Separations and Purifications

The separation of mixtures is important for two reasons. First, separatory techniques are required for analyzing any number of complex mixtures, from contaminants in well water to forensic DNA samples to pharmaceutical formulations. Second, it is often necessary to purify compounds for further use, for example, the isolation of morphine from poppy seeds or the purification of intermediates in a multistep organic synthesis. One of the simplest ways to separate out a desired product is through extraction, the transfer of a dissolved compound (the desired product) from a starting solvent into a solvent in which the product is more soluble. Liquid-liquid extraction is based on the fundamental concept that like dissolves like. This principle tells us that a polar substance will dissolve best in polar solvents, and a nonpolar substance will dissolve best in nonpolar solvents.1 These characteristics can be taken advantage of in order to extract only the desired product, leaving most of the impurities behind in the first solvent. When we perform extractions, it is important to make sure that the two solvents are immiscible, meaning that they form two layers that do not mix, like water and oil. The two layers are temporarily mixed by shaking so that solute can pass from one solvent to the other. For example, in a solution of isobutyric acid and diethyl ether, we can extract the isobutyric acid with water. Isobutyric acid, with its polar carboxyl group, is more soluble in a polar solvent like water than in a nonpolar solvent like ether. When the two solvents are mixed together, isobutyric acid will transfer to the water layer, which is called the aqueous phase (layer). The nonpolar ether layer is called the organic phase (layer). The aqueous and organic phases will separate on their own, given time to do so. In order to isolate these two phases, a piece of equipment called a separatory funnel is used.1

 

Gravitational forces causes the denser layer to sink to the bottom of the funnel, where it can then be removed by turning the stopcock at the bottom. It is more common for the organic layer to be on top, although the opposite can also occur like in the case of dichloromethane. Remember that the position of the layers is determined by their relative densities. Compounds that utilise hydrogen bonding, such as alcohols or acids, will move most easily into the aqueous layer. Compounds that utilise dipole–dipole interactions are less likely to move into the aqueous layer. Compounds that utilise Van der Waals forces are least likely to move into the aqueous layer. Let us assume in the above example with isobutyric acid that the aqueous layer is denser and settles to the bottom of the separatory funnel. Once we drain the aqueous layer from the separatory funnel, we repeat the extraction several times. Additional water is added to the separatory funnel which is shaken and allowed to settle, and the aqueous layer is once again drained off. This is done in order to extract as much of the isobutyric acid from the ether layer as possible because it does not completely transfer with the first extraction. Multiple extractions with fresh water are more effective for obtaining the most product, rather than a single extraction with a larger volume of water. Once the desired product has been isolated in the solvent, we can obtain the product alone by evaporating the solvent, usually by using a rotary evaporator.1 You can use the properties of acids and bases to your advantage in extraction:

 

HA + base → A +  H–base+

When the acid dissociates, the anion formed will be more soluble in the aqueous layer than the original protonated acid because it is charged. Thus, adding a base will help to extract an acid into the aqueous phase. Another way to take advantage of solubility properties is to perform the reverse of the extraction we just described in order to remove unwanted impurities. In this case, a small amount of solute is used to extract and remove impurities, rather than the compound of interest. This process is called a wash.

Extraction requires two solvents that are immiscible in order to separate the product. But what happens when the product itself is a liquid that is soluble in the solvent? This is where distillation is useful. Distillation takes advantage of differences in boiling point to separate two liquids by evaporation and condensation. The liquid with the lower boiling point will vaporize first, and the vapors will rise up the distillation column to condense in a water-cooled condenser. This condensate then drips down into a vessel. The end product is called the distillate. The heating temperature is kept low so that the liquid with the higher boiling point will not be able to boil and therefore will remain liquid in the initial container. This is the process that is used to make liquor at a distillery. Because ethanol boils at a lower temperature than water, we can use distillation to make beverages with high ethanol contents. Simple distillation is the least complex version of distillation. This technique should only be used to separate liquids that boil below 150°C and have at least a 25°C difference in boiling points. These restrictions prevent the temperature from becoming so high that the compounds degrade and provide a large enough difference in boiling points that the second compound won’t accidentally boil off into the distillate. The apparatus for this technique consists of a distilling flask containing the combined liquid solution, a distillation column consisting of a thermometer and a condenser, and a receiving flask to collect the distillate. Vacuum distillation is used whenever we want to distill a liquid with a boiling point over 150°C. By using a vacuum, we lower the pressure, thereby decreasing the temperature that the liquid must reach in order to boil.2 This allows us to distill compounds with higher boiling points at lower temperatures so that we do not have to worry about degrading the product.

 

Vacuum Distillation. Source: https://www.equilibar.com/2016/10/application-spotlight-vacuum-distillation/

 

The initial solution is placed in the heated distilling flask, where the components of the solution with the lowest boiling points will vaporize first. The vapor then condenses in the water-cooled condenser, and this distillate drips into the receiving flask. To separate two liquids with similar boiling points (less than 25°C apart), we use fractional distillation. In this technique, a fractionation column connects the distillation flask to the condenser. A fractionation column is a column in which the surface area is increased by the inclusion of inert objects like glass beads or steel wool. As the vapor rises up the column, it condenses on these surfaces and refluxes back down until rising heat causes it to evaporate again, only to condense again higher in the column. Each time the condensate evaporates, the vapor consists of a higher proportion of the compound with the lower boiling point. By the time the top of the column is reached, only the desired product drips down to the receiving flask.

Fractional Distillation

 

With increased surface area in the distillation column, the distillate has more places to condense on its way up the column. This allows for more refined separation of liquids with fairly close boiling points.

Chromatography is another tool that uses physical and chemical properties to separate and identify compounds from a complex mixture.3 In all forms of chromatography discussed here, the concept is identical: the more similar a compound is to its surroundings (whether by polarity, charge, or other characteristics), the more it will stick to and move slowly through its surroundings. Chromatography separates compounds based on how strongly they adhere to the solid, or stationary phase (or in other words, how easily they come off into the mobile phase). The process begins by placing the sample onto a solid medium called the stationary phase, or adsorbent. We then run the mobile phase, usually a liquid (or a gas in gas chromatography) through the stationary phase. This will displace (elute) the sample and carry it through the stationary phase. Depending on the characteristics of the substances in the sample and the polarity of the mobile phase, it will adhere to the stationary phase with differing strengths, causing the different substances to migrate at different speeds. This is called partitioning, and it represents an equilibrium between the two phases. Different compounds will have different partitioning coefficients and will elute at different rates. This results in separation within the stationary phase, allowing us to isolate each substance individually. Thin-layer chromatography and paper chromatography are extremely similar techniques, varying only in the medium used for the stationary phase. For thin-layer chromatography, a thin layer of silica gel or alumina adherent to an inert carrier sheet is used. For paper chromatography, as the name suggests, the medium used is paper, which is composed of cellulose.

For these techniques, the sample that we want to separate is placed directly onto the adsorbent itself and this is called spotting. It is called spotting because we apply a small, well-defined spot of the sample directly onto the silica or paper plate. The plate is then developed, which involves placing the adsorbent upright in a developing chamber, usually a beaker with a lid or a wide-mouthed jar. At the bottom of this jar is a shallow pool of solvent, called the eluent. The spots of sample must be above the level of the solvent, or else they will dissolve into the pool of solvent rather than running up the plate. When set up correctly, the solvent will creep up the plate by capillary action, carrying the various compounds in the sample with it at varying rates. When the solvent front nears the top of the plate, the plate is removed from the chamber and allowed to dry. As mentioned before, TLC is often done with silica gel, which is polar and hydrophilic. The mobile phase, on the other hand, is usually an organic solvent of weak to moderate polarity, so it doesn’t bind well to the gel. Because of this, nonpolar compounds dissolve in the organic solvent and move quickly as the solvent moves up the plate, whereas the more polar molecules stick to the gel. Thus, the more nonpolar the sample is, the further up the plate it will move. Reverse-phase chromatography is the exact opposite. In this technique, the stationary phase used is nonpolar, so polar molecules move up the plate quickly, while nonpolar molecules stick more tightly to the stationary phase. The spots of individual compounds are usually white, which makes them difficult or impossible to see on the white paper or TLC plate. To get around this problem, the developed plate can be placed under ultraviolet light, which will show any compounds that are ultraviolet-sensitive. Alternatively, iodine, phosphomolybdic acid, or vanillincan be used to stain the spots, although this will destroy the compounds such that they cannot be recovered. When TLC is performed, compounds are generally identified using the retention factor (Rf), which is relatively constant for a particular compound in a given solvent. The Rf is calculated using the equation:

Because its value is relatively constant, the Rf value can be used to identify unknown compounds. This technique is most frequently performed on a small scale to identify unknown compounds. It can also be used on a larger scale as a means of purification, a technique called preparative TLC. As the large plate develops, the larger spot of sample splits into bands of individual compounds, which can then be scraped off and washed to yield pure compounds. The principles behind column chromatography are the same as for thin-layer chromatography, although there are some differences. First, column chromatography uses an entire column filled with silica or aluminium beads as an adsorbent, allowing for much greater separation.3 In addition, thin-layer chromatography uses capillary action to move the solvent up the plate, whereas column chromatography uses gravity to move the solvent and compounds down the column. To speed up the process, one can force the solvent through the column using gas pressure, a technique called flash column chromatography. In column chromatography, the solvent polarity can also be changed to help elute the desired compound.

Column Chromatography

 

The sample is added to the top of the column, and a solvent is poured over it. The more similar the sample is to the mobile phase, the faster it elutes; the more similar it is to the stationary phase, the more slowly it will elute (if at all). Eventually, the solvent drips out of the end of the column, and the different fractions that leave the column can be collected over time. Each fraction will contain different compounds. After collection, the solvent can be evaporated, leaving behind the compounds of interest. Column chromatography is particularly useful in biochemistry because it can be used to separate and collect macromolecules such as proteins or nucleic acids.3 Gas chromatography (GC) is another method that can be used for qualitative separation. GC is similar to the other types of chromatography. The main conceptual difference is that the eluent is a gas (usually helium or nitrogen) instead of a liquid. The adsorbent is a crushed metal or polymer inside a 30-foot column.4 This column is coiled and kept inside an oven to control its temperature. The mixture is then injected into the column and vaporized. The gaseous compounds travel through the column at different rates because they adhere to the adsorbent in the column to different degrees and will separate in space by the time they reach the end of the column. The injected compounds must be volatile (low melting-point, sublimable solids or vaporizable liquids).4 The compounds are registered by a detector, which records them as a peak on a chart. It is common to separate molecules using GC and then to inject the pure molecules into a mass spectrometer for molecular weight determination, which is referred to as GC–mass spectrometry.5 

Gc-Ms Setup

 

The sample is injected into the column and moves with the gaseous mobile phase through a stationary liquid or solid phase; a computer identifies the sample components. In high-performance liquid chromatography (HPLC), the eluent is a liquid, and it travels through a column of a defined composition.6 There are a variety of stationary phases that can be chosen depending on the target molecule and the quantity of material that needs to be purified. This is fairly similar to column chromatography because the various compounds in solution will react differently with the adsorbent material. In the past, very high pressures were used, but recent advances allow for much lower pressures. In HPLC, a small sample is injected into the column, and separation occurs as it flows through. The compounds pass through a detector and are collected as the solvent flows out of the end of the apparatus. The interface is similar to that used for GC because the entire process is computerized, but uses liquid under pressure instead of gas. Because the whole process is under computer control, sophisticated solvent gradients as well as temperature can be applied to the column to help resolve the various compounds in the sample, hence the higher performance of HPLC over regular column chromatography.6

One important methodology for the separation and characterization of amino acids and oligopeptides is electrophoresis. In this analytical technique, a mixture of amino acids (or oligopeptides) is spotted onto the center of a conductive gel, across which is applied an electrical potential.7 The amino acids then migrate toward the anode or cathode, depending upon the net charge on each molecule (positively-charged species move to the cathode and negatively-charged species move to the anode). The mobility of each ion depends upon its net charge and its mass. Since amino acids are polyfunctional ionizable molecules, their net charge is a function of pH. In the case of alanine, for example, at very low pH the net charge is +1 (both the carboxylic acid and amine functionalities are protonated), thus at pH 2, alanine would move toward the cathode. Conversely, at very high pH the net charge is −1 (both the carboxylic acid and amine functionalities are deprotonated), thus at pH 10, alanine would move toward the anode. At pH 6, there is no net charge on the alanine molecule (positive and negative charges exactly cancel each other) therefore, alanine would not migrate on an electrophoresis gel at pH 6. This is known as the isoelectric point.7 A common contemporary method for electrophoretic analysis is known as capillary electrophoresis. The separatory medium in this technique is not a gel, but rather a capillary with either end immersed in a buffer solution. Application of an electrical potential across the capillary induces a flow of buffer toward the cathode, a phenomenon known as electroosmotic flow (EOF). The sample is introduced either by temporarily replacing the anode-side buffer with a solution of the sample or via a more sophisticated in-line injection system. While the finer details regarding the mode of separation are different for CE versus gel electrophoresis, the general principle of separation based on charge still holds.

In ion-exchange chromatography, the beads in the column are coated with charged substances so that they attract or bind compounds that have an opposite charge.8 For instance, a positively charged compound will attract and hold a negatively charged backbone of DNA or protein as it passes through the column, either increasing its retention time or retaining it completely. After all other compounds have moved through the column, a salt gradient is used to elute the charged molecules that have stuck to the column. In size-exclusion chromatography, the beads used in the column contain tiny pores of varying sizes. These tiny pores allow small compounds to enter the beads, thus slowing them down. Large compounds can’t fit into the pores, so they will move around them and travel through the column faster. It is important to remember that in this type of chromatography, the small compounds are slowed down and retained longer—which may be counterintuitive. The size of the pores may be varied so that molecules with different molecular weights can be fractionated. A common approach in protein purification is to use an ion-exchange column followed by a size-exclusion column. In affinity chromatography, a protein of interest is bound by creating a column with high affinity for that protein.9 This can be accomplished by coating beads with a receptor that binds the protein or a specific antibody to the protein; in either case, the protein is retained in the column. Common stationary phase molecules include nickel, which is used in separation of genetically engineered proteins with histidine tags, antibodies or antigens, and enzyme substrate analogues, which mimic the natural substrate for an enzyme of interest. Once the protein is retained in the column, it can be eluted by washing the column with a free receptor (or target or antibody), which will compete with the bead-bound receptor and ultimately free the protein from the column. Eluents can also be created with a varying pH or salinity level that disrupts the bonds between the ligand and the protein of interest.9 The only drawback of the elution step is that the recovered substance can be bound to the eluent. If, for example, the eluent was an inhibitor of an enzyme, it could be difficult to remove.

Stereoisomers have the same molecular formula, same connectivity, but have different 3-D arrangements across one or more asymmetric (chiral) centers.10 A chiral center is any atom with 4 different entities attached to it. Enantiomers are mirror images of each other. That means all chiral centers in one enantiomer is reversed in the other. You can’t have stereoisomers if you don’t have a chiral center. Racemic mixtures, or racemate, is one that has equal amounts of left- and right-handed enantiomers of a chiral molecule.10 Racemic mixtures do not rotate polarized light, so they are optically inactive. It is often desirable to prepare compounds as a single enantiomer. There are two general approaches to this problem: (1) by designing a synthesis that results only in a single enantiomer (chiral synthesis), or (2) by synthesizing the product in racemic form and then separating the enantiomers from one another (chiral resolution). The second approach can be accomplished using one of two methods: (1) by preferential crystallization and (2) by chiral HPLC. In preferential crystallization, a racemic mixture is treated with an optically pure compound that can coordinate very tightly with the racemate. Very often this is done with salt formation. The second general approach to chiral resolution is through HPLC with the use of a chiral stationary phase. In this technique, silica gel is derivatized with an enantiomerically pure chiral species. When a mixture of enantiomers travels through such a column, one enantiomer tends to interact more strongly with the chiral stationary phase, thereby slowing down its progress through the column. Separation of enantiomers can also be done by biological means, such as using enzymes. Enzymes are highly specific and can differentiate between enantiomers. For example, if an enzyme digests or modifies all L-amino acids, then you’d be able to use that enzyme to separate a D/L racemic mixture. In nature, all proteins are made up of L-amino acids.11 Chromatographic methods, whereby the stationary phase is a chiral reagent that adsorbs one enantiomer more strongly than the other, have been used to resolve racemic compounds, but such resolutions seldom have led to both pure enantiomers on a preparative scale. Other methods, called kinetic resolutions, are excellent when applicable. The procedure takes advantage of differences in reaction rates of enantiomers with chiral reagents. One enantiomer may react more rapidly, thereby leaving an excess of the other enantiomer behind. For example, racemic tartaric acid can be resolved with the aid of certain penicillin molds that consume the dextrorotatory enantiomer faster than the levorotatory enantiomer. As a result, almost pure (-)-tartaric acid can be recovered from the mixture:

(±)-tartaric acid + mold  →  (-)-tartaric acid + more mold

 

A disadvantage of resolutions of this type is that the more reactive enantiomer usually is not recoverable from the reaction mixture. A common way of separation of enantiomers uses the conversion into diastereomers, which are not mirror images of each other. The following example demonstrates this principle. The two enantiomers 1-phenylethylamine (S-(-), R-(+)) are separated using (L)-(+)-tartaric acid (also known as the (R,R)-form) as resolving agent. The two salts formed possess different cation ions are therefore not enantiomers of each other anymore. The (S)-(1)-phenylethyl ammonium-(R,R)-tartrate salt crystallizes faster than the (R)-(1)-phenylethyl ammonium-(R,R)-tartrate salt.

The free amine can be recovered by reaction of the ammonium salt with a strong base i.e., sodium hydroxide or sodium carbonate. While the ammonium salt is soluble in aqueous solution, the free amine is better soluble in organic solvents. The charged tartrate ion stays behind in the aqueous solution. In order to isolate the (R)-form of the amine, the (S,S)-form of tartaric acid should be used. The resolution of 1-phenylethylamine can also be accomplished using L-malic acid.

 

References

1) Jack D. Law, T. A. (2009, February 3). Liquid-Liquid Extraction Equipment. Retrieved from Idaho National Laboratory: http://www.cresp.org/NuclearChemCourse/monographs/11_Law_Liquid-liquid%20extraction%20equipment%20jdl_3_2_09.pdf

2) Vac Distillation – Harwood, L. M. (1989, June 13). In Experimental organic chemistry: Principles and Practice (Illustrated ed.) (pp. 147–149). Wiley Blackwell.

3) Laurence M. Harwood, C. J. (1989). In Experimental organic chemistry: Principles and Practice (Illustrated ed. (pp. 180–185).

4) Pavia, D. L. (2006). In Introduction to Organic Laboratory Techniques (4th Ed.) (pp. 797–817). Thomson Brooks/Cole.

5) AG, L. (2012, March 11). Gas Chromatography.

6) Gerber, F., Krummen, M., Potgeter, H., Roth, A., Siffrin, C., & Spoendlin, C. (2004). Practical aspects of fast reversed-phase high-performance liquid chromatography using 3μm particle packed columns and monolithic columns in pharmaceutical development and production working under current good manufacturing practice”. Journal of Chromatography , 127–133.

7) Lyklema, J. (1995). In Fundamentals of Interface and Colloid Science (p. 3).

8) Luqman, M. I. (2012). Ion Exchange Technology II. Springer Netherlands.

9) Ninfa, A. J., Ballou, D. P., & Benore, M. (2009). In Fundamental Laboratory Approaches for Biochemistry and Biotechnology (2 ed.) (p. 133). Wiley.

10) Dietmar Kennepohl, S. F. (2016, November 18). Racemic Mixtures and the Resolution of Enantiomers. Retrieved from Chemistry Libretexts: https://chem.libretexts.org/Textbook_Maps/Organic_Chemistry_Textbook_Maps/Map%3A_Organic_Chemistry_(McMurry)/Chapter_05%3A_Stereochemistry_at_Tetrahedral_Centers/5.08_Racemic_Mixtures_and_the_Resolution_of_Enantiomers

11) California, U. o. (2016, January 15). Separation of Enantiomers (Resolution). Retrieved from http://www.chem.ucla.edu/~bacher/Specialtopics/Resolution.html

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