His Tag Labeling and Purification: A Practical Guide
By Dr. Zubair Khalid, DVM, MS, PhD ·
Introduction to His Tag Labeling and Purification
Recombinant protein production is a cornerstone of modern molecular biology, enabling the study of individual proteins in isolation. However, expressing a protein of interest is only half the battle; the other half is isolating it from the thousands of other proteins present in a cell lysate. The polyhistidine tag, commonly called a His tag, is one of the most widely used tools for this purpose. A His tag is a short amino acid sequence, typically six to ten consecutive histidine residues, genetically fused to a target protein. This tag allows the protein to be purified using a technique called immobilized metal affinity chromatography (IMAC), which exploits the high affinity of histidine side chains for certain metal ions.
What is a His Tag?
A His tag is an engineered peptide sequence composed of repeating histidine residues, most commonly six (6xHis), though tags of eight or ten residues are also used. The tag is added to the gene encoding the target protein at the DNA level, so that when the gene is expressed, the resulting protein carries the tag at either its N-terminus or C-terminus. The histidine side chain contains an imidazole ring, a five-membered aromatic ring with two nitrogen atoms. One of these nitrogen atoms has a lone pair of electrons that can coordinate with transition metal ions such as nickel (Ni²⁺) or cobalt (Co²⁺). This coordination forms the basis of the purification strategy.
The tag is small, typically adding only 0.8–1.5 kDa to the molecular weight of the protein, which minimizes the risk of interfering with protein folding or function. It is also immunogenic enough to be recognized by commercial antibodies, enabling detection via Western blot, yet small enough that it often does not require removal for downstream applications such as enzyme activity assays or structural studies.
Why Use His Tags for Protein Purification?
His tags offer several advantages over other affinity tags, such as glutathione S-transferase (GST) or maltose-binding protein (MBP). First, the purification is based on a simple, well-characterized chemical interaction rather than a biological one, making it robust and reproducible. Second, the resin used for purification is relatively inexpensive and can be regenerated multiple times. Third, the elution conditions are mild—typically using imidazole, a histidine analog, at concentrations of 100–500 mM—which preserves protein activity. Fourth, the small size of the tag means it rarely needs to be cleaved off for functional studies, although protocols exist for tag removal if required.
The technique is applicable to a wide range of proteins, from soluble cytoplasmic proteins to membrane proteins, and is compatible with denaturing conditions, which is useful for purifying proteins from inclusion bodies. For a broader overview of the technique, see His Tag Protein Purification.
The Mechanism of His Tag Binding to Nickel or Cobalt Resins
Understanding the chemistry behind His tag purification is essential for troubleshooting and optimizing the process. The interaction between the His tag and the resin is not a simple electrostatic attraction; it is a coordination complex formed between the imidazole nitrogen atoms and a transition metal ion.
Immobilized Metal Affinity Chromatography (IMAC)
IMAC relies on the immobilization of a metal ion, usually Ni²⁺ or Co²⁺, onto a solid support such as agarose or magnetic beads. The metal ion is held in place by a chelating ligand, most commonly nitrilotriacetic acid (NTA) for nickel or carboxymethyl aspartate for cobalt. NTA occupies four of the six coordination sites on the Ni²⁺ ion, leaving two sites free to interact with the histidine residues of the tag. When a cell lysate is passed over the resin, the His-tagged protein binds to the immobilized metal ions, while untagged proteins flow through.
The affinity of the interaction depends on the number of histidines in the tag and the metal used. A 6xHis tag binds to Ni-NTA with a dissociation constant (Kd) in the low micromolar range, typically 10–20 µM. Cobalt resins, such as Talon, have a lower affinity for His tags (Kd around 50–100 µM) but offer higher specificity, meaning fewer contaminating proteins bind. This trade-off between binding strength and specificity is a key consideration when choosing a resin. For a detailed comparison of available resins, refer to His-tag Purification Resin.
Role of Imidazole in Binding and Elution
Imidazole is a small molecule that mimics the histidine side chain. It competes with the His tag for binding to the metal ion. At low concentrations (10–20 mM), imidazole is included in the binding and wash buffers to prevent non-specific binding of host proteins that contain surface-exposed histidines. At high concentrations (250–500 mM), imidazole outcompetes the His tag for the metal coordination sites, causing the tagged protein to elute from the resin.
The elution can be performed either as a step gradient (single high concentration of imidazole) or a linear gradient (gradually increasing imidazole concentration), which separates proteins based on their binding affinity. Proteins with a 10xHis tag, which binds more tightly, require higher imidazole concentrations for elution than those with a 6xHis tag.
Designing a His-Tagged Protein Construct
The design of the expression construct is a critical determinant of purification success. Decisions made at the cloning stage—tag position, length, and linker sequence—can profoundly affect protein yield, solubility, and activity.
Choosing the Tag Location
The His tag can be placed at the N-terminus, C-terminus, or even internally, though the former two are most common. The choice depends on the protein's structure and function.
- N-terminal tag: This is the default choice for many applications. The tag is translated first and is usually fully exposed on the protein surface, making it readily accessible to the resin. However, if the protein's N-terminus is buried in the folded structure or is involved in function, the tag may be inaccessible or disruptive. Additionally, the N-terminal methionine is often cleaved in E. coli, which can remove part of the tag if it is placed immediately after the start codon.
- C-terminal tag: This position is less likely to interfere with the signal peptide or the N-terminal domain, which is important for secreted proteins or proteins with cleavable signal sequences. However, if the C-terminus is buried or if translation terminates prematurely, the tag may be absent from truncated products.
A general rule is to try both positions if initial attempts fail. For membrane proteins, the C-terminal tag is often preferred because the N-terminus may be translocated across the membrane during biosynthesis.
Optimizing Tag Length and Linkers
A 6xHis tag is the standard for most applications. It provides sufficient affinity for purification without being overly immunogenic or disruptive. However, some proteins require a longer tag, such as 8xHis or 10xHis, to achieve efficient binding, particularly if the tag is partially buried or if the protein is expressed at low levels. Longer tags also allow for purification under more stringent wash conditions, reducing contamination.
The linker sequence between the tag and the protein is equally important. A flexible linker, such as Gly-Ser-Gly-Ser or a longer (GGGGS)₃ sequence, ensures that the tag is not constrained by the protein's tertiary structure. Without a linker, the tag may be sterically hindered and unable to reach the metal ions on the resin. A linker of 2–10 amino acids is typically sufficient.
If the tag must be removed after purification, a protease cleavage site, such as that for tobacco etch virus (TEV) protease (ENLYFQG) or PreScission protease (LEVLFQGP), should be inserted between the tag and the protein. TEV protease is particularly popular because it is highly specific and active at 4°C.
Expression of His-Tagged Proteins in Host Systems
The choice of expression host depends on the protein's origin, complexity, and required post-translational modifications. Each system has its own advantages and limitations for producing His-tagged proteins.
Bacterial Expression Systems
Escherichia coli is the most common host for recombinant protein expression due to its fast growth, low cost, and well-characterized genetics. The most widely used system is the T7 RNA polymerase-based pET vector system. In this system, the gene of interest is cloned downstream of a T7 promoter, and expression is induced by adding isopropyl β-D-1-thiogalactopyranoside (IPTG) at a final concentration of 0.1–1 mM. IPTG inactivates the lac repressor, allowing T7 RNA polymerase to transcribe the target gene.
Typical expression conditions involve growing the culture at 37°C to an optical density at 600 nm (OD₆₀₀) of 0.6–0.8, then inducing with IPTG and shifting the temperature to 18–25°C for 4–16 hours. The lower temperature slows protein synthesis, giving the protein more time to fold correctly and reducing the formation of inclusion bodies.
For proteins that are toxic to E. coli, the expression strain BL21(DE3)pLysS or Rosetta(DE3) can be used. The pLysS plasmid encodes T7 lysozyme, which inhibits basal T7 RNA polymerase activity, while Rosetta strains supply tRNAs for rare codons.
Eukaryotic Expression Systems
Proteins that require post-translational modifications, such as glycosylation or disulfide bond formation, may not fold correctly in E. coli. In such cases, yeast (Saccharomyces cerevisiae or Pichia pastoris), insect cells (Sf9 or High Five), or mammalian cells (HEK293 or CHO) are used.
- Yeast: P. pastoris is a popular choice for secreted proteins. The gene is cloned downstream of the alcohol oxidase 1 (AOX1) promoter, and expression is induced by adding methanol to a final concentration of 0.5–1%. The protein is secreted into the culture medium, simplifying purification.
- Mammalian cells: HEK293 cells are commonly used for proteins requiring human-like glycosylation. Expression is typically transient, using polyethyleneimine (PEI) or a viral vector to deliver the plasmid. The cells are cultured at 37°C in a 5% CO₂ atmosphere, and the protein is harvested from the medium 48–72 hours post-transfection.
For a comprehensive overview of expression strategies, see Protein Expression and Purification.
Cell Lysis and Sample Preparation for Purification
The quality of the cell lysate directly determines the efficiency of the purification. Poor lysis or inappropriate buffer conditions can result in low yield, high contamination, or protein degradation.
Lysis Methods
The choice of lysis method depends on the host organism and the scale of the preparation.
- Sonication: This is the most common method for E. coli. Cells are resuspended in lysis buffer and subjected to high-frequency sound waves, which create cavitation bubbles that disrupt the cell membrane. A typical protocol involves 6–10 cycles of 15–30 seconds of sonication at 40–60% amplitude, with 30–60 seconds of cooling on ice between cycles. Over-sonication can denature proteins, so it is important to monitor the process.
- French press: This method uses high pressure (10,000–20,000 psi) to force cells through a small orifice, shearing the cell walls. It is gentler than sonication and is suitable for larger volumes.
- Enzymatic lysis: Lysozyme (1 mg/mL) can be used to digest the peptidoglycan layer of bacterial cell walls. This is often combined with a freeze-thaw cycle or mild detergent to enhance lysis.
- Detergent lysis: For mammalian cells, which lack a cell wall, lysis is achieved using a buffer containing a mild detergent such as Triton X-100 (0.1–1%) or NP-40. The detergent solubilizes the lipid bilayer while preserving protein-protein interactions.
Buffer Conditions and Additives
The lysis buffer should be compatible with the downstream IMAC step. A typical buffer contains 50 mM sodium phosphate (pH 7.4–8.0), 300 mM sodium chloride, and 10–20 mM imidazole. The salt is included to reduce non-specific ionic interactions, while the low imidazole concentration blocks weak binding of contaminating proteins.
Protease inhibitors are essential to prevent degradation of the target protein. A cocktail containing phenylmethylsulfonyl fluoride (PMSF, 1 mM), leupeptin (1 µg/mL), and pepstatin (1 µg/mL) is commonly used. PMSF is unstable in aqueous solution and must be added fresh from a stock solution in isopropanol.
If the protein is expressed as inclusion bodies, the lysis buffer should include a denaturant such as 8 M urea or 6 M guanidine hydrochloride. Under these conditions, the His tag is fully exposed, and purification proceeds under denaturing conditions. The protein can be refolded after elution by dialyzing against a buffer with decreasing denaturant concentration.
Purification Protocol Using Ni-NTA or Cobalt Resins
Once the lysate is clarified by centrifugation (20,000 × g for 30 minutes at 4°C) and filtration through a 0.45 µm filter, the purification can proceed. The two main formats are batch purification and column chromatography.
Batch Purification vs Column Chromatography
Batch purification is ideal for small volumes and for proteins that are prone to precipitation. The clarified lysate is mixed with the resin (typically 1–2 mL of settled resin per liter of culture) and incubated with gentle rotation for 30–60 minutes at 4°C. The mixture is then centrifuged or placed on a magnetic stand (for magnetic beads) to separate the resin from the supernatant. The resin is washed several times, and the protein is eluted.
Column chromatography is preferred for larger volumes and for achieving higher purity. The resin is packed into a column, and the lysate is passed through by gravity flow or using a peristaltic pump. The column format allows for better control of flow rate and more efficient washing.
The choice between Ni-NTA and cobalt resins depends on the required purity and yield. Ni-NTA has a higher binding capacity (5–10 mg of protein per mL of resin) but also binds more contaminants. Cobalt resins have a lower capacity (1–3 mg/mL) but produce cleaner preparations. For proteins expressed at low levels or for applications requiring high purity, cobalt resin is often the better choice.
Optimizing Wash and Elution Steps
A standard purification protocol using Ni-NTA resin is as follows:
- Equilibrate the resin with 5 column volumes of binding buffer (50 mM sodium phosphate, 300 mM NaCl, 10 mM imidazole, pH 8.0).
- Apply the clarified lysate to the column at a flow rate of 0.5–1 mL/min.
- Wash with 10–20 column volumes of binding buffer to remove unbound proteins.
- Wash with 5–10 column volumes of wash buffer (binding buffer with 20–50 mM imidazole) to remove weakly bound contaminants.
- Elute with 5–10 column volumes of elution buffer (binding buffer with 250–500 mM imidazole). Collect fractions of 0.5–1 mL.
The elution can be performed as a step gradient or a linear gradient from 20 to 500 mM imidazole. A linear gradient provides better resolution and can separate the target protein from contaminants that bind with slightly different affinities.
After elution, the resin should be regenerated by washing with 5 column volumes of 500 mM imidazole, followed by 5 column volumes of water, and then stored in 20% ethanol at 4°C. For a detailed protocol, see His Tagged Protein Purification.
Analyzing Purity and Yield of Purified His-Tagged Proteins
After elution, it is essential to assess the success of the purification. This involves determining both the purity (the fraction of the total protein that is your target) and the yield (the total amount of target protein recovered).
SDS-PAGE and Coomassie Staining
Sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) is the standard method for assessing purity. Samples from each step of the purification—lysate, flow-through, washes, and elutions—are mixed with SDS loading buffer, boiled for 5 minutes, and loaded onto a polyacrylamide gel. After electrophoresis, the gel is stained with Coomassie Brilliant Blue R-250.
A successful purification should show a single dominant band at the expected molecular weight of the His-tagged protein in the elution fractions. The presence of multiple bands indicates contamination. The expected molecular weight can be calculated from the amino acid sequence using online tools such as ExPASy ProtParam.
Western Blot with Anti-His Antibodies
If the protein is expressed at low levels or if Coomassie staining is not sensitive enough, a Western blot can be performed. The proteins are transferred from the SDS-PAGE gel to a nitrocellulose or PVDF membrane, which is then probed with an anti-His antibody conjugated to horseradish peroxidase (HRP). The signal is detected using a chemiluminescent substrate.
This method is highly sensitive and can detect picogram amounts of protein. It is also useful for confirming that the purified protein is full-length, as degraded fragments will appear as lower molecular weight bands.
Protein concentration can be determined using the Bradford assay, the bicinchoninic acid (BCA) assay, or by measuring absorbance at 280 nm. The Bradford assay is quick and compatible with imidazole, while the BCA assay is more sensitive but incompatible with reducing agents. For a protein with a known extinction coefficient, A₂₈₀ measurement is the most accurate.
Troubleshooting and Common Pitfalls in His Tag Purification
Even with careful planning, purification can fail. The following are the most common problems and their solutions.
Low Yield or No Binding
If the His-tagged protein does not bind to the resin, the first thing to check is the accessibility of the tag. If the tag is buried in the folded protein, it may not be able to coordinate with the metal ions. Solutions include:
- Adding a longer linker between the tag and the protein.
- Moving the tag to the other terminus.
- Purifying under denaturing conditions (8 M urea) to expose the tag.
Another common cause is that the imidazole concentration in the binding buffer is too high. If the protein binds weakly (e.g., a 6xHis tag on a protein that dimerizes), reducing the imidazole to 5 mM or omitting it entirely may help.
The metal ions may also have leached from the resin. This can happen if the resin is old or if chelating agents such as EDTA or EGTA are present in the lysis buffer. Always check that the lysis buffer does not contain EDTA.
Contaminating Proteins
Contaminants are usually host proteins that have surface-exposed histidines and bind to the resin. To reduce contamination:
- Increase the imidazole concentration in the wash buffer (from 20 to 40–50 mM).
- Use a cobalt resin instead of Ni-NTA, as it has higher specificity.
- Add 0.1–0.5% Triton X-100 or Tween-20 to the binding and wash buffers to disrupt hydrophobic interactions.
- Include 5–10 mM β-mercaptoethanol to reduce disulfide bonds that may cause non-specific binding.
Protein Precipitation or Degradation
If the protein precipitates during elution, it may be due to the high protein concentration or the removal of stabilizing cofactors. Solutions include:
- Eluting with a lower imidazole concentration (150–250 mM) to reduce the ionic strength.
- Adding glycerol (5–10%) to the elution buffer to stabilize the protein.
- Eluting into a tube containing a high-pH buffer (e.g., 1 M Tris, pH 8.0) to immediately neutralize the imidazole.
Degradation is usually caused by proteases in the lysate. Ensure that protease inhibitors are added to the lysis buffer and that all steps are performed at 4°C. If degradation persists, use a protease-deficient strain such as BL21(DE3)pLysS.
For more advanced troubleshooting, see Custom Protein Purification.
Practical Summary and Best Practices
His tag purification is a robust and versatile technique, but success depends on attention to detail at every step, from construct design to final analysis.
Key Takeaways
- The His tag binds to Ni²⁺ or Co²⁺ ions via coordination chemistry involving the imidazole ring of histidine.
- Imidazole is used to prevent non-specific binding at low concentrations and to elute the target protein at high concentrations.
- Tag position (N- vs C-terminal) and length (6xHis vs 10xHis) should be optimized for each protein.
- E. coli is the default expression host, but eukaryotic systems are required for proteins needing post-translational modifications.
- Cell lysis must be thorough but gentle, and the buffer must be compatible with IMAC (no EDTA, low imidazole).
- Batch purification is suitable for small volumes; column chromatography is better for larger scales and higher purity.
- SDS-PAGE and Western blot are essential for assessing purity and confirming the identity of the purified protein.
Checklist for a Successful Purification
- Confirm the His tag is in-frame and accessible.
- Use a protease inhibitor cocktail in the lysis buffer.
- Clarify the lysate thoroughly by centrifugation and filtration.
- Equilibrate the resin before loading.
- Wash extensively to remove contaminants.
- Elute with a step or linear imidazole gradient.
- Analyze every fraction by SDS-PAGE.
- Regenerate the resin immediately after use.
For a deeper dive into the chemistry of the resins, see His-tag Purification Resin. If you are planning to use the purified protein for structural studies, you may also find X Ray Crystallography useful.
Frequently Asked Questions
What is a His tag and how does it work?
A His tag is a short sequence of 6–10 histidine residues fused to a recombinant protein. The imidazole side chains of histidine coordinate with transition metal ions (Ni²⁺ or Co²⁺) immobilized on a chromatography resin. This allows the tagged protein to bind to the resin while other proteins flow through. The protein is then eluted by adding imidazole, which competes for the metal binding sites.
Why is imidazole used in His tag purification?
Imidazole mimics the histidine side chain. At low concentrations (10–20 mM), it blocks weak, non-specific binding of host proteins. At high concentrations (250–500 mM), it outcompetes the His tag for the metal coordination sites, causing the tagged protein to elute.
Should I put the His tag at the N-terminus or C-terminus?
It depends on the protein. The N-terminus is the default choice, but if the N-terminus is buried or involved in function, use the C-terminus. For secreted proteins, the C-terminal tag is often preferred to avoid interference with the signal peptide. In many cases, both positions should be tested.
How many histidines should be in a His tag?
Six histidines (6xHis) is the standard and works for most proteins. If binding is weak or if you need to use stringent wash conditions, an 8xHis or 10xHis tag provides stronger binding. Longer tags also allow purification under denaturing conditions.
What is the difference between Ni-NTA and cobalt resins?
Ni-NTA has a higher binding capacity and binds His tags more tightly, but it also binds more contaminating proteins. Cobalt resins have a lower capacity but higher specificity, producing cleaner preparations. The choice depends on the required purity and the expression level of the protein.
Why is my His-tagged protein not binding to the resin?
Common causes include a buried or inaccessible tag, imidazole concentration too high in the binding buffer, EDTA in the lysis buffer stripping the metal ions, or the protein being expressed as inclusion bodies. Try moving the tag, reducing imidazole, omitting EDTA, or purifying under denaturing conditions.
How do I remove the His tag after purification?
If a protease cleavage site was included in the construct, the tag can be removed by incubating the purified protein with the appropriate protease (e.g., TEV protease) at 4°C overnight. The protease and the cleaved tag can then be removed by passing the mixture over the IMAC resin again; the tag binds, while the untagged protein flows through.
Can His tag purification be used for membrane proteins?
Yes, but with modifications. Membrane proteins require detergents to maintain solubility. The lysis and purification buffers must contain a suitable detergent, such as n-dodecyl-β-D-maltoside (DDM) at 0.03–0.1% or lauryl maltose neopentyl glycol (LMNG). The His tag is often placed at the C-terminus, which is typically cytoplasmic and accessible. Purification is usually performed at 4°C to minimize protein degradation.
Key Takeaways
- His tag purification relies on the coordination of histidine imidazole rings with immobilized Ni²⁺ or Co²⁺ ions.
- Imidazole is a dual-purpose reagent: low concentrations prevent contamination, high concentrations elute the target protein.
- Tag position, length, and linker design are critical for efficient binding and should be optimized empirically.
- E. coli with the pET system is the most common expression platform, but eukaryotic hosts are necessary for complex proteins.
- Proper lysis, buffer composition, and clarification are essential for reproducible purification.
- SDS-PAGE and Western blot are mandatory for validating purity and protein integrity.
- Troubleshooting should focus on tag accessibility, imidazole concentration, and the presence of chelating agents or proteases.