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Jun 2017

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Relative Quantitation of Polymerized Actin in Suspension Cells by Flow Cytometry
利用流式细胞术相对定量分析悬浮细胞中的聚合肌动蛋白   

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Abstract

The amount of polymerized actin within a cell can vary widely due to natural processes and/or experimentally induced perturbations. We routinely use this protocol to measure relative polymerized actin content between cell populations by staining cells in suspension with fluorescent phalloidin, then measuring total cell fluorescence using flow cytometry. Although developed for human cells, we have easily adapted this method for use with diverse eukaryotic cell types.

Keywords: Actin (肌动蛋白), F-actin (F-肌动蛋白), Quantitation (定量), Polymer (聚合物), Cells (细胞), Cytometry (细胞计数法), FACS (FACS)

Background

Biologists have long studied actin polymers using biochemical assays and fluorescence microscopy. The latter has been facilitated by the discovery of phalloidin, a small molecule toxin produced by poisonous mushrooms that stabilizes actin structures by specifically binding adjacent monomers within actin polymers (Vandekerckhove et al., 1985). Fluorescent phalloidin conjugates stain complex actin structures and facilitate detailed visualization by fluorescence microscopy. However, quantitating this fluorescence can be painstaking, particularly when tracing population-level changes. In contrast, flow cytometry quantitates the total fluorescence intensity of thousands of cells per minute.

Here we describe how we estimate differences in the amount of actin polymer between cell populations by staining cells with fluorescent phalloidin, measuring the total cell fluorescence of individual cells using flow cytometry, and finally comparing the fluorescence values of the populations. It is important to note that this approach is comparative and cannot be used to measure the absolute concentrations of cellular actin polymer. This approach requires cells to be in suspension; if your cells require adhesion for proper actin polymerization, we suggest using quantitative fluorescence microscopy as described above.

Although rapid and quantitative, flow cytometry provides no information about cell morphology. Therefore, we routinely save an aliquot of stained cells for visualization using fluorescence microscopy. We highly recommend doing this to ensure proper preservation of your structures of interest (Figure 1). Fluorescence microscopy can also reveal variation in experimental samples that are not apparent by flow cytometry.

This protocol is optimized for differentiated HL-60 cells that can be stimulated with fMLP to induce rapid actin polymerization (Figure 1, for additional information about these cells, see Fritz-Laylin et al., 2017), it can be modified for use with a wide variety of cell types by testing a variety of fixation conditions and buffers. For example, we have adapted this protocol for chytrid fungi (Batrachochytrium dendrobatidis), and the amoeba Naegleria gruberi (unpublished). To do this, we chose conditions that: 1) best preserve the overall morphology reasonably similar to live cells as visualized using DIC or phase contrast microscopy; 2) keep the polymerized actin intact through fixation; 3) when imaged by fluorescence microscopy, stain the appropriate structures. For information on buffers and fixatives (see Note 1) commonly used to preserve actin structures, we recommend the methods resource Fluorescence Procedures for the Actin and Tubulin Cytoskeleton in Fixed Cells from the Mitchison lab at Harvard University (L. Cramer and A. Desai).


Figure 1. An example of actin polymerization in response to cell stimulation. Maximum projection of AlexaFluor® 488 phalloidin stained HL-60 cells before and after treatment with 20 nM fMLP, which induces actin polymerization and results in brighter cells.

Materials and Reagents

  1. Standard uncoated 1.5 ml microcentrifuge tubes (USA Scientific, catalog number: 1615-5510)
  2. 15 and 50 ml conical tubes (any supplier)
  3. FACS tubes: sterile 5 ml polystyrene round-bottom tubes with a cell-strainer cap for filtering samples prior to flow cytometry (BD Falcon, catalog number: 352235), external filters are also available
    Note: Check your cytometer requirements and/or your flow cytometry facility's recommendation for tubes.
  4. Micropipette tips (Pipette.com, catalog numbers: UE-1000, UE-200, UE-10)
  5. Aluminum foil
  6. Razor blades
  7. Differentiated HL-60 cells (grown in medium RPMI supplemented with 10% Fetal Bovine Serum, and treated with 1.3% DMSO for 5 days; for additional details see Fritz-Laylin et al., 2017), or any other cell type for which you wish to quantitate relative amounts of polymerized actin 
  8. RPMI (Gibco, catalog number: 11875)
  9. AlexaFluor® 488 conjugated Phalloidin (Thermo Fisher Scientific, catalog number: A12379) (see Note 3)
  10. Sterile Milli-Q water
  11. TWEEN-20 (Sigma-Aldrich, catalog number: P9416)
  12. Triton X-100 (Sigma-Aldrich, catalog number: X100)
  13. MES (Sigma-Aldrich, catalog number: M3671)
  14. KCl (Fisher Scientific, catalog number: P217-100)
  15. MgCl2 (Sigma-Aldrich, catalog number: M9272)
  16. EGTA (Sigma-Aldrich, catalog number: E4378)
  17. NaCl (Fisher Scientific, catalog number: BP3581)
  18. Na2HPO4 (Sigma-Aldrich, catalog number: 795410)
  19. KH2PO4 (Sigma-Aldrich, catalog number: P0662)
  20. BSA (Fisher Scientific, catalog number: BP1600-100)
  21. 16% Paraformaldehyde (Electron Microscopy Sciences, catalog number: 15710)
  22. DMSO (Sigma-Aldrich, catalog number: D2650)
  23. fMLP (Sigma-Aldrich, catalog number: 47729)
  24. Culture medium for HL-60 cells (see Recipes)
  25. Fixation Buffer (see Recipes)
  26. Cytoskeleton Buffer (see Recipes)
  27. Permeabilization Buffer (see Recipes)
  28. Staining solution (see Recipes)
  29. PBS (10x) (see Recipes)
  30. Wash solution (see Recipes)

Equipment

  1. Pipettes (Eppendorf, P1000, P100, P20, P1)
  2. Microcentrifuge with rotor suitable for 1.5 ml centrifuge tubes (e.g., Eppendorf, model: 5415)
  3. Flow cytometer (e.g., FACSCalibur analyzer [BD, model: FACSCaliburTM] or LSRFortessa Dual [BD, model: LSRFortessaTM])
  4. Microscope equipped for fluorescence and transmitted light (we prefer phase and/or DIC) (e.g., Nikon, model: Eclipse Ti-E)
  5. Incubator for growing cells, for HL-60 cells you will need a chamber with continuous 37 °C and 5% CO2
  6. Autoclave
  7. Microscope filters approprisate for the phalloidin conjugate of your choice (see Note 3)
  8. Standard hemocytometer (e.g., Sigma-Aldrich, catalog number: Z359629)
  9. Vortexer 
  10. -20 °C Freezer

Software

  1. FACSDiva, BD (Becton, Dickinson & Company)
  2. FlowJo version 10, BD (Becton, Dickinson & Company)
  3. GraphPad Prism version 7, GraphPad Software

Procedure

  1. Fixing cells
    1. Prepare fresh fixation buffer and keep on ice until needed. 
    2. Label 1.5 ml tubes for each sample, including untreated and treated cells that will be stained with phalloidin, as well as matched control samples that will be processed identically but will not be stained with phalloidin (unstained controls).
    3. Prepare cells by concentrating to desired density by centrifugation and suspension in 100 μl per sample of fresh media or buffer; the minimum number of cells typically required for FACS analysis is 10,000 per sample (see Note 6). For stimulation of HL-60 cells, resuspend in serum-free RPMI supplemented with 2% BSA, and incubate for 1 h at 37 °C and 5% CO2 to depolarize the cells. 
    4. Stimulate cells by adding 10 ul of 200 nM fMLP, gently tapping tube to mix, and incubating at 37 °C for 3 min (treated sample), followed by immediate fixation (do not do this for the control, untreated samples).
    5. Immediately fix cells in 1.5 ml centrifuge tubes by adding 400 µl of cold fixation buffer to each sample. 
    6. Mix the sample by gently pipetting up and down. 
    7. Incubate on ice for 20 min (see Note 5).

  2. Staining cells
    1. Centrifuge cells at 4 °C to form a pellet (see Note 6). 
    2. Remove the supernatant and resuspend cells in 100 µl of permeabilization buffer (see Note 2), and mix gently by pipetting up and down. 
    3. Incubate for 10 min at room temperature. 
    4. Centrifuge to pellet cells at either room temperature or 4 °C.
    5. Remove supernatant and resuspend cells in 100 µl of staining solution (see Recipes and Note D). 
    6. Gently resuspend cells by pipetting up and down. 
    7. Cover samples with aluminum foil and incubate at room temperature for 20 min then transfer to 4 °C. The length of time that samples can be stored in phalloidin can vary depending on the cell type; test this empirically when working with a new cell type (see Note 5). 
    8. When ready to analyze cells by flow cytometry, centrifuge to form a pellet and remove the phalloidin supernatant. 
    9. Wash several times by gently resuspending cells with 500 µl of PBS supplemented with 0.1% TWEEN-20 (a mild detergent used to break up the cells and avoid clumping) and centrifuge at 4 °C to form a pellet. After the last wash, resuspend in at least 500 µl of pre-chilled PBS and store samples on ice. 
    10. Filter samples before flow cytometry (see Note 7).
    11. Check stained samples via fluorescence microscopy to ensure target structures are stained and that the overall cell morphology has not been compromised (the cells should look approximately similar to how they looked when alive).
    12. To ensure there are enough cells for the experiment, take an aliquot of stained cells and enumerate using a hemocytometer.

  3. Flow Cytometry
    This is not intended to be a complete guide to flow cytometry, and we suggest that readers unfamiliar with this technology see: the Flow Cytometry Basics Guide from Bio-Rad, FlowJo video tutorials such as Cytometry 101 (https://www.flowjo.com/learn/flowjo-university/flowjo/before-flowjo/2) or BD Biosciences Introduction to Flow Cytometry Web-Based Training (https://www.bdbiosciences.com/us/support/s/itf_launch). For the HL-60 cell data presented here, we used a FACSCalibur analyzer (BD).
    1. Setting Parameters
      For this analysis, we measure FSC-height, SSC-height, and FIT-C: height, log. We also recommend collecting either FSC-width, SSC-width, and/or FSC-area if the cytometer has those capabilities, as these values will allow for elimination of doublets (see Note 9). The differences in sample fluorescence are large, so we display the fluorescence intensities in log scale.
        FACSDiva software displays a worksheet populating data into histograms, scatter plots, and provides population statistics based on gating. We recommend setting up a global worksheet to visualize data during the adjustments and acquisition.
        Depending on your cytometer, adjust the laser intensity and/or the detector to 1) capture forward and side scatter (to eliminate dead cells, clumps, doublets, etc. see Note 9 for doublet elimination) and 2) make sure that your samples are fluorescing within the detectable range. Adjust these parameters as your control sample is running through the flow cytometer at a low speed. Typically, a scatterplot with forward scatter and side scatter will show the population in real time as cytometer collects data. The target population should be well separated from the axes to avoid exclusion of data. Using a selection tool, create an initial gate by setting points around the entire population and adjust the cytometer to center the fluorescence intensity of the entire cells in the population. Do this very carefully because you cannot later recover data outside of the range of the detector.
    2. Data acquisition
      Once the lasers and detector are properly adjusted, use the same laser intensities on all data collected for all samples in the replicate. The cytometer can (and should) be adjusted for each experimental replicate, but once set for your experiment, analyze all samples using the same cytometer settings (see Note 8). If you have an unstained sample, a low-level background fluorescence is normal as dead cells autofluoresce.

Data analysis

We use FlowJo software (Tree Star) for analysis of flow cytometry data. Here, we briefly describe our analysis workflow. For more detailed discussion of analysis of cytometry data, see the data analysis tutorials for BD FACSDiva (https://www.bdbiosciences.com/us/support/s/itf_launch) or FlowJo (https://www.flowjo.com/learn/flowjo-university/flowjo).


  1. Groups
    Importing files into FlowJo should automatically populate groups according to file names. If you need to create groups for analysis, there is a tool to select groups based on keywords in the file name ($fil) and other parameters.

  2. Gating
    Apply gates uniformly to all samples in the entire replicate. The gates described here 1) select intact cells and exclude cell debris; 2) establish populations; 3) allow for comparative analyses between samples (see Note 8).
    1. Open the graphing window and create a scatterplot with SSC-height (SSC-H) on the y-axis and FSC-height (FSC-H) on the x-axis. We like to use the pseudocolor option for the scatterplot since the monochrome dot plot reduces the number of events to represent only some of the data. Also, pseudocolor is easy to visualize the concentrations of events to establish gating.
    2. If the cytometer has the capacity to collect width and/or area data, gate the cell population to eliminate doublets (see Note 9).
    3. Select a cell population, being careful to not exclude data from the main cell population, using the ellipse tool (Figure 2). 
    4. Open this gated population as a histogram. Display the x-axis as “FIT-C” and the y-axis as a histogram.
    5. Add statistics such as median.
    6. Copy the analysis to the entire group. This creates a group gating hierarchy to allow for analysis of relative actin intensities within that group.


      Figure 2. Flow cytometry and analysis of actin polymerization. As an example of the experiment outlined in this protocol, we analyzed untreated and stimulated HL-60 cells as shown in Figure 1. To exclude cell debris and/or clumps, first set a forward and side scatter gate around the concentrated cell population (shown in pseudocolor) and apply it to all samples uniformly. Note: When differentiated with DMSO, not all of the cells fully differentiate, resulting in two populations of cells seen here. The histograms show the relative intensity of cells within the gates. The fluorescence intensity of the untreated cells (top right) has a single peak while the treated cells (middle right) have 2 peaks. A shift in actin intensity is easily seen in the overlay of histograms (bottom left). The median intensities of the untreated samples are set to 100%, and treated samples median intensities are normalized to the untreated controls.

  3. Histograms
    Histograms will automatically populate in FlowJo with “count” (the total number of cells under that gate) on the y-axis and arbitrary units of intensity in log scale on the x-axis. We normalize data on the y-axis to unit area and not to the number of cells (Figure 2) since the number of cells within a gate can vary by sample. FlowJo defines unit area as the maximum area equalling 1 unit with fluorescence intensity as a percentage of that area. FlowJo histograms place each cell (“event”) in one of 265 bins based on its intensity. After analyzing all samples, compare relative actin intensities by layering the histograms in the graphing layout window of FlowJo (Figure 2).

  4. Statistical analysis
    We use FlowJo to calculate the median fluorescence intensity of each cell sample. We normalize data to compensate for variation in fluorescence intensity between replicates due to drift in the lasers and detector (see Note 8). The untreated samples are set equal to 100%, and the treated samples are relative to the untreated samples (Table 1). We use Prism to draw scatterplots showing means and standard deviations (Figure 2).

    Table 1. Data Normalization

Notes

  1. Fixative: There are a number of fixatives commonly used to preserve the cytoskeleton, including methanol, glutaraldehyde, and paraformaldehyde. For this analysis, we use paraformaldehyde because methanol destroys the phalloidin binding site and, although glutaraldehyde is the fixative of choice for maintaining cell morphology it results in high autofluorescence that is difficult to quench when cells are in suspension.
  2. Permeabilization Buffer: Triton X-100 is a mild detergent that is commonly used to permeabilize cell membranes. We make a fresh dilution on the day of the experiment. Triton is viscous and can be difficult to pipet. We recommend using wide bore tips or cutting off the end of a regular pipette tip using a clean razor blade.
  3. Fluorescently labeled phalloidin: There are many fluorescently conjugated phalloidins that are commercially available. Choose one with a fluorophore that is compatible with both your flow cytometer and microscope filters. We typically use AlexaFluor® 488 conjugated phalloidin (Invitrogen), as it is bright and relatively photo-stable. Although the manufacturer recommends dissolving the phalloidin in methanol, we routinely resuspend the lyophilized phalloidin in DMSO, to a final concentration of 66 mM.
  4. Staining solution: Prepare the staining solution fresh and keep it cold until use, then allow it to come to room temperature before staining. The fluorescent dye is photosensitive so keep this out of direct light and/or cover in foil until use. You may need to optimize and empirically test concentrations of phalloidin that work best for your cell type.
  5. Storing samples: Although fixed, the actin cytoskeleton of cells is labile for two reasons: 1) the actin cytoskeleton is inherently dynamic. Because phalloidin binds multiple adjacent actin monomers, it serves to stabilize actin polymers, and drastically increase their lifetimes, even in fixed cells. It is best to stain your fixed samples as soon as possible. 2) Paraformaldehyde is a reversible fixative, and samples will not persist indefinitely. It is best to fix, stain, and analyze your samples on the same day. If you want to store samples for any length of time, we recommend testing this approach by imaging the phalloidin stained samples before and after storage.
  6. Cell loss: Statistical analysis of flow cytometry data requires a minimum of 10,000 recorded events (or cells) per sample. Some cell loss is unavoidable, but significant cell loss can occur at several points in this experiment. This can be avoided by:
    1. Carefully removing supernatant from tubes: It is crucial to leave a small volume around the pelleted cells to avoid disturbing the pellet and losing your sample. Do this by holding the tube at an angle with the pellet towards the ground and pipetting the supernatant off slowly and from the opposite side of the tube. If successful, the pellet should look unchanged after each step (before permeabilization). You should empirically test at which speeds and how long it takes for your cells to form a pellet without compromising cell morphology.
    2. Testing centrifugation speeds: Permeabilization creates holes in the cells’ membranes changing the density of the cell relative to the buffer. For most eukaryotic cells, centrifuging for 5 min at 1,000-2,000 x g is a fairly safe starting point. We recommend increasing centrifugation to get the best pellet without compromising overall cell morphology. Again, this should be empirically tested. Check samples on a microscope after each spin (e.g., 2 µl on a slide). If there is an increase in cell debris, you may be centrifuging too fast.
    3. Allowing sufficient cells for adjusting FSC/SSC laser intensities: Adjusting parameters can take time and requires samples to run during these adjustments. To avoid running out, prepare at least 3 times the volume needed for your controls careful not to dilute control samples as they should all roughly have the same cell densities.
    We recommend in the first trial experiment that you check the initial density of cells before fixing and record the cell density as well as the final density; you can use these densities to calculate the percent loss. Modify your starting cell densities according to the percent loss to guarantee you have enough cells to analyze. If cell loss is an issue, consider recording cell density after each centrifugation step to determine where you are losing cells.
  7. Cell clumping: After permeabilization, cells are generally stickier, and easily form clumps that are difficult to break apart. This is problematic for cytometry for several reasons: 1) Clumps clog flow cytometers, and therefore you must filter your samples before acquisition, greatly decreasing the number of cells in your sample. 2) Small cell clumps can be measured together as one larger cell with the combined intensity and a false population. To break up cell clumps gently mix by pipetting the full volume up and down and/or gently vortex before transferring samples to FACS tubes. We recommend checking a sample for clumps on a microscope (e.g., 2 µl on a slide).
  8. Comparing samples prepared/analyzed on different days: It is critical to process and analyze all samples of each biological replicate in parallel due to variation in phalloidin staining and fluorescence detection by cytometry. Measurements from samples within a replicate stained or analyzed on different days cannot be meaningfully compared.
  9. Gating for single cells: To ensure that only single cells are included in the analysis, the data may be gated in several ways to exclude doublets (described in detail in the FlowJo tutorials available from: https://www.flowjo.com/learn/flowjo-university/flowjo/getting-started-with-flowjo/58). We did not do this gating on the data set shown here because our cytometer does not have the capacity to detect area or width. Depending on the capabilities of the flow cytometer, data can be gated by any one of the following three methods:
    1. FSC-H vs. FSC-A: Display the data on a scatterplot with FSC-height (FSC-H) on the y-axis and FSC-area (FSC-A) on the x-axis. Select the polygon tool to establish the first gate around the population along the diagonal line. This gate selects “singlets” or single cells of similar size compared to total cell area and removes cell clumps or “doublets” from the data set.
    2. FSC-W vs. FSC-H: Display the data with FSC-W on the y-axis and FSC-H on the x-axis. Gate the population to exclude data points with FSC-W at higher values than the main population. 
    3. SSC-W vs. SSC-H: Display the data with SSC-W on the y-axis and SSC-H on the x-axis. Gate the population to exclude data points with SSC-W at higher values than the main population. 

Recipes

Note: All recipes below are made at room temperature and aseptically with Milli-Q water.

  1. Culture medium for HL-60 cells
    RPMI supplemented with 10% Fetal Bovine Serum
  2. Fixation Buffer (4% PFA; For 400 µl)
    100 µl of 16% PFA
    300 µl cytoskeleton buffer
    Does not need to be sterilized
    Shelf-life: < 24 h kept at 4 °C
    Note: PFA degrades over time, so a fresh ampule is best. We routinely use 16% Paraformaldehyde sold in 10 ml vials (Electron Microscopy Sciences). Once opened, we typically will use an open ampule for up to two weeks stored at 4 °C.
  3. Cytoskeleton Buffer
    10 mM MES, pH 6.1
    138 mM KCl
    3 mM MgCl2
    2 mM EGTA
    Sterile filter and store at 4 °C and supplement with sucrose on the day of use to a final concentration of 320 mM
  4. Permeabilization Buffer
    0.1% Triton X-100 in 1x PBS
    Does not need to be sterilized
    Prepare fresh on the day of the experiment
    Store at 20 °C
  5. Staining Solution
    66 nM AlexaFluor® 488 phalloidin diluted in 1x PBS supplemented with 2% BSA
    Does not need to be sterilized
    Prepare Fresh
  6. PBS (10x)
    1.37 M NaCl
    27 mM KCl
    100 mM Na2HPO4
    18 mM KH2PO4
    Adjust pH to 7.4
    Autoclave to sterilize
  7. Wash Solution
    1x PBS supplemented with 0.1% TWEEN-20
    Make fresh, and does not need to be sterilized

Acknowledgments

This protocol was adapted from previously published work (Fritz-Laylin et al., 2017). We would like to thank Amy Burnside at the University of Massachusetts Amherst Flow Cytometer (FACS) Core Facility for assistance with data analysis. We are funded by the National Institutes of Health (NIH NIAID) Grant 1R21AI139363.

Competing interests

We have no conflict of interest or competing interest to declare.

References

  1. Fritz-Laylin, L. K., Lord, S. J. and Mullins, R. D. (2017). WASP and SCAR are evolutionarily conserved in actin-filled pseudopod-based motility. J Cell Biol 216(6): 1673-1688.
  2. Vandekerckhove, J., Deboben, A., Nassal, M. and Wieland, T. (1985). The phalloidin binding site of F-actin. EMBO J 4(11): 2815-2818.

简介

由于天然过程和/或实验诱导的扰动,细胞内聚合的肌动蛋白的量可以广泛变化。 我们常规使用该方案通过用荧光鬼笔环肽染色悬浮细胞,然后使用流式细胞术测量总细胞荧光来测量细胞群之间的相对聚合肌动蛋白含量。 虽然是为人类细胞开发的,但我们很容易将这种方法用于各种真核细胞类型。

【背景】生物学家长期以来一直使用生化分析和荧光显微镜研究肌动蛋白聚合物。后者通过鬼笔环肽的发现而得到促进,鬼笔环肽是一种由有毒蘑菇产生的小分子毒素,通过特异性结合肌动蛋白聚合物内的相邻单体来稳定肌动蛋白结构(Vandekerckhove et al。,1985)。荧光鬼笔环肽缀合物染色复杂的肌动蛋白结构并通过荧光显微镜促进详细的可视化。然而,定量这种荧光可能是艰苦的,特别是在追踪人口水平的变化时。相反,流式细胞术定量每分钟数千个细胞的总荧光强度。

在这里,我们描述我们如何通过用荧光鬼笔环肽染色细胞,使用流式细胞术测量单个细胞的总细胞荧光,并最终比较群体的荧光值来估计细胞群之间的肌动蛋白聚合物量的差异。重要的是要注意,这种方法是比较性的,不能用于测量细胞肌动蛋白聚合物的绝对浓度。这种方法要求细胞处于悬浮状态;如果您的细胞需要粘附以进行适当的肌动蛋白聚合,我们建议使用如上所述的定量荧光显微镜。

尽管快速和定量,但流式细胞术未提供关于细胞形态的信息。因此,我们通常使用荧光显微镜保存等份的染色细胞用于可视化。我们强烈建议这样做,以确保您感兴趣的结构得到妥善保存(图1)。荧光显微镜还可以揭示流式细胞术不明显的实验样品的变化。

该方案针对分化的HL-60细胞进行了优化,这些细胞可以用fMLP刺激以诱导快速肌动蛋白聚合(图1,关于这些细胞的其他信息,参见Fritz-Laylin et al。,2017),通过测试各种固定条件和缓冲液,可以对其进行修改以用于各种细胞类型。例如,我们已经针对壶菌真菌( Batrachochytrium dendrobatidis )和变形虫 Naegleria gruberi (未发表)修改了该方案。为此,我们选择了以下条件:1)最好地保留与使用DIC或相差显微镜可视化的活细胞相似的整体形态; 2)通过固定保持聚合的肌动蛋白完整; 3)当通过荧光显微镜成像时,染色适当的结构。有关常用于保存肌动蛋白结构的缓冲液和固定剂(参见注释1)的信息,我们建议使用方法resource 固定细胞中肌动蛋白和微管蛋白细胞骨架的荧光程序 来自哈佛大学米奇森实验室(L. Cramer和A. Desai)。


图1.响应细胞刺激的肌动蛋白聚合的一个例子。用20nM fMLP处理之前和之后AlexaFluor® 488鬼笔环肽染色的HL-60细胞的最大投射,其中诱导肌动蛋白聚合并产生更亮的细胞。

关键字:肌动蛋白, F-肌动蛋白, 定量, 聚合物, 细胞, 细胞计数法, FACS

材料和试剂

  1. 标准无涂层1.5 ml微量离心管(USA Scientific,目录号:1615-5510)
  2. 15和50毫升锥形管(任何供应商)
  3. FACS管:无菌5 ml聚苯乙烯圆底管,带有细胞过滤器盖,用于在流式细胞仪(BD Falcon,目录号:352235)之前过滤样品,也可使用外部过滤器
    注意:检查您的细胞计数器要求和/或您的流式细胞仪设施对管的建议。
  4. 微量移液器吸头(Pipette.com,目录号:UE-1000,UE-200,UE-10)
  5. 铝箔
  6. 剃刀片
  7. 分化的HL-60细胞(在补充有10%胎牛血清的中等RPMI中生长,并用1.3%DMSO处理5天;有关其他细节,参见Fritz-Laylin et al。,2017),或您希望定量聚合肌动蛋白相对量的任何其他细胞类型&nbsp;
  8. RPMI(Gibco,目录号:11875)
  9. AlexaFluor® 488共轭鬼笔环素(Thermo Fisher Scientific,目录号:A12379)(见注3)
  10. 无菌Milli-Q水
  11. TWEEN-20(西格玛奥德里奇,目录号:P9416)
  12. Triton X-100(Sigma-Aldrich,目录号:X100)
  13. MES(西格玛奥德里奇,目录号:M3671)
  14. KCl(Fisher Scientific,目录号:P217-100)
  15. MgCl 2(Sigma-Aldrich,目录号:M9272)
  16. EGTA(西格玛奥德里奇,目录号:E4378)
  17. NaCl(Fisher Scientific,目录号:BP3581)
  18. Na 2 HPO 4(Sigma-Aldrich,目录号:795410)
  19. KH 2 PO 4(Sigma-Aldrich,目录号:P0662)
  20. BSA(Fisher Scientific,目录号:BP1600-100)
  21. 16%多聚甲醛(电子显微镜科学,目录号:15710)
  22. DMSO(Sigma-Aldrich,目录号:D2650)
  23. fMLP(Sigma-Aldrich,目录号:47729)
  24. HL-60细胞培养基(参见食谱)
  25. 固定缓冲液(见食谱)
  26. 细胞骨架缓冲液(见食谱)
  27. 透化缓冲液(见食谱)
  28. 染色液(见食谱)
  29. PBS(10x)(见食谱)
  30. 洗涤液(见食谱)

设备

  1. 移液器(Eppendorf,P1000,P100,P20,P1)
  2. 带转子的微量离心机适用于1.5 ml离心管(例如,Eppendorf,型号:5415)
  3. 流式细胞仪(例如,FACSCalibur分析仪[BD,型号:FACSCalibur TM]或LSRFortessa Dual [BD,型号:LSRFortessa TM])
  4. 配备荧光和透射光的显微镜(我们更喜欢相位和/或DIC)(例如,尼康,型号:Eclipse Ti-E)
  5. 用于培养细胞的培养箱,对于HL-60细胞,您需要一个连续37°C和5%CO 2的室
  6. 高压灭菌器
  7. 显微镜过滤器适用于您选择的鬼笔环肽结合物(见注3)
  8. 标准血细胞计数器(例如,Sigma-Aldrich,目录号:Z359629)
  9. 涡流混合器&NBSP;
  10. -20°C冰柜

软件

  1. FACSDiva,BD(Becton,Dickinson&amp; Company)
  2. FlowJo版本10,BD(Becton,Dickinson&amp; Company)
  3. GraphPad Prism版本7,GraphPad软件

程序

  1. 固定细胞
    1. 准备新鲜的固定缓冲液并保持在冰上直到需要为止。&nbsp;
    2. 标记每个样品的1.5ml管,包括将用鬼笔环肽染色的未处理和处理的细胞,以及将被相同处理但不会被鬼笔环肽染色的未匹配对照的匹配对照样品。
    3. 通过离心浓缩至所需密度来制备细胞,并在每个新鲜培养基或缓冲液样品中悬浮100μl; FACS分析通常需要的最小细胞数为每个样品10,000个(见注6)。为了刺激HL-60细胞,重悬于补充有2%BSA的无血清RPMI中,并在37℃和5%CO 2中孵育1小时以使细胞去极化。&nbsp;
    4. 通过加入10ul 200nM fMLP刺激细胞,轻轻敲打管混合,并在37℃下孵育3分钟(处理过的样品),然后立即固定(对于对照,未处理的样品不要这样做)。
    5. 通过向每个样品中加入400μl冷固定缓冲液,立即将细胞固定在1.5 ml离心管中。&nbsp;
    6. 轻轻地上下移液混合样品。&nbsp;
    7. 在冰上孵育20分钟(见注5)。

  2. 染色细胞
    1. 在4°C离心细胞以形成沉淀(参见注释6)。&nbsp;
    2. 去除上清液,将细胞重悬于100μl透化缓冲液中(见注2),上下移液轻轻混匀。&nbsp;
    3. 在室温下孵育10分钟。&nbsp;
    4. 在室温或4℃下离心沉淀细胞。
    5. 去除上清液并将细胞重悬于100μl染色溶液中(参见食谱和注释D)。&nbsp;
    6. 通过上下移液轻轻重悬细胞。&nbsp;
    7. 用铝箔覆盖样品并在室温下孵育20分钟,然后转移至4℃。样品可以存储在鬼笔环肽中的时间长短可以根据细胞类型而变化。使用新的细胞类型时,根据经验进行测试(参见注释5)。&nbsp;
    8. 当准备通过流式细胞术分析细胞时,离心形成沉淀并除去鬼笔环肽上清液。&nbsp;
    9. 通过用500μl补充有0.1%TWEEN-20(用于破碎细胞并避免结块的温和洗涤剂)的PBS轻轻重悬细胞数次洗涤,并在4℃下离心以形成沉淀。最后一次洗涤后,重悬于至少500μl预冷的PBS中,并将样品储存在冰上。&nbsp;
    10. 在流式细胞术之前过滤样品(见注7)。
    11. 通过荧光显微镜检查染色的样品,以确保目标结构被染色,并且整体细胞形态没有受到损害(细胞应该看起来与它们活着时的外观大致相似)。
    12. 为确保有足够的细胞进行实验,取一等份染色细胞并用血细胞计数器进行计数。

  3. 流式细胞仪
    这不是流式细胞仪的完整指南,我们建议不熟悉该技术的读者可以看到:流式细胞术基础指南,FlowJo视频教程,如 Cytometry 101 ( https://www.flowjo.com/learn/flowjo-university / flowjo / getting-started-with-flowjo / 2 )或BD Biosciences流式细胞仪网络培训简介( https://www.bdbiosciences.com/us/support/s/itf_launch )。对于此处提供的HL-60细胞数据,我们使用了FACSCalibur分析仪(BD)。

    1. 设置参数
      对于此分析,我们测量FSC高度,SSC高度和FIT-C:高度,对数。如果细胞计数器具有这些能力,我们还建议收集FSC宽度,SSC宽度和/或FSC区域,因为这些值将允许消除双峰(见注9)。样品荧光的差异很大,因此我们以对数标度显示荧光强度。
      &NBSP; FACSDiva软件显示工作表,将数据填充到直方图,散点图中,并根据门控提供人口统计数据。我们建议设置一个全局工作表,以便在调整和获取期间可视化数据。
      &NBSP;根据您的细胞计数器,调整激光强度和/或检测器1)捕获正向和侧向散射(以消除死细胞,团块,双峰,等参见注释9以进行双重消除)和2 )确保样品在可检测范围内发荧光。当您的对照样品以低速通过流式细胞仪时,调整这些参数。通常,具有前向散射和侧向散射的散点图将在细胞计数器收集数据时实时显示群体。目标人群应与轴完全分开,以避免排除数据。使用选择工具,通过设置整个群体周围的点来创建初始门,并调整细胞计数器以使群体中整个细胞的荧光强度居中。这样做非常小心,因为以后无法恢复探测器范围之外的数据。
    2. 数据采集
      一旦激光器和检测器得到适当调整,对复制中所有样品收集的所有数据使用相同的激光强度。可以(并且应该)针对每个实验重复调整细胞计数器,但是一旦设置用于实验,使用相同的细胞计数器设置分析所有样品(参见注释8)。如果你有一个未染色的样本,由于死细胞自发荧光,低水平的背景荧光是正常的。

数据分析

我们使用FlowJo软件(Tree Star)分析流式细胞仪数据。在这里,我们简要介绍一下我们的分析工有关细胞计数数据分析的更详细讨论,请参阅BD FACSDiva的数据分析教程( https: //www.bdbiosciences.com/us/support/s/itf_launch )或FlowJo( https://www.flowjo.com/learn/flowjo-university/flowjo )。


  1. 团体
    将文件导入FlowJo应根据文件名自动填充组。如果您需要创建用于分析的组,可以使用工具根据文件名($ fil)中的关键字和其他参数选择组。

  2. 门控
    将闸门均匀地应用于整个复制中的所有样品。这里描述的门1)选择完整的细胞并排除细胞碎片; 2)建立人口; 3)允许样品之间的比较分析(见注8)。
    1. 打开图形窗口并创建一个散点图,其中y轴为SSC高度(SSC-H),x轴为FSC高度(FSC-H)。我们喜欢使用伪彩色选项作为散点图,因为单色点图减少了仅表示某些数据的事件数。此外,伪彩色很容易可视化事件的浓度以建立门控。
    2. 如果细胞计数器具有收集宽度和/或面积数据的能力,则对细胞群进行门控以消除双峰(参见注释9)。
    3. 选择一个细胞群,小心不要使用椭圆工具从主细胞群中排除数据(图2)。&nbsp;
    4. 打开这个门控人口作为直方图。将x轴显示为“FIT-C”,将y轴显示为直方图。
    5. 添加中位数等统计信息。
    6. 将分析复制到整个组。这创建了一个组门控层次结构,以便分析该组内的相对肌动蛋白强度。


      图2.流式细胞术和肌动蛋白聚合分析。 作为本协议中概述的实验的一个例子,我们分析了未处理和刺激的HL-60细胞,如图1所示。为了排除细胞碎片和/或团块,首先在浓缩物周围设置前向和侧向散射门。细胞群(以伪彩色显示)并均匀地应用于所有样品。注意:当与DMSO分化时,并非所有细胞都完全分化,导致这里看到两个细胞群。直方图显示了门内细胞的相对强度。未处理细胞的荧光强度(右上)具有单峰,而处理细胞(右中)具有2个峰。在直方图的叠加中容易看到肌动蛋白强度的变化(左下)。将未处理的样品的中值强度设定为100%,并将处理的样品中值强度标准化为未处理的对照。

  3. 直方图
    直方图将在FlowJo中自动填充y轴上的“count”(该门下的单元格总数)和x轴上对数刻度的任意强度单位。我们将y轴上的数据标准化为单位面积而不是单元数量(图2),因为门内的单元数量可能因样本而异。 FlowJo将单位面积定义为最大面积等于1个单位,荧光强度占该面积的百分比。 FlowJo直方图根据其强度将每个单元格(“事件”)放置在265个区域之一中。在分析了所有样本后,通过在FlowJo的图形布局窗口中对直方图进行分层来比较相对肌动蛋白强度(图2)。

  4. 统计分析
    我们使用FlowJo计算每个细胞样本的中值荧光强度。我们将数据标准化以补偿由于激光和检测器中的漂移引起的重复之间荧光强度的变化(见注8)。未处理的样品设定为等于100%,处理的样品相对于未处理的样品(表1)。我们使用Prism绘制散点图,显示平均值和标准偏差(图2)。

    表1.数据规范化

笔记

  1. 固定剂:通常使用许多固定剂来保存细胞骨架,包括甲醇,戊二醛和多聚甲醛。对于该分析,我们使用多聚甲醛,因为甲醇破坏了鬼笔环肽结合位点,尽管戊二醛是维持细胞形态的固定剂,但它导致高自发荧光,当细胞处于悬浮状态时难以猝灭。
  2. 透化缓冲液 Triton X-100是一种温和的清洁剂,通常用于透过细胞膜。我们在实验当天进行了新鲜稀释。 Triton粘稠,很难移液。我们建议使用大口径尖端或使用干净的剃须刀片切断常规移液器吸头的末端。
  3. 荧光标记的鬼笔环肽:有许多可商购的荧光缀合的鬼笔环肽。选择一个荧光团,与您的流式细胞仪和显微镜过滤器兼容。我们通常使用AlexaFluor® 488缀合的鬼笔环肽(Invitrogen),因为它是明亮且相对光稳定的。尽管制造商建议将鬼笔环肽溶解在甲醇中,但我们通常将冻干的鬼笔环肽重新悬浮于DMSO中,最终浓度为66mM。
  4. 染色溶液:将染色溶液新鲜制备并保持冷却直至使用,然后在染色前使其达到室温。荧光染料是光敏的,因此请将其保持在直射光下和/或覆盖在箔中直至使用。您可能需要优化并凭经验测试最适合您细胞类型的鬼笔环肽浓度。
  5. 存储样本虽然固定,细胞的肌动蛋白细胞骨架是不稳定的,原因有两个:1)肌动蛋白细胞骨架本身就是动态的。因为鬼笔环肽结合多个相邻的肌动蛋白单体,它可以稳定肌动蛋白聚合物,并且即使在固定细胞中也可以显着延长它们的寿命。最好尽快染色固定样品。 2)多聚甲醛是一种可逆的固定剂,样品不会无限期地存在。最好在同一天对样品进行修复,染色和分析。如果您想存放样品任何时间长度,我们建议您在储存前后对鬼笔环肽染色的样品进行成像测试。
  6. 细胞丢失:流式细胞仪数据的统计分析要求每个样本至少记录10,000个事件(或细胞)。一些细胞损失是不可避免的,但在该实验中的几个点可能发生显着的细胞损失。这可以通过以下方式避免:
    1. 小心地从管中去除上清液:在颗粒细胞周围留下一小块体积是至关重要的,以避免干扰颗粒并丢失样品。通过将管子与颗粒保持一定角度朝向地面并且从管子的相对侧缓慢地移出上清液来这样做。如果成功,每个步骤后(透化前)颗粒应该看起来没有变化。您应该凭经验测试细胞形成颗粒所需的速度和时间,而不会影响细胞形态。
    2. 测试离心速度:渗透性在细胞膜上产生孔,改变细胞相对于缓冲液的密度。对于大多数真核细胞,在1,000-2,000 x g 下离心5分钟是一个相当安全的起点。我们建议增加离心以获得最佳颗粒,而不会影响整体细胞形态。同样,这应该是经验测试。每次旋转后在显微镜上检查样品(例如,载玻片上2μl)。如果细胞碎片增加,您可能离心过快。
    3. 允许足够的单元调整FSC / SSC激光强度:调整参数可能需要一些时间,并且需要在这些调整期间运行样本。为了避免耗尽,准备至少3倍的对照所需的体积小心不要稀释对照样品,因为它们应该大致具有相同的细胞密度。
    我们建议在第一次试验实验中检查固定前细胞的初始密度,并记录细胞密度和最终密度;您可以使用这些密度来计算损失百分比。根据损失百分比修改起始细胞密度,以确保您有足够的细胞进行分析。如果细胞丢失是一个问题,请考虑在每次离心步骤后记录细胞密度,以确定丢失细胞的位置。
  7. 细胞聚集:透化后,细胞通常更粘,容易形成难以分解的团块。这对于细胞计数是有问题的,原因如下:1)Clumps阻塞流式细胞仪,因此您必须在采集前过滤样品,大大减少样品中的细胞数量。 2)小细胞团块可以作为一个较大的细胞一起测量,具有组合的强度和错误的群体。在将样品转移到FACS管之前,通过向上和向下移液全部体积和/或轻轻涡旋来轻轻地混合细胞团。我们建议在显微镜上检查样品是否为团块(例如,载玻片上为2μl)。
  8. 比较在不同日期制备/分析的样品:由于鬼笔环肽染色的变化和细胞计数的荧光检测,并行处理和分析每个生物复制品的所有样品是至关重要的。在不同日期染色或分析的重复样品中的测量值不能进行有意义的比较。
  9. 单个细胞的门控:为了确保分析中只包含单个细胞,可以通过多种方式对数据进行门控以排除双峰(详见FlowJo教程中的以下内容: https://www.flowjo.com/learn/flowjo-university /的FlowJo /工具入门与 - 的FlowJo / 58 )。我们没有对这里显示的数据集进行门控,因为我们的细胞计数器没有检测区域或宽度的能力。根据流式细胞仪的功能,可以通过以下三种方法中的任何一种来控制数据:
    1. SSC-W与SSC-H:在y轴上显示SSC-W,在x轴上显示SSC-H。对人口进行门控以排除SSC-W的数据点,其数值高于主要人口。&nbsp;

食谱

注意:以下所有配方均在室温下进行,无菌使用Milli-Q水。

  1. HL-60细胞培养基
    RPMI补充10%胎牛血清
  2. 固定缓冲液(4%PFA;400μl)
    100μl16%PFA
    300μl细胞骨架缓冲液
    不需要消毒
    保质期:&lt; 24小时保持在4°C
    注意:PFA会随着时间的推移而降解,因此最好使用新的安瓿。我们通常使用在10ml小瓶中出售的16%多聚甲醛(Electron Microscopy Sciences)。一旦打开,我们通常会在4°C下使用开放的安瓿长达两周。
  3. 细胞骨架缓冲液
    10 mM MES,pH 6.1
    138 mM KCl
    3mM MgCl 2
    2 mM EGTA
    无菌过滤并在4°C下储存,并在使用当天补充蔗糖至终浓度为320 mM
  4. 透化缓冲液
    在1x PBS中0.1%Triton X-100
    不需要消毒
    在实验当天准备新鲜的
    储存在20°C
  5. 染色液
    66 nM AlexaFluor® 488鬼笔环肽在1x PBS中稀释,补充2%BSA
    不需要消毒
    准备新鲜
  6. PBS(10x)
    1.37 M NaCl
    27 mM KCl
    100mM Na 2 HPO 4
    18mM KH 2 PO 4
    将pH调节至7.4
    高压灭菌消毒
  7. 洗涤液
    1x PBS补充0.1%TWEEN-20
    新鲜,不需要消毒

致谢

该协议改编自先前发表的工作(Fritz-Laylin et al。,2017)。我们要感谢马萨诸塞大学阿默斯特流式细胞仪(FACS)核心设施的Amy Burnside对数据分析的帮助。我们由美国国立卫生研究院(NIH NIAID)Grant 1R21AI139363资助。

利益争夺这可以通过以下方式避免

我们没有利益冲突或竞争利益申报。

参考

  1. Fritz-Laylin,L。K.,Lord,S。J.和Mullins,R。D.(2017)。 WASP和SCAR在肌动蛋白填充的基于伪足的运动中具有进化保守性。 J Cell Biol 216(6):1673-1688。
  2. Vandekerckhove,J.,Deboben,A.,Nassal,M。和Wieland,T。(1985)。 F-actin的鬼笔环肽结合位点。 EMBO J 4(11):2815-2818。
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Copyright: © 2018 The Authors; exclusive licensee Bio-protocol LLC.
引用: Readers should cite both the Bio-protocol article and the original research article where this protocol was used:
  1. Kakley, M. R., Velle, K. B. and Fritz-Laylin, L. K. (2018). Relative Quantitation of Polymerized Actin in Suspension Cells by Flow Cytometry. Bio-protocol 8(22): e3094. DOI: 10.21769/BioProtoc.3094.
  2. Fritz-Laylin, L. K., Lord, S. J. and Mullins, R. D. (2017). WASP and SCAR are evolutionarily conserved in actin-filled pseudopod-based motility. J Cell Biol 216(6): 1673-1688.
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