This document provides an optimized protocol for preparing inexpensive Nextera mate pair libraries with improved performance for scaffolding genomes. Key optimizations include adjusting the tagmentation reaction to maximize DNA in the target size range, performing multiple rounds of Covaris shearing to achieve a narrower insert size distribution peaked at 450-500bp, and reducing PCR cycles to 10 or less to limit duplicates. The protocol aims to improve scaffolding by focusing on read pairs that contain junction adaptors through these adjustments to yield and read length.

1. iMate Protocol (version 2.0) by GRAS – April 11, 2016
NGS and Phyloinfo in Kobe
http://www.clst.riken.jp/phylo/
1
iMate Protocol: Improved and Inexpensive NexteraTM
Mate Pair Library Preparation
Authorized by Kaori Tatsumi, Osamu Nishimura, Kazu Itomi, Chiharu Tanegashima & Shigehiro Kuraku
Genome Resource & Analysis Station (GRAS)
operated by Phyloinformatics Unit
RIKEN Center for Life Science Technologies (CLST)
Notice: A benchmark paper introducing this protocol has been published in the journal Biotechniques. When
you present or publish data based on technical guidance in this protocol, you could think about citing this
protocol at our web site and the benchmark paper (Tatsumi et al., 2015) published in Biotechniques.
This protocol outlines the modifications to the ‘Gel-plus’ version of the standard protocol for
Nextera Mate Pair Library Preparation and the logical background for them. Focusing on how to
best improve scaffolding performance, we optimized the protocol under the possibly
conservative policy that only read pairs with junction adaptors (bona fide ‘mates’) should be
passed on to scaffolding. The keys were optimizing the 1) tagmentation condition, 2) Covaris
shearing condition, and 3) sequence read length, in order to enhance the yield of libraries and
the capability of detecting the junction adaptor in reads.
Basically, we understand that 4μg of starting genomic DNA, as formulated in the standard
protocol, is enough for preparation of mate-pair libraries with mate distance of >10kb. Ideally, we
could optimize the tagmentation condition so that as much DNA as possible fall into the targeted
size range. For this purpose, perform tagment reaction with multiple conditions; for example, in
three tubes with 4, 8 and 12 μl of kit supplied tagment enzyme respectively. The tagment buffer
can be self-made [1], which leads to cost-saving, if other limiting reagents are also saved.
Size distribution of tagmented DNA molecules should be analyzed with a trustworthy method,
such as pulse field electrophoresis (e.g., PippinPulse) or the Agilent TapeStation―the Agilent
Bioanalyzer does not perform well for this purpose. With comparable results from multiple
tagment reactions, you could figure out which tagment condition allows you to retrieve the
largest amount of DNA for the targeted size range.
Like the previous tagmentation step, the amounts of the supplied reagents used in this step are
the limiting factor in terms of how many libraries can be prepared with one purchased kit. Thus,
it would be preferable to find a way to decrease the amount of kit-supplied reagents required to
perform this step. We previously suggested (in the iMate protocol versions 1.X) to perform
strand displacement with 1/4 volume of all reaction components after size selection with
BluePippin. However, we have recently found that this can result in contamination by read pairs
with untargeted mate distances. Therefore, we currently do not recommend reversing the order
of strand displacement and size selection. We are now looking into alternative ways to save kit-
supplied reagents in the strand displacement step.
Do as instructed in the standard protocol.
We use a BluePippin in this step and usually set a width size range of 4kb (for example, from 6
kb to 10 kb), although this is a matter of further consideration. After strand displacement and
size selection, it is ideal to retain at least 100 ng of DNA. Although the standard protocol
mentions ‘150-400 ng’ (on page 27), 100-200ng is realistic and still promising in our experience.

2. iMate Protocol (version 2.0) by GRAS – April 11, 2016
NGS and Phyloinfo in Kobe
http://www.clst.riken.jp/phylo/
2
Do as instructed in the standard protocol.
Shearing determines the length of library inserts, which should ideally be coordinated with the
selected sequencing read length. If you regard only reads with an adaptor junction as true mate
pairs, we propose a shearing condition which will ultimately produce a library possessing an
insert size distribution of 300 – 700bp, with the peak at 450-500bp. Note that this is markedly
different from the size distribution illustrated in the standard protocol (300-1200bp; on page 49).
To achieve our proposed size distribution, we recommend performing successive shearing via
multiple executions of the Covaris condition instructed in the standard protocol. In our
experience, shearing the genomes of different species with the same condition can result in
markedly different fragment size distributions. Thus, you need to optimize the condition
specifically for your species of interest. For one of the species we worked on, we performed as
many as 7 runs of Covaris shearing with the condition specified in the standard protocol.
You may feel an urge to perform QC with Bioanalyzer immediately after the Covaris shearing,
but it will not give you a fair assessment of shearing results unless you use a large quantity of
your sheared DNA for QC, which is undesirable. Thus, we recommend to save as much DNA as
possible at this stage, and to instead measure the size distribution later in the ‘
’ step.
Do as instructed in the standard protocol.
To get as many unique mate-pair reads as possible, it is strongly recommended to reduce PCR
cycles and avoid excessive amplification. We suggest performing no more than 10 cycles of
PCR. This warning is supported by our experience of getting a sufficient amount of products with
10 PCR cycles, even for samples that are supposed to require 15 cycles according to the
standard protocol (for example, 100ng for libraries with mate distant ranges of 6-10kb; see [2] for
details of cycle number estimates). In fact, we normally perform 8 PCR cycles, and only when
we find the yield too low after AMPure clean-up do we perform additional PCR cycling (still, no
more than 10 cycles in total). If you do not get enough products within 10 cycles, you had better
first optimize the tagment condition to increase the yield for the targeted size range.
With the illumina system, it seems that the insert lengths of many reads actually sequenced are
shorter than the most frequent insert length of a library. Thus, be sure to perform greedy size
selection with AMPure to get rid of molecules with short inserts, as instructed in the standard
protocol (x0.67 AMPure to get rid of <300bp molecules), no matter what the size distribution of
library inserts is. Modest size selection can result in high proportion of read pairs with too small
lengths, and they may not suffice for effective scaffolding.
Use Bioanalyzer or equivalent in this final QC before sequencing. Keep in mind that the size
distribution is determined mostly by shearing condition and AMPure clean-up, rather than the
choice of size range of mate distance.

3. iMate Protocol (version 2.0) by GRAS – April 11, 2016
NGS and Phyloinfo in Kobe
http://www.clst.riken.jp/phylo/
3
We use KAPA Library Quantification Kit (KK4835) in this step. Quantification should not be tricky
if the library should have an ordinary unimodal size distribution. The standard protocol says that
you need 1.5nM-20nM of the synthesized library, but we think that 2nM is enough unless the
sequencing facility you are working with requests much more than required in an actual
sequencing run.
In your first trial, it is advised to run a MiSeq for small-scale pilot sequencing to get 300nt-long
paired-end reads from prepared libraries―sequencing as many as 10 libraries per MiSeq run
should provide you with fair validation of the libraries. Obtained 300nt-long paired-end reads
could also be used for simulating which read length yields the highest proportion of reads with
junction adaptor; for example, by chopping them at 100nt, 127nt and 171nt (if sequencing with
HiSeq is planned next).
The lengths of 127nt and 171nt may sound unusual, but with Rapid Run on HiSeq one can
obtain reads of these lengths by making the best use of the extra cycles inherently assigned for
Nextera dual indexing, which we do not need in mate-pair sequencing. This trick allows you to
get 127nt and 171nt by using three and four of the TruSeq Rapid SBS Kit for 50 cycles,
respectively (see page 6 of the official manual for TruSeq Rapid SBS Kit). Please consult with
the sequencing facility that you plan to work with about the possibility of this extra-cycle
sequencing. The intention of getting 127nt or 171nt is to increase the proportion of reads with
the junction adaptor inside, but if one plans to use all obtained reads regardless of junction
adaptor inclusion, it may be wiser to respect cost-saving and go for 100nt or shorter reads.
In our experience, Rapid Run mode with v1 chemistry on older HiSeq Control Software (HCS) is
vulnerable to suboptimal library pooling, such as the ‘low plex pooling’ issue (see this document
by illumina). In the course of your mate pair sequencing, you may encounter a situation in which
you have only 4 or fewer libraries to be sequenced in a Rapid Run. In this case there is a high
chance that the base composition of index reads will be too homogeneous, and you will get
lower QV in index reads, resulting in a larger proportion of reads that failed in demultiplexing. To
reduce this unfavorable effect, you could introduce multiple indices per library in the step above
. As long as demultiplexing between libraries works out without any
overlap of indices, this strategy is supposed to produce as many valid reads as possible, with
the only drawback being the handling of more data files in post-sequencing informatics steps.
The latest versions of HCS (version 2.2.38 or higher) seems to be robust against low diversity
samples, so you are suggested to contact the sequencing facility you are working with in
advance to check if you need to be concerned with the low plex pooling issue.
We recommend to first run a recent version of FastQC (v0.11 or higher) on raw fastq files to
monitor some standard metrics, including the frequency of junction adaptor appearance along
base positions (in the ‘Adapter Content’ view newly added in FastQC v0.11).
After the primary QC, run a read processing program, such as NextClip [2], and assess PCR
duplicate rate and what proportion of reads has the junction adaptors. After read processing, be
sure to rerun FastQC on the processed fastq files, in order to confirm that junction/external
adaptors and low-quality bases were properly trimmed.
1. Wang Q, Gu L, Adey A, Radlwimmer B, Wang W, Hovestadt V, Bahr M, Wolf S, Shendure J, Eils R et al:
Tagmentation-based whole-genome bisulfite sequencing. Nature protocols 2013, 8(10):2022-2032.
2. Heavens D, Garcia Accinelli G, Clavijo B, and Derek Clark M: A method to simultaneously construct up to
12 differently sized Illumina Nextera long mate pair libraries with reduced DNA input, time, and cost.
BioTechniques 2015, 59(1):42-45.
3. Leggett RM, Clavijo BJ, Clissold L, Clark MD, Caccamo M: NextClip: an analysis and read preparation tool
for Nextera Long Mate Pair libraries. Bioinformatics 2014, 30(4):566-568.