth. This stage is called the confluence stage (contact inhibition), when a secondary culture or a subculture needs to be established to ensure continuous cell growth.

Secondary cell culture. This simply refers to the first passaging of cells, a switch to a different kind of culture system, or the first culture obtained from a primary culture. This is usually carried out when cells in adherent cultures occupy all the available substrate or when cells in suspension cultures surpass the capacity of the medium to support further growth, and cell proliferation begins to decrease or ceases completely. So as to maintain optimal cell density for continued growth and to encourage further proliferation, the primary culture has to be subcultured. This process is known as secondary cell culture. Major differences between primary and secondary cell cultures are highlighted in table 1.1.

Cell line. Once a primary culture is subcultured or passaged it represents a cell line. A cell line that experiences indefinite growth of cells during subsequent subculturing is called a continuous cell line, whereas finite cell lines experience the death of cells after several subcultures.

Cell strain. A cell line is a permanently established cell culture which will proliferate forever if a suitable fresh medium is provided continuously, whereas cell strains have been adapted to culture but, unlike cell lines, have a finite division potential. A cell strain is obtained either from a primary culture or a cell line. This is done by selection or cloning of those particular cells having specific properties or characteristics (e.g. specific function or karyotype) which must be defined.

In summary, the first culture that is established from the in vivo environment is called the primary culture. This primary culture can be subcultured many times to develop cell lines. Cell lines are generally immortalized or transformed cells, i.e. cells that have lost control over division, because of mutations or genetic alterations, or because a primary cell was transfected with some genes that immortalized the cells. Most cell lines are tumorigenic as they originated from tumors. Cells derived from a primary cell line do not have this concern, however, it is challenging to maintain these cells. In usual practice, primary cell cultures require a nutrient medium containing a high amount of different amino acids, micronutrients and, occasionally, some types of hormones or growth factors. Primary cell cultures can be efficiently utilized up to a few passages, about two to four, afterwards their risk of contamination is higher than for cell lines. However, primary cell cultures have their own advantages. The biological response received from a primary culture will be closer to that in an in vivo environment than the response obtained from cell lines. From many years, several cell lines have been established and tested under different environmental conditions. This vast research has resulted in a good amount of data supporting the use of specific cell lines as models of primary cells. It has been suggested that cell lines that have been well tested under different conditions should be used instead of primary cultures, in the case that the latter are expensive.

As discussed above, the primary cell culture is the first culture of cells, tissues or organs derived directly from an organism; in other words it is the culture before the first subculture, whereas the cell line is for maintenance or propagation of a culture after subculture. There are certain techniques available for the development of primary cell cultures, such as:

• Mechanical disaggregation.
• Enzymatic disaggregation.
• Primary explant techniques.

1.4.1. Mechanical disaggregation

1.4.2. Enzymatic disaggregation

This approach involves efficient disaggregation of cells with high yield by using enzymes such as trypsin, collagenase and others. Enzyme based disaggregation allows hydrolysis of fibrous connective tissue and the extracellular matrix. Currently, the enzymatic method is extensively used as it offers high recovery of cells without affecting the viability of cells.

1.4.2.1. Trypsin based disaggregation or trypsinization

This allows disaggregation of tissue using trypsin, usually crude trypsin because this trypsin contains other proteases. In addition, cells can tolerate crude trypsin well and the ultimate effect of crude trypsin can easily be neutralized by serum or trypsin inhibitor (supplementation of trypsin inhibitor is required in the case of serum-free media). Pure trypsin can also be utilized for disaggregation of cells, provided that it is less toxic and very specific in its action. An overview of primary cell culture development is shown in figure 1.1. Two common approaches, namely warm and cold trypsinization, are described in the following.

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Warm trypsinization

This approach is extensively utilized for the disaggregation of cells. During the initial step, sliced tissue is washed with dissection basal salt solution and is subsequently transferred to a container of warm trypsin (37 °C). At regular intervals of 30 min the contents are stirred properly. Then, the supernatant having dissociated, the cells are separated to disperse in a suitable medium. Efficient dispersion of cells can be achieved by placing the container over ice.

Cold trypsinization

This method is also called trypsinization with cold pre-exposure. In this process the chance of cellular damage due to constant exposure to trypsin is reduced, which results in a high yield of viable cells with an improved survival rate for the cells (after 24 h of incubation). Since this method does not involve frequent stirring or centrifugation, it can be conveniently adopted in the research laboratory. During this process, after washing and chopping, tissue pieces are kept over ice in a vial and then subjected to treatment with cold trypsin for 6–24 h. Then, after the cold trypsin treatment the trypsin is removed and discarded. However, the tissue fragments still contain residual trypsin. These fragments are incubated at 37 °C (for 20–30 min) followed by repeated pipetting. This will encourage the dispersion of cells. The fully dispersed cells can be counted using a cell counter and properly diluted, and then further utilized.

Drawbacks of trypsin disaggregation

Trypsinization of cells can damage some cells, such as epithelial cells, and sometimes it is not effective for certain tissues, such as fibrous connective tissue, thus other enzymes are also recommended for dissociation of cells.

1.4.2.2. Collagenase based disaggregation

Collagenase is an enzyme which is responsible for the cleavage of peptide bonds in collagen. Collagen is a structural protein which is abundantly found in higher animals, mainly in the extracellular matrix of connective tissue and muscle. Collagenase, mainly crude collagenase, can be successfully used for the disaggregation of several tissues that may or may not be sensitive to trypsin. Purified collagenase has also been experimented with, but has shown poor results in comparison to crude collagenase. So far collagenase disaggregation has be carried out on several human tumors, epithelial tissues, the brain, lungs and other mammalian tissue. The combination of collagenase with hyaluronidase offers better results in disaggregating rat or rabbit liver, which can be achieved by perfusing the whole organ in situ. Several researchers have also utilized trypsin and collagenase in combination to dissociate cells to develop chick serum.

This process involves an initial transfer of the desired tissue into a basal salt solution which contains antibiotics. This is followed by washing with settling and then transfer into a medium containing collagenase. The solution is incubated for 1–5 days, followed by repeated pipetting for uniform dispersal of cells. Separation of these dispersed cells is encouraged by keeping the solution in a stationary phase to further encourage the settling of cells, as shown in figure 1.2.

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1.4.2.3. Other enzymes

In addition to the above mentioned enzymes, certain other enzymes such as bacterial proteases (e.g. dispase, pronase) have been tested, but unfortunately have not shown significant results. However, enzymes such as hyaluronidase and neuraminidase have received attention due to their significant results, and thus can potentially be utilized in conjugation with the enzymes discussed above.

1.4.3. Primary explant technique

After the development of a primary cell culture, it is essential to remove the non-viable cells from the disaggregated cells, which can be achieved by repeatedly changing the medium. Only a few will be left after dilution of the medium, and finally will gradually disappear when viable cells start proliferating. The alternative approach of centrifugation, mixing cells with ficoll and sodium metrizoate, can also be utilized to remove non-viable cells from the primary cell culture. Dead cells form a pellet at the bottom which can easily be removed from the solution.

Animal tissue culture techniques involve the frequent utilization of animal or human tissues, which raises the need for safety and ethics guidelines for using animals in research, also known as medical ethics. Handling animals raises numerous issues that are typically not faced when using animal tissue. In addition to the consent of local ethical committees, the consent of the patient or his/her relatives is required to initiate research or to study a human sample in the form of fetal materials or biopsy samples. Samples collected from a human donor should be accompanied by a donor consent form in a prescribed format. When dealing with human tissue, the following issues should be considered [16]:

• The patient's or relatives' consent for using tissue for research purposes.
• Ownership of specimens, in particular cell lines and their derivatives, i.e. the recipient will not trade or transfer the cell lines and their derivatives.
• Consent for genetic modification, in particular in the case of cell lines.
• Patent or intellectual rights for the commercial use of cell lines.
• Guidelines should be refined to meet the requirements of the latest ongoing developments in animal tissue culture science. These guidelines are framed to offer knowledge to those new to the field and others involved in training and instruction, with the data required to improve their awareness of issues and to allow them to deal with them more efficiently. The primary areas of focus in guidelines are:
  • i.  
    Acquisition of cell line.
  • ii.  
    Authentication of cell line.
  • iii.  
    Characterization of cell line.
  • iv.  
    Cryopreservation of cell line.
  • v.  
    Development of cell line.
  • vi.  
    Instability of cell line.
  • vii.  
    Legal and ethical requirements when deriving cell lines from human and animal tissues.
  • viii.  
    Microbial contamination of cell line.
  • ix.  
    Misidentification of cell line.
  • x.  
    Selection and maintenance of equipment.
  • xi.  
    Transfer of cell lines between laboratories.

Generally, when dealing with human tissue, the donor/relative is asked to sign a disclaimer statement in a prescribed format before tissue is collected. Doing this can reduce the chances of legal problems [16].

Handling human tissue involves a high risk of exposure to various infections, thus it is essential to handle human material in a biohazard cabinet. Before their use, tissues must be screened properly for various infections such as hepatitis, tuberculosis and HIV. In addition, media, apparatus and glass wares should be properly sterilized (autoclaved) to considerably reduce the chances of spreading any infections.

A cell line can be defined as a permanently established cell culture which will propagate forever, provided the continuous supply of suitable fresh medium and the availability of space for the cells to propagate. Thus, generally, a cell line can be defined as the propagation of a culture after the first subculture. In other words, when primary culture is subcultured it results in the development of a cell line. Cell lines differ from cell strains in that they become immortalized. A cell line contains several cell lineages, either similar or different in their phenotypical characteristics, and such cells can be selected by cloning or cell separation or by any other suitable procedure. The cell line obtained after selection or cloning is called a cell strain, which does not have a infinite life, since they die after a number of divisions.

1.8.1. Types of cell lines

1.8.2. Standard nomenclature of cell lines

1.8.3. Cell line selection

1.8.4. Verification of a cell line

1.8.5. Characterization of cell lines

1.8.6. Misidentification of cell lines

1.8.7. Maintenance of a cell line

Subculture is defined as the transfer of cells from one culture to start another culture. During this process proliferating cells are subdivided, which allows the development of new cell lines. This step is referred to as a passage, and a passage number is the recorded number of times a cell culture has been subcultured. Numerous adherent cell cultures will stop proliferating when they reach the confluent stage (i.e. when they completely cover the surface of the cell culture vessel), and certainly will die if they are left in the confluent stage for a prolonged period. Thus adherent cell cultures should be regularly passaged, that is, when cells reach the confluent stage a portion of the cells need to be passaged or subcultured to a new cell culture vessel. However, it is not recommended to regularly subculture adherent cells (no more than once every 48 h) as they must be trypsinized. In contrast, suspension cultures with high cell density require routine passaging as they use medium rapidly.

The standard growth curve of cells in a culture is shown in figure 1.2.

During the initial lag phase there is less growth as the cells are not adapted to the environment. Once they start adapting to the environment they proliferate exponentially, which is why this is called the exponential or log phase. This is the time when all cells actively grow and consume medium. During this time the medium should be changed, otherwise growth will stop. As discussed above, the confluent phase is reached when the culture exceeds the capacity of the medium. At this stage the culture has to be divided into subcultures. There are two types of subcultures: monolayer and suspension subcultures.

1.9.1. Monolayer cultures

1.9.2. Procedures for cell detachment

There are various means of cellular detachment from the culture vessel interface, such as physical and chemical methods (figure 1.3).

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The utilization of proteases is not recommended when cultures are loosely adhered, thus mechanical shaking and scraping are more appropriate in such cases. Due to its advantages, trypsin is often used for cell dissociation, however, other enzymes such as pronase, dispase and collagenase are used when monolayers cannot be disaggregated with trypsin. Prior treatment with EDTA is required to remove Ca⁺² so that it will not interfere with the action of enzymatic dissociation, and eventually uniform dispersion can be achieved [30]. As one-cell-thick monolayers are the simplest tissues in multicellular organisms, they act as a suitable model for development and normal physiology. It has been determined that the extracellular material (ECM) should be carefully considered when selecting the dissociation approach, as this helps in determining the effects of the dissociating agent on the cytoskeleton, adherent junctions and desmosomes. Usually monolayers can withstand the different mechanical stresses exerted by the interface itself under in vitro conditions, and can shield the internal environment from harmful external elements. Since dissociating elements or external environmental factors can affect ECM synthesis, optimization of the dissociating element is required before treatment in order to estimate a suitable dose for dissociation [30].

As mentioned in figure 1.4, subculturing is usually carried out between the middle log and plateau phases; it is not recommended to start subculturing during the lag phase.

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Understanding of growth patterns is necessary for:

• Designing culture experiments.
• Regular maintenance of a culture.
• Monitoring cell proliferation.
• Evaluating a culture's response to external factors.

There are certain considerations in subculturing monolayers, as follows.

Cellular density

Subculturing time is dependent on the cellular density. Cellular density is usually found to be high at the confluence stage. So, whenever normal or transformed cells reach the confluence stage it is advisable to perform subculturing as this can maintain the balance between nutrient supplementation and consumption by the cells/micro-organisms. In the confluence stage, when all the growth area is utilized and cells start coming closer to each other, growth may be hampered due to the negative force (contact inhibition) developed between the cells striving for nutrients to further meet their energy demands.

Exhaustion of nutrients

Usually, in microbiology a sudden drop in pH represents an increase in cellular density which again signifies the confluence stage, thus a drop in pH often necessitates subculturing.

Reason for subculture

Subculturing is also done in those cases when cells are to be used for any specific purpose other than routine propagation, in order to obtain high yield or stock, or to change the type of medium. In such cases the cell has to be subcultured frequently.

Scheduled timings for subculture

As we know, regular subculturing is generally performed as per a strict schedule to obtain significant results. Seeding density should be increased in the case that the cell will not reach the confluence stage at a suitable time, and seeding density should be reduced when the cells will reach the confluence stage a little too early. It is now possible to determine the correct seeding density and subculture interval by studying standard growth curves. In most cases medium change is performed after 3–4 days and subculturing after 7 days.

The steps involved in monolayer subculture are shown in figure 1.5. Monolayer subculturing involves multiple steps:

• Medium removed and monolayer washed.
• Treatment of cell with trypsin.
• Trypsin removed leaving a residual film.
• Incubation (37 °C for 30 min).
• Cell rounding up after incubation.
• Resuspension of the cells in the medium.
• Reseeding of cells.
• Confluent stage of monolayer.

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For the majority of continuous cell lines, the seeding concentration for subculturing lies between 1 × 10⁴ to 5 × 10⁴ ml. However, for developing a new culture, the initial concentration should be high and should then be reduced to meet the culture's requirements.

Most cell lines are grown as monolayer adherent cells, which grow only on the surfaces of culture vessels, however, certain cells are not adhesive, such as cells derived from leukemic tissue. Moreover, certain cells do not require support for their growth. These cells can be mechanically kept in suspension, and such cultures are referred to as suspension cultures. Transformed cells are usually subcultured using this method. Suspension culture of animal tissues is similar to the method used to subculture bacteria or yeast. There are a number of advantages to suspension cultures over monolayer cultures:

• Bulk production or production in mass can be achieved.
• The cultured cell has access to nutrition from all directions.
• Easy to maintain.
• Frequent replacement of medium is not required.
• The lag period is short.
• The process of propagation is fast.
• Scale-up is convenient.
• Trypsin treatment, or any other enzyme treatment, is not required.

Similar parameters have been reported for suspension cultures as for monoculture, i.e. culture density, fluctuations in pH, schedule timings, the purpose of the subculture, etc. During this process cells are suspended in a culture flask (figure 1.6) which contains culture medium. In the stirred flask technique, the medium is continuously stirred with the help of a magnetic pendulum, in order to offer homogeneous stirring and to avoid aggregation. This magnetic pendulum is allowed to rotate at the base of the flask and the suspension of cells should be regularly monitored for contamination, aggregate formation or any signs of deterioration.

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1.10.1. Cell synchronization

Synchronized cells have the same growth rate in all generations, whereas unsynchronized growth means different growth rates of cells, as shown in figure 1.7. The cell culture has to be synchronized so that the cells will be at the same phase at the same time, which makes it easier to determine the growth rate. Cell synchrony is essential to study the development of cells through the cell cycle, which needs to be monitored at periodic intervals. Several methods have been introduced to achieve cell synchronization. These approaches are broadly divided into two categories:

• Cell synchronization by physical means.
• Cell synchronization by chemical means.

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Cell synchronization using physical methods is more effective than using chemical methods, as the latter can cause toxicity to the cells. Deprivation of nutritional resources (part of the chemical approach) cannot be utilized to synchronize transformed cells. As the cell cycle is composed of different development or growth phases, these determine the synchrony of the cells, and more synchrony can be obtained at the first cycle than at the second or third cycles.

1.10.2. Cell synchronization by chemical means

1.10.3. Cell synchronization by physical means

Algal extracts contain high amounts of potential secondary metabolites which can be utilized to either elicit or promote growth of cells under in vitro conditions. These metabolites can be utilized in media to further increase the growth of the cells. Our recent research on the red algae, Porphyra vietnamensis, found among its diverse chemical compounds some with significant pharmacological properties [31–50]. Such types of algae can be utilized to elicit the growth of animal cells. Shinohara et al observed that algal phycocyanins are responsible for the growth of human cells in culture [50]. In their study, growth-promoting substances derived from blue-green algae, Synechococcus elongatus var., were separated to produce a biliprotein fraction that promoted the growth of RPMI 8226 cells; allophycocyanin was found to be more active than phycocyanin.

Tissue culture is the art of growing cells outside a living body. As we have already discussed the historical background and current innovations of the field of biotechnology in the first two volumes, it is well understood that there are direct and indirect relationships between the developmental biology of plant and animal cells [47]. There are certain challenges involved in culturing both plant and animal cells, such as exhaustion of nutrients in the growth medium, apoptotic/necrotic cell accumulation, cell cycle arrest (or senescence) due to intercellular communication or contact inhibition, etc, which should be investigated further [47]. Various approaches can be utilized to manipulate cell cultures of both plant and animal cells. Subculturing is a common practice which is adopted to replace old medium with new, nutrient enriched, medium. Subculture can also be used to prevent the major problem of senescence. This involves transferring a small number of cells into a new culture dish. The animal–plant co-culture system has not been explored due to its greater vulnerability to contamination of the culture. There is, however, scope to maintain suitable aseptic conditions and encourage a co-culture system to further study the impacts of their growth on each other [47].

The success of animal tissue culture products depends on their efficacy, cost effectiveness and the potential for scale-up. Recent and current advances in tissue culture science have enhanced the complexity in the design of biomaterials which have either been proposed or utilized to grow animal cells [51]. This complexity generally increases the difficulty for manufacturing industries in designing suitable fabrication techniques. It is worth noting that most of the features that are suitable for designing biomaterials originate in the structure and function of plants [51]. Several investigations have shown that decellularized plant tissues can be utilized as a suitable scaffold for the culture of human cells. It has been observed that through an approach of simple biofunctionalization it is possible to achieve the adhesion of human cells on various sets of plant tissues. The increased water transport efficiency and hydrophilicity of plant tissues facilitate increases in cell number over prolonged periods of culture [51]. In addition, animal cells are able to adapt well to the microstructure of plant frameworks without breaking any physiological conditions. This results in perfect cell positioning and formation of a perfect pattern over the feeding layer of plant cells. This supportive plant tissue based micro-framework can be utilized as an alternative potential scaffold for mammalian cells [51].

Recently, there has been a boom in biopolymers of natural and synthetic origin [51–53]. It is essential to understand the possible interactions between cells and these materials in order to develop new materials [48, 54]. When isolated cells from suitable tissues are cultured on a plastic culture dish, this transition of cells from an in vivo to in vitro environment results in the loss of several functions and cells usually begin dedifferentiation, for unknown reasons. Identification of the microenvironmental signals that are responsible for changes in cellular phenotype and function will help in understanding cell behavior under in vitro conditions. Most of the current research that deals with tissue-engineered constructs involves suitable scaffolds that not only act as anchorage cells, but that also help in studying cell behavior and developmental stages in more detail [48, 54]. Several scaffolds have been developed that offer suitable architecture, in particular initial structural integrity and support or a backbone in the form of a matrix in which the cells arrange themselves and form a mass of functioning tissue [51–53]. Several techniques have also been developed, such as three-dimensional matrices for culturing animal cells, to help in understanding how cells probe their surroundings [51–53]. The biomaterial matrix is designed in such a way that it will be able to control the cell position and function inside artificial environments [48, 54]. For most of the materials developed so far, we lack understanding of the effects of the biomaterial or surrounding microenvironment on cell development, behavior and functions.

Physiologically, cells are always surrounded by a sophisticated and dynamic microenvironment which includes the extracellular matrix, growth factors and cytokines, as well as neighboring cells. The extracellular matrix helps in connecting the cells' extracellular matrix proteins through specific cell surface receptors such as integrins [55–57]. Such receptors are responsible for connecting the intracellular cytoskeleton to the extracellular matrix [48, 54]. It is important to understand the interaction between the ligands present in the extracellular matrix and the receptors of the cell. Such an interaction allows multiple intracellular signaling processes that can result in the alteration of cellular behaviors, such as growth, migration and differentiation. Natural extracellular matrices such as collagen offer natural adhesive ligands that encourage cellular connection with integrins. Such biomaterials can be considered as potential resources for engineering new biomaterials [55–57]. One of the major shortcomings of such natural extracellular matrices is our failure to control their physicochemical properties. Several natural biomaterials have been explored recently. In one of the recent innovations in ligand chemistry, a short peptide sequence (arginine–glycine–aspartic acid) which is responsible for cellular adhesion was discovered [55–57]. This peptide can be conjugated with other biologically inert polymers to study their effect on cultured animal cells [55–57]. These peptides, once conjugated over the matrix of inert biomaterial, can initiate cellular adhesion which can further allow researchers to develop suitable matrices on the surface of which these peptides can be conjugated. This approach allows the development of suitable matrices the adhesion potential and chemistry of which can be controlled. Additionally, the incorporation of growth factors in the matrix facilitates their distribution to cells in a controlled fashion. Therefore, cellular adhesion and growth of the cell can be controlled by changing the chemical nature of the polymeric network. One of the most common approaches is functionalization. Growth factors can also be immoblized over polymeric networks to study and manipulate cells. One of the common examples of such immobilization is used in the study of insulin and epidermal growth factor [55–57].

Pattern-immobilization is a more reliable approach as in this process cells are allowed to culture in a matrix with an architecture that produces a three-dimensional structure mimicking the in vivo environment [55–57]. In such a systematic spatial arrangement, growth factor proteins or other proteins, such as those carrying the ligands, are embedded to encourage interaction between cell receptors and ligands. Such artificial frameworks help in the development of tissue through non-diffusion mechanisms, in which the movement of proteins is not dependent on the concentration gradient. This type of stimulation by immobilized growth factors mimics the in vivo environment of membrane-anchored growth factors such as heparin-binding epidermal growth factor, transforming growth factor and tumor necrosis factor. Furthermore, cellular growth can also be enhanced by co-immobilization with adhesion factors and by means of thermosensitive polymers, which allows the development of cells and, most importantly, allows the recovery of the cell through reducing the temperature [55–57].

Three dimensional biomaterials with large pore size (greater than 100 μm) carry a high number of functional units essential for the regeneration of various tissues. Pore size greater than 100 μm is essential for the cell adhesion and proliferation, whereas biomaterials with 325 μm pore size encourage migration of cells through the scaffolds. Scaffolds with pore size less 85 μm showed lowest intensity of cell adhesion and migration. So far, most tissue engineering studies have focused on macro-sized frameworks for cells greater in size than 100 μm (subcellular size) or cellular arrangements larger than 10 μm (cellular size). Such massive structures are required to produce real-sized organ systems [49]. However, to design functional units of tissue, not only are the subcellular and cellular scales required, but also nanostructures, 1–100 nm in size. This type of structural arrangement is essential to control cell behavior, in particular cell–cell interactions, cell–molecular interactions and the cellular environment [49]. The recovery of cell characteristics, in particular structure and function, can only be achieved by reconstruction of the nanostructures of the tissue itself. The current prospects of tissue engineering are very dependent on understanding the interaction of cells with these nanostructures. These tiny structures with three-dimensional arrangements can directly or indirectly affect the cell functions [49]. The trend to fabricate ever smaller structures (called miniaturization), mainly to regenerate the components inside the targeted tissue, has proven a reliable approach for researchers. Thus far, several nano-materials have been developed to mimic native tissues. These nanostructures are tissue-engineered grafts, biomaterial scaffolds that are engineered and then manufactured at the molecular level [49]. Several techniques are available to optimize materials, even at the level of atoms, molecules and supermolecules, 1–100 nm in scale. By means of nanotechnology, various materials or devices can be fabricated or designed to offer a product with high biocompatibility, and most importantly highly predictable biological and physical properties [49]. Animal biotechnology is a broad discipline that includes DNA science, genetic engineering, transgenic science and stem cell research. DNA research involves DNA isolation and screening methods, whereas genetic engineering involves the manipulation of the genetic makeup of an organism to either synthesize a product or alter the character of the organism [58, 59]. Currently, all these fields are being utilized to derive biopharmaceuticals. Moreover, enzyme research, mainly protein engineering, immobilization and biotransformation, has several applications in animal tissue culture science [58, 59].