Biology Unit 3 AOS1: Nucleic Acids and Proteins — Full Solutions

RNA polymerase II (or simply RNA polymerase). This enzyme catalyses the formation of the mRNA copy from the DNA template. It is specific to protein-coding genes in eukaryotes.

A codon is a sequence of 3 nucleotides on mRNA that specifies which amino acid should be added to the growing protein chain. An anticodon is a sequence of 3 nucleotides on tRNA that is complementary and antiparallel to the codon. They pair via base-pairing rules (A-U, G-C), ensuring the correct tRNA (carrying the correct amino acid) arrives at the ribosome.

Ribosomes are located in the cytoplasm (on rough endoplasmic reticulum or free in the cytoplasm), not in the nucleus. mRNA must exit the nucleus through nuclear pores so that it can encounter ribosomes and be translated into protein. Additionally, in eukaryotes, transcription (DNA → mRNA) occurs in the nucleus, but translation (mRNA → protein) occurs in the cytoplasm, so physical separation requires mRNA transport.

The mRNA codon is 5'-GUC-3' (antiparallel and complementary to 3'-CAG-5': C-G, A-U, G-C).

The tRNA carries glutamine (Gln). Wait—the hint says the codon for glutamine is CAG. Let me reconsider: if the anticodon is 3'-CAG-5', it pairs with mRNA codon 5'-GUC-3'. But this doesn't match the hint.

Correction: If the anticodon is 3'-CAG-5' and pairs with mRNA 5'-GUC-3', the amino acid is valine (Val, codon GUC). The hint suggesting glutamine would require the anticodon to be 3'-CTG-5' (pairing with 5'-GAC-3'... no, that's still not CAG). The student should state: anticodon 3'-CAG-5' pairs with codon 5'-GUC-3', which codes for valine.

Each amino acid is specified by a codon (3 nucleotides). So: 309 amino acids × 3 nucleotides/codon = 927 nucleotides in the ANP32A gene (exons only). In reality, the gene also contains introns (non-coding sequences that are spliced out) and regulatory regions (promoter, terminator), so the complete ANP32A gene is much longer—over 3 kb in the human genome—but the exon sequence that codes for the protein would be ~927 nucleotides.

Peptide bond. A peptide bond is a covalent bond between the carboxyl group (−COOH) of one amino acid and the amino group (−NH₂) of the next, releasing a water molecule (condensation reaction).

Alpha-helix (a right-handed spiral) and beta-pleated sheet (an extended zigzag pattern). Both are stabilised by hydrogen bonds between the backbone atoms of the polypeptide chain.

Heat disrupts the non-covalent interactions (hydrogen bonds, hydrophobic interactions, ionic bonds) that stabilise secondary and tertiary structure. These interactions are weak and temperature-sensitive. However, the peptide bonds linking amino acids in the primary structure are covalent and very strong; they resist breaking at moderate temperatures. So secondary and tertiary structures unfold, but the primary sequence stays intact. If the protein cools, it may refold correctly (or misfold if it gets stuck).

Disulphide bonds are covalent bonds (−S−S−), formed by an oxidation reaction between the R groups of two cysteines. Hydrogen bonds are non-covalent electrostatic interactions. Disulphide bonds are much stronger (~350 kJ/mol) than hydrogen bonds (~4−20 kJ/mol). Disulphide bonds are not easily disrupted by temperature, pH, or solvents, making them essential for stabilising proteins in harsh environments or providing permanent structural support. ANP32A uses disulphide bonds to "lock in" its 3D shape and maintain function under cellular stress.

Effect: The protein would likely misfold or become less stable.

Reasoning: In the native ANP32A protein, leucine (hydrophobic) is buried in the protein's hydrophobic core, away from water. A serine (hydrophilic, polar) R group cannot stay in the core—it would try to move to the protein surface to interact with water. This would disrupt the hydrophobic core's architecture, forcing the protein to refold. The loss of a hydrophobic residue from the core weakens hydrophobic interactions. Additionally, serine might form new hydrogen bonds or expose buried hydrophobic regions to water, destabilising the entire tertiary structure. The protein might misfold, aggregate, or be degraded by the cell, resulting in loss of ANP32A function and phenotypic consequences (e.g., impaired tumour suppression).

Silent mutation (also called synonymous mutation). This occurs when a nucleotide change in the DNA results in a codon that still codes for the same amino acid, usually because the change is in the wobble (third) position.

The genetic code is degenerate or redundant. Most amino acids are coded by more than one codon. This redundancy provides a buffer: many nucleotide mutations do not change the amino acid sequence because they result in synonymous codons (different codons for the same amino acid).

First codon pair (UUU vs UUC) is a silent mutation. Both code for phenylalanine, even though position 3 differs (U vs C). This is because of genetic code redundancy—the wobble position allows both codons to specify the same amino acid.

Second codon pair (GAA vs GUA) is a missense mutation. Position 1 differs (G vs G... wait, let me reread: GAA vs GUA). Actually, position 2 differs (A vs U). This change results in different amino acids: glutamic acid (GAA) vs valine (GUA). This is a missense mutation that changes the protein sequence.

All 12 mutations likely occurred in the wobble position (position 3 of codons), or they resulted in synonymous codons (different codons for the same amino acids). These would be silent mutations. Specifically, if the 12 mutations were distributed as 4 changes in position 3 of 4 different codons (accounting for wobble), or 6 codon pairs where the change is synonymous, then the 309-amino-acid protein would remain unchanged. This demonstrates the robustness of genetic code redundancy.

This is a silent mutation. The codon changes from UUA to CUU, but both code for leucine. The amino acid sequence is unchanged. This demonstrates that some nucleotide changes can be quite dramatic (changing multiple positions) but still result in the same amino acid, because leucine has multiple codons. However, a change from UUA (leucine) to UAA (stop codon) would be a nonsense mutation—a severe type of missense mutation.

The backbone, which consists of the central carbon (alpha carbon), amino group (−NH₂), carboxyl group (−COOH), and hydrogen atom (−H). These are identical in all amino acids.

Their R groups differ. Glycine has R = −H (just a hydrogen), while alanine has R = −CH₃ (a methyl group). Both have the same backbone, but the R group makes them different amino acids.

The R group determines the amino acid's polarity (polar vs nonpolar), charge (positive, negative, or neutral), and size. Nonpolar R groups cluster together in the protein core to avoid water. Polar R groups point to the surface. Charged R groups form ionic bonds, stabilising structure. By having 20 different amino acids with different R groups, proteins can be built with diverse properties—some proteins are enzymes, some are structural, some bind DNA. The sequence of different R groups determines the protein's 3D shape and function.

Although both leucine and lysine are large, their R groups have different properties: Leucine is nonpolar and hydrophobic. In a protein, it would cluster in the nonpolar core, away from water. Lysine is polar and positively charged. In a protein, it would be exposed on the surface where it can interact with water or form ionic bonds with negatively charged amino acids (e.g., aspartate). The nature of the R group determines the amino acid's location and function in the protein, not just its size.

The thiol group (−SH) in cysteine's R group can form a disulphide bond (−S−S−) with another cysteine's thiol group. This is a covalent bond that covalently links two different parts of the protein chain (or different protein molecules). Other amino acids cannot form disulphide bonds because their R groups don't have this reactive thiol functionality. Disulphide bonds are crucial for stabilising protein 3D structure, especially in extracellular proteins or proteins under stress.

A gene knockout refers to the complete inactivation or removal of a gene so that it no longer produces functional protein. The gene is disabled (via mutation, deletion, or other means) and has no biological activity in the cell.

CRISPR-Cas9 (the most common modern method). Other answers include: gene deletion, frameshift mutation, insertion of a stop codon, or homologous recombination. CRISPR-Cas9 uses molecular scissors (Cas9 enzyme) to cut the gene at a precise location; the cell's DNA repair machinery introduces mutations that inactivate the gene.

Because the three ANP32 genes are paralogs (similar genes with overlapping functions). Knocking out only one gene might not eliminate ANP32 protein production if the other genes can compensate (functional redundancy). By knocking out all three simultaneously, the researchers ensured that no functional ANP32 protein is produced, allowing them to definitively test whether ANP32 is essential for viral replication.

This implies that ANP32 proteins have essential roles in normal cellular processes, including:

• Regulation of gene transcription (development requires coordinated changes in gene expression)
• Control of cell cycle progression (necessary for growth and tissue formation)
• Possible roles in DNA repair or genome stability
• Reproduction/fecundity (ANP32 may regulate genes essential for gametogenesis or gamete function)

These are the protein's normal, non-viral functions. A knockout eliminates ANP32, disrupting these processes and harming the chicken.

Advantages of partial inhibition (knockdown) over complete knockout:

• Residual ANP32 function: The chicken retains some ANP32 protein, allowing partial normal development and reproduction (not completely harmed).
• Virus replication impaired but not blocked: Even partial ANP32 inhibition might reduce viral replication rate below the threshold needed for infection, while not eliminating all cellular ANP32 function.
• Tissue-specific targeting: A drug could be delivered to respiratory tissues (where influenza infects) to inhibit ANP32 locally, while leaving ANP32 active in other tissues (sparing development and reproduction).
• Dose titration: The drug dose could be adjusted to balance viral suppression with acceptable harm to the host.

This approach addresses the trade-off: gain some virus resistance without complete loss of normal ANP32 function.

Based on the avian influenza study pattern, we would predict that at very high doses of fungal spores, the edited plant would eventually become infected, but it would likely show slower disease progression and lower pathogen load compared to the wild-type plant. Complete resistance is unlikely at extreme exposure levels.

Reduced transmission means edited chickens shed less virus, so they spread infection more slowly to other birds. On a farm, this could slow the spread of an outbreak, giving farmers more time to isolate birds or implement other control measures, reducing total losses.

Viral load correlates with disease severity — more virus means more tissue damage, stronger symptoms, and longer recovery time. A lower viral load means less virus to damage cells, so the bird suffers less severe disease and may recover faster. Additionally, lower viral load reduces transmission to other birds (key to farm biosecurity).

It suggests genetic defences are not permanent solutions. Just as bacteria evolve antibiotic resistance over time, viruses can evolve mutations that bypass genetic edits. Single-gene edits may be vulnerable if the pathogen can mutate that one target. Long-term disease control might require multiple genetic modifications, rotating interventions, or combining genetic resistance with other strategies (vaccines, hygiene).

Expose edited chickens to the virus for many generations (successive infections in a population). If polymerase mutations arise that allow the virus to replicate despite the edit, you would see: increased infection rates over generations, increased viral loads, or evolution of virus strains that can infect ANP32A-edited cells. A control group of wild-type chickens should show consistent high infection rates throughout. Evidence of variant viruses with altered polymerase that can evade the edit would confirm the escape mutation risk.

DNA: A, T, G, C. mRNA: A, U, G, C. The difference is thymine (T) in DNA vs. uracil (U) in mRNA. Both pair with adenine, so the genetic code is preserved during transcription.

DNA is synthesised and stored in the nucleus (the genetic vault). mRNA is synthesised in the nucleus (by transcription of DNA) but must be transported to the cytoplasm where ribosomes are located (to translate it into protein). After translation, mRNA is degraded.

DNA persists for the lifetime of the cell (and is replicated to pass to daughter cells). mRNA typically lasts minutes to hours before being degraded by enzymes. This reflects their roles: DNA is permanent storage requiring protection; mRNA is a temporary working copy that allows the cell to regulate protein production. When an mRNA is degraded, the cell can stop making that protein, providing flexibility.

Less stability is actually advantageous for mRNA because it allows dynamic gene regulation. If mRNA were as stable as DNA, it would persist indefinitely and the cell could not respond to changing conditions — it couldn't quickly stop making a protein it no longer needs. The temporary nature of mRNA (minutes to hours lifespan) allows cells to fine-tune protein production. DNA, being permanent, is worth protecting; mRNA, being temporary, benefits from rapid degradation and recycling.

If mRNA used thymine, the cell would lose the ability to distinguish mRNA from DNA by base composition alone. This could cause multiple problems: (1) Enzymes that degrade mRNA (RNases) might also degrade DNA, destroying the genetic code; (2) DNA repair mechanisms designed to protect DNA (e.g. mismatch repair) might waste energy 'fixing' mRNA; (3) The cell would lose the ability to specifically regulate mRNA levels — mRNA could not be targeted for degradation without risking DNA damage. The use of uracil instead of thymine is thus a regulatory mechanism as much as a structural feature.

The two components are: (1) Cas9 protein — an enzyme that cuts DNA, and (2) Guide RNA (gRNA) — an RNA molecule that directs Cas9 to the target location. Together, they form a ribonucleoprotein complex.

The guide RNA must be complementary to the target ANP32A sequence because it uses Watson-Crick base pairing (A-U, G-C) to recognise and bind to the target DNA. This complementarity ensures Cas9 cuts at the correct location and not at other similar sequences in the genome.

Guide RNA (rather than DNA) is used because RNA is temporary and degradable. After the gRNA delivers Cas9 to the target site and the cut is made, cellular RNases break down the gRNA. This prevents Cas9 from continuing to cut at the same site repeatedly, which would cause more DNA damage than intended. DNA would persist and could lead to repeated or off-target cutting.

The guide RNA might direct Cas9 to cut at both locations — the intended ANP32A site and the unintended (off-target) site. This would cause 'off-target effects': unintended edits at the wrong location, potentially disrupting other genes and causing harmful mutations. This is why guide RNA design is carefully checked for specificity before use — scientists use bioinformatic tools to ensure the gRNA matches only the target, not similar sequences elsewhere.

The sgRNA simplifies the system by requiring synthesis and delivery of only one RNA molecule instead of two. This is more practical for scientists (easier to make, transport, and deliver into cells) and may reduce synthesis errors or incomplete complexes. Additionally, a single molecule may be more stable or more efficiently incorporated into the Cas9 ribonucleoprotein complex. The single-molecule design has become the standard because of these practical advantages.

The main components are: (1) Guide RNA (gRNA) — directs Cas9 to the target DNA sequence via complementary base pairing; (2) Cas9 protein — an endonuclease enzyme that cuts both strands of the DNA double helix when directed by the gRNA; (3) HDR template DNA — provides the corrected codon sequence that is inserted during DNA repair.

Each guide RNA is about 20 nucleotides long and must be complementary to the target DNA sequence. Different codons require different guide RNAs to ensure specificity — using the same gRNA for two different target sites would result in cuts at both sites (or off-target cuts). Each gRNA targets only one specific location via Watson-Crick base pairing.

NHEJ is the cell's natural quick repair pathway, but it re-glues the broken DNA ends somewhat randomly, often introducing small insertions or deletions. This would likely damage the ANP32A gene rather than replace the specific codons. HDR uses a provided DNA template containing the corrected codons (flanked by homology arms), so the cell accurately replaces the old sequence with the new one. For precise amino acid changes, HDR is essential.

Somatic cells are body cells that die and are replaced; an edit in a somatic cell affects only that cell and its daughters but is not passed to offspring. Germ cells (sperm/egg precursors) and early embryos are different: they undergo mitosis to create all cells of the body, including future germ cells. An edit at this stage is copied to every cell, including germ cells, so when the chicken reproduces, it passes the edited ANP32A to all offspring. Thus, only germ-line or early embryo edits are heritable.

Molecular steps: (1) Editing changes the DNA codons in the ANP32A gene (e.g., GAA → GAT); (2) During transcription, mRNA is synthesised with the new codon sequence (GAU in mRNA); (3) During translation, a tRNA with an anticodon complementary to the new codon (AUA) delivers a different amino acid; (4) The resulting protein has the new amino acids, altering its 3D structure. Functional consequence: The virus evolved to recognise and bind the original ANP32A protein structure as a 'key' to unlock cells. The 2 changed amino acids alter the protein's folding and shape, making it unrecognisable to the virus — the lock has changed and the virus's key no longer fits.

A promoter is a DNA regulatory region that is located upstream (before) the coding sequence of a gene. It contains consensus sequences (like the TATA box in eukaryotes) where RNA polymerase binds. The promoter signals where transcription should begin, at the transcription start site (+1).

If the TATA box were deleted, RNA polymerase II would not recognise the promoter and could not bind. Transcription of the ANP32A gene would not be initiated, so no mRNA would be made from the ANP32A gene. Without mRNA, ribosomes could not translate the ANP32A protein, resulting in little or no ANP32A protein being produced.

The promoter is essential for transcription initiation. If it is damaged, RNA polymerase cannot bind and transcription cannot begin. Without transcription, no mRNA is made. Without mRNA, no translation occurs and no protein is synthesised. Following the central dogma (DNA → mRNA → Protein), breaking the first step (transcription) prevents all subsequent steps. The edited coding sequence is irrelevant if the gene cannot be expressed — the cell produces no protein and gains no viral resistance benefit.

Promoters are critical regulatory regions; any damage to them severely disrupts or silences gene expression. An off-target cut in a promoter would prevent RNA polymerase binding and stop transcription of that entire gene, potentially disabling a functionally important protein. In contrast, introns are non-coding sequences that are removed during mRNA splicing and do not code for protein. An off-target cut in an intron might be repaired without consequence or might cause mild splicing errors. Promoters are thus more vulnerable and their disruption more consequential than intron damage.

The second guide RNA (5 bp downstream of TATA box) poses a higher risk. The promoter and TATA box region extend for several hundred base pairs, and regulatory sequences like the CAAT box can be located 75+ bp upstream of the transcription start site. A cut only 5 bp away from the TATA box is dangerously close and could easily damage important regulatory elements if the cut or subsequent repair extends beyond the target. The first guide RNA (100 bp downstream) is further from the promoter core and poses less risk of collateral promoter damage. Scientists use bioinformatic tools to map promoter boundaries and ensure guide RNAs target well beyond regulatory regions to minimise off-target effects.

Beneficence is the ethical principle of actively promoting the welfare and well-being of others by choosing actions that benefit them and reduce harm.

Developing a vaccine to prevent a serious infectious disease is a beneficent action because it reduces suffering and mortality in the population.

Antibiotics treat bacterial infections by killing the pathogen or inhibiting its growth. This aligns with beneficence because it relieves suffering, prevents complications, and promotes the patient's recovery and health—thereby advancing their welfare.

Example: Developing disease-resistant crops that can grow in drought-prone regions. This aligns with beneficence because it increases food security, reduces crop failure and farmer hardship, and improves nutritional outcomes in vulnerable populations—all of which promote human welfare and well-being.

Non-maleficence means 'do no harm' (avoiding negative outcomes). Beneficence means 'actively promote welfare' (pursuing positive outcomes). An example: prescribing pain relief medication after surgery is beneficent because it actively improves comfort and recovery, not just because it avoids harm.

Non-maleficence is the ethical principle of avoiding harm, injury, or suffering to others. It requires that interventions do not cause damage or negative consequences.

A surgeon performing an operation without obtaining informed consent from the patient violates non-maleficence because it causes physical harm (the surgery) without the patient's agreement. Similarly, prescribing a medication with severe side effects without exploring safer alternatives violates non-maleficence.

Widespread use of antibiotics in livestock promotes the development of antibiotic-resistant bacteria. These resistant strains can transmit to human populations, reducing treatment options and causing harm through untreatable infections. This harm is caused by the intervention itself, violating non-maleficence.

An intervention violates non-maleficence if it knowingly or negligently causes harm. Risk-taking involves the possibility of harm but not certainty. Some risks are justified if the potential benefits significantly outweigh the potential harms (e.g., surgery with risks but lifesaving outcomes). However, risks should be minimized, disclosed, and justified—not ignored.

The incomplete resistance at high doses means the virus still replicates in highly exposed birds. This creates a situation where the virus is under selective pressure (from partial resistance) but still able to replicate, accelerating the evolution of escape mutants. This violates non-maleficence because the intervention creates conditions that favour dangerous viral adaptation—a harm triggered by the edit itself.

Benefit (beneficence): Drought-resistant crops reduce food insecurity and improve agricultural yields in vulnerable regions, promoting human welfare. Risk (non-maleficence): Edited crops could cross-pollinate with wild relatives, creating invasive plant species or reducing biodiversity, causing ecological harm.

Non-maleficence (avoiding harm) is often prioritized over beneficence (promoting good) because preventing known harms is a foundational duty. It is generally more important to avoid causing suffering than to pursue additional benefits, especially when the harm risk is significant and well-understood.

Genetic screening aligns strongly with both principles. Beneficence: screening identifies and prevents genetic diseases, improving child health and welfare. Non-maleficence: screening avoids the harm of allowing disease to develop. In this case, both principles support the intervention, making it ethically stronger than cases where they conflict. The alignment of both principles increases ethical justification.

The incomplete protection weakens both beneficence and non-maleficence arguments: (1) Beneficence is undermined because the protective benefit is unreliable—it fails precisely when most needed (high viral exposure). (2) Non-maleficence is compromised because failing protection under pressure creates ideal conditions for viral selection and mutation. The combination of weak benefit and increased mutation risk strengthens the conclusion that risks outweigh benefits.

Proposal: Use CRISPR to edit chickens for avian flu resistance, but combine the edit with strict biosecurity protocols that prevent exposure to wild virus populations. Justification: Beneficence is maintained (protection against avian flu). Non-maleficence is strengthened (preventing virus exposure eliminates selective pressure for escape mutants, reducing pandemic risk). This hybrid approach aligns with both principles by gaining the benefit while managing the identified harm risk, making it ethically superior to the uncontrolled edit.

Consequence-based ethics judges whether an action is right or wrong based solely on its outcomes and consequences—good outcomes make it ethically justified, bad outcomes make it wrong, regardless of intentions.

If the consequence is that the patient survives and recovers due to the life-saving surgery, the action is justified from a consequence-based view, because the outcome (survival) is positive. Intentions or rule-breaking are irrelevant; only the outcome matters.

From a consequence-based perspective, developing antibiotics is justified because the consequences are positive: antibiotics kill bacteria, reduce infection, prevent death and suffering, and improve human health. Since the outcomes are beneficial, the action is ethically right.

Consequence-based ethics would weigh outcomes: benefit (disease elimination) vs. risk (5% chance of harmful mutation). If the disease is severe and widespread, the benefit may outweigh the 5% risk. But if the disease is rare or mild, a 5% harm risk is unacceptable. The decision depends on the actual consequences, not good intentions. The framework requires careful calculation of likely outcomes.

From a consequence-based view, incomplete protection is a negative consequence in two ways: (1) it fails to achieve the intended outcome (bird protection under realistic high-dose conditions), and (2) it creates selective pressure that favours viral mutation and escape. Both consequences are harmful. The failed protection means the edit cannot justify itself even on its own terms, strengthening the consequence-based argument against performing it.

Social factors are aspects of society that influence research development, including public perception, ethical attitudes, cultural values, regulatory environment, and social acceptance of technologies. They reflect how society views and responds to scientific innovation.

A large global market demand for the vaccine (e.g., for a common infectious disease) and the potential for significant profit through vaccine sales would accelerate research. Companies are more likely to invest if they expect a profitable return.

Public concern (social factor) led to strict EU regulations and consumer resistance to GM products. This created regulatory barriers and reduced market demand, deterring investment in GM crop research. Despite scientific support, social resistance slowed European GM agriculture development.

Social impact: Public awareness of pandemic risk would likely increase ethical concern and regulatory scrutiny, creating barriers to research approval and consumer acceptance. Economic impact: If regulatory approval becomes difficult and consumers avoid gene-edited chicken, profitability decreases, reducing industry investment. Conversely, if industry believes they can manage the risk safely, they may invest more to develop safeguards, accelerating research. The outcome depends on how convincingly industry can address public concerns.

Aligned factors (acceleration): High public acceptance of gene editing + profitable disease prevention creates strong research momentum. Consumers embrace gene-edited chicken; industry invests heavily; governments approve research readily. Research accelerates rapidly. Conflicting factors (slowdown): High public ethical concern about gene-edited food + regulatory barriers reduce investment despite high profit potential. Industry wants to develop the technology (economic incentive) but faces public resistance and regulatory delays (social barriers). Research slows despite economic incentive.

Restriction enzymes are endonucleases that cut double-stranded DNA at specific recognition sequences (e.g., EcoRI cuts at GAATTC). When the same restriction enzyme is used to cut both the foreign gene and the plasmid vector, it creates compatible sticky ends on both DNA molecules. These sticky ends are short, single-stranded overhangs with complementary sequences that can base-pair. This allows the gene and plasmid to align and be joined together by DNA ligase, forming the recombinant plasmid. Without restriction enzymes, there would be no way to precisely insert the gene into the plasmid.

DNA ligase is an enzyme that catalyses the formation of covalent phosphodiester bonds between the sugar-phosphate backbones of adjacent DNA strands. In the context of making a recombinant plasmid, after the foreign gene and plasmid are brought together by complementary sticky ends, DNA ligase 'glues' them together by joining the sugar-phosphate backbone. This creates one continuous DNA molecule (the recombinant plasmid) and seals the nick between the two DNA fragments.

Possible reasons: (1) The plasmid may lack an appropriate promoter sequence upstream of the human growth hormone gene. Bacterial RNA polymerase needs a bacterial-type promoter (e.g., a -10 and -35 box) to recognise where to start transcription. A human promoter would not work in bacteria. (2) The human growth hormone gene may contain introns that bacterial cells cannot splice out. If the gene still contains introns, it would be transcribed into an mRNA containing non-coding sequences, which would disrupt translation and produce a non-functional (or absent) protein. Alternatively, the human protein may be toxic to E. coli, or the ribosome binding site (Shine-Dalgarno sequence) may be missing or incorrect.

Why selection is essential: During transformation, only a small fraction of bacterial cells take up the plasmid (typically <1%). Additionally, some cells may take up the non-recombinant (original, non-cut) plasmid rather than the recombinant plasmid. Without selection, the culture would be a mixture of transformed and non-transformed bacteria, and of cells carrying different plasmids. There would be no way to distinguish which cells carry the desired recombinant plasmid. If selection were omitted: The researcher would culture a mixed population of bacteria. Most would not be producing the desired protein (they lack the plasmid entirely). Some might carry non-recombinant plasmid (wrong insert). Very few would actually have the recombinant plasmid and produce the target protein. The culture would be diluted with useless cells, making it inefficient and impossible to purify the desired protein to useful quantities.

Differences in process: A viral vector uses a modified virus (rather than a plasmid) to deliver the ANP32A gene into cells. The virus genome is engineered to remove genes responsible for viral replication and pathogenicity, but retain genes for packaging and cell entry. The ANP32A gene is inserted into the viral genome in place of the removed genes. The recombinant virus is then used to infect target cells, and the ANP32A gene is expressed inside them. Advantages of viral vectors: (1) Higher transduction efficiency — viruses have evolved powerful mechanisms to enter cells, so a much higher percentage of cells are successfully infected compared to transformation (which is low efficiency). (2) Can infect diverse cell types — depending on the viral vector, it can target specific tissues or cell types more effectively than plasmids. (3) Larger insert capacity — some viral vectors can carry larger foreign genes than small plasmids. Disadvantages: (1) Safety concerns — careful engineering is needed to prevent the virus from becoming pathogenic. (2) Immune response — cells may recognise the viral vector as foreign and trigger an immune response. (3) More complex to produce. For bacterial protein production (ANP32A from a chicken gene in bacteria), plasmids are typically sufficient and preferred due to simplicity and cost.

A transgenic organism is an organism that contains functional, heritable DNA from a different species. The foreign DNA was introduced through genetic engineering and is stable in the organism's genome (or, in this case, plasmid), passing to daughter cells during reproduction or cell division.

'Genetically modified' is a broad term referring to any organism whose DNA has been artificially altered by humans, including any insertion, deletion, or substitution. 'Transgenic' is a more specific term referring to organisms that contain heritable DNA from a different species. All transgenic organisms are genetically modified, but not all genetically modified organisms are transgenic — for example, a bacterium with a CRISPR-edited native gene is modified but not transgenic.

The mouse is transgenic because it contains functional, heritable DNA from a different species (humans). Specifically, it carries a human cancer-resistance gene that was artificially inserted into its genome and is stably inherited by its offspring. Since the foreign DNA comes from a different species, the definition of transgenic is met. The mouse is also genetically modified, but 'transgenic' is the more precise term.

This plant should be described as genetically modified only, not transgenic. Although the DNA has been artificially altered, all the edited DNA comes from the same plant species (no foreign DNA from another species is present). Transgenic specifically refers to organisms containing DNA from a different species. This is an important distinction because 'transgenic' organisms contain foreign genetic material, while this CRISPR-edited plant contains only rearranged native genes. (Note: This plant would be classified as 'CRISPR-edited' or 'gene-edited' within genetically modified, but not transgenic.)

This organism is best described as transgenic (and also genetically modified). It is transgenic because it contains functional, heritable DNA from two different species: the chicken ANP32A gene and the human insulin gene. Even though it also has CRISPR-edited native DNA, the presence of foreign DNA from two different species makes it transgenic. The term 'transgenic' applies — in fact, this is sometimes called 'multiply transgenic' because it contains DNA from multiple foreign species. The organism is also genetically modified (all genetic engineering makes an organism GM), but transgenic is more precise because it captures the key feature: the presence of foreign DNA from other species.

A DNA ladder (also called a size marker or molecular weight marker) is a commercial mixture of double-stranded DNA fragments of known sizes. It is typically composed of 5-15 fragments ranging from about 100 bp (base pairs) to 10,000+ bp, spaced at regular intervals (e.g., 100 bp, 250 bp, 500 bp, 1000 bp, 2000 bp, etc.). The exact composition varies by commercial ladder product.

Smaller DNA fragments move through the gel matrix more easily because they have less mass and encounter less resistance from the gel pores. Larger fragments are bulkier and move more slowly through the gel. Under the same electric field and gel conditions, smaller fragments reach farther distances, while larger fragments remain closer to the wells. This separation by size is the basis of gel electrophoresis.

No, you cannot safely conclude the sample is exactly 750 bp, even though 750 is mathematically halfway between 500 and 1000. Migration through a gel is not perfectly linear with size — the relationship between size and migration distance is logarithmic. A band exactly halfway in distance between 500 and 1000 bp ladder bands might be 750 bp, but it could also be 600 bp, 800 bp, or another value nearby. To estimate the size more accurately, you would need a ladder with more closely spaced bands (e.g., 500 bp, 600 bp, 700 bp, 800 bp, 900 bp, 1000 bp) to narrow down the estimate.

Method: (1) Load the ladder in one well. (2) Load uncut (unrestricted) plasmid in a second well — this should show a single band at the full plasmid size (e.g., 3000 bp). (3) Load plasmid that was cut with the restriction enzyme in a third well. (4) Run the gel and compare. If cutting was successful, the restricted plasmid lane should show two bands, each at approximately half the original size (e.g., two bands around 1500 bp each, using the ladder as reference). The sum of the two fragment sizes should approximately equal the original plasmid size. If only one band is visible in the restricted lane at the original size, the enzyme did not cut; if multiple bands appear, the enzyme may have cut at more than one site or the sample may be contaminated.

No, the ladder would not be reliable in this scenario. The reliability of a ladder depends on both the ladder and samples experiencing identical electrophoresis conditions — same voltage, same gel matrix, same buffer, same temperature. If different lanes experience different voltages, they will migrate at different rates, and the ladder's migration pattern will not match the samples' pattern. A sample that appears to align with a 500 bp ladder band might actually be 600 bp or 400 bp because the current difference caused different migration rates. To fix this, the gel would need to be re-run with proper equipment to ensure uniform current flow across all lanes.

You would expect to see 1 band located near the top of the gel (high position). Unrestricted plasmids remain circular and supercoiled in their native form. Supercoiled circular DNA is compact and migrates very quickly through the gel, reaching a position near the wells despite its large molecular weight. This makes unrestricted plasmids appear to be much smaller than they actually are.

Unrestricted plasmids are circular and supercoiled (tightly wound), making them compact and able to navigate through gel pores easily, so they migrate quickly (appear high on the gel). Restricted plasmids are linearised (no longer circular) and therefore extended in shape, encountering more resistance as they pass through the gel matrix. They migrate more slowly and reach a position lower on the gel based on their actual size. The same DNA molecule in different physical forms (circular vs. linear) has different migration rates because the gel separates based on both size and shape.

Lane B contains the recombinant plasmid. When a plasmid carries an insert (recombinant), restriction enzymes cut at the sites flanking the insert, releasing two separate linear pieces: the plasmid backbone and the insert. These appear as 2 bands on the gel. Lanes A and C each show 1 band, indicating non-recombinant plasmids (the plasmid alone, with no insert to be released). A single restriction cut linearises a non-recombinant plasmid into one piece.

The plasmid backbone is approximately 2000 bp and the insert is approximately 800 bp. When these two fragments are ligated together in the original recombinant plasmid, the total size would be 2000 + 800 = 2800 bp. This is the size of the uncut recombinant plasmid. (Note: the actual size may be slightly different from the estimate due to the approximate nature of gel size estimates, but 2800 bp is the expected calculated value.)

Both unrestricted plasmids (with and without insert) remain circular and supercoiled, so both migrate abnormally fast and appear as a single band near the top. The insert does not change the migration pattern significantly when the plasmid is still circular — the plasmid remains supercoiled and compact regardless. To distinguish them, you would need to: (1) Restrict (cut) both samples with the appropriate restriction enzyme. The recombinant would show 2 bands (backbone + insert), while the non-recombinant would show 1 band. Alternatively, (2) use a different detection method such as PCR to amplify and confirm the presence of the insert, or (3) use restriction mapping with multiple enzymes to establish different cutting patterns. Gel electrophoresis alone cannot distinguish unrestricted recombinant from unrestricted non-recombinant; they require restriction digestion for differentiation.

The unknown fragment is 500 base pairs (bp) or 0.5 kilobases (kb). Since the band aligns exactly with the 500 bp ladder mark, the fragment is 500 bp. This is the simplest type of size estimation — direct alignment with a known ladder band.

750 bp = 750 ÷ 1000 = 0.75 kb (or 0.75 kilobases). Since 1 kilobase = 1000 base pairs, divide the number of base pairs by 1000 to convert to kilobases.

The total size = insert size + plasmid backbone size = 750 bp + 2200 bp = 2950 bp or approximately 3.0 kb. This is the size of the original recombinant plasmid before it was cut with restriction enzymes. Note: The actual value may vary slightly from 3000 bp due to the positions of restriction sites and accuracy of gel estimates.

Plasmid 1 insert = 750 bp; Plasmid 2 insert = 1500 bp. The insert in Plasmid 2 is twice as large as the insert in Plasmid 1 (1500 ÷ 750 = 2). This could indicate that Plasmid 2 carries a larger version of the same gene (e.g., due to inclusion of introns, regulatory regions, or a longer gene), or a different gene altogether. The identity of the plasmid backbone (2200 bp in both) is confirmed by the identical 2200 bp band. If the experiment was comparing two versions of ANP32A, Plasmid 2 contains a longer ANP32A sequence.

Percentage error = |gel estimate - true value| ÷ true value × 100% = |750 - 780| ÷ 780 × 100% = 30 ÷ 780 × 100% = 3.8% (approximately 4%). The gel estimate of 750 bp is within 4% of the true 780 bp value. This small error is typical and acceptable for gel-based size estimates, which have inherent limitations due to interpolation between ladder bands. Gel electrophoresis is a useful quick screening method, but sequencing provides the exact size.

The plasmid contains 3 EcoRI restriction sites. For circular DNA, the number of restriction sites equals the number of fragments produced after complete digestion. Three fragments = three EcoRI sites.

For circular DNA with N restriction sites, N fragments are produced. For linear DNA with N restriction sites, N+1 fragments are produced. The difference is that linear DNA has 1 more fragment. This is because cutting a linear DNA at N sites divides it into N+1 segments, whereas cutting circular DNA at N sites divides it into N segments (each cut releases a piece). Example: 2 sites in circular = 2 fragments; 2 sites in linear = 3 fragments.

The recombinant plasmid contains 2 EcoRI restriction sites and 3 BamHI restriction sites. EcoRI produces 2 fragments, so there are 2 EcoRI sites (for circular DNA). BamHI produces 3 fragments, so there are 3 BamHI sites. The two enzymes recognise different sequences and have independent cutting patterns.

The non-recombinant vector has 1 EcoRI site (1 fragment when cut). When the ANP32A insert is ligated at this EcoRI site, the recombinant plasmid now has 2 EcoRI sites: one on each side of the insert (flanking it). Cutting with EcoRI releases 2 fragments: the insert and the vector backbone. This indicates that the insert was integrated at the single EcoRI site in the vector, and both ends of the insert have EcoRI-compatible sticky ends, creating two flanking sites. The insert likely also has an internal EcoRI site, or was prepared with EcoRI-sticky ends such that ligation created the second site.

Possible explanations: (1) The ANP32A insert contains an internal EcoRI site. When the insert is cut, it breaks into 2 pieces, so you see the plasmid backbone band + 2 insert fragments = 3 bands total. (2) The recombinant plasmid has more than 2 EcoRI sites — perhaps contaminating DNA or multiple inserts were incorporated. (3) Incomplete digestion — some EcoRI sites were not cut, producing unexpected fragments. (4) Contamination with another plasmid carrying different EcoRI sites. To clarify, compare the sizes of all 3 bands to the ladder. If two of the bands together equal the expected insert size, then the insert was cut internally. Use the sum of all band sizes to estimate the total plasmid size and verify against the expected recombinant plasmid size.

DNA migrates toward the positive electrode (anode). This is because DNA is negatively charged due to the phosphate groups in its sugar-phosphate backbone. In an electric field, negatively charged molecules move toward the positive pole and away from the negative pole (cathode).

Smaller DNA fragments travel farther because the gel matrix (agarose or polyacrylamide) is composed of pores, and smaller fragments navigate through these pores more easily with less resistance. Larger fragments are bulkier and encounter more resistance from the gel, so they move more slowly and don't travel as far in the same time period.

The gel matrix pore size determines the degree of sieving (size-based separation). A gel with smaller pores creates more resistance and provides better separation of DNA fragments because the size-dependent differences in migration rate are magnified — small fragments move much faster than large ones. A gel with larger pores creates less resistance and provides less separation because fragments of different sizes migrate more similarly. Gel concentration controls pore size: higher concentration → smaller pores → better separation of small-to-medium fragments. Lower concentration → larger pores → better for separating large fragments. The agarose concentration is typically chosen based on the expected size range of fragments being analysed (e.g., 0.8% for large fragments, 2% for small fragments).

All DNA molecules have approximately the same charge-to-mass ratio — the charge per unit mass is roughly constant regardless of DNA sequence or source. This is because the negative charge comes entirely from the phosphate groups in the backbone, and the number of phosphates is proportional to the DNA length. Therefore, charge alone does not differentiate DNA fragments; instead, size determines migration rate through the gel matrix due to differential resistance from pores. In contrast, proteins have variable charge-to-mass ratios (due to different amino acid compositions), so proteins can be separated by charge via isoelectric focusing. DNA's uniform charge-to-mass ratio makes gel electrophoresis an ideal size-separation technique for DNA, while other electrophoresis variants are needed for other molecules.

If all fragments travelled approximately equal distances, the gel pores are too large — the gel concentration is too low. When pores are large, even very different-sized fragments can migrate through with similar ease, eliminating the size-based separation. Correction: Increase the agarose concentration (e.g., from 0.5% to 1.0% or higher) to reduce pore size and increase resistance. This will increase the separation between fragments of different sizes. Additionally, run the gel longer (apply voltage for a longer time) to allow more time for separation to occur. Monitor the progress during the run to ensure bands are beginning to separate before they run off the gel.