This study was funded by JDRF 1-FAC-2014-243-A-N, ADA 7-14-JF-07, NIH 5 P30 DK079638-05 PILOT PJ, and The Robert and Janice McNair Foundation.
We thank Sandra Pena and Andrene McDonald for patient recruitment, Samuel Blum for technical assistance, Dr. George Makedonas for the gift of antibodies and control DNA used for PCR optimization.

1. Identify the T cell population of interest.
NOTE: Antigen specific proliferation in combination with cell division dye (like Carboxyfluorescein succinimidyl ester, CFSE) can be used to isolate T cells based on their proliferation in response to antigenic stimulation. If starting from PBMCs, a 7-day in vitro expansion should be sufficient to identify an antigen specific CFSE low population12.
<ol>
	<li>Mix PBMCs 1:1 with MHC-matched feeder cells, and label with 5 &#181;M CFSE cell division dye.</li>
	<li>Plate 2-2.5 x 105 cells per well in a 96-well round-bottom culture plate in 200 &#181;L 10% FCS complete RPMI media. 
	NOTE: Complete RPMI contains 2 mM l-glutamine, 1 mM sodium pyruvate, 100 &#181;M MEM nonessential amino acids, 5 mM HEPES, 5.5 &#215; 10&#8722; 5 units of 2-mercaptoethanol, 100 U mL&#8722; 1 penicillin and 100 &#181;g mL&#8722; 1 streptomycin.</li>
	<li>Stimulate cells with 25 &#181;M peptide antigen.</li>
	<li>On the fourth day of culture, carefully remove and replace 100 &#181;L of media. 
	NOTE: A more optimal approach for antigen-specific T cell identification is the use of peptide-HLA tetramers13. Tetramer staining allows for simultaneous assessment of antigen specificity, as well as HLA-restriction.</li>
</ol>
2. Set up the single cell sort.
<ol>
	<li>Perform all procedures in a hood or a designated area free of amplified template. Aliquot the reagents to avoid contamination, clean all work surfaces and, if feasible, equipment with 10% bleach prior to PCR setup. Use filter tips, RNAse and DNAse free reagents and plastics in all the PCR and vector cloning steps. Key reagents can be found in the Table of Materials. 
	NOTE: The multiplex PCR reaction is highly sensitive, and low levels of contamination can result in amplified product.</li>
	<li>Generate the reverse transcription master mix by combining 10x RT reaction buffer, 25X DNTPs, 9 U RT enzyme, 0.01% Triton-X, 0.7 U RNAse inhibitor, and 383 nM of each RT-TCR primer (listed in Table 1) in a sterile pipetting reservoir. For each 96-well plate, make up enough master mix for 10 additional reactions (total 106), to compensate for loss of liquid during pipetting. See Table 2 for reaction components and amounts per well.</li>
	<li>Prepare a 96-well PCR collection plate for cell sorting by pipetting 6 &#181;L of the reverse transcription master mix per well with a multichannel pipette. Keep the plate on ice or in a cold block.</li>
	<li>Label cells with antibodies to cell surface markers (such as CD4 and CD3, or CD8 and CD3, each at 2 &#181;g/mL, in combination with cell division dye or tetramer staining).</li>
	<li>Sort T cells at one cell per well, skipping the twelfth row, which will serve as a negative control. 
	NOTE: The efficiency of the sort will depend on precise collection plate alignment within the plate holder, as well as maintaining the cells and plates at a low temperature prior to cDNA reaction. Collection plates should be kept on ice at all times, except during sorting, when they are fixed within the cooling plate. Not all sorters are equipped with a temperature regulated plate holder; however, the use of one is strongly recommended. Since the sort generally takes about 10 - 15 min per plate, there is an increased potential for RNA degradation if the plate is not maintained at a low temperature.</li>
	<li>As a positive control, add a larger number of cells (100 cells) manually with a pipette to one of the wells at this stage. Alternatively, previously isolated RNA from pooled cells can be used as a positive control at 10 ng per well. 
	NOTE: Pipette purified control RNA with precision and care in order to avoid cross-contamination of other wells and false positive results.</li>
	<li>Seal the plate with adhesive film, and spin down at 1000 x g for 5 min at 4 &#176;C.</li>
	<li>Immediately transcribe RNA to cDNA by incubating the plate at 25 &#176;C for 10 min, 37 &#176;C for 45 min, and 85 &#176;C for 10 min. 
	NOTE: To ensure efficient PCR reaction, it is recommended that the first PCR reaction be performed the same day. Alternatively, the transcribed plates can be kept in -80&#176;C for up to 2 months.</li>
</ol>
3. Perform multiplex-nested PCR
NOTE: Prepare all master mixes in a clean template-free area to avoid contamination.
<ol>
	<li>Reconstitute each variable region primer to 100 &#181;M with DNAse and RNAse free H2O. Next, combine 20 &#181;L of each of the 44 primers to generate the V-alpha primer pool. To generate the V-beta primer pool combine 20 &#181;L of each of the 40 V-beta primers plus 80 &#181;L of DNAse and RNAse free H2O. Primer sequences are listed in Table 1. Once pooled, the concentration of each primer in the mix is 2.3 &#181;M.</li>
	<li>Reconstitute the constant region primers to 100 &#181;M stock solution with DNAse and RNAse free H2O. Dilute the constant region primers to 10 &#181;M working solution and aliquot for use.</li>
	<li>Prepare two separate master mixes for TCR-alpha and TCR-beta amplification. Generate the master mixes by combining 5X buffer, 80 &#181;M dNTPs, 3% DMSO, 2 &#181;M primer pool (final concentration of 46 nM for each primer), 200 nM TCR-constant region reverse primer, 1 U DNA polymerase, and DNAse and RNAse free H2O (Table 2). For each 96-well plate, make up enough master mix for 10 additional reactions (total 106), in order to compensate for loss of liquid during pipetting. 
	NOTE: The calculations are based on the final PCR volume of 25 &#181;L per well, where 22.5 &#181;L master PCR mix is combined with 2.5 &#181;L cDNA template.</li>
	<li>Keep the plate with cDNA on ice or in a cold block. Use a multichannel to pipette 22.5 &#181;L of TCR-beta reaction into a new 96-well PCR plate. Then use a multichannel to transfer 2.5 &#181;L of cDNA into the PCR plate.</li>
	<li>Use a multichannel to pipette 22.5 &#181;L of TCR-alpha reaction directly into the plate containing the remainder of the cDNA.</li>
	<li>Seal the plates with adhesive, gently vortex, and spin down at 1000 x g for 5 min at 4 &#176;C.</li>
	<li>Run the multiplex PCR reaction with the following cycles: 5 min at 95 &#176;C followed by 34 cycles of 20 s at 95 &#176;C, 30 s at 54 &#176;C, and 1 min at 72 &#176;C, followed by 7 min at 72 &#176;C.</li>
	<li>Carry out the nested PCR reaction in a final volume of 25 &#181;L, using 2.5 &#181;L of the multiplex PCR mixture as template.
	<ol>
		<li>Reconstitute the internal and adaptor primers to 100 &#181;M with DNAse and RNAse free H2O. Make 10 &#181;M aliquots for the working stock.</li>
		<li>Prepare two separate master mixes for TCR-alpha and TCR-beta amplification. Mix 5X buffer, 25X DNTP, 3% DMSO, 200 nM forward adaptor primer, 200 nM reverse nested primer, 1 U DNA polymerase, and DNA polymerase and nuclease-free H2O for a final volume of 22.5 &#181;L (Table 2). For each 96-well plate, make up enough master mix for 10 additional reactions (total 106), to compensate for loss of liquid during pipetting. 
		NOTE: In the second PCR reaction use the 5X PCR buffer containing loading dye to facilitate agarose gel loading.</li>
		<li>Using a multichannel, pipette the master mixes for the TCR-alpha and TCR-beta reactions at 22.5 &#181;L per well into two new 96-well PCR plates.</li>
		<li>Add the multiplex PCR reaction template at 2.5 &#181;L per well. 
		NOTE: The remaining first PCR reaction should be sealed, and stored at -20 &#176;C to serve as a source of additional template, if it becomes necessary (see step 4.3 Note).</li>
		<li>Seal the plates, gently vortex, and spin down at 1000 x g for 5 min at 4 &#176;C.</li>
		<li>Run the nested PCR reaction with the following cycles: 5 min at 95 &#176;C followed by 34 cycles of 20 s at 95 &#176;C, 30 s at 56 &#176;C, and 1 min at 72 &#176;C, followed by 7 min at 72 &#176;C.</li>
		<li>Run 5 &#181;L of the final reaction out on a 1% agarose gel containing ethidium bromide (including the negative and positive control wells). 
		NOTE: The expected band should run at around 500 bp size. An example reaction is shown in Figure 2.</li>
		<li>Purify the remainder of the second reaction (20 &#181;L) using a 96-well format PCR purification kit, elute with 15 &#181;L of 70 &#176;C H2O per well, and perform Sanger sequencing. Use the appropriate TCR-alpha or TCR-beta adaptor forward primer for sequencing (Table 1). 
		NOTE: Sequences can be analyzed with online available immunogenetics software.</li>
	</ol>
	</li>
</ol>
4. Sub-cloning alpha and beta PCR chains.
NOTE: The pool of the forward primers utilized in the first reaction, and the reverse primers in the second PCR reaction are used to incorporate restriction sites. Therefore, the obtained alpha and beta chain PCR products are ready to be sequentially sub-cloned into the template retroviral vector (Figure 1).
<ol>
	<li>Digest the purified PCR products from the beta nested reaction with BstbI and MfeI (Table 3). 
	NOTE: Do not exceed 5 &#181;g insert.</li>
	<li>In parallel, digest the template vector with BstbI and MfeI (Table 3). Use 1-2 &#181;g of the template vector per TCR. Run the digested vector on an agarose gel, and isolate the larger band (~7500 bp). Purify the vector with a gel DNA purification kit. 
	NOTE: During the restriction enzyme digest, the template murine TCR-beta variable region is removed, allowing for the addition of the human TCR-beta variable region.</li>
	<li>Purify the digested PCR products using a PCR purification kit. 
	NOTE: Verify the DNA purification capacity of the columns included in the kit; use several columns if necessary. The minimum amount of a purified insert necessary for a successful ligation reaction is 50 ng at a minimum concentration of 10 ng/&#181;L. If the purified insert amount is not sufficient for a successful ligation, the second PCR reaction can be repeated.</li>
	<li>Treat the template vector with 10 U CIP enzyme for 1 h at 37 &#176;C to prevent self-ligation, followed by heat inactivation for 10 min at 75 &#176;C. Purify the DNA with a PCR purification kit (Table 3).</li>
	<li>Ligate TCR-beta insert and vector using DNA ligase in a 20 &#181;L total volume at 6 molar excess of the insert for 30 min at room temperature. 
	NOTE: In total, the ligation reaction should contain about 150 ng of DNA (Table 3).</li>
	<li>For transformations, mix 8 &#181;L of the ligation reaction with 80 &#181;L of chemically competent E.coli DH5&#945; cells (Table of Materials) and incubate on ice for 30 min. Heat shock at 42 &#176;C for 45 s. Add 500 &#181;L Super Optimal broth with Catabolite suppression (S.O.C.) medium and shake for 1.5 h at 37 &#176;C.</li>
	<li>Plate out 250 &#181;L of bacterial culture on an ampicillin containing Lysogeny Broth (LB) agar plate. Incubate the plate overnight (~16 h) at 37 &#176;C.</li>
	<li>The next day pick bacterial colonies, and grow them up in 3 mL LB medium containing ampicillin overnight. Isolate the plasmid DNA using DNA plasmid purification kit.</li>
	<li>Confirm the successful insertion of the beta chain by a small-scale test-digest with the restriction enzymes BstbI and MfeI. In a 12 &#181;L total reaction volume, combine 200-500 &#181;g plasmid DNA, 0.25 &#181;L of each enzyme, and 10X enzyme buffer (Table 3). Run the reaction out on a 1% agarose gel containing ethidium bromide to confirm the size of the insert band (~500 bp).</li>
	<li>After confirming the presence of the insert, sequence the TCR-beta section of the vector using the IRES reverse primer (Table 1).</li>
	<li>Following sequence confirmation of the TCR-beta insert, perform a similar process for insertion of the alpha variable region. Digest the purified PCR products from the alpha nested reaction, and the newly generated TCR-beta insert containing vector, with SnaBI and SacII. 
	NOTE: During the restriction enzyme digest, the template murine TCR-alpha variable region is removed, allowing for the addition of the human TCR-alpha variable region.</li>
	<li>Ligate alpha inserts into TCR vectors encoding the corresponding beta variable region using DNA ligase in a 20 &#181;L total volume at 6 molar excess of the insert for 30 min at room temperature. 
	NOTE: In total, the ligation reaction should contain about 150 ng of DNA (Table 3).</li>
	<li>For transformations, mix 8 &#181;L of the ligation reaction with 80 &#181;L of chemically competent E.coli DH5&#945; cells (Table of Materials) and incubate on ice for 30 min. Heat shock at 42 &#176;C for 45 s. Add 500 &#181;L S.O.C. medium and shake for 1.5 h at 37 &#176;C.</li>
	<li>Plate out 250 &#181;L of bacterial culture on an ampicillin containing Lysogeny Broth (LB) agar plate. Incubate the plate overnight (~16 h) at 37 &#176;C.</li>
	<li>The next day pick bacterial colonies, and grow them up in 3 mL LB medium containing ampicillin overnight. Isolate the plasmid DNA using DNA plasmid purification kit.</li>
	<li>Confirm the successful insertion of the alpha chain by a small-scale test-digest with the restriction enzymes SnaBI and SacII. In a 12 &#181;L total reaction volume combine 200-500 &#181;g plasmid DNA, 0.25 &#181;L of each enzyme, and 10x enzyme buffer (Table 3). Run the reaction out on a 1% agarose gel containing ethidium bromide to confirm the size of the insert band (~500 bp).</li>
	<li>After confirming the presence of the insert, sequence the TCR-alpha section of the vector using the MSCV2-forward primer (Table 1).</li>
	<li>Once the alpha chain sequence is validated, verify the complete TCR insert by sequencing with TCR-beta-reverse and IRES-reverse primer (Table 1).</li>
</ol>
5. Verify TCR cell surface expression and specificity.
Note: HEK293T cells are used to test successful TCR chain paring and cell surface expression (Figure 3A)8. Peptide-MHC tetramer staining can also be performed on the transfected HEK293T cells to test for retention of antigenic specificity post TCR gene isolation (Figure 3B).
<ol>
	<li>Plate out HEK293T cells in 12-well tissue culture plates at 1 x 105 cells per well in 1 mL complete DMEM media containing 5% FCS, and culture overnight at 37 &#176;C. Plate out enough cells to have designated separate wells for each new TCR, control template TCR vector, CD3 vector only, control TCR vector only, and a non-transfected well. 
	NOTE: Transfection with a fully mouse template TCR vector (mTCR-pMIA) in combination with mCD3&#949;&#947;&#948;&#950;-pMIG vector, mTCR-pMIA vector alone, and mCD3&#949;&#947;&#948;&#950;-pMIG vector alone are used as positive and negative controls. 
	NOTE: Complete DMEM contains 2 mM L-glutamine, 1 mM Sodium Pyruvate, 100 &#181;M Non-essential Amino Acids, 5 mM HEPES buffer, 5.5 x 10-5 units of 2-mercaptoethanol, 100 U/ml Penicillin/Streptomycin.</li>
	<li>The following day, transfect the cells with 1 &#181;g chimeric h/mTCR-pMIA vector, 1 &#181;g mouse CD3&#949;&#947;&#948;&#950;-pMSCVII-GFP (pMIG) vector, and a liposome-based transfection reagent following manufacturer instructions.</li>
	<li>For each TCR, add 6 &#181;L transfection reagent to 100 &#181;L room temperature, serum-free DMEM. Briefly vortex, and incubate at room temperature for 15 min.</li>
	<li>In separate 1.5 mL tubes combine 1 &#181;g of TCR-pMIA vector with 1 &#181;g mouse CD3&#949;&#947;&#948;&#950;-pMSCVII-GFP (pMIG) vector, or 1 &#181;g of each vector separate for controls.</li>
	<li>Dropwise, add the transfection reagent/DMEM solution to the tubes containing the vector DNA. Incubate at room temperature for 15 min.</li>
	<li>Dropwise, add the transfection reagent/DMEM/DNA solution to the HEK293T cells. Gently tap the plate to mix. Incubate at 37 &#176;C for 24 h.</li>
	<li>After 24 h replace the media, and keep cells in culture for additional 24 h.</li>
	<li>Remove cells from the plate by vigorous pipetting, and stain the cells with anti-mouse CD3 antibody (2.5 &#181;g/mL) to verify surface expression of the chimeric-TCR by flow cytometry.</li>
	<li>In order to compare CD3 expression between cells that were transfected to a similar level, gate the analysis on cells expressing a similar high level of fluorescent reporter molecules (GFP for CD3 complex, and Ametrine for TCR expression vector). Optimally, the analysis should be gated on GFP+Ametrine+ cells within 104 to 105 fluorescent intensity (Figure 3A). 
	NOTE: The level of CD3 expression in cells transfected with the chimeric construct should be comparable to the control mouse (original template) vector (Figure 3A). The generated TCR retroviral vectors are compatible with in vivo expression systems as previously described4.</li>
</ol>

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The authors have nothing to disclose.

In the current protocol, we describe an efficient method for single cell TCR amplification and subsequent sub-cloning of paired TCR alpha and beta chains into a template retroviral expression vector. Although, several single cell PCR protocols have been developed, none so far have been compatible with immediate sub-cloning into an expression vector. In most cases, a partial sequence encompassing the highly variable CDR3 regions is amplified, with enough sequence to extrapolate the variable region. This smaller PCR product size supports higher PCR efficiency, but necessitates de novo TCR gene synthesis for functional studies. The current protocol maintains the efficiency of previously published CDR3-focused single cell TCR sequencing protocols, while at the same time yielding a longer amplicon containing the whole variable region suitable for cloning. Therefore, the system described has an increased flexibility in giving a quantitative sequence, as well as functional output.
Depending on the scientific question addressed, the source and specificity of T cells analyzed can vary. However, if the end goal of the project is the analysis of the in vivo TCR function, it is necessary to know the HLA restriction of isolated T cells. The current protocol has been specifically designed to be compatible with in vivo TCR expression in HLA-humanized mice. Many of these mice have been generated and are now commercially available, including HLA-A2.1, HLA-DQ8 (HLA-DQA1*301,HLA-DQB1*302), HLA-DQ6 (DQA1*0102,HLADQB1*0602), HLA-DR4 (DRB1*0401), HLA-DR1 (DRA*0101, DRB1*0101), HLA-A11 (A*11:01/K), HLA-B2705, HLA-A24, and HLA-B*0702 transgenic strains. This protocol can be successfully combined with the previously published protocol for retroviral mediated bone marrow stem cell expression of human TCRs4,15,16. We expect that the humanized TCR retrogenic system will be applicable and highly useful for functional analysis of human TCRs in various autoimmune, cancer, and infectious models. 
The current single cell PCR protocol describes the critical steps and gives suggestions necessary for successful amplification of paired TCR-alpha and TCR-beta sequences. The first critical step is a timely performance of cDNA transcription, and subsequent first round of multiplex PCR. All the steps involved should be performed in a timely manner, and all the reagents and cells kept on ice to prevent RNA and cDNA degradation. Secondly, because the protocol is designed to be highly sensitive to low levels of the template, it is highly susceptible to contamination. Therefore, it is imperative to work in a template-free area, away from amplified PCR products or TCR vector cloning. A separate room or hood should be used to set up cDNA and PCR reactions. Template from the first PCR reaction should be added in a different area from where the PCR is set up. It is advisable that a separate set of aliquoted reagents be used for the single cell PCR experiments.
This protocol has been specifically optimized using the reagents listed in Table of Materials, and any substitutions in reagents will require additional optimization. The initial identification of antigen-specific cells can be modified to use peptide-MHC tetramers. Tetramer staining allows for simultaneous assessment of antigen specificity, as well as HLA-restriction. Alternative methods for the detection of antigen reactivity, such as upregulation of early activation markers or secretion of inflammatory cytokines can be considered in the absence of tetramer reagents. For example, dual specificity fusion antibodies specific for a T cell antigen and IFN&#947; can be combined with magnetic bead isolation to enrich for antigen reactive IFN&#947; producing cells. 
While the chimeric human/mouse construct described in our protocol ensures compatibility with mouse cells, the level of its compatibility with human CD3 complex is unknown. Although not formally tested by us, others have shown that chimeric TCR constructs with murine constant regions can be expressed on the surface of human lymphocytes17. This indicates that murine TCR constant regions can support interaction with the human CD3 signaling complex. However, if the goal is to express the identified TCRs in human primary cells or human cell lines, such as Jurkat cells, a more optimal approach may be to use a fully human construct. This can be accomplished by swapping the murine constant regions within the vector with human constant regions. 
If an identified TCR variable region contains one of the restriction sites used for cloning, the TCR should be inserted into a shuttle vector and subsequently mutated using a site directed mutagenesis kit to induce a silent nucleotide change within the restriction cut site. An alternative approach is de novo synthesis of the TCR variable region of interest modified to exclude the unwanted restriction site. 
The main limitation of this technique is the inability to eliminate amplification of secondary 'cytoplasmic' alpha chains. TCR-alpha genes do not go through allelic exclusion like TCR-beta genes, thus a significant proportion of T cells will express a secondary 'cytoplasmic' alpha chain14. The described PCR protocol only results in amplification of one TCR alpha chain, but if the amplified alpha chain is the 'cytoplasmic' alpha chain it may not pair with the amplified beta chain. Therefore, it is imperative to test TCR cell surface expression in HEK293T cells to ensure successful chain pairing.

The T cell receptor (TCR) dictates T cell fate decision during development, steady state/homeostasis, and antigenic stimulation in periphery1,2,3. Recent expansion of deep sequencing technologies has uncovered a previously underappreciated TCR diversity within antigen specific T cell responses. TCR diversity suggests a potential for functionally broad T cell responses. In order to integrate the TCR sequence repertoire analysis with TCR functional assays, the sequencing approaches should be designed to be compatible with experimental systems and in vivo models utilized for subsequent functional analysis of select TCRs. We have developed an efficient approach for human TCR sequence isolation and streamlined sub-cloning into a chimeric human/mouse TCR template vector compatible with TCR expression in HLA-humanized mice4. Isolation of corresponding alpha and beta chains of heterodimeric TCRs requires PCR amplification of both chains from a single cell. Although, several single-cell TCR cloning protocols have been developed and utilized, none so far have been easily compatible with high-throughput streamlined direct cloning of unknown V&#945;/V&#946; TCRs into retroviral vectors necessary for re-expression in vivo4,5,6,7. Previous studies have utilized two major approaches, either to selectively amplify a limited portion of the TCR sufficient to extrapolate the sequence, or to amplify the entire TCR sequence4,5,6,7. Downstream functional analysis of TCR sequences obtained via the first approach requires costly in silico assembly and de novo construction of the TCR. While the second approach provides the complete TCR sequence, the human constant region is not compatible with in vivo expression of the cloned TCRs in mouse models. Our approach is specifically designed to be compatible with in vivo functional analysis of TCRs in mouse models. We developed an efficient and streamlined single cell PCR protocol that allows for direct sub-cloning of PCR fragments into the template expression vector.
Our approach utilizes a highly sensitive multiplex-nested PCR reaction that is performed in two steps. In the first multiplex reaction step a pool of 40 primers specific for all the V-beta chains and a pool of 44 primers specific for all the V-alpha chains are used to amplify the TCR-alpha or TCR-beta without the prior knowledge of the sequence (Table 1). The forward primer has an adaptor sequence, which is incorporated into the 5&#39; of the PCR product. The reverse primer is based on the constant region of the TCR. We have identified unique restriction sites that are absent from human variable or junction TCR regions, and incorporated these into newly redesigned TCR&#945; and TCR&#946; primers (Figure 1). In the second nested reaction, a primer specific for the adaptor sequence and a nested reverse primer within the constant region are used to further amplify the TCR chains with increased specificity (Table 1, Figure 1). After single cell isolation, two rounds of PCR (first reaction with a pool of both V&#945; and V&#946; primers, and second with adaptor primers) result in a PCR product that can be directly sub-cloned into the template retroviral vector. The final TCR construct will encode, in a single open reading frame (ORF), human variable regions combined with mouse constant regions connected by the &#39;self-cleaving&#39; protein sequence, P2A (hV&#945;-mC&#945;-P2A-hV&#946;-mC&#946;)8. The P2A sequence has been used in multiple systems, and has been extensively tested specifically for the expression of TCRs8,9,10,11. Although, after translation most of the P2A sequence remains attached to the C-terminus of the alpha chain connected by a flexible linker, while the beta chain signal sequence has an additional proline, this modification has no detrimental effect on TCR function. The mouse constant region is used in the construct instead of human to avoid potential altered interactions with downstream signaling components when re-expressed in mouse cells. The single ORF will result in stoichiometric separate expression of alpha and beta chains9,11. The current protocol is based on reagents and approaches that are widely available, and is designed to be performed in a streamlined and efficient fashion. Although we have specifically used this technique to assay TCRs from self-reactive T cells implicated in autoimmune diabetes, we anticipate that this protocol can be widely applicable for identification and functional assessment of human TCRs specific for autoimmune epitopes, cancer epitopes, or responses to pathogens and vaccines.

<table><tbody><tr><td>CFSE&amp;nbsp;</td><td>eBioscience&amp;nbsp;</td><td>65-0850-84</td><td>5 mM</td></tr><tr><td>anti-human CD4 (OKT4) BV421</td><td>Biolegend&amp;nbsp;</td><td>317433</td><td>2 mg/mL</td></tr><tr><td>anti-human CD3 (OKT3) PerCP/Cy5.5</td><td>Biolegend&amp;nbsp;</td><td>317336</td><td>2 mg/mL</td></tr><tr><td>anti-mouse CD3 AF647</td><td>Biolegend&amp;nbsp;</td><td>100322</td><td>2.5 mg/mL</td></tr><tr><td>Recombinant RNAse inhibitor biotech grd 2.5KU&amp;nbsp;&amp;nbsp;</td><td>VWR</td><td>97068-158</td><td>0.7 U</td></tr><tr><td>Applied Biosystems High capaity cDNA RT kit&amp;nbsp;</td><td>Invirtogen&amp;nbsp;</td><td>4368814</td><td>9 U (RT enzyme)&amp;nbsp;</td></tr><tr><td>NF-H2O: Nuclease-free water 1000mL</td><td>Invirtogen&amp;nbsp;</td><td>AM9932</td><td></td></tr><tr><td>Gotaq DNA polymerase 2500U</td><td>VWR</td><td>PAM3008</td><td>1 U</td></tr><tr><td>96 Well Half Skirt PCR Plate</td><td>Phenix</td><td>MPX-96M2</td><td></td></tr><tr><td>MicroAmp Clear Adhesive Film&amp;nbsp;</td><td>ThermoFisher&amp;nbsp;</td><td>4306311</td><td></td></tr><tr><td>UltraPure Agarose&amp;nbsp;</td><td>Invirtogen&amp;nbsp;</td><td>15510-027</td><td></td></tr><tr><td>SnaBI</td><td>New England Biolabs&amp;nbsp;</td><td>R0130</td><td>15 U (insert) 30 U (vector)</td></tr><tr><td>SacII</td><td>New England Biolabs&amp;nbsp;</td><td>R0157</td><td>40 U (insert) 80 U (vector)</td></tr><tr><td>MfeI</td><td>New England Biolabs&amp;nbsp;</td><td>R3589S</td><td>40 U (insert) 80 U (vector)</td></tr><tr><td>BstBI</td><td>New England Biolabs&amp;nbsp;</td><td>R0519</td><td>40 U (insert) 80 U (vector)</td></tr><tr><td>Calf Intestinal Phosphatase (CIP)</td><td>New England Biolabs&amp;nbsp;</td><td>M0290S</td><td>10 U</td></tr><tr><td>Quick ligase&amp;nbsp;</td><td>New England Biolabs&amp;nbsp;</td><td>M2200S</td><td></td></tr><tr><td>Wizard Plus minipreps DNA purification systems&amp;nbsp;</td><td>Promega&amp;nbsp;</td><td>A1460</td><td></td></tr><tr><td>Wizard Plus midipreps DNA purification systems&amp;nbsp;</td><td>Promega&amp;nbsp;</td><td>A7640</td><td></td></tr><tr><td>DNA clean &amp;amp; concentrator-25 kit</td><td>Genesee Scientific&amp;nbsp;</td><td>11-304C</td><td></td></tr><tr><td>ZR-96 DNA clean &amp;amp; concentrator-5 kit</td><td>Genesee Scientific&amp;nbsp;</td><td>11-306A</td><td></td></tr><tr><td>QIAquick gel extraction kit</td><td>Qiagen</td><td>28704</td><td></td></tr><tr><td>QIAquick PCR purification kit</td><td>Qiagen</td><td>28104</td><td></td></tr><tr><td>Subcloning Efficiency DH5a Competent Cells&amp;nbsp;</td><td>ThermoFisher&amp;nbsp;</td><td>18265017</td><td></td></tr><tr><td>Transit LT-1 Reagent 1mL (transfection reagent)</td><td>Mirus&amp;nbsp;</td><td>MIR2300</td><td></td></tr><tr><td>BD FACS Aria II (sorter)</td><td>BD</td><td></td><td></td></tr><tr><td>BD Fortessa (flow cytometer)</td><td>BD&amp;nbsp;</td><td></td><td></td></tr></tbody></table>

streamlined single cell tcr isolation and generation of retroviral vectors for in vitro and in vivo expression of human tcrs

Although, several methods for sequencing of paired T cell receptor (TCR) alpha and beta chains from single T cells have been developed, none so far have been conducive to downstream in vivo functional analysis of TCR heterodimers. We have developed an improved protocol based on a two-step multiplex-nested PCR, which results in a PCR product that spans entire variable regions of a human TCR alpha and beta chains. By identifying unique restriction sites and incorporating them into the PCR primers, we have made the PCR product compatible with direct sub-cloning into the template retroviral vector. The resulting retroviral construct encodes a chimeric human/mouse TCR with a mouse intracellular domain, which is functional in mouse cells or in in vivo mouse models. Overall, the protocol described here combines human single cell paired TCR alpha and beta chain identification with streamlined generation of retroviral vectors readily adaptable for in vitro and in vivo TCR expression. The video and the accompanying material are designed to give a highly detailed description of the single cell PCR, so that the critical steps can be followed and potential pitfalls avoided. Additionally, we provide a detailed description of the cloning steps necessary to generate the expression vector. Once mastered, the whole procedure from single cell sorting to TCR expression could be performed in a short two-week period.

Watch this Scientific Journal Video about Streamlined Single Cell TCR Isolation and Generation of Retroviral Vectors for In Vitro and In Vivo Expression of Human TCRs at JoVE.com

Streamlined Single Cell TCR Isolation and Generation of Retroviral Vectors for In Vitro and In Vivo Expression of Human TCRs

The current protocol combines single cell paired human TCR alpha and beta chain sequencing with streamlined generation of retroviral vectors compatible with in vitro and in vivo TCR expression.

Streamlined Single Cell TCR Isolation and Generation of Retroviral Vectors for In Vitro and In Vivo Expression of Human TCRs

Although, several methods for sequencing of paired T cell receptor (TCR) alpha and beta chains from single T cells have been developed, none so far have ...

streamlined-single-cell-tcr-isolation-generation-retroviral-vectors

Universidad Científica del Sur

Research

JoVE Journal

Immunology and Infection

9.4K Views.  Baylor College of Medicine, Texas Children's Hospital. The current protocol combines single cell paired human TCR alpha and beta chain sequencing with streamlined generation of retroviral vectors compatible with in vitro and in vivo TCR expression.

Video: Streamlined Single Cell TCR Isolation and Generation of Retroviral Vectors for In Vitro and In Vivo Expression of Human TCRs

Retroviral Transduction of T-cell Receptors in Mouse T-cells

T-cell receptors (TCRs) play a central role in the immune system. TCRs on T-cell surfaces can specifically recognize peptide antigens presented by antigen presenting cells (APCs)1. This recognition leads to the activation of T-cells and a series of functional outcomes (e.g. cytokine production, killing of the target cells). Understanding the functional role of TCRs is critical to harness the power of the immune system to treat a variety of immunology related diseases (e.g. cancer or autoimmunity).
It is convenient to study TCRs in mouse models, which can be accomplished in several ways. Making TCR transgenic mouse models is costly and time-consuming and currently there are only a limited number of them available2-4. Alternatively, mice with antigen-specific T-cells can be generated by bone marrow chimera. This method also takes several weeks and requires expertise5. Retroviral transduction of TCRs into in vitro activated mouse T-cells is a quick and relatively easy method to obtain T-cells of desired peptide-MHC specificity. Antigen-specific T-cells can be generated in one week and used in any downstream applications. Studying transduced T-cells also has direct application to human immunotherapy, as adoptive transfer of human T-cells transduced with antigen-specific TCRs is an emerging strategy for cancer treatment6.
Here we present a protocol to retrovirally transduce TCRs into in vitro activated mouse T-cells. Both human and mouse TCR genes can be used. Retroviruses carrying specific TCR genes are generated and used to infect mouse T-cells activated with anti-CD3 and anti-CD28 antibodies. After in vitro expansion, transduced T-cells are analyzed by flow cytometry.

We present a protocol to produce antigen-specific mouse T-cells using retroviral transduction

T-cell receptors (TCRs) play a central role in the immune system. TCRs on T-cell surfaces can specifically recognize peptide antigens presented by antigen ...

Retroviral Transduction of Bone Marrow Progenitor Cells to Generate T-cell Receptor Retrogenic Mice

T cell receptor (TCR) signaling is essential in the development and differentiation of T cells in the thymus and periphery, respectively. The vast array of TCRs proves studying a specific antigenic response difficult. Therefore, TCR transgenic mice were made to study positive and negative selection in the thymus as well as peripheral T cell activation, proliferation and tolerance. However, relatively few TCR transgenic mice have been generated specific to any given antigen. Thus, studies involving TCRs of varying affinities for the same antigenic peptide have been lacking. The generation of a new TCR transgenic line can take six or more months. Additionally, any specific backcrosses can take an additional six months. In order to allow faster generation and screening of multiple TCRs, a protocol for retroviral transduction of bone marrow was established with stoichiometric expression of the TCR&#945; and TCR&#946; chains and the generation of retrogenic mice. Each retrogenic mouse is essentially a founder, virtually negating a founder effect, while the length of time to generate a TCR retrogenic is cut from six months to approximately six weeks. Here we present a rapid and flexible alternative to TCR transgenic mice that can be expressed on any chosen background with any particular TCR.

We present a rapid and flexible protocol for a single T cell receptor (TCR) retroviral-based in vivo expression system. Retroviral vectors are used to transduce bone marrow progenitor cells to study T cell development and function of a single TCR in vivo as an alternative to TCR transgenic mice.

T cell receptor (TCR) signaling is essential in the development and differentiation of T cells in the thymus and periphery, respectively.  The vast array ...

Generating De Novo Antigen-specific Human T Cell Receptors by Retroviral Transduction of Centric Hemichain

T cell receptors (TCRs) are used clinically to direct the specificity of T cells to target tumors as a promising modality of immunotherapy. Therefore, cloning TCRs specific for various tumor-associated antigens has been the goal of many studies. To elicit an effective T cell response, the TCR must recognize the target antigen with optimal affinity. However, cloning such TCRs has been a challenge and many available TCRs possess sub-optimal affinity for the cognate antigen. In this protocol, we describe a method of cloning de novo high affinity antigen-specific TCRs using existing TCRs by exploiting hemichain centricity. It is known that for some TCRs, each TCR&#945; or TCR&#946; hemichain do not contribute equally to antigen recognition, and the dominant hemichain is referred to as the centric hemichain. We have shown that by pairing the centric hemichain with counter-chains differing from the original counter-chain, we are able to maintain the antigen specificity, while modulating its interaction strength for the cognate antigen. Thus, the therapeutic potential of a given TCR can be improved by optimizing the pairing between the centric and counter hemichains.

Generating De Novo Antigen-specific Human T Cell Receptors by Retroviral Transduction of Centric Hemichain

Herein we describe a novel method to generate antigen-specific T cell receptors (TCRs) by pairing the TCR&#945; or TCR&#946; of an existing TCR, possessing the antigen-specificity of interest, with complementary hemichain of the peripheral T cell receptor repertoire. The de novo generated TCRs retain antigen-specificity with varying affinity.

T cell receptors (TCRs) are used clinically to direct the specificity of T cells to target tumors as a promising modality of immunotherapy. Therefore, ...

Retroviral Transduction of Helper T Cells as a Genetic Approach to Study Mechanisms Controlling their Differentiation and Function

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents autoimmunity. Many approaches involving different model organisms have been utilized to understand the mechanisms controlling helper T cell development and function. However, studies using mouse models have proven to be highly informative due to the availability of genetic, cellular, and biochemical systems. One genetic approach in mice used by many labs involves retroviral transduction of primary helper T cells. This is a powerful approach due to its relative ease, making it accessible to almost any laboratory with basic skills in molecular biology and immunology. Therefore, multiple genes in wild type or mutant forms can readily be tested for function in helper T cells to understand their importance and mechanisms of action. We have optimized this approach and describe here the protocols for production of high titer retroviruses, isolation of primary murine helper T cells, and their transduction by retroviruses and differentiation toward the different helper subsets. Finally, the use of this approach is described in uncovering mechanisms utilized by microRNAs (miRNAs) to regulate pathways controlling helper T cell development and function.

Many experimental systems have been utilized to understand the mechanisms regulating T cell development and function in an immune response. Here a genetic approach using retroviral transduction is described, which is economic, time efficient, and most importantly, highly informative in identifying regulatory pathways.

Helper T cell development and function must be tightly regulated to induce an appropriate immune response that eliminates specific pathogens yet prevents ...

Generation of Human Regulatory T Cell Clones

Human regulatory T cells (Treg) are notoriously difficult to isolate in high purity given the current methods of Treg enrichment. These methods are based on the identification of Treg through several activation-dependent cellular surface markers with varying expression levels in different physiologic and pathologic conditions. Populations isolated as &#8220;Treg&#8221; therefore often contain considerable numbers of non-Treg effector cells (i.e., Teff) which hamper the precise phenotypic and functional characterization of these cells, their genomic and proteomic characterization, their reliable enumeration in different states of health and disease, as well as their isolation and expansion for therapeutic purposes. The latter, in particular, remains a major hurdle, as the inadvertent expansion of effector cells homing in Treg-relevant cellular compartments (e.g., CD4+CD25+ T cells) may render Treg-based immunotherapy ineffective, or even harmful. This work presents a method that circumvents the problems associated with population-based isolation and expansion of Treg and shows that the generation of Treg candidate clones with the subsequent selection, culture, and expansion of only carefully vetted, monoclonal cells, enables the generation of an ultrapure Treg cell product that can be kept in culture for many months, enabling downstream investigation of these cells, including for possible therapeutic applications.

This protocol describes the cloning and expansion of human regulatory T cells for the generation of ultra-high purity viable human Treg with stable demethylation at the Treg-specific demethylated region (TSDR) and Treg-specific phenotypic features.

Human regulatory T cells (Treg) are notoriously difficult to isolate in high purity given the current methods of Treg enrichment. These methods are based ...

Production of Human CRISPR-Engineered CAR-T Cells

Adoptive cell therapies using chimeric antigen receptor T cells (CAR-T cells) have demonstrated remarkable clinical efficacy in patients with hematological malignancies and are currently being investigated for various solid tumors. CAR-T cells are generated by removing T cells from a patient's blood and engineering them to express a synthetic immune receptor that redirects the T-cells to recognize and eliminate target tumor cells. Gene editing of CAR-T cells has the potential to improve safety of current CAR-T cell therapies and further increase the efficacy of CAR-T cells. Here, we describe methods for the activation, expansion, and characterization of human CRISPR-engineered CD19 directed CAR-T cells. This comprises transduction of the CAR lentiviral vector and use of single guide RNA (sgRNA) and Cas9 endonuclease to target genes of interest in T cells. The methods described in this protocol can be universally applied to other CAR constructs and target genes beyond the ones used for this study. Furthermore, this protocol discusses strategies for gRNA design, lead gRNA selection and target gene knockout validation to reproducibly achieve high-efficiency, multiplex CRISPR-Cas9 engineering of clinical grade human T cells.

Here, we present a protocol for gene editing in primary human T cells using CRISPR Cas Technology to modify CAR-T cells.

Adoptive cell therapies using chimeric antigen receptor T cells (CAR-T cells) have demonstrated remarkable clinical efficacy in patients with ...

Streamlined Protocol for Mouse CAR-T Cell Generation in Syngeneic Models

Efficient Generation of Murine Chimeric Antigen Receptor (CAR)-T Cells

Engineered cell therapies utilizing chimeric antigen receptor (CAR)-T cells have achieved remarkable effectiveness in individuals with hematological malignancies and are presently undergoing development for the treatment of diverse solid tumors. So far, the preliminary evaluation of novel CAR-T cell products has predominantly taken place in xenograft tumor models using immunodeficient mice. This approach is chosen to facilitate the successful engraftment of human CAR-T cells in the experimental setting. However, syngeneic mouse models, in which tumors and CAR-T cells are derived from the same mouse strain, allow evaluation of new CAR technologies in the context of a functional immune system and comprehensive tumor microenvironment (TME). The protocol described here aims to streamline the process of mouse CAR-T cell generation by presenting standardized methods for retroviral transduction and ex vivo T cell culture. The methods described in this protocol can be applied to other CAR constructs beyond the ones used in this study to enable routine evaluation of new CAR technologies in immune-competent systems.

Author Spotlight: Enhancing CAR-T Cell Function in Syngeneic Tumor Models 

This protocol streamlines retroviral vector production and murine T cell transduction, facilitating the efficient generation of mouse CAR-T cells.

Engineered cell therapies utilizing chimeric antigen receptor (CAR)-T cells have achieved remarkable effectiveness in individuals with hematological ...

Bioengineering

Retroviral Transduction of Guide RNA for Gene Editing in Th17 Differentiation

Retroviral CRISPR/Cas9-Mediated Gene Targeting for the Study of Th17 Differentiation in Vitro

T helper cells that produce IL-17A, known as Th17 cells, play a critical role in immune defense and are implicated in autoimmune disorders. CD4 T cells can be stimulated with antigens and well-defined cytokine cocktails in vitro to mimic Th17 cell differentiation in vivo. Research has been conducted extensively on the Th17 differentiation regulation mechanisms using the in vitro Th17 polarization assay.
Conventional Th17 polarization methods typically involve obtaining na&#239;ve CD4 T cells from genetically modified mice to study the effects of specific genes on Th17 differentiation and function. These methods can be time-consuming and costly and may be influenced by cell-extrinsic factors from the knockout animals. Thus, a protocol using retroviral transduction of guide RNA to introduce gene knockout in CRISPR/Cas9 knockin primary mouse T cells serves as a very useful alternative approach. This paper presents a protocol to differentiate na&#239;ve primary T cells into Th17 cells following retroviral-mediated gene targeting, as well as the subsequent flow cytometry analysis methods for assaying infection and differentiation efficiency.

We present a protocol for retroviral transduction of guide RNA into primary T cells from Cas9 transgenic mice, providing an efficient alternative for gene editing in studying Th17 differentiation.

T helper cells that produce IL-17A, known as Th17 cells, play a critical role in immune defense and are implicated in autoimmune disorders. CD4 T cells ...

Engineering and Expansion of Human CAR Tregs for Immune Regulation Therapies

Generation of Human Chimeric Antigen Receptor Regulatory T Cells

Chimeric antigen receptor (CAR) T-cell therapy has reshaped the face of cancer treatment, leading to record remission rates in previously incurable hematological cancers. These successes have spurred interest in adapting the CAR platform to a small yet pivotal subset of CD4+ T cells primarily responsible for regulating and inhibiting the immune response, regulatory T cells (Tregs). The ability to redirect Tregs&#39; immunosuppressive activity to any extracellular target has enormous implications for creating cell therapies for autoimmune disease, organ transplant rejection, and graft-versus-host disease. Here, we describe in detail methodologies for bona fide Treg isolation from human peripheral blood, genetic modification of human Tregs utilizing either lentivirus or CRISPR/Cas9-aided knock-in using adeno-associated virus-mediated homologous directed repair (HDR) template delivery, and ex vivo expansion of stable human CAR Tregs. Lastly, we describe the assessment of human CAR Treg phenotypic stability and in vitro suppressive function, which provides insights into how the human CAR Tregs will behave in preclinical and clinical applications.

This protocol provides a streamlined workflow to generate and test human chimeric antigen receptor regulatory T cells (CAR Tregs).

Chimeric antigen receptor (CAR) T-cell therapy has reshaped the face of cancer treatment, leading to record remission rates in previously incurable ...

Non-Viral Engineering of Primary Human T Cells via Homology-Mediated End-Joining Targeted Integration of Large DNA Templates

Many current adoptive cellular therapies rely on lenti- or retroviral vectors to engineer T cells for the expression of a chimeric antigen receptor (CAR) or exogenous T cell receptor (TCR) to target a specific tumor-associated antigen. Reliance on viral vectors for the production of therapeutic T cells significantly increases the timeline, cost, and complexity of manufacturing while limiting the translation of new therapies, particularly in the academic setting. A process is presented for efficient non-viral engineering of T cells using CRISPR/Cas9 and homology-mediated end joining to achieve targeted integration of large, multicistronic DNA cargo. This approach has achieved integration frequencies comparable to those of viral vectors while yielding highly functional T cells capable of potent anti-tumor efficacy both in vitro and in vivo. Notably, this method is rapidly adaptable to current good manufacturing practices (cGMP) and clinical scale-up, providing a near-term option for the manufacturing of therapeutic T cells for use in clinical trials.

A detailed protocol is provided for using CRISPR/Cas9 technology to achieve highly efficient targeted knock-in of large, multicistronic constructs in primary human T cells via the homology-mediated end joining (HMEJ) DNA repair pathway. T cells engineered with this cGMP-adaptable protocol maintain excellent cell expansion, cytotoxicity, and cytokine production.

Many current adoptive cellular therapies rely on lenti- or retroviral vectors to engineer T cells for the expression of a chimeric antigen receptor (CAR) ...

Streamlined Single Cell TCR Isolation and Generation of Retroviral Vectors for In Vitro and In Vivo Expression of Human TCRs

Summary

Explore More Videos

Retroviral Transduction of T-cell Receptors in Mouse T-cells

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Retroviral Transduction of Helper T Cells as a Genetic Approach to Study Mechanisms Controlling their Differentiation and Function

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Retroviral CRISPR/Cas9-Mediated Gene Targeting for the Study of Th17 Differentiation in Vitro

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