In this chapter, we provide a brief overview of the RNA interference (RNAi)
process and discuss technologies and products that can be used for RNAi experiments.
A protocol is provided for the enzymatic synthesis of double-stranded RNA in vitro
that provides an inexpensive alternative to chemical synthesis of RNAs. We also
discuss design and selection of short interfering RNA (siRNA) sequences and describe
a vector system, the psiCHECK™ Vectors, that can be used to screen potential siRNA
target sequences for effectiveness during RNAi optimization. In addition, the
GeneClip™ U1 Hairpin Cloning Systems, a DNA-directed RNAi (ddRNAi) system
specifically designed for expression of small hairpin RNAs in mammalian cells,
provide a cloning-based approach to allow fast, easy ligation and expression of
hairpin oligonucleotides. Various strategies for delivery of siRNA to target cells
are discussed, and example protocols for transient and stable transfection of
mammalian cells are provided. Finally, methods for quantitating target gene
suppression are briefly summarized.
RNA interference (RNAi) is a phenomenon in which double-stranded RNA (dsRNA)
suppresses expression of a target protein by stimulating the specific degradation of
the target mRNA (for reviews, see Hannon, 2003; Caplen, 2004; Fuchs et
al. 2004; Betz, 2003a). RNAi has been used to study loss of function for a
variety of genes in several organisms including various plants,
Caenorhabditis elegans and Drosophila,
and permits loss-of-function genetic screens and rapid tests for genetic interactions
in mammalian cells (Hannon, 2002; Williams et al. 2003).
RNAi involves a multistep process (Figure 2.1). dsRNA is recognized by an RNase
III family member (e.g., Dicer in Drosophila) and cleaved into
siRNAs of 21–23 nucleotides (Agrawal et al. 2003; Elbashir
et al. 2001b; Bernstein et al. 2001;
Hammond et al. 2000). These siRNAs are incorporated into an RNAi targeting complex
known as RISC (RNA-induced silencing complex), which destroys mRNAs homologous to the
integral siRNA (Hammond et al. 2000; Bernstein et
al. 2001). The target mRNA is cleaved in the center of the region
complementary to the siRNA (Elbashir et al. 2001c), with the net
result being rapid degradation of the target mRNA and decreased protein expression.
RNAi has revolutionized the study of gene function, and is being explored as a
therapeutic tool (for reviews, see Dorsett and Tuschl, 2004; Hannon and Rossi, 2004).
For example, RNAi has been used to identify gene products essential for cell growth
(Harborth et al. 2001), to cause subtype and species-specific
knockdown of various protein kinase C (PKC) isoforms in both human and rat cells
(Irie et al. 2002), and to specifically target degradation of an
oncogene product (Wilda et al. 2002). RNAi has also been used to
specifically target and prevent viral infections by HIV-1 and HCV in cell culture
(Park et al. 2002) and intact animals (McCaffrey et
al. 2002). These observations open the field for further studies toward
novel gene therapy approaches for anti-cancer or anti-viral treatments using siRNAs
Figure 2.1. Simplified schematic diagram of the proposed RNA interference mechanism.
dsRNA processing proteins (RNase III-like enzymes) bind to and cleave
dsRNA into siRNA. The siRNA forms a multicomponent nuclease complex, the
RNA-induced silencing complex (RISC). The target mRNA recognized by RISC is
cleaved in the center of the region complementary to the siRNA and quickly
degraded. An animated version of this illustration is also available.
presentation illustrating the entire RNAi process is available on the
Nature web site.
The use of long dsRNAs (>400bp) has been successful in generating RNA
interference effects in many organisms including Drosophila
(Misquitta and Paterson, 1999), zebrafish (Wargelius et al.
1999), Planaria (Sanchez-Alvarado et al.
1999) and numerous plants (Jorgensen, 1990, Fukusaki et al.
2004, Jensen et al. 2004). In mammalian systems, siRNA molecules
of 21–22 nucleotides or short hairpin RNAs (shRNAs) are used to avoid endogenous
nonspecific antiviral responses that target longer dsRNAs (Caplen
et al. 2001, Elbashir et al. 2001a). Yu
et al. (2002) and others (Brummelkamp et
al. 2002b; McManus et al. 2002; Sui et
al. 2002; Xia et al. 2002; Barton and Medzhitov,
2002) demonstrated that shRNAs bearing a fold-back, stem-loop structure of
approximately 19 perfectly matched nucleotides connected by various spacer regions
and ending in a 2-nucleotide 3′-overhang can be as efficient as siRNAs at inducing
RNA interference. si/shRNAs can induce specific gene silencing in a wide range of
mammalian cell lines without leading to global inhibition of mRNA translation (Caplen
et al. 2001; Elbashir et al. 2001a;
Paddison et al. 2002).
Generation of Short Interfering RNAs
siRNAs are the main effectors of the RNAi process. These molecules can be
synthesized chemically or enzymatically in vitro (Micura, 2002; Betz, 2003b;
Paddison et al. 2002) or endogenously expressed inside the
cells in the form of shRNAs (Yu et al. 2002; McManus
et al. 2002). Plasmid-based expression systems using RNA
polymerase III U6 or H1, or RNA polymerase II U1, small nuclear RNA promoters,
have been used for endogenous expression of shRNAs (Brummelkamp et
al. 2002b; Sui et al. 2002, Novarino et
Rational Design of Effective siRNA Probes
Design of the siRNA sequence is crucial for effective gene silencing. Rational
design strategies for effective siRNAs are being developed based on an
understanding of RNAi biochemistry and of naturally occurring microRNA (miRNA)
function. Several groups have proposed basic empirical guidelines for designing
effective siRNAs that can be applied to the selection of potential target
sequences (Chiu and Rana, 2002; Khvorova et al. 2003;
Schwarz et al. 2003; Hseih et al.
2004; Reynolds et al. 2004; Ui-Tei et
al. 2004). In addition, strategies for experimentally screening
effective siRNAs from pools of potential siRNAs are being developed (Kumar
et al. 2003; Vidugiriene et al. 2004)
and will remain a useful tool until potent siRNAs can be predicted accurately for
each target gene.
Delivery of siRNA
The efficient delivery of siRNAs is a vital step in RNAi-based gene silencing
experiments. Synthetic siRNAs can be delivered by electroporation or by using
lipophilic agents (McManus et al. 2002; Kishida et
al. 2004). siRNAs have been used successfully to silence target
genes; however, these approaches are limited by the transient nature of the
response. The use of plasmid systems to express small hairpin RNAs helps overcomes
this limitation by allowing stable suppression of target genes (Dykxhoorn
et al. 2003). Various viral delivery systems have also
been developed to deliver shRNA-expressing cassettes into cells that are difficult
to transfect, creating new possibilities for RNAi usage (Brummelkamp et
al. 2002a; Rubinson et al. 2003). Successful
delivery of siRNAs in live animals has also been reported (Hasuwa et
al. 2002; Carmell et al. 2003; Kobayashi
et al. 2004).
return to top of page
Identifying an optimal target sequence is critical to the success of RNA
interference experiments. Since it is not possible to predict the optimal siRNA
sequence for a given target, multiple siRNAs will usually need to be evaluated.
Recommendations for the design of siRNAs are constantly being improved upon as
knowledge of the RNAi process continues to expand. At the time of writing this
chapter, the recommendations are as follows: siRNA target sequences should be
specific to the gene of interest and have ~20–50% GC content (Henshel et
al. 2004). Ui-Tei
et al. (2004) report that siRNAs satisfying the following
conditions are capable of effective gene silencing in mammalian cells: 1) G/C at the
5′ end of the sense strand; 2) A/U at the 5′ end of the antisense strand; 3)
at least 5 A/U residues in the first 7 bases of the 5′ terminal of the antisense
strand; 4) no runs of more than 9 G/C residues. Additionally, primer design rules
specific to the RNA polymerase used will apply. For example, for RNA polymerase III,
the polymerase that transcribes from the U6 promoter, the preferred target sequence
is 5′-GN18-3′. Runs of 4 or more Ts (or As on the
other strand) serve as terminator sequences for RNA polymerase III and should be
avoided. In addition, regions with a run of any single base should be avoided
(Czauderna et al. 2003). It is generally recommended that the
mRNA target site be at least 50–200 bases downstream of the start codon (Sui
et al. 2002; Elbashir et al. 2002,
Duxbury and Whang, 2004) to avoid regions in which regulatory proteins might bind.
Not all siRNAs directed against a target gene are equally effective in suppressing
expression of that target in mammalian cells. Therefore, it is important to identify
siRNA sequences that are effective inhibitors of target gene expression. Although
rational designs for selection of potential target sequences have been encouraging in
generating effective siRNAs, accurate prediction of the most effective siRNAs still
remains to be achieved. Current screening technologies are based on semiquantitative,
time-consuming methods and are not easily modified to perform rapid, simultaneous
screening of multiple siRNA/shRNA sequences. However, as the field of RNAi advances,
and more high-throughput applications are adopted, there is a growing need for rapid,
quantitative screening to confirm siRNA effectiveness (Kumar et
al. 2003; Mousses et al. 2003).
Recently, several quantifiable procedures that use reporter genes to help rapidly
identify effective siRNAs have been developed. In these approaches, the change in
expression of a reporter gene fused to a target gene is used as an indicator of the
effectiveness of an RNAi methodology. Here, we describe the psiCHECK™ Vector system,
which is based on use of the bioluminescent Renilla luciferase
reporter gene. The psiCHECK™ Vectors offer several advantages compared to other
fusion approaches such as green fluorescent protein (GFP)- or Flag-tag-based methods.
Measurement of net fluorescence from GFP in cell culture can be difficult and, in
most cases, a flow cytometer is required for quantitation. Flag-tag quantitation
requires Western blot analysis, which can be time-consuming. The high sensitivity of
bioluminescence detection can readily tolerate lower expression levels, and
introduction of a second reporter gene, firefly luciferase, allows normalization of
changes in Renilla luciferase expression, making the psiCHECK™
Vector approach more robust and giving greater reproducibility of results.
The psiCHECK™-1 and -2 Vectors allow quantitative selection of optimal siRNA
target sites and can be adapted for use in high-throughput applications. Figure 2.2
provides a basic illustration of how the psiCHECK™ Vectors are used. Both vectors
contain a synthetic version of the Renilla luciferase
(hRluc) reporter gene for monitoring RNAi activity. Several
restriction sites are included 3′ of the luciferase translational stop codon,
allowing creation of transcriptional fusions between the gene of interest and the
Renilla luciferase reporter gene. Because of the presence of
a stop codon in-frame with the Renilla luciferase open reading
frame, no fusion protein is produced. Consequently, there is no need to maintain
frames when inserting the target gene. Also, toxic genes or gene fragments can be
analyzed using this design without the danger of these genes killing the transfected
The psiCHECK™-1 Vector (Cat.# C8011) is recommended
for monitoring RNAi effects in live cells. Changes in Renilla
luciferase activity can be measured with the EnduRen™ Live Cell Substrate
(Cat.# E6481). This approach permits continuous
monitoring of intracellular luminescence. Renilla luciferase
expression can be monitored continuously for 2 days without interfering with
normal cell physiology.
The psiCHECK™-2 Vector (Cat.# C8021) contains an
additional reporter gene, a synthetic firefly luciferase gene
(hluc+), and is designed for endpoint lytic assays. Inclusion of
the firefly luciferase gene permits normalization of changes in
Renilla luciferase expression to firefly luciferase expression.
Renilla and firefly luciferase activities can be measured
using either the Dual-Luciferase® Reporter Assay System
(Cat.# E1910) or the
Dual-Glo® Luciferase Assay System (Cat.#
To use the psiCHECK™ Vectors for screening siRNA targets, the gene of interest is
cloned into the multiple cloning region located 3´ to the synthetic
Renilla luciferase gene and its translational stop codon. After
cloning, the vector is transfected into a mammalian cell line, and a fusion of the
Renilla gene and the target gene is transcribed. Functional
Renilla luciferase is translated from the intact transcript.
Depending on your experimental design, vectors expressing shRNA or synthetic siRNA
can be either cotransfected simultaneously or sequentially. If a specific shRNA/siRNA
effectively initiates the RNAi process on the target RNA, the fused
Renilla target gene mRNA sequence will be degraded, resulting in
reduced Renilla luciferase activity.
Additional Resources for psiCHECK™ Vectors
Technical Bulletins and Manuals
psiCHECK™ Vectors Technical Bulletin
The use of bioluminescent reporter genes for RNAi optimization
return to top of page
siRNA synthesis in vitro provides a useful alternative to the potentially expensive
chemical synthesis of RNA (Figure 2.3). The method relies on T7 phage RNA
polymerase to produce individual sense and antisense strands that are annealed in vitro
prior to delivery into the cells of choice (Fire et al. 1998; Donze
and Picard, 2002; Yu et al. 2002, Shim et al.
The T7 RiboMAX™ Express RNAi System (Cat.# P1700) is
an in vitro transcription system designed for rapid production of milligram amounts of
double-stranded RNA (dsRNA). The system can be used to synthesize siRNAs for use in
mammalian systems (Figure 2.4; Betz, 2003b, Hwang et al. 2004) or
longer interfering RNAs for nonmammalian systems (Betz and Worzella, 2003; Betz, 2003c).
The DNA templates for in vitro transcription of siRNAs are a pair of short, duplex
oligonucleotides that contain T7 RNA polymerase promoters upstream of the sense and
antisense RNA sequences. Each oligonucleotide of the duplex is a separate template for
the synthesis of one strand of the siRNA. The separate short RNA strands that are
synthesized are then annealed to form siRNA.
Figure 2.4. Suppression of endogenous p53 protein using siRNA prepared using the T7
RiboMAX™ Express RNAi System.
Twenty-four hours after plating in a 12-well plate, 293T cells were
transfected with 200ng scrambled siRNA (lane 1), 200ng in vitro synthesized p53
siRNA (lane 2), or 200ng chemically synthesized p53 siRNA (lane 3). Twenty-four
hours after transfection, cells were lysed using 1X Reporter Lysis Buffer
(Cat.# E3971) containing protease inhibitors,
and the protein was quantitated using the BCA Protein Assay (Pierce). Equal
amounts of each lysate (10µg) were separated on a 4–12% polyacrylamide Bis-Tris
gel (Invitrogen) and transferred to Hybond®-C
membrane (Amersham). The blot was probed with both a p53 antibody (Calbiochem)
and a β-actin antibody (Abcam). Detection was performed using Goat Anti-Mouse
HRP Conjugate (Cat.# W4021) and the Transcend™
Chemiluminescent Non-Radioactive Translation Detection System
(Cat.# L5080). The blot was exposed to Kodak
X-OMAT® film for approximately 4 minutes. The
simultaneous detection of the β-actin protein controlled for loading and
transfer. The p53 and β-actin bands are indicated and are of the expected
RNAi experiments in nonmammalian systems are typically performed with dsRNA of
400bp or larger (Elbashir et al. 2001b; Yang et
al. 2000, Hammond et al. 2000). The minimum size of
dsRNA recommended for RNAi in these systems is ~200bp. In general, templates for
transcription of dsRNA for use in RNAi experiments correspond to most or all of the
target message sequence. Data suggests that longer dsRNA molecules are more effective
on a molar basis at silencing protein expression, but higher concentrations of
smaller dsRNA molecules may have similar silencing effects. Data generated at Promega
suggests that smaller dsRNAs can be as effective and efficient at inducing RNAi in
nonmammalian systems (Betz, 2003c).
In the T7 RiboMAX™ Express RNAi System, dsRNA production requires a T7 RNA
polymerase promoter at the 5′-ends of both DNA target sequence strands. To achieve
this, separate DNA templates, each containing the target sequence in a different
orientation relative to the T7 promoter, are transcribed in two separate reactions.
The resulting transcripts are mixed and annealed post-transcriptionally. DNA
templates can be created by PCR or by using two linearized plasmid templates, each
containing the T7 polymerase promoter at a different end of the target sequence.
See the T7 RiboMAX™ Express RNAi System Technical Bulletin
#TB316 for a protocol for in vitro synthesis of dsRNA for RNAi in nonmammalian
Figure 2.5 outlines the protocol for synthesis of siRNA using the T7 RiboMAX™
Express RNAi System. The initial step is generating the DNA template, which consists
of two DNA oligonucleotides annealed to form a duplex. Generally 20pmol of duplex
oligonucleotides are required per 20µl in vitro transcription reaction. Using the
RiboMAX™ Express T7 Buffer and Enzyme Mix allows efficient synthesis of RNA in as
little as 30 minutes. The annealed DNA oligonucleotide template is removed by a DNase
digestion step, and the separate small RNA strands (sense and antisense) are annealed
to form siRNA. The siRNA is precipitated using sodium acetate and isopropanol, and
the resuspended product can be analyzed on polyacrylamide gels for size and
integrity. Quantitation of the siRNA can be accomplished by either gel analysis or
RiboGreen® analysis (Molecular Probes).
- T7 RiboMAX™ Express RNAi System (Cat.#
- 2X oligo annealing buffer (20mM Tris-HCl [pH 7.5], 100mM NaCl)
- nuclease-free water
- gene-specific oligonucleotides
- 70% ethanol
Designing DNA Oligonucleotides
The target mRNA sequence selected must be screened for the sequence
5′-GN17C-3′. The generation of 3–5 different siRNA
sequences for a particular target is recommended to allow screening for the
optimal target site. The oligonucleotides consist of the target sequence plus the
T7 RNA polymerase promoter sequence and 6 extra nucleotides upstream of the
minimal promoter sequence to allow for efficient T7 RNA polymerase binding.
Details on design of oligonucleotides for use with this system are found in the
T7 RiboMAX™ Express RNAi System Technical Bulletin #TB316.
Annealing DNA Oligonucleotides
- Resuspend DNA oligonucleotides in nuclease-free water to a final
concentration of 100pmol/µl.
- Combine each pair of DNA oligonucleotides to generate either the sense
strand RNA or antisense strand RNA templates as follows:
|oligonucleotide 1 (100pmol/µl)
|oligonucleotide 2 (100pmol/µl)
|2X oligo annealing buffer
- Heat at 90–95°C for 3–5 minutes, then allow the mixture to cool slowly to
room temperature. The final concentration of annealed oligonucleotide is
10pmol/µl. Store annealed oligonucleotide DNA template at either 4°C or
Synthesizing Large Quantities of siRNA
- Set up the reaction at room temperature. The 20µl reaction may be scaled
as necessary (up to 500µl total volume; use multiple tubes for reaction
volumes >500µl). Add the components in the order shown below. For
each siRNA, two separate reactions must be assembled as each RNA strand is
synthesized separately, and then mixed following transcription.
|T7 Reaction Components
|RiboMAX™ Express 2X Buffer
|annealed oligonucleotide template DNA (10pmol/µl)
|pGEM® Express Positive Control
|Enzyme Mix, T7 Express
- Incubate for 30 minutes at 37°C.
Removing the DNA Template and Annealing siRNA
The DNA template can be removed by digestion with DNase following the
transcription reaction. RQ1 RNase-Free DNase (Cat.#
M6101) has been tested for its ability to degrade DNA while
maintaining the integrity of RNA. If accurate RNA concentration determination is
desired, the RNA should be DNase-treated and purified to remove potentially
inhibitory or interfering components.
- To each 20µl transcription reaction, add 1µl of RQ1 RNase-Free DNase and
incubate for 30 minutes at 37°C.
- Combine separate sense and antisense reactions and incubate for 10
minutes at 70°C, then allow the tubes to cool to room temperature
(approximately 20 minutes). This step anneals the separate short sense
and antisense RNA strands, generating siRNA.
- Add 0.1 volume of 3M Sodium Acetate (pH 5.2) and 1 volume of
isopropanol. Mix and place on ice for 5 minutes. The reaction will
appear cloudy. Spin at top speed in a microcentrifuge for 10
- Carefully aspirate the supernatant, and wash the pellet with 0.5ml of
cold 70% ethanol, removing all ethanol following the wash. Air-dry the
pellet for 15 minutes at room temperature, and resuspend the RNA sample in
nuclease-free water in a volume 2–5 times the original reaction volume (at
least 2 volumes are required for adequate resuspension). Store at –20°C or
Additional Resources for T7 RiboMAX™ Express RNAi System
Technical Bulletins and Manuals
T7 RiboMAX™ Express RNAi System Technical Bulletin
RNAi in Drosophila S2 cells: Effect of dsRNA size,
concentration, and exposure time
The T7 RiboMAX™ Express RNAi System: Efficient synthesis of dsRNA for
Produce functional siRNAs and hairpin siRNAs using the T7 RiboMAX™
Express RNAi System
Hwang, C.K. et al.
(2004) Transcriptional regulation of mouse µ opioid receptor gene by PU.1. J. Biol. Chem. 279
In this article, siRNA was used to reduce the level of the PU.1
transcription factor. The siRNA was generated using the T7 RiboMAX™
Express RNAi System. Annealed siRNA was purified by isopropanol
precipitation. Forty-eight hours after transfecting 2.5µg of siRNA into
RAW264.7 cells, RNA and protein were isolated from the cells, and the
siRNA effect was analyzed by RT-PCR and Western blot.
Kim, C.S. et al.
(2004) Neuron-restrictive silencer factor (NRSF) functions as a repressor in
neuronal cells to regulate µ opioid receptor gene. J. Biol. Chem. 279
In this article, siRNA was used to silence endogenous mouse and human
NRSF expression in NS20Y and HeLa cells. siRNAs were generated using the
T7 RiboMAX™ Express RNAi System.
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DNA-directed RNA interference (ddRNAi) involves the use of DNA templates to
synthesize si/shRNA in vivo. ddRNAi relies on U6 or H1 [RNA polymerase III], or U1
[RNA polymerase II]) promoters for the expression of siRNA target sequences that have
been transfected into mammalian cells (Miyagishi and Taira, 2002; Brummelkamp
et al. 2002b; Novarino et al. 2004).
si/shRNA target sequences can be generated by PCR, creating “expression cassettes”
that can be transfected directly into cells (Csiszar et al.
2004; Castanotto et al. 2002) or cloned into expression vectors
(Sui et al. 2002; Paul et al. 2002; Gou
et al. 2003; Yu et al. 2002). PCR
generation is recommended when rapid screening of numerous siRNAs is desired.
Cloning-based approaches that allow direct ligation of hairpin oligonucleotides into
a ddRNAi vector provide another method for quickly and easily screening various
targets (Yeager et al. 2005). Screening can also be performed
using synthetic RNAs, but this can become expensive for numerous targets.
Vector-based approaches also offer the potential of stable, long-term inhibition
of gene expression by providing siRNAs on plasmids that allow selection of
transfected cells. Vectors with markers, such as puromycin, neomycin or hygromycin,
can be used for suppression of target genes for several weeks or longer. Transfection
with synthetic siRNAs allows for only a transient measurement (usually 48–72 hours)
of the RNAi effect.
The success of ddRNAi depends on several parameters including generation of
vectors containing full-length sh/siRNA sequences, delivery of those vectors into
cells, and expression of the si/shRNA constructs. Most strategies for cloning siRNA
target sequences into expression vectors utilize the design of a hairpin structure.
This design consists of two inverted repeats separated by a short spacer sequence
(loop sequence). After transcription by RNA polymerase, the inverted repeats anneal
and form a hairpin, which is then cleaved by Dicer to form an siRNA.
The GeneClip™ U1 Hairpin Cloning Systems facilitate easy expression of shRNAs in
vivo by ddRNAi-based methods. This system provides a simple, cloning-based approach,
allowing ligation of potential shRNA target sequences into vectors that allow
transient or stable expression in mammalian cells.
The GeneClip™ U1 Hairpin Cloning Systems (Cat.# C8750, C8760,
C8770, C8780, C8790) are designed for rapid and efficient cloning
of hairpin sequences for expression of shRNAs in vivo. The U1 promoter provides the
benefit of production of shRNA with a defined termination site, mimicking siRNA
produced in vivo when the Dicer complex cleaves the loop off the hairpin. Using the
GeneClip™ Systems, complementary oligonucleotides supplied by the user are annealed
and cloned into the predigested vector downstream of the U1 promoter. The linearized
vectors contain overhangs for increased cloning efficiency. The GeneClip™ Systems are
provided in five formats. The GeneClip™ U1 Hairpin Cloning System—Basic is designed
for transient suppression of the gene of interest. The GeneClip™ U1 Hairpin Cloning
System—hMGFP contains the green fluorescent protein gene from Montastrea
cavernosa and can therefore be used to determine transfection
efficiency; it also allows separation of transfected cells by fluorescent-activated
cell sorting (FACS®). The three remaining GeneClip™ U1
Hairpin Cloning Systems provide the ability to stably select transfected cells using
either neomycin, hygromycin or puromycin, so that experimental results do not depend
on transfection efficiency.
An overview of the GeneClip™ U1 Hairpin Cloning System protocol is given in Figure
Once target sites have been selected, the hairpin oligonucleotides are
annealed and ligated into the pGeneClip™ Vector, which is then screened for the
proper insert (Figure 2.7). These two oligonucleotides form a DNA insert that
contains a hairpin siRNA target sequence. Standard desalting of the
oligonucleotides is necessary, but gel purification and 5′ phosphorylation are not
required. Because the oligonucleotides inserted into the vector are short (~60nt),
detection by standard agarose gel electrophoresis can be difficult. To make
detection of inserts more convenient, a PstI site is engineered into the vector
backbone (Figure 2.7). Upon successful ligation of the annealed
oligonucleotides into the vector, a second PstI site is created. Digestion with
PstI therefore results in two easily separated bands of approximately 3kb and 1kb.
If no insert is present, PstI digestion will result in a single band. In our
experiments, ligation of the insert typically results in over 50-fold more
colonies than control ligations containing vector alone. In addition, the PstI
digestion showed over 90% of the colonies contained insert.
Target Site Selection
The introduction of long strands of dsRNA into mammalian cells induces a strong
interferon response that can lead to an overall shutdown of protein synthesis and
cell death. Short dsRNA molecules (<30nt) can bypass the interferon
response and still function in the RNAi pathway (Elbashir et
al. 2001a). Since the entire mRNA sequence cannot be used in mammalian
cells, various target sites must be selected for use in RNAi experiments.
Unfortunately not all target sequences in an mRNA will work equally well to
suppress target gene expression. Experimental testing of the optimal sequences is
required to confirm a high level of inhibition.
Transient and Stable In Vivo Suppression
The pGeneClip™ Basic and pGeneClip™ hMGFP Vectors are recommended for use in
transient transfection assays. For many target sequences, transient transfection
can quickly yield cells that can be assessed for the effects of gene suppression,
including changes in phenotype, protein expression levels or other effects.
One consideration in evaluating transient transfection experiments is
transfection efficiency. For example, if only 30% of the cells are transfected, it
may be difficult to detect inhibition of expression, since 70% of the cells were
not successfully transfected and still express the target mRNA. The GeneClip™ U1
Hairpin Cloning System—hMGFP (Cat.# C8790) provides
an shRNA expression vector that contains an internal fluorescent marker for
monitoring delivery efficiency of shRNA-expressing constructs. In addition to
allowing transcription of hairpin target sequences and generation of siRNAs in
vivo, the vector contains an improved, synthetic version of the green fluorescent
protein (hMGFP) gene. The hMGFP gene encodes a 26kDa protein that gives improved
fluorescence and reduced cytotoxicity compared with other GFP proteins. The
presence of GFP allows easy determination of transfection efficiency and allows
selection of transfected cells by FACS® (Cormack
et al. 1996; Sorensen et al. 1999;
Galbraith et al. 1999). Importantly, the expression of hMGFP
does not affect gene silencing by shRNA molecules expressed from the same vector.
The pGeneClip™ Puromycin, pGeneClip™ Hygromycin and pGeneClip™ Neomycin Vectors
allow selection of stably transfected cells. Thus, the results are no longer
dependent on transfection efficiency. If necessary, clonal lines of transfected
cells can be generated if the population of selected cells does not show the
expected inhibition levels. Because integration of the vector into different
positions in the genome can affect expression of the RNAi hairpin, a population of
cells may not show suppression. A clonal cell line that sufficiently expresses the
RNAi hairpin may be required to demonstrate suppression with the pGeneClip™
To test the effectiveness of the GeneClip™ U1 promoter, we monitored reduction
in p53 protein levels after targeting p53 mRNA. p53 expression was targeted using
shRNAs cloned into a pGeneClip™ Basic Vector. More than 90% inhibition was
observed for the protein compared to a nonspecific control (data not shown). To
test p53 reduction in vivo, 293T cells, which contain elevated levels of p53, were
transfected with a pGeneClip™ Basic Vector expressing p53 shRNA. Compared to
nontransfected cells, the cells containing the pGeneClip™ Vector expressing p53
shRNA exhibited greater than 85% reduction in p53 protein levels (Figure 2.8,
Figure 2.8. Inhibition of p53 expression in transient transfections and in stable
293T cells were transfected with pGeneClip™ Basic Vectors containing
either a hairpin target sequence directed against p53 or a nonspecific
target sequence. After 48 hours, cells were collected, lysed, and protein
levels quantitated by BCA assay (Pierce). For stable transfections, 293T
cells were transfected with a pGeneClip™ Puromycin Vector containing
either a nonspecific or a p53-specific target sequence. Cells were
selected with puromycin for 7 days, and clones were assayed after passage
4 and passage 11. For Western analysis, 2μg protein per lane was loaded
on an 8% Tris-Glycine gel and then transferred to nitrocellulose. Blots
were probed with monoclonal antibodies to p53 (Oncogene Research
Products, Ab-2, 1:1,000 dilution) and β-actin (Abcam, AC-12, 1:5,000
dilution). Detection was performed using ECL™ Plus (Amersham). Protein
levels were quantitated by densitometry after exposure to film.
Panel A. shRNA suppression of p53 expression in
transiently transfected cells.
Panel B. Stable reduction of p53 at passage 4. Panel
C. Stable reduction of p53 at passage 11.
Additional Resources for the GeneClip™ U1 Hairpin Cloning Systems
Technical Bulletins and Manuals
GeneClip™ U1 Hairpin Cloning Systems Technical Manual
GeneClip™ U1 Hairpin Cloning Systems for expression of short hairpin
RNAs in vivo
return to top of page
Successful RNAi experiments are dependent on both siRNA design and effective delivery
of siRNA duplexes into cells. RNAi delivery strategies vary depending on the target
cells or organism. For example, C. elegans may be injected (Fire
et al. 1998; Grishok et al. 2000), soaked
in (Tabara et al. 1998), or fed (Timmons and Fire, 1998; Kamath
et al. 2001; Fraser et al. 2000) dsRNA.
Successful delivery of interfering RNA has also been achieved by microinjection of RNA
into Drosophila embryos (Kennerdell and Carthew, 1998) and mouse
oocytes (Wianny and Zernicka-Goetz, 2000). Delivery to Drosophila
S2 cells in culture can be achieved by incubating the cells with the chosen RNA (Clemens
et al. 2000; Betz and Worzella, 2003). Use of DNA-based
approaches like ddRNAi vectors allows use of standard transfection reagents/methods; for
example, cationic lipids, calcium phosphate, DEAE-dextran, polybrene-DMSO or
electroporation (Caplen et al. 2001; Elbashir et
Once annealed, hairpin oligonucleotides are ligated to the appropriate pGeneClip™
Vector, the resulting constructs can be used for transient or stable transfection.
The GeneClip™ U1 Hairpin Cloning Systems provide a choice of vectors containing
various selectable markers (neomycin, hygromycin or puromycin) that can be used for
stable expression of a pool of cells or individual clones. Transfection of the
plasmid DNA into human cells may be mediated by cationic lipids, calcium phosphate,
DEAE-Dextran, polybrene-DMSO or electroporation. Transfection conditions will need to
be optimized for your particular system. Guidelines for transfection of the
pGeneClip™ Vectors are provided in Technical Manual #TM256. General considerations
for transient and stable transfection are given below.
High transfection efficiency is essential for achieving substantial suppression
levels using a transient transfection approach. Prior to testing for suppression
of the target protein, optimize the transfection conditions for maximum efficiency
in the system to be tested. The optimal conditions will vary with cell type,
transfection method used and the amount of DNA. When using the pGeneClip™ Basic,
Puromycin, Hygromycin or Neomycin Vectors, optimization can be performed using a
GFP reporter such as the Monster Green® Fluorescent
Protein phMGFP Vector (Cat.# E6421). The pGeneClip™
hMGFP Vector already contains the GFP reporter. The GFP reporter can also be used
to determine transfection efficiency for the assay. To test the effectiveness of
the pGeneClip™ Vector constructs (screening various sequences for levels of
inhibition), the use of a reporter, such as GFP, is highly recommended. This
control can be performed as a separate transfection to determine the percentage of
the cell population transfected or as a cotransfection where flow cytometry can be
used to sort GFP-positive cells. The level of target RNA suppression in
transfected cells can then be determined by taking the transfection efficiency
Obtaining maximum suppression requires optimizing specific assay conditions. We
have observed variations in suppression efficiency as a result of the cell line,
cell culture conditions, target sequence and transfection conditions. Varying the
amount of transfection reagent, amount of DNA and cell density can influence
transfection efficiency. Obtaining the highest transfection efficiency with low
toxicity is essential for maximizing the siRNA interference (suppression) effect
in a transient assay. Additionally, maintaining healthy cell cultures is essential
for this application. The key considerations are discussed more fully below.
Cell Density (Confluence) at Transfection: The recommended cell
density for most cell types at transfection is approximately 30–50%; this level is
lower than standard transfection experiments where cells are plated at 50–70%
confluency. The optimal cell density should be determined for each cell type.
Continued proliferation and the need to passage cells should be considered when
determining the number of cells to plate.
Cell Proliferation: The successful suppression of gene expression
requires actively proliferating and dividing cells, so it is essential to maintain
healthy cell cultures. It also is important to minimize the decrease in cell
growth associated with nonspecific transfection effects and to maintain cell
culture under subconfluent conditions to assure rapid cell division. We recommend
using the CellTiter-Glo® Luminescent Cell Viability
Assay (Cat.# G7570) to monitor cell viability and
Time: The optimal time after transfection for analyzing interference
effects must be determined empirically by testing a range of incubation times.
Typically little inhibition is seen after 24 hours, but the maximal suppression
time can vary from 48–96 hours, depending on the cells used and the experimental
For stable expression, antibiotic selection must be applied following
transfection. Cell lines vary in the level of resistance to antibiotics, so the
level of resistance of a particular cell line must be tested before attempting
stable selection of the cells. A "kill curve" will determine the minimum
concentration of the antibiotic needed to kill nontransfected cells. The
antibiotic concentration for selection will vary depending on the cell type and
the growth rate. In addition, cells that are confluent are more resistant to
antibiotics, so it is important to keep the cells subconfluent. The typical
effective ranges and lengths of time needed for selection are given in Table 2.1.
|Table 2.1. Typical Conditions for Selection of Stable Transfectants.
||Time Needed for Selection
|pGeneClip™ Puromycin Vector
|pGeneClip™ Hygromycin Vector
|pGeneClip™ Neomycin Vector
For example, to generate a kill curve for G-418 selection, test G-418
concentrations of 0, 100, 200, 400, 600, 800 and 1,000µg/ml to determine the
concentration that is toxic to nontransfected cells. Once the effective
concentration of antibiotic has been determined, transfected cells can be selected
Once the effective concentration of antibiotic has been determined, transfected
cells can be selected for resistance, as outlined in the protocol below.
- Following transfection, seed cells at a low cell density.
- Apply antibiotic to the medium at the effective concentration determined
from the kill curve.
- Prepare a control plate for all selection experiments by treating
nontransfected cells with antibiotic in medium under the experimental
conditions. This control plate will confirm whether the conditions of
antibiotic selection were sufficiently stringent to eliminate cells not
expressing the resistance gene.
- Change the medium every 2–3 days until drug-resistant clones
- Once clones (or pools of cells) are selected, grow the cells in media
containing the antibiotic at a reduced antibiotic concentration, typically
25–50% of the level used during selection.
See Figure 2.8, Panels B and C for stably transfected cells that suppressed
expression of p53.
The protocol outlined below was used to successfully deliver PCR products of
various sizes (180bp or 505bp) generated either from the 778bp ERK-A target or from a
control plasmid containing the Renilla luciferase gene
(phRL-null Vector; 500bp or 1,000bp) to Drosophila S2 cells in
culture (Figure 2.9; Betz and Worzella, 2003). Purified, in vitro-synthesized ERK-A
dsRNA was introduced into Drosophila S2 cells using the method
described by Clemens et al. (2000) following the protocol
- Incubate 1 x 106 S2 cells in 1ml of
Drosophila expression system (DES) serum-free medium
(Invitrogen) in triplicate wells of a six-well culture dish in the presence or
absence of various amounts (0, 9.5, 38, or 190nM) of the test (ERK-A) dsRNA or
a nonspecific (Renilla luciferase) dsRNA.
- Incubate the S2 cells at room temperature with the dsRNA for 1
hour, then add 2ml of complete growth medium.
- Incubate the cells at room temperature for an additional 3 days to allow for
turnover of the target protein.
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Reduction of the targeted gene expression can be measured by 1) monitoring phenotypic
changes of the cell, 2) measuring changes in mRNA levels (e.g., using RT-PCR), or 3)
detecting changes in protein levels by immunocytochemistry or Western blot analysis
(Huang et al. 2003; Kullmann et al. 2002; Lang
et al. 2003). The suppression effect will vary depending on the
target, cell line and experimental conditions.
Controlling for nonspecific effects on other targets is very important. As a negative
control, cells can be transfected with either a nonspecific or scrambled target
sequence. This will show that suppression of gene expression is specific to the
expression of the hairpin siRNA target sequences. When suppression is determined by
Western analysis, positive controls for other genes (e.g., tubulin or actin) should be
included (Huang et al. 2003). Additional controls may also be
desirable (Editorial (2003) Nat. Cell Biol. 5,
Several summary articles are available that suggest various options for controls
that should be incorporated into RNAi experimental design to ensure accuracy and
correct identification of an RNAi effect (Editorial (2003) Nat. Cell
Biol. 5, 489–90; Duxbury and Whang, 2004). The preferred
control is to show restoration of functionality of a gene through artificial
overexpression of the target gene in a form that is not affected by RNAi. For
example, the target gene can be engineered to contain silent mutations that render
the mRNA invulnerable to the RNAi effect and introduced into the cell on a plasmid
vector. If such constructs “rescue” the original function of the gene, this is a good
indication that the observed suppression is mediated by RNAi. Use of siRNAs targeting
several different areas of the same gene to suppress expression may also be used to
provide evidence that an effect is mediated by RNAi. The observation of the same
suppression effect using more that one target RNA can confirm that the observed
effect is indeed RNAi.
For experiments using in vitro-synthesized siRNAs, the minimum concentration of
RNAi showing an effect should be used to avoid nonspecific effects due to the
introduction of large quantities of RNA into the cell. Ideally, any observed
suppression should be confirmed at both the mRNA and protein levels. Northern
blotting and quantitative, real-time RT-PCR can be used to demonstrate reduction of
expression at the RNA level. Quantitative Western blotting, phenotypic and functional
assays are some of the options available to show protein knockdown.
Scrambled siRNAs and siRNAs containing a single mismatch can be used as negative
controls. However, the latter are regarded as more informative. Positive controls
with RNAs known to exhibit an RNAi effect may also be useful.
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J. Biol. Chem.
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J. Biol. Chem.
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Biochem. Biophys. Res. Commun.
- Jensen, P.E. et al. (2004) The PSI-O subunit of plant photosystem I is involved in balancing the
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J. Biol. Chem.
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Products may be covered by pending or issued patents. Please visit our Web site
for more information. For additional patent and licensing information on the
psiCHECK™ Vectors, see statements (a) through (c), below.
(a)For research use only. This product and/or its use is
subject to one or more of the following Promega patents: U.S. Pat. Appln. Ser. Nos.
09/645,706, 10/664,341, 10/943,508, U.S. Pat. Nos. 5,670,356, 6,387,675 and
6,552,179, Australian Pat. No. 698424 and various corresponding patent applications
and issued patents. The terms of the limited license conveyed with the purchase of
this product are as follows: Researchers may use this product in their own research
and they may transfer derivatives to others for such research use provided that at
the time of transfer a copy of this label license is given to the recipients and the
recipients agree to be bound by the conditions of this label license. Researchers
shall have no right to modify or otherwise create variations of the nucleotide
sequence of the luciferase gene except that Researchers may: (1) clone heterologous
DNA sequences at either or both ends of said luciferase gene so as to create fused
gene sequences provided that the coding sequence of the resulting luciferase gene has
no more than four deoxynucleotides missing at the affected terminus when compared to
the intact luciferase gene sequence, and (2) insert and remove nucleic acid sequences
in furtherance of splicing research predicated on the inactivation or reconstitution
of the luminescent activity of the encoded luciferase. In addition, Researchers must
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