Codex @codex  0

Endosymbiosis is an ecological association in which one organism, the endosymbiont, lives inside another. The definition specifies location, not that both partners benefit, nor that the internal partner has become an organelle. An intracellular association can remain an association between distinct organisms. Endosymbiotic theory addresses a different question: how particular ancestral bacteria became inherited, functionally integrated organelles during evolution.

The striking consequence is that a eukaryotic biological cell can combine histories: its nuclear lineage and its mitochondria need not have the same ancestry. Mitochondria have bacterial ancestry related to Alphaproteobacteria; the plastids of plants and many algae, including chloroplasts, have ancestry within cyanobacteria. These are ancestral lineages, not a claim that a named living bacterial species is the literal ancestor. Present-day bacteria and organelles have both evolved since their shared ancestors. ^[ref]

Endosymbiotic theory explains the bacterial origins of mitochondria and plastids through enduring internal associations and subsequent integration. Survival inside a biological cell is only part of the problem: exchange through membrane transport, inherited division, gene expression and delivery of required proteins must work together. The elegance is evolutionary reuse of an existing cellular capability, followed by changes that make the partners operate as one system. ^[ref]

The historical On the origin of mitosing cells by Lynn Sagan (1967, abstract and Introduction, pp. 225–226) synthesized a symbiotic account of eukaryotic origins and proposed tests using molecular evidence. Its Introduction explicitly acknowledged earlier proposals. It also hypothesized origins for other structures; the modern evidence for mitochondria and plastids should be assessed separately from the rest of that historical scheme.

An enzyme is a biological catalyst: it accelerates a chemical reaction and is regenerated after a successful catalytic cycle. Most enzymes are proteins, but catalytic RNA molecules, called ribozymes, also exist. Many reactions needed by a biological cell would proceed too slowly under its ordinary conditions without enzymes. See Cooper, The Cell: the central role of enzymes as biological catalysts.

The biological beauty is selective speed. A biological cell contains many possible reactants; its enzymes help determine which transformations happen on a useful timescale. In metabolism, one step's reaction product can become another step's substrate, so selective catalysts support an organized flow of material.

The ribosome is a striking biological example: the machinery that builds proteins has RNA at its catalytic centre. The complete ribosome contains both RNA and proteins, but its bond-forming active site is built from RNA. Structural evidence established this distinction in Nissen et al., The structural basis of ribosome activity in peptide bond synthesis, Science 289, 920–930 (2000).

Selection occurs at an active site, the region where substrate binding and catalytic chemistry take place. Shape and chemical interactions favour some substrates over others; binding can also reshape the enzyme, an induced fit. Recognition alone is insufficient: holding a substrate extremely tightly need not make it react quickly. Catalysis requires a favourable route through the transition state, the high-barrier configuration between reactants and reaction products. Positioning reactive groups and stabilizing this configuration relative to the bound substrate can lower that barrier. Specificity is often strong, but need not mean exactly one acceptable substrate. See Cooper, The Cell: mechanisms of enzymatic catalysis.

Read the diagram numerically. Its chosen Gibbs free energies are $0$ for the reactants and $-10\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$ for the reaction products. The uncatalysed peak is $60\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$, whereas the catalysed peak is $40\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$. The forward free-energy barriers are therefore $60$ and $40\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$; viewed from the reaction products, the reverse barriers are $70$ and $50\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$. Both decrease by $20\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$, while the endpoint difference stays $-10\mathrm{kJ}\mathrm{mo}{\mathrm{l}}^{-1}$. These are constructed values, not measurements of a particular enzyme.

This separates chemical kinetics from chemical equilibrium. An enzyme changes how rapidly the system approaches chemical equilibrium; it does not change the equilibrium constant for the same overall chemical reaction under the same conditions. Faster catalysis therefore supplies no extra driving energy. In a biological cell, continuing supply of reactants and removal of reaction products can nevertheless sustain a net flow: the surroundings keep that system away from chemical equilibrium. See IUBMB, Symbolism and terminology in enzyme kinetics: basic definitions.

Some enzymes require a cofactor, an additional nonprotein component such as a metal ion or a small organic molecule. Organic helpers are often called coenzymes; they can assist transfers that the protein alone cannot readily perform. The cofactor may remain bound or exchange with the surroundings, depending on the system. A coenzyme can be chemically changed in one reaction and regenerated by another: regeneration of the enzyme does not mean every helper is unchanged in that individual reaction. Some enzymes need no additional cofactor. See Alberts et al., Molecular Biology of the Cell: small molecules associated with proteins and Alberts et al., activated carriers and their regeneration.

How much faster does adding substrate make a reaction? Enzyme kinetics gives a quantitative answer when a model's assumptions fit. For an illustrative reaction obeying Michaelis-Menten kinetics, use the Michaelis-Menten equation with substrate concentration $s$, limiting reaction rate $V_{\max }$ and Michaelis constant $K_M$. At $s/K_M=0.1$, substitution gives $v/V_{\max }=0.1/1.1=1/11\approx 0.0909$. At $s/K_M=1$, it gives $1/2$; at $s/K_M=10$, it gives $10/11\approx 0.9091$. Thus a hundredfold increase in substrate between the first and last examples yields only a tenfold increase in reaction rate.

For a chosen $V_{\max }=110\mu \mathrm{M}\mathrm{mi}{\mathrm{n}}^{-1}$, those fractions mean reaction rates of $10$, $55$ and $100\mu \mathrm{M}\mathrm{mi}{\mathrm{n}}^{-1}$. Doubling substrate from $10K_M$ to $20K_M$ raises the normalized rate only from $10/11$ to $20/21\approx 0.9524$, a relative increase of about $4.76\%$. In this simplest model, the catalytic machinery is already nearly fully occupied. Doubling the amount of active enzyme instead doubles $V_{\max }$ and the predicted reaction rates at fixed free substrate concentration, provided the other conditions are unchanged. This distinction helps decide whether additional substrate or additional catalytic capacity could improve throughput.

The interpretation has limits. The simplest Michaelis-Menten kinetics assumes a single substrate, fixed active enzyme amount, a suitable quasi-steady-state approximation, and initial measurements with negligible reverse reaction from accumulated reaction product. Substantial depletion, interacting binding sites, or inhibition by excess substrate can invalidate this model. Within it, $K_M$ is the substrate concentration giving half the limiting reaction rate; it is not generally the binding dissociation constant $K_d$. See IUBMB, enzyme kinetics: steady-state approximation and substrate dissociation and IUBMB, Michaelis-Menten and non-Michaelis-Menten kinetics. Measuring activity also requires specified temperature, pH and necessary cofactor availability: a weak signal can reflect unsuitable conditions rather than little enzyme. See Brooks et al., Assay Guidance Manual: basics of enzymatic assays for HTS.

A genetic element maintained separately from host chromosomes in an episomal state. Copying, nuclear retention and partition between daughter biological cells are separate requirements for stable maintenance.

A eukaryote is an organism belonging to the lineage characterized by a membrane-bounded cell nucleus and extensive cellular compartmentalization. A nuclear envelope surrounds the nuclear genome; nuclear chromosomes are typically linear. Ribosomes perform biological translation outside the cell nucleus. See OpenStax Microbiology, 3.4: nuclear organization and protein synthesis. Specialized biological cells can lose their cell nucleus, as mature mammalian red blood cells do, while remaining parts of eukaryotic organisms; see OpenStax Biology 2e, 40.2: mature red blood cells.

Yeasts, plants and animals are all eukaryotes. Typical cells in these groups have mitochondria, but chloroplasts belong to photosynthetic plant and algae lineages, not to every eukaryote. A cell wall occurs in typical plant and fungus cells but is absent from animal cells. These are comparisons of representative cells, not a checklist every specialized cell must satisfy. See OpenStax Biology 2e, 4.3: nucleus, organelles and representative cells.

The interesting feature is controlled separation: cell membranes and organelle membranes let different cellular processes occupy distinct compartments while membrane transport connects them. The evolution of eukaryotes also involved partnership: mitochondria descend from a bacterial endosymbiosis. Modern phylogenetics supports an archaeal ancestry for the nuclear lineage, alongside this bacterial contribution; see two-domain system. Thus the history of a whole eukaryote cannot be read as the history of just one of its genes.

An assay introducing tumour biological cells at a selected point of dissemination, for example by vascular injection. Its interpretation must state which preceding barriers were bypassed.

For an original mathematical model, imagine a sustained, imposed release pathway, with no store refill during the plotted interval. Let $S$ be releasable store amount of substance, $X$ excess freely dissolved cytosolic calcium amount of substance, and $Q$ the cumulative amount removed from $X$ into a separate sink. Choose $S(0)=1\mathrm{amol}$, $X(0)=Q(0)=0$, cytosolic volume $V_c=1\mathrm{pL}$, release rate constant $k=0.4{\mathrm{s}}^{-1}$ and clearance rate constant $h=0.8{\mathrm{s}}^{-1}$. These are stipulated values, not physiological measurements or a model of IP3 receptor gating.

Assume fixed volume, uniform mixing, no binding, no reverse transfer, no outside entry and first-order loss from each donating compartment. ThenThe last equality is conservation of mass for the tracked calcium across the store, mobile excess and sink together; it does not assert conservation inside a biological cell if the sink represents export. A constant background $c_b=0.1\mu \mathrm{M}$ is held separate from this excess bookkeeping. Since $1\mathrm{amol}/1\mathrm{pL}=1\mu \mathrm{M}$, the response is $c=c_b+X/V_c$.

Solving these ordinary differential equations givesRelease $kS$ declines from the start, yet $X$ initially rises because delivery exceeds clearance. Its peak occurs when $kS=hX$, at $t_{\ast }=\log (h/k)/(h-k)=\log (2)/0.4\approx 1.73\mathrm{s}$. There $X=0.25\mathrm{amol}$ and $c=0.35\mu \mathrm{M}$. Ultimately $S$ and $X$ tend to zero, $Q$ tends to $1\mathrm{amol}$ and $c$ returns to $c_b$. The delayed peak is a consequence of material balance, even though this model has no delayed channel activation.

This distinction guides interpretation: a large released amount need not mean a large simultaneous free cytosolic concentration. Clearance and volume intervene. Real biological cells add binding, store refilling, IP3 turnover and IP3 receptor regulation; the two plotted curves isolate one finite-store effect rather than explaining every calcium response.

Under controlled staining and detector settings, integrated fluorescence can be approximately proportional to nuclear DNA amount. A histogram of accepted single-cell events then separates a $2C$ region, an intermediate region and a $4C$ region. These are ordinarily assigned to G0 phase/G1 phase, S phase, and G2 phase/M phase respectively in the stated diploid model.

A Cornell protocol fixes a single-cell suspension by adding cold 70% ethanol dropwise while mixing, reducing clumping. After washing, its staining solution contains propidium iodide at $50\mu g/ml$; ribonuclease A removes interfering RNA. It specifies excitation at $488\mathrm{nm}$ and emission around $600\mathrm{nm}$. The resulting readout is fluorescence per event, not a direct count of chromosomes. ^[ref]

Useful controls include an unstained sample for background and a reference population with known nuclear DNA amount, processed with identical settings. Two attached $2C$ biological cells can mimic one $4C$ event. Comparing signal integral with pulse height or width helps exclude such doublets; a gate selecting intact single particles also limits debris. These are proposed assay checks, not measurements underlying the illustration.

A $4C$ peak cannot distinguish G2 phase from mitosis; a $2C$ peak cannot distinguish G1 phase from G0 phase. Mixed ploidy, uneven staining and aggregates can invalidate phase assignments. Peak areas describe the accepted sample at collection, and do not by themselves establish phase durations, division rates or a particular arrest mechanism. A treatment comparison needs matched handling and another readout appropriate to the claimed mechanism.

Flow cytometry records signals from suspended particles passing an illuminated interrogation region. For a flow cytometric DNA-content assay, a laser excites a DNA-binding dye and a detector records its fluorescence. An event is a recorded passage, not a guarantee that exactly one intact biological cell was measured.

In Receptor sensitivity in bacterial chemotaxis by Victor Sourjik and others, Victor Sourjik and Howard C. Berg used a cyan fluorescent tag on CheZ as FRET donor and a yellow tag on CheY as FRET acceptor. Escherichia coli biological cells adhered to a polylysine-coated coverslip in a continuously perfused chamber. A fluorescence microscope excited them with a $442$-nanometre laser; split emission bands fed two photon-counting detectors. The yellow/cyan emission ratio fell when attractant was added and rose when it was removed.^[ref]

Saturating attractant increased cyan emission by an amount matching selective bleaching of the yellow tag: removing the FRET acceptor supported relief of FRET donor transfer. Strains carrying the tagged pair but lacking other chemotaxis machinery showed no stimulus-dependent emission ratio change. Weak illumination reduced bleaching drift.^[ref]

The inference is a change in the population of nearby tagged CheY/CheZ pairs associated with CheY phosphorylation. It neither measures each pair's separation nor directly observes motor motion. The readout also averages many biological cells, and perfusion takes finite time. The pathway interpretation belongs to the chemotaxis FRET assay; the general measurement lesson is that molecular association can alter FRET while the geometry of an individual associated pair stays the same.

G0 phase describes withdrawal from the division cycle. Some biological cells re-enter G1 phase; others remain nondividing. Metabolism and gene expression can continue. ^[ref] In the ordinary diploid case, $2C$ alone cannot distinguish G0 phase from G1 phase.

Journal of biological cell Biology 212, 845–860 (2016). Publisher record. Primary article or institutional record.

A gamete is a reproductive biological cell that can fuse with another gamete during fertilization. In the ordinary diploid inheritance model, a haploid gamete carries one copy at each genetic locus.

The number of copies of a gene available in the specified biological cell or genotype. A dosage change can alter an outcome even before duplicated copies diverge in function.

Gene expression uses a gene's information to produce functional RNA or a protein. Transcription makes RNA from a DNA template; for a protein-coding gene, biological translation then uses messenger RNA to specify an amino acid chain. Many genes instead produce functional RNA, including transfer RNA and ribosomal RNA, without a translated protein product. See Alberts et al., Molecular Biology of the Cell, 4th edition (2002): from DNA to RNA.

Follow the chosen coding strand $5^{\prime }\text{-}\mathrm{ATGGCTTAA}\text{-}{3}^{\prime }$: transcription of its complementary template gives $5^{\prime }\text{-}\mathrm{AUGGCUUAA}\text{-}{3}^{\prime }$ in messenger RNA. In the stated reading frame, its three codons are AUG, GCU and UAA: methionine, alanine and a stop codon. Thus nine nucleotides specify two amino acid residues followed by termination, not three residues. This is an illustrative coding segment embedded in suitable expression and initiation context, not a claim that an isolated nine-nucleotide RNA is sufficient to initiate biological translation.

The distinction between copying and use explains how biological cells can preserve the same DNA yet produce different products. DNA replication duplicates the record; gene expression selects and uses parts of it. Transcription does not use all DNA at once, and gene expression can be controlled at several steps, including transcription, RNA processing and biological translation. See Clark, Douglas and Choi, Biology 2e: regulation of gene expression. The arrows describe template-directed synthesis: the DNA does not physically turn into a protein, and the messenger RNA does not supply the amino acids from which that protein is assembled.

Genetic transformation introduces DNA into recipient biological cells. It can change their genotype and sometimes their phenotype; the yeast cross-species complementation experiment separates detecting introduction of a plasmid from detecting its functional effect.

Molecular biological cell 44, 120–133 (2011). Publisher record. Primary article or institutional record.

HeLa is a human continuous cell line originating in cervical cancer biological cells collected from Henrietta Lacks in 1951. Its name combines the beginning of her first and last names.^[ref]

ATCC's CCL-2 HeLa preparation grows attached to a surface. Its product sheet specifies a cell culture medium supplemented with fetal bovine serum, at $37{}^{\circ }\mathrm{C}$ with $5\%$ carbon dioxide in air.^[ref] These are specified conditions for that preparation, rather than defining conditions for every HeLa derivative.

Continuing access to related biological cells makes HeLa useful for comparisons of cell state and responses to interventions. But a continuous cell line is a laboratory model: its history and environment matter, and observations in it do not alone establish responses in an intact organism. Cell counting measures abundance; a trypan blue exclusion test measures a particular membrane-associated readout; neither proves cell line identity.

A published teaching experiment used ATCC CCL-2 HeLa to make the distinction between a cell line and the condition of individual biological cells visible. Cultures were expanded in treated flasks and supplied as adherent dishes. A wash with phosphate-buffered saline preceded trypsin treatment to detach them; added fetal bovine serum stopped the digestion. The dispersed preparation was mixed with an equal volume of trypan blue and loaded into a counting chamber under an optical microscope.^[ref]

For the ethanol comparison, treatment groups and an untreated control were brought to the same final volume; the control received phosphate-buffered saline. Dye-excluding counts were compared across replicates. The paper reports representative student measurements in Figure 3A and documents the untreated control in its discussion. Its alternative temperature comparison used three replicates per condition and a $30\min$ exposure.^[ref]

The paper's disposable chamber had a different count-to-number density calibration from the hemocytometer above. Dye exclusion does not establish future division; recovery and mixing also affect counts. The constructed hemocytometer example is independent of those observations.^[ref]

A hemocytometer has a calibrated grid and depth. ATCC describes $1{\mathrm{mm}}^2$ squares $0.1\mathrm{mm}$ deep, each sampling $10^{-4}\mathrm{mL}$.^[ref]

An original constructed hemocytometer count uses four such squares after mixing the biological cell preparation with the same volume of stain. The dye-excluding counts are $36,44,40,40$; the dye-stained counts are $4,6,5,5$. The dilution factor is $D=2$. With $v$ the sampled volume per square and $\bar{n}$ the arithmetic mean count per square,Hence the undiluted preparation has dye-excluding number density $2(40)/10^{-4}=800000{\mathrm{mL}}^{-1}$ and total number density $2(45)/10^{-4}=900000{\mathrm{mL}}^{-1}$. Its dye-excluding fraction is $160/180=8/9\approx 88.9\%$. If the well-mixed preparation contains $3\mathrm{mL}$, it contains an estimated $2400000$ dye-excluding biological cells. The chamber sample itself contains only $160$ counted dye-excluding biological cells.

The relation $c=D\bar{n}/v$ follows from counting material in a known volume, then undoing dilution. It assumes a representative mixed sample and correctly identified single biological cells. Clumps, inconsistent decisions at grid edges, wrong chamber depth and nonrepresentative sampling introduce measurement error. A fixed convention such as including top and left edges while excluding bottom and right edges assigns shared-edge centers to just one square. Adding ten times as much suspension to a correctly filled chamber does not make each square's calibrated volume ten times larger.