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4.3: Testing cell viability - Biology

4.3: Testing cell viability - Biology


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Introduction

Promoting appropriate cell life and death is a key part of tissue engineering. For example, immune cells called naïve T cells require the cytokine IL-7 and contact with self-MHC proteins for survival.

Many assays are available to monitor the numbers of live and dead cells in a culture. The kit you will use today is made by Molecular Probes, a company (now partnered with Invitrogen) that makes a plethora of fluorescent cell stains for various purposes. The principle exploited by the LIVE/DEAD® kit is the relative permeability of cell membranes when the cell is live (intact membrane) or dead (damaged membrane). Ethidium is a nucleic acid stain that you are familiar with from running agarose gels in modules 1 and 2; the ethidium homodimer-2 variant emits red fluorescence, and cannot diffuse past intact cell membranes. The dye SYTO 10, on the other hand, is membrane-permeant, and thus enters both live and dead cells; it emits fluorescence in the green channel. SYTO 10 has lower affinity for nucleic acids than does ethidium, and thus is excluded from dead cells over time, enabling one to distinguish between live (green) and dead (red) cells. Viability can be inferred by monitoring parameters other than cell permeability. For example, some membrane-permeable dyes are only activated to a fluorescent form inside cells that have active esterase enzymes, thus indicating their metabolic activity. Assays that measure cell potentials or redox activity are also available. In general, fluorescence assays are more sensitive than colorimetric assays. Along with sensitivity, stability, toxicity, and ease of scale-up are important factors to consider when choosing an assay.

Cell vitality (or lack thereof) tells only one part of a cell culture's story. For example, kits like the one we are using today cannot determine whether the cells assayed have divided or not. However, other dyes are available that specifically test for cell proliferation, or even distinguish cells based on what part of the cell cycle they are presently in. Proliferation assays are important for drug development, cancer research, and in tissue engineering. Total nucleic acid content is sometimes used as a measure of proliferation – Hoechst is a popular dye for this purpose. Active proliferation can be monitored by addition of 5-bromo-2'-deoxyuridine (BrdU) to cell cultures. BrdU will be incorporated only in recently synthesized DNA (S-phase cells), and can be assessed by antibody-detection after a time lag. For tracking multiple cell divisions, long-lived fluorescent dyes such as the fluorescein derivative CFDA-SE are used: about 6-10 divisions can be seen by flow cytometry (see figure at right).

Remember that cell death is just as important as cell life, and that the type of death also matters. Cells that die due to acute trauma or other tissue damage typically die by necrosis: they swell and finally burst, releasing their contents and often promoting inflammation. Under other circumstances, particularly in development and immunity, many cells undergo a programmed death called apoptosis. Unlike the more disruptive necrotic cells, apoptotic cells condense and then fragment, finally releasing membrane-contained cell bodies. Apoptosis gone awry is implicated in many diseases, and thus researchers are very interested in tracking apoptotic cells in various culture systems. Special dyes can be used to track nuclear fragmentation and other changes in early and late apoptotic cells.

Your objective today is to determine the viabilities of your two different cell cultures, and to gain experience with fluorescence assays. You are likely to encounter fluorescence and other microscopy techniques in many fields of biological engineering research.

Protocols

Today you can stagger your arrivals to lab. Only one group at a time will be able to work on the microscope, and assuming that cell culture setup takes ~ 1 hour, you will each have ~25 minutes to spend on the microscope. Please be respectful of your labmates' time. Reading the protocol in advance will help you work more quickly, and is strongly recommended.

Before or after performing the viability assay, and/or during incubation steps, you should work on Part 3 of today's protocol.

Part 1: Bead Preparation for LIVE/DEAD Luorescence Assay

  1. Retrieve your 2 six-well dishes from the incubator.
  2. Begin by counting your beads by eye, and decide how many (1-3 beads per sample) you can spare. Ideally, for RNA and protein isolation, you want at least 10-20 beads remaining.
    • Also take this time to describe bead uniformity in your notebook, as this feature may affect your eventual experimental outcomes. Some groups had more luck than others in keeping bead size consistent between their two samples.
    • During a later incubation step, you might also take a look at your plate under the microscope, and focus in on cells within the beads. What is cell morphology and density like in each sample?
  3. Using a sterile spatula, remove the beads (keeping the two samples separate) to two labeled Petri dishes.
  4. Within the Petri dish, cut your whole beads in half using a spatula or razor blade.
  5. Per dish, rinse the beads with 3 mL of warm HEPES buffered saline solution (HBSS).
  6. Aspirate the HBSS - this may be easiest/safest to do with a P1000 - then pipet 200 μL of dye solution right on the beads.
  7. Incubate for 15 min. with the TC hood light off.
  8. Remove the entire supernatant with a pipet, and expel it in the conical tube labeled Dye Collection. The dye waste will be disposed of by the teaching faculty. You should also throw the pipet tip into the beaker in your hood; tips will later be moved to the solid waste container in the chemical fume hood.
  9. Rinse the cells with 3 mL HBSS buffer again. Aspirate off as much liquid as possible, again into the Dye Collection tube.
  10. Soak in 3 mL of 4% glutaraldehyde solution for 15 minutes.
  11. Aspirate the solution, then bring your Petri dish to the fluorescent microscope bench in the lab.
  12. For observation, place the half-bead on a glass slide and then cover with a coverslip.
    • You will probably want to look at the beads both flat side up (to see the core) and flat side down (to see the surface).

Part 2: Microscopy

When observing your cells under fluorescence excitation, you should work with the room lights off for best results. You can turn on the working lamp at the microscope bench as you set up your samples, and otherwise when you need to see what you are doing.

  1. Prior to the first group using the microscope, the teaching faculty will turn on the microscope and allow it to warm up for 15-20 min. First, on the mercury lamp that is next to the microscope, the 'POWER' switch will be flipped. Next, the 'Ignition' button will be held down for about a second, then released.
  2. When you arrive, the lamp ready and power indicators should both be lit up – talk to the teaching faculty if this is not the case.
  3. Place your first sample slide on the microscope, coverslip-side up, by pulling away the left side of the metal sample holder for a moment.
  4. Begin your observations with the 10X objective.
  5. Turn on the illumination using the button at the bottom left of the microscope body (on the right-hand side is a light intensity slider).
  6. Next, turn the excitation light slider at the top of the microscope to 'DIA-ILL' (position 4).
  7. Try to focus your sample. However, be aware that the contrast is not great for your cells, and you might not be able to focus unless you find a piece of debris. Whether or not you find focus, after a minute or two, switch over to fluorescence. Your cells will be easier to find this way.
    • First, turn the white light illumination off.
    • Next, move the excitation slider to 'FITC' (position 3). You should see a blue light coming from the bottom part of the microscope.
      • This light can excite both the green and the red dye in the viability kit, and the associated filter allows you to view both colours at once.
    • Finally, you must slide the light shield (labeled 'SHUTTER') to the right to unblock it. Now you can look in the microscope, and use the coarse focus to find your cells (which should primarily be bright green), then the fine focus to get a clearer view.
    • You can also switch the excitation slider over to 'EthD-1' (position 2) to see only the red-stained cells. Some of your cells may appear to be dimly red, but the dead ones are usually obviously/brightly stained.
    • Be aware that the dyes do fade upon prolonged exposure to the excitation light, so don't stay in one place too long, and when you are not actively looking in the microscope, slide the light shield back into place.
  8. You can try looking at your cells with the 40X objective as well if you have time. As you move between objectives and samples, choose a few representative fields to take pictures of. As a minimal data set, try to get 3 fields at 10X of both of your alginate samples.
    • To take a picture, remove one eyepiece from the microscope, and replace it with the camera adaptor. Be sure to keep the light shield in place until you are ready to take the picture (to avoid photobleaching)!
    • Note that 10X images will reveal a broader field, but 40X images may have better contrast.
    • Check with the teaching faculty if you are having difficulty getting clear pictures.
    • Later in the module, you will compare the average cell numbers in each sample using the statistical methods we discussed in lecture today.
  9. Post two well-captioned pictures to the wiki before leaving (one of each sample), so we can discuss the class data in our next lecture. Be sure to note whether the image is at the surface or core of the bead.

Sample results from a student group, showing clustering of cells within the bead:

A related pair of images by Agi Stachowiak, showing cell suspensions isolated from the beads:

Part 3: Research Idea Discussion

Before or after your fluorescence assay work, find a place (across the hall, in a coffeeshop, etc.) to discuss the five research results you wrote up for homework with your lab partner, guided by the instructions below.

Writing a research proposal requires that you identify an interesting topic, spend lots of time learning about it, and then design some clever experiments to advance the field. It also requires that you articulate your ideas so any reader is convinced of your expertise, your creativity and the significance of your findings, should you have the opportunity to carry out the experiments you've proposed. To begin you must identify your research question. This may be the hardest part and the most fun. Fortunately you started by finding a handful of topics to share with your lab partner. Today you should discuss and evaluate the topics you've gathered. Consider them based on:

  • your interest in the topic
  • the availability of good background information
  • your likelihood of successfully advancing current understanding
  • the possibility of advancing foundational technologies or finding practical applications
  • if your proposal could be carried out in a reasonable amount of time and with non-infinite resources

It might be that not one of the topics you've identified is really suitable, in which case you should find some new ideas. It's also possible that through discussion with your lab partner, you've found something new to consider. Both of these outcomes are fine but by the end of today's lab you should have settled on a general topic or two so you can begin the next step in your proposal writing, namely background reading and critical thinking about the topic.

A few ground rules that are 20.109 specific:

  • You should not propose any research question that has been the subject of your UROP or research experience outside of 20.109. This proposal must be original.
  • You should keep in mind that this proposal will be presented to the class, so try to limit your scope to an idea that can be convincingly presented in a twelve minute oral presentation.

Once you and your partner have decided on a suitable research problem, it's time to become an expert on the topic. This will mean searching the literature, talking with people, generating some ideas and critically evaluating them. To keep track of your efforts, you should start a wiki catalog on your OpenWetWare user page. How you format the page is up to you but check out the "yeast rebuild" or the "T7.2" wiki pages on OpenWetWare for examples of research ideas in process. As part of your For Next Time assignment, you will have to print out your wiki page specifying your topic, your research goal and at least two helpful references that you've read and summarized.

For Next Time

  1. The first time this module was run, students created single-cell suspensions from their alginate beads by dissolving said beads in EDTA-citrate buffer, and only then stained the cells.
    • Given what you have learned in Modules 2 and 3, why does EDTA dissolve alginate beads?
    • What additional information about cell viability do you gain by staining whole constructs rather than cell isolates?
  2. Read this Tissue Engineering editorial by Professor Alan Russell about standards, and come to lecture next time prepared to discuss and/or write about your thoughts.
    • Russell, A. J. "Editorial: Standardized Experimental Procedures in Tissue Engineering: Cure or Curse." Tissue Engineering 11, nos. 9-10 (Sept-Oct 2005): vii-ix.

    You may find other helpful articles in this essay assignment from the Spring 2008 20.109 course.

  3. Begin to define your research proposal by making a wiki page to collect your ideas and resources (you can do this on one page with your partner or split the effort and each turn in an individual page). Keep in mind that your presentation to the class will ultimately need:
    • a brief project overview
    • sufficient background information for everyone to understand your proposal
    • a statement of the research problem and goals
    • project details and methods
    • predicted outcomes if everything goes according to plan and if nothing does
    • needed resources to complete the work

You can organize your wiki page along these lines or however you feel is most helpful. For now, focus on coming up with a research problem and giving a little background about it. Print your user page(s) for next time, making sure it defines your topic, your idea and two or more references you've collected and summarized. Keep in mind that you're not committed to this idea - if you come up with something that you like better later on, that's fine.

Reagent List

  • HEPES-buffered saline solution (HBSS)
    • 135 mM NaCl
    • 5 mM KCl
    • 1 mM MgSO4
    • 1.8 mM CaCl2
    • 10 mM HEPES
    • pH 7.4
  • 4% glutaraldehyde (GAH) in HBSS
    • original stock GAH at 50%, reagent grade
  • Live/Dead Reduced Biohazard Viability/Cytotoxicity kit from Invitrogen
    • SYTO 10 green nucleic acid stain
    • ethidium homodimer-2 red nucleic acid stain
    • original dye stocks in DMSO, diluted in HBSS 1:500

Contents

The first impedance-based flow cytometry device, using the Coulter principle, was disclosed in U.S. Patent 2,656,508, issued in 1953, to Wallace H. Coulter. Mack Fulwyler was the inventor of the forerunner to today's flow cytometers - particularly the cell sorter. [5] Fulwyler developed this in 1965 with his publication in Science. [6] The first fluorescence-based flow cytometry device (ICP 11) was developed in 1968 by Wolfgang Göhde from the University of Münster, filed for patent on 18 December 1968 [7] and first commercialized in 1968/69 by German developer and manufacturer Partec through Phywe AG in Göttingen. At that time, absorption methods were still widely favored by other scientists over fluorescence methods. [8] Soon after, flow cytometry instruments were developed, including the Cytofluorograph (1971) from Bio/Physics Systems Inc. (later: Ortho Diagnostics), the PAS 8000 (1973) from Partec, the first FACS (fluorescence-activated cell sorting) instrument from Becton Dickinson (1974), the ICP 22 (1975) from Partec/Phywe and the Epics from Coulter (1977/78). The first label-free high-frequency impedance flow cytometer based on a patented microfluidic "lab-on-chip", Ampha Z30, was introduced by Amphasys (2012). [ citation needed ]

Name of the technology Edit

The original name of the fluorescence-based flow cytometry technology was "pulse cytophotometry" (German: Impulszytophotometrie), based on the first patent application on fluorescence-based flow cytometry. At the 5th American Engineering Foundation Conference on Automated Cytology in Pensacola (Florida) in 1976 - eight years after the introduction of the first fluorescence-based flow cytometer (1968) - it was agreed to commonly use the name "flow cytometry", a term that quickly became popular. [9]

Modern flow cytometers are able to analyze many thousands of particles per second, in "real time" and, if configured as cell sorters, can actively separate and isolate particles with specified optical properties at similar rates. A flow cytometer is similar to a microscope, except that, instead of producing an image of the cell, flow cytometry offers high-throughput, automated quantification of specified optical parameters on a cell-by-cell basis. To analyze solid tissues, a single-cell suspension must first be prepared.

A flow cytometer has five main components: a flow cell, a measuring system, a detector, an amplification system, and a computer for analysis of the signals. The flow cell has a liquid stream (sheath fluid), which carries and aligns the cells so that they pass single file through the light beam for sensing. The measuring system commonly uses measurement of impedance (or conductivity) and optical systems - lamps (mercury, xenon) high-power water-cooled lasers (argon, krypton, dye laser) low-power air-cooled lasers (argon (488 nm), red-HeNe (633 nm), green-HeNe, HeCd (UV)) diode lasers (blue, green, red, violet) resulting in light signals. The detector and analog-to-digital conversion (ADC) system converts analog measurements of forward-scattered light (FSC) and side-scattered light (SSC) as well as dye-specific fluorescence signals into digital signals that can be processed by a computer. The amplification system can be linear or logarithmic.

The process of collecting data from samples using the flow cytometer is termed "acquisition". Acquisition is mediated by a computer physically connected to the flow cytometer, and the software which handles the digital interface with the cytometer. The software is capable of adjusting parameters (e.g., voltage, compensation) for the sample being tested, and also assists in displaying initial sample information while acquiring sample data to ensure that parameters are set correctly. Early flow cytometers were, in general, experimental devices, but technological advances have enabled widespread applications for use in a variety of both clinical and research purposes. Due to these developments, a considerable market for instrumentation, analysis software, as well as the reagents used in acquisition such as fluorescently labeled antibodies have been developed.

Modern instruments usually have multiple lasers and fluorescence detectors. The current record for a commercial instrument is ten lasers [10] and 30 fluorescence detectors. [11] Increasing the number of lasers and detectors allows for multiple antibody labeling, and can more precisely identify a target population by their phenotypic markers. Certain instruments can even take digital images of individual cells, allowing for the analysis of fluorescent signal location within or on the surface of cells.

Fluidics system of a flow cytometer Edit

Cells must pass uniformly through the center of focused laser beams to accurately measure optical properties of cells in any flow cytometer. [12] [13] [14] The purpose of the fluidic system is to move the cells one by one through the lasers beam and throughout the instrument. Fluidics in a flow cytometer with cell sorting capabilities also use the stream to carry sorted cells into collection tubes or wells. [12]

Hydrodynamic focusing Edit

For precise positioning of cells in a liquid jet, hydrodynamic focusing is used in most cytometers. [12] [13] [14] The cells in suspension enter into the instrument enclosed by an outer sheath fluid. The sample core is maintained in the center of the sheath fluid. The sample input rate or how fast the cells flow through to the laser interrogation can be controlled by the pressure of the sheath fluid on the sample core. Under optimal conditions, the central fluid stream and sheath fluid do not mix.

Acoustic-assisted hydrodynamic focusing Edit

Acoustic focusing technology is used in some flow cytometers to support hydrodynamic focusing. [12] [14] Acoustic waves (>2 MHz) pre-focus the sample before introduction to sheath fluid. The pre-focused sample is then injected into the hydrodynamic core and flowed through the instrument. This may help with increasing data accuracy under high sample input rates.

Optics and electronics Edit

Optical filters Edit

Light emitted from fluorophores are in a spectrum of wavelengths, so combining multiple fluorophores may cause overlap. To add specificity, optical filters and dichroic mirrors are used to filter and move light to the detectors such as photomultiplier tubes (PMTs) or avalanche photodiodes (APD). [12] Optical filters are designed as band pass (BP), long pass (LP), or short pass (SP) filters. Most flow cytometers uses dichroic mirrors and band pass filters to select specific bands of the optical spectrum.

Prisms, gratings, and spectral flow cytometry Edit

Spectral flow cytometry uses prisms or diffraction gratings to disperse the emitted light of a marker across a detector array. [12] [15] This allows for the full spectra from each particle to be measured. The measured spectra from single cells are subsequently unmixed by using reference spectra of all used dyes and the autofluorescence spectrum. This may allow for a wider panel design and the application of new biological markers.

Imaging flow cytometry Edit

Imaging flow cytometry (IFC) captures multichannel images of cells. [12] [16] Detectors used in imaging platforms can be equipped with charge-coupled device (CCD) or complementary metal-oxide-semiconductor (CMOS) to capture images of individual cells.

Compensation Edit

Each fluorochrome has a broad fluorescence spectrum. When more than one fluorochrome is used, the overlap between fluorochromes can occur. This situation is called spectrum overlap. This situation needs to be overcome. For example, the emission spectrum for FITC and PE is that the light emitted by the fluorescein overlaps the same wavelength as it passes through the filter used for PE. This spectral overlap is corrected by removing a portion of the FITC signal from the PE signals or vice versa. This process is called color compensation, which calculates a fluorochrome as a percentage to measure itself. [17]

Compensation is the mathematical process by which spectral overlap of multiparameter flow cytometric data is corrected. Since fluorochromes can have wide-ranging spectrum, they can overlap, causing the undesirable result of confusion during the analysis of data. This overlap, known as spillover and quantified in the spillover coefficient, is usually caused by detectors for a certain fluorochrome measuring a significant peak in wavelength from a different fluorochrome. Linear algebra is most often used to make this correction. [17]

In general, when graphs of one or more parameters are displayed, it is to show that the other parameters do not contribute to the distribution shown. Especially when using the parameters which are more than double, this problem is more severe. Currently, no tools have been discovered to efficiently display multidimensional parameters. Compensation is very important to see the distinction between cells.

Gating Edit

The data generated by flow-cytometers can be plotted in a single dimension, to produce a histogram, or in two-dimensional dot plots, or even in three dimensions. The regions on these plots can be sequentially separated, based on fluorescence intensity, by creating a series of subset extractions, termed "gates." Specific gating protocols exist for diagnostic and clinical purposes, especially in relation to hematology. Individual single cells are often distinguished from cell doublets or higher aggregates by their "time-of-flight" (denoted also as a "pulse-width") through the narrowly focused laser beam [18]

The plots are often made on logarithmic scales. Because different fluorescent dyes' emission spectra overlap, [19] [20] signals at the detectors have to be compensated electronically as well as computationally. Data accumulated using the flow cytometer can be analyzed using software. Once the data is collected, there is no need to stay connected to the flow cytometer and analysis is most often performed on a separate computer. [ citation needed ] This is especially necessary in core facilities where usage of these machines is in high demand. [ citation needed ]

Computational analysis Edit

Recent progress on automated population identification using computational methods has offered an alternative to traditional gating strategies. Automated identification systems could potentially help findings of rare and hidden populations. Representative automated methods include FLOCK [21] in Immunology Database and Analysis Portal (ImmPort), [22] SamSPECTRAL [23] and flowClust [24] [25] [26] in Bioconductor, and FLAME [27] in GenePattern. T-Distributed Stochastic Neighbor Embedding (tSNE) is an algorithm designed to perform dimensionality reduction, to allow visualization of complex multi-dimensional data in a two-dimensional "map". [28] Collaborative efforts have resulted in an open project called FlowCAP (Flow Cytometry: Critical Assessment of Population Identification Methods, [29] ) to provide an objective way to compare and evaluate the flow cytometry data clustering methods, and also to establish guidance about appropriate use and application of these methods.

FMO controls Edit

Fluorescence minus one (FMO) controls are important for data interpretation when building multi-color panels - in which a cell is stained with multiple fluorochromes simultaneously. FMO controls provide a measure of fluorescence spillover in a given channel and allow for compensation. To generate a FMO control, a sample is stained with all the fluorochromes except the one that is being tested - meaning if you are using 4 different fluorochromes your FMO control must contain only 3 of them (example: fluorochromes - A, B, C, D FMOs - ABC_, AB_D, A_CD, _BCD).

Cell sorting is a method to purify cell populations based on the presence or absence of specific physical characteristics. [12] [14] [30] In flow cytometers with sorting capabilities, the instrument detects cells using parameters including cell size, morphology, and protein expression, and then droplet technology to sort cells and recover the subsets for post-experimental use. [12] [14]

The first prototype sorter was built at the Los Alamos National Laboratory (LANL) in 1965 by physicist Mack J. Fulwyler by joining a Coulter volume sensor with the newly invented ink jet printer. [31] Live cell cell sorter or fluorescence-activated cell sorter (FACS) [a] was generated by Len Herzenberg, who subsequently won the Kyoto Prize in 2006 for his seminal work. [33]

Flow cytometry cell sorters have a collection system unlike flow cytometry analyzers. The collection process starts when a sample is injected into a stream of sheath fluid that passes through the flow cell and laser intercepts. [34] The stream then carries the cell through a vibrating nozzle which generates droplets with most containing either one cell or no cells. An electrical charging ring is placed just at the point where the stream breaks into droplets and a charge is placed on the ring based immediately prior to fluorescence intensity being measured the opposite charge is trapped on the droplet as it breaks from the stream and the droplets are therefore charged. The charged droplets then fall through an electrostatic deflection system that diverts droplets into containers based on their charge. In some systems, the charge is applied directly to the stream, and the droplet breaking off retains charge of the same sign as the stream. The stream is then returned to neutral after the droplet breaks off. After collecting, these cells can be further cultured, manipulated, and studied.

Flow cytometry uses the light properties scattered from cells or particles for identification or quantitative measurement of physical properties. Labels, dyes, and stains can be used for multi-parametric analysis (understand more properties about a cell). Immunophenotyping is the analysis of heterogeneous populations of cells using labeled antibodies [35] and other fluorophore containing reagents such as dyes and stains.

Fluorescent labels Edit

A wide range of fluorophores can be used as labels in flow cytometry. [19] Fluorophores, or simply "fluors", [ citation needed ] are typically attached to an antibody that recognizes a target feature on or in the cell they may also be attached to a chemical entity with affinity for the cell membrane or another cellular structure. Each fluorophore has a characteristic peak excitation and emission wavelength, and the emission spectra often overlap. Consequently, the combination of labels which can be used depends on the wavelength of the lamp(s) or laser(s) used to excite the fluorochromes and on the detectors available. [36] The maximum number of distinguishable fluorescent labels is thought to be 17 or 18, and this level of complexity necessitates laborious optimization to limit artifacts, as well as complex deconvolution algorithms to separate overlapping spectra. [37] Flow cytometry uses fluorescence as a quantitative tool the utmost sensitivity of flow cytometry is unmatched by other fluorescent detection platforms such as confocal microscopy. Absolute fluorescence sensitivity is generally lower in confocal microscopy because out-of-focus signals are rejected by the confocal optical system and because the image is built up serially from individual measurements at every location across the cell, reducing the amount of time available to collect signal. [38]

Quantum dots Edit

Quantum dots are sometimes used in place of traditional fluorophores because of their narrower emission peaks.

Isotope labeling Edit

Mass cytometry overcomes the fluorescent labeling limit by utilizing lanthanide isotopes attached to antibodies. This method could theoretically allow the use of 40 to 60 distinguishable labels and has been demonstrated for 30 labels. [37] Mass cytometry is fundamentally different from flow cytometry: cells are introduced into a plasma, ionized, and associated isotopes are quantified via time-of-flight mass spectrometry. Although this method permits the use of a large number of labels, it currently has lower throughput capacity than flow cytometry. It also destroys the analysed cells, precluding their recovery by sorting. [37]

In addition to the ability to label and identify individual cells via fluorescent antibodies, cellular products such as cytokines, proteins, and other factors may be measured as well. Similar to ELISA sandwich assays, cytometric bead array (CBA) assays use multiple bead populations typically differentiated by size and different levels of fluorescence intensity to distinguish multiple analytes in a single assay. The amount of the analyte captured is detected via a biotinylated antibody against a secondary epitope of the protein, followed by a streptavidin-R-phycoerythrin treatment. The fluorescent intensity of R-phycoerythrin on the beads is quantified on a flow cytometer equipped with a 488 nm excitation source. Concentrations of a protein of interest in the samples can be obtained by comparing the fluorescent signals to those of a standard curve generated from a serial dilution of a known concentration of the analyte. Commonly also referred to as cytokine bead array (CBA).

Impedance-based single cell analysis systems are commonly known as Coulter counters. They represent a well-established method for counting and sizing virtually any kind of cells and particles. The label-free technology has recently been enhanced by a "lab-on-a-chip" based approach and by applying high frequency alternating current (AC) in the radio frequency range (from 100 kHz to 30 MHz) instead of a static direct current (DC) or low frequency AC field. [39] [40] This patented technology allows a highly accurate cell analysis and provides additional information like membrane capacitance and viability. The relatively small size and robustness allow battery powered on-site use in the field.

    (quantification, measurement of DNA degradation, mitochondrial membrane potential, permeability changes, caspase activity)
  • Cell adherence (for instance, pathogen-host cell adherence)
  • Cell pigments such as chlorophyll or phycoerythrin
  • Cell surface antigens (Cluster of differentiation (CD) markers)
  • Cell viability : isolation and purification
  • Characterising multidrug resistance (MDR) in cancer cells and sorting (library construction, chromosome paint) copy number variation (by Flow-FISH or BACs-on-Beads technology) activity antigens (various cytokines, secondary mediators, etc.)
  • Monitoring electropermeabilization of cells
  • Nuclear antigens , intracellular ionizedcalcium, magnesium, membrane potential expression and localization
  • Protein modifications, phospho-proteins
  • Scattering of light can be used to measure volume (by forward scatter) and morphological complexity (by side scatter) of cells or other particles, even those that are non-fluorescent. These are conventionally abbreviated as FSC and SSC respectively.
  • Total DNA content (cell cycle analysis, cell kinetics, proliferation, ploidy, aneuploidy, endoreduplication, etc.)
  • Total RNA content
  • Transgenic products in vivo, particularly the green fluorescent protein or related fluorescent proteins
  • Various combinations (DNA/surface antigens, etc.)

The technology has applications in a number of fields, including molecular biology, pathology, immunology, virology, [41] plant biology and marine biology. [42] It has broad application in medicine especially in transplantation, hematology, tumor immunology and chemotherapy, prenatal diagnosis, genetics and sperm sorting for sex preselection. Flow cytometry is widely applied to detect sperm cells abnormality associated with DNA fragmentation [43] in male fertility assays. [44] Also, it is extensively used in research for the detection of DNA damage, [45] [46] caspase cleavage and apoptosis. [47] Photoacoustic flow cytometry is used in the study of multi-drug-resistant bacteria (most commonly MRSA) to detect, differentiate, and quantify bacteria in the blood marked with dyed bacteriophages. [48] In neuroscience, co-expression of cell surface and intracellular antigens can also be analyzed. [49] In microbiology, it can be used to screen and sort transposon mutant libraries constructed with a GFP-encoding transposon (TnMHA), [50] or to assess viability. [51] In protein engineering, flow cytometry is used in conjunction with yeast display and bacterial display to identify cell surface-displayed protein variants with desired properties. The main advantages of flow cytometry over histology and IHC is the possibility to precisely measure the quantities of antigens and the possibility to stain each cell with multiple antibodies-fluorophores, in current laboratories around 10 antibodies can be bound to each cell. This is much less than mass cytometer where up to 40 can be currently measured, but at a higher price and a slower pace.

Aquatic research Edit

In aquatic systems, flow cytometry is used for the analysis of autofluorescing cells or cells that are fluorescently-labeled with added stains. This research started in 1981 when Clarice Yentsch used flow cytometry to measure the fluorescence in a red tide producing dinoflagellate. [52] The next year researchers published flow cytometric measurements of multiple algal species which could be distinguished based on their fluorescence characteristics. [53] By 1983, marine researchers were assembling their own flow cytometers [54] or using commercially available flow cytometers on seawater samples collected off Bermuda to demonstrate that phytoplankton cells could be distinguished from non-living material and that cyanobacteria could be sorted from a mixed community and subsequently cultured in the lab. [55] Flow cytometry also allowed marine researchers to distinguish between dimly-fluorescing Prochlorococcus and heterotrophic microorganisms, a distinction that is difficult with microscopy-based assessments. [56] Advances in technology now allow aquatic scientists to use flow cytometers continuously during research cruises [57] and flow cytometers are used to provide images of individual phytoplankton cells. [58] [59] Marine scientists use the sorting ability of flow cytometers to make discrete measurements of cellular activity and diversity, [60] [61] to conduct investigations into the mutualistic relationships between microorganisms that live in close proximity, [62] and to measure biogeochemical rates of multiple processes in the ocean. [63]

Cell proliferation assay Edit

Cell proliferation is the major function in the immune system. Often it is required to analyse the proliferative nature of the cells in order to make some conclusions. One such assay to determine the cell proliferation is the tracking dye carboxyfluorescein diacetate succinimidyl ester (CFSE). It helps to monitor proliferative cells. This assay gives quantitative as well as qualitative data during time-series experiments. [64] This dye binds covalently with the long-lived molecules present inside the cell. When the cells divide, the molecules divide too and, the daughter cells possess half the dye than the parent population. This decrease in the intensity can be visualized by flow cytometry. [65] In literature, this powerful technique of flow cytometry and CFSE has been used to find the efficiency of T-cells in killing the target cells in cancer such as leukemia. In order to visualize the target cell death, both rapid and slow, scientists have used CFSE labelling with antibody staining of certain kinds of cells and fluorescently labelled microbeads. This also gave information regarding the proliferation of the target cells upon the treatment of certain cytokines. [66]


1 INTRODUCTION

Cell viability is defined as the number of healthy cells in a sample. The measurement of cell viability plays an important role for all forms of cell culture. Sometimes it is the main purpose of the experiment as in toxicity assays, or it can be used to correlate cell behavior to the number of cells (Stoddart, 2011 ). Cell viability assays are essentially used for screening the response of the cells against a drug or a chemical agent. In particular, pharmaceutical industry widely uses viability assays to evaluate the influence of developed agents on the cells. Researchers apply various types of assays in order to screen the outcome of a developed therapeutics that often target cancer cells (Adan, Kiraz, & Baran, 2016 ).

There are several types of assays that can be used to determine the number of viable cells. These assays are based on various functions of cells including enzyme activity, cell membrane permeability, cell adherence, adenosine triphosphate (ATP) production, co-enzyme production, and nucleotide uptake activity (Thangaraj, 2016 ). Although there are different classifications, cell viability assays may be broadly categorized as (a) dye exclusion assays, (b) colorimetric assays, (c) fluorometric assays, (d) luminometric assays, and (e) flow cytometric assays. Dye exclusion assays are the simplest methods that are based on utilization of different dyes such as trypan blue, eosin, congo red, and erythrosine B, which are excluded by the living cells, but not by dead cells. For these assays, although staining procedure is quite straightforward, experimental procedure may be time-consuming in case of large sample sizes. Colorimetric assays are based on the measurement of a biochemical marker to determine the metabolic activity of the cells. In these assays, the colorimetric measurement of cell viability is carried out spectrophotometrically. 3-[4,5-Dimethylthiazol-2-yl]-2,5 diphenyl tetrazolium bromide (MTT), 3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium (MTS), 2,3-bis-(2-methoxy-4-nitro-5-sulfophenyl)-2H-tetrazolium-5-carboxanilide (XTT), 2-(4-iodophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H tetrazolium, monosodium salt (WST-1), 2-(2-methoxy-4-nitrophenyl)-3-(4-nitrophenyl)-5-(2,4-disulfophenyl)-2H-tetrazolium, monosodium salt (WST-8), lactate dehydrogenase (LDH), sulforhodamine B (SRB), neutral red uptake (NRU), and crystal violet stain (CVS) assays are among the most widely applied colorimetric assays. These assays are simple and economical, and can be applied to both cell suspensions and adherent cells. Fluorometric assays including resazurin and 5-carboxyfluorescein diacetate acetoxymethyl ester (5-CFDA-AM) assays may be performed with a fluorometer, fluorescence microplate reader, fluorescence microscope, or flow cytometer. These assays are advantageous over dye exclusion and colorimetric assays as they are more sensitive. In luminometric assays, a persistent and stable glow-type signal is produced following the addition of reagent. These methods comprise ATP and real-time viability assays (Aslantürk, 2018 ). Flow cytometry allows simultaneous measurement of the changes in cell morphology by forward and side light scatter, which makes this technology uniquely suited to measuring the complex progression of cell death (Telford, 2012 ). Major flow cytometric assays include membrane asymmetry (e.g., annexin V and F2N12S staining assays), membrane permeability (e.g., nucleic acid and inclusion and exclusion dyes), and mitochondria assays.

When selecting the appropriate cell viability assay, factors including cost, speed, sensitivity, and the required equipment should be considered in order to obtain reliable results (Shokrzadeh & Modanloo, 2017 ). An ideal cell viability assay should be safe, rapid, reliable, efficient, and time- and cost-effective, and should not interfere with the test compound (Aslantürk, 2018 ). On the other hand, regardless of the assay chosen, the most critical factors for accurate and reproducible measurements include (a) the use of a controlled and consistent source of cells to set up experiments and (b) performing suitable characterization of reagent concentration and incubation time for each experimental model system (Riss et al., 2016 ). Considering the above, in this guideline, the mechanisms and the practice of assessment of the most common cell viability assays applied in research labs are discussed in detail.


Assays and Tools to Monitor Biology in 3D Cell Cultures

The Assay Guidance Manual: A Guide for In Vitro and In Vivo Assays in Early Drug Discovery

Cell Viability Assay Guide


Volume 1

Cell viability and proliferation

Cell viability and proliferation are important metrics of a material or construct for tissue repair. They can provide important feedback on the suitability of surface modifications, three-dimensional architecture, oxygen transport, degradation product compatibility, and many other aspects of the material or structure. While often used interchangeably cell viability and proliferation are similar to each other in that both are measures of live cells but distinct in that viability is a measure of cellular activity and overall health, while proliferation is a measure of the rate of growth and production of daughter cells of a cell population. Either may be suitable as a tool to measure the health of a cell population and the suitability of a construct or material for cell survival but the assays and techniques used to assess each can vary and care should be taken to ensure that the method of analysis or choice of assay is appropriate. Generally speaking, proliferation is required to have a construct fully populated by cells which in turn mandates a fully interconnected pore structure or other means of adequate nutrient and waste exchange to allow the cells to remain viable within the depths of the construct. If the construct has insufficient nutrient transport cells will not proliferate to the center of the construct and new tissue will not be deposited. Measuring proliferation is one measure of how well a construct can support the residence of cells throughout its structure. Common assays used to test cellular viability include the MTT and MTS assays, which measure the viability of a cell through its metabolic activity, DNA synthesis assays which can translate to actual cell numbers, and manual cell counts using a dye such as trypan blue, which, as discussed earlier, will permeate a damaged cell membrane as an indicator of a nonviable, or dead, cell.


Cell Proliferation and Cytotoxicity Assays

Cell viability is defined as the number of healthy cells in a sample and proliferation of cells is a vital indicator for understanding the mechanisms in action of certain genes, proteins and pathways involved cell survival or death after exposing to toxic agents. Generally, methods used to determine viability are also common for the detection of cell proliferation. Cell cytotoxicity and proliferation assays are generally used for drug screening to detect whether the test molecules have effects on cell proliferation or display direct cytotoxic effects. Regardless of the type of cell-based assay being used, it is important to know how many viable cells are remaining at the end of the experiment. There are a variety of assay methods based on various cell functions such as enzyme activity, cell membrane permeability, cell adherence, ATP production, co-enzyme production, and nucleotide uptake activity. These methods could be basically classified into different categories: (I) dye exclusion methods such as trypan blue dye exclusion assay, (II) methods based on metabolic activity, (III) ATP assay, (IV) sulforhodamine B assay, (V) protease viability marker assay, (VI) clonogenic cell survival assay, (VII) DNA synthesis cell proliferation assays and (V) raman micro-spectroscopy. In order to choose the optimal viability assay, the cell type, applied culture conditions, and the specific questions being asked should be considered in detail. This particular review aims to provide an overview of common cell proliferation and cytotoxicity assays together with their own advantages and disadvantages, their methodologies, comparisons and intended purposes.


Background

The evaluation of cell population density (i.e. the total number of living cells in the culture) and cell viability (i.e. the percentage of living cells in the sample) is fundamental during biology studies [1]. The majority of laboratories engaged in cell biology routinely perform cell viability and counting analysis for different purposes, ranging from ecosystem investigation [2] to proliferation studies [3], in both 2D (two-dimensional) [4] and 3D (three-dimensional) cell cultures [5].

Among the various typologies of 3D cell cultures, multicellular tumour spheroids are those typically used for testing drugs and radiation treatments [6]. The measurement of viability and the reduction of cancer culture population are fundamental parameters for evaluating the efficacy of the treatments under investigation [7]. Accordingly, the reliability of the method used to estimate these parameters plays a key role in this analysis [8]. In addition, cell counting and viability assessment often need to be performed for other 3D cell cultures, such as stem cell spheroids generated for regenerative medicine purposes [9], and organoids used to study (some) organ characteristics [10].

Many different methods (e.g. AlamarBlue ® and MMT assay) and systems (e.g. Bio-Rad TC20™ Automated Cell Counter, ChemoMetec NucleoCounter ® , Beckman Coulter Vi-CELL™ XR Cell Viability Analyzer [11]) can be used to analyse cell viability [12]. Most of these share the same approach: the cells are stained using a light (or a fluorescent) dye to highlight dead cells (or living cells), and a detection system counts the number of cells highlighted, in addition to the total number of cells. Finally, cell viability is computed as the percentage of healthy cells in the sample [13]. However, the Trypan Blue (TB) dye exclusion assay [14] ,the first method proposed in the literature, is considered the standard cell viability measurement method [15] and is still the most widely used approach [16]. Furthermore, TB paired with a haemocytometer grid (Fig. 1) is regarded as the standard approach for estimating the cell population density [17], i.e. the total number of living cells in the culture [18].

Haemocytometer grid containing cells stained with TB. a Picture of a Kova glasstic slide with grids (Hycor Biomedical Inc.). Each slide contains 10 counting chambers. b Schematic representation of the grid of a counting chamber. c Cells in brightfield are characterized by very low contrast. This magnified real-world detail shows some living and dead cells. In particular: a and b show the typical appearance of a living and a dead cell (stained with TB), respectively

TB was synthesised for the first time in 1904 by Paul Ehrlich (Nobel prize in medicine, 1908) and was first used for clinical analysis before becoming a standard probe in biology. Today it is still widely used for several medical purposes such as the visualization of the lymph-associated primo vascular system [19] and of the anterior capsule during cataract surgery [20]. Chemically, TB is defined as toluidine-derived dye characterized by a molecular weight of 960 Da [15]. Its chemical construction is C 34 H 28 N 6 O 14 S 4. Azidine Blue, Benzamine Blue, Chlorazol Blue, Diamine Blue, and Niagara Blue are synonyms for TB. TB is a cell membrane-impermeable molecule and therefore only enters cells having compromised membrane. From a practical point of view, with TB the cell viability is determined indirectly by detecting cell membrane integrity [21]. Upon entry into the cell, TB binds to intracellular proteins and in brightfield the dead cells appear blue (apoptotic and necrotic cells are not distinguished [1]), whereas the colour of living cells remains unchanged (Fig. 1c).

Over the past two decades a number of studies comparing TB with other assays have been published [15] and several methods have proven more efficient than TB [22], especially those using fluorescent dyes [23]. The use of TB has, in fact, several drawbacks [24]: (a) TB exerts a toxic effect on cells after a short exposure period, thus limiting cell counting to only a brief period after staining [25] (b) As TB binds to cellular proteins, there is a potential for binding to non-specific cellular artifacts, especially in primary cells from clinical samples (c) There is a large number of false positives, i.e. “dead cells” resulting from irreversible damage to their membrane, and false negatives from cells that have already initiated the apoptotic pathway but still have intact membranes (d) There is no standardized TB concentration for the measurement of cell viability (e) Manual counting using a haemocytometer and a light microscope is time-consuming and operator-dependent. Although the TB assay requires the use of a fluorescence microscope, it has long been known that several fluorescent dyes are more reliable indicators of cell viability than the more traditional coloured dyes [26]. For example, Acridine Orange (AO) and Propidium Iodide (PI) stainings have been shown to be more accurate in detecting live and dead cells than TB [27]. AO is a membrane-permeable cationic dye that binds to nucleic acids of viable cells. At low concentrations it causes a green fluorescence. PI is impermeable to intact membranes but readily penetrates the membranes of nonviable cells and binds to DNA or RNA, causing orange fluorescence. When AO and PI are used simultaneously, viable cells fluoresce green and nonviable cells fluoresce orange under fluorescence microscopy. Notwithstanding, TB is still the most commonly used dye for cell viability analysis because it is inexpensive, easy to use, it reacts quickly, and can be visualized with a standard brightfield microscope available in all biological laboratories [2]. TB is also used in several automatic counters [28] and as the reference method for comparing customized cell-counting algorithms [29]. However, in-depth validation studies of the TB assay used in combination with a haemocytometer in viability and counting measurements are lacking. Several articles have provided statistical analyses on its reliability. In 1964, Tennant [30] and Hathaway et al. [31] performed preliminary studies comparing TB, eosin Y and AO for the determination of the viability of in vitro and in vivo cultures. Twenty years later, Jones and Senft [26] also considered fluorescein diacetase (FDA) and PI. In 1999, Leite et al. [32] extended the research into this area, comparing the reliability of TB, AO and six other methods (i.e. Giemsa staining, ethidium bromide, PI, Annexin V, TUNEL assay and DNA ladder). In 2000, Mascotti et al. [27] published an in-depth comparison between AO/PI and TB assays in which the viability of 7 aliquots of hematopoietic progenitor cells (HPC) and the percentage of viable cells was calculated as the average of 5 viability measurements performed by two operators. However, as the raw counting data was not reported, it was not possible to quantitatively infer the repeatability (intra-rater reliability) and reproducibility (inter-rater reliability) of the counts. The first study on the repeatability and reproducibility of the TB assay appeared in 2011 when Sanfilippo et al. [33] assessed the reliability of TB and calcein AM/ethidium homodimer-1 (CaAM/EthD-1) staining in fresh and thawed human ovarian follicles. Measurements were performed by two independent operators. Reliability was evaluated by the intraclass correlation coefficient (ICC) and the differences between paired measurements were tested by the Wilcoxon signed-rank test. TB proved to be the more reliable staining method to evaluate follicle viability. However, the operators only evaluated 10 samples simultaneously. Finally, in 2015 Cadena-Herrera et al. [34] validated a manual, semi-automated, and fully automated TB exclusion-based methods. A single operator counted several samples in triplicate and the results obtained did not reveal a significant difference between the automated methods and the manual assay. However, 3D cell cultures were not taken into account and no considerations about measurement errors between different operators were made.

In this work we studied repeatability and reproducibility with the specific aim of assessing measurement errors occurring when TB is used in counting and viability applications in 2D and 3D cell cultures. Repeatability is the closeness of the agreement among subsequent measurements of the same object carried out under the same measurement conditions. Reproducibility is defined as the closeness of the agreement among measurements of the same object carried out under different measurement conditions [35]. In particular, the viability and total number of living cells of the culture were the “objects” being measured in our experiments. Thus, the operators performing the measurements represented the changing “condition” when assessing reproducibility. In practical terms, each operator generated and analysed 5 different samples from the same 13 2D cell cultures and 8 3D cell cultures (i.e. multicellular spheroids), making a total of 10 samples considered for each culture. Repeatability for each culture was evaluated by calculating the variability of the measurements obtained by the single operator. Conversely, reproducibility for each culture was estimated by comparing the measurements obtained by two operators. Overall, 210 samples were analysed (Table 1).

The main aim of this work was to make researchers aware of the measurement errors that can occur when the TB assay is used to evaluate population and viability of 2D and 3D cell cultures. Given that this is a preliminary study, global accurate overall accuracy values of assay reliability used in different contexts and with different cell lines cannot be provided. However, we believe that our findings can help researchers to evaluate whether the expected repeatability and reproducibility of the TB assay are compliant with those required by their own application.


Cell Form and Function

Cells with different functions often have varying shapes. The cells pictured below (Figure 4.3.3) are just a few examples of the many different shapes that human cells may have. Each type of cell has characteristics that help it do its job. The job of the nerve cell, for example, is to carry messages to other cells. The nerve cell has many long extensions that reach out in all directions, allowing it to pass messages to many other cells at once. Do you see the tail of each tiny sperm cell? Its tail helps a sperm cell “swim” through fluids in the female reproductive tract in order to reach an egg cell. The white blood cell has the job of destroying bacteria and other pathogens. It is a large cell that can engulf foreign invaders.

Figure 4.3.3 Human cells may have many different shapes that help them to do their jobs.


Assay Protocol to Measure Cell Growth

Additional Reagents Required:

Culture medium, e.g., DMEM (D5671) containing 10% heat inactivated FCS (fetal calf serum, 12106C), 2 mM glutamine (G6392), 0.55 mM L-arginine (A8094), 0.24 mM L-asparagine-monohydrate (A4284), 50 μM 2-mercaptoethanol (M3148), HT-media supplement (H0137) (1×), containing 0.1 mM hypoxanthine and 16 μM thymidine. If an antibiotic is to be used, additionally supplement media with penicillin/streptomycin or gentamicin. Interleukin-6, human (hIL-6, SRP3096) (200,000 U/ml, 2 μg/ml) sterile.

Protocol:

For the determination of human interleukin-6 (hIL-6) activity on 7TD1 cells (mouse-mouse hybridoma, DSMZ, ACC 23) (see fig. 3).

  1. Seed 7TD1 cells at a concentration of 2 × 10 3 cells/well in 100 μl culture medium containing various amounts of IL-6 [final concentration e.g., 0.1-10 U/ml (0.001-0.1 ng/ml)] into microplates (tissue culture grade, 96 wells, flat bottom).
  2. Incubate cell cultures for 4 days at +37 °C and 5-6.5% CO2.
  3. After the incubation period, add 10 μl of the MTT labeling reagent (final concentration 0.5 mg/ml) to each well.
  4. Incubate the microplate for 4 h in a humidified atmosphere (e.g., +37 °C, 5-6.5% CO2).
  5. Add 100 μl of the Solubilization solution into each well.
  6. Allow the plate to stand overnight in the incubator in a humidified atmosphere (e.g., +37 °C, 5-6.5% CO2).
  7. Check for complete solubilization of the purple formazan crystals and measure the spectrophotometrical absorbance of the samples using a microplate (ELISA) reader. The wavelength to measure absorbance of the formazan product is between 550 and 600 nm according to the filters available for the ELISA reader, used. The reference wavelength should be more than 650 nm.

Figure 3. Proliferation of 7TD1 cells (mouse-mouse hybridoma) in response to recombinant human interleukin-6 (hIL-6) using MTT assay.

Similar Assays

XTT assay and WST-1 assay can also be used for measuring cell viability and proliferation.


Watch the video: Cell Viability Assays (June 2022).


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