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Файл:Ординатура / Хирургия / Библиотека им академика М.И. Перельмана / Книга_5847_Библиотеки_им_академика_М_И_Перельмана
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Isolate tissue aseptically using sterile equipment
Treat the organ/tissue with 7X-detergent for 5 min
Immerse in a freshly prepared solution of chlorinated lime for 20 min
Wash the explant with sterile distilled water many times
Cut it into smallpieces of tissue
Incubate in the culture medium
Shoot formation
Root formation
Acclimatization
Fig. 1.6 Schematic representation of organ culture method
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Large-scale plant cell, tissue and organ cultures provide an even and controlled
supply of phytochemical products irrespective of the plant availability. In the past,
organ culture did not prove very successful as organ tissue culture occurs in a continuous batch. Any change in conditions would disturb the equilibrium and the cultures would not grow. But the advancement of technology and the development of
specic bioreactors for organ culture has dramatically improved the situation.
Researchers are even more motivated to develop bioreactors to improve the procurement of secondary metabolites from organ micropropagation [25, 26].
1.3.3 Single Cell Culture
In this technique, isolated single cells are grown on a suitable nutrient medium in a
controlled sterile environment. Single cell culture has numerous applications.
Sources of the single cell can be a callus, cell suspension, variety of tissues and
organs. Single cells can be obtained by treating intact plant tissue (leaf, stem, etc.)
with enzymes or mechanical separation. The single isolated cell can be cultured in
a suitable liquid or solid medium. In the culture medium, the isolated cells multiply
to form a callus tissue. This callus tissue can also grow into a plantlet through organogenesis and embryogenesis.
Diffuse liquid cultures can also be used for single cells cultures. Single cells can
grow effectively on lter paper over established cultures. When the cultures are
moved to agar, they grow very well. Some cultures can grow well even after being
transferred 25 times or more within 5–6weeks without lessening their growth. This
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technique has proven successful for the clone establishment of grapes of crown gall
origin and marigold. This method can also be applied to Nicotiana tabacum (strain
WTI), N. glutinosa×N. tabacum (strain WTH). Researchers have observed altera-
tions among clones in growth rate, texture, and color resulting from the same parent
plant. Different media can be used to grow these single cells to establish shoots or
roots in the culture [28].
The rst callus is produced and maintained at 24°C on a suitable medium in the
dark to produce single cell cultures. One-third of the calli are refreshed and recultured into the fresh medium plate. The cell suspension can be initiated after
14days of sub-culturing the calli. To initiate single-cell suspension, take 2g of the
callus and transfer it to 50ml medium in a 250ml conical ask. The ask is gently
shaken to break open the callus at 100rpm. The ask is kept in a shaking incubator
at 23°C for 5days in dark. To refresh the medium, top it with 50ml of the freshly
prepared medium. After some days, a mixture of micro calli and single cells is
formed. To get the desired product, the ask is allowed to stand by for some time to
separate the micro callus, leaving a compound mixture of dead and viable cells in
the suspension. Decant the two-thirds of the supernatant gently and only keep the
one third of the supernatant. This residual mixture contains both single cells and
micro calli. Refresh the inoculum by adding 100ml of freshly prepared medium.
Incubate this ask in the dark at 23°C for 12days. Repeat this procedure 3–4 times.
This procedure allows an average of 6×10−5cell/mL with 65% viable single cells
in the supernatant. This suspension can be maintained and utilized for 6months. A
owsheet diagram is shown in the Fig.1.7 [29].
Single cell culture is a very valuable technique that can be used not only to produce plant clones but also to produce resistant cell lines and as a research tool
(Fig.1.7). It can be used to study photosynthesis, ion transport, secondary metabolite production, cell growth, differentiation, and apoptosis. For example, the cell
suspension cultures of Arabidopsis and Zinnia cells were used to detect multiple
genes and a wide range of gene expression studies [29]. Single cell cultures are very
helpful in producing resistant cell lines. Single cells are grown on a medium with
mutagenic compounds and the proliferating cell lines are isolated. The cultures
which survive these mutagens are selected and grown into whole plants to conrm
and compare their phenotypes with normal plants. This simple method has assisted
in selecting cell lines and plants resistant to antibiotics, herbicides, fungal toxins,
etc. By using this technique rather than growing the whole plant, a greater amount
of commercially and medically important compounds can be produced [29–32]
(Fig.1.8).
I. Fatima etal.
1.3.4 Suspension Culture
In this micropropagation technique, a single cell or minute masses of cells (explants)
are cultured in a liquid medium on a shaking incubator [33]. Preferably, cells should
be suspended singly in cell suspension cultures through continuous agitation. But
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produce callus and
maintain them at
medium 24
○
C
reculture them on a
medium plate
transfer it to 50ml
medium in a 250ml
conical flask.
refresh the calli with
50ml fresh medium
on a suitable
medium in dark
initiate cell
suspension after 14
days of
Sub-culrturing
shake the flask at
rpm to break 100
the callus
A mixture of micro
calli and single cells
is formed
refresh one third of
calli
take 2 g of the
callus
keep the flask at
o
C for 5 days in
23
dark
separate them by
decanting two third
and keeping the one
third suspension
refresh the
Inoculum with
100ml medium
Fig. 1.7 Schematic representation of single cell culture
such culture exists only in rare cases. Researchers strive to obtain a ne culture by
increased cell dissociation, which can lead to increased culture uniformity. But even
after many measures are taken, some cell aggregates are formed. The cell aggregates can be divided into two groups. One is ‘ne’ suspension cultures which consist of micro- to sub-macroscopic colonies consisting of approximately 5–200 cells.
Other group contains aggregates of cells about 0.5–1.0mm in diameter (Fig.1.9).
The second group is often readily achievable in which cells grow perfectly well and
meet all the investigation requirements depending on the purpose of the research. It
is preferable to have some degree of cell aggregates in the suspension as cells in this
repeat the procedure
2 to 3 times
single cell
suspension is
obtained
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I. Fatima etal.
• photosynthesis
•apoptosis
•ion transport
•cell growth
•genes andgenome
•cell differentiation
Study and
research
purposes
Clones of
plants
•production of wholeplants
from single cells
Fig. 1.8 Applications of single cell culture
Production
of Cell lines
resistant to
Production
of valuable
compounds
•antibiotics
•herbicides
•fungaltoxinss
•secondary metabolites
•alkaloids
•steroids
Actively growing cells are secured.
Initial pH is set 4.6 to 7.0.
Isolation of single cells from in vitro culture of callus
Growth and development of cells on filter paper
Cultures are transferred to fresh agar plates many times
Fig. 1.9 Schematic representation of suspension culture
state can retain the totipotent character and enhance the production of desired
metabolites [34].
Plant cell suspension has wide applications in research because it bypasses the
structural complexity of the whole plant. Suspension cultures provide homogenous
cell populations with high growth rates and reproducibility of conditions which help
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scientists to study complex physiological processes at molecular and cellular processes. It can also be used to produce high-value secondary metabolites of commercial interest [35]. Cultures growing homogeneously in the liquid medium are
benecial for metabolic engineering and targeting metabolite production. Metabolite
production can be commercialized using a bioreactor system. Other advantages of
suspension culture are the study of in vitro mutagenesis and genetic transformation
[33, 35].
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1.3.5 Embryo Culture
In this process, an embryo (in different developmental stages) is aseptically isolated
from mature seeds and cultured under aseptic invitro controlled conditions on a
suitable medium. In most situations, sterilization of embryos is not required after
collection because they are protected by the aseptic environment of the ovule.
Therefore, entire ovules or ovaries are sterilized exteriorly, and embryos are removed
aseptically from the adjacent tissues. Researchers also use harsh methods to disinfect the surface due to hard protecting tissues. Direct decontamination is only necessary if the seed coat is cracked or there is any sign of endophytic pathogens.
Endophytic infections can occur in the seeds of fescue (Festuca spp. L.), corn (Zea
mays L.), and dogwood (Cornus spp. L.) [36].
Embryos of larger size are not hard to excise, but small embryos pose dissection
problems because researchers need microdissection tools and a dissecting microscope for excision without injury. It is necessary to ensure that the embryo is not
desiccated or injured during culturing. The process may vary from specie to specie.
Still, it is generally performed by making an incision at the micropylar end of the
young ovule and applying pressure at opposite ends to force the embryo out through
the opening. If a liquid endosperm surrounds the embryo, the pressure must be
applied carefully because it may damage the fragile embryonic tissue[36].
Figure1.10 shows the process of embryo culture.
Similarly, in somatic embryogenesis, a single or group of cells can regenerate
from non-zygotic embryos and germinate to form complete plants. Immature
zygotic embryos and seedlings are appropriate explants for somatic embryogenesis.
Somatic embryogenesis can be either direct or indirect. In direct embryogenesis,
embryos are developed from the exterior of explants, whereas during indirect
embryogenesis, embryo formation involves the intermediate callus phase. This
method is benecial for metabolic engineering. Mature organs (leaves or roots)
have a low capacity for somatic embryogenesis [12]. Embryo culture is useful in
studying the factors involved in the dormancy of seeds and crosses between plants
of different ploidy within the same species, e.g., Iris and Zea. This technique also
helps to procure inter-specic hybrids, e.g., Gossypium, Datura, and Lycopersicum.
This tissue culturing technique has shortened breeding cycles. This method has
served as a tool for studying the effect of specic substances on the morphology of
embryos [37].
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I. Fatima etal.
Take siliques
containing at the
early bent-
cotyledon stage
Acclimatization in
green house
Surface
sterilized for 10
sec in
%alcohol70
Transfer to
shooting and
rooting
medium
10 min
incubation in
commercial
bleach
Embryogenic cell
lines are
maintained by
subculturing
Wash 3 times
with distilled
water
Refresh medium
after 2 weeks
Isolate immature
zygotic embryos
under dissecting
microscope
Incubate
embryos in
shaking
•
incubator
(100rpm)
autoclaved B5-4
•
medium
25°C
•
3000lux
•
16h light/8hours
•
darkness
Fig. 1.10 Schematic representation of embryo culture
1.3.6 Anther Culture
Anther culture is also known as “Androgenesis.” In this method, haploid plants are
developed invitro from effective pollen grains through a series of cell divisions and
differentiation. Androgenesis can be performed by both in vivo and in vitro methods. In vivo androgenesis is induced by irradiation or temperature-shock treatments,
as reported in Crepis tectorum and Antirrhinum majus [38].
In vitro androgenesis involves the separation of anthers from the bud. The bud
from a young ower is surface sterilized and rinsed with water when the pollens
reach a suitable development phase. The bud is dissected after removing the sepals
and petals. Each anther is gently detached from the lament and placed horizontally
on the nutrient medium (either liquid or agar medium). Only intact and uninjured
anthers are selected for culturing. Figure1.11 represents the schematic diagrams for
anther culture. Androgenesis has many advantages. It creates haploid and spontaneous diploid plants in vitro. These plants have genetic potential that is rapidly
expressed phenotypically. This technique lessens the time needed to grow selfpollinated crops, reduces the breeding cycle, and requires less space and labour
[39, 40].
1.3.7 Protoplast Culture
A cell without a cell wall is known as a protoplast. A protoplast can regenerate its
cell wall, divide, and grow into a whole plant if a suitable medium is provided under
aseptic conditions. The technique which uses protoplast as an explant is known as
protoplast culture. Protoplasts can be obtained by removing the cell wall by
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Surface sterilization
Anther are cultured
horizontally in nutrient
medium
Microspore isolation from
pretreated anthers
Multicellular structures
start to form in 14 days
Acclimatization
Separation of anthers
from the bud
Pretreatment of the plant
material with cold or heat
shock to start mitosis in
embryos or embryo-like
cell
Microspore culture Cell division can be
These multicellular
structures develop into
structures
Anthers are placed in petri
developmental stages are
Further development into
plate
Anther at the right
selected
observed after 4 days
young plants
Fig. 1.11 Schematic representation of anther culture
mechanical disruption or treating the plant tissue or cultured cells with enzymes
[41]. In order to get protoplasts, fresh-weight tissues such as leaves or embryos are
chopped and incubated in 0.5 M mannitol for an hour. After this, mannitol is
replaced with an enzymatic mixture containing cellulase, macerase and driselase.
The suspension is then incubated in the dark for 16h with gentle shaking (50rpm).
The digest is then passed through a 50mm nylon sieve. After repeated centrifugation, the pellet is resuspended in 0.5 M mannitol agar. These protoplasts can be
cultured after protoplast fusion or without fusion. For culturing, both liquid and
solid media can be used. Figure1.12 shows the ow chart of the process [42].
Protoplasts are used when selection or hybridization at the cellular level is
required. Commonly protoplasts are fused by either chemical or electrical means.
The mostly employed chemical is polyethylene glycol (PEG). Specic equipment
that can induce the electrical fusion of cells is needed for electrical fusion. It is
equipped with an AC eld or a DC source to apply the AC eld or DC pulses and a
suitable fusion chamber [41].
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Chop leaf or embryo tissue
Incubate it in 0.5M mannitol for 1 hour
I. Fatima etal.
Incubate the enzymatic mixture in dark for 16 hours at 50
Pass the digest through 50mm nylon sieve
Centrifuge it several times
Resuspend the palette in 0.5M mannitol
Inoculate it in the medium
either fused or without fusion
Fig. 1.12 Schematic diagram of Protoplast culture
rpm
1.3.8 Meristem Culture
Meristem refers to the tips of shoots that contain the shoot apical meristem. Meristem
tip comprises the meristematic dome and often a pair of leaf primordia which can
be used as an explant. This explant is cultured into a suitable nutrient medium under
aseptic conditions and can develop into whole plants. Occasionally, a liquid medium
is used. This method is used for contaminant-free axillary shoot multiplication and
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sustaining diverse superior mother plants that amass a greater number of metabolites of interest. Meristem cultures are an excellent method for virus-free cultivation [12].
About 10mm long plant tips are cut off and used as explants. These are sterilized
carefully by washing them with running tap water and then detergent for 5min.
After this, the explant is washed with water to eliminate the detergent, then treated
with 1% sodium hypochlorite solution for 15minutes and rinsed three times with
sterile distilled water to clean all sodium hypochlorite. The explants should be
0.5–1.0cm when inoculated on the medium. The medium must be supplemented
with suitable growth hormones along with sugar. The cultures are then incubated at
25°C at 3000lux and provided with cool white uorescent lamps. When cultures
show growth, they should be re-cultured into fresh medium. After 2–4weeks of
further growth, plantlets are transferred for rooting and acclimatization. To acclimatize plantlets, the soil should be disinfected and saturated with water in a greenhouse. Figure1.13 shows the procedure in a schematic diagram [43, 44].
Fig. 1.13 Schematic
representation of meristem
culture
Wash the explants with water
Treat then with detergent for 5minutes
Wash to remove the detergent
Apply1%NaOCl for 15min
Wash three timeswithsterile distilledwater
Cut explantin0.5 to 1.0 cm size
Incubate in culture medium
Incubated:
at 25◦C 3000 lux light.
Re- culture the explants in fresh medium
Rooting
Acclimatization
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I. Fatima etal.
1.3.9 Pollen Culture
In pollen culture, pollen grains are separated from intact anther at microscopic
stages and transferred to a suitable medium to produce microspores without producing male gametes. This technique enables scientists to culture haploid plants for
propagating haploid plants and for genetic studies [45]. The details of the procedure
are given in Fig.1.14. First, the pollen grains are isolated from anthers by crushing.
The pollen grains are ltered by passing them through a sieve of a pore size of
100pm. Isolated pollen grains are washed with distilled water twice and centrifuged
at 100× for 2min. Pollen suspension of known concentration is prepared and inoculated into the nutrient medium. The medium used by Matsushima and colleagues
was supplemented with 1% sucrose and glucose, 0.1% yeast extract and casein
hydrolysate, and 10–5M 2, 4-dichlorophenoxyacetic acid (pH5.8). They got the
best results with the medium diluted multiple times. A concentrated medium does
not give good results with pollen culture [45].
Crush the anthers to isolate pollen grains
Filter the pollen grains through a sieve of pore size 100 pm
Wash the isolated pollen grains with distilled water
Centrifuge then at 100X for 2 min
Prepare a pollen suspension of known concentration
Dilute the suspension eight times
Inoculate it into the nutrient medium
Callus formation
Shoot formation
Root formation
Acclimatization
Fig. 1.14 Schematic representation of pollen culture
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