His-Tagged Protein Purification: Principles and Methods

By Dr. Zubair Khalid, DVM, MS, PhD · 2026-08-11

Introduction to His-Tagged Protein Purification

Recombinant protein production is a cornerstone of modern molecular biology, enabling the study of protein structure, function, and interactions. A critical bottleneck in this workflow is the purification of the target protein from the complex mixture of thousands of cellular proteins present in a crude lysate. To address this, researchers commonly fuse a short, engineered peptide sequence—the polyhistidine tag, or His-tag—to the protein of interest. This tag exploits a well-characterized biochemical interaction to achieve rapid, single-step purification.

A His-tag typically consists of six consecutive histidine residues (6xHis), though variations with eight or ten histidines exist. The tag is genetically encoded, meaning it is added to the gene of interest at the DNA level, either at the N-terminus or C-terminus of the coding sequence. When the recombinant gene is expressed in a host organism—most commonly Escherichia coli, but also yeast, insect, or mammalian cells—the resulting protein carries the His-tag as an integral part of its polypeptide chain.

The power of the His-tag lies in its ability to bind reversibly to immobilized metal ions. This forms the basis of Immobilized Metal Affinity Chromatography (IMAC), the standard method for His Tag Protein Purification. The interaction is strong enough to capture the protein from a crude lysate but can be gently reversed by adding a competitive molecule, allowing the purified protein to be eluted in a functional state. This combination of simplicity, versatility, and cost-effectiveness has made His-tag purification the most widely used affinity purification method in biochemistry.

What is a His-Tag?

A His-tag is a short amino acid sequence, usually six histidines (HHHHHH), engineered onto the N- or C-terminus of a recombinant protein. Histidine is an amino acid with a side chain (an imidazole ring) that has a unique chemical property: it can coordinate with certain transition metal ions, including nickel (Ni²⁺) and cobalt (Co²⁺). This coordination is the basis of the entire purification strategy.

The tag is introduced via standard molecular cloning. The DNA sequence encoding the histidines is included in the primer used for PCR amplification of the gene of interest, or it may already be present in a commercial expression vector. The choice of tag position (N-terminal vs. C-terminal) can affect protein folding and function, and is often determined empirically. For example, an N-terminal tag may interfere with signal peptide processing in secreted proteins, while a C-terminal tag may be inaccessible if the protein's C-terminus is buried in the folded structure.

Applications in Research and Industry

His-tagged proteins are used across a vast range of applications. In research, they are essential for:

• Structural biology: Producing milligram quantities of pure protein for X-ray crystallography, NMR spectroscopy, and cryo-electron microscopy. The purity achieved by IMAC is often sufficient for initial crystallization screens.
• Enzyme kinetics: Studying the catalytic activity of purified enzymes in defined in vitro systems.
• Protein-protein interaction studies: Using the purified tagged protein as bait in pull-down assays or for surface plasmon resonance (SPR) analysis.
• Antibody production: Generating antigens for immunization.

In industry, His-tagged proteins are used in the production of therapeutic proteins, diagnostic reagents, and industrial enzymes. The scalability of IMAC, from microcentrifuge tubes to large-scale chromatography columns, makes it suitable for both laboratory and manufacturing settings. The Custom Protein Purification services offered by many biotech companies are frequently built around His-tag technology.

The Mechanism of Immobilized Metal Affinity Chromatography

IMAC is a type of affinity chromatography that separates proteins based on their affinity for metal ions. The principle is straightforward: a chromatographic resin is chemically modified to carry a chelating group that tightly holds a metal ion. The His-tagged protein in a complex mixture binds to this immobilized metal, while untagged proteins flow through. The bound protein is then released by adding a competing molecule.

Histidine-Metal Coordination

The interaction between histidine and metal ions is a form of coordination chemistry. The imidazole side chain of histidine contains a nitrogen atom (Nε2) that has a lone pair of electrons. This lone pair can be donated to the empty d-orbitals of a transition metal ion, forming a coordinate covalent bond.

Nickel (Ni²⁺) and cobalt (Co²⁺) are the most commonly used metals because they have a high affinity for imidazole and are relatively inexpensive. The metal ion is not free in solution; it is itself held by a chelator that is covalently attached to the resin. The most common chelator is nitrilotriacetic acid (NTA), which occupies four of the six coordination sites on the Ni²⁺ ion. This leaves two coordination sites available for interaction with the histidine residues of the tag. A single 6xHis tag can occupy both of these sites, forming a stable complex. The geometry of this interaction is such that two consecutive histidines in the tag are sufficient for binding, but six provide much higher affinity and stability.

Cobalt-based resins, such as TALON, use a different chelator (carboxymethylaspartate) that coordinates Co²⁺ differently, leaving more coordination sites available. This results in a weaker but more specific interaction, which can lead to higher purity of the target protein, albeit with a lower binding capacity.

Role of Imidazole in Binding and Elution

Imidazole is a small molecule that is structurally identical to the histidine side chain. It acts as a competitive inhibitor of the His-tag–metal interaction. When imidazole is present in the buffer, it competes with the histidine residues for the coordination sites on the metal ion.

• Low imidazole concentration (e.g., 10–20 mM) is included in the binding and wash buffers. This concentration is sufficient to prevent non-specific binding of host proteins that have surface-exposed histidines, but low enough that the 6xHis tag (which presents six histidines in close proximity) can still bind effectively.
• High imidazole concentration (e.g., 200–500 mM) is used in the elution buffer. At this concentration, imidazole outcompetes the His-tag for the metal coordination sites, causing the tagged protein to dissociate from the resin and elute in the collected fractions.

The concentration of imidazole required for elution depends on the affinity of the tag for the resin. A 6xHis tag typically elutes at 150–250 mM imidazole from Ni-NTA, while a 10xHis tag, which has higher avidity, may require higher concentrations.

Key Components of His-Tag Purification

A successful His-tag purification requires careful selection of several components. Each plays a specific role in the overall efficiency and purity of the final product.

Expression Vectors with 6xHis Tag

The expression vector is the DNA construct that drives production of the His-tagged protein. Key features include:

• Promoter: A strong, inducible promoter such as the T7 promoter (in pET vectors) or the araBAD promoter (in pBAD vectors). The T7 system, which uses T7 RNA polymerase, is the most common for high-level expression in E. coli. Induction is typically achieved with isopropyl β-D-1-thiogalactopyranoside (IPTG) at a final concentration of 0.1–1.0 mM.
• Multiple Cloning Site (MCS): A region with unique restriction sites for inserting the gene of interest.
• Tag sequence: The coding sequence for the 6xHis tag, often followed by a linker sequence and a protease cleavage site (e.g., for thrombin, TEV protease, or Factor Xa) to allow tag removal after purification.
• Selectable marker: An antibiotic resistance gene (e.g., ampicillin, kanamycin) for maintaining the plasmid in the host cells.

The choice of vector also determines the fusion context. Some vectors place the His-tag at the N-terminus, others at the C-terminus, and some have dual tags. The Protein Expression and Purification workflow begins with choosing the right vector for the target protein's properties.

Nickel-NTA vs. Cobalt Resins

The resin is the solid support to which the metal ion is attached. Two main types are used:

Ni-NTA is the default choice for most applications due to its high capacity and low cost. Cobalt resins are preferred when purity is critical and the target protein is expressed at moderate levels, as they tend to bind fewer host proteins non-specifically. The choice of resin is a key decision in His-tag Purification Resin selection.

Buffers and Additives

The composition of the buffers used throughout the purification is critical. A standard set of buffers includes:

• Lysis buffer: 50 mM sodium phosphate (pH 8.0), 300 mM NaCl, 10 mM imidazole, plus protease inhibitors (e.g., 1 mM phenylmethylsulfonyl fluoride, PMSF) and often lysozyme (1 mg/mL) for bacterial cell lysis.
• Wash buffer: Same as lysis buffer, but with a slightly higher imidazole concentration (20–50 mM).
• Elution buffer: Same as lysis buffer, but with 200–500 mM imidazole.

The pH is typically maintained at 7.5–8.0. At this pH, the histidine residues (pKa ~6.0) are largely unprotonated, which is required for efficient coordination with the metal ion. Sodium chloride (NaCl) at 300 mM is included to reduce ionic interactions between the resin and non-specific proteins. Additives such as glycerol (5–10%) can stabilize proteins, and non-ionic detergents like Triton X-100 (0.1%) or Tween-20 can help prevent aggregation and reduce non-specific binding.

Step-by-Step Purification Protocol

The following is a standard protocol for purifying a His-tagged protein from E. coli under native conditions. All steps are performed at 4°C unless otherwise noted, to minimize proteolysis.

Cell Lysis and Clarification

1. Harvest cells: After induction and expression, pellet the bacterial culture by centrifugation at 6,000 × g for 15 minutes at 4°C. Discard the supernatant.
2. Resuspend the pellet in lysis buffer (typically 5 mL per gram of wet cell paste). Ensure the pellet is fully resuspended to avoid clumps.
3. Lyse the cells: The most common method is sonication. Use a probe sonicator on ice, with cycles of 10 seconds on, 10 seconds off, for a total of 3–5 minutes of sonication time. Alternatively, use a French press or enzymatic lysis with lysozyme (incubate at 4°C for 30 minutes with gentle shaking).
4. Clarify the lysate: Centrifuge at 20,000 × g for 30 minutes at 4°C to pellet cell debris and insoluble material. Carefully collect the supernatant, which contains the soluble His-tagged protein. Filter through a 0.45 µm syringe filter if the lysate is viscous or cloudy.

Binding to Resin

1. Equilibrate the resin: Add the appropriate volume of Ni-NTA resin (typically 1 mL of settled resin per 5–10 mg of target protein) to a gravity-flow column or a tube. Wash with 5–10 column volumes of lysis buffer to equilibrate.
2. Apply the clarified lysate: Add the lysate to the equilibrated resin. For batch binding, mix gently on a rotating platform for 30–60 minutes at 4°C. For gravity-flow columns, allow the lysate to flow through by gravity, collecting the flow-through.
3. Collect the flow-through: This fraction contains proteins that did not bind. It can be saved for analysis or troubleshooting.

Washing Steps

1. Wash with lysis buffer: Add 10–20 column volumes of lysis buffer (with 10 mM imidazole) to remove weakly bound proteins. Collect the wash fractions.
2. Wash with wash buffer: Add 10–20 column volumes of wash buffer (with 20–50 mM imidazole). This step removes proteins that bind non-specifically to the resin or the metal ion. Collect the wash fractions.

The goal of the washing steps is to remove all contaminating proteins while retaining the His-tagged protein on the resin. The imidazole concentration in the wash buffer is a critical parameter that must be optimized for each protein.

Elution with Imidazole

1. Apply elution buffer: Add 5–10 column volumes of elution buffer (with 200–500 mM imidazole). Collect fractions of 0.5–1 column volume each.
2. Monitor the elution: The His-tagged protein will typically elute in the first 2–3 fractions. The presence of protein can be monitored by measuring the absorbance at 280 nm (A₂₈₀) or by a rapid protein assay.
3. Pool the peak fractions: Combine the fractions containing the highest protein concentration.

After elution, the resin can be regenerated by washing with 5 column volumes of 0.5 M NaOH, followed by extensive washing with water and re-equilibration with lysis buffer. The purified protein can be further processed, such as by dialysis to remove imidazole, or by His Tag Labeling Purification for downstream applications.

Optimizing Binding and Elution Conditions

The standard protocol provides a starting point, but optimal conditions vary between proteins. Systematic optimization is often required to maximize yield and purity.

Imidazole Concentration in Wash and Elution

The imidazole concentration in the wash buffer is the most important variable to optimize. Too low a concentration (e.g., 10 mM) may not remove all non-specifically bound proteins. Too high a concentration (e.g., 100 mM) may cause the His-tagged protein itself to elute prematurely, reducing yield.

A common approach is to perform a small-scale test. Bind the lysate to a small amount of resin, then wash with increasing concentrations of imidazole (e.g., 10, 20, 40, 60, 80 mM), collecting each wash separately. Analyze the fractions by SDS-PAGE to determine the highest imidazole concentration that does not elute the target protein. This concentration is then used for the large-scale wash.

For elution, a gradient of imidazole (e.g., 50–500 mM) can be used to separate the target protein from contaminants that elute at slightly different concentrations. However, a step elution with a single high concentration is simpler and often sufficient.

Effect of pH on Binding

The binding of histidine to Ni²⁺ is pH-dependent. At low pH, the imidazole ring becomes protonated (positively charged) and loses its ability to donate electrons to the metal ion. Therefore, binding is favored at higher pH.

• pH 8.0: Standard for most purifications. Histidine is largely deprotonated, and binding is efficient.
• pH 7.0: Binding is weaker, which can be useful for reducing non-specific binding but may also reduce yield.
• pH 6.0–6.5: The His-tagged protein will begin to elute, as the histidines become protonated. A pH gradient can be used as an alternative elution method, though imidazole is more common.

For proteins that are unstable at pH 8.0, a lower pH can be used, but the imidazole concentration in the binding buffer may need to be reduced to maintain binding.

Salt and Detergent Additives

Salt (NaCl) is included to reduce ionic interactions. A concentration of 300 mM is standard, but higher salt (up to 1 M) can be used to reduce non-specific binding of nucleic acids and other proteins. However, very high salt can weaken the His-tag–metal interaction.

Detergents are added to prevent hydrophobic interactions and protein aggregation. Non-ionic detergents (Triton X-100, Tween-20) at 0.1–1% are compatible with IMAC. Ionic detergents like SDS should be avoided in native purifications, as they denature proteins and disrupt the interaction.

For membrane proteins, which are hydrophobic and tend to aggregate, the purification is performed in the presence of mild detergents such as n-dodecyl-β-D-maltoside (DDM) at 0.03–0.1%. The detergent is included in all buffers to maintain the protein in a soluble, monodisperse state.

Analyzing Purity and Yield

After purification, it is essential to assess the quality of the preparation. This involves determining both the purity (what fraction of the total protein is the target) and the yield (how much target protein was obtained).

SDS-PAGE and Coomassie Staining

Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is the standard method for assessing purity. The protein sample is denatured with SDS, which binds to the protein and gives it a uniform negative charge, so that proteins separate by molecular weight alone.

1. Prepare samples: Mix an aliquot of each fraction (lysate, flow-through, washes, elutions) with SDS-PAGE loading buffer (containing SDS, β-mercaptoethanol, and glycerol). Heat at 95°C for 5 minutes.
2. Run the gel: Load the samples onto a polyacrylamide gel (typically 12% for proteins in the 20–100 kDa range). Run at 150–200 V until the dye front reaches the bottom.
3. Stain with Coomassie Blue: After electrophoresis, stain the gel with Coomassie Brilliant Blue R-250 (0.1% in 40% methanol, 10% acetic acid) for 30–60 minutes, then destain in the same solution without dye.

The target protein should appear as a prominent band at its expected molecular weight in the elution fractions. The purity can be estimated by the relative intensity of the target band compared to all other bands. A single band indicates high purity (>95%).

Western Blot with Anti-His Antibody

If the target protein is expressed at low levels or is difficult to detect by Coomassie staining, a Western blot can be used. The proteins are transferred from the SDS-PAGE gel to a nitrocellulose or PVDF membrane, which is then probed with an antibody that recognizes the His-tag (anti-His antibody). This is a highly sensitive method that can detect picogram quantities of protein. It is also useful for confirming that the purified protein is indeed the His-tagged target, and for detecting degradation products that retain the tag.

Quantification (Bradford, BCA)

The protein concentration in the pooled elution fractions is determined using a colorimetric assay.

• Bradford assay: Uses Coomassie G-250 dye, which shifts its absorbance from 465 nm to 595 nm upon binding to protein. It is quick and simple but can be affected by the presence of detergents.
• BCA assay: Uses bicinchoninic acid, which forms a purple complex with Cu⁺ produced by the reduction of Cu²⁺ by protein. It is more tolerant of detergents but is slower and less compatible with reducing agents.

For more precise quantification, especially for downstream applications like structural studies, the A₂₈₀ absorbance can be used if the protein's extinction coefficient is known. The yield is calculated as the total amount of protein in the pooled elution fractions. The specific yield (mg of protein per liter of culture) is a useful metric for comparing expression conditions. For high-throughput workflows, Automated Protein Quantification systems can streamline this analysis.

Common Pitfalls and Troubleshooting

Despite the simplicity of His-tag purification, several problems can arise. The following are the most common issues and their solutions.

Low Binding or Yield

Symptom: The target protein is found in the flow-through rather than the elution fractions.

Possible causes and solutions:

• Tag is not accessible: The His-tag may be buried in the folded protein. Try purifying under denaturing conditions (see below), or move the tag to the other terminus.
• Imidazole concentration too high in the binding buffer: Reduce the imidazole concentration to 5–10 mM or omit it entirely.
• pH too low: Ensure the lysis buffer is at pH 8.0.
• Resin capacity exceeded: The amount of target protein may exceed the binding capacity of the resin. Use more resin or reduce the amount of lysate.
• Metal ion leached from the resin: This can happen if the resin is old or was exposed to chelating agents like EDTA. Use fresh resin.
• Protein is insoluble: The target protein may be in the pellet after centrifugation. Check the insoluble fraction by SDS-PAGE. If so, consider lower expression temperature (e.g., 16°C overnight) or denaturing purification.

Contaminating Proteins

Symptom: Multiple bands are visible on the SDS-PAGE gel in the elution fractions.

Possible causes and solutions:

• Insufficient washing: Increase the number of wash steps or the imidazole concentration in the wash buffer.
• Non-specific binding: Host proteins with surface-exposed histidines can bind to the resin. Increase the salt concentration (to 500 mM–1 M NaCl) or add a low concentration of imidazole (20–40 mM) to the binding buffer.
• Use a cobalt resin: Cobalt resins have higher specificity and may reduce contamination.
• Protein degradation: Contaminating bands may be degradation products of the target protein. Add more protease inhibitors to the lysis buffer and work quickly at 4°C.

Protein Precipitation

Symptom: The protein precipitates during elution or after dialysis, resulting in a cloudy solution or a loss of protein.

Possible causes and solutions:

• Protein is unstable at high concentration: Elute into a buffer containing glycerol (10–20%) or a stabilizing agent.
• Imidazole causes precipitation: Some proteins are sensitive to high imidazole concentrations. Elute with a lower imidazole concentration (e.g., 150 mM) or use a pH gradient for elution.
• Protein aggregates during dialysis: Add a reducing agent (e.g., 1 mM DTT) or a mild detergent to the dialysis buffer.

Tag Cleavage

Symptom: The purified protein has a lower molecular weight than expected, or multiple bands are present.

Possible causes and solutions:

• Proteolytic cleavage: The His-tag may be cleaved by host proteases during expression or purification. Use protease-deficient E. coli strains (e.g., BL21(DE3) pLysS) and include protease inhibitors in all buffers.
• Cleavage during storage: If the protein is stored for long periods, the tag may be cleaved by residual proteases. Store at -80°C in small aliquots.

Alternative Methods and Advanced Considerations

While the 6xHis tag is the most common, it is not the only option. For challenging proteins, alternative tags and strategies may be necessary.

Other Affinity Tags (GST, MBP)

• GST (Glutathione S-Transferase) tag: A 26 kDa protein that binds to glutathione immobilized on a resin. It is larger than a His-tag, which can improve the solubility of the fusion protein, but it must be cleaved off for many applications, and the larger tag can interfere with structure-function studies.
• MBP (Maltose-Binding Protein) tag: A 40 kDa protein that binds to amylose resin. It is highly effective at promoting solubility of difficult proteins, but its large size requires removal for most downstream applications.

These tags are often used in combination with a His-tag, allowing sequential purification steps. The choice of tag depends on the specific requirements of the target protein and the downstream application.

Purification of Insoluble Proteins

If the target protein forms inclusion bodies (insoluble aggregates) in E. coli, it can be purified under denaturing conditions.

1. Lyse cells in a buffer containing 8 M urea or 6 M guanidine hydrochloride. These chaotropes denature all proteins and solubilize the inclusion bodies.
2. Bind to the resin in the presence of the denaturant. The His-tag can still bind to Ni-NTA under these conditions, as the interaction is based on coordination chemistry, not on the protein's native structure.
3. Wash with denaturing buffer containing imidazole.
4. Elute with denaturing buffer containing high imidazole.

The purified protein is denatured and must be refolded to regain its native structure. Refolding is often performed by dialysis against a buffer without denaturant, or by rapid dilution into a refolding buffer. This is a challenging process that often requires optimization.

On-Column Refolding

An alternative to refolding in solution is on-column refolding. The denatured protein is bound to the resin, and the denaturant is gradually removed by washing with a decreasing gradient of urea or guanidine. This allows the protein to refold while immobilized, which can prevent aggregation. The refolded protein is then eluted with imidazole. This method is particularly useful for proteins that are prone to aggregation during refolding in solution.

For proteins destined for structural studies, the purity and monodispersity achieved by IMAC may be sufficient for initial Protein Crystallization screens. If higher purity is required, a second purification step, such as size-exclusion chromatography (SEC), can be added.

Practical Summary and Best Practices

Successful His-tagged protein purification requires attention to detail at every step, from cloning to final analysis. The following checklist summarizes the key considerations.

Quick Protocol Checklist

1. Design the construct: Choose the tag position (N- or C-terminal) and include a protease cleavage site if tag removal is required.
2. Optimize expression: Test different induction temperatures (16°C, 25°C, 37°C) and IPTG concentrations (0.1–1.0 mM) to maximize soluble expression.
3. Prepare buffers fresh: Include protease inhibitors and degas the buffers if using a gravity column.
4. Use the correct resin: Select Ni-NTA for high capacity or cobalt for higher purity.
5. Bind efficiently: Use batch binding for large volumes and ensure adequate mixing time (30–60 minutes).
6. Wash thoroughly: Use 10–20 column volumes of wash buffer, and optimize the imidazole concentration if purity is insufficient.
7. Elute in small fractions: Collect 0.5–1 column volume fractions to concentrate the protein and avoid dilution.
8. Analyze immediately: Run an SDS-PAGE gel on the same day to assess purity and yield.
9. Store properly: Dialyze to remove imidazole, then aliquot and freeze at -80°C.

Safety and Cost Considerations

• Imidazole: Handle with care; it is a mild irritant. Use in a fume hood when weighing.
• Ni-NTA resin: Nickel compounds are potential carcinogens. Wear gloves and dispose of spent resin as hazardous waste.
• Sonication: Always use ear protection and keep the probe immersed to avoid aerosolization.
• Cost: Ni-NTA resin is relatively inexpensive and can be regenerated multiple times. The main costs are the expression reagents (IPTG, antibiotics) and the resin itself.

Frequently Asked Questions

What is the principle behind his-tagged protein purification?

The principle is based on the coordination chemistry between histidine residues and transition metal ions. A 6xHis tag fused to the target protein binds to Ni²⁺ or Co²⁺ ions that are chelated on a chromatographic resin. The binding is reversible: it can be disrupted by adding imidazole, which competes with the histidines for the metal coordination sites, or by lowering the pH, which protonates the histidines and abolishes their ability to coordinate the metal.

Why is imidazole used in his-tag purification?

Imidazole is used because it is structurally identical to the histidine side chain. It acts as a competitive inhibitor of the His-tag–metal interaction. At low concentrations (10–50 mM), it prevents non-specific binding of host proteins that have surface-exposed histidines. At high concentrations (200–500 mM), it outcompetes the His-tag for the metal binding sites, causing the tagged protein to elute from the resin.

What is the difference between Ni-NTA and cobalt resins?

Ni-NTA uses nickel ions chelated by nitrilotriacetic acid. It has a high binding capacity and high affinity for His-tags, but it also binds more contaminating proteins. Cobalt resins (e.g., TALON) use cobalt ions chelated by carboxymethylaspartate. They have a lower binding capacity and weaker affinity, but they are more specific, resulting in higher purity. Cobalt resins are also more tolerant of reducing agents like DTT.

How do I choose the right imidazole concentration for washing?

The optimal wash concentration is the highest concentration that does not elute the target protein. This is determined empirically. Perform a small-scale test: bind the lysate to a small amount of resin, then wash with increasing imidazole concentrations (e.g., 10, 20, 40, 60, 80 mM). Analyze the washes by SDS-PAGE to find the concentration at which the target protein begins to appear in the wash. Use a concentration slightly below this for the large-scale wash.

Why is my his-tagged protein not binding to the resin?

Common causes include: the His-tag is not accessible (buried in the folded protein), the imidazole concentration in the binding buffer is too high, the pH is too low, the resin capacity is exceeded, or the metal ion has leached from the resin. Check each of these parameters. If the tag is inaccessible, consider purifying under denaturing conditions or moving the tag to the other terminus.

Can I purify his-tagged proteins under denaturing conditions?

Yes. The His-tag–metal interaction is based on coordination chemistry and does not require the protein to be folded. Purification can be performed in the presence of 8 M urea or 6 M guanidine hydrochloride. The protein is bound, washed, and eluted in the denaturing buffer. The purified protein must then be refolded by removing the denaturant, either by dialysis or by on-column refolding.

How do I remove the his-tag after purification?

The His-tag can be removed by incorporating a protease cleavage site between the tag and the protein. Common proteases include thrombin, TEV protease, and Factor Xa. After purification, the protease is added to the pooled elution fractions, and the mixture is incubated under appropriate conditions (e.g., 4°C overnight for TEV). The cleaved His-tag and the protease (if it is also His-tagged) can be removed by passing the mixture through a Ni-NTA column again; the untagged protein flows through, while the tag and protease bind.

Key Takeaways

• His-tagged protein purification relies on the reversible coordination of histidine residues to immobilized metal ions (Ni²⁺ or Co²⁺), a technique known as Immobilized Metal Affinity Chromatography (IMAC).
• The 6xHis tag is small, genetically encodable, and can be fused to either terminus of a target protein without usually affecting its function.
• Imidazole is the key reagent: low concentrations prevent non-specific binding, while high concentrations elute the target protein by competitive displacement.
• Ni-NTA resin offers high binding capacity and low cost, while cobalt resins provide higher specificity and purity at the expense of capacity.
• The standard workflow involves cell lysis, binding, washing, and elution, with buffer composition (pH, salt, imidazole) being the primary variables for optimization.
• Purity and yield are assessed by SDS-PAGE, Western blotting, and colorimetric protein assays; a single band on a Coomassie-stained gel indicates high purity.
• Common problems—low yield, contamination, precipitation, and degradation—can usually be solved by systematic troubleshooting of buffer conditions, resin choice, and expression parameters.

Further Reading

• Luo X et al. Ultrafast His-Tagged Protein Purification. Current protocols. 2024. PubMed 39301792
• Le Thi HN et al. Novel melanin-derived stationary phase for immobilized metal ion affinity chromatography in recombinant His-tagged protein purification. Protein expression and purification. 2024. PubMed 38365166
• Xu F et al. Polymer brush-modified magnetic nanoparticles for His-tagged protein purification. Langmuir : the ACS journal of surfaces and colloids. 2011. PubMed 21338107
• Dong XY, Feng XD, Sun Y. His-tagged protein purification by metal-chelate affinity extraction with nickel-chelate reverse micelles. Biotechnology progress. 2010. PubMed 20730766
• Li S et al. Surface sieving coordinated IMAC material for purification of His-tagged proteins. Analytica chimica acta. 2018. PubMed 29149997
• Qi X et al. NiCoMnO4: A Bifunctional Affinity Probe for His-Tagged Protein Purification and Phosphorylation Sites Recognition. ACS applied materials & interfaces. 2016. PubMed 27381638

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