Adaptive immunity relies on the ability of the body to recognize and respond to a nearly endless number of molecules, a complex feat achieved through T-cell and B-cell receptors (TCRs and BCRs, respectively). Receptor-antigen interactions are specific, meaning a tremendous amount of diversity in TCRs and BCRs is required to recognize the wide assortment of antigens one might encounter. To this end, the adaptive immune system has evolved a system for somatic diversification, commonly referred to as V(D)J recombination. Ultimately, this process yields cell populations with sufficient TCR and BCR diversity to recognize any peptide imaginable. However, it is this very diversity of receptor sequences that can make it difficult to analyze them, and a myriad of research applications—from examining the mechanisms behind autoimmune diseases and cancers to developing models for immunotherapy—rely on being successful in this first step.
Traditionally, immune repertoire profiling has relied on cloning and Sanger sequencing or functional methods for identifying antigen-specificity (e.g., tetramer assays/stains). Although these methods have yielded valuable insights, the continued development of next-generation sequencing (NGS) has dramatically expanded the scale of repertoire profiling research. Along with this added technology in the researcher's toolkit comes several key decision points on how to proceed with your studies.
Using genomic DNA as a template can make it easier to determine the relative abundance of clonotypes, as each cell only has a single template. However, this comes at a cost. Because each cell has multiple copies of the TCR or BCR transcript, RNA-seq approaches are more sensitive. Thus, using RNA allows for a more comprehensive identification of unique receptor variants, including those that are present in a very small proportion of cells.
Additionally, sequencing mRNA reveals the sequence of expressed receptors that have undergone splicing and post-transcriptional processing, and thus are more likely to yield functional proteins. DNA-based approaches, by contrast, do not identify receptor sequences in their translated forms, and will unavoidably yield many functionally irrelevant sequences. Additionally, template choice can determine your options for NGS library prep (see below).
Most immune profiling experiments have focused on the CDR3 region of TCR and BCR sequences. The CDR3 region contains the core site of interaction between antigen and receptor and is, therefore, the most variable, so this is not surprising. However, full-length sequencing will include additional regions of the receptor, including CDR1 and CDR2, which play roles in the affinity of antigen-receptor binding and/or downstream signaling. Full-length sequencing also makes it easier to directly clone and express those receptors identified in immune repertoire sequencing studies. This is particularly important for antibody therapy or T-cell therapy applications.
The choice to use bulk or single-cell analysis depends on the goal of your experiment. Generally, analysis of immune repertoire diversity tends to rely on bulk approaches, while studies focused on identifying TCR or BCR sequences specific to a particular antigen use single-cell analysis. Bulk analyses can sample many more sequences in a single experiment but do so at the cost of pairing information. Functional TCRs are made up of paired alpha and beta chains, while BCRs have paired heavy and light chains. Thus, this pairing information is required for the identification of the actual TCR or BCR molecule in question. Since individual T cells or B cells express a single receptor sequence, single-cell analysis makes it possible to capture both the sequence and the pairing information. However, analyzing single cells generally decreases the number of samples that can be analyzed. Researchers often start with bulk analysis and move to single cells after selecting for binding affinity or other phenotypes of interest.
The most common method of preparing sequencing libraries for immune profiling is multiplex PCR. Multiplex PCR is appropriate for both DNA and RNA templates and generally consists of two rounds of PCR (after cDNA synthesis, if RNA is used as a template). The first round of PCR amplifies the receptor locus and adds known sequences that act as priming sites for the second round, in which sequencing adaptors and indexes are incorporated. However, the priming sites at the 3′ and especially the 5′ end of the first PCR overlap sites with significant sequence diversity, which means many degenerate primers are needed to amplify different TCR/BCR sequences (see the right panel in the figure below). Of course, this can also introduce significant PCR bias where receptor sequences that are more similar to the primers are amplified more efficiently.
Another approach is 5′ RACE (Rapid Amplification of cDNA Ends) combined with semi-nested PCR (see the left panel in the figure below). This method is only appropriate for RNA inputs, but does avoid the issue of bias introduced by multiplex PCR. During cDNA synthesis, the reverse transcriptase (RT) adds nontemplated nucleotides to the 5′ end of the first-strand cDNA. A template-switching oligo complementary to these nontemplated nucleotides then hybridizes to the first-strand cDNA. This enables the RT to switch templates and incorporate an adapter sequence on the 5′ end. The two subsequent rounds of PCR use this adapter sequence instead of relying on degenerate primers within the TCR or BCR cDNA. This greatly reduces PCR bias and ensures the immune repertoire profile reflects the original sample, not the primer design. In addition, the second PCR is semi-nested, which ensures a high on-target rate, thus reducing sequencing costs.
Antibodies are one of the most specific tools for the detection and capture of molecular targets. The right antibody can play a key role in revealing the presence, quantity, dynamics, and even binding interactions of a target protein. However, finding the right antibody from over 2 million commercial antibodies and 300+ suppliers can be an overwhelming and confusing task.1 We’ve compiled 7 essential tips to help jump-start your search and hone in on the perfect antibody for your experiment.
Identifying the most effective target protein, or antigen, and understanding its complexity is key in choosing the right antibody. Homology with closely related proteins can result in cross-reactivity. Proteins may have multiple names, isoforms, splice variants, and post-translational modifications. If your research depends on the antibody recognizing a specific portion of your target, note the epitope the antibody was raised against and confirm it’s within the domain of interest. Fixation methods can alter an antigen as well, and permeabilization or antigen retrieval treatments may be required to make the target or epitope accessible to the antibody. However, both fixation and permeabilization can affect immune affinity reactions. Protein databases, like UniProt and GeneCards, are great resources for more information.
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With “direct” labeling, primary antibodies that bind to the antigen are directly detected. “Indirect” labeling is when labeled secondary antibodies are used to detect the primary antibody against your chosen antigen.
The use of directly-labeled primary antibodies requires fewer steps and reagents and is the preferred method for some applications that require multiplexing (i.e., flow cytometry), but the signal intensity is lower as only the fluorophores bound to the primary antibody are present. Depending on your method of detection and the degree of expression of the target, this may be acceptable. Some dyes, such as our CF® Dyes, can help minimize the issue of low signal-to-noise as they allow for a higher concentration of dye molecules per antibody and may produce lower background due to improved solubility and less non-specific binding. Directly-labeled primary antibodies also allow researchers to multiplex using a combination of antibodies from the same host. For more information on this, see Biotium’s Tech Tip on Combining Direct and Indirect IF Using Primary Antibodies from the Same Host.
Using secondary antibodies requires more steps and reagents, but increases sensitivity due to the signal amplification from multiple secondary antibodies binding to a single primary antibody. Secondary antibodies can also be used in Tyramide Signal Amplification (TSA) to further enhance the signal of low-abundance antigens and provide opportunity for multiplexing in microscopy. See this helpful Tech Tip to learn more about multi-color labeling using Tyramide Amplification Kits.
Due to target variation between species, you should confirm that your antibody has been validated for your sample species. In most cases, you will want to select an antibody created in a different host species from your sample species, particularly if using secondary antibodies. Validated antibodies are usually available for widely researched model organisms. If you’re unable to find an antibody validated for your sample species, you will need to assess the antibody’s performance and specificity yourself. This is worthwhile as antibodies are often raised against relatively preserved domains and may recognize your target even if the sequence homology of the epitope is as low as 75%.2
The different types of immunoglobulins (IgG, IgM, IgY, etc.) may also impact cross-reactivity and multiplexing. For example, secondary antibodies that react with IgG (H+L) will react with epitopes on both heavy and light chains, so they will react with other isotypes of primary antibody or different subtypes of IgG. Secondary antibodies that specify a specific isotype for their reactivity (e.g., IgG2a) are cross-adsorbed against other isotypes for specific binding.3 If you seek signal strength and broadness of detection over specificity, consider polyclonal antibodies, which are a mixture of antibodies that recognize different epitopes on a given target. If your experiment requires a higher degree of specificity and consistency, you may want to consider monoclonal antibodies.
Antibodies are available in various formulations and degrees of purification. Antibodies may be supplied as pure IgG in PBS but may also come with stabilizers such as BSA, gelatin, glycerol, or amino acids. Crude preparations of antibody in serum, ascites fluid, or supernatant may contain unwanted immunoglobulins or other components that may need to be removed to avoid skewing your results. This is especially important if you plan to label your own antibody with a fluorescent dye or other type of label, as additives like amino acids, glycerol, or Tris can affect labeling efficiency.
Selecting an antibody that has been independently validated can help you get successful results. Suppliers often list validated applications online or in the product documents. Never assume that antibodies validated for one application, such as western blotting, will work well in another, such as flow cytometry, as the state of the antigen may change with the technique (e.g., the antigen will typically be denatured in western blots while the native form is usually present for microscopy or flow cytometry). Finding published examples of an antibody being used in an experiment similar to yours is one of the best predictors of success. PubMed and CiteAb are great resources for determining which antibodies other researchers are using. It is also worthwhile to look for antibodies validated by the seller. Biotium offers a selection of monoclonal Biotium Choice Antibodies that have been carefully curated and extensively validated in-house for flow cytometry.
Multiplexing allows for simultaneous detection of multiple markers in a single sample. Careful consideration of species and immunoglobulin classes is essential to minimize cross-reactivity and background. Highly cross-adsorbed antibodies work best, and most researchers prefer to use directly-conjugated primary antibodies, as some experiments can require up to dozens of different antibodies. If using fluorescently labeled antibodies, remember to choose conjugates that don’t have overlapping emission spectra!
Most experiments require troubleshooting and optimization. Choosing a supplier with knowledgeable technical support and helpful resources will save valuable time and reagents. Refer to these materials to determine how to best reconstitute, store, and aliquot your antibody, and take note of any special considerations listed. These documents typically provide suggestions for optimal concentrations for different applications as well. In addition to product information documents, Biotium offers an antibody selection tool and a variety of tech tips and general protocols for antibody-based detection. Our Technical Support Scientists are always happy to help with experimental troubleshooting once your antibodies have been put to use, as well.
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