DNA Metabolism

Explain the molecular basis of bidirectional replication, how it was experimentally demonstrated, and its significance.

Molecular basis of bidirectional replication:

• Replication initiates at a specific origin
• Two replication forks form and move in opposite directions
• Both DNA strands are replicated simultaneously at each fork
• Produces a characteristic theta (θ) structure in circular chromosomes

Experimental demonstration:

• John Cairns used autoradiography with tritium-labeled thymidine
• Visualization showed replication loops in E. coli chromosome
• Pulse-labeling experiments (adding ³H briefly before stopping replication)
• Label appeared at both replication forks, proving bidirectional movement
• Denaturation mapping with bacteriophage λ confirmed single origin point

Significance:

• Maximizes replication efficiency by covering the chromosome in minimum time
• Two forks meet approximately halfway around a circular chromosome
• Allows complete replication to occur twice as fast as unidirectional replication
• Universal in most bacterial and eukaryotic replication systems

Why was isotope labeling with heavy nitrogen (¹⁵N) crucial for the Meselson-Stahl experiment, and what property allowed the separation of different DNA molecules?

Isotope labeling with ¹⁵N was crucial because:

• It created a detectable physical difference between original and newly synthesized DNA without altering chemical properties or function • ¹⁵N is 7% heavier than the common ¹⁴N isotope, making DNA density ~1% greater • This small but significant difference allowed separation of DNA molecules based solely on their nitrogen composition

Separation principle: • Cesium chloride (CsCl) density gradient centrifugation separates molecules based on buoyant density • When centrifuged at high speed (>100,000 × g), CsCl forms a stable density gradient • DNA molecules migrate to positions where their density matches the surrounding CsCl solution • This enables separation of molecules differing by as little as 0.1% in density

The technique's remarkable resolution allowed researchers to distinguish between heavy DNA (¹⁵N-¹⁵N), hybrid DNA (¹⁵N-¹⁴N), and light DNA (¹⁴N-¹⁴N), providing direct physical evidence for the semiconservative model.

What distinguishes DNA polymerase III from other E. coli DNA polymerases in terms of replication efficiency?

DNA polymerase III is distinguished by:

• Highest polymerization rate (250-1,000 nucleotides/second)
• Exceptional processivity (≥500,000 nucleotides added before dissociation)
• Most complex structure (≥10 different subunit types; Mr 791,500)
• Lacks 5'→3' exonuclease activity (unlike DNA polymerase I)

This makes DNA polymerase III the primary replication enzyme in E. coli, capable of synthesizing DNA at rates necessary to keep pace with replication fork movement in the bacterial cell.

Explain the complete architecture and function of the primosome complex in the replication fork, showing how it integrates with other components of the replisome.

Primosome architecture and function:

Core components:

• DnaB helicase: Hexameric ring-shaped enzyme that unwinds DNA
• DnaG primase: Synthesizes RNA primers (10-60 nucleotides)

Integration with replisome:

1. Physical connection:

   • DnaB helicase interacts with τ subunit of DNA polymerase III
   • Places primosome in optimal position relative to polymerase

2. Functional coordination:

   • DnaB moves 5'→3' on lagging strand template
   • Creates single-stranded DNA template
   • Periodically recruits primase to synthesize primers
   • Primase association with DnaB is transient and repeated

3. Activity cycle:

   • DnaB continuously unwinds DNA ahead of replication fork
   • Primase occasionally associates to create primers
   • Primers become starting points for Okazaki fragments
   • New β sliding clamps loaded at each primer

4. DNA structure organization:

   • Works with topoisomerases to relieve torsional stress
   • SSB proteins coat exposed single strands between primers

This integration creates a functional unit that coordinates the discontinuous synthesis of the lagging strand with continuous leading strand synthesis.

Explain the functional roles of each protein component at the replication fork during lagging strand synthesis.

Key protein functions during lagging strand synthesis:

• DnaB helicase: Unwinds DNA duplex ahead of the replication fork, traveling 5'→3' on the lagging strand template

• DNA topoisomerase II (DNA gyrase): Relieves topological stress created by DNA unwinding

• DNA primase: Synthesizes short RNA primers (10-60 nucleotides) that provide a 3'-OH group for DNA polymerase III

• DNA polymerase III: Extends RNA primers by adding deoxyribonucleotides to create Okazaki fragments

• Single-stranded DNA-binding protein (SSB): Stabilizes exposed single-stranded DNA, preventing secondary structure formation and reannealing

• DNA polymerase I: Later removes RNA primers and fills the gaps with DNA

• DNA ligase: Seals nicks between adjacent Okazaki fragments after primer removal

This coordinated protein assembly ensures orderly synthesis despite the need for discontinuous replication on the lagging strand.

What is the functional significance of the β sliding clamp in DNA replication, and how does the clamp recycling process work?

Function of β sliding clamp: • Forms a ring-shaped dimer that encircles the DNA double helix • Tethers DNA polymerase III to the template, dramatically increasing processivity • Prevents polymerase dissociation, enabling addition of thousands of nucleotides without detachment • Serves as a mobile platform that slides along DNA as replication proceeds

Clamp recycling process:

1. When an Okazaki fragment is completed, the polymerase core dissociates from its β clamp
2. The clamp-loading complex (γ complex) prepares and positions a new β clamp at a newly synthesized RNA primer
3. The core polymerase transfers to this new clamp to begin synthesis of the next Okazaki fragment
4. The old β clamp is left behind and eventually recycled

This dynamic loading and unloading of sliding clamps allows efficient discontinuous synthesis on the lagging strand while maintaining the integrity of the replisome complex.

How would you interpret different patterns of bacterial colony growth in the Ames test, and what information can be derived about the test compound's properties?

Interpretation of colony growth patterns in Ames test:

• Few scattered colonies (control plate): Represents spontaneous back-mutation rate • Concentration-dependent ring of colonies: Confirms mutagenic activity • Clear zone immediately around application point: Indicates toxicity at high concentrations • Gradual increase in colony density moving away from center: Shows dose-response relationship • No increase in colonies compared to control: Suggests non-mutagenic compound • Colony numbers proportional to mutagen concentration: Allows quantitative comparison between compounds

Additional interpretations: • Mutagenicity present only after liver enzyme treatment: Compound is a "pro-mutagen" requiring metabolic activation • Different colony patterns with different Salmonella strains: Reveals specificity of mutagenic mechanism • Size of mutagenic zone: Indicates relative potency and diffusion properties of the compound

Example: A strong mutagen produces a clear ring pattern with dense colonies at optimal concentrations and a toxic clear zone at the center, while a weak mutagen shows only slightly elevated colony counts compared to background.

How would defects in the Dam methylase system affect mutation rates in E. coli, and through what specific mechanism?

Defects in Dam methylase would increase mutation rates through disruption of strand discrimination:

• Without proper methylation patterns:

• Repair machinery cannot reliably identify the template strand
• Mismatches may be repaired using the incorrect strand as template
• Some mismatches go unrepaired when strand identity is ambiguous

Specific mechanisms:

1. Undermethylation: New strand may be repaired correctly, but many mismatches escape repair
2. Overmethylation: If both strands are methylated immediately, repair is inhibited
3. Random methylation: Repair may target either strand arbitrarily, causing mutations

Experimental evidence shows: • Dam-deficient strains show 10-100× higher mutation rates • These mutations arise primarily from inability to direct repair to the proper strand • Similar mutation patterns occur in bacteria lacking functional MutH protein

This system illustrates why post-replicative modifications are essential for genomic stability.

Compare the direct repair mechanism used for O6-methylguanine with excision repair mechanisms in terms of efficiency, molecular consequences, and energy cost.

Direct repair of O6-methylguanine vs. excision repair:

Efficiency:

• Direct repair: Immediate correction without creating intermediate structures
• Excision repair: Requires multiple steps (recognition, excision, synthesis, ligation)

Molecular consequences:

• Direct repair: DNA structure remains intact; only the methyl group is removed
• Excision repair: Creates temporary single-strand breaks and gaps in DNA

Energy cost:

• Direct repair: High protein cost (one methyltransferase consumed per lesion) but no ATP required
• Excision repair: Lower protein cost (enzymes can catalyze multiple repairs) but requires ATP for steps like DNA synthesis and ligation

Evolutionary significance:

• Direct repair evolved for specific high-priority lesions that have severe mutagenic potential
• Excision repair evolved as a versatile system capable of removing diverse types of damage

In both cases, the repair strategy reflects the biological importance of maintaining genomic integrity, but direct repair represents an extreme case where an entire protein is sacrificed to prevent a single mutation.

What are the key differences between integration and excision processes in bacteriophage λ site-specific recombination?

Integration and excision in bacteriophage λ differ in several critical ways:

1. Required sites:

   • Integration: Occurs between attP (phage) and attB (bacterial) sites
   • Excision: Occurs between attL and attR sites that flank the integrated prophage

2. Protein requirements:

   • Integration: Requires λ integrase (INT) and IHF (integration host factor)
   • Excision: Requires λ integrase (INT), IHF, plus additional proteins: • XIS (excisionase) - encoded by bacteriophage • FIS (factor for inversion stimulation) - encoded by bacterium

3. Directionality:

   • Each process has distinct regulatory controls preventing unwanted reactions
   • The additional proteins for excision ensure the reaction is not accidentally reversed

Note: The specificity of these processes allows bacteriophage λ to precisely control its lifecycle, alternating between lytic growth and lysogenic integration in the host chromosome.