Types Of Tissue Culture - Types Of Plant Tissue Culture
· Cultures are generally initiated from sterile pieces of a whole plant.
· These pieces are termed ‘explants’, and may consist of pieces of organs, such as leaves or roots, or may be specific cell types, such as pollen or endosperm.
· Many features of the explant are known to affect the efficiency of culture initiation.
· Generally, younger, more rapidly growing tissue (or tissue at an early stage of development) is most effective.
· Several different culture types most commonly used in plant transformation studies will now be examined in more detail.
1. Callus
Explants, when cultured on the appropriate medium, usually with both an auxin and a cytokinin, can give rise to an unorganised, growing and dividing mass of cells.
· It is thought that any plant tissue can be used as an explant, if the correct conditions are found.
· In culture, this proliferation can be maintained more or less indefinitely, provided that the callus is subcultured on to fresh medium periodically.
· During callus formation there is some degree of dedifferentiation (i.e. the changes that occur during development and specialization are, to some extent, reversed), both in morphology (callus is usually composed of unspecialised parenchyma cells) and metabolism.
· One major consequence of this dedifferentiation is that most plant cultures lose the ability to photosynthesise.
· This has important consequences for the culture of callus tissue, as the metabolic profile will probably not match that of the donor plant.
· This necessitates the addition of other components—such as vitamins and, most importantly, a carbon source—to the culture medium, in addition to the usual mineral nutrients.
· Callus culture is often performed in the dark (the lack of photosynthetic capability being no drawback) as light can encourage differentiation of the callus.
· During long-term culture, the culture may lose the requirement for auxin and/or cytokinin.
· This process, known as ‘habituation’, is common in callus cultures from some plant species (such as sugar beet).
· Callus cultures are extremely important in plant biotechnology. Manipulation of the auxin to cytokinin ratio in the medium can lead to the
· development of shoots, roots or somatic embryos from which whole plants can subsequently be produced.
· Callus cultures can also be used to initiate cell suspensions, which are used in a variety of ways in plant transformation studies.
2. Cell-suspension cultures
· Callus cultures, broadly speaking, fall into one of two categories: compact or friable.
· In compact callus the cells are densely aggregated, whereas in friable callus the cells are only loosely associated with each other and the callus becomes soft and breaks apart easily.
· Friable callus provides the inoculum to form cell-suspension cultures. Explants from some plant species or particular cell types tend not to form friable callus, making cell-suspension initiation a difficult task.
· The friability of callus can sometimes be improved by manipulating the medium components or by repeated sub-culturing.
· The friability of the callus can also sometimes be improved by culturing it on ‘semi-solid’ medium (medium with a low concentration of gelling agent).
· When friable callus is placed into a liquid medium (usually the same composition as the solid medium used for the callus culture) and then agitated, single cells and/or small clumps of cells are released into the medium.
· Under the correct conditions, these released cells continue to grow and divide, eventually producing a cell-suspension culture.A relatively -large inoculum should be used when initiating cell suspensions so that the released cell numbers build up quickly.
· The inoculum should not be too large though, as toxic products released from damaged or stressed cells can build up to lethal levels.
· Large cell clumps can be removed during subculture of the cell suspension.
· Cell suspensions can be maintained relatively simply as batch cultures in conical flasks.
· They are continually cultured by repeated subculturing into fresh medium.
· This results in dilution of the suspension and the initiation of another batch growth cycle.
· The degree of dilution during subculture should be determined empirically for each culture.
· Too great a degree of dilution will result in a greatly extended lag period or, in extreme cases, death of the transferred cells.
· After subculture, the cells divide and the biomass of the culture increases in a characteristic fashion, until nutrients in the medium are exhausted and/or toxic by-products build up to inhibitory levels—this is called the ‘stationary phase’.
· If cells are left in the stationary phase for too long, they will die and the culture will be lost.
· Therefore, cells should be transferred as they enter the stationary phase.
· It is therefore important that the batch growth-cycle parameters are determined for each cel
· suspension culture.
3. Protoplasts
· Protoplasts are plant cells with the cell wall removed.
· Protoplasts are most commonly isolated from either leaf mesophyll cells or cell suspensions, although other sources can be used to advantage.
· Two general approaches to removing the cell wall (a difficult task without damaging the protoplast) can be taken—mechanical or enzymatic isolation.
· Mechanical isolation, although possible, often results in low yields, poor quality and poor performance in culture due to substances released from damaged cells.
· Enzymatic isolation is usually carried out in a simple salt solution with a high osmoticum, plus the cell wall degrading enzymes.
· It is usual to use a mix 44 2 : Plant tissue culture of both cellulase and pectinase enzymes, which must be of high quality and purity.
· Protoplasts are fragile and easily damaged, and therefore must be cultured carefully.
· Liquid medium is not agitated and a high osmotic potential is maintained, at least in the initial stages.
· The liquid medium must be shallow enough to allow aeration in the absence of agitation.
· Protoplasts can be plated out on to solid medium and callus produced.
· Whole plants can be regenerated by organogenesis or somatic embryogenesis from this callus.
· Protoplasts are ideal targets for transformation by a variety of means.
4. Root cultures
· Root cultures can be established in vitro from explants of the root tip of either primary or lateral roots and can be cultured on fairly simple media.
· The growth of roots in vitro is potentially unlimited, as roots are indeterminate organs.
· Although the establishment of root cultures was one of the first achievements of modern plant tissue culture, they are not widely used in plant transformation studies.
5. Shoot tip and meristem culture
· The tips of shoots (which contain the shoot apical meristem) can be cultured in vitro, producing clumps of shoots from either axillary or adventitious buds.
· This method can be used for clonal propagation.
· Shoot meristem cultures are potential alternatives to the more commonly used methods for cereal regeneration (see the Case study below) as they are less genotype-dependent and more efficient (seedlings can be used as donor material).
6. Embryo culture
· Embryos can be used as explants to generate callus cultures or somatic embryos.
· Both immature and mature embryos can be used as explants. Immature, embryo-derived embryogenic callus is the most popular method of monocot plant regeneration.
7. Microspore culture
· Haploid tissue can be cultured in vitro by using pollen or anthers as an explant.
· Pollen contains the male gametophyte, which is termed the ‘microspore’.
· Both callus and embryos can be produced from pollen.
· Two main approaches can be taken to produce in vitro cultures from haploid tissue.
· The first method depends on using the anther as the explant. Anthers (somatic tissue that surrounds and contains the pollen) can be cultured on solid medium (agar should not be used to solidify the medium as it contains Culture types 45 inhibitory substances).
· Pollen-derived embryos are subsequently produced via dehiscence of the mature anthers.
· The dehiscence of the anther depends both on its isolation at the correct stage and on the correct culture conditions.
· In some species, the reliance on natural dehiscence can be circumvented by cutting the wall of the anther, although this does, of course, take a considerable amount of time.
· Anthers can also be cultured in liquid medium, and pollen released from the anthers can be induced to form embryos, although the efficiency of plant regeneration is often very low.
· Immature pollen can also be extracted from developing anthers and cultured directly, although this is a very time-consuming process.
· Both methods have advantages and disadvantages. Some beneficial effects to the culture are observed when anthers are used as the explant material.
· There is, however, the danger that some of the embryos produced from anther culture will originate from the somatic anther tissue rather than the haploid microspore cells.
· If isolated pollen is used there is no danger of mixed embryo formation, but the efficiency is low and the process is time-consuming.
· In microspore culture, the condition of the donor plant is of critical importance, as is the timing of isolation.
· Pretreatments, such as a cold treatment, are often found to increase the efficiency.
· These pretreatments can be applied before culture, or, in some species, after placing the anthers in culture.
· Plant species can be divided into two groups, depending on whether they require the addition of plant growth regulators to the medium for pollen/anther culture; those that do also often require organic supplements, e.g. amino acids.
· Many of the cereals (rice, wheat, barley and maize) require medium supplemented with plant growth regulators for pollen/anther culture.
· Regeneration from microspore explants can be obtained by direct embryogenesis, or via a callus stage and subsequent embryogenesis.
· Haploid tissue cultures can also be initiated from the female gametophyte (the ovule).
· In some cases, this is a more efficient method than using pollen or anthers.
· The ploidy of the plants obtained from haploid cultures may not be haploid.
· This can be a consequence of chromosome doubling during the culture period.
· Chromosome doubling (which often has to be induced by treatment with chemicals such as colchicine) may be an advantage, as in many cases haploid plants are not the desired outcome of regeneration from haploid tissues.
· Such plants are often referred to as ‘di-haploids’, because they contain two copies of the same haploid genome.
8. Anther culture
· Obtain two buds at the appropriate stage. This occurs in tobacco when the sepals and the petals in the bud are the same length.
· Holding the bud by the pedicel between the thumb and first finger, dip the entire bud in 95% ethanol for 15 seconds Remove bud and allow excess alcohol to drip off.
· With a pair of sterile forceps, remove the outer layer of tissue, the sepals.
· Next, remove the inner layer of tissue, the petals, exposing the anthers.
· Open the petri dish containing the medium for the induction of haploids.
· Remove each anther from the bud and drop it onto the medium.
· Do not damage the anther or include any filament tissue.
· Repeat for another bud. When finished, seal the plates and place in incubator (25°C).
· In 2–3 weeks examine for somatic embryo initiation. Embryoid-forming cells are characterizedby dense cytoplasmic contents, large starch grains and a relatively large nucleus.
· Embryoids appear opaque among translucent cells.
· Embryoids also exhibit high dehydrogenase activity and can be detected by tetrazolium staining (Dodds and Roberts, 1985).
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