CHARACTERISTICS OF JUVENILE PLANTS

There are numerous substantive phenotypic traits associated with juvenility, but they vary onsiderably among species.  Commonly, the leaves on young plants are of a different shape than those on mature parts and may be simple rather than compound (or occasionally the reverse); juvenile leaves may also have a special type of cuticle and be arranged with a distinct phyllotaxy. Compared to their adult counterparts, young plants may have a modified resistance to pests and diseases.  Juvenility in woody plants is often manifested by prolonged vigorous shoot growth.  With Citrus and Gleditsia triacanthos, juvenile forms are thorny, whereas adult forms lack the thorniness and with some tree species, such as those of  Quercus and  Fagus, juvenile forms and older, more juvenile parts of the trees, hold their senesced leaves throughout the winter.

1.1.1.      Vegetative propagation

            To the plant propagator, the most important attribute of juvenile shoots is their ability to rovide cuttings that readily form adventitious roots or explants that respond and grow well  in vitro. Cuttings taken from adult shoots of plants can be rooted, but the frequency of success is often low, especially with woody plants.  Likewise, researchers have had great challenges when attempting to micropropagate adult forms of many woody species. The change from the juvenile to adult phase is the most serious constraint to rooting in shrubs and trees (Howard, 1990).  Most of the difficulty experienced in rooting mature shoots seems to be caused by their altered physiology, but can also be related to greater contamination with microorganisms and viruses.

            Hedera helix has been used widely to study juvenility because it has a distinctively different morphology between the juvenile and adult phases. Juvenile plants have a different growth habit, leaf shape, and an enhanced ability to form adventitious roots.  When petioles from the juvenile form were excised and treated with auxin  in vitro, cortical parenchyma cells adjacent to the vascular bundles divided and formed root primordia (Geneve  et al., 1988).  However, when  petioles from adult leaves were treated in a similar manner, callus formed and some callus cells divided to form root primordia.  The juvenile form had pre-existing competent cells that were able to respond to auxin and become determined to form roots.  However, the adult form appeared to lack cells with pre-existing competence to form roots, but competence was acquired by some callus cells once they had been initiated.

            Explants taken from mature shoots are frequently more liable than juvenile material to uffer necrosis, especially when surface disinfested and placed in culture (Hanus and Rohr, 1987).  For example, shoot tip explant death can occur within a few hours for adult  Juglans nigra whereas healthy growth was evident on seedling explants when both sources were compared in different vessels containing the same medium (Preece and Van Sambeek unpublished).  It was only by changing the medium and culture conditions that adult J. nigra shoot cultures have been maintained for years  (Pearson and Preece un-published). However, adult origin  J. nigra micro-shoots still cannot be rooted.

For tissue culture, juvenile explants are usually more readily established  in vitro and grow and proliferate at a more rapid rate than adult material. This is particularly true with tree species where micropropagation of adult material is often difficult.

1.1.2.      Plant propagation dilemma 

            It is well known that it is easier to propagate vegetatively, juvenile forms of plants than adult forms of plants.  When breeding and selecting new, superior plants for clonal propagation, it is sually necessary to wait until the plant reaches maturity.  This allows for evaluation of important features, such as ultimate form and size, flowering and fruiting characteristics, autumnal coloration, and other traits.  At the point that the mature phenotype is known, the plant is an adult and often becomes difficult to propagate clonally.  Libby and Hood (1976) showed that juvenility can be maintained by hedging radiata pine. They rooted cuttings of many juvenile selections and  by allowing some individuals to grow to maturity for evaluation, other, hedged members of the same clone could be maintained as juvenile plants for propagation.  This technique also has potential for micropropagation.

PHASES OF GROWTH

The sequence of stages through which a higher plant has passed in its development from a ertilized egg to an adult organism, is known as its ontogeny or ontogenesis.

In the juvenile phase, a young seedling plant displays one or more distinctive characteristics of both a morphological and physiological nature. These distinguish a juvenile plant from an adult.  The juvenile phenotype gradually disappears during subsequent growth, and under natural conditions it is replaced by the adult (mature) phase in those parts of capable of changing from vegetative to flowering meristems under “normal” inductive conditions.  In some plants, a transitional phase of development can be distinguished between the juvenile and adult phases during which the potential to flower gradually increases. the plant in which maturation has occurred. Maturation occurs when the apical meristems become capable of changing from vegetative to flowering meristems under “normal” inductive conditions.  In some plants, a transitional phase of development can be distinguished between the juvenile and adult phases during which the potential to flower gradually increases.

When a plant is induced to flower, the shoot apical meristem changes from forming vegetative structures to forming reproductive organs (Hackett and Murray, 1997).  Typically, it is an external environmental stimulus, such as day length or chilling that will trigger this change.  Under such a normal environmental stimulus the apical meristem of a juvenile plant is not capable of perceiving or responding to the signal and remains vegetative. Under certain circumstances,  juvenile plants can be induced to initiate flowers but will revert to the non-flowering juvenile state for several years (Hackett, 1987).  However, it is often difficult to devise methods to promote flowering that will not also advance maturation (Hackett, 1985).  In many conifers, flowering can be induced on juvenile shoots by gibberellins or auxin/gibberellin combinations (Pharis and King, 1985; Pharis and Kuo, 1977; Pharis et al., 1976, 1980; Ross et al., 1981).

In general, the duration of the juvenile phase is proportional to the potential ultimate size of the plant, being shortest in annual herbaceous species and progressively longer and more noticeable in perennial and woody plants.  In trees it may sometimes persist for many years.

Phase change from juvenile to adult is an epigenetic change, in that there are phenotypical changes that are the result of changing gene expression, not mutation.  When an adult plant is regenerated from an embryo (either zygotic or somatic), juvenile traits are again expressed.  Gene expression for dihydroflavonol reductase (DFR) was studied in adult and juvenile leaf lamina tissue of English Ivy (Hackett and Murray, 1997).   DFR activity was detected in juvenile leaves treated with sucrose and light, but not in mature phase discs.  It was determined that the lack of DFR activity was because there was no accumulation of DFR mRNA because of a lack of transcription of the DFR gene in mature phase discs.  Lack of expression of this gene was also reported for adult phase stem tissue.  This gene was expressed in juvenile leaves and stems. The specific reason why the  DFR locus was not transcribed was unknown.  In Arabidopsis, both light and temperature independently regulate the floral promoter (FT) gene (Poethig, 2003).  However, there are other genes that also regulate temperature sensitivity of flowering time.  These genes (FCA and FVE) act upstream from the FT gene and seem to be involved in phase change.

EPIGENETIC EXPRESSION AND CELL DETERMINATION

Plants cells have an amazing ability to respond to a signal, which almost invariably is a plant growth substance.  When the cells can respond, they can change from a previously determined developmental pathway.  For example, a leaf cell can change so that it will begin to divide and a root or shoot will form.

            In tissue culture and plant propagation, the ratio of cytokinin : auxin is important (Chapter 6).  When cytokinin is in excess, adventitious shoots will form, when auxin is in excess, adventitious roots or somatic embryos will form, and when there are moderate to high levels of both, callus will develop.  This is the normal wound healing process in plants.  Many plant species have the remarkable ability to replace wounded or lost organs.  Following wounding, the endogenous hormone levels change and callus formation and organogenesis are often the result.  Similarly, with some species, because of high auxin levels associated with sexual fertilization in the seed, adjacent somatic cells change and produce embryos through the process of apomixis.  In plant propagation and plant tissue culture, we exploit this healing process and the apomictic ability of cells.

            Different cells within a tissue will not all respond the same way to a change in the cytokinin : auxin ratio.  Understanding what distinguishes certain cells from others in their ability to respond to the plant growth substance is important.  As our knowledge of the molecular biology of cells increases we may understand why there are these cellular differences.  With such knowledge, we may gain the ability to regenerate recalcitrant genotypes, such as adult forms of many plants.

Christianson (1987) and Christianson and Warnick (1983, 1988) stated that there are competent cells in a plant or explant (see Chapter 10).  These are the cells that are capable  of recognizing the plant growth regulator signal  and changing, or becoming determined.  When the plant growth regulator signal is removed, the determined cells will continue to respond.  They will dedifferentiate (become meristematic again), and their daughter cells will differentiate into the new shoot, root, or somatic embryo.

Not all cells in a tissue  are competent.  Exactly what makes one cell competent to perceive a plant growth regulator signal and its adjacent cell not to be competent is not well understood.  It is known that applied plant growth regulators elicit specific mRNA molecules (Christianson and Warnick, 1988).  This is evidence of specific genes being expressed in response to the exogenous plant growth substances.  However, this specific gene expression is more clearly related to changes in determined cells than to what makes cells competent.

A problem with trying to understand what makes a cell competent is that it is difficult to ascertain that a specific cell is competent until after it responds to the plant growth substances and becomes determined.  Once determined, the cell changes and its gene expression can be studied.  However, it is difficult to know which cell is competent and which one is not before the cell becomes determined.

            It is likely that epigenetic expression is the reason that a cell is competent.  Perhaps there is a gene or genes expressed for the receptor molecule of the elicitor plant growth regulator, which would allow that competent cell to recognize the plant growth substance.  However, there are other possibilities that could lead to competence, such as a expression of an important gene in the signal transduction pathway in a competent cell.

The predetermination of cells and tissues can often be changed in culture by cell differentiation and cell division, leading to the direct formation of organogenic or embryogenic meristems or callus. But if the tissue is not competent to undergo these changes, or the growth regulators in the medium are not suitable, previously predicated developmental pathways may not be interrupted.  Some aspects of determination may nevertheless survive in newly-established cultures and can occasionally persist for many callus transfers.

Results and Discussion

One of the most popular gateways is NowSMS since it can work very well with mobile phone that soon
will operate as a GSM modem where Sony Ericsson k800i had been preferred. Configuration process is
important to make sure that SMS gateway can work well with the mobile phone. 
Alert system will be activated only if the normal parameter range is exceeded. Normally, the temperature
and humidity of tissue culture did not change rapidly. In many cases, the next reading will be more or less
than previous reading. It is quite troublesome  since monitoring process is done manually by using
thermometer and hydrometer. The problem typically occurred during weekend or holidays since there is no
one available to do the monitoring process. Worst case scenario is electrical supply failure when its blackout,
the tissue culture will definitely contaminate be due to the inconsistency of temperature and humidity. Fig. 3
shows the data of temperature and humidity in tissue culture growth room at MPOB.

The suitable parameter for temperature is between 25°C to 30°C while for humidity is between 45% until
90%. If the parameter range is exceeded, alert system will notify the user by sending SMS and email
notification. Fig. 4 shows the temperature and humidity graphs of tissue culture in growth room.

SMS and email notifications display shows in Fig. 5. The SMS information are including the subject,
parameters range, date and time as well as the current value of parameters. The email notification shows the
sender’s address, date and time, recipient’s address, subject, the parameters range as well as the current value
of temperature and humidity that had been triggered by alert system.

Methodology

Besides expanding the range of applications in wireless technology, this development also focusing on
alert system that provide a new application of SMS in order to quickly provide urgent information to the user.
A mobile phone used to connect with the computer and act as GSM modems to replace a common modem
that requires a higher cost. Fig. 1 shows the process involved in monitoring the oil palm tissue culture which
is begin with hardware development to collect the data, data monitoring and acquisition and finally, an alert
system. In addition to the SMS service, the user also will receive an email notification about the parameter
data that can be kept as the track record.

This alert system is connected with database that contains data of temperature and humidity in the tissue
culture vessel or in growth room. It begins with comparison of the current data and the parameters range. In
condition of current data is within normal range, the alert system will be in idle stage. Nevertheless, if the
current data is beyond the parameters range, the alert system will be activated where notification will be send
to user’s mobile phone via SMS and email to user’s email address. 
The intention of this implementation alert system into the technology of oil palm tissue culture is to
provide efficient monitoring methods. The combination between Apache, PHP and MySQL is the most
important tools to develop the alert system. Apache is used as a web server that is connected with MySQL
database and PHP scripting language.
SMS alert system is developed by using SMS Gateway software called NowSMS. NowSMS is a scalable
solution that is affordable for development and testing, with scalability to support mobile operator systems.
The configurations for email and SMS are including the recipient’s address, sender’s address, present date
and time, parameters range and also current value of temperature and humidity that triggered the alert system.
Fig. 2 shows a diagram on how SMS and emails are sent from PHP scripting command through the MySQL
database manager. 

Theoretical Background

1.  Tissue Culture
Tissue culture is the propagation of plants through the placement of small amounts of undifferentiated
tissue or single cells in an artificial environment. The tissue is placed in a nutrient medium that favours the
production of roots and shoots, and is later planted normally. By using tissue culture, the favourable qualities
of plants can be precisely controlled, so that each plant is identical for the particular quality. The progress in
the development of the technologies of plant tissue and cell culture over the past decades has been
remarkable. Tissue culture is the most fundamental technique in any plant cell biologist's toolbox [6].
Basically the technique consists of taking a piece of a plant such as a stem tip, node, embryo, or even a seed
and placing it in a sterile, (usually gel-based) nutrient medium where it multiplies. Plant tissue culture
comprises a set of in vitro techniques, methods and strategies that are part of the group of technologies called
plant biotechnology. Tissue culture has been exploited to create genetic variability from which crop plants
can be improved, to improve the state of health of the planted material and to increase the number of
desirable germplasms available to the plant breeder [7]. This technology  can be expected to have an ever
increasing impact on crop improvement as an approach to the new millennium.

2.  Alert System 
Alert system is originally based on telemetry system. Telemetry is a highly automated communications
technique with the help of which measurements and  data collection are done at remote locations and
transmitted for monitoring [8]. The most important uses of telemetry include weather data collection,
monitoring power generation plants and keeping track of space flights. A telemetry system typically consists
of a transducer as an input device, a transmission medium in the form of wired lines or radio waves, signal
processing devices, and devices for recording or displaying data. The transducer converts a physical quantity
into a corresponding electrical signal, which is then transmitted over a distance for the purpose of
measurement and recording. There is normally a sound or vibration that will indicate that the message has
come in. The receiver may be able to see the sender’s telephone number, name and also date and time [9].

3.  SMS in GSM Networking 
SMS is a method by which messages can be sent  to a cell phone via another cell phone, a computer
connected to the Internet or a regular land line [10]. SMS is a communication service component of the GSM
(Global System for Mobile Communications) system, using standardized communications protocols that
allow the exchange of short text messages between mobile phone devices.

4.  Email
Email systems are based on a store and forward model in which email server computer systems accept,
forward, deliver and store messages on behalf of users, who only need to connect to the email infrastructure,
typically an email server, with a network-enabled device for the duration of message submission or retrieval
[11]. Email messages are posted electronically to individuals at specific. The address denotes the computer
that the individual employs as a mail server. A mail server is like a computer that sends and receives
electronic mail for a specific network. 

TYPES OF TISSUE CULTURE

3.1. ORGAN CULTURES
Differentiated plant organs can usually be grown in culture without loss of integrity. They can be of two types:
• Determinate organs which are destined to have only a defined size and shape (e.g. leaves, flowers and fruits);
• Indeterminate organs, where growth is potentially unlimited (apical meristems of roots and nonflowering
shoots).

In the past, it has been thought that the meristematic cells within root or shoot apices were not committed to a particular kind of development. It is now accepted that, like the primordia of determinate organs such as leaves, apical meristems also become inherently programmed (or determined) into either root or shoot pathways (see Chapter 8). The eventual pattern of development of both indeterminate and determinate organs is often established at a very early stage. For example, the meristematic protrusions in a shoot apex become
programmed to develop as either lateral buds or leaves after only a few cell divisions have taken place (see Chapter 10).
3.1.1. Culture of determinate organs An organ arises from a group of meristematic cells. In an indeterminate organ, such cells are theoretically able to continue in the same pattern of growth indefinitely. The situation is different in the primordium of a determinate organ. Here, as meristematic cells receive instructions on how to differentiate, their capacity for further division becomes limited.
If the primordium of a determinate organ is excised and transferred to culture, it will sometimes continue to grow to maturity. The organ obtained in vitro may be smaller than that which would have
developed on the original plant in vivo, but otherwise is likely to be normal. The growth of determinate
organs cannot be extended by subculture as growth ceases when they have reached their maximum size.

THAWING AND RECOVERING HUMAN CELLS

When cryopreserved cells are needed for study, they should be thawed rapidly and plated
at high density to optimize recovery.
CAUTION: Protective clothing, particularly insulated gloves and goggles, should be worn
when removing frozen vials or ampules from the liquid nitrogen freezer. The room
containing the liquid nitrogen freezer should be well-ventilated. Care should be taken not
to spill liquid nitrogen on the skin.
Additional Materials (also see Basic Protocol)
70% (v/v) ethanol
Complete medium/20% FBS (e.g., supplemented DMEM-20, APPENDIX 2A), 37°C
NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2
incubator unless otherwise specified. Some media (e.g., DMEM) may require altered
levels of CO2 to maintain pH 7.4.
1. Remove vial from liquid nitrogen freezer and immediately place it into a 37°C water
bath. Agitate vial continuously until medium is thawed.
The medium usually thaws in <60 sec.
Cells should be thawed as quickly as possible to prevent formation of ice crystals that can
cause cell lysis. Try to avoid getting water around the cap of the vial.
2. Wipe top of vial with 70% ethanol before opening.
Some labs prefer to submerge the vial in 70% ethanol and air dry before opening.
3. Transfer thawed cell suspension into a sterile centrifuge tube containing 2 ml warm
complete medium/20% FBS. Centrifuge 10 min at 150 to 200 × g (∼1000 rpm in
Fisher Centrific), room temperature. Discard supernatant.
Cells are washed with fresh medium to remove residual DMSO.
4. Gently resuspend cell pellet in small amount (∼1 ml) of complete medium/20% FBS
and transfer to properly labeled culture plate containing the appropriate amount of
medium.
Cultures are reestablished at a higher cell density than that used for original cultures
because there is some cell death associated with freezing. Generally, 1 ml of cell suspension
is reseeded in 5 to 20 ml medium.
5. Check cultures after ∼24 hr to ensure that cells have attached to the plate.
6. Change medium after 5 to 7 days or when pH indicator (e.g., phenol red) in medium
changes color. Keep cultures in medium with 20% FBS until cell line is reestablished.
If recovery rate is extremely low, only a subpopulation of the original culture may be
growing; be especially careful of this when working with cell lines known to be mosaic.

FREEZING CELLS GROWN IN SUSPENSION CULTURE

Freezing cells from suspension culture is similar in principle to freezing cells from monolayer. The major difference is that suspension cultures need not be trypsinized.
1. Transfer cell suspension to a centrifuge tube and spin 10 min at 300 to 350 × g (∼1500 rpm in Fisher Centrific centrifuge), room temperature.
2. Remove supernatant and resuspend pellet in 4°C freezing medium at a density of 106 to 107 cells/ml.
Some laboratories freeze lymphoblastoid lines at the higher cell density because they plan to recover them in a larger volume of medium and because there may be a greater loss of cell viability upon recovery as compared to other types of cells (e.g., fibroblasts).
3. Transfer 1-ml aliquots of cell suspension into cryovials and freeze as for monolayer

FREEZING HUMAN CELLS GROWN IN MONOLAYER CULTURES

It is sometimes desirable to store cell lines for future study. To preserve cells, avoid
senescence, reduce the risk of contamination, and minimize effects of genetic drift, cell
lines may be frozen for long-term storage. Without the use of a cryoprotective agent
freezing would be lethal to the cells in most cases. Generally, a cryoprotective substance
such as dimethylsulfoxide (DMSO) is used in conjunction with complete medium for
preserving cells at −70°C or lower. DMSO acts to reduce the freezing point and allows a
slower cooling rate. Gradual freezing reduces the risk of ice crystal formation and cell
damage.

Materials
Log-phase monolayer culture of cells in petri plate
Complete medium (e.g., supplemented DMEM, APPENDIX 2A)
Freezing medium: complete medium supplemented with 10% to 20% (v/v)
FBS and 5% to 10% (v/v) DMSO, 4°C
Benchtop clinical centrifuge (e.g., Fisher Centrific or Clay Adams Dynac)
with 45°C fixed-angle or swinging-bucket rotor
1. Trypsinize cells from plate (see Basic Protocol, steps 1 to 4).
It is best to use cells in log-phase growth for cryopreservation.
2. Transfer cell suspension to a sterile centrifuge tube and add 2 ml complete medium
with serum. Centrifuge 5 min at 300 to 350 × g (∼1500 rpm in Fisher Centrific rotor),
room temperature.
Cells from three or more dishes from the same subculture of the same source can be
combined in one tube.
3. Remove supernatant and add 1 ml of 4°C freezing medium. Resuspend pellet.
4. Add 4 ml of 4°C freezing medium, mix cells thoroughly, and place on wet ice.
5. Count cells using a hemacytometer (see Support Protocol 3). Dilute with more
freezing medium as necessary to get a final cell concentration of 106 or 107 cells/ml.
To freeze cells from a nearly confluent 25-cm2 flask, resuspend in ∼3 ml freezing medium.
6. Pipet 1-ml aliquots of cell suspension into labeled 2-ml cryovials. Tighten caps on
vials.
7. Place vials 1 hr to overnight in a −70°C freezer, then transfer to liquid nitrogen storage
freezer.
Alternatively, freeze cells in a freezing chamber in the neck of a Dewar flask according to
manufacturer’s instructions. Some laboratories place vials directly into the liquid nitrogen
freezer, omitting the gradual temperature drop. Although this is contrary to the general
recommendation to gradually reduce the temperature, laboratories that routinely use a
direct-freezing technique report no loss of cell viability on recovery.
Keep accurate records of the identity and location of cells stored in liquid nitrogen freezers.
Cells may be stored for many years and proper information is imperative for locating a
particular line for future use.

PASSAGING CELLS IN SUSPENSION CULTURE

A suspension culture is grown in culture flasks in a humidified 37°C, 5% CO2 incubator.
Passaging of suspension cultures is somewhat less complicated than passaging of monolayer
cultures. Because the cells are suspended in medium rather than attached to a surface,
it is not necessary to disperse them enzymatically before passaging. However, before
passaging, cells must be maintained in culture by feeding every 2 to 3 days until they
reach confluency (i.e., until the cells clump together in the suspension and the medium
appears turbid when the flask is swirled).
NOTE: All culture incubations should be performed in a humidified 37°C, 5% CO2
incubator unless otherwise specified. Some media (e.g., DMEM) may require altered
levels of CO2 to maintain pH 7.4.
1. Feed cells as follows every 2 to 3 days until the cultures are confluent:
a. Remove flask of suspension cells from incubator, taking care not to disturb those
that have settled to the flask bottom.
b. Aseptically remove and discard about one-third of the medium from flask and
replace with an equal volume of prewarmed (37°C) medium. If the cells are
growing rapidly, add an additional 10% medium by volume in order to maintain
optimum concentration of 1 × 106 cells/ml. Gently swirl flask to resuspend cells.
c. Return flask to incubator. If there is <15 ml of medium in the flask, incubate flask
in horizontal position to enhance cell/medium contact.
At higher volumes of medium the flask can be incubated in the vertical position.
If using a 25-cm2 flask, there should be 20 to 30 ml of medium in the flask at confluency.
2. On the days cultures are not being fed, check them by swirling flask to resuspend
cells and observing color changes in the medium that indicate good metabolic growth.
3. When cultures are confluent (∼2.5 × 106 cells/ml), passage culture as follows:
a. Remove flask from incubator and swirl flask so that cells are evenly distributed in
the medium.
b. Aseptically remove half of the volume of cell suspension and place into a fresh
flask.
c. Feed each flask with 7 to 10 ml prewarmed medium and return flask to incubator.
Some labs prefer to split the cells 1:3 or 1:4, although increasing the split ratio will
result in a longer interval before subcultures reach confluency.

Callus Formation and Multiplication

Callus is defined as an unorganized tissue mass growing on solid substrate. Callus forms
naturally on plants in response to wounding, infestations, or at graft unions (Bottino, 1981). Since
extensive callus formation can be induced by elevated hormone levels, tissue culture media
designed to produce callus contain pharmacological additions of cytokinins and auxins.
Callus formation is central to many investigative and applied tissue culture procedures. Callus
can be multiplied and later used to clone numerous whole plants. Additionally, various genetic
engineering protocols employ callus initiation procedures after DNA has been inserted into cells;
transgenic plants are then regenerated from transformed callus. In other protocols callus is
generated for use in biotechnological procedures such as the formation of suspension cultures from
which valuable plant products can be harvested.

Callus Formation
Explants from several parts of large intact plants can be used to form callus. The most
successful explants are often young tissues of one or a few cell types. Pith cells of young stem are
usually a good source of explant material. Initially, callus cells proliferate without differentiating,
but eventually differentiation occurs within the tissue mass. Actively dividing cells are those
uppermost and peripheral in the callus. The extent of overall differentiation usually depends on the
hormone balance of the support medium and the physiological state of the tissue.

Callus Multiplication
Actively growing callus can be initiated on culture media with an even physiological balance of
cytokinin and auxin (Tobacco Callus Initiation Medium; Appendix E). After callus biomass
increases two to four times (after 2–4 weeks of growth), callus can be divided and placed on fresh
Tobacco Callus Initiation Medium for callus multiplication. Multiplication procedures can be
repeated several times (up to eight sequential transfers) before gross chromosome instability (or
contamination) occurs.

Differentiation and Plant Regeneration
Multiplied callus can be stimulated to form shoots by increasing the cytokinin concentration and
decreasing auxin content of culture media (Tobacco Shoot Development Medium; Appendix E).
Shoot masses can be cut apart and transferred to rooting medium. Once rooted, regenerated plants
can be acclimatized to natural rather than "in vitro" growth conditions. Regenerated plants are
especially valuable if the parent plant was itself unique or if the plants were genetically engineered.
If, for example, multiplied callus was first used to form suspension cultures on which genetic
engineering or cell selection was accomplished, resultant regenerated plants via tissue culture could
possess special traits or capabilities.

Materials and Methods
Callus Formation (Bottino, 1981)
1. Obtain a 5-cm section of tobacco stem.
2. Cut off all leaves.
3. Immerse it in a beaker of 95% ethanol for 15 seconds.
4. In the laminar flow hood, expose the pith by cutting away epidermis, cortex, and vascular
tissue with a sterile scalpel (Figure 9.5).
5. Slice the exposed length of pith into a sterile petri dish.
6. Cover the dish to keep pith sterile.
7. Aseptically slice 5-mm cross-sections of pith.
8. Transfer one cross-section to each plate of Tobacco Callus Initiation Medium.
9. Cover the dishes, seal with parafilm, and place in an incubator at 22–25°C.

Callus Multiplication
1. Obtain a plate of tobacco callus.
2. Aseptically divide the callus into smaller pieces.
3. Transfer divided callus pieces to fresh Tobacco Callus Initiation Medium.

Effects of Hormone Balance on Explant Growth and Morphogenesis

Background
Plant hormones, like animal hormones, are relatively small molecules that are effective at low
tissue concentrations. The two types of plant hormones used in this experiment are cytokinins and
auxins.
Cytokinins are derived from adenine and produce two immediate effects on undifferentiated
cells: the stimulation of DNA synthesis and increased cell division (Ting, 1982). Cytokinins also
produce a delayed response in undifferentiated tissue which is the formation of shoot primordia.
Both naturally occurring cytokinins, such as zeatin and synthetic analogs, such as kinetin,
demonstrate cytokinin effects (Figure 9.3). Although low tissue concentrations of cytokinins (e.g., 1
× 10-8 M zeatin) have noticeable effects, higher concentrations are found in actively dividing tissues
such as those of plant embryos and developing fruits.

Auxins are indole or indole-like compounds that stimulate cell expansion, particularly cell
elongation. Auxins also promote adventitious root development. Indoleacetic acid (IAA), a
naturally occurring auxin, and napthaleneacetic acid (NAA), a synthetic auxin, are depicted in
Figure 9.3. Only small amounts of auxin (1 × 10-6 M) are required to demonstrate an IAA response
and even smaller amounts of synthetic auxin (e.g., NAA) are required for a tissue response. The
likely reason for potency of synthetic auxins is their stability in plant tissue (i.e., the enzymes and
processes that degrade IAA do not "recognize" synthetic auxins). Synthetic auxins, then, are more
effective hormones that also last for an extended length of time. Furthermore, light influences the
physiological activity of IAA while synthetic auxins are not as light sensitive.
Plant hormones do not function in isolation within the plant body, but, instead, function in
relation to each other. Hormone balance is apparently more important than the absolute
concentration of any one hormone. Both cell division and cell expansion occur in actively dividing
tissue, therefore cytokinin and auxin balance plays a role in the overall growth of plant tissue. Since
hormone balance is presumably important to the overall effect on growth and morphological
changes, then the hormone differentials in each of the experimental media (A, B, and C) should
produce somewhat different effects on the growth and development of excised explants.

Source of Aseptic Explant Material
During seed development the embryos are formed with a placenta-like interface of intervening
tissues between parental vascular supply and the embryo proper. This circumstance depresses
passive migration of most foreign bodies and microorganisms into the developing embryo. If the
embryo which often develops aseptically is released from the seed by aseptic germination
procedures, then aseptic seedlings result. Any part of the aseptic seedling can be used as "in vitro"
experimental material. In this experiment three explant types will be used: hypocotyl (undeveloped
lower stem), epicotyl (shoot apex), and cotyledons.


Media Formulae
Media A, B, C, D, and E each contain the same complement of minerals, that is, salt base as in
Medium D (Appendix D). The effect of minerals alone on explant growth and development
constitute "basal growth rate" against which the effects of other media constituents can be
measured. Medium D, then, serves as the base-line control for endogenous growth. Medium E,
containing both essential minerals plus sucrose, constitutes the organic and inorganic control which
can be used as the base-line indicator of explant growth when both minerals and sucrose are
supplied. In addition to the substrate, sucrose, Medium E contains two organic growth factors,
inositol and thiamine, which promote sugar metabolism and general anabolic growth processes.
Medium E also contains additional phosphate thereby matching the phosphate concentrations of the
experimental media (A, B, C). The experimental media contain similar inorganic and organic
complements, but differ in hormone content.
Since cytokinins are derived from adenine, adenine sulfate has been added to each of the
experimental media (A, B, C). In addition, either kinetin or 2iP ([2-isopentenyl]-adenine), both of
which are synthetic cytokinins having immediate hormone activity, are supplemental cytokinins in
media A, B, and C. Of the three experimental media, Medium A contains the highest amount of
active cytokinin (30 mg/liter), while media B and C contain much lower amounts (2 mg/liter and 1
mg/liter, respectively).
Conversely, Medium A contains only a small amount of auxin (0.3 mg IAA/liter), while
Medium B contains a higher amount (2 mgIAA/liter). Medium C contains the lowest absolute
concentration of auxin (0.1 mg NAA/liter), but this synthetic auxin is more efficient in promoting
cell expansion and root formation than the naturally occurring auxin, IAA. Medium C, then, may
actually represent the formula with the highest physiological auxin activity.
Since cytokinin/auxin balance is reportedly important to the final overall effect on growth and
development, the results for each experimental media may be expected to differ. The balance

represented by Medium A is decidedly skewed towards a high cytokinin/low auxin ratio. Medium
B represents a more even distribution of cytokinin and auxin, while Medium C may have an
effectively higher auxin than cytokinin ratio because of the "in vivo" stability of NAA as well as its
effectiveness as an auxin.

Methods: Week 1 (Aseptic Seed Germination)
Materials:
cucumber seeds (tomato may also be used; sterilization procedure, Experiment 1)
95% ethanol
sterile jars, sterile water
25% chlorox, freshly prepared
sterile forceps or spatula
Procedure:
1. Sterilize seeds (cucumber) for 1 minute with 95% EtOH.
2. Rinse in sterile water
3. Sterilize in 25% bleach for 5 minutes and rinse three times with sterile water.
4. Transfer 10 seeds with a sterile forceps or spatula to a nutrient agar plate.
5. Incubate for 1 week (20–23°C).
Methods: Week 2
Seedling explants (approximately 1 cm in length) of aseptically germinated cucumber (or
tomato) can be cut from the seedlings as shown in Figure 9.4. Aseptic techniques including the use
of the laminar flow hood are necessary to evaluate growth experiments in which nutrient rich media
are used. An explant (hypocotyl, epicotyl, or cotyledon) should be placed on each of the following
experimental media:
A) Murashige Shoot Multiplication, Medium A,
B) Murashige Shoot Multiplication, Medium B, and
C) Murashige Shoot Multiplication, Medium C,
which contain different concentrations of growth hormones (Appendix D). For comparison, one
explant of each type should be placed on each control media:
D) Murashige and Skoog Salt Base, Medium D, and
E) Murashige Minimal Organic Medium + Sucrose with NaH2PO4⋅H20, Medium E.
Seal the petri dishes with parafilm to prevent desiccation and incubate at low light intensity until
next week. Record incubation conditions.


Discussion Questions
What effect does a hormone balance that is applied pharmacologically "in vitro" have on
seedling explant growth and morphogenesis? Which media formulations produce callus? To what
extent? On which explants? Were these results predictable? What media produce anomalous

Demonstration of "in vitro" Morphogenesis and Totipotency of Seedling Explants

A simple exercise demonstrating plant totipotency as well as the nutritional requirements of
different plant organs employs shoot tip and root tip explants cut from aseptically germinated
seedlings. Each type of explant (excised part of the intact organism) is transferred to three simple
tissue culture media.

Background
During seed formation, the developing embryo and associated tissues tend to exclude pathogens
and foreign materials that may be in the parent plant. Contents of the seed, then, are essentially
aseptic and the resultant seedlings can be maintained in the aseptic condition if the outer surface of
the seed (seed coat) is sterilized with sodium hypochlorite (or other surface sterilant) prior to
germinating the seeds in a sterile petri dish.

Methods: Week 1
The manipulations that are required for the germination of aseptic seedlings are outlined below
and illustrated in Figure 9.1.

1. Outside the laminar flow hood: Place several (5 to 10) tomato or lettuce seeds in a small petri
dish. Fill the petri dish with a 7% chlorox solution to which a drop of wetting agent has been
added. The soapy chlorox solution is usually a good surface sterilant. Swirl the seeds
intermittently during the 10- or 15-minute chlorox treatment.

2. Preparation for aseptic transfers: Begin by washing your hands and forearms with soap,
followed by swabbing with 70% ethyl alcohol (EtOH). Sterilize the laminar flow hood by
wiping the inside (top, sides, and bottom) with EtOH. Turn on the hood; 10–15 minute
operation of the hood before use insures aseptic conditions within the work area of the hood.
Continue to swirl the seeds intermittently during the chlorox treatment. Prior to actual aseptic
transfers inside the chamber, swab hands and forearms with EtOH again; also wipe the external
surface of the petri dish before placing it inside the hood. The hood should contain the
following: a large jar which can be used as a "sink," flasks of sterile water, forceps in a beaker
of ethanol, sterile filter paper (5–7 cm diameter filter paper can be sterilized in glass petri
dishes), and sterile petri dishes which can be used as the seed germination dishes.

3. Inside the laminar flow hood: Decant chlorox and replace with sterile H2O. Rinse this way
twice. Each rinse should rest 10 minutes. Prepare the sterile germinating petri dish by
retrieving a forceps from the 70% EtOH beaker. Using the sterile forceps remove three (3)
rounds of sterile filter paper from a sterile container and place them in the germinating dish
(sterile plastic petri dish). Finally, add 5–10 ml of sterile H2O to the seeds; decant seeds and
water into the sterile germinating dish and incubate at 25°C until the next laboratory. (Both
tomato and light-insensitive lettuce seeds germinate in the light. Since shoots become green but
roots remain white under these conditions, seedling morphology is recognized more easily when
light-germinated.)

Methods: Week 2
Examine the contents of the aseptic germinating dish without opening the lid. If there is no
fungal or bacterial contamination around the seedlings, proceed; if contamination exists, request a
dish of aseptic seedlings from the instructor. The manipulation required for the transfer of seedling
explants to Mineral Salts (M) and Minimal Organic (O) growth media are outlined below and
illustrated in Figure 9.2.
1. Swab chamber, hands, and upper/lower surfaces of petri dish with 70% ethanol.
2. Place germinating dish in transfer chamber.
3. Remove scalpel or scissors from the ETOH beaker already in the hood. Slip instrument between
sheets of sterile toweling to remove ETOH (ethanol).
4. Lift one edge of lid and cut off no more than 10 mm of root tip. Excise two root tips. Lower
lid. Place scalpel back into ETOH beaker.
5. Place tubes with sterile media into the transfer chamber. (Media formulae are given in
Appendix B.) Use one tube of Minimal Organic Medium (O) and one of Mineral Salts Medium
(M). Loosen these caps.

6. Remove forceps or inoculating loop from ETOH. Slip between sterile toweling to remove
ETOH.
7. Remove excised root tip from germinating dish and transfer to the surface of the Minimal
Organic Medium (O). Transfer second root tip to the surface of the Mineral Salts Medium (M).
Caution: pick up root tip by the severed end; damage to the apical meristem disrupts mitosis!
Measure or estimate length of root tips. Record.
8. Using aseptic technique as above, prepare and transfer one shoot tip into each type of media.
Pick up the shoot tip by the severed end and insert it part way into the medium with an overall
vertical orientation of the cotyledons and shoot tip. Record size and shape of shoot tip.
9. Place the four tubes in a slant rack under lights.
10. Examine cultures each week. Record observations on the amount of growth and morphogenesis
of both root and shoot cultures.


Observations
As cultures progress it should be possible to correlate size/shape changes with the nutrient
content of the medium. A third medium, the B-deficient Medium, contains the same mineral
constituents as does the Mineral Salts Medium (M) and the same amount of sucrose as the Minimal
Organic Medium (O), but is devoid of B vitamins. Thus, this medium is referred to as B-deficient (-
B). Shoot tips and root tips have been transferred to this demonstration medium. Growth on this
medium can be evaluated and compared with growth on student experimental media M and O. The
Mineral Salts Medium (M) is the basal growth medium, supplying essential mineral nutrients for
autotrophic plant growth (Appendix C).
Predict the resultant growth in each circumstance, then monitor the growth and development of
root and shoot explants in each medium (M, O, and -B) and evaluate the following (record
observations in Table 9.1):
1. Effect of B-vitamins on:
a) shoot growth (increase in size) and morphology (change of shape), and
b) root growth and morphology.
2. Effect of organic medium containing sucrose on:
a) shoot growth and morphology, and
b) root growth and morphology.
Optional: media M, O, and -B are set up with root tip and shoot tip explants in darkness. This
set of samples can be observed along with those in the light to evaluate the effect of light as well as
media contents on the growth and development of plant organs. A row labeled "etiolation" would
be added to the bottom of Table 9.1.

Experimental observations should include the following:
1. Parameters: temperature; light quality, duration, and intensity.
2. Drawings to scale.
3. Gross measurements (length, biomass accumulation, extent of morphogenesis, totipotency,
primordia, number of branches) after 1 week and 2 weeks.
4. Net changes (Table 9.1).
5. Does the irregular orientation of the shoot explant change the growth pattern? How can these
observations be explained?

Laboratory Requirements for Tissue Cultur

General Organization
Localize each portion of the tissue culture procedure in a specified place in the laboratory. An
assembly-line arrangement of work areas (such as, media preparation, glassware washing,
sterilization, microscopy, and aseptic transfers) facilitates all operations and enhances cleanliness.
Media (tissue culture and nutrient agar) are available from Carolina Biological Supply Co.,
Burlington, NC. Laminar flow hoods are available from several suppliers.

Glassware
Use glassware that has only been used for tissue culture and not other experiments. Toxic metal
ions absorbed on glassware can be especially troublesome. Wash glassware with laboratory
detergent, then rinse several times with tap water and, finally, rinse with purified water.

High-purity Water
Use only high-purity water in tissue culture procedures. Double glass distilled water or
deionized water from an ion-exchanger are acceptable. Water should not be stored, but used
immediately. Regular maintenance and monitoring of water purification equipment are necessary.
Purified water for tissue culture can also be purchased.

Plant Material
Plants used in tissue culture need to be healthy and actively growing. Stressed plants,
particularly water-stressed plants, usually do not grow as tissue cultures. Insect and disease-free
greenhouse plants are rendered aseptic more readily, so contamination rate is lower when these
plants are used in tissue culture procedures. Seeds that can be easily surface sterilized usually
produce contamination-free plants that can be grown under clean greenhouse conditions for later
experimental use.

Aseptic Technique
The essence of aseptic technique is the exclusion of invading microorganisms during
experimental procedures. If sterile tissues are available, then the exclusion of microorganisms is
accomplished by using sterile instruments and culture media concurrently with standard
bacteriological transfer procedures to avoid extraneous contamination.
Media and apparatus are rendered sterile by autoclaving at 15 lbs/inch2 (121°C) for 15 minutes.
The use of disposable sterile plasticware reduces the need for some autoclaving. Alternative
sterilization techniques such as filter sterilization must be employed for heat-labile substances like
cytokinins.
Aseptic transfers can be made on the laboratory bench top by using standard bacteriological
techniques (i.e., flaming instruments prior to use and flaming the opening of receiving vessels prior
to transfer). Aseptic transfers are more easily performed in a transfer chamber such as a laminar
flow hood, which is also preferably equipped with a bunsen burner (Bottino, 1981).
If experimental tissues are not aseptic, then surface sterilization procedures specific to the
tissues are employed. Common sterilants are ethyl alcohol and/or chlorox with an added surfactant.
Concentration of sterilants and exposure time are determined empirically.

Plant Tissue Culture Procedure - Background

1. INTRODUCTION
Plant tissue culture is the science of growing plant cells, tissues or organs isolated from the mother plant, on artificial media. It includes techniques and methods used to research into many botanical disciplines and has several practical objectives. Before beginning to propagate plants by tissue culture methods, it is necessary to have a clear understanding of the ways in which plant material can be grown and manipulated in ‘test tubes’. This chapter therefore describes the techniques that have been developed for the isolation and in vitro culture of plant material, and shows where further information can be obtained. Both organised and unorganised growth are possible in vitro.

1.1. ORGANISED GROWTH
Organised growth contributes towards the creation or maintenance of a defined structure. Itoccurs when plant organs such as the growing points of shoots or roots (apical meristems), leaf initials, young flower buds or small fruits, are transferred to culture and continue to grow with their structure preserved. Growth that is coherently organised also occurs when organs are induced. This may occur in vitro either directly upon an organ or upon a piece of tissue placed in culture (an explant), or during the culture of previously unorganised tissues. The process of de novo organ formation is called organogenesis or morphogenesis (the development of
form).

1.2. UNORGANISED GROWTH
The growth of higher plants depends on the organised allocation of functions to organs which in consequence become differentiated, that is to say, modified and specialised to enable them undertake their essential roles. Unorganised growth is seldom found in nature, but occurs fairly frequently when pieces of whole plants are cultured in vitro. The cell aggregates, which are then formed, typically lack any recognisable structure and contain only a limited number of the many kinds of specialised and differentiated cells found in an intact plant. A differentiated cell is one that has developed a specialised form (morphology) and/or function (physiology). A differentiated tissue (e.g. xylem or epidermis) is an aggregation of differentiated cells. So far, the formation of differentiated cell types can only be controlled to a limited extent in culture. It is not possible, for example, to maintain and multiply a culture composed entirely of epidermal cells. By contrast, unorganised tissues can be increased in volume by subculture and can be maintained on semisolid or liquid media for long periods. They can often also be used to commence cell suspension cultures. Differentiation is also used botanically to
describe the formation of distinct organs through morphogenesis.

2. TISSUE CULTURE

2.1. CULTURES OF ORGANISED STRUCTURES
Organ culture is used as a general term for those types of culture in which an organised form of growth can be continuously maintained. It includes the aseptic isolation from whole plants of such definite structures as leaf primordia, immature flowers and fruits, and their growth in vitro. For the purposes of plant propagation, the most important kinds of organ culture are:
• Meristem cultures, in which are grown very small excised shoot apices, each consisting of the apical
meristematic dome with or without one or two leaf primordia. The shoot apex is typically grown to give one single shoot.
• Shoot tip, or shoot cultures, started from excised shoot tips, or buds, larger than the shoot apices employed to establish meristem cultures, having several leaf primordia. These shoot apices are usually cultured in such a way that each produces multiple shoots.
• Node cultures of separate lateral buds, each carried on a small piece of stem tissue; stem pieces carrying either single or multiple nodes may be cultured. Each bud is grown to provide a single shoot.
• Isolated root cultures. The growth of roots, unconnected to shoots: a branched root system may be obtained.

• Embryo cultures, where fertilised or unfertilised zygotic (seed) embryos are dissected out of developing seeds or fruits and cultured in vitro until they have grown into seedlings. Embryo culture is quite distinct from somatic embryogenesis (see below). These types of cultures are described in more detail later in this chapter.

2.2. CULTURES OF UNORGANISED TISSUES
‘Tissue culture’ is commonly used as a collective term to describe all kinds of in vitro plant cultures although strictly it should refer only to cultures of unorganised aggregates of cells. In practice the following kinds of cultures are most generally recognised:
• Callus (or tissue) cultures. The growth and maintenance of largely unorganised cell masses,  which arise from the uncoordinated and disorganised growth of small plant organs, pieces of plant tissue or previously cultured cells.
• Suspension (or cell) cultures. Populations of plant cells and small cell clumps, dispersed in an agitated, that is aerated, liquid medium.
• Protoplast cultures. The culture of plant cells that have been isolated without a cell wall.
• Anther cultures. The culture of complete anthers containing immature pollen microspores. The objective is usually to obtain haploid plants by the formation of somatic embryos (see below) directly from the pollen, or sometimes by organogenesis via callus. Pollen cultures are those initiated from pollen that has been removed from anthers.

2.3. USING TISSUE CULTURES FOR PLANT PROPAGATION
The objective of plant propagation via tissue culture, termed micropropagation, is to propagate plants true-to-type, that is, as clones. Plants obtained from tissue culture are called microplants and can be derived from tissue cultures in three ways:
• from pre-existing shoot buds or primordial buds (meristems) which are encouraged to grow and proliferate;
• following shoot morphogenesis when new shoots are induced to form in unorganised tissues or directly
upon explanted tissues of the mother plant;
• through the formation of somatic embryos which resemble the seed embryos of intact plants, and which
can grow into seedlings in the same way. This process is called somatic embryogenesis. To obtain plants by the first two of these methods, it is necessary to treat shoots of an adequate size as miniature cuttings and induce them to produce roots. The derivation of new plants from cells, which would not normally have taken part in the process of regeneration, shows that living, differentiated plant cells may express totipotency, i.e. they each retain a latent capacity to produce a whole plant. Totipotency is a special characteristic of cells in young tissues and meristems. It can be exhibited by some differentiated cells, e.g. cambial cells and leaf palisade cells but not those which have developed into terminally differentiated structures (e.g. sieve tubes or
tracheids). Theoretically, plant cells, organs, or plants, can all be cloned, i.e., produced in large numbers as a
population where all the individuals have the same genetic constitution as the parent. Present tissue culture techniques do not permit this in every case and irregularities do sometimes occur, resulting in ‘somaclonal variants’ (Larkin and Scowcroft, 1981). Nevertheless, as will be described in the chapters, which follow, a very large measure of success can be achieved and cultures of various kinds can be used to
propagate plants.