TALEN/CRISPR-mediated engineering of a promoterless anti-viral RNAi hairpin into an endogenous miRNA locus
Nucleic Acids Research
TALEN/CRISPR-mediated engineering of a promoterless anti-viral RNAi hairpin into an endogenous miRNA locus
Elena Sen´ıs 1 3
Stefan Mockenhaupt 1 3
Daniel Rupp 0 7
Tobias Bauer 6
Nagarajan Paramasivam 5 6
Bettina Knapp 4
Jan Gronych 9
Stefanie Grosse 1 3
P. Windisch 0
Florian Schmidt 1 3
Fabian J. Theis 4 8
Roland Eils 1 2 6
Peter Lichter 9
Matthias Schlesner 6
Ralf Bartenschlager 0 7
Dirk Grimm 1 3
0 Department of Infectious Diseases , Molecular Virology , Heidelberg University Hospital , Heidelberg, 69120 , Germany
1 BioQuant Center, University of Heidelberg , Heidelberg, 69120 , Germany
2 Department for Bioinformatics and Functional Genomics, Institute for Pharmacy and Molecular Biotechnology (IPMB), Heidelberg University , Heidelberg, 69120 , Germany
3 Department of Infectious Diseases, Virology, Heidelberg University Hospital, Cluster of Excellence CellNetworks , Heidelberg, 69120 , Germany
4 Institute of Computational Biology, Helmholtz Zentrum M u ̈nchen , Neuherberg, 85764 , Germany
5 Medical Faculty Heidelberg, Heidelberg University , Heidelberg, 69120 , Germany
6 Division of Theoretical Bioinformatics (B080), German Cancer Research Center (DKFZ) , Heidelberg, 69120 , Germany
7 Division of Virus-Associated Carcinogenesis (F170), German Cancer Research Center (DKFZ) , Heidelberg, 69120 , Germany
8 Department of Mathematics, Technische Universit a ̈t M u ̈nchen , Garching, 85748 , Germany
9 Division of Molecular Genetics (B060), German Cancer Research Center (DKFZ) and German Cancer Consortium (DKTK) , Heidelberg, 69120 , Germany
*To whom correspondence should be addressed. Tel: +49 6221 5451339; Fax: +49 6221 5451481; Email: Present address: Marc P. Windisch, Hepatitis Research Laboratory, Institut Pasteur Korea, Seongnam-si, Gyeonggi-do, Rep. of Korea.
Successful RNAi applications depend on strategies
allowing robust and persistent expression of
minimal gene silencing triggers without perturbing
endogenous gene expression. Here, we propose a
novel avenue which is integration of a
promoterless shmiRNA, i.e. a shRNA embedded in a
microRNA (miRNA) scaffold, into an engineered genomic
miRNA locus. For proof-of-concept, we used TALE or
CRISPR/Cas9 nucleases to site-specifically integrate
an anti-hepatitis C virus (HCV) shmiRNA into the
liver-specific miR-122/hcr locus in hepatoma cells,
with the aim to obtain cellular clones that are
genetically protected against HCV infection. Using
reporter assays, Northern blotting and qRT-PCR, we
confirmed anti-HCV shmiRNA expression as well as
miR-122 integrity and functionality in selected
cellular progeny. Moreover, we employed a
comprehensive battery of PCR, cDNA/miRNA profiling and
whole genome sequencing analyses to validate
targeted integration of a single shmiRNA molecule at
the expected position, and to rule out deleterious
effects on the genomes or transcriptomes of the
engineered cells. Importantly, a subgenomic HCV
replicon and a full-length reporter virus, but not a Dengue
virus control, were significantly impaired in the
modified cells. Our original combination of DNA
engineering and RNAi expression technologies benefits
numerous applications, from miRNA, genome and
transgenesis research, to human gene therapy.
Originally discovered in plants and worms, RNA
interference (RNAi) has become a powerful and versatile
technology for gene regulation in biology and medicine (1,2). A
major part of its popularity is due to the ease with which it is
triggered in mammalian cells, as it merely requires the
delivery of double-stranded RNA mimics of endogenous
microRNAs (miRNAs), such as short hairpin RNAs, which
engage the intrinsic cellular RNAi machinery for
processing and target mRNA inhibition. Particularly appealing is
that artificial RNAi triggers can be expressed from
various promoters and encoded by non-viral or viral gene
delivery vectors, yielding a comprehensive toolbox for
applications ranging from fundamental gene annotation in
cultured cells, to therapeutic (m)RNA suppression in humans.
Very attractive clinical targets are viral pathogens such as
the hepatitis B or C viruses (HBV or HCV, respectively)
since they go through an obligatory RNA phase that is
vulnerable to RNAi (3,4), and since many established
treatments have limited efficacy and/or adverse side effects.
Indeed, we and others have previously exemplified the
tremendous potential of RNAi for robust and stable in vivo
suppression of hepatitis viruses, including in HBV-transgenic
mice that were infused with liver-specific Adeno-associated
viral vectors of type 8 (AAV8) encoding anti-HBV shRNAs
Still, RNAi applications and their clinical translation
remain severely hampered by concerns over specificity, safety
and longevity (Figure 1A). Particularly alarming are
findings in different animal species, from mice to primates, that
shRNA over-expression from strong RNA polymerase III
promoters can cause cytotoxicity, organ failure and
lethality (8,10–13). Possible reasons include adverse off-targeting
and dose-dependent saturation of the cellular RNAi
machinery which is needed for processing and functionality
of miRNAs and other endogenous RNAi triggers. This has
tempted us and others to improve RNAi expression
strategies, for instance by pre-selecting inherently safe and potent
shRNAs (8,14), or by placing shRNAs under weak and/or
tissue-specific promoters (7). Another strategy to diminish
in vivo RNAi toxicity is to embed an ectopic RNAi sequence
within a cellular miRNA, by replacing one of the two arms
of the double-stranded miRNA with the shRNA antisense
arm, resulting in a so-called shmiRNA (15–19).
While all these avenues can mitigate adverse RNAi effects
to some extent, a major remaining challenge is to combine
these advances with equally improved strategies for safe and
specific ex or in vivo RNAi delivery. A promising option are
vectors derived from AAV as they are non-pathogenic,
easyto-engineer and retargetable to tissues and cells of choice
(20–24). However, because their genomes remain
predominantly episomal and are rapidly lost in dividing cells (25)
(Figure 1A), the usefulness of AAV/RNAi vectors is highest
in quiescent tissues, such as the liver, or in applications
requiring short-term RNAi expression. Conversely,
integrating vectors derived from retro- or lentiviruses permit
longterm RNAi expression, but due to their promiscuity bear
a risk of insertional mutagenesis and progression to clonal
expansion or oncogenesis (26–29). Mechanisms include
disruption of tumor suppressor genes, or activation of
neighboring oncogenes by vector-borne promoter/enhancer
elements. The latter was observed in retro- or lentiviral gene
therapy trials, and it was also noted in mice that developed
hepatocellular carcinoma due to AAV vector integration
into a certain locus (Rian) and activation of proximal small
RNAs and genes (30–32). Additional concerns about
imprecise vector integration include insufficient control over
the number of insertion events and over local effects from
the integration site, and consequently limited options to
govern RNAi expression levels. Finally, ectopic promoters
can vary in their strength between different cell types or
become silenced over time, further complicating the
finetuning of sh(mi)RNA expression levels and the
establishment of lifelong RNAi phenotypes, even in non-dividing
Here, we developed and validated a novel RNAi
expression strategy that alleviates these vector-associated
concerns and uncertainties, and that concurrently exploits the
benefit of higher safety of miRNA-embedded shmiRNAs.
To this end, we devised the original concept to stably
integrate a minimal promoterless shmiRNA hairpin into an
endogenous miRNA locus, in order to hijack the
cellular miRNA promoter for long-term shmiRNA expression
at robust yet non-toxic levels (Figure 1B). For
proof-ofprinciple and convenient read-out, we harnessed TALE
and CRISPR/Cas9 nucleases to insert a shmiRNA against
HCV, a human pathogen that is susceptible to RNAi
(36,37), into the genomic miR-122 locus (hcr) .The latter
is abundantly and specifically expressed in liver cells, where
HCV infection occurs (38). Using a comprehensive battery
of assays including whole genome sequencing, we show that
it is indeed possible to obtain engineered cellular clones that
still largely resemble the parental cells, but that are stably
protected against HCV replication and infection. As the
identical strategy can likely be customized for many further
miRNA loci and for any RNAi target, we expect the new
concept introduced here to hold significant potential for a
wide scope of basic or therapeutic RNAi applications.
MATERIALS AND METHODS
Details on plasmids, vectors, cell culture, transfections,
PCRs, Northern blotting, luciferase assays, FISH and
statistical analysis are found in Supplementary Methods.
Generation of stable shmiRNA cell lines
Huh7 cells were transfected as described in Supplementary
Methods with TALEN or CRISPR/Cas9 expression
plasmids together with the homologous recombination
template pSSV9-hcr-donor-shmiRHCV318. Fourty-eight to 72
h post-transfection, cells were trypsinized and transferred
to 15 cm dishes. They were then cultured for at least 15 days
in Huh7 media supplemented with 500 g/ml G418, before
single colonies were picked and transferred to 96-well plates.
Genomic DNA was extracted from these colonies using
DirectPCR Lysis Reagent Cell (PeqLab, Erlangen, Germany)
and analyzed by polymerase chain reaction (PCR) to
assess the occurrence of proper homologous recombination.
Positive colonies were expanded and eventually re-seeded in
24-well plates. Two wells of each cell line were transduced
with an AAV vector expressing Cre recombinase, while a
third well was treated with an AAV vector expressing YFP
as a transduction control (see next chapter for details). Two
days after transduction, genomic DNA was extracted from
the cells of one of the Cre-treated wells and analyzed by
PCR, to monitor the formation of minicircle DNA derived
from Cre-mediated excision of the floxed NeoR cassette. The
cells from the other well were trypsinized and transferred
to 15 cm dishes to allow formation of single colonies for
subcloning. The cells were then cultured for approximately
seven days in Huh7 media without G418 (and diluted again,
in case the initial cell density turned out to be too high for
colony formation from a single cell). Afterward, colonies
were again picked and analyzed by PCR to validate that the
floxed NeoR cassette had been excised properly. Finally, the
PCR product was sub-cloned and sequenced, and positive
clones were characterized further.
To generate the stable HEK293 cells used in
Supplementary Figure S9, no colonies were picked after treatment with
the Cre-expressing AAV vector. Instead, based on the 100%
efficiency of the AAV/Cre vector, the entire cell pool was
used for further analysis.
To eliminate the floxed NeoR cassette from the
shmiRNAengineered clones, they were transduced with AAV vectors
expressing Cre recombinase. To this end, the different Huh7
or HEK293 clones were seeded at 5 × 104 cells per well in
24-well plates and 24 h later transduced with AAV vectors,
expressing either Cre recombinase or a YFP control, at a
multiplicity of infection (MOI) of 4 × 104 viral genomes per
cell. Cells were re-seeded for colony picking (Huh7) or for
luciferase assays (HEK293), or harvested for PCR analysis
48 h post-transduction.
Whole genome sequencing analysis
Whole genome sequencing was performed on Huh7 T2 31.3
and Huh7 wildtype (WT) cells to validate the specificity
of shmiRHCV318 integration and to identify possible
offtarget integration sites. After library preparation,
sequencing was performed using the Illumina HiSeq platform.
Sequencing reads were aligned against the human
reference genome hs37d5 (http://www.1000genomes.org/home)
and the shmiRHCV318 as separate contig using the
BWAMEM (v0.7.8) software (39). Resulting BAM files were
coordinate-sorted, merged and PCR duplicates marked by
biobambam (v0.0.148) software (40). The resulting BAM
files have an average coverage of 19.82x and 21.69x for
WT and T2 31.3 cells, respectively (0x and 12.69x on the
shmiRHCV318 contig). The integration site at chr18 and
the shmiRHCV318 contig were visualized by using IGV
Gene and miRNA expression profiling
Prior to using total RNA samples for gene and miRNA
expression profiling, their quality was assessed via two
methods. First, RNA concentration as well as A260/280 and
A260/230 ratios were measured using a Nanovue Plus
Spectrophotometer (GE Healthcare, Pasching, Austria).
Second, the RNA integrity number (RIN) was determined
using the Agilent 2100 Bioanalyzer and the Agilent RNA 6000
Nano Kit (both Agilent Technologies, Waldbronn,
Germany) following the manufacturer’s instructions. Samples
with a high RNA quality (RIN near or equal to 10) were
then subjected to gene and miRNA expression profiling at
the German Cancer Research Center (DKFZ, Heidelberg,
Germany), Genomics and Proteomics Core Facility. Three
biological replicates (different cell passages) were provided
for each cell clone.
Gene expression profiling was performed using
Affymetrix Human Genome U133 Plus 2.0 arrays.
Raw expression values were quantile-normalized and
log2-transformed. A principal component analysis showed
a cumulative R2 of 0.99 within the first two principal
components. For hypothesis testing, log2 fold-changes
of each sample (T2 31.3, TS 30.20 and U6 20.16) as
compared to the WT samples and pairwise between the
samples (after correction for the WT) were computed,
and a fixed-effects linear model was fit for each individual
gene to estimate expression differences. An empirical
Bayes approach was used to moderate the standard errors
of the normalized log2 fold-changes. Finally, we used a
two-sided moderated t-test to compute P-values controlled
by Benjamini–Hochberg for multiple testing. Genes with
an absolute log2 fold-change larger than 1 and a corrected
P-value < alpha = 0.05 were assigned to be differentially
expressed with high significance.
MiRNA expression profiling was performed using the
Agilent-046064 Unrestricted Human miRNA V19.0 Mic
roarray. Raw miRNA microarray data were pre-processed
with background subtraction using the ‘normexp’
convolution standard method with subsequent quantile
normalization as implemented in the Bioconductor limma
package (v3.22.1) (42) for R statistical software (v3.1.2).
Expression values were log2-transformed, and probe duplicates
were averaged and quantile normalized to ensure
comparability between samples. Potential batch effects between
Agilent chips conducted at different dates were reduced
by the ComBat-function in the Bioconductor sva-package
(v3.12.0) on the basis of an empirical Bayesian framework
We calculated differential gene expression on
quantilenormalized Affymetrix mRNA expression profiles with the
R statistical software (44) (v3.1.0). Data were filtered for
low expression and variance with standard parameters,
retaining 12 680 (of 54 675) probe IDs. Significance
Analysis of Microarrays ((45), R-package ‘siggenes’ v1.40.0 (46))
was performed and differentially expressed probe IDs were
called at a false discovery rate threshold of <0.05. The same
procedure was applied to the Agilent microarray data to
identify differentially expressed miRNAs.
HCV inhibition experiments
In vitro transcription of plasmids
pFKi341PiLucNS33 -JFH1-dg, pFKi341PiLucNS3-3 -JFH1-dg-dGDD,
pFKi389-JCR2a-dGDDdg-JC1 (all 10 g, MluI-linearized) was used to generate
subgenomic or full-length HCV RNA, respectively.
Likewise, in vitro transcription of 10 g of XbaI-linearized
plasmid pFK-sgDVs-R2A (47) resulted in subgenomic
Dengue virus replicon RNA. DNA was purified by using
the NucleoSpin Gel and PCR Clean up Kit (Macherey
Nagel, D u¨ren, Germany) according to the manufacturer’s
instructions. In vitro transcription reaction mixtures (total
volume 100 l) for use with T7 polymerase contained 80
mM HEPES (pH 7.5), 12 mM MgCl2, 2 mM spermidine,
40 mM dithiothreitol (DTT), 3.125 mM of each nucleoside
triphosphate, 1 U/ l RNasin (Promega), 0.1 g/ l of
plasmid DNA and 0.6 U/ l of T7 RNA polymerase (Promega).
For HCV transcripts and use with SP6 polymerase, the mix
contained 80 mM HEPES (pH 7.5), 16 mM MgCl2, 2 mM
spermidine, 40 mM DTT, 3.125 mM each of rATP, rCTP
and rUTP, 1.5625 mM of rGTP, 1 mM m7G(5 )ppp(5 )G
RNA cap structure analog, 1 U/ l of RNasin, 0.1 g/ l
of plasmid DNA and 0.6 U/ l of SP6 RNA polymerase
(Promega). After a 2 h incubation at 37◦C (HCV) or 40◦C
(Dengue virus), 0.3 U of T7 RNA polymerase or 0.4 U of
SP6 RNA polymerase, respectively, were added per l of
reaction mixture, and the reaction mixture was incubated
overnight. Transcription was terminated by adding 1.2 U
of RNase-free DNase (Promega) per g plasmid DNA and
30 min incubation at 37◦C. RNA was extracted with acidic
phenol and chloroform, precipitated with isopropanol at
room temperature and dissolved in RNase-free water.
For electroporations, single-cell suspensions of WT or
engineered Huh7 clones were prepared by trypsinization,
washing with 1x phosphate buffered saline (PBS) and
resuspension at a concentration of 1 × 107 cells per ml in
Cytomix (48) supplemented with 2 mM adenosine
triphosphate (ATP) and 5 mM glutathione. Next, 2.5 g in vitro
transcripts were mixed with 100 l cell suspension and
transfected by electroporation using a GenePulser system
(Bio-Rad, Hercules, CA, USA) and a 0.2 cm gap cuvette
(Bio-Rad) at 500 F and 166 V. Cells were diluted
immediately in complete Dulbecco’s modified Eagle’s medium and
JCR2a virus was produced in Huh7.5 cells by
electroporation of HCV RNA. Supernatants were harvested every 24
h between 48 and 96 h after electroporation. Infectious titer
(median tissue culture infective dose, TCID50) was
determined by limited dilution. For infection, Huh7 cells were
seeded at 5 × 104 cells per well in a 24-well plate 1 day prior
to infection with a MOI of 0.5.
For luciferase assays, cells were lysed in luciferase lysis
buffer (1% [v/v] Triton X-100, 10% [v/v] glycerol, 25 mM
glycyl-glycine pH 7.8, 15 mM l MgSO4, 4 mM EGTA,
kept at 4◦C and freshly supplied with 1 mM DTT
immediately before use). For luciferase measurements, cells were
washed once with PBS, lysed directly in the 96-well plate
with 30 l lysis buffer per well and frozen at −80◦C. Shortly
before measurement, lysates were allowed to thaw at room
temperature for 30 to 60 min. Firefly luciferase activity was
measured for 3 s in a Mithras LB940 multimode microplate
reader (Berthold Technologies, Bad Wildbad, Germany).
Renilla luciferase activity was measured for 10 s. Firefly
assay buffer (25 mM glycyl-glycine pH 7.8, 15 mM K2PO4
pH 7.8, 15 mM MgSO4, 4 mM EGTA, 1 mM DTT, 2 mM
ATP) supplemented with 70 M D-luciferin (PJK GmbH,
Kleinblittersdorf, Germany), or Renilla assay buffer (25
mM glycyl-glycine pH 7.8, 15 mM K2PO4 pH 7.8, 15 mM
MgSO4, 4 mM EGTA, 1 mM DTT) supplemented with 1.5
M Coelenterazine (PJK GmbH), was added to each well
automatically prior to each measurement.
All microarray data are available in the NCBI Gene
Expression Omnibus database (GEO; http://www.ncbi.nlm.nih.
gov/geo/) under accession number GSE68638. The whole
genome sequencing data have been deposited in the
European Genome-phenome Archive (EGA, https://www.ebi.ac.
uk/ega/home) under accession number EGAS00001001252.
Design and validation of an anti-HCV shmiRNA for
integration into the hcr locus
To create an effective anti-HCV shmiRNA, we harnessed
the antisense sequence of the published HCV-specific
shRNA HCV-321 (36) that binds a highly conserved
target in the HCV 5 NTR (non-translated region) starting at
nucleotide 321. For adaptation to the precursor of human
miR-122 (used as scaffold for our shmiRNA design), we
added three nucleotides to the 3 end of the shRNA, yielding
an extended 22 nucleotides long antisense sequence starting
at HCV position 318 (accordingly named shmiRHCV318,
Figure 2A). Importantly, we have recently validated
antiHCV efficacy of an analogous shRNA (shHCV318),
proving that this part of the HCV genome is well accessible (49).
While considering a suitable integration site within the
miR-122 locus (hcr), we realized that the precursor
(pre)miR-122 hairpin is located close to the putative hcr
polyadenylation signal (Supplementary Figure S1A). It
thus appeared reasonable to integrate the shmiRNA
upstream of the miR-122 hairpin, to ensure its proper
inclusion into the full transcript. We moreover argued that
targeting a site inherently lacking a local RNA structure would
increase the likelihood of preserving folding and processing
of the miR-122 stem-loop. We therefore computationally
predicted the RNA secondary structure of the hcr 3 end
including the precursor (pre-)miR-122 hairpin, and then
selected one exposed site 63 nt upstream of pre-miR-122
(or 76 nt upstream of the miR-122 5p arm, respectively)
for shmiRNA integration (Supplementary Figures S1B and
To verify that this position permits miR-122 and
shmiRNA co-expression, we generated plasmids expressing
1.2 kb of the hcr gene (hg38: chr18: 58450012–58451248)
comprising miR-122 alone or co-encoding shmiRHCV318
(manually cloned into position hg38: chr18: 58451012).
Two further constructs contained a shmiRNA against a
different target (human alpha-1-antitrypsin, hAAT), either
in place of shmiRHCV318 or in combination, the
latter resulting in a triple hairpin (Figure 2B). These
constructs served as control for the anti-HCV shmiRNA, and
to assess whether our strategy would enable
multiplexing of three hairpins. Co-transfections of HEK293T cells
(human embryonic kidney cells lacking miR-122
expression) with these expression plasmids and luciferase
reporters carrying perfect binding sites for the different
hairpins confirmed that each shmiRNA potently and
specifically inhibited its cognate target (Figure 2C). Notably,
this was observed for both configurations, double (either
shmiRHCV318 or shmiRhAAT plus miR-122) and triple
hairpins (shmiRHCV318, shmiRhAAT and miR-122).
Activity of the co-encoded miR-122 hairpin was comparably
high in all four plasmids, and only marginally and
nonsignificantly reduced in the triple versus the single miR-122
expression plasmid (Figure 2C, blue bars; best visible in the
right graph). Collectively, this demonstrates the feasibility
to co-express at least two exogenous shmiRNAs together
with miR-122 without significantly compromising its
Efficient and specific anti-HCV shmiRNA integration into
the miR-122 locus via homologous recombination
To site-specifically and efficiently integrate shmiRHCV318
into the hcr locus of liver cells, we devised a
homologous recombination strategy based on potent TALEN and
CRISPR constructs against human miR-122 from our lab
(50) (Supplementary Figure S3 and Figure 3A). We chose
the human hepatocarcinoma cell line Huh7 as target since
it is susceptible to HCV infection and abundantly expresses
miR-122 (38). Notably, despite being a tumor-derived cell
line, Huh7 carries only two copies of the long arm of
chromosome 18 where the hcr/miR-122 locus resides, as
demonstrated by multiplex fluorescence-in-situ-hybridization
(MFISH) (Supplementary Figure S4). The recombination
template contained the shmiRHCV318 hairpin plus a floxed
neomycin resistance (NeoR) cassette for selection, flanked
on each side by 1.25 kb of DNA homologous to the
hcr locus (the 3 homology arm comprised the miR-122
hairpin). Following nuclease and template co-transfection
and G418 selection, single colonies were picked and
analyzed by PCR, to detect and distinguish random
versus directed shmiRHCV318 integration. Results from the
TALEN-based approaches are shown in Figure 3B (for
primer sequences see Supplementary Table S1), illustrating
that roughly 90% of all clones that were negative for random
integration showed proper insertion of the exogenous
hairpin into the hcr locus. If we factor in that we pre-selected
for, and excluded, negative clones (random integration), we
still calculate around 65% positive clones (correct insertion)
for the best TALEN pair T2 (Figure 3C).
Curiously, in our first experiments, the efficiencies of
homologous integration were substantially lower with the TS
TALEN pair as well as the CRISPR strategy, reaching only
up to 7.1% or 3.3%, respectively (Figure 3B and C, and data
not shown). We then realized that both, the TS TALEN
pair and the anti-miR-122 gRNA, bind and inadvertently
cleave a sequence that partially overlaps with the miR-122
hairpin and is thus fully conserved in the
shmiRHCV318containing repair template (see also Supplementary
Figure S3), providing a likely explanation for their inferior
performance. In line with this, the much more effective
TALEN pairs T1 and T2 exclusively recognize and cleave
the endogenous hcr locus, but not the repair DNA. For
the gRNA-based strategy, a possible solution was to
mutate the protospacer-adjacent motif (PAM, a critical three
nucleotide sequence directly downstream of the actual
target sequence) in the repair template. This should abolish its
adverse Cas9-mediated cutting before, during or after
recombination, and instead restrict Cas9 cleavage to the
endogenous locus. Accordingly, we generated an improved
repair template in which we point-mutated the gRNA PAM
from the canonical TGG to a TGC that is not recognized by
Streptococcus pyogenes Cas9 (Supplementary Figure S5A).
Importantly, because the point mutation occurred in a small
bulge of the predicted pre-miR-122 structure, it did not
affect miR-122 functionality (Supplementary Figure S5B). In
addition, we replaced the Cas9 cDNA in our original
construct (50) with a more potently expressed variant (see
Supplementary Methods). These two modifications boosted the
efficiency by 2-fold (new repair template) or 5-fold (new
hcr + HCV318
hcr + hAAT
hcr + HCV318 + hAAT
Cas9 cDNA), respectively, or by roughly 10-fold altogether
(Supplementary Figure S5C). Notably, the resulting 33%
clones that were positive for homologous integration using
the improved CRISPR strategy now approached the results
obtained with our second-best TALEN pair T1 (43.8%,
Figure 3B and C).
To remove the floxed NeoR cassette from positive clones,
selected candidates were treated with an AAV vector
expressing Cre recombinase under the strong CAG (chicken
beta-actin) promoter (Figure 3D). Successful excision was
then confirmed by PCR-based detection of resulting 0.7 kb
minicircles (Figure 3E top). From positive pools of cells,
individual cellular clones were expanded further and
reanalyzed using PCR primers that spanned the entire
integration site (red arrows in Figure 3D). As exemplified
for clone T2 31 in Figure 3E (bottom), this procedure was
highly efficient, resulting in elimination of the NeoR cassette
in all of the tested AAV/Cre-treated progeny clones (10 out
of 10, 100%). As expected, this restored G418 sensitivity of
these cells, further evidencing complete removal of the
selection marker (Supplementary Figure S6).
Random integration (RI) PCR
Homologous integration (HI) PCR
-Minicircle (0.7 kb)
-NeoR+ (4.3 kb)
-NeoR- (2.7 kb)
-WT (2.5 kb)
Sequencing analysis of integration events and
nucleaseinduced modifications in the miR-122 locus
To verify proper integration of the shmiRHCV318 hairpin,
PCR products spanning the targeted region were
amplified from genomic DNA from some of the engineered
cellular clones and then sequenced. Surprisingly, we observed
a broad spectrum of nuclease-induced modifications within
the edited miR-122 hairpin, as illustrated for seven clones in
Figure 4 (complete sequences are shown in Supplementary
Figure S7). Importantly, successful integration of the
exogenous shmiRHCV318 hairpin into at least one miR-122
allele was detected in all clones. The only exception was the
Cas9/gRNA-derived clone U6 6.21 which carried a
deletion of 4 nucleotides in the WT allele, and a 195 nucleotide
deletion in the edited allele that spanned the shmiRHCV318
hairpin (but not the NeoR cassette, explaining why this
clone was G418-resistant).
Amongst the six other clones, two (T2 4.37 and U6 20.16)
were homozygous for shmiRHCV318 integration, while the
remaining four were heterozygous. This was concluded from
detection of either one (homozygous) or two (heterozygous)
PCR products after AAV/Cre treatment (see Figure 3E
above for an example for a heterozygous clone), and from
sequencing of at least five independent bacterial colonies
after subcloning of each product. The miR-122 hairpin had
remained fully intact in three of these six clones, including
one of the clones with a homozygous shmiRHCV318
integration (T2 4.37). The other homozygous clone, U6 20.16,
carried an identical 66 nucleotide insertion right upstream
of the PAM sequence in both alleles. Two of the four
heterozygous clones (T2 31.3 and U6 6.30) had intact miR-122
alleles, while the other two (TS 30.20 and U6 7.16) showed
modifications ranging from a 1 nucleotide insertion, to a 13
nucleotide deletion. Possible explanations for this spectrum
of on-target mutations are provided in Supplementary
Discussion part A.
Originally, we had intended to derive and characterize a
clone with minimal modification of the hcr locus, i.e.
heterozygous shmiRHCV318 integration and preserved
miR122 sequence, as is indeed the case with clones T2 31.3
and U6 6.30. However, as miR-122 is a host factor for
HCV (38), we reasoned that partial miR-122 disruption
might in fact inhibit viral replication synergistically, and
thus decided to also study clones with miR-122
modifications. Overall, we had to restrict our selection to three
different clones in order to maintain feasibility of the
comprehensive downstream analysis, including HCV infection
experiments, miRNA profiling and whole genome sequencing
(see below). We therefore selected the following three
representative clones for all further studies: (i) T2 31.3
(heterozygous for shmiRHCV318, both miR-122 alleles intact), (ii)
TS 30.20 (heterozygous for shmiRHCV318, minor
modifications in both miR-122 alleles) and (iii) U6 20.16
(homozygous for shmiRHCV318, also homozygous for a 66
nucleotide insertion upstream of miR-122).
Characterization of anti-HCV shmiRNA levels and of the
different miR-122 phenotypes
Steady-state levels and functionality of the two hairpins
(shmiRHCV318 and miR-122) were first assessed by
quantitative real-time PCR (qRT-PCR), Northern blotting and
luciferase reporter assays (Figure 5). All three clones
expressed shmiRHCV318 at different levels as detected by
qRT-PCR that correlated with the degree of miR-122
mutation (compare Figure 4 and Supplementary Figure S7),
from about 200 copies per cell for the fully intact clone T2
31.3, to about 25 for the largely miR-122-mutated clone
U6 20.16 (Figure 5A). These numbers were mirrored by
different relative levels of mature shmiRHCV318 as
measured by Northern blotting (Figure 5B top), and they
moreover corresponded well to the extent of inhibition
of a shmiRHCV318-sensitive luciferase reporter, reaching
around 80% for the best clone T2 31.3 (Figure 5C). A
similar pattern was noted for endogenous miR-122, which was
most prominently detected in clone T2 31.3 but almost
absent in clone U6 20.16 (Figure 5B bottom). Mature miR-122
steady-state levels were reduced by 2.1- or 4.8-fold (by
qRTPCR) in clones T2 31.3 and TS 30.20, respectively (Figure
5D), congruent with 2-fold lower levels of precursor
miR122 in clone T2 31.3 (Supplementary Figure S8). See
Supplementary Discussion part B for possible reasons for the
mild reduction in clone T2 31.3 despite the absence of
mutations in both miR-122 alleles. Notably, this barely affected
knockdown of a miR-122-sensitive luciferase reporter
(Figure 5E). In contrast, clone U6 20.16 which showed 188-fold
lower miR-122 expression in qRT-PCR (Figure 5D) and no
detectable expression by Northern blotting (Figure 5B
bottom) also exhibited no miR-122 reporter inhibition (Figure
5E), characterizing this clone as a complete miR-122
As a control experiment to verify the liver lineage
specificity of our strategy, we also integrated the shmiRHCV318
hairpin into human HEK293 kidney cells in which the
miR122 locus is inherently silent. With up to 81%
(Supplementary Figure S9A and B), the efficiencies of
homologous integration even surpassed those observed in Huh7
cells (Figure 3C), perhaps related to the higher
transfectability of HEK293 cells. Interestingly, we noted a
moderate read-through effect from the Rous Sarcoma Virus
(RSV) promoter driving the NeoR cassette that resulted
in shmiRHCV318 and miR-122 expression, as detected
by luciferase reporter assays (Supplementary Figure S9C
and D). Most importantly, however, this effect was
eliminated once the NeoR cassette and hence the ectopic
promoter were floxed out (compare dark and light blue bars
in Supplementary Figure S9D). Accordingly, the fully
engineered HEK293 cells no longer exhibited suppression of the
shmiRHCV318 or miR-122 luciferase reporters, identical
to the parental HEK293 cells and in contrast to the Huh7
clone T2 31.3 that was included as positive control. We thus
conclude that expression of integrated shmiRHCV318 is
strictly dependent on the activity of the endogenous
miR122 promoter (silent in HEK293, but active in Huh7 cells),
and that, vice versa, integration of the exogenous hairpin
does not trans-activate the silent hcr locus in non-hepatic
cells, both adding to the stringency of our approach.
heterozygous editing, large mutations
Comprehensive gene and miRNA expression analysis to verify
the similarity of lead clone T2 31.3 to WT Huh7 cells
To further assess the similarity of the three selected cellular
clones and WT Huh7 cells, gene expression profiling of all
cells was performed at three different passages. Analyses of
significantly differentially expressed genes (DEG) revealed
a minor number of DEG in clones T2 31.3 (24 DEG) and
TS 30.20 (16 DEG) (all genes are listed in the
Supplementary Expression profiles). Conversely, a much larger
number of genes were affected in clone U6 20.16 (1797 DEG).
This is evident from Supplementary Figure S10 that
visualizes the quantiles of the observed versus the expected
statistics. The higher similarity of the two TALEN-derived clones
to WT Huh7 cells, especially of clone T2 31.3, is moreover
illustrated by principal component analysis (PCA) which
showed that clones T2 31.3 and TS 30.20 cluster together
and resemble WT Huh7 cells, but are distinct from clone
U6 20.16 (Figure 6A).
Congruent with this, global miRNA expression
profiling confirmed a notable similarity between clone T2 31.3
TALEN SEED BS
homozygous editing, no mutations
homozygous editing, large mutations
heterozygous editing, no mutations
heterozygous editing, small mutations
Wild-type allele (WT)
Integration allele (I)
tRNAQ1 (loading control)
1.00 0.58 0.28 0.00
and WT Huh7, as opposed to clone U6 20.16 in which
numerous miRNAs were expressed aberrantly (Figure 6B; see
Supplementary Discussion part C for possible reasons). The
largest discrepancies in miRNA expression between this
clone and the parental cells were noted for the two strands
of miR-122 (Figure 6C, arrows), concordant with the
qRTPCR, Northern blotting, sequencing and functional data
(see above). In clone U6 20.16, miR-122 is strongly
downregulated (fold-change FC = 0.06), while the least difference
was found in T2 31.3 (FC = 0.48); clone TS 30.20 was in
between (FC = 0.18).
Altogether, these data consistently indicate that
shmiRHCV318 expression per se (even the high levels
in clone T2 31.3), and also the additional mutations of
miR-122 in clone TS 30.20, only mildly affected global
gene and miRNA expression. This is in contrast to the
largely miR-122-mutated clone U6 20.16 in which we
noted 100-fold more dysregulated genes (Supplementary
Expression profiles) and poorer clustering (Figure 6) with
the parental cells.
Next, we performed whole genome sequencing of T2
31.3, which had emerged as our lead clone at this point
based on its superior similarity to WT Huh7 cells. We
mapped the whole genome sequencing reads of T2 31.3
and WT Huh7 cells to a concatenated reference genome
consisting of GRCh37 version hs37d5 and the shmiRNA
contig. This identified 37 read pairs which mapped to
shmiRHCV318 and the expected integration locus on
chromosome 18 (Supplementary Figure S11). In addition, all
reads overlapping the shmiRNA contig borders support the
intended integration site. Importantly, we detected no read
pair or read overhang mapping to a different genomic
position or being unmappable, which would have indicated
offtarget integration in the Huh7 genome.
Furthermore, we identified single nucleotide variants and
short indels via Platypus (51), and structural variants via
Manta (52), again using GRCh37 version hs37d5 as
reference genome. Only the variants that passed all internal
filters within Platypus and Manta were then used in
comparisons of WT Huh7 cells and our lead clone T2 31.3.
Gene expression profiling
miRNA expression profiling
plementary Table S2 shows that in all categories, parental
Huh7 cells had even more private genetic variants than
clone T2 31.3. This suggests that the private variants in
T2 31.3 accumulated during Huh7 clonal derivation rather
than representing specific nuclease-induced variations.
As a whole, these complementary analyses suggested the
presence of a single shmiRHCV318 integration within an
intact miR-122 locus at the expected position in clone T2
31.3. This independently verifies our sequencing data in
Figure 4 and Supplementary Figure S7, and illustrates the
possibility to juxtapose high precision with low genotoxicity
through our nuclease-mediated miRNA engineering
Validation of efficient and specific impairment of HCV
replication in engineered Huh7 cells
Finally, we challenged all three cell clones with HCV to
study whether their genetic engineering had induced the
expected resistance against HCV. Fourty-eight and 72 h after
electroporation of a subgenomic HCV replicon encoding a
luciferase reporter (surrogate marker for HCV replication
(53)), we noted about 10- (T2 31.3) to 100-fold (TS 30.20)
reduced viral replication in all clones as compared to WT
Huh7 (Figure 7A and Supplementary Figure S12A). HCV
inhibition was likewise pronounced when we electroporated
a full-length reporter virus genome encoding luciferase and
able to form infectious HCV particles (54) (Figure 7B). In
fact, remaining luciferase levels were indistinguishable from
mock-electroporated cells (dashed line in Figure 7B) or a
replication-deficient full-length HCV control
Tough decoy (TuD)
tary Figure S12B). Identical findings were made after
infection with purified full-length reporter HCV, which was also
completely blocked in our engineered cells (Figure 7C).
Curiously, a Dengue virus (DV) subgenomic replicon used as
control also amplified inefficiently in the TS 30.20 cells,
albeit it still expressed above the background levels observed
in the mock control (Figure 7D) or expected with fully
replication-deficient DV (55). This indicates that this clone,
which also showed the highest inhibition of the HCV
subgenomic replicon (Figure 7A), has a defect that broadly
impairs replication of flaviviruses. Importantly, the same DV
control behaved normally in clone T2 31.3 and replicated
even slightly better in clone U6 20.16, verifying that the
HCV inhibition observed in these two clones is truly specific
and mediated by the integrated anti-HCV hairpin. This was
further validated through sequestration of the anti-HCV
shmiRHCV318 in clone T2 31.3 with a ‘tough decoy’
inhibitor (56), which rescued the inhibition of HCV
replication (Figure 7E and F and Supplementary Figure S13).
Here, we demonstrated the feasibility to usurp an
endogenous miRNA locus for stable, precise and potent expression
of an exogenous promoterless RNAi hairpin, using
nucleases for its site-specific integration. Our lead cellular clone,
T2 31.3, (i) carries a single targeted shmiRHCV318
integration, (ii) shows neither mutations in the miR-122 alleles
nor substantial changes in gene or miRNA expression
(including miR-122), (iii) stably expresses shmiRHCV318 for
at least 20 passages (data not shown) and (iv) specifically
and robustly inhibits HCV, together illustrating the great
potential of our new RNAi expression strategy.
Compared to conventional stable transfection or
transduction of RNAi cassettes, our unique approach essentially
differs in that it obviates the need for ectopic regulatory
elements for RNAi hairpin transcription, especially
vectorborne RNA polymerase II or III promoters. This is highly
advantageous considering that exogenous promoters can
dysregulate cellular genes via cis- or trans-activation,
compete for transcription factors, get silenced over time and/or
vary in strength between different cell types (35). A recent
example strikingly illustrating these concerns are findings
in mice that AAV vector integration into the Rian locus
can cause hepatocellular carcinoma (HCC), depending on
vector dose and promoter/enhancer elements (31).
Accordingly, HCC incidences reached over 70% when the transgene
was expressed from a chicken beta-actin or a
thyroxinebinding globuline promoter, but 0% with a human
alpha-1antitrypsin promoter (the same used in clinical AAV trials
(57)). For reasons unknown, the first two promoters but not
the third had induced an upregulation of various small
noncoding RNAs including miR-543 as well as of the Rtl1 gene
(associated with HCC (58)). Importantly, these adverse
localized transcriptional perturbations were not due to the
integration per se since all three vector variants were found
in the Rian locus. Similarly, another study provided
evidence that integration of the WT AAV2 3 inverted terminal
repeat (ITR), believed to have intrinsic enhancer activity,
might be associated with HCC in humans (59). It must be
noted that to date, over 100 clinical trials with AAV vectors
yielded no evidence that this putative tumorigenic
property is conserved in recombinant ITRs (30,60). Nonetheless,
these latest studies highlight dangers and malignancy risks
that can be associated with ectopic DNA cassettes
comprising strong promoters/enhancers and capable of
chromosomal integration. Our new strategy, where the shmiRNA
lacks its own promoter and is placed under an endogenous
miRNA promoter, helps to alleviate such concerns about cis
or trans perturbation of cellular gene expression, and
concomitantly promises persistent and cell-specific expression
at physiological levels (depending on the selected miRNA).
With these features, our work complements and expands
on recent efforts to integrate exogenous promoterless
cDNAs into tissue-specific genes that likewise used
homologous recombination and followed the same rationale. In one
example, the Porteus lab integrated fluorescent reporters
under the - or -globin promoters in human K562 cells
(61), to screen small compounds for effects on transcription
of the endogenous -/ -globin genes. A second example
comes from the Kay lab who integrated the human factor
IX cDNA into the abundantly expressed albumin locus in
mouse livers, using AAV8 vectors for cDNA delivery (33).
Another study also used AAV vectors and harnessed the
same locus, but integrated other (partial) therapeutic
cDNAs (62). Moreover, in this work, the efficiency of cDNA
integration was boosted by co-expressing Zinc-finger
nucleases (ZFNs) against the albumin locus from AAV vectors.
Finally noteworthy are two studies that generated
transgenic animals via CRISPR-mediated integration of
promoterless ectopic sequences into cellular genes. In one, the
human albumin cDNA was inserted into the albumin locus
in swine zygotes, creating transgenic pigs secreting human
albumin in their blood (63). In the other study, transgenic
mice were created by inserting an artificial miRNA into an
intron of the eEF-2 gene, via microinjection of mouse
embryos and subsequent PCR-based selection of positive
Our own work adds a unique and versatile concept to this
rapidly growing list of strategies that harness endogenous
elements to control exogenous sequences, by exemplifying
the possibility to engineer and repurpose a human miRNA
locus for shmiRNA expression. Like some of the
aforementioned studies, we employed TALEN and CRISPR
nucleases to induce double-stranded DNA breaks and thus boost
the efficiency of homology-directed repair. This benefit of
including nucleases was also highlighted by Sharma et al.
who failed to obtain detectable factor IX levels in mice when
trying to integrate the cDNA in the absence of co-delivered
ZFNs (62). Further informative is that Barzel et al. achieved
only 0.5% gene editing in their nuclease-free in vivo targeting
approach (33). Concurrently, the Zhang lab reported over
40% gene editing with AAV/CRISPR vectors in mouse
livers (64), the same organ targeted by Sharma or Barzel et al.
(33,62). Importantly, this high number only reflects gene
cleavage and faulty repair, leaving open what the
frequencies of targeted integration of foreign DNA will be in an in
Generally, one may conclude that a common asset of
promoterless cDNA or RNAi hairpin integration strategies
is a gain in specificity and safety, compared to
promiscuous chromosomal insertion of vectors. Concomitantly, we
note that our use of current nuclease generations resulted
in sporadic insertions or deletions within the targeted
region. Moreover, we found a slight, 2-fold reduction of
mature miR-122 in our lead clone T2 31.3 despite the absence
of mutations in both miR-122 alleles (see Supplementary
Discussion parts A and B for likely explanations). Together,
this illustrates the spectrum of possible on-target
modifications that can be found and that should be considered
in forthcoming iterations of our new approach. Whether
or not such extra alterations can be tolerated will depend
on the exact target and application. In the case of
miR122 and HCV, they might in fact be beneficial in view of
this miRNA’s critical role as viral dependency factor and of
in vivo data that miR-122 inhibition suppresses viremia in
HCV-infected chimpanzees and patients (65–67).
Congruent with this, clone U6 20.16 with a large mutation in both
miR-122 alleles even outperformed our lead clone T2 31.3 at
inhibiting subgenomic HCV replicons. However, U6 20.16
also differed in global gene and miRNA expression profiles
(see Supplementary Discussion part C for possible reasons),
hampering direct comparison. One should moreover
consider that miR-122 plays various essential roles in the liver
and acts as tumor suppressor (68,69), suggesting the
importance of keeping at least one allele intact when engineering
this locus. As noted, this was indeed the case in clone T2
31.3 and in three others: T2 4.37 and U6 6.30 (both
miR122 alleles intact), and U6 7.16 (one allele intact); i.e. 4 out
of 7 (57%).
Next to on-target effects, one must consider possible
adverse off-target modifications, although they seem rare in
our lead clone T2 31.3 within the detection limits of our
assays. Fortunately, nuclease specificity is an area of highly
active inquiry, and there are already numerous
encouraging options to minimize off-targeting. Most notable are
the latest generations of CRISPR/Cas9 systems with
improved specificity and safety, including truncated gRNAs,
double-nicking approaches, Cas9 orthologs, high-fidelity
Cas9 mutants, the Cpf1 nuclease or split Cas9 variants (70–
80). Looming efforts to enhance CRISPR specificity will
concomitantly benefit from the expanding repertoire of
experimental options to detect these adverse events, such as
ChiP-Seq, Digenome-Seq or GUIDE-Seq (81–86). While
our present focus was to provide proof-of-concept for our
novel strategy including a first characterization of possible
outcomes, we readily anticipate that future iterations will
greatly profit from these tools and hence see an increase in
specificity and thus safety.
These improved CRISPR systems and their compatibility
with gene delivery vehicles (87,88) also imply a broad
applicability of our new RNAi expression strategy beyond our
feasibility study in cultured liver cells. One intriguing use
could be ex vivo engineering of induced pluripotent stem
cells before differentiation into liver cells, or direct
modification of primary human hepatocytes. In both cases,
integration of RNAi hairpins could genetically ‘immunize’
these cells against infection with hepatitis viruses prior to
transplantation into a patient with hepatitis. Likewise,
engineering an anti-HIV RNAi hairpin into a miRNA locus
in hematopoietic progenitor cells could yield HIV-resistant
cells for autologous transplantation. This would be
similar to, but probably genetically safer than, clinical trials
in which such hairpins were stably delivered with
lentiviral vectors under exogenous promoters (89). Interestingly,
miR-122 is induced during HIV infection (90), implying
that our miR-122-specific tools and strategies could be
directly applied. Moreover, notable is our evidence that the
miR-122 locus (and likely others as well) tolerates at least
two different additional hairpins, which could be useful
to inhibit highly mutagenic viruses such as HCV or HIV
with combinatorial RNAi (91,92). Such attempts will profit
from the vast experience in multiplexing of RNA hairpins
gained in other contexts, including long hairpin RNA
precursors, engineered miRNA polycistrons, extended short
hairpin RNAs or clusters of TuD inhibitors, to name a few
(92–96). A third attractive example for clinical applications
are human cancers which are frequently characterized by
dysregulation of cellular miRNAs (97), suggesting the
possibility to exploit viral CRISPR vectors (50,64,87,88) to
integrate cytotoxic RNAi hairpins into miRNA genes that are
We moreover see great potential of our new approach
for generation of transgenic knockdown animals that
stably express RNAi hairpins, to study gene function in normal
physiology or disease. For these models to be stringent, it is
essential to avoid local effects of integration sites, multiple
single or tandem insertions, or transgene silencing. All these
side effects are typical for conventional transgenesis
technologies, but may be overcome by repurposing cellular
promoters for expression of minimal RNAi hairpins. One study
exploited a ubiquitously expressed mouse gene for this
purpose (34), but our own strategy that uses a tissue-specific
miRNA locus may be even more beneficial as it combines
robust expression with high lineage specificity. Finally, our
approach to link exogenous to endogenous RNAi triggers,
or, generally, to precisely engineer and harness miRNA loci
for expression of detectable foreign RNA sequences, should
prove valuable for investigations into fundamental cellular
RNAi and genome biology. This could include a direct
replacement of a target miRNA with another artificial or
natural miRNA, to study effects on physiology, differentiation
or other basic cellular parameters.
We acknowledge that all these future applications,
especially direct in vivo use and other advances toward clinical
translation, require improvements in efficiency and
accuracy of nuclease-mediated DNA modification. Ideally, these
will alleviate the necessity for clonal selection, which
results from the possibility of undesirable on- and off-target
modifications that is inherent to all current nucleases. We
are aware of these restrictions but reiterate that these are
not intrinsic to our original strategy, and that some
concerns about efficiency and specificity will be resolved with
newer generations of nucleases and delivery systems.
Particularly encouraging is the aforementioned report of greater
than 40% targeted gene editing in adult mouse livers, using
moderate doses of an AAV8 vector expressing Cas9 from
S. aureus and delivered via peripheral infusion (64). Similar
optimism is raised by the successful AAV/ZFN-mediated
in vivo integration of promoterless cDNA in mouse livers
(62). We further note an intriguing study showing
cell-tocell transfer of small RNAs in livers of mice (98),
implying that therapeutic benefit may be achieved from editing
only a fraction of cells in the liver (and potentially other
organs). It may also be rewarding to replace Cre recombinase
with alternatives posing a lower inherent risk of
chromosomal translocation (albeit not observed in our study), e.g.
hyperactive piggyBac transposase delivered as protein (99).
Finally promising is that homology-directed repair can be
promoted in mammalian cells and mice by suppressing key
factors in the non-homologous end joining pathway, such
as DNA ligase IV, which expands the options to enhance
frequencies of targeted modification of miRNA loci (100–
102). In light of these and other advances, and of the
extreme pace with which the entire genome engineering field
is currently progressing, we are optimistic that the way will
soon be paved for a wealth of exciting ex or in vivo
applications of our new concept for safe and accurate expression
of exogenous RNAi hairpins.
Supplementary Data are available at NAR Online.
The authors thank the microarray and sequencing units of
the DKFZ Genomics and Proteomics Core Facility for
providing gene expression, miRNA expression as well as whole
genome sequencing analysis. The authors are grateful to
Sarah Klinnert for help with cloning, as well as to members
of the Grimm lab for critical reading.
German Research Foundation [DFG, EXC81 (Cluster of
Excellence CellNetworks) to E.S., S.G., S.M. and D.G.,
SFB1129 (Collaborative Research Center 1129) to D.G.
(TP2), TRR179 (Transregional Collaborative Research
Center 179) to R.B. (TP9) and D.G. (TP18), SPP1395
(InKoMBio, TH 900/6-1) to B.K.]; Helmholtz Initiative
for Synthetic Biology [E.S and D.G.]; Heidelberg
University Graduate Academy [PhD completion grant to E.S.];
HBIGS International Graduate School [MD/PhD
program to D.R.]; European Research Council [Starting grant
Latent Causes (259294) to F.J.T.]; Federal Ministry of
Education and Research [BMBF, e:Med program for systems
biology (PANC-STRAT consortium, 01ZX1305) to T.B.].
Funding for open access charge: Heidelberg University
Conflict of interest statement. None declared.
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