Obtaining soluble and biologically active recombinant proteins from inclusion bodies

In prokaryotic hosts such as E. coli, formation of inclusion bodies is common during recombinant protein expression. For many insoluble or structurally complex proteins—especially human proteins rich in disulfide bonds or intrinsically disordered regions—inclusion bodies are often nearly unavoidable. On the surface, inclusion bodies signify misfolding and insolubility; however, with appropriate strategies, they can serve as a “protective form” and a favorable starting point for crude purification. Through proper solubilization and refolding, it is still possible to obtain high-purity target proteins with biological activity.

I. Formation and Characteristics of Inclusion Bodies

Inclusion bodies are insoluble protein aggregates formed under high-level expression, particularly common in E. coli. Compared with eukaryotes, the bacterial cytosol is more reducing, which affects protein folding and disulfide bond formation. When the expression level of a heterologous protein exceeds a certain proportion of the total endogenous bacterial protein, the folding load increases, greatly enhancing misfolding and aggregation and thus promoting inclusion body formation.

Inclusion bodies typically show the following features:

1) They exist as high-density particles inside cells and pellet with the precipitate upon centrifugation;

2) They consist of highly concentrated misfolded or partially folded proteins, often accompanied by intermolecular disulfide crosslinks;

3) Under microscopy, they are often located at the cell poles and appear as dense particulate distributions.

Although inclusion bodies represent folding failure, their highly enriched and relatively homogeneous composition also provides a basis for subsequent crude purification and for protecting protein stability.

II. Buffering Host Toxicity via Inclusion Bodies

Many recombinant proteins carry potential toxicity to the host cells. For example, when human transcription factors are highly expressed in bacteria, they may bind nonspecifically to the bacterial genomic DNA, disrupting finely tuned transcriptional networks, inhibiting growth, or even causing cell death. In such cases, high-expressing cells often die rapidly during culture, leaving only low-expressing cells to proliferate, thereby markedly reducing the total obtainable target protein.

Forcing such proteins into inclusion bodies can mitigate toxicity. Inclusion bodies physically isolate the protein, making it difficult to contact DNA or other critical cellular components, thereby avoiding perturbation of normal host physiology. In practice, by reducing early “leaky expression,” inducing rapidly at an appropriate cell density, and pushing expression strength to drive the protein into inclusion bodies, one can obtain higher total protein yield without markedly suppressing bacterial growth.

III. Suppressing Proteolytic Degradation via Inclusion Bodies

For proteins that are highly disordered or contain long disordered segments, protease-mediated degradation is particularly prominent. Even with protease inhibitors added during lysis and purification, it is difficult to completely prevent proteolysis occurring during the expression phase in bacteria. The typical outcomes are:

1) The product contains both full-length target protein and numerous truncation fragments;

2) Degradation fragments have physicochemical properties similar to the full-length protein, making separation difficult;

3) Both purity and yield are limited.

When the protein exists as inclusion bodies, proteins are tightly packed within insoluble particles and are spatially isolated from most soluble proteases. As long as proteases in the supernatant are removed by multiple wash-and-spin cycles after lysis, subsequent inclusion-body solubilization can yield the target protein under conditions that are nearly free of protease background, thereby markedly reducing degradation produced during expression.

IV. Solubilizing and Purifying Protein from Inclusion Bodies

1.Inclusion bodies as a “crude purification” step

After cell lysis, inclusion bodies can be collected by centrifugation into the pellet, while most endogenous soluble proteins (including proteases) remain in the supernatant. Subsequent multiple resuspension and centrifugation washes of the inclusion-body pellet can effectively remove soluble contaminants. This process essentially uses inclusion bodies as an initial separation step, creating a simpler starting point for downstream solubilization and purification.

2.Inclusion-body solubilization: strong denaturants and reducing agents

Releasing protein from inclusion bodies typically relies on a combination of strong denaturants and reducing agents.

Common schemes include:

6–8 M urea or 6 M guanidine hydrochloride, supplemented with reducing agents such as DTT or TCEP to cleave inter- and intramolecular disulfide bonds.

Typical steps:

1) Resuspend the washed inclusion-body pellet in buffer containing urea or guanidine hydrochloride and a reducing agent;

2) Fully disperse the pellet by sonication, stirring, or repeated pipetting;

3) Incubate at room temperature or low temperature for 30–60 minutes to allow denaturant and reducing agent to act fully;

4) Centrifuge again to remove insoluble material; the target protein is now mainly in the supernatant.

If the target protein carries a His tag, Ni²⁺ affinity chromatography can be performed directly under high-urea or high-guanidine conditions to further purify. This step leverages the inherent enrichment of inclusion bodies and helps quickly obtain a high concentration of the target protein.

V. Protein Refolding: Returning from the Denatured State to the Native Conformation

Using urea or guanidine hydrochloride to solubilize inclusion bodies inevitably renders the protein fully or highly denatured, losing its secondary and tertiary structures. Thus, removing the denaturant and restoring the native fold without aggregation is the most challenging part of the workflow.

1.Dialysis refolding

Dialysis is one of the most common and straightforward refolding methods. The solution containing the denatured protein is loaded into a dialysis membrane and slowly dialyzed against buffer with low or no denaturant so that urea or guanidine gradually diffuses out.

Its advantages are simplicity and controllable conditions; however, two potential issues exist: the refolding process is slow, and the protein may linger in partially folded intermediates for a long time; these intermediates often have highly exposed hydrophobic surfaces and readily associate to form aggregates, lowering the yield of soluble protein. By optimizing dialysis rate, protein concentration, salt concentration, pH, and adding chaperones or small-molecule folding aids, success rates can be improved to some extent, but there remains considerable uncertainty for many aggregation-prone proteins.

2.On-column refolding

For His-tagged proteins, refolding on a Ni²⁺ affinity column is a common and effective strategy. General workflow:

1) In binding buffer containing high urea or guanidine, load the denatured protein onto a Ni²⁺ column to bind via the His tag;

2) Keep the protein bound to the resin and gradually or directly switch the mobile phase to buffer lacking denaturant;

3) During this process, the immobilized protein rapidly sheds the denaturant and completes folding;

4) After refolding, elute the protein with imidazole-containing buffer.

Advantages include: short refolding time—completion within seconds to minutes—reducing the opportunity for intermediate aggregation; the protein is immobilized on the resin with restricted spatial freedom and is less likely to contact and aggregate with other molecules; refolding and purification occur on the same support, making the workflow compact.

VI. Functional Validation of Refolded Protein

Even if biochemical and biophysical parameters appear reasonable, whether the refolded protein truly regains function must be confirmed by downstream functional assays. Common validation methods depend on protein type and include, but are not limited to:

1) Enzymatic activity assays;

2) Receptor–ligand binding assays;

3) DNA or RNA binding assays;

4) Cell-based functional recovery or signaling activation assays, etc.

For example, for DNA-binding transcription factors, electrophoretic mobility shift assays (EMSA) or quantitative binding assays can be used to compare the affinity of proteins refolded from inclusion bodies with those purified directly from soluble expression for specific DNA sequences. If the binding curves are highly consistent under the same conditions, it indicates that the inclusion-body-derived protein has been restored to the correct folded and functional state.

At higher resolution, structural methods such as X-ray crystallography or NMR can be used to verify whether the 3D structure of the refolded protein matches that produced by other preparation routes. However, for most routine applications, appropriately designed functional assays are sufficient to judge refolding success without resorting to structural biology.

By rationally designing expression strategies, choosing suitable denaturation and solubilization conditions, and combining dialysis or on-column refolding, many proteins that are originally insoluble or aggregation-prone can still be successfully refolded into soluble, biologically active forms. Although not all proteins are amenable to preparation via the inclusion-body route, when dealing with hard-to-express or highly toxic proteins, treating inclusion bodies as a usable intermediate form—rather than a mere by-product—helps broaden experimental design and increases the likelihood of obtaining high-quality recombinant protein.

Aladdin: https://www.aladdinsci.com/