Characterizing Mutational Load and Clonal Composition of Human Blood

Samples must be obtained in accordance with appropriate ethics protocols, and donors must give informed consent prior to the procedure.

1. Preparation of Sample Material

NOTE: When working with freshly obtained material, start with step 1.1. When working with frozen material, start with step 1.2.

1. Preparing fresh bone marrow, cord blood, or peripheral blood
   1. Isolate the mononuclear fraction from the sample using density gradient separation by following the manufacturers’ instructions (see Table of Materials), and count the mononuclear cells using a hemocytometer. After isolation of the mononuclear cells, continue with step 1.3.
   2. OPTIONAL: The recommended number of cells required to sort a full 384 well plate of HSPCs is 1–2 x 10⁷. If more cells are isolated during density gradient centrifugation, store the surplus of cells in liquid nitrogen.
   3. Resuspend cells in 500 μL of IMDM + 10% FBS per 1 x 10⁷ cells, and add drop-by-drop an equal volume of IMDM + 30% FBS + 20% DMSO to achieve a suspension of 1 x 10⁷ cells in 1 mL of IMDM + 20% FBS + 10% DMSO.
   4. Immediately transfer the mononuclear cells to 1 mL cryogenic vials and freeze cells at -80 °C in a controlled-rate cell freezing container overnight. Transfer the cells the next day to a liquid nitrogen storage upon further processing.
2. Preparing frozen mononuclear cells from bone marrow, cord blood, or peripheral blood
   1. Prepare 50 mL of cell thawing medium containing 45 mL of Iscove’s Modified Eagle’s Medium (IMDM) and 5 mL of fetal bovine serum (FBS), and warm in 37 °C water bath.
   2. Take the vial containing the sample from liquid nitrogen storage, transfer the sample to dry ice, and thaw as quickly as possible in a 37 °C water bath.
   3. When the sample is almost thawed, wipe the vial with 70% ethanol and transfer its contents to a 50 mL conical tube. Rinse the vial with 1 mL of pre-warmed IMDM + 10% FBS to collect the remaining cells, and add this solution dropwise (5 s per drop) to the thawed sample while gently swirling the tube.
   4. Add an additional pre-warmed 15 mL of IMDM + 10% FBS dropwise to the sample while gently swirling the tube.
   5. Pellet the cells by centrifugation for 5 min at 350 x g.
   6. Remove all but ±3 mL of the supernatant. Resuspend the cells in the remaining supernatant and dilute by adding 20 mL of IMDM + 10% FBS drop-by-drop while gently shaking the tube.
   7. Take 10 μL of the cell suspension for cell counting. Dilute these 10 μL by adding 20 μL of 0.4% trypan blue solution and count the cells using a hemocytometer. The cell number can decrease upon thawing, with up to 50% cell loss after thawing. Cell viability should range between 70% and 90%.
3. If working with bone marrow or umbilical cord blood cells, take up 5 x 10⁶ mononuclear cells for MSC culture (step 2.1). If working with peripheral blood, take up 2–5 x 10⁶ cells for T-cell isolation (step 2.2)
4. Pellet remaining cells 5 min at 350 x g and resuspend in 3 mL of FACS buffer (0.05% BSA + 1 mM EDTA in PBS).
5. Transfer 1 x 10⁵ cells to a microtube filled with 200 μL of FACS buffer, which will serve as a negative control for flow cytometry (step 3.8), and keep on ice.

2. Cell Culture

NOTE: To obtain catalogues of somatically acquired mutations, donor-specific germline variation needs to be filtered out. When starting with bone marrow biopsies or umbilical cord blood, mesenchymal stromal cells (MSCs) can be used as matched control to filter for germline variation. In this case, follow section 2.1. When using (mobilized) peripheral blood follow step 2.2 to isolate and use T-cells as matched control sample to filter for germline variation (Figure 1). The bulk T-cell population will share the same lineage relationship as HSPCs.

1. MSC culture
   1. Prepare 50 mL of MSC medium containing 45 mL of DMEM/F12 medium, 10% FBS, 500 μL of of 200 mM L-glutamine or L-glutamine alternative, and 500 μL of penicillin/streptomycin solution.
   2. Plate approximately 5 x 10⁵ mononuclear cells in 1.5 mL of MSC medium per well. Place the cells in a humidified incubator at 37 °C with 5% CO₂.
   3. Replace the medium after 24 h, and subsequently replace medium every 3 days to ensure that all hematopoietic cells are washed off. Continue to culture until the confluency is 100%.
   4. If the MSCs are confluent, wash cells with 1 mL of PBS and harvest the MSCs by adding 200 μL of trypsin or trypsin alternative per well. Incubate cells for 5 min at 37 °C. Add 800 μL of MSC medium, and pipet the cells up and down to loosen cells from the well plate.
   5. Transfer MSCs to microcentrifuge tube and pellet the cells by centrifugation for 5 min at 350 x g. Remove the supernatant and continue with DNA isolation or store the pellet at -20 °C for later DNA isolation (section 4).
2. T-cell isolation
   NOTE: If using (mobilized) peripheral blood, T-cells can be isolated and used as germline control.
   1. Resuspend the cell pellet in 100 μL of anti-CD3 staining solution (1:100 dilution of anti-CD antibody in FACS buffer).
   2. Wash the cells by adding 1 mL of FACS buffer. Pellet the cells by centrifugation for 5 min at 350 x g and resuspend in 300 μL of FACS buffer.
   3. Isolate at least 5 x 10⁵ CD3+ cells using a FACS-sorter in a 5 mL polystyrene tube pre-filled with 1 mL of FBS.
   4. Pellet the sorted cells using centrifugation for 5 min at 350 x g, remove the supernatant, and continue directly with DNA isolation (section 4) or store the pellet at -20 °C for later DNA isolation.

3. HSPC Isolation, Sorting, and Culture

1. Spin down at 1–2 x 10⁷ mononuclear cells for 5 min at 350 x g and resuspend in 50 μL of FACS buffer (see step 2.2.1). Transfer the cells to a microcentrifuge tube.
   NOTE: When sorting with >2 x 10⁷ cells, increase the antibody mix and FACS buffer volumes accordingly.
2. Prepare 50 μL of 2x HSC staining mix according to the recipe seen in Table 1.

Table 1: HSC sorting mix. Shown is a table indicating the dilutions of antibodies used to sort the HSCs.

3. Mix 50 μL of cell solution with the prepared HSC staining mix and incubate the cells for 15 min at room temperature (RT) or for 1 h on ice for the antibodies to bind.
4. Wash the cells by adding 1 mL of FACS buffer and pellet by centrifuging for 5 min at 350 x g.
5. Resuspend the cells in 300 μL of FACS buffer and filter the cell suspension through a 35 µm cell strainer-capped 5 mL polystyrene tube to remove cell clumps before fluorescence-activated cell sorting (FACS).
6. Prepare 25 mL of HSPC culture medium, consisting of 1x SFEM medium supplemented with 100 ng/mL SCF, 100 ng/mL Flt3, 50 ng/mL TPO, 10 ng/mL IL-3, 20 ng/mL IL- 6, and 100 ng/mL antibiotic formulation (see Table of Materials).
7. Fill a 384 well cell culture plate with 75 μL of HSPC culture medium in each well.
   NOTE: To prevent evaporation of the medium in the outer wells, fill the outer wells with 75 μL of sterile water or PBS, and do not use these wells for cell sorting.
8. Sorting single HSPCs
   1. Set gates for the HSPC sorting based on an unstained control (step 1.9) and 10,000 cells from the stained sample. A representative result for setting gates is depicted in Figure 1. Gate single cells by drawing a gate around the linear FSC-height vs. FSC-area fraction. Use unstained control fraction to draw gate for lineage^– fraction. Draw gates for CD34⁺ cells and further characterize this subset by setting a specific gate for CD38^– CD45RA^– cells.
   2. Load the 384 well plate on the FACS machine and sort single cells.
      NOTE: If applicable to the FACS-machine, toggle on the option to keep index sorting data to enable re-tracing of the sorted cells.
9. Culturing singly-sorted HSCs
   1. Directly transfer the 384 well plate to a humidified 37 °C incubator with 5% CO₂.
      NOTE: To prevent evaporation during culture, wrap the 384 well culture plate (with lid) in transparent polyethylene wrap.
   2. Keep the 384 well plate in the incubator for 3–4 weeks until visible clones appear. Representative images of clonal culture are depicted in Figure 2. Based on the condition of the input material 5%–30% of sorted cells will clonally expand.

4. Harvesting HSPC Clones

1. After 4 weeks of culturing, determine which wells have a confluency of 30% or higher.
2. Pre-fill (for each clonal outgrowth) 1.5 mL microtubes with 1 mL of 1% BSA in PBS and label the tube according to the corresponding well.
3. Pre-wet a pipette tip with 1% BSA in PBS to minimize the number of cells sticking to the pipette tip.
4. Pipet up/down the medium in the well fiercely (at least 5 times) with a 200 μL pipette (set at 75 μL) and scrape the bottom of the well to loosen cells in the well, and collect the cell suspension in the labeled microtube corresponding to the well.
5. Take up 75 μL of fresh 1% BSA in PBS and repeat pipetting in the well to ensure maximum uptake of cells.
   NOTE: Clonally cultured cells can stick to the bottom of well. Inspect the wells using a standard inverted light microscope to ensure whether all cells have been collected.
6. If all wells with >30% confluency have been harvested, place the 384 well plate back in incubator. Clonal cultures can proliferate for up to 5 weeks.
7. Spin down the cell suspension for 5 min at 350 x g. A small pellet should be visible.
8. Carefully remove all but about 5 μL of supernatant. Cell pellets can be frozen at -20 °C and stored for multiple months before DNA isolation.

5. DNA Isolation

1. Isolate HSPC and MSC/T-cell DNA using a micro-scale DNA isolation kit according to the manufacturer’s instruction with the following adjustments:
   1. Add 2 μL of RNase A after addition of buffer AL during section 2. Incubate for 2 min before adding proteinase K.
   2. Incubate for 30 min at 56 °C instead of 10 min.
   3. Elute the DNA by loading the column with 50 μL of TE buffer with low EDTA (10 mM Tris, 0.1 mM EDTA). For optimal elution, reload the eluate again on the column and spin again.
2. Determine the DNA concentration using a DNA measuring 2 μL per clone. The DNA yield typically varies between 0.5–3 ng/μL.

6. Sequencing

1. Perform DNA sequencing as described by Jager et al.¹³

7. Mapping and Somatic Mutation Calling

1. Map the output of sequencing (FASTQ files) to the reference genome and call mutations as described in Jager et al.¹³
2. Inspect the data for aberrant karyotypic changes in sequenced clone and bulk data using a copy number analysis tool, such as Control-FreeC¹⁴. Until now, we have not reported any HSPCs with karyotypic aberrances.
3. Generate a blacklist, which consists of a panel of unmatched normal samples for filtering purposes from an own set of samples, as previously described¹³, or use the following uploaded blacklist: <https://data.mendeley.com/datasets/9y4yhwt5rp/3>.
4. Filter single nucleotide variations using SNVFI <https://github.com/ToolsVanBox/SNVFI>.
   1. Preset SNVFI.config file, such that all paths to helper functions are correct.
   2. Run SNVFI with .ini file configured according to the settings seen in Supplemental File 1 (SNVFI.ini). To exclude in vitro induced mutations we filter for a VAF ≥0.3⁹.
5. Check the variant allele fraction (VAF) output of SNVFI (Figure 4). Check whether the peak of the density plot is near 0.5, indicating the sample is clonal.
6. OPTIONAL: To determine the part of the genome which is covered during filtering, determine the callable regions along germline and control using CallableLoci from GATK (Genome Analysis ToolKit):
   java -jar GenomeAnalysisTK.jar
           -T CallableLoci
           -R reference.fasta
           -I myreads.bam
           -summary table.txt
           -o callable_status.bed
7. OPTIONAL: Retrieve the Callable regions from the CallableLoci output and perform pairwise intersections between the samples and the bulk using the python script CallableLoci_processor.py present at https://github.com/ToolsVanBox/CallableLoci_processor. The resulting bed files can be used to further filter the output of SNVFI and to inspect the mutational profile in section 9:
   CallableLoci_processor.py dir_in dir_out sample_name bulk_name –samples sample1 sample2 sample3

8. Indel Calling

1. Select all insertions and deletions (Indels) in the raw_variants.vcf file using GATK SelectVariants:
   java -Xmx12G
           -jar GenomeAnalysisTK.jar
           -T SelectVariants
           -R reference_genome.fasta
           -V raw_variants.vcf
           -o raw_INDELs.vcf
           -selectType INDEL
2. Filter raw_INDELS.vcf list using INDELFI <https://github.com/ToolsVanBox/INDELFI>:
   perl INDELFI.pl -i input.vcf (from step 8.1)
           -s column test sample
           -c column control sample

9. Mutational Profile Inspection

1. Use the resulting .vcf files from SNVFI output from step 7.6 (or from step 7.9 with optional callable loci analysis) to analyze the genomic mutational profile, mutation types, and signature analysis using the R package MutationalPatterns¹⁵: <http://bioconductor.org/packages/release/bioc/html/MutationalPatterns.html>. For representative output that can be produced with the resulting .vcf file such as a 96-trinucleotide mutational spectrum, see Figure 5.

10. Construction of a Developmental Lineage Tree Using Base Substitutions

1. To construct a developmental lineage tree, detect shared mutations between clones. Mutations present in the first branches of the lineage tree can also be subclonally present in the bulk sample (MSCs/T-cells). Later branching lineages will be defined by mutations shared by HSPC clones only.
2. To identify mutations that are present in a subset of the clones and subclonally present in the bulk, perform the following steps.
3. In order to filter for somatic mutations shared between clones, run the filterSomatic.py script in a Unix-based terminal. The script can be found at https://github.com/ToolsVanBox/filterSomatic. Before running this script, edit the filterSomatic.ini file (see Supplemental File 2) to set the paths and adjust the other parameters.
4. Run filterSomatic.py (python3 filterSomatic.py -i filterSomatic.ini).
5. Filter for mutations that are subclonally present in the bulk using the Determine_lowVAF_bulk.R script in a Unix-based terminal. The script can be found at https://github.com/ToolsVanBox/Identify_lowVAF_bulk_muts. This will generate separate .vcf files for shared and unique SNVs:
   Rscript Determine_lowVAF_bulk.R
           –vcf Path/To/Filter_somatic_output.vcf
           –bulk bulk_name
           –sample_name sample-name
           –gender [M|F]
           –out_dir out_dir
6. Determine all mutations shared between clones which are not present in the bulk sample by overlapping all mutation positions (concatenate column 1 and 2 of SNVFI output).
7. Exclude false positives obtained during steps 10.5 and 10.6 by manual inspection using IGV¹⁶. Mutations are considered false when not present, when the mutation is present in the germline or when present in poorly mapped regions, see Figure 7.
   NOTE: We highly recommend to re-sequence all shared loci independently using targeted or sanger sequencing.
8. Use the shared mutations obtained during steps 10.1 and 10.2 to build a binary table of mutations versus sequenced clones, with 0 indicating that the mutation is not present and 1 indicating presence of the mutation.
9. Output the mutation binary table as in a heatmap together with a dendrogram indicating lineage relationships between cells using R. The heatmap indicates mutations status for each cell. See the output of this function (Figure 6).
           
   Clones <- read.table(“Path/To/BinaryTable”)
           
   my_palette <- colorRampPalette(c("#cccccc", "#333333"))(n = 2)
   col_breaks <- c(0,0.5,1)
           
   heatmap.2(clones, distfun=function(x) dist(x,method = 'binary'),
           hclustfun=function(x) hclust(x,method = average),
           dendrogram = "column", Rowv = F,
           col=my_palette, breaks=col_breaks,
           trace="none", density.info="none")