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Chromatography

Chromatography is a biophysical technique that enables the separation, identification, and purification of the components of a mixture for qualitative and quantitative analysis.^[1] The separation is achieved based on the differential partitioning of the analytes between a mobile phase, which flows through the system, and a stationary phase, with components interacting differently due to factors such as polarity, size, charge, or affinity.^[2] The technique was first developed in 1903 by Russian botanist Mikhail Tswett, who applied adsorption chromatography using columns packed with calcium carbonate to separate colorful plant pigments like chlorophyll and carotenoids, thereby coining the term "chromatography" from the Greek words for "color" and "to write" in his 1906 publication.^[2] Although initially overlooked, the method gained prominence in the 1930s and 1940s through advancements in partition chromatography by chemists like Archer Martin and Richard Synge, who earned the 1952 Nobel Prize in Chemistry for their contributions.^[2] At its core, chromatography operates on the principle of differential partitioning between the mobile and stationary phases, involving repeated equilibria where the mobile phase transports the sample mixture over or through the stationary phase, causing components to elute at different rates based on their relative affinities.^[2] Types of chromatography are broadly categorized as planar (e.g., paper chromatography and thin-layer chromatography) or column-based (e.g., gas chromatography and liquid chromatography), with further subdivisions including adsorption, partition, ion-exchange, size-exclusion, and affinity chromatography, each suited to specific analyte properties and sample types.^[2]^[1] Chromatography is indispensable across multiple disciplines, including pharmaceuticals for assessing drug purity and monitoring therapeutic levels, clinical diagnostics for analyzing metabolites in biological fluids like newborn screening for metabolic disorders, and toxicology for identifying poisons or drugs in patient samples.^[2] In biochemistry, it facilitates protein purification, determination of molecular weights, and analysis of biomolecules such as amino acids, carbohydrates, lipids, nucleic acids, and steroids.^[1] Additional applications extend to environmental science for detecting pollutants, food industry for quality control of additives and contaminants, and forensics for substance identification in criminal investigations.^[2]

Fundamentals

Etymology and Pronunciation

The term "chromatography" was coined in 1906 by Russian botanist Mikhail S. Tswett (also spelled Tsvett) in his foundational publications on separating plant pigments, derived from the Greek words chroma (χρῶμα), meaning "color," and graphein (γράφειν), meaning "to write" or "to record."^[3]^[4] This etymology evokes the technique's origin in visually recording separations as distinct colored bands, as Tswett observed when passing chlorophyll extracts through a column of adsorbent material like calcium carbonate powder packed in glass tubes.^[3] In standard English pronunciation, "chromatography" is rendered in American English as /ˌkroʊməˈtɑːɡrəfi/ (approximated phonetically as kroh-muh-TAH-gruh-fee), with stress on the third syllable and a broad "a" sound in "tog."^[5] In British English, it is /ˌkrɒməˈtɒɡrəfi/ (krom-uh-TOG-ruh-fee), featuring a short "o" in the first syllable and a short "o" in "tog," also with primary stress on the third syllable.^[6] These variations align with broader differences in vowel quality between the dialects, while the word's five-syllable structure—chro-ma-tog-ra-phy—remains consistent.^[5]

Historical Development

The invention of chromatography is credited to Mikhail Tswett, a Russian botanist, who developed the technique between 1903 and 1906 to separate plant pigments such as chlorophylls and carotenoids.^[7] In 1903, Tswett presented his initial findings in a lecture in Warsaw, describing the use of a vertical glass column packed with calcium carbonate as the stationary phase and petroleum ether as the mobile phase to achieve adsorption-based separation, resulting in colored bands he termed "chromatogram."^[7] He formalized the method in two seminal 1906 papers published in Berichte der Deutschen Botanischen Gesellschaft, where he coined the term "chromatography" from the Greek words for "color" and "to write," emphasizing its application to colored substances while noting its potential for colorless compounds as well.^[7] Despite its promise, Tswett's work was largely overlooked until the 1930s, when it was rediscovered and expanded by biochemists Richard Kuhn and Edgar Lederer for separating biochemical compounds.^[8] In 1931, Kuhn's group in Heidelberg applied adsorption chromatography to isolate and characterize isomers of carotene from carrots and other natural sources, demonstrating its utility in resolving complex mixtures of vitamins and pigments that were previously inseparable.^[8] This revival marked chromatography's transition from botanical research to broader biochemical applications, with Lederer's contributions highlighting its effectiveness in handling small sample quantities and achieving high resolution for labile biomolecules.^[8] Post-World War II advancements propelled chromatography into a cornerstone analytical technique, beginning with the 1941 development of partition chromatography by Archer J.P. Martin and Richard L.M. Synge at the Wool Industries Research Association in Leeds. Their method utilized a liquid stationary phase coated on an inert solid support, with a liquid mobile phase creating partitions based on solute distribution coefficients, enabling efficient separation of amino acids from protein hydrolysates using silica gel columns and water-phenol systems. For this innovation, which dramatically improved speed and sensitivity over adsorption methods, Martin and Synge shared the 1952 Nobel Prize in Chemistry.^[9] Building on partition principles, Martin and Anthony T. James introduced gas chromatography in 1952 while at the National Institute for Medical Research in London.^[10] Their gas-liquid partition chromatography employed an inert gas as the mobile phase and a nonvolatile liquid coated on a solid support as the stationary phase, allowing rapid separation and micro-estimation of volatile fatty acids from formic to dodecanoic acid using a thermal conductivity detector.^[10] This technique extended chromatography to gaseous and volatile samples, revolutionizing analyses in organic and biochemical fields. In 1956, Egon Stahl at the University of Saarbrücken pioneered thin-layer chromatography (TLC) as a faster, more accessible alternative to paper chromatography. Stahl's method involved spreading a thin adsorbent layer, such as alumina or silica gel, on glass plates for separations driven by capillary action, enabling quick visualization and isolation of compounds from microgram quantities in under an hour. High-performance liquid chromatography (HPLC) emerged in the 1960s and 1970s, transforming liquid chromatography into a high-speed, high-resolution tool through advancements in pressure-resistant columns and fine particles. In 1967, Csaba Horváth and colleagues at Yale University demonstrated the first modern HPLC system using pellicular beads under high pressure to separate nucleotides rapidly, achieving resolutions unattainable with classical methods. By 1969, Joseph J. Kirkland at DuPont developed bonded-phase columns with octadecylsilane stationary phases, enabling reversed-phase separations under pressures up to 1000 psi and broadening HPLC's applicability to nonpolar compounds in pharmaceuticals and biochemistry. These innovations, commercialized in the 1970s, established HPLC as the dominant chromatographic technique for quantitative analysis.

Basic Terminology

In chromatography, the stationary phase refers to the immobile material, typically a solid, gel, or liquid supported on a solid, that interacts with sample components to facilitate separation.^[11] The mobile phase, in contrast, is the fluid—such as a liquid, gas, or supercritical fluid—that flows through or over the stationary phase, carrying the sample components along a defined path.^[11] The analyte denotes the specific component or components of interest within the sample being separated and analyzed.^[11] In elution-based methods, the mobile phase is often termed the eluent, emphasizing its role in displacing and transporting analytes from the stationary phase.^[11] Central to interpreting chromatographic results are several time-based parameters. The retention time (t_R) is the duration from sample injection to the maximum of the corresponding peak in the output, representing the total time an analyte spends in both phases.^[11] The void time (t_0), also known as hold-up time, is the time required for an unretained solute to traverse the system, equivalent to the transit time through the mobile phase alone.^[11] From these, the capacity factor (or retention factor, k) is derived as k = \frac{t_R - t_0}{t_0}, quantifying the relative affinity of the analyte for the stationary phase over the mobile phase.^[12] A chromatogram is the graphical record of the separation, plotting detector response (proportional to analyte concentration in the eluent) against time or volume, where distinct peaks indicate separated components. To assess separation quality, resolution (R_s) measures how well two adjacent peaks are distinguished, calculated as R_s = \frac{2(t_{R2} - t_{R1})}{w_1 + w_2}, where t_{R1} and t_{R2} are retention times of the earlier and later peaks, and w_1 and w_2 are their baseline widths; values greater than 1.5 typically ensure baseline separation.^[13]

Core Principles

Chromatographic Separation Process

The chromatographic separation process involves the differential migration of sample components through a system comprising a stationary phase and a mobile phase, enabling the isolation of mixtures based on their interactions with these phases.^[2] This technique is widely applied in analytical chemistry for qualitative and quantitative analysis, purification, and characterization of compounds across various fields such as pharmaceuticals and environmental monitoring.^[14] The process relies on repeated equilibrium partitioning of analytes between the mobile phase, which carries the sample, and the stationary phase, which selectively retains components based on their affinity differences.^[15] Components with stronger interactions with the stationary phase move slower, while those favoring the mobile phase elute faster, resulting in spatial or temporal separation.^[2] The workflow typically begins with sample preparation, where the analyte mixture is dissolved or suspended in a suitable mobile phase to ensure compatibility and minimize interferences; this may include extraction, filtration, or concentration steps to enhance resolution.^[2] Next, the prepared sample is introduced into the system, often via injection using a syringe or autosampler for precise volume control, typically in microliter quantities to avoid overloading the column.^[14] The sample then undergoes migration through the stationary phase as the mobile phase flows, driven by pressure or gravity, allowing analytes to partition repeatedly and separate based on their retention behaviors.^[15] Upon reaching the end of the separation medium, components are detected using instruments such as ultraviolet (UV) absorbance, mass spectrometry (MS), or fluorescence detectors, which generate signals proportional to concentration as peaks in a chromatogram.^[2] Finally, data analysis involves interpreting the chromatogram to identify analytes by retention time—the duration from injection to peak maximum—and quantify them via peak area or height integration, often using software for calibration against standards.^[14] Several operational factors qualitatively influence the separation efficiency and speed. Flow rate of the mobile phase affects the time for partitioning and peak broadening; optimal rates, such as 0.1–5 mL/min in liquid systems, balance resolution and analysis duration without excessive diffusion.^[2] Temperature modulates analyte volatility and interactions, generally increasing elution speed in gas-based systems while potentially altering selectivity in liquid ones.^[14] Pressure, applied up to several hundred bars in pressurized systems, ensures consistent flow through resistive media and enhances resolution for complex mixtures.^[2] Elution modes dictate how the mobile phase composition is managed during separation. In isocratic elution, the mobile phase remains constant throughout, simplifying operation but potentially leading to long retention for late-eluting peaks in diverse samples.^[2] Conversely, gradient elution involves systematically varying the mobile phase composition—such as increasing organic solvent strength in reversed-phase liquid chromatography—to maintain consistent retention factors, improving resolution and shortening overall run times for samples with wide polarity ranges.^[14]

Retention Mechanisms and Factors

Retention in chromatography refers to the process by which analytes are temporarily held by the stationary phase, delaying their migration through the system relative to the mobile phase. This differential retention is fundamental to separation and arises from specific physical and chemical interactions between the analyte, stationary phase, and mobile phase. The primary retention mechanisms include adsorption, partition, ion exchange, and size exclusion, each exploiting distinct molecular properties to achieve selectivity.^[1] Adsorption involves the adherence of analyte molecules to the surface of a solid stationary phase through physical forces such as van der Waals interactions or hydrogen bonding, or chemical bonds in some cases; this mechanism is prominent in techniques using polar surfaces like silica gel, where non-polar analytes exhibit weaker retention compared to polar ones.^[1] Partition chromatography relies on the solubility differences of analytes between a liquid stationary phase (often coated on a solid support) and the mobile phase, allowing analytes with higher affinity for the stationary phase to spend more time there and elute later; this was first theoretically described by Martin and Synge in their seminal 1941 work on liquid-liquid distribution systems. Ion exchange separation occurs via electrostatic attractions between charged analytes and oppositely charged functional groups on the stationary phase, enabling the exchange of ions based on charge affinity.^[1] Size exclusion, in contrast, is a steric mechanism where analytes are separated purely by molecular size, with larger molecules unable to enter the pores of the stationary phase and thus eluting faster, while smaller ones are retained longer within the porous matrix. Several factors influence the extent and selectivity of retention across these mechanisms. The polarity of the stationary and mobile phases determines the strength of interactions, with matching polarities enhancing retention for analytes of similar polarity; for instance, a polar stationary phase retains polar analytes more strongly in non-polar mobile phases.^[15] pH affects retention particularly in ion exchange and partition modes by altering the ionization state of analytes and the stationary phase, thereby changing charge-based interactions—acidic conditions may protonate analytes, reducing their retention on cation exchangers.^[16] Ionic strength modulates electrostatic forces in ion exchange chromatography, where higher salt concentrations shield charges and decrease retention by competing for binding sites.^[17] Temperature impacts retention by influencing the viscosity of the mobile phase, which decreases with rising temperature to allow faster flow and reduced retention times, and by altering equilibrium constants governing analyte-phase distributions, often following the van't Hoff relationship where higher temperatures weaken enthalpic interactions.^[18] The distribution coefficient, K_d, quantifies retention by representing the equilibrium ratio of analyte concentration in the stationary phase to that in the mobile phase (K_d = \frac{[\text{analyte}]_{\text{stationary}}}{[\text{analyte}]_{\text{mobile}}}); higher K_d values indicate stronger retention and longer elution times, serving as a foundational parameter for predicting separation behavior.^[2] Selectivity, denoted as \alpha = \frac{k_2}{k_1}, measures the differential retention between two analytes (where k is the retention factor for each), with values greater than 1 enabling effective separation; it arises from variations in K_d due to molecular differences and is crucial for resolving closely related compounds.^[19]

Theoretical Models

Theoretical models in chromatography provide mathematical frameworks to predict and optimize separation efficiency, primarily through the concepts of theoretical plates and rate theory, which account for band broadening and peak shapes.^[14] The plate theory, introduced by Martin and Synge, conceptualizes the chromatographic column as a series of discrete equilibrium stages or "theoretical plates," where each plate represents a local equilibration of solute between the mobile and stationary phases.^[14] The number of theoretical plates, N, quantifies column efficiency and is calculated for Gaussian peaks approximated as triangular using the formula N = 16 \left( \frac{t_R}{w} \right)^2, where t_R is the retention time and w is the baseline peak width. The height equivalent to a theoretical plate (HETP), denoted as H, measures efficiency per unit column length and is given by H = \frac{L}{N}, with L as the column length; lower H values indicate higher efficiency.^[14] In contrast, rate theory addresses the continuous nature of mass transfer and diffusion processes, offering a more dynamic view of band broadening. The seminal Van Deemter equation describes HETP as a function of linear mobile phase velocity u:

H = A + \frac{B}{u} + C u

where A represents eddy diffusion due to inhomogeneous packing, B accounts for longitudinal molecular diffusion, and C encompasses resistance to mass transfer between phases.^[14] This equation predicts an optimal velocity minimizing H, balancing diffusive spreading at low speeds against mass transfer limitations at high speeds.^[14] Band broadening in chromatography arises from multiple sources integrated in rate theory: eddy diffusion from variable flow paths around particles, longitudinal diffusion causing solute spread along the flow direction, and resistance to mass transfer due to finite equilibration rates across phases.^[14] These contributions degrade resolution by increasing peak width relative to retention differences. Peak asymmetry, particularly tailing where the trailing edge extends beyond the leading edge, results from non-uniform interactions or secondary equilibria, leading to distorted elution profiles that reduce efficiency and complicate quantification; the tailing factor qualitatively assesses this by comparing front and rear peak widths at 5% height, with values near 1 indicating symmetry and greater than 1 signaling tailing.^[14]

Techniques by Bed Shape

Column Chromatography

Column chromatography is a foundational separation technique where the mobile phase, typically a liquid, percolates through a stationary phase packed in a vertical tube, allowing components of a mixture to separate based on their differential interactions with the two phases. The setup involves a cylindrical column, often made of glass or sometimes metal for larger scales, filled with a granular stationary phase such as silica gel or alumina, which is slurried in a solvent and packed to ensure even distribution and minimal channeling. The flow is driven either by gravity in classical setups or by low external pressure in modern variants, with the column diameter and length adjusted according to the sample size and desired resolution—narrower columns for analytical purposes and wider ones for preparative work.^[20]^[21] In the sample loading and elution process, the mixture is first dissolved in a minimal volume of compatible solvent and applied uniformly to the top of the stationary phase bed, often after adding a layer of inert sand to prevent disturbance. Elution then proceeds by continuously adding the mobile phase solvent or solvent gradient from the top, causing the sample components to migrate downward at rates determined by their relative affinities for the stationary and mobile phases—more strongly adsorbed compounds elute later. Fractions of the eluate are collected sequentially at the column bottom, monitored via TLC or other methods, and combined based on purity, enabling the isolation of target compounds from complex mixtures. This process adheres to the general chromatographic separation outlined in core principles, where retention is governed by adsorption or partition mechanisms.^[22]^[23] In classical adsorption column chromatography, coarser silica gel (70-230 mesh) is commonly used as the stationary phase, with non-polar solvents like hexane serving as the initial mobile phase to elute less polar impurities first. It is particularly valued in organic synthesis and phytochemistry for purifying extracts from plant or microbial sources, where large sample loads can be processed without specialized equipment.^[20] A key advancement, flash chromatography, was introduced by Still, Kahn, and Mitra in 1978 as a faster alternative for preparative separations with moderate resolution, employing air or nitrogen pressure (typically 10-15 psi) to drive the mobile phase through finer silica gel (230-400 mesh) for improved flow rates and band sharpness. Unlike classical methods, flash setups often include a fritted disc at the bottom and solvent reservoirs connected to a pressure source, allowing separations in minutes rather than hours while maintaining scalability. This variant is widely adopted in laboratories for rapid purification of synthetic intermediates or natural products, using solvent systems optimized via prior TLC analysis.^[24]^[22]^[25] Column chromatography excels in scalability for preparative applications, handling sample sizes from milligrams to kilograms with relatively low cost and disposability of materials, making it ideal for isolating substantial quantities of natural products or reaction products. However, it suffers from lower efficiency compared to high-performance liquid chromatography (HPLC), as the larger particle sizes (typically 40-60 μm) and reliance on gravity or low pressure result in broader peaks, reduced resolution, and longer run times for complex mixtures. These limitations stem from the inherent low plate heights in low-pressure systems, restricting its use to scenarios where high throughput and simplicity outweigh the need for ultra-fine separations.^[26]^[27]

Planar Chromatography

Planar chromatography encompasses techniques where the stationary phase is applied as a thin layer on a flat support, such as paper or a coated plate, allowing separation to occur in a two-dimensional plane. The mobile phase, typically a liquid, migrates across the stationary phase primarily through capillary action, enabling the differential partitioning of analytes based on their affinities. This setup is particularly advantageous for qualitative analysis due to its simplicity, low cost, and ability to handle multiple samples simultaneously on a single plate.^[28]^[2] Paper chromatography, one of the earliest planar methods, utilizes filter paper as the stationary phase, where the hydrophilic cellulose fibers retain polar compounds while the mobile phase—a solvent or solvent mixture—advances by capillary forces. Samples are applied as spots near the base of the paper strip or sheet, which is then placed in a developing chamber; separation occurs as components migrate at rates dependent on their solubility and interaction with the paper. The retention factor, or Rf value, quantifies this migration and is calculated as the ratio of the distance traveled by the compound to the distance traveled by the solvent front, yielding values between 0 and 1 that aid in compound identification.^[2]^[28] Thin-layer chromatography (TLC), an advancement over paper methods, employs a thin, uniform layer of adsorbent material, such as silica gel, alumina, or cellulose, coated onto a rigid support like glass, plastic, or aluminum plates, typically 20 cm × 10 cm in size. The sample is spotted or streaked near the bottom edge, and development proceeds similarly via capillary action in a solvent chamber, often allowing for faster separations (5–15 minutes) due to the finer particle size (5–20 μm) of the stationary phase compared to paper. Visualization of separated bands, which may be colorless, is achieved through ultraviolet (UV) light absorption, chemical staining (e.g., iodine vapor or ninhydrin), or fluorescence indicators; quantitative analysis can be performed using densitometric scanning, where peak areas correlate with analyte concentrations via calibration curves. High-performance TLC (HPTLC) variants enhance resolution with finer sorbents (≤10 μm) and automated application, supporting limits of detection as low as 6–900 ng per band.^[28]^[29] In forensics, planar chromatography facilitates rapid screening of substances like drugs of abuse (e.g., cocaine and amphetamines in urine or tablets) and pesticides in poisoning cases, often coupled with techniques like surface-enhanced Raman spectroscopy for confirmation, achieving limits of detection around 0.2 mg/kg.^[30] In pharmaceuticals, it serves for quality control and purity assessment of active ingredients, such as simultaneous quantification of ofloxacin and ornidazole in tablets (recoveries of 99.61–100.23%) or fingerprinting herbal extracts like ginseng for adulteration detection, enabling efficient monitoring of stability and formulation integrity.^[29]^[28]

Techniques by Mobile Phase State

Gas Chromatography

Gas chromatography (GC) is an analytical separation technique that employs a gaseous mobile phase to separate and analyze volatile, thermally stable compounds based on their differential partitioning between the gas phase and a stationary liquid phase. The method is particularly suited for samples that can be vaporized without decomposition, such as hydrocarbons, pesticides, and essential oils. Separation occurs as the sample components travel through a column at different rates depending on their volatility and interaction with the stationary phase, allowing for high-resolution analysis of complex mixtures. The typical GC setup includes an injection port where the liquid or gaseous sample is introduced and vaporized, a temperature-controlled oven housing the separation column, and a detector at the column outlet. Columns are either packed with a solid support coated in liquid stationary phase or, more commonly, open-tubular capillary columns with an inner wall coating of the stationary phase for enhanced efficiency. An inert carrier gas, such as helium or nitrogen, serves as the mobile phase, flowing continuously through the system at a controlled pressure and linear velocity to transport the vaporized sample components. Helium is preferred for its optimal balance of safety, efficiency, and wide linear velocity range, while nitrogen offers a cost-effective alternative with slightly lower performance. The oven temperature is programmed to increase gradually, accelerating elution of higher-boiling-point compounds.^[31] In GC, separation relies on the vapor pressure of analytes and their equilibrium distribution between the inert carrier gas and the non-volatile liquid stationary phase immobilized on the column. Analytes with higher vapor pressure and lower affinity for the stationary phase elute faster, while those that partition more favorably into the liquid phase are retained longer, resulting in distinct retention times. This gas-liquid partitioning mechanism enables precise qualitative and quantitative analysis, often with resolution exceeding 10,000 theoretical plates in capillary columns. For identification, retention times are compared to standards, though coupling with mass spectrometry enhances specificity.^[32] Common detectors in GC include the flame ionization detector (FID), electron capture detector (ECD), and mass spectrometry (MS) in GC-MS configurations. The FID, a universal detector for organic compounds, ionizes carbon-containing molecules in a hydrogen-air flame, producing a current proportional to the analyte mass, with a detection limit around 5 pg of carbon per second and a linear range over six orders of magnitude. The ECD selectively detects electronegative compounds like halocarbons by measuring electron capture in a radioactive nickel-63 source, offering femtogram sensitivity but requiring careful handling due to radioactivity. GC-MS combines chromatographic separation with mass spectral identification, providing structural information via fragmentation patterns and enabling trace-level detection in environmental and forensic applications.^[33]^[34] GC offers significant advantages, including high separation efficiency, rapid analysis times (often under 30 minutes), and excellent sensitivity for volatile organics, making it indispensable in fields like petrochemicals, pharmaceuticals, and food safety. Its inert mobile phase minimizes interference, and automated systems allow high-throughput processing. However, limitations include restriction to thermally stable, volatile samples; non-volatile or polar compounds often require chemical derivatization to enhance volatility, adding complexity and potential artifacts. Additionally, carrier gas supply costs, particularly for helium, can be a practical constraint.^[35]^[33]

Liquid Chromatography

Liquid chromatography refers to a family of analytical techniques that employ a liquid mobile phase to separate components of a mixture based on their differential partitioning between the mobile and stationary phases. It excels in the separation of polar, ionic, and thermally sensitive analytes that cannot be volatilized for gas-phase analysis, making it indispensable in fields such as pharmaceuticals, environmental monitoring, and biochemistry. Unlike gaseous mobile phases, liquids provide greater solvating power for a broader range of compounds, enabling the analysis of non-volatile substances at ambient temperatures.^[2] The versatility of liquid chromatography stems from its two primary modes: normal-phase and reversed-phase, which differ in the relative polarities of their stationary and mobile phases. In normal-phase chromatography, a polar stationary phase—typically unmodified silica or alumina—is used with a non-polar mobile phase such as hexane or a hexane-ethyl acetate mixture. Analytes separate primarily through adsorption, where more polar compounds interact more strongly with the stationary phase's polar groups (e.g., silanol on silica), resulting in longer retention times; this mode is particularly effective for separating isomers and fat-soluble vitamins.^[36] Conversely, reversed-phase chromatography, the dominant mode accounting for over 80% of applications, features a non-polar stationary phase like octadecyl (C18) or octyl (C8)-bonded silica and a polar mobile phase, commonly water mixed with acetonitrile or methanol. Retention here is governed by hydrophobic interactions, with non-polar analytes eluting later than polar ones; it is widely used for drug assays, peptide separations, and environmental pollutant detection due to its robustness and compatibility with aqueous samples.^[37]^[2] High-performance liquid chromatography (HPLC), the pressurized evolution of classical liquid chromatography, achieves superior resolution and speed through the use of high-pressure systems and finely divided stationary phases. Pioneered in the 1960s, with Csaba Horváth constructing the first modern instrument in 1966 to separate amino acids under pressures up to 500 psi, HPLC marked a shift from low-pressure gravity-fed columns to engineered systems capable of handling backpressures of 50–600 bar.^[38]^[39] These pressures propel the mobile phase through narrow columns (typically 4.6 mm inner diameter and 150–250 mm length) packed with spherical particles of 3–10 μm diameter, which minimize eddy diffusion and maximize plate efficiency (often >10,000 theoretical plates per meter).^[40]^[41] Reciprocating pumps, such as binary or quaternary gradient models, maintain precise flow rates of 0.1–5 mL/min with low pulsation (<1%), ensuring reproducible elution profiles.^[36] Autosamplers enhance throughput by automating sample introduction via fixed-volume loops (1–100 μL injections), reducing manual error and enabling high-sample workloads in routine labs.^[37] Detection in HPLC relies on flow-through sensors positioned post-column to monitor eluate composition continuously. Ultraviolet-visible (UV-Vis) detectors, the most prevalent, quantify analytes by absorbance at fixed (e.g., 254 nm) or variable wavelengths (190–700 nm), ideal for aromatic or conjugated compounds like pharmaceuticals and proteins, though limited to UV-absorbing species.^[37] Refractive index (RI) detectors measure changes in light refraction caused by differing solute concentrations, providing universal detection for non-chromophoric analytes such as carbohydrates and alcohols, but they require isocratic conditions and are less sensitive (detection limits ~0.1–1 μg).^[2] Fluorescence detectors offer enhanced selectivity and sensitivity (down to picograms) by exciting analytes at 200–650 nm and measuring emission up to 900 nm, commonly applied to derivatized amino acids or native fluorophores like PAHs, though only suitable for fluorescent compounds.^[37] For resolving complex mixtures spanning wide polarity ranges, gradient elution adjusts the mobile phase composition dynamically during the run, often increasing the organic solvent fraction in reversed-phase (e.g., from 5% to 95% acetonitrile in water over 20–60 minutes) to compress retention times and improve peak spacing. This approach, first demonstrated in 1952 for inorganic separations on paper, extends general elution principles by preventing early peaks from eluting too quickly and late ones from broadening excessively, thus optimizing resolution for samples like protein digests or pesticide residues without excessive run times.^[42]^[36]^[2]

Supercritical Fluid Chromatography

Supercritical fluid chromatography (SFC) utilizes a supercritical fluid as the mobile phase to achieve separations that leverage properties intermediate between those of gases and liquids, enabling enhanced diffusivity and solvating power for efficient analyte transport through the stationary phase.^[43] The most commonly employed supercritical fluid is carbon dioxide (CO₂), which reaches its critical state above 31.1°C and 73.8 bar, where its density can be tuned by adjusting temperature and pressure to optimize solvating capabilities for a wide range of compounds.^[44] This tunability allows the mobile phase to bridge the limitations of gas and liquid chromatography, providing low viscosity for rapid mass transfer while maintaining sufficient density for solubility.^[45] The instrumental setup for SFC closely resembles that of high-performance liquid chromatography (HPLC), featuring binary pumps to deliver CO₂ and organic modifiers (such as methanol at 5-50% concentration), a column oven for temperature control (typically 40-60°C), and an automated back-pressure regulator to sustain supercritical conditions at 150-400 bar post-column.^[43] Columns are usually packed with silica-based particles (3-5 μm diameter, e.g., 4.6 × 150 mm dimensions) modified for normal-phase interactions, with flow rates of 1.8-5 mL/min to minimize pressure drops and ensure consistent performance.^[43] Additives like 0.1-2% ammonium acetate or trifluoroacetic acid may be included in the modifier to enhance peak shapes for polar analytes, particularly when interfacing with mass spectrometry.^[43] SFC offers distinct advantages, including separation speeds 3-5 times faster than HPLC due to the high diffusion coefficients of supercritical fluids (e.g., 16 × 10⁻⁵ cm²/s for benzoic acid in CO₂), which reduce analysis times to under 2-4 minutes for many samples.^[43] It is greener than traditional liquid chromatography, consuming up to 90% less solvent through the use of recyclable CO₂, which exhibits low toxicity and flammability, thereby lowering environmental impact and operational costs.^[46] Additionally, SFC excels in chiral separations with success rates of 95-98% and is particularly effective for lipid analysis, enabling high-resolution profiling of complex mixtures like triglycerides and phospholipids.^[43]^[47] Detection in SFC is versatile and compatible with techniques similar to those in gas and liquid chromatography, including ultraviolet (UV) absorbance with flow cells optimized for high pressure (e.g., 10-mm path length, noise <0.1 mAU), mass spectrometry (MS) for structural elucidation when using volatile additives, and flame ionization detection (FID) for nonpolar solutes with pure CO₂ mobile phases.^[43] Evaporative light scattering detection (ELSD) is also employed for compounds lacking chromophores, providing broad applicability across diverse analyte classes.^[43] The tunable properties of the supercritical fluid briefly tie to retention mechanisms by allowing density adjustments that modulate analyte-stationary phase interactions, akin to mobile phase variations in other techniques.^[45]

Techniques by Separation Mechanism

Affinity Chromatography

Affinity chromatography is a separation technique that exploits highly specific, reversible biospecific interactions between a target biomolecule and a complementary ligand immobilized on a stationary phase support, enabling the purification of proteins, enzymes, and other biomolecules from complex mixtures.^[48] Introduced in 1968 by Cuatrecasas, Wilchek, and Anfinsen, it revolutionized biomolecule isolation by achieving high purity in a single step through selective binding, often yielding recoveries exceeding 90% for targeted analytes.^[49] This method operates under mild conditions, preserving the biological activity of sensitive molecules like enzymes and antibodies.^[50] The core mechanism involves attaching a ligand—such as an antibody, enzyme substrate, or metal ion chelate—to an insoluble matrix like agarose or silica beads, which serves as the stationary phase.^[48] The target molecule, present in the mobile phase (typically an aqueous buffer), binds reversibly to the ligand via non-covalent interactions including hydrogen bonding, ionic forces, and hydrophobic effects, forming a stable complex while non-specific components pass through unbound.^[51] This selectivity stems from the ligand's affinity for a unique structural feature of the target, such as an active site or epitope, distinguishing it from other separation methods.^[52] The purification process consists of three main stages: loading, washing, and elution. During loading, the sample is applied to the column in a binding buffer optimized for pH and ionic strength to promote target-ligand association, often at neutral pH (6-8) for protein targets.^[53] Washing follows with the same or similar buffer to remove unbound contaminants, minimizing non-specific adsorption through additives like salts or detergents.^[48] Elution disrupts the interaction by introducing a competitor ligand, altering pH (e.g., to 2-3 for acid-labile bonds), or changing ionic strength, releasing the purified target in a concentrated form.^[54] Key applications include the purification of recombinant proteins using histidine (His)-tags, where a polyhistidine sequence binds to nickel-nitrilotriacetic acid (Ni-NTA) resins via coordination chemistry, allowing facile isolation from bacterial lysates with purities often >95%.^[55] Another prominent use is antibody isolation, leveraging ligands like Protein A or G that specifically bind the Fc region of immunoglobulins from serum or culture supernatants.^[56] These techniques are integral in biotechnology for producing therapeutic proteins and in research for studying biomolecular interactions. Variants of affinity chromatography expand its utility to diverse biomolecules. Immunoaffinity chromatography employs antibodies as ligands to capture antigens with exquisite specificity, commonly used for purifying cytokines or hormones from biological fluids.^[57] Lectin affinity chromatography utilizes carbohydrate-binding proteins (lectins) like concanavalin A to isolate glycoproteins by recognizing glycan moieties, facilitating the enrichment of membrane proteins or viral glycoproteins.^[58] These adaptations maintain the core principle of biospecific binding while tailoring to structural features like charge or glycosylation.

Ion-Exchange Chromatography

Ion-exchange chromatography is a separation technique that exploits differences in the net charge of molecules to achieve fractionation, primarily through reversible electrostatic interactions between charged analytes and an oppositely charged stationary phase composed of ion-exchange resins.^[16] The method is particularly effective for ionizable biomolecules such as proteins and nucleic acids, where the charge is modulated by the buffer pH relative to the molecule's isoelectric point (pI).^[59] There are two primary types of ion-exchange resins: cation-exchange and anion-exchange. Cation-exchange resins, which bear negatively charged functional groups such as sulfonic acid (e.g., SP or S types), bind positively charged analytes when the buffer pH is below the analyte's pI, attracting cations like protons or metal ions.^[16] Anion-exchange resins, featuring positively charged groups such as quaternary ammonium (e.g., Q type) or diethylaminoethyl (DEAE), interact with negatively charged species at pH values above the pI, facilitating the binding of anions.^[59] These resins are typically cross-linked polymers like polystyrene-divinylbenzene or agarose, with functional groups covalently attached to provide stable charge throughout a wide pH range (e.g., 2–14 for strong exchangers).^[16] The separation process begins with sample loading in a low-ionic-strength buffer at a pH selected to ensure analyte binding—typically 0.5–1 unit away from the pI—while unbound components pass through.^[16] Elution is achieved by gradually increasing ionic strength, often via a salt gradient (e.g., 0–0.5 M NaCl), which competes with bound ions and displaces analytes in order of their charge affinity; alternatively, pH shifts can be used, though salt gradients are more common for preserving biomolecule activity.^[59] Retention depends on factors like net charge density and ionic environment, with weakly charged molecules eluting first.^[16] Applications of ion-exchange chromatography span analytical and preparative scales, including protein fractionation where it resolves isoforms differing by a single charged residue, such as separating human serum proteins on Q-based columns.^[16] It is also employed in water purification to remove charged impurities like heavy metals or ions via selective binding and regeneration.^[16] For nucleic acid separation, anion exchangers capture negatively charged DNA or RNA, enabling purification of oligonucleotides from complex mixtures.^[16] The ion-exchange capacity of resins, a key performance metric, is quantified in milliequivalents per gram (meq/g), typically ranging from 1.5–4.0 meq/g for common strong exchangers, reflecting the amount of exchangeable ions available.^[60]

Size-Exclusion Chromatography

Size-exclusion chromatography (SEC), also referred to as gel filtration in aqueous systems or gel permeation chromatography in organic solvents, separates molecules passively based on their hydrodynamic volume without relying on adsorption, ion-exchange, or other interactive mechanisms. In this technique, a sample is applied to a column packed with porous beads, and as the mobile phase carries the mixture through, larger molecules are excluded from the internal pores of the beads, traveling only through the interstitial void volume and eluting first. Smaller molecules, in contrast, can diffuse into the pores, thereby extending their effective path length and resulting in later elution. This size-dependent separation produces an inverse elution order relative to molecular size, governed primarily by entropic effects in solution rather than enthalpic interactions.^[61]^[62]^[63] The technique was pioneered in 1959 by Jerker Porath and Per Flodin, who developed the first practical implementation using cross-linked dextran gels for aqueous separations of proteins and salts, demonstrating effective desalting and group fractionation. These early dextran gels, commercialized as Sephadex by Pharmacia, feature a controlled pore size distribution that defines the exclusion limit—the molecular weight above which species are fully excluded from pores—and the fractionation range, where partial penetration occurs. For biomolecular applications, agarose-based media such as Superdex provide enhanced rigidity and larger pore sizes suitable for high-molecular-weight species like antibodies (up to 300,000 Da), while maintaining biocompatibility and low non-specific binding. Pore sizes are engineered to span ranges from small molecules (e.g., Sephadex G-10, exclusion limit ~700 Da) to macromolecules (e.g., Sephadex G-200, up to 800,000 Da), ensuring versatility across sample types.^[64]^[63]^[61] SEC finds primary use in determining molecular weights and size distributions of polymers, proteins, and other macromolecules by establishing a baseline for hydrodynamic behavior. It is particularly valuable for desalting, where small ions or buffer components are separated from larger biomolecules in volumes up to 30% of the column bed, and for analyzing oligomer distributions, such as distinguishing protein monomers from dimers or aggregates in biopharmaceutical quality control. In molecular weight estimation, samples are compared against elution profiles of known standards, enabling assessment of conformational changes or conjugation effects without absolute mass spectrometry. These applications leverage SEC's gentle, non-denaturing conditions to preserve biological activity, making it indispensable in biochemistry and polymer science.^[62]^[63]^[61] Calibration of SEC columns relies on linear standards of defined molecular weights, such as globular proteins (e.g., thyroglobulin at 670 kDa or cytochrome c at 12.4 kDa), which are chromatographed to generate a sigmoidal calibration curve. Within the linear portion of this curve—typically spanning the media's fractionation range—the logarithm of molecular weight is plotted against elution volume (Ve), often normalized as Ve/Vo (where Vo is the void volume), yielding a straight line for interpolating unknown sample sizes. This relative calibration method, while dependent on standard similarity to the analyte, provides reliable size estimates and is routinely used to validate column performance and pore accessibility. For more precise absolute measurements, advanced setups couple SEC with detectors like light scattering, though standard-based plots remain the foundational approach.^[62]^[61]^[63]

Specialized Techniques

Displacement and Frontal Chromatography

Displacement chromatography is a non-elution technique where a carrier or displacer molecule with high affinity for the stationary phase is used to sequentially displace bound analytes from the column in order of their decreasing affinity, resulting in the formation of adjacent, rectangular zones rather than Gaussian peaks.^[65] This mode operates under overloaded conditions, maximizing column capacity by leveraging competitive adsorption, where the displacer saturates the stationary phase and pushes analytes forward based on their relative binding strengths.^[66] Unlike elution methods, which rely on mobile phase changes to desorb analytes individually, displacement chromatography produces sharp, square-wave fronts with minimal band broadening, enabling efficient separation of complex mixtures without the need for gradient programming.^[65] In practice, the process begins with loading the sample onto the column, followed by introduction of the displacer, which displaces the analytes in a train of contiguous zones; each zone corresponds to a component ordered by affinity, with the least affine eluting first.^[65] This technique, originally conceptualized by Arne Tiselius in 1943, has been refined for biomolecule purification, particularly through sample displacement chromatography (SDC), where the sample components themselves act as mutual displacers without an external carrier.^[65] Key advantages include higher productivity and reduced solvent consumption compared to elution, as up to 80% of the column's binding capacity can be utilized.^[66] Frontal chromatography, or frontal analysis, involves the continuous application of a sample solution to the column until the stationary phase reaches saturation, allowing measurement of breakthrough curves to determine adsorption isotherms and binding capacities.^[67] In this method, the sample is fed at a constant concentration, and the effluent is monitored for the appearance of the analyte (breakthrough), after which the concentration stabilizes at the feed level; the retained amount is calculated from the integral of the breakthrough curve, providing insights into adsorption behavior such as Langmuir isotherms.^[67] Distinct from displacement, frontal analysis does not require a separate displacer and is primarily analytical, focusing on the least-retained component's elution in the flow-through until saturation by more strongly bound species.^[67] Applications of displacement chromatography are prominent in large-scale purification of therapeutic proteins, such as monoclonal antibodies and clotting factors from plasma, where it facilitates high-throughput processing with smaller columns and straightforward scaling by parameter multiplication.^[66] For instance, SDC has been employed to separate charge variants of monoclonal antibodies and enrich low-abundance proteins from complex biological matrices like renal tissue extracts, supporting proteomics and biomarker discovery in drug development.^[66]^[65] Frontal analysis complements these by characterizing adsorption properties essential for optimizing displacement systems, with uses in evaluating protein binding on ion-exchange resins like POROS Q/M for bovine serum albumin.^[67]

Countercurrent Chromatography

Countercurrent chromatography (CCC) is a liquid-liquid partitioning technique that separates compounds based on their differential distribution between two immiscible liquid phases, one serving as the stationary phase retained by centrifugal force and the other as the mobile phase flowing through the system. Unlike traditional chromatography, CCC employs no solid support, relying instead on the density difference and J-force (a combination of centrifugal and Coriolis forces) to hold the stationary phase in place while allowing continuous mixing and settling of the phases. This principle enables high sample recovery and avoids irreversible adsorption, making it particularly suitable for sensitive biomolecules.^[68] In hydrodynamic CCC, such as high-speed countercurrent chromatography (HSCCC), the separation occurs within a coiled column mounted on a planetary centrifuge that rotates around two axes, generating a variable centrifugal field to promote phase distribution. Elution can proceed in head-to-tail mode, where the denser stationary phase occupies the head of the coil and the mobile phase flows from head to tail, or tail-to-head mode for reverse-phase operation, enhancing versatility for polar and non-polar samples. Centrifugal partition chromatography (CPC), a related variant, uses a series of interconnected chambers fixed in a rotor subjected to a constant centrifugal field along a single axis, providing more stable phase retention but requiring rotary seals. These configurations, pioneered by Yoichiro Ito in the 1970s and 1980s, allow for efficient separations under low pressure (0.1–10 kg/cm² for HSCCC).^[68] CCC finds extensive applications in the isolation of natural products from complex plant and marine extracts, where it excels in purifying compounds like ginsenosides from Panax ginseng and salvianolic acid B from Salvia miltiorrhiza with high throughput (up to 5 g/h) and purity exceeding 95%. It is also effective for chiral separations, employing specialized elution modes such as recycling or multiple dual-mode to resolve enantiomers via biphasic chiral recognition, offering advantages in preparative-scale enantioseparation with low solvent consumption. The absence of solid supports prevents analyte denaturation and contamination, supporting scalability from analytical to industrial levels in pharmaceutical and food analysis.^[69]^[70]

Chiral Chromatography

Chiral chromatography is a specialized separation technique designed to resolve enantiomers, which are mirror-image isomers of chiral molecules that exhibit identical physical properties but differ in biological activity. This method relies on the use of chiral selectors that interact differently with each enantiomer, forming transient diastereomeric complexes that lead to differential retention times.^[71] The technique is essential in fields where enantiomeric purity is critical, as the wrong enantiomer can be ineffective or harmful.^[72] The core mechanism of chiral chromatography involves chiral stationary phases (CSPs), which are immobilized selectors covalently bonded to a solid support within the column. These CSPs exploit the three-point interaction model, where the chiral selector forms diastereomeric complexes with the analyte enantiomers through non-covalent forces such as hydrogen bonding, π-π interactions, and steric repulsion, resulting in distinct affinities for each enantiomer.^[71] Common CSPs include cyclodextrins, which are cyclic oligosaccharides that encapsulate hydrophobic portions of enantiomers in their toroidal cavity, forming inclusion complexes stabilized by dipole-dipole and van der Waals forces; this approach is particularly effective for separating small polar molecules.^[73] Crown ethers, another prominent class, function via ion-dipole interactions and hydrogen bonding with primary amino groups of analytes, creating host-guest complexes that differentiate enantiomers based on spatial fit within the ether's ring structure.^[74] Various modes of chiral chromatography adapt these mechanisms to different mobile phases and analytes. High-performance liquid chromatography (HPLC)-based chiral separations typically employ packed columns with CSPs, offering robust resolution for a wide range of non-volatile compounds under normal or reversed-phase conditions.^[71] Supercritical fluid chromatography (SFC) enhances enantioselectivity compared to traditional liquid mobile phases by using CO₂-based fluids, which provide lower viscosity, higher diffusivity, and tunable density, leading to faster separations and higher efficiency for preparative-scale enantioseparations.^[75] For volatile analytes, gas chromatography (GC) utilizes CSPs coated on capillary columns, leveraging vapor-phase interactions to separate enantiomers of thermally stable compounds like essential oils or pheromones.^[76] In pharmaceutical applications, chiral chromatography ensures the purity of enantiomers, such as separating the (R)- and (S)-forms of drugs like thalidomide or ibuprofen, where the bioactive enantiomer must predominate to avoid adverse effects.^[72] Similarly, in agrochemicals, it facilitates the analysis and purification of chiral pesticides, such as the enantiomers of metalaxyl fungicide, to optimize efficacy while minimizing environmental persistence of inactive forms.^[77] The success of these separations is validated by measuring enantiomeric excess (ee), defined as:

\text{ee} = \frac{|\%R - \%S|}{(\%R + \%S)} \times 100

where %R and %S represent the percentages of the respective enantiomers, providing a quantitative indicator of optical purity typically targeted above 99% in regulatory contexts.^[78]

Applications and Advances

Analytical and Preparative Uses

Chromatography serves as a cornerstone in analytical applications across various fields, enabling the detection and quantification of trace compounds in complex matrices. In environmental monitoring, gas chromatography (GC) is extensively employed to analyze pesticides in water, soil, and food samples, adhering to stringent regulatory limits such as the European Union's maximum of 0.1 μg/L per pesticide in drinking water. This technique facilitates screening for over 300 pesticides, supporting efforts to assess contamination levels and ensure ecosystem safety. In food safety, high-performance liquid chromatography (HPLC) methods are routinely used to determine synthetic additives like preservatives and colorants in beverages and processed foods, verifying compliance with regulatory thresholds to prevent health risks from overconsumption. For clinical analysis, liquid chromatography coupled with mass spectrometry (LC-MS) plays a vital role in therapeutic drug monitoring by quantifying drug metabolites in biological fluids such as plasma, aiding in personalized dosing for medications like digoxin and warfarin to optimize efficacy and minimize toxicity. Preparative chromatography, in contrast, focuses on large-scale isolation and purification for downstream applications. In biopharmaceutical production, affinity chromatography is indispensable for purifying recombinant proteins, leveraging specific ligand interactions to achieve high selectivity and yield, as seen in the isolation of histidine-tagged proteins and glycoproteins essential for therapeutic development. This method supports the manufacturing of biologics by removing impurities and enabling scalable processes critical for drug formulation. For natural products research, column chromatography, often using silica gel or alumina as stationary phases, is a primary technique for isolating bioactive compounds from plant extracts, such as alkaloids from Kopsia arborea or taxol from Taxus cuspidata, providing pure isolates for pharmacological evaluation and potential drug discovery. Hyphenated techniques enhance the analytical power of chromatography by integrating separation with spectroscopic identification. Gas chromatography-mass spectrometry (GC-MS) excels in analyzing volatile compounds, combining chromatographic retention times with mass spectral fragmentation for precise structural confirmation and quantification, as applied in profiling essential oils and alkaloids in herbal medicines. Similarly, LC-MS, particularly with electrospray ionization, handles non-volatile and polar analytes, enabling the identification and measurement of metabolites in complex samples like citrus-derived coumarins, thereby improving accuracy in both qualitative and quantitative assessments across pharmaceutical and environmental contexts. Quantitative performance in chromatographic methods is characterized by metrics such as the limit of detection (LOD) and linearity ranges, which define sensitivity and reliable measurement scopes. The LOD, often calculated as 3.3 times the standard deviation of the response divided by the calibration curve slope, allows detection of analytes at concentrations as low as 1 ng/mL in HPLC assays for trace contaminants. Linearity ranges typically extend from the limit of quantification (LOQ, around 3 ng/mL) to 150% of expected levels, ensuring proportional responses; for instance, HPLC analysis of food additives like carmoisine exhibits linearity from 2–10 μg/mL with LODs of 0.10–0.19 μg/mL, supporting robust validation for routine monitoring.^[79]

Recent Developments and Instrumentation

Recent advancements in chromatography have focused on miniaturization to enhance portability, efficiency, and sensitivity, particularly through the development of ultra-high-performance liquid chromatography (UHPLC) and microfluidic systems. UHPLC systems utilize sub-2 μm particle columns, enabling separations at pressures exceeding 1000 bar, which significantly improves resolution and reduces analysis times compared to traditional HPLC.^[80]^[81] This technology has been widely adopted since the early 2000s for applications requiring high throughput, such as pharmaceutical analysis.^[82] Complementing UHPLC, microfluidic chips have revolutionized liquid chromatography by integrating sample preparation, separation, and detection on a single platform, often using channels less than 100 μm in width to minimize sample volumes and enable point-of-care diagnostics.^[83]^[84] These chips, fabricated via photolithography or 3D printing, have advanced since 2010 to support portable systems for environmental and biological monitoring.^[85]^[86] Multidimensional chromatography setups have emerged as powerful tools for resolving complex mixtures, particularly in proteomics and metabolomics. Comprehensive two-dimensional liquid chromatography (LC×LC) couples orthogonal separation mechanisms, such as size-exclusion and reversed-phase, to analyze proteomes with peak capacities over 1000, far surpassing one-dimensional methods.^[87]^[88] Since the 2010s, LC×LC has been optimized with automated modulation interfaces to handle high-resolution mass spectrometry detection, enabling the identification of thousands of proteins in single runs.^[89]^[90] Similarly, comprehensive two-dimensional gas chromatography (2D-GC) has seen innovations in cryogenic and thermal modulation techniques, improving separation of volatile compounds in environmental samples with enhanced sensitivity via fast detectors.^[91] Recent developments include valve-based modulators that reduce band broadening, making 2D-GC suitable for high-throughput petrochemical and food analyses since 2020.^[92]^[93] Efforts toward greener chromatography have prioritized reducing solvent use and waste, with supercritical fluid chromatography (SFC) leading through its reliance on carbon dioxide (CO₂) as a primary mobile phase. SFC operates under supercritical conditions to achieve faster separations with significantly less organic solvent than HPLC, with reductions of 75-90% in collected fractions, aligning with green chemistry principles established in the 2010s.^[94]^[95]^[96] Innovations in CO₂ recycling systems, such as closed-loop setups that recapture and reuse the fluid post-separation, have further minimized environmental impact, particularly in preparative-scale purifications.^[97]^[98] Solventless variants, including gas-solid chromatography adaptations, have also gained traction for volatile analyte separations without liquid phases.^[96] The integration of artificial intelligence (AI) and machine learning has transformed chromatographic data handling and method development. AI-driven algorithms now automate peak deconvolution in complex chromatograms, using convolutional neural networks to identify overlapping peaks with over 95% accuracy in LC-MS datasets.^[99]^[100] Since 2020, tools like PeakBot and NeatMS have reduced false positives in peak detection by learning from labeled spectra, enhancing reproducibility in high-throughput proteomics.^[101] For method optimization, machine learning models predict optimal gradients and column conditions, shortening development times by up to 50% in UHPLC workflows.^[102]^[103] Emerging techniques address challenges in processing crude samples and real-time oversight. Expanded bed adsorption (EBA) chromatography allows direct loading of unclarified feeds, such as cell lysates, by fluidizing high-density beads to prevent clogging and achieve up to 80% recovery in a single step.^[104]^[105] Developed in the late 1990s but refined post-2010 for biopharmaceuticals, EBA integrates clarification and capture, reducing process steps.^[106] Online monitoring via Raman spectroscopy has advanced process analytical technology, enabling non-invasive, real-time quantification of analytes and impurities during elution with sub-minute resolution.^[107]^[108] Fiber-optic Raman probes, integrated since 2020, facilitate in-line control in bioprocess chromatography, improving yield and compliance.^[109]^[110] As of 2025, further innovations include new high-performance liquid chromatography (HPLC) columns and accessories released in 2024-2025, enhancing resolution and throughput in analytical separations.^[111] Covalent organic frameworks (COFs) have emerged as advanced stationary phases, offering superior selectivity and stability for biomedical and environmental analyses.^[112] Additionally, green chromatography techniques continue to evolve, incorporating sustainable solvents and AI-optimized methods to minimize environmental impact.^[113]

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