Assembly Parameters

The parameters contained in a params file affect the actions that are performed during each step of an ipyrad assembly. The defaults that we chose are fairly reasonable values for most assemblies, however, you will always need to modify at least a few of them (for example, to indicate the location of your data), and often times you will want to modify many of the parameters. The ability to easily assemble your data set under a range of parameter settings is one of the main features of ipyrad.

Below is an explanation of each parameter setting, the steps of the assembly that it effects, and example entries for the parameter into a params.txt file.

0. Assembly name

The Assembly name is used as the prefix for all output files. It should be a unique identifier for the assembly, meaning the set of parameters you are using for the current data set. When I assemble multiple data with different parameter combinations I usually either name them consecutively (e.g., data1, data2), or with names indicating their parameter combinations (e.g., data_clust90, data_clust85). The Assembly name cannot be changed after an Assembly is created with the -n flag, but a new Assembly with a different name can be created by branching the Assembly (see branching workflow).

Affected steps: 1-7 Example: new Assemblies are created with the -n or -b options to ipyrad:

>>> ipyrad -n data1                       ## create a new assembly named data1
>>> ipyrad -p params-data1.txt -b data2   ## create a branch assembly named data2

1. Project dir

A project directory can be used to group together multiple related Assemblies. A new directory will be created at the given path if it does not already exist. A good name for Project_dir will generally be the name of the organism being studied. The project dir path should generally not be changed after an analysis is initiated, unless the entire directory is moved to a different location/machine.

Affected steps: 1-7 Example entries into params.txt:

/home/deren/ipyrad/tests/finches   ## [1] create/use project dir called finches
finches                            ## [1] create/use project dir called finches

2. Raw fastq path

This is a path to the location of raw (non-demultiplexed) fastq data files. If your data are already demultiplexed then this should be left blank. The input files can be gzip compressed (i.e., have name-endings with .gz). If you enter a path for raw data files then you should also enter a path to a barcodes file. To select multiple files, or all files in a directory, use a wildcard character (*).

Affected steps = 1 Example entries into params.txt:

/home/deren/ipyrad/tests/data/*.fastq.gz     ## [2] select all gzip data files
~/ipyrad/tests/data/*.fastq.gz               ## [2] select all gzip data files
./ipsimdata/rad_example*.fastq.gz            ## [2] select files w/ `rad_example` in name

3. Barcodes path

This is a path to the location of a barcodes_file_. This is used in step1 for demuliplexing, and can also be used in step2 to improve the detection of adapter/primer sequences that should be filtered out. If your data are already demultiplexed the barcodes path can be left blank.

Affected steps = 1-2. Example entries into params.txt:

/home/deren/ipsimdata/rad_example_barcodes.txt    ## [3] select barcode file
./ipsimdata/rad_example_barcodes.txt              ## [3] select barcode file

4. Sorted fastq path

This is a path to the location of sorted fastq data. If your data are already demultiplexed then this is the location from which data will be loaded when you run step 1. A wildcard character can be used to select multiple files in directory.

Affected steps = 1 Example entries into params.txt:

/home/deren/ipyrad/tests/ipsimdata/*.fastq.gz    ## [4] select all gzip data files
~/ipyrad/tests/ipsimdata/*.fastq                 ## [4] select all fastq data files
./ipsimdata/rad_example*.fastq.gz                ## [4] select files w/ `rad_example` in name

5. Assembly method

There are four Assembly_methods options in ipyrad: denovo, reference, denovo+reference, and denovo-reference. The latter three all require a reference sequence file (param #6) in fasta format. See the Tutorials (running ipyrad) for an example.

Affected steps = 3, 6 Example entries into params.txt:

denovo                            ## [5] denovo assembly
reference                         ## [5] reference assembly
denovo+reference                  ## [5] reference addition assembly
denovo-reference                  ## [5] reference subtraction assembly

6. Reference sequence

The reference sequence file should be in fasta format. It does not need to be a complete nuclear genome, but could also be any other type of data that you wish to map RAD data to; for example plastome or transcriptome data.

~/ipyrad/tests/ipsimdata/rad_example_genome.fa   ## [6] select fasta file
./data/finch_full_genome.fasta                   ## [6] select fasta file

7. Datatype

There are now many forms of restriction-site associated DNA library preparation methods and thus many differently named data types. Currently, we categorize these into six data types. Follow the link to deteremine the appropriate category for your data type.

rad                       ## [7] rad data type (1 cutter, sonication)
pairddrad                 ## [7] paired ddrad type (2 different cutters)
pairgbs                   ## [7] paired gbs type (1 cutter cuts both ends)

8. Restriction_overhang

The restriction overhang is used during demultiplexing (step1) and also to detect and filter out adapters/primers (in step2), if the filter_adapters parameter is turned on. Identifying the correct sequence to enter for the restriction_overhang can be tricky. You do not enter the restriction recognition sequence, but rather the portion of this sequence that is left attached to the sequenced read after digestion. For example, the enzyme PstI has the following palindromic sequence, with ^ indicating the cut position.

5'...C TGCA^G...'3
3'...G^ACGT C...'5

Digestion with this enzyme results in DNA fragments with the sequence CTGCA adjacent to the cut site, which when sequenced results in the reverse complement TGCAG as the restriction overhang at the beginning of each read. The easiest way to identify the restriction overhang is simply to look at the raw (or demultiplexed) data files yourself. The restriction overhang will be the (mostly) invariant sequence that occurs at the very beginning of each read if the data are already demultiplexed, or right after the barcode in the case of non-demultplexed data. Use the command below to peek at the first few lines of your fastQ files to find the invariant sequence.

## gunzip decompresses the file,
## the flag -c means print to screen,
## and `less` tells it to only print the first 100 lines
zless 100 my_R1_input_file.fastq.gz

This will print something like the following. You can see that each of the lines of sequence data begins with TGCAG followed by variable sequence data. For data that used two-cutters (ddrad), you will likely not be able to see the second cutter overhang for single-end reads, but if your data are paired-end, then the _R2_ files will begin with the second restriction_overhang. The second restriction_overhang is only used to detect adapters/primers if the filter_adapters parameter is set > 1. The second restriction_overhang can optionally be left blank.

@HWI-ST609:152:D0RDLACXX:2:2202:18249:93964 1:N:0:
TGCAGCAGCAAGTGCTATTCGTACAGTCATCGATCAGGGTATGCAACGAGCAGAAGTCATGATAAAGGGTCCCGGTCTAGGAAGAGACGCAGCATTA
+
BDFFHHHHHJJJHIJJJJJIJIGJJJJJJJJJJJJJJJJDGGIJJJJJHIIIJJJJHIIHIGIGHHHHFFFFEDDDDDACCDCDDDDDDDDDBBDC:
@HWI-ST609:152:D0RDLACXX:2:2202:18428:93944 1:N:0:
TGCAGGATATATAAAGAATATACCAATCCTAAGGATCCATAGATTTAATTGTGGATCCAACAATAGAAACATCGGCTCAACCCTTTTAGTAAAAGAT
+
ADEFGHFHGIJJJJJJIJJIIJJJIJJIJGIJJJJJJJJIJJIJJJJIIIGGHIEGHJJJJJJG@@CG@@DDHHEFF>?A@;>CCCDC?CDDCCDDC
@HWI-ST609:152:D0RDLACXX:2:2202:18489:93970 1:N:0:
TGCAGCCATTATGTGGCATAGGGGTTACATCCCGTACAAAAGTTAATAGTATACCACTCCTACGAATAGCTCGTAATGCTGCGTCTCTTCCTAGACC
+
BDFFHHHHHJJJJIJJIJJJJJJJHIJJJJJJJJHIIJJJJIFHIJJJJFGIIJFHIJJIJJIFBHHFFDFEBACEDCDDDDBBBDCCCDDDCDDC:
@HWI-ST609:152:D0RDLACXX:2:2202:18714:93960 1:N:0:
TGCAGCATCTGGAAATTATGGGGTTATTTCACAGAAGCTGGAATCTCTTGGGCAATTTCACAGAATCTGGGAATATCTGGGGTAAATCTGCAAGATC
+
BDEFHHHHHJJJIJJJJJJJJJJCGIIJJGHJJJJJJJJJJIJJIJJJIHIJJJJJJJHHIIJJJJJJJHGHHHFEFFFDEDABDDFDDEDDDDDDA
@HWI-ST609:152:D0RDLACXX:2:2202:20484:93905 1:N:0:
TGCAGAGGGGGATTTTCTGGAGTTCTGAGCATGGACTCGTCCCGTTTGTGCTGCTCGAACACTGACGTTACTTCGTTGATCCCTATGGACTTGGTCA
+
ADEEHHHHGJJHIIJJJJJJIJFHJIIIJJJJIIIJJJJEHHIIGHIGGHGHHHHFFEEEDDDD;CBBDDDACC@DC?<CDDCCCCCCA@CCD>>@:
@HWI-ST609:152:D0RDLACXX:2:2202:20938:93852 1:N:0:
TGCAGAAGCTGGAGATTCTGGGGCAGCTTTGCAGCAAGCTGAAAATTCTGGGGGTCGATCTGCAAGATCGGAAGAGCGGTTCAGCAGGAATGCCGAG

In some cases restriction enzymes can bind to more than one specific sequence, for example ApoI will bind to AATTY (i.e. AATTC and AATTT). If you used an enzyme with reduced specificity you can include ambiguity codes in the restriction overhang sequence.

Affected steps = 1,2. Example entries to params.txt file:

TGCAG                     ## [8] single cutter (e.g., rad, gbs)
TGCAG, AATT               ## [8] double digest (e.g., ddrad, pairddrad)
CWGC                      ## [8] single cutter w/ degenerate base

NB: 3RAD and SeqCap data can use up to 4 restriction enzymes. If you have this kind of data, simply list all the restriction overhangs for all your cutters.

CTAGA, CTAGC, AATTC               ## [8] 3rad data (multiple cutters)

9. max_low_qual_bases

During step 2 bases are trimmed from the 3’ end of reads when the quality score is consistently below 20 (which can be modified by modifying phred_Qscore_offset). However, your reads may still contain some number of ambiguous (N) sites that were not trimmed based on quality scores, and these will affect the efficiency and accuracy of clustering downstream. This parameter sets the upper limit on the number of Ns allowed in reads. The default value for max_low_qual_bases is 5. I would generally recommend against increasing this value greatly.

Affected steps = 2. Example entries to params.txt:

0                      ## [9] allow zero low quality bases in a read
5                      ## [9] allow up to five low quality bases in a read

10. Phred_Qscore_offset

Bases are trimmed from the 3’ end of reads if their quality scores is below this 20. The default offset for quality scores is 33. Some older data use a qscore offset of 64, but this is increasingly rare. You can toggle the offset number to change the threshold for trimming. For example, reducing the offset from 33 to 23 is equivalent to changing the minimum quality score from 20 to 10, which is approximately 95% probability of a correct base call.

Affected steps = 2. Example entries to params.txt:

33                 ## [10] default offset of 33, converts to min score=20
43                 ## [10] offset increased by 10, converts to min score=30
64                 ## [10] offset used by older data, converts to min score=20.

11. mindepth_statistical

This is the minimum depth at which statistical base calls will be made during step 5 consensus base calling. By default this is set to 6, which for most reasonable error rates estimates is approximately the minimum depth at which a heterozygous base call can be distinguished from a sequencing error.

Affected steps = 4, 5. Example entries to params.txt

6                 ## [11] set mindepth statistical to 6
10                ## [11] set to 10

12. mindepth_majrule

This is the minimum depth at which majority rule base calls are made during step 5 consensus base calling. By default this is set to the same value as mindepth_statistical, such that only statistical base calls are made. This value must be <= mindepth_statistical. If lower, then sites with coverage >= mindepth_majrule and < mindepth_statistical will make majority rule calls. If your data set is very low coverage such that many clusters are excluded due to low sequencing depth then lowering mindepth_majrule can be an effective way to increase the amount of usable information in your data set. However, you should be aware the majority rule consensus base calls will underestimate heterozygosity.

Affected steps = 4, 5. Example entries to params.txt:

6                 ## [12] set to relatively high value similar to mindepth_stat
2                 ## [12] set below the statistical limit for base calls.

13. maxdepth

Sequencing coverage is often highly uneven among due to differences in the rate at which fragments are amplified during library preparation, the extent to which varies across different library prep methods. Moreover, repetitive regions of the genome may appear highly similar and thus cluster as high depth clusters. Setting a maxdepth helps to remove the latter problem, but at the expense of potentially removing good clusters that simply were sequenced to high depth. The default maxdepth is set quite high (10,000), but you may change it as you see fit.

Affected steps = 4, 5. Example entries to params.txt:

10000             ## [13] maxdepth above which clusters are excluded.

14. clust_threshold

This the level of sequence similarity at which two sequences are identified as being homologous, and thus cluster together. The value should be entered as a decimal (e.g., 0.90). We do not recommend using values higher than 0.95, as homologous sequences may not cluster together at such high threshold due to the presence of Ns, indels, sequencing errors, or polymorphisms.

Affected steps = 3, 6. Example entries to params.txt:

0.90              ## [14] clust threshold set to 90%
0.85              ## [14] clust threshold set to 85%

15. max_barcodes_mismatch

The maximum number of allowed mismatches between the barcodes in the barcodes file and those found in the sequenced reads. Default is 0. Barcodes usually differ by a minimum of 2 bases, so I would not generally recommend using a value >2.

Affected steps = 1. Example entries to params.txt:

0              ## [15] allow no mismatches
1              ## [15] allow 1 mismatched base

16. filter_adapters

It is important to remove Illumina adapters from your data if present. Depending on the fidelity of the size selection procedure implemented during library preparation there is often at least some small proportion of sequences in which the read length is longer than the actual DNA fragment, such that the primer/adapter sequence ends up in the read. This occurs more commonly in double-digest (GBS, ddRAD) data sets that use a common cutter, and can be especially problematic for GBS data sets, in which short fragments are sequenced from either end. The filter_adapters parameter has three settings (0, 1, or 2). If 0, then reads are only removed if they contain more Ns than allowed by the max_low_qual_bases parameter. If 1, then reads are trimmed to the first base which has a Qscore < 20 (on either read for paired data), and also removed if there are too many Ns. If 2, then reads are searched for the common Illumina adapter, plus the reverse complement of the second cut site (if present), plus the barcode (if present), and this part of the read is trimmed. This filter is applied using code from the software cutadapt, which allows for errors within the adapter sequence.

Affected steps = 2. Example entries to params.txt:

0                ## [16] No adapter filtering
1                ## [16] filter based on quality scores
2                ## [16] strict filter for adapters

17. filter_min_trim_len

During step 2 if filter_adapters is > 0 reads may be trimmed to a shorter length if they are either low quality or contain Illumina adapter sequences. By default ipyrad will keep trimmed reads down to a minimum length of 35bp. If you want to set a higher limit you can do so here.

Affected steps = 2. Example entries to params.txt

50                ## [17] minimum trimmed seqlen of 50
75                ## [17] minimum trimmed seqlen of 75

18. max_alleles_consens:

This is the maximum number of unique alleles allowed in (individual) consens reads after accounting for sequencing errors. Default=2, which is fitting for diploids. At this setting any locus which has a sample with more than 2 alleles detected will be excluded/filtered out. If set to max_alleles_consens = 1 (haploid) then error-rate and heterozygosity are estimated with H fixed to 0.0 in step 4, and base calls are made with the estimated error rate, and any consensus reads with more than 1 allele present are excluded. If max_alleles_consens is set > 2 then more alleles are allowed, however, heterozygous base calls are still made under the assumption of diploidy i.e., hetero allele frequency=50%.

Affected steps = 4, 7. Example entries to params.txt

2                ## [18] diploid base calls, exclude if >2 alleles
1                ## [18] haploid base calls, exclude if >1 allele
4                ## [18] diploid-base calls, exclude if >4 alleles

19. max_Ns_consens:

The maximum number of uncalled bases allowed in consens seqs (R1, R2). If a base call cannot be made confidently (statistically) then it is called as ambiguous (N). You do not want to allow too many Ns in consensus reads or it will affect their ability to cluster with consensus reads from other Samples, and it may represent a poor alignment. Default is 5, 5. For single end data only the first value is used, for paired data the first value affects R1s and the second value affects R2s.

Affected steps = 5. Example entries to params.txt

2                ## [19] allow max of 2 Ns in a consensus seq
5, 5             ## [19] allow max of 5 Ns in a consensus seq (R1, R2)

20. max_Hs_consens:

The maximum number of heterozygous bases allowed in consens seqs (R1, R2). This filter helps to remove poor alignments which will tend to have an excess of Hs. Default is 8, 8. For single end data only the first value is used, for paired data the first value affects R1s and the second value affects R2s.

Affected steps = 5. Example entries to params.txt

2                ## [20] allow max of 2 Hs in a consensus seq
8, 8             ## [20] allow max of 8 Hs in a consensus seq (R1, R2)

21. min_samples_locus

The minimum number of Samples that must have data at a given locus for it to be retained in the final data set. If you enter a number equal to the full number of samples in your data set then it will return only loci that have data shared across all samples. Whereas if you enter a lower value, like 4, it will return a more sparse matrix, including any loci for which at least four samples contain data. This parameter is overridden if a min_samples values are entered in the popfile. Default value is 4.

Affected steps = 7. Example entries to params.txt

4                ## [21] create a min4 assembly
12               ## [21] create a min12 assembly

22. max_SNPs_locus

Maximum number of SNPs allowed in a final locus. This can remove potential effects of poor alignments in repetitive regions in a final data set by excluding loci with more than N snps. The default is 20, 20. Setting lower values is likely only helpful for extra filtering of very messy data sets. For single end data only the first value is used, for paired data the first value affects R1s and the second value affects R2s.

Affected steps = 7. Example entries to params.txt

20                  ## [22] allow max of 20 SNPs at a single-end locus.
20, 30              ## [22] allow max of 20 and 30 SNPs in paired locus.

23. max_Indels_locus

The maximum number of Indels allowed in a final locus. This helps to filter out poor final alignments, particularly for paired-end data. The default is 8,8. For single end data only the first value is used, for paired data the first value affects R1s and the second value affects R2s.

Affected steps = 7. Example entries to params.txt

5                ## [23] allow max of 5 indels at a single-end locus.
5, 10            ## [23] allow max of 5 and 10 indels in paired locus.

24. max_shared_Hs_locus

Maximum number (or proportion) of shared polymorphic sites in a locus. This option is used to detect potential paralogs, as a shared heterozygous site across many samples likely represents clustering of paralogs with a fixed difference rather than a true heterozygous site. Default is 0.5. For single end data only the first value is used, for paired data the first value affects R1s and the second value affects R2s.

Affected steps = 7. Example entries to params.txt

0.25             ## [24] allow hetero site to occur across max of 25% of Samples
10               ## [24] allow hetero site to occur across max of 10 Samples

25. trim_reads

Sometimes you can look at your fastq data files and see that there was a problem with the sequencing such that the cut site which should occur at the beginning of your reads is either offset by one or more bases, or contains many errors. You can trim off N bases from the beginning or end of R1 and R2 reads during step 2 by setting the number of bases here. This could similarly be used to trim all reads to a uniform length (though uniform read lengths are not required in ipyrad).

Affected steps = 2. Example entries to params.txt

0, 0, 0, 0       ## [25] does nothing
5, 0, 0, 0       ## [25] trims first 5 bases from R1s
5, -5, 0, 0      ## [25] trims first 5 bases and last five based from R1s
5, 80, 0, 0      ## [25] trims first 5 bases from R1s and trims maxlen to 80
5, 75, 5, 75     ## [25] trims first 5 from R1 and R1, and maxlen to 75.

26. trim_loci

Trim N bases from the edges of final aligned loci. This can be useful in denovo data sets in particular, where the 3’ edge of reads is less well aligned than the 5’ edge, and thus error rates are sometimes higher at the ends of reads.

Affected steps = 7. Example entries to params.txt

0, 0, 0, 0     ## [26] no locus edge trimming
0, 5, 0, 0     ## [26] trims first 5 bases from R1s in aligned locus
0, 5, 5, 0     ## [26] trims last 5 bases from R1s and first 5 from R2s

27. output_formats

Disk space is cheap, and these are quick to make, so by default we make all formats. More are coming (alleles, treemix, migrate-n, finestructure). The short list of available options is below but see output formats section for full descriptions of the available formats.

p: PHYLIP (Full dataset)
s: PHYLIP (SNPs only)
u: PHYLIP (One SNP per locus)
n: NEXUS
k: STRUCTURE
g: EIGENSTRAT .geno
G: G-PhoCS
v: VCF (SNPs only)

Affected steps = 7. Example entries to params.txt

*                     ## [27] Make all output datatypes
n, v, g               ## [27] Only write out nexus, vcf and geno formats
u,k                   ## [27] Only write out unlinked snps in phylip, and structure

28. pop_assign_file

Population assignment file for creating population output files, or assigning min_samples_locus value to each population. Enter a path to the file. (see below for details of the file).

Affected step: 7. Example entries to params.txt

/home/user/ipyrad/popfile.txt        ## [28] example...

The population assignment file should be formatted as a plain-txt, whitespace delimited list of individuals and population assignments. Care should be taken with spelling and capitalization. Each line should contain a sample name followed by a population name to which that sample is assigned. One or more additional lines should be included that start with one or more “#” characters. These special lines tell ipyrad how many samples must have data within each population for the locus to be retained in the final assembly, and thus assign different min_samples_locus values to each population. This will override the global min_samples_locus value.

See the example below.

Sample1 pop1
Sample2 pop1
Sample3 pop1
Sample4 pop2
Sample5 pop2

# pop1:2 pop2:2