Virus Cultivation: Purposes and Methods
Viruses are obligate intracellular parasites, meaning they are incapable of replication or survival on their own. Unlike bacteria, they cannot be grown in non-living culture media or on agar plates, as they absolutely require living host cells to support their replication machinery. The ability to grow or ‘cultivate’ viruses in a controlled laboratory setting is a fundamental technique in virology and has been crucial for advancements in medicine and public health.
The primary purposes of virus cultivation are threefold. Firstly, it is essential for the isolation and identification of viruses from clinical samples, which is the ‘gold standard’ for establishing the viral etiology of a disease. Secondly, cultivation is indispensable for basic research, allowing scientists to study viral structure, replication cycles, genetics, and the effects a virus has on its host cell (pathogenesis). Thirdly, and most significantly for public health, cultivating viruses in large quantities is a critical step in the preparation and mass production of viral vaccines and diagnostic antigens.
Method 1: Animal Inoculation
Historically, and occasionally still, live, susceptible experimental animals were used for the cultivation of viruses that were difficult or impossible to grow via other means. Laboratory animals such as mice, guinea pigs, rabbits, and primates are selected, ensuring they are healthy and free from communicable diseases. Viruses are inoculated into the animal using various routes, including intraperitoneal, subcutaneous, intracerebral, or intranasal injection. After inoculation, the animals are observed for clinical symptoms of disease or death, which indicates viral multiplication, and the virus is then isolated and purified from the animal’s tissue.
While animal models are valuable for studying disease pathogenesis, immunology, and chemotherapy, they present several significant disadvantages. They are expensive, require difficult maintenance and handling, and raise numerous ethical and animal welfare concerns. Furthermore, some human viruses cannot be grown in animals, or if they do, they may not cause a clinically relevant disease, limiting their utility for specific research or diagnostic purposes.
Method 2: Inoculation into Embryonated Eggs
Inoculation into embryonated hen’s eggs, a technique pioneered by Woodruff and Goodpasture in 1931, is a simpler, less expensive, and widely used method compared to animal inoculation. Fertilized chicken eggs, typically 7-12 days old, provide a sterile, self-contained, and living substrate with a wide range of embryonic tissues and fluids suitable for viral growth and replication. They are particularly important for producing large quantities of viruses for vaccine manufacturing, notably for the annual influenza vaccine.
The site of inoculation depends on the specific virus being cultivated, as each virus has a tropism for different membranes or cavities. Common routes include the Chorioallantoic Membrane (CAM), where viral growth results in visible lesions called pocks (used for poxviruses and herpes simplex virus); the Allantoic Cavity, which is often used for the high-yield production of viruses like influenza and yellow fever; the Amniotic Sac, primarily used for the primary isolation of influenza and mumps virus; and the Yolk Sac, a simpler method for the growth of certain viruses and bacteria like Chlamydia and Rickettsiae. Viral multiplication in the egg embryo can be detected by the death of the embryo, by embryo cell damage, changes in fluids, or by the formation of typical pocks or lesions on the egg membranes.
This method is advantageous because embryonated eggs are readily available, cost-effective, and generally free from contaminating bacteria or many latent viruses, and their immune defense factors are less involved. The inoculation process involves disinfecting the shell, drilling a small sterile hole, injecting the virus into the desired site (e.g., allantoic cavity inoculation usually occurs at 8–10 days of embryonation), sealing the hole, and incubating the egg for several days before harvesting the virus.
Method 3: Cell Culture (Tissue Culture)
Cell culture, also known as tissue culture, is the most versatile and widely used method today, and is considered the ‘gold standard’ for virus isolation in modern diagnostic virology. It involves growing cells *in vitro* (in glass or treated plastic vessels) under controlled conditions using a suitable nutrient medium. The cells are grown either in suspension cultures or, more commonly, as a **monolayer**, a single, confluent layer on the surface of the vessel, which they achieve through anchorage-dependent division until contact inhibition is reached.
Cell cultures are broadly classified into three types. **Primary cell cultures** are derived directly from animal or human organs or tissues (e.g., monkey kidney cell culture) and are able to grow for only a limited number of divisions (typically 5-10 times) before they undergo senescence. They are used for the primary isolation of a broad spectrum of viruses due to their varied cell mosaic. **Diploid cell strains** are of a single cell type, usually human fibroblasts (e.g., MRC-5), that retain their normal chromosome number and can be passaged for a maximum of 50 times. They are used for the isolation of some fastidious viruses and the production of certain viral vaccines. **Continuous cell lines** are derived from cancerous or transformed cells (e.g., HeLa, HEp-2) and have acquired the ability to divide indefinitely, providing a stable, readily available source of cells for various diagnostic and research purposes, though they may have altered characteristics.
Once the host cells are cultured, the viral sample is inoculated. Viral growth is typically detected by observing the **Cytopathic Effect (CPE)** under an inverted microscope—characteristic morphological changes in the host cells like rounding, lysis, inclusion bodies, or syncytia formation. The CPE is crucial for preliminary identification. Cell culture is more convenient and cost-effective than using animals, but its disadvantage lies in the necessity for trained technicians and specialized, sterile facilities, which is why clinical specimens are often sent to central laboratories for testing.
Interconnections and Comprehensive Significance
In addition to microscopic observation of CPE, virus growth can be quantified using assays. The **plaque assay**, a standard procedure, involves inoculating a viral dilution onto a cell monolayer, overlaying it with a semi-solid medium to restrict spread, and staining the cells. Each infective particle gives rise to a clear area of cell death (a plaque) against the stained background, allowing the infectivity titer to be expressed in Plaque-Forming Units (PFU) per milliliter. Other detection methods include transformation assays for oncogenic viruses and serological techniques like haemagglutination assay (in eggs) and Enzyme Immunoassays (EIAs).
The choice of cultivation technique—animal inoculation, embryonated eggs, or cell culture—is governed by the specific virus and the purpose of the study (e.g., diagnosis, vaccine production, or basic research on replication). While in vivo systems like animal models and embryonated eggs still play specialized roles, the in vitro system of cell culture has become the dominant technology, providing a versatile, controlled, and sensitive environment essential for the advancement of modern virology.