Traditionally, pan-genomes for bacteria are gene based. That is, you study the set of genes found at a certain taxonomic unit, see Tettelin et al., 2015 for a review.

In this practical, we explore ways of implementing such a gene-level pan-genome in the vg software ecosystem. This allows us to detect the presence/absence of genes as well as to model sequence variation within genes. This is important since key traits, such as drug resistance, can be conferred by both transfer of whole genes but also by specific mutations (Blair et al., 2015).

Data

Today, we will mostly work on E. coli data. On your workstations you find the following data sets:

• /media/gtpb_shared_drive/To_Participant/bacteria/ncbi-whole-genomes one file for each complete E. coli strain present at NCBI, containing the complete reference genome for that strain,
• /media/gtpb_shared_drive/To_Participant/bacteria/ncbi-genes one file for each of these NCBI E. coli strain containing one line per gene,
• /media/gtpb_shared_drive/To_Participant/bacteria/reads whole genome sequence data for a (random) subset of 10 E.coli strains for this study: Earle et al., 2015,
• /media/gtpb_shared_drive/To_Participant/bacteria/contigs the result of running the Minia3 assembler on the reads provided in the above directory.

Objectives

• Build a graph-based representation of a single gene (of your choice),
• Come up with ways of determining whether a gene is absent/present in a given sample,
• Build a whole bacterial pan-genome.
• Use this representation to find a set of essential genes (present in all strains) as opposed to accessory genes (missing in some strains).

Ideas

• Do a pooled assembly of the 10 strains
• Pool the contigs after assembly, potentially iterate the assembly process, i.e. build a cDBG from contigs
• Use variant calls to linear reference genome and use vg construct to build a pan-genome
• Use vg vectorize to distinguish core /accessory genome after mapping contigs into the graph (i.e. constructed from all of the assemblies)
• Use an existing polished reference genome as a starting point (e.g. to align contigs to and then augment)
• Start from all the known gene sequences and use vg msga to build gene models. The expectation is that the different genes would end up in different components. But this process might be slow, DBG-based methods might be faster. Maybe start from a subset.
• Place contigs along a reference genome to avoid spurious contig-to-contig alignments
• Map reads to the pan-genome model
• Select one gene (gyrA) and align all sequences of this gene (from NCBI) from all strains progressively
• Use this gene model for genotyping
• Count gene identify in NCBI gene set and take the most frequent one, map reads to it (make sure to filter out false positive mappings)

Hints

GFA input to vg from minia and bcalm

You can read the minia3 assemblies into GFA and then vg using these commands. (The same is true for bcalm, and can be done on the unitig or contig sets from both of these assemblers.)

First we convert the graph to GFA:

convertToGFA.py SRR3050857_merged.fastq.contigs.fa SRR3050857_merged.fastq.contigs.gfa 51

Note that we used kmer size 51 for the assemblies, and you will need to change this parameter to convertToGFA.py if you use a different k.

Now we fix up the ID space to make vg happy (it can’t handle node id == 0) and feed the result into vg:

cat SRR3050857_merged.fastq.contigs.gfa \
    | awk '$1=="L" { $2 +=1 ; $4+=1 } $1=="S" { $2+=1 } { print }' | tr ' ' '\t' \
    | vg view -Fv - >SRR3050857_merged.fastq.contigs.vg

It’s now possible to view the graph in GFA format in Bandage to ensure that the conversion worked.

vg view SRR3050857_merged.fastq.contigs.vg >SRR3050857_merged.fastq.contigs.+1.gfa
Bandage &

Pruning assembly graphs

Assembly graphs can have rather complicated regions in them. We need to get rid of these to run alignment against the graph. In contrast to typical pruning, we may need to do some pruning to the base reference graph that we feed to vg map, and further pruning to the graph that’s given to GCSA indexing. If this is a problem, we can remove high-degree nodes (nodes with many edges) from the graph:

vg mod -M 8 raw.vg >out.vg

In any event, we also need to “chop” nodes to be shorter than a given length, so that GCSA2 can index the graph. We also and sort the output to make the ids space increase through the linear components of the graph, which helps the mapper.

vg mod -X 32 raw.vg | vg sort - >out.vg

Bringing it together, we want to first remove the high-degree nodes from what will become our “reference” graph for a particular analysis, build the xg index from that, then prune and build the GCSA index on the pruned graph. For mapping, we bring these two results back together:

vg view -Fv base.gfa | vg mod -X 32 - | vg mod -M 8 - | vg sort - >base.vg
vg index -x base.xg base.vg
vg prune -k 16 -e 3 base.vg >prune.vg
vg index -g base.gcsa -k 16 prune.vg
vg map -f reads.fq.gz -x base.xg -g base.gcsa >aln.gam

Another consideration is that it’s often very difficult to map paired end reads against the graph. Single end mapping is fine.

Viewing paths with Bandage

You can view the position of the nodes along the paths in the graph using Bandage. First use these commands to insert the paths into the graph:

xg -g graph.gfa -o graph.gfa.xg
xg -i graph.gfa.xg -G > graph.withpaths.gfa

Then create a CSV file with the node labels:

grep -P '^S' < graph.withpaths.gfa | cut -f 2,4 | sed '1iNode\tPath\' | sed 's/\\t/\t/g' > graph.paths.csv

Then load the CSV into Bandage using File->Load CSV data and select the file. Check the box “CSV data:” in the left panel under “Node labels”. The label for each node will show the paths that pass through the node: name, orientation (+ forward / - backward) and position along the path.

Filtering alignments

You may want to filter alignments to only keep those with a positive score (that are mapped). One easy way to do this is to use jq. If we want to keep alignments that are aligned we can do the following:

vg view -a aln.gam | jq -cr 'select(.score > 0)' | vg view -JaG - >aln.filt.gam

Combining graphs together

vg has the join command, but this is probably not what you want as it does the strange operation of linking all the subgraphs to a single head node.

Instead, it’s possible to combine graphs by concatenating them via cat. However, they must have separate id spaces or they will overwrite each other. To make a single joint id space, use vg ids -j, passing the files you want to join together on the command line:

vg construct -v tiny/tiny.vcf.gz -r tiny/tiny.fa >tiny.vg
cp tiny.vg 1.vg; cp tiny.vg 2.vg; cp tiny.vg 3.vg
# these are the same
vg stats -lz 1.vg
cat 1.vg 2.vg 3.vg | vg stats -lz -
# joining the id space allows us to merge the graphs safely
vg ids -j 1.vg 2.vg 3.vg
# now we get a different, bigger graph that contains three copies of input graph
cat 1.vg 2.vg 3.vg | vg stats -lz -

How fast is my alignment going?

You can check how many reads per second you are aligning using:

vg map -d x -f reads.fq.gz -j | pv -l >/dev/null

Extrace gene sequences

for f in *.gff.gz; do b=$(echo $f | cut -d '.' -f 1,2); \
awk '{if ($3 == "gene") print;}' < $f | sed 's/ID=[^;]*;\(Dbxref=[^;]*;\)\?Name=\([^;]*\);gbkey=[^;]*;\(gene=[^;]*;\)\?gene_biotype=[^;]*;locus_tag=.*/\2/g' | awk '{print $1 "\t" $2 "\t" $9 "\t" $4 "\t" $5 "\t" $6 "\t" $7 "\t" $8}' > fix.gff; \
bedtools getfasta -fi $b.fna -bed fix.gff -name > $b.fa; \
done;

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