Chapter 15—
Plant Cell Culture
15.1—
Introduction
Almost every chapter of this book illustrates that research in plant cell physiology and biochemistry is, at each moment in time, limited by the resolving power of current experimental techniques and by the availability of appropriate experimental plant material. Here we are concerned with the development of new experimental materials (cultured organs, tissues and cells), with culture systems being evolved for their exploitation and with recent experimental work which illustrates that these culture systems are adding a new dimension to studies in plant cell physiology.
The complexity of higher plants has inevitably led workers, concerned with investigating particular aspects of plant physiology, to the use of systems which are of reduced complexity. Hence the early and continuing use by plant physiologists of isolated plant organs (e.g. seedling roots, leaves), complex tissue systems (e.g. discs cut from leaves and storage organs) organ segments (e.g. segments of coleoptiles and hypocotyls) and isolated cell organelles. However, isolated organs and organ fragments are still systems of very considerable complexity and they are, from the beginning, systems of declining viability and favourable sites for colonization by microorganisms.
The concept that the aseptic culture of isolated organs, tissues and cells would 'give some interesting insight into the properties and potentialities which the cell as an elementary organism possesses' and 'would provide information about the inter-relationships and complementary influences to which cells within the multicellular whole organism are exposed' was formulated as early as 1902 by Haberlandt. Then, after a lapse of more than 30 years, White (1934) described successful root cultures initiated from the root tips of tomato seedlings, and White (1939) and Gautheret (1939) demonstrated that the parenchymatous wound callus which frequently forms at the exposed surfaces of organ segments could be removed and grown indefinitely as a relatively undifferentiated tissue (callus) culture. From this period there has been rapid progress in organ and tissue culture. Root cultures have been developed from many species (Butcher & Street, 1964). Cultures of stem apices (meristem culture) and of leaf, flower and fruit primordia have been successfully established (Street, 1969). These organ cultures as experimental systems differ from isolated organs in two important respects; they are sterile (free from microorganism contamination) and are handled aseptically, and they are sytems where unimpaired viability is evidenced by growth involving both cell division and cell expansion. Such
cultures have contributed to our knowledge of the specific nutritional and hormonal requirements essential for the growth and development of the separate organs of the whole plant and of their specific physiology and biosynthetic activities (Street, 1969). Callus cultures have also now been established from a very wide range of species and have been used in studies on the initiation of root and shoot primordia (Street, 1975a), on cytodifferentiation (Wetmore & Rier, 1963, Gautheret, 1966; Torrey, 1971), on the induction of division in quiescent tissue cells (Yeomann & Aitchison, 1973), on the nature of plant tumour cells (Butcher, 1973) and on the synthesis of a diversity of secondary plant products (Yeomann & Aitchison, 1973).
In 1953, Muir reported that if fragments of callus cultures of Tagetes erecta or Nicotiana tabacum were transferred to liquid medium and agitated on a reciprocal shaker, then the callus fragments broke up to give a suspension of single cells and small aggregates of cells and that this suspension contained actively dividing cells and hence could be propagated by serial subculture (Muir, Hilderbrandt & Riker, 1954). Such liquid cell suspension cultures have now been obtained from calluses of a number of species and this chapter will outline the development of more sophisticated techniques for their culture and assess their value for studies on the control of growth, metabolism and differentiation in higher plant cells.
15.2—
Changes in Growth and Metabolism of Plant Cells in Batch Culture-Cytodifferentiation
Plant cells in batch culture, i.e. cultures in a fixed volume of culture medium, increase in biomass by cell division and cell growth until a factor in the culture environment becomes limiting and sends them into a stationary phase. When such stationary phase cells are subcultured they pass in succession through a lag phase, a short-lived period of exponential growth, a period of declining relative growth rate and then again enter stationary phase (Fig. 15.1A). Traditionally such cultures are initiated by an inoculum establishing a relatively high initial cell density and only accomplish a very limited number of divisions before entering stationary phase. For example cell cultures of sycamore (Acer pseudoplatanus ) initiated at ca. 2 × 105 cells ml–1 will reach a final cell density of ca. 3 × 106 cells ml–1 corresponding to 4 successive doublings of the initial population.
The degree of cellular aggregation in these cell cultures depends upon the species of cell, or cell line within a species, and the culture conditions, but always shows a basically similar pattern of change during the growth cycle of a batch culture. The culture at stationary phase contains the highest proportion of free cells and mean cell volume is at its maximum value. When subcultured to new
medium, the cells first embark upon a massive synthesis of new cytoplasm and associated organelles and then begin to divide. For a short time cell division proceeds at a specific growth rate (µ) which is constant and maximal (µmax ) for that culture environment. During this phase mean cell volume declines sharply and the proportion of cells in aggregates rises. Then the specific growth rate begins to decline (slowly at first and later at an ever increasing rate) mean cell volume increases and, associated with this cell expansion, the aggregates break up and release free cells. Associated with these growth and structural changes are changes in physiological activity. Measurements, on a per cell basis, of respiration, of the levels of individual cell constituents and of the activities of individual enzymes show that peaks of activity occur. These may be quite sharp and are not coincidental (Fig. 15.1A–D). Different metabolic patterns emerge and decline during the progress of batch culture. Thus RNA synthesis is initiated prior to cell division, proceeds for a time at a greater rate than cell number increase and then ceases whilst cell division is still proceeding (Short, Brown & Street, 1969; Nash & Davies, 1972). Free nucleotides (mainly UDP-glucose and ATP in sycamore cells) are synthesized rapidly during lag phase, presumably an essential preparation for subsequent synthesis of cell-wall polysaccharides and as an energy source for the endergonic processes of cell division, but their net synthesis ceases very shortly after the onset of division (Brown & Short, 1969). Similar transient high activity in carbohydrate oxidation by the pentose phosphate pathway during lag phase has been interpreted as providing the necessary NADPH for the massive biosynthesis achieved during the lag phase of the growth cycle (Fowler, 1971). By contrast, the very sharp peak in ethylene production in sycamore cell cultures occurs late in the cell division phase when the cells are beginning to increase in mean cell volume and may be responsible for initiating aggregate breakdown (Mackenzie & Street, 1970). An essentially similar pattern of ethylene production has been reported for cell cultures of Rosa spp., Glycine max, Triticum monococcum, Melilotus alba, Haplopappus gracilis and Ruta graveolens (La Rue & Gamborg, 1971). Large changes in the activity of phenylalanine ammonia-lyase (PAL) and in p -coumarate: CoA ligase occur prior to stationary phase in cultures of Glycine max (Hahlbrock, Kühlen & Lindl, 1971). A similar peak of PAL activity has been reported in cell cultures of Rosa sp. (Davies, 1972). These changes coincide with maximum production of total phenols by the cultures. Other secondary products are produced by cell cultures, the time of maximum synthesis being restricted to a phase in the growth cycle and often being markedly influenced in its intensity by the plant growth hormone composition of the culture medium (e.g. hemicelluloses and lignin by sycamore; see Carcellar, Davey, Fowler & Street, 1971; visnagin (a physiologically active furanochromone) by Ammi visnaga; see Kaul & Staba, 1967; caffeine by tea cell cultures; see Ogutuga & Northcote, 1970; various alkaloids, by cell cultures of solanaceous plants; see Tabata, Yamanito & Hiroaka, 1971). Thus during the progress of batch culture the cells pass through a series of contrasted physiological states which encompass cells
becoming meristematic, cells expressing high meristematic activity, cells undergoing expansion and becoming either metabolically quiescent or in which certain restricted metabolic pathways are emphasized.
Are these large changes in cellular structure and metabolic activity observed in batch-propagated cell cultures examples of cellular differentiation? This important question cannot at present be satisfactorily answered. Certainly cultured
Figure 15.1
Changes in metabolic activity during the progress of growth of sycamore
(Acer pseudoplatanus L.) cells in batch culture. (a) Growth curve and data for
total DNA and RNA per cell (from Short, Brown & Street, 1969.) (b) Data for total
cellular nitrogen and respiration (from Givan & Collin, 1969) and for ethylene production
(from MacKenzie & Street, 1970.) (c) Data for levels of nucleotides (ADP=adenosine
diphosphate, UMP=uridine monophosphate, UDP-G= uridine diphosphate glucose)
(from Brown & Short, 1969.) (d) Data for activities of glucose-6-phosphate dehydrogenase
(G-6-PD) and phosphofuctokinase (PFK) (from Fowler, 1971). (e) Data for cell wall invertase
assayed at pH 4.5 ( 
) and for soluble invertase assayed at pH 4.5 
and pH 7.0 
(from Copping & Street, 1972.)
cells do not correspond closely, either structurally or physiologically, with particular tissue cells of the plant body. To reproduce in culture the complete pattern of differentiation of selected specialized tissue cells is thus an objective not yet realized. Nevertheless the study of the origin in culture of particular physiological states and their cytological basis may advance our understanding of the molecular basis of cytodifferentiation in plants.
A recent book on cytodifferentiation in animal cells advances the widely accepted concept that the changes involved are consequent upon the activation of different sets of genes in different cell types and that this activation is expressed in terms of the synthesis of enzymes and other cellular proteins (Truman, 1974). Further, the differentiation process is regarded as a relatively permanent and irreversible change. In support of this it is possible to quote studies such as those of Cahn and Cahn (1966) on the culture of retina cells; such cells continue to produce the characteristic pigment granules during prolonged culture under conditions conducive to active cell division whereas in vivo such cells are non-dividing. If under certain secondary conditions of culture pigment granules were
lost, nevertheless when the cells were returned to the primary conditions of culture the pigmentation returned. To distinguish more minor and readily reversible changes in physiology from cytodifferentiation the former are described as modulations and are considered to reflect the operation of allosteric and other 'fine' processes of metabolic regulation. However, Truman (1974) concedes that 'it is not at all easy to draw a distinction between those enzymes which are fundamental to differentiation and those which represent very short-term modulations in the activity of cells' and in considering cytodifferentiation in liver cells states that 'differentiation and modulation do not represent distinct processes but are merely the extreme ends of a spectrum of changes that can occur'. Although from a number of plant species, cell cultures can be readily initiated from different organs (roots, stems, leaf petiole or lamina, cotyledons etc.) or from different living tissues within an organ (parenchymatous cells of pith, cortex, or mesophyll, cambial and other meristematic cells, immature vascular cells etc.) there is no very convincing evidence that they retain, in culture, characteristics of their in vivo origin although, as will be mentioned later in this chapter, they may not have undergone, during culture induction, the required degree of dedifferentiation necessary to express their totipotency (i.e. the capacity to generate a new plant in the way normally achieved from the fertilized egg). Such observations suggest that cytodifferentiation in higher plants, provided it has not proceeded to the point where cell death is inevitable, is a more readily reversible process than in the cells of higher animals. Hence the readily reversible physiological states observed in plant cell cultures may be basically identical with the states involved in normal cytodifferentiation. Further, it raises the possibility that cytodifferentiation does not depend on the transcriptional activity of different sets of genes for each kind of tissue cell but that the 'specialized' physiology of such cells may represent the influence of cytoplasmic factors (plant hormones?) on the stability and transport of RNA species and other aspects of the translational steps in gene expression. As discussed below, it may be possible to examine such hypotheses experimentally with plant cell cultures.
15.3—
Steady States of Growth and Metabolism of Plant Cells in Continuous Culture
Reference was made above to the short period of exponential growth observed in batch cultures. However, even during this phase of the growth cycle, the cells do not achieve a steady state (a state of balanced growth); cell division is uncoupled from increase in cell dry weight and protein content so that the cells are changing in size and composition despite the constancy of the double time of the culture (Fig. 15.2). Such observations raised the question of whether balanced growth could be achieved in plant cell cultures if a constant culture environment could be established by developing open continuous culture
systems (systems in which inflow of fresh medium is balanced by outflow of an equal volume of culture). Wilson, King & Street (1971) have developed such a system providing conditions of aeration and agitation appropriate to plant cell cultures, capable of long-term aseptic operation and functional either as a chemostat or as a turbidostat.
Figure 15.2
Unbalanced growth of sycamore cells (Acer pseudoplatanus L)
during the transient exponential growth phase achieved in batch culture.
(a) Semilogarithmic plots showing rate of change in cell number, total protein
and cell dry weight per unit volume of culture. The slopes of the lines of best
fit (calculated by linear regression analysis, P < 0.01) were used to determine
the specific growth rates (µ) for each parameter.
where x0 = initial value of parameter, x = value after time t(days). When
log10x plotted against t , slope = µ/2.303. (b) Changes in total protein
and cell dry weight per 106 cells with time calculated from data in A.
(From King & Street, 1973.)
15.3.1—
Chemostat Cultures
Subsequent work (King & Street, 1973; King Mansfield & Street, 1973) with this system operated as a chemostat (where equilibrium is established at a fixed rate of imput of a growth-limiting nutrient) has shown that long-term steady states of growth can be achieved with plant cell cultures and that such cultures conform to the chemostat theory developed from work with microorganisms (Monod, 1950; Novick & Sziland, 1950). In a chemostat, the relationship between cell density, x (cells per unit volume of culture), dilution rate, D (volume
of new medium added per unit time expressed as a fraction of the total culture volume), specific growth rate, µ (increase in biomass per unit biomass per unit time) and time (t ) is given by the equation
When equilibrium is reached and a steady state established dx/dt = 0, µ = D and x has a value characteristic of the dilution rate. Further, the nutritive environment remains constant, each nutrient achieving an equilibrium concentration which is related to its imput concentration and the rate of its consumption by the culture. As defined above, the equilibrium achieved in a chemostat culture results from one particular nutrient (depending upon the composition of the culture medium) becoming the limiting nutrient and determining the specific growth rate (µ) of the cells.
The nature of these steady states can be illustrated from work involving sycamore cell cultures growing in a synthetic medium (Stuart & Street, 1969) in which the supply of nitrogen is the limiting factor. Data for one such steady state (D = 0.194 day–1 ) is presented in Fig. 15.3. This shows that the cells are in a balanced state of growth and metabolism (as illustrated by the values for cell number, packed cell volume, cell dry weight, protein, DNA and RNA, and oxygen demand) and that the nutrient medium within the culture vessel is constant in composition (as illustrated by constancy of culture pH and the levels of glucose, phosphate and nitrate). Such steady state cells also display constant levels of metabolites (e.g. amino acids; Street, Gould & King, 1975) and constant levels of activity of individual enzymes (e.g. enzymes concerned with carbohydrate respiration; Fowler & Clifton, 1974; and with nitrogen assimilation; Young, 1973).
Such chemostat cultures can be operated from very low growth rates (i.e. low dilution rates) to growth rates approaching the maximum growth rate (µmax ) for the culture medium chosen, provided dilution rate is such that the cells can still achieve a matching growth rate. Of course if dilution rate is further increased, cell density does not stabilize and the culture suffers wash-out. Cells in balanced growth but highly contrasted in growth rate (and hence in cytology and metabolism) can therefore be obtained by chemostat culture. The range of change in certain cell parameters in sycamore cell cultures at different dilution rates (and hence specific growth rates) over the range D = 0.06–0.236 day–1 (corresponding to double times over the range 280–70 hr) is illustrated in Fig. 15.4. What this means is that it is possible to stabilize at will, by fixing dilution rate at an appropriate level, the individual physiological states which have only a transient existence in batch culture.
Work with chemostat cultures has shown that the same cell population can be taken through a series of steady states and then if the dilution rate is returned to that of an earlier steady state the cells again achieve not only the new predictable growth rate but also the physiological activities earlier recorded as
Figure 15.3
A steady state established in a 4-litre chemostat culture or A. pseudoplatanus cells. The
culture was diluted for 400 hrs. at a rate of 0.194 culture volumes per day. Samples were withdrawn
at intervals for biomass measurements, determinations of nutrient levels in the culture medium and for
respiration rate measurements. Culture opacity and pH were monitored continuously in the culture vessel.
(From King & Street, 1973.)
Figure 15.4
Steady state values for parameters of cell composition and physiological activities recorded
over a range of growth rates (as expressed by dilution rates) established in chemostat cultures
of sycamore cells (A. pseudoplatanus ) with nitrate/N as the limiting nutrient. The relationship
between dilution rate, D (fraction of culture volume displaced per unit time), specific growth rate
(m ) and doubling time (td) of the cell population in a steady state is given by 
(From King & Street, 1973.)
characteristic. Such experiments demonstate the full reversibility of the cytological and physiological changes invoked. Interest will now focus on detailed studies of the kinetics of the transition between steady states in terms of enzyme activities and metabolite levels. Preliminary studies along these lines have already shown that during these transitions there are pronounced oscillations in enzyme activity levels (characteristic for each enzyme monitored) which gradually decline in amplitude as the new steady state is established. How far this behaviour is to be explained in terms of changes in rates of enzyme synthesis and degradation has yet to be determined. Clearly study of these transitions will yield entirely new data on metabolic regulation in higher plant cells; whether it will yield the key to expose the changes underlying cytodifferentiation is less certain.
15.3.2—
Turbidostat Cultures
The constancy of culture opacity in the steady state described by Fig. 15.3 has formed the basis for a second form of continuous culture—the turbidostat system (a continuous system in which inflow of new medium occurs in response to an increase in the opacity-decrease in the light transmission of the culture). The turbidostat culture system developed for work with sycamore cell cultures is shown in Fig. 15.5 (Wilson, King & Street, 1971). Here, each time the population density exceeds a pre-selected value as determined by the optical density continuously monitored by the photocell, an electronically operated valve opens to admit a pulse of new medium. This reduces the optical density of the culture and the valve closes. These imputs of new medium are balanced by periodic harvesting of small volumes of culture in response to an electronically controlled level detector which controls the output valve. The effect is to produce a culture of constant volume and constant population density growing at a constant rate. Whereas in the chemostat growth rate must always be below µmax (the culture growth being limited by the supply of a chosen nutrient) here in the turbidostat growth can safely proceed under non-limiting nutrient conditions and one can study the effect of physical factors (e.g. temperature, light regime, CO2 tension) and growth regulators, in particular plant growth hormones, on growth rate (King, 1976), and by appropriate techniques (Gould, Baylis & Street, 1974) on the duration of the different phases of the cell cycle. This is the system where one can attempt to achieve conditions under which biomass increase expresses the maximum genetic potential for cell growth. Such a system offers an entirely new approach to studies on the molecular basis of the hormonal control of cell growth.
15.4—
Synchronous Cell Cultures—Study of the Cell Cycle
As indicated earlier in this chapter, plant cell cultures are routinely propagated by batch cultures initiated at a relatively high initial cell density (2 × 105 cells
Figure 15.5
The turbidostat culture system of Wilson, King & Street (1971). The 4-litre culture vessel is mounted
over a magnetic stirrer and is controlled in temperature by an internal water circulating coil. The culture
flows through an external circulation loop (exit and entry lines of this loop labelled CL) by the action of
the flow inducer (FI). This circulation loop flows through a cuvette (Fig. C. CU) in the density detector
(DD). As the cells divide in the culture its opacity increases and this alters the light transmission between
the lamp (L) and the light sensitive resistor (LSR) in the density detector (see Fig. C and D). When this
transmission falls below a preset value the optical monitoring unit (OMC) sends an impulse to the medium
imput solenoid value (MIS) and a pulse of new medium flows from the intermediate medium reservoir (IMR)
fed from the main medium reservoir (MR) via a filter unit (MFU). The size of this imput of new medium is
controlled by the observation chamber (OC) in the circulating loop since the imput solenoid valve closes only
when new medium displaces culture in the cuvette within the density detector. These imputs of new medium
are balanced by release of culture into the culture receiving vessel (CRV) via the outlet solenoid valve (OS)
operated by an outlet solenoid control unit (OSC) responding to the electrodes in the constant level device
(CLD). Thus the culture is maintained at a predetermined optical density (corresponding to a fixed cell number
per unit volume) and the rate of entry of new medium (and balancing harvest of culture) is a measure of the
growth rate of the culture. Samples of culture can at any time be withdrawn for cell counting and biochemical
analysis via a sample tube located to the right of the culture vessel and after collecting the aliquot of culture
this sample collector can be washed with sterile water from the reservoir WR. Periodically the excess culture
collected in CRV is withdrawn and its volume measured and the outlet protected by washing with mercuric
chloride solution from the reservoir MCR. The various ports in the lid of the culture vessel provide for
introduction of the initial inoculum of cells, for aeration ( via a glass sinter tube) for withdrawal of samples
into the sample collector, and for flow of water through the internal temperature-controlling glass coil.
ml–1 ) and subcultured when they enter stationary phase. It is, however, possible to initiate such batch cultures of sycamore cells at much lower cell densities (the minimum inocular density being ca. 1.5–4.0 × 104 cells ml–1 in the standard medium; see Stuart & Street, 1969) and to use as inoculum cells maintained in stationary phase as long as possible without suffering decline in viability. When this is done using an enriched synthetic medium (Stuart & Street, 1971) the cultures show a more extended lag phase and then embark upon a succession of highly synchronous divisions as evidenced by the data for cell counts (Fig. 15.6), mitotic index (percentage cells in a recognizable stage of mitosis) determinations and estimations of nuclear DNA content (by microdensity following Feulgen staining) (Fig. 15.7; see also Street, King & Mansfield, 1971; Gould & Street, 1975). By using for such synchronous batch cultures the 4-litre culture vessel developed for the continuous culture systems but modified by adding a stainless-steel sampling valve automatically operated by a timing device (Wilson, King &
Figure 15.6
Growth curve of a synchronous cell culture of sycamore ( A. pseudoplatanus )
showing log cell number ml–1 with time. The interphase plateaux separating
the synchronous increases in cell number are labelled P1 –P4 . The lag phase of this
culture was of 9 days duration and the culture reached stationary phase on day 24.
Street, 1971) it has been possible to show the synchrony of a number of metabolic events in the cell cycle (King et al., 1974).
The period during which DNA is doubled takes place during a restricted period of the interphase between successive mitoses. Howard and Pelc (1953) termed this the S phase and distinguished the interphase period before S phase as G1 (the first gap) and the period after completion of S phase and before mitosis (M) as G2 (the second gap). Cell cleavage (cytokinesis) usually follows directly upon mitosis so that G1 can be timed from the origin of the daughter cells although, as originally defined, G1 begins as soon as nuclear division is complete. These stages in the cell cycle can be determined in exponential asynchronous cultures by determinations of doubling time (cell count data), mitotic index, labelling index (per cent nuclei and mitoses labelled following flash label with tritiated thymidine—3 H-Tdr), fraction of labelled mitoses (following pulse-labelling with 3 H-Tdr; see Quastler & Sherman, 1959; and continuous labelling with 3 H-Tdr; see Cleaver, 1967), microdensitomitry and autoradiography, achieved by the single slide technique of Mak (1965). These methods have been applied to cultures of sycamore cell lines (Gould, Bayliss & Street, 1974) during the phase of exponential growth in batch cultures and to steady state chemostat cultures growing at different rates (doubling times ranging from 22 to 85 hr). This has revealed that the phases S (7.0 ± 0.2 hr), G2 (8.7 ± 0.6 hr) and M (2.9 ± 0.3 hr) are relatively constant whereas, according to the cell doubling time, G1 varies widely (4–60 hr). This observation that G1 varies with different cycle times, whereas S + G2 is relatively constant has also been observed in work with mammalian cells both in culture and in vivo (Mitchison,
1971). Certain critical events in G1 may be essential to the initiation of S phase and may be rate limiting. In the work with sycamore cells growing at reduced rates, nitrogen supply is the limiting factor and when cells enter stationary phase in batch culture, they are arrested in G1 (this arrest is important in relation to the synchrony which can be achieved by regrowth of nitrogen-starved cells; see Gould & Street, 1975).
Figure15.7
(A) Mitotic index fluctuations for P2 and P3 of Fig. 15.6. (B) Average nuclear
DNA contents over the same period. G1 and G2 values for nuclear DNA
content of the cell line are shown for comparison. [C] represents the
durations of the two successive periods of cell number increase.
(Data from Gould & Street, 1975.)
The value of synchronous cultures for studies on the cell cycle is that they enable particular metabolic events to be monitored. Work along these lines with plant cells has only recently been undertaken. In a study involving synchronous sycamore cell cultures (King et al., 1974) it was shown that total extractable protein and RNA rise throughout interphase but at an increased rate during S + G2. This was supported by studies of the rate of incorporation of labelled amino acids and uridine into these fractions. Respiration rate similarly rose throughout interphase but with two peaks of activity during S phase and cytokinesis. When, however, changes in the activity of extracted enzymes were studied different patterns emerged. Thymidine kinase and aspartate transcarbamoylase showed single and separated peaks of activity (Fig. 15.8), succinic dehydrogenase showed two well separated peaks of activity, and glucose-6-phosphate dehydrogenase activity rose continuously throughout interphase.
Whilst the content of total extractable histone rose parallel with total protein, study of the rates of incorporation of 3 H-labelled lysine and 14 C-labelled arginine into this fraction pointed to greatly enhanced histone turn-over associated with S-phase (Street, Gould & King, 1975). It is not known whether the changes in enzyme activity detected in these studies reflected changes in rates of enzyme synthesis. Yeoman (1974) in work on cell synchrony in explants of Jerusalem artichoke tubers has shown however, by using the deuterium labelling technique (Hu, Bock & Halvorson, 1962), that changes in the activity of glucose-6-phosphate dehydrogenase during interphase do result from changes in rate of synthesis of the enzyme.
Figure 15.8
Changes in the activities of thymidine kinase (TK). and aspartate
transcamoylase (ATC) during a cell cycle in a synchronized cell
culture of sycamore (A. pseudoplatanus ). C = duration of cytokinesis
from the cell count data. M = duration of mitosis from the mitotic index data.
(From King, Cox, Fowler & Street, 1974.)
If future work along these lines enables the many separate metabolic events of the cell cycle of plant cells to be chartered, we will then have a number of 'markers' (points where particular metabolic events are initiated or terminated). This will enable us to identify those events along the interphase plateau whose initiation or pace is affected by nutritional factors and plant hormones, to determine whether certain events only occur when their controlling genes (which could be 'mapped') are exposed during DNA replication and whether other processes which proceed continuously show gene dosage effects as
evidenced by increases in their rate after the points in DNA replication when their controlling genes are duplicated.
15.5—
Morphogenesis in Cell Cultures—Concepts of Totipotency and Determination
Various forms of morphogenesis have been observed in cell suspension cultures; this morphogenesis is apparently dependent upon the development in the cultures of relative large aggregates of cells, often several hundred in number, associated in a symplast. The morphogenesis can take the form of root initiation, shoot bud initiation or the development of somatic embryos (often referred to as embryoids) (Konar, Thomas & Street, 1972). Usually only one form of morphogenesis is expressed; sometimes the situation is more complex but in these cases one form is usually dominant—carrot cell cultures can be manipulated to show predominantly root initiation or exclusively embryoid development (Kessel & Carr, 1972).
15.5.1—
Somatic Embryogenesis
Work on embryogenesis in carrot cultures illustrates clearly the problems in cell physiology raised by the morphogenetic potential of plant cell cultures. These studies date from the report in 1958 (Steward, Mapes & Mears, 1958; Steward, 1958) of embryo-like plantlets in liquid carrot cultures and their continuing development when the cultures were plated out on agar-solidified medium. Further work quickly established the presence in such cultures of structures strikingly similar to the globular, heart-shaped and torpedo-shaped stages of normal embryology from the zygote.
Halperin and coworkers (Halperin & Wetherell, 1964; Halperin, 1967; Halperin & Jensen, 1967) and Street and coworkers (Smith & Street, 1974; McWilliam, Smith & Street, 1974) have shown that the embryos in carrot cultures arise from single cells at the surface of the cellular aggregates and are released, at various stages of development, as free-floating structures which, if they have already reached the advanced globular stage, are capable of completing their development in isolation from the parent embryogenic clump. These cultures grow actively, the embryogenic clumps proliferating and fragmenting due to enlargement and separation of cells in their interior, and do not form embryos in a synthetic medium containing sucrose, inorganic salts, thiamine, meso -inositol, kinetin and auxin (2, 4-dichlorophenoxyacetic acid). To initiate embryogenesis they are transferred to a similar medium, lacking auxin, which contains nitrate and ammonia (or urea or glutamine) and is at pH 5.0–5.4. Despite earlier claims, coconut milk is neither essential for growth nor for embryogenesis. Evidence that immature embryos have exacting nutritional
requirements (isolated embryo culture—see Street, 1969) and the contrasted simplicity of the culture medium which supports prolific embryogenesis in carrot cultures suggests that the embryogenic cell aggregate fulfils a 'nurse' role to the embryogenic surface cells and that the embryos must remain attached to the aggregate to complete their early development to the point where they can survive and mature into plantlets when released.
This phenomenon of somatic embryogenesis raises a number of questions of cellular physiology. The term totipotency has been introduced to describe the embryogenic competence of the single cells from which the embryos arise. At present it is only possible to establish new plants, whether via embryogenesis of via shoot bud initiation, followed by adventitious root development, from the cell cultures of a limited (if now quite large) number of species or varieties within species. Cultures which show this morphogenetic potential support the view that the pathways of cytodifferentiation which result in living tissue cells do not involve any loss or permanent inactivation of the genome and that they are, under appropriate environmental stimuli, completely reversible. Whilst however some cell cultures remain recalcitrant, this cannot be established as a universal principle. It may be that in such cases the conditions of culture fail to provide or permit the synthesis of an essential morphogen. Halperin (1967) has, on the other hand, advanced the very interesting hypothesis that the achievement of totipotency occurs during the initiation of the carrot culture from the primary explant (storage root or seedling organ) and that embryogenesis is expressed by cell clumps derived from such 'induced' cells, the primary culture consisting of both these and 'non-induced' cells. Retention of high embryogenic capacity in the cultures will then depend upon culture conditions favouring the active proliferation of the induced cells. On this hypothesis, failure to obtain embryogenesis in culture would have as its primary cause inappropriate conditions of callus initiation; the conditions of initiation would have effectively activated cell division and growth in the explant cells but failed to achieve the necessary 'dedifferentiation' to obtain cells with the competence of the zygote (a concept already raised here in previously discussing cytodifferentiation in cell cultures).
In classical plant embryology it has been considered that the early segmentations of the zygote (at least up to the 16-celled proembryo) follow a precise and species specific sequence which has phylogenetic significance (Johansen, 1950). During these divisions the cells of the proembryo are considered to inherit different cytoplasmic potentialities from the different regions of the zygote and these differences are regarded as determining from the beginning the exact role they and their daughter cells will play in constructing the embryo and its parts. This concept is often referred to as the theory of precise mosaic organization. The early segmentations involved in somatic embryogenesis have now been followed in a limited number of species (Street, 1976). Figure 15.9 illustrates the sequence of early embryology in carrot cultures. These segmentation sequences involved in somatic embryogenesis show more uniformity one with another than is depicted in the published accounts of the zygote embryology
of the species concerned. In summary, early development of somatic embryos involves enlargement and regular segmentation in a subspherical cell mass by walls of minimum surface. This supports the concept of D'Arcy Thompson (1942), based upon his studies in animal embryology, that surface tensions are important in determining the early segmentations of embryology and that only at a later stage do localized growth centres emerge whose functioning gives rise to the divergences in the morphology and anatomy of embryos of different species. This regulative theory of organization regards the early segmentations as being controlled by physical factors and as not involving any 'determination' of the early formed cells. With this background, the greater diversity and specificity recorded for zygotic embryogenesis can be interpreted as resulting from the physical restrictions and polar chemical gradients imposed upon the embryo as it develops within the ovule; when these influences are removed, as in cell cultures, the embryology reverts to a more basic or 'primitive' type of segmentation. This interpretation is supported by observations on the segmentations observed in natural polyembryony and by recent reports that indeed much more variable patterns of segmentation occur in ovule embryology than has hitherto been recognized (Jensen, 1965; Brown & Morgensen, 1972).
15.5.2—
Polarity of Embryogenic Cells
The recognition that the embryos arising in cell culture have their origin in superficial cells of the cell aggregates raises the question of whether such cells have any unique cytological characters and whether their observation can yield information on the physical basis of cell polarity. In carrot cultures proliferating in the presence of 2, 4-D (Street & Withers, 1974) these cells are small, rich in cytoplasm and with a large diffusely-staining nucleus containing a prominent nucleolus. Small vacuoles are clustered round the nucleus and each cell contains several amyloplasts containing prominent starch grains (Fig. 15.10). Study of these cells in the electrorn microscope shows the presence of numerous round and oval mitochondria and Golgi bodies and the regular presence of small numbers of lipid bodies (spherosomes). As might be expected from their meristematic activity, these cells are frequently observed in mitosis (in contrast to the expanded interior cells of the aggregates) and show numerous wall microtubules and limited arrays of cytoplasmic and nuclear microfibrils. When the cultures are transferred to auxin-free medium there is a transient increase in proliferation prior to the initiation of embryogenesis and the superficial cells show a change in segmentation pattern leading to the origin of 4-celled groups (Fig. 15.10). The individual cells in these groups either initiate an embryo, or by their further division promote the growth of the aggregate or undergo expansion (and senescence?) and become involved either in the release of the developing embryo or the break-up of the proliferating embryogenic aggregate. Associated with this changed segmentation pattern the densely cytoplasmic superficial cells
Figure 15.9
Earlier stage in the development of somatic embryos as observed in a cell suspension
culture of carrot (Daucus carota, L ). Scale lines on 1–4 = 20 m m, on 5–7 = 25 m m. Stages 1, 3,
5, 6 stained with periodic acid—Schiff (PAS), stage 2, 4 and 7 stained with toluidine blue (TB).
(From Street & Withers, 1974.)
show changes in fine structure. They have increased numbers of E.R. profiles (parallel arrays of rough E.R. profiles become particularly prominent) and of ribosomes. The Golgi bodies are more numerous and more compact. Additional large mitochondria (discs with swollen rims in outline) make their appearance and may come to occupy a considerable volume of the cytoplasm. The cells of the young proembryos show similar fine structure (Fig. 15.11). Although the first division of the embryogenic cells is by a wall parallel to the surface of the aggregate and at right angles to the longer axis of the cell (giving rise to an apical and a basal cell; the former being the first cell of the proembryo proper and the latter of the very variable suspensor), nevertheless fine structure studies do not reveal any prior asymmetry in the distribution of cytoplasm or cell organelles. Rather disappointingly, these studies have not revealed any unique features of the embryogenic cells or exposed any structural polarity. Perhaps this is not unexpected when we bear in mind the lack of any uniformity of fine structure in those angiosperm zygotes which have been studied (Jensen, 1965; Schulze & Jensen, 1969; van Went, 1970; Morgensen, 1972). Such studies only serve to emphasize that the special nature of embryogenic cells must now be approached at the level of molecular biology.
15.6—
Pollen Grains as Isolated Embryogenic Cells and as a Source of Haploid Cell Lines for Mutagenesis
This field of study was opened up by the pioneering studies on anther culture by Guha and Maheshwari (1964) working with Datura, and Bourghin & Nitsch (1967) working with Nicotiana. They showed that anthers, excised at an appropriate stage in pollen grain development and cultured in a simple medium gave rise to haploid embryos derived from individual pollen grains. Haploid plantlets can now be obtained by this technique from many species within the family Solanaceae. This approach extended to species in other angiosperm families has in a few cases yielded haploid callus but more frequently given a negative result. If we assume that immature pollen grains can, under appropriate stimuli, embark upon embryogenesis (sporophyte development) then the difficulty of achieving this with most excised anthers may be because conditions within the anther are too strongly promotive of the gametophyte pathway of development (i.e. that which gives functional pollen grains containing a tube nucleus and a generative cell). Recently however Nitsch (1974) in work with two Solanaceous species Nicotiana tabacum and N. sylvestris has shown that if anthers are excised at the peak of the first pollen mitosis, kept at 5°C for 48 hours and cultured for 4 days in a simple medium, then the immature pollen grains can be extracted. If this pollen grain suspension is cultured in a thin layer of liquid medium containing IAA, zeatin, glutamine and serine as supplements, some 10% of the grains embark upon embryogenesis. Although this work has so far
Figure 15.10
Embryogenic cell aggregates in suspension cultures of carrot ( D. carota ).
A and B from cultures in medium containing 0.1 mg 1–1 2,4-dichlorophenoxyacetic
acid (2,4-D), C and D from cultures in media with 2,4-D omitted. (A) Section through a
cell clump stained with PAS. Scale line = 200 µm. (B) Section stained with TB to show the
densely cytoplasmic superficial cells with nuclei surrounded with small vacuoles and more
internal cells with a large central vacuole. Scale line = 25 µm. (C) Localized superficial region
of active cell division. Arrows indicate characteristic pattern of cell segmentation which appears
following subculture to the auxin-free medium. Scale line = 10 µm. (D) General view of section through
a cell aggregate bearing numerous globular embryos at its surface. Stained with TB. Scale line = 200 µm.
(A & B from Smith & Street, 1974; C & D from Street & Withers, 1974.) (See also legend to Fig. 15.9.)
been restricted to Solanaceous plants, it could prove to be the key to inducing embryogenesis or haploid callus initiation in pollen grains of the many species resistant to the culture of whole anthers.
The importance of these studies is twofold. First is their potential for providing haploid plants (and by appropriate treatment homozygous diploids) and haploid cell cultures from many species and varieties. The haploid cell lines could be used as a source of mutant cell lines extending dramatically the field of the biochemical genetics of higher plants. Such mutant haploid lines could be rendered diploid by colchicine treatment and hence used to generate fertile mutants, provided their potential for morphogenesis was not impaired by the mutation. They could be preserved as cell culture lines by freezing preservation (Nag & Street, 1973, 1975a,b) as is already routinely done with mutant lines of bacteria. The use of such mutant cell lines in the continuous and synchronous culture systems previously described would permit more critical studies to be undertaken on particular metabolic sequences and their control and on the biochemical nature of the controls which operate in cell growth and division. Work along these lines is already actively proceeding (Widholm, 1974; Street, 1975b) but is currently handicapped by the need for more effective means of selecting the desired mutant cells which are present in very small numbers in the cell populations after mutagen treatment (e.g. 1 in 107 ).
The second important feature of these studies is in relation to embryogenesis. If within the pollen suspension, the pollen grains destined to embark upon embryogenesis are of characteristic size and/or density, it may be possible to isolate them by appropriate density gradient centrifugation. This would enable us to characterize embryogenic cells by the techniques of molecular biology, and to study in greater detail the segmentation patterns of early ermbryogenesis and the spontaneous expression of polarity in the proembryonal cell mass. Since the early stages of embryogenesis in pollen occurs within the enclosing pollen grain wall, study of the food reserves of such grains and of their biosynthetic activity may enable us to determine the special requirements of the proembryo.
This chapter has drawn attention to the availability of new experimental material and of new techniques in the field of plant cell culture. The exploitation of this approach in the molecular physiology of plants is still in its infancy. It is therefore important that the new generation of plant physiologists and biochemists should remain well informed of its present transient limitations and assured future prospects.
Figure 15.11
Light (A and B) and electron micrographs (C,D,E) of a four-celled somatic
embryo of D. carota. (A) The embryo located within a characteristic small
cell group (compare Fig. 10, C) at the surface of an embryogenic cell aggregate.
Scale line = 10 µm. (B) Enlarged view of embryo from A. The nucleus (n) of the basal
cell is seen to be surrounded with optically dense plastids (p). Scale line = 2.5 µm. (C)
The nucleus of the middle cell of the embryo surrounded by amyloplasts (p). Scale line
= 1 µm. (D) The two terminal (apical) cells of the embryo showing large rounded nuclei,
amyloplasts and the newly-formed thin dividing wall between them. Scale line = 2 µm. E.
The basal cell of the embryo separated from adjacent cells of the aggregate by a thick
plasmodesmata -free wall (cw). Parallel arrays of endoplasmic reticulum (er) can be seen
at the cell periphery. The nucleus (note glancing section showing nuclear pores–np)
is again surrounded by amyloplasts (p). Scale line = 1 µm. (From Street & Withers, 1974.)
Further Reading
King P.J., Mansfield K.J. & Street H.E. (1973) Control of growth and cell division in plant cell suspension cultures. Canad. J. Bot.51, 1807–23.
Ledoux L. (Ed.) (1975) Genetic Manipulations with Plant Material. Plenum Press, London. Les Cultures de Tissus de Plantes. (1971) Colloques Internationaux du C.N.R.S. No. 193, Paris.
Steward F.C. (Ed.) (1969) Plant Physiology. Vol. 5B. Academic Press, New York.
Street H.E. (Ed.) (1977) Plant Tissue and Cell Culture. Blackwell Scientific Publications, Oxford. (Second Edition.)
Street H.E. (Ed.) 1974) Tissue Culture and Plant Science, 1974. Academic Press, London.
Street H.E. (1976) Experimental embryogenesis—the totipotency of cultured plant cells. In The Developmental Biology of Plants and Animals. (eds Wareing, P.F. and Graham, C.F.). Blackwell Scientific Publications, Oxford.
Wilson S.B., King P.J. & Street, H.E. (1971) Studies on the growth in culture of plant cells. XII. A versatile system for the large scale batch and continuous cultures of plant cell suspensions. J. exp. Bot.21, 177–207.