June 15, 2014 (Vol. 34, No. 12)
Every immunoassay exploits the ability of an antibody (referred to as a primary antibody) to bind selectively to a particular target antigen, which is usually presented in a complex fluid such as serum, blood or urine, or in a solid matrix such as a slice of tissue, a cell monolayer, a nitrocellulose blot or a plastic microplate. The antibody, in conjunction with a label that provides measurability, allows the quantity and/or distribution of the antigen to be determined. There are many types of labels but fluorescent proteins, enzymes or organic dyes are perhaps the most common ones. Immunoassay applications in which labels are employed include, among others, flow cytometry, ELISA, Western blotting, and immunohistochemistry.
There are two main classes of immunoassays: (i) direct assays, in which the label is covalently attached to the primary antibody; (ii) indirect assays, where the label is attached to a secondary reagent, which in turn can bind to the primary antibody noncovalently and indirectly report the presence of antigen. Indirect assays are more laborious as they have two antibody incubation steps and twice as many wash steps as the direct assay format. However, in the direct assay format the label must first be covalently attached to the primary antibody to form a conjugate. While some primary antibody conjugates are commercially available, the vast majority of antibodies are offered only in their native unlabeled form.
Direct assay formats, apart from their simplicity, also have the advantage that several primary antibody conjugates can be mixed to provide a profile of expression of multiple antigens in a single sample. With the indirect format, it is not possible to mix unlabeled primary antibodies that come from the same species, thus restricting choice and forcing compromise. Moreover, the secondary reagents have to be completely species specific; any crossover binding of a labeled secondary reagent onto the wrong primary antibody will generate a false assay signal.
One of the often-stated advantages of the indirect method is that amplification of the assay signal takes place. This arises because bound primary antibodies may accommodate more than one molecule of labeled secondary antibody. However, the off-rate of the primary antibody is crucially important and the two formats are rarely compared. Much of the primary antibody can dissociate during the extra incubation and wash steps in the indirect procedure, and what is actually amplified is a diminishing amount of primary antibody. Figure 1 shows immunohistochemical data in which the direct and indirect approaches give essentially the same result.
Ideally, the researcher should have at his/her disposal whatever reagents are required to get the best performance when developing a new immunoassay. The main obstacle to wider adoption of the direct assay format is the lack of commercially available primary antibody conjugates. There are several reasons for this situation: (i) The chemicals that are found in antibodies are not always compatible with conjugation methodologies; (ii) antibody conjugation processes are too complex and require specialist knowledge; (iii) antibody companies cannot anticipate which conjugates and labels will be required.
All antibodies are produced initially as crude mixtures, e.g., immune serum, ascites fluid or hybridoma tissue culture supernatant. The chemicals used in some antibody purification processes may interfere in conjugation reactions. Glycine and tris buffer are best avoided, but if they are present the simple operation of dialysis (if performed carefully) circumvents any potential issues. The benefits of changing the dialysis buffer should not be underestimated; for example, it is easy to show mathematically that dialysing 10 mL of antibody against 3 x 1.5 L of buffer (i.e., 4.5 L overall) instead of against 1 x 5 L gives >1,000-fold more efficient removal of unwanted low-molecular-weight substances. Antibodies, once purified, should be stored at 4ºC, preferably at a high concentration (>2 mg/mL) without additives. If any preservatives are deemed necessary they should be used at their lowest effective concentrations. Most commercial antibodies are dialyzed against PBS, which is perfect for conjugation work.
A standard method for conjugating a protein label to an antibody is depicted in Figure 2 (upper panel). The method requires the use of two tags, “X” and “Y”, one of which is attached to the antibody and the other to the label. The label (but not the antibody) can be sometimes be purchased with the Y tag already attached. After removal of excess reactants from the antibody using spin columns, the two entities are mixed and X and Y react to form a covalent X–Y bond that connects the antibody and label. There are a number of related two-tag methods and although the chemistry varies they all have the same advantages and limitations. The key advantage is that the formation of homodimers is prevented because X must react with Y. The major disadvantage is that the purified antibody has to be chemically tagged and separated from excess reactants before the label can be attached. These operations lead to: (i) losses of antibody on spin columns; (ii) batch-to-batch variation; (iii) difficulty in scaling up; and (iv) difficulty in scaling down (which is highly desirable when making trial conjugates with expensive antibody reagents).
To overcome the limitations of the traditional approaches to conjugation, Innova Biosciences has developed Lightning-Link® technology, which has the same control as the two-tag methods but with none of the tags or process complexity (Figure 2, lower panel). Lightning-Link technology allows direct one-step attachment of the antibody to specially modified labels. No knowledge of chemistry is required; the primary antibody is simply pipetted into a vial of lyophilized Lightning-Link mixture. The necessary reactive groups are created in situ while unwanted side reactions are suppressed. Because there are no spin column separation steps, the concentrations and ratios of reactants are tightly controlled by the operator and very easy to replicate. As little as 10 micrograms of antibody can be conjugated, and the hands-on time is always less than 30 seconds, irrespective of the scale. Presently over fifty labels are available in the Lightning-Link one-step format including fluorescein, other organic dyes (e.g., DyLight®, Cy®, and Atto), fluorescent proteins, tandem dyes, enzymes, biotin, and streptavidin.
There are hundreds of suppliers of antibodies and some have over 100,000 primary antibody reagents, the vast majority of course with no labels. The need to apply different chemistries with different types of labels, the sheer number of labels and the complexity of the two-tag systems deter suppliers from investing in the production of thousands of conjugates. It is easier and less risky for suppliers to maintain collections of antibodies in their unlabeled form. There are ~500,000 commercial antibodies (and perhaps a similar number around the world generated by individual researchers) and all of them are compatible in their native forms with Lightning-Link. This opens up some exciting possibilities and Lightning-Link has become the driving force behind the virtual library concept—a vast collection of some 50 million research tools (1 million primary antibodies x 50 Lightning-Link labels), any one of which can become real with under 30 seconds of effort. This is a fabulous position from an assay developer’s perspective and also attractive to antibody companies that can effectively expand their portfolios with zero risk.
Finally, while antibody conjugation was once practiced only by those with specialist knowledge of chemistry, today everyone is a competent conjugation scientist and making antibody conjugates has never been easier.