Work in the Barton lab was supported by a grant from the Cancer Prevention and Research Institute of Texas (CPRIT RP110471). This research was supported in part, by a training grant fellowship for Ryan L. McCarthy from the National Institutes of Health Training Program in Molecular Genetics 5 T32 CA009299. The core facility that maintains and runs the mass cytometry machine is supported by CPRIT grant (RP121010) and NIH core grant (CA016672).

1. Cell Harvesting
<ol>
	<li>Harvesting cells from primary tissue 
	NOTE: The process described for harvesting cells from primary tissue is specifically applicable to mouse breast tumor tissue and may not be applicable as is to primary tissues from other sources.
	<ol>
		<li>Prepare digestion buffer by dissolving 5 mg hyaluronidase and 30 mg collagenase in 10 mL DMEM/F12 media per gram of tissue to be processed. Filter sterilize digestion buffer.</li>
		<li>Isolate primary mammary gland tumor from mouse and record tumor weight.</li>
		<li>Mince tissue with #10 scalpel for at least 5 min.</li>
		<li>Add minced tissue to appropriate volume of digestion buffer (10 mL buffer per gram of tissue).</li>
		<li>Shake minced tissue in digestion buffer at 100 rpm for 1-3 h at 37 &#176;C.</li>
		<li>Obtain single cell suspension by passing cells through 0.4 &#956;m filter into a 50 ml tube.</li>
		<li>Centrifuge cells at 300 x g for 10 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Resuspend pellet in 4 mL DMEM/F12 and transfer to 5 mL round bottom tube.</li>
	</ol>
	</li>
	<li>Harvesting cells from tissue culture
	<ol>
		<li>Wash plate or flask of adherent cells with calcium and magnesium free phosphate buffered saline (PBS) at room temperature. 
		NOTE: The technique described for harvesting cells is specifically applicable to adherent human embryonic stem cell (hESC) culture cells. However, the remainder of the described protocol is broadly applicable to other cultured cell lines, non-adherent lines and primary cells.</li>
		<li>Add warmed (37 &#176;C) trypsin or other enzymatic digestion reagent optimized for achieving a single cell suspension from the adherent cells. Follow manufacturer&#39;s protocol. 
		NOTE: Trypsin and other enzymatic dissociation reagents have the potential to affect cellular markers.</li>
		<li>Incubate the plate at 37 &#176;C for 2-5 min to detach cells. For high confluency cultures, gently tap the plate to help detach the cells.</li>
		<li>Transfer fully detached cells from the plate into a 5 mL round bottom tube and gently pipet using a 1 mL pipet to dissociate cells. 
		NOTE: For processing large numbers of samples all steps can alternatively be performed in a 96 well round bottom plate. For processing samples in a 96 well format all washes can be performed at a volume of 250 &#956;L. The volumes described in other steps can be adjusted so the total volume does not exceed 300 &#956;L.</li>
		<li>Quench the enzymatic digestion reagent with an equal volume wash buffer (0.5% BSA and 0.02% sodium azide in PBS). 
		NOTE: Sodium azide is a hazardous chemical. Proper safety precautions must be observed for its preparation and handling.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 37 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Resuspend cells in 1 mL serum free media.</li>
		<li>Count cells by transferring 10 &#956;L of the cell suspension to a 1.5 mL microcentrifuge tube. Dilute aliquot to a known ratio in trypan blue and transfer to a hemocytometer or automated cell counting system.</li>
	</ol>
	</li>
	<li>Labeling of S-phase population 
	NOTE: If S-phase labeling is not to be performed proceed to stem 1.4.
	<ol>
		<li>Prepare 1 mL of 10 &#956;M 5-Iodo-2&#39;-deoxyuridine (IdU) in serum free DMEM F12 or other appropriate media for each 2 x 106 cells to be analyzed.</li>
		<li>Add 500 &#956;L of 10 &#956;M IdU in serum free media for each 2 x 106 cells. 
		NOTE: While IdU labeling can be performed in media containing serum, free protein will react with cisplatin, preventing proper labeling of dead cells. If media containing serum is used during IdU labeling an additional wash step is serum free media is necessary to remove serum proteins before cisplatin live/dead staining.</li>
		<li>Gently resuspend pellets with 1 mL pipet.</li>
		<li>Incubate for 10 min at 37 &#176;C with gentle rocking.</li>
	</ol>
	</li>
	<li>Live/Dead labeling
	<ol>
		<li>For each sample prepare 500 &#956;L of 50 &#956;M cisplatin in serum free DMEM-F12 or cell type appropriate media. 
		NOTE: If labeling of the S-phase population by IdU is not to be performed, resuspend samples at a concentration of 4 x 106 cells/mL. Resuspending the cells before adding cisplatin enables quicker mixing of solutions for uniform live/dead labeling.</li>
		<li>Add 500 &#956;L of 50 &#956;M cisplatin per 2 x 106 cells to samples.</li>
		<li>Incubate at room temperature (RT) for 1 min on an orbital shaker with constant mixing.</li>
		<li>Quench the cisplatin with an equal volume of wash buffer.</li>
		<li>Centrifuge cells at 300 x g for 5 min at RT. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Wash samples twice with 500 &#956;L wash buffer to remove excess cisplatin.</li>
		<li>Wash samples twice with 500 &#956;L PBS to remove excess protein.</li>
	</ol>
	</li>
	<li>Sample fixation
	<ol>
		<li>Resuspend sample in 500 &#956;L PBS.</li>
		<li>Add equal volume of 4% PFA in PBS for a final concentration of 2%; pipette to mix. 
		NOTE: Prepare 4% PFA fresh from powder, adjust pH to 6.9 and pass through a 0.44 &#956;m filter. 4% PFA in PBS can be stored at 4 &#176;C for up to two weeks. Avoid premade formalin supplied in ampules, unless specifically tested, as it often contains metal contaminants which may interfere with the measured mass channels. Lower concentration of PFA may be sufficient and help to minimize the effect of fixation on antibody binding but should be tested empirically.</li>
		<li>Incubate for 15 min at RT on an orbital shaker with constant mixing.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Wash samples with PBS to remove PFA solution. 
		NOTE: At this step samples can be stored briefly at 4 &#176;C. Prolonged storage at 4 &#176;C may result in degradation of the sample and decreased staining.</li>
	</ol>
	</li>
</ol>
2. Barcoding
NOTE: While methodology for utilizing monoisotopic cisplatin barcoding and palladium barcoding is presented simultaneously, either can be used independently or avoided entirely if not required by the experiment.

<ol>
	<li>Monoisotopic Cisplatin Barcoding 
	NOTE: Since cisplatin barcoding and cisplatin Live/Dead staining rely on overlapping channels care should be taken to ensure that background cisplatin Live/Dead staining in the live cells is minimized as this may interfere with the accuracy of the cisplatin barcode.
	<ol>
		<li>Dilute 1 mM monoisotopic cisplatin stock solutions to 200 &#956;M in PBS.</li>
		<li>For each unique cisplatin barcode prepare a 1.5 mL tube with 500 &#956;l PBS for each 2 x 106 cells receiving that barcode.</li>
		<li>To prepare each unique barcode add each applicable monoisotopic cisplatin to a final concentration of 200 nM.</li>
		<li>Resuspend cells in 500 &#956;l of PBS per 2 x 106 cells.</li>
		<li>Add an equal volume of the corresponding barcode solution to each sample.</li>
		<li>Incubate samples at RT for 5 min on an orbital shaker with constant mixing.</li>
		<li>Quench cisplatin with an equal volume of wash buffer.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Wash samples twice with 500 &#956;L wash buffer to remove excess cisplatin.</li>
	</ol>
	</li>
	<li>Palladium Barcoding 
	NOTE: Palladium barcoding reagents can be either prepared5 or purchased commercially.
	<ol>
		<li>Transient partial permeabilization19 
		NOTE: Palladium chelation barcoding requires partial permeabilization for the reagents to pass through the cell membrane and robustly label the cell.
		<ol>
			<li>Wash samples with 500 &#956;L PBS.</li>
			<li>Wash samples with 500 &#956;L PBS plus 0.02% saponin.</li>
			<li>Resuspend pellet in residual volume.</li>
		</ol>
		</li>
		<li>Apply palladium barcoding.
		<ol>
			<li>Dilute 100x stock of palladium barcoding reagent in 1 mL ice-cold PBS plus 0.02% saponin.</li>
			<li>Rapidly add diluted barcoding reagent to resuspended sample.</li>
			<li>Incubate for 15 min at RT on an orbital shaker at with constant mixing.</li>
			<li>Wash samples twice with 500 &#956;L wash buffer.</li>
		</ol>
		</li>
	</ol>
	</li>
</ol>
3. Cell Surface Staining
NOTE: Cell surface staining can be performed on live cells prior to fixation. This may be necessary for antibodies that lose affinity for their epitope after fixation. Some cell samples may benefit from additional blocking, such as Fc blocking for leukocytes, prior to antibody labeling to reduce background. Optimal blocking should be empirically determined for specific sample type.

<ol>
	<li>Prepare cell surface antibody staining panel.
	<ol>
		<li>Combine 0.5 &#956;L, or empirically determined quantity, of each 0.5 mg/mL cell surface antibody per sample into a 1.5 mL tube. Add washing buffer if necessary to bring volume up to 10 &#956;L per sample. Mix by pipetting. 
		NOTE: Determine working dilution for each antibody prior to experiment empirically by titration experiment.</li>
		<li>Split cell surface antibody pool equally into one 1.5 mL tube for each sample to be stained.</li>
		<li>Prepare 500 &#956;l of wash buffer in a 1.5 mL tube for each sample.</li>
	</ol>
	</li>
	<li>Apply cell surface staining to samples in normalized volume for all samples. 
	NOTE: To ensure consistent antibody labeling across multiple samples it is vital to carefully control the sample volume and antibody concentration. The following steps aim to achieve this consistency by ensuring that the final sample volumes after the antibody has been added are uniform.
	<ol>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>With a 200 &#956;L pipette set to 50 &#956;L, remove remaining solution from sample pellet and transfer to a tube of the aliquoted cell surface antibody pool.</li>
		<li>Pipette up all liquid in the aliquot tube without drawing air into the pipette tip.</li>
		<li>While holding the pipette plunger to avoid drawing air move the tip into a prepared tube of 500 &#956;L wash buffer and release the plunger.</li>
		<li>Add back the contents of the pipette tip to the sample and resuspend the sample in the liquid with gentle pipetting.</li>
		<li>Incubate the samples for 1 h at RT on an orbital shaker with constant mixing.</li>
		<li>Wash samples twice with 500 &#956;L wash buffer to ensure that all free antibody is washed away.</li>
		<li>Wash samples with 500 &#956;L PBS.</li>
		<li>Resuspend the cell pellet in 500 &#956;l PBS.</li>
		<li>Add equal volume of 4% PFA in PBS; pipette to mix.</li>
		<li>Incubate for 10 min on an orbital shaker.</li>
	</ol>
	</li>
</ol>
4. Cell Permeabilization and Intracellular Staining
<ol>
	<li>Cell permeabilization
	<ol>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Resuspend cells in residual volume by gently vortexing until the pellet is dissociated. 
		NOTE: Failure to dissociate cells prior to adding MeOH will result in cells adhering together and producing a thin film which will result in partial sample loss.</li>
		<li>Immediately add 1 mL of 100% MeOH per 2 x 106 cells and gently pipette to mix. 
		NOTE: Other permeabilization methods can be substituted for MeOH and may be necessary to achieve optimal permeabilization for some cell types. For some cell sources a 0.1% Triton X-100, 0.2% Tween-20 or 0.1% saponin in PBS solution for permeabilization may maintain cell integrity better while still providing consistent permeabilization.</li>
		<li>Incubate samples for at least 1 h or up to a maximum of 24 h at 4 &#176;C for permeabilization. 
		NOTE: At this step samples in MeOH can be stored for up to 1 month at -80 &#176;C.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Add 1 mL of washing buffer for each ml MeOH used to permeabilize the sample. Gently resuspend cell pellet.</li>
		<li>Wash sample twice with 500 &#956;L wash buffer.</li>
	</ol>
	</li>
	<li>Prepare intracellular antibody staining panel.
	<ol>
		<li>Combine 0.5 &#956;L, or empirically determined quantity, of each 0.5 mg/mL intracellular antibody per sample into a 1.5 mL tube. Add washing buffer if necessary to bring volume up to 10 &#956;L per sample. Mix by pipetting.</li>
		<li>Split intracellular antibody pool equally into one 1.5 mL tube for each sample.</li>
		<li>Prepare 500 &#956;L of wash buffer in a 1.5 mL tube for each sample.</li>
	</ol>
	</li>
	<li>Apply intracellular staining to samples in normalized volume for all samples. 
	NOTE: To&#160;ensure consistent antibody labeling across multiple samples it is vital to carefully control the sample volume and antibody concentration.
	<ol>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>With 200 &#956;L pipette set to 50 &#956;L remove remaining solution from sample pellet and transfer to a tube of the aliquoted cell surface antibody pool.</li>
		<li>Pipette up all liquid in the aliquot tube without drawing air into the pipette tip.</li>
		<li>While holding the pipette plunger to avoid drawing air move the tip into a prepared tube of 500 &#956;L wash buffer and release the plunger, drawing up wash buffer for a total sample volume of 50 &#956;L.</li>
		<li>Add back the contents of the pipette tip to the sample and resuspend the sample in the liquid with gentle pipetting.</li>
		<li>Incubate the samples for 1 hr at RT on an orbital shaker with constant mixing.</li>
		<li>Wash samples twice with 500 &#956;L wash buffer.</li>
		<li>Wash samples with 500 &#956;L PBS.</li>
	</ol>
	</li>
</ol>
5. Iridium Labeling
<ol>
	<li>Fix samples
	<ol>
		<li>Resuspend samples in 500 &#956;L of PBS.</li>
		<li>Add 500 &#956;L of 4% PFA in PBS.</li>
		<li>Incubate for a minimum of 15 min at RT on an orbital shaker with constant mixing.</li>
	</ol>
	</li>
	<li>Apply Iridium labeling of DNA
	<ol>
		<li>Dilute the 125 &#956;M 500x iridium intercalating reagent in 4% PFA in PBS to a concentration of 62.5 nM.</li>
		<li>Add 500 &#956;L 62.5 nM iridium PFA solution to samples. 
		NOTE: For experiments using permeabilized cells, dilute the 500x&#160;Ir191/193 stock for a final dilution of 1:2,000 to avoid overstaining.</li>
		<li>Incubate for 15 min at RT on an orbital shaker with constant mixing. 
		NOTE: If performing monoisotopic cisplatin barcoding do not incubate samples with iridium for longer than 15 min since strong Ir193 signals can contribute to the&#160;Pt194 mass channel and negatively impact barcoding quality.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
		<li>Wash samples twice with 500 &#956;L 0.1% BSA in filtered deionized water. 
		NOTE: Once prepared we recommend running samples within 24 h if possible. Prepared samples may be stored short term at 4 &#176;C, but care should be taken to avoid degradation. If possible the final washes should be performed in filtered deionized water without BSA as this will decrease the deposition on the spray chamber and sampler cones of the machine which will decrease instrument cleaning. For hESCs it is necessary to perform these washes while including BSA to avoid sample loss from sticking to the tube surface.</li>
	</ol>
	</li>
</ol>
6. Prepare Samples for Acquisition
<ol>
	<li>Count cells
	<ol>
		<li>Resuspend cells in 500 &#956;L 0.1% BSA in filtered deionized water and filter into a fresh tube through a 35 &#956;m cell strainer cap. 
		NOTE: Lowering the BSA level for the final washes decreases the residue left on the Mass cytometry nebulizer. Non-adherent cells can be washed and resuspended in filtered deionized water.</li>
		<li>Count cells by transferring 10 &#956;L of the cell suspension to a microcentrifuge tube. Dilute aliquot to a known ratio in trypan blue (to aid with cell visualization) and transfer to a hemocytometer or automated cell counting system.</li>
		<li>Centrifuge cells at 300 x g for 5 min at 4 &#176;C. Decant or carefully aspirate the supernatant without disturbing the pellet.</li>
	</ol>
	</li>
	<li>Prepare and load samples
	<ol>
		<li>To prepare calibration beads remove from refrigerator and shake vigorously to resuspend.</li>
		<li>If running samples using a mass cytometry autosampler, add 50 &#956;L of calibration beads to each well of sample plate then add the desired number of cells to be analyzed. If running samples by manual injection add 50 &#956;L of calibration beads to sample, dilute sample in filtered deionized water, mix well and load sample into Mass cytometry sample injection port. The appropriate sample dilution may vary depending upon the cell type being analyzed, but a dilution of 106 is generally appropriate1.</li>
	</ol>
	</li>
</ol>

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K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Transient+partial+permeabilization+with+saponin+enables+cellular+barcoding+prior+to+surface+marker+staining.">Transient partial permeabilization with saponin enables cellular barcoding prior to surface marker staining.</a> Cytometry A. 85, 1011-1019 (2014).</li><li>Finck, R., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Normalization+of+mass+cytometry+data+with+bead+standards.">Normalization of mass cytometry data with bead standards.</a> Cytometry A. 83, 483-494 (2013).</li><li>Chester, C., Maecker, H. T. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Algorithmic+Tools+for+Mining+High-Dimensional+Cytometry+Data.">Algorithmic Tools for Mining High-Dimensional Cytometry Data.</a> J Immunol. 195, 773-779 (2015).</li><li>Diggins, K. E., Ferrell, P. B., Irish, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Methods+for+discovery+and+characterization+of+cell+subsets+in+high+dimensional+mass+cytometry+data.">Methods for discovery and characterization of cell subsets in high dimensional mass cytometry data.</a> Methods. 82, 55-63 (2015).</li><li>Leelatian, N., Diggins, K. E., Irish, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Characterizing+Phenotypes+and+Signaling+Networks+of+Single+Human+Cells+by+Mass+Cytometry.">Characterizing Phenotypes and Signaling Networks of Single Human Cells by Mass Cytometry.</a> Methods Mol Biol. 1346, 99-113 (2015).</li><li>van der Maaten, L. J. P., Hinton, G. E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizaing+Data+using+t-SNE.">Visualizaing Data using t-SNE.</a> J. Mach. Learn. Res. 9, 2431-2456 (2008).</li><li>Kling, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Cytometry:+Measure+for+measure.">Cytometry: Measure for measure.</a> Nature. 518, 439-443 (2015).</li><li>Gregori, G., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Hyperspectral+cytometry.">Hyperspectral cytometry.</a> Curr Top Microbiol Immunol. 377, 191-210 (2014).</li><li>Chattopadhyay, P. K., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Brilliant+violet+fluorophores:+a+new+class+of+ultrabright+fluorescent+compounds+for+immunofluorescence+experiments.">Brilliant violet fluorophores: a new class of ultrabright fluorescent compounds for immunofluorescence experiments.</a> Cytometry A. 81, 456-466 (2012).</li><li>Rieger, A. M., Havixbeck, J. J., Barreda, D. R. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=X-FISH:+Analysis+of+cellular+RNA+expression+patterns+using+flow+cytometry.">X-FISH: Analysis of cellular RNA expression patterns using flow cytometry.</a> J Immunol Methods. 423, 111-119 (2015).</li><li>Frei, A. P., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Highly+multiplexed+simultaneous+detection+of+RNAs+and+proteins+in+single+cells.">Highly multiplexed simultaneous detection of RNAs and proteins in single cells.</a> Nat Methods. 13, 269-275 (2016).</li></ol>

The authors have nothing to disclose.

The protocol presented here has been successfully employed for the processing of various cultured cell lines (H1 and H9 hESCs, mESCs, MCF7, HEK 293, KBM5, HMEC, MCF10a) and primary tissue samples (mouse bone marrow, mouse embryonic liver, mouse adult liver, mouse tumor). Irrespective of the source, any tissue that can be dissociated into single cells while preserving the cellular state should be amenable to analysis by mass cytometry, but may require some protocol optimization. It is important to note that some epitopes ...

Cytometry enables the simultaneous measurement of multiple antibody targets at a single cell level across large populations of cells. In traditional fluorescence-based flow cytometry, the number of parameters that can be quantified is limited by spectral overlap between the emission spectrum of multiple fluorophores, which requires increasingly complex compensation calculations as the number of parameters increases. These limitations are addressed by mass cytometry, where heavy metal-conjugated antibodies are detected and quantified by time of flight (TOF) mass spectrometry to greatly expand the number of parameters collected simultaneously and yield a high dimensional protein-abundance profile for each individual cell.
A basic understanding of the workings of the mass cytometry instrument is beneficial to the user and may be essential for troubleshooting. For a more thorough description of tuning and running a mass cytometry machine, see the related manuscript1. Briefly, a cell sample is labeled with a panel of metal conjugated antibodies targeting cell surface markers, cytoplasmic proteins, nuclear proteins, chromatin-bound proteins or other epitopes of interest (Figure 1i). The labeled cells are loaded onto the machine, either one at a time by manual injection or using an autosampler that samples from a 96-well plate. The loaded cells are injected through a nebulizer, which generates a spray of liquid droplets encapsulating the cells (Figure 1ii). This spray is positioned so that the cells are ionized by an argon plasma torch. This ionization generates a particle cloud comprised of all the constituent atoms of each cell (Figure 1iii). As this particle cloud travels to the detector, low atomic mass atoms are separated from the high mass ions by a quadrupole mass filter. The remaining high mass ions continue to the detector, where the abundance of each isotope is quantified (Figure 1iv). The raw data collected by the detector, are analyzed by the mass cytometry instrument software to identify cell events. For each identified cell event, the signal detected in each channel is quantified and saved to an output .fcs file. The mass channels used for detecting the heavy metal conjugated antibodies exhibit minimal spectral overlap, which ranges from 0-4% with most contributions below 1%. Because of this low cross-talk between channels, it is generally unnecessary to transform the data with a compensation matrix before analysis2. However, care should be taken during the design of the experimental panel of antibodies to ensure that an antibody with a high-abundance epitope is not assigned to a mass channel that contributes signal to a channel assigned to a low-abundance antibody, as this may create an artificially high population in the channel receiving the signal contribution.
The exact preparation of samples for mass cytometry analysis is highly dependent upon the types of samples being collected and the experimental hypothesis being tested. We provide examples of two protocols for harvesting different types of cells (human embryonic stem cells) and primary cells (mouse breast tumor epithelial cell isolate). When beginning a mass cytometry study, it is advisable to first carry out a small pilot experiment to ensure that all protocols and antibodies to be utilized are working well in the investigator&#39;s hands. This is especially essential when custom conjugated antibodies are to be used, as the partial reduction reaction during the antibody conjugation has the potential to disrupt antibody affinity; therefore, each custom antibody needs to be validated empirically to ensure data accuracy. Other considerations include determining if cell surface antibodies to be used in the assay will recognize their epitopes after fixation or if they need to be applied to live cells. Some fixation may prevent proper staining with cell surface antibodies as is known to occur with peptide-MHC staining3,4. Performing cell surface antibody staining on live cells may be necessary for some antibodies, but this precludes the option of barcoding prior to antibody labeling as most barcoding methods are performed after fixation5 although antibody-based barcoding6,7 is an exception.
When determining the number of cells to be prepared for analysis, it is important to know that only a fraction of the cells loaded onto the machine will be detected and produce data. This loss is primarily due to inefficiencies when the nebulizer sprays the sample at the torch. The exact percent loss depends upon the mass cytometry machine setup and cell type but can range from 50-85% and should be considered when designing the experiment. This protocol has been optimized for 2x106 cells per sample but can be adapted for processing smaller or larger sample sizes. This protocol can be used with minimal modifications for sample sizes from 1 x 106 to 4 x 106, however it is critical that sample size be kept consistent within an experiment. Changes in cell number can affect the intensity of barcoding and antibody staining and should be kept roughly consistent across samples to be directly compared.
Some procedures, such as dead cell labeling and DNA labeling, should be carried out in the vast majority, if not all experimental designs. The labeling of dead cells in experiments analyzing only surface markers can be achieved using Rh103, but Rh103 should be avoided for experiments where permeabilization is required as it results in staining loss. For experiments involving permeabilization, it is advisable to utilize cisplatin8 and, when possible, an antibody recognizing cleaved PARP or cleaved Caspase to detect apoptotic and dead cells.
Barcoding samples can reduce antibody consumption, decrease acquisition time and eliminate sample-to-sample variability9. The process of barcoding involves applying a unique sample-specific code to all cells, which is used to assign each cell to its sample of origin. After barcoding the samples may be pooled prior to antibody staining, a process that ensures all samples are stained equally. To minimize experimental error, it is prudent to barcode and pool any samples that are to be directly compared to each other. Currently there exist several methods for barcoding samples for mass cytometry analysis5,6,7, two of which are presented here (see below).
With currently available reagents it is possible to quantify the abundance of 38 antibody targets, identify cells in S-phase10, distinguish live and dead cells8 and measure cellular levels of hypoxia11 in a multiplexed experiment of multiple samples. This ability to collect high parameter single cell data for large cell populations enables improved profiling of highly heterogeneous cell populations and has already produced a number of novel biological insights12,13,14,15,16. Distilling the resulting complex data to interpretable information has required the use of computational algorithms including Spanning Tree Progression of Density Normalized Events (SPADE)17 and viSNE18.
This article presents an accessible overview of how to design and carry out a mass cytometry experiment and introduces basic analysis of mass cytometry data.

<table><tbody><tr><td>mTeSR1 medium kit</td><td>Stem Cell technologies</td><td>05850</td><td>Warm at room temperature before use</td></tr><tr><td>DMEM F-12</td><td>ThermoFisher</td><td>11330-032</td><td>Warm at 37&amp;nbsp;&amp;deg;C before use</td></tr><tr><td>Accutase</td><td>Stem Cell technologies</td><td>07920</td><td>Warm at 37&amp;nbsp;&amp;deg;C before use</td></tr><tr><td>Bovine serum albumin</td><td>Equitech</td><td>BAH62</td><td></td></tr><tr><td>phosphate buffered saline</td><td>Hyclone</td><td>SH30256.01</td><td></td></tr><tr><td>Saponin</td><td>Sigma-Aldrich</td><td>47036</td><td></td></tr><tr><td>Cell ID Pt194</td><td>Fluidigm</td><td></td><td>Provided at 1&amp;nbsp;mM</td></tr><tr><td>Cell ID Pt195</td><td>Fluidigm</td><td></td><td>Provided at 1&amp;nbsp;mM</td></tr><tr><td>Cell ID Pt196</td><td>Fluidigm</td><td></td><td>Provided at 1&amp;nbsp;mM</td></tr><tr><td>Cisplatin</td><td>Enzo Life Sciences</td><td>ALX-400-040-M250</td><td>Soluble to 25&amp;nbsp;mg/mL in DMSO</td></tr><tr><td>Cell-ID 20-plex Pd Barcoding Kit</td><td>Fluidigm</td><td>201060</td><td></td></tr><tr><td>5-Iodo-2&#039;-deoxyuridine</td><td>Sigma-Aldrich</td><td>I7125</td><td>Soluble to 74&amp;nbsp;mg/mL in 0.2&amp;nbsp;N NaOH</td></tr><tr><td>Rhodium 103 intercelating agent</td><td>Fluidigm</td><td>201103A</td><td></td></tr><tr><td>Methanol</td><td>Fisher</td><td>BP1105-4</td><td>Chill at -20&amp;nbsp;&amp;deg;C before use</td></tr><tr><td>Sodium Azide</td><td>Sigma-Aldrich</td><td>S2002</td><td></td></tr><tr><td>5 mL round bottom tubes</td><td>Falcon</td><td>352058</td><td></td></tr><tr><td>5 mL 35 &amp;#956;m filter cap tubes</td><td>Falcon</td><td>352235</td><td></td></tr><tr><td>EQ four element calibration beads</td><td>Fluidigm</td><td>201078</td><td></td></tr><tr><td>Hyaluronidase Type I-S</td><td>Sigma-Aldrich</td><td>H3506</td><td></td></tr><tr><td>Collagenase from Clostridium histolyticum</td><td>Sigma-Aldrich</td><td>C9891</td><td></td></tr><tr><td>Disposable Scalpel #10</td><td>Sigma-Aldrich</td><td>Z69239</td><td></td></tr></tbody></table>

sample preparation for mass cytometry analysis

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities for deep analysis well beyond what is possible with conventional fluorescence-based flow cytometry. As with any new technology, there are critical steps that help ensure the reliable generation of high-quality data. Presented here is an optimized protocol that incorporates multiple techniques for the processing of cell samples for mass cytometry analysis. The methods described here will help the user avoid common pitfalls and achieve consistent results by minimizing variability, which can lead to inaccurate data. To inform experimental design, the rationale behind optional or alternative steps in the protocol and their efficacy in uncovering new findings in the biology of the system being investigated is covered. Lastly, representative data is presented to illustrate expected results from the techniques presented here.

The protocol presented here can be broadly applied to a wide variety of cultured and primary cell samples with only minor modifications. Depending upon the requirements of the experiment, it can be performed in a modular manner when certain elements, such as barcoding or cell surface staining, are not necessary. Utilized in its entirety this protocol enables the preparation of multiplexed mass cytometry samples labeled for dead cell exclusion, S-phase population identification and stained...

Watch this Scientific Journal Video about Sample Preparation for Mass Cytometry Analysis at JoVE.com

Sample Preparation for Mass Cytometry Analysis

This article describes the collection and processing of samples for mass cytometry analysis.

Mass cytometry utilizes antibodies conjugated with heavy metal labels, an approach that has greatly increased the number of parameters and opportunities ...

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Biology

16.0K Views.  The University of Texas MD Anderson Cancer Center. This article describes the collection and processing of samples for mass cytometry analysis.

Video: Sample Preparation for Mass Cytometry Analysis

Mass Cytometry: Protocol for Daily Tuning and Running Cell Samples on a CyTOF Mass Cytometer

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the issue of spectral overlap of fluorophore emissions has limited the number of simultaneous probes. In contrast, the new CyTOF mass cytometer by DVS Sciences couples a liquid single-cell introduction system to an ICP-MS.1 Rather than fluorophores, chelating polymers containing highly-enriched metal isotopes are coupled to antibodies or other specific probes.2-5 Because of the metal purity and mass resolution of the mass cytometer, there is no "spectral overlap" from neighboring isotopes, and therefore no need for compensation matrices. Additionally, due to the use of lanthanide metals, there is no biological background and therefore no equivalent of autofluorescence. With a mass window spanning atomic mass 103-203, theoretically up to 100 labels could be distinguished simultaneously. Currently, more than 35 channels are available using the chelating reagents available from DVS Sciences, allowing unprecedented dissection of the immunological profile of samples.6-7
Disadvantages to mass cytometry include the strict requirement for a separate metal isotope per probe (no equivalent of forward or side scatter), and the fact that it is a destructive technique (no possibility of sorting recovery). The current configuration of the mass cytometer also has a cell transmission rate of only ~25%, thus requiring a higher input number of cells.
Optimal daily performance of the mass cytometer requires several steps. The basic goal of the optimization is to maximize the measured signal intensity of the desired metal isotopes (M) while minimizing the formation of oxides (M+16) that will decrease the M signal intensity and interfere with any desired signal at M+16. The first step is to warm up the machine so a hot, stable ICP plasma has been established. Second, the settings for current and make-up gas flow rate must be optimized on a daily basis. During sample collection, the maximum cell event rate is limited by detector efficiency and processing speed to 1000 cells/sec. However, depending on the sample quality, a slower cell event rate (300-500 cells/sec) is usually desirable to allow better resolution between cells events and thus maximize intact singlets over doublets and debris. Finally, adequate cleaning of the machine at the end of the day helps minimize background signal due to free metal.

The steps necessary for daily tuning and optimization of the performance of a CyTOF mass cytometer are described. Comments on optimal sample preparation and flow rate are discussed

In recent years, the rapid analysis of single cells has commonly been performed using flow cytometry and fluorescently-labeled antibodies. However, the ...

Bioengineering

Isolation and Staining of Mouse Skin Keratinocytes for Cell Cycle Specific Analysis of Cellular Protein Expression by Mass Cytometry

The goal of this protocol is to detect and quantify protein expression changes in a cell cycle-dependent manner using single cells isolated from the epidermis of mouse skin. There are seven important steps: separation of the epidermis from the dermis, digestion of the epidermis, staining of the epidermal cell populations with cisplatin, sample barcoding, staining with metal tagged antibodies for cell cycle markers and proteins of interest, detection of metal-tagged antibodies by mass cytometry, and the analysis of expression in the various cell cycle phases. The advantage of this approach over histological methods is the potential to assay the expression pattern of &#62;40 different markers in a single cell at different phases of the cell cycle. This approach also allows for the multivariate correlation analysis of protein expression that is more quantifiable than histological/imaging methods. The disadvantage of this protocol is that a suspension of single cells is needed, which results in the loss of location information provided by the staining of tissue sections. This approach may also require the inclusion of additional markers to identify different cell types in crude cell suspensions. The application of this protocol is evident in the analysis of hyperplastic skin disease models. Moreover, this protocol can be adapted for the analysis of specific sub-type of cells (e.g., stem cells) by the addition of lineage-specific antibodies. This protocol can also be adapted for the analysis of skin cells in other experimental species.

This protocol describes how to isolate skin keratinocytes from mouse models, to stain with metal-tagged antibodies, and to analyze stained cells by mass cytometry in order to profile the expression pattern of proteins of interest in the different cell cycle phases.

The goal of this protocol is to detect and quantify protein expression changes in a cell cycle-dependent manner using single cells isolated from the ...

Developmental Biology

Collection, Isolation, and Flow Cytometric Analysis of Human Endocervical Samples

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female genital tract (FGT). Because heterosexual transmission now represents the dominant mechanism of HIV transmission, and given the continual spread of sexually transmitted infections (STIs), it is critical to understand the interplay between host and pathogen at the genital mucosa. The substantial gaps in knowledge around FGT immunity are partially due to the difficulty in successfully collecting and processing mucosal samples. In order to facilitate studies with sufficient sample size, collection techniques must be minimally invasive and efficient. To this end, a protocol for the collection of cervical cytobrush samples and subsequent isolation of cervical mononuclear cells (CMC) has been optimized. Using ex vivo flow cytometry-based immunophenotyping, it is possible to accurately and reliably quantify CMC lymphocyte/monocyte population frequencies and phenotypes. This technique can be coupled with the collection of cervical-vaginal lavage (CVL), which contains soluble immune mediators including cytokines, chemokines and anti-proteases, all of which can be used to determine the anti- or pro-inflammatory environment in the vagina.

The use of cytobrush sampling to collect lymphocytes and monocytes from the endocervix is a minimally invasive technique that provides samples for analysis of female genital tract immunity. In this protocol, we describe the collection of cytobrush samples and immune cell isolation for flow cytometry assays.

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female ...

Medicine

Identification and Isolation of Burst-Forming Unit and Colony-Forming Unit Erythroid Progenitors from Mouse Tissue by Flow Cytometry

Early erythroid progenitors were originally defined by their colony-forming potential in vitro and classified into burst-forming and colony-forming &#34;units&#34; known as BFU-e and CFU-e. Until recently, methods for the direct prospective and complete isolation of pure BFU-e and CFU-e progenitors from freshly isolated adult mouse bone marrow were not available. To address this gap, a single-cell RNA-seq (scRNAseq) dataset of mouse bone marrow was analyzed for the expression of genes coding for cell surface markers. This analysis was combined with cell fate assays, allowing the development of a novel flow cytometric approach that identifies and allows the isolation of complete and pure subsets of BFU-e and CFU-e progenitors in mouse bone marrow or spleen. This approach also identifies other progenitor subsets, including subsets enriched for basophil/mast cell and megakaryocytic potentials. The method consists of labeling fresh bone marrow or spleen cells with antibodies directed at Kit and CD55. Progenitors that express both these markers are then subdivided into five principal populations. Population 1 (P1 or CFU-e, Kit+ CD55+ CD49fmed/low CD105med/high CD71med/high) contains all of the CFU-e progenitors and may be further subdivided into P1-low (CD71med CD150high) and P1-hi (CD71high CD150low), corresponding to early and late CFU-e, respectively; Population 2 (P2 or BFU-e, Kit+ CD55+ CD49fmed/low CD105med/high CD71low CD150high) contains all of the BFU-e progenitors; Population P3 (P3, Kit+ CD55+ CD49fmed/high CD105med/low CD150low CD41low) is enriched for basophil/mast cell progenitors; Population 4 (P4, Kit+ CD55+ CD49fmed/high CD105med/low CD150high CD41+) is enriched for megakaryocytic progenitors; and Population 5 (P5, Kit+ CD55+ CD49fmed/high CD105med/low CD150high CD41-) contains progenitors with erythroid, basophil/mast cell, and megakaryocytic potential (EBMP) and erythroid/ megakaryocytic/ basophil-biased multipotential progenitors (MPPs). This novel approach allows greater precision when analyzing erythroid and other hematopoietic progenitors and also allows for reference to transcriptome information for each flow cytometrically defined population.

Here, we describe a novel flow cytometric method for prospective isolation of early burst-forming unit erythroid (BFU-e) and colony-forming unit erythroid (CFU-e) progenitors directly from fresh mouse bone marrow and spleen. This protocol, developed based on single-cell transcriptomic data, is the first to isolate all the tissue's erythroid progenitors with high purity.

Early erythroid progenitors were originally defined by their colony-forming potential in vitro and classified into burst-forming and colony-forming ...

Single-cell Analysis of Immunophenotype and Cytokine Production in Peripheral Whole Blood via Mass Cytometry

Cytokines play a pivotal role in the pathogenesis of autoimmune diseases. Hence, the measurement of cytokine levels has been the focus of multiple studies in an attempt to understand the precise mechanisms that lead to the breakdown of self-tolerance and subsequent autoimmunity. Approaches thus far have been based on the study of one specific aspect of the immune system (a single or few cell types or cytokines), and do not offer a global assessment of complex autoimmune disease. While patient sera-based studies have afforded important insights into autoimmunity, they do not provide the specific cellular source of the dysregulated cytokines detected. A comprehensive single-cell approach to evaluate cytokine production in multiple immune cell subsets, within the context of &#34;intrinsic&#34; patient-specific plasma circulating factors, is described here. This approach enables monitoring of the patient-specific immune phenotype (surface markers) and function (cytokines), either in its native &#34;intrinsic pathogenic&#34; disease state, or in the presence of therapeutic agents (in vivo or ex vivo).

Here we describe a single-cell proteomic approach to evaluate immune phenotypic and functional (intracellular cytokine induction) alterations in peripheral whole blood samples, analyzed via mass cytometry.

Cytokines play a pivotal role in the pathogenesis of autoimmune diseases. Hence, the measurement of cytokine levels has been the focus of multiple studies ...

Immunology and Infection

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...

Low-Input Metabolic Studies in Mice Cell Types

Targeted Metabolomics on Rare Primary Cells

Cellular function critically depends on metabolism, and the function of the underlying metabolic networks can be studied by measuring small molecule intermediates. However, obtaining accurate and reliable measurements of cellular metabolism, particularly in rare cell types like hematopoietic stem cells, has traditionally required pooling cells from multiple animals. A protocol now enables researchers to measure metabolites in rare cell types using only one mouse per sample while generating multiple replicates for more abundant cell types. This reduces the number of animals that are required for a given project. The protocol presented here involves several key differences over traditional metabolomics protocols, such as using 5 g/L NaCl as a sheath fluid, sorting directly into acetonitrile, and utilizing targeted quantification with rigorous use of internal standards, allowing for more accurate and comprehensive measurements of cellular metabolism. Despite the time required for the isolation of single cells, fluorescent staining, and sorting, the protocol can preserve differences among cell types and drug treatments to a large extent.

Author Spotlight: Advancing Metabolomics Analysis of Rare Hematopoietic Stem Cells 

Here, we present a protocol to accurately and reliably measure metabolites in rare cell types. Technical improvements, including a modified sheath fluid for cell sorting and the generation of relevant blank samples, enable a comprehensive quantification of metabolites with an input of only 5000 cells per sample.

Cellular function critically depends on metabolism, and the function of the underlying metabolic networks can be studied by measuring small molecule ...

Measurement of Mitochondrial Mass and Membrane Potential in Hematopoietic Stem Cells and T-cells by Flow Cytometry

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production of all mature blood cells. In recent years cellular metabolism has emerged as a crucial regulator of HSC function and fate. We have previously demonstrated that modulation of mitochondrial metabolism influences HSC fate. Specifically, by chemically uncoupling the electron transport chain we were able to maintain HSC function in culture conditions that normally induce rapid differentiation. However, limiting HSC numbers often precludes the use of standard assays to measure HSC metabolism and therefore predict their function. Here, we report a simple flow cytometry assay that allows reliable measurement of mitochondrial membrane potential and mitochondrial mass in scarce cells such as HSCs. We discuss the isolation of HSCs from mouse bone marrow and measurement of mitochondrial mass and membrane potential post ex vivo culture. As an example, we show the modulation of these parameters in HSCs via treatment with a metabolic modulator. Moreover, we extend the application of this methodology on human peripheral blood-derived T cells and human tumor infiltrating lymphocytes (TILs), showing dramatic differences in their mitochondrial profiles, possibly reflecting different T cell functionality. We believe this assay can be employed in screenings to identify modulators of mitochondrial metabolism in various cell types in different contexts.

Here we describe a reliable method to measure mitochondrial mass and membrane potential in ex vivo cultured hematopoietic stem cells and T cells.

A fine balance of quiescence, self-renewal, and differentiation is key to preserve the hematopoietic stem cell (HSC) pool and maintain lifelong production ...

Single-Cell Proteomics Preparation for Mass Spectrometry Analysis Using Freeze-Heat Lysis and an Isobaric Carrier

Single-cell proteomics analysis requires sensitive, quantitatively accurate, widely accessible, and robust methods. To meet these requirements, the Single-Cell ProtEomics (SCoPE2) protocol was developed as a second-generation method for quantifying hundreds to thousands of proteins from limited samples, down to the level of a single cell. Experiments using this method have achieved quantifying over 3,000 proteins across 1,500 single mammalian cells (500-1,000 proteins per cell) in 10 days of mass spectrometer instrument time. SCoPE2 leverages a freeze-heat cycle for cell lysis, obviating the need for clean-up of single cells and consequently reducing sample losses, while expediting sample preparation and simplifying its automation. Additionally, the method uses an isobaric carrier, which aids protein identification and reduces sample losses.
This video protocol provides detailed guidance to enable the adoption of automated single-cell protein analysis using only equipment and reagents that are widely accessible. We demonstrate critical steps in the procedure of preparing single cells for proteomic analysis, from harvesting up to injection to liquid chromatography-tandem mass spectrometry (LC-MS/MS) analysis. Additionally, viewers are guided through the principles of experimental design with the isobaric carrier, quality control for both isobaric carrier and single-cell preparations, and representative results with a discussion of limitations of the approach.

In this protocol, we describe how to prepare mammalian cells for single-cell proteomics analysis via mass spectrometry using commercially available reagents and equipment, with options for both manual and automatic pipetting.

Single-cell proteomics analysis requires sensitive, quantitatively accurate, widely accessible, and robust methods. To meet these requirements, the ...

High-Quality Sample Preparation for Multiome 10X Genomics Analysis

High-Quality Brain and Bone Marrow Nuclei Preparation for Single Nuclei Multiome Assays

Single-cell analysis has become the approach of choice for unraveling the complexity of biological processes that require assessing the variability of individual cellular responses to treatment or infection with single-cell resolution. 
Many techniques for single-cell molecular profiling have been developed over the past 10 years, and several dedicated technologies have been commercialized. The 10X Genomics droplet-based single-cell profiling is a widespread technology that offers ready-to-use reagents for transcriptomic and multi-omic single-cell profiling. The technology includes workflows for single-cell and single-nuclei RNA sequencing (scRNA-Seq and snRNA-Seq, respectively), scATAC-Seq, single-cell immune profiling (BCR/TCR sequencing), and multiome. The latter combines transcriptional (scRNA-Seq) and epigenetic information (scATAC-Seq) coming from the same cell. 
The quality (viability, integrity, purity) of single-cell or single-nuclei suspensions isolated from tissues and analyzed by any of these approaches is critical for generating high-quality data. Therefore, the sample preparation protocols should be adapted to the particularities of each biological tissue and ensure the generation of high-quality cell and nuclei suspensions.
This article describes two protocols for preparing brain and bone marrow samples for the downstream multiome 10X Genomics pipeline. The protocols are performed stepwise and cover tissue dissociation, cell sorting, nuclei isolation, and quality control of prepared nuclei suspension that is used as starting material for cell partitioning and barcoding, library preparation, and sequencing. These standardized protocols produce high-quality nuclei libraries and robust and reliable data.

Author Spotlight: Advancing Biomedical Research Through Single Cell Analysis 

The success of single-cell/single-nuclei transcriptomics and multi-omics largely depends on the quality of cells/nuclei. Therefore, isolating cells/nuclei from tissue and their purification must be highly standardized. This protocol describes the preparation of nuclei from the brain and bone marrow for downstream single-nuclei multiome assay.

Single-cell analysis has become the approach of choice for unraveling the complexity of biological processes that require assessing the variability of ...

Sample Preparation for Mass Cytometry Analysis

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