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An active site is the region of an enzyme where a substrate binds and undergoes a chemical reaction. In a protein enzyme, it includes the amino acid residues that contact the substrate and those that directly assist catalysis. These roles can overlap: an amino acid side chain can both orient a substrate and participate in chemistry. The active site occupies only part of the folded protein; the remaining amino acid residues help establish its protein structure, stability and movement. An active site need not be a consecutive stretch of protein primary structure: protein folding can bring distant amino acid residues together. A ribozyme instead supplies an RNA active site. See Alberts et al., Molecular Biology of the Cell, Protein Function: binding sites and catalytic sites.

Binding and catalysis answer different questions. A protein binding site recognizes shape and chemistry through such interactions as hydrogen bonds, electrostatics and the hydrophobic effect. At an active site, appropriate positioning also helps a substrate reach the transition state of a chemical reaction. Some amino acid side chains transfer a proton; others participate transiently in a covalent bond. The enzyme-substrate complex is the associated enzyme and substrate, before conversion to a reaction product. Merely accumulating this complex does not establish rapid catalysis: stabilizing the bound substrate without sufficiently stabilizing the transition state can leave a large barrier. The important complementarity is chemical as well as geometric. See Geoffrey M. Cooper, The Cell, The Central Role of Enzymes as Biological Catalysts: substrate binding and transition-state stabilization.

The cartoon makes a useful experimental distinction. Changing a contacting amino acid residue might weaken substrate binding, change the reaction rate, or do both. Changing a distant amino acid residue can also disrupt the active site by changing protein folding. Therefore a loss of activity after a mutation alone does not locate a catalytic group: compare binding, protein structure and enzyme kinetics as well. This is an interpretation guide, not a report of a particular mutation experiment.

A concrete example is hexokinase: it transfers the terminal phosphate group of ATP to the $6$-position hydroxyl group of a sugar such as glucose. Specificity need not mean only one acceptable substrate: the hexokinase reaction class includes several possible acceptor sugars. Which are used, and at what reaction rates, depends on the particular enzyme and conditions. See Swiss Institute of Bioinformatics, ENZYME entry EC 2.7.1.1: hexokinase reaction and acceptor range.

Bennett and Steitz compared yeast hexokinase A crystallized with glucose against hexokinase B crystallized without glucose. X-rays scattered by the crystals supplied diffraction measurements from which the authors reconstructed protein structure; comparing coordinates located movement, rather than measuring a reaction rate. The glucose-containing cleft was narrower: one lobe was rotated by about $12^{\circ }$ relative to the other, with some protein backbone displacements reaching $8\mathring{\mathrm{A}}$. This makes the active site a moving arrangement of interacting groups, rather than a permanent hole. See William S. Bennett Jr. and Thomas A. Steitz, Glucose-induced conformational change in yeast hexokinase, PNAS 75 (1978), 4848–4852.

The structural comparison has two explicit limitations: different forms of hexokinase and different crystal environments were compared, and static coordinates do not record the order of binding and movement. Closure is consistent with induced fit, but those structures alone cannot exclude conformational selection. The solution experiment under induced fit addresses whether compaction also occurs outside a crystal.

The amino acid composition lists amounts or proportions of the different amino acid residues in a protein or specified polypeptide chain, without giving their order. The toy MAGAMG and MMAAGG each have counts M:2, A:2, G:2, or a proportion $1/3$ of each. Knowing composition therefore does not supply protein primary structure. Sanger's account begins by distinguishing the already available composition measurements from the largely unknown residue order. See Sanger's authored 1985 account of the move from composition to terminal-residue analysis.

An amino acid residue is the portion of an amino acid incorporated into a polypeptide chain. For example, the A in the illustrative protein primary structure is an alanine residue, rather than a free alanine molecule floating beside the chain. Counting amino acid residues counts chain positions; repeated letters are separate positions. See IUPAC-IUB, section 3AA-16: symbols for amino-acid residues.

A proteinogenic amino acid represented by R or Arg in protein primary structure. Its amino acid side chain carries predominantly positive electric charge near neutral pH.^[ref]

Trypsin preferentially cleaves after its amino acid residues.

A beta sheet consists of extended stretches of protein backbone lying alongside one another, joined by backbone hydrogen bonds. The stretches can belong to different portions of the same polypeptide chain or to different chains.

Each extended stretch is a beta strand. Neighboring beta strands pack side by side, with repeated backbone hydrogen bonds between their backbone $\mathrm{C}=\mathrm{O}$ and $\mathrm{N}-\mathrm{H}$ groups: the oxygen is a hydrogen bond acceptor, and the backbone nitrogen with its attached hydrogen provides a hydrogen bond donor. These are interactions of the shared protein backbone, not merely contacts between amino acid side chains. Unlike an alpha helix, a beta sheet joins adjacent extended stretches rather than coiling one stretch around an axis. See Clark, Douglas and Choi, Biology 2e, 3.4: backbone groups in secondary structure.

Read each beta strand in the sequence direction from the N-terminus toward the C-terminus, even when it is an internal part of a longer polypeptide chain. In a parallel beta sheet, neighboring beta strands run in the same N→C direction; in an antiparallel beta sheet, they run in opposite N→C directions. A three-strand example can therefore have arrows right–right–right or right–left–right. A beta sheet can also mix the two arrangements. Direction does not identify chain membership: a single polypeptide chain can fold back to supply several beta strands, and different polypeptide chains can contribute to one beta sheet. See Alberts et al., Molecular Biology of the Cell: beta-sheet strand arrangements.

The pleated appearance comes from the geometry of the protein backbone. Successive amino acid residues carry amino acid side chains on alternating faces of an ideal beta strand, conventionally drawn above and below the sheet. The attachment positions are the amino acid alpha carbons. Thus a six-position cartoon has groups 1, 3 and 5 on one face and 2, 4 and 6 on the other. This alternation is a spatial pattern, not a rule that alternating positions must contain identical amino acids. See Clark, Douglas and Choi: side chains above and below the pleats.

Real beta sheets commonly twist and curve; pleating does not make them perfectly flat plates. Which beta strands are neighbors, how connecting regions join them, and how their faces pack are features of the three-dimensional protein structure. An alternating pattern in the protein primary structure alone neither establishes a beta sheet nor specifies a unique fold independently of surroundings and interacting molecules. See Breda et al., Protein Structure, Modelling and Applications: extended strands, sheet twist and three-dimensional organization and protein folding.

Why this arrangement is useful: repeated backbone hydrogen bonds organize extended chain regions into a framework that can contribute to the interior of a folded protein. Amino acid side chains and connecting loops give that framework chemical variety and can shape a protein binding site for a ligand. Its attractive feature is the combination of a reusable backbone pattern with variable exposed groups: similar beta sheets can support different recognition surfaces. See Alberts et al.: beta-sheet cores and loops that form binding sites.

Historically, Linus Pauling and Robert B. Corey published The Pleated Sheet, a New Layer Configuration of Polypeptide Chains in PNAS 37, 251–256 (1951). They reasoned from peptide bond geometry and possible backbone hydrogen bonds, proposing an idealized layer rather than measuring every modern beta sheet. Their initial model should not be identified with all present-day beta sheet geometries; they published further pleated-sheet configurations later in 1951. See Pauling and Corey: original 1951 paper and scan, ^[ref], and Pauling and Corey, Configurations of Polypeptide Chains With Favored Orientations Around Single Bonds: Two New Pleated Sheets, PNAS 37, 729–740 (1951).

A CDK1 phosphorylation state identifies modified amino acid residues. Different sites have opposite effects: in studied Xenopus laevis Cdc2, Thr14/Tyr15 are inhibitory, while Thr161 phosphorylation is activating. T denotes a threonine residue, Y a tyrosine residue, and numbers are protein primary structure positions. Do not assume this numbering for every species.^[ref]

The 1992 study supplies a useful separation of mechanisms: replacing Thr14, Tyr15 and Thr161 still permitted cyclin B binding, but replacing Thr161 prevented activation of histone H1 phosphorylation. Binding therefore did not suffice for catalytic activity in that preparation. Activating phosphorylation also occurred with catalytically inactive Cdc2, supporting a separate CDK-activating kinase rather than a requirement for intramolecular autophosphorylation.^[ref]

In this deliberately coarse classification, bound cyclin B, activating phosphorylation, and cleared inhibitory sites must coincide for the activity-permissive state. Removing inhibitory phosphate groups does not supply missing cyclin B or activating phosphorylation. Conversely, adding activating phosphorylation does not erase inhibitory modification. Actual preparations contain mixtures of states and additional regulators; this classification gives no numerical reaction rate.

A molecular chaperone is a protein that assists other proteins in reaching or maintaining useful arrangements, for example by limiting unwanted association during protein folding. Assistance does not rewrite the client's protein primary structure. In a crowded cytoplasm, reducing protein aggregation can make successful protein folding more reliable. See Cooper: molecular chaperones and folding assistance.

Cotranslational protein translocation is passage of a growing polypeptide chain across a membrane during biological translation. SRP recognizes an exposed hydrophobic ER signal sequence. The SRP receptor enables handoff to the Sec61 complex. Biological translation can slow during delivery; the ribosome remains on the cytosol side.^[ref]

The useful engineering feature is coordination: a targeting instruction can become accessible while the rest of its cargo is still being made. Recognition, docking and crossing remain distinct operations. A defect in any one can prevent successful import even when the protein primary structure contains an appropriate signal.

This route is not universal. Many tail-anchored proteins use delivery after biological translation, and their membrane-spanning segment lies near the C-terminus. Thus protein targeting signals need not be at the N-terminus.^[ref]

Protein denaturation is the loss of a protein's native spatial organization, often with loss of biological function. Changes in temperature, pH or surroundings can disrupt protein secondary structure, protein tertiary structure or protein quaternary structure. Protein denaturation ordinarily preserves the peptide bonds and protein primary structure; it is different from cutting the chain. Recovery by protein folding is possible in some cases, but removing the denaturing conditions does not guarantee that every protein recovers. See Clark, Douglas and Choi: denaturation, environmental conditions and loss of function.

In the diagram, unfolding preserves all eleven peptide bonds. Cleaving the connection between positions 6 and 7 leaves two six-residue fragments: each contains five peptide bonds, so there are $5+5=10$ in total. Simply cooling or otherwise restoring suitable conditions cannot recover the original chain by protein folding alone; the missing covalent bond would have to be remade. Conversely, protein aggregation can impede recovery without breaking any peptide bond. These distinctions explain why loss of function does not by itself reveal whether a protein unfolded, associated with other chains, or suffered chemical damage.

A disulfide bond is the single covalent bond between the two adjacent sulfur chemical atoms in a disulfide, conventionally written $\mathrm{R}-\mathrm{S}-\mathrm{S}-{\mathrm{R}}^{\prime }$. Here $R$ and $R^{\prime }$ stand for the attached organic groups; they may belong to the same molecule or to initially separate molecules. A disulfide bond is a chemical bridge, rather than a hydrogen bond or a mere contact between nearby groups. Two thiols, $\mathrm{R}-\mathrm{SH}$ and ${\mathrm{R}}^{\prime }-\mathrm{SH}$, can supply its two sulfur endpoints. ^[ref]

The stoichiometry of forming a disulfide bond is especially revealing. Written as an oxidation half-reaction, it isThe reverse reduction half-reaction consumes two electrons and two hydrogen ions to restore two thiols. Both sides contain two sulfur chemical atoms; the two hydrogen chemical atoms have moved into the two hydrogen ions, and the net electric charge remains zero because $+2-2=0$. Thus one disulfide bond corresponds to two thiol groups and two transferred electrons. These half-reactions describe redox bookkeeping, not a complete standalone chemical reaction: an accompanying reactant must accept or supply the electrons, and pH affects the actual protonation of the groups. Mercaptoethanol can promote reduction through exchange of disulfide bonds, but adequate excess and appropriate conditions are needed; adding it does not by itself prove complete reduction. ^[ref]

In a protein disulfide bond, the endpoints are normally the amino acid side chains of two cysteine amino acid residues. An intrachain protein disulfide bond closes an intramolecular bridge within one polypeptide chain; an interchain protein disulfide bond connects different polypeptide chains, and can arise by intermolecular crosslinking of initially separate molecules. Forming or reducing these bridges changes covalent bond connectivity and can change protein structure, while preserving the order of amino acid residues and the peptide bonds in the protein backbone. A complete chemical account of protein primary structure includes crosslinks as well as sequence, so preservation of sequence does not imply preservation of every covalent bond. Mature insulin has two interchain protein disulfide bonds and one intrachain protein disulfide bond. ^[ref]

A bridge is useful because it constrains which parts of a polypeptide chain can move apart without breaking a covalent bond. Its placement therefore matters as well as its presence. The ribonuclease A refolding experiment provides the existing experimental example: breaking and reforming four protein disulfide bonds can accompany loss and recovery of function. Counting four bridges alone cannot tell whether their endpoints have the native pairing. This is an attractive connection between simple chemical counting and the richer problem of protein folding.

How can an experiment reveal an initially hidden disulfide bond? Ellman's reagent, also called DTNB, measures free thiols by producing a yellow absorbing species. Its reactive disulfide bond undergoes exchange with the conjugate base $\mathrm{R}-{\mathrm{S}}^{-}$ of a thiol, producing a mixed disulfide attached to $R$ and one released 2-nitro-5-thiobenzoate species, abbreviated TNB. At the assay pH, the strongly colored form is the dianion $\mathrm{TN}{\mathrm{B}}^{2-}$: one negative charge is on sulfur and the other is on the carboxylic acid-derived $\mathrm{CO}{\mathrm{O}}^{-}$ group. With excess DTNB and complete reaction, one released TNB reports one reacted thiol. George L. Ellman's 1959 paper introduced the tissue assay; it was subsequently reexamined quantitatively. ^{[ref] [ref]}

The modern apparatus can be simple: tubes, pipettes, a buffer solution, cysteine standards, DTNB, and an instrument that measures absorbance through a cuvette at wavelength $412\mathrm{nm}$. The Pierce protocol uses $50\mu \mathrm{L}$ of a $4\mathrm{mg}\mathrm{m}{\mathrm{L}}^{-1}$ DTNB stock, $2.50\mathrm{mL}$ of buffer solution, and $250\mu \mathrm{L}$ of sample. Its buffer solution is $0.1\mathrm{mol}{\mathrm{L}}^{-1}$ sodium phosphate at pH 8.0 with a metal-binding additive. After 15 minutes at room temperature, measure absorbance against a buffer solution plus DTNB blank; run known cysteine concentrations in parallel. The observable is increased yellow color and increased blank-corrected absorbance as more accessible thiols react, rather than direct imaging of disulfide bonds. These are published protocol conditions, not measurements performed for this article. ^[ref]

For a before/after test, split a purified protein sample into matched aliquots. Measure accessible free thiols in one aliquot; subject the other to complete reduction of its disulfide bonds, remove the reducing reagent, then measure the resulting thiols with fresh DTNB. Process the unreduced aliquot through a matching cleanup. Normalize to recovered protein and known dilution, since cleanup can change both. To estimate all reducible protein disulfide bonds, expose buried groups using matching protein denaturation conditions in both aliquots and calibrate in that same chemical solution; a native-accessibility measurement answers a different question. A larger corrected reading after reduction supports conversion of previously unreactive groups into detectable thiols. Attribution to disulfide bonds requires excluding other reducible sulfur modifications. This is a proposed comparison based on the detection principles, not a reconstruction of Ellman's original tissue measurements. ^[ref]

Under the Beer-Lambert law, the blank-corrected absorbance is $A=\epsilon lc$, where $l$ is the cuvette path length, $c$ is the released TNB concentration, and $\epsilon$ is its molar absorption coefficient. Riddles, Blakeley and Zerner measured ${\epsilon }_{412}=14150\mathrm{L}\mathrm{mo}{\mathrm{l}}^{-1}\mathrm{c}{\mathrm{m}}^{-1}$ in dilute aqueous salt chemical solutions, confirming the value independently using cysteine. It must not be applied indiscriminately to other solvents or protein denaturation mixtures. ^[ref] For aliquots compared on the same final volume and protein concentration basis, the ideal estimate isThe factor of two follows from the reduction stoichiometry; ${\bar{n}}_{\mathrm{S}-\mathrm{S}}$ is the number of disulfide bonds per protein molecule, or an average for a mixed population. If the aliquots have different dilutions or recoveries, convert each reading to a common sample basis before subtracting.

As a hypothetical calculation, choose $c_{\mathrm{protein}}=10\mu \mathrm{mol}{\mathrm{L}}^{-1}$ in each final cuvette, two free thiols and two disulfide bonds per protein molecule, and $l=1\mathrm{cm}$. Before reduction, $c_{\mathrm{SH}}=20\mu \mathrm{mol}{\mathrm{L}}^{-1}$ gives $A=0.283$; complete reduction adds four thiols per protein molecule, so $c_{\mathrm{SH}}=60\mu \mathrm{mol}{\mathrm{L}}^{-1}$ gives $A=0.849$. The difference $0.566$ gives $c_{\mathrm{S}-\mathrm{S}}=20\mu \mathrm{mol}{\mathrm{L}}^{-1}$ and ${\bar{n}}_{\mathrm{S}-\mathrm{S}}=2$. The published tube volumes imply a dilution factor $2.80/0.250=11.2$; reversing that dilution multiplies both thiol and protein concentrations by 11.2 and leaves the bridge count per protein molecule unchanged.

Controls determine whether that arithmetic has chemical meaning. Use fresh cysteine standards and a processed blank without protein to detect color generated by residual reducing reagent. Even a reducing reagent containing no thiols can interfere: Han and Han showed that tris(2-carboxyethyl)phosphine, abbreviated TCEP, reduces DTNB to two TNB equivalents per TCEP. ^[ref] In the hypothetical model, $5\mu \mathrm{mol}{\mathrm{L}}^{-1}$ residual TCEP in only the reduced cuvette would add $A=0.1415$ and falsely add $0.5$ bridge per protein molecule. A stable endpoint and agreement after a further reduction treatment help check completion. A sample without DTNB checks its own color; cloudy samples can also attenuate light without producing TNB. Match pH, temperature, accessibility and calibration chemical solution, and minimize delay that permits reoxidation. If two already-free but buried thiols become accessible only in the reduced aliquot, they mimic one new disulfide bond per protein molecule. Finally, bulk absorbance supplies a count under these assumptions: it does not locate the cysteine endpoints, distinguish intrachain from interchain bridges, or establish the native pairing of a protein disulfide bond.

Gerald M. Edelman reduced IgG disulfide bonds and used dissociating chemical solutions, including $6\mathrm{mol}{\mathrm{L}}^{-1}$ urea. Ultracentrifugal analysis showed a decrease in molar mass, supporting a multichain protein. In the comparison reproduced in his lecture, starch gel electrophoresis in urea separated serum IgG and a urinary protein from the same patient, alongside their reduced and alkylation-treated preparations. The untreated samples provided the treatment comparison; the urinary material provided a chain-identity comparison. The treated serum sample revealed heavy- and light-chain bands. Two-dimensional peptide mapping compared the urinary protein with the isolated immunoglobulin light chain. Together these readouts supported a shared chain identity.^[ref]

The controls address distinct alternatives. Treatment-dependent separation supports chains joined in the starting protein; a matching gel electrophoresis position alone would not uniquely identify a chain. Agreement of peptide mapping adds an independent chemical comparison. Neither readout gives atomic coordinates, and this patient's preparation does not establish a common protein primary structure for all antibodies.

Reduction targets protein disulfide bonds, while protein denaturation weakens the folded assembly. Neither operation is the proteolysis that cuts peptide bonds. This explains why chain separation and Papain digestion of IgG give different inventories: four intact chain backbones in the first case, three assembled fragments in the second.

Immunoglobulin G is an antibody class whose conventional four-chain architecture supplies the example above.

An ordinary IgG has two matching immunoglobulin heavy chains and two matching immunoglobulin light chains, abbreviated $H_2{L}_2$. Its gamma-type immunoglobulin heavy chains contribute both to recognition and to interactions with immune partners.^[ref]

A useful way to read this picture is to follow a single polypeptide chain continuously from its N-terminus to its C-terminus, then examine the crosslinks separately. The same immunoglobulin heavy chain contributes to a Fab fragment and to the Fc fragment; these named fragments are not pre-existing separate chains. Conversely, one Fab fragment contains material from two different chains. The protein primary structure and the fragment inventory therefore answer different questions about the same protein.

In induced fit, a substrate binds to an enzyme and the resulting enzyme-substrate complex subsequently changes its protein structure. This adjustment can position interacting groups at the active site for catalysis. The mechanistic claim concerns order: binding precedes the relevant change. In conformational selection, the unbound enzyme already visits an alternative protein structure, which the substrate preferentially binds. Both processes can occur in one system. Let $E$ and $E^{\ast }$ denote two structures of the same enzyme, and $S$ a substrate. The limiting routes areThe star labels a structure, not a different protein primary structure, and does not by itself mean catalytic activity. Each reversible arrow abbreviates a step, not the whole catalytic cycle. See Gordon G. Hammes, Yu-Chu Chang and Terrence G. Oas, Conformational selection or induced fit: a flux description of reaction mechanism, PNAS 106 (2009), 13737–13741.

A rigid lock-and-key cartoon usefully suggests selective substrate recognition, but it suppresses movement of the protein and chemical changes at the active site. Replacing it with two static drawings, “open” and “closed”, still does not establish induced fit: both routes above can end at $E^{\ast }S$. Also, if complete closure prevents a substrate entering, the pre-existing structure selected in conformational selection might be partly open and undergo further induced fit after binding. The usefulness of the distinction is that it turns an attractive shape story into questions about enzyme kinetics. Relative traffic through the routes depends on populations, rate constants and free substrate concentration, rather than affinity alone. See Hammes, Chang and Oas, 2009: coupled pathways, their concentrations and the limitations of endpoint structures.

For this analytical illustration, hold the free substrate concentration $s$ constant and assume independent waiting times with exponential distributions of constant rates. The law of mass action gives the binding rate $a=k_{\mathrm{on}}s$; the competing change has rate $b$. The probability density function for binding to happen first at time $t$ is $a{e}^{-(a+b)t}$, because neither event has happened earlier. Its integral isWith the stated invented rate constants, $s=0.1,1,10\mu \mathrm{M}$ gives $1/11,1/2,10/11$, respectively. Raising free substrate concentration makes binding more likely to win this initial competition. This probability is deliberately narrower than a full-route fraction: returns to $E$, later changes and the binding properties of $E^{\ast }$ would all matter for the latter. The illustration explains why timing and concentration must accompany structural evidence, without assigning a binding mechanism to a real enzyme.

A motivating experiment tested the crystal interpretation in solution. McDonald, Steitz and Engelman prepared solutions of monomeric yeast hexokinase B and compared the unbound preparation with sugar-bound preparations. Their apparatus measured small-angle scattering of X-rays; the spatial extent inferred from that readout was the radius of gyration, rather than a catalytic reaction rate. Binding glucose reduced the radius of gyration by $0.95\pm 0.24\mathring{\mathrm{A}}$; binding the phosphorylated glucose product reduced it by $1.25\pm 0.28\mathring{\mathrm{A}}$, as reported by the authors. The glucose change agreed with the reduction calculated from the crystal coordinates. See Robert C. McDonald, Thomas A. Steitz and Donald M. Engelman, Yeast hexokinase in solution exhibits a large conformational change upon binding glucose or glucose 6-phosphate, Biochemistry 18 (1979), 338–342: primary abstract.

The comparison without added substrate supplies a baseline within the same form of hexokinase; the second bound preparation tests another binding condition. Together with the crystal comparison, solution compaction supports closure beyond crystal packing. It still gives an average spatial readout, not a movie of individual binding events, and product-associated compaction does not demonstrate ongoing catalysis. Neither experiment alone proves that binding precedes movement. To distinguish induced fit from conformational selection, structural comparisons must be combined with measurements sensitive to timing and with a model of the relevant populations and enzyme kinetics.

A proteinogenic amino acid represented by K or Lys in protein primary structure. Its amino acid side chain carries predominantly positive electric charge near neutral pH.^[ref]

Trypsin preferentially cleaves after its amino acid residues.

A paratope is the recognizing surface on an antibody; the epitope is the recognized feature on its antigen. Contacts depend on spatial arrangement and chemistry, including hydrogen bonds and Van der Waals forces. Thus a paratope is not simply a slot for a protein primary structure string. A folded protein can bring distant amino acid residues into one epitope. ^[ref]

The three resistant substitutions map near one another in the folded actin monomer, near the protein binding site for the nucleotide. This cluster identifies a candidate region involved in drug recognition or in the conformational response to binding. Neighbouring residues in three-dimensional structure can help form a pocket even when they are not adjacent in the protein primary structure. Reduced drug binding at this region would connect the structural map to the resistant phenotype.

The map does not show a bound ToxB molecule. Substitutions could act indirectly by changing folding, nucleotide exchange or the assembly-competent conformation. Proximity to ATP does not prove competition for the ATP-binding site, and the mutant's abnormal baseline actin polymerization makes an allosteric explanation particularly relevant. Direct binding measurements on comparably folded wild-type and mutant actin monomers, together with nucleotide-binding controls and a ligand-bound structure, would distinguish these models. The allele/residue labels are interpreted as defined in this paper; no identification of their original experimental sequence substitutions is required.

A stable native conformation is favoured when its Gibbs free energy, including the protein and its surrounding solvent, is lower than that of competing conformational ensembles under the specified conditions:Folding greatly reduces the chain's conformational entropy, an unfavourable contribution. It can nevertheless gain favourable hydrogen-bonding and packing interactions, van der Waals contacts, screened electrostatic interactions and solvent free energy. In aqueous globular proteins, burying nonpolar surface reduces the cost of exposing it to water; the hydrophobic effect is not an extra covalent attraction between all hydrophobic residues. Hydrogen bonds and ionic contacts must be judged relative to the interactions those groups made with solvent before folding, so merely counting bonds in the folded structure does not determine stability.

The protein primary structure determines which contacts and protein secondary structures can cooperate. A free-energy landscape biased towards compatible contacts permits many routes towards the native ensemble, rather than an exhaustive random search through every conformation. Folding funnels describe this bias, while barriers explain metastability and folding rates. Reversible denaturation/refolding experiments show that some proteins can recover native structure and activity when suitable conditions are restored, supporting a sequence-encoded thermodynamic tendency without asserting that every protein folds unaided.

“Stable” means an equilibrium preference and a useful lifetime, not one perfectly rigid shape. Native proteins fluctuate and can have several functional conformations; the unfolded ensemble remains populated with relative weight depending on its full free energy. Changes of temperature, pH, salt, solvent, ligands or pressure alter the balance, and molecular chaperones can prevent protein aggregation or assist kinetic access without supplying a different primary sequence. Membrane proteins require their appropriate environment, and intrinsically disordered proteins need not possess one stable folded state in isolation.

Thus a released stretched chain can gain configurational entropy on first recoiling yet later favour a more ordered native structure because the total free energy balance, including solvent, is different. Equilibrium stability, barriers to unfolding and resistance to irreversible protein aggregation are distinct properties. The existence of a native minimum alone does not guarantee rapid, reversible folding under every experimental condition.

Protein primary structure is the ordered amino acid sequence of a polypeptide, linked by covalent peptide bonds. The approximately planar peptide bond restricts local geometry, while backbone torsion angles give conformational freedom subject to steric exclusion. Protein secondary structure comprises local backbone arrangements, especially the alpha helix, with backbone hydrogen bonds between residues $i$ and $i+4$, and the beta sheet, whose strands are joined by backbone hydrogen bonds. Turns and loops connect these elements. Protein tertiary structure is the full three-dimensional organization of a single chain, often including several protein domains. Protein quaternary structure is the arrangement of multiple chains in a protein complex.

Protein folding is governed by the Gibbs free energy of the chain together with its solvent, not by the chain's energy alone. Folding reduces the chain's conformational entropy, but can gain solvent entropy through the hydrophobic effect, burying nonpolar side chains away from water. Favorable hydrogen bonds, Van der Waals forces, electrostatic interactions and sometimes covalent disulfide bonds stabilize particular structures. Steric repulsion prevents atomic overlap. At fixed temperature and pressure, the native ensemble is favored when its total $\Delta G=\Delta H-T\Delta S$ is lower than competing ensembles. The balance depends on the solvent, pH, salt and ligands. Molecular chaperones help avoid aggregation or kinetic traps; a stable fold is not a perfectly rigid conformation, and some proteins have disordered regions.

For a soluble globular protein, a typical pattern is a nonpolar core and a polar, hydrated surface. A transmembrane protein instead presents nonpolar side chains to the hydrocarbon interior of the lipid bilayer. Membrane-spanning alpha helices satisfy their polar backbone groups through internal hydrogen bonds; a roughly membrane-thickness helix commonly contains about twenty residues, depending on tilt and bilayer thickness. Multiple helices can enclose polar channels or binding sites. Membrane-spanning beta barrels likewise expose mainly nonpolar groups outward towards lipid while arranging backbone hydrogen bonds between strands and often lining an aqueous pore with polar groups. Exposed loops and domains contacting water can resemble soluble proteins. Not every membrane-associated protein spans the membrane: peripheral attachment and lipid anchoring are distinct possibilities.

The organizing distinction is the environment of each surface: water favors accessible polar groups, while the membrane core favors outward-facing nonpolar groups. Buried polar groups can still be essential to a transport pathway or catalytic site.

The protein primary structure is the ordered protein sequence along a polypeptide, from its N-terminus to its C-terminus. In cells, information in a DNA coding sequence is copied into messenger RNA by transcription. During translation, the ribosome reads successive three-base codons; transfer RNA molecules match these by their anticodons and supply the corresponding amino acids. The genetic code is degenerate: different codons can encode the same amino acid. A start establishes the reading frame and a stop terminates translation. Peptide bonds link the amino acids into the backbone, leaving chemically different side chains. Thus the sequence specifies the order of side-chain interactions, rather than a literal coordinate list for the folded molecule.

Protein secondary structure describes local backbone arrangements. An alpha helix is stabilized by a regular pattern of backbone hydrogen bonds, typically between the carbonyl of residue $i$ and the amide of residue $i+4$. Beta sheets assemble extended strands through inter-strand hydrogen bonding, with parallel or antiparallel strand orientations. Turns and loops connect these elements; not every region belongs to a helix or sheet. The peptide bond is approximately planar, while backbone torsion angles provide conformational freedom subject to steric restrictions.

Protein tertiary structure is the complete three-dimensional organization of one chain, often arranged into protein domains. Protein quaternary structure describes the organization of multiple chains in an assembly. Soluble globular proteins often bury nonpolar side chains and expose polar ones; membrane proteins can instead expose hydrophobic surfaces to the surrounding lipid. Other stabilizing interactions include hydrogen bonds, electrostatic interactions, Van der Waals forces and sometimes covalent disulfide bonds. These competing contributions define active sites, binding pockets and mechanically useful structures; folded proteins are dynamic ensembles, not perfectly rigid objects.

The connection from sequence to structure is thermodynamic and kinetic. Folding usually decreases the chain's conformational entropy, but favorable interactions and changes in solvent entropy, including the hydrophobic effect, can lower the total Gibbs free energy. Under suitable conditions the native ensemble is favored over unfolded alternatives. A change in sequence can therefore change the balance of accessible conformations and alter folding or function. Reversible refolding of a denatured protein in an appropriate environment demonstrates that the sequence can contain the information needed for its native structure.^[ref]

The environment is part of the specification: temperature, pH, ionic conditions, ligands and post-translational modifications matter. Molecular chaperones can prevent aggregation and help chains escape kinetic traps without providing a residue-by-residue structural template. Some sequences are intrinsically disordered proteins or disordered regions, and some require partners for a stable fold. Thus primary structure strongly constrains structure, but does not guarantee one unique rigid shape independent of conditions.

A phosphotyrosine residue is a tyrosine residue modified by phosphorylation. Its location and surrounding protein primary structure help determine which SH2 domains can recognize it.