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18.2 Plant Tissue Culture 351
Zinc: Zinc is added in the medium in the form of zinc sul-
fate in the concentration of 5–70 . Zinc is absorbed as
a divalent cation Zn2+. Zinc influences the activities of
carbonic anhydrase and hydrogenase, as well as the production of cytochrome and the stabilization of ribosomal fractions. Zinc triggers plant enzymes that are
involved in protein synthesis, glucose metabolism,
membrane integrity maintenance, and auxin production
control. Zinc is also necessary for the production of tryptophan. It also plays an important part in the creation of
Auxin. Plants deprived of zinc exhibit shortened internodes and smaller leaves [23, 24].
Boron: It is added in the form of boric acid in the concen-
tration of 25–100 µ. It is necessary for phenolic acid
metabolism. Boron regulates the activities of phenolase
enzymes and is thus involved in lignin biosynthesis. It is
required for plasma membrane integrity and functioning. Boron is essential for meristematic activity to be
maintained because it is involved in the production of
uracil, which is required for RNA synthesis. Boron deficiency causes the demise of shoot tip meristems. In the
absence of boron, cell division is inhibited due to reduced
nuclear RNA production [25, 26].
Copper: With a concentration ranging from 0.1 to 1.0 µ,
copper is introduced to the medium as copper sulfate
(or, on rare occasions, cupric chloride, or cupric nitrate).
The cytochrome oxidase system, like many other enzyme
processes, depends on copper [27, 28].
Cobalt: Plant physiologists do not consider cobalt to be an
essential component; however, it is found in many of the
most popular culture mediums. It is added into the
medium at a 0.1 µ concentration. Cobalt is a vitamin
component; however, in plant tissue cultures, it has
B
12
no stimulatory effects on growth or morphogenesis [29].
Chloride: For plant development, chloride ion is required.
The medium is supplemented with chloride at a concentration of 3–6 µ. The main functions of chloride appear
to be turgor maintenance and balancing abrupt fluctuations in the concentration of free cations such as Mg
+
Na
, and K+. Plants absorb chloride ions best at a pH
2+
that is somewhat acidic [30–32].
Molybdenum: Molybdenum is added to the medium in the
form of sodium molybdate in the concentration of 1 µ.
Molybdenum is found in nitrogen-utilizing enzymes such
as nitrate reductase and nitrogenase. It converts ammonia
from nitrate [33].
Iron: Iron is usually added as ferrous or ferric salts. Iron is
believed to be absorbed by plants in ferrous form. It is generally present in media at a concentration of approximately
1 µ. Iron is utilized by plants for oxidation/reduction processes in the mitochondria, chloroplasts, and peroxisomes.
It is also found in the protein ferredoxin, which functions
as an electron transporter during photosynthesis. Because
it precipitates in alkaline pH and produces insoluble compounds, iron is difficult to feed in-vitro. In order to prevent
this, Fe is provided as a chelated EDTA (ethylene diamine
tetra acetic acid) complex. The use of chelating agents,
which bind metal ions, increases the stability and availability of iron to plant tissues over a larger pH range (up to
a pH of 8.0) [34–36].
Iodine: Iodine is not considered as an essential compo-
nent, but it is frequently added to the medium in the
form of potassium iodide or methylene iodide at a concentration of 5 . Iodine has been demonstrated to
increase callus and root development in-vitro [37, 38].
Nickel: Even though nickel is crucial for plant metabo-
lism, it is poisonous to most plants if present in large
amounts. Plant transpiration and photosynthesis are
inhibited by high nickel concentrations. In the form of
nickel chloride or nickel sulfate, it is given to the medium
up to 0.1 m concentration. Several metalloenzymes
have nickel as a component. Plants with nickel deficits
have decreased enzyme activity and have problems
assimilating nitrogen [39].
Silicon: Although silicon is usually present in significant
quantities in soils and may be readily absorbed by plant
roots, it is not thought to be a necessary nutrient for
plants. It has been found that silicon helps plants thrive
and reduces biotic and abiotic stress. Typically, it is
added as sodium silicate up to a concentration of 0.4 .
Plant osmotic adjustment may involve silicon. It can
provide plant cell walls rigidity and roughness. Silicon
supplementation improves organogenesis, physiological
traits of leaves, protects cells from metal toxicity,
increases tolerance to low temperature and salinity, prevents oxidative phenolic browning, and decreases hyperhydricity in a variety of plants [40].
(ii) Carbon and Energy Source
Since in-vitro plant cells, tissues, and organ cultures are not
fully autotrophic (capable of fixing carbon through photo-
,
synthesis), carbohydrates in the culture medium are required
to maintain osmotic potential and to provide energy and carbon for various energy-intensive developmental processes
such as organogenesis, root induction, embryogenesis, and
shoot proliferation. Due to the growth occurring under tissue-culture settings unfavorable for photosynthesis, photosynthesis is insufficient, making carbon sources important.
Carbohydrates are used as a source of carbon. Sucrose,
which is employed at a concentration of 2–5%, is the most
popular source. Good growth is also supported by glucose.
Occasionally, other sugars including glycerin, lactose, galactose, maltose, sorbitol, and raffinose are also employed.
Depending on the genotypes and particular stages of growth,
352 18 Medicinal Plant Biotechnology
different carbon sources are utilized in culture media. Young
embryos require a high sugar content. Growth and development generally rise with sugar concentration until an optimum is attained, then decline at high concentrations, e.g.
the Callus culture of easter lilly at 5% sucrose concentration
was found superior to the 2 and 10% sucrose concentration.
When autoclaved, sucrose in the culture medium is normally hydrolyzed completely or partly into the components
glucose and fructose. Cultures grow better on autoclaved
sucrose media as ready glucose and fructose are available for
their growth [41–42].
(iii) Organic Supplements
Vitamins: Plants are a major source of essential vitamins
for animals and humans. Vitamins are produced
endogenously by plants, and they are mostly used as
catalysts in various ways or as necessary intermediates
in biochemical activities. In-vitro grown plant tissues or
cells produce extremely little or no vitamin production.
Vitamins must therefore be added to the medium for
tissue to develop. Depending on the plant species and
culture type, different plant cells have different requirements for vitamin concentration. Nearly all plant tissue
cultures require thiamine, often known as vitamin B
or aneurin. Thiamine’s function in plants is diverse and
serves as a cofactor in enzymatic reactions. Thiamine
pyrophosphate is an important coenzyme in glucose
metabolism and has a direct role in the production of
certain amino acids. Concentration range of 0.1–
1
10 mg L
) is used for Thiamine. In some culture mediums, other vitamins are provided as well, including
nicotinic acid (niacin), pyridoxine (Vit B6), folic acid,
biotin, ascorbic acid (vitamin C), and vitamin E
(tocopherol). The concentration of vitamins in the culture medium is very low. E.g. Murashige and Skoog
(MS) medium contains thiamine, nicotinic acid, pyridoxine, and myoinositol at 0.1, 0.5, 0.5, and 100 mg L
1
respectively [43].
Amino Acids: Plant cells can ordinarily synthesize all the
necessary amino acids for metabolic functions; however,
the addition of some amino acids can help to accelerate
the growth of plant cells or tissues. Nitrogen provided by
amino acids is more easily digested by plant cells than
inorganic nitrogen sources. Casein hydrolysate (0.25–
1
1 g L
), cysteine (10 mg L1), glutamine (8 m),
L-arginine (10 mg L1), asparagine (100 mg L1), glycine
1
(2 mg L
), and L-tyrosine (100 mg L1) are the common
sources of organic nitrogen for culture media. Threonine
and valine sometimes reduce ammonium utilization by
inactivating glutamate synthase.
Complex Organic Supplements: Complex organic sub-
stances such as fruit juice (banana, papaya, orange,
tomato, and watermelon), fruit pulp, yeast extract, malt
extract, and casein hydrolysates are occasionally added
to the culture medium.
(iv) Solidifying or Gelling Agent
Gelling agents are used to prepare the solid medium. The
most commonly used gelling agent is agar (0.5–1.0%)
because it is tolerant to enzymes and has no interaction
with medium elements. Gellan gums and agarose, a pure
kind of gel, are also employed. It is exceedingly stable and
does not precipitate in the presence of some cations, unlike
calcium-containing alginates. It withstands heat treatments
quite well, even at temperatures above 100 °C, allowing for
effective sterilization. Because of agar’s exceptional reversibility, it may be repeatedly gelled and melted without losing
any of its original qualities. Agar is a seaweed polysaccharide, whereas gellan gum (such as phytagel and gelrite) is a
bacterial polysaccharide [44].
(v) Plant Growth Regulators/Phytohormones
The term “phytohormones” often refers to endogenous
(naturally occurring) growth agents, whereas the term
“growth regulator” generally refers to artificial growth
agents. An organic substance known as a plant hormone is
1
synthesized in a particular part of the plant and then transported to another, where it triggers a physiological response
in very low concentrations. Plant growth regulators are a
class of chemical substances that influence how plant cells,
tissues, and organs develop and differentiate. From plant to
plant, the concentration and ratio may differ. The hormones found in plants are classified as either promoters
(auxins, gibberellin, and cytokinin) or inhibitors (ethylene
and abscisic acid). Cell division can only begin and be
maintained with the help of phytohormones or their synthetic analogs, either alone or in combination [45].
Auxin: The first plant hormones to be identified were aux-
,
ins. Auxin means to grow, it is derived from the Greek
word auxein. For a group of hormones that are involved
in growth responses (induce callus division of cells,
elongate cells, etc.) auxin is a generic term. It is made in
the root and shoot apices, and it moves from the apex to
the zone of elongation. The coordination of numerous
growth and behavioral processes in plants is crucially
aided by auxin. Fritz Went, a Dutch scientist, was the
first to notice this behavior in auxin [46]. Indole-3-acetic
acid (IAA) is the most significant auxin present in plants.
Indole-3-butyric acid, indole acetonitrile, phenylacetic
acid, and 4-chloro-indoleacetic acid are more auxins
that have been identified from plants. These are probably in-vivo transformed to IAA.
Cytokinins (Kinetin): Cytokinin promotes cytokinesis
(cell division), which leads to its given name. Kinetin
(Skoog and Miller 1950), also known as 6-furfuryl
18.2 Plant Tissue Culture 353
aminopurine, was the first identified as cytokinin.
Zeatin, derived from corn (Zea mays), is the most common form of naturally occurring cytokinin in plants
today. Cytokinin levels are high in the shoot apex, root
tip, and immature seeds. Some plant pathogenic bacteria, such as Agrobacterium tumefaciens, also produce
cytokinin. Cytokinins are N6-substituted purine derivatives that occur naturally. Based on the structure of
the N6-substituent, cytokinins are classed as isoprenoid or aromatic. Cytokinins are of two types, (i) adenine type cytokinin e.g. kinetin, zeatin, and 6 benzyl
aminopurine (BAP; (ii) phenyl urea type cytokinin,
e.g. diphenyl urea and thidiazuron. The most common
cytokinins discovered in higher plants are isopentenyladenine (iP), zeatin (Z), and dihydrozeatin (DZ).
The amount of shoot and/or root production in tissue
culture depends on the auxin-cytokinin ratio being utilized in the culture media. A high auxin-to-cytokinin ratio
promotes root growth, while a high cytokinin-to-auxin
ratio promotes shoot growth. Callus development is facilitated by both hormones at intermediate levels [47].
Gibberellins (GAs): Elichi Kurosawa, a Japanese scien-
tist, discovered in 1926 that infected rice seedlings of the
fungus Gibberella fujikuroi grow higher and become
extremely thin and pale. From the infected seedlings, an
active compound was identified and given the name gibberellin. The first gibberellin to have its structural characteristics determined was gibberellic acid, also known
as GA3. A total of 136 GAs have currently been found in
plants, fungi, and bacteria. Gibberellins are found in
plants in two forms: free gibberellins and bound gibberellins. Gibberellin-glycosides are the most common form
of bound gibberellins. The bioactive GAs are GA1, GA3,
GA4, and GA7 [48].
Abscisic Acid: It is a plant hormone that controls the
early stages of embryo development, abscission, and
dormancy. It encourages morphogenesis and is necessary for the healthy growth and development of somatic
embryos. Scientists isolated a chemical from cotton
balls in the 1960s that was subsequently identified as an
abscission factor (abscisin II). During the same period,
Cornforth et al. (1965) were successful in isolating a
compound called dormin from sycamore leaves, which
was shown to be involved in bud dormancy. Later, it was
found that they were the same compounds and it was
renamed as abscisic acid [49]. A high concentration of
abscisic acid in the leaves leads the stomata to close
allowing the plant to conserve water during droughts.
Roots control ion and water uptake. It promotes seed
maturity by stimulating the production of storage proteins in developing seeds.
Ethylene: Ethylene is the only naturally occurring plant
growth hormone in gaseous form. All higher plants produce ethylene from methionine in nearly all tissues. The
synthesis of ethylene usually occurs under both abiotic
and biotic stresses.
18.2.3 Plant Tissue Culture Laboratory Requirements
Regardless of the precise aim, several basic facilities must
be available in each laboratory in which tissue culture
procedures are carried out. A general washing space, sterilization, and storage area, media preparation, environmentally controlled incubators, transfer area for aseptic
preparation, culture rooms, plant growth chambers, and
observation/data gathering area are frequently included
(Figure 18.2).
Washing Area: In the washing room, there should be
large, acid- and alkali-resistant sinks, flowing hot and
cold water, brushes of various sizes and shapes for cleaning glassware, access to deionized/distilled water, and
draining boards and racks. Several plastic or steel buckets are required for soaking and washing the culture
vials and other laboratory glass wares. Before cleaning, a
bucket with a cover is also necessary for disposal of discarded or diseased media. Drying ovens or racks, pipette
washers and dryers, acid baths, and storage cabinets are
required in the washing room. The washed labware
o
should be placed in a hot air oven at 70
C for drying and
stored in a dust-proof cupboard.
Washing room
Change room
Figure 18.2 Layout of tissue culture laboratory.
Dark room
Storage room
for glasswares
and chemicals
Media preparation room
Aseptic area for
transfer
Lab work and
data collection
Aseptic are for
shakers and
storage
(controlled
room)
354 18 Medicinal Plant Biotechnology
Media Preparation: The area used for media preparation
should have enough bench space and space for the storage
of the chemicals, culture vessels, labware, and equipment
(pH meters, hot plates, balances, and water baths) is
needed for media preparation and dispensing. Additionally
requirementsare a microwave or conventional oven, laminar air flow, vacuum pump, autoclave, distilled water unit,
dissecting microscope, shaker, magnetic stirrer, and refrigerator or deep freezer. Chemicals of an analytical grade
ought to be used while producing media. High-quality
pure water is required while producing media. Tap water
should not be used as it contains impurities such as organic
and inorganic compounds, dissolved gases, particulate
debris, and microorganisms.
Sterilization Area: Plant sources (explants), culture
medium, labware, laboratory environment, and instruments are sources of microorganisms and contaminants.
The development of plant tissue will be substantially
hampered by the growth of microorganisms in the culture media. Sterilization is therefore a crucial component of plant tissue culture. Glassware may be sterilized
using an autoclave (121 °C and 15 psi for 15 minutes of
moist heat sterilization) or a hot air oven (160–180 °C for
three hours of dry heat sterilization). It is possible to
repeatedly autoclave plastic products made of polypropylene, polymethyl pentene, polyallomer, Tefzel ETFE,
and Teflon FEP. By using a filter sterilizer (thermolabile
chemicals) or an autoclave, culture media can be sterilized. The culture vials that hold the media should be
closed with a cotton plug, aluminum foil, and sterilized
at 121 °C and 15 pressure for 15–40 minutes. Some substances lose their activity or break down when autoclaved, such as sucrose (which breaks down into glucose
and fructose), gibberellic acid, which loses 90% of its
action, and vitamin B
(which breaks down into pyrimi-
1
dine and thiazole). Such solutions should be sterilized
using a filter sterilization technique with bacteria-proof
membranes with pores between 0.22 and 0.45 m in
size. Autoclaving is used to sterilize devices used for
aseptic manipulation such as needles, forceps, and spatulas, whereas 95% ethanol and flame heating is used to
sterilize inoculum loops. Surface sterilization is the preferred method for explant sterilization because moist
heat sterilization and dry heat sterilization are not suitable due to the fragility of explants and the possibility of
viability loss. Surface sterilization comprises washing
with detergent, sterilant/disinfectant and washing with
sterile distilled water. The sterilant or disinfectant can be
harmful to the plant tissue, therefore the disinfectant, its
concentration and treatment duration should be chosen
to minimize the tissue death. Bromine water, calcium
hypochlorite, silver nitrate, hydrogen peroxide, ethanol,
mercuric chloride, and sodium hypochlorite are a few
disinfectants that are used for sterilizing plant material.
Transfer Area for Aseptic Manipulation: To keep the
area dust-free, there should not be any windows or ventilation in the transfer area. There should be an automatic
door closing in the room. The appropriate material
should be installed on the floor to make cleaning easier.
The simplest type of transfer area is an enclosed wooden/
plastic box. The upper half of the box’s side walls are constructed of huge glass sheets. This chamber is sterilized
by UV light and the floor is sterilized by ethanol. The controls to activate the UV light are located outside the
chamber, allowing the lamp to be safely turned on and
off. This box can be placed on a small table. This box is
suitable only for a few transfers. The most ideal, practical,
and dependable tool for aseptic transfer is laminar airflow or a biosafety cabinet. Several small bower motors
are used in laminar air flow to blow air that passes
through high-efficiency particulate air (HEPA) filters.
These filters exclude material that is larger than 0.3 m.
Culture Rooms: A culture room refers to a space used to
maintain or incubate a culture under conditions of controlled temperature, humidity, and light. This space
needs to be kept clean. There should be positive air pressure in the room or an overhead air curtain at the
entrance. Both of these things will clear away surface
dust. To keep this space dust-free, it should be devoid of
windows and ventilation. The internal light cycle should
not be disrupted by windows; hence, they should be
avoided. Additionally, there are shelves in this area.
Shelves can be made up of glass. Plant growth chambers
are also used to maintain controlled conditions of temperature, light, and humidity.
Air coolers or heaters are employed in the culture
room or plant growth chamber to keep the temperature
at 25 ±2 °C or to set at a specific temperature depending
on the plant’s requirement. Relative humidity needs to
be kept at or above 50% in most of the cases. A hygrometer and thermometer are mounted on the wall to measure the relative humidity and temperature in the culture
room. Cultures may thrive in both light and darkness.
Fluorescent lamps are included in each culture rack for
providing light of specific ‘“Lux” intensity. Since they
generate uniform light intensity, white, fluorescent
lamps with electronic ballasts are usually used. Specific
conditions for light:dark can be maintained using the
light panel controls. Blue and red light are very important for plant growth. For dark culture incubation, racks
with black curtains function effectively. A shaker for
suspending culture should also be included in the culture room. Shakers with speed, light, and temperature
controls should also be available. A generator backup or
18.2 Plant Tissue Culture 355
UPS should be provided which will be helpful in power
failure.
Observation or Data Collection Area: The cultures
should be observed at regular intervals, and they must be
collected in the aseptic area. Data can be collected in the
culture room. If any microscopical work is required,
then it should be done in the lab space.
18.2.4 Micropropagation
Only mitotic cell division occurs during vegetative plant
propagation techniques like cutting, budding, grafting, etc.
A clone is the offspring produced via vegetative multiplication of a single plant. Micropropagation refers to the tissue
culture-based in-vitro clonal propagation of plants.
Explants from a mother plant that is robust and healthy are
chosen for the process. Explants can be any plant component, including a leaf, bud, apical meristem, or root. The
primary goal of micropropagation is to create plants with
identical genotypes to the parent plants. The following
three pathways help to do this [50–54].
i) Axillary bud proliferation (Proliferation from pre-
existing meristems)
ii) Organogenesis, and
iii) Somatic embryogenesis
ferentiation. Some species do not grow new shoots from
their axillary buds. In these cases, the explant’s shoot
bud is excised and broken into tiny pieces to create nodal
explants, which are then subcultured to start a fresh
cycle of micropropagation. This is known as a singlenode culture.
Organogenesis: It is characterized as the formation of
organs such as branches, roots, and flowers from either
an explant or a callus culture. Organogenesis is classified
into two types: direct organogenesis and indirect organogenesis. Direct organogenesis, also known as adventitious regeneration, is the direct formation of organs on
an explant, skipping the callus stage, such as shoots,
roots, flowers, buds, etc. Shoots and roots are produced
in tissues that do not normally produce these organs
(Figure 18.3). In the indirect organogenesis process, the
explant initiates the formation of the callus from which
branches and roots grow (Figure 18.4).
Somatic Embryogenesis: A zygote typically develops
after a sperm has fertilized an egg. The zygote subsequently undergoes zygotic embryogenesis to become an
embryo. The process by which embryos are formed from
somatic cells, organs, or tissues is known as somatic
embryogenesis. These embryos are also called as nonzygotic embryos. [55-60].
Proliferation from Pre-existing Meristems (axillary
bud proliferation): This technique uses an already pre-
sent meristem to start an in-vitro culture (such as a
shoot-tip or nodal explant). The shoot has already been
differentiated when employing the proliferation of axillary bud from a bud or node; the only thing that remains
to do is to complete the shoot’s elongation and root dif-
Cytokinin
Explant
Figure 18.3 Direct organogenesis.
Cytokinin Cytokinin
Explant
Callus
Shoots Formed
Shoots Formed
18.2.5 Types of Culture
A sterile piece of the entire plant is used to initiate cultures.
These pieces, known as explants, could be specific cell
types, like pollen or endosperm, or they could be parts of
organs like leaves or roots. The general process of plant tissue culture is summarized in Figure 18.5. It includes the
Divide
Shootlets
Divide Auxin
Shootlets
Auxin
Rooted
Plantlets
Rooted
Plantlets
Figure 18.4 Indirect organogenesis.
356 18 Medicinal Plant Biotechnology
Selection of source plant
Incubation under aseptic conditions
Initiation of callus
Transfer of Plantlet to green house or field
Figure 18.5 General process of plant tissue culture.
Isolation of explant
Subculturing
proliferation of axillary shoots, initiation of aseptic culture,
selection of explant, rooting, and transfer of the plant to
the field or greenhouse [61–64].
18.2.5.1 Callus Culture
The word callus is derived from the Latin word Callum,
which means hard. A callus is an unorganized tissue mass
that grows on a solid substrate and occurs spontaneously
on plants in response to wounding. The callus cells are parenchymatous in nature. A single differentiated cell may
make callus, and numerous callus cells are totipotent,
capable of generating a full plant. Plant regeneration from
calli is conceivable by de novo organogenesis or somatic
embryogenesis. Callus is divided into categories depending
on macroscopic properties. Friable (poorly associated cells
and crumpy appearance) or compact callus (cells are
strongly associated with each other). A liquid cell suspension culture is frequently started with a friable callus. A
friable callus is a source of protoplasts. The production of
callus is influenced by genotype, nutritional medium composition, and physical growth factors. The size and shape
of the explants are also crucial considerations. Because of
cell proliferation, nutrient depletion, and medium drying,
callus cultures should be subcultured every three to five
weeks.
18.2.5.2 Cell Suspension Culture
To establish a cell suspension culture, a friable, soft callus
is transferred to a liquid medium with the same callus culture’s original composition. It is set up on a shaker and
shaken at speeds ranging from 90 to 150 rpm. Shaking
helps to distribute the cells in the medium and to supply
4
oxygen. Inoculating 10
cells per mL is required to create a
well-growing cell suspension culture; otherwise, the cells
might not divide. Suspension cultures develop more
quickly than callus cultures, and they should be subcultured once a week. The suspension cultures are broadly
Surface sterilization of explant
Inoculation of explant in nutrition media
Plantlet regeneration
grouped as follows: batch cultures; continuous cultures,
and immobilized cell cultures.
Batch Culture: When cells are cultivated in a fixed vol-
ume of nutritional culture medium, it is known as a
batch culture. In this type of culture, cell growth and
division increase the biomass of the cell suspension until
a culture medium constituent becomes restrictive, at
which point growth stops. The growth cycle of the cells
in batch culture goes through five stages: (i) the lag
phase, during which the division of cells occurs, (ii) the
log or exponential phase, during which cell division
occurs at the fastest pace, (iii) linear phase, during which
the rate of cell expansion quickens but cell division
decreases, (iv) deceleration phase, during which cell
growth and division rates slow down, and (v) stationary
phase, during which both cell size and number are
constant.
Continuous Culture: The cell population is kept stable by
replacing a part of the old or spent media with fresh
media regularly in continuous culture.
Immobilized Cell Culture: A suitable substance, such as
calcium alginate gels and agarose, or a membrane or
stainless-steel screen, can be used to enclose plant cells
and cell groups. Cell-containing gel beads or cells can be
placed in an appropriate column, such as a membrane or
wire cloth column. To give cells nourishment and aeration, the liquid medium continues to flow through the
column. Cells that are immobilized have different cellular physiologies compared to cells in suspension
culture.
18.2.5.3 Single-cell Culture
From cell suspension culture or friable callus culture, a single cell can be separated. A friable callus is immersed in an
agitated liquid media. The callus becomes detached by
movement and the cells disperse. Filtration is used to
18.2 Plant Tissue Culture 357
remove cell clumping from the resulting cell suspension.
To separate the single cell from the pellet, the resultant filtrate is centrifuged. A solid or liquid media can be used to
cultivate the isolated single cell. Single-cell culture is crucial for the fundamental investigation and analysis of
mutations.
18.2.5.4 Protoplast Culture
A protoplast is a plant cell that does not have a cell wall.
Because the cell wall has been eliminated via mechanical
or enzymatic means, it is known as a “naked plant cell.”
Almost all plant components, including the roots, leaves,
fruits, tubers, endosperm, and pollen, can yield protoplast.
The aseptic isolation and in-vitro culture of protoplasts are
referred to as protoplast culture.
18.2.5.4.1 Methods of Protoplast Isolation
Mechanical Method: Cells are maintained in an appro-
priate plasmolyticum (13% w/v mannitol) and the cell
wall is cut with a sharp knife before being deplasmolyzed. During cutting, some protoplasts may be damaged, while intact ones with properly cut cell walls will
emerge.This approach may be used to extract highly
vacuolated cells from protoplast storage tissues like radish roots and onion bulbs.
Enzymatic Method: Cellulose, hemicelluloses, and pec-
tin are the three main elements that make up a cell wall.
Cellulase, hemicellulase and pectinase break down the
cell wall’s cellulosic, hemicellulosic and pectin constituents, respectively. pH 4.7–6.0 should be maintained
since enzymes are pH-dependent. There are two methods for performing the enzymatic isolation: (i) sequential (two-step) method that involves treating plant tissues
with pectinase before treating them with cellulase and
hemicellulase to form protoplast, and (ii) direct (onestep) approach in which plant tissues are plasmolyzed in
the presence of a mixture of cellulases, hemicellulases
and pectinases, resulting in simultaneous cell division
and protoplast collapse.
18.2.5.4.2 Purification of Protoplast
Purification and separation of undesired material are crucial because during protoplast isolation certain cell organelles may be released and cell wall debris may also remain
with the protoplast. Filtration, sedimentation, and washing
are used in combination to complete the process. To remove
undigested cell clumps, the suspension is passed through a
nylon mesh (50–100 m) or a metal sieve to filter out the
protoplasts and other debris. After adding an appropriate
volume of osmoticum to the filtered protoplast-enzyme
solution, the combination is centrifuged (at 50–100 g for
five minutes) to pellet the protoplasts, which will then settle to the bottom. By using a Pasteur pipette, the protoplast
pellets are pulled into another centrifuge where they
receive three washings. The protoplast pellet should be
resuspended in new, fresh media. Gradients (mannitol,
sorbitol) may be used to separate low-density protoplast
from other cell debris, allowing the protoplast to float while
the waste settles to the bottom. The isolated protoplast are
washed after removing the supernatant fluid.
18.2.5.4.3 Protoplast Viability
Protoplast viability is estimated using the following
methods.
i) Fluorescein Diacetate: Viable protoplasts accumulate
fluorescein diacetate inside their plasmalemma, which
allows fluorescence microscopy to identify it. It is used
at a concentration of 0.01% dissolved in acetone.
Esterases in viable protoplasts break the fluorescein
diacetate (FDA), releasing fluorescein, which fluoresces yellowish-green in five minutes. After about 15
minutes, the FDA separates from the membrane.
ii) Calcofluor White: By recognizing the onset of cell wall
construction, this staining approach ensures protoplast viability. A luminous ring can be seen encircling
the membrane when calcofluor (0.1% w/v solution) in
newly synthesized cell walls attaches to beta-linked
glucosides.
iii) Phenosafranin (0.1%): It identifies dead protoplasts,
which become red when they come into contact with
it. A viable protoplast stays unstained.
iv) Monitoring cells’ oxygen uptake via an oxygen elec-
trode, which reveals respiration, is another method for
determining protoplast viability.
18.2.5.4.4 Protocol for Protoplast Culture
The explants are surface sterilized with an appropriate sterilizing agent and washed in distilled water to remove any
remaining sterilizing residues. The explant is cut into tiny
pieces and given a set amount of time to plasmolyze. The
explant fragments are then kept in an enzyme solution and
incubated for a specified amount of time (16–18 hours).
The protoplast is separated and cleaned using a filtration
and washing combination. Protoplast pellets are suspended
in an appropriate nutritive medium under carefully monitored conditions. Small cell colonies will be visible after
three to four weeks. Within five to six weeks, colonies will
attain a diameter of about 1 mm. They are transferred to an
osmotic-free medium to generate callus when small colonies have formed. Organogenic or embryogenic differentiation of callus takes place, resulting in the production of
plants.
358 18 Medicinal Plant Biotechnology
The protoplast fusion process involves fusing protoplasts
with two different genomes, choosing the required somatic
hybrid cells and then growing new hybrid plants. Protoplast
fusion can be carried out by following techniques:
i) Spontaneous fusion: When protoplasts are isolated for
culture, some of them in proximity fuse spontaneously
to form homokaryons or homokaryocytes.
ii) Mechanical fusion which involves joining two proto-
plasts without utilizing a substance that induces
fusion.
iii) Induced fusion: Fusion-inducing agent (fusogen), such
as NaNO
or polyethylene glycol, is used to fuse proto-
3
plasts from two different species (interspecific fusion)
or from two separate sources that are members of the
same species.
iv) Electrofusion: The protoplast is inserted in a small fusion
chamber that has parallel electrodes made of wires or
plates. To align the protoplast between electrodes, a low
voltage (5–12 amp) is supplied. The voltage is then
increased to cause the protoplasts to fuse where they
have touched, which will result in fusion. It is possible to
manipulate hybrid production and incorporate the
desired characteristics such as disease resistance, cold
tolerance, and nitrogen fixation by protoplast fusion.
Cybrids or Cytoplasmic Hybrids: Cybrids are somatic
hybrids that contain one parent’s nuclear DNA but both
parent’s cytoplasm. The following procedures are
involved in the development of somatic hybrids:
i) Protoplast isolation from two distinct species,
ii) Fusion of two distinct species protoplasts,
iii) Fused protoplast isolation, and
iv) Fertile hybrid plants regeneration from fused
protoplasts.
18.2.5.5 Organ Culture
In this culture, a specific organ is isolated and aseptically
grown in a nutritive medium with a predefined chemical
composition. It does not cause callus formation and grows in
the same way as its intact counterpart. In-vitro culture and
maintenance of an excised organ or full portion of an organ
in a method that allows differentiation and preservation of
an organ’s structure is referred to as organ culture. Based on
the explant, organ culture is categorized into different types
such as root culture, shoot-tip culture, leaf culture, etc.
18.2.5.6 Root Culture
Since the roots are located deep in the soil, the whole
plant’s root tip is unusable. Young seedlings’ root tips are
also unsuitable because of their extreme sensitivity to
sterilants. So, prevention of surface sterilization of the
root tip is very important. Alternatively, root culture can
be initiated from the excised radicle tips of aseptically
germinated seed. The seeds are surface sterilized before
being placed on moist filter paper or nutritional media to
germinate. When seedlings are 20–30 mm long, excise the
8–10 mm long apical tip, place it aseptically into a nutritional medium, and then incubate it.
18.2.5.7 Leaf Culture
It is the in-vitro development and growth of an excised
young leaf or immature young leaf of the shoot apex aseptically on a suitable nutritional medium under controlled
conditions. The growth potential of the young leaf is more
than the matured leaf. The young leaf is excised and surface sterilized or leaves of aseptically grown plants can be
taken. The young leaf from the shoot apex is detached,
after washing and sterilizing it with sterilant (ethanol/
sodium hypochlorite solution), inoculate the excised leaf
on a nutritional medium.
18.2.5.8 Flower Bud Culture
It is the in-vitro development and growth of an excised
flower bud aseptically on a suitable nutritional medium
under controlled conditions. Flowers can be cultured at
different stages of development such as the bud stage, preand post-pollination stage. Pre- and post-pollination stage
flower explants need a simpler medium than the primordial or bud stage.
18.2.5.9 Ovary Culture
It is the formation and growth of pollinated or unpollinated flower-isolated ovaries in a controlled environment
on an appropriate nutritional media. Compared to the ovaries of unpollinated flowers, the ovaries of pollinated flowers need a simple medium. The flowers are washed with
tap water (whether they are pollinated or not), surface
sterilized with an appropriate sterilant (5% sodium
hypochlorite solution), and then washed once more with
distilled water. Gently remove the calyx, corolla, and
anthers without harming the ovaries, then isolate the
gynoecium and grow it on a nutrient medium.. Incubate
the ovaries at the appropriate temperature after inoculating them with nutritive media.
18.2.5.10 Ovule Culture
It is the development and proliferation of ovules that
have been removed from the ovary in-vitro on a suitable
nutritional medium under controlled circumstances.
The nucellus (megasporangium), the female gametophyte (megagametophyte), and the integument, which is
the outer layer, make up the ovule. After collecting
healthy flowers, the ovaries are separated from all other
components, including sepals, petals, and androecium.
18.2 Plant Tissue Culture 359
Wash with distilled water after sterilizing the ovaries
with the appropriate sterilant. Remove the ovules by
breaking the funicles and thereafter incubate them on a
suitable nutritional medium.
18.2.5.11 Embryo Culture
It is in-vitro development and growth of embryos of different developmental stages isolated aseptically from tissues
of ovules, seed on suitable nutritional medium under controlled conditions.
18.2.5.12 Anther and Pollen Culture (Microspore
Culture)
Anther culture: It is the in-vitro development and growth
of excised anthers obtained from unopened flower buds on
a suitable nutritional medium under controlled conditions.
Microspores within the cultivated anther develop into callus tissue or embryoid, resulting in the formation of haploid plantlets by organogenesis or embryogenesis.
Pollen culture (Microspores culture): It is the in-vitro
development and growth of pollen grains (at the uninucleate stage) obtained from the intact anther on a suitable nutritional medium under controlled conditions.
Microspores mature into haploid embryoid or callus tissue, which gives rise to haploid plantlets via organogenesis or embryogenesis without producing male gametes.
Haploid plant production through another culture is
called androgenesis. Androgenesis can be direct or indirect. Microspores create embryoids, which give birth to
plantlets during direct androgenesis. The microspore splits
repeatedly in indirect androgenesis to create a callus tissue
that develops into haploid plantlets.
18.2.6 Synthetic Seed or Artificial Seed
A chemical membrane is put over (encapsulated) somatic
embryos (bipolar structures with apical and basal meristematic areas capable of developing shoot and root, respectively), shoot buds, or any other plant material produced
in-vitro. Such materials behave like seeds when enclosed.
These are referred to as synthetic or artificial seeds. The
synthetic coating serves as a seed coat substitute. Such
seeds resemble beads, and they can germinate and produce
plantlets. Artificial seed coverings can be made from a variety of materials. Among them are agar, polyacrylamide,
agarose, ethyl cellulose, carrageenin, nitrocellulose, and
sodium alginate. Most frequently, sodium alginate is used
[65–70].
The advantages of synthetic seeds are as follows:
1. Compared to the plant’s natural seeds, artificial seeds
are smaller in size.
2. Such seeds are simpler to transport and store.
3. All the seeds are 100% viable.
4. On a suitable substrate, artificial seeds can be made to
germinate consistently.
5. These seeds don’t appear to be dormant.
6. The plant grower has the option of growing the desired
plant at any moment, regardless of season.
7. It is feasible to produce seeds on a large scale from any
type of plant component.
The disadvantages of synthetic seeds are as follows:
1. Since they are temperature-sensitive, artificial seeds
cannot be kept for an extended period.
2. The initial expense of generating artificial seed exceeds
the cost of producing natural seed.
3. Aseptic conditions are necessary for the creation and
germination of artificial seeds. Any divergence will
have an impact on the seeds’ quality and future growth.
18.2.7 In-Vitro Plant Germplasm Conservation
Germplasm is the whole set of genetic material of a species
of plant. The germplasm storage or preservation is critical.
Seeds were traditionally used to preserve germplasm.
However, it is critical to retain seeds that cannot be utilized
for plant regeneration or that have unstable shoot and root
tissue. It is also essential to protect endangered and uncommon plant species; otherwise, some of the valuable genetic
features found in current and primitive plants would be
lost [71]. This can be achieved in the following ways:
Cryopreservation (Greek-krayos-frost): Any plant tissue
may be cryopreserved, such as seeds, protoplasts, calluses, endosperms, meristems, embryos, and ovules.
Cells are kept in a frozen form during cryopreservation.
The germplasm is preserved at extremely low temperatures (80 °C), solid carbon dioxide (79 °C), liquid
nitrogen (196 °C), and vapor nitrogen (at 150 °C).
The cells remain fully inert and may thus be stored for
lengthy periods of time. DMSO (dimethyl sulfoxide),
acetamide, praline, mannose, glycerol, sucrose, propylene, ethylene, glucose, and other chemicals are introduced during cryopreservation. These are known as
cryoprotectants, and they work by lowering the freezing
and supercooling points of water to protect cells from
harm caused by freezing or thawing.
Slow Growth Culture: Slow development of cultures
entails restricting growth circumstances such that the
culture does not expand and propagate at a normal rate.
This can be accomplished by reducing the elements
influencing development. Lowering the temperature
below the ideal threshold was discovered to have an
effect on the cultures by slowing their development rate.
360 18 Medicinal Plant Biotechnology
The limiting of certain nutrients which is vital for
growth and differentiation helped in achieving the slow
growth culture.
18.2.8 Plant Cell Immobilization
The ability of cells to synthesize important substances,
such as energy, nutrients, and pharmaceuticals, is well
established. Most biomanufacturing procedures have frequently used submerged cultures with suspended free
cells. Cells can readily be suspended in cultures but are
challenging to remove from the culture medium because
they are typically tiny (1–10 µm) and have densities that
are similar to that of the culture medium. Cell separation
procedures sometimes involve significant capital expenditure, high energy usage, and contamination risks when the
cells are recycled into the bioreactor [72]. Immobilizing the
cells in cultures is a viable strategy to overcome these problems. Cells can be immobilized by confining or anchoring
them in or on an inert support to ensure their stability and
functional reuse [73]. Cells are improved for their industrial application by using this technology, which also makes
them more affordable. The idea of cell immobilization
originated by the immobilization of enzymes. Actually, the
entire cell and enzymes are comparable in that they are
both seen as biological catalysts, and they share the same
techniques for immobilization.
Compared to free cell systems, cell immobilization technology has numerous potential benefits, which comprises
maintaining higher cell densities in a bioreactor and easy
and inexpensive cell isolation from a culture medium; simple to establish continuous cell growth at high dilution
rates; cell development lag phase reduction; volumetric
production increased; improved substrate utilization; and
reduced the risk of microbial contamination [74, 75].
18.2.8.1 Methods of Immobilization
a binding agent (crosslinking agent) such as glutaraldehyde or carbodiimide, covalent bonds are formed between
the cell and the activated inorganic substrate. Although
covalent binding may have higher immobilization effectiveness than adsorption, it is still less effective than other
immobilization techniques like entrapment [76].
b) Entrapment: Cells can be entrapped in porous parti-
cles or gel matrixes.
Entrapment in Gel Matrixes: To encapsulate the cells,
cells are suspended in gel solutions, which are subsequently gelled into beads or sheets. Polymeric matrices
(such as gelatin, polyvinyl alcohol, and collagen) or polysaccharide gels (such as alginates, -carrageenan, agar,
chitosan, and polygalacturonic acid) can be used as support materials. While the cells are trapped within the gel
beads, substances and products can diffuse in and out of
them [77-79]. Alginate is the most often utilized gel
matrix-forming substance for cell entrapment. Alginate
is a polysaccharide composed of mannuronic and guluronic acids that is obtained from marine brown algae.
Entrapment in Porous Particles: Cells can also become
entangled in the inner pore of prefabricated support
materials. Cells will get confined inside the porous materials once they have grown and reached a particular cell
density. This immobilization technique uses porous
glasses, clays, zeolite, and ceramics materials [80]. The
secondary metabolite Capsaicin production is improved
by entrapment in reticulated polyurethane foam [81].
Diosgenin production is also improved by entrapment in
polyurethane foam [82].
c) Encapsulation: It is a method of producing spherical
particles in which a liquid or solution remains trapped
within a semipermeable membrane.The membrane
may be polymeric, lipoidal, lipoprotein-based or nonionic in nature.
a) Adsorption: Electrostatic interactions or adsorption
are based on Van der Waals forces between charged
supports and immobilized cells. Hydrophobic surfaces
tend to be more sticky than those that are hydrophilic.
Minor changes in pH, ionic strength, or temperature
can quickly dislodge the attached cell. The low immobilization efficacy of this method is its main drawback.
Depending on the mechanism of immobilization, an
unstable balance between cell attachment and detachment may occur, and in most cases, freely suspended
cells coexist alongside immobilized cells. As a result,
the technology’s potential applications for many bioproduction systems are limited.
Covalent binding can also be used to attach whole cells
to the surfaces of the support material. In the presence of
18.2.9 Biotransformation
Chemical processes that are catalyzed by cells, organs, or
enzymes are called biotransformation. . Microbial, plant or
animal cells or pure enzymes can be utilized as catalysts in
biotransformation to perform particular conversions of
complicated substrates. An exogenously provided material is
chemically transformed in this procedure via a living cell
culture. Plant cells can transform a variety of substrates and
may therefore carry out several processes, including oxidation, reduction, amino-acylation, methylation, hydroxylation, and glucosylation-acylation. There is a significant
metabolic potential for the generation of particular secondary metabolites in plant cell cultures. In plant cell cultures,