packbacks #7 to #9 Flashcards

The catabolic oxidation of methanol to carbon dioxide can be represented by the following overall reaction:

CH3OH + 1.5O2 → CO2 + 2H2O

In this process, methanol (CH₃OH) acts as the electron donor, and oxygen (O₂) serves as the terminal electron acceptor.

Using the given standard redox potentials (E₀′):

• CO₂/methanol redox couple: −0.38 V
• ½ O₂/H₂O redox couple: +0.82 V

The overall cell potential (ΔE₀′) is determined by subtracting the donor potential from the acceptor potential:

ΔE0′ = E0′ (acceptor) − E0′ (donor) = (+0.82) − (−0.38) = + 1.20V

This large positive potential indicates a highly exergonic (energy-releasing) redox reaction. The released free energy drives electron transport, which pumps protons (H⁺) across the membrane, forming a proton motive force (PMF). The PMF then powers ATP synthase, producing ATP via oxidative phosphorylation.

Simultaneously, some of the electrons from methanol oxidation can be used to reduce NAD⁺ to NADH, providing reducing power for biosynthetic reactions.

In summary:

• Electron donor: methanol (CH₃OH → CO₂)
• Electron acceptor: oxygen (O₂ → H₂O)
• ΔE₀′ = +1.20 V, supporting efficient production of ATP and NADH through aerobic respiration.

The expression of the lac operon in E. coli is regulated by the availability of glucose and lactose in the environment, allowing the cell to efficiently manage energy resources. The operon contains genes responsible for the breakdown of lactose, but these genes are expressed only when lactose is available and glucose is scarce. When glucose levels are high, the cell prioritizes its use as an energy source, and cAMP levels remain low, preventing the cAMP–CAP complex from binding to the promoter. This reduces or prevents transcription, even if lactose is present. On the other hand, when lactose is absent, a repressor protein binds to the operator region of the operon, blocking RNA polymerase and preventing gene expression. However, when lactose is present, it is converted into allolactose, which binds to the repressor and inactivates it, allowing RNA polymerase to access the promoter. At the same time, if glucose levels are low, cAMP concentrations rise, enabling the cAMP–CAP complex to bind to the promoter and enhance transcription. This dual control ensures that the lac operon is fully activated only when lactose is available and glucose is absent, a phenomenon known as catabolite repression. Overall, the lac operon demonstrates both negative control (by the repressor) and positive control (by the CAP–cAMP complex), illustrating a precise and efficient system of gene regulation in prokaryotes.

Bacteria regulate gene expression through operons that can be either inducible, repressible, or positively controlled, depending on nutrient availability. The lac, arg, and mal operons illustrate how activators and corepressors fine-tune transcriptional control in response to the cell’s metabolic needs.

The lac operon is an inducible operon responsible for lactose metabolism. Its normal state is off, since a repressor protein binds to the operator, blocking transcription. When lactose is present, it is converted into allolactose, which serves as an inducer by binding to and inactivating the repressor, allowing RNA polymerase to transcribe the genes needed for lactose utilization. The lac operon also involves positive control: when glucose levels are low, cAMP levels rise, allowing the CAP–cAMP complex to bind to the promoter and enhance transcription. Together, these mechanisms ensure that lactose is metabolized only when lactose is available and glucose is scarce.

The arg operon is a repressible operon used for arginine biosynthesis. It is normally on, producing enzymes that synthesize arginine. When intracellular arginine levels become high, arginine acts as a corepressor by binding to the ArgR repressor protein, activating it. The ArgR–arginine complex then binds to the operator region, blocking transcription. This prevents overproduction of arginine and conserves cellular resources.

The mal operon controls maltose metabolism and is regulated by positive control. When maltose is present, it binds to and activates the MalT activator protein, which then promotes RNA polymerase binding to the promoter, initiating transcription of the maltose-utilizing genes. In the absence of maltose, MalT remains inactive, and transcription does not occur.

Overall, these three operons demonstrate distinct modes of regulation: the lac operon is an inducible system that uses both repressors and activators, the arg operon is a repressible system regulated by a corepressor, and the mal operon is a positively controlled inducible system that depends on an activator. Together, they illustrate how bacteria efficiently balance gene expression, conserving energy by producing enzymes only when they are needed.

In bacterial gene regulation, several key components work together to control when and how genes are expressed. These include activators, repressors, inducers, corepressors, effectors, operators, operons, and regulons.

An activator is a regulatory protein that promotes transcription by helping RNA polymerase bind to the promoter region of DNA. Activators exert positive control, increasing transcription rates when bound to their target DNA sites. For example, in the lac operon, the CAP–cAMP complex acts as an activator when glucose is scarce, enhancing transcription.

A repressor is a protein that inhibits transcription by binding to a DNA sequence called the operator, which is usually located near or overlapping the promoter. When bound, the repressor physically blocks RNA polymerase from initiating transcription, exerting negative control.

An inducer is a small molecule that turns on gene expression, often by binding to a repressor and inactivating it or by activating an activator protein. In the lac operon, for instance, allolactose serves as an inducer by binding to the lac repressor, releasing it from the operator, and allowing transcription to occur.

A corepressor is a small molecule that works with a repressor to stop gene expression. When it binds to an inactive repressor, it causes a conformational change that activates the repressor and enables it to bind DNA. In the arg operon, arginine functions as a corepressor, helping the ArgR repressor shut down transcription when arginine levels are high.

An effector is a general term for any small molecule—such as an inducer or corepressor—that binds to a regulatory protein (activator or repressor) and modifies its activity, either enhancing or inhibiting transcription.

The operator is a specific DNA sequence adjacent to the promoter that serves as the binding site for regulatory proteins such as repressors or activators. Its role is to control access of RNA polymerase to the structural genes.

An operon is a cluster of functionally related genes that are transcribed together as a single mRNA from one promoter and operator. This arrangement allows coordinated regulation of genes involved in the same biological pathway—for example, the lac operon for lactose metabolism or the trp operon for tryptophan biosynthesis.

Finally, a regulon is a collection of multiple operons or genes scattered throughout the genome that are all regulated by the same regulatory protein or signal. For example, the LexA regulon controls numerous genes involved in the bacterial SOS response.

a) Electron flow that generates proton motive force (downhill to O₂)

Fe²⁺ (E₀′ = +0.20) → cytochrome c (≈ +0.25) → cytochrome a (≈ +0.39) → ½O₂/H₂O (+0.82)

This downhill flow moves electrons from a lower to a higher redox potential, releasing energy that is used to pump protons across the membrane, generating a proton motive force (PMF). The PMF then drives ATP synthase, producing ATP through oxidative phosphorylation.

b) Electron flow that generates NADH (uphill via reverse electron transport)

Fe²⁺ (+0.20) → cytochrome c (+0.25) ⇢ (PMF-driven, uphill) ⇢ cytochrome b (+0.035) → ubiquinone/ubiquinol (+0.11) ⇢ (reverse Complex I/NDH-1) ⇢ NAD⁺ (−0.32) → NADH

Because the NAD⁺/NADH couple (−0.32 V) has a much more negative potential than Fe²⁺ (+0.20 V), this transfer is energetically unfavorable and must be driven uphill using energy from the PMF. This process, called reverse electron transport, allows the cell to produce NADH for biosynthetic reactions.

Bacteria can produce ATP and NADH by oxidizing molecular hydrogen (H₂) through a process known as chemolithotrophy, which uses inorganic compounds as energy sources. This process relies on specialized enzymes called hydrogenases, which catalyze the reaction
H₂ → 2H⁺ + 2e⁻, splitting hydrogen into protons and electrons.

The released electrons are transferred from hydrogenase to the electron transport chain (ETC) embedded in the bacterial membrane, where they are passed through a series of redox carriers such as quinones and cytochromes. As electrons flow through the ETC, energy is released and used to pump protons (H⁺) from the cytoplasm to the periplasmic space, creating a proton motive force (PMF) across the membrane. When the protons flow back through ATP synthase, the enzyme harnesses this gradient to synthesize ATP from ADP and inorganic phosphate — a process called chemiosmosis.

At the same time, some of the electrons generated from hydrogen oxidation are used to reduce NAD⁺ to NADH, either directly by soluble hydrogenases or indirectly via reverse electron transport driven by the proton gradient. The resulting NADH serves as an electron carrier and provides reducing power for biosynthetic reactions such as carbon fixation or anabolic metabolism.

This mechanism is particularly important in chemolithoautotrophic bacteria, which use hydrogen as their primary energy source and can fix CO₂ through pathways like the Calvin cycle. By oxidizing hydrogen, these bacteria efficiently generate both ATP for energy and NADH for biosynthesis, allowing them to thrive in environments with few organic nutrients.

Bacterial Sox systems (sulfur oxidation systems) enable sulfur-oxidizing bacteria to extract energy by oxidizing reduced sulfur compounds such as hydrogen sulfide (H₂S), elemental sulfur (S⁰), and thiosulfate (S₂O₃²⁻) to sulfate (SO₄²⁻). These reactions are chemolithotrophic processes that use sulfur compounds as electron donors and couple their oxidation to ATP and NADH production through the electron transport chain (ETC).

According to the redox tower, reduced sulfur compounds have relatively high-energy (more negative) reduction potentials, while oxygen and NAD⁺ are lower-energy (more positive) electron acceptors. As electrons flow “down” the redox tower—from high-energy sulfur donors to low-energy acceptors—energy is released. The Sox system captures this energy by transferring electrons from sulfur compounds to cytochromes and quinones in the bacterial membrane. These carriers pass the electrons through the ETC, releasing free energy that drives proton pumping across the membrane. The resulting proton motive force (PMF) powers ATP synthase, which converts ADP and inorganic phosphate into ATP via chemiosmosis.

To generate NADH, some of the electrons from sulfur oxidation are pushed “uphill” through reverse electron transport, an energy-requiring process that uses the PMF to reduce NAD⁺ to NADH. The NADH produced provides the reducing power necessary for CO₂ fixation through the Calvin cycle, supporting autotrophic growth.

The Sox pathway involves several key periplasmic proteins—SoxXA, SoxYZ, SoxB, and SoxCD—that work together to oxidize sulfur stepwise while keeping intermediates bound to the carrier protein SoxYZ. Electrons released from this pathway are passed to cytochrome c, which connects the Sox reactions to the electron transport chain. Ultimately, the oxidation of reduced sulfur to sulfate provides both ATP (energy conservation) and NADH (reducing power), allowing sulfur-oxidizing bacteria to thrive in environments rich in inorganic sulfur compounds.

Mutating both tryptophan codons (TGG → GGG) in the leader sequence of the trp operon disrupts the normal attenuation mechanism that regulates transcription of the structural genes. Under normal conditions, the leader peptide acts as a sensor of intracellular tryptophan levels. When tryptophan is abundant, the ribosome quickly translates the two tryptophan codons, allowing formation of the terminator hairpin (3–4 stem-loop) in the mRNA, which causes transcription to stop before reaching the structural genes. When tryptophan is scarce, the ribosome stalls at those tryptophan codons, allowing an antiterminator structure (2–3 loop) to form instead, enabling RNA polymerase to continue transcription of the genes needed for tryptophan biosynthesis. However, when the tryptophan codons are replaced with glycine codons, the ribosome no longer pauses in response to tryptophan scarcity, since glycine is readily available. As a result, the terminator hairpin forms constantly, causing premature termination of transcription regardless of tryptophan levels. This means the operon cannot respond to low tryptophan conditions, and the bacterium would fail to synthesize tryptophan when it is needed, impairing its ability to survive in tryptophan-poor environments. Overall, these mutations lock the trp operon in a state of constant repression, eliminating its regulatory flexibility.

Attenuation does not occur in mammalian cells because it relies on the coupling of transcription and translation, a process unique to prokaryotes. In bacteria, transcription and translation happen simultaneously in the cytoplasm, allowing the ribosome to sense amino acid availability and influence transcription in real time. For example, in the trp operon of E. coli, the ribosome stalls at tryptophan codons when tryptophan is scarce, permitting the formation of an antiterminator structure so transcription continues. When tryptophan is abundant, the ribosome moves quickly, forming a terminator hairpin that stops transcription early—this is the essence of attenuation.

In contrast, mammalian (eukaryotic) cells separate these processes: transcription occurs in the nucleus, while translation takes place in the cytoplasm. Because they are compartmentalized by the nuclear membrane, transcription and translation cannot occur simultaneously, making attenuation impossible. Instead, mammalian cells regulate gene expression through more complex mechanisms such as transcription factors, chromatin remodeling, RNA splicing, epigenetic regulation, and signaling pathways that control RNA synthesis and processing. These alternative regulatory systems allow precise control of gene expression without relying on attenuation.

Bacteria rapidly activate and deactivate stress responses like the SOS and heat-shock systems using positive and negative feedback loops to ensure survival while conserving energy.

The SOS response is triggered by DNA damage, which produces single-stranded DNA that activates the RecA protein. Activated RecA stimulates the self-cleavage of LexA, a repressor that normally inhibits SOS genes. As LexA levels drop, genes responsible for DNA repair, cell division inhibition, and mutagenesis are expressed. This forms a positive feedback loop, because more DNA damage leads to more RecA activation, more LexA cleavage, and faster gene induction. Once the DNA is repaired, there is less single-stranded DNA to activate RecA, allowing LexA to accumulate again and repress the SOS genes. This restoration of LexA activity acts as negative feedback, shutting down the response and returning the cell to normal conditions.

The heat-shock response works similarly through the sigma factor σ³² (RpoH). Under normal conditions, σ³² is rapidly degraded, but when temperature rises and proteins misfold, chaperones like DnaK and DnaJ become occupied with refolding these proteins. This stabilizes σ³², allowing it to accumulate and activate heat-shock genes that produce more chaperones and proteases. This creates a positive feedback loop that quickly boosts the cell’s defense against heat stress. As conditions improve and fewer misfolded proteins remain, newly made chaperones bind to σ³² again, promoting its degradation. This negative feedback turns off the heat-shock response once the cell’s proteins are stabilized.

Together, these systems illustrate how bacteria use positive feedback for rapid activation under stress and negative feedback for precise shutdown once homeostasis is restored — ensuring efficient, adaptable control of gene expression in changing environments.

Chemotaxis is a temporal, not a spatial, response because bacteria like E. coli are too small to detect concentration differences across their cell body at a single moment in space. Instead of comparing chemical levels at two points on their surface, they compare changes over time as they move. While swimming through a chemical gradient, bacteria use methyl-accepting chemotaxis proteins (MCPs)—specialized chemoreceptors embedded in the cell membrane—to continually sense whether concentrations of attractants (such as nutrients) or repellents are increasing or decreasing.

When the bacterium detects that conditions are improving (a higher attractant concentration than a few seconds earlier), it maintains a smooth, straight “run” by keeping its flagella rotating counterclockwise. If the concentration worsens, it increases the frequency of “tumbles”, randomly reorienting itself to find a better direction. This behavior depends on temporal sampling, meaning the cell measures changes between past and present signals rather than comparing two points in space simultaneously.

In contrast, spatial sensing—the ability to detect different concentrations at the same time across a single cell—is characteristic of larger or slower-moving eukaryotic cells, not bacteria. Because prokaryotic cells lack the physical size to sense gradients spatially, their chemotactic navigation relies entirely on temporal feedback, allowing them to effectively climb or descend chemical gradients despite their microscopic dimensions.

Bacteria respond to population density through a communication process known as quorum sensing, which allows them to coordinate behavior collectively once a critical number of cells is reached. Each bacterium produces and releases small signaling molecules called autoinducers into the surrounding environment. As the bacterial population grows, these autoinducers accumulate. When their concentration surpasses a specific threshold, they bind to receptor proteins—either transcriptional activators or sensor kinases—that trigger changes in gene expression across the entire population.

This synchronized regulation enables bacteria to act like a multicellular community, carrying out activities that are only effective when performed by many cells at once. Through quorum sensing, bacteria can initiate biofilm formation, virulence factor production, antibiotic resistance, and even bioluminescence. A classic example is Vibrio fischeri, which produces light only when a high population density is reached inside the light organ of its squid host.

Quorum sensing involves different signaling systems depending on the bacterial type. In Gram-negative bacteria, autoinducers are often small molecules like acyl homoserine lactones (AHLs) that freely diffuse across the cell membrane, while Gram-positive bacteria typically use small peptides recognized by membrane-bound receptors. In both cases, the response ensures that certain genes are expressed only when the bacterial community is large enough for the resulting behavior to be beneficial.

Overall, quorum sensing represents a density-dependent regulatory mechanism that lets bacteria sense their environment, conserve energy, and behave cooperatively—enhancing their survival, adaptability, and ability to interact with hosts or competitors.