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J Biomed Biotechnol. 2006; 2006: 87340. Published online 2006 May 28. doi: 10.1155/JBB/2006/87340. | PMCID: PMC1559937 |
RNA-Mediated Gene Silencing in Hematopoietic Cells Letizia Venturini, Matthias Eder,* and Michaela Scherr* Department of Hematology, Hemostasis, and Oncology, Hannover Medical School, 30625 Hannover, Germany Received February 8, 2006; Accepted April 3, 2006. This is an open access article distributed under the Creative Commons
Attribution License, which permits unrestricted use, distribution, and reproduction
in any medium, provided the original work is properly cited. In the past few years, the discovery of RNA-mediated gene
silencing mechanisms, like RNA interference (RNAi), has
revolutionized our understanding of eukaryotic gene expression.
These mechanisms are activated by double-stranded RNA (dsRNA) and
mediate gene silencing either by inducing the sequence-specific
degradation of complementary mRNA or by inhibiting mRNA
translation. RNAi now provides a powerful experimental tool to
elucidate gene function in vitro and in vivo, thereby opening new
exciting perspectives in the fields of molecular analysis and
eventually therapy of several diseases such as infections and
cancer. In hematology, numerous studies have described the
successful application of RNAi to better define the role of
oncogenic fusion proteins in leukemogenesis and to explore
therapeutic approaches in hematological malignancies. In this
review, we highlight recent advances and caveats relating to the
application of this powerful new methodology to hematopoiesis. The discovery of RNAi In 1998 Andrew Fire and Craig Mello discovered in a series of
experiments in Caenorhabditis elegans that injection of
sense or antisense RNAs led to negligible decreases of target RNA,
whereas introduction of dsRNA resulted in effective and specific
degradation of cytoplasmic mRNA. Furthermore, these silencing
effects of dsRNA in C elegans were systemic and heritable
[ 1]. Later on, the sequence-specific gene
silencing capacity of dsRNA, now known as RNA interference (RNAi), has been linked to
previously described gene silencing phenomena such as
posttranscriptional gene silencing (PTGS), co-suppression in
plants, and quelling in fungi [ 2– 7]. Following this
exciting and fundamental discovery, intense studies were
undertaken with the purpose to dissect the molecular mechanism of
RNAi. Indeed, in the past few years, many details of the
biogenesis of small dsRNAs have been elucidated, and components,
structure, and function of protein complexes of the RNAi machinery
have been identified. RNAi has thus emerged as an evolutionarily
highly conserved and fundamental mechanism for the regulation of
gene expression and has rapidly been developed into a tool to
analyze gene function. RNAi molecular pathways The detailed molecular mechanism of RNAi has been the subject of
numerous recent reviews [ 8– 16]. As this review concentrates on delivery of RNAi in the hematopoietic system and
discusses its potential diagnostic and therapeutic applications in hematology,
we will only briefly resume the principal phases of dsRNA-mediated
gene silencing pathways as schematically represented in
. Two types of small noncoding dsRNA can serve as effector molecules
and trigger RNAi: small interfering RNA (siRNA) and micro RNA
(miRNA). Long dsRNAs, introduced experimentally into the cell or
generated as intermediates during viral infections
[ 17] or as
aberrant transcripts derived from inverted tandem repeats and
transposons [ 18], are processed
in the cytoplasm by a ribonuclease (RNase) III-like enzyme called Dicer
[ 19, 20]
into siRNA duplexes of 21–25 nt in length with 3′ dinucleotide overhangs, 5′-phosphates, and 3′-hydroxyl termini
[ 21]. With the discovery of micro RNAs (miRNAs) the important
physiological role of RNA-mediated gene silencing for regulating
gene expression during development [ 22],
differentiation
[ 23], and apoptosis
[ 9, 24],
as well as its possible involvement in diseases like cancer, soon became manifest (see
[ 8, 9]
for review). miRNAs constitute a large class of
endogenously expressed, highly conserved, noncoding small RNA
molecules which act as negative regulators of gene expression in a
variety of organisms ranging from plants to mammals
[ 9].
Usually miRNAs are transcribed from endogenous genes by RNA
polymerase II [ 25,
26] as long primary transcripts
(pri-miRNAs) (see [ 27] for review)
possessing 5′-caps and
3′-poly A tails [ 25,
28] (). Recent
studies suggest that miRNA expression is regulated at
the level of transcription [ 29– 31], similarly to mRNA and
likely involving a similar or identical molecular machinery. The
pri-miRNAs undergo a two-step processing to give rise to
the mature miRNAs: in the nucleus, the characteristic
stem-loop structure of the precursor-miRNA (pre-miRNA) of
~ 70 bp is released from the primary transcript by another
RNase III-like enzyme called Drosha [ 32– 36]. The pre-mRNAs are then transported from the nucleus to the cytoplasm
by exportin-5 [ 37– 40], where its 2 nt 3′-hydroxyl
terminus is recognized by Dicer, which generates the mature miRNA
of 21–23 nt length [ 41– 43]. Dicer thus represents the
core of the RNAi machinery where the different RNA-mediated gene
silencing processes converge (). Only one strand of the mature si/miRNAs, distinguished by the
lower thermodynamic stability at its 5′ terminus
[ 44, 45],
is retained in the active ribonucleoprotein effector complexes and
acts as a guide to target the homologous mRNA
(). Normally, siRNAs are fully complementary
to their cognate mRNAs and guide the RNA-induced silencing complex
(RISC) [ 46,
47] to the target transcripts, followed by
cleavage and degradation. A member of the Argonaute
(Ago) family of proteins [ 48,
49], which contain a small
RNA-binding domain (PAZ [ 50])
and an RNase activity (PIWI
domain [ 51]),
constitutes the catalytic ribonuclease
component of RISC responsible for cleaving the target mRNA at the
center of the region complementary to the guide RNA
[ 48, 49].
RISC is then released and the cleaved mRNA is degraded by cellular
exonucleases. Except in plants where they have perfect homology to
their targets and act exactly as siRNAs inducing mRNA degradation
[ 52] upon integration into complexes
usually referred to as
miRNA-ribonucleoproteins (miRNPs), most miRNAs bind only with
partial complementarity to the 3′ untranslated region (UTR) and
lead to translational inhibition downstream of the initiation step
[ 53]. In fact, apart from the
distinction based on their
biogenesis, a precise boundary between si- and miRNA functionality
cannot always be established, since both have been shown to be
capable to target mRNA for degradation or translational
inhibition, depending on the cellular context and the degree of
complementarity between the RNA effector and the mRNA target
[ 46, 54– 58]. In addition to posttranscriptional gene silencing, RNAi has also
been demonstrated to induce transcriptional repression through
RNA-directed DNA methylation in plants [ 59],
and recent studies have reported similar effects even in human cells
[ 60, 61],
inducing heterochromatin formation (see [ 62,
63] for review). RNAi IN MAMMALIAN CELLS: DESIGN, DELIVERY, AND TARGET VALIDATION After its discovery, RNAi was rapidly employed as a powerful tool
for large-scale reverse genetic screens in C elegans,
where it can be easily induced by direct injection of dsRNAs, by
feeding worms with bacteria engineered to express si-RNAs,
or simply by soaking the animals in medium containing
siRNAs. Furthermore, in nematodes, RNAi acts systemically and is a
long-lasting heritable event due to the presence of RNA-dependent
RNA polymerases (RdRPs) which allow the amplification of the
trigger and the perpetuation of the silencing process
[ 64– 66]. In mammals, however, initial efforts to use RNAi for specific gene
silencing encountered more difficulties, especially due to the
induction of nonspecific inhibition of gene expression resulting
from the activation of the interferon (IFN) response pathway by
dsRNAs longer than 30 bp [ 67].
The discovery from
Elbashir et al [ 68] and Caplen et al
[ 69] that RNA
duplexes of 21 nt in length, mimicking the Dicer cleavage
products, were able to mediate efficient and specific RNAi upon
transfection into mammalian cells without eliciting the INF
response, finally enabled loss-of-function studies of specific
target genes in mammalian systems as well. Designing RNAi effector molecules To design efficient siRNAs several parameters should
be considered. Although duplex RNAs of 21 nt in length were
shown to be the most effective RNAi triggers, recent studies
suggest that 27- to 29-mers may be more active than 21-mers
[ 70– 72]. This could be due to the fact that these dsRNAs
are processed by Dicer and thus may be incorporated directly and
more efficiently into RISC [ 73].
Reynolds et al [ 74]
conducted systematic analyses to evaluate physicochemical
characteristics associated with highly functional siRNAs and set
up an algorithm with several criteria which significantly improved
selection of potent siRNAs. To comply with the rules promoting
asymmetric incorporation into RISC, the base pair at the 5′ end
of the siRNA antisense (guide) strand should have a lower
thermodynamic stability compared with the 3′-end
[ 44, 45].
Accordingly, the presence of three or more A/U nucleotides at the
3′-terminus of the sense strand was defined as a criterion for
siRNA functionality [ 74].
Low G-C content (30%–52%),
lack of internal inverted repeats which can form secondary
structures, and specific nucleotide preferences at positions 3
(A), 10 (U), 13 (absence of G), and 19 (A, absence of G or C) on
the sense strand also increased the probability of selecting a
potent siRNA in this study [ 74].
Concerning the specificity
of the sequence match necessary to achieve efficient gene
silencing, different groups have reported varying degrees of
mismatch tolerance for siRNA-mediated silencing
[ 75– 81]. Mismatched small RNAs may still be
competent and can function like miRNAs which may bind to mRNAs
with multiple target sites in the 3′-UTR
[ 82]. miRNA
binding is not limited to the 3′-UTR but can include the coding
sequence of an mRNA as well [ 83].
These results demonstrate
the possibility of off-target effects by siRNAs used for
experimental or therapeutic purposes. Therefore, for prevention of
cross-reactive silencing, a BLAST search of potential target
sequences should be performed in order to exclude candidate siRNAs
with some degree of homology with other genes
[ 74, 84].
For the efficacy of gene silencing, the overall stability of the
antisense RNA/RISC-mRNA complex is probably more important than
the absolute number of mismatches. Accordingly, base pairing at
the center of the duplex is critical, while one or two mismatches
located at the 3′- or 5′-end of the siRNA may be well
tolerated [ 85].
Since regions which are not involved in
intramolecular folding have been demonstrated to be optimal
targets [ 86],
computational approaches have been reported to
analyze the secondary structure and the local folding of the
target mRNA. However, all predictions based on physicochemical
characteristics, sequence homologies, or secondary structure can
not guarantee the generation of an efficient siRNA, and functional
testing is still always required. Similarly, different
si- RNAs targeting the same gene may have different
silencing efficacies not always predictable by the parameters
discussed above. Therefore, more than one target sequence should
be tested empirically to identify the optimal small RNA for
efficient and specific silencing of a given target mRNA. RNAi delivery in mammalian cells RNAi in mammalian cells can be triggered by direct introduction
through injection, electroporation, lipid-mediated transfection,
nanoparticles, or antibody bound enzymatically
generated or chemically synthesized siRNAs, among others.
Alternatively, siRNAs or small hairpin RNAs (shRNAs) can be
delivered by vector-based intracellular expression. Synthetic siRNA-mediated RNAi siRNAs can be synthesized chemically [ 75,
87, 88],
generated enzymatically through in vitro transcription by T7 phage
polymerase [ 89,
90], or through endonuclease digestion by
recombinant Dicer of in vitro transcribed long dsRNA
[ 91, 92].
In mammalian cells, direct delivery of siRNAs can only induce
transient silencing due to their limited half-life and to their
dilution during cell division. Vector-based RNAi Transient downregulation of gene expression may not be sufficient
for many applications, for example, for studies of proteins with
long half-lives in rapidly dividing cells. In order to produce
long-lasting RNAi in mammalian cells, plasmids and viral
expression vectors have been developed to drive continuous
intracellular expression of siRNA or shRNAs under the control of
highly active RNA polymerase III promoters such as U6
or H1 [ 93, 94]. As represented in , the two strands of an
siRNA can be transcribed from distinct expression units, either
cloned in tandem or in two separate vectors
[ 95, 96],
or can result from bidirectional transcription of a single 19- to 29-mer
DNA fragment under the control of two opposite promoters
[ 97, 98].
The intracellular expression and hybridization of
the two strands gives rise to functionally active siRNA duplexes.
However, the most commonly used approach involves the
intracellular expression of shRNAs. They are transcribed as
single-stranded RNAs from an expression cassette inserted
immediately downstream of the pol III promoter, which contains, in
the following order, a 19 nt sequence homologous to the target
mRNA, a spacer 6 to 9 nucleotides in length, the antisense
sequence, and the RNA pol III terminator signal composed of a
stretch of about 6 thymidines. After transcription, the resulting
stem-loop RNA structure, like miRNA precursors, is cleaved by
Dicer to yield a functionally active siRNA [ 93,
99, 100]
(). A second generation of RNAi-delivering
vectors is based on RNA pol II promoters driving transcription of
shRNAs incorporated into a miRNA chimeric transcript, comprising
flanking sequences optimized for Drosha/Dicer processing
[ 100, 101]. A major advantage of vector-dependent RNAi is the usage of
selectable markers to generate stable transfectants or of reporter
genes such as green fluorescent protein (GFP) or red fluorescent
protein (RFP) to identify and eventually isolate the
si/shRNA expressing cells in a quantitative manner. When using
shRNA-expressing vectors, however, it is absolutely necessary to
confirm the sequence of the shRNA expression cassette, because
genetic recombination and/or introduction of point mutations are
very frequent and can occur in almost every step of the cloning
strategy. Finally, when the target gene to be silenced is essential for cell
survival and/or proliferation and the constitutive knockdown of
its expression is even lethal, employing inducible or
tissue-specific RNAi could be especially important
[ 100]. In
the last years, various methods for inducible expression of shRNAs
have been described. Tetracycline-inducible [ 94,
102, 103]
and ecdysone-inducible [ 104] expression systems have been
reported which mediate induced and reversible downregulation of
gene expression. However, no standard technique for inducible RNAi
has been established so far. Many suspension and primary cells are difficult to transfect
efficiently. However, viral transduction strategies allow stable
induction of RNAi in these cells. Particularly, the capacity of
lentiviruses to integrate into the genome of noncycling cells, such as stem cells or terminally differentiated
cells, renders lentiviral vectors much more efficient than
retroviral vectors in inducing RNAi in these cells
[ 105, 106]. Finally, transgenic technology has also been adapted for in vivo
delivered RNAi in mice. Transgenic animals have been reported
which produce siRNAs constitutively [ 107] or conditionally,
in a stage- or tissue-specific manner [ 108], to repress
selected target genes. Tiscornia et al [ 106] efficiently
used a lentiviral vector system to express siRNA in
preimplantation mouse embryos. In vivo systemic delivery of siRNAs Chemical modifications are required to potentiate siRNA nuclease
and thermodynamic stability in vivo without compromising their
efficacy. Recently, several groups reported different approaches
for systemic in vivo delivery of siRNAs. Soutschek et al
[ 109] described
intravenous injection in mice of chemically
modified naked siRNAs coupled to a cholesterol group chemically
linked to the terminal hydroxyl group of the sense strand to
promote entry into the cells. In vivo delivery of chemically
modified siRNAs encapsulated into liposome particles has been
recently reported by Morrisey et al [ 110],
and Song et al [ 111]
described an antibody-based delivery system which could
offer a possibility for systemic, cell-type-specific siRNA
delivery. Specificity of RNAi-induced gene silencing Prior to functional analysis, monitoring the level of target mRNA
expression is necessary for siRNA target validation. Quantitative
RT-PCR is a fast and reliable method to measure target transcript
levels in specific versus control siRNA-treated cells, to
ascertain that any phenotypic changes are really due to
specific gene knockdown and not to nonspecific effects of the RNAi
strategy. This issue can be further addressed by verifying that
the same phenotype is induced by siRNAs homologous to
different regions of the target transcript. The correlation
between the extent of gene silencing and dose of the delivered
si/shRNA can offer some evidence of specificity. Finally, the
availability of a rescue experiment able to revert the
loss-of-function phenotype can ensure RNAi specificity. RNAi in the HEMATOPOIETIC SYSTEM siRNAs and hematopoietic cells Apart from its transient nature in mammals, the use of RNAi in
primary hematopoietic cells is limited by the difficulty to
deliver siRNA through conventional transfection methods. In
contrast, lentiviruses have been shown to efficiently transduce
human hematopoietic stem and NOD/SCID repopulating cells (HSCs and
SRCs) as well as more committed colony forming progenitors
[ 112– 114]
and can offer a useful means for effective and
stable delivery of RNAi triggers in the hematopoietic system.
Lentiviral transduction was successfully employed by our group to
induce RNAi against the common β chain of the receptors for
granulocyte-macrophage colony-stimulating factor (GM-CSF),
interleukin (IL)-3, and IL-5 in human CD34 + SRCs and
colony forming cells [ 115].
Similarly, Schomber et al [ 116]
reported efficient and stable silencing of the
p53 gene in human cord blood-derived CD34 + cells
through lentivirus-mediated RNAi. Application of siRNAs to target leukemia-associated oncogenes Chromosomal translocations leading to the expression of chimeric
oncoproteins are frequently involved in malignant transformation
in leukemias and lymphomas. In the past few years, RNAi technology
has been used to specifically silence the expression of
translocation products ( Table 1). For example, the
bcr-abl oncogene resulting from the t(9;22) translocation
characterizes chronic myelogenous leukemia (CML) and some variants
of acute lymphoblastic leukemia (ALL). It codes for a
constitutively active cytoplasmatic tyrosine kinase which is both
necessary and sufficient for leukemic transformation in several
models [ 129– 131].
Different research groups
[ 117– 120]
have demonstrated the feasibility of
specific interference with the bcr-abl expression without
affecting the expression of wild-type c-abl or c-bcr using
breakpoint-specific siRNAs delivered by electroporation in bcr-abl
positive hematopoietic cell lines and primary CD34 +
cells from CML patients. Upon siRNA treatment, inhibition of
survival and proliferation, increased sensitivity to the
abl-specific tyrosine kinase inhibitor imatinib mesylate (STI571)
[ 123, 132],
and increased apoptosis were observed
[ 119, 120]. | Table 1RNAi targeting hematopoietic fusion genes. |
In a recent study, our group used lentiviral gene transfer of
shRNAs to trigger stable RNAi targeting the bcr-abl oncogene
[ 133]. Stable, but not transient,
RNAi was demonstrated to induce depletion of bcr-abl positive cells from suspension
cultures. This depletion, as well as the degree of bcr-abl gene
silencing, correlates with the multiplicity of lentiviral
infection (MOI), the number of lentiviral integration into the
host cell genome, and the expression level of the RFP reporter
gene: cells with lower RFP expression and fewer lentiviral
integrations could survive and were selected in suspension
cultures but still showed reduced bcr-abl expression, aberrant
proliferation kinetics, and enhanced sensitivity to STI571 as
compared to controls. Furthermore, in contrast to transient RNAi
[ 118], stable RNAi-induced silencing
of bcr-abl inhibited the
colony forming capacity of primary CD34 + cells from
CML patients. Heidenreich et al [ 123]
used siRNAs to specifically repress
the AML1/MTG8 fusion product resulting from the t(8;21) translocation [ 134]
found in about 10% to 15% of all
cases of de novo AML. Electroporation of siRNAs specific for the
fusion site of the AML1/MTG8 into the Kasumi-1 t(8;21)-positive cell line specifically suppressed the expression of the fusion
product, without impairing the expression of the two respective
wild-type genes. Despite the transient character of the RNAi
strategy employed, the authors described some functional effects
due to the suppression of the fusion protein such as increased
myeloid differentiation and reduced clonogenic potential upon
TGFβ/vitamin D treatment. Stable retroviral delivery of shRNAs was used by Chen et al
[ 124] to target the fusion
sequence of the TEL-PDGFβR fusion product derived from the t(5;12) translocation, a
recurrent cytogenetic aberration associated with chronic
myelomonocytic leukemia (CMML) [ 135].
Stable RNAi-mediated inhibition of TEL-PDGFβR significantly reduced the
proliferation of TEL-PDGFβR-transformed Ba/F3 cells, but did
not restore IL-3 dependence, concordant with a marked decrease,
but not abrogation of TEL-PDGFβR expression and selection of
TEL-PDGFβR expressing cells. The authors also reported a
significantly prolonged disease latency and survival of nude mice
or Balb/C mice injected with TEL-PDGFβR-transormed Ba/F3
cells coexpressing siRNA as compared with injection of
TEL-PDGFβR-transformed cells not expressing siRNA. However,
as observed in cell culture, the expression of siRNA alone was not
sufficient to completely abrogate TEL-PDGFβR-induced
transformation in these murine models. A synergistic effect
between si- RNAs and small molecule inhibitors of
tyrosine kinase activity, such as imatinib, was also demonstrated
in TEL-PDGFβR-transformed Ba/F3 cells. The mixed-lineage leukemia (MLL) gene is involved in numerous
translocations in a variety of leukemias [ 136].
Most frequently, the MLL gene is fused to the AF4 gene as a consequence
of the t(4;11) translocation [ 137– 139],
found in acute lymphoblastic leukemia (ALL) with poor prognosis in infants
[ 140]. Thomas et al
[ 125] applied siRNAs to silence
MLL-AF4 and demonstrated decreased proliferation and clonogenicity
of t(4;11)-positive leukemic cells as well as induction of
apoptosis through caspase-3 activation and repression of the
BCL-X L anti-apoptotic gene. They also observed that
MLL-AF4 depletion resulted in a reduced expression of the homeotic
genes HoxA9, MEIS1, and HoxA7, known to be upregulated by MLL
fusion proteins, and of the CD133 marker for hematopoietic stem
cell and early progenitors, which may suggest a reactivation of
hematopoietic differentiation. Finally, using a NOD/SCID mouse
xenotransplantation model, the authors showed that siRNA-mediated
repression of MLL-AF4 compromised leukemic engraftment and the
development of leukemia in vivo. The t(2;5) chromosome translocation fuses the ALK (anaplastic
lymphoma kinase) gene on chromosome 2 to the nucleophosmin (NPM1)
gene on chromosome 5 and is associated with anaplastic large-cell
lymphomas (ALCLs) [ 141].
In a recent study, Piva et al
[ 127] reported that silencing of
NPM1-ALK induced by shRNAs
directed against the 3′ sequences encoding the cytoplasmatic
domain of ALK-R, which is retained in all oncogenic fusion
proteins involving ALK, leads to abrogation of NPM1-ALK-mediated
transformation of MEF cells and inhibition of cell growth in
several human NPM1-ALK-positive cell lines. Moreover, an increased
number of apoptotic cells together with caspase activation and
downregulation of the anti-apoptotic protein survivin were
detected in ALCL cells 4 days after lentivirus-mediated RNAi.
Similar results were seen in vivo: shRNA-expressing ALCL cells
injected into NOD/SCID mice revealed a reduction in tumor
formation as compared to control cells. Furthermore, in a second
series of experiments, injection of lentiviruses driving shRNA
expression directly into ALCL tumor masses showed growth
inhibition of neoplastic cells, and histologic sections of the
tumors demonstrated the presence of large necrotic regions and, in
areas with retained viability, many apoptotic cells. The receptor FMS-like tyrosine kinase 3 (FLT3) is the single most
frequently mutated gene in AML. It is constitutively activated by
internal tandem duplications (ITDs) within the juxtamembrane
domain or by point mutations within the catalytic kinase domain in
approximately 30% of AML patients
[ 142, 143]
and appears to confer an unfavourable prognosis. RNAi-mediated silencing of FLT3
was reported by Walters et al [ 128].
The authors used an siRNA pool to effectively downregulate the expression of FLT3 in
FLT3-ITD-positive human leukemia cells and showed diminished
phosphorylation of downstream signalling molecules, comprising
STAT5, MAPK, and Akt, inhibition of cell proliferation, and
induction of apoptosis. In addition, upon siRNA treatment in these
cells, they found increased sensitivity to treatment with the FLT3
inhibitor MLN518, further demonstrating the potential benefit of
such combined therapeutic approaches. miRNAs in the hematopoietic system miRNAs associated with hematopoietic differentiation Fine modulation of gene expression is essential for the correct
realization of differentiation programs. Consistent with this,
several groups recently demonstrated the implication of miRNAs in
controlling hematopoietic differentiation. Chen et al [ 23] described three miRNAs,
miR-181, miR-223, and miR-142s, which are differentially expressed in the murine
hematopoietic system, and showed that miR-181 plays a specific
role in B-cell differentiation. They found that miR-181 is
normally expressed at low levels in murine hematopoietic
progenitors and becomes upregulated during B-cell differentiation.
Overexpression of miR-181 in hematopoietic progenitors gives rise
to a greater fraction of B-lymphoid cells than in wild-type
progenitors, in vitro as well as in vivo. In a recent publication, Felli et al [ 144]
described miR-221 and miR-222 as inhibitors of normal erythropoiesis and indicated
the kit receptor mRNA as a major target of these two miRNAs. Using
microarray chip and Northern blot-analysis, they showed that
miR-221 and miR-222 are downregulated in erythropoietic cultures
of cord blood CD34 + progenitors. In addition, they
observed an impairment of human CD34 + cell engraftment
in NOD/SCID mice as well as an inhibition of cell growth in the
c-kit+ TF-1 erythroleukemic cell line upon overexpression of
miR-221 and miR-222. Recent studies conducted by Fazi et al [ 29]
revealed the implication of miR-223 in human myeloid differentiation: miR-223
expression increases during retinoic acid- (RA-) induced
granulocytic differentiation of the NB4 promyelocytic cell line as
well as of blasts from patients with acute promyelocytic leukemia
(APL) undergoing RA treatment. The authors depicted a finely
regulated network involving miR-223 and the transcription factors
C/EBPα, well known for its implication in granulocytic
differentiation [ 145– 147],
and NFI-A [ 148,
149].
C/EBPα and NFI-A, which can induce or repress miR-223
expression, respectively, are in competition for an overlapping
binding site on the miR-223 promoter. In undifferentiated cells,
NFI-A maintains miR-223 and consequently its translation
inhibitory effect at low levels. Upon RA treatment, C/EBPα displaces NFI-A from the miR-223 promoter, thus activating its
expression. Interestingly, NFI-A is a target of miR-223 which,
through a positive feedback, represses NFI-A translation, reduces
the competition with C/EBPα, and maintains sustained levels
of its own expression. miRNA alterations in hematological malignancies The expression of about one-third of human mRNAs appears to be
regulated by miRNAs, each of which, according to computational
analysis, is predicted to regulate a broad spectrum of different
mRNAs [ 150], revealing a very complex regulatory network. As
cancers essentially derive from alteration of gene expression
and/or gene function, it is not surprising that several recent
publications supported the direct involvement of miRNAs in
tumorigenesis. Approximately 50 percent of the known human miRNA
genes are located at fragile sites and cancer-associated regions
of the genome [ 151].
Dysregulation of various human miRNAs
has been associated with leukemias and lymphomas: the precursor of
miR-155 was found to be overexpressed in the majority of childhood
Burkitt lymphoma [ 152];
the miR-15a/miR-16 cluster at locus
13q14 is frequently deleted or downregulated in patients with
B-cell chronic lymphocytic leukemia (CLL), mantle cell lymphoma,
and multiple myeloma [ 153].
As miR-15/16 was demonstrated to
induce apoptosis by targeting the apoptosis inhibitor protein BCL2
in CLL cells [ 154],
downregulation of miR-15a and miR-16-1
can contribute to malignant transformation through BCL2
upregulation and inhibition of apoptosis. The miR-17-92
polycistron, located at 13q31, is amplified in human B-cell
lymphomas [ 155]. He et al
[ 156] found that enforced
expression of the miR-17-92 cluster can augment the oncogenic
potential of c-myc in a mouse B-cell lymphoma model,
offering the first evidence of a miRNA to act as an oncogene.
O'Donnell et al [ 30]
further confirmed the relationship
between this miR cluster and cancer. Using a lymphoma cell line
with inducible c-myc expression, they demonstrated that
the miR-17 cluster is specifically and directly upregulated by
c-myc, but at least miR-17-5p and miR-20a
downregulate E2F1-protein expression, a target of c-myc
which promotes cell cycle progression. These findings reveal a
feedback mechanism through which c-Myc activates E2F1
transcription and simultaneously induces inhibition of its
translation. Recent microarray-based studies have provided evidence that
specific alterations in human miRNA expression profiles are
associated with specific types of cancers. Lu et al [ 157]
established a sensitive method to analyze the expression profiles
of 217 miRNAs in a panel of 334 samples representing diverse human
normal tissues and corresponding tumors. They observed a general
downregulation of miRNAs in tumors compared with normal tissues
and demonstrated that miRNA expression profiles correlate with the
developmental origins of specific cancers. Furthermore, even
within a single developmental lineage, distinct patterns of miRNA
expression seem to reflect the mechanism of transformation.
Indeed, clustering of miRNA profiles of bone marrow samples from
patients with acute lymphoblastic leukemia (ALL) showed a
nonrandom distribution into three major groups in correlation with
previously characterized molecular alterations and phenotypic
classifications (BCR-ABL-positive and TEL-AML1-positive samples,
T-cell acute lymphoblastic leukemias, and mixed lineage
leukemias). Their results suggest that miRNA expression profiles
could be more accurate for the classification and diagnosis of
human cancers than mRNA microarrays. Using miRNA profiling, Calin et al [ 158]
found that different patterns of miRNA expression distinguish CLL cells from
normal CD5+ B cells. In a recent study based on genome-wide
expression profiling of a large number of samples from CLL
patients [ 159],
the same group showed that a miRNA signature
is associated with the presence of other known prognostic factors
(levels of ZAP-70 expression and the mutational status of the
immunoglobulin heavy-chain (IgV H) gene) and with
disease progression in CLL: a molecular signature composed of 13
miRNAs differentiated CLL patients with high levels of ZAP-70
expression and unmutated IgV H from patients with low
ZAP-70 expression and mutated IgV H. Nine of these
miRNAs were significantly overexpressed in the first group of
patients, associated with a poor prognosis. Furthermore, 9 miRNAs
of the prognostic signature were able to discriminate between
patients with a short interval from diagnosis to therapy and
patients with a longer interval: in the first group, 8 of the 9
miRNAs were upregulated, suggesting their involvement in disease
progression. The authors also showed some functionally relevant
mutations in miRNA genes in CLL. Some of these mutations are
located in the flanking sequences of the pre-miRNA, compromising
the correct processing and expression of the mature miRNA. CONCLUSIONS AND FUTURE PERSPECTIVES Despite many remaining technical problems, current advances in
strategies to extend genome-wide screens with siRNA or shRNA
libraries to mammalian cells [ 160– 167]
as well as specific gene
silencing approaches may finally facilitate the identification of
essential genes involved in human diseases and may identify new
potential therapeutic targets. On the other hand, disease-
and stage-specific systematic analysis of miRNA gene-expression
profiles may help to establish new diagnostic and prognostic
markers. Employing RNAi in a therapeutic setting may still encounter
numerous obstacles: the issue of efficient delivery in a clinical
setting, as well as problems deriving from toxicity, and possible
off-target effects. While viral delivery systems are certainly of great utility for
experimental models, further studies are necessary before their
possible therapeutic application may become possible in the
future. If all these obstacles can be overcome, cancer-specific oncogenes,
such as the fusion genes produced by chromosomal translocations
involved in several types of leukemia, could be suitable
candidates for tumor cell-specific targeting in RNAi-mediated
therapeutic approaches. Beside those, gene expression by
infectious organisms may be targeted by RNAi. Indeed, early
clinical trials are under way or being started targeting vascular
epidermal growth factor (VEGF) receptor in age-related macular
degeneration or genes expressed by respiratory syncytial virus
(RSV). Finally, as suggested by recent reports demonstrating the
cooperative effects of RNAi and selective molecular inhibitors,
such as imatinib for Bcr-Abl [ 119,
120] and TEL-PDGFβR [ 124],
and the kinase inhibitor MLN518 for FLT3 [ 128],
combination of targeted therapies including small molecules and
RNAi could be taken into consideration, especially when drug
resistance becomes a problem. We thank Michael Morgan and Michael Stadler for critical reading
of the manuscript. M. Eder and M. Scherr are supported by the H
& W Hector-Stiftung, the Wilhelm Sanders-Stiftung, and the
Deutsche Forschungsgemeinschaft (SFB566). 1. Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC. Potent and specific genetic interference by double-stranded
RNA in Caenorhabditis elegans. Nature. 1998;391(6669):806–811. [PubMed] 2. Covey SN, Al-Kaff NS, Lángara A, Turner DS. Plants combat infection by gene silencing. Nature. 1997;385(6619):781–782. 3. Lindbo JA, Silva-Rosales L, Proebsting WM, Dougherty WG. Induction of a highly specific antiviral state in transgenic
plants: implications for regulation of gene expression
and virus resistance. Plant Cell. 1993;5(12):1749–1759. [PubMed] 4. Napoli C, Lemieux C, Jorgensen R. Introduction of a chimeric chalcone synthase gene into
petunia results in reversible co-suppression of homologous
genes in trans. Plant Cell. 1990;2(4):279–289. [PubMed] 5. Ratcliff FG, MacFarlane SA, Baulcombe DC. Gene silencing without DNA: RNA-mediated cross-protection
between viruses. Plant Cell. 1999;11(7):1207–1216. [PubMed] 6. van Blokland R, ten Lohuis M, Meyer P. Condensation of chromatin in transcriptional regions of an
inactivated plant transgene: evidence for an active role of
transcription in gene silencing. Molecular and General Genetics. 1997;257(1):1–13. [PubMed] 7. Van Der Krol AR, Mur LA, Beld M, Mol JNM, Stuitje AR. Flavonoid genes in petunia: addition of a limited number
of gene copies may lead to a suppression of gene expression. Plant Cell. 1990;2(4):291–299. [PubMed] 8. Ambros V. The functions of animal microRNAs. Nature. 2004;431(7006):350–355. [PubMed] 9. Bartel DP. MicroRNAs: genomics, biogenesis, mechanism, and function. Cell. 2004;116(2):281–297. [PubMed] 10. Filipowicz W. RNAi: the nuts and bolts of the RISC machine. Cell. 2005;122(1):17–20. [PubMed] 11. Hannon GJ, Rossi JJ. Unlocking the potential of the human genome with RNA
interference. Nature. 2004;431(7006):371–378. [PubMed] 12. He L, Hannon GJ. MicroRNAs: small RNAs with a big role in gene regulation. Nature Reviews Genetics. 2004;5(7):522–531. 13. Matzke MA, Birchler JA. RNAi-mediated pathways in the nucleus. Nature Reviews Genetics. 2005;6(1):24–35. 14. Meister G, Tuschl T. Mechanisms of gene silencing by doublestranded RNA. Nature. 2004;431(7006):343–349. [PubMed] 15. Tomari Y, Zamore PD. Perspective: machines for RNAi. Genes and Development. 2005;19(5):517–529. [PubMed] 16. Zamore PD, Haley B. Ribo-gnome: the big world of small RNAs. Science. 2005;309(5740):1519–1524. [PubMed] 17. Voinnet O. Induction and suppression of RNA silencing: insights from
viral infections. Nature Reviews Genetics. 2005;6(3):206–220. 18. Aravin AA, Naumova NM, Tulin AV, Vagin VV, Rozovsky YM, Gvozdev VA. Double-stranded RNA-mediated silencing of genomic tandem
repeats and transposable elements in the D. melanogaster
germline. Current Biology. 2001;11(13):1017–1027. [PubMed] 19. Bernstein E, Caudy AA, Hammond SM, Hannon GJ. Role for a bidentate ribonuclease in the initiation step of
RNA interference. Nature. 2001;409(6818):363–366. [PubMed] 20. Tabara H, Yigit E, Siomi H, Mello CC. The dsRNA binding protein RDE-4 interacts with RDE-1,
DCR-1, and a DExH-Box helicase to direct RNAi in C. elegans. Cell. 2002;109(7):861–871. [PubMed] 21. Elbashir SM, Lendeckel W, Tuschl T. RNA interference is mediated by 21- and 22-nucleotide RNAs. Genes and Development. 2001;15(2):188–200. [PubMed] 22. Reinhart BJ, Slack FJ, Basson M, et al. The 21-nucleotide let-7 RNA regulates developmental
timing in Caenorhabditis elegans. Nature. 2000;403(6772):901–906. [PubMed] 23. Chen C-Z, Li L, Lodish HF, Bartel DP. MicroRNAs modulate hematopoietic lineage differentiation. Science. 2004;303(5654):83–86. [PubMed] 24. Brennecke J, Hipfner DR, Stark A, Russell RB, Cohen SM. bantam encodes a developmentally regulated microRNA that
controls cell proliferation and regulates the proapoptotic
gene hid in Drosophila. Cell. 2003;113(1):25–36. [PubMed] 25. Cai X, Hagedorn CH, Cullen BR. Human microRNAs are processed from capped, polyadenylated
transcripts that can also function as mRNAs. RNA. 2004;10(12):1957–1966. [PubMed] 26. Lee Y, Kim M, Han J, et al. MicroRNA genes are transcribed by RNA polymerase II. EMBO Journal. 2004;23(20):4051–4060. [PubMed] 27. Kim VN. MicroRNA biogenesis: coordinated cropping and dicing. Nature Reviews. Molecular Cell Biology. 2005;6(5):376–385. 28. Smalheiser NR. EST analyses predict the existence of a population of
chimeric microRNA precursor-mRNA transcripts expressed
in normal human and mouse tissues. Genome Biology. 2003;4(7):403. [PubMed] 29. Fazi F, Rosa A, Fatica A, et al. A minicircuitry comprised of microRNA-223 and transcription
factors NFI-A and C/EBPα regulates human granulopoiesis. Cell. 2005;123(5):819–831. [PubMed] 30. O'Donnell KA, Wentzel EA, Zeller KI, Dang CV, Mendell JT. c-Myc-regulated microRNAs modulate E2F1 expression. Nature. 2005;435(7043):839–843. [PubMed] 31. Zhao Y, Samal E, Srivastava D. Serum response factor regulates a muscle-specific microRNA
that targets Hand2 during cardiogenesis. Nature. 2005;436(7048):214–220. [PubMed] 32. Denli AM, Tops BBJ, Plasterk RHA, Ketting RF, Hannon GJ. Processing of primary microRNAs by the Microprocessor complex. Nature. 2004;432(7014):231–235. [PubMed] 33. Gregory RI, Yan K-P, Amuthan G, et al. The Microprocessor complex mediates the genesis of microRNAs. Nature. 2004;432(7014):235–240. [PubMed] 34. Han J, Lee Y, Yeom K-H, Kim Y-K, Jin H, Kim VN. The Drosha-DGCR8 complex in primary microRNA processing. Genes and Development. 2004;18(24):3016–3027. [PubMed] 35. Lee Y, Ahn C, Han J, et al. The nuclear RNase III Drosha initiates microRNA processing. Nature. 2003;425(6956):415–419. [PubMed] 36. Zeng Y, Yi R, Cullen BR. Recognition and cleavage of primary microRNA precursors
by the nuclear processing enzyme Drosha. EMBO Journal. 2005;24(1):138–148. [PubMed] 37. Bohnsack MT, Czaplinski K, Görlich D. Exportin 5 is a RanGTP-dependent dsRNA-binding protein that
mediates nuclear export of pre-miRNAs. RNA. 2004;10(2):185–191. [PubMed] 38. Lund E, Güttinger S, Calado A, Dahlberg JE, Kutay U. Nuclear export of microRNA precursors. Science. 2004;303(5654):95–98. [PubMed] 39. Park MY, Wu G, Gonzalez-Sulser A, Vaucheret H, Poethig RS. Nuclear processing and export of microRNAs in Arabidopsis. Proceedings of the National Academy of Sciences of the
United States of America. 2005;102(10):3691–3696. [PubMed] 40. Yi R, Qin Y, Macara IG, Cullen BR. Exportin-5 mediates the nuclear export of pre-microRNAs and
short hairpin RNAs. Genes and Development. 2003;17(24):3011–3016. [PubMed] 41. Hutvägner G, McLachlan J, Pasquinelli AE, Bälint E, Tuschl T, Zamore PD. A cellular function for the RNA-interference enzyme Dicer
in the maturation of the let-7 small temporal RNA. Science. 2001;293(5531):834–838. [PubMed] 42. Ketting RF, Fischer SEJ, Bernstein E, Sijen T, Hannon GJ, Plasterk RHA. Dicer functions in RNA interference and in synthesis of
small RNA involved in developmental timing in C. elegans. Genes and Development. 2001;15(20):2654–2659. [PubMed] 43. Knight SW, Bass BL. A role for the RNase III enzyme DCR-1 in RNA interference
and germ line development in Caenorhabditis elegans. Science. 2001;293(5538):2269–2271. [PubMed] 44. Khvorova A, Reynolds A, Jayasena SD. Functional siRNAs and miRNAs exhibit strand bias. Cell. 2003;115(2):209–216. [PubMed] 45. Schwarz DS, Hutvágner G, Du T, Xu Z, Aronin N, Zamore PD. Asymmetry in the assembly of the RNAi enzyme complex. Cell. 2003;115(2):199–208. [PubMed] 46. Hutvágner G, Zamore PD. A microRNA in a multiple-turnover RNAi enzyme complex. Science. 2002;297(5589):2056–2060. [PubMed] 47. Maniataki E, Mourelatos Z. A human, ATP-independent, RISC assembly machine fueled by
pre-miRNA. Genes and Development. 2005;19(24):2979–2990. [PubMed] 48. Liu J, Carmell MA, Rivas FV, et al. Argonaute2 is the catalytic engine of mammalian RNAi. Science. 2004;305(5689):1437–1441. [PubMed] 49. Meister G, Landthaler M, Patkaniowska A, Dorsett Y, Teng G, Tuschl T. Human Argonaute2 mediates RNA cleavage targeted by miRNAs
and siRNAs. Molecular Cell. 2004;15(2):185–197. [PubMed] 50. Ma JB, Ye K, Patel DJ. Structural basis for overhang-specific small interfering
RNA recognition by the PAZ domain. Nature. 2004;429(6989):318–322. [PubMed] 51. Song JJ, Smith SK, Hannon GJ, Joshua-Tor L. Crystal structure of argonaute and its implications for
RISC slicer activity. Science. 2004;305(5689):1434–1437. [PubMed] 52. Rhoades MW, Reinhart BJ, Lim LP, Burge CB, Bartel B, Bartel DP. Prediction of plant microRNA targets. Cell. 2002;110(4):513–520. [PubMed] 53. Kim J, Krichevsky A, Grad Y, et al. Identification of many microRNAs that copurify with
polyribosomes in mammalian neurons. Proceedings of the National Academy of Sciences of
the United States of America. 2004;101(1):360–365. [PubMed] 54. Doench JG, Petersen CP, Sharp PA. siRNAs can function as miRNAs. Genes and Development. 2003;17(4):438–442. [PubMed] 55. Llave C, Xie Z, Kasschau KD, Carrington JC. Cleavage of Scarecrow-like mRNA targets directed by a
class of Arabidopsis miRNA. Science. 2002;297(5589):2053–2056. [PubMed] 56. Mansfield JH, Harfe BD, Nissen R, et al. MicroRNA-responsive ‘sensor’ transgenes uncover
Hox-like and other developmentally regulated patterns of vertebrate
microRNA expression. Nature Genetics. 2004;36(10):1079–1083. [PubMed] 57. Yekta S, Shih IH, Bartel DP. MicroRNA-directed cleavage of HOXB8 mRNA. Science. 2004;304(5670):594–596. [PubMed] 58. Zeng Y, Yi R, Cullen BR. MicroRNAs and small interfering RNAs can inhibit mRNA
expression by similar mechanisms. Proceedings of the National Academy of Sciences of
the United States of America. 2003;100(17):9779–9784. [PubMed] 59. Mette MF, Aufsatz W, Van der Winden J, Matzke MA, Matzke AJM. Transcriptional silencing and promoter methylation triggered
by double-stranded RNA. EMBO Journal. 2000;19(19):5194–5201. [PubMed] 60. Lippman Z, Martienssen R. The role of RNA interference in heterochromatic silencing. Nature. 2004;431(7006):364–370. [PubMed] 61. Morris KV, Chan SWL, Jacobsen SE, Looney DJ. Small interfering RNA-induced transcriptional gene silencing
in human cells. Science. 2004;305(5688):1289–1292. [PubMed] 62. Bernstein E, Allis CD. RNA meets chromatin. Genes and Development. 2005;19(14):1635–1655. [PubMed] 63. Matzke MA, Birchler JA. RNAi-mediated pathways in the nucleus. Nature Reviews Genetics. 2005;6(1):24–35. 64. Dalmay T, Hamilton A, Rudd S, Angell S, Baulcombe DC. An RNA-dependent RNA polymerase gene in arabidopsis is
required for posttranscriptional gene silencing mediated
by a transgene but not by a virus. Cell. 2000;101(5):543–553. [PubMed] 65. Sijen T, Fleenor J, Simmer F, et al. On the role of RNA amplification in dsRNA-triggered gene
silencing. Cell. 2001;107(4):465–476. [PubMed] 66. Volpe TA, Kidner C, Hall IM, Teng G, Grewal SIS, Martienssen RA. Regulation of heterochromatic silencing and histone
H3 lysine-9 methylation by RNAi. Science. 2002;297(5588):1833–1837. [PubMed] 67. Stark GR, Kerr IM, Williams BRG, Silverman RH, Schreiber RD. How cells respond to interferons. Annual Review of Biochemistry. 1998;67:227–264. 68. Elbashir SM, Harborth J, Lendeckel W, Yalcin A, Weber K, Tuschl T. Duplexes of 21-nucleotide RNAs mediate RNA interference
in cultured mammalian cells. Nature. 2001;411(6836):494–498. [PubMed] 69. Caplen NJ, Parrish S, Imani F, Fire A, Morgan RA. Specific inhibition of gene expression by small
double-stranded RNAs in invertebrate and vertebrate systems. Proceedings of the National Academy of Sciences of the
United States of America. 2001;98(17):9742–9747. [PubMed] 70. Kim DH, Behlke MA, Rose SD, Chang MS, Choi S, Rossi JJ. Synthetic dsRNA Dicer substrates enhance RNAi potency and efficacy. Nature Biotechnology. 2005;23(2):222–226. 71. Rose SD, Kim DH, Amarzguioui M, et al. Functional polarity is introduced by Dicer processing of
short substrate RNAs. Nucleic Acids Research. 2005;33(13):4140–4156. [PubMed] 72. Siolas D, Lerner C, Burchard J, et al. Synthetic shRNAs as potent RNAi triggers. Nature Biotechnology. 2005;23(2):227–231. 73. Lee YS, Nakahara K, Pham JW, et al. Distinct roles for Drosophila Dicer-1 and Dicer-2 in the
siRNA/miRNA silencing pathways. Cell. 2004;117(1):69–81. [PubMed] 74. Reynolds A, Leake D, Boese Q, Scaringe S, Marshall WS, Khvorova A. Rational siRNA design for RNA interference. Nature Biotechnology. 2004;22(3):326–330. 75. Amarzguioui M, Holen T, Babaie E, Prydz H. Tolerance for mutations and chemical modifications in a
siRNA. Nucleic Acids Research. 2003;31(2):589–595. [PubMed] 76. Chi JT, Chang HY, Wang NN, Chang DS, Dunphy N, Brown PO. Genomewide view of gene silencing by small interfering RNAs. Proceedings of the National Academy of Sciences of
the United States of America. 2003;100(11):6343–6346. [PubMed] 77. Elbashir SM, Martinez J, Patkaniowska A, Lendeckel W, Tuschl T. Functional anatomy of siRNAs for mediating efficient RNAi
in Drosophila melanogaster embryo lysate. EMBO Journal. 2001;20(23):6877–6888. [PubMed] 78. Holen T, Amarzguioui M, Wiiger MT, Babaie E, Prydz H. Positional effects of short interfering RNAs targeting
the human coagulation trigger Tissue Factor. Nucleic Acids Research. 2002;30(8):1757–1766. [PubMed] 79. Jackson AL, Bartz SR, Schelter J, et al. Expression profiling reveals off-target gene regulation by RNAi. Nature Biotechnology. 2003;21(6):635–637. 80. Semizarov D, Frost L, Sarthy A, Kroeger P, Halbert DN, Fesik SW. Specificity of short interfering RNA determined through
gene expression signatures. Proceedings of the National Academy of Sciences of
the United States of America. 2003;100(11):6347–6352. [PubMed] 81. Zeng Y, Cullen BR. Sequence requirements for micro RNA processing and
function in human cells. RNA. 2003;9(1):112–123. [PubMed] 82. Doench JG, Sharp PA. Specificity of microRNA target selection in translational
repression. Genes and Development. 2004;18(5):504–511. [PubMed] 83. Saxena S, Jonsson ZO, Dutta A. Small RNAs with imperfect match to endogenous mRNA repress
translation. Implications for off-target activity of small
inhibitory RNA in mammalian cells. Journal of Biological Chemistry. 2003;278(45):44312–44319. [PubMed] 84. Huppi K, Martin SE, Caplen NJ. Defining and assaying RNAi in mammalian cells. Molecular Cell. 2005;17(1):1–10. [PubMed] 85. Holen T, Moe SE, Sorbo JG, Meza TJ, Ottersen OP, Klungland A. Tolerated wobble mutations in siRNAs decrease specificity,
but can enhance activity in vivo. Nucleic Acids Research. 2005;33(15):4704–4710. [PubMed] 86. Kretschmer-Kazemi Far R, Sczakiel G. The activity of siRNA in mammalian cells is related to
structural target accessibility: a comparison with antisense
oligonucleotides. Nucleic Acids Research. 2003;31(15):4417–4424. [PubMed] 87. Braasch DA, Jensen S, Liu Y, et al. RNA interference in mammalian cells by chemically-modified
RNA. Biochemistry. 2003;42(26):7967–7975. [PubMed] 88. Chiu Y-L, Rana TM. siRNA function in RNAi: a chemical modification analysis. RNA. 2003;9(9):1034–1048. [PubMed] 89. Donze O, Picard D. RNA interference in mammalian cells using siRNAs synthesized
with T7 RNA polymerase. Nucleic Acids Research. 2002;30(10):e46. [PubMed] 90. Sohail M, Doran G, Riedemann J, Macaulay V, Southern EM. A simple and cost-effective method for producing small
interfering RNAs with high efficacy. Nucleic Acids Research. 2003;31(7):e38. [PubMed] 91. Myers JW, Ferrell JE. Silencing gene expression with Dicergenerated siRNA pools. Methods in Molecular Biology. 2005;309:93–196. [PubMed] 92. Zhang H, Kolb FA, Brondani V, Billy E, Filipowicz W. Human Dicer preferentially cleaves dsRNAs at their termini
without a requirement for ATP. EMBO Journal. 2002;21(21):5875–5885. [PubMed] 93. Brummelkamp TR, Bernards R, Agami R. A system for stable expression of short interfering RNAs
in mammalian cells. Science. 2002;296(5567):550–553. [PubMed] 94. Sui G, Soohoo C, Affar EB, et al. A DNA vector-based RNAi technology to suppress gene
expression in mammalian cells. Proceedings of the National Academy of Sciences
of the United States of America. 2002;99(8):5515–5520. [PubMed] 95. Lee NS, Dohjima T, Bauer G, et al. Expression of small interfering RNAs targeted against
HIV-1 rev transcripts in human cells. Nature Biotechnology. 2002;20(5):500–505. 96. Yu J-Y, DeRuiter SL, Turner DL. RNA interference by expression of short-interfering RNAs
and hairpin RNAs in mammalian cells. Proceedings of the National Academy of Sciences of
the United States of America. 2002;99(9):6047–6052. [PubMed] 97. Chen M, Zhang L, Zhang H-Y, et al. A universal plasmid library encoding all permutations of
small interfering RNA. Proceedings of the National Academy of Sciences
of the United States of America. 2005;102(7):2356–2361. [PubMed] 98. Zheng L, Liu J, Batalov S, et al. An approach to genomewide screens of expressed small
interfering RNAs in mammalian cells. Proceedings of the National Academy of Sciences of
the United States of America. 2004;101(1):135–140. [PubMed] 99. Paddison PJ, Caudy AA, Bernstein E, Hannon GJ, Conklin DS. Short hairpin RNAs (shRNAs) induce sequence-specific
silencing in mammalian cells. Genes and Development. 2002;16(8):948–958. [PubMed] 100. Paddison PJ, Caudy AA, Sachidanandam R, Hannon GJ. Short hairpin activated gene silencing in mammalian cells. Methods in Molecular Biology. 2004;265:85–100. [PubMed] 101. Zeng Y, Cullen BR. Efficient processing of primary microRNA hairpins by Drosha
requires flanking nonstructured RNA sequences. Journal of Biological Chemistry. 2005;280(30):27595–27603. [PubMed] 102. Matsukura S, Jones PA, Takai D. Establishment of conditional vectors for hairpin siRNA
knockdowns. Nucleic Acids Research. 2003;31(15):e77. [PubMed] 103. van de Wetering M, Oving I, Muncan V, et al. Specific inhibition of gene expression using a stably
integrated, inducible small-interfering-RNA vector. EMBO Reports. 2003;4(6):609–615. [PubMed] 104. Gupta S, Schoer RA, Egan JE, Hannon GJ, Mittal V. Inducible, reversible, and stable RNA interference in
mammalian cells. Proceedings of the National Academy of Sciences of
the United States of America. 2004;101(7):1927–1932. [PubMed] 105. Scherr M, Battmer K, Ganser A, Eder M. Modulation of gene expression by lentiviral-mediated
delivery of small interfering RNA. Cell Cycle. 2003;2(3):251–257. [PubMed] 106. Tiscornia G, Singer O, Ikawa M, Verma IM. A general method for gene knockdown in mice by using
lentiviral vectors expressing small interfering RNA. Proceedings of the National Academy of Sciences of
the United States of America. 2003;100(4):1844–1848. [PubMed] 107. Kunath T, Gish G, Lickert H, Jones N, Pawson T, Rossant J. Transgenic RNA interference in ES cell-derived embryos
recapitulates a genetic null phenotype. Nature Biotechnology. 2003;21(5):559–561. 108. Fritsch L, Martinez LA, Sekhri R, et al. Conditional gene knock-down by CRE-dependent short
interfering RNAs. EMBO Reports. 2004;5(2):178–182. [PubMed] 109. Soutschek J, Akinc A, Bramlage B, et al. Therapeutic silencing of an endogenous gene by systemic
administration of modified siRNAs. Nature. 2004;432(7014):173–178. [PubMed] 110. Morrissey DV, Lockridge JA, Shaw L, et al. Potent and persistent in vivo anti-HBV activity of
chemically modified siRNAs. Nature Biotechnology. 2005;23(8):1002–1007. 111. Song E, Zhu P, Lee S-K, et al. Antibody mediated in vivo delivery of small interfering
RNAs via cell-surface receptors. Nature Biotechnology. 2005;23(6):709–717. 112. Case SS, Price MA, Jordan CT, et al. Stable transduction of quiescent
CD34+CD38− human hematopoietic
cells by HIV-1-based lentiviral vectors. Proceedings of the National Academy of Sciences of
the United States of America. 1999;96(6):2988–2993. [PubMed] 113. Luther-Wyrsch A, Costello E, Thali M, et al. Stable transduction with lentiviral vectors and
amplification of immature hematopoietic progenitors from
cord blood of preterm human fetuses. Human Gene Therapy. 2001;12(4):377–389. [PubMed] 114. Uchida N, Sutton RE, Friera AM, et al. HIV, but not murine leukemia virus, vectors mediate high
efficiency gene transfer into freshly isolated G0/G1
human hematopoietic stem cells. Proceedings of the National Academy of Sciences
of the United States of America. 1998;95(20):11939–11944. [PubMed] 115. Scherr M, Battmer K, Dallmann I, Ganser A, Eder M. Inhibition of GM-CSF receptor function by stable RNA
interference in a NOD/SCID mouse hematopoietic
stem cell transplantation model. Oligonucleotides. 2003;13(5):353–363. [PubMed] 116. Schomber T, Kalberer CP, Wodnar-Filipowicz A, Skoda RC. Gene silencing by lentivirus-mediated delivery of siRNA
in human CD34+ cells. Blood. 2004;103(12):4511–4513. [PubMed] 117. Wilda M, Fuchs U, Wossmann W, Borkhardt A. Killing of leukemic cells with a BCR/ABL fusion gene by
RNA interference (RNAi). Oncogene. 2002;21(37):5716–5724. [PubMed] 118. Scherr M, Battmer K, Winkler T, Heidenreich O, Ganser A, Eder M. Specific inhibition of bcr-abl gene expression by small
interfering RNA. Blood. 2003;101(4):1566–1569. [PubMed] 119. Wohlbold L, Van der Kuip H, Miething C, et al. Inhibition of bcr-abl gene expression by small interfering
RNA sensitizes for imatinib mesylate (STI571). Blood. 2003;102(6):2236–2239. [PubMed] 120. Scherr M, Battmer K, Schultheis B, Ganser A, Eder M. Stable RNA interference (RNAi) as an option for anti-bcr-abl
therapy. Gene Therapy. 2005;12(1):12–21. [PubMed] 121. Wohlbold L, van der Kuip H, Moehring A, Granot G, Oren M, Vornlocher H-P, Aulitzky WE. All common p210 and p190 Bcr-abl variants can be targeted
by RNA interference [3]. Leukemia. 2005;19(2):290–292. [PubMed] 122. Rangatia J, Bonnet D. Transient or long-term silencing of BCR-ABL alone induces
cell cycle and proliferation arrest, apoptosis and
differentiation. Leukemia. 2006;20(1):68–76. [PubMed] 123. Heidenreich O, Krauter J, Riehle H, et al. AML1/MTG8 oncogene suppression by small interfering RNAs
supports myeloid differentiation of t(8;21)-positive leukemic
cells. Blood. 2003;101(8):3157–3163. [PubMed] 124. Chen J, Wall NR, Kocher K, et al. Stable expression of small interfering RNA sensitizes
TEL-PDGFβR to inhibition with imatinib or rapamycin. Journal of Clinical Investigation. 2004;113(12):1784–1791. [PubMed] 125. Thomas M, Geßner A, Vornlocher H-P, Hadwiger P, Greil J, Heidenreich O. Targeting MLL-AF4 with short interfering RNAs inhibits
clonogenicity and engraftment of t(4;11)-positive human
leukemic cells. Blood. 2005;106(10):3559–3566. [PubMed] 126. Ritter U, Damm-Welk C, Fuchs U, Bohle RM, Borkhardt A, Woessmann W. Design and evaluation of chemically synthesized siRNA
targeting the NPM-ALK fusion site in anaplastic large cell
lymphoma (ALCL). Oligonucleotides. 2003;13(5):365–373. [PubMed] 127. Piva R, Chiarle R, Manazza AD, et al. Ablation of oncogenic ALK is a viable therapeutic approach
for anaplastic large-cell lymphomas. Blood. 2006;107(2):689–697. [PubMed] 128. Walters DK, Stoffregen EP, Heinrich MC, Deininger MW, Druker BJ. RNAi-induced down-regulation of FLT3 expression in AML
cell lines increases sensitivity to MLN518. Blood. 2005;105(7):2952–2954. [PubMed] 129. Daley GQ, Van Etten RA, Baltimore D. Induction of chronic myelogenous leukemia in mice by the
P210(bcr/abl) gene of the Philadelphia chromosome. Science. 1990;247(4944):824–830. [PubMed] 130. Huettner CS, Zhang P, Van Etten RA, Tenen DG. Reversibility of acute B-cell leukaemia induced by BCR-ABL1. Nature Genetics. 2000;24(1):57–60. [PubMed] 131. Lugo TG, Pendergast A-M, Muller AJ, Witte ON. Tyrosine kinase activity and transformation potency of
bcr-abl oncogene products. Science. 1990;247(4946):1079–1082. [PubMed] 132. Kantarjian H, Sawyers C, Hochhaus A, et al. International STI571 CML Study Group. Hematologic and
cytogenetic responses to imatinib mesylate in chronic
myelogenous leukemia. New England Journal of Medicine. 2002;346(9):645–652. [PubMed] 133. Druker BJ, Sawyers CL, Kantarjian H, et al. Activity of a specific inhibitor of the BCR-ABL tyrosine
kinase in the blast crisis of chronic myeloid leukemia
and acute lymphoblastic leukemia with the Philadelphia
chromosome. New England Journal of Medicine. 2001;344(14):1038–1042. Erratum in: New England Journal of Medicine.
2001;345(3):232. [PubMed] 134. Miyoshi H, Kozu T, Shimizu K, et al. The t(8;21) translocation in acute myeloid leukemia
results in production of an AML1-MTG8 fusion transcript. EMBO Journal. 1993;12(7):2715–2721. [PubMed] 135. Golub TR, Barker GF, Lovett M, Gilliland DG. Fusion of PDGF receptor β to a novel ets-like gene,
tel, in chronic myelomonocytic leukemia with t(5;12)
chromosomal translocation. Cell. 1994;77(2):307–316. [PubMed] 136. Ernst P, Wang J, Korsmeyer SJ. The role of MLL in hematopoiesis and leukemia. Current Opinion in Hematology. 2002;9(4):282–287. [PubMed] 137. Domer PH, Fakharzadeh SS, Chen C-S, et al. Acute mixed-lineage leukemia t(4;11)(q21;q23) generates an
MLL-AF4 fusion product. Proceedings of the National Academy of Sciences of
the United States of America. 1993;90(16):7884–7888. [PubMed] 138. Gu Y, Nakamura T, Alder H, et al. The t(4;11) chromosome translocation of human acute
leukemias fuses the ALL-1 gene, related to Drosophila
trithorax, to the AF-4 gene. Cell. 1992;71(4):701–708. [PubMed] 139. McCabe NR, Burnett RC, Gill HJ, et al. Cloning of cDNAs of the MLL gene that detect DNA
rearrangements and altered RNA transcripts in human
leukemic cells with 11q23 translocations. Proceedings of the National Academy of Sciences of
the United States of America. 1992;89(24):11794–11798. [PubMed] 140. Pui C-H, Gaynon PS, Boyett JM, et al. Outcome of treatment in childhood acute lymphoblastic
leukaemia with rearrangements of the 11q23 chromosomal
region. Lancet. 2002;359(9321):1909–1915. [PubMed] 141. Morris SW, Kirstein MN, Valentine MB, et al. Fusion of a kinase gene, ALK, to a nucleolar protein gene,
NPM, in non-Hodgkin's lymphoma. Science. 1994;263(5151):1281–1284. [PubMed] 142. Yamamoto Y, Kiyoi H, Nakano Y, et al. Activating mutation of D835 within the activation loop of
FLT3 in human hematologic malignancies. Blood. 2001;97(8):2434–2439. [PubMed] 143. Yokota S, Kiyoi H, Nakao M, et al. Internal tandem duplication of the FLT3 gene is
preferentially seen in acute myeloid leukemia and
myelodysplastic syndrome among various hematological
malignancies. A study on a large series of patients and
cell lines. Leukemia. 1997;11(10):1605–1609. [PubMed] 144. Felli N, Fontana L, Pelosi E, et al. MicroRNAs 221 and 222 inhibit normal erythropoiesis and
erythroleukemic cell growth via kit receptor down-modulation. Proceedings of the National Academy of Sciences of the
United States of America. 2005;102(50):18081–18086. [PubMed] 145. Radomska HS, Huettner CS, Zhang P, Cheng T, Scadden DT, Tenen DG. CCAAT/enhancer binding protein α is a regulatory
switch sufficient for induction of granulocytic development
from bipotential myeloid progenitors. Molecular and Cellular Biology. 1998;18(7):4301–4314. [PubMed] 146. Tenen DG. Disruption of differentiation in human cancer: AML shows
the way. Nature Reviews Cancer. 2003;3(2):89–101. 147. Wang X, Scott E, Sawyers CL, Friedman AD. C/EBPα bypasses granulocyte colony-stimulating
factor signals to rapidly induce PU.1 gene expression,
stimulate granulocytic differentiation, and limit
proliferation in 32D cl3 myeloblasts. Blood. 1999;94(2):560–571. [PubMed] 148. Gronostajski RM. Roles of the NFI/CTF gene family in transcription and
development. Gene. 2000;249(1-2):31–45. [PubMed] 149. Santoro C, Mermod N, Andrews PC, Tjian R. A family of human CCAAT-box-binding proteins active in
transcription and DNA replication: cloning and expression
of multiple cDNAs. Nature. 1988;334(6179):218–224. [PubMed] 150. Lewis BP, Shih I-H, Jones-Rhoades MW, Bartel DP, Burge CB. Prediction of mammalian microRNA targets. Cell. 2003;115(7):787–798. [PubMed] 151. Calin GA, Sevignani C, Dumitru CD, et al. Human microRNA genes are frequently located at fragile
sites and genomic regions involved in cancers. Proceedings of the National Academy of Sciences of the
United States of America. 2004;101(9):2999–3004. [PubMed] 152. Metzler M, Strissel PL, Strick R, et al. Emergence of translocation t(9;11)-positive leukemia
during treatment of childhood acute lymphoblastic leukemia. Genes Chromosomes and Cancer. 2004;41(3):291–296. [PubMed] 153. Calin GA, Dumitru CD, Shimizu M, et al. Frequent deletions and down-regulation of micro-RNA genes
miR15 and miR16 at 13q14 in chronic lymphocytic leukemia. Proceedings of the National Academy of Sciences of the
United States of America. 2002;99(24):15524–15529. [PubMed] 154. Cimmino A, Calin GA, Fabbri M, et al. miR-15 and miR-16 induce apoptosis by targeting BCL2. Proceedings of the National Academy of Sciences of the
United States of America. 2005;102(39):13944–13949. [PubMed] 155. Ota A, Tagawa H, Karnan S, et al. Identification and characterization of a novel gene,
C13orf25, as a target for 13q31-q32 amplification in
malignant lymphoma. Cancer Research. 2004;64(9):3087–3095. [PubMed] 156. He L, Thomson JM, Hemann MT, et al. A microRNA polycistron as a potential human oncogene. Nature. 2005;435(7043):828–833. [PubMed] 157. Lu J, Getz G, Miska EA, et al. MicroRNA expression profiles classify human cancers. Nature. 2005;435(7043):834–838. [PubMed] 158. Calin GA, Liu C-G, Sevignani C, et al. MicroRNA profiling reveals distinct signatures in B cell
chronic lymphocytic leukemias. Proceedings of the National Academy of Sciences
of the United States of America. 2004;101(32):11755–11760. [PubMed] 159. Calin GA, Ferracin M, Cimmino A, et al. A microRNA signature associated with prognosis and
progression in chronic lymphocytic leukemia. New England Journal of Medicin. 2005;353(17):1793–1801. 160. Berns K, Hijmans EM, Mullenders J, et al. A large-scale RNAi screen in human cells identifies new
components of the p53 pathway. Nature. 2004;428(6981):431–437. [PubMed] 161. Futami T, Miyagishi M, Taira K. Identification of a network involved in thapsigargin-induced
apoptosis using a library of small interfering RNA expression
vectors. Journal of Biological Chemistry. 2005;280(1):826–831. [PubMed] 162. Kittler R, Putz G, Pelletier L, et al. An endoribonuclease-prepared siRNA screen in human cells
identifies genes essential for cell division. Nature. 2004;432(7020):1036–1040. [PubMed] 163. Luo B, Heard AD, Lodish HF. Small interfering RNA production by enzymatic engineering
of DNA (SPEED). Proceedings of the National Academy of Sciences of
the United States of America. 2004;101(15):5494–5499. [PubMed] 164. Matsumoto S, Miyagishi M, Akashi H, Nagai R, Taira K. Analysis of double-stranded RNA-induced apoptosis pathways
using interferon-response noninducible small interfering
RNA expression vector library. Journal of Biological Chemistry. 2005;280(27):25687–25696. [PubMed] 165. Paddison PJ, Silva JM, Conklin DS, et al. A resource for large-scale RNA-interference-based screens
in mammals. Nature. 2004;428(6981):427–431. [PubMed] 166. Sen G, Wehrman TS, Myers JW, Blau HM. Restriction enzyme-generated siRNA (REGS) vectors and
libraries. Nature Genetics. 2004;36(2):183–189. [PubMed] 167. Shirane D, Sugao K, Namiki S, Tanabe M, Iino M, Hirose K. Enzymatic production of RNAi libraries from cDNAs. Nature Genetics. 2004;36(2):190–196. [PubMed]
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