The marriage between immunology and cytometry is one of the most stable and productive in the recent history of science. A rapid search in PubMed shows that, as of July 2017, using “flow cytometry immunology” as a search term yields more than 68 000 articles, the first of which, interestingly, is not about lymphocytes. It might be stated that, after a short engagement, the exchange of the wedding rings between immunology and cytometry officially occurred when the idea to link fluorochromes to monoclonal antibodies came about. After this, recognizing different types of cells became relatively easy and feasible not only by using a simple fluorescence microscope, but also by a complex and sometimes esoteric instrument, the flow cytometer that is able to count hundreds of cells in a single second, and can provide repetitive results in a tireless manner. Given this, the possibility to analyse immune phenotypes in a variety of clinical conditions has changed the use of the flow cytometer, which was incidentally invented in the late 1960s to measure cellular DNA by using intercalating dyes, such as ethidium bromide. The epidemics of HIV/AIDS in the 1980s then gave a dramatic impulse to the technology of counting specific cells, since it became clear that the quantification of the number of peripheral blood CD4+ T cells was crucial to follow the course of the infection, and eventually for monitoring the therapy. As a consequence, the development of flow cytometers that had to be easy-to-use in all clinical laboratories helped to widely disseminate this technology. Nowadays, it is rare to find an immunological paper or read a conference abstract in which the authors did not use flow cytometry as the main tool to dissect the immune system and identify its fine and complex functions. Of note, recent developments have created the sophisticated technology of mass cytometry, which is able to simultaneously identify dozens of molecules at the single cell level and allows us to better understand the complexity and beauty of the immune system. However, the moon has a dark side. The main strengths of this technology, i.e. the fact that it is relatively easy to use and that often only a brief training is sufficient to use a flow cytometer and start producing data, is also its main weakness. Indeed, in several (too many) papers, the eye of a well-trained cytometrist can identify aspects that would need, to be polite, a “little” improvement. Not to mention the cases in which technical mistakes are performed, involving, among others, the use of (in)adequate controls, the (lack of appropriate) compensation, sorting strategies, or even the description of the methods used. For this reason, the editorial team of the European Journal of Immunology feels it is worthwhile to offer our community guidelines for the correct use of cytometric techniques in the field of immunology. Thus, starting at the European Congress of Immunology (ECI 2015) in Vienna (Austria) and under the guidance of Professor Andreas Radbruch, we asked colleagues and friends, all renowned in this field, to contribute by sharing their knowledge in their particular areas of expertise, in order to present a collection of protocols of great interest. Such information includes, among others, suggestions and tricks regarding how to study cell phenotypes, the type or amount of molecules produced or secreted after stimulation by a cell population of interest, signalling processes, differentiation, proliferation or cell death, cytotoxic activities, cell-cell interactions, activity of intracellular organelles such as mitochondria, different types of response induced against tumours or by anticancer or immunosuppressive drugs, transcription factor activity, the quantification of soluble molecules, drug uptake, and rare events. Today's challenges also involve the choice of reagents, the preparation and eventual storage of the cells under analysis, the overall experimental plan and, last but not least, data analyses. We are no longer limited by complex instrumentation, but by our creativity to ask the critical questions. These “Guidelines for the use of flow cytometry and cell sorting in immunological studies” thus represent a community effort to collect the currently accepted best methods for monitoring most of the variation of the major players of immune system (along with their organelles and functionality) and include standards for data interpretation, as well as cautions about technical issues. One aspect of the guidelines concerns data reproducibility, a topic that has recently attracted considerable attention. Therefore, the guidelines are meant to help researchers avoid potential pitfalls that could drastically alter the interpretation of their data. While preparing the guidelines, feedback was received that we feel should be highlighted in this Introduction. Firstly, “FACS” (fluorescence activated cell sorting) should only be used for Becton Dickinson (BD) technologies as it is a BD trademark (FACSTM); the more general term “flow cytometry cell sorting” should be used to be company agnostic. Secondly, CD mAbs and not anti-CD mAbs (in other words CD1 mAb and not anti-CD1 mAb, for example) should be used. This is because the CD nomenclature is primarily a system to cluster/characterize mAbs and it was only later accepted to use this system to also describe the respective CD molecules. Thirdly, although the guidelines are as comprehensive as possible, there are naturally limitations e.g. only a subset of antibodies and antigens are shown and, at times, only certain reagents/companies are used as examples. It is our opinion that all efforts must be improved—this is how science works! Thus, we would be glad to receive from readers of the European Journal of Immunology critical comments, new ideas, and even suggestions for new articles for possible future updates of the Guidelines. Before closing, we would like to thank four people who played a major role in ensuring that Andreas Radbruch's and Andrea Cossarizza's vision became a reality. These are Hyun-Dong Chang and Ute Hoffman, both at the DRFZ, and Karen Chu, former Associate Editor, and Cate Livingstone, Managing Editor of the European Journal of Immunology. Together this core team coordinated author invitations, and the submission, peer review and revision of all the sections and proofs, as well as ensuring that community feedback was sought and incorporated. We would also like to thank the full editorial team of the European Journal of Immunology for their invaluable work on this project. To accurately measure optical properties of cells with a flow cytometer, cells have to pass through the uniformly bright center of focused laser beams. Light collection optics is focused on the intersection point of cells with the laser beams to pick up fluorescence and scattered light from cells. This is the sensing zone of a flow cytometer, here the measurements of cell parameters are taken. In a stream-in-air cell sorters, the sensing zone is located around 0.3 mm under the nozzle tip, in other cytometers it is located inside a cuvette. One purpose of the fluidic system is to move the cells one by one precisely through the sensing region in a liquid stream in such a way that each cell is illuminated by the same amount of light from the lasers. In cytometers with sort capabilities or cell sorters, the fluidic system has to establish a stable break off of the liquid stream in small uniform droplets. Droplets containing the cells of interest can be charged and deflected in an electric field for sorting. This kind of cell sorting technique was invented by Mack J. Fulwyler in 1965 at Los Alamos National Laboratory 1. Mack Fulwyler needed a machine for testing the performance of Coulter counters, so the first particle separator was used for sorting of particles with different Coulter volumes. Len Herzenberg was interested in a machine that can sort living cells on the basis of fluorescence, he got the design plans of the particle separator from Mack Fulwyler and found a little group at Stanford University to build the first FACS in the late 1960s (see the video Inventing the Cell Sorter, Herzenberg Lab, https://www.youtube.com/watch?v=Ro8P3w9BPhg). For precise positioning of cells in a liquid jet the hydrodynamic focusing technique is used in most cytometers and cell counters 2. The cells in suspension are injected by a thin tubing in a laminar flow of a sheath fluid that enters from a wide tubing into a narrow tubing or small orifice. The sheath flow speeds up when it enters the narrow tubing and the diameter of sheath and sample flow (sample core) is decreased (Fig. 1). Crosland-Taylor described this technique first in Nature 1953 3 and used it in a device for counting small particles suspended in a fluid. Some years before in 1947, F.T. Gucker used a similar technique for detecting bacteria in a laminar sheath stream of air 4. The hydrodynamic focusing takes place in the so-called flow chamber or flow cell of a cytometer. A detailed description of an optimized flow chamber for a stream-in-air cell sorter can be found in the patent applications from Gerrit van den Engh 5, 6 and a flow chamber of a cuvette system is found in another patent application from BD 7. In addition to flow chambers for laser based cytometers, flow chambers with hydrodynamic focusing for cytometers with an arc lamp light source were developed. These early cytometers are based on a standard fluorescence microscope with epi-fluorescence setup. Here the same microscope lens is used to bring excitation light to the cells and take fluorescence emission from the cells. Excitation and emission light is separated by a dichroic mirror and special filters. With an immersion microscope lens of high numerical aperture, a stabilized arc lamp and optimized staining protocol, DNA histograms with coefficient of variations (CVs) lower than 1% (0.50–0.7%) were achieved 8, 9. With the hydrodynamic focusing technique, cells can be aligned to a precision of one micrometer. With high sample flow rates the sample core is increased, however, and cells in the sample core can move out of the focus center of the laser. Thus, not all cells get the same amount of laser illumination. This means that the accuracy of measurements is lost. To avoid loss of measurement precision when the sample core increases and to maintain laser intensity, cytometers use elliptical laser focus spots. Typical sizes of focus spot are 60–150 micrometers horizontally and 5–20 micrometers vertically. Recently, beam shaping optics for flat top focused laser beams were introduced in flow cytometers by the manufacturer. The intensity profile of a gaussian laser beam with 60 100, and 150 micrometer focus diameters is shown in Fig. 2. The approximation of the sample core diameter calculation shows that for a ten times lower sample concentration a more than three times bigger sample core diameter is necessary to keep the particle measurement rate. For the sheath fluid, PBS (phosphate buffered saline) filtered through a 0.22 or 0.1 micrometer filter is often used. The sheath fluid should be compatible with cells or species that have to be sorted. An acoustic focusing technology was developed by Gregory Kaduchak and co-workers at the Los Alamos National Laboratory in 2001 and introduced to flow cytometry 11, 12. Recently, the acoustic focusing technique was implemented into a flow cytometer to support hydrodynamic focusing. This technique helps to increase measurement precision in particular if wide sample cores are used. According to the manufacturer, cytometers with acoustic-assisted hydrodynamic focusing can run samples with low concentrations of cells up to 10 times faster as compared with cytometers without and still maintain the precision of the measurements. The fundamentals of acoustic cytometry are given in 13. Based on the invention from Richard Sweet 14, droplet formation of the liquid jet of a cell sorter is stabilized by vibrations of an ultrasonic transducer. Little disturbances on the surface of the liquid jet at the exit of the nozzle orifice are generated by the transducer. The disturbances grow exponentially and lead to break up of the jet in little droplets 2, 10. A cell of interest that should be sorted is measured at the sensing zone and moves down the stream to the breakoff point. During the separation of the droplet with the cell in it from the liquid jet, a voltage pulse is given to the liquid jet. So electrons are caught with the cell in a droplet and cannot go back when the droplet is separated from the liquid stream and the voltage pulse is shut off. The droplet with the cell is charged and can be deflected in a static electric field of two deflection plates for sorting (Fig. 3). It is important for the sorting process that the cell of interest is at the right place when a voltage pulse is given to the liquid jet to charge a droplet. The delay from the measurements of cell parameters to the charging pulse is determined by the cell sorter operator or by the cell sorter electronics. This is done with the help of fluorescence beads and a laser beam under the deflection plates. The laser beam illuminates the streams of deflected and un-deflected droplets. The fluorescence beads are sorted all in one direction, and with a camera, the fluorescence in the droplet streams is observed on a monitor. During observation of the fluorescence spots the drop delay is changed so that the brightness of the fluorescence spot of the deflected droplet stream is maximized and the brightness of the fluorescence spot of the un-deflected droplet stream is minimized. The distance from the sensing zone to the break off point is controlled by a microscope and held constant. The delay setting is fixed during sorting and in general the break off distance is kept constant by the operator. If the velocity of the liquid jet is constant during sorting the sorting works fine, but in practice this is not always the case. Small changes of sheath pressure for example due to partial clogging of the sheath filter can alter jet velocity during sorting. Timothy Petersen and Gerrit van den Engh have examined the problem and showed how little variations of sheath pressure can disturb the sorting process and how the operator can handle it 16. Toralf Kaiser examined how temperature changes of sheath fluid alters sorting performance and gives a solution for stabilizing sheath fluid temperature 17. A schematic of a typical fluid system of a cell stream-in-air sorter is shown in Fig. 4. From a technical point of view a flow cytometer is a light detection device capable of detecting photons of different wavelengths over a high dynamic range. In order to achieve a high dynamic range, the optics, signal detection, and processing units must be carefully designed. In flow cytometers, lenses are used to collect light emitted from the cell of interest, i.e. due to their spatial resolution they collect light only from the point of interest. Furthermore, they are used to make the collected light parallel in order to direct it through the optical bench to the detectors. A flow cytometer employs collection and collimation lenses. Collection lenses (convex lenses) are used to focus the light from the interrogation point either to the end of an optical fiber or directly to a collimation lens (e.g. aspheric condenser lenses). Some instruments use optical fibers to route the detected light to detectors which are installed in an octagon. In this case a collimation lens is installed at the other end of the fiber to ensure that all light is routed parallel through the octagon. Inside the octagon another collimation lens is placed in front of each detector to focus the parallel light onto the photocathode. In instruments without fiber optics the parallel light is routed through the optical bench and then focused onto the photocathode by a collimation lens. The photodetectors used in flow cytometers are spectrally broadband and therefore unable to generate a signal exclusively from specific wavelengths and thus specific markers. To add specificity, optical filters and dichroic mirrors are used in a well defined manner to route the light to the detectors. Optical filters are designed as band pass (BP), long pass (LP), or short pass (SP) filters and are mostly installed in front of the light detectors. The common property of the filters is that they transmit light only within a spectral range. A BP filter transmits light in a certain range. For example, if the BP is named as 660/20, this means that light between 650 and 670 nm will pass through the filter to the photomultiplier tube and all other wavelengths will be reflected to the next filter set within the specified laser configuration. SP filters will pass short wavelengths and block longer ones whereas LP filters will do the opposite meaning that SP and LP filters transmit light below (SP) or above (LP) a certain wavelength. For example, a LP of 660 nm will transmit all light above 660 nm. Due to aging, quality of coating, and contamination, the actual parameter of an optical filter can differ from the technical description. Therefore, it is recommended to check the transmission spectra of new filters provided by the manufacturer and always keep filters dust free. Sometimes mirrors (usually silver mirrors) are used in the optical bench of a flow cytometer in order to deflect light for geometrical or constructive reasons. These filters are >99%, reflective over a wide range of wavelengths. In contrast, a dichroic mirror deflects light of a certain wavelength while the rest pass-through. The effect of the dichroic is dependent on the operating angle. In some instruments, the dichroics employed have a working angle of 45° whereas others have a working angle of 12.5°. Recently, commercial cytometers have become available which use spatially dispersing elements instead of or in combination with optical filters in order to deflect light wavelength specific to a detector array. The rationale behind this is the measurement of the entire emission spectra of a cell (see Section I.3: Flow cytometry, including flow cytometry cell sorting). A dispersing element can be a dispersive prism or a grating. Prisms have a higher light efficiency over gratings and they are not sensitive for polarized light. This maybe the reason why they are employed in the spectral flow cytometer from Sony. A dispersing element is installed between the interrogation point and a detector array. Lasers employed for flow cytometers are mainly solid-state, continuous wave lasers. Such lasers have a small footprint and a typical output power range from 20 to 100 mW. Lasers are coherent light sources which allow a high photon density at the illumination point, and therefore an efficient energy transfer to the fluorochrome. Modern cytometers are equipped with up to seven different lasers in a typical laser line ranging from 355 to 650 nm. This gives high flexibility in choosing the fluorophores. As a flow cytometer measures the biological information of a particle (e.g. a cell) via photons, this light needs to be converted to electrons and processed by an amplifier, filter, analog to digital converter (ADC), and baseline restorer in order to visualize and store the biological information of the cells or other particles. In this section, the main components of cytometer electronics are briefly described. From a technical point of view, the detection of cell related light is difficult due to (i) the low light level, (ii) the high analysis rate, and (iii) the high dynamic range of the light level. Photomultiplier tubes (PMTs) meet these requirements and are therefore employed in almost all flow cytometers. PMTs are vacuum tubes containing a photocathode, electron focusing electrodes, and a series of dynodes for electron multiplication. The photocathode converts photons to photoelectrons which are then multiplied by a series of dynodes driven by a high voltage (Fig. 5). Photocathodes of PMTs employed in flow cytometers are made from bialkali material which determines the spectral quantum efficiency η of the PMT, which is the ratio of emitted electrons to incident photons. The quantum efficiency of the photocathode is always 0< η <1 and is a function of the light quantum energy (h*f). A typical PMT (R9220, Hamamatsu) of a cytometer has a quantum efficiency η = 0.2 at 500 nm and η = 0.09 at 700 nm which is a reduction in sensitivity of about 7 dB. This means that the detection of PE-Cy7 is always less sensitive as the detection of FITC, for example. In many applications, PMTs are increasingly being replaced, e.g. by avalanche photodiodes due to their higher quantum efficiency. However, in flow cytometry, only one commercial instrument (CytoFlex, Beckman Coulter) employs APDs in order to improve the sensitivity for wavelengths >700 nm 18. All amplifiers in a cytometer are analogue hardware devices which must be very well designed for optimal signal to noise ratios (SNRs). In a typical cytometer such amplifiers have an SNR of >86 dB. Once the signals are processed by the pre-amplifiers, the main amplifier moves the signal level to a suitable range for the ADC (Fig. 5). In modern cytometers, the conversion of the continuous analog voltage signal into discrete digital values is done by ADCs which are defined by their sampling frequency and sample resolution. The required dynamic detection range (DNR) of a flow cytometer can be defined as the intensity range of stained and unstained cells, for example. A stained cell can be 10 000 times brighter than an unstained cell which gives a DNR of 4 log or 80 dB (DNR[dB]=20log(104)). The DNR of an ideal ADC is given by: DNR = 6.02*N + 1.76 dB 19. This means that in theory an ADC with N = 14 bit will have a DNR of 86.04 dB. In practice, the effective number of bits of an ADC is, due to noise and distortion of the circuit, some decibels below the theoretical value (e.g. the ADC AD9240AS of the BD Diva electronic has 78.5 dB 20). This limits the dynamic range to less than 4 decades and, more importantly, shrinks the resolution of dim signals. The sampling frequency of the AD9240AS is 10 MHz which results in 30 samples per measured pulse of a high speed cell sorter (pulse length = 3 μs). This results in a peak detection error of 1–2% 21. Modern ADCs have a resolution of 16 bit and a sampling frequency of 250 MHz which allows the design of flow cytometers with dynamic range of >4 decades and a peak detection error of <0.1%. In the digital domain the signals are processed by filters, baseline restorer, pulse height, pulse width algorithms, and trigger (see Section I.3: Flow cytometry, including flow cytometry cell sorting). Filtering is done to smoothen the raw PMT signal in order to improve the SNR. The resulting signal consists of an unwanted DC part due to laser scatter light and electronic noise (among others) and a specific AC part. Hence, the DC part is subtracted by baseline restorers to increase the SNR and the DNR of the cytometer. The baseline restorer attempts to keep the baseline at zero. In practise however, baseline restoring is not perfect and can lead to negative values on the histogram axis or introduce a slight distortion of low signals and therefore to a increased CV of dim signals. After baseline restoring, the pulse parameters (height, width, and area) are extracted and converted into a *.fcs file. Taken together, the analogue and digital components of a flow cytometer in combination with the baseline and pulse shaping algorithms need to be well adjusted in order to maximize SNR and DNR. Since the invention of the first prototype of a Fluorescence Activated Cell Sorter in 1968 at Stanford University, the technology has become a powerful tool to analyze and sort individual cells based on their functional status. Moreover, flow cytometry provides a robust statistic of thousands of individual cells and can detect rare events at a frequency below 10–4 cells. The sample uptake by the instrument can be done from tubes or multi-well plates at an acquisition rate of thousands of cells per second. In a typical cytometer, the sensitivity decreases with increasing flow rate due to the increasing diameter of the cell stream within the flow cell. Alternatively, the AttuneNXT (ThermoFisher) uses acoustic-assisted hydrodynamic focusing which helps keeping the core stream tight and therefore gives accurate results even at a much higher sample throughput. Furthermore, the serial acquisition of multiple cell samples can be automated by using high-throughput platforms (HyperCyt®). Today, instruments are available designed to detect up to 27 different bio-markers on an individual cell. Typically these markers are fluorescently tagged antibodies, molecular sensors, as well as genetically encoded reporters. For instance, the FACSymphony™ (Becton Dickinson) is technically capable of detecting up to 50 parameters of an individual cell. In practice, this high number of parameters is not achievable because at the moment the range of appropriate fluorescent dyes is limited. Technical limitations regarding the maximum number of detectable markers are also given by the overlap of the emission spectra of the different fluorescent tags, since each fluorescence detection channel is correlated to a biological marker. To overcome this, fluorescent tags became available which have different excitation wavelengths. Currently, up to seven lasers with emission wavelengths from 325 to 650 nm are used in order to achieve a high flexibility in the choice of the fluorescent tags. Furthermore, tunable lasers are used for special applications like fluorescent life time measurements (FLIMs). Flow cytometers use either photomultipliers (PMTs) or avalanche diodes to convert the emitted or scattered light into amplified electrical pulses which are processed by appropriate electronics to extract information like pulse height, area, length, and time. The electronics of the cytometer consist basically of a preamp circuit, baseline restoration circuit, and an analog to digital converter (ADC). In most modern cytometers, the data post-processing (i.e. pulse integration, compensation, log-transformation) and data analysis is done in a computer by software. All components together must have a low noise level (i.e. a high SNR) to achieve high instrument sensitivity (Q) and low background (B) detection. Avalanche diodes have better detection efficiency in long wavelengths and thus a better SNR in that range over PMTs. Furthermore, they open new possibilities for the application of fluorescent tags with long-wave emission spectra. Avalanche diodes are implemented in the CytoFLEX (Beckman Coulter) cytometer. Within this instrument, the emitted fluorescence light is divided by a wavelength division multiplexer (WDM) through a series of band pass filters and integrated optics, onto an array of avalanche diodes which enables a high sensitivity in the detection of e.g. PE-Cy7. Avalanche diodes or PMTs itself are light detectors which are unsuitable for wavelength detection, hence the fluorescent light needs to be filtered by optical filters and mirrors. These filters must be carefully chosen because a multiparameter experiment, i.e. an experiment in which multiple parameters (markers) are analyzed, requires that multiple fluorophores are used simultaneously; a consequence of this is spectral overlap or spillover (see Section III.1: Compensation). Conventional flow cytometers circumvent this problem by compensation (see Section III.1: Compensation) in order to accurately correlate the physical light properties with the biological properties of the cell. Following this, the data are analyzed in a multivariate fashion in combination with a hierarchical gating strategy (see Section VI.1: Data analysis—An overview, and Section VI.2: Data analysis—Automated analysis: Automated flow cytometry cell population identification and visualization). It is essential to adapt the combination of fluorescent tags to the given optical, laser, and electronic setup of the instrument to minimize spillover, increase Q, and lower B signals. For instance, by choosing the right concentration of a certain reagent (see Section IV.2: Titration—Determining optimal reagent concentration), the fluorochrome related B can be optimized such that it contributes ideally nothing to the B given by the instrument. This can help to increase the separation (the distance between the means) between a blank and a fluorescent population which is a function of Q and B. Thus, it requires the characterization of Q and B of the used instrument. Mostly polystyrene particles (beads) are used for this purpose in combination with software based protocols implemented in the instruments e.g. MACSQuant, Fortessa, Yeti, Cytoflex to name just a few. Beads are small particles and so to say “cell dummies” of well defined fluorescent intensity and sizes which also can be used for PMT voltage optimization, compensation setup, cell counting, scale calibration and so on. Scale calibration is an especially useful approach to measure absolute values (e.g. number of binding antibodies, amount of fluorescent molecules or photoelectrons) instead of relative mean fluorescent intensities (MFIs) which leads to quantitative flow cytometry (see Section VII: Cytometric parameters). Beside beads, scale calibration can also be achieved by using LED light pulses. Recently, the quantiFlash™ (APE) tool has become available which provides ultra stable LED light pulses. Furthermore, by using this tool, instruments can be compared within or between labs regarding their Q and B values. Up to this point, analytical cytometers have been described but cells can, in addition, be sorted based on specific marker expression for downstream analysis (molecular biology, sequ
