Tips For Optimal Flow Cytometry

Posted by L.Flintham on 12th Nov 2020

Tips For Optimal Flow Cytometry

Tips For Optimal Flow Cytometry

Flow cytometry is a widely utilised laser-based technique which lends itself to the analysis of various physical and chemical characteristics in a particular cell population. The method has grown to be a popular technique within the realm of antibody research through its proven use in efficient antibody labelling and phenotypic marker identification. Furthermore, flow cytometry offers a highly time-efficient process where tens of thousands of cells can be analysed in just a short space of time. Below, we have assembled our top tips, from reducing the risk of non-specific binding to optimising the permeabilising stage, to help you get the most out of the flow cytometry technique.

Can cell preparation affect the quality of flow cytometry results?

Yes, poorly prepared samples could lead to inaccurate results. It is important to store cells on ice before experimentation and, if possible, reduce the development of aggregates which can block the flow cell. Similarly, vortexing the cells should be avoided when forming cell suspensions. Ultimately, the procedures used in cell preparation should not hinder cell viability. Inaccurate results can develop due to ineffective gating and, resultantly, a greater degree of background autofluorescence.

Should dead cells be identified before analysis?

Yes, while using healthy cells is vital to generate accurate experimental results, staining dead cells can be a highly useful method of reinforcing meaningful data. As dead cells can bind non-specifically to any antibody there are simple ways of locating them during sample preparation. The most effective way to stain the dead cells is to pass a fluorescent dye through the plasma membrane. In a further tip, use a dye that covalently binds to the dead cells as it reduces leakage and ensures that the cells remain stained after permeabilization.

How can the risk of non-specific binding be reduced?

Various aggravating factors increase the chances of non-specific binding, one of which is using a surplus of antibody. This can cause a reduction in both the separation of positive cells and a decrease in the signal: noise ratio. This factor and its influence upon the research can be reduced by titrating the antibody sample. Likewise, the extracellular matrix is a further common source of non-specific binding. This risk could be reduced through the addition of a protein, commonly Bovine Serum Antibody (BSA) is used, to the wash and staining procedures in order to complement an array of binding sites.

How can the high intensity fluorescent signal be maintained to improve the quality of results?

Maintaining a high intensity fluorescent signal is imperative to the optimisation of experimental results, but this can be problematic if extracellular antigens are internalised during antibody binding. Therefore, by following tips such as reducing the number of aggregates and, where possible, using monovalent antibodies with fragmented antigen binding, a high intensity signal can be maintained.

How can the cell permeabilization process be optimised?

It can be challenging to pass antibody-based probes through the plasma membrane and therefore cell fixation and permeabilization are fundamental to stain intracellular antigens effectively. Various methods can optimise this process. Firstly, when staining secreted proteins such as cytokines, it can be beneficial to add a protein transport inhibitor before the permeabilization stage to confine the cytokine. Also, in intracellular staining procedures, we advise staining for surface markers before commencing the permeabilization phase. This is recommended as the process can alter certain epitopes and subsequently hinder the validity of observations. Finally, we recommend completing sample stimulation before surface staining. This is endorsed as antibody-antigen surface binding can incite cells and alter the expression of intracellular signalling proteins.

How can the choice of permeabilising agent affect the results?

Using the correct permeabilising agent for the research experiment is central to correct data interpretation. We suggest avoiding agents such as Tween if possible, as Tween can lyse cells during prolonged incubation. Comparatively, Saponin does not change the antigen epitopes and can, therefore, be used in later staining. Agents such as Methanol are compatible with a majority of antigens, but not all fluorochromes can tolerate Methanol, highlighting the need to thoroughly research the appropriate agent for your experiment before commencing the research.

Are there ways to reduce the risk of loss of epitope?

Yes, there are a few different tips which can be used to help prevent a loss of epitope. Firstly, fixing the sample for too long can increase the risk; therefore, stringent time management during the fixing stage is recommended. In most cases, the sample should not be fixed for more than 15 minutes. Similarly, keeping the sample on ice can help to reduce the chances of epitope loss by preserving activity at a low temperature. Finally, using too much paraformaldehyde, which releases methanol in its breakdown, is also associated with a loss of epitope. We advise using only 1% paraformaldehyde.

Is there a way to determine the amount of background signal?

The extent of background noise can be ascertained by always using an appropriate experimental control. It is advised that a negative control, which is of the same isotype as the labelled antibody, is included as a comparison against which the background signal can be based. Consequently, we recommend including a sample of unstained cells which have been incubated and analysed in tandem with the experimental sample.

Is there an optimal time period between stimulation and experimentation?

In order to limit the variability and staining intensity, stimulate the cell populations at the same time. It is recommended that a buffer is used to freeze the sample and all populations are thawed as one, approximately two hours before analysis. Likewise, running all of the samples in one batch can limit degradation and variability.

Ultimately, whether it is used as a single process or in combination with other analytical techniques, flow cytometry represents a swift and efficient means of antibody analysis. Through appropriate sample preparation, agent choice and time management, researchers can utilise this favoured technique in a way that significantly reduces the risk of experimental error. In doing so, both time and finances can be saved in the face of reliable and dependable research.