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Absorbance is the dimensionless quantity $A={\log }_{10}(P_0/P)$, where $P_0$ and $P$ are reference and transmitted radiant powers at the chosen wavelength. For example, $A=1$ means one tenth of the reference power is transmitted. Under the Beer-Lambert law, absorbance can determine an absorbing species' concentration; attenuation by cloudiness or unrelated colored solutes needs separate controls. ^[ref]

The acid dissociation constant quantifies the chemical equilibrium for loss of one proton from a chemical acid. In water, write the actual proton transfer as $\mathrm{HA}+{\mathrm{H}}_2\mathrm{O}\rightleftharpoons {\mathrm{H}}_3{\mathrm{O}}^{+}+{\mathrm{A}}^{-}$. Its full dimensionless equilibrium constant isThe usual aqueous acid dissociation constant uses the solvent convention with water thermodynamic activity approximated by one:Here hydrated hydrogen ions are understood; ${\mathrm{H}}^{+}$ is shorthand, not an isolated bare proton in the chemical solution. All thermodynamic activities and $K_a$ are dimensionless quantities, using consistent standard states. If water thermodynamic activity differs appreciably from one, retain it in the full equilibrium constant instead of silently mixing conventions.

With activity coefficients ${\gamma }_i$ and reference concentration $c^{\circ }=1\mathrm{mol}{\mathrm{L}}^{-1}$, put $a_i={\gamma }_i[i]/c^{\circ }$ for each solute. ThenThe last expression is the dilute ideal concentration approximation, not the definition in a nonideal chemical solution. Omitting $c^{\circ }$ instead produces a quotient with units of concentration; its numerical value depends on the chosen units. Do not take a logarithm of that dimensional quotient. Conditional concentration quotients can change with ionic strength even at fixed temperature, whereas the thermodynamic acid dissociation constant retains the activity coefficient corrections. The acid dissociation constant entry distinguishes these conventions.

At the same temperature, solvent and standard states, a larger acid dissociation constant, equivalently a smaller pKa, indicates a stronger tendency to donate a proton. It does not specify an analytical concentration, a pH, or a reaction rate. Composition and the weak acid mass and charge balances are needed to predict pH; chemical kinetics addresses how quickly the chemical equilibrium is approached.

Active transport moves a substance uphill in its electrochemical gradient by coupling the transfer to a downhill process. Some carrier proteins use ATP hydrolysis; others couple uphill transfer of one solute to downhill transfer of another solute, often an ion. The latter mechanism uses a stored electrochemical gradient and need not consume ATP directly at that carrier protein. Maintaining the driving electrochemical gradient still requires a source of energy. See Alberts et al.: carrier proteins and active membrane transport.

For the neutral 8:2 example, B-to-A transfer costs $RT\ln 4\simeq 1.38629RT$ per mole. As an explicitly assumed bookkeeping example, suppose a tightly coupled cycle supplies a downhill free energy change of $-2RT$ per mole of transferred solute. The total is then $\Delta G_{\mathrm{cycle}}=RT\ln 4-2RT\simeq -0.61371RT$. This makes forward cycling energetically possible under those assumptions; it specifies neither a molecular mechanism nor a rate. Merely adding a carrier protein cannot supply the missing free energy. Conversely, as the transported concentration ratio rises, the required uphill cost rises, so a fixed driving process has a finite thermodynamic limit.

Adsorption is accumulation of a substance at an interface. Binding a solute to a packing surface can delay its passage through a chromatography column. Ideal size-exclusion chromatography separates through SEC pore access without such binding; extra retention from adsorption requires a different interpretation.

For a gas contacting a solid, adsorption concentrates molecules at the boundary; absorption takes material into the bulk. A porous solid can have a large internal surface area: binding on pore walls is still adsorption, even though those walls lie inside the particle. Both processes can occur together. Desorption returns adsorbed material to the surrounding phase. The useful question is therefore not just how much material disappeared from the gas, but where it went and whether it returns when the pressure falls.

An adsorption site model turns this spatial distinction into counting. With twelve identical adsorption sites and three bound molecules, the surface coverage is $\theta =3/12=1/4$. Filling six gives $\theta =1/2$; filling twelve gives a complete monolayer. These are configurations illustrating the count, rather than measured arrangements or claims that all real surfaces have twelve sites.

An actual teaching experiment prepares a Kool-Aid stock by weighing powder, transferring it quantitatively into a volumetric flask, and filling to the mark with water. A Genesys 30 spectrophotometer, blanked with water in a cuvette, scans for the peak absorbance wavelength. A Genesys 10 then measures absorbance at that wavelength, with a water reference blank. Each serial dilution transfers $50\mathrm{mL}$ of the preceding chemical solution to a $100\mathrm{mL}$ volumetric flask and fills to the mark; the protocol records the blank and sample readouts and repeats the halving sequence. These are prescribed experimental steps, not a published numerical data set.^[ref]

The experiment tests whether absorbance tracks the prepared concentration. Its concentration is powder mass per final volume, so the calibration does not by itself supply a single dye's molar absorption coefficient. The manual identifies loss of discrimination for very concentrated, very dilute, or very similar samples.^[ref]

To interpret such a calibration, hold cuvette orientation fixed with its clear faces in the light path, keep fingerprints and bubbles out of that path, and keep the geometric length unchanged. Mix each dilution completely. Re-read the reference blank during the sequence and re-read one standard to test drift. A separate preparation of a standard checks errors propagated through serial dilution; repeating only its instrumental reading does not test the transferred volumes. These controls follow from keeping $P_0$, $l$, and $c$ comparable between readings.

Three mechanisms explain why a straight line can fail. First, scattering of light can divert radiant power away from the photodetector, so cloudiness can raise apparent absorbance without the intended absorbing solute. A reference blank removes only losses shared with the sample. Second, stray light reaching the photodetector can make a strongly absorbing sample appear too transmissive. In an original model with extra detected radiant power $s{P}_0$ in both readings,As the ideal transmittance approaches zero, the reported absorbance approaches ${\log }_{10}((1+s)/s)$ for $s>0$. This finite ceiling is a model prediction, not measured instrument performance.

Third, chemical equilibrium can redistribute the absorbing chemical species when dilution, pH, or temperature changes. Even if each chemical species separately follows the Beer-Lambert law, their combined absorbance is $A=l{\sum }_i{\epsilon }_i{c}_i$, where $c_i$ is each species' molar concentration; changing those fractions need not preserve one effective molar absorption coefficient. Thus an unknown should be compared with standards under matching chemical solution conditions, within the tested concentration range. A good linear regression alone does not establish chemical selectivity.

Buffer exchange changes the small components surrounding a retained macromolecule toward those of a desired buffer solution. Components absent from fresh dialysate tend to leave; supplied components can enter. Each permeant solute has its own exchange problem.^[ref]

For a dilute, nonbinding permeant solute, an ideal equilibrium calculation approaches the bath's specified concentration. That approximation need not describe a solute bound to a protein or a charged mixture with additional electrical constraints. Retaining a protein also does not by itself establish that its protein structure or catalytic function survives the change of buffer solution.

A membrane carrier protein binds a solute and changes its exposed binding site from one side of a cell membrane to the other. Depending on its mechanism and coupling, it can mediate passive transport or active transport.

The cell membrane, also called the plasma membrane, is the selective boundary separating a biological cell's cytoplasm from its surroundings. Both prokaryotes and eukaryotes have a cell membrane. Where a cell wall is present, it is a distinct structure outside the cell membrane; the two names are not synonyms. This separation lets a biological cell maintain internal conditions while exchanging material through membrane transport. See NHGRI: cell membrane.

The familiar architecture is a lipid bilayer containing membrane proteins. In a phospholipid-based lipid bilayer, two membrane leaflets face opposite ways: hydrophilic heads face the water on each side, while hydrophobic tails face one another in the interior. The heads are therefore exposed both outside and inside the biological cell. This arrangement combines a water-facing surface with a barrier to many water-soluble solutes; the schematic represents organization rather than an atomistic structure. See Alberts et al., Molecular Biology of the Cell: the lipid bilayer.

Selective permeability means that different substances cross at different rates under comparable conditions. Small nonpolar molecules can pass through a lipid bilayer by diffusion, while ions encounter a strong barrier without a suitable route. An ion channel supplies a selective water-filled pathway when open; a carrier protein binds a solute and changes shape to expose it to the other side. These membrane proteins therefore add specific routes to the barrier. An open ion channel permits passive transport, whereas a carrier protein can mediate either passive transport or active transport. See Alberts et al.: principles of membrane transport.

The useful combination is retention plus controlled exchange: a completely impermeable boundary would prevent uptake, whereas unrestricted mixing would erase differences in concentration and membrane potential. The fluid mosaic model connects these functions to an organized but mobile lipid bilayer containing proteins.

A cell wall is a supporting structure outside a cell membrane. It helps preserve shape and bears the load when osmosis raises internal pressure. The cell membrane supplies the selective permeability described in its own article; the cell wall supplies mechanical reinforcement. These functions cooperate: a porous supporting layer can restrain expansion without being the boundary that retains dissolved solutes. Turgor pressure makes this cooperation visible in a firm leaf; plasmolysis and a lysozyme wall-removal experiment separate the two functions experimentally.^[ref]

The material is not universal. Most bacteria use peptidoglycan, whose linked sugar chains and amino acid cross-links are explained below. A plant cell wall uses cellulose; a fungal cell wall contains chitin and glucans. Animals lack a cell wall, some bacteria are naturally wall-free, and archaea have different wall chemistry. Thus a cell wall is neither a universal cellular component nor one chemical recipe.^{[ref][ref][ref]}

The bacterial outer membrane in typical Gram-negative bacteria is an additional membrane outside the peptidoglycan, not another name for that supporting mesh. The cell envelope includes the boundary layers together. An extra barrier can change access to the peptidoglycan, so response to lysozyme cannot be inferred from the mere presence of a cell wall.^[ref]

The chromatographic mobile phase carries solutes through the apparatus. In liquid chromatography it is a liquid solvent or chemical solution.

The eluent is the supplied chromatographic mobile phase. Its composition can alter both solute solubility and competition for adsorption sites, so changing the eluent can change retention even with the same chromatographic stationary phase.

For a dilute solute distributed between two liquid regions at local chemical equilibrium, define the chromatographic partition coefficient asHere $c_m,c_s$ are concentrations expressed in the same units, $V_m,V_s$ specify accessible volume in the chromatographic mobile phase and immobilized liquid chromatographic stationary phase, respectively, $n$ is total solute amount of substance, and $f_m$ is the fraction of that amount of substance in the chromatographic mobile phase. Assume a constant chromatographic partition coefficient, rapid exchange, unchanged phases and temperature, and no chemical reaction. A chemical equilibrium population still exchanges molecules; it is not permanently divided into two sets.

The amount of substance balance proves the expression for $f_m$. Increasing either $K$ or $V_s/V_m$ reduces the mobile fraction, illustrating why a concentration ratio alone cannot determine retention. For surface adsorption, specify an adsorption storage law instead of assigning an arbitrary liquid volume to the solid.

An original discrete model makes the separation visible. In each identical cell, first establish local chemical equilibrium, then advance only the mobile fraction by one cell; let the stationary fraction wait. Repeat twelve times, without any additional spreading. With $V_s=V_m$, stipulated chromatographic partition coefficients $K_A=1/3$ and $K_B=3$ give $f_A=3/4$ and $f_B=1/4$. Each cycle contributes mean displacement $f_m$ cells, so the mean positions after twelve cycles are nine cells and three cells. Redistribution itself broadens each band: the cell positions have a binomial distribution with parameters $12,f_m$. These dimensionless chosen values illustrate transport, not measurements of any particular substances.

The chromatographic stationary phase remains in place during separation. A porous packing in size-exclusion chromatography supplies internal liquid regions with size-dependent accessibility; ideal separation requires no adsorption of the solute to the packing.

A chromatographic stationary phase may instead expose a solid surface for adsorption, or retain an immobilized liquid into which solutes partition. In ordinary silica gel thin-layer chromatography, interactions with the surface delay migration. This mechanism differs from ideal SEC pore access.

Chromatography separates components of a mixture through different migration rates in contact with a chromatographic stationary phase while a chromatographic mobile phase transports them. Liquid chromatography describes the physical state of the transporting phase; size-exclusion chromatography describes a separation mechanism. Classification by phase and classification by mechanism answer different questions. In ideal size-exclusion chromatography, differential access to liquid-filled pores replaces binding as the source of differential migration.

In many other forms of chromatography, a solute repeatedly leaves the chromatographic mobile phase and returns to it. Reversible adsorption holds it at a surface; partition distributes it between the chromatographic mobile phase and an immobilized liquid chromatographic stationary phase. Spending more time stationary means less progress with the moving fluid. Two initially overlapping components can therefore emerge as separate bands without either being permanently trapped. Chromatography is useful both for comparing a sample with reference substances and for collecting separated components; a visible band need not contain just one substance.

Relative retention belongs to the sample together with the chosen chromatographic stationary phase, chromatographic mobile phase, eluent composition, and temperature. Changing these conditions can change the separation order. Chemical polarity helps explain interactions in some systems but is not a universal ranking rule. In thin-layer chromatography, a spot's TLC retardation factor is consequently conditional evidence about identity, rather than a fixed identifier intrinsic to a molecule.

The external chemical solution on the other side of a dialysis semipermeable membrane. Its composition supplies the boundary condition for exchange: it can receive unwanted small solutes or supply desired small solutes. A bath replacement discards the old dialysate and introduces a new reservoir with a specified composition.

Dialysis exchanges small solutes between a sample and an external bath through a semipermeable membrane, while retaining suitably large macromolecules, such as proteins. The transport is passive diffusion: random motion carries permeant molecules both ways, with net transport down their concentration gradient. The bath is the dialysate. At equal permeant solute concentrations, microscopic exchange continues but net exchange stops.^[ref]

For example, a protein sample containing excess sodium chloride can lose the small ions to a low-salt dialysate without deliberately transferring the protein. Conversely, components of a new buffer solution can enter from the bath. This is useful before studying enzyme kinetics: changing the surrounding chemical solution can remove an interfering small solute while keeping the enzyme available for the next experiment.

The striking feature is that the retained protein does not have to be captured and released again to change its surroundings. However, retention is conditional on the semipermeable membrane and the particular macromolecule, and dialysis is poorly suited to separating proteins of nearly the same size. The molecular weight cut-off is a nominal retention rating, not a sharp geometrical boundary.

The manufacturer-issued Protein Preparation Handbook reports a paired dialysis comparison: $2\mathrm{mL}$ of bovine serum albumin, $0.25mg/mL$ in $1\mathrm{M}$ sodium chloride, was held behind a $10\mathrm{kDa}$ MWCO membrane against $45\mathrm{mL}$ of water in a conical tube, shaken at room temperature. The bath was replaced after two hours. A conventional stirred-beaker treatment against $2\mathrm{L}$ of water supplied the comparison control. Residual salt was followed by sample electrical conductivity; results averaged two samples. This readout tests small-ion removal, not protein retention.^[ref]

Its separate retention experiment used individual proteins at $1mg/mL$ in saline or carbonate-bicarbonate buffer solution, pH $9.4$, with membranes rated $2$, $3.5$, $7$, $10$ and $20\mathrm{kDa}$. After seventeen hours at $4{}^{\circ }\mathrm{C}$, retained protein was estimated by a bicinchoninic acid assay. The size series and membrane series compare retention; they do not isolate shape independently of mass. The figure caption supplies neither a no-membrane recovery control nor a complete uncertainty analysis. These are manufacturer comparisons, not proof of universal recovery or unchanged enzyme function.^[ref]

The two experiments answer complementary questions: did the small solute leave, and did the desired macromolecule remain? In a new experiment, a bath blank and a starting salt standard would help interpret electrical conductivity, while a matched undialyzed protein aliquot and recovered volume would help distinguish handling loss from dilution. Those are proposed additional controls, not controls claimed for the published figures. Other ions also contribute to electrical conductivity, so the readout cannot generally identify sodium chloride alone in an unfamiliar buffer solution.

Diffusion is net transport associated with random microscopic motion. A localized patch of a dilute solute can spread through a stationary solvent: individual molecules move in both directions, but more cross from the crowded side of an imaginary surface than from the less crowded side. This imbalance gives a net diffusive flux down the concentration gradient; the molecules do not need to sense which direction has lower concentration. At uniform concentration, microscopic exchanges continue even though the net diffusive flux vanishes. ^[ref]

The distinction from advection is easiest to see in a dye patch. Uniform bulk motion carries its centre downstream; diffusion broadens it relative to that centre. Both can happen together. A moving or spreading patch by itself does not identify the mechanism: a nonuniform velocity field can also stretch a patch. ^[ref]

For a dilute nonreacting solute in a uniform medium at fixed temperature, Fick's first law models the relative diffusive flux as $\mathbf{j}=-D\nabla C$, with scalar diffusion coefficient $D>0$ of units ${\mathrm{m}}^2{\mathrm{s}}^{-1}$. Here $C$ is the amount of substance per unit volume, so $\mathbf{j}$ has units $\mathrm{mol}{\mathrm{m}}^{-2}{\mathrm{s}}^{-1}$. With carrier velocity field $\mathbf{u}$, the total flux is $C\mathbf{u}+\mathbf{j}$. The continuity equation, with no sources or chemical reactions, givesFor constant $D$ and incompressible flow, this reduces to $C_t+\mathbf{u}\cdot \nabla C=D{\nabla }^{2}C$, where ${\nabla }^2$ is the Laplacian. In a stationary carrier it is the diffusion equation. These assumptions matter: variable $D$ must remain inside the divergence, and changing sources require extra terms. ^[ref]

The useful surprise is the common mathematics: thermal conduction smooths temperature through the analogous heat equation, while molecular diffusion smooths composition. This analogy already motivated the original formulation of Fick's laws. ^[ref] However, the thermal diffusivity and the solutal diffusivity need not be equal. A diffusive time scale $t_D\sim L^2/D$ means that crossing ten times the distance requires roughly one hundred times the time. This is why molecular diffusion is especially effective across thin layers, and why chaotic advection can assist mixing by making those layers thinner. Physical diffusion is a transport mechanism, distinct from the broader probabilistic class of Markov diffusion.

Dilution lowers a solute's concentration by adding solvent. For conserved amount of substance, molar concentration satisfiesBoth $V_1$ and $V_2$ are chemical solution volumes; $V_2$ is the final total, not the added solvent.^[ref]

The proof is simply the definition $n=cV$ applied before and after. Its useful assumptions are full transfer of the selected portion, no loss or gain of the counted solute, and complete mixing. An ideal chemical solution approximation is not required for this inventory equation. If chemical equilibrium redistributes dissolved chemical species, use a conserved analytical concentration or chemical element inventory rather than assuming each individual species is conserved. This conservation statement does not by itself preserve the pH: a chemical equilibrium may change as the chemical solution is diluted.

The molar electrochemical potential combines the molar chemical potential ${\mu }_{\mathrm{m}}$ with the electrical contribution $z{F}_{\mathrm{Far}}\psi$, where $z$ is the signed charge number of an ion, $F_{\mathrm{Far}}=N_{\mathrm{A}}e$ is the Faraday constant and $\psi$ is electric potential. Both terms have units $\mathrm{J}\mathrm{mo}{\mathrm{l}}^{-1}$. In the particle convention the corresponding expression is ${\tilde{\mu }}_{\mathrm{particle}}={\mu }_{\mathrm{particle}}+ze\psi$, with elementary charge $e$; multiplying it by the Avogadro constant gives the molar form. Transfer between locations at the same temperature and pressure is driven by differences in electrochemical potential. The membrane transport examples use it to distinguish neutral solute transfer from ion transfer.

The enantiomeric excess of an enantiomer pair is the magnitude of the difference between their relative mole fractions, $\mathrm{ee}=|x_{+}-x_{-}|$, often reported as a percentage. Here $x_{+}+x_{-}=1$ counts the pair only, excluding solvent; the signs label optical rotation under the specified conditions, not R and S configuration.

In the model above, $x_{+}=0.75$ and $x_{-}=0.25$ give $\mathrm{ee}=0.50=50\%$. Keep the same total pair concentration $c=0.200\mathrm{g}\mathrm{m}{\mathrm{L}}^{-1}$, path length, solvent, temperature and wavelength. The two contributions areThe $+8^{\circ }$ pure-member reference therefore gives $\alpha /{\alpha }_{\mathrm{pure},+}=+0.50$. This ratio measures signed population imbalance only for a pure pair with additive linear responses under matching conditions. Adding unrelated solutes, changing total concentration, or changing chemical equilibrium can invalidate that inference. For $x_{+}=x_{-}=0.50$, the racemate gives zero net optical rotation by cancellation; each molecule remains chiral. A zero reading alone proves neither a racemate nor molecular achirality.