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Jun 2019

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Quantitative Kinetic Analyses of Histone Turnover Using Imaging and Flow Cytometry
组蛋白转换的定量动力学成像及流式细胞术研究   

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Abstract

Dynamic histone changes occur as a central part of chromatin regulation. Deposition of histone variants and post-translational modifications of histones are strongly associated with properties of chromatin status. Characterizing the kinetics of histone variants allows important insights into transcription regulation, chromatin maintenance and other chromatin properties. Here we provide a protocol of quantitative and sensitive approaches to test the timing of incorporation and dissociation of histones using a two-color SNAP-labeling system, labelling pre-existing and newly-incorporated histones distinctly. Together with cell cycle synchronization methods and cell cycle markers, this approach enables a pulse-chase analysis to determine the turnover of histone variants during the cell cycle, detected using imaging or flow cytometry methods at single cell resolution. As well as testing global histone turnover, cell cycle-dependent cellular localization of histone variants can be also addressed using imaging approaches.

Keywords: Chromatin dynamics (染色质动力学), Histone (组蛋白), Cell cycle (细胞周期), Imaging (成像), Flow Cytometry (流式细胞仪)

Background

Chromatin remodeling is part of the numerous fundamental cellular activities in eukaryotic cells (Geiman and Robertson, 2002; Clapier and Cairns, 2009). Accessibility by transcription factors and RNA polymerases are generally associated with changes in DNA methylation and chromatin states, including accessibility, post-translational histone modifications and deposition of histone variants. Histone variants differentially coordinate the gene expression that regulates development, cell differentiation or other physiological activities (Banaszynski et al., 2010). They also play diverse roles in DNA repair, telomere maintenance, heterochromatin formation and chromatin segregation (Henikoff and Smith, 2015; Zink and Hake, 2016). Moreover, dysregulation of a histone variant’s incorporation is associated with cancer (Vardabasso et al., 2014), indicating a significant role in human disease. To understand the kinetics of histone variants, including incorporation into and dissociation from specific chromatin regions during the cell cycle, is crucial since it tightly links regulatory properties with the mitotic maintenance of epigenetic (heritability from parent to daughter cells). DNA replication involves major chromatin remodeling to duplicate the entire chromatin structure following mitosis. Histones that associate with chromatin prior to DNA replication are transiently dissociated from DNA by the access of the DNA polymerase complex. Release of pre-existing histones randomly re-associate at newly synthesizing replication forks together with newly synthesized histones (Balhorn et al., 1975; Alabert and Groth, 2012; Annunziato, 2012). Pre-existing post-translational modifications on histones and some histone variants also re-associate on the newly synthesized DNA at the replication fork, explaining re-association of pre-existing histone at the replication fork is one part of the maintenance of mitotic inheritance of chromatin states. To determine the timing of post-translational modifications or incorporation of histone variants, a sensitive pulse-chase system that can distinguish the detection of newly incorporated histone from pre-existing histones is required. Newly synthesized canonical histones are unmodified when they are incorporated into chromatin during DNA replication. Unlike de novo DNA methylation, which occurs together with DNA replication (Tillo et al., 2016), most histone marks are not established on newly incorporated histones on the replicating fork. Proteomic based pulse-chase approaches (see in Alternative methods section below) have been used to determine the global kinetics of histone post-translational modifications by detecting the pre-existing and new deposition of histone acetylation and methylation at lysine residues (Pesavento et al., 2008; Scharf et al., 2009; Martinez-Garcia et al., 2011; Xu et al., 2011; Zee et al., 2012; Alabert et al., 2015). This approach identified two distinctive kinetic patterns of histone modifications. One group, such as histone H3 acetyl-lysine at 27 (H3 K27 ac), exhibits rapid turnover to equalize, and notably they are not maintained through the cell cycle (Scharf et al., 2009). This rapid acetylation kinetics probably represent temporally active transcription dynamics (Stasevich et al., 2014). While another group including H3 lysine tri-methylation at 9 (K9me3) or H3 K27me3 is acquired more slowly and step-wise from mono and di to tri-methylation, established during G1 phase following the cell cycle instead of before mitosis (Pesavento et al., 2008; Scharf et al., 2009; Martinez-Garcia et al., 2011; Xu et al., 2011; Zee et al., 2012; Alabert et al., 2015). Importantly, this group has the property of mitotic chromosome memory. The potential mechanism of maintenance of histone marks over cell division has been addressed by imaging approaches. In situ proximity ligation assays using specific antibodies against histone lysine methylations and their methyltransferases detected that histone modifiers continuously associate with the replicating DNA component in Drosophila embryos, suggesting the association of modifiers on replicating DNA may provide a “tag” to be methylated (Petruk et al., 2012).

Recent advances in chemical protein labeling technologies provide us with a powerful tool for protein tagging applications in living cells (e.g., SNAP (New England Biolabs), CLIP (New England Biolabs), Halo (Promega) and TMP (Active Motif) tag). These technologies are based on the covalent labeling of genetically encoded tags that bind with specific ligands conjugated to cell permeable substrates such as synthetic fluorescent dyes or biotin, which can mediate affinity purification in biochemical applications. In contrast with common genetically encoded tags, this chemical labeling of protein can be utilized in timing-dependent labeling with many choices of fluorophores, which allows the pulse-chase labeling of specific protein. This labeling technology has revealed the deposition timing of histone variants at specific chromatin architectures using imaging detection (e.g., CENP-A at centromeres [Dunleavy et al., 2009] and macroH2A at heterochromatin [Sato et al., 2019]).

Here, we provide a detailed protocol of a pulse-chase method using the SNAP-tag labeling system which has utilized quantitative histone variant detection with single cell resolution. Using this protocol, distinct histone kinetics, dissociation of pre-existing histones and association of newly synthesized histones, can be detected simultaneously. We describe two detection approaches, fluorescence microscopy and flow cytometry, as well as the detail of imaging analysis using FIJI/ImageJ software which is freely available (https://fiji.sc/).

Advantages and Limitations
The pulse-chase method using a chemical protein labeling system is an easy, non-hazardous and sensitive approach compared with a conventional pulse-chase approach using radioactive molecules (see in Alternative methods section). Most of the required reagents and fluorophores are commercially available. Global turnover of histones can be addressed using imaging and flow cytometric applications and notably, timing-specific localization at specific chromatin architectures also can be characterized with imaging approaches. Using this protocol, both pre-existing and newly incorporated histone variants can be detected simultaneously in the same cells with single cell resolution. Unlike radioisotope or metabolic labeling approaches, which detect endogenous histones, this labeling approach relies on the genetically encoded tags (e.g., SNAP-, CLIP-, Halo-tag) that can be linked with specific substrates. Therefore, a plasmid construct that expresses the target histones with SNAP-tag and its use in a stably expressing cell line are required. In addition, the localization and other biological functions of desired tagging histones must be examined to determine whether it remains functioning as an endogenous histone. Optimization of the construct (e.g., N-terminus or C-terminus tagging, changing the choice of promoter) and levels of expression might be necessary for obtaining accurate observations. Another limitation of this approach is that it is not applicable for detection of post-translational modifications of histones.

Alternative methods
Isotope labelling with proteomic detection
SILAC (Stable Isotope Labelling with Amino acids in Culture) followed by mass spectrometry is a powerful approach to investigate global turnover of endogenous histone variants and post-translational modifications (Yuan et al., 2014). In this approach, newly synthesized histones are labeled with radioactive heavy isotope and chase the turnover of labeled his tones compared with pre-existing histones containing light amino acids. Following mass spectrometry analysis determines the pre-existing and deposition of post-translational modifications or variants. This approach is suitable to detect global histone turnover, but is not able to detect histone marks at specific chromatin architecture or genomic loci.

Metabolic labeling with genome-wide approaches
Non-radioactive metabolic labeling of nascent proteins can be an alternative approach to label global newly synthesized histones. The approach, “Covalent Attachment of Tags to Capture Histones and Identify Turnover”, also called ‘CATCH-IT’, enables genome-wide investigation to characterize active histone replacement (Deal et al., 2010). This approach is based on the labeling scheme of nascent peptide by incorporation of methionine homolog, azidohomoalanine (Aha), which is generally used for the detection of active translation in cells. In this approach, the nucleosomes containing Aha-labeled newly synthesized histones are bioconjugated with biotin by a cycloaddition reaction (as known as “click” chemistry), and pulled down with streptavidin beads. Isolated DNA from pull-down was applied on a tiling microarray to determine the genomic loci that exhibit active histone replacement in Drosophila S2 cells. The characterized genomic sites that have active histone turnover correspond with the site of incorporation of histone H3 variant, H3.3, which is detected at transcriptionally active loci (Henikoff et al., 2009). This approach enables the detection of genomic loci with active turnover of histones.

Chemical labeling approaches such as the SNAP-tagging system can also be the alternative option to investigate genome-wide histone variant incorporation (Sato et al., 2019). In this approach, newly incorporated SNAP-tagged histones are linked with SNAP-biotin after the treatment of SNAP-Cell® Block (bromothenylpteridine, BTP), a non-fluorescent substrate to mask the reactivity of pre-existing histones. Then, biotin-linked, newly incorporated histones can be pulled down with streptavidin beads. The purified DNA fraction from pull-down samples can be sequenced with massive parallel sequencing. This approach might be useful to detect timing and genomic loci dependent incorporation of histone variants but unable to detect post-translational modifications.

Materials and Reagents

  1. 50-ml Falcon tubes
  2. 15-ml Falcon tubes
  3. 1.5-ml tubes
  4. Tissue culture plates (12 or 24-well; Falcon, catalog number: 353043 or 353047 )
  5. Tissue culture dishes (35 x 10 mm or 60 x 15 mm, and 100 x 20 mm, Falcon, catalog numbers: 353001 or 353002 and 353003 )
  6. Coverslips (18 or 12 mm No.1.5)
  7. Microscope slides (e.g., Fisher Scientific, catalog number: 12-544-2 )
  8. 5-ml FACS tubes (Fisher Scientific, catalog number: 149595 )
  9. Adherent cells of interest (e.g., mouse embryonic fibroblasts [MEFs], HEK293 cells)
  10. SNAP-Cell Oregon Green (New England BioLabs, catalog number: S9104S )
  11. SNAP-Cell Block (New England BioLabs, catalog number: S9106S )
  12. SNAP-Cell TMR-Star (New England BioLabs, catalog number: S9105S )
  13. Thymidine (Sigma, catalog number: T1895 )
  14. RO-3306 (Santa Cruz Biotechnology, catalog number: sc-358700 )
  15. Nocodazole (Sigma, catalog number: M1404 )
  16. Prolong Diamond Antifade Mountant (Thermo Fisher Scientific, catalog number: P36970 )
  17. Prolong Diamond Antifade Mountant with DAPI (Thermo Fisher Scientific, catalog number: P36962 )
  18. Drug for selection of stably expressing cells [e.g., 600-1,200 µg/ml G418 (geneticin), Zeocin (Thermo Fisher Scientific, catalog number: R25001 )]
  19. Click-iTTM EdU Cell Proliferation Kit for Imaging, Alexa FluorTM 647 dye (Thermo Fisher Scientific, catalog number: C10340
  20. pSNAPf Vector (New England BioLabs, catalog number: N9183S )
  21. pCCL-CellCycle (Sato et al., 2019)
  22. Suitable culture media and supplements [e.g., DMEM (Sigma, catalog number: D6429 ) supplemented with 10% FBS ( Atlanta Biologicals, Inc., catalog number: S11150H ) and 100 U/ml Penicillin-Streptomycin (Sigma, catalog number: 15140148 )]
  23. EDTA, 0.5 M, pH 8.0, Molecular Biology Grade (Sigma, catalog number: 324506 )
  24. HEPES solution, 1 M, pH 7.0-7.6, sterile-filtered (Sigma, catalog number: H0887 )
  25. Coverslips coating solution, e.g., 0.01% Poly-L-lysine solution (Sigma, catalog number: P4707 ), collagen coating solution (according to the manufacturer's instruction, Sigma, catalog number: SAFC-125-50 )
  26. Sterile 1x PBS pH 7.4, no calcium, no magnesium for cell culture (e.g., Corning, catalog number: 21-031-CV )
  27. 10× DPBS, (e.g., Roche, catalog number: 11666789001 )
  28. 32% (wt/vol) paraformaldehyde (PFA) (Electron Microscopy Sciences, catalog number: 15714 )
    !CAUTION: Paraformaldehyde is a hazardous solution and a cross-linking agent. Wear protective gloves and handle it under a fume hood.
  29. Triton X-100, 0.1% (vol/vol) (Thermo Fisher Scientific, catalog number: BD 151500 )
  30. Nuclease-free water (Thermo Fisher Scientific, catalog number: 10977-015 )
  31. Immersion oil 1.518, for the microscope/objective
  32. Bovine serum albumin (BSA) (Sigma, catalog number: A7030 )
  33. Cloning of SNAP-histone expression vector and its stably expressing cell line (see Recipes)
  34. Fixation Buffer (see Recipes)
  35. Permeabilization buffer (see Recipes)
  36. Blocking buffer (see Recipes)
  37. Sorting Buffer (see Recipes)

Equipment

  1. Vortex mixer
  2. Tabletop centrifuge
  3. Pointed tip tweezer
  4. Water bath
  5. Biological hood/biosafety cabinet (for cell culture work)
  6. Cell culture incubator suitable for cell culture of your choice (e.g., 37 °C, 5% CO2)
  7. Wide-field fluorescent microscope (e.g., Olympus, model: BX-61 , equipped with four filter sets for DAPI (Semrock, model: DAPI-5060C-Zero ), Cy3 (Chroma, model: 41007 ), FITC (Semrock, model: FITC-5050A-Zero), and Cy5 (Semrock, model: Cy5-4040C-Zero ), an EXFO X-Cite Series 120 PC metal halide light source, Photometrics Cool SNAP HQ CCD camera, and molecular Devices Metamorph acquisition software or equivalent microscope set-ups or equivalent microscope system
  8. Flow cytometer (BD Biosciences, model: LSR II )

Software

  1. Metamorph software for image acquisition (https://www.moleculardevices.com/systems/metamorph-research-imaging/metamorph-microscopy-automation-and-image-analysis-software)
  2. ImageJ/FIJI (Schindelin et al., 2012) (avalable at https://fiji.sc/)
  3. FlowJo (https://www.flowjo.com/solutions/flowjo/downloads)

Procedure

The protocol contains five main processes: (i) Generation of the SNAP-tagged histone expression vector and stable expressing cell lines, (ii) Optimization of cell cycle synchronization, (iii) Detection of global and local histone incorporation using imaging, (iv) Detection of global histone incorporation using flow cytometry, (v) Analysis. Although the first process (i) explains general procedures for cloning and establishing stably expressing cell lines, we emphasize the tips on how to design the SNAP-tagged histone expression vector and isolation of stably expressing cells. This protocol mainly describes the procedures for the fluorescent labeling approaches to investigate histone turnover using microscope (iii) and flow cytometry (iv) (Figure 1).


Figure 1. Workflow of protocol. Three sections of pipelines, Generation of constructs, Sample preparation for imaging and Sample preparation for flow cytometry, are shown as boxes.

(i) Generation of the SNAP-tagged histone expression vector and stable expressing cell lines
Protein tagging with a small epitope is valuable for detection of the protein of interest in various biochemical approaches. However, tagging a small peptide sometimes interferes with the biological function of the target protein and localization of the protein. In general, it is desirable to test whether the N-terminal or C-terminal tagging of protein alters its functions. The localization of the SNAP-tagged histone variant can be confirmed using immunofluorescence by comparing with the endogenous histone variant whether it is localized at the expected chromatin sites. Additionally, it may be necessary to test if the SNAP-tagged construct retains its specific function in chromatin.
  To determine the precise turnover of histone variants, the generation of stably expressing cell lines is highly recommended, since expression level is expected to be altered after mitosis when a transiently transfected SNAP-tagged histone is used. Inserting the SNAP-tag sequence into the endogenous histone locus using CRISPR/Cas9 technology is ideal but not necessary. The SNAP-expressing vector is commercially available from NEB, which contains a neomycin selection gene. In the process of establishing stably expressing cell lines, titration of the drug concentration in your cells is required for the isolation of positive cells. If your cell line already obtains neomycin resistance, such as HEK 293T cell which is immortalized with the large T antigen with neomycin resistance, the neomycin selection marker needs to be replaced.
  The expression level of the SNAP-tagged histone may also influence the timing of incorporation of histones. Since overexpression of histone variants may cause undesirable non-specific incorporation into chromatin, isolating a cell population with a moderate to lower expression of SNAP-tagging histones by Fluorescence-activated cell sorting (FACS) before the pulse-chase experiment is recommended. We also recommend testing the level of SNAP-histone variants compared with endogenous histone variants in purified chromatin fractions by western blotting after establishment of the cell line.

(ii) Optimization of cell cycle synchronization
Cell cycle synchronization is a common method to arrest the whole cell population into a particular cell cycle phase. Most cell cycle synchronization methods rely on the use of a drug which blocks a specific function required for cell cycle progression. While various cell cycle synchronization methods were established in the past (Table 1), the efficiency of synchronization with drug concentration might depend on the cell type used. Any cell synchronization methods can be used in this protocol after the optimization of drug treatment and staining scheme of pre-existing and newly incorporated histones. Here we introduce the protocol in HEK293T cells using a double thymidine block for synchronization at G1/S phase and mitotic shake-off for synchronization at G2/M phase (Figure 2). Both are widely utilized as general cell cycle synchronization methods. Successful cell synchronization and release should be confirmed by flow cytometry, western blotting or immunofluorescence (IF) with cell cycle indicators or markers such as co-expressing the Fucci cell cycle reporter (Sakaue-Sawano et al., 2008), DNA staining with Hoechst 33342, anti-phosphorylated Histone-3 at Serine-28 (M phase marker) or any other cell cycle markers.


Figure 2. Workflow of cell synchronization and labeling of histones. Each step of cell synchronization methods (A. double thymidine block and B. mitotic shake-off) and timing of labeling of pre-existing histones are shown.

Table 1. Common drug list for cell synchronization


(iii) Detection of global and local histone incorporation using imaging
A key procedure for imaging detection is to grow cells on coverslips. Specific procedures for coating of coverslips might be required for different cell types. Careful handling is important to keep the adherent cells attached to the coverslips during the entire procedure. Localization of the histone variant with specific chromatin structure can be addressed by colocalization analysis with a marker of desired chromatin structure using IF after the fixation of cells. In this case, carefully consider the choice of fluorophores and the filter setting of your microscope to avoid the incompatible bleed-through detection of fluorophores.


  1. Labeling of histone incorporation during S-G2 phase ● Timing 5 days
    1. Place coverslips in 12-well or 24-well plate and perform coating coverslips (e.g., 0.01% poly-L lysine, 0.01% collagen) at room temperature for 1 h. After washing 3 times with double distilled water, spread cells on coated coverslips and grow cells at least 24 h.
    2. Cell synchronization at the G1/S can be performed using double thymidine block methods (Jackman and O'Connor, 2001). Incubation time and drug concentrations must be optimized for each cell type. Synchronize the cells at the G1/S transition with a treatment of 2 mM thymidine for 16 h, release from thymidine for 9 h and treat again with 2 mM thymidine again for 17 h.
      ? TROUBLESHOOTING (Table 2)

      Table 2. Troubleshooting Table


    3. Label pre-existing SNAP-tagged histones with SNAP-Cell Oregon Green (1 µM) in culture medium for 30 min at 37 °C in 5% CO2.
    4. Block unlabeled pre-existing SNAP-tagged histones with non-fluorescent SNAP-substrates, SNAP-cell Block (10 µM) for 30 min in culture medium at 37 °C in 5% CO2.
      CRITICAL STEP: This step is important to avoid insufficient initial labeling of pre-existing histones that will result in inaccurate detection of newly incorporated histones.
    5. Add culture medium containing 10 μM of EdU and 10 μM RO-3306 and incubate for 12 h at 37 °C in 5% CO2.
      ? TROUBLESHOOTING (Table 2)
    6. Label newly incorporated SNAP histones using SNAP-Cell TMR (1 µM) in culture medium with RO-3306 for 30 min at 37 °C in 5% CO2.

  2. Labeling of histone incorporation during G1 phase ● Timing 5 days
    To collect mitotic cells, a mitotic shake off can be performed.
    1. Spread cells in 100 mm dish.
    2. Synchronize the cells at S/G1 border with 2 mM thymidine for 24 h, then treat with 20-500 nM nocodazole for 12 h. Incubation time and drug concentrations must be optimized for each cell type.
    3. Check the cells forming rounding shape and weak attachment using light microscopy.
      CRITICAL STEP: If cells are not forming rounded shape, synchronizing at M phase is not successful.
      ? TROUBLESHOOTING (Table 2)
    4. Gently shake the dish to collect mitotic cells into 1.5 ml Eppendorf tube or 15 ml Falcon tubes.
    5. Centrifuge the cells at 500 x g for 4 min and wash cells with a culture medium.
    6. Wash cells twice with culture medium and spread cells on pre-coated coverslips as described in Step 1.
    7. One to two hours later, check the cells if they attached to the coverslips.
      CRITICAL STEP: The cells must be attached to the cover glass. If not, releasing from nocodazole treatment is not successful.
      ? TROUBLESHOOTING (Table 2)
    8. Label pre-existing SNAP-tagged histones with SNAP-Cell Oregon Green (1 µM) in culture medium for 30 min at 37 °C in 5% CO2.
    9. Block unlabeled pre-existing SNAP-tagged histones with non-fluorescent SNAP-substrates, SNAP-cell Block (10 µM) for 30 min in culture medium at 37 °C in 5% CO2.
      CRITICAL STEP: This step is important to avoid insufficient initial labeling of pre-existing histones that will result in inaccurate detection of newly incorporated histones.
    10. Add 10 µM of EdU and 2 mM thymidine in culture medium for 12-18 h at 37 °C in 5% CO2.
    11. Label newly incorporated SNAP histones using SNAP-Cell TMR (1 µM) in culture medium with 2 mM thymidine for 30 min at 37 °C in 5% CO2.

  3. Cell fixation, Permeabilization and Click-it EdU labeling ● Timing 2 days
    1. Fix cells with fixation buffer for 15 min at room temperature.
      PAUSE POINT: The fixed cells can be stored in PBS at 4 °C for up to 1 month.
      ! CAUTION Paraformaldehyde is a hazardous solution and a cross-linking agent. Wear protective gloves and handle it under a fume hood. 
    2. Permeabilize the cells with permeabilization buffer for 15 min at room temperature.
    3. Block cells using 1x PBS containing 3% BSA for 1 h and perform click-it EdU labeling as described in the manufacturer's instruction.
    4. (Optional) Immunofluorescence (IF) can be performed using the desired antibody if co-localization with histones needs to be addressed. However, cautious optimization is required for the microscope filter setup and wavelength range to avoid bleed-through from other fluorophores that co-labeled in the same cells.
    5. DNA staining with DAPI or Hoechst 33342.
    6. Mount coverslips using anti-Prolong Diamond Antifade Mountant.

  4. Detection using microscope ● Timing 1-2 days
    1. Images can be acquired with an Olympus BX61 widefield, epifluorescent microscope using a 60x 1.4 PlanApo objective or equivalent. Filter sets for DAPI (Semrock), Cy3 (Chroma), FITC (Semrock), and Cy5 (Semrock), with an EXFO X-Cite Series 120 PC metal halide light source, Photometrics Cool SNAP HQ CCD camera, Olympus Type-F immersion oil (nd 1.516) and Molecular Devices Metamorph acquisition software or equivalent microscope set-ups are required. Images can be taken with optically sectioned using a 0.5 μm Z step, spanning a 6~10.0 μm Z depth in total depending on the thickness of each cell type. Exposure times of 10 to 200 ms are typically used to acquire each plane in the Cy3, Cy5, FITC and DAPI channels.

(iv) Detection of global incorporation using flow cytometry

We also introduce the protocol addressing the global histone turnover using flow cytometry detection. Although this approach can determine the global histone kinetics, it is unable to detect the histone kinetics at specific genomic loci or chromatin architectures.


  1. Labeling of histone incorporation
    1. Spread cells in 12 of 35 mm or 60 mm dishes.
    2. Synchronize the cells at the G1/S transition by double thymidine block. Briefly, treated cells with 2 mM thymidine for 16 h, wash 3 times with 1x PBS, and release from thymidine by replacing culture medium for 9 h and treat with 2 mM thymidine again for 17 h.
    3. Label pre-existing SNAP-tagged histones with SNAP-Cell Oregon Green (1 µM) in culture medium for 30 min at 37 °C in 5% CO2.
    4. Treat cells with SNAP-cell Block (10 µM) for 30 min in culture medium at 37 °C in 5% CO2 to mask insufficient labeling of pre-existing SNAP-tagged histones with non-fluorescent SNAP-substrates.
      CRITICAL STEP: This step is important to avoid insufficient initial labeling of pre-existing histones that will result in inaccurate detection of newly incorporated histones.
    5. Release 6 dishes from cell synchronization for detection of 12- to 22-hour time points. Keep synchronizing other 6 dishes for detection of 2- to 12-hour time point.
    6. Incubate cells for 12 h.
    7. Release the other 6 dishes from cell synchronization for detection of 2- to 12-hour time point.
    8. Harvest cells every 2 h and store cells at -80 °C.
      PAUSE POINT: The frozen medium and protocol are depending on cell types.
    9. Thaw cells and wash one time with a pre-warmed medium.
    10. Label newly incorporated SNAP histones by adding culture medium with SNAP-JF646 (1 µM) to cell pellets. Pipet cells and incubate for 30 min at 37 °C water bath.
    11. Centrifuge the cells at 500 x g for 4 min at room temperature and wash twice with pre-warmed culture medium.
    12. (Optional) Commercially available Dyes detecting cell cycle (e.g., Hoechst 33342) are also optional instead of the Fucci cell cycle indicator.
    13. Detection of expression levels of Oregon green (Pre-existing histones), JF646 (newly incorporated histones), mCherry (S-G2 phase cell cycle indicator) and TagBFP (G1 phase cell cycle indicator) using Flow cytometry.

    BOX 1. Manual imaging Analysis workflow of global pre-existing and newly incorporated histones (see also Figure 3)


    Figure 3. Imaging analysis workflow. Outlines of imaging analysis to detect intensity of pre-existing, newly incorporated histones, nuclear staining (e.g., DAPI and Hoechst 33342) and EdU (S phase marker) are shown.

    1. Process all image sets to Maximum Intensity Z-projection using the batch process function as follows:
      1. Process/Batch/Macro.
      2. Choose the Input folder in "Input…".
      3. Choose the output folder in "Output…".
      4. (Optional) Put specific name in “File name contains:” if specific files in selected folder need to be processed.
      5. Put macro code “run("Z Project...", "projection=[Max Intensity]");” in the script box.
      6. Click “Process”.
      It can be manually processed as follows:
      1. Open four images by dragging the files into the main interface of FIJI or following:
        File/Open/, then choose your image.
      2. Image/Stacks/Z Project/, then choose projection type “Max Intensity”.
    2. Define the region of interest (ROI) to measure the intensities of pre-existing and newly incorporated histones, and EdU incorporation by making a mask using nuclear staining such as DNA staining with DAPI.
      Duplicate the image displaying DNA staining such as DAPI.
      1. Select duplicated DAPI channel image.
      2. Image/Duplicate… 
      Define the nuclear area using the intensity of nuclear staining.
      1. Image/Adjust/Threshold; then choose suitable threshold method (Default can be chosen as a start), check “Dark background” and Apply.
      2. Process/Binary/Convert to Mask.
      3. (Optional) Process/Binary/Fill holes.
      4. (Optional) Process/Binary/Watershed (https://imagej.net/Nuclei_Watershed_Separation).
      5. Analyze/Set Measurements, check “Area”, “Mean gray value” and “Display label”, then OK.
      6. Analyze/Analyze Particles, add proper number to accomplish the selection of proper area into “Size (pixel^2)” and “Circularity”, check “Add to Manager”, then OK.
    3. Measure DAPI signal (DNA) in the masked area using Region of interest (ROI).
      1. Select channel image.
      2. Click “Measure” in ROI Manager.
    4. Measure Oregon green signal (Pre-existing histones) using ROI.
      1. Select Oregon green channel image.
      2. Click “Measure” in ROI Manager.
    5. Measure TMR-STAR signal (newly incorporated histones) signal using ROI.
      1. Select TMR-STAR channel image.
      2. Click “Measure” in ROI Manager.
    6. Measure EdU signal (Nascent DNA) using ROI.
      1. Select EdU channel image.
      2. Click “Measure” in ROI Manager.
    7. Save results and analyze.

    BOX 2. Analysis workflow of local pre-existing and newly incorporated histones using color profiler or RBG profiler (see also Figure 5A)
    1. Install the plugin in Fiji/ImageJ software.
    2. Open all image sets that will be compared by dragging the files into the main interface of FIJI or following:
      File/Open/, then choose your image.
    3. Merge these images into a single image.
      Image/Color/Merge Channels.
    4. Change the image type to RGB color.
      Image/Type/RGB color.
    5. Make a line to make a profile.
      1. Choose a line tool.
      2. Make a line.
      3. Plugins/RGB Profiler.

Imaging analysis (~1 week)
Steps 1-6, cell synchronization using double thymidine block and labeling pre-existing histones for imaging analysis: 4 days.
Steps 7-17, cell synchronization using mitotic shake off, and labeling pre-existing and newly incorporated histones for imaging analysis: 4 days.
Steps 18- 19, fixation and permeabilization of the cells: 1 h.
Step 20, labeling of EdU using click-it kit: 1 h.
Step 21, (optional) IF using the desired antibody to determine the histone deposition at specific chromatin architecture or factors: 3 h.
Steps 22- 23, DNA staining and mount coverslips: Overnight.
Step 24, microscopy imaging: 1 day.

Flow cytometry analysis (~1 week)
Step 25, Spread cells and cell synchronization using double thymidine block and labeling pre-existing histones: 4 days.
Step 32, Release from cell synchronization and harvest cells: 2 days.
Steps 34-37, labeling of newly incorporated cells and Flow cytometry analysis: 2 h.

Data analysis

We introduce a basic analytical pipeline for imaging approach using FIJI/ImageJ which is a free open source image processing package. An automated analysis using ImageJ macro code would be time-saving for large data sets if you are familiar with programming. Additionally, many other imaging processing programs (e.g., Matlab) can be also a great option to process enormous data sets.
  Data analysis for global histone turnover using flow cytometry detection can be easily accomplished using available flow cytometers such as LSR II (BD Bioscience). The software such as FlowJo (BD Bioscience) can assess the FCS data sets and permit visualizing complex cytometric data sets. In this protocol, we can use a mean or median of the total intensity of fluorophore in the entire cell population with single-cell resolution to investigate the timing of incorporation and deposition of histones. Displaying histograms of fluorescent intensities from labeling with pre-existing and newly incorporated histones as well as cell cycle markers in this protocol, can help to visualize the timing of global histone kinetics including the dissociation of pre-existing histones and incorporation of newly synthesized histones during the cell cycle.

Anticipated results
In this protocol, we summarized our previously published results for the cell cycle-specific dynamics of histone H3.1 and macroH2A1.2 in HEK293T cells to show that the SNAP-labeling pulse-chase system works reliably and to illustrate how to analyze the imaging data sets (Sato et al., 2019).
  Although we illustrate three parts in this protocol (Figure 1) including generation of constructs, sample preparation for imaging and sample preparation for flow cytometry, we mainly described the detailed protocol for the latter two. We also illustrate how to analyze the imaging data sets using ImageJ/FIJI (Figure 3 and Box 1). Here we present an example of the SNAP pulse-chase detection during the cell cycle using an imaging approach detecting SNAP-histone H3.1 (Figure 4). We demonstrated two types of cell synchronization, double thymidine block and mitotic shake off, to investigate histone incorporation and dissociation during the S-G2 or G1 phase. The pre-existing and newly incorporated SNAP-H3.1 stably expressed in HEK 293T were labeled as shown in Figures 4A-4B. EdU click-it detection was also performed to confirm successful cell synchronization. The global histone incorporation rates in individual cells were determined by the mean of pixel intensity of newly incorporated histones (TMR-Star) normalized by the mean of pixel intensity of pre-existing histones (Oregon Green) in the nucleus. Using this approach, we detected H3.1 incorporation during S-G2 phase as it is known as DNA replication-dependent deposition (Figures 4C-4D).
  We also illustrate the detection of colocalization in imaging analysis using RGB Profiler (Figure 5A and Box 2). Here we present the colocalization of newly incorporated macroH2A histone variant at inactivate X chromosomes (Xi) in HEK 293T cells (Figures 5A-5B). The pre-existing and newly incorporated SNAP-macroH2A1.2 were labeled as shown in Figure 4A and incorporation of SNAP-macroH2A1.2 in Xi during the cell cycle was determined by the colocalization with pre-existing SNAP-macroH2A1.2. As shown in Figure 5B, macroH2A1.2 is incorporated into the Xi during the G1 phase.
  Global histone incorporation and dissociation also can be detected by flow cytometry. We demonstrated the SNAP-macroH2A1.2 pulse-chase approach and detected the global SNAP-macroH2A dynamics by flow cytometry as shown in Figures 5C-5D. Here we synchronized cells at G1/S border and labeled pre-existing SNAP-macroH2A1.2 with Oregon green, then detected newly incorporated SNAP-macroH2A1.2 every two h after releasing from cell synchronization. Although we determined the cell cycle transition using the Fucci cell cycle sensor (Figure 5C [two right panels] and 5D [top]), other cell cycle indicator such as Hoechst 33342 DNA staining also could be the option (Figure 6). A histogram shows the fluorescent intensities of cell populations. The transitions of signal intensity of pre-existing SNAP-macroH2A1.2 (Oregon green, Figure 5C [left] and 5D [middle]) show that the signal intensity decreased when cells entered mitosis but little pre-existing SNAP-macroH2A dissociate from chromatin through the entire cell cycle. The transitions of newly incorporated SNAP-macroH2A1.2 (JF646, Figure 5C [second left], and 5D [bottom]).


Figure 4. Detection of global pre-existing and newly incorporated histone H3.1 during specific cell cycle. A. The illustration of labelling histones at S/G2 and G1 phases for the analysis in B-D. (Left panel) To label newly incorporated histones in S/G2 phase, HEK 293T cells stably expressing SNAP-tagged H3 were synchronized at the G1/S phase border by double thymidine block as shown as TH. Pre-existing histones were labeled with SNAP-Oregon Green (green arrow), subsequently blocked using the non-fluorescent SNAP-Block reagent to avoid non-sufficient labeling. The cells were released from synchronization to progress to the G2/M transition until they were synchronized at the G2/M border using RO-3306 (a CDK1/cyclin B1 and CDK1/cyclin A inhibitor, shown as RO), then newly-incorporated SNAP-tagged histones were labeled with SNAP-TMR Star (red arrow). (Right panel) To detect newly incorporated histones in the G1 phase, mitotic cells were collected by shake-off following nocodazole treatment for 12 h and spread onto coverslips (shown as Noc). After 2 h, pre-existing SNAP-H3 were labelled with Oregon Green and treated with the blocking reagent. Cells were released from synchronization and progressed to the G1/S transition using double thymidine block, then newly-incorporated SNAP-tagged histones were labeled with SNAP-TMR Star. After being released from the first synchronization, the cells were also treated with EdU until the second synchronization, allowing cells that had undergone DNA synthesis to be identified. B. Representative images of pre-existing and newly-synthesized SNAP-tagged H3.1 in S/G2 or G1 phase. Scale bar = 10 µm. Image analysis showing a higher incorporation of histone H3 during the S-G2 phase (C-D). C. Frequency distribution of histone incorporation rate in S-G2 and G1 cell cycle phase. Histone incorporation rate was calculated by the mean pixel intensity of red (newly incorporated histones) normalized by mean pixel intensity of green (Pre-existing histones) in the nucleus during the S/G2 and in G1 phases. Successful cell synchronization was confirmed by EdU labeling. Hoechst 33342 labeling was performed to define the area of a nucleus in the following analysis. D. Quantification of histone incorporation rate during S-G2 and G2 phase. Each dot denotes the incorporation rate as described in Figure 4C from single cells. The error bars represent one standard deviation from the number of single cells (n = that indicated on each data set. *P < 0.0001. The P values were determined using two-tailed unpaired t-tests.


Figure 5. Detection of local and global pre-existing and newly incorporated histone macroH2A1.2 during specific cell cycle. A. The outline of image analysis using RBG Profiler plugin (https://imagej.nih.gov/ij/plugins/rgb-profiler.html, Color Profiler plugin (https://imagej.nih.gov/ij/plugins/color-profiler.html) also provides the same functionality) to address the colocalization of newly incorporated histones with pre-existing histones. Cell cycle-specific histone incorporation at specific chromatin architecture can also be addressed with the detection of chromatin structure using IF. Here we show the example detecting colocalization with pre-existing and newly incorporating histones. B. the example of image analysis using cell profiler. The left images of pre-existing (Oregon Green: green), newly incorporated (TMR: red) SNAP-tagged macroH2A during S-G2 (upper) and G1 (lower) phase were merged into single images and colocalization on the two inactive X chromosomes (Xi) were determined in HEK 293T cells. In the upper and lower right panels, the signal intensities measured along the white lines in the images of pre-existing and newly incorporated are shown. C. Flow cytometric analysis of pre-existing (left: Oregon Green) and newly incorporated macroH2A (second left: JF646), with S-G2 (second right: mCherry-tagged hGeminin) and G1 (right: TagBFP-tagged hCdt1) cell cycle markers derived from Fucci sensor. Histograms indicate the fluorescent intensity at each time point (2-22 hour) after releasing from synchronization at the G1/S border. The cell cycles at each time point were estimated from the Fucci cell cycle sensor. D. the median of fluorescent intensity of pre-existing (middle: Oregon Green) and newly incorporated macroH2A (bottom: JF646), with S-G2 and G1 (top: mCherry-tagged hGeminin and TagBFP-tagged hCdt1) cell cycle markers from Flow cytometric analysis are plotted. Error bars indicate 95% confident intervals.


Figure 6. Cell cycle determination using Hoechst 33342 staining and Flow Cytometry detection. A. Flow cytometric analysis of cell cycle (left: Hoechst 33342) and pre-existing SNAP-macroH2A (right: Oregon Green) labeled and released from cell synchronization as described in Steps 25-36. Histograms indicate the fluorescent intensity at each time point (2-22 h) after releasing from synchronization at the G1/S border. The cell cycles at each time point could be determined by the intensity transitions of Hoechst 33342 in the cell population. B. the median of fluorescent intensity of DNA staining (top: Hoechst 33342) and pre-existing SNAP-macorH2A1.2 (middle: Oregon Green) from flow cytometric analysis are plotted. Error bars indicate 95% confident intervals.

Recipes

  1. Cloning of SNAP-histone expression vector and its stably expressing cell line
    A plasmid that encodes desired histone with SNAP-tag needs to be cloned by general cloning methods. pSNAPf Vector (catalog # N9183S from NEB) is useful to establish your construct and stably expressing cell line since it contains neomycin resistance gene for efficient drug selection.
  2. Fixation Buffer
    4% (wt/vol) PFA in 1x PBS
    Note: This solution can be prepared from 32% (wt/vol) PFA. Protected from light for up to 2 years.
  3. Permeabilization buffer
    0.5% Triton in 1x PBS
  4. Blocking buffer
    3% BSA in 1x PBS
  5. Sorting Buffer
    1 mM EDTA
    25 mM HEPES pH 7.0
    1% FBS (Heat inactivated) in 1x DPBS

Acknowledgments

This work was supported by NIH grant R01 DA030317 (JMG). We thank members of the Greally and Singer laboratories for discussions, the Einstein FACS and Genomics cores. We also thank L. Lavis for SNAP-JF646. This protocol was adopted from previous work (Sato et al., 2019).
  Author contributions statement: H.S. designed and performed the experiments and analyzed the data. H.S., J.M.G. and R.H.S. wrote the manuscript. J.M.G. and R.H.S. supervised the research.

Competing interests

The authors declare no competing financial interests.

References

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简介

[摘要] 动态的组蛋白变化是染色质调节的核心部分。组蛋白变体的沉积和组蛋白的翻译后修饰与染色质状态的属性密切相关。表征组蛋白变体的动力学特性可为深入了解转录调控,染色质维持和其他染色质特性提供重要信息。在这里,我们提供了一种定量和敏感方法的协议,以使用双色SNAP标记系统测试组蛋白的结合和解离时间,分别标记预先存在的和新结合的组蛋白。结合细胞周期同步方法和细胞周期标志物,这种方法可以进行脉冲追踪分析,以确定在细胞周期内使用成像或流式细胞仪方法以单细胞分辨率检测到的组蛋白变体的周转率。除了测试整体组蛋白更新,还可以使用成像方法解决组蛋白变体的细胞周期依赖性细胞定位。

[背景] 染色质重塑是真核细胞众多基本细胞活动的一部分(Geiman 和Robertson,2002;Clapier 和Cairns ,2009)。转录因子和RNA聚合酶的可及性通常与DNA甲基化和染色质状态的变化相关,包括可及性,翻译后的组蛋白修饰和组蛋白变体的沉积。组蛋白变体差异性地调节调节发育,细胞分化或其他生理活动的基因表达(Banaszynski 等,2010)。它们在DNA修复,端粒维护,异染色质形成和染色质分离中也发挥着不同的作用(Henikoff 和Smith,2015; Zink和Hake,2016)。此外,组蛋白变体的掺入失调与癌症有关(Vardabasso et al。,2014),表明其在人类疾病中的重要作用。了解组蛋白变体的动力学,包括在细胞周期中掺入特定的染色质区域或从中分离,是至关重要的,因为它将调节特性与表观遗传的有丝分裂维持紧密联系在一起(从亲代细胞到子代细胞的遗传力)。DNA复制涉及主要的染色质重塑,以在有丝分裂后复制整个染色质结构。在DNA复制之前与染色质相关的组蛋白通过DNA聚合酶复合物的进入而从DNA瞬时解离。新合成的复制叉与新合成的组蛋白一起随机重新释放先前存在的组蛋白(Balhorn 等,1975;Alabert 和Groth ,2012;Annunziato ,2012)。组蛋白和某些组蛋白变体的预先存在的翻译后修饰也在复制叉处重新与新合成的DNA结合,这说明复制叉处预先存在的组蛋白的重新结合是维持有丝分裂遗传的一部分。染色质状态。为了确定翻译后修饰或组蛋白变体掺入的时间,需要一种灵敏的脉冲追踪系统,该系统可以将新掺入的组蛋白的检测与先前存在的组蛋白区分开。当新合成的规范组蛋白在DNA复制过程中被整合到染色质中时,它们不会被修饰。与DNA复制同时发生的从头DNA甲基化不同(Tillo 等人,2016),大多数组蛋白标记未在复制叉上新掺入的组蛋白上建立。基于蛋白质组学的脉冲追踪方法(请参见下面的替代方法部分)已用于通过检测赖氨酸残基上已存在和新沉积的组蛋白乙酰化和甲基化来确定组蛋白翻译后修饰的整体动力学(Pesavento 等。,2008; Scharf 等,2009; Martinez-Garcia 等,2011; Xu 等,2011; Zee 等,2012;Alabert 等,2015)。这种方法确定了组蛋白修饰的两个独特的动力学模式。一组,例如在27(H3 K27 ac)处的组蛋白H3乙酰赖氨酸,表现出快速的周转以平衡,并且值得注意的是,它们在整个细胞周期中都没有得到维持(Scharf et al。,2009)。这种快速的乙酰化动力学可能代表了暂时活跃的转录动力学(Stasevich et al。,2014)。而包括在图9(K9me3)H3赖氨酸三甲基化或H3 K27me3另一组是在细胞周期的代替以下更缓慢地获取并逐步从单和二到三甲基化,建立期间G1期有丝分裂之前(Pesavento 等人。,2008;沙尔夫等人,2009; Martinez的加西亚。等人,2011;徐。等人,2011; Zee的。等人,2012; Alabert 。等人,2015) 。重要的是,该组具有有丝分裂染色体记忆的特性。通过成像方法已经解决了在细胞分裂过程中维持组蛋白标记的潜在机制。原位 使用抗组蛋白赖氨酸甲基化的特异性抗体及其甲基转移酶进行的近距离连接测定法检测到,组蛋白修饰剂与果蝇胚胎中复制的DNA成分连续缔合,表明复制DNA上修饰剂的缔合可能提供了被甲基化的`` 标签'' (Petruk 等。,2012)。

化学蛋白质标记技术的最新进展为我们提供了一种在活细胞中进行蛋白质标记应用的强大工具(例如SNAP(新英格兰生物实验室),CLIP(新英格兰生物实验室),Halo(普罗mega)和TMP(活性母体)标签)。这些技术基于与与细胞渗透性底物(例如合成荧光染料或生物素)缀合的特定配体结合的遗传编码标签的共价标记,可以在生化应用中介导亲和纯化。与常见的遗传编码标签相反,蛋白质的这种化学标记可用于具有多种荧光团选择的时序依赖性标记中,从而可以对特定蛋白质进行脉冲追踪标记。这项标记技术通过成像检测(例如,着丝粒处的CENP-A [Dunleavy 等,2009 ] 和异染色质处的macroH2A [ Sato 等,2019 ] )揭示了特定染色质结构上组蛋白变体的沉积时间。

在这里,我们提供了使用SNAP标签标记系统的脉冲追踪方法的详细协议,该系统利用了具有单细胞分辨率的定量组蛋白变异检测。使用此协议,可以同时检测到不同的组蛋白动力学,既有组蛋白的解离和新合成的组蛋白的缔合。我们描述了两种检测方法,荧光显微镜和流式细胞仪,以及使用FIJI / ImageJ软件(免费提供)的成像分析的详细信息(https://fiji.sc/)。



优点和局限性

与使用放射性分子的常规脉冲追踪方法相比,使用化学蛋白质标记系统的脉冲追踪方法是一种简单,无危险且敏感的方法(请参阅替代方法部分)。大多数所需的试剂和荧光团是可商购的。可以使用成像和流式细胞仪应用解决组蛋白的全球营业额,特别是,特定的染色质结构上的时序特异性定位也可以通过成像方法来表征。使用此协议,可以在同一细胞中以单个细胞分辨率同时检测既存的和新合并的组蛋白变体。与检测内源性组蛋白的放射性同位素或代谢标记方法不同,此标记方法依赖于可以与特定底物相连的遗传编码标签(例如SNAP-,CLIP-,Halo标签)。因此,需要表达具有SNAP标签的靶组蛋白的质粒构建体及其在稳定表达的细胞系中的用途。另外,必须检查所需标签组蛋白的定位和其他生物学功能,以确定其是否仍起内源性组蛋白的作用。为了获得准确的观察结果,可能需要优化构建体(例如,N末端或C末端标记,更改启动子的选择)和表达水平。该方法的另一个局限性在于它不适用于检测组蛋白的翻译后修饰。



替代方法

蛋白质组学检测同位素标记

SILAC(稳定的同位素在文化中用氨基酸标记),然后进行质谱分析是研究内源性组蛋白变体和翻译后修饰的全球营业额的有力方法(Yuan 等人,2014 )。在这种方法中,新合成的组蛋白被放射性重同位素标记,与已存在的含轻氨基酸的组蛋白相比,追踪了被标记的组蛋白的转换。质谱分析后确定翻译后修饰或变体的预先存在和沉积。这种方法适用于检测整体组蛋白更新,但不能检测特定染色质结构或基因组位点处的组蛋白标记。



全基因组方法的代谢标记

新生蛋白质的非放射性代谢标记可以是标记全局新合成的组蛋白的替代方法。也称为“ CATCH-IT”的方法“将标签共价附着以捕获组蛋白和识别周转率”,可以进行全基因组研究以表征活性组蛋白置换(Deal 等,2010)。此方法基于通过结合蛋氨酸同系物叠氮高丙氨酸(Aha)来新生肽的标记方案,该方法通常用于检测细胞中的活性翻译。在这种方法中,将含有Aha标记的新合成组蛋白的核小体通过环加成反应(称为“喀哒”化学)与生物素生物缀合,并用抗生蛋白链菌素珠粒拉下。将来自下拉的分离的DNA 应用于平铺微阵列,以确定在果蝇S2细胞中表现出活性组蛋白置换的基因组位点。具有活性组蛋白更新的特征基因组位点与组蛋白H3变体H3.3的掺入位点相对应,该位点在转录活性位点被检测到(Henikoff 等,2009)。这种方法能够检测具有组蛋白活跃更新的基因组位点。

化学标记方法(例如SNAP标记系统)也可以是研究全基因组组蛋白变体掺入的替代选择(Sato 等人,2019)。在这种方法中,新组成SNAP标签组蛋白与SNAP-生物素治疗SNAP-细胞后联® 块(bromothenylpteridine ,BTP),非荧光底物,以掩盖预先存在的组蛋白的反应性。然后,可以用抗生蛋白链菌素珠将生物素连接的新结合的组蛋白拉下来。从下拉样品中纯化的DNA片段可通过大规模平行测序进行测序。该方法对于检测组蛋白变体的时间和基因组基因座依赖性掺入但不能检测翻译后修饰可能是有用的。

关键字:染色质动力学, 组蛋白, 细胞周期, 成像, 流式细胞仪

材料和试剂


 


50 ml猎鹰管
15毫升猎鹰管
1.5 ml管
组织培养板(12或24孔; Falcon,目录号:353043或353047)
组织培养皿(35×10mm或60×15毫米和100×20毫米,隼,目录NU MBER 小号:353001或353002和353003)
盖玻片(18或12毫米1.5号)
显微镜载玻片(例如Fisher Fisher,目录号:12-544-2)
5 ml FACS管(Fisher Scientific,目录号:149595)
感兴趣的贴壁细胞(例如,小鼠胚胎成纤维细胞[MEF],HEK293细胞)
SNAP-Cell俄勒冈绿色(New England BioLabs ,目录号:S9104S)
SNAP细胞块(New England BioLabs ,目录号:S9106S)
SNAP-细胞TMR明星(新英格兰生物实验室,目录号:S9105S)
胸苷(Sigma,目录号:T1895)
RO-3306(圣ç 鲁斯乙艾讴,目录号:SC-358700)
诺考达唑(Sigma,目录号:M1404)
延长钻石抗荧光淬灭封固(赛默飞世尔科技,目录号:P36970)             
延长钻石抗淬灭封固用DAPI (赛默飞世尔科技,目录号:P36962)             
药物用于稳定表达细胞的选择[ 例如,600 - 1200微克/毫升G418(遗传霉素),博来霉素(赛默飞世尔科技,产品目录号:R25001)]
Click- iT TM EdU 细胞增殖试剂盒,用于成像,Alexa Fluor TM 647染料(Thermo Fisher Scientific,目录号:C10340)                            
pSNAPf 载体(新英格兰生物实验室,目录号:N9183S)
pCCL-CellCycle (Sato et al。,2019)
合适的培养基和补充剂[ 例如,DMEM(Sigma ,目录号:D6429)补充有10%FBS(Atlanta Biologicals,Inc. ,目录号:S11150H)和100 U / m l 青霉素-链霉素(Sigma ,目录号:15140148) )]             
EDTA,0.5 M,pH 8.0,分子生物学等级(Sigma ,目录号:324506)
HEPES 溶液,1 M,pH 7.0-7.6,无菌过滤(Sigma ,目录号:H0887)
盖玻片涂层溶液,例如0.01%聚L-赖氨酸溶液(Sigma ,目录号:P4707),胶原蛋白涂层溶液(根据制造商的说明,Sigma ,目录号:SAFC-125-50)
无菌1 x PBS pH 7.4,无钙,无镁用于细胞培养(例如,C 序列号:21-031-CV)
10×DPBS,(例如Roche,目录号:11666789001)
32%(wt / vol)多聚甲醛(PFA)(电子显微镜科学,目录号:15714)
!小心:多聚甲醛是危险溶液和交联剂。戴上防护手套并在通风橱中操作。


Triton X- 100,0.1 %(vol / vol)(Thermo Fisher Scientific,目录号:BD 151500)
无核酸酶的水(Thermo Fisher Scientific,目录号:10977-015)
浸油1.518,对于显微镜/ objec 略去
牛血清白蛋白(BSA)(西格玛(Sigma),目录号:A7030)
SNAP组蛋白表达载体及其稳定表达的细胞系的克隆(请参见食谱)
固定缓冲液(请参见配方)
透化缓冲液(请参见配方)
阻塞缓冲区(请参见食谱)
排序缓冲区(请参见食谱)
 


设备


 


涡旋混合器
台式离心机
尖头镊子
水浴
生物罩/生物安全柜(用于细胞培养工作)
细胞培养培养箱适合您选择的细胞培养(例如,37°C,5%CO 2 )
宽网络视场荧光显微镜(例如,Olymp酒店我们,型号:BX-61,配备˚F 我们的过滤器集DAPI(Semrock ,型号:DAPI-50-60℃零),Cy3的(色度,型号:41007),FITC(Semrock ,型号:FITC-5050A零),和Cy5(小号emrock ,型号:Cy5-4040C零),一个ñEXFO X-引用系列120 PC金属卤化物光源,Photometri CS冷却SNAP HQ CCD照相机,和米olecular设备Metamorph 采集软件或等效的显微镜设置或等效的显微镜系统
流式细胞仪(BD Biosciences,型号:LSR II)
 


软件


 


用于图像采集的Metamorph 软件(https://www.moleculardevices.com/systems/metamorph-research-imaging/metamorph-microscopy-automation-and-image-analysis-software)
ImageJ的/斐济(Schindelin 等人,2012) (avalable 在https://fiji.sc/)
FlowJo (https://www.flowjo.com/solutions/flowjo/downloads)
 


程序


 


该协议包含五个主要过程:(我的SNAP-标记的组蛋白的表达载体和稳定表达细胞系的)代,(ⅱ)细胞周期同步的优化,(ⅲ)d 使用成像,全局和局部组蛋白掺入etection(ⅳ )d 使用流式细胞仪整体组蛋白掺入etection,(v)的分析。尽管第一个过程(i )解释了克隆和建立稳定表达细胞系的一般程序,但我们强调了有关如何设计SNAP标记组蛋白表达载体和稳定表达细胞分离的技巧。此协议的主要描述了程序第ë荧光标记方法使用显微镜来研究组蛋白周转(iii)和流式细胞术(ⅳ)(FI 克URE 1) 。


 


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig1 Workflow.jpg


图1 。协议的工作流程。方框显示了管道的三个部分,即构建体的生成,用于成像的样品制备和用于流式细胞术的样品制备。


 


(我)ģ 的SNAP标签组蛋白表达载体的生成及稳定表达细胞系


带有小表位的蛋白质标签对于在各种生化方法中检测目标蛋白质非常有价值。但是,标记小肽有时会干扰目标蛋白的生物学功能和蛋白的定位。通常,期望测试蛋白质的N-末端或C-末端标签改变其功能。在SNAP标记的组蛋白变体的定位可以用免疫荧光通过比较来确认与内源性组蛋白杂物NT 无论是禄在预期的染色质部位alized。另外,可能有必要测试SNAP标签的构建体是否在染色质中保留其特定功能。


  为了确定组蛋白变体的精确周转率,强烈建议生成稳定表达的细胞系,因为当使用瞬时转染的SNAP标记的组蛋白时,有望在有丝分裂后改变表达水平。使用CRISPR / Cas9技术将SNAP-tag序列插入内源性组蛋白基因座是理想的,但不是必需的。SNAP表达载体可从NEB商购,其含有新霉素选择基因。在建立稳定表达的细胞系的过程中,需要滴定细胞中的药物浓度才能分离出阳性细胞。如果您的细胞系已经获得了新霉素抗性,例如用具有新霉素抗性的大T抗原永生化的HEK 293T细胞,则需要更换新霉素选择标记。


  SNAP标记的组蛋白的表达水平也可能影响组蛋白掺入的时机。由于组蛋白变体的过表达可能导致不希望的非特异性结合到染色质中,因此建议在进行脉冲追踪实验之前通过荧光激活细胞分选(FACS)分离具有中等至较低SNAP标签组蛋白表达的细胞群。我们还建议在建立细胞系后通过蛋白质印迹法在纯化的染色质组分中测试SNAP组蛋白变体与内源性组蛋白变体的水平。


 


(ⅱ)ø 细胞周期同步的ptimization


细胞周期同步是使整个细胞群体停滞到特定细胞周期阶段的一种常用方法。大多数细胞周期同步方法依赖于药物的使用,该药物阻断了细胞周期进程所需的特定功能。尽管过去建立了各种细胞周期同步方法(表1 ),但与药物浓度同步的效率可能取决于所用细胞的类型。优化现有药物和新掺入的组蛋白的药物处理和染色方案后,可以在该方案中使用任何细胞同步方法。在这里,我们介绍了在HEK293T细胞中使用双胸腺嘧啶核苷在G1 / S阶段进行同步,并在M2 / G阶段进行有丝分裂摇动以进行同步的协议(图2)。两者都被广泛用作一般的细胞周期同步方法。成功的细胞同步和释放应通过流式细胞术,蛋白质印迹或带有细胞周期指示剂或标记物的免疫荧光(IF)来证实,例如共同表达Fucci 细胞周期报告子(Sakaue-Sawano 等人,2008),用Hoechst进行DNA染色33342,丝氨酸28的抗磷酸化组蛋白3(M期标记)或任何其他细胞周期标记。


 


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig2单元格和label.jpg的同步


图2 。细胞同步和组蛋白标记的工作流程。显示了细胞同步方法的每个步骤(A. 双胸腺嘧啶核苷和B. 有丝分裂),以及预先标记组蛋白的时间。


 


表1.用于细胞同步的常用药物清单


名称





机制


参考文献


胸苷


G1 / S


过量的胸腺嘧啶核苷对DNA复制的反馈抑制作用


(Whitfield 等,2002)


蚜虫


G1 / S


DNA聚合酶抑制剂α


(Wang,1991年)


羟基脲


G1 / S


核糖核苷酸还原酶抑制剂


(Biegel 等,1987;Gallo 等,198 4 ) 


甲氨蝶呤


G1 / S


胸苷生物合成抑制剂


(Yunis 等,1981)


氟脱氧尿苷


G1 / S


胸苷生物合成抑制剂


(韦伯和加森,1983年)


丁酸盐


G1 / S


组蛋白脱乙酰基酶抑制剂


(Lampkin et al。,1971 ; Li,2011 )


洛伐他汀


G1


导致抑制CDK2活性的CDK抑制剂的积累


(Keyomarsi 等,1991)


诺考达唑


G2 / M


微管聚合的抑制剂


(Hoebeke et al。,1976)


RO-3306


前期


CDK1 / cyclin B1和CDK1 / cyclin A的抑制剂


(Vassilev,2006年)


秋水仙碱


中期


微管中的微管蛋白解聚


(Boquest 等,1999)


 






(ⅲ)挪威全局和局部组蛋白掺入使用成像的挠度


成像检测的关键步骤是在盖玻片上生长细胞。用于盖玻片的涂覆具体程序可能需要FO ř 不同CE LL类型。在整个过程中,小心处理对于使粘附细胞保持附着在盖玻片上非常重要。具有特定染色质结构的组蛋白变体的定位可以通过在细胞固定后使用IF用所需染色质结构的标记进行共定位分析来解决。在这种情况下,请仔细考虑荧光团的选择和显微镜的滤光片设置,以避免对荧光团进行不兼容的渗漏检测。


 


在S-G2阶段标记组蛋白掺入● 时间5天
1 将盖玻片放在12孔或24孔板中,并在室温下进行盖玻片(例如0.01%的聚L赖氨酸,0.01%的胶原蛋白)包被1小时。用两次蒸馏水洗涤3次后,将细胞铺在带涂层的盖玻片上,并使细胞生长至少24小时。        


2 G1 / S处的细胞同步可使用双胸苷阻断方法进行(Jackman和O'Connor,2001)。必须针对每种细胞类型优化孵育时间和药物浓度。使细胞在G1 / S转换时与2 mM胸腺嘧啶核苷同步处理16 h,从胸腺嘧啶核苷释放9 h,然后再次用2 mM胸腺嘧啶核苷处理17 h。        


?故障排除(表2)


 


表2.故障排除表





问题


可能的原因





2 5 5


单元不同步


药物浓度或孵育时间未优化。


必须优化药物浓度或孵育时间,或尝试使用其他细胞同步方法。


5、13


单元未从同步释放


药物浓度或孵育时间未优化。


必须优化药物浓度或孵育时间,或尝试使用其他细胞同步方法。


13


细胞未附着在培养皿上


药物浓度或孵育时间未优化。


必须优化药物浓度或孵育时间,或尝试使用其他细胞同步方法。


 


3 用SNAP-Cell Oregon Green(1 µM)在培养基中在5%CO 2中于37°C孵育30分钟,以对预先存在SNAP标签的组蛋白进行标记。        


4 用非荧光SNAP底物封闭SNAP标签的未标记的组蛋白,将SNAP细胞封闭(10 µM)在37°C的5%CO 2 培养基中处理30分钟。        


关键步骤:该步骤对于避免对预先存在的组蛋白进行充分的初始标记是很重要的,因为这将导致新结合的组蛋白的检测不准确。


5 添加含有10的培养基         μ中号的的EdU 和10 μ 中号RO-3306孵育12小时,在37℃在5%CO 2 。


?故障排除(表2)


6 使用SNAP-Cell TMR(1 µM)在含RO-3306的培养基中于37°C于5%CO 2 中培养30分钟,以标记新掺入的SNAP组蛋白。        


 


在G1阶段标记组蛋白掺入● 时间5天
为了收集有丝分裂细胞,可以进行有丝分裂摆脱。


7将细胞铺在100毫米的培养皿中。        


8 将S / G1边界的细胞与2 mM胸腺嘧啶核苷同步24 h,然后用20-500 nM Nocodazole处理12 h。必须针对每种细胞类型优化孵育时间和药物浓度。        


9 使用光学显微镜检查细胞是否形成圆形和附着力弱。        


关键步骤:如果单元未形成圆形,则M相位同步将失败。


?故障排除(表2)


10 轻轻摇动培养皿,将有丝分裂细胞收集到1.5 ml Eppendorf管或15 ml F alcon管中。     


11 以500 xg 离心细胞4分钟,并用培养基洗涤细胞。     


12 用培养基洗涤细胞两次,然后按照步骤1 所述将细胞铺在预涂的盖玻片上。     


13 一到两个小时后,检查细胞是否附着在盖玻片上。     


关键步骤:电池必须固定在玻璃盖上。如果不是这样,从诺考达唑治疗中释放就不会成功。


?故障排除(表2)


14 用SNAP-Cell Oregon Green(1 µM)在培养基中在5%CO 2中于37°C的条件下,用SNAP-Cell Oregon Green(1 µM)标记预先存在的SNAP标签的组蛋白。     


15 用非荧光SNAP底物封闭未标记的SNAP标签组蛋白,SNAP细胞封闭(10 µM)在37 °C的5%CO 2 培养基中30分钟。     


关键步骤:该步骤对于避免对预先存在的组蛋白进行充分的初始标记是很重要的,因为这将导致新结合的组蛋白的检测不准确。


16 在37°C的5%CO 2中于培养基中加入10 µM EdU 和2 mM胸腺嘧啶核苷12-18 h 。     


17 使用SNAP-Cell TMR(1 µM)在含2 mM胸苷的培养基中于37°C于5%CO 2中标记30分钟,以标记新掺入的SNAP组蛋白。     


 


细胞固定,通透性和Click- it EdU 标记● 时间2天
18 在室温下用固定缓冲液固定细胞15分钟。     


暂停点:固定的细胞可以在PBS中于4°C下保存1个月。


!小心多聚甲醛是危险溶液和交联剂。戴上防护手套并在通风橱中操作。


19 用透化缓冲液在室温下透化细胞15分钟。     


20使用1x含3%BSA的PBS 封闭细胞1 h,并按照制造商的说明进行点击EdU 标记。     


21 (可选)如果需要解决与组蛋白的共定位问题,可以使用所需抗体进行免疫荧光(IF)。但是,需要对显微镜滤光片的设置和波长范围进行谨慎的优化,以避免从同一细胞中共同标记的其他荧光团渗出。     


22 用DAPI或Hoechst 33342进行DNA染色。     


23 摩盖玻片使用抗时延钻石抗荧光淬灭封固。     


 


使用显微镜进行检测● 时间为1-2天
24 可以使用60 x 1.4 PlanApo 物镜或等效物,通过Olympus BX61宽视野落射荧光显微镜获取图像。DAPI(Semrock ),Cy3(Chroma),FITC(Semrock )和Cy5(Semrock )的滤光片组,带有EXFO X-Cite系列120 PC金属卤化物光源,Photometrics Cool SNAP HQ CCD相机,Olympus Type-F浸入式需要使用油(nd 1.516)和Molecular Devices Metamorph 采集软件或等效的显微镜设置。图像可以采取使用0.5光学切片微米Z步长,跨越6〜10.0 微米的总取决于每种细胞类型的厚度Z向深度。È 的10至200倍xposure MS 通常被用于获取所述的Cy3,Cy5的,FITC每个平面和DAPIÇ hannels。     


 


(ⅳ)挪威使用流式细胞术全球掺入挠度


我们还介绍了使用流式细胞仪检测解决全球组蛋白更新的协议。尽管这种方法可以确定整体组蛋白动力学,但它无法在特定的基因组位点或染色质结构上检测组蛋白动力学。


 


组蛋白掺入的标签
25 在35毫米或60毫米的皿中的12个中散布细胞。     


26 通过双胸腺嘧啶核苷使细胞在G1 / S转变处同步。简而言之,用2 mM胸腺嘧啶核苷处理细胞16小时,用1 x PBS 洗涤3次,并通过更换培养基9 h从胸腺嘧啶核苷中释放出来,并再次用2 mM胸腺嘧啶核苷处理17 h。     


27 用SNAP-Cell Oregon Green(1 µM)在培养基中在5%CO 2中于37°C放置30分钟的标签,将预先存在SNAP标签的组蛋白标记。     


28 在37°C的5%CO 2 中的培养基中,用SNAP细胞块(10 µM)处理细胞30分钟,以掩盖不存在SNAP标记的组蛋白对非荧光SNAP底物的标记不足。     


关键步骤:该步骤对于避免对预先存在的组蛋白进行充分的初始标记是很重要的,因为这将导致新结合的组蛋白的检测不准确。


29 从细胞同步中释放6个培养皿,以检测12到22小时的时间点。保持同步其他6道菜以检测2到12小时的时间点。     


30 孵育细胞12小时。     


31 从细胞同步中释放其他6个培养皿,以检测2到12小时的时间点。     


每2 h收获32个细胞,并将细胞储存在-80°C下。     


暂停点:冻结的培养基和协议取决于细胞类型。


33 解冻细胞,并用预热的培养基洗涤一次。     


34 通过将含有SNAP-JF646(1 µM)的培养基添加到细胞沉淀中,标记新掺入的SNAP组蛋白。吸移细胞并在37°C水浴中孵育30分钟。     


35 在室温下将细胞以500 xg离心4分钟,并用预热的培养基洗涤两次。     


36 (可选)检测细胞周期的染料(例如,Hoechst 33342)也可以替代Fucci 细胞周期指示剂。     


37 使用流式细胞仪检测俄勒冈绿色(已有组蛋白),JF646(新加入的组蛋白),mCherry (S-G2期细胞周期指示剂)和TagBFP (G1期细胞周期指示剂)的表达水平     






方框1.手动成像分析全球现有和新合并的组蛋白的工作流程(另请参见图3)
 


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig3成像分析工作流程.jpg


图3.成像分析工作流程。显示了成像分析的概述,以检测预先存在的,新引入的组蛋白的强度,核染色(例如,DAPI 和Hoechst 33342)和EdU (S相标记)。


 


使用batch处理功能将所有图像集处理为最大强度Z投影,如下所示:
处理/批处理/宏。
在“输入...”中选择输入文件夹。
在“输出...”中选择输出文件夹。
(Ø 在ptional)将特定的名称为“文件名包含:”如果选择的文件夹需要特定的文件进行处理。
输入宏代码“ run(“ Z Project ...”,“ projection = [Max Intensity]”);“ 在脚本框中。
点击“处理”。
可以按以下方式进行手动处理:


通过将文件拖到FIJI 的主界面中或以下方式来打开四个图像:
File / Open / ,然后选择您的图像。


图像/堆栈/ Z项目/ ,然后选择投影类型“最大强度”。
定义目标区域(ROI),以通过使用核染色(如DAPI的DNA染色)制作掩模来测量先前存在的和新结合的组蛋白的强度,以及EdU 结合的强度。
复制显示DNA染色的图像,例如DAPI。


选择复制的DAPI 通道映像。
图片/副本... 
使用核染色强度定义核区域。


图像/调整/阈值;然后选择合适的阈值方法(可以选择默认值作为开始),选中“深色背景” 并应用。
处理/二进制/转换为遮罩。
(可选)处理/二进制/填充孔。
(可选)流程/二进制/分水岭(https://imagej.net/Nuclei_Watershed_Separation)。
分析/设置测量值,检查“面积”,“平均灰度值” 和“显示标签”,然后单击确定。
分析/分析粒子,在“大小(pixel ^ 2)” 和“圆度”中添加适当的数字以完成适当区域的选择,选中“ 添加到管理器” ,然后单击确定。
使用目标区域(ROI)在遮罩区域中测量DAPI 信号(DNA)。
选择频道图片。
在ROI Manager中单击“ 测量” 。
使用ROI测量俄勒冈州的绿色信号(预先存在的组蛋白)。
选择俄勒冈绿色通道图像。
在ROI Manager中单击“ 测量” 。
使用ROI测量TMR-STAR信号(新加入的组蛋白)信号。
  选择“ TMR-STAR”频道图像。
在ROI Manager中单击“ 测量” 。
使用ROI 测量EdU 信号(新生DNA)。
选择的EdU Ç hannel图像。
在ROI Manager中单击“ 测量” 。
保存结果并进行分析。
 


的BOX 2.分析工作流局部预先存在的,并使用颜色分析器或分析器RBG新掺入的组蛋白(小号EE也图5A)


在Fiji / ImageJ软件中安装插件。
通过将文件拖到FIJI 的主界面中或以下方式来打开所有要比较的图像集:
File / Open /,然后选择您的图像。


将这些图像合并为一个图像。
图像/颜色/合并通道。


将图像类型更改为RGB颜色。
图像/类型/ RGB颜色。


进行线条制作轮廓。
选择线工具。
排队。
插件/ RGB Profiler 。
 


影像分析(约1周)


步骤1-6,使用双胸腺嘧啶核苷和标记预先存在的组蛋白进行细胞同步以进行成像分析:4天。


步骤7-17,使用有丝分裂摇动细胞同步,并标记先前存在的和新掺入的组蛋白进行成像分析:4天。


步骤18-19,细胞的固定和通透性:1 h 。


步骤20,使用点击工具包标记EdU :1 h 。


步骤21,(可选)IF使用所需抗体确定特定染色质结构或因子下的组蛋白沉积:3 h 。


步骤22-23 ,DNA染色和盖玻片:隔夜。


步骤24,显微镜成像:1天。


 


              流式细胞仪分析(约1周)


步骤25,使用双胸腺嘧啶阻断剂扩散细胞并同步细胞,并标记预先存在的组蛋白:4天。


步骤32,从细胞同步释放并收获细胞:2天。


步骤s 34-37,新合并细胞的标记和流式细胞仪分析:2 h 。


 


数据一nalysis


 


我们介绍了使用FIJI / ImageJ(一种免费的开源图像处理软件包)进行成像的基本分析流程。如果您熟悉编程,那么使用ImageJ宏代码进行的自动分析将为大型数据集节省时间。此外,许多其它的成像处理方案(例如,Matlab的)可以是也有很大的选项处理巨大的数据集。


  使用流式细胞仪,例如LSR II(BD Bioscience),可以轻松完成使用流式细胞仪检测进行全局组蛋白更新的数据分析。FlowJo (BD Bioscience)等软件可以评估FCS数据集并允许可视化复杂的细胞计数数据集。在此协议中,我们可以使用整个细胞群体中具有单个细胞分辨率的荧光团总强度的平均值或中值来研究组蛋白掺入和沉积的时间。在此方案中显示通过标记预先存在的和新掺入的组蛋白以及细胞周期标记物标记的荧光强度的直方图,可以帮助可视化全局组蛋白动力学的时间,包括解散先前存在的组蛋白和合并新合成的组蛋白。细胞周期。


 


预期结果


在此协议中,我们总结了先前发表的关于HEK293T细胞中组蛋白H3.1和macroH2A1.2的细胞周期特异性动力学的结果,以表明SNAP标记脉冲追踪系统可靠地工作,并说明了如何分析成像数据集(Sato 等人,2019)。


  尽管我们在此方案中说明了三个部分(图1 ),包括构建体的生成,用于成像的样品制备和用于流式细胞术的样品制备,但我们主要描述了后两个的详细方案。我们还将说明如何使用ImageJ / FIJI分析成像数据集(图3 和方框1 )。在这里,我们介绍了使用成像方法检测SNAP组蛋白H3.1(图4 )的细胞周期中SNAP脉冲追踪检测的示例。我们展示了两种类型的细胞同步性,双胸苷阻滞和有丝分裂摆脱,以调查在S-G2或G1期的组蛋白掺入和解离。如图4A-4B所示,在HEK 293T中稳定表达的先前存在的和新掺入的SNAP-H3.1进行了标记。还执行了EdU 单击检测以确认成功的单元同步。单个细胞中的整体组蛋白掺入率取决于新掺入的组蛋白(TMR-Star)的像素强度平均值,该平均值通过细胞核中先前存在的组蛋白(俄勒冈州绿)的像素强度平均值进行标准化。使用这种方法,我们在S-G2阶段检测到H3.1掺入,这被称为DNA复制依赖性沉积(图4C-4D )。


  我们还说明了使用RGB Profiler在成像分析中共定位的检测(图5A 和方框2 )。在这里,我们在HEK在293T灭活X染色体(Xi)的C存在新掺入的macroH2A组蛋白变体的共定位厄尔(图小号5A-5B )。如图4A 所示标记先前存在的和新掺入的SNAP-macroH2A1.2,并且通过与先前存在的SNAP-macroH2A1.2共定位来确定Xi中SNAP-macroH2A1.2在细胞周期中的掺入。如图5B 所示,在G1阶段将macroH2A1.2结合到Xi中。


  整体组蛋白的结合和解离也可以通过流式细胞仪检测。我们展示了SNAP-macroH2A1.2脉冲追踪方法,并通过流式细胞仪检测了全球SNAP-macroH2A动态,如图5C-5D所示。在这里,我们同步了G1 / S边界处的细胞,并用俄勒冈绿色标记了先前存在的SNAP-macroH2A1.2,然后从细胞同步释放后每两小时检测到新并入的SNAP-macroH2A1.2。尽管我们使用Fucci 细胞周期传感器确定了细胞周期的过渡(图5C [ 两个右图] 和5D [ 顶部] ),但也可以选择其他细胞周期指示剂,例如Hoechst 33342 DNA染色(图6 )。直方图显示细胞群体的荧光强度。先前存在的SNAP-macroH2A1.2(俄勒冈绿色,图5C [ 左] 和5D [ 中] )的信号强度转变表明,当细胞进入有丝分裂时,信号强度下降,但是几乎没有SNAP-macroH2A与染色质解离在整个细胞周期中 新合并的SNAP-macroH2A1.2(JF646,图5C [ 左下] 和5D [ 下] )的过渡。


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig4 Histone detection.jpg


图4.在特定细胞周期中检测整体先前存在的和新掺入的组蛋白H3.1。A. 在BD中分析S / G2和G1相标记组蛋白的图解。(左图)为了标记新掺入的S / G2相中的组蛋白,稳定表达SNAP标签的H3的HEK 293T细胞在G1 / S相边界处被双胸腺嘧啶核苷同步化,如TH所示。预先存在的组蛋白用SNAP-俄勒冈绿色(绿色箭头)标记,随后使用非荧光SNAP-Block试剂封闭,以避免标记不足。使细胞从同步释放,前进到G2 / M转变,直到使用RO-3306(CDK1 / cyclin B1和CDK1 / cyclin A抑制剂,显示为RO)在G2 / M边界同步,然后重新整合SNAP标记的组蛋白用SNAP-TMR Star(红色箭头)标记。(右图)为检测G1期新掺入的组蛋白,在诺考达唑处理12小时后通过摇晃收集有丝分裂细胞,并铺在盖玻片上(显示为Noc )。后2 小时,预先存在的SNAP-H3分别标记有俄勒冈绿和与阻挡试剂处理。从同步释放细胞,并使用双胸腺嘧啶阻断进行到G1 / S过渡,然后用SNAP-TMR Star标记新掺入SNAP标签的组蛋白。从第一次同步释放后,还用EdU 处理细胞,直到第二次同步,从而鉴定出已进行DNA合成的细胞。B. S / G2或G1阶段中已存在和新合成的带有SNAP标签的H3.1的代表性图像。标度b ar = 10 µm。图像分析显示在S-G2相(CD )期间组蛋白H3的掺入率更高。C. 在S-G2和G1细胞周期阶段中组蛋白结合率的频率分布。组蛋白掺入率是通过在S / G2和G1阶段中核中红色(新掺入的组蛋白)的平均像素强度和绿色(已存在的组蛋白)的平均像素强度进行归一化计算的。EdU 标记确认单元同步成功。在以下分析中,进行了Hoechst 33342标记以定义核的面积。D. 在S-G2和G2阶段中组蛋白掺入率的定量。每个点表示来自单个细胞的掺入率,如图4C中所述。误差线表示与单个单元格数量的一个标准偏差(n =每个数据集上指示的数量。* P < 0.0001。P 值是使用两尾不成对t 检验确定的。


 


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig5 mH2A local incorporation2.jpg


图5.在特定细胞周期中检测局部和全局预先存在的和新掺入的组蛋白macroH2A1.2。A. 使用RBG Profiler插件(https://imagej.nih.gov/ij/plugins/rgb-profiler.html,Color Profiler插件(https://imagej.nih.gov/ij/plugins/ color-profiler.html)还提供了相同的功能),以解决新合并的组蛋白与预先存在的组蛋白的共定位问题。也可以通过使用IF检测染色质结构来解决特定染色质结构上细胞周期特异性组蛋白的掺入。在这里,我们展示了检测与现有和新加入的组蛋白共定位的示例。B. 使用Cell Profiler进行图像分析的示例。在S-G2(上部)和G1(下部)阶段中,将预先存在的(俄勒冈绿色:绿色),新合并的(TMR:红色)SNAP标签的macroH2A的左侧图像合并为单个图像,并在两个非活动X上进行共定位在HEK 293T细胞中确定了染色体(Xi)。在右上方和右下方的面板中,显示了沿既有图像和新合并图像中的白线测量的信号强度。C. 现有细胞(左:俄勒冈州绿)和新整合的macroH2A(左第二:JF646),S-G2(右第二:mCherry 标记的hGeminin )和G1(右:TagBFP 标记的hCdt1)细胞的流式细胞仪分析循环标记来自Fucci 传感器。直方图表示从G1 / S边界同步释放后的每个时间点(2-22小时)的荧光强度。从Fucci 细胞周期传感器估计每个时间点的细胞周期。D. 之前存在的(中间:俄勒冈州绿色)和新加入的macroH2A(底部:JF646)以及S-G2和G1(顶部:mCherry 标签的hGeminin 和TagBFP 标签的hCdt1)的荧光强度的中值绘制流式细胞仪分析。误差线表示95%的置信区间。


 


D:\ Reformatting \ 2020-6-1 \ 2003151--1462 Hanae Sato 799386 \ Figs jpg \ Fig6 Hoechst.jpg


图6 。使用Hoechst 33342染色和流式细胞术检测确定细胞周期。A. 如步骤25-36所述,对细胞周期(左:Hoechst 33342)和已存在并标记并从细胞同步释放的SNAP-macroH2A(右:俄勒冈绿色)进行流式细胞仪分析。直方图表示从G1 / S边界同步释放后的每个时间点(2-22 h)的荧光强度。每个时间点的细胞周期可以通过细胞群中Hoechst 33342的强度转变来确定。B. 绘制流式细胞术分析的DNA染色的荧光强度的中值(上图:Hoechst 33342)和预先存在的SNAP-macorH2A1.2(中:俄勒冈州绿)。误差线表示95%的置信区间。


 


菜谱


 


SNAP组蛋白表达载体的克隆及其稳定表达的细胞系
需要通过常规克隆方法克隆编码带有SNAP标签的所需组蛋白的质粒。pSNAPf 载体(NEB的产品目录号N9183S)可用于建立您的构建体并稳定表达细胞系,因为它含有新霉素抗性基因,可进行有效的药物选择。


固定缓冲液
1 x PBS中4%(wt / vol)PFA


注意:该溶液可以由32%(wt / vol)PFA 制备。避光长达2年。


通透缓冲液
0.5%Triton在1 x PBS中


阻塞缓冲区
1 x PBS中的3%BSA


排序缓冲区
1毫米EDTA


25 mM HEPES pH 7.0


1 x DPBS中的1%FBS(热灭活)


 


致谢


 


这项工作得到了美国国立卫生研究院(NIH)资助R01 DA030317(JMG)的支持。我们感谢Greally 和Singer实验室的成员进行的讨论,爱因斯坦FACS和Genomics核心。我们也感谢L. Lavis 的SNAP-JF646。该协议是从以前的工作中采用的(Sato et al。,2019)。


  作者贡献声明:HS设计并进行了实验并分析了数据。HS,JMG和RHS撰写了手稿。JMG和RHS监督了这项研究。


 


利益争夺


 


作者宣称没有相互竞争的经济利益。


 


参考文献


 


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Copyright: © 2020 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. Sato, H., Singer, R. H. and Greally, J. M. (2020). Quantitative Kinetic Analyses of Histone Turnover Using Imaging and Flow Cytometry. Bio-protocol 10(17): e3738. DOI: 10.21769/BioProtoc.3738.
  2. Sato, H., Wu, B., Delahaye, F., Singer, R. H. and Greally, J. M. (2019). Retargeting of macroH2A following mitosis to cytogenetic-scale heterochromatic domains. J Cell Biol 218(6): 1810-1823.
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