Silencing genes of interest at the RNA stage has proven to be a very useful tool for ascertaining the role of genes in live paradigms. Techniques for gene silencing, though, rely on the normal regulatory processes of cells and therefore have not always been wholly predictable or even effective. As scientists have found out more about the endogenous siRNA control of gene expression, so we have been able to ‘piggy-back’ more effectively on these natural processes. As a result, RNAi techniques can now be more effectively designed with surprising accuracy and much better potency. One of the key developments introduced via an academic/commercial collaboration has been a move from ‘21mers’ to ‘27mers’. Here, we look at the advantages of this move and some of the necessary optimisation processes to ensure that we combine nature and our growing knowledge to full effect.
by Dr Mark Behlke
In mammalian cells, small interfering RNAs (siRNAs) are produced by the enzymatic processing of long double-stranded RNAs (dsRNAs) by the RNase III class endoribonuclease Dicer. Newly formed siRNAs (usually around 21 bases in length) associate with Dicer, TRBP and Ago2 to form the RNA-induced silencing complex (RISC). Only one strand of the RNA is needed and therefore once the RISC has been formed, one strand of the siRNA (the ‘passenger’ strand) is degraded or discarded, whereas the other strand (the ‘guide’ strand) is retained and directs the sequence specificity of gene silencing. This natural pathway is what is exploited by researchers as an experimental tool for the selective reduction in expression of target genes.
SILENT RESEARCH
For research purposes, siRNAs have traditionally been chemically synthesised as 21-mer RNA duplexes that mimic the products of natural Dicer processing and assemble directly into the RISC. Alternative approaches to trigger RNAi exist that may have certain advantages, since they enter the endogenous process earlier on than traditional siRNAs. Dicer-substrate RNAs for example, are chemically synthesised 27-mer RNA duplexes that are optimised for Dicer processing. These are cleaved by Dicer into active, well defined 21-mers. Dicer-substrate RNAi methods take advantage of the link between Dicer and RISC loading that occurs when RNAs are processed by Dicer, and can boost potency by tenfold or more compared with 21-mer siRNAs at the same site [1]. A further advantage to using Dicer-substrate RNAs is that the direction of Dicer processing can favour loading of the antisense strand into RISC [2]. Correct design of the 27-mers is therefore crucial to realize the full benefits of this approach.
USING THE RIGHT KIT
Dicer-substrate RNAi methods were developed as a collaborative project between John Rossi and Dongho Kim at the Beckman Research Institute, City of Hope National Medical Center and Integrated DNA Technologies (IDT) [1, 2]. IDT has commercialised Dicer-substrate methods and reagents for research applications and offers synthesis of custom Dicer-substrate RNA duplexes, pre-made libraries of Dicer-substrate RNA duplexes (including a 550-target, 2,200-duplex human kinome library) and pre-designed sets of Dicer-substrate RNA duplexes for individual genes in the form of the TriFECTa kits. These contain three Dicer-substrate 27-mer RNA duplexes specific for a single target gene. The sequences used are selected from a pre-designed library of optimised 27-mers based upon the latest release of the RefSeq database in GenBank (http://www.ncbi.nlm.nih.gov/RefSeq/). The TriFECTa sequence library currently includes seven of the genomes in the RefSeq collection, including human, mouse and rat. TriFECTa duplexes are selected using a rational design algorithm that integrates both traditional 21-mer siRNA design rules as well as new 27-mer–specific criteria. Many other aspects of molecular interaction have to be considered as well, and it is therefore important to perform analysis to ensure that the chosen sites do not target alternatively spliced exons, do not include known single-nucleotide polymorphisms and share minimal homology to other genes in that species’ transcriptome (Smith-Waterman analysis).
MAINTAINING CONTROL
In addition to the target-specific duplexes, each kit has been designed to contain three controls including a fluorescent dye–labelled duplex (the Cy3 DS transfection control), a ‘universal’ negative control duplex (‘DS scrambled neg’, which targets a site that is absent from human, mouse and rat genomes) and a positive control duplex (‘HPRT-S1 DS positive control’), which targets a site in the hypoxanthine phosphoriblosy-transferase, a gene that is common between human, mouse and rat). Control reagents should be used to optimise the RNAi experimental system before undertaking studies on new targets. Furthermore, it is considered good practice to optimise transfection conditions for each different cell line studied as well as for each different form of nucleic acid used. This is especially important considering that large DNA plasmids often require different transfection conditions from short dsRNA oligonucleotides, for example. Dicer-substrate RNA duplexes can be used with all commonly used transfection methods, such as cationic lipids, liposomes and electroporation.
OPTIMISING RNAI EXPERIMENTS
Optimising the design of the siRNA is only one part of the overall RNAi process. It is important to remember that ‘A successful RNAi experiment always starts with a good transfection.’ With this in mind, it may be necessary to empirically test several different cationic lipids (or test other approaches) to establish a protocol that performs optimally with each cell line to be used. By using a dye-labelled transfection control oligo this can be reasonably quick and makes screening of many reagents in parallel very easy. Ideally the transfection method should result in >90% of the cells transfected with minimal cell death. At best, a transfection with 70% efficiency still leaves 30% of the target RNA intact if the RNAi duplex is 100% effective. Non-transfected cells will lead to underestimation of the potency of a siRNA duplex and may obscure important biological effects.
ENOUGH BUT NOT TOO MUCH!
When optimising transfection methods, it is recommended that dye-labelled oligos be used at 10 nM or less; higher concentrations can increase the amount of non-specific binding, which can cause high background and falsely elevate the apparent success of transfection. To remove any un-transfected material and provide the cells with ‘normal’ growing conditions, the culture medium should be changed and the cells washed before visualising the fluorescence. The ideal incubation time after transfection before cells are examined does vary with the target and the assays that are used, and is usually done less than a day after transfection (the optimal range is 6–24). An example of successful transfection using the TriFECTa Cy3 DS transfection control duplex is shown in Figure 1.
A GOOD START BUT NOT THE WHOLE STORY…
The use of dye-labelled controls is not always sufficient to optimise transfection protocols as they only indicate that the duplexes have entered the cell, but not whether they are in the correct cytoplasmic location for effective RNAi. Transfection conditions that succeed in the dye-labelled study should then be examined for functional knockdown using a positive control siRNA such as the HPRT-S1 DS positive control duplex. Once optimal transfection methods have been established, target-specific duplexes can be tested for relative potency and also used to establish a dose-response curve: It is usually adequate to test 10 nM, 1 nM and 0.1 nM concentrations.
KNOCKDOWN EFFICACY MRNA
Off-target effects are dose dependent and it is best to minimise the risk of adverse events by using the lowest concentration of RNA duplex that achieves the desired level of knockdown of the target mRNA. RNA assays (quantitative reverse-transcriptase PCR (qRT-PCR), northern blots, RNase protection assays, among others) are commonly performed 24–48 h after transfection, but a decrease in the amount of mRNA can often be detected in only a few hours after transfection.
To illustrate this, four TriFECTa duplexes specific for human HPRT1, SSB, STAT1 and HNRPH1 targets were prepared and transfected in HeLa cells. Response curves for these duplexes are displayed in Figure 2 and show that it would be advisable to use the HPRT1, STAT1 and HNRPH1 duplexes in the 1–5 nM range while the SSB duplex should be used at 10 nM concentration in any experiments.
PROTEINS
The downstream effect of gene knockdown is to reduce the production of the target protein and therefore protein assays (or assessment of phenotypes) are usually performed 48–72 h after transfection. Measurements of protein concentrations, enzyme activity or phenotype can vary widely due to a number of different factors such as protein half-life and rate of cell division (dilution), such that it is possible to observe a seemingly negative result even when mRNA concentrations have been substantially suppressed. Protein concentrations and phenotypic effects are best assessed after first establishing that mRNA levels are adequately reduced.
To give an idea of the different timescales of mRNA and protein knockdown for the same experiment, an HPRT-S1 DS positive control duplex was transfected into HeLa cells at 10 nM concentration using a cationic lipid. HPRT mRNA levels were measured at different time points using qRT-PCR [Figure 3 mRNA plot]. A reduction in mRNA levels was seen as early as 3 h after transfection and maximum knockdown was reached by 12 h. The average levels of protein knockdown for this series of experiments are also shown in Figure 3 [protein plot]. HPRT mRNA levels were reduced by >90% by 12 h after transfection, whereas protein levels did not reach this level of knockdown until the 72-h time point. In all of the experiments, we used the ‘scrambled neg’ duplex as negative control and levels of the unknown were normalised against an internal control standard.
OPTIMISATION SUMMARY
It is worth summarising here some of the key steps for an optimisation strategy:
1) Optimise transfection methods
a) Dye-labelled control
b) Positive-control knockdown
2) Test target-specific duplexes
a) Validate that duplexes actually work in your cell line
b) Titrate dose response
3) Knockdown studies of targeted gene
a) Assess changes in mRNA levels
b) Assess changes in protein levels
c) Assess changes in phenotype
CLOSING NOTE
It is important to consider, especially in whole organism studies, that RNA duplexes can trigger interferon responses. Stimulation of the innate immune system can occur through a variety of receptor molecules, but it is possible to mitigate this effect by modifying the RNA duplex with 2’-O-methyl RNA bases [5]. Methods for use of Dicer-substrate RNA duplexes in vivo are under development [6], and modified TriFECTa duplexes are available to support these needs.
REFERENCES
1. Kim DH et al. Synthetic dsRNA Dicer substrates enhance RNAi potency and efficacy. Nat Biotechnol 2005; 23: 222–226.
2. Rose SD et al. Functional polarity is introduced by Dicer processing of short substrate RNAs. Nucleic Acids Res 2005; 33: 4140–4156.
3. Marques JT & Williams BR. Activation of the mammalian immune system by siRNAs. Nat Biotechnol 2005; 23: 1399–1405.
4. Marques JT et al. A structural basis for discriminating between self and nonself double-stranded RNAs in mammalian cells. Nat Biotechnol 2006; 24: 559–565.
5. Judge AD et al. Design of noninflammatory synthetic siRNA mediating potent gene silencing in vivo. Mol Ther 2006; 13: 494–505.
6. Amarzguioui M et al. Rational design and in vitro and in vivo delivery of Dicer substrate siRNA. Nat Protocols 2006; 1: 508–517.
THE AUTHOR
Mark Behlke, M.D., Ph.D., Chief Scientific Officer
Integrated DNA Technologies, Inc.
1710 Commercial Park
Coralville, IA 52241, USA
