Fresh Versus Frozen – Sample Considerations for Clinical Flow Cytometry (a Flowmetric blog)

Optimal sample matrix selection and processing are the foundation of good experimental design in flow cytometry. Starting with high-quality samples will not only help to ensure the scientific quality and rigor of your data, but also ensure reproducibility of the data across time, different laboratories, and research teams. However, the selection of the sample matrix is highly nuanced and should be experimentally determined before the start of any clinical flow cytometry program. This is because different cell types, surface biomarkers, and activation markers are all impacted differently by sample collection, processing, and preservation techniques; and so, understanding what your critical endpoints are and how these may be affected post sample collection is central to the quality of the analysis. In this blog, we explore the considerations for fresh versus frozen samples for clinical flow analysis and the subtleties that should be considered when selecting the sample matrix for a flow cytometry assay.


Fresh Samples


Fresh samples are nearly always preferred for clinical trials; however, their use requires careful coordination in sample collection, shipping, and receipt at the testing laboratory to ensure the samples can be processed within predetermined stability windows. This is not easily achieved and as a result, critical samples may be lost or have to be discarded from the study, so these risk factors must be taken into consideration when establishing clinical trial protocols.


One flow cytometry assay format that is almost always performed using fresh samples is Receptor Occupancy Assays (ROA). There are many challenges in the design and execution of ROAs, and these are compounded when the target-antigen is expressed at low levels, or on rare cell populations and when the therapeutic molecules are bi-specific. The selection of anticoagulant, stabilizing agent, and the storage/shipping conditions (e.g. cold packs versus ambient temperature) are all critical parameters and should be evaluated independently to determine their influences on bound drug and receptor levels on the target cell population (Liang, M. et. al. 2016).


If fresh samples are required, it is important to take time to identify the optimal anticoagulant for the clinical assay. All anticoagulants have some impact on sample quality, biomarker stability, and cell profiles over time (Karai et. al. 2018), and so all these factors need to be assessed with the use of different anticoagulants. This is particularly important when examining rare cell populations or highly variable markers that pose some challenges to preserve. It is therefore wise to select the anticoagulant based on cell population of interest, since the lability and auto fluorescence of different cell types can be influenced by the anticoagulant used.


Since EDTA is a solid, it is the anticoagulant of choice for Complete Blood Count (CBC) and White Blood Cell Differential by flow cytometry. But there are numerous examples of suboptimal CD marker staining in samples collected into EDTA, the most characterized of which is CD11b staining (Repo et. al. 1995). Since EDTA binds calcium, it may impact conformational epitopes of a number of markers including CD41, CD44 and CD49d, and is therefore typically not the anticoagulant of choice under these circumstances. Furthermore, EDTA impacts cytokine-induced killer (CIK) cell proliferation, diminishes NK cytotoxicity and can increase proinflammatory cytokine expression, so should be used with caution for functional assay systems.


Table 1. Recommended Anticoagulants and Storage Times for Commonly Performed Assays


Sample Preservation


There are many obvious advantages for preserving clinical samples, particularly when multiple clinical sites are involved in sample collection, and studies are run over significant periods of time. The simpler the preservation methods employed, the more likely that compliance and consistency will be achieved. Several commercial sample preservatives contain fixatives and/or permeabilization agents, and these may facilitate the downstream staining of extra – and intracellular epitopes for flow cytometry analysis. However, there are limitations to their use, and the detection of each critical biomarker should be evaluated with the use of these reagents. One of the most significant advantages to sample preservation is the ability to batch process samples – this enhances consistency and quality of the analysis and can help in the management of programs costs and timelines.


Fixation of Samples


Depending on the type of assay and clinical endpoints, it might be possible to fix the cells prior to staining and acquisition. One basic protocol involves fixing the cells in 4% formaldehyde for 30 minutes, followed by washing and storage at 2-8°C in an appropriate buffer. In some instances, it maybe be preferable to stain the cells first and then fix and store in the dark before acquiring on the flow cytometer. In both scenarios, long term exposure to formaldehyde should be avoided and test to ensure that antibody conjugates can still bind their targets in fixed samples.


There are a number of commercially available sample preservation solutions on the market that have been successfully adapted for flow cytometry analysis.


Cyto-Chex™ (Streck Laboratories) vacuum tubes preserve fresh whole blood for up to 7 days for routine WBC enumeration, using a cross-linking, formalin-free method with K3-EDTA anticoagulant. For some clinical applications, this method has been demonstrated to preserve sample integrity for up to 30 days in samples from patient cohorts with hematological disorders including leukemia, lymphomas and HIV.


The SmartTube system. Blood drawn into vacutainers containing sodium heparin can be preserved over an extended period using the SmartTube system. This utilizes a base station in which the blood is stimulated and stabilized with partial fixation before cooling to 4°C for transportation. Samples are reportedly stable for up to 11 days.


TransFix Reagent is simply to use requiring the only addition of the TransFix reagent into the blood collection tube followed by gentle mixing and stored between 2-25°C for as long as 10 days. For optimal preservation, blood samples should be treated with TransFix (1:5 ratio of reagent to blood) as soon as possible after collection, but no more than 6 hours. There are some nuances to its use, including that the blood sample should not be kept on ice or in the refrigerator before treatment with TransFix.


It is important to note that any fixation process can affect the relative distribution of the cell populations within a sample. Typically, granulocytes and neutrophils are the most labile, resulting in the over-representation of lymphocytes and monocytes.


Preparation of PBMCs and Freezing Down


In some instances, the clinical sites are equipped to prepare Peripheral Blood Mononuclear Cells (PBMCs) from the fresh blood samples, and these can be frozen down and shipped to clinical testing labs for analysis. The interval between blood collection and processing is a critical parameter for many functional immunological assays. Granulocytes can become activated, and this results in oxidative stress in lymphocytes that can impact their functionality. In addition, increased storage time before processing can result in the contamination of the PBMC fraction with granulocytes which can impact downstream analysis. Several commercial products have been developed to help mitigate the contamination of the PBMC fraction with granulocytes including T Cell Xtend™ (Oxford Immunotec), a reagent that cross-links granulocytes with RBCs, and therefore pellets the granulocytes during density centrifugation. Alternatively, EasySep Human Whole Blood CD66b Positive Selection kit and EasySep Magnet (StemCell Technologies Inc) can be employed to deplete the sample of granulocytes prior to processing.


Cryopreservation is the optimal method for long term PBMCs storage. The cryopreservation mechanism requires freezing the cells without instigating cryogenic damage. This is achieved through slow, controlled rates of cooling that create intracellular ice crystals but discourage the formation of extracellular ice crystals. In addition, the use of cryoprotectants such as DMSO or Glycerol that help prevent water-deprivation related cellular damage, coupled with very low storage temperatures (-80°C and LN2 vapor-phase) work to maintain cellular viability throughout storage and thawing. However, since DMSO is toxic to cells, it is typically used at 10-15% within freezing media and rapidly removed post-thaw to prevent loss in cell viability.


Final Thoughts


For every flow cytometry assay, labs should conduct their own assessment of optimal sample collection and storage and maintain these across any given study.


There are key points that should be applied across all studies. These include minimizing the delay between sample collection and processing, and assessing the impact of anticoagulant, and sample storage and processing for each cell type and every biomarker under evaluation.


Whereas fresh samples are often preferred, there are challenges to their use; and with clinical studies, there are advantages to sample preservation and batch processing to support consistency across the study. In these instances, the generation of frozen PBMCs can provide a suitable compromise to fresh samples.


Determining sample stability should be included in every clinical validation plan. Since it is unlikely that every reportable will pass the predetermined stability test script acceptance criteria, it may be necessary to identify the most critical clinical endpoints and assess these independently.


Working with a CRO who offers expertise in flow cytometry assay development will ensure that the optimal sample matrices are identified for any given study. Our team can help establish validation test scripts to fully understand the limitations of the experimental design and deliver the highest quality of scientific rigor for your translational and clinical programs.


Other commonly used anticoagulants include sodium heparin and acid citrate dextrose (ACD). These support different sample stability windows – Sodium Heparin and ACD treated whole blood samples are reported stable for up to 72 hours, whereas EDTA sample stability has been demonstrated up to 48 hours (Davis et. al. 2013). It is important to note that since ACD is a liquid, it is not suitable for use in quantitative flow cytometry applications. Another commonly found anticoagulant is heparin. Heparin functions though antithrombin III to prevent clot formation, but it can result in white blood cell clumping, and so precautions should be made to remove clumps before processing and acquisition. (Diks, A. M., et. al 2019).




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