What is Dr. Zubair Khalid's research focus?

Dr. Zubair Khalid specializes in molecular virology, mRNA vaccine development, and computational biology, with a focus on avian pathogens like IBDV and Avian Reovirus.

Where is Dr. Zubair Khalid currently working?

Dr. Zubair Khalid is a Postdoctoral Research Associate at the University of Maryland (UMD), specifically within the Department of Animal and Avian Sciences.

CRISPR in T Cells: Methods, Applications, and Pitfalls

By Dr. Zubair Khalid, DVM, MS, PhD · 2026-08-11

Introduction to CRISPR in T Cells

What is CRISPR-Cas9?

CRISPR-Cas9 is an adaptive immune system found naturally in bacteria and archaea that has been repurposed as a programmable genome-editing tool. The acronym CRISPR stands for Clustered Regularly Interspaced Short Palindromic Repeats, which are genomic loci containing short repeated sequences interspersed with "spacer" sequences derived from previous viral invaders. The Cas9 protein is an RNA-guided endonuclease—an enzyme that introduces a double-strand break (DSB) at a specific DNA sequence determined by a short RNA molecule.

The system works because Cas9 can be programmed with a single-guide RNA (sgRNA) that contains a 20-nucleotide sequence complementary to the target genomic locus. When the sgRNA pairs with the target DNA, Cas9 undergoes a conformational change and cleaves both strands of the DNA, creating a DSB. This break is then repaired by the cell's endogenous DNA repair machinery, which can be exploited to introduce specific genetic modifications.

For a deeper dive into the core components and variants of this system, see CRISPR Cas 9.

Why edit T cells?

T cells are central effectors of the adaptive immune system. They recognize antigens presented on major histocompatibility complex (MHC) molecules, become activated, proliferate, and execute cytotoxic or helper functions. Their role in immune surveillance makes them attractive targets for both basic research and therapeutic intervention.

Several features make T cells particularly amenable to CRISPR editing. First, they can be isolated from peripheral blood, expanded to large numbers ex vivo, and returned to a patient—a process called adoptive cell transfer. Second, many T cell functions are controlled by single genes whose knockout can produce dramatic phenotypic changes. Third, T cells are relatively resistant to some forms of genetic manipulation, which has driven the development of specialized delivery methods.

The therapeutic potential is substantial. CRISPR-edited T cells are already in clinical trials for cancer immunotherapy, where they are engineered to express chimeric antigen receptors (CARs) or to lack immune checkpoint molecules such as PD-1. Beyond oncology, edited T cells are being explored for autoimmune disease, HIV infection (by disrupting the CCR5 co-receptor), and primary immunodeficiency disorders. The broader landscape of therapeutic applications is covered in CRISPR in Medicine.

Mechanism of CRISPR-Cas9 Gene Editing

Cas9 and guide RNA

The most commonly used Cas9 variant is Streptococcus pyogenes Cas9 (SpCas9), a 1,368-amino-acid protein with two nuclease domains: the HNH domain, which cleaves the strand complementary to the sgRNA, and the RuvC domain, which cleaves the non-complementary strand. SpCas9 recognizes a protospacer adjacent motif (PAM) of 5′-NGG-3′ immediately downstream of the target sequence. The PAM is essential for target recognition; without it, Cas9 will not bind or cleave.

The sgRNA is a chimeric RNA molecule that combines two naturally occurring RNAs from the bacterial system: the CRISPR RNA (crRNA), which contains the 20-nucleotide spacer that directs target recognition, and the trans-activating crRNA (tracrRNA), which is required for Cas9 loading and activation. In practice, the sgRNA is expressed from a plasmid or delivered as in vitro-transcribed RNA, or it can be complexed with Cas9 protein to form a ribonucleoprotein (RNP) complex.

The specificity of Cas9 is determined primarily by the 20-nucleotide guide sequence, but mismatches are tolerated to varying degrees depending on their position. Mismatches in the "seed region" (the 8–12 nucleotides proximal to the PAM) are generally poorly tolerated, whereas mismatches in the distal region are more permissive. This has important implications for off-target editing, discussed later.

Double-strand break repair: NHEJ vs HDR

Once Cas9 introduces a DSB, the cell repairs it through one of two major pathways:

Non-homologous end joining (NHEJ) is the dominant repair pathway in most cell types, including T cells. NHEJ directly ligates the broken ends together, often with the insertion or deletion of a few nucleotides (indels) at the break site. These indels frequently cause frameshift mutations that introduce premature stop codons, effectively knocking out the target gene. NHEJ is active throughout the cell cycle and is fast, but it is error-prone by design.

Homology-directed repair (HDR) uses a homologous DNA template to repair the break with high fidelity. If an exogenous donor template—either a single-stranded oligodeoxynucleotide (ssODN) or a plasmid with homology arms—is provided, HDR can be used to introduce precise insertions, point mutations, or reporter genes. However, HDR is restricted to the S and G2 phases of the cell cycle and is much less efficient than NHEJ in T cells, often by an order of magnitude or more.

The choice between NHEJ and HDR is a central consideration in experimental design. For gene knockout, NHEJ is sufficient and preferred. For knock-in applications—such as inserting a CAR construct into the TRAC locus—HDR must be stimulated, which typically requires optimization of delivery timing, donor template design, and cell cycle synchronization. The distinction between these two approaches is elaborated in CRISPR Knockout and CRISPR Knock.

Delivery Methods for CRISPR in T Cells

Electroporation of ribonucleoprotein complexes

Electroporation is the most widely used method for delivering CRISPR components into primary T cells. The approach involves applying a brief electrical pulse to cells suspended in a conductive buffer, which transiently permeabilizes the plasma membrane and allows macromolecules to enter the cytoplasm.

The preferred format is delivery of pre-formed Cas9-sgRNA ribonucleoprotein (RNP) complexes. RNPs offer several advantages over plasmid-based delivery: they are active immediately upon entry, they are degraded within 24–72 hours (reducing off-target editing), and they avoid the risk of genomic integration of plasmid DNA. A typical protocol uses 1–5 µM Cas9 protein complexed with 1.2–6 µM sgRNA, electroporated in a commercial buffer such as P3 Primary Cell 4D-Nucleofector solution (Lonza) or Ingenio solution (Mirus).

Electroporation parameters must be optimized for each T cell source. For human peripheral blood T cells, the Lonza 4D-Nucleofector with program EO-115 is commonly used, but optimization is recommended because different donors and activation states respond differently. Cell viability after electroporation typically drops to 50–80%, and editing efficiency ranges from 40–90% depending on the target locus and guide RNA quality.

Viral vector delivery

Viral vectors are an alternative delivery method, particularly when stable expression of the CRISPR machinery is desired or when the target cells are difficult to transfect.

Lentiviral vectors are derived from HIV-1 and can transduce both dividing and non-dividing cells. They integrate into the host genome, providing stable expression of Cas9 and sgRNA. This is useful for generating stable knockout cell lines or for in vivo delivery. However, stable Cas9 expression increases the cumulative risk of off-target editing and can trigger an immune response against Cas9 in therapeutic applications. For T cells, lentiviral transduction typically requires activation with anti-CD3/anti-CD28 antibodies to make cells permissive to infection.

Adeno-associated virus (AAV) vectors are non-integrating and have a small packaging capacity (~4.7 kb), which is sufficient for an sgRNA expression cassette or an HDR donor template but not for the full SpCas9 coding sequence. AAV is therefore often used in combination with RNP electroporation: the RNP provides the nuclease, and AAV delivers the HDR template. This "two-step" approach has been used successfully for knock-in editing in T cells.

Viral delivery methods are discussed in more detail in CRISPR Delivered to Cells.

Non-viral delivery

Beyond electroporation and viral vectors, several non-viral approaches are being developed. Lipid nanoparticles (LNPs) encapsulating Cas9 mRNA and sgRNA have been used to edit T cells in vivo in preclinical models. Cationic lipid formulations such as those used for mRNA vaccines can deliver Cas9 mRNA, which is translated into protein inside the cell, along with chemically modified sgRNA.

Nanoparticle-based delivery offers the advantage of being non-integrating, non-immunogenic (relative to viral vectors), and scalable for clinical manufacturing. However, efficiency in primary T cells remains lower than electroporation, and the technology is still maturing.

The table below summarizes the key characteristics of the major delivery methods:

Method	Cargo	Integration	Editing efficiency	Viability	Best use case
RNP electroporation	Cas9 protein + sgRNA	No	40–90%	50–80%	Knockout; ex vivo editing
Lentivirus	Cas9 + sgRNA DNA	Yes	30–70%	70–90%	Stable knockout lines
AAV	sgRNA or HDR donor	No	10–40% (with RNP)	60–80%	Knock-in with HDR
Lipid nanoparticles	Cas9 mRNA + sgRNA	No	10–50%	60–80%	In vivo editing

Applications of CRISPR in T Cell Research and Therapy

Engineering CAR-T cells

Chimeric antigen receptor (CAR)-T cell therapy is the most clinically advanced application of CRISPR in T cells. A CAR is a synthetic receptor consisting of an extracellular antigen-binding domain (typically a single-chain variable fragment, scFv), a hinge region, a transmembrane domain, and intracellular signaling domains (usually CD3ζ plus a costimulatory domain such as 4-1BB or CD28).

The standard manufacturing process uses viral vectors to integrate the CAR transgene randomly into the genome. CRISPR offers two improvements. First, CRISPR can be used to knock the CAR construct into a specific locus, most commonly the T cell receptor alpha constant (TRAC) locus. This achieves two goals simultaneously: it places the CAR under the control of the endogenous TCR promoter, ensuring physiological expression levels, and it disrupts the endogenous TCR, reducing the risk of graft-versus-host disease in allogeneic settings. Second, CRISPR can be used to eliminate additional genes that impair CAR-T function, such as PDCD1 (encoding PD-1) or CIITA (to reduce alloreactivity).

The process of generating a CRISPR-edited CAR-T cell involves the following steps:

Isolate T cells from peripheral blood by density gradient centrifugation or apheresis.
Activate T cells with anti-CD3/anti-CD28 antibody-coated beads for 24–48 hours.
Electroporate Cas9 RNP complexes targeting the TRAC locus along with an HDR donor template encoding the CAR.
Expand edited cells for 7–14 days in media containing IL-2 (typically 100–300 IU/mL) or other cytokines.
Enrich for edited cells by magnetic selection or flow cytometry sorting.
Cryopreserve the final product for later infusion, following standard Cryopreservation of Animal Cells protocols.

Knockout of PD-1 and other checkpoints

Immune checkpoint molecules negatively regulate T cell activation and are frequently upregulated in the tumor microenvironment, contributing to T cell exhaustion. PD-1 (encoded by PDCD1) is the most studied checkpoint; its ligands PD-L1 and PD-L2 are often overexpressed on tumor cells. Disrupting PDCD1 in tumor-infiltrating lymphocytes or in adoptively transferred T cells can restore effector function.

CRISPR-mediated knockout of PDCD1 is straightforward: a single sgRNA targeting exon 1 or exon 2 of the gene, delivered as an RNP, typically yields 60–90% knockout efficiency in activated T cells. Other checkpoints targeted by CRISPR include CTLA4, LAG3, TIM3 (encoded by HAVCR2), and TIGIT. In some experimental designs, multiple checkpoints are knocked out simultaneously by delivering several sgRNAs in a single electroporation.

Functional genomics in T cells

CRISPR has revolutionized functional genomics by enabling systematic loss-of-function screens in primary T cells. Pooled CRISPR screens involve transducing a population of T cells with a lentiviral library of sgRNAs targeting thousands of genes, then applying a selective pressure (e.g., a pathogen, a cytokine, or a tumor co-culture) and identifying sgRNAs that are enriched or depleted by next-generation sequencing.

These screens have identified genes that regulate T cell activation, differentiation, exhaustion, and cytotoxicity. For example, genome-wide screens have revealed novel regulators of the transcription factor FOXP3 in regulatory T cells and identified genes whose knockout enhances CAR-T cell antitumor activity. The ability to perform such screens in primary human T cells, rather than immortalized cell lines, has been a major advance in immunology.

Optimizing CRISPR Efficiency in T Cells

Guide RNA design considerations

Guide RNA design is the single most important determinant of editing efficiency. Several rules apply:

Target sequence: The 20-nucleotide spacer should be followed by an NGG PAM. Guides targeting the first coding exon or a conserved functional domain are more likely to produce a null phenotype.
GC content: Guides with 40–70% GC content generally perform better, as they form more stable RNA-DNA hybrids.
Position within the gene: For knockout, targeting the 5′ end of the coding sequence increases the likelihood of a frameshift mutation that truncates the protein.
Off-target prediction: Use algorithms such as CRISPOR, CHOPCHOP, or Benchling to score guides for predicted off-target sites. Guides with high specificity scores should be prioritized.
Validation: Always test at least two or three guides per gene, as individual guides can vary dramatically in efficiency.

For knock-in experiments, the guide should cut within 10–20 nucleotides of the intended insertion site. The HDR donor template should have homology arms of 40–80 nucleotides for ssODNs or 500–800 nucleotides for plasmid donors.

T cell activation and expansion

Primary T cells are quiescent when isolated from blood. They must be activated before they will divide, and cell division is required for efficient editing—particularly for HDR, which is restricted to S/G2 phase. Activation also upregulates DNA repair machinery and makes cells more permissive to electroporation.

Standard activation uses anti-CD3 and anti-CD28 antibodies, either immobilized on tissue culture plates or conjugated to magnetic beads (e.g., Dynabeads Human T-Activator CD3/CD28). The bead-to-cell ratio is typically 1:1 to 3:1. Activation for 24–72 hours before electroporation is common; longer activation times improve HDR efficiency but can lead to T cell differentiation and reduced persistence in vivo.

Culture medium is typically RPMI-1640 or AIM-V supplemented with 10% fetal bovine serum (FBS) or human serum, plus IL-2 at 100–300 IU/mL. Some protocols use IL-7 and IL-15 instead of or in addition to IL-2, which promotes a less differentiated, more stem-like phenotype associated with better antitumor activity.

Enhancing HDR for knock-in

HDR efficiency in T cells is notoriously low, often 5–20% compared to 60–90% for NHEJ-mediated knockout. Several strategies can improve HDR:

Timing: Deliver the HDR donor template at the same time as the RNP, or within a few hours. The donor must be present when the DSB occurs.
Donor format: For small insertions (<100 bp), ssODNs are more efficient than double-stranded plasmid donors. For larger insertions (e.g., a CAR construct of 1–2 kb), a double-stranded plasmid with long homology arms or an AAV donor is required.
Cell cycle synchronization: Arresting cells in S phase with agents such as nocodazole or using cell cycle-synchronized populations can increase HDR.
Chemical inhibitors of NHEJ: Small molecules such as Scr7 (an inhibitor of DNA ligase IV) or M3814 (a DNA-PK inhibitor) can shift the repair balance toward HDR, though they may increase toxicity.
Cas9 variants: High-fidelity Cas9 variants (e.g., HiFi Cas9) or Cas9 fusion proteins with HDR-enhancing factors (e.g., Cas9-CT4) are being developed.

Analyzing CRISPR-Edited T Cells

Genotypic analysis

Verification of editing at the DNA level is essential. The most common methods are:

T7 endonuclease I (T7E1) assay: This assay detects heteroduplex DNA formed when wild-type and mutant alleles are hybridized. Genomic DNA is PCR-amplified across the target site, denatured, and re-annealed. T7E1 cleaves at mismatches, producing fragments that can be resolved by agarose gel electrophoresis. The fraction of cleaved DNA approximates the editing efficiency. This method is simple and inexpensive but underestimates editing efficiency and cannot identify specific mutations.

Sanger sequencing with peak decomposition: PCR products are sequenced, and software such as ICE (Inference of CRISPR Edits) or TIDE (Tracking of Indels by Decomposition) deconvolutes the mixed sequencing traces to estimate the frequency and types of indels.

Next-generation sequencing (NGS): Amplicon sequencing of the target locus provides the most accurate quantification of editing efficiency and the spectrum of mutations. This is the gold standard for clinical applications.

Phenotypic analysis

Genotypic confirmation is necessary but not sufficient; you must also confirm that the intended phenotype has been achieved.

Flow cytometry is the primary tool for analyzing protein expression. For knockout of a surface protein such as PD-1, stain with a fluorophore-conjugated antibody and compare to unedited controls. For knock-in of a CAR, the CAR can be detected with a recombinant antigen conjugated to a fluorophore or with an anti-Fab antibody.

Functional assays depend on the gene of interest. For knockout of a checkpoint, measure T cell proliferation or cytokine production (e.g., IFN-γ by ELISA or intracellular flow cytometry) after stimulation. For CAR-T cells, co-culture with antigen-expressing target cells and measure cytotoxicity using a lactate dehydrogenase (LDH) release assay or real-time impedance-based systems.

Challenges and Limitations

Off-target editing

Off-target effects are the most significant safety concern for CRISPR-based therapies. Cas9 can tolerate mismatches between the sgRNA and genomic DNA, particularly in the PAM-distal region, leading to cleavage at unintended loci. The consequences range from benign to catastrophic—including disruption of tumor suppressor genes or oncogenic translocations.

Several strategies mitigate off-target risk:

Use high-fidelity Cas9 variants (e.g., SpCas9-HF1, eSpCas9, HiFi Cas9) that have reduced tolerance for mismatches.
Deliver Cas9 as an RNP rather than as a plasmid, so that the nuclease is present only transiently.
Use truncated sgRNAs (17–18 nucleotides) that retain on-target activity but have fewer off-target sites.
Perform whole-genome sequencing or targeted amplicon sequencing of predicted off-target sites to verify specificity.

Cell viability and expansion issues

Primary T cells are sensitive to manipulation. Electroporation alone can kill 20–50% of cells, and the addition of Cas9 RNP can increase toxicity. Cells that survive may fail to expand, particularly if the edited gene is essential for proliferation or survival.

The choice of electroporation buffer, pulse program, and cell density all affect viability. Keeping cells at 1–2 × 10⁶ cells per 20 µL reaction volume is standard. Reducing the amount of Cas9 protein to the minimum effective concentration can also improve viability. After electroporation, cells should be transferred immediately to pre-warmed complete medium and allowed to recover for at least 2–4 hours before any further manipulation.

Ethical and Safety Considerations

The use of CRISPR-edited T cells in humans raises several ethical and regulatory issues. In most jurisdictions, ex vivo gene editing of somatic cells is regulated similarly to other gene therapies, requiring approval from national regulatory agencies (e.g., the FDA in the United States, EMA in Europe). Germline editing is prohibited in most countries and is not relevant to T cell therapies, since edited T cells are not inherited.

Key safety considerations include:

Off-target effects: As discussed, these must be minimized and characterized before clinical use.
Insertional mutagenesis: If viral vectors are used, integration site analysis is required to rule out disruption of oncogenes.
Long-term persistence: Edited T cells may persist for years in patients; the long-term consequences of gene disruption are unknown.
Informed consent: Patients must be informed that gene editing is permanent and that long-term risks are not fully characterized.

For a broader discussion of clinical applications and regulatory frameworks, see CRISPR in Medicine.

Common Pitfalls and Troubleshooting

Low editing efficiency

If editing efficiency is below expectations, consider the following:

Poor guide RNA design: Verify that the guide sequence is correct and that the PAM is present. Test multiple guides.
Degraded sgRNA: sgRNA is susceptible to RNase degradation. Store at -80°C in small aliquots and avoid repeated freeze-thaw cycles.
Inactive Cas9: Confirm that the Cas9 protein is functional by testing on a control locus with a known efficient guide.
Insufficient RNP complex formation: The Cas9 and sgRNA must be complexed at a 1:1 molar ratio. Incubate at room temperature for 10–15 minutes before electroporation.
Cells not activated: Quiescent T cells edit poorly. Ensure activation is confirmed by checking CD25 or CD69 expression by flow cytometry before electroporation.

High cell death after electroporation

Cell death after electroporation is common but can be minimized:

Optimize the electroporation program: Different programs trade off efficiency against viability. Test several programs and choose the one with the best balance.
Reduce Cas9 concentration: Use the lowest concentration that achieves acceptable editing.
Use fresh cells: Cells that have been in culture for more than 7 days are more fragile.
Handle cells gently: Avoid vigorous pipetting and vortexing. Use wide-bore pipette tips.
Recovery medium: Use pre-warmed medium with extra IL-2 (300 IU/mL) for the first 24 hours after electroporation.

Inconsistent results

Inconsistency between experiments is often due to donor variability. Primary T cells from different donors have different activation kinetics, proliferation rates, and editing efficiencies. To reduce variability:

Use cells from the same donor for replicate experiments.
Standardize the activation protocol, including the bead-to-cell ratio and activation duration.
Use a single lot of Cas9 protein and sgRNA.
Include a positive control guide (e.g., targeting TRAC or AAVS1) in every experiment to normalize for technical variation.

If cells must be stored between isolation and editing, follow proper Freeze Cells for Cryopreservation protocols and label all cryovials with Cryopreservation Labels for Cells to avoid mix-ups.

Frequently Asked Questions

What is the best way to deliver CRISPR into T cells?

For most laboratory applications, electroporation of Cas9-sgRNA ribonucleoprotein complexes is the best method. It offers high editing efficiency, low off-target effects due to transient Cas9 activity, and avoids the risk of genomic integration. Viral vectors are preferred when stable expression is needed or when transducing large numbers of cells is impractical by electroporation.

How do you make a CAR-T cell using CRISPR?

To make a CAR-T cell using CRISPR, you activate T cells with anti-CD3/anti-CD28 beads, then electroporate Cas9 RNP targeting the TRAC locus along with an HDR donor template encoding the CAR. The donor template has homology arms flanking the CAR sequence, allowing it to be inserted precisely at the TRAC locus. This disrupts the endogenous TCR while placing the CAR under the control of the TCR promoter.

Why is CRISPR editing in T cells inefficient?

CRISPR editing in primary T cells is less efficient than in immortalized cell lines for several reasons: T cells are quiescent and must be activated before editing; they are sensitive to electroporation-induced toxicity; and the HDR pathway is poorly active in these cells. Additionally, primary cells have variable responses across donors.

What are off-target effects in CRISPR?

Off-target effects are unintended edits at genomic sites that share sequence homology with the target site. Cas9 can tolerate mismatches between the sgRNA and DNA, particularly in the PAM-distal region, leading to cleavage at these off-target loci. These edits can disrupt genes and cause unintended phenotypes or safety risks.

How do you verify CRISPR editing in T cells?

Editing is verified at the DNA level using T7E1 assay, Sanger sequencing with peak decomposition (ICE/TIDE), or next-generation sequencing. At the protein level, flow cytometry is used to confirm loss or gain of surface markers. Functional assays, such as cytokine production or cytotoxicity, confirm that the edit has the expected biological effect.

Can CRISPR be used to treat T cell cancers?

Yes. CRISPR-edited T cells are being developed for the treatment of T cell malignancies. One approach is to knock out the TRAC locus to eliminate the endogenous TCR, preventing fratricide (killing of edited T cells by each other) and graft-versus-host disease. Another approach is to knock out CD7, a surface marker expressed on malignant T cells, to prevent CAR-T cells from targeting each other.

What is the difference between NHEJ and HDR?

NHEJ (non-homologous end joining) is an error-prone repair pathway that directly ligates broken DNA ends, often introducing small insertions or deletions. It is active throughout the cell cycle and is used for gene knockout. HDR (homology-directed repair) uses a homologous template to repair the break precisely and is active only in S/G2 phase. It is used for knock-in applications where precise insertion or replacement is required.

Key Takeaways

CRISPR-Cas9 introduces double-strand breaks at specific genomic loci, which are repaired by NHEJ (for knockout) or HDR (for knock-in).
Electroporation of Cas9-sgRNA ribonucleoprotein complexes is the preferred delivery method for primary T cells due to high efficiency and low off-target risk.
T cells must be activated before editing; activation status, guide RNA design, and culture conditions are the main determinants of editing efficiency.
CRISPR is used to engineer CAR-T cells, knock out immune checkpoints like PD-1, and perform functional genomic screens.
Off-target editing and cell toxicity are the major challenges; they are mitigated by using high-fidelity Cas9 variants, transient delivery, and optimized electroporation conditions.
Verification of editing requires both genotypic analysis (T7E1, sequencing) and phenotypic analysis (flow cytometry, functional assays).
Clinical applications of CRISPR-edited T cells are advancing rapidly but require rigorous safety assessment and regulatory oversight.