Background

Single-cell techniques have revolutionized biological and clinical research by quantitatively capturing the genomic and transcriptomic state of individual cells at unprecedented resolution and scope [1]. This approach moves beyond averaging the bulk population, which consists of various cell types. However, the technical and logistical challenges of acquiring or generating high-viability single-cell suspensions have limited the application of single-cell genomics and transcriptomics.

Multi-center clinical studies often involve sample collection from remote areas lacking processing infrastructure [2,3]; additionally, longitudinal and cohort recruitment of patients occurs over the years, subject to patient availability and logistics constraints. Moreover, even in advanced clinical centers, the urgency of tissue harvesting during surgical procedures, biopsies, or autopsies can depend on factors such as the type of surgery, the purpose, and the specific protocol followed, as well as the transfer procedures between surgery and histopathology [4,5]. In basic research using organoids, embryos, or rare specimens, the limited cell number poses a significant bottleneck for conducting scientifically meaningful and economically feasible single-cell analysis.

In many experimental setups typical for clinical research, the freeze-thaw process remains the predominant option for sample collection en masse. However, freezing will physically damage the cell membrane, impact the post-thaw viability, and disrupt intracellular compartments where endogenous RNases are sequestered, leading to RNA degradation. In addition, cryopreservation may lead to the loss of certain cell types and induce cellular stress response [6]. Moreover, variably staggered freezing and thawing through aliquoting cells for parallel orthogonal assays, for example, single-cell mass cytometry (CyTOF), single-cell RNA-Seq, single-cell ATAC-Seq, and additional multiomic assays, can lead to inconsistency, introducing technical artifacts and batch-wise noise to the data. Recently, the fixed RNA (flex) assay has been gaining popularity due to its robustness; however, so far probes are only available for humans and mice and do not target immunologically and clinically paramount transcripts such as joining and variable regions in B and T-cell receptor (BCR and TCR), Killer cell immunoglobulin-like receptors (KIRs), and human leukocyte antigen (HLA), nor polyadenylated non-coding RNA.

Notably, many important immune cells—such as neutrophils, dendritic cells, monocytes, macrophages, and lymphocytes (B cell and T cells), as well as cells with higher intrinsic mechanical integrity such as epithelial cells—are considered fragile for single-cell applications due to their susceptibility to mechanical damage during sample preparation, including cell sorting or tissue dissociation [7–9]. Loss of these cells often leads to underrepresentation in downstream analysis. Therefore, identifying a method allowing the stabilization of fragile cells without perturbing RNA integrity will enhance the recovery of these important populations.

The ideal fixative must possess several characteristics: first, it should be small and able to penetrate the cell membrane and tissues. Second, it should preserve the structure and integrity of cells. Third, it should not destroy RNA integrity and should, ideally, inhibit RNase activity at least temporarily. Currently, one of the most common options for single-cell assays is using either standard or modified methanol fixation [10–14]. Methanol works by dehydrating samples and denaturing proteins in a gentle manner, similar to histology fixation. However, methanol as a standalone fixative has raised concerns regarding biased results or ambient RNA leakage in single-cell RNA-Seq [6]. A possible mechanistic explanation is the irreversible intracellular compartment disruption resulting in the loss of normal lipid and protein structure during dehydration–rehydration cycles. In addition, incomplete reverse transcription of mRNAs with more complex secondary structures was suggested as a major caveat [15].

In contrast, dithio-bis-succinimidyl propionate (DSP), known as Lomant's reagent [16], has been traditionally used for histological tissue fixation to stabilize cellular integrity and structures through covalent crosslinking of free amine groups found at the N-terminus of polypeptide chains (diagrammatic molecular basis is depicted in Figure 1 of Akaki et al. [17]). It has been repurposed for single-cell RNA sequencing by maintaining RNA integrity and yield in bulk RNA extractions [18–20]. The cell and membrane permeability have been proven to be particularly useful in detecting rare cell populations such as tissue-resident immune cells. Certain T-cell populations can exist in very low numbers upon activation, making them difficult to isolate and analyze with conventional single-cell methods. A protocol called CLInt-Seq (crosslinker regulated intracellular phenotype sequencing) was developed to overcome the hurdle by crosslinking intracellular cytokines [21]. This technique combines and improves upon existing techniques to collect and genetically sequence rare T cells. The reversibility of DSP-induced crosslinking is a key feature allowing cells and tissue to be further processed or dissociated for downstream applications. However, a recent study concluded that de-crosslinking the DSP-fixed samples is optional, and additional handling steps of de-crosslinking may contribute counterproductively to cell loss [22].

Recently, we developed the further optimized FixNCut protocol and demonstrated its robustness and benefits systematically by comparing fresh and fixed lung, colon, and pancreas samples from different species even under cryopreservation for an extended period [23,24]. Here, we provide an end-to-end solution in the form of a practical guide on the implementation of this method, to remove practical barriers by streamlining the transport of samples and scheduling of shared instruments for downstream single-cell isolation and processing.