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Next-generation mammalian genetics toward organism-levelsystems biology
Etsuo A. Susaki1,2,3, Hideki Ukai2 and Hiroki R. Ueda1,2
Organism-level systems biology in mammals aims to identify, analyze, control, and design molecular and cellular networks
executing various biological functions in mammals. In particular, system-level identiﬁcation and analysis of molecular and cellular
networks can be accelerated by next-generation mammalian genetics. Mammalian genetics without crossing, where all production
and phenotyping studies of genome-edited animals are completed within a single generation drastically reduce the time, space,and effort of conducting the systems research. Next-generation mammalian genetics is based on recent technologicaladvancements in genome editing and developmental engineering. The process begins with introduction of double-strand breaks
into genomic DNA by using site-speciﬁc endonucleases, which results in highly efﬁcient genome editing in mammalian zygotes orembryonic stem cells. By using nuclease-mediated genome editing in zygotes, or ~100% embryonic stem cell-derived mousetechnology, whole-body knock-out and knock-in mice can be produced within a single generation. These emerging technologies
allow us to produce multiple knock-out or knock-in strains in high-throughput manner. In this review, we discuss the basic concepts
and related technologies as well as current challenges and future opportunities for next-generation mammalian genetics inorganism-level systems biology.
npj Systems Biology and Applications (2017)3:15 ; doi:10.1038/s41540-017-0015-2

INTRODUCTION
Systems Biology is a natural extension of molecular and cellularbiology,1–3 which consists of multi-stage processes beginning witha (1) comprehensive identiﬁcation and (2) quantitative analysis ofindividual system components and their networked interaction,
which leads to the ability to (3) control existing systems toward
the desired state and (4) design new systems based on anunderstanding of the underlying structural and dynamical
principles. After identiﬁcation of key genes by classical forward
and reverse genetics, systems biology in mammals has beenfurther accelerated by a series of genome projects, especially at
the molecular-to-cellular levels, where in vitro cell culture systems
allow system-level identiﬁcation, analysis, control, and design ofmolecular networks. On the other hand, organism-level systems
biology in mammals still remains an important challenge inbiology.4
In order to identify and analyze molecular networks and/or
cellular circuits in organisms, gene knock-out (KO) or knock-in (KI)are powerful technologies often used in mammalian reverse
genetics. However, this classical genetics requires several generations of crosses to produce mutant animals of sufﬁcient quality andquantity for phenotype analysis. The time consuming conventional
methods for producing KO or KI mice usually involve targetingvector construction (2 weeks to a few months; depending on thecomplexity of constructs), the introduction of target mutations into
embryonic stem cells (ESCs) by homologous recombination (a few
weeks), and the injection of the mutant ESCs into wild-type
blastocysts to produce chimera mice (~3 weeks). If the mutant
ESCs contribute to the germ-line of the newborn chimera mice,

their next-generation offspring will possess a heterozygous
mutation (~3 months). Further crossings of the offspring (several
months to years; at least 3 months per generation) will produce
mice with completely homozygous KO or KI mutations on an
inbred genomic background, which is required for reliablephenotype analysis. Thus, conventional methods require substantial amounts of time, space, and effort to knock out or knock ineven a single gene. Therefore, to comprehensively identify andquantitatively analyze molecular networks and/or cellular circuits inorganisms in an efﬁcient manner will require next-generation
genetics, i.e., genetic alterations without crossing. In this review, we
discuss the basic concepts and related technologies as well as
current challenges and future opportunities for next-generation
mammalian genetics in organism-level systems biology.

CONVENTIONAL MAMMALIAN GENETICS
Mammalian genetics (particularly in mice) has been widely
exploited in order to investigate complex and dynamic biologicalprocesses executed by molecular networks and/or cellular circuits
in organisms. Forward genetics (germline mutagenesis and genetrap) and reverse genetics (targeted KO or KI) are available inmouse genetics as in other model organisms such as yeast,nematode and ﬂy. Especially, developmental engineering basedon the establishment of cultured ESCs was often used to generate
KO and/or KI mice.5–7 Various genetic tools can be also introducedby transgenic (Tg) mice techniques.8
However, the production of genome-edited mice has been
generally low-throughput, and needed huge time and effort in the1
Department of Systems Pharmacology, Graduate School of Medicine, The University of Tokyo, 7-3-1 Hongo, Bunkyo-ku, Tokyo 113-0033, Japan; 2Laboratory for Synthetic
Biology, RIKEN Quantitative Biology Center, 1-3 Yamadaoka, Suita, Osaka 565-0871, Japan and 3PRESTO, Japan Science and Technology Agency (JST), 4-1-8 Honcho, Kawaguchi,
Saitama 332-0012, Japan
Correspondence: Hiroki R. Ueda (uedah-tky@umin.ac.jp)

Received: 26 October 2016 Revised: 1 May 2017 Accepted: 4 May 2017

Published in partnership with the Systems Biology InstituteNext-generation mammalian genetics
EA Susaki et al.

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Published in partnership with the Systems Biology InstituteNext-generation mammalian genetics
EA Susaki et al.

Fig. 1 Conventional and next-generation mammalian genetics. a A typical procedure for conventional mouse genetics. Upper panel:
generation of a transgenic mouse, lower panel: gene targeting in ESCs and generation of the mutated mouse. An inbred strain such as C57BL/6(B6) is widely used for ﬁnal analysis, while hybrid or other inbred strains are used in the production stages for practical reasons. Therefore, aprolonged backcross procedure is needed in many cases. In addition, gene targeting in ESCs is dependent on a spontaneous DSB andfollowing HDR in the cells, causing an inefﬁcient targeting rate. b In next-generation mouse genetics, all of the crossing procedures are notneeded because of the use of an inbred strain for analysis, efﬁcient genome editing in zygotes or ESCs mediated by site-speciﬁcendonucleases, and one-step generation of the genome-edited bi-allelic KO mouse or KI ES mouse. These F0 animals can be used insubsequent phenotyping experiments

conventional ways (Fig. 1a). For example, a Tg mouse strain is
produced by pronuclear injection of a DNA fragment harboring atransgene, which is randomly integrated. Therefore, non-speciﬁc
expressions of the transgene are usually observed in the resultant
strain and the F0 founders must be selected and further expanded
for use as the strains for the subsequent research. In case of genetargeting in ESCs, a chimera mouse (mouse having both ESC andhost embryo-derived cells) is ﬁrst produced by injection of the
ESCs into blastocyst-stage embryos. If the injected ESCs by chancecontribute to germ-line cells, the resultant F0 chimera can transfer
the introduced mutation to the next F1 generation. Therefore, thehomozygous mutants can be obtained, in principle, at least in thethird (F2) generation, which takes 9 months after ESC injection.
However, these procedures are not robust and it usually takes
longer because of low targeting rates in ESCs, low germ-line
transmission rate in chimera, or unexpected infertility of thecreated mutant strain. Furthermore, because a mixed genetic
background can cause phenotypical alterations which make theexperimental results difﬁcult to interpret,9 the generated strains
additionally need to be backcrossed to a ‘standard’ inbred strain
such as C57BL/6 (hereafter denoted as B6) several times. This
labor-intensive step is practically required in most cases because
F1 hybrid strains or 129 strain-derived ESCs are commonly used inTg zygote production or targeting in ESCs, respectively due totheir higher viability or efﬁcient germ-line transmission in F0
chimera.
Despite the limitations of conventional mammalian genetics,
systematic, large-scale mouse genetics projects have been
performed. For example, ethyl-nitrosourea mutagenesis in micewas exploited to screen mammalian circadian clock genes10–12
and for systematic gene function studies.13, 14 The gene-trap
strategy has more recently been applied to such forward-genetics
approaches, and >100,000 of trapped ESC lines have been
established and kept in international organizations (e.g., International gene trap consortium or IGTC, http://www.genetrap.org).15
Other systematic international efforts to collect, prepare andmaintain mutant mice and ESCs have also been performed, suchas the International Knockout Mouse Consortium/International
Mouse Phenotype Consortium (http://www.mousephenotype.
org).16–19 Multiple Cre Tg/KI strains have also been establishedby individual researchers, institutes and international consortiums.20–22 However, to carry out organism-level systems biology,
these large-scale efforts should be scaled down to the singlelaboratory scale or even to the single-researcher scale. To address
this technological challenge, next-generation mammalian genetics
without crossing is proposed here to allow completion of KO or KI
mouse production and phenotyping analysis within the F0
generation (Fig. 1b). This can be realized by the application ofhighly efﬁcient genome editing by site-speciﬁc nucleases for onestep generation of whole-body genome-edited inbred animals
within a single generation. Recently, there has been rapid progressin next-generation mammalian genetics, as introduced below,
which will form an essential platform for organism-level systems
biology.
Published in partnership with the Systems Biology Institute

CURRENT TECHNOLOGIES FOR EFFICIENT GENOME EDITINGBY SITE-SPECIFIC ENDONUCLEASES
Double-strand breaks (DSBs) lead to several DNA repair pathways,
such as (1) homology-directed repair (HDR) where a homologous
DNA sequence is used for recombination or annealing,23 (2) nonhomologous end joining (NHEJ), where the broken ends are
directly reconnected with a frequent insertion or deletion of arandom number of bases (denoted as “indels”),24 (3)microhomology-mediated end joining (MMEJ), where a small
microhomology fragment is used for the end-joining.25 Induced
DSB and the following repair processes enable highly efﬁcientgenome editing (both KO or KI) at the broken locus. Although the
genome editing induced by DSBs was previously investigated byintroduction of I-SceI (a mitochondrial endonuclease from S.
cerevisiae) into cultured mammalian cells, which stimulated
extrachromosomal homologous recombination,26 DSB-induced
genome editing has been practically used in recent years. We
review such efﬁcient genome editing methods using site-speciﬁc
nucleases, which can accelerate the production of KO and KI mice
via relatively simple steps and thus help realize next-generation
mammalian genetics (Fig. 2).
Site-speciﬁc endonucleases used in modern genetics
Three major classes of site-speciﬁc endonucleases have been used
for genome-editing,27 zinc-ﬁnger nuclease (ZFN),28 transcription
activator-like effector nuclease (TALEN)29 and clustered regularlyinterspaced short palindromic repeats (CRISPR)-associated Protein9 (Cas9).30 ZFN and TALEN are categorized into customizable
endonucleases because they are composed of a customizable
sequence-speciﬁc DNA-binding domain fused to a nonspeciﬁc
DNA catalytic domain of FokI endonuclease.31 On the other hand,
Cas9 is a RNA-guided endonuclease and recruited to speciﬁc DNA
sequences by a short RNA guide molecule that recognizes target
DNA via base-pairing.32
The DNA-binding domain of ZFN contains 3–6 arrays of Cys2His2 zinc ﬁnger motif.33, 34 The individual zinc ﬁnger recognizes 3
bp in the major groove of DNA.35 Selected zinc-ﬁnger modules
that recognize nearly all of the 64 possible nucleotide triplets havebeen developed. Therefore, an assembled custom array of six zincﬁngers can be constructed to recognize a unique 18 bp
sequence.34, 36–38 This length of target sequence covers a possible
68 billion unique DNA sequences.
TALEN contains another type of customizable DNA-binding
domain, which is composed of a tandem 33–35-amino acid repeat(TALE repeat) derived from the plant pathogenic bacteria genus
Xanthomonas.39, 40 The individual TALE repeat recognizes a single
base pair via two hypervariable amino acid residues inside therepeat (repeat-variable di-residues or RVDs). The four mostcommon RVDs (HD, NG, NI, and NN) are known to recognize
each of the four nucleotides (C, T, A, and G). Therefore, the tandem
TALE-repeat is usually constructed with approximately 18 TALE
repeats of different base pair-binding speciﬁcities, under consideration of its limitation that TALE-binding sites should start witha T base. The TALE repeat domain generally gives similar DNAbinding speciﬁcity and more ﬂexibly when compared with ZFNs.41
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EA Susaki et al.

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DSB induction by site-specificendonucleases

Null mutation(with NHEJ)

ZFNIndel insertion

FokI
Genome DNA

TALEN
IndelsCRISPR/Cas9

Large fragment deletion

DSB

IndelsFragment insertion
Large fragment insertion(with HDR)

Large fragment insertion(with NHEJ)

Targeting vector(with long homology arm)
Donor plasmid(digested in vivo)
Inserted fragment

IndelsDonor fragment(PCR products,
double-strand ODN)
IndelsInserted fragment

Small fragment insertion(with HDR)

Large fragment insertion(with MMEJ)

ssODN

Inserted sequence(Intended mutation, protein tag, LoxP etc.)

Donor plasmid or fragment(with microhomology sequences)

Inserted fragment

Susaki, Ukai and Ueda
Fig. 2 DSB-mediated genome editing. Upper left: type of site-speciﬁcendonucleases which are recently used for efﬁcient genome editing
purposes. Upper right: introduction of a null mutation by DSB. Whenrepaired by NHEJ pathway, small deletion or insertion of nucleotides(indels) occurred at the joint site, which cause a nonsense or
missense mutation in the targeted ORF. Long deletions can also be
introduced by multiple DSBs. Lower panels: strategies of fragment
insertion. Homology-directed repair (HDR) supports insertion of alarge or a small fragment with homology sequences. NHEJ also
supports the insertion of a large fragment without homology
sequence, although inserted direction is not controllable and indels
are introduced at the joint regions. Microhomology-mediated end
joining (MMEJ) mediates fragment insertion with very short (10–40bp) microhomology arms and thus potentially ameliorates drawbacks in the other two pathways

npj Systems Biology and Applications (2017) 15

Dimerization of the FokI endonuclease catalytic domain isessential for cleavage of DNA by ZFN and TALEN.31 This meansthat two ZFN or TALEN molecules must bind on both right and left
sides of the target site with an appropriate orientation andspacing. Therefore, the dimer recognizes 2-fold longer sequenceat the target site than single ZFN or TALEN molecules. This
molecular property gives higher speciﬁcity and reduced off-target
effect.
Unlike the former molecules, Cas9 is an RNA-guided DNA
endonuclease derived from the type II bacterial adaptive immune
system CRISPR, and is recruited to speciﬁc target sequences by
two short RNA molecules:32 the CRISPR RNA (crRNA) whichanneals with the target sequence, and the trans-activating crRNA(tracrRNA) which is partially complementary to the crRNA andanneals to the crRNA. This two-component RNA system wasfurther simpliﬁed to synthetic single-guide RNA (sgRNA) consistingof a fusion of crRNA and tracrRNA.42 The target sequence in the
CRISPR/Cas9 system can be readily changed by simply redesigning a part (around 20 bp) of the crRNA or sgRNA. This
simplicity is in contrast to the much more burdensome proceduresin ZFN and TALEN vector construction. This simplicity endows the
CRISPR/Cas9 system with a signiﬁcant advantage for use as a sitespeciﬁc endonuclease for various genome editing purposes,
including multiple gene KO,43, 44 or even genome-wide gene
perturbations.45, 46
Many studies have tried to increase the ﬂexibility and decrease
any off-target effect of the CRISPR/Cas9 system for practical use.
The DNA cleavage activity of Cas9 molecules is dependent on thepresence of a short (around 2–6 nucleotides) protospacer adjacentmotif (PAM), which is located beside the complemental sequenceof crRNA/sgRNA-targeted region.47 PAM sequence varies according to the CRISPR-based systems and organisms, and restricts the
ﬂexibility of the target sequence. However, the sequence
dependency of the Cas9 molecule can be artiﬁcially modiﬁed
and such PAM engineering can expand the target range of thesystem.48, 49 Other modiﬁcation of Cas9 molecules contribute to
off-target suppression. Cas9 induces DSBs at approximately threebases upstream of the PAM by two endonuclease domains, a
RuvC-like endonuclease domain (RuvC domain) and a HNH-likeendonuclease domain (HNH domain), which are located at theamino terminus and the mid-region of the Cas9, respectively.50
The RuvC domain cleaves the non-complementary strand while
the HNH domain cleaves the crRNA-complementary strand.
Inactivation of these endonuclease domains via point mutations
can convert Cas9 endonuclease into a DNA “nickase” that creates asingle-stranded break, which reduces off-target activity by 50-fold
to 1500-fold in cell lines and zygotes without sacriﬁcing on-target
cleavage efﬁciency.51, 52 Others have tried FokI-dCas9 fusion
protein as a dimer to improve targeting speciﬁcity by theirrecognition of distinct sites.53, 54 Use of truncated sgRNA can also
suppress undesired off-target activity by >5000-fold without
sacriﬁcing on-target genome editing efﬁciency, possibly bydecreasing the sgRNA-DNA interface.55
Introduction of null mutation
DSBs induced by site-speciﬁc endonucleases activate an internal
DSB-repair pathway, which is exploited for efﬁcient genome
editing. Among them, NHEJ-dependent indel insertion is thesimplest and the most effective method for gene KO24, 25 (Fig. 2).
Indels in the open reading frame (ORF) of targeted gene lead toloss-of-function mutation by creating frame-shift mutations or an
accidental stop codon at the cleavage site. The NHEJ-dependent
gene KO works stably in mammalian cultured cells,28, 30, 56 ESCs
and other pluripotent cells, or even mammalian zygotes.57–59
Furthermore, simultaneous use of multiple sgRNAs can introduce
mutations in multiple genes and create a large deletion between
the targeted loci, as well as increase the KO efﬁciency43, 59–62
Published in partnership with the Systems Biology InstituteNext-generation mammalian genetics
EA Susaki et al.

5(Fig. 2). In a recent study, the improvement in KO efﬁciency withthe use of three sgRNAs (triple-CRISPR) was examined in depth.
Based on simulation, the average KO efﬁciency expected withsingle sgRNA was around 60% and triple-CRISPR would increase
the rate to over 80%. The actual rate reached to over 95% due tolong deletions between CRISPR targeted sites.62 A set of triple
sgRNAs which cover ~80% of all genes in the mouse genome has
been created as an open database (http://crispr.riken.jp/).
Potential off-target effects can be also excluded by using thesecond set of triple sgRNAs that covers ~70% of all mousegenes.62
Introduction of DNA fragments
Targeted insertion or KI of a DNA fragment with mutatedsequence, short functional sequence (restriction enzyme site,
recombinase recognition site, or protein tag etc.), or functional
expression cassette can be also facilitated via HDR, NHEJ andMMEJ by co-transfer of linear or circular donor vector, PCR
fragment or single-stranded oligo DNA nucleotide (ssODN)
together with the site-speciﬁc endonucleases (Fig. 2).
Homologous recombination (HR)-dependent targeting is
mediated by a form of HDR. This pathway has been widely used
for a large fragment insertion or KI both in cultured cells andzygotes by using a donor targeting vector with long homology
arms.43, 44, 63–69 The targeting rate is relatively low but efﬁciently
enriched by antibiotic drug selection in the culture. A shorteraB6, Tyr -/- (Triple-CRISPR KO)

functional sequence or small mutation can be more simply
introduced by using ssODN.51, 55, 59, 66, 70–75
NHEJ-mediated fragment insertion/KI is easier and more efﬁcientthan the HR pathway, because the NHEJ-repair reaction is thought
to predominate over the HR reaction for DSB repair.76, 77 In the
NHEJ-mediated insertion, both the donor plasmid and the target
genome loci are digested simultaneously. And then, the digested
donor plasmid is integrated into the digested genome loci. A PCR
fragment or double-stranded ODN can be also applied as anintegrated fragment without digestion. This pathway works not onlyin cultured mammalian cells (including ESCs) but also in zebraﬁsh,and does not necessarily require antibiotic selection.56, 78–81
In addition, there is no need to prepare a targeting vector withlong homology arms, which is generally a time-consuming
process. On the other hand, it is of note that the direction of
the inserted fragment is not controllable, and indels are usually
introduced at the junction site. Therefore, the method is
inappropriate for some KI purposes, such as in-frame KI of an
exogenous ORF into an endogenous gene.
MMEJ-mediated editing provides more simpliﬁed KI strategy
with precise direction and junction sequence. Instead of theconventionally used long homology arms for HR-mediated KI, thispathway uses only extremely short microhomology sequences(10–40 bp) for the precise fragment insertion. MMEJ-mediated KI
also works in mammalian cells, and the inserted fragment can besupplied as an in vivo digested plasmid or a PCR fragment.82, 83b

B6-derived ES mouse

GFP
DAPI

c

Host cell-derived

ESC-derived (GFP+)

Susaki, Ukai and Ueda
Fig. 3 One-step generation of genome-edited mice. a An example of triple-CRISPR KO littermates (before weaning) in B6 strain. Tyr genecoding tyrosinase (an enzyme involving black coat color) was knocked out by the triple-CRISPR method.62 All littermates had white coat color,
indicating ~100% bi-allelic KO rate of the targeted gene. b An example of B6 ES mouse littermates (before weaning) by 3i + LIF culture and 8cell injection.141 All littermates had black coat color, indicating efﬁcient generation of ~100% ESC-derived mouse. c An ES mouse embryo(E13.5) derived from an H2B-EGFP KI ESC clone.170 Only the embryo (but not the extraembryonic tissues) expresses EGFP, suggesting theunique contribution of ESC-derived cells. Entire section of the ES mouse is shown in the right panel. All animal experiments here wereapproved by the Institutional Animal Care and Use Committee of RIKEN Kobe Branch, and all of the animal care was in accordance with theInstitutional Guidelines
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Therefore, the editing pathway potentially overcomes problems in
HR-mediated or NHEJ-mediated KI.
CURRENT TECHNOLOGIES FOR ONE-STEP PRODUCTION OF
GENOME-EDITED MICE
Direct genome editing in one-cell zygotes
The compelling advantages of the site-speciﬁc endonucleases in
efﬁcient genome-editing has been examined in recent years. In
particular, zygotic genome editing enables one-step production of
genome-edited animals, skipping the in vitro targeting step in
ESCs. Introduction of components into one-cell zygotes are
relatively simple and easy, particularly for the CRISPR/
Cas9 system, which just requires cytoplasmic microinjection orelectroporation.84–87
KO animals can be generated in a one-step manner by
exploiting endonuclease-mediated DSBs followed by NHEJ withindel insertion in zygotes. In an earlier study, ZFN was tested in rat
zygotes,57 where up to 75% of live-born F0 founders were
harboring mutations. TALEN has been similarly tested,88 while
CRISPR/Cas9 was mainly used in the most recent studies, since2013.59, 60, 89, 90 This method accelerates the generation of KO animals
via the co-injection of RNA encoding the Cas9 protein and targetlocus-speciﬁc guide RNAs into embryos. Long deletions of a genomic
region (10–100 kb) were induced by using two sgRNAs.60, 91, 92
Others reported F0 phenotyping of CRISPR/Cas9 KO animals,93
suggesting the potential of this method for use in next-generation
genetics schemes. Several modiﬁcations of the CRISPR/
Cas9 system have been also introduced to improve the efﬁciency
and speciﬁcity of targeted mutations in a genome.51, 52, 55, 61
However, two problems have remained: (1) ﬁrst-generation mice
often contain a mosaic of wild-type and KO cells, and (2) the rateof whole-body bi-allelic mutant mice generated is relatively low(usually ~60–80% at best). Therefore, the highly efﬁcient (>90%)
production of whole-body bi-allelic KO in a single generation
remained a fundamental challenge for next-generation mammalian genetics. To realize this vision of next-generation mammalian
genetics, the triple-CRISPR method signiﬁcantly improved bi-allelicmodiﬁcation efﬁciency and further elicited almost perfect (~100%)whole-body bi-allelic KO mice62 (Fig. 3a). It is of note that this was
performed with B6 zygotes so that the resulted KO animals could
be used for the subsequent experiments without backcross. Taken
together, next-generation mammalian genetics has beenachieved, at least for the production of KO mice (Fig. 1b).
On the other hand, one-step production of KI mice (zygotic KI) is
still under development. An earlier study using ZFN reported that
the KI mice were generated in one-step manner at the production
rate of 1.7–4.5% (% of KI pups/all pups).65 This was consideredoutstanding given that a spontaneous recombination rate is~0.1% in zygotes.94 In more recent studies, introducing mutations(including multiplexed editing), short functional sequences oreven a large reporter cassette were tested mainly by using CRISPR/
Cas9 system.59–61, 66, 85, 91, 92, 95–101 In contrast to the improved KO
rates in the one-step production scheme, zygotic KI by HDR still
remains inefﬁcient, particularly in the case of long fragment
insertion by homologous recombination (initially ~20%84). Several
studies have tried to improve the genome-editing (KI) rate. For
example, inhibition of the NHEJ pathway by administration of DNA
ligase IV inhibitor (Scr7) gives a 2 to 4-fold increase of HDR rate inmouse zygotes,97 although another study debated the capacity ofthis inhibitor in human models.102 Similarly, the treatment with anactin polymerization inhibitor (cytochalasin B or D) increases the
HDR targeting rate presumably due to the delayed DSB repair.99
The use of Cas9 protein rather than synthesized mRNA also
increases HDR rate.92, 96, 98 One of these studies showed anincrease of the genome-editing (KI) rate, up to ~45% KI efﬁciencyof live-born pups by injecting Cas9 protein complex with
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synthesized dual-crRNA:tracrRNA into pronuclei.98 The use of
Cas9 protein also reduces mosaicism when introduced withproper timing in early zygotes.101 Another study reported thegeneration of two KI newborns out of 123 injected embryos,
where one was heterozygous KI while another was homozygous
KI.92 ssODN-mediated KI, in which the cut sites of the targeted
genome and the inserted fragment are ligated with 80-bp ssODNs
homologous to the two cut ends (Fig. 2), was also shown tointegrate up to ~200 kb of bacterial artiﬁcial chromosome (BAC)
into the targeted locus.100 Finally, it was recently reported that
HDR accuracy is dramatically increased by incorporating silent
CRISPR/Cas-blocking mutations in sgRNA or Cas9-targeted
sequence (CORRECT method)103 although testing this in zygotes
still remains to be done. Please note however, that the
reproducibility of these recent studies should be further examinedbecause some of the conclusions are based on small number ofexperiments. Further improvements in the targeting rate andspeciﬁcity will be needed for one-step generation of KI mice.
ESC-derived mouse (ES mouse)
Although one-step generation of KI mice would be ideal for nextgeneration mammalian genetics, a number of issue have yet to be
overcome, speciﬁcally inefﬁcient editing and live-birth rate(particularly with a large fragment KI) and undesirable indel
insertion and mosaicism in resultant animals. Alternatively, twostep generation of KI mice by almost completely ESC-derived
mouse (“ES mouse”) is an attractive alternative at present. The
advantages of using ESCs are in its selectivity of sex, easier storageand ﬂexibility for more complex genome editing (e.g. multi-gene
KO and KI) in in vitro culture. Furthermore, genome editing in ESCs
is becoming easier by the site-speciﬁc nucleases.65–67, 84, 91, 104, 105
In recent studies, an HDR targeting rate over 10% was achieved in
ESCs even using very short (0.5 kb) homologous arms together
with CRISPR/Cas9.104 Multiple KO ESCs were also prepared in onestep manner, at the rate of approximately 20% in a triple KOexperiment.59 The other modern genome-editing methods
discussed above may potentially to be applicable as well.
Validity of ES mouse production and phenotyping analysis
within a single generation was ﬁrst proposed and tested by using
the tetraploid complementation method.106–113 However, several
possible drawbacks of the method are known. First, substantial
contamination of host cells was often observed in chimera mice
produced by this method, which can cause developmental
abnormalities.113–115 Second, the genetic heterozygosity of bothtetraploid embryos and ESCs seems to be crucial for survival of theresultant ES mice,110, 112, 114 which means that the use of inbred
ESCs does not seem possible and further backcrossing is required.
In addition, preparing hundreds of tetraploid embryos every time
does not seem practical for routine generation of many ES mice.
For these reasons, few reports have used tetraploid complementation in a large-scale phenotyping assay of ES mice.
Alternatively, ES mice can be generated by ESCs injected into or
aggregated with eight-cell embryos rather than the conventionally used blastocyst embryos.116–118 The eight-cell injection/
aggregation, in which totipotent host cells and ESCs as epiblast
precursors are used, produces almost completely ESC-derived
mice with ~0.1% contamination of host embryo cells. Furthermore, it is striking that ES mice from inbred strains, such as B6 andBALB/C were efﬁciently generated at the live-birth rates of
11~29% (ES mice/embryos transferred), which was comparable
to the 129 strain (9~15%) and hybrid ES mice (6~40%) andsigniﬁcantly higher than in blastocyst injection in any case (all0%).117 Therefore, F0 phenotyping of inbred ES mice generated
with 8-cell injection/aggregation was considered plausible in the
study. Possible drawbacks would be that the ES mouse production
rate might depend on the ‘quality’ of cultured ESCs. A further
Published in partnership with the Systems Biology InstituteNext-generation mammalian genetics
EA Susaki et al.

optimized method to stably maintain ESCs in their naive
pluripotent state was thus required.
Secretory regulatory factors and their downstream mechanisms
for maintaining ESC’s naive pluripotency have been well studied(recently reviewed in Huang et al.119 and others). Historically, theleukemia inhibitory factor–signal transducer and activator oftranscription 3 (LIF/Stat3) pathway has been found to beindispensable for maintaining pluripotency and self-renewal
ability of ESC.120–123 More recently, additional pathways essential
for ESC maintenance or differentiation were uncovered. One is theWnt–β-catenin pathway which supports ESC propagation andmaintenance of the pluripotent naive state, and is antagonized byglycogen synthase kinase-3 (Gsk3)–Tcf3 (also known as Tcf7l1).124–128
The other is ﬁbroblast growth factor 4–mitogen-activated protein
kinase (Mek)–mitogen-activated protein kinase (Mapk or Erk)pathway which leads ESC differentiation and thus its inhibition
suppresses ESC differentiation.129, 130 Therefore, naive pluripotent
ESCs can be stably maintained by shielding the cells from thesedifferentiation triggers, and by addition of inhibitors (3i: SU5402
for FGF receptor, PD184352 for Mek, CHIR99021 for Gsk3, or 2i:
PD0325901 for Mek and CHIR99021 for Gsk3) and in serum-free
conditions.131–133 3i/2i-cultured ESCs exhibit restricted expressionsof lineage-afﬁliated genes and stabilized gene expressions
involving a naive pluripotent state via epigenetic modulations
and appropriate control of the pluripotency factors.134–139
The administration of 3i/2i enabled more efﬁcient creation andmaintenance of ESCs from even inbred mouse strains (including
B6) or rat,133, 140–142 and increased germ-line transmission of B6derived ESC chimera.116 The stable establishment and maintenance of B6 ESCs in ES mouse production is particularly critical
for the next-generation mammalian genetics without crossing.
Previously, B6 ESCs were suggested to have problems of
maintenance, less efﬁcient chimera formation and germ-line
transmission, and genomic instability in standard culture conditions.143, 144 To overcome these problems, B6 ESCs were
established and maintained in serum-free 3i/2i + LIF medium,
which demonstrates signiﬁcantly higher success rate (67 vs. 3% inmedia containing serum)141 (Fig. 3b, c). In addition, ~100% ESCderived mice from the B6-3i ESCs can be stably generated witheight-cell injection, even after many passages and traditional
homologous-recombined targeting, at a production rate of30~100% (ES mice/live-born mice). Therefore, 3i/2i + LIF culture
and eight-cell injection/aggregation of ESCs enables the efﬁcient
one-step generation of ES mice, and subsequent F0 phenotyping
can be performed once the genome-edited ES mice are created.
Indeed, production and data acquisition of a novel ES mousestrain expressing a bright ﬂuorescent protein was completed
within a few months.145, 146 CRISPR/Cas9-mediated knock-in in 2i+ LIF-cultured ESCs followed by eight-cell injection have also been
performed.147 These examples support the potential of ES mouse
schemes. Note that ICR/CD-1 host embryos can be used for the ES
mouse production and there is no need for maintaining a speciﬁc
Tg colony for host embryos. Furthermore, experimental procedures (such as an operation for implantation) are similar to, oreven lesser than conventional chimera mouse production.
To further improve the efﬁciency of ES mouse production,better culture methods and ESC quality control will be needed.
Genomic instability, in particular, should be avoided during culture
because chromosomal aneuploidy can cause embryonic death.
Telomere extension seems important for maintaining normal
karyotype of ESCs, and frequent activation of telomere maintenance factor Zscan4 restores and maintains the ESC’s potency inlong-term culture.148, 149 Aneuploidy detection in cell culture
populations is also important for ESC’s quality control. This can be
performed by not only a conventional karyotyping but also by adroplet digital PCR-based screening.150 Furthermore, additional
chemical treatments can possibly ameliorate ESC culture conditions. So far, a variety of chemicals including ROCK inhibitor,151, 152
Published in partnership with the Systems Biology Institute

PKC inhibitor,153, 154 ERK/p38 inhibitor,155 HDAC inhibitors [e.g.,trichostatin A, sodium butylate or valproic acid156–158 or Vitamin
C159] may potentially contribute to improved potency of ESCs.
Therefore, applications of these chemicals and routine quality
control may help accelerate next-generation genetics based on ES
mouse production.
PRACTICE OF NEXT-GENERATION MAMMALIAN GENETICSAs discussed above, high-throughput KO or KI mouse production
is pivotal for accelerating system-level identiﬁcation, and analysis
of molecular networks and cellular circuits in organisms. Given
that various genetic tools, such as optogenetics and chemogenetic tools160, 161 are developing rapidly in recent years, highthroughput genome-edited mouse production is required for theirin vivo implementation. Next-generation mammalian genetics
potentially enables a single laboratory or a single researcher togenerate, maintain and analyze multiple genome-edited strains
rather than institutes or consortiums for production, deposit anddistribution of various strains and ESCs. Because sgRNAs fortargeted sites can be readily designed and prepared, the CRISPR/
Cas9 system, in particular, makes such large-scale genetics
feasible. Indeed, a recent study generated 31 novel CRISPR-KO
mice lacking testis-expressing genes.162 Since whole-body biallelic KO rates were not sufﬁciently high for the next-generation
scheme, the authors performed the crossing of selected F0
founders based on sequence and PCR screening data. In order torealize almost perfect (~100%) whole-body bi-allelic KO rate fornext-generation mammalian genetics, we recently performed
triple-CRISPR-based large-scale reverse genetics for sleep research.
To identify genes involving neural electrophysiological activities
during sleep or wake, we ﬁrst developed an average neuronmodel in silico and found that genes involved in intracellular Ca2+
regulation (Ca2+ channels, Ca2+-dependent channels, Ca2+-pumps
or Ca2+-dependent enzymes) are important for electrophysiological slow-wave-oscillation patterns during sleep. To further assess
the roles of these genes in vivo, we next produced KO mice for 33
genes with the triple-CRISPR methods and eventually identiﬁed 8
genes important for regulating sleep duration.62, 163
ES mouse technology can deal with more complex genome
editing, which would be difﬁcult, if not impossible, with theconventional crossing-based genetics. For example, we also
produced ~20 KO-rescue ES mice in order to perform systemlevel analysis of circadian clock-gene circuits in organisms.164 In
this experiment, a 3i + LIF-cultured ESC clone derived from adouble-KO mouse lacking two core clock genes (Cry1 and Cry2)
was established. A rescue expression cassette of the wild-type ormutated Cry1 gene was then homologously knocked-in in the ESC
clone (thus ﬁve alleles had been edited). Double-KO ES mice andKO-rescue ES mice were then generated and used for F0
phenotyping to measure in vivo 24 h rhythmicity. As explicitly
indicated by these examples, next-generation mammalian genetics enables large-scale organism-level experiments within reasonable time, space and labor.
Next-generation genetics is also important for improving animal
welfare and 3R principles, particularly contributing to “reduction”
of animal use. In our triple-CRISPR experiments,62 the yields of biallelic KO mice lacking tyrosinase gene (judged by the white coatcolor) were 17% on average, and 36% in the best case, of theinjected and transferred B6 zygotes. Therefore, between 60(average) and 30 (best case) of host embryos would be enoughfor generating a sufﬁcient number (around 10) of bi-allelic KO
mice. The rate of bi-allelic tyrosinase KO mice among the F0
littermates was 97.5% on average and 100% at best. Similarly, atleast in our ES-mouse experiments of Cry1 rescue in the Cry1/Cry2
DKO background,164 the yield of ES mice available for phenotyping was 5.5% on average, and 35% in the best case, of the injected8-cell embryos. Therefore, between 170 (average) and 30 (bestnpj Systems Biology and Applications (2017) 15

7Next-generation mammalian genetics
EA Susaki et al.

8case) of host embryos would be enough for generating a sufﬁcientnumber (around 10) of ES mice. The rate of ES mice among the F0
littermates was 43% as average and 91% at the best. Only thelittermates of embryonic lethal, non-KO or non-ES mice weresacriﬁced and no further animals are needed. The number ofanimals used is thus much smaller than the conventional
methods, in which a similar number of host embryos are used
for injection, and only a part of the founders or chimera mice areused for further crossing. In the conventional case, dozens oflittermates are produced and sacriﬁced during crossing to selectmice with an expected genotype. With conventional methods thenumber needed exponentially increases when a more complicated genetic background (e.g., double KO) is desired, while with
next-generation genetics the number of used animals is notdependent on genetic complexity.
On the other hand, researchers need to take special careregarding some issues with the use of F0 animals for phenotype
studies. In particular, researchers should carefully consider to what
extent potential mosaicism (e.g., mutational variations in the
triple-CRISPR method, or undetectable contamination of wild-type
cells in the ES mouse method) would affect the ﬁnal results of ascientiﬁc study. In our above experiments, the phenotypicvariations of F0 mice were comparable with those in wild-type
or suitable control animals,62, 163, 164 suggesting that mutationalvariations (triple-CRISPR) or undetectable contamination of wildtype cells (ES mouse) do not seem problematic at lease in thesecases. To further exclude the possibility of artifact phenotypes due
to mutational variations or undetectable contamination of wildtype cells, we recommend that researchers independently
generate a whole-body bi-allelic KO mice by using a second setof triple-CRISPR for the same gene, or to independently generate awhole-body bi-allelic KI mice using an independent clone of ES
cells. Such stringent criteria (the production of independent KO or
KI mice to conﬁrm the observed phenotype) is sometimes difﬁcult
to fulﬁll with conventional mouse genetics because it takes acouple of years for another round of production. On the other
hand, this step is feasible with next-generation genetics because it
only takes a couple of months. In this sense, the quality ofscientiﬁc studies using next-generation genetics should exceedthat of scientiﬁc studies using conventional genetics.
Another point that researchers need to consider carefully is
experimental design. This includes (1) preparation of appropriate
control animals without any obvious defects in the focused
phenotype (e.g., ES mice from wild-type ESCs or ESCs without
genome editing for a clock gene rescue study,164 and tyrosinase
triple-CRISPR KO mice for a sleep study62), (2) evaluation of genetic
composition of the F0 animals with strict criteria (e.g., detection of agenomic deletion as a proof of efﬁcient triple-CRISPR method62, 163
or a highly sensitive detection of contaminated host embryoderived cells164). In addition, it is also useful, if necessary, toevaluate undesirable mutations in the coding sequences by exon
sequencing163 of triple-CRISPR KO mice. It is also useful to avoid
cumulative mutations165 by using ESCs with minimal passage
numbers for the production of ES mice. Moreover, theseexperimental procedures should be described according to a
general guideline for animal experiments (e.g., the ARRIVE
guidelines166). Further efforts to minimize problems from theseissues should be expanded in future studies.
PERSPECTIVES
Next-generation mammalian genetics will facilitate system-level
analysis of molecular and cellular circuits in organisms. To further
improve the throughput for genome-edited animal production,
additional developments of new technologies related to nextgeneration mammalian genetics will be required. For example,
one-step production of whole-body bi-allelic KI mice by more
efﬁcient genome editing of zygotes still remains unachieved. For
npj Systems Biology and Applications (2017) 15

the two-step production of KI mice using ES mouse technology,
the preparation of host embryos and surrogate mothers are stilllabor intensive and pose limitations on its throughput. New
technologies which overcome these limitations will further
accelerate next-generation mammalian genetics and also reduce
the number of experimental animals used to obtain the same
information.
Next-generation mammalian genetics together with efﬁcient,quantitative and non-invasive phenotyping methods will provide
an attractive platform for investigating the organism-level
functions of the molecular and cellular circuits of interests.
Genome-wide phenome analysis has been performed in international KO mouse phenotyping efforts18, 167 to systematicallysurvey the functions of molecular networks in organisms.
Although most large-scale organism-level phenotyping projects
are usually labor-intensive, the development of more facilealternatives is possible. Recently, whole-body clearing andimaging methods with single-cell resolution have been developed146, 168, 169 and started to provide comprehensive andquantitative experimental data at the cell-to-organism level,
further facilitating organism-level systems biology. The development of non-invasive phenotyping will be also an attractive
direction. For example, sleep phenotyping represents such a
recent attempt where a non-invasive sleep phenotyping system
was used instead of a conventional invasive EEG/EMG-based
measurement system. In fact, these non-invasive methods havealready enabled sleep phenotyping of dozens of triple-CRISPR KO
mice.62, 163 Organism-level systems biology is thus coming tofruition with next-generation mammalian genetics, whole-bodyclearing and imaging with single-cell resolution, and non-invasive
and quantitative phenotyping methods. Organism-level systems
biology based on such new technologies will accelerate ourunderstanding of complex and dynamic molecular and cellular
circuits in the near future.
ACKNOWLEDGEMENTS
We thank all the lab members at RIKEN CDB, RIKEN QBiC and the University of Tokyo,
in particular, Dr. Kenta Sumiyama, Dr. Kazuki Tainaka, Dr. Naoshi Koide, Dr. Genshiro
Sunagawa, Ms. Junko Garçon and Dr. Kylius Wilkins for preparing ﬁgure materials, for
critical reading and for valuable comments on the manuscript; We also thank Dr.
Hiroshi Kiyonari (RIKEN CLST) for collaborations of ES mouse generation in Fig. 3. We
apologize to researchers whose work is not cited due to space limitations. This work
was supported by a grant from AMED-CREST (AMED/MEXT, H.R.U.), CREST (JST/MEXT,
H.R.U.), Brain/MINDS (AMED/MEXT, H.R.U.), Basic Science and Platform Technology
Program for Innovative Biological Medicine (AMED/MEXT, H.R.U.), a Grant-in-Aid for
Scientiﬁc Research (S) (JSPS KAKENHI Grant Number 25221004, H.R.U.), the strategic
programs for R&D (President’s discretionary fund) of RIKEN (H.R.U.), an intramural
Grant-in-Aid from the RIKEN QBiC (H.R.U.), Grant-in-Aids from Takeda Science
Foundation (H.R.U. and E.A.S.), a Grant-in-Aid for Young Scientists (A) (JSPS KAKENHI
Grant Number 15H05650, E.A.S.), a Grant-in-Aid by PRESTO from JST (JPMJPR15F4, E.
A.S.), a Grant-in-Aid from the Japan Foundation for Applied Enzymology (E.A.S.), and a
grant of the Brain Sciences Project of the Center for Novel Science Initiatives of the
National Institutes of Natural Sciences (BS281004, E.A.S.).

AUTHOR CONTRIBUTIONS
E.A.S., H.U., and H.R.U. discussed the contents and wrote the manuscript text.

COMPETING INTERESTS
The authors declare that they have no competing ﬁnancial interests.

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