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This protocol describes the use of a template-switch anchored reverse transcription-polymerase chain reaction (RT-PCR) to quantify and characterize all expressed T cell receptor (TR) gene products within any defined T cell population. The approach is based on a switching mechanism at the 5'-terminus of the RNA transcript (SMART) (Chenchik, 1998) and the rapid amplification of cDNA ends (RACE) (Matz et al., 1999), which subsequently enable the unbiased and linear amplification of rearranged TR-specific transcripts using primers that bind an anchor sequence and a conserved 3' region within the TR constant (C) domain. Combined with amplicon subcloning and high throughput sequencing, a comprehensive dataset is generated that defines all constituent clonotypes and their relative frequencies within the template pool (Douek et al., 2002). The procedure is optimized for direct ex vivo analysis, such that in vitro amplification is unnecessary and outgrowth frequency bias is avoided (Price et al., 2004). Materials and methods are provided for each of the following steps: (1) isolation of antigen-specific T cell populations for analysis of TR gene expression; (2) mRNA extraction; (3) cDNA synthesis; (4) cDNA clean-up; (5) PCR amplification of rearranged TR products; (6) gel extraction and purification of amplicons; (7) amplicon ligation into TA cloning vector; (8) vector transformation into competent E. coli; and, (9) colony PCR and sequencing.
The success of this protocol depends greatly on high quality mRNA, which in turn depends greatly on cell preparation and handling. Cells must be isolated viably and not subjected to fixation buffers or azide; wash steps are best conducted using standard cell medium with a serum component. If fixation is required for safety reasons, then an alternative protocol should be followed to preserve RNA template quality (van Bockel et al., 2007). A high degree of purity (>98%) is essential to prevent the artefactual detection of non-specific clonotypes and overestimation of clonality. For this reason, high definition flow cytometric sorting with purity checking is imperative. Other methods for cell enrichment, such as magnetic beadbased procedures, should be avoided in conjunction with this protocol. In addition, detailed phenotypic information can be collected during the flow cytometric sorting procedure. Specificity can be determined either physically with pMHC multimers (Chattopadhyay et al., 2008; Wooldridge et al., 2009) or functionally using the detection of cell surface activation markers after stimulation with the antigen of choice (Casazza et al., 2006). Clean separation from non-specific background is the primary consideration in the selection of an appropriate sorting strategy. In all cases, cells should be handled carefully, processed as rapidly as possible and sorted immediately after surface staining into an RNA protectant solution. CAUTION: Biosafety practices should be followed rigorously when working with viable blood or tissues.
Purification of mRNA is carried out using the Oligotex Direct mRNA Mini Kit (Qiagen), with small alterations to the manufacturer’s protocol. This method is based on the isolation of pure polyadenylated RNA (polyA+ RNA) from cell lysates in an RNase-free environment using particles coated with oligo(dT), which binds to the polyA+ tail. The beads are then used to pull down the mRNA for washing.
This section describes the synthesis of cDNA with anchor sequence incorporation. The method is based on the template-switching ability of RNAse H- point mutants of MMLV reverse transcriptase (Figure 1).
The SMARTer™ PCR cDNA Synthesis Kit (Clontech) contains several components required for cDNA synthesis, including a proprietary modified SMART oligo, which can be successfully used in this method. However, the 5’CDS oligodT should be synthesized as described above. The 5' PCR primer II A from the SMARTer™ PCR cDNA Synthesis Kit (Clontech) can be used as an alternative to the Universal Primer Mix in Section 5 – PCR amplification of rearranged TR products.
|SMART II oligo||1 µl|
|Add:||5× RT Buffer||2.2 µl|
|dNTP mix||1 µl|
|Superscript II RT||1 µl|
Approximately 20 µl of cDNA is made; it can be stored at −80°C or used immediately in the clean-up procedure prior to PCR.
In the NucleoSpin® Extract II procedure, cDNA binds to a silica membrane in the presence of chaotropic salt added by the NT binding buffer. The binding mixture is loaded directly on to NucleoSpin® Extract II Columns. Contaminants, such as salts and soluble macromolecular components, are removed by a simple washing step with ethanolic wash buffer NT3. Pure cDNA is finally eluted under low ionic strength conditions with the slightly alkaline elution buffer NE (5 mM Tris-HCl, pH 8.5).
Touchdown PCR is performed on clean single-stranded cDNA using TRC-specific and anchor-complementary primers. The TRC-specific primers are designed to enable optimal sequencing across the CDR3 including unequivocal identification of the proximal J region. In the protocol described below, the human TRBC primer MBC2 is used; this primer binds both C1 and C2. Universal primer mix contains two oligonucleotides of different lengths that allow an initial specific extension from the adapter to reduce subsequent non-specific binding. The PCR product is then run on a 1% agarose gel with a 100 bp DNA ladder and excised. Bands containing amplified TR genes are 500–700 bp in length.
The Advantage®2 PCR Kit contains all necessary reagents except the TRC-specific primer and the 10× UPM. In this protocol, MBC2 is used for illustration; all TRC-specific α and β primer sequences for mouse, macaque and human are detailed in Table 1. The 10× UPM can be prepared by mixing the following primers at the indicated concentrations:
|Long (0.4 µM):||5’-CTAATACGACTCACTATAGGGCAAGCAGTGGTATCAACGCAGAGT|
|Short (2 µM):||5’-CTAATACGACTCACTATAGGGC|
|10× PCR buffer||5 µl|
|10× UPM||5 µl|
|MBC2 primer||1 µl|
|dNTP mix||1 µl|
|Water to total 50 µl (depending on template volume)|
|For 1 cycle:||95°C||30 sec|
|For 5 cycles:||95°C||5 sec|
|For 5 cycles:||95°C||5 sec|
|For 30 cycles:||95°C||5 sec|
Amplified TR products are extracted from the gel and cleaned up for sub-cloning.
Amplicons are ligated into plasmids designed to capture adenine overhangs for sub-cloning and sequencing.
The pGEM-T Easy vector contains an ampicillin resistance gene; thus, only transformed bacterial colonies will grow on agar plates in the presence of ampicillin. Blue/white colony screening is used to identify transformants that contain insert-positive vectors; these appear white due to insertional inactivation of the beta-galactosidase gene in the vector.
Single bacterial colony plasmid inserts are amplified in a 96-well plate format using primers flanking the insertion site. White colonies contain the insert and should be selected for amplification and sequencing. It is important to pick only single colonies and to avoid splashing between wells.
|10× HiFi buffer||250µl|
|M13F primer||100 µl|
|M13R primer||100 µl|
|HiFi Taq||14 µl|
|For 1 cycle||95°C||5 min|
|For 35 cycles||95°C||30 sec|
Sequences can be aligned, checked and compiled manually using specific analysis software such as Sequencher (Gene Codes Corporation). Any samples with base ambiguities or sequence quality <80% should be discarded. Sequences encoding pseudogenes, frame-shifts or premature stop codons should also be excluded from the analysis; the latter two scenarios originate from non-productive rearrangements that can maintain basal levels of expression. In addition, erroneous sequences (e.g. GIRLS or GINAEYA) can arise from the universal primer; these should similarly be disregarded. A minimum of 50 functional sequences should be compiled per sorted T cell population. In all cases, chromatograms should be checked visually. The V and J gene segments are identified from their 3' and 5' ends, respectively, using the tables from http://imgt.cines.fr/. As an alternative, the concatenated sequences can be exported as a FASTA file into IMGT/V-QUEST, which can identify the V and J segments. The CDR3 should be displayed as a sequence, as TRBD gene usage is difficult to assign due to N-diversity. Grouped sequences can then be ordered according to their frequency, which reflects the original clonotypic hierarchy in the sorted T cell population due to the linear and unbiased nature of the initial PCR. Further details on bioinformatics tools can be found in Appendix 1W.
The αβ T cell receptor (TR) is a membrane-anchored heterodimer expressed on the surface of T cells that mediates specific recognition of antigen in the form of major histocompatibility complex (MHC) molecules presenting endogenously-derived peptides bound in the α1α2 groove. Surface expression of such peptide-MHC (pMHC) antigens provides a display library that enables T cells to detect abnormal intracellular processes through TR-mediated surveillance, which in turn is the pivotal event that dictates the initiation of a T cell response. Each α and β chain comprises three hypervariable complementarity-determining regions (CDRs), which determine the antigen specificity of the heterodimeric TR. Three mechanisms govern the generation of diversity within the TR repertoire that is necessary to recognize the myriad potential antigenic peptides that a host may encounter (Nikolich-Zugich et al., 2004). First, each TR chain is formed by the genetic rearrangement of variable (V), diversity (D; β chain only) and joining (J) germline gene segments, which vary in number between species, with a TRC gene. Second, nucleotide insertions and deletions at the V(D)J junctions (N-diversity) provide an additional level of variability within the CDR3 of each chain; the CDR1 and CDR2 regions are germline-encoded by the V genes. Third, pairing of individual α and β chains further amplifies the potential number of TRs that can be generated. After thymic selection, which operates to ensure a degree of MHC bias and delete autoreactive TRs, it has been estimated that approximately 2.5 × 107 unique human (Arstila et al., 1999) and 2×106 murine (Casrouge et al., 2000) TRs populate the periphery and are available to respond to antigenic challenges. However, these numbers are dwarfed by the potential number of antigenic pMHC combinations. By necessity, then, an intrinsic degree of cross-reactivity is incorporated within the TR recognition system to enable sufficient coverage and also to increase the likelihood that a given antigen will be recognized by a cognate TR within a time frame that facilitates an effective response (Mason, 1998). Thus, an individual clonotype, i.e. a T cell defined by the singular expressed TR, can recognize multiple ligands; similarly, individual pMHC ligands can productively engage multiple TRs. As a consequence, any antigen-specific T cell response can comprise multiple clonotypes, which in turn dictate the functional qualities of the T cell population. The role of this protocol is to enable the accurate and quantitative analysis of constituent clonotypes within T cell populations specific for defined antigens. A detailed understanding of clonal selection within the memory and effector T cell pools is essential to further our understanding of the factors that influence effective T cell immunity and has direct implications for the rational design of vaccines and immunotherapies (Appay et al., 2008; Price et al., 2009).
Meticulous attention to detail is essential for the successful and reliable application of this protocol. Reagents should be DNase/RNase-free and PCR-grade plastics should be used throughout. Dedicated hoods in a "clean room" free from high copy number plasmids should be used for mRNA extraction, cDNA synthesis and PCR set-up. Benches and pipettes should be cleaned with 10% bleach solution and an RNase-inactivator (e.g. RNase AWAY; Sigma-Aldrich) both before and after each procedure to minimize contamination. Disposable sleeves and gloves should be used throughout. Refer to UNIT 10.20 for detailed recommendations on optimal work practices in a molecular biology laboratory.
As discussed in section (1) above, this protocol is entirely dependent on the isolation of good quality RNA from viable, healthy and pure T cell populations. The sequence output, due to its sensitivity and linearity, will reflect the nature of the starting template. Thus, erroneously captured T cells will lead to the overestimatation of clonality and diversity. Similarly, exclusion of specific T cells, for example due to antigen-induced cell death, can lead to the loss of true clonotypes from the starting template and subsequent underestimation of clonality and diversity.
Related to template quality, adequate sampling of the repertoire under investigation is necessary to ensure reproducibility. Thus, as many cells as possible should be sorted from the initial pool of T cells and the number of sequences generated should reflect the heterogeneity of constituent TRs within the pool. In practice, based on empirical replicate analyses, adequate sampling is achieved for largely oligoclonal pMHC tetramer-sorted T cell populations with >500 cells and >50 sequences. These minimal guidelines should be adjusted, however, according to the nature of the population under investigation. Thus, lower frequency clonotypes will be detected as more sequences are generated; however, the intrinsic sequence error rate will limit the reliable detection of very low frequency clonotypes. Sequence comparisons should account for such issues (Venturi et al., 2008a; Venturi et al., 2007).
If the 5'-RACE PCR does not generate bands of the expected size, repeat the procedure using more cDNA (e.g. 13 µl). Further cDNA can be made from the stored RNA for additional attempts. However, unsuccessful amplification generally reflects a poor starting template, in which case RNA concentration can be useful (see below). Do not use additional PCR cycles or nested approaches to amplification, as these procedures increase the error rate.
If the 5'-RACE PCR gave a distinct band of the expected size but few bacterial colonies contained an insert, product may have been lost during gel extraction. Most notably, ensure that buffer NT3 contains the appropriate quantity of ethanol; this can be reduced due to differential evaporation after reconstitution, especially if the bottle is opened frequently. Ligation efficiency can also be compromised due to loss of adenine overhangs if the PCR product is not processed promptly; thus, always proceed to ligation as quickly as possible once the 5'-RACE PCR is completed. Finally, the quality of the competent bacteria should be verified.
This protocol allows the rapid and comprehensive analysis of all expressed TRs within any given T cell population and offers significant advantages over previous approaches (Rufer, 2005). Multiple facets of TR usage can be assessed depending on the particular issues under investigation. Indeed, fundamental aspects of T cell immunobiology have already been illuminated with this approach (Davenport et al., 2007; Venturi et al., 2008b). It should be noted that allelic exclusion does not operate at the α locus; thus, clonotypic assessment is defined by molecular analysis of TRB gene products.
With experience and good organization, samples can be processed from mRNA extraction to overnight ligation in one day. An approximate time-guide is: mRNA extraction, 90 min; cDNA synthesis and clean-up, 3 hr; RACE-PCR amplification of rearranged TR products, 2 hr 45 min; gel extraction and ligation, 2 hr plus overnight incubation; transformation, 3 hr; colony PCR, 4 hr; preparation of samples for sequencing, 1.5 hr. Up to 4 samples can comfortably be handled in parallel. However, caution should be exercised as scaling up increases the risk of cross-contamination.