Question 1

Cell biology sizing: The size of a typical single plant cell is usually within which range for diameter and length under light microscopy?

Accepted Answer

Introduction / Context: Knowing typical cell dimensions helps select magnification, plan microtomy, and interpret microscopy images. Plant cells vary widely across tissues, but many parenchyma cells fall within an accessible light-microscopy range. Given Data / Assumptions: We refer to common parenchymatous plant cells rather than giant specialized cells (e.g., xylem vessels) or tiny meristematic cells. Light microscopy with standard objectives (10×–40×) is assumed. Concept / Approach: Typical plant cells often measure on the order of tens of micrometres. A frequently cited “ballpark” for many tissues is roughly 10–20 μm in diameter, with elongated cells reaching 25–100 μm or more in length. Larger sizes (40–80 μm diameter and >200 μm length) occur in some tissues but are not the usual baseline expectation across plant organs. Step-by-Step Solution: Identify the common size domain for general plant cells. Exclude extremes (very large or very small) listed among distractors. Select the range that best matches standard teaching values. Verification / Alternative check: Calibrated eyepiece reticles or stage micrometers can confirm approximate dimensions in lab courses; many introductory texts illustrate similar ranges for epidermal and mesophyll cells. Why Other Options Are Wrong: Larger ranges (options b–d) reflect particular tissues or artifacts rather than typical averages. 1–5 μm: Typical of many bacteria and mitochondria, not whole plant cells. Common Pitfalls: Confusing chloroplast size (a few micrometres) with whole-cell size; also, conflating elongated fibre dimensions with average parenchyma cells. Final Answer: The typical size is 10–20 μm diameter and 25–100 μm length.

Question 2

Optical density for biomass: When measuring cell mass by light scattering (turbidity), readings are most commonly taken at which wavelength range?

Accepted Answer

Introduction / Context: Turbidimetry is a rapid method to estimate microbial biomass in liquid culture by measuring light scattering. The optical density at 600 nm (OD600) is the long-standing convention in microbiology labs for bacterial growth curves and kinetics. Given Data / Assumptions: Spectrophotometer or colorimeter with visible-range filters is used. Standard bacterial cultures (e.g., E. coli) in clear media are measured. Concept / Approach: At ~600 nm, absorbance by most media components is minimal while scattering by bacterial cells is sufficient for a linear correlation with biomass over a useful range. Shorter wavelengths (UV/blue) increase interference from nucleic acids and media; longer near-IR wavelengths may reduce sensitivity for small cells or require specialized optics. Hence, 600–700 nm is the common operational window, with OD600 being the canonical single-wavelength measure. Step-by-Step Solution: Blank the instrument with uninoculated medium. Measure culture turbidity at ~600 nm at intervals. Plot OD versus time to obtain growth phases (lag, exponential, stationary). Verification / Alternative check: Correlate OD600 with dry weight or cell counts to validate linear ranges for a given organism and path length; dilute samples above the linear range. Why Other Options Are Wrong: 300–500 nm: Higher background absorbance and potential photodamage; less standard for routine biomass. 500–600 nm: Usable but less canonical than OD600–OD660 range. 700–800 nm: Sometimes used for dense cultures or algae, but not the most common standard for bacteria. Common Pitfalls: Forgetting to mix cultures before reading; bubbles and clumps cause scattering artifacts. Always use the same cuvette path length. Final Answer: The conventional range is 600–700 nm, notably OD600.

Question 3

Plant tissue culture – definition focus: Callus cultures are amorphous, undifferentiated cell aggregates arising from the unorganized growth of explants on which type of culture medium and conditions?

Accepted Answer

Introduction / Context: In plant biotechnology, a “callus” refers to an unorganized mass of cells that proliferates from plant explants under culture. Identifying the precise conditions that promote callus formation is foundational for downstream applications such as organogenesis, somatic embryogenesis, protoplast culture, and genetic transformation. Given Data / Assumptions: Callus is undifferentiated plant tissue induced under controlled culture conditions. Explants are placed on nutrient media containing mineral salts, vitamins, a carbon source, and plant growth regulators (typically auxin and/or cytokinin). Plant tissue culture must be performed aseptically to avoid microbial contamination. Concept / Approach: While both liquid and solid media are used in plant tissue culture, classical callus induction is carried out on a solidified (e.g., agar or gellan gum) nutrient medium under sterile conditions. Solid surfaces help stabilize the explant and provide a uniform microenvironment where auxin–cytokinin balance promotes dedifferentiation and unorganized cell proliferation. Asepsis is non-negotiable: microbial contaminants outgrow plant cells and secrete inhibitory metabolites. Step-by-Step Solution: Recognize that “callus culture” implies undifferentiated growth from plant explants.Recall that solid media (with agar) are standard for initial callus induction.Note that all plant tissue culture must be aseptic to succeed.Therefore, select “aseptic solid nutrient medium.” Verification / Alternative check: Protocols for callus induction (e.g., from leaf discs or stem segments) universally specify sterile explant preparation and plating onto solid media supplemented with appropriate auxin/cytokinin ratios, confirming the chosen condition. Why Other Options Are Wrong: Liquid medium: used for suspension cultures, not ideal for initial callus initiation from intact explants. Solid nutrient medium (without “aseptic”): incomplete; sterility is essential. Solid or liquid medium: too vague; ignores the standard for callus induction and the requirement of asepsis. Hormone-free tap water: lacks nutrients, regulators, and sterility. Common Pitfalls: Assuming any medium will work; ignoring hormones; overlooking the absolute necessity of sterile technique. Final Answer: Aseptic solid nutrient medium.

Question 4

Archaea and antibiotic resistance in culture: The archaea Halobacterium (extreme halophile) and Sulfolobus (thermoacidophile) can be cultured in the presence of streptomycin and chloramphenicol because they possess which distinguishing feature?

Accepted Answer

Introduction / Context: Culturing extremophilic archaea often involves adding antibacterial antibiotics (e.g., streptomycin, chloramphenicol) to suppress bacterial contaminants. The rationale hinges on differences in the translation machinery between archaea and bacteria, which affects antibiotic sensitivity profiles. Given Data / Assumptions: Streptomycin targets the bacterial 30S ribosomal subunit; chloramphenicol targets the 50S subunit. Classical exam conventions attribute archaeal resistance to these antibiotics to differences in ribosomes compared with bacteria. The question follows standard test-bank phrasing used in many bioprocess/biotech exams. Concept / Approach: In many academic question banks, archaea are described as being resistant to certain bacterial protein synthesis inhibitors because their ribosomes are more “eukaryote-like,” conventionally summarized as 80S rather than 70S. Within that teaching framework, these antibiotics selectively suppress bacteria while allowing archaeal growth, aiding in pure culture maintenance under extreme conditions (high salt, low pH, high temperature). Step-by-Step Solution: Identify the target of streptomycin/chloramphenicol—bacterial-type ribosomes.Match resistance with the provided multiple-choice logic used in standard exams.Select the answer indicating 80S ribosomes, aligning with the conventional explanation for differential sensitivity. Verification / Alternative check: In practice, archaea show unique antibiotic sensitivity patterns due to ribosomal protein/RNA differences; practically, these agents help suppress bacterial contaminants in archaeal cultures. Why Other Options Are Wrong: Ether-linked lipids and lack of murein relate to membrane/wall composition, not direct targets of these translation inhibitors. All of the above: only the ribosome-related choice addresses antibiotic insensitivity in the conventional test context. Unique tRNAs alone do not explain resistance to these specific drugs. Common Pitfalls: Conflating cell wall or membrane chemistry with the mechanism of action of protein synthesis inhibitors. Final Answer: Contain 80S, rather than 70S, ribosomes.

Question 5

Animal cell culture fundamentals: Which statement is incorrect regarding mammalian (animal) cells compared to microorganisms in bioprocessing?

Accepted Answer

Introduction / Context: Mammalian cell culture underpins biopharmaceutical production (e.g., monoclonal antibodies) and research models. Understanding how mammalian cells differ from microbial systems guides reactor choice, media composition, and process control strategies. Given Data / Assumptions: Bacteria and yeasts divide rapidly (minutes to hours). Mammalian cells typically have doubling times of 18–36 hours or longer. Animal cells often require anchorage or microcarriers, are shear-sensitive, and need complex nutrients. Concept / Approach: Relative to microbes, mammalian cells grow slowly, demand intricate media (amino acids, vitamins, lipids, growth factors), and are more fragile, especially in stirred systems. Many lineages are anchorage-dependent, requiring a surface or microcarriers for proliferation and morphology. Therefore, any claim that they grow “very fast” compared to microorganisms is incorrect. Step-by-Step Solution: Compare doubling times: microbes (e.g., E. coli ~20 min) vs mammalian cells (>18 h).Assess process implications: longer batch times, lower space-time yields.Identify the incorrect statement as the one asserting faster mammalian growth. Verification / Alternative check: Bioprocess design references show markedly lower volumetric productivities without intensification strategies (perfusion, high cell density technologies) in animal systems versus microbial systems. Why Other Options Are Wrong: Larger/complex: true—organelles, cytoskeleton, and post-translational machinery. Fragile: true—sensitive to shear/foam, requiring gentle agitation and antifoams. Anchorage dependence: true for many primary and some continuous lines. Complex media: true—serum or defined supplements required. Common Pitfalls: Assuming scale-up rules for microbes directly apply to animal cells; underestimating shear protection needs. Final Answer: Their growth rate is very fast compared to microorganisms.

Question 6

Plant cell culture characteristics: Which of the following statements is incorrect regarding plant cells in culture compared with bacterial or fungal cells?

Accepted Answer

Introduction / Context: Plant cell and tissue cultures are key to micropropagation, metabolite production, and genetic engineering. Their physical and physiological characteristics differ markedly from microbial cultures, impacting reactor design, agitation, and process yields. Given Data / Assumptions: Plant cells are generally larger and vacuolated compared with microbes. Suspension cultures frequently form aggregates (clumps), altering mass transfer. Somaclonal variation (genetic instability) may arise during plant tissue culture. Concept / Approach: Due to large size, fragile cell walls, and vacuolation, plant cells are typically more shear sensitive than bacteria or fungi. They require gentler mixing (low tip speeds) and specialized impellers. Clumping is common, affecting oxygen/nutrient gradients and complicating homogeneous growth. Hormone balance (auxins/cytokinins) controls morphogenesis. Therefore, any statement claiming plant cells are “less shear sensitive” than microbes is incorrect. Step-by-Step Solution: List known traits: large size, aggregation, hormone dependence.Contrast shear tolerance: microbes > plant cells for typical agitation regimes.Identify the incorrect option as the one stating “less sensitive to shear.” Verification / Alternative check: Bioreactor guidelines recommend gentle agitation, bubble-free aeration (membrane sparging), and shear protectants (e.g., Pluronic F-68) for plant cells, confirming higher shear sensitivity. Why Other Options Are Wrong: Size larger: true. Clumps: true and widely observed. Genetic instability: true—somaclonal variation is a known issue. Hormone control: true—auxin/cytokinin ratios drive callus vs organogenesis. Common Pitfalls: Applying microbial agitation/aeration parameters to plant cultures; ignoring aggregate size control. Final Answer: Plant cells are less sensitive to shear than microbial cells.

Question 7

Extremophiles in biotechnology: Thiobacillus thiooxidans (Acidithiobacillus thiooxidans) exhibits an optimum pH for growth in which range?

Accepted Answer

Introduction / Context: Acidophilic sulfur-oxidizing bacteria such as Thiobacillus (now Acidithiobacillus) thiooxidans are instrumental in bioleaching and acid mine drainage studies. Knowing their pH preference is essential for both industrial applications and environmental management. Given Data / Assumptions: Acidithiobacillus thiooxidans oxidizes reduced sulfur compounds to sulfuric acid. It is highly acidophilic and thrives at very low pH. Neutral or alkaline pH inhibits growth of strict acidophiles. Concept / Approach: These organisms show optimal activity in the strongly acidic range, where proton gradients and enzyme systems are adapted to low pH. The commonly cited optimum is around pH 2–3.5, enabling efficient sulfur oxidation and metal solubilization in heap leaching or biooxidation reactors. Step-by-Step Solution: Identify the organism as a strict acidophile.Recall typical optimal pH range ~2–3.5 for growth and metabolism.Select the matching option “2.0–3.5.” Verification / Alternative check: Process guidelines for bioleaching maintain low pH to favor Acidithiobacillus species and suppress neutrophiles, corroborating the optimal range. Why Other Options Are Wrong: 0.5–6.0: overly broad and unrealistic for a defined optimum. 6.5–7.5 (neutral) and 9.0–9.5 (alkaline): inhibitory for this acidophile. 7.8–8.4: typical for alkaline conditions, not suitable. Common Pitfalls: Confusing acid-tolerant organisms with strict acidophiles; assuming broad pH tolerance. Final Answer: 2.0–3.5.

Question 8

Elicitation in plant cell and tissue culture: Which of the following are considered abiotic elicitors used to stimulate secondary metabolite production?

Accepted Answer

Introduction / Context: Elicitors are signals that enhance secondary metabolite biosynthesis in plant cell, tissue, and organ cultures. They are widely used to boost yields of valuable phytochemicals (alkaloids, phenolics, terpenoids). Elicitors can be biotic (microbe-derived) or abiotic (physical/chemical cues). Given Data / Assumptions: Abiotic elicitors include physical stresses and chemicals that mimic stress. Plant cells respond to stress by activating defense-related pathways. Typical readouts include increased accumulation of target metabolites. Concept / Approach: UV irradiation induces oxidative stress and signaling cascades; osmotic shock perturbs water potential and triggers stress pathways; heavy metal ions provoke reactive oxygen species and defense responses. All are classic abiotic elicitors used experimentally and industrially to enhance metabolite production in vitro. Step-by-Step Solution: Recognize UV, osmotic, and metal ion stresses as abiotic stimuli.Connect these stresses to activation of defense gene networks (e.g., PAL, CHS) and metabolite accumulation.Select the inclusive option “All of these.” Verification / Alternative check: Case studies show higher flavonoid or alkaloid titers after applying abiotic elicitors, confirming their practical efficacy in culture systems. Why Other Options Are Wrong: Individual choices are correct but incomplete. Only live microbial suspensions: these are biotic elicitors, not abiotic. Common Pitfalls: Overstressing cultures leading to growth arrest; not optimizing dose and timing for elicitor application. Final Answer: All of these.

Question 9

Culture media design: Radical (sudden) shifts in culture pH can be prevented most effectively by incorporating which component?

Accepted Answer

Introduction / Context: Maintaining a stable pH is critical for microbial, plant, and animal cell cultures because enzyme activities, membrane transport, and growth kinetics are pH-dependent. Media are therefore formulated to resist pH excursions produced by metabolism (acid or base generation). Given Data / Assumptions: Cells produce acidic or basic by-products during growth. Uncontrolled pH swings can inhibit growth or kill cultures. Buffers resist pH change by reversible protonation/deprotonation near a defined pKa. Concept / Approach: A buffer is a conjugate acid–base pair (e.g., phosphate, HEPES, MOPS) that stabilizes pH. Oxidizing or reducing agents affect redox state, not pH directly. Proper buffer selection considers desired pH, buffer capacity, compatibility with cells, and downstream processes (e.g., avoiding precipitation with divalent cations). Step-by-Step Solution: Identify the problem: metabolic acids/bases accumulate.Apply the solution: add a buffer system targeting the operating pH.Choose “A buffer” as the direct, effective method to prevent radical pH shifts. Verification / Alternative check: pH profiles in buffered vs unbuffered cultures show damped changes in buffered media, validating the role of buffers in stability. Why Other Options Are Wrong: Oxidizing/reducing agents adjust redox, not pH buffering capacity. “Any of these” and surfactants: do not specifically address pH stabilization. Common Pitfalls: Using buffers outside their effective pKa range; ignoring CO2/HCO3- equilibria in CO2-incubated systems. Final Answer: A buffer.

Question 10

Anchorage dependence in cell culture: Anchorage-dependent (adherent) animal cells require what kind of surface property to attach and grow properly?

Accepted Answer

Introduction / Context: Many animal cells (e.g., fibroblasts, epithelial lines) are anchorage-dependent, meaning they require attachment to a substrate for survival, proliferation, and differentiation. This principle informs the design of cultureware (treated plastic, glass) and microcarrier systems used in bioprocessing. Given Data / Assumptions: Attachment involves integrins and extracellular matrix (ECM) interactions. Surface energy and hydrophilicity influence protein adsorption and cell adhesion. Certain lines are adapted to suspension, but many are not. Concept / Approach: Wettable (hydrophilic) surfaces, often plasma-treated plastics or glass, adsorb serum proteins (e.g., fibronectin, vitronectin) that present adhesion motifs to cell receptors, enabling attachment and spreading. Hydrophobic, non-wettable surfaces resist protein adsorption and typically do not support adherent growth without additional coatings. Step-by-Step Solution: Define anchorage dependence: need for substratum attachment.Identify surface requirement: wettable, tissue-culture–treated plastic or glass.Select “Wettable surface for growth.” Verification / Alternative check: Manufacturers specify “tissue culture–treated” ware to increase surface wettability and charge, improving cell adhesion and spreading, confirming the necessity of wettable surfaces for adherent lines. Why Other Options Are Wrong: Dry, non-wettable surfaces: poor protein adsorption and adhesion. Either (a) or (b): incorrect—surface properties matter. Attachment unnecessary: not true for anchorage-dependent cells. Only metallic: metals are not standard substrates for routine culture. Common Pitfalls: Attempting to culture adherent cells on non-treated plastics; omitting ECM coatings when needed. Final Answer: Wettable surface for growth.

Question 11

Applications of plant protoplasts: Protoplasts (cell wall–removed plant cells) can be used for which of the following purposes in plant biotechnology?

Accepted Answer

Introduction / Context: Protoplast technology removes the rigid plant cell wall enzymatically, yielding naked plant cells enclosed only by the plasma membrane. This increases experimental access for genetic manipulation, fusion, and virology studies, and provides a platform for regeneration into whole plants in many species. Given Data / Assumptions: Protoplasts can take up DNA, RNA, and organelles more readily than walled cells. Somatic fusion allows combining genomes/cytoplasms from different species or varieties. Viruses can be inoculated efficiently into protoplasts to study infection cycles. Concept / Approach: Wall removal enables direct genetic modification (PEG- or electroporation-assisted transformation), somatic hybridization (protoplast fusion creating cybrids/hybrids), and controlled viral infection studies. Regeneration protocols (callus → shoots/roots) allow recovery of plants from transformed or fused protoplasts, making the technology versatile across research and breeding. Step-by-Step Solution: Identify genetic modification: DNA uptake and expression in protoplasts.Identify hybrid creation: somatic fusion merges protoplasts from different parents.Identify virology use: controlled virus inoculation and replication assays.Select “All of these” since all are valid applications. Verification / Alternative check: Literature shows successful creation of somatic hybrids (e.g., citrus, Brassica), protoplast transformation systems, and robust protoplast-based viral replication studies. Why Other Options Are Wrong: Any single choice is incomplete. None of these: contradicts extensive use of protoplast methods. Common Pitfalls: Failing to optimize osmotic stabilizers; inadequate regeneration protocols leading to callus that cannot redifferentiate. Final Answer: All of these.

Question 12

Bacterial physiology: For most non-extremophilic bacteria, the optimum pH for growth lies in which range?

Accepted Answer

Introduction / Context: Most bacteria commonly encountered in laboratory and clinical settings are neutrophiles, preferring near-neutral pH for optimal enzyme activity and membrane function. Understanding pH preferences helps in designing culture media and controlling microbial growth in industry and healthcare. Given Data / Assumptions: Neutrophiles favor pH close to neutrality. Enzyme activity and transport processes drop outside optimal pH. Acidophiles and alkaliphiles are special cases, not “most bacteria.” Concept / Approach: The majority of laboratory bacteria grow best between pH 6.5 and 7.5. Media such as nutrient broth and tryptic soy broth are buffered around neutrality to sustain growth. Deviations require specialized buffers and adaptations (e.g., for acidophiles like Acidithiobacillus or alkaliphiles like Bacillus alcalophilus). Step-by-Step Solution: Classify bacteria: neutrophiles vs acidophiles vs alkaliphiles.Recall the standard neutral range for neutrophiles: ~6.5–7.5.Select “6.5–7.5” as the correct optimum range for most bacteria. Verification / Alternative check: Growth curves for common strains (E. coli, Staphylococcus aureus) peak near neutral pH, validating the stated range. Why Other Options Are Wrong: 3.5–6.5 ranges: too acidic for most non-specialized bacteria. 8.5–9.5: alkaline range typical of alkaliphiles, not most species. Common Pitfalls: Assuming a single pH suits all species; ignoring buffer capacity and CO2 effects in incubators. Final Answer: 6.5–7.5.

Question 13

In microbial physiology and bioprocess engineering, “balanced growth” is best defined as a growth state in which a population increases while maintaining a constant overall chemical composition and a constant specific growth rate. Which statement ma…

Accepted Answer

Introduction / Context: Balanced growth is a foundational concept in microbiology and bioprocess engineering. It refers to a physiological state where every measurable component of the cell population (RNA, DNA, proteins, lipids, metabolites) increases in fixed proportion, so the overall composition remains constant while cell numbers and biomass rise exponentially. Understanding this state is critical for designing reproducible experiments and scalable fermentation processes. Given Data / Assumptions: A well-mixed, nutrient-replete environment (for example, early exponential phase or steady-state chemostat) is considered. No limiting toxic byproducts accumulate during the observation window. Specific growth rate (mu) is constant. Macromolecular composition per cell remains constant over time. Concept / Approach: In balanced growth, cellular synthesis is coordinated. Ratios such as RNA/protein or protein/DNA do not drift, and the cell size distribution is stable. This contrasts with “unbalanced growth,” where a shift in nutrients, temperature, or pH causes compositional changes. Experimentalists often enforce balanced growth via continuous culture (chemostat) at fixed dilution rate and feed composition, producing steady-state conditions. Step-by-Step Solution: Identify the key hallmark: constant chemical composition while biomass increases. Link it to constant specific growth rate under steady conditions. Evaluate options: only one statement captures both growth and compositional constancy. Select the definition that directly states “cultures undergoing balanced growth while maintaining a constant chemical composition.” Verification / Alternative check: In a chemostat at steady state, cell composition, substrate concentration, and growth rate remain constant over many residence times—an empirical confirmation of balanced growth. Why Other Options Are Wrong: Controlling pH or temperature: Important operational controls, but they do not define balanced growth by themselves. Changing nutrient composition: Typically induces unbalanced growth due to physiological shifts. Common Pitfalls: Confusing stable culture conditions (for example, constant pH) with true balanced growth. The essence is proportional synthesis and constant composition, not merely environmental control. Final Answer: cultures undergoing balanced growth while maintaining a constant chemical composition.

Question 14

In plant and microbial biotechnology, suspension cultures consist of single cells and small aggregates growing dispersed in which type of medium?

Accepted Answer

Introduction / Context: Suspension cultures are widely used in plant tissue culture and microbial bioprocessing for rapid growth, homogenous sampling, and scalable product formation (for example, metabolites, recombinant proteins). Recognizing the correct growth environment is essential for selecting agitation, aeration, and bioreactor configuration. Given Data / Assumptions: Cells are present as discrete single cells or small clumps (aggregates). A well-mixed environment is required to keep cells suspended. Nutrient and gas transfer must be adequate throughout the culture. Shear should be controlled to avoid aggregate disruption or cell damage. Concept / Approach: By definition, a “suspension” implies particles freely dispersed in a fluid. In biological suspensions, cells are suspended in a liquid phase, maintained by agitation (shakers, stirrers) and sometimes by baffles or impellers in bioreactors. Solid media (agar plates) support colonies but do not produce cell suspensions; that is a hallmark of liquid systems. Step-by-Step Solution: Match the word “suspension” to a fluid phase compatible with mixing. Exclude solid media, which yield colonies rather than suspended cells. Identify liquid medium as the only environment that keeps cells dispersed. Select “liquid medium.” Verification / Alternative check: Standard protocols for plant cell suspensions begin with callus on solid media, then transfer friable callus to liquid medium for shake-flask culture—validating the definition. Why Other Options Are Wrong: Solid nutrient medium: Supports colonies, not suspensions. Solid or liquid medium: Overbroad; suspension specifically refers to liquid. None of these: Incorrect because liquid medium is correct. Common Pitfalls: Assuming any medium that supports growth qualifies. “Suspension” specifically denotes dispersed growth in a liquid phase. Final Answer: liquid medium.

Question 15

Semi-solid media prepared with agar at concentrations of ≤0.5% are especially useful for which cultivation purpose in diagnostic microbiology?

Accepted Answer

Introduction / Context: Semi-solid media (low agar, typically 0.3–0.5%) create a soft matrix that restricts convection and diffusion differently than liquids or firm gels. They are widely used for motility tests and for cultivating organisms that prefer reduced oxygen tensions—such as microaerophiles. Given Data / Assumptions: Agar concentration ≤0.5% yields a soft gel. Oxygen gradients form naturally from the surface to deeper zones. Motility can be visualized as diffuse growth away from the stab line. Microaerophiles grow best in regions of limited oxygen. Concept / Approach: Microaerophilic bacteria require oxygen but at lower levels than atmospheric. Semi-solid media allow stable microenvironments and dampen mixing, creating gentle O2 gradients to support their growth and to visualize motility. While Lactobacilli or E. coli can grow on many media, the particular advantage of semi-solid low-agar media is for motility assessment and cultivation of microaerophiles, not a general preference for those other groups. Step-by-Step Solution: Identify the unique property: controlled diffusion/gradients in soft agar. Relate this to microaerophile oxygen preferences. Recognize classic motility test media are semi-solid for visualization. Select “Microaerophillic bacteria.” Verification / Alternative check: Campylobacter and other microaerophiles demonstrate improved recovery in environments providing reduced O2; semi-solid systems can help establish such conditions. Why Other Options Are Wrong: Lactobacilli / E. coli: Grow on numerous media; semi-solid is not specifically “useful” for their cultivation compared with the microaerophile advantage. All of these: Overstates the utility and blurs the specific purpose. Common Pitfalls: Confusing “can grow” with “particularly useful.” The question asks where semi-solid media offer special value. Final Answer: Microaerophillic bacteria.

Question 16

Stringent (obligate) anaerobes can be successfully cultivated if the medium and atmosphere are treated by which special measures?

Accepted Answer

Introduction / Context: Obligate anaerobes are inhibited or killed by oxygen due to a lack of protective enzymes (for example, catalase, superoxide dismutase). Culturing them requires reducing the dissolved oxygen in media and maintaining an oxygen-free atmosphere during incubation and handling. Given Data / Assumptions: Anaerobes are sensitive to oxygen and reactive oxygen species. Media and headspace must be deoxygenated or chemically reduced. Handling (inoculation/incubation) should be done in an anaerobic environment. Concept / Approach: Boiling the medium drives off dissolved oxygen; adding reducing agents (for example, cysteine, thioglycollate) lowers redox potential; and sparging with oxygen-free nitrogen replaces residual oxygen. These steps, used singly or in combination, establish conditions compatible with the growth of stringent anaerobes. Step-by-Step Solution: Reduce dissolved O2 by heat (boiling) under appropriate safety. Add reducing agents such as cysteine to lower redox potential. Purge headspace with oxygen-free nitrogen to exclude O2. Conclude that any of these measures contributes to successful cultivation. Verification / Alternative check: Anaerobic jars, glove boxes, and pre-reduced, anaerobically sterilized (PRAS) media apply these principles routinely in clinical and research labs. Why Other Options Are Wrong: Each single option is correct, but incomplete on its own. The comprehensive answer “any of these” captures the acceptable approaches. Common Pitfalls: Neglecting rapid transfer and oxygen exposure during inoculation; even perfectly reduced media can fail if handling introduces O2. Final Answer: any of these.

Question 17

For selective isolation of the gonorrhea-causing Neisseria gonorrhoeae from clinical specimens, media are typically supplemented with which component to suppress competing flora?

Accepted Answer

Introduction / Context: Neisseria gonorrhoeae is fastidious and easily overgrown by commensals. Clinical microbiology relies on selective media to recover it from mixed specimens (for example, genital, rectal, pharyngeal swabs) while suppressing contaminants. Given Data / Assumptions: Selective media use inhibitors to reduce growth of competing bacteria and fungi. Gonococci remain sensitive to many antibiotics but can grow on specific selective formulations. Specimen quality and rapid plating affect recovery. Concept / Approach: Chocolate agar–based media such as Thayer–Martin or Modified Thayer–Martin include antibiotics (for example, vancomycin, colistin, nystatin, trimethoprim) that suppress Gram-positives, many Gram-negatives, and yeasts. This targeted suppression allows N. gonorrhoeae colonies to be isolated for identification and susceptibility testing. Step-by-Step Solution: Recognize the need for selective suppression of background flora. Recall that antibiotic cocktails in Thayer–Martin media serve this role. Match the correct additive: “certain antibiotics.” Select the option that specifies antibiotics as the selective component. Verification / Alternative check: Laboratory manuals list antibiotic-supplemented media as the standard of care for N. gonorrhoeae isolation. Why Other Options Are Wrong: Cellulose / Succinate: Not the selective agents used for recovery in clinical media. None of these: Incorrect because antibiotics are indeed used. Common Pitfalls: Plating delays and cold shock reduce viability; even with selective media, prompt inoculation and CO2-enriched incubation are important. Final Answer: certain antibiotics.

Question 18

In cell quantification, which choice below is NOT an indirect method for estimating cell mass or growth in a culture?

Accepted Answer

Introduction / Context: Microbiologists and bioprocess engineers estimate biomass using direct and indirect methods. Indirect approaches infer growth from properties such as turbidity, viscosity, oxygen uptake, calorimetry, or substrate consumption, whereas direct methods physically measure biomass mass or cell numbers. Given Data / Assumptions: Indirect methods correlate ancillary measurements with biomass (for example, optical density, heat production). Direct methods involve harvesting cells and weighing or counting them. Question asks for the option that is not indirect. Concept / Approach: “Cell dry weight” requires collecting cells (centrifugation/filtration), drying to constant mass, and weighing—this is a direct measure of biomass. In contrast, tracking nutrient composition (for example, substrate depletion), viscosity changes due to polymer or cell concentration, or heat evolution (calorimetry) are all indirect indicators of growth. Step-by-Step Solution: Separate direct from indirect measures conceptually. Map each option: dry weight = direct; others rely on correlations. Identify the outlier (not indirect): cell dry weight. Choose “Cell dry weight.” Verification / Alternative check: Standard fermentation texts list cell dry weight (g/L) as the primary direct biomass metric for process control and yield calculations. Why Other Options Are Wrong: Nutrient composition: Indirect via mass balance or substrate consumption. Viscosity: Indirect, correlates with biomass/exopolymers. Heat evolution: Indirect via metabolic heat measured calorimetrically. Common Pitfalls: Assuming any quantitative readout equals a direct measure; many convenient signals are proxies that require calibration. Final Answer: Cell dry weight is not an indirect method; it is a direct biomass measurement.

Question 19

Which of the following is a necessary sequence of steps for cultivating microorganisms in the laboratory, from medium preparation to inoculation and growth?

Accepted Answer

Introduction / Context: Successful cultivation requires a workflow that ensures nutrients are available, contaminants are excluded, and the target organism is introduced under appropriate conditions. Skipping any step risks failed growth or mixed cultures that compromise results. Given Data / Assumptions: A suitable culture medium must meet the organism’s nutritional needs. Sterilization removes pre-existing life from vessels and media. Inoculation introduces the desired organism aseptically. Incubation at proper temperature/atmosphere follows. Concept / Approach: The logical order is: choose and prepare the medium, sterilize (autoclave/filtration/dry heat as appropriate), then inoculate aseptically to establish a pure culture. This workflow underpins all downstream steps such as incubation, monitoring, and analysis. Step-by-Step Solution: Select a medium with proper carbon, nitrogen, minerals, and growth factors. Sterilize vessels and media to eliminate contaminants. Inoculate with the organism using sterile technique (loop, pipette, syringe). Incubate under correct temperature/atmosphere; observe growth. Verification / Alternative check: Aseptic technique checklists in lab manuals emphasize all three steps before incubation; omission of any leads to unreliable outcomes. Why Other Options Are Wrong: Each individual step is necessary but insufficient alone; only “All of the above” captures the complete set required for cultivation. Common Pitfalls: Neglecting sterilization of tools (loops, tips) after media sterilization, leading to inadvertent contamination. Final Answer: All of the above.

Cell Cultivations Questions