Unit 1: Introduction; Callus and Suspension Culture

Plant tissue culture is the aseptic growth of plant cells, tissues or organs on a defined nutrient medium under controlled physical conditions. The discipline rests on cellular totipotency — the idea, proposed by Gottlieb Haberlandt (1902), that every living plant cell retains the full genetic capacity to regenerate a complete organism.

• Totipotency: every nucleated somatic cell carries the entire genome and can, under the right signals, develop into a whole plant.
• Aseptic condition: cultures must be free of bacteria, fungi and other contaminants that outcompete slow-growing plant cells.
• Defined medium: growth depends on a chemically known supply of macro- and micronutrients, carbon source, vitamins and growth regulators.
• Controlled environment: temperature (usually 25 ± 2 °C), photoperiod (often 16 h light) and humidity are held constant.
• Dedifferentiation and redifferentiation: mature cells first lose specialisation to divide, then reorganise into new structures.

II. Introduction to Plant Tissue Culture

Origins, techniques and the culture systems that follow from totipotency.

The section establishes the historical arc, the working vocabulary and the standard laboratory operations on which all later culture work depends.

A. Historical background

The field advanced through a sequence of enabling discoveries.

• Haberlandt (1902): first cultured isolated palisade cells; failed to divide because he lacked growth hormones, but proposed totipotency.
• Kotte & Robbins (1922): achieved limited root-tip growth in culture.
• White (1934): established indefinite tomato root cultures on a defined medium (White's medium).
• Gautheret & Nobécourt (1939): obtained the first continuously growing callus of carrot and tobacco.
• Skoog & Miller (1957): showed the auxin : cytokinin ratio controls organogenesis — high cytokinin favours shoots, high auxin favours roots.
• Reinert & Steward (1958): demonstrated somatic embryogenesis from carrot cells, direct proof of totipotency.
• Murashige & Skoog (1962): formulated the high-salt MS medium, still the most widely used.

B. Concepts and basic techniques in tissue culture

The core operations convert a plant fragment into a sterile, growing culture.

• Inoculation: transfer of a sterilised explant onto medium under a laminar-flow hood.
• Subculturing: periodic transfer of growing tissue to fresh medium (every 3–4 weeks) to renew nutrients and remove toxic exudates.
• Aseptic technique: flaming of instruments, alcohol-swabbed surfaces and sterile transfer prevent contamination.
• Acclimatisation (hardening): gradual exposure of regenerated plantlets to greenhouse conditions to develop cuticle and functional stomata.

C. Media preparation and optimization

The medium supplies every nutrient the excised tissue can no longer obtain from the parent plant.

• Macronutrients: N, P, K, Ca, Mg, S supplied as nitrate, ammonium and phosphate salts at mM levels.
• Micronutrients: Fe (as Fe-EDTA chelate), Mn, Zn, B, Cu, Mo, Co at µM levels.
• Carbon source: sucrose (2–3 %) provides energy since cultures are largely heterotrophic.
• Vitamins: thiamine (essential), plus nicotinic acid, pyridoxine and myo-inositol.
• Growth regulators: auxins (2,4-D, IAA, NAA) and cytokinins (BAP, kinetin) direct division and morphogenesis.
• Gelling agent: agar (0.6–0.8 %) for solid medium; omitted for liquid medium.
• Optimization steps:
  • pH adjustment: set to 5.6–5.8 before autoclaving; higher pH stops agar setting.
  • Sterilisation: autoclave at 121 °C, 15 psi, 15–20 min; heat-labile substances (e.g. GA₃) are filter-sterilised and added after cooling.

D. Cell, tissue and organ culture

Culture systems are classified by the level of organisation of the starting material.

• Cell culture: free cells or small aggregates grown in agitated liquid (suspension), used for biomass and secondary metabolites.
• Tissue (callus) culture: unorganised proliferating cell mass grown on solid medium.
• Organ culture: intact organs maintained in vitro.
  • Meristem culture: shoot apical dome (0.1–0.5 mm) grown to produce virus-free plants.
  • Root, anther and embryo culture: cultivation of a specific organ to study or exploit its development.

E. Organogenesis

Organogenesis is the de novo formation of shoots or roots from cultured cells or callus.

• Hormonal control: governed by the Skoog–Miller principle of the auxin : cytokinin ratio.
  • High cytokinin : auxin → shoot bud (caulogenesis).
  • High auxin : cytokinin → root formation (rhizogenesis).
  • Balanced ratio → callus proliferation without organs.
• Direct organogenesis: organs arise straight from the explant with little intervening callus.
• Indirect organogenesis: organs form from an intermediate callus phase.
• Contrast with somatic embryogenesis: organogenesis produces a unipolar structure (only shoot or root) attached to parent tissue, whereas a somatic embryo is bipolar (shoot and root poles together) and independent.

F. Explant preparation & sterilization techniques

The explant is the excised plant part used to start a culture, and its decontamination is the critical first step.

• Explant choice: young, actively dividing tissue (nodal segment, shoot tip, leaf, cotyledon) regenerates best.
• Washing: running tap water plus a drop of detergent (Tween-20) removes surface soil and dust.
• Surface sterilisation: sequential chemical treatment in the laminar-flow hood.
  • Ethanol (70 %): 30–60 s to dissolve waxy cuticle and kill surface microbes.
  • Sodium hypochlorite (0.5–1 % available Cl) or mercuric chloride (0.1 %): 5–15 min as the main disinfectant.
  • Sterile-water rinses: three to four changes to remove residual sterilant.
• Trimming: cut off damaged, chemically burnt edges before inoculation.
• Contamination control: antioxidants (ascorbic/citric acid) or activated charcoal counter browning from oxidised phenolics released at cut surfaces.

III. Callus and Suspension Culture

Unorganised cell masses and their dispersion into free-cell suspensions.

Callus is a disorganised, proliferating mass of parenchyma-like cells formed when an explant dedifferentiates; suspension culture disperses that mass into liquid medium for rapid, uniform growth.

A. Initiation and maintenance of callus cultures

Callus is initiated by placing an explant on solid medium with the right growth regulators until dedifferentiated cells proliferate.

• Induction: an auxin such as 2,4-D (0.5–2 mg/L), often with a cytokinin, triggers dedifferentiation within 2–3 weeks.
• Growth phases: callus follows a sigmoid curve — lag, exponential (log), linear, deceleration and stationary phases.
• Maintenance: subculture every 3–4 weeks onto fresh medium; without transfer the tissue exhausts nutrients and accumulates toxic phenolics.
• Practical anchor: a tobacco pith explant on MS + 2 mg/L 2,4-D yields visible pale callus in about 14 days.
• Genetic caution: prolonged callus culture causes somaclonal variation — chromosomal and point changes that make old callus unreliable for clonal propagation.

B. Types of callus culture

Callus is categorised by texture and morphogenic capacity, which predicts its usefulness.

• Compact callus: hard, tightly packed cells; poor for making suspensions.
• Friable callus: loose, crumbly cells that separate easily; the preferred source for suspension cultures.
• Embryogenic callus: nodular tissue competent to form somatic embryos.
• Organogenic callus: capable of forming shoots or roots.
• Non-morphogenic callus: proliferates but has lost regeneration capacity.
• Pigmented callus: green (chlorophyll-bearing) or coloured from anthocyanins/other metabolites, used in secondary-metabolite work.

C. Initiation and maintenance of suspension cultures

A suspension culture is a population of single cells and small aggregates grown dispersed in agitated liquid medium.

• Initiation: transfer friable callus into liquid medium (same formulation minus agar) in a flask.
• Agitation: rotary shaker at 80–120 rpm breaks aggregates, aerates the medium and keeps cells suspended.
• Maintenance modes:
  1. Batch culture: a fixed volume of medium; cells grow through the sigmoid phases until nutrients are exhausted, then are subcultured — simple but growth conditions change continuously.
  2. Continuous culture: fresh medium is added and spent medium removed to hold cells in steady exponential growth.
     • Open: cells and medium both harvested, biomass held constant.
     • Closed: medium replaced but cells retained, allowing accumulation.
• Monitoring growth:
  • Packed cell volume (PCV): volume of cells after centrifugation, per unit culture volume.
  • Cell viability: stained by fluorescein diacetate (living cells fluoresce green).
• Applications: large-scale production of secondary metabolites (e.g. shikonin, berberine), protoplast isolation and biotransformation studies.
• Limitation: cells tend to re-aggregate and vary genetically, so scale-up requires filtering and regular subculture.