It may seem odd that one would want to regulate the expression of a protein, if the entire purpose is to "make buckets of it." Many proteins turn out to be toxic when they are overexpressed in a cell, however. We need to have some ability to turn off expression, simply so that the cells stay alive long enough.

We are already very familiar with the regulation of the lac operon, and its derepression with IPTG. A derivative of the lac operon is also commonly used to control expression from the plasmid vectors pGEX and pET.

• In the case of pGEX vectors, the regulation is direct, in that the gene of interest is cloned directly in front of a lac promoter and operator region.
• In the case of pET vectors, the regulation is indirect. The gene of interest is under the control of a T7 RNA polymerase promoter, and IPTG induces expression of the T7 RNA polymerase.

Purification of the protein product

Here is a good place to have already planned ahead! You really have two choices when expressing a protein in a host. You may either express it as a fusion protein, meaning that your sequence of interest is fused to a "tag" or handle of some sort, or you may express your protein in its native state, meaning that the ribosome is initiating translation at the AUG start codon of your gene. Some cloning vectors offer both scenarios as possibilities, and you make your choice at the time you clone your gene.

What are the advantages of having a gene expressed, starting with its own AUG start codon?

• If you are interested in the function of the gene, you may be relieved not to have "uninvited" residues attached. They might confound your interpretation of the function of your gene product.

What are the advantages of fusion proteins?

• The chimera may be more stable or more soluble in the host cell, than would be the native protein.
• The fusion partner may be a convenient "tag" for detection or purification.
• The fusion partner segment may be cut off using a sequence specific protease such as thrombin, factor Xa, or enterokinase.
• If you are dealing with only small segments of a coding region in a gene, a fusion protein prepared with those segments can make the products large enough to see on a gel. They may also act as a hapten to ensure that the segments you care about are immunogenic.

Here is an example of a purification method, based on the GST (glutathione S transferase) fusion system of pGEX, as we discussed previously.

Suppose this schematic diagram represents the fusion protein, with the N-terminal GST part shown in orange and the protein of interest shown in green.

Then we can follow the purification of the chimeric protein on glutathione sepharose, the so-called "RediPack" method from Pharmacia.

Expressed fusion proteins are easily purified from bacterial lysates by affinity chromatography using Glutathione Sepharose 4B. Cleavage of the desired protein from the fusion product is achieved using a site-specific protease whose recognition sequence is located immediately upstream from the multiple cloning site (MCS). The GST System has been used successfully in many applications such as molecular immunology, the production of vaccines and studies involving protein-protein and DNA-protein interactions. Glutathione Sepharose 4B consists of the ligand glutathione, coupled via a 10-carbon spacer arm to the oxirane group of epoxy-activated Sepharose 4B. http://www.biotech.pharmacia.se/gene_xpr/gene_xpr.htm

Detection

The glutathione S transferase also provides a convenient "tag" for detection. That is handy because the cell is making lots of different products, and it can be difficult to find your "needle in a haystack" if you don't have a specific tool to help you look. There are two ways of detecting the glutathione S transferase activity: by enzymatic assay or by immunologic assay.

A colorimetric assay for GST enzyme:

The GST Detection Module is designed to identify GST fusion proteins using either a biochemical or immunological assay. The biochemical assay utilizes glutathione and 1-chloro-2-4-dinitrobenzene (CDNB) as substrates for GST. This reaction yields a yellow product detectable at 340 nm. http://www.biotech.pharmacia.se/gene_xpr/gene_xpr.htm

If you would rather see your fusion protein "light up" on a western blot, there's always an immunological method based on an anti-GST antibody:

Anti-GST Antibody is a polyclonal antibody purified from the sera of goats immunized with purified schistosomal glutathione S-transferase (GST). Because of its polyclonal nature, it can recognize more than one epitope on GST, thereby improving its capacity for recognizing GST fusion proteins even if some binding sites are masked due to recombinant protein folding. http://www.biotech.pharmacia.se/gene_xpr/gene_xpr.htm

The choice of methods for breaking open the cells can be critical, especially with E. coli systems such as pET and pGEX. The overall expression can be determined by simple Coomassie blue staining, after polyacrylamide gel electrophoresis of samples boiled in SDS sample buffer. An amount equal to about 15 µliters of culture volume (at an OD600 of 1.5) is approximately in the right ballpark for a single lane on the gel, though if a Western analysis is to be performed then 1/20 to 1/100 of that amount could suffice.

Some proteins are expressed in a soluble fraction of a cell, some are secreted into a periplasmic space (if your vector has provided ompT or pelB leader sequences), and some form what are called "inclusion bodies" in the cell. Inclusion bodies are an indication that the protein is not soluble, which may be problematic for some purposes. On the other hand, for other purposes (such as the preparation of immunogens) there is no problem at all, and having proteins in these insoluble "inclusion bodies" gives you an easy way to purify them!

One very popular method for purifying proteins is the "His tag" system. Your gene of interest is cloned as a fusion protein with 6 to 10 consecutive histidine residues as a "tag" on the amino or carboxy terminus. When you are ready to purify your protein product, you take advantage of the fact that consecutive histidines can join forces to bind divalent cations such as nickel. You can apply your lysate to a nickel chelation resin, where the his-tagged protein will stick, and wash away the unbound proteins that you don't want. You then elute your tagged protein with imidazole as a competitor. This system is popular because it works under a variety of conditions. You may bind and release a protein from the column under gentle conditions that maintain structure and function, or you alternatively under harsh denaturing conditions (as when you are trying to "persuade" inclusion bodies to go into solution). A small 2.5 ml metal chelation resin column has a capacity of 20 mg recombinant protein.

An example of a purification method using a His tagged protein Purification of (His)10-tagged protein from inclusion bodies in 8 M urea using HisTrap and syringe operation. SDS electrophoresis on PhastSystem using PhastGel 10-15 and silver staining. http://www.novagen.com

Cleavage of fusion proteins

You have several choices, when working with fusion proteins, for how to separate the fusion partner from the peptide sequence of interest. With some vectors, there are sites engineered just upstream of the point of fusion, that allow digestion with specific proteases such as thrombin, factor Xa, or enterokinase. The pGEX vectors have these capabilities.

It's easy to imagine that these new methods of purifying recombinant proteins will revolutionize the field of biochemistry, and will offer a "fast track" for the generation of peptide pharmaceuticals.

Protein display systems

While those biochemists were busy, watching their columns go "drip drip drip", the molecular biologists did another favor for them! They created phage display libraries.

Think for a moment, about the problem of working with proteins. Aside from the bone-chilling time you have to spend in the cold-room, the molecules you work on don't even carry their genetic information with them. Wouldn't it be terrific if a protein just carried its nucleic acid coding sequence with itself, like a suitcase? Then if you found a protein you were particularly interested in, you could just look into the suitcase and pull out the gene sequence that encoded the protein.

That's essentially what we have with phage display libraries (and similar pili display libraries in E. coli). If you clone a random collection of coding sequences into a T7 phage vector coat protein (i.e. as a fusion between the coat protein of T7 phage and your random collection of sequences) then the protein encoded by the inserted sequence will be displayed on the outside of the phage. Why? Because the coat protein, which is assembled on the outside of the phage capsid, is now fused to the peptide encoded by the inserted sequence.

Why is this any help? Because now we can screen a library for phage that are displaying the very protein we are interested in, by any sort of binding assay. In the schematic below, phage are allowed to interact with a ligand bound to a solid support. Those that don't bind are washed away. Those that do bind are isolated, and their fusion gene is sequenced. After several sequential rounds of isolation, a pattern may emerge.

This method is called the Ph.D. kit (for "Phage Display") by New England Biolabs, Incorporated.

Here's an example of how this has worked in epitope mapping. In the given example, the phage display library contains random short segments of amino acids. Larger sequences may also be cloned into the phage display system, to assay native cDNAs for example, but a different T7 vector must be used.

Epitope Mapping of an Anti-Beta-Endorphin Monoclonal Antibody The Ph.D.-12 library was panned against anti-beta-endorphin antibody in solution (10 nM antibody), followed by affinity capture of the antibody-phage complexes onto Protein A-agarose (rounds 1 and 3) or Protein G-agarose (round 2). Bound phage were eluted with 0.2 M glycine-HCl, pH 2.2. Selected 12-mer sequences from each round are shown aligned with the first 12 resides of beta-endorphin; consensus elements are boxed. The results clearly show that the epitope for this antibody spans the first 7 residues of beta-endorphin, and that the bulk of the antibody-antigen binding energy is contributed by the first 4 residues (YGGF), with some flexibility allowed in the third position. Additionally, the conserved position of the selected sequences within the 12 residue window suggests that the free alpha-amino group of the N-terminal tyrosine is part of the epitope. http://www.neb.com/neb/products/phd/phd.html

Description of Phage Display (from New England Biolabs) Phage display describes a selection technique in which a peptide or protein is expressed as a fusion with a coat protein of a bacteriophage, resulting in display of the fused protein on the exterior surface of the phage virion, while the DNA encoding the fusion resides within the virion. Phage display has been used to create a physical linkage between a vast library of random peptide sequences to the DNA encoding each sequence, allowing rapid identification of peptide ligands for a variety of target molecules (antibodies, enzymes, cell-surface receptors, etc.) by an in vitro selection process called biopanning. In its simplest form, biopanning is carried out by incubating a library of phage-displayed peptides with a plate (or bead) coated with the target, washing away the unbound phage, and eluting the specifically-bound phage. (Alternatively the phage can be reacted with the target in solution, followed by affinity capture of the phage-target complexes onto a plate or bead that specifically binds the target.) The eluted phage is then amplified and taken through additional cycles of biopanning and amplification to successively enrich the pool of phage in favor of the tightest binding sequences. After 3-4 rounds, individual clones are characterized by DNA sequencing and ELISA. The Ph.D.-7 linear 7-mer library contains 2.0 x 109 independent clones, while the Ph.D.-C7C disulfide-constrained library contains 3.7 x 109 independent clones. Both libraries are sufficiently complex to contain most if not all of the 207 = 1.28 x 109 possible 7-mer sequences. In contrast, the Ph.D.-12 library, with 1.9 x 109 independent clones, represents only a very small sampling of the potential sequence space of 2012 = 4.1 x 1015 12-mer sequences. http://www.neb.com/neb/products/phd/phd.html