An Efficient Protocol for CUT&RUN Analysis of FACS-Isolated

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An Ecient Protocol for CUT&RUN Analysis of

FACS-Isolated Mouse Satellite Cells

Kamar Ghaibour, Joe Rizk, Claudine Ebel, Tao Ye, Muriel Philipps, Valérie

Schreiber, Daniel Metzger, Delphine Duteil

To cite this version:

Kamar Ghaibour, Joe Rizk, Claudine Ebel, Tao Ye, Muriel Philipps, et al.. An Ecient Protocol

for CUT&RUN Analysis of FACS-Isolated Mouse Satellite Cells. Journal of visualized experiments :

JoVE, 2023, 197 (e65215), pp.1-21. �10.3791/65215�. �hal-04192479�

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An Efficient Protocol for CUT&RUN Analysis of FACS-

Isolated Mouse Satellite Cells

Kamar Ghaibour

*,1

, Joe Rizk

*,1

, Claudine Ebel

, Tao Ye

, Muriel Philipps

, Valérie Schreiber

, Daniel Metzger

, Delphine

Duteil

CNRS, Inserm, IGBMC UMR 7104- UMR-S 1258, Université de Strasbourg

These authors contributed equally

Corresponding Author

Delphine Duteil

[email protected]

Citation

Ghaibour, K., Rizk, J., Ebel, C., Ye, T.,

Philipps, M., Schreiber, V., Metzger, D.,

Duteil, D. An Efficient Protocol for

CUT&RUN Analysis of FACS-Isolated

Mouse Satellite Cells. J. Vis. Exp. (197),

e65215, doi:10.3791/65215 (2023).

Date Published

July 7, 2023

DOI

10.3791/65215

URL

jove.com/video/65215

Abstract

Genome-wide analyses with small cell populations are a major constraint for studies,

particularly in the stem cell field. This work describes an efficient protocol for the

fluorescence-activated cell sorting (FACS) isolation of satellite cells from the limb

muscle, a tissue with a high content of structural proteins. Dissected limb muscles

from adult mice were mechanically disrupted by mincing in medium supplemented with

dispase and type I collagenase. Upon digestion, the homogenate was filtered through

cell strainers, and cells were suspended in FACS buffer. Viability was determined

with fixable viability stain, and immunostained satellite cells were isolated by FACS.

Cells were lysed with Triton X-100 and released nuclei were bound to concanavalin

A magnetic beads. Nucleus/bead complexes were incubated with antibodies against

the transcription factor or histone modifications of interest. After washes, nucleus/

bead complexes were incubated with protein A-micrococcal nuclease, and chromatin

cleavage was initiated with CaCl

. After DNA extraction, libraries were generated and

sequenced, and the profiles for genome-wide transcription factor binding and covalent

histone modifications were obtained by bioinformatic analysis. The peaks obtained for

the various histone marks showed that the binding events were specific for satellite

cells. Moreover, known motif analysis unveiled that the transcription factor was bound

to chromatin via its cognate response element. This protocol is therefore adapted to

study gene regulation in adult mice limb muscle satellite cells.

Introduction

Skeletal striated muscles represent on average 40% of the

weight of the total human body

. Muscle fibers exhibit a

remarkable capacity for regeneration upon injury, which is

described by the fusion of newly formed myocytes and

the generation of new myofibers that replace the damaged

ones

. In 1961, Alexander Mauro reported a population of

mononuclear cells that he termed as satellite cells

. These

stem cells express the transcription factor paired box 7

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(PAX7), and are located between the basal lamina and the

sarcolemma of muscle fibers

. They were reported to express

the cluster of differentiation 34 (CD34; a hematopoietic,

endothelial progenitor and mesenchymal stem cell marker),

integrin alpha 7 (ITGA7; a smooth, cardiac and skeletal

muscle marker), as well as the C-X-C chemokine receptor

type 4 (CXCR4; a lymphocyte, hematopoietic, and satellite

cell marker)

. In basal conditions, satellite cells reside in a

particular microenvironment that keeps them in a quiescent

state

. Upon muscle damage, they become activated,

proliferate, and undergo myogenesis

. However, contributing

only to a minor fraction of the total number of muscle cells,

their genome-wide analyses are particularly challenging,

especially under physiological settings (<1% of total cells).

Various methods for chromatin isolation from satellite

cells have been described, which involve chromatin

immunoprecipitation followed by massive parallel sequencing

(ChIP-seq) or cleavage under targets and tagmentation

(CUT&Tag) experiments. Nevertheless, these two techniques

present some significant limitations that remain unchallenged.

Indeed, ChIP-seq requires a high amount of starting

material to generate enough chromatin, a large proportion

of which is lost during the sonication step. CUT&Tag is

more appropriate for low cell number, but generates more

off-target cleavage sites than ChIP-seq due to the Tn5

transposase activity. In addition, since this enzyme has

a high affinity for open-chromatin regions, the CUT&Tag

approach might be preferentially used for analyzing histone

modifications or transcription factors associated with actively

transcribed regions of the genome, instead of silenced

heterochromatin

8 , 9

Presented here is a detailed protocol that allows the

isolation of mouse limb muscle satellite cells by FACS

for cleavage under targets and release using nuclease

(CUT&RUN)

10 , 11

analysis. The various steps involve the

mechanical disruption of tissue, cell sorting, and nuclei

isolation. The method's efficiency, regarding the preparation

of a viable cell suspension, was demonstrated by performing

CUT&RUN analysis for covalent histone modifications and

transcription factors. The quality of isolated cells makes

the described method particularly attractive for preparing

chromatin that captures the native genomic occupancy

state faithfully, and is likely to be suitable for capturing

the chromosome conformation in combination with high-

throughput sequencing at specific loci (4C-seq) or at genome-

wide levels (Hi-C).

Protocol

Mice were kept in an accredited animal house, in compliance

with National Animal Care Guidelines (European Commission

directive 86/609/CEE; French decree no.87-848) on the use

of laboratory animals for research. Intended manipulations

were submitted to the Ethical committee (Com'Eth,

Strasbourg, France) and to the French Research Ministry

(MESR) for ethical evaluation and authorization according to

the 2010/63/EU directive under the APAFIS number #22281.

1. Preparation of cell suspension for isolation

of satellite cells by fluorescence-activated cell

sorting (FACS) (Figure 1 )

1. Isolation of muscle tissue

1. Decontaminate the tools for muscle dissection,

including forceps, scalpels, and scissors, using a

cleaning agent (Table 1), and rinse thoroughly with

distilled water.

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2. Prepare two 2 mL tubes (Table 1), each containing

1 mL of muscle isolation buffer, and place them on

ice for collecting harvested muscles.

3. Sacrifice two 10-week-old C57/Bl6J male mice by

asphyxiation followed by cervical dislocation.

Spray 70% ethanol over each entire mouse. Peel off

the skin from the hind limb using forceps. Dissect out

all the limb muscles surrounding the femur, tibia, and

fibula (approximately 1 mg of muscles per mouse).

NOTE: The satellite cells number decreases after 15

weeks of age.

4. Place the harvested limb muscles into 2 mL tubes

containing 1 mL of muscle isolation buffer prepared

in step 1.1.2. Collect the muscles from the second

mouse following the same procedure. Mince the

harvested muscles with scissors on ice until smaller

than 1 mm

fragments are obtained.

NOTE: Muscles were collected and minced mainly

as described

. Perform tissue digestion either

following step 1.2 or 1.3.

2. Tissue digestion with collagenase enzyme

1. Transfer the minced muscle suspension from the

two mice by pouring them into a 50 mL tube

(Table 1) containing 18 mL of muscle isolation buffer

(supplemented with 5 mL [5 U/mL] of dispase and 5

mg of type I collagenase) (Table 1).

2. Close the tube tightly and seal it with laboratory film

(Table 1). Place it horizontally in a shaking water

bath (Table 1) at 37 °C at 100 rpm for 30 min.

3. After 30 min, add 5 mg of type I collagenase. Keep

tubes for another 30 min under agitation in a shaking

water bath at 37 °C at 100 rpm.

4. Upon digestion, pipette the muscle suspension up

and down 10 times with a 10 mL pipette to improve

the efficiency of dissociation. Centrifuge at 4 °C at

400 x g for 5 min. A clear pellet will be visible at the

bottom of the tube. Discard the supernatant using a

10 mL pipette, leaving 5 mL of medium in the tube.

NOTE: Leaving the medium helps avoid stressing

the cells. Add 10 mL of fresh muscle isolation buffer

and resuspend the pellet by pipetting up and down

with a 10 mL pipette.

5. Place 100 µm, 70 µm, and 40 µm cell strainers (one

of each kind) (Table 1) on open 50 mL tubes. Pipette

the suspension onto the successive cell strainers

(100 µm, 70 µm, 40 µm) and collect the flowthrough

into the 50 mL tubes, containing cells below 40 µm.

6. Centrifuge the suspension at 4 °C at 400 x g for 5

min. Discard the supernatant using a 10 mL pipette

until 2 mL remain, and then use a 0.2-1 mL pipette

until 100-200 µL remain. Resuspend the pellet in 2

mL of red blood cell lysis buffer (Table 2). Incubate

on ice for 3 min.

7. Centrifuge at 4 °C at 400 x g for 5 min and discard the

supernatant using a 20-200 µL pipette. Resuspend

the cells in 100 µL of cold FACS buffer (Table 2).

Place on ice.

3. Alternative method for tissue digestion with Liberase

thermolysin low (TL) enzyme

1. Follow the descriptions in step 1.1. for tissue

isolation.

2. For Liberase-mediated tissue disaggregation,

harvest muscles in 2 mL of Roswell Park Memorial

Institute (RPMI) isolation buffer (Table 2), instead of

muscle isolation buffer described in step 1.1.2.

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3. Transfer the minced muscle suspension from the

two mice by pouring them into a 50 mL tube

(Table 1) containing 18 mL of RPMI isolation buffer

supplemented with 300 or 600 µL of Liberase TL at

5 mg/mL (Table 1) (i.e., 0.083 mg/mL and 0.167 mg/

mL final concentrations, respectively)

4. Close the tube tightly and seal it with laboratory film

(Table 1). Place it horizontally in a shaking water

bath (Table 1) at 37 °C at 100 rpm for 30 min.

5. Upon digestion, pipette the muscle suspension up

and down 10 times with a 10 mL pipette to dissociate

and improve the efficiency of dissociation.

6. Centrifuge at 4 °C at 400 x g for 5 min. A clear pellet

will be visible at the bottom of the tube. Discard the

supernatant using a 10 mL pipette, leaving 5 mL

of medium in the tube. Leaving the medium helps

avoid stressing the cells. Add 10 mL of fresh RPMI

isolation buffer, and resuspend the pellet by pipetting

up and down with a 10 mL pipette.

7. Place 100 µm, 70 µm, and 40 µm cell strainers (one

of each kind) (Table 1) on open 50 mL tubes.

8. Pipette the suspension onto successive cell

strainers (100 µm, 70 µm, 40 µm) and collect the

flowthrough into the 50 mL tubes, containing cells

below 40 µm.

9. Centrifuge the suspension at 4 °C at 400 x g for 5

min. Discard the supernatant using a 10 mL pipette

until 2 mL remain, and then use a 0.2-1 mL pipette

until 100-200 µL remain.

10. Resuspend the pellet in 2 mL of red blood cell lysis

buffer (Table 2). Incubate on ice for 3 min.

11. Centrifuge at 4 °C at 400 x g for 5 min and discard

the supernatant using a 20-200 µL pipette.

12. Resuspend the cells in 100 µL of cold FACS buffer

(Table 2). Place on ice.

4. Preparation of cell suspension for FACS isolation

1. Transfer 10 µL of the cell suspension obtained in

step 1.2.12 into a fresh 1.5 mL tube. This sample

will constitute the unstained control or the negative

control (Figure 2). Add 190 µL of FACS buffer,

transfer to a 5 mL tube (Table 1), and store on ice.

2. Centrifuge the remaining 90 µL of the cell

suspension obtained in step 1.2.12 at 4 °C at 400 x g

for 5 min and discard the supernatant using a pipette

(20-200 µL tip volume). Incubate the cells with 400

µL of fixable viability stain (Table 3) diluted in serum-

free Dulbecco's modified Eagle medium (DMEM) for

15 min at room temperature (RT).

3. Wash the cells by centrifugation at 4 °C at 400 x g for

5 min and add 100 µL of FACS buffer. Gently invert

the tubes three times and centrifuge again at 4 °C at

400 x g for 5 min.

4. During the centrifugation time, prepare 100 µL of

a master mix of primary antibodies coupled to

fluorophores and directed against CD11b, CD31,

CD45, TER119, CD34, ITGA7, and CXCR4 (Table

3), diluted in FACS buffer.

5. Centrifuge at 400 x g for 5 min at 4 °C, discard

the cell supernatant using a pipette (20-200 µL tip

volume), and add the 100 µL antibody mix. Gently

invert the tube three times. Do not vortex. Incubate

in the dark on ice for 30 min.

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6. Centrifuge at 400 x g for 5 min at 4 °C. Discard the

supernatant using a 20-200 µL pipette and add 500

µL of 1x phosphate-buffered saline (PBS) to wash

the cells. Gently invert the tube three times. Re-

centrifuge at 400 x g for 5 min at 4 °C and discard

the supernatant using a 20-200 µL pipette.

7. Resuspend the cell pellet in 500 µL of FACS buffer

and transfer the suspension to a 5 mL tube.

NOTE: the cell suspension obtained from Liberase

digestion is processed in the same way.

5. Satellite cells selection by FACS

1. Briefly vortex the cell suspension (2-5 sec) and

process the cells on a flow cytometer equipped with

a 100 µm nozzle (Table 1).

2. Determine the various gate sizes based on the

unstained sample stored at step 1.4.1 (Figure 2).

3. Coat a 5 mL tube with 1 mL of pure fetal calf serum

(FCS) to improve cell collection and add 500 µL of

FACS buffer.

4. Exchange the unstained sample with the antibody-

labelled sample.

5. Select the population of interest according to the

forward scatter area (FSC-A) and side scatter area

(SSC-A) (Figure 3A), and remove doublet cells

with the FSC-A and forward scatter height (FSC-H)

(Figure 3B)

6. Identify living cells with fixable viability stain negative

staining (Figure 3C).

7. Select negative cells for CD31, CD45, TER119, and

CD11b (Figure 3D).

8. To identify satellite cells, select first the cells that are

positive for CD34 and ITGA7 (Figure 3E), and then

select the CXCR4-positive cells on the CD34- and

ITGA7-selected population (Figure 3F).

9. Collect the selected cells (between 40,000 and

80,000 cells, according to the quality of the

preparation) in the 5 mL coated tube containing 500

µL of FACS buffer.

2. Validation of the isolated population in tissue

culture

1. Slide coating with hydrogel

1. Dilute 280 µL of a pure hydrogel human embryonic

stem cell (hESC) qualified matrix (Table 1) in 12 mL

of serum-free DMEM/F12 medium.

2. Coat a chamber slide (Table 1) with the hydrogel

solution, and incubate it overnight at 4 °C.

3. The next day, incubate the chamber slide at 37 °C

and 5% CO

for 1 h before cell seeding.

2. Cell growth and differentiation

1. Plate out approximately 20,000 cells, obtained from

step 1.5.9, per well, and grow them in growth

medium (Table 2) for 5 days. Take phase-contrast

images using a brightfield microscope (Figure 4A),

before processing them for immunofluorescence

analysis to ensure the quality of the preparation

(Figure 4B).

2. To induce myogenesis, grow amplified satellite cells

from step 2.2.1 in myogenic medium (Table 2) for

an additional 7 days. Take phase-contrast images

using brightfield microscope (Figure 4C) before

processing them for immunofluorescence analysis

to ensure the quality of the preparation (Figure 4D).

3. Immunocytofluorescence analysis

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1. Gently remove the medium, wash the cells that

were cultured on chamber slide with 100 µL of

1x PBS twice, and fix them with 100 µL of 4 %

paraformaldehyde (PFA) at RT for 1 h.

NOTE: This step should be performed with care.

A small volume of medium should always be kept

in the chamber to prevent stressing the cells, and

the PBS should be poured through the walls of the

chamber.

2. Wash the cells three times with 100 µL of 1x

PBS, supplemented with 0.1% Tween 20 (PBST) to

permeabilize the cell membranes.

3. Block unspecific signals by incubation in 100 µL of

1x PBST supplemented with 5% FCS (PBST-FCS)

at RT for 1 h.

4. Incubate the cells with 100 µL of a master mix of anti-

PAX7 and anti-dystrophin (DMD) antibodies (diluted

in 1x PBST-FCS) at 4 °C overnight, to detect satellite

cells and myofibers, respectively.

5. Wash the cells three times with 100 µL of 1x

PBST, and incubate them with 100 µL of goat anti-

mouse Cy3 or goat anti-rabbit Alexa 488 secondary

antibodies (Table 3) diluted in 1x PBST-FCS at RT

for 1 h.

6. Dissociate the chamber wells from the slide using

the equipment provided by the supplier, add 20 µL

of aqueous mounting medium with 4′,6-diamidino-2-

phenylindole (DAPI), and cover the slide with a

coverslip (Table 1).

7. Observe and capture the image of the stained cells

with a confocal microscope.

8. Process the images using image analysis software

(Figures 4B,D).

3. CUT&RUN analysis

1. Sample preparation for CUT&RUN analysis on FACS-

isolated satellite cells

NOTE: CUT&RUN was performed essentially as

described

10 , 15

. The buffer composition is presented in

Table 2.

1. For the CUT&RUN assay, use approximately 40,000

of the cells obtained in method 1, step 1.5.9, per

sample/antibody that must be tested.

2. Centrifuge the FACS-isolated satellite cells at RT at

500 x g for 10 min, then discard the supernatant

using a pipette (20-200 µL tip volume).

3. Wash the cells with 1 mL of 1x PBS, centrifuge at RT

at 500 x g for 5 min, discard the supernatant using a

pipette (0.2-1 mL tip volume), and resuspend them

in 1 mL of cold nuclear extraction buffer (Table 2).

Incubate on ice for 20 min.

4. During incubation, prepare one 1.5 mL tube

containing 850 µL of cold binding buffer (Table 2)

and add 20 µL of concanavalin A-coated magnetic

beads per sample (Table 1).

5. Wash the beads twice with 1 mL of cold binding

buffer using a magnetic rack (Table 1). For each

wash or buffer change throughout the procedure, let

the beads accumulate at the side of the tube on the

magnetic rack for 5 min before removing the cleared

supernatant with a pipette (0.2-1 mL tip volume).

Then, resuspend gently in 300 µL of cold binding

buffer.

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6. Centrifuge the nuclei at 4 °C at 600 x g for 5 min

and resuspend them gently in 600 µL of nuclear

extraction buffer. Gently mix the 600 µL of extracted

nuclei with the 300 µL of concanavalin A bead slurry,

and incubate at 4 °C for 10 min.

7. Remove the supernatant using a magnetic rack, as

described in step 3.1.4, and resuspend the bead-

bound nuclei gently with 1 mL of cold blocking buffer

(Table 2). Incubate at RT for 5 min.

8. Remove the supernatant using a magnetic rack

and wash the bead-bound nuclei twice with 1 mL

of cold wash buffer (Table 2). During the second

wash, equally split the bead-bound nuclei into 1.5

mL tubes. Each tube will be treated with a specific

antibody in the following step.

NOTE: In this example, 250 µL of bead-bound nuclei

were split in four 1.5 mL tubes.

9. Separate the supernatant using a magnetic rack, as

described in step 3.1.4, and aspirate with a pipette.

Gently resuspend the nucleus/bead complexes with

a specific primary antibody (Table 3), or an IgG of

another species (here rabbit) diluted in 250 µL of

cold wash buffer. Incubate at 4 °C overnight with

gentle agitation.

NOTE: The antibodies used here are directed

against AR, H3K4me2, and H3K27ac.

10. Remove the supernatant with a magnetic rack, as

described in step 3.1.4, wash the bead-bound nuclei

twice with 1 mL of cold wash buffer, and resuspend

in 100 µL of cold wash buffer.

11. Dilute protein A-micrococcal nuclease at 1.4 ng/µL

in 100 µL per sample of cold wash buffer.

12. Add 100 µL of protein A-micrococcal nuclease to the

100 µL of sample obtained in 3.1.11 and incubate at

4 °C for 1 h with agitation.

13. Remove the supernatant with a magnetic rack, as

described in step 3.1.4, wash twice with 1 mL of cold

wash buffer, and resuspend the bead-bound nuclei

in 150 µL of cold wash buffer.

14. To initiate DNA cleavage, add 3 µL of 100 mM

of CaCl

to the 150 µL of sample, mix quickly

by flicking, and incubate on ice for 30 min. Stop

the reaction by adding 150 µL of stop buffer and

incubate at 37 °C for 20 min to digest the RNA and

release the DNA fragments.

15. For DNA extraction, centrifuge the samples at

16,000 x g at 4 °C for 5 min.

16. Transfer the supernatant to a new microfuge tube

and discard the pellet and beads.

17. Add 3 µL of 10% sodium dodecyl sulfate (SDS) and

2.5 µL of 20 mg/mL proteinase K. Mix by inversion.

Incubate for 10 min at 70 °C (no shaking).

18. Add 300 µL of phenol/chloroform/isoamyl alcohol,

vortex, transfer to 2 mL phase-lock tubes (pre-

spinned for 5 min at 16,000 x g), and centrifuge for

5 min at 16,000 x g at 4 °C.

19. Add 300 µL of chloroform to the same tube and

centrifuge for 5 min at 16,000 x g at 4 °C. Collect the

supernatant (~300 µL) with a pipette (0.2-1 mL tip

volume) and transfer into a new 1.5 mL tube.

20. Add 1 µL of glycogen (20 mg/mL concentration).

21. Add 750 µL of 100% ethanol and precipitate

overnight at -20 °C.

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22. Pellet the DNA by centrifugation for 15 min at 16,000

x g at 4 °C. Wash the pellet with 1 mL of 100%

ethanol, centrifuge for 5 min at 16,000 x g, discard

the supernatant, centrifuge for 30 s at 16,000 x g,

and remove the liquid with a pipette (20-200 µL tip

volume).

23. Air-dry the pellet for ~5 min. Resuspend in 25

µL of 1 mM Tris-HCl (pH 8) and 0.1 mM

ethylenediaminetetraacetic acid (EDTA; pH 8).

2. Bioinformatics analysis

1. Prepare libraries from immunocleaved DNA and

sequence them as paired-end 100 bp reads with the

help of the genomic platform as described

2. Remove reads overlapping with the ENCODE

blacklist region (V2) and separate the remaining

reads into two groups: fragment size <120 bp

(without nucleosome, in general for transcription

factors) and fragment size >150 bp (with

nucleosomes, normally for histone marks). Map

to the mm10 reference genome using Bowtie 2

(v2.3.4.3)

3. Generate bigwig files with bamCoverage (deeptools

3.3.0: bamCoverage --normalizeUsing RPKM --

binSize 20).

4. Retain uniquely mapped reads for further analysis.

5. Generate raw bedgraph files with

genomeCoverageBed (bedtools v2.26.0).

6. Use the SEACR 1.3 algorithm (stringent option) for

the peak calling. Load the target data bedgraph

file in UCSC bedgraph format that omits regions

containing zero signal, and control (IgG) data

bedgraph file to generate an empirical threshold for

peak calling

7. Perform a Pearson correlation analysis with

deeptools to determine the similarity between

the samples

. Use the command line

multiBamSummary bins --bamfiles file1.bam

file2.bam -o results.npz, followed by

plotCorrelation -in results.npz --corMethod

pearson --skipZeros --plotTitle "Pearson

Correlation of Read Counts" --whatToPlot

heatmap --colorMap RdYlBu --plotNumbers

-o heatmap_PearsonCorr_readCounts.png --

outFileCorMatrix PearsonCorr_readCounts.tab.

8. Visualize the genome-wide intensity profiles with

IGV

using bedgraph files and the bed file peaks

obtained from SEACR.

9. Use HOMER for peak annotation and motif

10. Finally, compare the datasets with previously

published ones with ChIP-Atlas Peak browser to

visualize them on IGV, and/or enrichment analysis

using the SEACR-generated bed files as an input

dataset

Representative Results

Satellite cells from mouse skeletal muscles were isolated by

combining the protocols of Gunther et al. (hereafter Protocol

and of Liu et al.

(hereafter Protocol 2). Since non-

digested muscle fibers were observed after digestion when

using the concentration of collagenase and dispase proposed

in Protocol 1, the quantity of enzymes was increased to

improve muscle fiber dissociation, as described in steps 1.2.1

and 1.2.3. As indicated in Protocol 2, the samples were

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subjected to a gentle agitation in a water bath to maintain

cell viability. We performed filtration through cell strainers,

as mentioned in Protocol 1, and incubation with red blood

cell lysis buffer (see steps 1.2.7 to 1.2.10). In Protocol 1,

cells were loaded on a Percoll density gradient of 30%/70%

to isolate mononucleated cells at the interphase, which may

have led to the loss of cells of interest. Thus, this step was

omitted, as proposed in Protocol 2.

FSC-A versus SSC-A gating was used to identify

mononucleated cells based on size (FSC) and granularity

(SSC) (Figure 3A). Debris were excluded by ignoring events

below 40 K on the FSC-A axis, and 38.8% ± 3.6% of cells

were selected. FSC-A versus forward scatter height (FSC-H)

gating density plots were used for doublet exclusion (Figure

3B). After selecting the cells negative for the fixable viability

stain marker, an average of 34.3% ± 7.7% of live cells was

obtained (Figure 3C).

The percentage shown in the dot plot in Figure 3C, 75.4%,

corresponds to the percentage of single living cells, calculated

from the parent cell population, which is in this case the

singles. 95.7% of single cells are obtained from the live

events which make up 35% of the total events. Thus, the

percentage 34.3% obtained on average, is calculated from

the total events and not the parent populations.

About 3% of single living cells were negative for leukocyte-

(CD45), monocyte- (CD11b), endothelial- (CD31), and

erythroid-specific (TER119) markers (Figure 3D). CD11b/

CD45/CD31/TER119 negative cells were then selected

according to their expression of CD34 (hematopoietic,

endothelial progenitors, and mesenchymal stem cells) and

ITGA7 (cardiac, smooth, and skeletal muscle cells) markers

(Figure 3E). A final gating for CXCR4 (lymphocytes,

hematopoietic, and satellite cells) was performed to select

putative satellite cells (Figure 3F). From the CD34+/ITGA7+

cells, ~80% were found to be positive for CXCR4, which

represents an average of 1% ± 0.15% of total single living

cells, and an absolute number of 60,000 ± 14,000 putative

satellite cells per mouse limb muscles among 14 independent

experiments. An additional resorting assessment on the

CXCR4+ cell fraction revealed that about 80% of living cells

were obtained after post-sorting, of which almost 70% were

CXCR4+ (Figure S1), showing the high viability and purity of

this FACS-isolated cell population.

Since various studies compared the efficiency of collagenase

and Liberase TL enzymes for cell isolation

24 , 25

, these two

digestion methods have been processed in parallel. With 300

µL of Liberase TL, the digestion was less efficient than with

collagenase, as undigested fibers remained. In addition, more

cell debris and large events were observed (Figures S2A,B),

and only 17.3% of single living cells on average were obtained

after FVS 780 selection (Figure S2C). An additional concern

with Liberase digestion was the low number of CD34+/

ITGA7+ cells compared to collagenase (Figures S2D-S2E),

even though CXCR4 gating was similar (Figure S2F). With

600 µL of Liberase TL, the digestion was more efficient.

However, the amount of cell debris remained elevated and

the cell viability substandard (16.3%) (Figure S3). Thus,

digestion with Liberase TL was less efficient for satellite cell

isolation.

When seeded, more than 70% of the CD34+/ITGA7+/

CXCR4+ cells (Figure 4A) expressed PAX7 (Figure 4B),

contrary to CD34+/ITGA7-/CXCR4- cells that were PAX7-

negative (Figure 4C). CD34+/ITGA7+/CXCR4+ cells were

able to differentiate into myofibers when grown in a

myogenic medium for 7 additional days, as shown by

dystrophin (DMD) staining (Figure 4D), confirming their

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myogenic potential. Thus, combined tissue culture and

immunofluorescence analyses showed that FACS-isolated

CD34+/ITGA7+/CXCR4+ cells are satellite cells.

To determine whether isolated satellite cells are suitable

for CUT&RUN analysis, the genomic profile of acetylated

lysine 27 (H3K27ac) and dimethylated lysine 4 (H3K4me2)

of histone H3, two histone modifications found at active

promoter and enhancer regions, was determined. Our data

uncovered 68,694 and 13,514 peaks for H3K4me2 and

H3K27ac, respectively, with a similar genomic repartition

between the two histone marks (Figure S4A). In detail, we

found that one-quarter of the peaks were located ± 2 kb

from the transcription start site (TSS) of the nearest gene,

the majority being located either -100 kb to -10 kb, or 10

kb to 100 kb from the TSS (Figure S4B). Of note, Pearson

analysis showed an 80% correlation between H3K27ac and

H3K4me2 read profiles (Figure S4C). To assess that the

chromatin prepared from the abovementioned protocol was

isolated from satellite cells, the presence of H3K27ac and

H3K4me2 at the promoter of satellite cell-specific genes was

determined. H3K4me2 was enriched around the TSS of Pax7,

Itga7, Lamb2, Cxcr4, and Vcam1 (Figure 5A), but not at those

of Itgam (CD11b) and Ptprc (CD45) (immune cells), Pecam

(CD31; endothelial cells) (Figure 5B), Ckm (developing

muscle fibers), or Myh3 (myosin heavy chain fast embryonic,

myofibers) (Figure 5C). Similar results were obtained with

H3K27ac (Figure 5). Almost no signal was obtained with

the IgG sample (Figure 5). Moreover, comparison of the

H3K27ac peaks obtained from SEACR with those resulting

from MACS2 peak calling from published ChIP-seq datasets

showed a high correlation, as noted below each panel (ChIP-

Atlas track; Figure 5). Together, these results indicate that the

reads obtained after the bioinformatics analysis originate from

the chromatin of FACS-isolated satellite cells and correlate

with those previously identified by ChIP-seq analyses.

While single-cell RNA sequencing studies in mouse satellite

cells showed a striking stress response, caused by the

isolation procedure

, the presence of H3K4me2 or H3K27ac

was determined at stress response genes, as exemplified

with Atf3, Azin1, Gls, and Elf2. The results provide evidence

that the H3K27ac mark is not deposited at the promoter

of such genes, and that levels of H3K4me2 remain low

compared to those obtained for satellite cell-specific genes

(Figure S5). Thus, these data highlight how mild the isolation

procedure is.

Next, the suitability of our isolation method for transcription

factors was examined. CUT&RUN analysis for the androgen

receptor (AR), a transcription factor that belongs to the

nuclear receptors superfamily and that plays an important

role in myogenic differentiation

, unraveled 7,840 peaks.

These peaks were mainly located at intron and intergenic

regions (Figure S4A), either -100 kb to -10 kb, or 10

kb to 100 kb from the TSS (Figure S4B). Phylogenetic

analyses revealed that AR, alongside the glucocorticoid

(GR/Nr3c1), the mineralocorticoid (MR/Nr3c2), and the

progesterone (PR/Nr3c3) receptors, is a member of the

oxosteroid nuclear receptors subfamily

, that bind as

homodimers to DNA segments which are made up of

two 5′-RGAACA-3′ palindromic half-sites separated by

three base pairs

. Searching for a known motif by

means of hypergeometric optimization of motif enrichment

(HOMER, http://homer.ucsd.edu/homer/) revealed that AR is

bound to the 5′-RGRNCA-3′ AR half-site motif, to the typical

5′-RGNACAnnnTGTNC-3′ oxosteroid motif (referred in the

figure as PR), and to the 5′-RGNACAnnnTGTNCY-3′ AR

consensus motif in >32%, 17%, and 2% of the targeted

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regions, respectively (Figure 6A). It should be noted

that similar proportions were previously obtained for the

glucocorticoid receptor in skeletal muscle

SEACR analysis unveiled almost 500 peaks with a peak

score >50, with more than 200 of them with a score >100;

some of which are exemplified in Figure 6B. Our analysis

also uncovered a modest enrichment of AR at the previously

described binding sites of genes involved in polyamine

biosynthesis (Amd1, Oat, and Smox) and in prostate cancer

(Acox1, Fkbp5, and Tmprss2) (Figure S6). Interestingly, the

AR was found at the loci of genes involved in satellite cell

stemness (Pax7, Cxcr4, and Cd34) (Figure S7). Of note, the

oxosteroid response element was found at each of these AR-

enriched regions (Figure S6 and Figure S7), demonstrating

that the AR signal seen in the CUT&RUN data is highly

specific.

Figure 1: Schematic representation of the protocol used for satellite cell isolation from mouse limb muscles. The

protocol comprises four mains steps. Briefly, mice are sacrificed, and the hind limb muscles are harvested (steps 1.1.1-1.1.3)

and mechanically minced using scissors (steps 1.4-1.5). This is followed by steps 1.2.1-1.2.4, during which muscles are

dissociated in a water-bath using collagenase and dispase. In steps 1.2.5-1.2.8, the cell suspension is filtered through

100, 70, and 40 µm cell strainers, consecutively, and erythrocytes are eliminated using red blood cell lysis buffer (steps

1.2.9-1.2.12). The remaining cell populations are labelled in step 1.3 with antibodies coupled to fluorophores, which are

directed against specific cellular phenotypical markers. Step 1.4 includes samples passing through FACS to collect satellite

cells for further application. Please click here to view a larger version of this figure.

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Figure 2: Flow cytometry analysis of an unstained cell preparation. (A) Selection of the population of interest based on

FSC-A and SSC-A parameters. (B) Single cell identification based on FSC-A and FSC-H. (C) Characterization of the auto-

fluorescence threshold of collected cells for APC-Cy7 conjugated to the fixable viability stain (FVS 780) marking dead cells.

(D-F). Auto-fluorescence measurement for PE-Cy7 conjugated to CD11b antibody and PE-conjugated to TER119/CD45/

CD31 markers (D), as well as for Alexa fluor 488 conjugated to ITGA7, Alexa fluor 405 conjugated to CD34 (E), and APC

fluorochrome conjugated to CXCR4 (F). Gates are represented as black boxes. Please click here to view a larger version of

this figure.

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Figure 3: Flow cytometer gating strategy for satellite cell sorting. (A) Selection of the population of interest based on

FSC-A and SSC-A parameters. (B) Single cell identification based on FSC-A and FSC-H. (C) Identification of living cells

with FVS 780. (D) Negative cell selection based on CD11b, CD31, CD45, and TER119 antigens. (E-F) Positive cell selection

based on CD34 and ITGA7 (E), as well as CXCR4 (F) antigens. Gates are represented as black boxes. Please click here to

view a larger version of this figure.

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Figure 4: Analysis of FACS-isolated CD34+/ITGA7+/CXCR4+ and CD34+/ITGA7-/CXCR4- cells. (A) Phase contrast

images of CD34+/ITGA7+/CXCR4+ and CD34+/ITGA7-/CXCR4- cells cultured in growth medium. (B) Immunofluorescence

analysis of CD34+/ITGA7+/CXCR4+ and CD34+/ITGA7-/CXCR4- cells cultured in growth medium using antibodies directed

against PAX7 (red) and dystrophin (DMD, green). Nuclei were stained with DAPI (blue). Magenta arrows indicate PAX7-

positive cells. (C) Phase contrast images of CD34+/ITGA7+/CXCR4+ cells grown for 5 days in growth medium (left panel)

and for 7 additional days in myogenic medium (right panel). (D) Immunofluorescence analysis images of CD34+/ITGA7+/

CXCR4+ cells grown for 5 days in growth medium (left panel) and for 7 additional days in myogenic medium (right panel)

using antibodies directed against PAX7 (red) and dystrophin (DMD, green). Nuclei were stained with DAPI (blue). Magenta

arrows indicate PAX7-positive cells. Green arrows indicate DMD-positive cells. Scale bars: brightfield panels = 25 µm;

immunofluorescence panels = 50 µm. Please click here to view a larger version of this figure.

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Figure 5: Genomic profiles of satellite cell chromatin. Localization of H3K4me2 and H3K27ac at satellite cell-specific

genes (A), immune and endothelial cell-specific genes (B), and myofiber-specific genes (C) on the chromatin of satellite cells

by CUT&RUN. Active promoters are boxed in green, whereas inactive promoters are boxed in brown. CUT&RUN performed

with IgG was used as a negative control. H3K27ac enrichment analysis for histone marks is presented below each track.

Enrichment score showing the levels of confidence of published datasets is depicted as a heatmap. Brown stars (*) refer

to H3K27ac ChIP-seq peaks performed in muscle satellite cells, blue stars to H3K4me3, and the pink one to H4K16me1.

Please click here to view a larger version of this figure.

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Figure 6: AR genomic distribution on satellite cell chromatin. (A) HOMER-known motif analysis of AR peaks in satellite

cells. PR: progesterone receptor. Nb target refers to the number of peaks presenting a particular motif. (B) Localization of

H3K4me2 and AR at indicated genes on the chromatin of satellite cells determined by CUT&RUN. CUT&RUN performed

with IgG was used as a negative control. Please click here to view a larger version of this figure.

Table 1: List of materials, reagents, and software. Please

click here to download this Table.

Table 2: Buffer compositions. Please click here to

download this Table.

Table 3: Antibody references and concentrations. Please

click here to download this Table.

Supplementary Figure 1: Flow cytometry analysis

of the cell preparation post-sorting. (A) Selection of

the population of interest based on FSC-A and SSC-A

parameters. (B) Single cell identification based on FSC-

A and FSC-H. (C) Identification of living cells with fixable

viability stain (FVS 780). (D) Negative cell selection based on

CD11b, CD31, CD45, and TER119 antigens. (E-F) Positive

cell selection based on CD34 and ITGA7 (E) as well as

CXCR4 (F) antigens. Gates are represented as black boxes.

Please click here to download this File.

Supplementary Figure 2: Flow cytometer gating strategy

for satellite cell sorting after digestion with 300 µL of

Liberase TL. (A) Selection of the population of interest

based on FSC-A and SSC-A parameters. (B) Single cell

identification based on FSC-A and FSC-H. (C) Identification of

living cells with FVS 780. (D) Negative cell selection based on

CD11b, CD31, CD45, and TER119 antigens. (E-F) Positive

cell selection based on CD34 and ITGA7 (E) as well as

CXCR4 (F) antigens. Gates are represented as black boxes.

Please click here to download this File.

Supplementary Figure 3: Flow cytometer gating strategy

for satellite cell sorting after digestion with 600 µL of

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Liberase TL. (A) Selection of the population of interest

based on FSC-A and SSC-A parameters. (B) Single cell

identification based on FSC-A and FSC-H. (C) Identification of

living cells with FVS 780. (D) Negative cell selection based on

CD11b, CD31, CD45, and TER119 antigens. (E-F) Positive

cell selection based on CD34 and ITGA7 (E) as well as

CXCR4 (F) antigens. Gates are represented as black boxes.

Please click here to download this File.

Supplementary Figure 4: Characterization of H3K4me2,

H3K27ac, and AR genomic locations. (A) Pie charts

depicting the peak distribution of H3K4me2, H3K27ac, and

AR according to genome features in satellite cells. (B) Pie

charts depicting the peak distribution of H3K4me2, H3K27ac,

and AR according to their distance to the nearest TSS in

satellite cells. (C) Heatmap showing the Pearson correlation

between H3K4me2, H3K27ac, and IgG control. Please click

here to download this File.

Supplementary Figure 5: Genomic profiles of satellite cell

chromatin at stress-response induced genes. Localization

of H3K4me2 and H3K27ac at indicated genes on the

chromatin of satellite cells. Immunoprecipitation with IgG was

used as a negative control. Please click here to download this

File.

Supplementary Figure 6: Genomic profiles of satellite

cell chromatin at known AR target genes. Localization

of H3K4me2, H3K27ac, and AR at indicated genes on the

chromatin of satellite cells determined by CUT&RUN. AR

peaks are boxed in blue, and corresponding AR-responsive

elements are presented below. CUT&RUN performed with

IgG was used as a negative control. Please click here to

download this File.

Supplementary Figure 7: Genomic profiles of satellite

cell chromatin at their selectively expressed genes.

Localization of H3K4me2, H3K27ac, and AR at indicated

genes on the chromatin of satellite cells determined by

CUT&RUN. AR peaks are boxed in blue, and corresponding

AR-responsive elements are presented below. CUT&RUN

performed with IgG was used as a negative control. Please

click here to download this File.

Discussion

The present study reports a standardized, reliable, and easy-

to-perform method for the isolation and culture of mouse

satellite cells, as well as the assessment of transcriptional

regulation by the CUT&RUN method.

This protocol involves several critical steps. The first is

muscle disruption and fiber digestion to ensure a high

number of collected cells. Despite the increased enzyme

concentration, more living cells were obtained than using

Protocol 1. Satellite cells express a specific pattern of

various membrane proteins. To increase the stringency of

our sorting, we used a combination of previously described

negative (CD31, TER119, CD45, and CD11b) and positive

(CD34, ITGA7, and CXCR4) satellite cell markers

30 , 31

Using this strategy, an average of 1% of living putative

satellite cells was obtained. This outcome is in the range

of what was expected, since satellite cells represent 2%-7%

of muscle cell population at adulthood in mice

. As a

control for the satellite cell selection markers, CD34+/ITGA7-/

CXCR4- cells were isolated. Immunofluorescent staining

demonstrated that, whereas CD34+/ITGA7-/CXCR4- cells

did not express PAX7, 70% of CD34+/ITGA7+/CXCR4+

sorted cells were positive for this satellite cell-specific marker,

thereby demonstrating that the FACS isolated population

corresponds to satellite cells. In addition, 70% of satellite

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cells grown in myogenic medium for 7 days were PAX7-/DMD

+, as shown by immunostaining, and formed an elongated

multinucleated myofiber, demonstrating that isolated satellite

cells conserved their stemness potential.

The major limitation in sample preparation might be the use

of a high concentration of enzymes, which results in a large

amount of dissociated biological material, but may contribute

to increased cell death. To resolve this issue, an assay

requiring lower enzyme quantity was tested. In this context,

Liberase TL, a purified form of the traditional collagenase

was tested. Although digestion was apparently more efficient

with this enzyme, the amount of cell debris remained

elevated, and the cell viability was slightly lowered. These

observations are in agreement with previous reports that

compared Liberase-mediated tissue digestion to this with

recombinant collagenase or custom collagenase

24 , 25

. In

addition, even though the proportion of endothelial and

immune cells was similar between collagenase and Liberase

digestion, the percentage of satellite cells was lower

in the Liberase-processed sample, overall showing that

Liberase is not recommended for satellite cell isolation. The

use of this enzyme is suitable for the phenotypic and

quantitative analysis of immune cells and could eventually be

recommended in the context of muscle and/or other tissues.

This is in concordance with what has been reported on

Liberase TL as the most suited to effectively isolate viable

immune cells

34 , 35 , 36

Another potential issue is the stress caused by both

dissociating and sorting satellite cells. However, the absence

of H3K27ac and the low H3K4me2 levels at the promoter

region of stress response genes identified by single-cell

RNA sequencing in mouse satellite cells

show that the

procedure is mild enough not to induce the stress response

transcriptional repertoire. Other limitations to be noted

are the selected antigens that remain empiric. However,

studies show a high overlap between unique surface marker

combinations that are enriched for skeletal muscle satellite

cells

30 , 31

An additional critical step is the purification of nuclei from

sorted cells. For this, the sorting speed is maintained low in

order not to damage the cells, but fast enough that they are

not stored for too long in FACS buffer. Indeed, CUT&RUN

nuclei are not fixed, and a longer incubation time might

influence the read obtained from the protocol. A typical

preparation from the two hind limbs of one mouse allows

CUT&RUN with one antibody and one IgG control, with an

amount of 2.5 ng of chromatin for each.

CUT&RUN experiments, designed to assess transcription

factor binding and epigenetic modifications, provided strong

and robust read signals for H3K4me2 and H3K27ac. The

high read levels of H3K4me2 and H3K27ac on genes that

are known to be expressed in satellite cells, compared to

genes expressed in immune or endothelial cells as well

as in myofibers, showed that the signal was amplified

predominantly from the satellite cells nuclei. In addition, this

protocol made the identification of the cistrome of the AR in

muscle stem cells possible, which to date remains a technical

issue inherent to the poor quality of mouse AR antibodies

and the low AR expression levels in cells that are not highly

androgen-sensitive, such as epithelial prostate cells. The

data presented here unveiled AR-bound regions with high

confidence peak scores. As only one replicate was originally

assessed per antibody, the robustness of our datasets will

be improved by adding at least two additional biological

replicates per condition. However, AR known target genes

and respective histone marks, originally described as highly

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expressed and/or easily detectable in other tissue contexts

such as the prostate, present lower expression levels and are

very challenging to detect in satellite cells.

One limitation of the CUT&RUN approach is that it is

performed in unfixed cells. Thus, we might have missed

some AR targets because of the labile nature of the

interactions between transcription factors and their binding

site. Similarly, some post-translational modifications, such as

acetylation, can also be rather labile. Therefore, including a

light fixation step with formaldehyde or paraformaldehyde,

after FACS sorting, and before isolating the nuclei, could

be considered to stabilize the protein/DNA interactions and

histone modifications.

Although this paper shows that CUT&RUN assays can be

conducted on FACS-isolated satellite cells, other analyses

that use non-fixed chromatin such as ATAC-seq might also

be performed. In addition, even though here muscles were

harvested in basal physiological conditions, this protocol may

be applied to pathological contexts such as injury or aging.

All these protocol uses will allow a detailed understanding of

gene regulatory mechanisms in satellite cells, and may help

in understanding how these mechanisms might be altered by

pathophysiological conditions.

In summary, this low-cost and time-efficient protocol provides

an effective experimental setting to study transcription factor

recruitment and chromatin landscape in skeletal muscle

precursor cells. Chromatin prepared by this protocol has

provided the first genome-wide analysis of AR cistrome

in satellite cells and will facilitate future studies on gene

regulation.

Disclosures

The authors declare that they have no competing financial

interests.

Acknowledgments

We thank Anastasia Bannwarth for providing excellent

technical assistance. We thank the IGBMC animal house

facility, the cell culture, the Mouse Clinical Institute (ICS,

Illkirch, France), the imaging, the electron microscopy, the

flow cytometry, and the GenomEast platform, a member of

the 'France Génomique' consortium (ANR-10-INBS-0009).

This work of the Interdisciplinary Thematic Institute IMCBio,

as part of the ITI 2021-2028 program of the University

of Strasbourg, CNRS and Inserm, was supported by

IdEx Unistra (ANR-10-IDEX-0002) and by SFRI-STRAT'US

project (ANR 20-SFRI-0012) and EUR IMCBio (ANR-17-

EURE-0023) under the framework of the French Investments

for the Future Program. Additional funding was delivered

by INSERM, CNRS, Unistra, IGBMC, Agence Nationale

de la Recherche (ANR-16-CE11-0009, AR2GR), AFM-

Téléthon strategic program 24376 (to D.D.), INSERM young

researcher grant (to D.D.), ANR-10-LABX-0030-INRT, and a

French State fund managed by the ANR under the frame

program Investissements d'Avenir (ANR-10-IDEX-0002-02).

J.R. was supported by the Programme CDFA-07-22 from the

Université franco-allemande and Ministère de l'Enseignement

Supérieur de la Recherche et de l'Innovation, and K.G. by the

Association pour la Recherche à l'IGBMC (ARI).

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