3.2: Protein Purification

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Learning Goals (ChatGPT o1, 1/25/25)

• Explain the Rationale for Protein Purification:

  • Describe why isolating a single protein from complex cellular mixtures is essential for structural and functional studies.

• Differentiate Between Preparative and Analytical Purification:

  • Distinguish the objectives, techniques, and quality control measures used in preparative versus analytical purification.

• Outline Sample Processing Techniques:

  • Identify various cell lysis methods (e.g., freeze/thaw, sonication, detergent treatment) and discuss how centrifugation separates soluble proteins from cellular debris.

• Understand Precipitation and Dialysis Methods:

  • Explain the role of ammonium sulfate precipitation and dialysis in concentrating proteins and removing contaminants.

• Apply Chromatography Principles:

  • Compare and contrast different chromatographic techniques (size exclusion, ion exchange, hydrophobic interaction, and affinity chromatography) based on protein properties such as size, charge, hydrophobicity, and binding affinity.

• Interpret Chromatographic Data:

  • Analyze elution profiles, calculate distribution coefficients, and evaluate parameters like void volume and stationary phase volume.

• Quantitatively Evaluate Purification Efficiency:

  • Calculate and interpret metrics such as total protein, total activity, specific activity, yield, and fold-purification to assess the success of a purification scheme.

• Describe the Theory of Electrophoresis:

  • Explain how charged particles move under an electric field, and relate factors like electrophoretic mobility to charge, size, and shape (including concepts like the Stokes radius).

• Distinguish Native and Denaturing Electrophoresis:

  • Compare techniques such as native PAGE and SDS-PAGE, and understand how SDS alters protein charge and conformation to facilitate molecular weight-based separation.

• Integrate Isoelectric Focusing and Two-Dimensional Electrophoresis:

  • Understand how pH gradients in isoelectric focusing separate proteins by their isoelectric points and how 2-DE combines IEF with SDS-PAGE for high-resolution analysis.

• Interpret Gel Staining and Detection Methods:

  • Evaluate different protein detection methods (Coomassie Blue, silver staining, autoradiography, Western blotting) and their applications in visualizing protein bands.

• Relate Biochemical Principles to Instrumentation:

  • Explain how HPLC, FPLC, and mass spectrometry (e.g., MALDI-TOF MS) integrate with protein purification protocols to provide structural and functional insights.

• Understand the Role of Protein Tagging and Peptide Synthesis:

  • Describe how molecular tags (e.g., His, FLAG, GFP) and solid-phase peptide synthesis facilitate purification and identification of proteins.

• Critically Analyze Experimental Design:

  • Identify potential sources of error in purification protocols (e.g., proteolysis, non-specific binding) and propose strategies to optimize yield and purity.

• Apply Theoretical Concepts to Practical Scenarios:

  • Use mathematical relationships (such as U = Q/(6πηRs)) to predict and analyze protein migration behavior in electrophoretic systems.

These goals aim to provide a comprehensive understanding that links the theory and practice of protein purification and electrophoresis, preparing students for advanced research and practical applications in biochemistry.

Introduction

A protein or other biological macromolecule must be purified before rigorously being studied structurally and functionally. Purifying a single protein from a mixture of as many as 10,000 other cellular or tissue proteins, each containing the same 20 naturally occurring amino acids linked in different orders, is difficult. Proteins differ in size (how many amino acids), charge (how many positively and negatively charged amino acids), sequence, and presence of specific binding sites on the proteins. Any technique that could be used to purify protein must be based on these inherent differences. Once the protein is purified, it must be analyzed by a spectral or electrophoretic technique.

Protein purification is a series of processes intended to isolate and purify a single protein or complex from cells, tissues, or whole organisms. Protein purification is vital for characterizing the function, structure, and interactions of the protein of interest. Separation steps usually exploit differences in protein size, physical-chemical properties, binding affinity, and biological activity.

Protein purification is either preparative or analytical. Preparative purifications aim to produce a relatively large quantity of purified proteins for subsequent use. Examples include the preparation of commercial products such as enzymes (e.g., lactase), nutritional proteins (e.g., soy protein isolate), and certain biopharmaceuticals (e.g., insulin). Many steps and quality control are required to remove other host proteins and biomolecules that could threaten the patient's health. Analytical purification produces a relatively small amount of a protein for various research or analytical purposes, including identification, structural characterization, and studies of the protein's structure, post-translational modifications, and function.

The choice of a starting material is key to the design of a purification process. In plants or animals, a particular protein usually isn't distributed homogeneously throughout the organism; different organs or tissues have higher or lower protein concentrations. Using tissues or organs with the highest concentration decreases the volumes needed to produce a given amount of purified protein. A protein of low abundance or high commercial value is often made using recombinant DNA technology. These techniques will be discussed in greater detail in Chapter 5.

Sample Processing

If the organism does not secrete the protein of interest into the surrounding solution, the first step of each purification process disrupts the cells containing the protein. Depending on how fragile the protein is, one of several techniques could be used, including repeated freezing and thawing, sonication, homogenization by high pressure (French press), homogenization by grinding (bead mill), and permeabilization by detergents (e.g., Triton X-100) and/or enzymes (e.g., lysozyme). Finally, the cell debris can be removed by centrifugation so that the proteins and other soluble compounds remain in the supernatant.

Proteases released during cell lysis cleave the proteins in the solution. As the protein of interest may be sensitive to proteolysis, it is important to proceed quickly and conduct many steps at low temperatures to reduce unwanted proteolysis. Alternatively, one or more protease inhibitors can be added to the lysis buffer immediately before cell disruption. Sometimes, it is also necessary to add DNase to reduce the viscosity of the cell lysate caused by a high DNA content.

Centrifugation

Centrifugation is a process that uses centrifugal force to separate mixtures of particles of varying masses or densities suspended in a liquid. When a vessel (typically a tube or bottle) containing a mixture of proteins or other particulate matter, such as bacterial cells, is rotated at high speeds, the inertia of each particle yields a force in the direction of the particle's velocity proportional to its mass. The tendency of a given particle to move through the liquid because of this force is offset by the resistance the fluid exerts on the particle. The net effect of "spinning" the sample in a centrifuge is that massive, small, and dense particles move outward faster than less massive particles or particles with more "drag" in the liquid. When suspensions of particles are "spun" in a centrifuge, a "pellet" may form at the bottom of the vessel enriched for the most massive particles with low drag in the liquid.

Non-compacted particles remain mostly in the liquid "supernatant" and can be removed from the vessel, separating the supernatant from the pellet. The angular acceleration applied to the sample determines the centrifugation rate, typically measured compared to g, the gravitational constant. If samples are centrifuged long enough, the vessels' particles will reach equilibrium, wherein the particles accumulate specifically at a point in the vessel where their buoyant density is balanced with centrifugal force. Such an "equilibrium" centrifugation can allow extensive particle purification.

In sucrose gradient centrifugation, a linear concentration gradient of sugar (typically sucrose, glycerol, or a silica-based density gradient media, like Percoll) is generated in a tube such that the highest concentration is on the bottom and the lowest on top. Percoll is a trademark owned by GE Healthcare companies. A protein sample is then layered on the gradient and spun at high speeds in an ultracentrifuge. This causes heavy macromolecules to migrate toward the bottom of the tube faster than lighter material. During centrifugation, in the absence of sucrose, as particles move farther and farther from the center of rotation, they experience greater centrifugal forces (the further they move, the faster they move). However, the useful separation range within the vessel is restricted to a small observable window. A properly designed sucrose gradient will counteract the increasing centrifugal force so the particles move proportionately to their time in the centrifugal field. After separating the protein/particles, the gradient is fractionated and collected. These are described in Figure \(\PageIndex{1}\).

Figure \(\PageIndex{1}\): Sucrose Density Gradient. Image derived from Michel Awkal

Precipitation and Differential Solubilization

In bulk protein purification, precipitation is a common first step to isolate proteins using a salt such as ammonium sulfate (NH₄)₂SO₄. Ammonium sulfate is often used as it is highly soluble in water, has relative freedom from temperature effects, and typically is not harmful to most proteins. Proteins are precipitated by (NH₄)₂SO₄ in their native state, which is important if you need the protein for structure/function studies. Furthermore, dialysis can remove ammonium sulfate as described in Figure \(\PageIndex{2}\).

Figure \(\PageIndex{2}\): Dialysis of a macromolecule (blue dots). Image adapted from Gjk003

The process of dialysis separates dissolved molecules by their size. The biological sample is placed inside a closed membrane, where the protein of interest is too large to pass through the membrane's pores but through which smaller ions can easily pass. As the solution reaches equilibrium, the ions become evenly distributed throughout the solution while the protein remains concentrated in the membrane. This reduces the overall salt concentration of the suspension.

The mechanism underlying salt precipitation is quite complicated. High concentrations of sodium chloride don't precipitate protein. Other salts like guanidinium chloride unfold proteins and do not lead to precipitation. Salt ions interact with the protein and solvent water in somewhat complicated ways (which we will explore later). We will be satisfied with the empirical observation that ammonium sulfate is the salt of choice to precipitate and concentrate proteins from a solution. One advantage of (NH₄)₂SO₄ protein precipitation from solution is that it can be performed inexpensively with very large volumes, so it is used early in many purification proteins. Different proteins precipitate at different (NH₄)₂SO₄ concentrations, so differential precipitation is often used. (NH₄)₂SO₄ concentrations are increased stepwise until the protein of interest is precipitated.

Some proteins are not soluble in water. These include transmembrane proteins that span cell membranes and large fibrous proteins. Membrane proteins can be solubilized by adding detergents like sodium dodecyl sulfate (SDS), which unfolds the proteins, and octylglucoside or Triton X-100, which keeps the protein structure intact.

Chromatography

Chromatography is used in almost all protein purification methods and is the key to separating a given protein from the 1000s of different proteins in cells and tissues. The separation of proteins on a chromatography column depends on the type of column and the chemical/physical properties of the molecule. There are four main types of chromatographies used to separate proteins:

• size exclusion chromatography in which proteins can be separated according to their size/shape or molecular weight
• ion exchange chromatography in which proteins are separated by their charge/isoelectric point;
• hydrophobic interaction chromatography (similar to reverse phase columns for purifying organic molecules) in which they are separated based on their relative hydrophobicity
• affinity chromatography in which proteins are separated based on binding to a ligand covalently attached to a column bead.

The purification protocol generally contains one or more chromatographic steps for preparative protein purification. The basic procedure in chromatography is to flow the solution containing the protein through a column packed with a chromatography resin selected to separate proteins based on a specific property of the protein. Different proteins interact differently with the column material. They can thus be separated by the time required to pass the column or the conditions required to elute the protein from the column. Usually, proteins are detected as they are eluting from the column by measuring the absorbance at 280 nm, at which the aromatic amino acids absorb.

Size Exclusion Chromatography (also known as Gel Filtration Chromatography)

This method is used to separate proteins based on size and shape. The chromatography beads have tiny openings/pores into which proteins smaller than the pore diameter can enter. Large proteins that can't enter the pores flow around the beads and elute faster than small ones that enter the pores. They diffuse out of the pores and enter the rest of the moving solvent before getting "trapped" again for a short time in more pores. Eventually, they move through the column and elute at a much greater volume than larger proteins, which can't enter the pores. Thus, proteins will be separated based on size, as illustrated in Figure \(\PageIndex{3}\). The eluate is collected in sequential test tubes (or fractions). The figure below shows the pores as channels that go through the bead. The openings in resin beads should be considered tiny crevices and indentations, not channels.

Figure \(\PageIndex{3}\): Size Exclusion Chromatography. Image from Dr. Kevin Ahern and Indira Rajagopal

Also known as gel filtration chromatography, it is a low-resolution isolation method involving beads with tiny “pores" that have a precise size distribution. The size is called an “exclusion limit," meaning molecules above a certain molecular weight will not fit into the tunnels. Molecules with sizes larger than the exclusion limit do not enter the tunnels and pass through the column relatively quickly by making their way between the beads. Smaller molecules, which can enter the tunnels, do so and, thus, have a longer path in passing through the column. Because of this, molecules larger than the exclusion limit will leave the column earlier, while smaller molecules that pass through the beads will elute from the column later. This method allows the separation of molecules by their size.

In any chromatography system, there is a mobile and stationary phase. For size exclusion chromatography, the stationary phase is usually a polymerized agarose or acrylamide bead, which contains pores of various sizes filled with the solvent. Let's pretend that the solvent (typically aqueous buffered solution) inside the bead is trapped there and doesn't exchange with the solvent moving around the bead so that it would be part of the stationary phase. The mobile phase is the solvent used to elute the column, which flows around the bead. The chromatography beads are often supplied in dried form, which must be swollen in the solvent before they are packed in the column. The volume of the agarose or acrylamide bead is very small compared to the volume of solution within their hydrated forms.

Size and shape effects in size exclusion chromatography

Size-size exclusion chromatography is so common so that we will explore it in greater detail

Several different column volumes can be defined as shown in Figure \(\PageIndex{4}\), where the packed chromatography beads are shown as circles.

Figure \(\PageIndex{4}\): Define volumes in size exclusion chromatography

If we consider the mass of the beads to offer a negligible amount to the volume of the bead, the actual volume in the bead is mostly from the trapped solution, which can be considered the "stationary" phase. The volume around the bead is called the void volume, V_o. It should be apparent the volume inside the bead is given by

\begin{equation}
V_i=V_t-V_o
\end{equation}

A solute elutes from the column in a broad peak. If the sample volume applied to the column is very small compared to V_t, the volume at which a solute elutes, \(V_e\), is considered the center of the elution peak. This is true when \(V_{sample} \gg V_e\).

Suppose we view this chromatography as a solute partitioning between the mobile and stationary phases (the basis of all chromatography). In that case, we might be interested in what fraction of the stationary phase, V_i, a solute might partition into. Such a ratio would be given by:

\begin{equation}
K=\frac{V_e-V_o}{V_t-V_o}
\end{equation}

V_t-V_o (= V_inside) represents 100% of the stationary phase, where \(K\) is a distribution coefficient. Consider two cases:

1. A very large solute compared to the pore size of the bead: In this case, V_e-V_o = 0 since V_e would be equal to V_o. (The solute wouldn't "see" any of the V_i.) In this case, K = 0. The solute would elute in the column's void volume since it is too large to partition into the volume within the beads. All solutes of molecular weight greater than or equal to the smallest solute that can't enter the gel beads will all elute in the void volume. Hence, solutes greater than this minimal size will co-elute from the column and not be separated. V_o is usually about 30-40% of the V_t.
2. A very small solute compared to the pore size. In this case, V_e-V_o = V_t - V_o, since V_e would be equal to V_t. The solute would "see" all of the solvent within the bead. In this case, K = 1. Like above, all solutes of MW equal to or less than the largest solute that can partition into the entire volume within a bead will co-elute at a volume near V_t.

Hence \(K\) is a partition coefficient, which varies from 0 - 1 and represents that fraction of V_i into which a solute could partition. However, this K is not exactly a partition coefficient since the actual volume of the gel matrix is assumed to be zero above. The graph in Figure \(\PageIndex{5}\) shows typical V_e as a fraction of V_t for solutes of different sizes (the x-axis is V_e/V_t).

Figure \(\PageIndex{5}\): Elution volumes (V_e for different size macromolecules)

Large species that cannot enter the pores in the beads flow around it and elute in the void volume (V₀), which is about 35-40% of V_t (red bell-shaped curve). Very small species can partition into V₀ and V_i, so the elute near V_t (green bell-shaped curve). If a species adsorbs to the column bead through noncovalent interactions (such as hydrogen bonds or ion-ion interactions), it may elute after V_t (purple bell-shaped curve).

K depends on the size and shape of the solute. The size and shape of an object determine its flow properties in a fluid. Frictional resistance (itself a force that acts in the opposite direction to the velocity, another vector quantity) can be shown to be proportional to the velocity.

\begin{equation}
F_f \propto v
\end{equation}

or

\begin{equation}
F_f=-f v
\end{equation}

where \(f\) is the frictional coefficient, which depends on the shape. The bigger the object, the more frictional resistance to movement. For a sphere, it can be shown that:

\begin{equation}
f=6 \pi \eta R_s
\end{equation}

where η is the viscosity (a measure of the resistance to flow of a liquid - water has a low viscosity, real maple syrup a high viscosity), and R_s (Stokes radius) is the radius of the hydrated sphere (the larger R_s, the larger the frictional coefficient, the larger the F_f which resists motion). For an irregularly shaped object, the Stokes radius is the radius of a sphere that would have the same frictional coefficient as the object. Hence, the R_s for a protein molecule that was not spherical would be much larger than the R_s for another protein molecule of identical molecular weight that was spherical. Hence, the V_e and the K values for a solute on a gel filtration column would best be related to the Stokes radius since R_s values consider both size and shape.

If you separate two proteins of equal mass but one is highly elongated, and the other is spherical, the elongated one, with a large R_S, would elute first (assuming that both don't elute together in the void volume, V₀.

Gel filtration can determine the molecular weight of an unknown, spherical (globular) protein compared to a standard curve generated from other globular proteins of known molecule weight. To ensure the protein has the same "effective" shape, the proteins are eluted under denaturing conditions to remove shape contributions to the elution order.

Separation based on charge - Ion Exchange Chromatography

The chromatography resin in this type consists of an agarose, acrylamide, or cellulose resin or bead, which is derivatized to contain covalently linked positively or negatively charged groups. Proteins in the mobile phase will bind through electrostatic interactions to the charged group on the column. In a mixture of proteins, positively charged proteins will bind to a resin containing negatively charged groups, like the carboxymethyl group, CM (-OCH₂COO^-) or sulfopropyl, SP, (-OCH₂CH₂CH₂SO₃^-). In contrast, the negatively charged proteins will pass through the column. The positively charged proteins can be eluted from the column with a mobile phase containing either a gradient of increasing salt concentration or a single higher salt concentration (isocratic elution). The most positively charged protein will be eluted last, at the highest salt concentration. Likewise, negatively charged proteins will bind to a resin containing positively charged groups, like the diethylaminoethyl group, DEAE (-OCH₂CH₂NH(C₂H₅)₂⁺) or a quaternary ethyl amino group, QAE, and can be separated analogously.

Ion exchange chromatography separates compounds according to the nature and degree of their ionic charge. The column used is selected according to its type and charge strength. Anion exchange resins have a positive charge and are used to retain and separate negatively charged compounds (anions). In contrast, cation exchange resins have a negative charge and are used to separate positively charged molecules (cations).

Before the separation begins, a buffer is pumped through the column to equilibrate the opposing charged ions. Upon sample injection, solute molecules will exchange with the buffer ions as each competes for the binding sites on the resin. The length of retention for each solute depends upon the strength of its charge. The most weakly charged compounds will elute first, followed by those with successively stronger charges. Because of the nature of the separating mechanism, pH, buffer type, buffer concentration, and temperature all play important roles in controlling the separation.

Figure \(\PageIndex{6}\) shows a cation exchange column. The beads (brown) contain negatively charged functional groups that can bind positive protein (blue) or concentrated regions of positive charge on a protein.

Figure \(\PageIndex{6}\): Ion Exchange Chromatography https://upload.wikimedia.org/wikiped...n_exchange.jpg. Jspiteri at English Wikibooks, Public domain, via Wikimedia Commons

Before loading the column with protein, the negatively charged beads would interact with positively charged countercations (often Na⁺) from the column equilibration buffer. When the protein solution is introduced to the column, the positively charged protein will exchange with the bound Na⁺ ions (hence the name cation exchanger). Conversely, an anion exchanger consists of positively charged beads, which exchange anions. Proteins bound through ion-ion interactions can be eluted by increasing the Na⁺ concentration in the eluting solution stepwise or using a salt concentration gradient. Ion exchange chromatography is a powerful protein purification tool and is frequently used in analytical and preparative separations.

Affinity Chromatography

In this technique, the chromatography resin is derivatized with a group that binds to a specific site on a given protein of interest. It may be a group that binds to the active site of an enzyme (such as benzamidine-agarose, which is used to purify trypsin) or an antibody that recognizes a specific amino acid sequence (an epitope) on a protein. For example, an antibody can be made to a specific peptide from albumin, the antibody covalently linked to agarose, and the antibody-agarose column then used to purify albumin specifically. This is a powerful technique since antibodies can be made that will bind selectively to a single protein. Knowing only the DNA sequence of a protein that has never been previously isolated, the amino acid sequence of the unknown protein can be derived from the DNA sequence. A 10-12 amino acid peptide from that protein can be synthesized in the lab (see the last section below), and an antibody raised against the peptide. The antibody will most likely bind to the unknown protein and the peptide and could be used to purify the protein.

These features of affinity chromatography are illustrated in Figure \(\PageIndex{7}\).

Figure \(\PageIndex{7}\): Example of Affinity Chromatography.

In this example in Figure \(\PageIndex{7}\), protein P1 has an affinity for ligand Z and will bind to the column, while proteins P2 and P3 will pass through the column. Protein P1 can then be eluted from the column using high concentrations of free ligand Z.

In vitro peptide synthesis for antibody production

For more details on the chemistry of in vitro peptide synthesis, click the link below.

Answer

When making anti-peptide antibodies that recognize target proteins or studying an isolated peptide by itself, it is more difficult to isolate and purify a peptide from its original protein than to synthesize it in the lab using solid-phase synthesis. We describe this technique below.

Peptides are chemically synthesized by the condensation reaction of the carboxyl group of one amino acid to the amino group of another. Two chemical challenges must be addressed. The formation of an amide bond between the carboxylic acid of one amino acid and the amine of the other is thermodynamically unfavorable, so the carboxyl end must be activated typically by the reaction of the incoming amino acid with a reagent such as dicyclocarbodiimide. Secondly, reactive functional groups on the side chains and the amine of the carboxyl group-activated amino acid must be protected from unwanted reactions. Chemical peptide synthesis most commonly starts at the carboxyl end of the peptide (C-terminus) and proceeds toward the amino-terminus (N-terminus). Protein biosynthesis in living organisms occurs in the opposite direction. Chemical synthesis facilitates the production of peptides that incorporate unnatural amino acids, peptide/protein backbone modification, and the synthesis of D-amino acids.

The established method for producing synthetic peptides in the lab is known as solid-phase peptide synthesis (SPPS). Pioneered by Robert Bruce Merrifield, SPPS allows the rapid assembly of a peptide chain through successive reactions of amino acid derivatives on an insoluble porous support. The solid support consists of small, polymeric resin beads functionalized with reactive groups (such as amine or hydroxyl groups) that link to the nascent peptide chain. Since the peptide remains covalently attached to the support throughout the synthesis, excess reagents and side products can be removed by washing and filtration. This approach circumvents the comparatively time-consuming isolation of the product peptide from the solution after each reaction step, which would be required when using conventional solution-phase synthesis.

Each amino acid to be coupled to the peptide chain N-terminus must be protected on its N-terminus and side chain using appropriate protecting groups such as t-Boc (t-butyloxycarbonyl-, acid-labile) or fluorenylmethyloxycarbonyl (Fmoc, base-labile), depending on the side chain and the protection strategy used (see below).

The general SPPS procedure involves repeated cycles of alternate N-terminal deprotections and coupling reactions. The resin can be washed between each step to remove side products. Figure \(\PageIndex{8}\) shows the mechanism for the solid phase synthesis of a dipeptide.

Figure \(\PageIndex{8}\): Solid phase peptide/protein synthesis

A. Deprotection of AA₁: The first amino acid is coupled to the resin or purchased pre-coupled. The amine terminus contacting an FMOC group is deprotected with piperidine. The hydrogen abstracted from the FMOC is acidic as its negatively charged conjugated base is aromatic since the negative charge on that C becomes sp² hybridized to create the aromatic anion. The weak base piperidine is used to avoid side reactions.

B. Activation of AA₂: The carboxyl group of AA₂ reacts with a carbodiimide, which is attacked by the carboxylate of AA_2, leading to the formation of an isourea derivative. This can react with a second nucleophilic catalyst (which is regenerated in step C), hydrobenzotriazole (HBT), to form the activated HBT ester and the very stable urea derivative.

C. Coupling Reaction: The activated AA₂ now reacts with the amine of solid phase N-terminal deprotected AA1 to form the peptide bond.

This cycle repeats until the desired sequence has been synthesized. At the end of the synthesis, the crude peptide is cleaved from the solid support while simultaneously removing all protecting groups using a reagent strong acid like trifluoroacetic acid. The crude peptide can be precipitated from a non-polar solvent like diethyl ether to remove organic soluble by-products and then purified using reversed-phase HPLC. The purification process, especially of longer peptides, can be challenging because small amounts of several byproducts, which are very similar to the product, must be removed. For this reason, so-called continuous chromatography processes such as MCSGP are increasingly being used in commercial settings to maximize the yield without sacrificing purity levels.

Reaction yields limit SPPS, and typically, peptides and proteins with 70 amino acids or more push the limits of synthetic accessibility. Synthetic difficulty is also sequence-dependent; aggregation-prone sequences, such as amyloids, are typically difficult to make. Longer lengths can be accessed by using ligation approaches, such as native chemical ligation, where two shorter, fully deprotected synthetic peptides can be joined together in solution.

 

Cell proteins can be increasingly engineered by manipulating their gene to contain a molecular tag, either a small peptide or a protein for which antibodies are commercially available. The tag is expressed at either the target's N- or C-terminal end to not interfere with the folding of the expressed target protein. Examples of peptide tags include the His (sequence HHHHHH), FLAG (sequence DYKDDDDK), and HA (YPYDVPDYA) tags. The HA tags derive from the influenza hemagglutinin protein. A small protein, such as the green Fluorescent Protein - GFP), can also be used as a tag. The resulting fusion protein of GFP connected to the target protein can also allow the target protein to be localized and followed by confocal fluorescence microscopy within the cell. Chromatography resins with covalently attached antibodies to the His, FLAG, HA peptide tags, and GFP are commercially available as affinity chromatography resins as shown in the right-hand side of Figure \(\PageIndex{9}\): below.

Figure \(\PageIndex{9}\): Protein Tags and small molecule bead ligands for affinity chromatography

Affinity reagents other than antibodies can be attached to the beads, as shown in the left-hand side of Figure \(\PageIndex{9}\). Two, in particular, are Ni-Nitrilotriacetic acid (Ni-NTA) and the short peptide glutathione (γ-gluatmylcysteinylglycine). They also bind tagged proteins. The Ni-Nitrilotriacetic binds the His tag by chelating the nickel ion with the 6 histidine imidazole groups on the His-tagged protein. (Note that His tags can also be bound to anti-His tag antibody beads.) Glutathione binds a protein tag, Glutathione-S-Transferase (GST), linked in a fusion protein to the target.

The structure of the Ni²⁺-NTA complex attached to a bead and imidazoles (on a His₆ tag) are shown in Figure \(\PageIndex{10}\) below.

 

Figure \(\PageIndex{10}\): Ni²⁺-NTA - imidazole complexes (after Wegner and Spatz,  https://doi.org/10.1002/anie.201210317) 

Figure \(\PageIndex{11}\) shows an interactive iCn3D model of the Ni²⁺ NTA complex.

Figure \(\PageIndex{11}\): iCn3D model of the Ni²⁺-NTA complex. To see the Ni²⁺ in spacefill, follow these commands in the iCn3D window.  Click the = menu icon and then:

• Select, Select on 3D, choose Atom
• Alt-click the central Ni ion
• Style, Chemical, Sphere

Without a His₆ tag, two water molecules occupy adjacent axial and equatorial positions (shown interacting with imidazole rings of His in the images above) in the generally octahedral complex.  One ring C in the iCn3D models shows a carbon with an unfilled valence.  The bead is covalently attached to that carbon.

The His tag, probably the most widely used, binds strongly to divalent metal ions such as nickel and cobalt. The protein can be passed through a column containing Ni-nitrilotriacetic. All untagged proteins pass through the column. The protein can be eluted with imidazole, which competes with the imidazole side chain on the His tag for binding to the column, or by a decrease in pH (typically to 4.5), which decreases the affinity of the tag for the resin. While this procedure is generally used to purify recombinant proteins with an engineered affinity tag (such as a 6xHis tag), it can also be used for natural proteins with an inherent affinity for divalent cations.

Hydrophobic Interaction Chromatography (HIC)

HIC media is similar to reverse phase chromatography in which a matrix like silica (very polar with exposed OH groups) is derivatized with ester or ether links from the silica surface hydroxyl OHs to nonpolar molecules, usually containing 8 or 18 carbons in the acyl or alkyl chain. Proteins with exposed hydrophobic groups would preferentially bind to the bead. The interactions of the protein with the derivatized beads are increased by adding high concentrations of salt to the aqueous solution, making water effectively more polar. This would shift the equilibrium towards binding the surface-exposed nonpolar region on the protein to the nonpolar C8 or C18 chains. The ionic strength of the buffer is then reduced to elute proteins in order of increasing hydrophobicity, as shown in Figure \(\PageIndex{11}\).

Figure \(\PageIndex{11}\): Hydrophobic Interaction Chromatography.

The column matrix, shown in blue, has a hydrophobic ligand covalently attached. In high salt conditions, proteins will bind to the matrix with differing affinity, with more hydrophobic proteins (shown in yellow) binding more tightly than more hydrophilic proteins (shown in green) When the salt concentration is decreased, more hydrophilic proteins will be released first, followed more hydrophobic proteins.

High Performance Liquid Chromatography (HPLC) and Fast Protein Liquid Chromatography (FPLC)

High-performance liquid chromatography or high-pressure liquid chromatography (HPLC) applies high pressure to drive the solutes through the column faster than the gravity-forced flow of solvent. The small and close-packed packing beads allow greatly increased resolution. Because of the close packing of the small beads, no flow would occur with an external pump. The most common form of HPLC is "reversed phase" HPLC, where the column packing material is hydrophobic. The proteins are eluted by a gradient of water and increasing amounts of an organic solvent, such as acetonitrile. The proteins elute according to their hydrophobicity. After purification by HPLC, the protein is in a solution that only contains volatile compounds and can easily be lyophilized (freeze-dried). HPLC purification frequently results in the denaturation of the purified proteins and is thus not applicable to proteins that do not spontaneously refold.

Due to the drawbacks of HPLC, an alternative technique using lower pressure was developed called Fast protein liquid chromatography (FPLC). In FPLC, the mobile phase is an aqueous solution, or "buffer". A positive displacement pump controls the buffer flow rate and is normally kept constant. In contrast, the composition of the buffer can be varied by drawing fluids in different proportions from two or more external reservoirs. The stationary phase is a resin composed of beads, usually of cross-linked agarose, packed into a cylindrical glass or plastic column. Depending on the application, FPLC resins are available in a wide range of bead sizes and surface ligands.

An ion exchange resin is typically chosen in the most common FPLC purification systems, as shown in Figure \(\PageIndex{12}\).

Figure \(\PageIndex{12}\): Typical FPLC System. A. Scheme of basic components and typical flow path for a chromatography system. B. Picrue of GE Healthcare AKTA FPLC apparatus. Image provided by LaVerde, V., Dominici, P. and Astegno, A. (2017) Bio-protocol 7(8): e2230.

A mixture containing one or more proteins of interest is dissolved in 100% buffer A and pumped into the column. The proteins of interest bind to the resin while other components are carried out in the buffer. The total flow rate of the buffer is kept constant; however, the proportion of Buffer B (the "elution" buffer) is gradually increased from 0% to 100% according to a programmed change in concentration (the "gradient"). Buffer B contains high concentrations of the exchanger ion. Thus, as Buffer B's concentration gradually increases, bound proteins will dissociate depending on their ionic interactions with the column matrix and appear in the eluant. The eluant passes through two detectors, which measure salt concentration (by conductivity) and protein concentration (by absorption of ultraviolet light at a wavelength of 280 nm). As each protein is eluted, it appears in the eluant as a "peak" in protein concentration and can be collected for further use.

Purification Scheme

During the protein purification process, a quantitative system is necessary to determine the total amount and concentration of total and target protein at each step, the biological activity of the target protein, and its overall purity. This will help guide and optimize the purification method being developed. Ineffective separation techniques can be disregarded, and other techniques that give higher yields and retain the protein's biological activity can be adopted.

Thus, each step in the purification scheme is quantitatively evaluated for the following parameters: total protein, total activity, specific activity, yield, and purification level. The terms are described below.

Pretend you are a researcher who wants to isolate a novel, unknown protein from a bacterial culture. You grow 500 ml of the bacteria overnight at 37^oC and harvest the bacteria by centrifugation. You remove the culture broth and retain the bacterial pellet. You then lyse the bacteria using freeze/thaw in 10 mL of reaction buffer. You then centrifuge the lysed bacteria to remove the insoluble materials and retain the supernatant containing the soluble proteins. Your protein of interest has a biological activity you can measure using a simple assay that causes a color change in the reaction mixture, as illustrated in Figure \(\PageIndex{13}\). You also note that this reaction rate increases with increasing concentrations of your protein supernatant.

Figure \(\PageIndex{13}\): Example of a Chemical Reaction that causes a color change from orange to brown depending on increasing concentration. Image from: Ludwig, N., et. al. (2015) on Research Gate

At this point, you can measure your baseline concentrations for the first purification level (bacterial lysis and removal of insoluble proteins and other cellular debris by centrifugation).

Total Protein is calculated by measuring the concentration in a fraction of the sample and then multiplying that by the total volume of your sample. In this case, you are starting with 10 mL of supernatant. In a typical assay to measure protein concentration, you will use 50 - 200 μL of sample to determine the protein concentration. For example, if you calculate that there is 7.5 μg/μL in your initial assay, you would need to convert that value into mg/mL and then multiply it by 10 mL for a total of 75 mg of protein in 10 mL of supernatant (Table \(\PageIndex{1}\)).

Total Activity is measured as the enzyme activity within the assay multiplied by the total volume of the sample. For example, you might use 5 to 50 μL of the sample in your biological reaction. If you calculated the activity in your assay to be 2.5 units/μL, this would be equivalent to 2,500 units/mL or 25,000 units/10 mL of supernatant. Note that the enzyme unit, or international unit for the enzyme (symbol U, sometimes also IU), describes the enzyme's catalytic activity. 1 U (μmol/min) is defined as the amount of the enzyme that catalyzes the conversion of one micromole of substrate per minute under the specified conditions of the assay method.

Specific Activity is measured by dividing the Total Activity by the Total Protein. In our example, 25,000 units divided by 75 mg of protein = 333.3 units/mg.

Yield measures the biological activity retained in the sample after each purification step. The amount in the first step is set to be 100%. All subsequent yield steps will be evaluated using the first purification step. It is calculated by dividing the current step's total activity by the first step's total activity and multiplying it by 100.

The Purification level evaluates the purity of the protein of interest by dividing the specific activity calculated after each purification step by the specific activity of the first purification step. Thus, the first step always has a value of 1. A typical purification analysis scheme is shown below in Table \(\PageIndex{1}\).

Table \(\PageIndex{1}\): A typical purification analysis scheme.

Note that after each purification step, the total protein goes down, as you separate the target protein from other proteins in the mixture. Total Activity also goes down with each purification step, as some of your protein of interest is also lost at each purification step because (1) some protein will stick to the test tubes and glassware, (2) some protein won't bind with 100% efficiency to your column matrix, (3) some protein may bind too tightly to be removed from the column matrix during elution, and (4) some protein may be denatured or degraded during the purification process.

The overall percent yield (or recovery) for each purification step represents the amount of your protein of interest that is retained. Alternative purification methods should be explored if the percent yield is too low.

Note that in a good protein purification scheme, the specific activity should go up substantially with each level of purification, as the amount of your protein of interest makes up a greater percentage of the total protein within that fraction. If the specific activity only increases modestly within a purification step, or if it decreases during a purification step, this could indicate that (1) your protein of interest is being substantially lost at that step, (2) your protein of interest is being denatured or degraded and is no longer biologically active, or (3) that a required cofactor or binding protein is being reduced at that purification step. Additional experiments may need to be conducted to determine which causes predominate so that steps can be taken to reduce protein inactivation. For example, many proteins are temperature sensitive and will degrade or denature at room temperature. Completing purification steps on ice can often reduce degradation.

The fold-increase (specific activity at step n/specific activity initial) or purification level should increase during purification. A high-quality purification procedure results in a high fold increase or purification level AND a high recovery or yield.

Electrophoresis: Separation and Analysis

In column chromatography, flow through the column is driven by hydrostatic pressure, causing flow from higher regions of higher pressure at the top of the column reservoir to lower pressure (drops eluting from the bottom of the column). Ultimately, the hydrostatic pressure (in columns not driven by mechanical pumps) derives from the gravitational force. However, proteins are also charged particles and can be moved by an external electric field instead of a gravitational field. Electrophoresis is the movement of charged particles in an electric field. As we will show below, the movement of a charged protein within a static matrix in the presence of an external electric field depends on both size and shape. Electrophoresis can be used for both analytical and preparative separations of proteins. The most common uses are for analytical separations.

Theory

What determines how a protein moves in an electric field? Consider a charged particle (+Q) moving in an electric field (E) in a nonconducting medium, such as water. Suppose the particle is moving at a constant velocity toward the cathode (- electrode where cations go). In that case, the net force F_tot on the particle is 0 (since F=ma, and the acceleration (a) of the particle is zero at constant velocity). Two forces are exerted on the particle, one F_E, the force exerted on the charged particle by the field, which is in the direction of the motion (toward the cathode), and the other, F_f, the frictional force on the charged particle, which retards its motion toward the cathode, and hence is in the direction opposite to the motion (toward the anode (+) electrode). This is shown in the Figure \(\PageIndex{14}\):

Figure \(\PageIndex{14}\): Movement of charge Q⁺ in an electric field

Therefore:

\begin{equation}
\mathrm{F}_{\text {tot }}=\mathrm{F}_{\mathrm{E}}+\mathrm{F}_{\mathrm{f}}
\end{equation}

where F_E, the electrical force, is

\begin{equation}
\mathrm{F}_{\mathrm{E}}=\mathrm{QE}
\end{equation}

and

F_f, the frictional force, is

\begin{equation}
\mathrm{F}_{\mathrm{f}}=-\mathrm{fv}
\end{equation}

In the last equation, v is the particle's velocity, and f is a constant called the frictional coefficient. This equation shows that the force F_f hindering motion toward the cathode is proportional to the particle's velocity. This is intuitive since one would expect the higher the velocity, the greater the F_f, which would hinder the motion. The frictional coefficient depends on the size and shape of the molecule. The larger the molecule, the larger the frictional coefficient (i.e. more resistance to the molecule's motion). It can be shown that the frictional coefficient for a spherical particle is given by

\begin{equation}
\mathrm{f}=6 \pi \eta \mathrm{R}_{\mathrm{S}}
\end{equation}

where η is the viscosity (a measure of the resistance to flow of a liquid - water has a low viscosity, real maple syrup a high viscosity), and R_s (Stokes radius) is the radius of the hydrated sphere (the larger R_s, the larger the frictional coefficient, the larger the F_f which resists motion toward the cathode). This equation should be intuitive based on your experiences.

When the velocity of the charged particles is constant (i.e, there is no net force on the particle), F_e = F_f, and using equations 3.3.6-8 gives

\begin{equation}
\mathrm{QE}=\mathrm{fv}
\end{equation}

Hence v/E = Q/f = U = the electrophoretic mobility, or

\begin{equation}
\mathrm{U}=\frac{\mathrm{V}}{\mathrm{E}}=\frac{\mathrm{Q}}{6 \pi \eta \mathrm{R}_{\mathrm{S}}}
\end{equation}

Therefore, the electrophoretic mobility U is proportional to the particle's charge density (charge/size, Q/Rs), not just the size, as is the case for spherical proteins in size exclusion chromatography. Macromolecules of different charge densities can thus be separated by electrophoresis. This discussion deals with the simplest case since, in reality, there are counter ions in the solution (from salts), which would form a cloud around the charged macromolecule and partially shield the charged particle from the electric field E.

Modern-day electrophoresis is conducted in solid gels (such as polyacrylamide) formed from liquid acrylamide solutions after adding a polymerizing agent. The solid gel is porous to solute and solvent molecules. It serves as a medium for electrophoresis while helping to eliminate convection forces in the liquid that interfere with the separation. Electrophoretic experiments have been conducted on the space station in weightless conditions to prevent such perturbations.

One complication that affects this idealized description of electrophoresis in polyacrylamide gels is that the gels have pores through which the macromolecules move. Think of the protein moving under an electric force through a "spider web-like" matrix. As in gel chromatography, the smaller molecules can pass through the pores more readily than larger molecules, so an additional sieving mechanism contributes to the effective mobility. (Also, the gel could alter the local effective electric field). The sieving effect of the gel increases the resolving power of this technique.

It has been determined that the actual electrophoretic mobility of the protein, U, is a function of the mobility of the protein in a concentrated sucrose solution (Uo) and T, the total concentration of the acrylamide in the polymerized gel. The higher the concentration of acrylamide in the unpolymerized gel solution, the smaller the size of the pores in the polymerized gel. An equation showing the relationship between U, Uo, and T is shown below:

\begin{equation}
\log \mathrm{U}=\log \mathrm{U}_{0}-\mathrm{K}_{\mathrm{r}} \mathrm{T}
\end{equation}

where K_r is the slope of a log U vs T plot for a given protein. Since K_r is a function of the radius of the molecule, it is possible to determine the molecular weight of a protein molecule by performing several electrophoretic separations in gels of different acrylamide concentrations (T) and extrapolating results to T = 0, hence eliminating pore size effects. Problems arise, however, if the proteins are not spheroid in shape.

Is there any way to obtain molecular weight information, in addition to purity determination, on a single gel? What would result if two different proteins with the same molecular weight and total net charge but different shapes were run on a single acrylamide gel? The more elongated shape (large Stokes radius) would have lower electrophoretic mobility (U = Q/6πηR_s). A larger Rs would also cause the protein to enter the pores more slowly. Hence, electrophoretic mobility and sieving effects would cause this protein to run anomalously slowly and have a higher apparent molecular weight. Also, imagine two globular proteins of different sizes but with compensatory charge differences, which might allow the two proteins to migrate at the same speed in the gel.

An astute reader might quickly recognize a problem separating proteins by gel electrophoresis. Some proteins are negatively charged (pH > pI), some would be neutral (pH=pI), and the rest would be positive (pH < pI). Only some proteins would enter the gel and move to the electrode at the bottom of the gel. Luckily, there is a way to eliminate both charge and shape effects in the electrophoresis of proteins, and that is to run the gel under denaturing conditions when all proteins have the same charge density. The denaturant of choice for electrophoresis is usually sodium dodecyl sulfate (SDS), which is an ionic detergent with the structure CH₃(CH₂)₁₀CH₂OSO₃^- (a single chain amphiphile). This detergent binds to and denatures most proteins, with about 1.4 g SDS binding/g of protein (about 1 SDS/2 amino acids). Since there is 1 negative charge/SDS, the binding of SDS masks any of the charges on the protein and gives all proteins an overall large negative charge. Additionally, SDS-proteins complexes have been shown to have an elongated cylindrical-like shape generally. Since the amount of SDS bound per unit protein mass is constant, the overall charge density on all proteins is similar, so the electrophoretic mobility is only determined by sieving effects.

SDS also eliminates shape differences in the proteins as a variable since all proteins have the same general rod-like shape. (The use of SDS is analogous to using 8M urea in the gel chromatographic separation of proteins to determine molecular weights). Mobility becomes only a function of the molecular weight of the protein and not its shape. The molecular weight of an unknown protein can be determined by comparing the protein's position on an SDS polyacrylamide gel with a series of known molecular weight standards from which a linear plot of the ln Mr vs R_f can be used to calculate unknown molecular weights. This is similar to the analysis in gel chromatography, where ln Mr is a linear function of K_avg, the distribution coefficient when the gel is run under denaturing conditions. However, some proteins run anomalously on such gels (due to incomplete or excess binding SDS), so alternative techniques of molecular weight determination should be used in conjunction with this technique.

Proteins are usually heated in SDS to 100 ^oC for 3 minutes, in the presence of a reducing agent such as β-mercaptoethanol (βME), to denature the protein to a rod-shaped protein. Apparent molecular weight can be obtained under non-reducing conditions (without βME), but these should be considered just estimates. Running proteins, both in the presence and absence of the reducing agent, can provide important information on the subunit structure of a protein. A multimeric protein whose subunits are held together by disulfide bonds can be resolved into component subunits by adding the reducing agent. If the subunits are held together only by noncovalent interactions, they will separate under denaturing conditions (SDS), eliminating subunit interactions in the presence or absence of β-ME.

Electrolytic vs Galvanic Cells

Electrode nomenclature can be confusing. In the above example, in the presence of an existing electric field produced by a power source, the +Q particle, a cation, moved to the cathode, the negative electrode.  In protein or DNA electrophoresis, you know that the negatively charged proteins or DNA fragments, which are anions, move to the bottom of the gel towards the anode, the positive electrode.  However, when you studied galvanic cells (that spontaneously produce current without a power supply) in general chemistry courses, the cathode is the positive electrode, and the anode is the negative electrode.  Instead of remembering the charge state of the electrode, it is better to focus on the redox reactions that occur at each electrode: 

• It is always true that Oxidation occurs at the Anode - both start with a vowel - and Reduction occurs at the Cathode - both start with a consonant! 

In galvanic cells, an electrical current is generated from a spontaneous set of redox half-reactions.  In electrolytic cells, an external power supply is required to drive the motion of the particle and the nonspontaneous redox reactions that occur at the electrodes. In electrophoretic cells for protein analysis, the bubbles released at the electrodes are from the electrolysis of water - 2H₂O(l) → 2H₂ (g) + O₂(g).  The oxidation number of H goes from +1 to 0 in the reaction so H is reduced and is produced at the cathode. In SDS-PAGE, SDS-coated proteins (negatively charged) migrate to the anode, the positive electrode in this case.  Commercial electrolytic cells can produce Cl₂(g) and Mg(s) from the aqueous electrolyte MgCl₂(aq).  Figure \(\PageIndex{15}\) illustrates the differences between galvanic and electrolytic cells.

Figure \(\PageIndex{15}\): Galvanic vs electrolytic cells

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Lithium Ion Batteries

Optional:  To support the material in Chapter 32:  Biochemistry and Climate Change, to review your learning in previous courses, to prepare for more complex redox reactions (mitochondrial electron transport, photosynthesis, and nitrogen fixation) that we will encounter later in this book, and as we move to electric vehicles (EVs) to reduce greenhouse gases from burning fossil fuels, it's important to know how Li⁺ ion batteries that power EVs work.  All you need is basic general chemistry, whose ideas we need to understand biochemical redox reactions.

Click the link below if you wish to learn more about lithium ion batteries

The diagram below shows the components of a Li⁺ ion battery that discharges to power electronic devices (in the case of a light bulb) and electric cars.  Compare it to the spontaneously discharging galvanic/voltaic cell (left image above).

Schematic of the involved species/charge transports and electrochemical reactions in lithium-ion cells during discharge.  Lithium atoms are shown as red spheres, Li⁺ ions as light orange, and electrons as light green.   Jiang, F., Peng, P. Elucidating the Performance Limitations of Lithium-ion Batteries due to Species and Charge Transport through Five Characteristic Parameters. Sci Rep 6, 32639 (2016). https://doi.org/10.1038/srep32639.  Creative Commons Attribution 4.0 International License. http://creativecommons.org/licenses/by/4.0/

In a lithium-ion battery:

* The - electrode (anode, where oxidation occurs) is porous carbon graphite (parallel sheets of graphene) with Li atoms/ions on a conducting metal (Cu). This is where Li⁺ ions and free electrons (e^-) are formed.  The electrons move through the electric circuit to the + electrode (the cathode);

* A liquid electrolyte solution (replaces the "salt bridge" in the Cu/Zn²⁺ cell) allows the transfer of Li⁺ to the + cathode (the tricky part to remember) to preserve charge neutrality.

* The + electrode (cathode, where reduction occurs) is a metal oxide (such as CoO₂) on a conducting metal (Al), which reduces the arriving Li⁺ ions to neutral L and intercalation into the CoO₂ polymer. This is the reaction shown in the figure above.  Most references state that the Co(IV) in CoO₂ is reduced to Co(III), which is perhaps more likely given the low ionization energy of the gaseous form of Group 1 Li. 

Here are the actual chemical reactions that occur at the anode and cathode on the discharge of the battery to power a light or an electric car:

Li atoms (neutral, red spheres) in the graphite anode are oxidized to Li⁺ ions (yellow spheres) as shown in this reaction: 

Oxidation at anode:  LiC₆ → C₆ + Li⁺ + e^- 

The Li⁺ ions move from the anode through the electrolyte to the cathode side, where they get reduced (like Cu²⁺ in the left figure above).  Here is the reaction:

Reduction (at cathode): Li⁺ + e^- + CoO₂ → LiCoO₂

The last reaction could indicate that either Li⁺ or Co(IV) is reduced to maintain charge balance. The oxidation number of Co in CoCO₂ is +4 (as each O is -2).  In LiCoO₂, if Co is reduced, it becomes Co +3 as it gains an electron, with the +1 on Li balancing the net -4 on the two oxygens. If Li⁺ is reduced, it becomes atomic Li⁰.  This presumes that the interactions are all ionic. So with charge states/oxidation numbers, this could be written as:

Li⁺ + e^- + Co(IV)(O^2-)₂ → [Li⁺Co(III)(O^2-)₂ or Li⁰Co(IV)(O^2-)₂]

A better way might be to envision a sharing of the electron between the Li⁺ and Co(IV) ions.  We'll address this more below.

On charging (nonspontaneous, driven by plugging in the car), the reverse process occurs as reduced Li in LiCoO₂ gives up an electron (as Zn in the left figure above) to form Li^+, which moves to the graphite electrode. Here is an animation (which sometimes works on refreshing the browser) showing the structure and Li-ion diffusion out of a LiCoO2 cathode (click the Lithium Ion Diffusion Icon).

The figure below shows the 3D structures of different graphite forms with intercalated Li in the anode. 

The crystal structure of (a) and (b) graphite, (c) and (d) C₁₂Li, and (e) and (f) C₆Li. (b, d, and f) Show a c-axis view of graphite, C₁₂Li, and C₆Li, in which Li is located at the center of six-member carbon rings. Phys. Chem. Chem. Phys., 2017,19, 19058-19066. Creative Commons Attribution-NonCommercial 3.0 Unported

Here are two views of the LiC₆ anode structure (see this link for a 3D view).  Carbons in graphite are shown as gray spheres, while Li is shown as purple spheres.  Lithium migrates in a 2D plane between 2 graphene layers (graphene is a single layer of graphite).  Here is another link for the structure of LiC₆. 

 

 

The empirical formula for this structure is LiC_6, but it is also written as C₆Li.  If we consider LiC₆ as purely an ionic compound, the lithium would appear to have a +1 charge and oxidation state, and each of the Cs would be -1/6 = -0.166, with the electron from Li entering the pi cloud of graphite.  But didn't we say above that Li likely has a 0 oxidation number/charge (i.e., it's a neutral atom) in the LiC₆ structure?  

Here is some help for this problem.  Assigning electrons for charge state in metal ion complexes is arbitrary. It doesn't indicate the actual electron density around an atom (such as in the case of H₃O⁺,  in which O has a formal charge of +1 but has almost all of the electron density).  In the case of a transition metal complex, we can consider the bond between the metal and ligand to be a coordinate covalent bond. For a transition metal complex, the metal's oxidation number can be determined by first separating (in your mind) the ligand and its donating electrons from the metal.  There are 2 ways to do this:

* send both electrons in the coordinate covalent bond to the withdrawn ligand (ie. a heterolytic cleavage).  If the ligand is -NH₃, the removed ligand has a full lone pair (:NH₃) and is neutral.  For a -CH₃ ligand, the removed ligand would be :CH₃-, a carbanion with a negative charge.

* send one electron to the withdrawn ligand and one to the metal (i.e a homolytic cleavage).  If the ligand is -CH_3, the removed ligand is ^.CH₃ (free radical and neutral).

For the case of Li, a nontransition Group I metal, if we assign one electron in the bond to Li and the other to the separated C₆ electron cloud, the oxidation state of Li would be 0, consistent with the anode reaction shown above.  Given the delocalized electron density from the pi clouds of the graphite interacting with Li, a coordinate covalent bond is a better descriptor than a simple ionic one. At the same time, given how easily Li gives up an electron, it makes sense to show both Li and Li⁺ in the electrodes.

Here is a figure showing the LiCoO₂ cathode.

Lithium is shown in purple, oxygen in red, and Co in orange.  https://www.chemtube3d.com/lib_lco-2/

Much research is devoted to finding new electrons and metal substitutes to make them cheaper and more environmentally favorable.  Lithium manganese oxide (LiMn₂O₄) and lithium iron phosphate (LiFePO₄) are alternatives for the cathode, which don't rely on expensive and toxic cobalt.

 

---

Polyacrylamide Gel Electrophoresis - PAGE

Electrophoresis is performed in a porous yet solid medium to eliminate any problems associated with convection currents. Such media are formed from solidifying a liquid solution of agarose (used mostly for electrophoresis of DNA fragments and very large proteins) or the polymerization of a solution of acrylamide. Polymerization of acrylamide is initiated by the additions of ammonium persulfate in the presence of tetramethylenediamine (TEMED), along with a dimer of acrylamide (N,N'-methylene-bis(acrylamide) connected covalently between the amide nitrogens of the acrylamides by a methylene group. The structures of these compounds are shown in Figure \(\PageIndex{16}\).

Figure \(\PageIndex{16}\): Structures of reagents use to make PAGE gels

As shown above, the acrylamide's free radical polymerization is initiated by adding ammonium persulfate, which forms free radicals on dissolving in water.

The radical initiates polymerization of the acrylamide, as shown below. The TEMED, through its ability to exist as a free radical, acts as an additional catalyst for polymerization. A rigid gel is only formed, however, when N,N'-methylene-bis(acrylamide) is added to the mixture during the polymerization, which cross-links adjacent acrylamide polymers as shown in Figure \(\PageIndex{17}\).

Figure \(\PageIndex{17}\): Polymerization of acrylamide

The amount of bisacrylamide added during polymerization controls the degree of cross-linking and, hence, the pore size of the polymerized gel. The effect of pore size is OPPOSITE to that in gel chromatography. In both cases, large proteins have a difficult time entering the pore. In gel chromatography, large proteins partition preferentially into the mobile liquid phase (the void volume) and are eluted most QUICKLY from the column. In electrophoresis, large proteins, which can not readily enter the pores in the gel, are not as easily transported by the electric field through the gel and elute most SLOWLY. Pore size can not be controlled as accurately as in the manufacture of gel chromatography resins.

How do proteins migrate through the gel? A viscous protein solution is layered on the top of the gel in a small well molded into the gel during the polymerization process. The bottom and top parts of the gel are inserted into reservoirs containing a buffered solution and the appropriate electrode. The electric field is applied, and the proteins migrate through the hydrated gel. The nature of the buffer solution in the reservoir and the polymerized gel is important. The buffer components must not bind to the proteins to be separated. Additionally, for native (non-denatured gels), the pH of the medium must be such that the proteins have the appropriate charge so they will migrate in the expected direction.

There are many variations of electrophoresis commonly used. Gels can be polymerized in tubes or slabs and the presence or absence of denaturing agents. Additionally, a given slab might consist of two separate slabs polymerized on top of the other, each with a different acrylamide concentration and pH value. The top part is the stacking gel, the bottom is called the running gel. Other gels have a continuous gradient of acrylamide concentrations (from low at the top to high at the bottom). Most commercially available precast gels use continuous acrylamide concentration gradients. Figure \(\PageIndex{18}\) shows a gel placed in an electrophoresis chamber.

Figure \(\PageIndex{18}\): Electrophoresis gel and chamber https://openwetware.org/mediawiki/in...&oldid=1096467

Whether the gel has a continuous gradient or is discontinuous, the top part of the gel is a low concentration acrylamide (2-4%), often in a Tris HCl buffer solution (pH 6.5), usually 2 pH units below that used in the running gel. The lower part of the gel is 8-15% acrylamide, depending on the choice of gel, which is selected based on the molecular weight of the proteins to be separated. The upper buffer reservoir contains Tris-buffered with a weak acid such as glycine (pKa2 = 9.6) to the same pH as the running gel.

Proteins electrophorese quickly through the low-concentration stacking gel at the top of continuous gradient gels and effectively "stack" as they hit the interface between the stacking and running gels or before they enter too far into the continuous gradient gel. This increases the compactness of the proteins before they enter the "running" section of the gel and increases resolution.

For discontinuous gels, how does this stacking process work? When the electrophoresis is started, glycine ions from the upper reservoir (at pH 8.7) enter the stacking gel since, at that pH, they have an average partial negative charge. The stacking gel buffer ions continue moving in the stacking gel, but when the glycine ions enter the pH 6.5 of the stacking gel, they become zwitterions with a zero net charge and hence stop their motion toward the anode. The electrical resistance in the stacking gel increases since the number of ions moving through the stacking gel decreases. To maintain a constant current throughout the circuit, there will be a localized increase in the voltage in the stacking gel (from Ohms Law, V=iR). This will cause the proteins to migrate quickly and all stack in a single, very thin disc right behind the Cl- ions in the stacking gel (which are in front because they have the highest charge density and electrophoretic mobility of any ion in the stacking gel). The proteins will not pass the Cl- ions since, if they did, they would immediately slow down since they would no longer be in an area of diminished charged carriers and higher voltage. At the stacking gel/running gel interface, the proteins can not all migrate at the same speed due to the sieving effects of the more concentrated gel.  They will be separated in the running gel. The glycine eventually enters the running gel, assumes its fully charged state at that pH (8.7), passes the proteins, and restores the deficiency in charge that occurred in the stacking gel.

Detection of proteins in the gel:

Most proteins do not absorb at visible wavelengths of light and hence will not be visible during electrophoresis. To ensure that the proteins are not eluted from the gel into the lower buffer reservoir, a small molecular weight, anionic dye, and bromophenol blue are added to the protein before it is placed on the gel. The electrophoresis is halted when the dye reaches the bottom of the gel. The gel assembly is removed from the electrophoresis chamber, the glass plates separated, and the gel washed into a series of solutions to render the banded proteins visible.

• Coomassie Brilliant Blue dye is the most common stain in labs. It is dissolved in a methanol/acetic acid solution, generating significant waste. Proteins bind this dye, with a concomitant spectral shift in the absorbance properties of the bound dye. The methanol and acetic acid in the dye solution also help to "fix" the proteins in the gel and prevent their diffusion into the solution. After the gel is stained, the background stain is removed with acetic acid/methanol, leaving the blue-colored protein bands. Some proteins will not be stained with Coomassie blue.  A "colloidal Coomassie Blue" stain doesn't use methanol or acetic acid, so it is commonly used.  A simple water solution can destain the gel.
• Silver staining: This involves the reduction of Ag(I) to elemental silver and its deposition by protein in the appropriate reaction solutions, much as in a photographic process. (Remember, in the BCA assay, peptide bonds reduce Cu(II) to Cu(I), which is chelated to BCA.) A developer and fixer solution is required. This technique is 10-50 X more sensitive than Coomassie Blue staining. Figure \(\PageIndex{19}\) shows gels stained with Coomassie Blue (A)and silver staining (B).

Figure \(\PageIndex{19}\): PAGE gels stained with Coomassie Blue (A) and silver stain (B) Chen Y-Y, Chen J-C, Lin Y-C, Kitikiew S, Li H-F, Bai J-C, et al. (2014) Endogenous Molecules Induced by a Pathogen-Associated Molecular Pattern (PAMP) Elicit Innate Immunity in Shrimp. PLoS ONE 9(12): e115232. https://doi.org/10.1371/journal.pone.0115232. Creative Commons Attribution License

• Modifying proteins with fluorescent or radioactive tags before electrophoresis: This offers greater sensitivity. After the electrophoresis of a radiolabeled protein, the gel can be dried and overlaid with X-ray film for periods as long as months, if necessary, to allow sufficient film exposure by a low-concentration protein. This visualization technique is called autoradiography.

Variations on polyacrylamide gel electrophoresis:

Isoelectric focusing: In this technique, a pH gradient is set up within the polyacrylamide gel or strip. This is accomplished by pre-electrophoresing a series of low molecular weight molecules containing amino and carboxyl groups called ampholytes, each with a different isoelectric point. When subjected to an electric field, the most negative of the species will concentrate at the anode, while the most positive will concentrate near the cathode. The remaining ampholytes will migrate in between as ampholytes migrate to their isoelectric point and set up a linear pH gradient in the gel.

Proteins initially in regions with a pH below its isoelectric point are positively charged and migrate toward the cathode. In contrast, those in a media with pH lower than its pI will be negatively charged and migrate towards the anode as shown below in Figure \(\PageIndex{20}\). The migration will lead to a region where the pH coincides with its pI. There, the protein will have a zero net charge and stop. Thus, amphoteric molecules are located in narrow bands where the pI coincides with the pH. In this technique, the point of application is not critical, as molecules will always move to their pI region. The stable pH gradient between the electrodes is achieved using a mixture of low molecular weight ampholytes whose pIs cover a preset pH range.

Figure \(\PageIndex{20}\): Isoelectric Focusing. A pH gradient is established in a gel before loading the sample. After the sample is loaded a voltage is applied. The protein will migrate to its isoelectric pH, at which they have no net charge. Image from Magdeldin, S.

2D electrophoresis: Two-dimensional gel electrophoresis (2-DE) is based on separating a mixture of proteins according to two molecular properties, one in each dimension. The most used is based on a first dimension separation by isoelectric focusing (IEF) and a second dimension according to molecular weight by SDS-PAGE. A conditioning step is applied to proteins separated by IEF before the second-dimension run. This process reduces disulfide bonds and alkylates the resultant sulfhydryl groups of the cysteine residues. Concurrently, proteins are coated with SDS for separation based on molecular weight. After the IEF, the tube or strip is placed across the top of a slab gel and subjected to SDS-polyacrylamide gel electrophoresis in a direction 90^o from the initial isoelectric focusing experiment. Figure \(\PageIndex{21}\) shows a stained 2D electrophoresis gel.

Figure \(\PageIndex{21}\): Two-Dimentional Gel Electrophoresis. Image from Magdeldin, S

In Figure \(\PageIndex{21}\), proteins of Chlamydomonas reinhardtii are resolved by 2-DE from preparative gels stained with MALDI-MS compatible silver reagent for peptide mass fingerprinting analysis. First dimension: isoelectric focusing in a 3-11 pH gradient. Second dimension: SDS-PAGE in a 12% acrylamide (2.6% crosslinking) gel (1.0 mm thick). Numbered spots marked with a circle correspond to proteins compared to be subsequently identified by MALDI-TOF MS. The MALDI-TOF MS analysis of protein sequences is discussed in more detail in Chapter 3.4.

One of the biggest problems in 2-DE is the analysis and comparison of complex mixtures of proteins. Currently, there are databases capable of comparing two-dimensional gel patterns. These systems allow automatic comparison of spots to identify those needed in the quantitative analysis precisely. Once interesting proteins are identified, they can be excised from gels, destained, and digested to prepare for mass spectrometry. This technique is known as peptide mass fingerprinting. The ability to precisely determine molecular weight by matrix-assisted laser desorption/ionization-time of flight mass spectrometry (MALDI-TOF MS) and to search databases for peptide mass matches has made high-throughput protein identification possible. Proteins not identified by MALDI- TOF can be identified by sequence tagging or de novo sequencing using the Q-TOF electrospray LC-MS-MS.

Western blotting: After a standard SDS-slab electrophoresis experiment, the gel is overlaid with a piece of nitrocellulose membrane. The sandwich of gel and filter paper is placed back into an electrophoresis chamber such that the proteins migrate from the gel into the nitrocellulose, where they irreversibly bind. This is illustrated in the figure below. Note, however, that in the absence of staining, the protein bands in either the PAGE gel or Western blot would not be visible. Standards (lane 5) would be visible if they were labeled with chromophores, as shown in Figure \(\PageIndex{22}\).

Figure \(\PageIndex{22}\): Western blotting of proteins

If a cell lysate were applied to a lane of a PAGE gel, the bands would appear as overlapping smears on the stained gel after staining with any technique. Western blots are useful because specific bands can be visualized (stained) on the nitrocellulose membrane using a detection system linked to an antibody that recognizes a specific target protein. This is illustrated in Figure \(\PageIndex{23}\).

Figure \(\PageIndex{23}\): Developing a Western blot

3D electrophoresis: To detect specific proteins in a 2D electrophoresis experiment, a 3^rd dimension of separation, a Western blot, could be performed on the PAGE gel and the nitrocellulose stained with an antibody specific to a target protein. That is illustrated in Figure \(\PageIndex{24}\).

Figure \(\PageIndex{24}\): "3D" electrophoresis - Isoelectric focusing, PAGE and Western blotting

Part A, isoelectric focusing, is followed by a PAGE gel (B). The red dots represent proteins that have undergone a post-translational modification in which a phosphate group has been added to tyrosine side chains (for example). Western blotting is performed in panel C, and staining is performed in panel D. The left blot in D uses an antibody that recognizes phosphorylated tyrosine side chains on protein. The right blot is D is sometimes called a Far Western blot. If the protein on the nitrocellulose membrane retains some 3D native structure or can be induced to refold, it can be probed on the blot by a protein that binds to the native form of the protein on the blot. In the example shown in panel D above, the p-Tyr-protein target on the nitrocellulose membrane recognizes a fusion protein of PTP-GST. GST is a protein tag for detection. PTP is a protein tyrosine phosphatase, an enzyme that hydrolyzes p-Tyr on specific phosphorylated target proteins.

Summary

This chapter provides a comprehensive overview of protein purification and electrophoretic analysis, two essential methodologies in biochemistry for isolating and characterizing proteins. The chapter begins by emphasizing the necessity of protein purification, given the vast diversity of proteins in cells and tissues. It describes how inherent differences—such as size, charge, hydrophobicity, and binding affinity—are exploited to isolate a single protein from complex biological mixtures.

Sample Processing and Initial Fractionation:
The process starts with sample processing where cells or tissues are disrupted using methods such as freeze/thaw cycles, sonication, or detergent treatment. Centrifugation is then used to separate soluble proteins from cellular debris. Special attention is given to preventing proteolysis by performing procedures at low temperatures and adding protease inhibitors.

Bulk Purification Techniques:
Precipitation using ammonium sulfate is introduced as an economical initial step to concentrate proteins while maintaining their native state, followed by dialysis to remove excess salt. The chapter then details various chromatographic techniques:

• Size Exclusion Chromatography (Gel Filtration): Separates proteins based on size and shape by exploiting differences in how they access the internal pore volume of the resin beads.
• Ion Exchange Chromatography: Separates proteins based on their net charge; proteins bind to charged resins and are eluted by altering the ionic strength or pH of the buffer.
• Affinity Chromatography: Uses specific ligand-receptor interactions (e.g., His-tag binding to Ni-NTA or antibody-antigen recognition) to achieve high selectivity in isolating the protein of interest.
• Hydrophobic Interaction Chromatography (HIC): Exploits differences in protein hydrophobicity by modulating salt concentrations to promote binding to nonpolar ligands on the resin. Advanced methods such as HPLC and FPLC are also discussed as ways to increase resolution and throughput in protein separation.

Quantitative Evaluation of Purification:
The chapter stresses the importance of tracking purification efficiency through quantitative metrics like total protein, total activity, specific activity, yield, and fold-purification. These measurements are crucial for optimizing purification protocols and ensuring the integrity of the target protein.

Electrophoretic Analysis:
The latter portion of the chapter focuses on electrophoresis, a technique that separates proteins based on their size, charge, and shape under the influence of an electric field. Both native and denaturing methods are covered:

• SDS-PAGE: By binding to proteins and imparting a uniform negative charge, SDS standardizes the electrophoretic mobility of proteins so that separation is primarily based on molecular weight.
• Isoelectric Focusing (IEF): Separates proteins based on their isoelectric points (pI) by establishing a pH gradient.
• Two-Dimensional Electrophoresis (2-DE): Combines IEF and SDS-PAGE to achieve high-resolution separation of complex protein mixtures. Detection methods, including Coomassie Blue and silver staining, as well as Western blotting and autoradiography for enhanced sensitivity and specificity, are also described.

This chapter integrates the theoretical and practical aspects of protein purification and analysis. It highlights the sequential nature of the purification process—from cell lysis to chromatography and electrophoretic separation—and underscores the importance of quantitative evaluation at each step to ensure high purity and activity of the isolated protein. This foundation is critical for subsequent structural, functional, and proteomic studies in biochemistry.

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