Question 1

Plant cell fusion technology — Prior to fusion, researchers prepare large numbers of which cell type to enable somatic hybridization and cybrid formation?

Accepted Answer

Introduction / Context: Somatic hybridization allows fusion of plant cells from the same or different species to combine nuclear and cytoplasmic genomes, creating novel hybrids and cybrids. The prerequisite is removing rigid cell walls to permit membrane–membrane contact and fusion. Given Data / Assumptions: Plant cell walls prevent direct plasma membrane fusion. Enzymes such as cellulase and pectinase digest walls to yield protoplasts. Large populations of viable protoplasts improve fusion efficiency and regeneration success. Concept / Approach: Protoplasts are naked plant cells bounded only by the plasma membrane. They can be fused chemically (e.g., polyethylene glycol, PEG) or by electrofusion (short electric pulses). After fusion, hybrid cells are cultured to regenerate cell walls and whole plants via tissue culture, enabling transfer of traits like cytoplasmic male sterility or disease resistance. Step-by-Step Solution: Isolate target tissues; enzymatically digest walls to generate many protoplasts.Select and purify viable protoplasts by density or filtration.Induce fusion using PEG or electric fields.Culture fused cells on regeneration media to form callus and then plants. Verification / Alternative check: Microscopy confirms wall removal (spherical morphology) and fusion events (heterokaryons). Regenerated plants can be genotyped to verify hybrid status. Why Other Options Are Wrong: A single protoplast is insufficient for practical fusion work; large numbers are required. Intact-walled cells do not fuse efficiently. Different species can be fused, but only after wall removal. Bacterial spheroplasts are unrelated to plant somatic hybridization. Common Pitfalls: Neglecting the need for osmotic stabilization of protoplasts during handling; lysis is common without proper osmolarity control. Final Answer: Plant cells stripped of their cell wall (protoplasts)

Question 2

Clonal propagation—Which of the following are disadvantages of conventional (non-tissue-culture) methods compared with plant tissue culture? Consider how traditional nursery practices fare versus in vitro micropropagation in terms of speed, hygiene,…

Accepted Answer

Introduction / Context: Clonal propagation aims to multiply genetically uniform, high-performing plants. Conventional nursery methods (cuttings, layers, divisions) work, but they are slower and more exposed to pests, pathogens, and environmental constraints than plant tissue culture. This question asks you to pinpoint the disadvantages of conventional methods when compared with modern micropropagation. Given Data / Assumptions: Goal is rapid, large-scale clonal multiplication. Conventional methods rely on field or greenhouse conditions. Micropropagation (in vitro) offers asepsis, year-round production, and compact storage/transport. Concept / Approach: Identify which statements truly describe drawbacks of conventional (non-tissue-culture) propagation relative to micropropagation. Focus on disease status, multiplication rate, and logistics (storage/transport of planting material). Step-by-Step Solution: Check option (b): Conventional methods often have lower multiplication rates of disease-free plants because stock and environment are harder to sterilize and maintain pathogen-free.Check option (c): Conventional propagules (bulky cuttings, rooted plants) are heavier, require more space, and are harder to store/ship than small, clean in vitro plantlets.Option (a) is not a disadvantage of conventional methods; “multiplication of sexually derived sterile hybrids” is actually an advantage of micropropagation, not a negative for conventional methods.Therefore, the correct combined choice is (b) and (c). Verification / Alternative check: Commercial protocols for bananas, ornamentals, and forest trees favor micropropagation to achieve disease indexing, phytosanitary movement, and compact shipping—advantages that directly counter the limitations of conventional propagation. Why Other Options Are Wrong: (a) Misframed and not a disadvantage; sterile hybrids are better managed via tissue culture. (e) Incorrect because (b) and (c) are genuine drawbacks of conventional propagation. Common Pitfalls: Confusing “what tissue culture can do” with “what conventional methods lack.” Also, overlooking transport/storage economics when scaling nurseries. Final Answer: Both (b) and (c)

Question 3

Producing plant protoplasts—Which enzyme cocktail is required? Protoplasts can be released from suspension cultures, callus, or intact tissues by digesting the wall. Which enzymatic treatment is necessary to remove both cellulose and pectic componen…

Accepted Answer

Introduction / Context: Plant cell walls are composite structures rich in cellulose microfibrils embedded in a pectin-hemicellulose matrix. To generate protoplasts (cells devoid of walls but with intact plasma membranes), enzymes must dissolve both the cellulose fraction and the pectic matrix. Given Data / Assumptions: Target outcome: intact, viable protoplasts. Wall composition includes cellulose and pectin among other polymers. Osmotic stabilizers (e.g., mannitol) are present during digestion. Concept / Approach: Cellulases hydrolyze β-1,4-glucan chains in cellulose microfibrils; pectinases (pectolyic enzymes) depolymerize pectins in the middle lamella and primary wall matrix. Using both is standard to efficiently free protoplasts without excessive shear. Step-by-Step Solution: Prepare enzyme solution containing cellulase + pectinase in osmoticum and buffer at appropriate pH.Incubate finely cut tissue or callus under gentle agitation and controlled temperature.Filter, wash, and purify spherical protoplasts; assess viability with vital dyes.Culture in high-osmotic, nutrient-rich medium to permit wall regeneration and division. Verification / Alternative check: Microscopy confirms loss of wall boundaries and spherical morphology; yield and viability are highest when both enzyme classes are included. Why Other Options Are Wrong: (a) or (b) alone often yields poor release because the other matrix component remains intact. (d) Proteases do not target the main wall polysaccharides. (e) Amylases digest starch, not the structural wall. Common Pitfalls: Insufficient osmotic protection causes protoplast lysis; over-digestion decreases viability. Final Answer: Both cellulolytic and pectolytic enzymes

Question 4

Callus competence—What is the term for the ability of callus cells to regenerate a whole plant? Choose the precise biological term used in plant tissue culture for the capacity of component cells in a callus to form an entire plant under appropriate…

Accepted Answer

Introduction / Context: Plant tissue culture exploits the remarkable capacity of somatic cells to regenerate whole plants. The correct term for this inherent ability is “totipotency,” a foundational idea that underlies micropropagation, somatic embryogenesis, and organogenesis. Given Data / Assumptions: Callus cells can be induced to form shoots, roots, or somatic embryos. Appropriate hormonal regimes (auxin/cytokinin balance) and media are provided. Regenerated plantlets are genetically similar to the donor (barring somaclonal variation). Concept / Approach: Define and distinguish related terms. Totipotency is the potential to regenerate an entire organism. Dedifferentiation refers to the reversion of specialized cells to a meristematic state (often forming callus). Redifferentiation is the subsequent specialization into organs or embryos. Somaclonal variation denotes genetic/epigenetic changes arising during culture. Step-by-Step Solution: Identify the capability described: whole-plant regeneration from callus cells.Map that capability to the correct term: totipotency.Eliminate similar but distinct processes (de- and redifferentiation) and unrelated terms. Verification / Alternative check: Classic examples include regeneration from tobacco and carrot callus; single cells or small aggregates can yield complete, fertile plants. Why Other Options Are Wrong: Redifferentiation/dedifferentiation describe steps within regeneration, not the overarching capacity. Somaclonal variation is a by-product, not a capacity. Transdifferentiation is not the standard term in this context. Common Pitfalls: Using “redifferentiation” as a synonym for totipotency; they are not interchangeable. Final Answer: Totipotency

Question 5

Protoplast wall regeneration—Typical timeframe in nutrient medium After plating viable plant protoplasts on a suitable nutrient medium with osmoticum, approximately how long do they generally take to synthesize and deposit a new cell wall under stan…

Accepted Answer

Introduction / Context: Once plant protoplasts are isolated, the first milestone toward cell division is the regeneration of the cell wall. Knowing the typical timeframe helps practitioners schedule medium changes, monitor viability, and plan downstream steps like colony formation. Given Data / Assumptions: Healthy, enzyme-released protoplasts are plated in an osmoticum-balanced medium. Temperature, pH, and nutrients are appropriate for the species. No mechanical stress or contamination. Concept / Approach: Wall regeneration involves re-synthesis of cellulose and matrix polysaccharides at the plasma membrane, guided by cytoskeletal elements. Under standard conditions, visible wall deposition and rounding-to-polygonal transition are observed within a few days rather than hours or weeks. Step-by-Step Solution: Isolate protoplasts and culture under osmotic stabilization.Observe first signs of wall deposition with dyes (e.g., Calcofluor white) after several days.Expect most species to complete an initial wall sufficient for mitosis within 2–5 days, after which first divisions may occur. Verification / Alternative check: Microscopy (anisotropic birefringence, cellulose staining) confirms new wall formation typically in the 2–5 day window for many herbaceous species. Why Other Options Are Wrong: (b)–(d) are longer than usual for initial wall formation and risk culture decline. (e) A few hours is too short for robust wall re-assembly in most systems. Common Pitfalls: Underestimating the need for correct osmoticum; failure leads to lysis or delayed wall regeneration. Final Answer: 2–5 days

Question 6

Plant propagation techniques—Which method routinely employs girdling (ringing) as a step? Identify the propagation method in which a stem is partially girdled or wounded to induce adventitious roots while still attached to the parent plant.

Accepted Answer

Introduction / Context: Several vegetative propagation techniques exist in horticulture. Girdling (ringing) the bark to interrupt phloem transport is a hallmark step in air layering, a form of layering used to induce roots on stems still attached to the mother plant. Given Data / Assumptions: The method involves wounding or removing a bark ring. Rooting is encouraged at the wound site while the shoot remains attached. Moist medium (sphagnum/soil) is wrapped around the girdled area. Concept / Approach: Layering exploits the plant’s ability to form adventitious roots under hormonal and moisture cues. Girdling accumulates carbohydrates and auxins above the wound, improving rooting success. Once roots develop, the new plant is severed and potted. Step-by-Step Solution: Select a suitable stem; remove a narrow ring of bark or wound the stem.Apply rooting hormone and wrap moist medium; cover with plastic to retain moisture.Wait for root formation; then detach and plant the rooted layer. Verification / Alternative check: Standard horticulture manuals describe ringing/wounding specifically under layering, especially air layering (marcotting). Why Other Options Are Wrong: Grafting/budding transfer a scion or bud to a rootstock; no rooting at a girdled site. Cuttings are detached first; girdling is not typical. Micropropagation is in vitro and does not use bark girdling. Common Pitfalls: Confusing wounding for cuttings with true girdling used to initiate roots while attached. Final Answer: Layering (e.g., air layering/marcotting)

Question 7

Why subculturing resembles propagation by cuttings—choose the best reason In micropropagation, which statement best explains why subculturing is considered analogous to propagation by cuttings in conventional horticulture?

Accepted Answer

Introduction / Context: Subculturing (multiplication stage) in micropropagation parallels the logic of taking cuttings: you partition a parent shoot into smaller, autonomous units that then root or elongate into new plants. Understanding this analogy clarifies workflow design in the tissue culture lab. Given Data / Assumptions: Microshoot clusters are produced on cytokinin-rich media. Subculture involves dividing these clusters and transferring to fresh medium. Conventional cuttings also partition a plant into units that can root and grow independently. Concept / Approach: Focus on the operational similarity: physical separation of propagules and providing a growth-supportive environment. Unlike grafting/budding, no scion-rootstock union is created during subculture. Step-by-Step Solution: Identify the act of subdividing shoots (analogous to taking cuttings).Transfer each unit to optimized medium (analogous to rooting medium in cuttings).Allow independent development into plantlets, then root and acclimatize. Verification / Alternative check: Protocols describe regular subculture intervals where clusters are divided; this mirrors nursery schedules for taking and rooting cuttings. Why Other Options Are Wrong: (b) Refers to grafting terminology; not part of subculture. (c) Conventional cuttings are ex vitro, not in vitro. (d) and (e) overstate or misstate the practice. Common Pitfalls: Assuming subculturing equals grafting or that all propagation occurs in vitro; the analogy is about partitioning shoots, not method identity. Final Answer: It separates multiple microshoots and places each on fresh medium for independent growth

Question 8

Benefits of micropropagation (clonal propagation)—select the most comprehensive answer Which option best captures the major advantages of micropropagation for commercial and research applications, including speed, sanitation, and special genotypes?

Accepted Answer

Introduction / Context: Micropropagation is widely adopted in horticulture, forestry, and seed potato programs because it provides fast, clean, and scalable clonal production. The method is especially powerful for elite, sterile, or recalcitrant genotypes that cannot be propagated sexually. Given Data / Assumptions: Culture is performed under aseptic, controlled conditions. Source material can be indexed for pathogens to generate clean mother stocks. Multiplication stage can be iterated to achieve exponential plantlet numbers. Concept / Approach: Evaluate whether each listed benefit is a recognized strength of micropropagation. Speed (rapid multiplication), sanitation (disease-free plants), and the ability to propagate sterile hybrids are all hallmark advantages. Step-by-Step Solution: Confirm (a): Cytokinin-driven shoot proliferation enables rapid multiplication of elite clones.Confirm (b): Meristem culture and indexing help eliminate or avoid systemic pathogens, yielding clean stock.Confirm (c): Sterile or seedless cultivars (e.g., triploids) can be multiplied vegetatively in vitro.Therefore, the most comprehensive choice is “All of the above.” Verification / Alternative check: Commercial success stories (banana, sugarcane, ornamentals) consistently cite these advantages as the rationale for adopting tissue culture pipelines. Why Other Options Are Wrong: (e) omits the crucial benefit for sterile hybrids. Common Pitfalls: Assuming sterility prevents clonal propagation; micropropagation bypasses sexual reproduction entirely. Final Answer: All of the above

Question 9

Agrobacterium-mediated gene transfer—efficiency across plant groups With respect to natural transformation by Agrobacterium tumefaciens (Ti plasmid systems), which statement best reflects its inherent efficiency across major plant groups in c…

Accepted Answer

Introduction / Context: Agrobacterium naturally infects many dicotyledonous species, transferring T-DNA into the plant genome. Historically and inherently, this system was most efficient with dicots. Monocot transformation required later innovations (e.g., super-binary vectors, tissue wounding, specific genotypes), and biolistics was often used for cereals. Given Data / Assumptions: Classical textbooks emphasize dicot susceptibility in natural infections. The question asks about inherent efficiency, not modern engineered workarounds. Binary vectors and improved protocols have expanded host range post-hoc. Concept / Approach: Differentiate between the natural proclivity of Agrobacterium and the broad applicability achieved through advanced methods. The inherently efficient group is dicots. Step-by-Step Solution: Identify the naturally susceptible clade: dicots.Note that many monocots posed historical barriers (e.g., recalcitrant tissues).Conclude that “Only with dicots” captures the classical efficiency statement. Verification / Alternative check: Legacy transformation pipelines for tobacco, tomato, and petunia relied on Agrobacterium; cereal crops often required alternative methods until later protocol refinements. Why Other Options Are Wrong: (b) Contradicts the natural host range. (c) Overgeneralizes current capabilities as inherent. (d) Reverses the historical pattern. Common Pitfalls: Conflating today’s engineered versatility with the organism’s native propensity. Final Answer: Only with dicots

Question 10

Hormonal control in callus culture—Which regulator promotes shoots vs roots? In a plant callus culture, how do altered levels of cytokinins versus auxins typically influence organogenesis from undifferentiated tissue?

Accepted Answer

Introduction / Context: Organogenesis from callus depends largely on the ratio of two plant growth regulators: cytokinins and auxins. Mastering this balance is fundamental for predictable shoot and root induction in vitro. Given Data / Assumptions: Callus is competent to respond to hormone cues. Media components and culture conditions are otherwise suitable. Species differences exist, but the general trend holds widely. Concept / Approach: Classic Skoog and Miller findings established that a higher cytokinin/auxin ratio favors shoot primordia, whereas a higher auxin/cytokinin ratio favors root primordia. This hormonal antagonism is used throughout micropropagation protocols. Step-by-Step Solution: Start with callus on basal medium.Increase cytokinin (e.g., BAP, kinetin) to stimulate shoot bud formation.Alternatively, increase auxin (e.g., NAA, IBA, IAA) to promote rooting.Subculture regenerants to elongation/rooting media as needed. Verification / Alternative check: Histology shows meristem formation under cytokinin-rich conditions and root initials under auxin-rich conditions; this is standard across manuals and research articles. Why Other Options Are Wrong: (b) Reverses the well-known relationship. (c) and (d)/(e) ignore the synergistic requirement and balance between both hormones in most systems. Common Pitfalls: Using absolute concentrations instead of considering the ratio; species-specific fine-tuning is often necessary. Final Answer: Increasing cytokinin induces shoot formation; increasing auxin promotes root formation

Question 11

Definition—What is meant by “organ culture” in experimental biology? Choose the option that most accurately defines organ culture as used in tissue and organ culture techniques, emphasizing viability and partial immersion in nutritive media.

Accepted Answer

Introduction / Context: Organ culture is a classical technique in experimental biology distinct from cell or tissue culture. It preserves the three-dimensional architecture and many physiological interactions of an intact organ ex vivo, enabling pharmacological and developmental studies. Given Data / Assumptions: Whole organ (or large organ rudiment) is excised. Viability is maintained in vitro using nutrient media and appropriate gas exchange. Architecture and function are preserved better than in dissociated cell culture. Concept / Approach: Unlike dissociated cell culture, organ culture keeps tissue layers and extracellular matrices intact. Partial immersion (or interface methods) supports diffusion of nutrients while preventing anoxic conditions. This method sits between in vivo physiology and simplified cell culture models. Step-by-Step Solution: Excise the organ aseptically and place on a support at the air–liquid interface or partially immersed in medium.Maintain temperature, pH, and gas composition to mimic physiological conditions.Assess viability and function (e.g., electrical activity, secretion, development). Verification / Alternative check: Historical and modern protocols (e.g., embryonic organ rudiments) demonstrate maintained morphogenesis and responsiveness under organ culture conditions. Why Other Options Are Wrong: (b) Refers to transplantation and genetics, not organ culture. (c) Describes tissue engineering/organogenesis, not maintenance culture. (d) Social sciences; irrelevant. (e) Confuses cell culture with organ culture. Common Pitfalls: Assuming full immersion is optimal; many organs require an interface method to ensure oxygenation. Final Answer: Maintenance alive of a whole organ, after removal from the organism, by partial immersion in a nutrient fluid

Question 12

Plant tissue culture terminology — In micropropagation and in vitro culture, what does “organogenesis” specifically refer to?

Accepted Answer

Introduction / Context: In plant tissue culture and micropropagation, organogenesis describes the developmental process by which discrete organs such as shoots and roots are induced to form under controlled conditions. It is a core route for clonal propagation and regeneration after transformation. Given Data / Assumptions: Explants or callus are grown on media with defined auxin/cytokinin ratios. Callus refers to an undifferentiated cell mass. Organogenesis yields recognizable organs (shoots, roots) rather than whole embryos. Concept / Approach: Organogenesis can be direct (organs arise from the explant without an intervening callus phase) or indirect (organs arise from prior callus). The hormonal balance concept is key: high cytokinin relative to auxin favors shoot organogenesis, whereas higher auxin favors root organogenesis. Step-by-Step Solution: Recognize that “callus formation” alone is not organogenesis; it is a precursor in indirect routes.Identify organ formation (shoots/roots) as the defining feature of organogenesis.Match the correct statement to this definition: formation of roots and/or shoots from callus or directly from explants. Verification / Alternative check: Protocols demonstrate that altering the auxin:cytokinin ratio switches between callusing and organ formation. The presence of meristematic centers and subsequent organ primordia confirms organogenesis. Why Other Options Are Wrong: a) Callus formation is dedifferentiation, not organogenesis.c) Mixing stages obscures the definition; organogenesis is specifically organ formation.d) Too broad; not specific to in vitro regeneration.e) Somatic embryogenesis is a different pathway that produces embryo-like structures. Common Pitfalls: Equating callus induction with complete regeneration; forgetting that direct organogenesis can bypass callus. Final Answer: Formation of new roots and/or shoots from callus or directly from explant tissues.

Question 13

Plant micropropagation — In standard in vitro terminology, what is an “explant”? (Context: leaf or stem piece excised and placed onto culture medium)

Accepted Answer

Introduction / Context: An explant is the starting biological material for plant tissue culture. It can be taken from leaves, stems, roots, meristems, embryos, or other organs and placed onto sterile culture media to initiate regeneration. Given Data / Assumptions: Sterile technique is used to surface-sterilize plant material. Explants are plated on defined media containing macro-/micronutrients, vitamins, sugar, and plant growth regulators. Downstream outcomes can include callus, organogenesis, or somatic embryogenesis. Concept / Approach: Terminology precision is vital: “explant” refers to the initial plant piece; “medium” is the substrate; “microshoot” is a regenerated shoot; “scion” is a grafting term mostly outside micropropagation contexts. Step-by-Step Solution: Identify the role: the leaf or stem piece is the inoculum → explant.Differentiate from medium (nutrient base) and microshoot (a product of culture).Select the option that matches the definition: explant. Verification / Alternative check: Standard protocols label culture vessels with “explant type,” reinforcing that the tissue piece itself is the explant. Why Other Options Are Wrong: a) Microshoot is an outcome stage.b) Medium is the growth substrate.d) Scion is for grafting.e) Callus is a dedifferentiated mass, not the starting piece. Common Pitfalls: Confusing explant with callus or microshoot; the stage matters in SOPs and recordkeeping. Final Answer: Explant (the excised piece of plant tissue used to initiate culture).

Question 14

Feeder support in regeneration — What is used when culturing regenerating protoplasts, single cells, or very dilute plant cell suspensions?

Accepted Answer

Introduction / Context: Single plant cells and protoplasts are fragile and often require paracrine-like support to divide and regenerate. Nurse (feeder) systems provide conditioned media and/or living feeder cells that release hormones, peptides, and metabolites enhancing survival and morphogenesis. Given Data / Assumptions: Protoplasts lack cell walls initially and are osmotically sensitive. Very low cell densities reduce autocrine/cell–cell signaling support. Feeder layers or conditioned media can supply diffusible stimuli. Concept / Approach: Both a “nurse medium” (conditioned by an actively growing culture) and a “nurse/feeder culture” (living cells separated by a membrane or co-cultured) serve the same purpose: supplying growth factors and stabilizing the microenvironment for regeneration. Step-by-Step Solution: Identify the need: dilute or wall-less cells require support.Recognize two standard strategies: conditioned medium and feeder cells.Conclude that both approaches are valid solutions. Verification / Alternative check: Empirical data show increased plating efficiency and colony formation when using nurse systems versus unsupported minimal media. Why Other Options Are Wrong: d,e) Unsupported conditions typically yield poor survival/division for single cells or protoplasts. Common Pitfalls: Skipping osmotic stabilizers (e.g., mannitol, sorbitol) and appropriate Ca/Mg can negate nurse benefits. Final Answer: Both (a) and (b).

Question 15

Definition check — In plant biotechnology, “protoplasts” are cells that are devoid of which structural component?

Accepted Answer

Introduction / Context: Protoplasts are plant, fungal, or algal cells whose cell walls have been removed, leaving the plasma membrane intact. They are foundational for protoplast fusion, direct DNA uptake, and cellular studies requiring wall-less cells. Given Data / Assumptions: Wall removal is typically achieved using cellulases, pectinases, and hemicellulases. Osmotic stabilizers prevent lysis once the rigid wall is gone. Viable protoplasts retain functional plasma membranes. Concept / Approach: Differentiate between the cell wall (external, polysaccharide-rich) and the cell membrane (phospholipid bilayer). A protoplast must have an intact membrane to be living; therefore it cannot be missing the membrane. Step-by-Step Solution: Define protoplast = cell without wall.Confirm plasma membrane remains → cell is osmotically sensitive but viable.Select the option indicating absence of the cell wall only. Verification / Alternative check: Microscopy shows spherical morphology and plasmolysis sensitivity characteristic of wall-less cells. Why Other Options Are Wrong: a,c) Without a membrane the cell is non-viable; that is not a protoplast.d) Incorrect; protoplasts are specifically wall-less.e) Tonoplast may be present; its loss is not defining. Common Pitfalls: Using insufficient osmotic support leads to protoplast bursting; ensure isotonic conditions. Final Answer: Cell wall (removed enzymatically or chemically).

Question 16

Breeding and hybrid production — Which breeding method uses enzymatic/chemical removal of cell walls and fusion of protoplasts from two sexually incompatible plant species?

Accepted Answer

Introduction / Context: Protoplast fusion enables somatic hybridization between distantly related plants that cannot cross sexually. By removing cell walls, plasma membranes can be brought into close contact and fused to combine nuclear and cytoplasmic genomes. Given Data / Assumptions: Cell walls are removed using cellulases/pectinases. Fusion is induced by polyethylene glycol (PEG) and/or electric fields (electrofusion). Somatic hybrids and cybrids can be regenerated via tissue culture. Concept / Approach: The key distinction is wall removal plus fusion, which is unique to protoplast fusion. Mass selection and marker-assisted selection do not involve cell wall removal; transformation introduces DNA but does not fuse whole cells. Step-by-Step Solution: Generate viable protoplasts from two species.Induce fusion (PEG or electrofusion).Regenerate hybrid callus and select for hybrids. Verification / Alternative check: Hybrid cells can be confirmed by cytological markers, selectable markers, or organelle DNA profiling. Why Other Options Are Wrong: a,e) Selection strategies, not fusion methods.c) Transformation alters genomes but does not fuse cells.d) Irrelevant to breeding. Common Pitfalls: Losing viability due to osmotic shock; careful stabilization and plating density are critical. Final Answer: Protoplast fusion.

Question 17

Developmental biology concept — “Cellular totipotency” is best described as a property most broadly exhibited by which kingdom in practical biotechnology?

Accepted Answer

Introduction / Context: Totipotency refers to the inherent potential of a single cell to regenerate into a complete organism. In practical tissue culture, plant somatic cells frequently display totipotency, enabling mass propagation, rescue of hybrids, and regeneration after genetic transformation. Given Data / Assumptions: Plant cells can dedifferentiate and re-enter the cell cycle. Hormone regimes (auxin/cytokinin) guide differentiation into shoots and roots. Animal somatic cells are generally not totipotent (exceptions: very early embryonic cells). Concept / Approach: While the zygote of many organisms is totipotent by definition, the question targets practical, widely used totipotency in biotechnology. Plants uniquely and routinely express totipotency across diverse somatic tissues. Step-by-Step Solution: Identify the domain with routine whole-organism regeneration from somatic cells.Associate this with plant tissue culture successes across species.Select plants as the correct answer. Verification / Alternative check: Protocols for callus induction, organogenesis, and somatic embryogenesis in many crops (e.g., rice, tobacco) are standard and demonstrate totipotency. Why Other Options Are Wrong: b) Animal somatic cells are typically pluripotent at best when reprogrammed, not naturally totipotent.c) Single bacteria form colonies but not multicellular plants/animals.d,e) Incorrect generalizations. Common Pitfalls: Confusing pluripotency/induced pluripotency with totipotency; only totipotency yields a full organism including extra-embryonic tissues in animals. Final Answer: Plants (most somatic cells can regenerate a whole plant under the right conditions).

Question 18

Callus biology — The reversion of mature, specialized plant cells to a meristematic, dividing state that leads to callus formation is called what?

Accepted Answer

Introduction / Context: Dedifferentiation and redifferentiation are paired concepts in plant regeneration. Dedifferentiation first produces callus from mature tissues; subsequent redifferentiation gives rise to organs or embryos. Given Data / Assumptions: Explants originate from mature organs. Hormone regimes trigger cell cycle re-entry and loss of specialized features. Later stages direct formation of shoots/roots or somatic embryos. Concept / Approach: Define terms precisely: dedifferentiation = specialized to unspecialized; redifferentiation = unspecialized to specialized. The phenomenon described (mature cells returning to a meristematic state and forming callus) matches dedifferentiation. Step-by-Step Solution: Map description: mature → meristematic → callus.Select the term that fits: dedifferentiation.Note that redifferentiation occurs later during organ or embryo formation. Verification / Alternative check: Histological sections of induced callus show small, isodiametric, actively dividing cells typical of dedifferentiation. Why Other Options Are Wrong: a) Redifferentiation is the opposite stage.c) The terms are not interchangeable.d) Incorrect; a precise term exists.e) Transdifferentiation skips the callus-like stage. Common Pitfalls: Using “redifferentiation” for early callus; reserve it for organ/embryo formation. Final Answer: Dedifferentiation (mature cells reverting to a meristematic state).

Plant Cell Culture Questions