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Are all antibodies against a common antigen identical?

Are all antibodies against a common antigen identical?


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I understand that when some antigen (e.g., virus, bacteria, etc.) is recognized in the body, antibodies specific to this antigen are produced that, in turn, bind to the antigen and effectively neutralize them as a potential threat to the organism.

Are all the antibodies that are produced against a specific antigen necessarily identical? Either within a single organism (e.g., a single person), or between different organisms (e.g., between different people, different species, etc.)? If two people get sick with the same disease, develop antibodies, and recover, are these antibodies the same? If a human and some other animal get sick with the same disease, will the antibodies each develop match?

Key to the operation of the antibodies is their specific recognition/binding of the antigen. I assume that the structure and composition of the antibody is tuned to bind to specific exposed moieties on the antigen. I can imagine that there are many different potential binding sites on the antigen to target and many possible combinations of ways to recognize those sites by the antibodies. I would think that it would be a remarkable coincidence (highly unlikely) if separate organisms both, independently, happened to develop the same “molecular strategy” for recognizing a particular antigen. This outcome seems to be implied in this question, How are antibodies specific for a disease detected in the blood if everybody produces a different antibody for the same antigen?


Alright, there are two major questions:

(a) Are all the antibodies that are produced against a specific antigen necessarily identical?

Not in most cases. Antigens can either be monovalent or polyvalent, meaning that they can either have one epitope or various different epitopes (an epitope is the site of the antigen that binds to a T-cell receptor or a B-cell receptor). Most antigens are polyvalent, which implies that a single antigen may interact with several different T and B cell lineages. Each B cell lineage is only able to produce and export one type of antibody. Because the antigen will interact with different lineages, it will promote the production of different types of antibodies. These multiple antibodies produced against one antigen are called polyclonal antibodies, since they arise from multiple clonal lineages. When the antigen only has one kind of epitope, it will induce the production of only one type of antibody, which are called monoclonal antibodies, because they arise from only one clonal lineage.

(b) Do the antibodies produced against a certain antigen differ between members of the same species or between species?

Yes, in both cases. There is an important genetic variability surrounding immunoglobulins and T cell receptors.

Reference: Karp, G. (2009). Molecular and Cellular Biology: Methods and Concepts. Wiley. New York.


42.3 Antibodies

By the end of this section, you will be able to do the following:

  • Explain cross-reactivity
  • Describe the structure and function of antibodies
  • Discuss antibody production

An antibody , also known as an immunoglobulin (Ig), is a protein that is produced by plasma cells after stimulation by an antigen. Antibodies are the functional basis of humoral immunity. Antibodies occur in the blood, in gastric and mucus secretions, and in breast milk. Antibodies in these bodily fluids can bind pathogens and mark them for destruction by phagocytes before they can infect cells.

Antibody Structure

An antibody molecule is comprised of four polypeptides: two identical heavy chains (large peptide units) that are partially bound to each other in a “Y” formation, which are flanked by two identical light chains (small peptide units), as illustrated in Figure 42.22. Bonds between the cysteine amino acids in the antibody molecule attach the polypeptides to each other. The areas where the antigen is recognized on the antibody are variable domains and the antibody base is composed of constant domains.

In germ-line B cells, the variable region of the light chain gene has 40 variable (V) and five joining (J) segments. An enzyme called DNA recombinase randomly excises most of these segments out of the gene, and splices one V segment to one J segment. During RNA processing, all but one V and J segment are spliced out. Recombination and splicing may result in over 10 6 possible VJ combinations. As a result, each differentiated B cell in the human body typically has a unique variable chain. The constant domain, which does not bind antibody, is the same for all antibodies.

Similar to TCRs and BCRs, antibody diversity is produced by the mutation and recombination of approximately 300 different gene segments encoding the light and heavy chain variable domains in precursor cells that are destined to become B cells. The variable domains from the heavy and light chains interact to form the binding site through which an antibody can bind a specific epitope on an antigen. The numbers of repeated constant domains in Ig classes are the same for all antibodies corresponding to a specific class. Antibodies are structurally similar to the extracellular component of the BCRs, and B cell maturation to plasma cells can be visualized in simple terms as the cell acquires the ability to secrete the extracellular portion of its BCR in large quantities.

Antibody Classes

Antibodies can be divided into five classes—IgM, IgG, IgA, IgD, IgE—based on their physiochemical, structural, and immunological properties. IgGs, which make up about 80 percent of all antibodies, have heavy chains that consist of one variable domain and three identical constant domains. IgA and IgD also have three constant domains per heavy chain, whereas IgM and IgE each have four constant domains per heavy chain. The variable domain determines binding specificity and the constant domain of the heavy chain determines the immunological mechanism of action of the corresponding antibody class. It is possible for two antibodies to have the same binding specificities but be in different classes and, therefore, to be involved in different functions.

After an adaptive defense is produced against a pathogen, typically plasma cells first secrete IgM into the blood. BCRs on naïve B cells are of the IgM class and occasionally IgD class. IgM molecules make up approximately ten percent of all antibodies. Prior to antibody secretion, plasma cells assemble IgM molecules into pentamers (five individual antibodies) linked by a joining (J) chain, as shown in Figure 42.23. The pentamer arrangement means that these macromolecules can bind ten identical antigens. However, IgM molecules released early in the adaptive immune response do not bind to antigens as stably as IgGs, which are one of the possible types of antibodies secreted in large quantities upon reexposure to the same pathogen. Figure 42.23 summarizes the properties of immunoglobulins and illustrates their basic structures.

IgAs populate the saliva, tears, breast milk, and mucus secretions of the gastrointestinal, respiratory, and genitourinary tracts. Collectively, these bodily fluids coat and protect the extensive mucosa (4000 square feet in humans). The total number of IgA molecules in these bodily secretions is greater than the number of IgG molecules in the blood serum. A small amount of IgA is also secreted into the serum in monomeric form. Conversely, some IgM is secreted into bodily fluids of the mucosa. Similar to IgM, IgA molecules are secreted as polymeric structures linked with a J chain. However, IgAs are secreted mostly as dimeric molecules, not pentamers.

IgE is present in the serum in small quantities and is best characterized in its role as an allergy mediator. IgD is also present in small quantities. Similar to IgM, BCRs of the IgD class are found on the surface of naïve B cells. This class supports antigen recognition and maturation of B cells to plasma cells.

Antibody Functions

Differentiated plasma cells are crucial players in the humoral response, and the antibodies they secrete are particularly significant against extracellular pathogens and toxins. Antibodies circulate freely and act independently of plasma cells. Antibodies can be transferred from one individual to another to temporarily protect against infectious disease. For instance, a person who has recently produced a successful immune response against a particular disease agent can donate blood to a nonimmune recipient and confer temporary immunity through antibodies in the donor’s blood serum. This phenomenon is called passive immunity it also occurs naturally during breastfeeding, which makes breastfed infants highly resistant to infections during the first few months of life.

Antibodies coat extracellular pathogens and neutralize them, as illustrated in Figure 42.24, by blocking key sites on the pathogen that enhance their infectivity (such as receptors that “dock” pathogens on host cells). Antibody neutralization can prevent pathogens from entering and infecting host cells, as opposed to the CTL-mediated approach of killing cells that are already infected to prevent progression of an established infection. The neutralized antibody-coated pathogens can then be filtered by the spleen and eliminated in urine or feces.

Antibodies also mark pathogens for destruction by phagocytic cells, such as macrophages or neutrophils, because phagocytic cells are highly attracted to macromolecules complexed with antibodies. Phagocytic enhancement by antibodies is called opsonization. In a process called complement fixation, IgM and IgG in serum bind to antigens and provide docking sites onto which sequential complement proteins can bind. The combination of antibodies and complement enhances opsonization even further and promotes rapid clearing of pathogens.

Affinity, Avidity, and Cross Reactivity

Not all antibodies bind with the same strength, specificity, and stability. In fact, antibodies exhibit different affinities (attraction) depending on the molecular complementarity between antigen and antibody molecules, as illustrated in Figure 42.25. An antibody with a higher affinity for a particular antigen would bind more strongly and stably, and thus would be expected to present a more challenging defense against the pathogen corresponding to the specific antigen.

The term avidity describes binding by antibody classes that are secreted as joined, multivalent structures (such as IgM and IgA). Although avidity measures the strength of binding, just as affinity does, the avidity is not simply the sum of the affinities of the antibodies in a multimeric structure. The avidity depends on the number of identical binding sites on the antigen being detected, as well as other physical and chemical factors. Typically, multimeric antibodies, such as pentameric IgM, are classified as having lower affinity than monomeric antibodies, but high avidity. Essentially, the fact that multimeric antibodies can bind many antigens simultaneously balances their slightly lower binding strength for each antibody/antigen interaction.

Antibodies secreted after binding to one epitope on an antigen may exhibit cross reactivity for the same or similar epitopes on different antigens. Because an epitope corresponds to such a small region (the surface area of about four to six amino acids), it is possible for different macromolecules to exhibit the same molecular identities and orientations over short regions. Cross reactivity describes when an antibody binds not to the antigen that elicited its synthesis and secretion, but to a different antigen.

Cross reactivity can be beneficial if an individual develops immunity to several related pathogens despite having only been exposed to or vaccinated against one of them. For instance, antibody cross reactivity may occur against the similar surface structures of various Gram-negative bacteria. Conversely, antibodies raised against pathogenic molecular components that resemble self molecules may incorrectly mark host cells for destruction and cause autoimmune damage. Patients who develop systemic lupus erythematosus (SLE) commonly exhibit antibodies that react with their own DNA. These antibodies may have been initially raised against the nucleic acid of microorganisms but later cross-reacted with self-antigens. This phenomenon is also called molecular mimicry.

Antibodies of the Mucosal Immune System

Antibodies synthesized by the mucosal immune system include IgA and IgM. Activated B cells differentiate into mucosal plasma cells that synthesize and secrete dimeric IgA, and to a lesser extent, pentameric IgM. Secreted IgA is abundant in tears, saliva, breast milk, and in secretions of the gastrointestinal and respiratory tracts. Antibody secretion results in a local humoral response at epithelial surfaces and prevents infection of the mucosa by binding and neutralizing pathogens.


Antibody-Antigen Interaction Kinetics

The specific association of antigens and antibodies is dependent on hydrogen bonds, hydrophobic interactions, electrostatic forces, and Van der Waals forces. These are of a weak, noncovalent nature, yet some of the associations between antigen and antibody can be quite strong. Like antibodies, antigens can be multivalent, either through multiple copies of the same epitope, or through the presence of multiple epitopes that are recognized by multiple antibodies. Interactions involving multivalency can produce more stabilized complexes however, multivalency can also result in steric difficulties, thus reducing the possibility for binding. All antigen antibody binding is reversible and follows the basic thermodynamic principles of any reversible bimolecular interaction: where KA is the affinity constant, [Ab-Ag] is the molar concentration of the antibody-antigen complex, and [Ab] and [Ag] are the molar concentrations of unoccupied binding sites on the antibody (Ab) or antigen (Ag), respectively.

The time taken to reach equilibrium is dependent on the rate of diffusion and the affinity of the antibody for the antigen and can vary widely. The affinity constant for antibody-antigen binding can span a wide range, extending from below 105/mol to above 1012/mol. Affinity constants can be affected by temperature, pH, and solvent. Affinity constants can be determined for monoclonal antibodies, but not for polyclonal antibodies, as multiple bond formations take place between polyclonal antibodies and their antigens. Quantitative measurements of antibody affinity for antigen can be made by equilibrium dialysis. Repeated equilibrium dialyses with a constant antibody concentration, but varying ligand concentration are used to generate Scatchard plots, which give information about affinity valence and possible cross-reactivity.

When designing experimental procedures, it is important to differentiate between monoclonal and polyclonal antibodies, as these differences are the foundation of both advantages and limitations of their use.


Use of Antibodies in Testing and Identification

Enzyme immunoassays (EIAs) use antibodies to detect the presence of antigens. However, the assays are conducted in microtiter plates or in vivo. There are many different types of EIAs, but they all involve an antibody molecule whose constant region binds an enzyme, leaving the variable region free to bind its specific antigen. The addition of a substrate for the enzyme allows the antigen to be visualized or quantified (Figure (PageIndex<4>)).

In EIAs, the substrate for the enzyme is most often a chromogen, a colorless molecule that is converted into a colored end product. The most widely used enzymes are alkaline phosphatase and horseradish peroxidase for which appropriate substrates are readily available. In some EIAs, the substrate is a fluorogen, a nonfluorescent molecule that the enzyme converts into a fluorescent form. EIAs that utilize a fluorogen are called fluorescent enzyme immunoassays (FEIAs). Fluorescence can be detected by either a fluorescence microscope or a spectrophotometer.

Figure (PageIndex<4>): Enzyme immunoassays, such as the direct ELISA shown here, use an enzyme-antibody conjugate to deliver a detectable substrate to the site of an antigen. The substrate may be a colorless molecule that is converted into a colored end product or an inactive fluorescent molecule that fluoresces after enzyme activation. (credit: modification of work by &ldquoCavitri&rdquo/Wikimedia Commons)

The MMR vaccine is a combination vaccine that provides protection against measles, mumps, and rubella (German measles). Most people receive the MMR vaccine as children and thus have antibodies against these diseases. However, for various reasons, even vaccinated individuals may become susceptible to these diseases again later in life. For example, some children may receive only one round of the MMR vaccine instead of the recommended two. In addition, the titer of protective antibodies in an individual&rsquos body may begin to decline with age or as the result of some medical conditions.

To determine whether the titer of antibody in an individual&rsquos bloodstream is sufficient to provide protection, an MMR titer test can be performed. The test is a simple immunoassay that can be done quickly with a blood sample. The results of the test will indicate whether the individual still has immunity or needs another dose of the MMR vaccine.

Submitting to an MMR titer is often a pre-employment requirement for healthcare workers, especially those who will frequently be in contact with young children or immunocompromised patients. Were a healthcare worker to become infected with measles, mumps, or rubella, the individual could easily pass these diseases on to susceptible patients, leading to an outbreak. Depending on the results of the MMR titer, healthcare workers might need to be revaccinated prior to beginning work.

Immunostaining

One powerful use of EIA is immunostaining, in which antibody-enzyme conjugates enhance microscopy. Immunohistochemistry (IHC) is used for examining whole tissues. As seen in Figure (PageIndex<5>), a section of tissue can be stained to visualize the various cell types. In this example, a mAb against CD8 was used to stain CD8 cells in a section of tonsil tissue. It is now possible to count the number of CD8 cells, determine their relative numbers versus the other cell types present, and determine the location of these cells within this tissue. Such data would be useful for studying diseases such as AIDS, in which the normal function of CD8 cells is crucial for slowing disease progression.

Immunocytochemistry (ICC) is another valuable form of immunostaining. While similar to IHC, in ICC, extracellular matrix material is stripped away, and the cell membrane is etched with alcohol to make it permeable to antibodies. This allows antibodies to pass through the cell membrane and bind to specific targets inside the cell. Organelles, cytoskeletal components, and other intracellular structures can be visualized in this way. While some ICC techniques use EIA, the enzyme can be replaced with a fluorescent molecule, making it a fluorescent immunoassay.

Figure (PageIndex<5>): Enzyme-linked antibodies against CD8 were used to stain the CD8 cells in this preparation of bone marrow using a chromogen. (credit: modification of work by Yamashita M, Fujii Y, Ozaki K, Urano Y, Iwasa M, Nakamura S, Fujii S, Abe M, Sato Y, Yoshino T)

  1. What is the difference between immunohistochemistry and immunocytochemistry?
  2. What must be true of the product of the enzymatic reaction used in immunohistochemistry?

Enzyme-linked Immunosorbent Assays (ELISAs)

The enzyme-linked immunosorbent assays (ELISAs) are widely used EIAs. In the direct ELISA, antigens are immobilized in the well of a microtiter plate. An antibody that is specific for a particular antigen and is conjugated to an enzyme is added to each well. If the antigen is present, then the antibody will bind. After washing to remove any unbound antibodies, a colorless substrate (chromogen) is added. The presence of the enzyme converts the substrate into a colored end product (Figure (PageIndex<4>)). While this technique is faster because it only requires the use of one antibody, it has the disadvantage that the signal from a direct ELISA is lower (lower sensitivity).

In a sandwich ELISA, the goal is to use antibodies to precisely quantify specific antigen present in a solution, such as antigen from a pathogen, a serum protein, or a hormone from the blood or urine to list just a few examples. The first step of a sandwich ELISA is to add the primary antibody to all the wells of a microtiter plate (Figure (PageIndex<6>)). The antibody sticks to the plastic by hydrophobic interactions. After an appropriate incubation time, any unbound antibody is washed away. Comparable washes are used between each of the subsequent steps to ensure that only specifically bound molecules remain attached to the plate. A blocking protein is then added (e.g., albumin or the milk protein casein) to bind the remaining nonspecific protein-binding sites in the well. Some of the wells will receive known amounts of antigen to allow the construction of a standard curve, and unknown antigen solutions are added to the other wells. The primary antibody captures the antigen and, following a wash, the secondary antibody is added, which is a polyclonal antibody that is conjugated to an enzyme. After a final wash, a colorless substrate (chromogen) is added, and the enzyme converts it into a colored end product. The color intensity of the sample caused by the end product is measured with a spectrophotometer. The amount of color produced (measured as absorbance) is directly proportional to the amount of enzyme, which in turn is directly proportional to the captured antigen. ELISAs are extremely sensitive, allowing antigen to be quantified in the nanogram (10 &ndash9 g) per mL range.

In an indirect ELISA, we quantify antigen-specific antibody rather than antigen. We can use indirect ELISA to detect antibodies against many types of pathogens, including Borrelia burgdorferi (Lyme disease) and HIV. There are three important differences between indirect and direct ELISAs as shown in Figure (PageIndex<7>). Rather than using antibody to capture antigen, the indirect ELISA starts with attaching known antigen (e.g., peptides from HIV) to the bottom of the microtiter plate wells. After blocking the unbound sites on the plate, patient serum is added if antibodies are present (primary antibody), they will bind the antigen. After washing away any unbound proteins, the secondary antibody with its conjugated enzyme is directed against the primary antibody (e.g., antihuman immunoglobulin). The secondary antibody allows us to quantify how much antigen-specific antibody is present in the patient&rsquos serum by the intensity of the color produced from the conjugated enzyme-chromogen reaction.

As with several other tests for antibodies discussed in this chapter, there is always concern about cross-reactivity with antibodies directed against some other antigen, which can lead to false-positive results. Thus, we cannot definitively diagnose an HIV infection (or any other type of infection) based on a single indirect ELISA assay. We must confirm any suspected positive test, which is most often done using either an immunoblot that actually identifies the presence of specific peptides from the pathogen or a test to identify the nucleic acids associated with the pathogen, such as reverse transcriptase PCR (RT-PCR) or a nucleic acid antigen test.

Figure (PageIndex<6>): (a) In a sandwich ELISA, a primary antibody is used to first capture an antigen with the primary antibody. A secondary antibody conjugated to an enzyme that also recognizes epitopes on the antigen is added. After the addition of the chromogen, a spectrophotometer measures the absorbance of end product, which is directly proportional to the amount of captured antigen. (b) An ELISA plate shows dilutions of antibodies (left) and antigens (bottom). Higher concentrations result in a darker final color. (credit b: modification of work by U.S. Fish and Wildlife Service Pacific Region) Figure (PageIndex<7>): The indirect ELISA is used to quantify antigen-specific antibodies in patient serum for disease diagnosis. Antigen from the suspected disease agent is attached to microtiter plates. The primary antibody comes from the patient&rsquos serum, which is subsequently bound by the enzyme-conjugated secondary antibody. Measuring the production of end product allows us to detect or quantify the amount of antigen-specific antibody present in the patient&rsquos serum.


Producing Monoclonal Antibodies

Some types of assays require better antibody specificity and affinity than can be obtained using a polyclonal antiserum. To attain this high specificity, all of the antibodies must bind with high affinity to a single epitope. This high specificity can be provided by monoclonal antibodies (mAbs). Table 1 compares some of the important characteristics of monoclonal and polyclonal antibodies.

Table 1. Characteristics of Polyclonal and Monoclonal Antibodies
Monoclonal Antibodies Polyclonal Antibodies
Expensive production Inexpensive production
Long production time Rapid production
Large quantities of specific antibodies Large quantities of nonspecific antibodies
Recognize a single epitope on an antigen Recognize multiple epitopes on an antigen
Production is continuous and uniform once the hybridoma is made Different batches vary in composition

Unlike polyclonal antibodies, which are produced in live animals, monoclonal antibodies are produced in vitro using tissue-culture techniques. mAbs are produced by immunizing an animal, often a mouse, multiple times with a specific antigen. B cells from the spleen of the immunized animal are then removed. Since normal B cells are unable to proliferate forever, they are fused with immortal, cancerous B cells called myeloma cells, to yield hybridoma cells. All of the cells are then placed in a selective medium that allows only the hybridomas to grow unfused myeloma cells cannot grow, and any unfused B cells die off. The hybridomas, which are capable of growing continuously in culture while producing antibodies, are then screened for the desired mAb. Those producing the desired mAb are grown in tissue culture the culture medium is harvested periodically and mAbs are purified from the medium. This is a very expensive and time-consuming process. It may take weeks of culturing and many liters of media to provide enough mAbs for an experiment or to treat a single patient. mAbs are expensive (Figure 3).

Figure 3. Monoclonal antibodies (mAbs) are produced by introducing an antigen to a mouse and then fusing polyclonal B cells from the mouse’s spleen to myeloma cells. The resulting hybridoma cells are cultured and continue to produce antibodies to the antigen. Hybridomas producing the desired mAb are then grown in large numbers on a selective medium that is periodically harvested to obtain the desired mAbs.


Introduction

Antibody-Antigen (Ab-Ag) interactions are based on non-covalent binding between the antibody (Ab) and the antigen (Ag). Correct identification of the residues that mediate Ag recognition and binding would improve our understanding of antigenic interactions and may permit the modification and manipulation of Abs. For example, introducing mutations into the V-genes has been suggested as a way to improve Ab affinity [1]–[3]. However, mutations in the framework regions (FRs) rather than in the Ag binding residues themselves are more likely to evoke an undesired immune response [4]. Knowing which residues bind the Ag can help direct such mutations and be beneficial to Ab engineering [5]–[7]. It has been shown that Ag binding residues are primarily located in the so called complementarity determining regions (CDRs) [7]–[9]. Thus, the attempt to identify CDRs, and particularly the attempt to define their boundaries, has become the focus of extensive research over the last few decades [7], [8], [10]. Kabat and co-workers [9], [11] attempted to systematically identify CDRs in newly sequenced Abs. Their approach was based on the assumption that CDRs include the most variable positions in Abs and therefore could be identified by aligning the fairly limited number of Abs available then. Based on this alignment they introduced a numbering scheme for the residues in the hypervariable regions and determined which positions mark the beginning and the end of each CDR. The Kabat numbering scheme was developed when no structural information was available. Chothia et al. [12], [13] analyzed a small number of Ab structures and determined the relationship between the sequences of the Abs and the structures of their CDRs. The boundaries of the FRs and the CDRs were determined and the latter have been shown to adopt a restricted set of conformations based on the presence of certain residues at key positions in the CDRs and the flanking FRs. This analysis suggested that the sites of insertions and deletions in CDRs L1 and H1 are different than those suggested by Kabat. Thus, the Chothia numbering scheme is almost identical to the Kabat scheme, but based on structural considerations, places the insertions in CDRs L1 and H1 at different positions. As more experimental data became available, the analysis was performed anew, re-defining the boundaries of the CDRs. These definitions of CDRs are mostly based on manual analysis and may require adjustments as the structure of more Abs become available. Abhinandan et al. [14] aligned Ab sequences in the context of structure and found that approximately 10% of the sequences in the manually annotated Kabat database have erroneous numbering. A more recent attempt to define CDRs is that of the IMGT database [15] which curates nucleotide sequence information for immunoglobulins (IG), T-cell receptors (TcR) and Major Histocompatibility Complex (MHC) molecules. It proposes a uniform numbering system for IG and TcR sequences, based on aligning more than 5000 IG and TcR variable region sequences, taking into account and combining the Kabat definition of FRs and CDRs [16], structural data [17] and Chothia's characterization of the hypervariable loops [12]. Their numbering scheme does not differentiate between the various immunoglobulins (i.e., IG or TcR), the chain type (i.e., heavy or light) or the species.

A drawback of these numbering schemes is that CDRs length variability is accommodated with either annotation of insertion (Kabat and Chothia) or by providing excess numbers (IMGT). Abs with unusually long insertions may be hard to annotate this way, and therefore their CDRs may not be identified correctly. Honegger and Pluckthun [18] suggested a structurally improved version of the IMGT scheme. Instead of introducing unidirectional insertions and deletions as in the IMGT and Chothia schemes, they were placed symmetrically around a key position. MacCallum et al. [8] have proposed focusing on the specific notion of Ag binding residues rather than the more vague concept of CDRs. They suggested that these residues could be identified based on structural analysis of the binding patterns of canonical loops. Other studies have dubbed those Ag binding residues Specificity Determining Regions (SDRs) [5], [7]. Here, we analyze Ag-Ab complexes and show that virtually all Ag binding residues fall within regions of structural consensus. We refer to these regions as Ag Binding Regions (ABRs). We show that these regions can be identified from the Ab sequence as well. We used “Paratome”, an implementation of a structural approach for the identification of structural consensus in Abs [19]. While residues identified by Paratome cover virtually all the Ag binding sites, the CDRs (as identified by the commonly used CDR identification tools) miss significant portions of them. We refer to the Ag binding residues which are identified by Paratome but are not identified by any of the common CDR identification methods, as Paratome-unique residues. Similarly, Ag binding residues that are identified by any of the common CDR identification methods but are not identified by Paratome are referred to as CDRs-unique residues. We show that Paratome-unique residues make crucial energetic contribution to Ab-Ag interactions, while CDRs-unique residues have a rather minor contribution. These results allow for better identification of Ag binding sites and thus for better identification of B-cell epitopes. They may also help improve vaccine and Ab design.


Epitopes of an antigen

The actual portions or fragments of an antigen that react with receptors on B-lymphocytes and T-lymphocytes, as well as with free antibody molecules, are called epitopes or antigenic determinants. The size of an epitope is generally thought to be equivalent to 5-15 amino acids or 3-4 sugar residues.

Some antigens, such as polysaccharides, usually have many epitopes, but all of the same specificity. This is because polysaccharides may be composed of hundreds of sugars with branching sugar side chains, but usually contain only one or two different sugars. As a result, most "shapes" along the polysaccharide are the same (see Figure (PageIndex<1>)).

Other antigens such as proteins usually have many epitopes of different specificities. This is because proteins are usually hundreds of amino acids long and are composed of 20 different amino acids. Certain amino acids are able to interact with other amino acids in the protein chain and this causes the protein to fold over upon itself and assume a complex three-dimensional shape. As a result, there are many different "shapes" on the protein (see Figure (PageIndex<2>)). That is why proteins are more immunogenic than polysaccharides they are chemically more complex.

A microbe, such as a single bacterium, has many different proteins (and polysaccharides) on its surface that collectively form its various structures, and each different protein may have many different epitopes. Therefore, immune responses are directed against many different epitopes of many different antigens of the same microbe. (For example, a bacterial cell wall alone may contain over 100 different epitopes.) Even simple viruses possess many different epitopes. (see Figure (PageIndex<3>)).


Antigenic Characterization

&ldquoAntigens&rdquo are molecular structures on the surface of viruses that are recognized by the immune system and are capable of triggering an immune response (antibody production). On influenza viruses, the major antigens are found on the virus&rsquo surface proteins (see Figure 1).

When someone is exposed to an influenza virus (either through infection or vaccination) their immune system makes specific antibodies against the antigens (surface proteins) on that particular influenza virus. The term &ldquoantigenic properties&rdquo is used to describe the antibody or immune response triggered by the antigens on a particular virus. &ldquoAntigenic characterization&rdquo refers to the analysis of a virus&rsquo antigenic properties to help assess how related it is to another virus.

CDC antigenically characterizes about 2,000 influenza viruses every year to compare how similar currently circulating influenza viruses are to those that were included in the influenza vaccine and to monitor for changes in circulating influenza viruses. Antigenic characterization can give an indication of the flu vaccine&rsquos ability to produce an immune response against the influenza viruses circulating in people. This information also helps experts decide what viruses should be included in the upcoming season&rsquos influenza vaccine.

Other information that determines how similar a circulating virus is to a vaccine virus or another virus are the results of serology tests and genetic sequencing.

The above image shows the different features of an influenza virus, including the surface proteins hemagglutinin (HA) and neuraminidase (NA). Following influenza infection or receipt of the influenza vaccine, the body&rsquos immune system develops antibodies that recognize and bind to &ldquoantigenic sites,&rdquo which are regions found on an influenza virus&rsquo surface proteins. By binding to these antigenic sites, antibodies neutralize flu viruses, which prevents them from causing further infection.

The Hemagglutinin Inhibition Assay (HI Test)

Scientists use a test called the hemagglutinin inhibition (HI) assay to antigenically characterize influenza viruses. The HI test works by measuring how well antibodies bind to (and thus inactivate) influenza viruses.

Scientists use the HI test to assess the antigenic similarity between influenza viruses. This test is particularly useful for helping to select the vaccine viruses used in the seasonal flu vaccine. HI test results can tell us whether antibodies developed against vaccination with one virus are antigenically similar enough to another circulating influenza virus to produce an immune response against that circulating virus. Scientists also use the HI test to compare antigenic changes in currently circulating influenza viruses with influenza viruses that have circulated in the past.

The HI test involves three main components: antibodies, influenza virus, and red blood cells that are mixed together in the wells (i.e., cups) of a microtiter plate. (See Image 1.)

A microtiter plate is used to perform the HI test. The plate contains wells (i.e., cup-like depressions that can hold a small amount of liquid) where the solution of antibodies, influenza virus and red blood cells are inserted and allowed to interact. These wells are arranged according to rows and columns (which are identified on the microtiter plate by letters and numbers, respectively). The rows of the plate can be used to test different influenza viruses against the same set of antibodies. The columns can be used to differentiate between greater dilutions of antibodies, like a scale from low to high going from left to right (see Figures 3 and 4 for an example).

The antibodies used in the HI test are obtained by infecting an animal (usually a ferret) that is immunologically naïve (i.e., it has not been exposed to any influenza virus or vaccine previously in its lifetime). The animal&rsquos immune system creates antibodies in response to the antigens on the surface of the specific flu virus that was used to infect that animal. To study these antibodies, a sample of blood (serum) is drawn from the animal. The HI test measures how well these antibodies recognize and bind to other influenza viruses (for example, influenza viruses that have been isolated from flu patients). If the ferret antibodies (that resulted from exposure to the vaccine virus) recognize and bind to the influenza virus from a sick patient, this indicates that the vaccine virus is antigenically similar to the influenza virus obtained from the sick patient. This finding has implications for how well the vaccine might work in people. See Flu Vaccine Effectiveness: Questions and Answers for Health Professionals for more information.

As previously mentioned, the influenza viruses used in the HI test are taken from samples from sick people. CDC and other WHO collaborating centers collect specimens from people all over the world to track which influenza viruses are infecting humans and to monitor how these viruses are changing.

For the HI test, red blood cells (RBCs) are taken from animals (usually turkeys or guinea pigs). They are used in the HI test because influenza viruses bind to them. Normally, RBCs in a solution will sink to the bottom of the assay well and form a red dot at the bottom (Figure 2A). However, when an influenza virus is added to the RBC solution, the virus&rsquo hemagglutinin (HA) surface proteins will bind to multiple RBCs. When influenza viruses bind to the RBCs, the red cells form a lattice structure (Figure 2B). This keeps the RBCs suspended in solution instead of sinking to the bottom and forming the red dot. The process of the influenza virus binding to RBCs to form the lattice structure is called &ldquohemagglutination.&rdquo

The HI test involves the interaction of red blood cells (RBCs), antibody and influenza virus. Row A shows that in the absence of virus, RBCs in a solution will sink to the bottom of a microtiter plate well and look like a red dot. Row B shows that influenza viruses will bind to red blood cells when placed in the same solution. This is called hemagglutination and is represented by the formation of the lattice structure, depicted in the far right column under &ldquoMicrotiter Results.&rdquo Row C shows how antibodies that are antigenically similar to a virus being tested will recognize and bind to that influenza virus. This prevents the virus and RBCs from binding, and therefore, hemagglutination does not occur (i.e., hemagglutination inhibition occurs instead).

When antibodies are pre-mixed with influenza virus followed by RBCs, the antibodies will bind to influenza virus antigens that they recognize, covering the virus so that its HA surface proteins can no longer bind to RBCs (Figure 2C). The reaction between the antibody and the virus inhibits (i.e., prevents) hemagglutination from occurring, which results in hemagglutination inhibition (as shown in Figure 2C). This is why the assay is called a &ldquohemagglutinin inhibition (HI) test.&rdquo Hemagglutination (as depicted in Figure 2B) occurs when antibodies do not recognize and bind to the influenza viruses in the solution, and as a result, the influenza viruses bind to the red blood cells in the solution, forming the lattice structure. When the antibodies do recognize and bind to the influenza viruses in the solution, this shows that the vaccine virus (like the one the ferrets were infected with) has produced an immune response against the influenza virus obtained from the sick patient. When this happens, the influenza virus being tested is said to be &ldquoantigenically like&rdquo the influenza virus that created the antibodies (from ferrets).

When a circulating influenza virus is antigenically different from a vaccine or reference virus, the antibodies (developed in response to the vaccine or reference virus) will not recognize and bind to the circulating influenza virus&rsquo surface antigens. In the HI test this will cause hemagglutination to occur (see Figure 2B). This indicates that the vaccine virus or reference virus has not caused an immune response (i.e., the creation of antibodies) that recognizes and targets the circulating influenza virus. Circulating influenza viruses tested via the HI test are typically obtained from respiratory samples collected from ill patients.

Assessing Antigenic Similarity Using the HI Test

The HI test assesses the degree of antigenic similarity between two viruses using a scale based on greater dilutions of antibodies. As previously mentioned, the HI test is performed using a microtiter plate. The microtiter plate contains rows and columns of wells (i.e., cups) where RBCs, influenza virus and antibodies (developed against a comparison virus, such as a vaccine virus) are mixed. Dilutions are marked across the top of the microtiter plate. These dilutions function as a scale for assessing antigenic similarity and immune response. By testing the ability of greater dilutions of antibody to prevent hemagglutination, scientists measure how well those antibodies recognize and bind to (and therefore inactivate) an influenza virus. The higher the dilution, the fewer antibodies are needed to block hemagglutination and the more antigenically similar the two viruses being compared are to each other. The highest dilution of antibody that results in hemagglutinin inhibition is considered a virus&rsquos HI titer (Figure 3). Higher HI titers are associated with greater antigenic similarity. Greater antigenic similarity suggests that vaccination would produce an immune response against the test virus.

This virus sample has an HI titer of 1280, which means that the greatest dilution of antibody that still blocked hemagglutination from occurring was at 1280 dilution. At this dilution, the antibodies were still capable of recognizing and binding to the antigens on the virus.

When CDC antigenically characterizes influenza viruses to inform decisions on the formulation of the seasonal flu vaccine, the HI test is used to compare currently circulating viruses (B&C) with vaccine viruses (A). This allows scientists to quickly determine if a virus used in the seasonal flu vaccine is antigenically similar to circulating influenza viruses and therefore capable of producing an immune response against them.

Public health experts consider influenza viruses to be antigenically similar or &ldquolike&rdquo each other if their HI titers differ by two dilutions or less. (This is equivalent to a two-well (i.e., a four-fold dilution) or less difference). Using figure 4 as an example, when circulating virus 1 is compared to a vaccine virus, circulating virus 1 differs by one dilution (a 2-fold difference) and therefore is &ldquolike&rdquo the previous season&rsquos vaccine virus. However, circulating virus 2 differs by five dilutions (a 32-fold difference) and therefore is not like the previous season&rsquos vaccine virus. Circulating viruses that are antigenically dissimilar (i.e., not &ldquolike&rdquo) the reference panel are considered &ldquolow reactors.&rdquo

Limitations

Antigenic characterization gives important information about whether a vaccine made using a specific vaccine virus will protect against circulating influenza viruses, but there are several limitations to antigenic characterization test methodology, which are described below.

Egg Adaptations

Right now, most flu vaccines are made using viruses grown in eggs. As human influenza viruses adapt to grow in eggs, genetic changes can occur in the viruses. These are called &ldquoegg-adapted&rdquo changes. Some egg-adapted changes may change the virus&rsquo antigenic (or immunogenic) properties while others may not. Egg-adapted changes have become a particular problem for selection of candidate vaccine viruses (CVVs) for the influenza A(H3N2) virus component of the flu vaccine. Influenza A(H3N2) viruses tend to grow less well in chicken eggs than influenza A(H1N1) viruses and they also are more prone to egg-adapted changes. Such changes can reduce the immune protection provided by the flu vaccine against circulating A(H3N2) viruses.


Rhesus and Other Fetomaternal Incompatibilities

74.4.2 Molecular Basis of Rh Antigens

Studies of the molecular basis of the Rh antigens at the protein level were complicated by loss of Rh antigenic reactivity after membrane solubilization or immunoblotting.

Moore et al. (11) and Gahmberg (12) independently identified proteins of 28–32kDa that could be immunoprecipitated using anti-Rh antibodies. A team led by Cartron at Institut National de Transfusion Sanguine (INTS) in Paris, France virally transformed B cells from donors with circulating Rh antibodies to develop cell lines secreting monoclonal antibodies specific for RhD, c, and E antigens. These monoclonal antibodies allowed the number of Rh antigen sites per cell to be determined. There are approximately 105 Rh antigens to each erythrocyte, with C/c, D, and E each contributing about one-third of the total (13–15) . The Rh polypeptides could then be immunopurified by the addition of large quantities of the monoclonal or polyclonal anti-D to erythrocyte membranes that had been surface labeled with 125I (16,17) . Rh polypeptides were also isolated from surface-labeled RhD-positive erythrocytes by hydroxyapatite chromatography and electrophoresis of SDS-solubilized membrane skeleton vesicles (18) , leading to a nearly 200-fold purification. From these studies, the total number of Rh polypeptides per erythrocyte was calculated to be approximately 60,000. The 28- to 32-kDa Rh polypeptides were suspected to consist of several species, and one-dimensional SDS polyacrylamide gel electrophoresis showed that there was a small difference in the mobilities of the different polypeptides. The RhD polypeptide migrated with an apparent molecular weight of 31.9kDa, whereas c and E were 33.1kDa. When the isolated Rh polypeptides were digested and analyzed by electrophoresis, variations in the degradation patterns indicated that RhD is distinct from the C/c and E/e polypeptides. Antibodies raised to denatured, purified RhD polypeptide were found to cross-react with the Rhc and Rhe polypeptides (19) . Digestion of intact erythrocytes with phospholipase A2 and papain caused degradation of the RhD polypeptide but not the RhC/c or -E/e polypeptides (20) . These studies, therefore, showed that the Rhc, -D, and -E polypeptides are very similar although each is a distinct protein (21) . The c and E polypeptides were found to be nearly identical, whereas the D polypeptide was found to be related but less similar ( Figure 74-1 ).

FIGURE 74-1 . The genetic basis of the rhesus blood group system. RhD protein is encoded by a single gene that is deleted on RHD-negative chromosomes. The RhCcEe proteins are encoded by a single gene that is highly homologous to the RHD gene. Translation of a full-length mRNA transcript produces the RhEe polypeptide, which is of similar size and structure to the RhD polypeptide. The RhEe polymorphism is due to a single-point mutation, which changes amino acid 226. RhC polypeptides are produced from smaller splicing isoform transcripts of the same gene and are therefore smaller peptides than their RhD or RhEe counterparts. The RhCc polymorphism is due to four-point mutations, which make four amino acid substitutions. Associated with these substitutions are two further silent point mutations.

Almost all our current knowledge of this group of proteins has come from cloning studies of the Rh cDNA. Although antibodies to RhD and other Rh antigens were widely available, they were only specific for the Rh antigens. They were not suitable for the identification of Rh polypeptides produced from cDNA expression libraries. Oligonucleotide probes for isolating Rh cDNAs were designed from partial amino acid sequence data of the isolated polypeptides. A clone specific for one species of Rh polypeptide was isolated independently in 1990 by the Paris group (22) and a group in Bristol (23) , both having used the polymerase chain reaction (PCR) with oligonucleotide primers designed from segments of the N-terminal amino acid sequence to amplify cDNA templates prepared from either thalassemic spleen erythroblasts or peripheral reticulocytes. These PCR products were then hybridized to cDNA libraries to identify specific clones of approximately 1.4kb. Both groups used the same commercially acquired cDNA library, prepared from the marrow of the same donor. The sequence of the open reading frames reported by each group was identical. The isolated cDNA localized to chromosome 1p34.3–p36.1 by in situ hybridization, which was consistent with previous linkage evidence that localized the RH gene locus to chromosome 1 (24) . This first cDNA clone proved to encode both the C/c and E/e proteins. In 1992, the Paris group reported the isolation of the RhD polypeptide cDNA from a cDNA library (25) . The coding sequence of this clone exhibited 3.5% divergence at the amino acid level. To confirm that this clone encoded the RhD polypeptide, they demonstrated that its gene was present only in RhD-positive individuals. Restriction fragment length polymorphism analysis of DNA from individuals with RhD-positive and RhD-negative erythrocytes shows that RhD-positive individuals have two Rh polypeptide genes and that RhD-negative individuals have only one (25) . It therefore appears that the RH gene locus consists of two highly homologous, closely linked genes, one of which encodes both C/c and E/e proteins. The other gene encodes the RhD protein and is absent in RhD-negative individuals. Rh antigens are expressed on the erythrocyte surface only in the presence of RhAG. Both Rh and RhAG proteins have 12 transmembrane α-helices through the erythroid cell membrane, joined by six exoloops and five endoloops with N- and C-termini within the cytoplasm (26,27) . The Rh accessory proteins form a group of glycoproteins that are associated with the Rh protein family, and together the association is called the “Rh complex.”

The Rh polypeptides are 30-kDa unglycosylated proteins that can undergo reversible palmitoylation. It is likely that they play a fundamental role in the physiology of RBC membranes that is unrelated to their antigenicity. Similar observations have been made in the cases of other blood group antigens that are often functionally important structures. The multiple membrane-spanning domains of the Rh polypeptides suggest a transporter protein, as a channel, an exchanger or a pump, but the function of the polypeptide or the ligand has not yet been conclusively demonstrated. It has recently been postulated that they may have a function similar to methylamine permease (Mep) transporters (28) , that they may be related to the superfamily of ammonium transporters (29) , and that the Rh glycoproteins function as dual directional ammonium transporters (30) . However, the homology is about 20%, the Mep/Amt family of transporter proteins is present in bacteria, yeast, and plants but not in vertebrates, and there are conserved sequences within the Rh family not found in the Mep/Amt family. So, it remains unclear what may be transported across the membrane, if indeed anything at all. The study of uptake and excretion of metabolic molecules such as ammonium across normal and Rh-null erythrocyte membranes may help elucidate their biological function in mammals.

Erythrocytes from donors of all the common Rh phenotypes are normal, suggesting that the C/c or E/e polymorphisms have no effect on the function of the protein and that erythrocytes can function entirely normally without expression of the D polypeptide. Comparison of distantly related proteins isolated from erythrocytes of several nonhuman species show conservation of the fatty acylation characteristic, suggesting a common functional significance. The membrane defects seen on erythrocytes from individuals with the Rh-null phenotype provide clues to the functional roles of the Rh polypeptides. Rh-null patients have a mild to moderate chronic hemolytic anemia. Rh-null erythrocytes are pleomorphic but always have some degree of stomatocytosis and spherocytosis, and have increased sensitivity to osmotic lysis. Rh-null membranes have characteristically hyperactive membrane ATPases, reduced cation and water contents, and a deficiency in membrane cholesterol (31) . Physiologic roles in membrane stability and volume regulation have been suggested for the Rh polypeptides, but specific details have yet to be elucidated.

Certainly, the principal clinical interest in the Rh polypeptides is in their roles as antigens. The RhD protein expresses the D antigen. The RhCE protein expresses both the C (or c) and E (or e) antigens on the same protein, the C/c antigens being present on the second exoloop and the E/e antigens on the fourth exoloop (32,33) . Many of the current data on the primary structures of these polypeptides have come from cDNA studies (see previous discussion). They are 417 amino acids long, there being only about 8% difference in the sequence between RhD and RhCE, or by only 30–35 amino acids ( Figure 74-2 ). Despite such homology, the two proteins do not share antigens unless there is hybrid rearrangement between the two genes (34) . These antigens appear early during erythropoietic development and early in fetal life.

FIGURE 74-2 . The genetic basis of the RhCcEe polymorphisms. (A) The RhCE proteins are encoded by a single gene, which is highly homologous to RHD. (B) Common RhCE polymorphisms arise from point mutations within RHCE. Single amino acid substitutions (P, proline S, serine A, alanine) give rise to the antigenic differences on the exoloops observed between the various isoforms. Numbers start from first methionine residue.


Polyclonal and monoclonal antibodies

Antibodies (whatever their class or subclass) are produced and purified in two basic forms for use as reagents in immunoassays: polyclonal and monoclonal. Typically, the immunological response to an antigen is heterogeneous, resulting in many different cell lines of B-lymphocytes (precursors of plasma cells) producing antibodies to the same antigen. All of these cells originate from common stem cells, yet each develops the individual capacity to make an antibody that recognizes a particular determinant (epitope) on the same antigen. As a consequence of this heterogeneous response, serum from an immunized animal will contain numerous antigen-specific antibody clones, potentially of several different immuglobulin classes and subclasses comprising generally 2 to 5% of the total immunoglobulin. Because it contains this heterogeneous collection of antigen-binding immunoglobulins, an antibody purified from such a sample is called a polyclonal antibody. Polyclonal antibodies, which are generally purified directly from serum, are especially useful as labeled secondary antibodies in immunoassays.

Because an individual B-lymphocyte produces and secretes only one specific antibody molecule, clones of B-lymphocytes produce monoclonal antibodies. All antibodies secreted by a B-cell clone are identical, providing a source of homogeneous antibody having a single defined specificity. However, while B-lymphocytes can be isolated from suspensions of spleen or lymph node cells excised from immunized animals, they have a limited life span and cannot be cultured directly to produce antibody in useful amounts. Fortunately, this restriction has been overcome with the development of hybridoma technology, wherein isolated B-lymphocytes in suspension are fused with myeloma cells from the same species (usually mouse) to create monoclonal hybrid cell lines that are virtually immortal while still retaining their antibody-producing abilities. Such hybridomas may be stored frozen and cultured as needed to produce the specific monoclonal antibody. Monoclonal antibodies are especially useful as primary antibodies in applications that require single epitope specificity and an unchanging supply over many years of use. Hybridoma clones may be grown in cell culture for collection of antibodies from ascites fluid.



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