Are Monoclonal Antibodies Really More Specific?

It is common to see references to the “specificity” or “greater specificity” of monoclonal antibodies compared to polyclonal antibodies.  Is that claim justified?  No one would dispute the fact that a monoclonal is directed toward a single site or epitope of the target protein, while a polyclonal may contain antibodies directed toward multiple epitopes on the protein.  However, in my opinion, that fact alone does not make monoclonal antibodies more specific.  Indeed the single epitope toward which a monoclonal is directed may in fact be shared by many different proteins in addition to the protein of interest.   Such a monoclonal would not be specific even though it recognizes only a single epitope.  In contrast, a polyclonal antibody raised against the same protein may contain antibodies directed toward that same non-specific epitope as the monoclonal, as well as other epitopes that are more specific.  In such a situation the serum of the polyclonal would at least contain some antibodies that are specific and thus it would be “more specific” than the monoclonal.  Moreover, it may be possible using affinity purification to isolate the specific antibodies. This may seem to some as a trivial issue but it can be extremely important in IHC where it is quite difficult to control for cross reactivity.  Thus simply opting for a monoclonal is no guarantee of specificity and one must still utilize a full range of specificity controls. An example of such a non-specific monoclonal is shown in the figure 1.  Note that the monoclonal antibody recognizes the NET protein at ~50 kD but it also recognizes proteins at 75 and 95 kD.

Figure 1. The monoclonal raised against the NET protein labels three 

prominent bands at 50, 75 and 95 kD in a lysate of rat cortex.

One additional issue relating to this question is the difference in the purification/selection of monoclonal and polyclonal antibodies.  The process of purification and selection of monoclonal antibodies rarely involves screening for specificity.  Thus the hybridoma screens of hundreds or thousands of clones typically rely solely on the ability to recognize a target in a plate assay.  There is typically no selection for affinity for the target and or specificity.  This contrasts with polyclonal antibody affinity purification which, as its name suggests, can preferentially yield higher affinity and higher specificity antibodies. 

Taken together these comments are not at all meant to minimize the very real importance of monoclonal antibodies in clinical environments.  Rather these comments are meant to underscore the importance of specificity controls in using all antibodies and to show that, in some cases, polyclonal antibodies may be more specific than monoclonal antibodies.

Antibody specificity:  The use of a blocking control has only limited value

Antibody specificity is one of the key issues in determining whether you have an antibody that works.  How does one determine that the antibody specifically recognizes only the target of interest?  There are a number of control procedures one can use to be sure that the signal generated in the antibody based assay truly and quantitatively represents the presence of the target of interest. 

In western blots one can at least partially address this issue by determining that the relative molecular weight of the antibody signal matches that of the target. However in most other antibody based imaging assays (e.g. IHC and IF) no such information is available and thus determining specificity in such assays is even more critical.  One of the most common controls for antibody specificity utilizes the antigen that was used to make the antibody as a blocking control.

 Unfortunately the value of this control is often greatly overestimated.  For example take a case where an antibody raised against a protein antigen recognizes only a single epitope in the protein.  Assume for example that this antibody is non-specific and its epitope is also found in a number of other proteins. The antibody will thus recognize its epitope in all of those other proteins as well as in the target protein and thus in IHC it may give a very strong signal as it is detecting many proteins in the tissue.  When one adds the immunizing antigen (which contains the epitope) to the antibody labeling assay, the antigen blocks the antibody labeling of all the proteins which contain the epitope.  Thus it gives a complete block of all IHC signal.  Normally that is interpreted as indicating that the antibody is specific.   Clearly in this hypothetical case the blocking control failed because in fact the antibody was NOT specific.  

This effect can be seen in the Figure at right.  In this western blot as shown in lane 1, an antibody raised against synaptotagmin labels three unknown protein bands in addition to the 60k band representing synaptotagmin.  When the blocking control is used (lane 2) the labeling of the specific 60k band and all three non-specific bands is blocked.  So the blocking control eliminated all of the antibody signal but the antibody was clearly not specific for synaptotagmin.

 Thus anytime an antibody is non-specific and recognizes an epitope that is present in more than one target, the antigen blocking control is virtually useless. Since this type of cross reactivity or non-specificity is the one of the most troublesome types of antibody non-specificity, I would argue that antigen block is only one control to be used and that it is a relatively weak control for antibody specificity.

            One of the best controls for antibody specificity is recombinant tissue that has been engineered to lack the target antigen.  When using such tissue one should see no antibody signal in contrast to wild type tissue.  Phosphatase treated tissue is another one of the best controls is to use when testing phospho-specific antibodies.  Provided that the phosphatase can dephosphorylate the target, the signal from a phosphospecific antibody should be eliminated from the phosphatase treated tissue with no change in the total amount of the target protein compared to untreated tissue.

1% SDS is the lysis buffer of choice  for most western blots or

the case of the missing protein in western blots.

As mentioned in my opening blog, good antibodies sometimes do not work because of poor technique.  One of the most common problems of this type is the failure to solubilize cellular proteins in the lysis step prior to western blot analysis.  Thus,  after centrifugation of the cell lysate many cellular proteins are discarded with the pellet and are consequently missing (not detected) from the western blot.  This problem occurs principally because of the use of nonionic detergents such as NP-40 or triton for cell lysis.  These detergents fail to solubilize many cellular proteins involved in cell signaling.  This problem is particularly acute in brain where synaptic junctions are known to be insoluble in nonionic detergents.  To obviate this problem, the lysis buffer of choice for western blots is virtually always 1% SDS which completely solubilizes membrane and other hard to solubilize proteins and even synaptic junction proteins.  As an added advantage, SDS also inactivates many cellular proteases.  However, inclusion of protease inhibitors with the 1% SDS is often recommended as some proteases are insensitive to or even activated (e.g. proteinase K) by SDS. 

Phospho-specific antibodies

The use of nonionic detergents is made even more problematic when the phosphorylation state of a protein is assayed in western blots using phosphospecific antibodies. This is because nonionic detergents are ineffective in blocking protein phosphatase activity.  Virtually all cellular lysates contain high levels of phosphatase activity such that the lysate proteins can be completely dephosphorylated in a matter of minutes or even seconds.  This would make it impossible for a phosphospecific antibody to work in a western blot as its phosphorylation target has been removed by the phosphatases.  Fortunately, the 1% SDS lysate buffer described above has the added benefit that it completely denatures protein kinases and phosphatases.

Exceptions to the rule

1.    Subcellular fractionation and/or protein-protein interaction.

Because 1% SDS disrupts cell organelles, it is obviously NOT recommended if isolation of cellular organelles such as membranes, mitochondria and nuclei is required.  However, once the organelles have been isolated, it is essential that 1% SDS be used to lyse the organelle fraction to insure solubilization of all the proteins in the organelle. Similarly SDS solubilization is NOT recommended when analyzing protein-protein interactions as SDS disrupts these interactions.

2.    Immunoprecipitation.

Antibodies are inactivated by 1% SDS and this makes immunoprecipitation from the SDS lysis buffer difficult.  This effect can be overcome in some cases (Goebel-Goody et al. 2009) but in the absence of such procedures immunoprecipitation from 1% SDS is not recommended.  Non-ionic detergents do not typically inactivate antibodies and these detergents are commonly used prior to immunoprecipitation.  However, it must also be recognized that the lysate prepared by using nonionic detergent is missing a number of key proteins.  So immunoprecipitation from such lysates must be interpreted with this factor in mind.

Refs:

Davies, KD, Goebel-Goody, SM, Coultrap, SJ and Browning, MD (2008) Long-term synaptic depression that is associated with GluR1 dephosphorylation but not AMPA receptor internalization. J Biol Chem.283:33138-46.

Goebel-Goody, SM, Davies, KD, Linger, RA, Freund,R and Browning, MD. (2009) Phospho-regulation of synaptic and extrasynaptic NMDA receptors in adult hippocampal slices.  Neuroscience 158:1446-1459

Antibodies that work begins

Antibodies that work are extremely valuable research tools.   However, the search for such antibodies is made quite difficult by two different problems.  The first problem, “the good antibody problem” is that production of a good antibody is a difficult, time consuming process; and many researchers and antibody companies do not possess either the patience or the skill necessary to make good antibodies.  The second problem, “the bad technique problem” is that a good antibody may not “work” in all of the various antibody-based assays. This may occur because the antibody cannot detect its target in a particular type of assay (e.g. the antibody target may become denatured in formalin fixation, thus formalin fixation is a bad technique for this antibody).  Alternatively, incorrect technique (AKA  bad technique) in an assay may interfere with the antibody binding to its target. 

            I have been working with antibodies for quite some time and I have had many opportunities to observe antibodies that do and do not work.  For my students who used antibodies that worked, the outcome was often the development of important new insight into a specific protein’s role in normal or disease function.  In contrast, months of frustration and false leads were usually all that resulted when my students used antibodies that did not work.   In this blog I will attempt to discuss how to address both the good antibody problem and the bad technique problem so that more antibodies will work. 

            I am hoping that this will become a collaborative project and I invite anyone who cares about antibodies to contribute to the discussion by offering tips, advice and/or topics for discussion.  I look forward to talking with you on this blog in the coming months. 

            My next topic will be on a bad technique problem and is titled “Antibodies and westerns blots, the case of the missing protein." This is better known as what happens when you use a poor lysis buffer.