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RNA interference (RNAi) is a biological process in which RNA molecules are involved in sequence-specific suppression of gene expression by double-stranded RNA, through translational or transcriptional repression. Historically, RNAi was known by other names, including co-suppression, post-transcriptional gene silencing (PTGS), and quelling. The detailed study of each of these seemingly different processes elucidated that the identity of these phenomena were all actually RNAi. Andrew Fire and Craig C. Mello shared the 2006 Nobel Prize in Physiology or Medicine for their work on RNAi in the nematode worm Caenorhabditis elegans, which they published in 1998. Since the discovery of RNAi and its regulatory potentials, it has become evident that RNAi has immense potential in suppression of desired genes. RNAi is now known as precise, efficient, stable and better than antisense therapy for gene suppression.^[1] Antisense RNA produced intracellularly by an expression vector may be developed and find utility as novel therapeutic agents.^[2]

Two types of small ribonucleic acid (RNA) molecules, microRNA (miRNA) and small interfering RNA (siRNA), are central to components to the RNAi pathway. Once mRNA is degraded, post-transcriptional silencing occurs as protein translation is prevented. Transcription can be inhibited via the pre-transcriptional silencing mechanism of RNAi, through which an enzyme complex catalyzes DNA methylation at genomic positions complementary to complexed siRNA or miRNA. RNAi has an important role in defending cells against parasitic nucleotide sequences (e.g., viruses or transposons) and also influences development of organisms.

The RNAi pathway is a naturally occurring process found in many eukaryotes and animal cells. It is initiated by the enzyme Dicer, which cleaves long double-stranded RNA (dsRNA) molecules into short double-stranded fragments of approximately 21 to 23 nucleotide siRNAs. Each siRNA is unwound into two single-stranded RNAs (ssRNAs), the passenger (sense) strand and the guide (antisense) strand. The passenger strand is then cleaved by the protein Argonaute 2 (Ago2). The passenger strand is degraded and the guide strand is incorporated into the RNA-induced silencing complex (RISC). The RISC assembly then binds and degrades the target mRNA. Specifically, this is accomplished when the guide strand pairs with a complementary sequence in a mRNA molecule and induces cleavage by Ago2, a catalytic component of the RISC. In some organisms, this process spreads systemically, despite the initially limited molar concentrations of siRNA.^[3]

RNAi is a valuable research tool, both in cell culture and in living organisms, because synthetic dsRNA introduced into cells can selectively and robustly induce suppression of specific genes of interest. RNAi may be used for large-scale screens that systematically shut down each gene (and the subsequent proteins it codes for) in the cell, which can help to identify the components necessary for a particular cellular process or an event such as cell division. The pathway is also used as a practical tool for food, medicine and insecticides.^[4]

Cellular mechanism

The Dicer protein from *Giardia*, which catalyzes the cleavage of dsRNA to siRNAs. The RNase domains are colored green, the PAZ domain yellow, the platform domain red, and the connector helix blue.^[5]

RNAi is an RNA-dependent gene silencing process that is controlled by RISC and is initiated by short double-stranded RNA molecules in a cell's cytoplasm, where they interact with the catalytic RISC component Argonaute.^[6] When the dsRNA is exogenous (coming from infection by a virus with an RNA genome or laboratory manipulations), the RNA is imported directly into the cytoplasm and cleaved to short fragments by Dicer. The initiating dsRNA can also be endogenous (originating in the cell), as in pre-microRNAs expressed from RNA-coding genes in the genome. The primary transcripts from such genes are first processed to form the characteristic stem-loop structure of pre-miRNA in the nucleus, then exported to the cytoplasm. Thus, the two dsRNA pathways, exogenous and endogenous, converge at the RISC.^[7]

Exogenous dsRNA initiates RNAi by activating the ribonuclease protein Dicer,^[8] which binds and cleaves dsRNAs in plants, or short hairpin RNAs (shRNAs) in humans, to produce double-stranded fragments of 20–25 base pairs with a 2-nucleotide overhang at the 3′ end.^[9] Bioinformatics studies on the genomes of multiple organisms suggest this length maximizes target-gene specificity and minimizes non-specific effects.^[10] These short double-stranded fragments are called siRNAs. These siRNAs are then separated into single strands and integrated into an active RISC, by RISC-Loading Complex (RLC). RLC includes Dicer-2 and R2D2, and is crucial to unite Ago2 and RISC.^[11] TATA-binding protein-associated factor 11 (TAF11) assembles the RLC by facilitating Dcr-2-R2D2 tetramerization, which increases the binding affinity to siRNA by 10-fold. Association with TAF11 would convert the R2-D2-Initiator (RDI) complex into the RLC.^[12] R2D2 carries tandem double-stranded RNA-binding domains to recognize the thermodynamically stable terminus of siRNA duplexes, whereas Dicer-2 the other less stable extremity. Loading is asymmetric: the MID domain of Ago2 recognizes the thermodynamically stable end of the siRNA. Therefore, the "passenger" (sense) strand whose 5′ end is discarded by MID is ejected, while the saved "guide" (antisense) strand cooperates with AGO to form the RISC.^[11]

After integration into the RISC, siRNAs base-pair to their target mRNA and cleave it, thereby preventing it from being used as a translation template.^[13] Differently from siRNA, a miRNA-loaded RISC complex scans cytoplasmic mRNAs for potential complementarity. Instead of destructive cleavage (by Ago2), miRNAs rather target the 3′ untranslated region (UTR) regions of mRNAs where they typically bind with imperfect complementarity, thus blocking the access of ribosomes for translation.^[14]

Exogenous dsRNA is detected and bound by an effector protein, known as RDE-4 in C. elegans and R2D2 in Drosophila, that stimulates Dicer activity.^[15] The mechanism producing this length specificity is unknown and this protein only binds long dsRNAs.^[15]

In C. elegans this initiation response is amplified through the synthesis of a population of 'secondary' siRNAs during which the Dicer-produced initiating or 'primary' siRNAs are used as templates.^[16] These 'secondary' siRNAs are structurally distinct from Dicer-produced siRNAs and appear to be produced by an RNA-dependent RNA polymerase (RdRP).^[17]

MicroRNA

MicroRNAs (miRNAs) are genomically encoded non-coding RNAs that help regulate gene expression, particularly during development.^[18] The phenomenon of RNAi, broadly defined, includes the endogenously induced gene silencing effects of miRNAs as well as silencing triggered by foreign dsRNA. Mature miRNAs are structurally similar to siRNAs produced from exogenous dsRNA, but before reaching maturity, miRNAs must first undergo extensive post-transcriptional modification. A miRNA is expressed from a much longer RNA-coding gene as a primary transcript known as a pri-miRNA which is processed, in the cell nucleus, to a 70-nucleotide stem-loop structure called a pre-miRNA by the microprocessor complex. This complex consists of an RNase III enzyme called Drosha and a dsRNA-binding protein DGCR8. The dsRNA portion of this pre-miRNA is bound and cleaved by Dicer to produce the mature miRNA molecule that can be integrated into the RISC complex; thus, miRNA and siRNA share the same downstream cellular machinery.^[19] First, viral encoded miRNA was described in Epstein–Barr virus (EBV).^[20] Thereafter, an increasing number of microRNAs have been described in viruses. VIRmiRNA is a comprehensive catalogue covering viral microRNA, their targets and anti-viral miRNAs^[21] (see also VIRmiRNA resource: http://crdd.osdd.net/servers/virmirna/).

siRNAs derived from long dsRNA precursors differ from miRNAs in that miRNAs, especially those in animals, typically have incomplete base pairing to a target and inhibit the translation of many different mRNAs with similar sequences. In contrast, siRNAs typically base-pair perfectly and induce mRNA cleavage only in a single, specific target.^[22] In Drosophila and C. elegans, miRNA and siRNA are processed by distinct Argonaute proteins and Dicer enzymes.^[23]^[24]

Three prime untranslated regions and microRNAs

Three prime untranslated regions (3′UTRs) of mRNAs often contain regulatory sequences that post-transcriptionally cause RNAi. Such 3′-UTRs often contain both binding sites for miRNAs as well as for regulatory proteins. By binding to specific sites within the 3′-UTR, miRNAs can decrease gene expression of various mRNAs by either inhibiting translation or directly causing degradation of the transcript. The 3′-UTR also may have silencer regions that bind repressor proteins that inhibit the expression of a mRNA.

The 3′-UTR often contains microRNA response elements (MREs). MREs are sequences to which miRNAs bind. These are prevalent motifs within 3′-UTRs. Among all regulatory motifs within the 3′-UTRs (e.g. including silencer regions), MREs make up about half of the motifs.

As of 2023, the miRBase web site,^[25] an archive of miRNA sequences and annotations, listed 28,645 entries in 271 biologic species. Of these, 1,917 miRNAs were in annotated human miRNA loci. miRNAs were predicted to have an average of about four hundred target mRNAs (affecting expression of several hundred genes).^[26] Friedman et al.^[26] estimate that >45,000 miRNA target sites within human mRNA 3′UTRs are conserved above background levels, and >60% of human protein-coding genes have been under selective pressure to maintain pairing to miRNAs.

Direct experiments show that a single miRNA can reduce the stability of hundreds of unique mRNAs.^[27] Other experiments show that a single miRNA may repress the production of hundreds of proteins, but that this repression often is relatively mild (less than 2-fold).^[28]^[29]

The effects of miRNA dysregulation of gene expression seem to be important in cancer.^[30] For instance, in gastrointestinal cancers, nine miRNAs have been identified as epigenetically altered and effective in down regulating DNA repair enzymes.^[31]

The effects of miRNA dysregulation of gene expression also seem to be important in neuropsychiatric disorders, such as schizophrenia, bipolar disorder, major depression, Parkinson's disease, Alzheimer's disease and autism spectrum disorders.^[32]^[33]^[34]

RISC activation and catalysis

Exogenous dsRNA is detected and bound by an effector protein, known as RDE-4 in C. elegans and R2D2 in Drosophila, that stimulates Dicer activity.^[15] This protein only binds long dsRNAs, but the mechanism producing this length specificity is unknown.^[15] This RNA-binding protein then facilitates the transfer of cleaved siRNAs to the RISC complex.^[35]

In C. elegans this initiation response is amplified through the synthesis of a population of 'secondary' siRNAs during which the Dicer-produced initiating or 'primary' siRNAs are used as templates.^[16] These 'secondary' siRNAs are structurally distinct from Dicer-produced siRNAs and appear to be produced by an RNA-dependent RNA polymerase (RdRP).^[17]

The active components of an RNA-induced silencing complex (RISC) are endonucleases called Argonaute proteins, which cleave the target mRNA strand complementary to their bound siRNA.^[6] As the fragments produced by Dicer are double-stranded, they could each in theory produce a functional siRNA. However, only one of the two strands, which is known as the guide strand, binds Argonaute and directs gene silencing. The other anti-guide strand or passenger strand is degraded during RISC activation.^[36] Although it was first believed that an ATP-dependent helicase separated these two strands,^[37] the process proved to be ATP-independent and performed directly by the protein components of RISC.^[3]^[38] However, an in vitro kinetic analysis of RNAi in the presence and absence of ATP showed that ATP may be required to unwind and remove the cleaved mRNA strand from the RISC complex after catalysis.^[39] The guide strand tends to be the one whose 5′ end is less stably paired to its complement,^[40] but strand selection is unaffected by the direction in which Dicer cleaves the dsRNA before RISC incorporation.^[41] Instead, the R2D2 protein may serve as the differentiating factor by binding the more-stable 5′ end of the passenger strand.^[42]

The structural basis for binding of RNA to the Argonaute protein was examined by X-ray crystallography of the binding domain of an RNA-bound Argonaute. Here, the phosphorylated 5′ end of the RNA strand enters a conserved basic surface pocket and makes contacts through a divalent cation (an atom with two positive charges) such as magnesium and by aromatic stacking (a process that allows more than one atom to share an electron by passing it back and forth) between the 5′ nucleotide in the siRNA and a conserved tyrosine residue. This site is thought to form a nucleation site for the binding of the siRNA to its mRNA target.^[43] Analysis of the inhibitory effect of mismatches in either the 5’ or 3’ end of the guide strand has demonstrated that the 5’ end of the guide strand is likely responsible for matching and binding the target mRNA, while the 3’ end is responsible for physically arranging target mRNA into a cleavage-favorable RISC region.^[39]

It is not understood how the activated RISC complex locates complementary mRNAs within the cell. Although the cleavage process has been proposed to be linked to translation, translation of the mRNA target is not essential for RNAi-mediated degradation.^[44] Indeed, RNAi may be more effective against mRNA targets that are not translated.^[45] Argonaute proteins are localized to specific regions in the cytoplasm called P-bodies (also cytoplasmic bodies or GW bodies), which are regions with high rates of mRNA decay;^[46] miRNA activity is also clustered in P-bodies.^[47] Disruption of P-bodies decreases the efficiency of RNAi, suggesting that they are a critical site in the RNAi process.^[48]

Transcriptional silencing

The enzyme Dicer trims double stranded RNA, to form small interfering RNA or microRNA. These processed RNAs are incorporated into the RNA-induced silencing complex (RISC), which targets messenger RNA to prevent translation.^[49]

Components of the RNAi pathway are used in many eukaryotes in the maintenance of the organization and structure of their genomes. Modification of histones and associated induction of heterochromatin formation serves to downregulate genes pre-transcriptionally;^[50] this process is referred to as RNA-induced transcriptional silencing (RITS), and is carried out by a complex of proteins called the RITS complex. In fission yeast this complex contains Argonaute, a chromodomain protein Chp1, and a protein called Tas3 of unknown function.^[51] As a consequence, the induction and spread of heterochromatic regions requires the Argonaute and RdRP proteins.^[52] Indeed, deletion of these genes in the fission yeast S. pombe disrupts histone methylation and centromere formation,^[53] causing slow or stalled anaphase during cell division.^[54] In some cases, similar processes associated with histone modification have been observed to transcriptionally upregulate genes.^[55]

The mechanism by which the RITS complex induces heterochromatin formation and organization is not well understood. Most studies have focused on the mating-type region in fission yeast, which may not be representative of activities in other genomic regions/organisms. In maintenance of existing heterochromatin regions, RITS forms a complex with siRNAs complementary to the local genes and stably binds local methylated histones, acting co-transcriptionally to degrade any nascent pre-mRNA transcripts that are initiated by RNA polymerase. The formation of such a heterochromatin region, though not its maintenance, is Dicer-dependent, presumably because Dicer is required to generate the initial complement of siRNAs that target subsequent transcripts.^[56] Heterochromatin maintenance has been suggested to function as a self-reinforcing feedback loop, as new siRNAs are formed from the occasional nascent transcripts by RdRP for incorporation into local RITS complexes.^[57] The relevance of observations from fission yeast mating-type regions and centromeres to mammals is not clear, as heterochromatin maintenance in mammalian cells may be independent of the components of the RNAi pathway.^[58]

Crosstalk with RNA editing

The type of RNA editing that is most prevalent in higher eukaryotes converts adenosine nucleotides into inosine in dsRNAs via the enzyme adenosine deaminase (ADAR).^[59] It was originally proposed in 2000 that the RNAi and A→I RNA editing pathways might compete for a common dsRNA substrate.^[60] Some pre-miRNAs do undergo A→I RNA editing^[61]^[62] and this mechanism may regulate the processing and expression of mature miRNAs.^[62] Furthermore, at least one mammalian ADAR can sequester siRNAs from RNAi pathway components.^[63] Further support for this model comes from studies on ADAR-null C. elegans strains indicating that A→I RNA editing may counteract RNAi silencing of endogenous genes and transgenes.^[64]

Variation among organisms

Organisms vary in their ability to take up foreign dsRNA and use it in the RNAi pathway. The effects of RNAi can be both systemic and heritable in plants and C. elegans, although not in Drosophila or mammals. In plants, RNAi is thought to propagate by the transfer of siRNAs between cells through plasmodesmata (channels in the cell walls that enable communication and transport).^[37] Heritability comes from methylation of promoters targeted by RNAi; the new methylation pattern is copied in each new generation of the cell.^[66] A broad general distinction between plants and animals lies in the targeting of endogenously produced miRNAs; in plants, miRNAs are usually perfectly or nearly perfectly complementary to their target genes and induce direct mRNA cleavage by RISC, while animals' miRNAs tend to be more divergent in sequence and induce translational repression.^[65] This translational effect may be produced by inhibiting the interactions of translation initiation factors with the mRNA's polyadenine tail.^[67]

Some eukaryotic protozoa such as Leishmania major and Trypanosoma cruzi lack the RNAi pathway entirely.^[68]^[69] Most or all of the components are also missing in some fungi, most notably the model organism Saccharomyces cerevisiae.^[70] The presence of RNAi in other budding yeast species such as Saccharomyces castellii and Candida albicans, further demonstrates that inducing two RNAi-related proteins from S. castellii facilitates RNAi in S. cerevisiae.^[71] That certain ascomycetes and basidiomycetes are missing RNAi pathways indicates that proteins required for RNA silencing have been lost independently from many fungal lineages, possibly due to the evolution of a novel pathway with similar function, or to the lack of selective advantage in certain niches.^[72]

Related prokaryotic systems

Gene expression in prokaryotes is influenced by an RNA-based system similar in some respects to RNAi. Here, RNA-encoding genes control mRNA abundance or translation by producing a complementary RNA that anneals to an mRNA. However these regulatory RNAs are not generally considered to be analogous to miRNAs because the Dicer enzyme is not involved.^[73] It has been suggested that CRISPR interference systems in prokaryotes are analogous to eukaryotic RNAi systems, although none of the protein components are orthologous.^[74]

Biological functions

Immunity

RNAi is a vital part of the immune response to viruses and other foreign genetic material, especially in plants where it may also prevent the self-propagation of transposons.^[75] Plants such as Arabidopsis thaliana express multiple Dicer homologs that are specialized to react differently when the plant is exposed to different viruses.^[76] Even before the RNAi pathway was fully understood, it was known that induced gene silencing in plants could spread throughout the plant in a systemic effect and could be transferred from stock to scion plants via grafting.^[77] This phenomenon has since been recognized as a feature of the plant immune system which allows the entire plant to respond to a virus after an initial localized encounter.^[78] In response, many plant viruses have evolved elaborate mechanisms to suppress the RNAi response.^[79] These include viral proteins that bind short double-stranded RNA fragments with single-stranded overhang ends, such as those produced by Dicer.^[80] Some plant genomes also express endogenous siRNAs in response to infection by specific types of bacteria.^[81] These effects may be part of a generalized response to pathogens that downregulates any metabolic process in the host that aids the infection process.^[82]

Although animals generally express fewer variants of the Dicer enzyme than plants, RNAi in some animals produces an antiviral response. In both juvenile and adult Drosophila, RNAi is important in antiviral innate immunity and is active against pathogens such as Drosophila X virus.^[83]^[84] A similar role in immunity may operate in C. elegans, as Argonaute proteins are upregulated in response to viruses and worms that overexpress components of the RNAi pathway are resistant to viral infection.^[85]^[86]

The role of RNAi in mammalian innate immunity is poorly understood, and relatively little data is available. However, the existence of viruses that encode genes able to suppress the RNAi response in mammalian cells may be evidence in favour of an RNAi-dependent mammalian immune response,^[87]^[88] although this hypothesis has been challenged as poorly substantiated.^[89] Evidence for the existence of a functional antiviral RNAi pathway in mammalian cells has been presented.^[90]^[91]

Other functions for RNAi in mammalian viruses also exist, such as miRNAs expressed by the herpes virus that may act as heterochromatin organization triggers to mediate viral latency.^[92]

Downregulation of genes

Endogenously expressed miRNAs, including both intronic and intergenic miRNAs, are most important in translational repression^[65] and in the regulation of development, especially on the timing of morphogenesis and the maintenance of undifferentiated or incompletely differentiated cell types such as stem cells.^[93] The role of endogenously expressed miRNA in downregulating gene expression was first described in C. elegans in 1993.^[94] In plants this function was discovered when the "JAW microRNA" of Arabidopsis was shown to be involved in the regulation of several genes that control plant shape.^[95] In plants, the majority of genes regulated by miRNAs are transcription factors;^[96] thus miRNA activity is particularly wide-ranging and regulates entire gene networks during development by modulating the expression of key regulatory genes, including transcription factors as well as F-box proteins.^[97] In many organisms, including humans, miRNAs are linked to the formation of tumors and dysregulation of the cell cycle. Here, miRNAs can function as both oncogenes and tumor suppressors.^[98]

Evolution

Based on parsimony-based phylogenetic analysis, the most recent common ancestor of all eukaryotes most likely already possessed an early RNAi pathway; the absence of the pathway in certain eukaryotes is thought to be a derived characteristic.^[99] This ancestral RNAi system probably contained at least one Dicer-like protein, one Argonaute, one PIWI protein, and an RNA-dependent RNA polymerase that may also have played other cellular roles. A large-scale comparative genomics study likewise indicates that the eukaryotic crown group already possessed these components, which may then have had closer functional associations with generalized RNA degradation systems such as the exosome.^[100] This study also suggests that the RNA-binding Argonaute protein family, which is shared among eukaryotes, most archaea, and at least some bacteria (such as Aquifex aeolicus), is homologous to and originally evolved from components of the translation initiation system.

Applications

RNAi pathway for gene knockdown

Gene knockdown is a method used to reduce the expression of an organism’s specific genes. This is accomplished by using the naturally occurring process of RNAi.^[6] This gene knockdown technique uses a double-stranded siRNA molecule that is synthesized with a sequence complementary to the gene of interest. The RNAi cascade begins once the Dicer enzyme starts to process siRNA. The end result of the process leads to degradation of mRNA and destroys any instructions needed to build certain proteins. Using this method, researchers are able to decrease (but not completely eliminate) the expression of a targeted gene. Studying the effects of this decrease in expression may show the physiological role or impact of the targeted gene products.^[101]^[102]

Off-Target Effects of Gene Knockdown

Extensive efforts in computational biology have been directed toward the design of successful dsRNA reagents that maximize gene knockdown but minimize "off-target" effects. Off-target effects arise when an introduced RNA has a base sequence that can pair with and thus reduce the expression of multiple genes. Such problems occur more frequently when the dsRNA contains repetitive sequences. It has been estimated from studying the genomes of humans, C. elegans and S. pombe that about 10% of possible siRNAs have substantial off-target effects.^[10] A multitude of software tools have been developed implementing algorithms for the design of general^[103]^[104] mammal-specific,^[105] and virus-specific^[106] siRNAs that are automatically checked for possible cross-reactivity.

Depending on the organism and experimental system, the exogenous RNA may be a long strand designed to be cleaved by Dicer, or short RNAs designed to serve as siRNA substrates. In most mammalian cells, shorter RNAs are used because long double-stranded RNA molecules induce the mammalian interferon response, a form of innate immunity that reacts nonspecifically to foreign genetic material.^[107] Mouse oocytes and cells from early mouse embryos lack this reaction to exogenous dsRNA and are therefore a common model system for studying mammalian gene-knockdown effects.^[108] Specialized laboratory techniques have also been developed to improve the utility of RNAi in mammalian systems by avoiding the direct introduction of siRNA, for example, by stable transfection with a plasmid encoding the appropriate sequence from which siRNAs can be transcribed,^[109] or by more elaborate lentiviral vector systems allowing the inducible activation or deactivation of transcription, known as conditional RNAi.^[110]^[111]

Medications

The technique of knocking down genes using RNAi therapeutics has demonstrated success in randomized controlled clinical studies. These medications are a growing class of siRNA-based drugs that decrease the expression of proteins encoded by certain genes. To date, five RNAi medications have been approved by regulatory authorities in the US and Europe: patisiran (2018), givosiran (2019), lumasiran (2020), inclisiran (2020 in Europe with anticipated US approval in 2021), and vutrisiran (2022).^[112]^[113]^[114]^[115]

While all of the current regulatory body approved RNAi therapeutics focus on diseases that originate in the liver, additional medications under investigation target a host of disease areas including cardiovascular diseases, bleeding disorders, alcohol use disorders, cystic fibrosis, gout, carcinoma, and eye disorders.

Patisiran is the first double stranded siRNA-based medication approved in 2018 and developed by Alnylam Pharmaceuticals. Patisiran uses the RNAi cascade to suppress the gene that codes for TTR (transthryetin). Mutations in this gene may cause the misfolding of a protein responsible for hereditary ATTR amyloidosis. To achieve therapeutic response, patisiran is encased by a lipid nanoparticle membrane that facilitates crossover into the cytoplasm. Once inside the cell, the siRNA begins processing by the enzyme Dicer. Patisiran is administered by a healthcare professional through an intravenous infusion with dosing based on body weight. Warnings and precautions include risk of infusion-related reactions and reduced vitamin A levels (serum).^[116]

In 2019, the FDA and EMA approved givosiran for the treatment of adults with acute hepatic porphyria (AHP).^[117] The FDA also granted givosiran a breakthrough therapy designation, priority review designation, and orphan drug designation for the treatment of acute hepatic porphyria (AHP) in November 2019.^[118] By 2020, givosiran received EMA approval.^[119] Givosiran is an siRNA that breaks down aminolevulinic acid synthase 1 (ALAS1) mRNA in the liver. Breaking down ALAS1 mRNA prevents toxins (responsible for neurovisceral attacks and AHP disease) such as aminolevulinic acid (ALA) and porphobilinogen (PBG) from accumulating.^[120]^[121]^[122]^[123] To facilitate entry into the cytoplasm, givosiran uses GalNAc ligands and enters into liver cells. The medication is administered subcutaneously by a healthcare professional with dosing based on body weight. Warnings and precautions include risk of anaphylactic reactions, hepatic toxicity, renal toxicity and injection site reactions.^[124]

Lumasiran was approved as a siRNA-based medication in 2020 for use in both the European Union and the United States.^[125]^[126] This medication is used for the treatment of primary hyperoxaluria type 1 (PH1) in pediatric and adult populations. The drug is designed to reduce hepatic oxalate production and urinary oxalate levels through RNAi by targeting hydroxyacid oxidase 1 (HAO1) mRNA for breakdown. Lowering HAO1 enzyme levels reduces the oxidation of glycolate to glyoxylate (which is a substrate for oxalate). Lumasiran is administered subcutaneously by a healthcare professional with dosing based on body weight.^[127] Data from randomized controlled clinical trials indicate that the most common adverse reaction that was reported was injection site reactions. These reactions were mild and were present in 38 percent of patients treated with lumasiran.^[128]

In 2022, the FDA and EMA approved vutrisiran for the treatment of adults with hereditary transthyretin mediated amyloidosis with polyneuropathy stage 1 or 2.^[129]^[130] Vutrisiran is designed to break down the mRNA that codes for transthyretin.

Other investigational drugs using RNAi that are being developed by pharmaceutical companies such as Arrowhead Pharmaceuticals, Dicerna, Alnylam Pharmaceuticals, Amgen, and Sylentis. These medications cover a variety of targets via RNAi and diseases.

Investigational RNAi therapeutics in development:

Drug	Target	Delivery System	Disease	Phase	Status	Company	Identifier
ALN–VSP02	KSP and VEGF	LNP	Solid tumours	I	Completed	Alnylam Pharmaceuticals	NCT01158079
siRNA–EphA2–DOPC	EphA2	LNP	Advanced cancers	I	Recruiting	MD Anderson Cancer Center	NCT01591356
Atu027	PKN3	LNP	Solid tumours	I	Completed	Silence Therapeutics	NCT00938574
TKM–080301	PLK1	LNP	Cancer	I	Recruiting	Tekmira Pharmaceutical	NCT01262235
TKM–100201	VP24, VP35, Zaire Ebola L-polymerase	LNP	Ebola-virus infection	I	Recruiting	Tekmira Pharmaceutical	NCT01518881
ALN–RSV01	RSV nucleocapsid	Naked siRNA	Respiratory syncytial virus infections	II	Completed	Alnylam Pharmaceuticals	NCT00658086
PRO-040201	ApoB	LNP	Hypercholesterolaemia	I	Terminated	Tekmira Pharmaceutical	NCT00927459
ALN–PCS02	PCSK9	LNP	Hypercholesterolaemia	I	Completed	Alnylam Pharmaceuticals	NCT01437059
ALN–TTR02	TTR	LNP	Transthyretin-mediated amyloidosis	II	Recruiting	Alnylam Pharmaceuticals	NCT01617967
CALAA-01	RRM2	Cyclodextrin NP	Solid tumours	I	Active	Calando Pharmaceuticals	NCT00689065
TD101	K6a (N171K mutation)	Naked siRNA	Pachyonychia congenita	I	Completed	Pachyonychia Congenita Project	NCT00716014
AGN211745	VEGFR1	Naked siRNA	Age-related macular degeneration, choroidal neovascularization	II	Terminated	Allergan	NCT00395057
QPI-1007	CASP2	Naked siRNA	Optic atrophy, non-arteritic anterior ischaemic optic neuropathy	I	Completed	Quark Pharmaceuticals	NCT01064505
I5NP	p53	Naked siRNA	Kidney injury, acute renal failure	I	Completed	Quark Pharmaceuticals	NCT00554359
			Delayed graft function, complications of kidney transplant	I, II	Recruiting	Quark Pharmaceuticals	NCT00802347
PF-655 (PF-04523655)	RTP801 (Proprietary target)	Naked siRNA	Choroidal neovascularization, diabetic retinopathy, diabetic macular oedema	II	Active	Quark Pharmaceuticals	NCT01445899
siG12D LODER	KRAS	LODER polymer	Pancreatic cancer	II	Recruiting	Silenseed	NCT01676259
Bevasiranib	VEGF	Naked siRNA	Diabetic macular oedema, macular degeneration	II	Completed	Opko Health	NCT00306904
SYL1001	TRPV1	Naked siRNA	Ocular pain, dry-eye syndrome	I, II	Recruiting	Sylentis	NCT01776658
SYL040012	ADRB2	Naked siRNA	Ocular hypertension, open-angle glaucoma	II	Recruiting	Sylentis	NCT01739244
CEQ508	CTNNB1	Escherichia coli-carrying shRNA	Familial adenomatous polyposis	I, II	Recruiting	Marina Biotech	Unknown
RXi-109	CTGF	Self-delivering RNAi compound	Cicatrix scar prevention	I	Recruiting	RXi Pharmaceuticals	NCT01780077
ALN–TTRsc	TTR	siRNA–GalNAc conjugate	Transthyretin-mediated amyloidosis	I	Recruiting	Alnylam Pharmaceuticals	NCT01814839
ARC-520	Conserved regions of HBV	DPC	HBV	I	Recruiting	Arrowhead Research	NCT01872065

Legal categorization and legal issues in a near future

Currently, both miRNA and SiRNA are currently chemically synthesized and so, are legally categorized inside EU and in USA as "simple" medicinal products. But as bioengineered siRNA (BERAs) are in development, these would be classified as biological medicinal products, at least in EU. The development of the BERAs technology raises the question of the categorization of drugs having the same mechanism of action but being produced chemically or biologically. This lack of consistency should be addressed.^[131]

Delivery mechanisms

To achieve the clinical potential of RNAi, siRNA must be efficiently transported to the cells of target tissues. However, there are various barriers that must be fixed before it can be used clinically. For example, "naked" siRNA is susceptible to several obstacles that reduce its therapeutic efficacy.^[132] Additionally, once siRNA has entered the bloodstream, naked RNA can be degraded by serum nucleases and can stimulate the innate immune system.^[132] Due to its size and highly polyanionic (containing negative charges at several sites) nature, unmodified siRNA molecules cannot readily enter the cells through the cell membrane. Therefore, artificial or nanoparticle encapsulated siRNA must be used. If siRNA is transferred across the cell membrane, unintended toxicities can occur if therapeutic doses are not optimized, and siRNAs can exhibit off-target effects (e.g. unintended downregulation of genes with partial sequence complementarity).^[133] Even after entering the cells, repeated dosing is required since their effects are diluted at each cell division. In response to these potential issues and barriers, two approaches help facilitate siRNA delivery to target cells: lipid nanoparticles and conjugates.^[134]

Lipid nanoparticles

Lipid nanoparticles (LNPs) are based on liposome-like structures that are typically made of an aqueous center surrounded by a lipid shell.^[135] A subset of liposomal structures used for delivery drugs to tissues rest in large unilamellar vesicles (LUVs) which may be 100 nm in size. LNP delivery mechanisms have become an increasing source of encasing nucleic acids and may include plasmids, CRISPR and mRNA.^[136]

The first approved use of lipid nanoparticles as a drug delivery mechanism began in 2018 with the siRNA drug patisiran, developed by Alnylam Pharmaceuticals. Dicerna Pharmaceuticals, Persomics, Sanofi and Sirna Therapeutics also worked to bring RNAi therapies to market.^[137]^[138]

Other recent applications include two FDA approved COVID-19 vaccines: mRNA-1273, developed by Moderna. and BNT162b, developed by a collaboration between Pfizer and BioNtech.^[139] These two vaccines use lipid nanoparticles to deliver antigen mRNA. Encapsulating the mRNA molecule in lipid nanoparticles was a critical breakthrough for producing viable mRNA vaccines, solving a number of key technical barriers in delivering the mRNA molecule into the host cell as distributed through apolipoprotein E (apoE) in the low-density lipoprotein receptor (LDLR). In December 2020, Novartis announced that positive results from phase III efficacy studies deemed inclisiran was a treatment for heterozygous familial hypercholesterolemia (HeFH) and atherosclerotic cardiovascular disease (ASCVD).^[140]

Conjugates

In addition to LNPs, RNAi therapeutics have targeted delivery through siRNA conjugates (e.g., GalNAc, carbohydrates, peptides, aptamers, antibodies).^[141] Therapeutics using siRNA conjugates have been developed for rare or genetic diseases such as acute hepatic porphyria (AHP), hemophilia, primary hyperoxaluria (PH) and hereditary ATTR amyloidosis as well as other cardiometabolic diseases such as hypertension and non-alcoholic steatohepatitis (NASH).^[142]

Biotechnology

RNAi has been used for a variety of other applications including food, crops and insecticides. The use of the RNAi pathway has developed numerous products such as foods like Arctic apples, nicotine-free tobacco, decaffeinated coffee, nutrient fortified vegetation and hypoallergenic crops.^[143]^[144]^[145] The emerging use of RNAi has the potential to develop many other products for future use.

Viral infection

Antiviral treatment is one of the earliest proposed RNAi-based medical applications, and two different types have been developed. The first type is to target viral RNAs. Many studies have shown that targeting viral RNAs can suppress the replication of numerous viruses, including HIV,^[146] HPV,^[147] hepatitis A,^[148] hepatitis B,^[149] influenza virus,^[150]^[151]^[152]^[153] respiratory syncytial virus (RSV),^[153] SARS coronavirus (SARS-CoV),^[153] adenovirus^[153] and measles virus.^[154] The other strategy is to block the initial viral entries by targeting the host cell genes.^[155] For example, suppression of chemokine receptors (CXCR4 and CCR5) on host cells can prevent HIV viral entry.^[156]

Cancer

While traditional chemotherapy can effectively kill cancer cells, lack of specificity for discriminating normal cells and cancer cells in these treatments usually cause severe side effects. Numerous studies have demonstrated that RNAi can provide a more specific approach to inhibit tumor growth by targeting cancer-related genes (i.e., oncogene).^[157] It has also been proposed that RNAi can enhance the sensitivity of cancer cells to chemotherapeutic agents, providing a combinatorial therapeutic approach with chemotherapy.^[158] Another potential RNAi-based treatment is to inhibit cell invasion and migration.^[159]

Compared with chemotherapy or other anti-cancer drugs, there are a lot of advantages of siRNA drug.^[160] SiRNA acts on the post-transcriptional stage of gene expression, so it does not modify or change DNA in a deleterious effect.^[160] SiRNA can also be used to produce a specific response in a certain type of way, such as by downgrading suppression of gene expression.^[160] In a single cancer cell, siRNA can cause dramatic suppression of gene expression with just several copies.^[160] This happens by silencing cancer-promoting genes with RNAi, as well as targeting an mRNA sequence.^[160]

RNAi drugs treat cancer by silencing certain cancer promoting genes.^[160] This is done by complementing the cancer genes with the RNAi, such as keeping the mRNA sequences in accordance with the RNAi drug.^[160] Ideally, RNAi is should be injected and/or chemically modified so the RNAi can reach cancer cells more efficiently.^[160] RNAi uptake and regulation is controlled by the kidneys.^[160]

Neurological diseases

RNAi strategies also show potential for treating neurodegenerative diseases. Studies in cells and in mouse have shown that specifically targeting Amyloid beta-producing genes (e.g. BACE1 and APP) by RNAi can significantly reduced the amount of Aβ peptide which is correlated with the cause of Alzheimer's disease.^[161]^[162]^[163] In addition, this silencing-based approaches also provide promising results in treatment of Parkinson's disease and Polyglutamine disease.^[164]^[165]^[166]

Stimulation of immune response

The human immune system is divided into two separate branches: the innate immune system and the adaptive immune system.^[167] The innate immune system is the first defense against infection and responds to pathogens in a generic fashion.^[167] On the other hand, the adaptive immune system, a system that was evolved later than the innate, is composed mainly of highly specialized B and T cells that are trained to react to specific portions of pathogenic molecules.^[167]

The challenge between old pathogens and new has helped create a system of guarded cells and particles that are called safe framework.^[167] This framework has given humans an army of systems that search out and destroy invader particles, such as pathogens, microscopic organisms, parasites, and infections.^[167] The mammalian safe framework has developed to incorporate siRNA as a tool to indicate viral contamination, which has allowed siRNA is create an intense innate immune response.^[167]

siRNA is controlled by the innate immune system, which can be divided into the acute inflammatory responses and antiviral responses.^[167] The inflammatory response is created with signals from small signaling molecules, or cytokines.^[167] These include interleukin-1 (IL-1), interleukin-6 (IL-6), interleukin-12 (IL-12) and tumor necrosis factor α (TNF-α).^[167] The innate immune system generates inflammation and antiviral responses, which cause the release pattern recognition receptors (PRRs).^[167] These receptors help in labeling which pathogens are viruses, fungi, or bacteria.^[167] Moreover, the importance of siRNA and the innate immune system is to include more PRRs to help recognize different RNA structures.^[167] This makes it more likely for the siRNA to cause an immunostimulant response in the event of the pathogen.^[167]

Food

RNAi has been used to genetically engineer plants to produce lower levels of natural plant toxins. Such techniques take advantage of the stable and heritable RNAi phenotype in plant stocks. Cotton seeds are rich in dietary protein but naturally contain the toxic terpenoid product gossypol, making them unsuitable for human consumption. RNAi has been used to produce cotton stocks whose seeds contain reduced levels of delta-cadinene synthase, a key enzyme in gossypol production, without affecting the enzyme's production in other parts of the plant, where gossypol is itself important in preventing damage from plant pests.^[168]

Development efforts have successfully reduced the levels of allergens in tomato plants^[169] and fortification of plants such as tomatoes with dietary antioxidants.^[170] RNAi silencing of alpha-amylase have also been used to decrease Aspergillus flavus fungal growth in maize which would have otherwise contaminated the kernels with dangerous aflatoxins.^[171] Silencing lachrymatory factor synthase in onions have produced tearless onions and RNAi has been used in BP1 genes in rapeseeds to improve photosynthesis.^[172] SBEIIa and SBEIIb genes in wheat have been targeted in wheat in order to produce higher levels of amylose in order to improve bowel function,^[173] and Travella et al. 2006 employed RNAi for functional genomics an investigation of hexaploid bread races, while virus-induced gene silencing (VIGS, a subtype of RNAi) was used by Scofield et al. 2005 to investigate the mechanism of resistance provided by Lr21 against wheat leaf rust in hexaploid wheat.^[174]

Insecticides

RNAi is under development as an insecticide, employing multiple approaches, including genetic engineering and topical application.^[4] Cells in the midgut of some insects take up the dsRNA molecules in the process referred to as environmental RNAi.^[175] In some insects the effect is systemic as the signal spreads throughout the insect's body (referred to as systemic RNAi).^[176]

Animals exposed to RNAi at doses millions of times higher than anticipated human exposure levels show no adverse effects.^[177] RNAi has varying effects in different species of Lepidoptera (butterflies and moths).^[178]

Drosophila spp., Bombyx mori, Locusta spp., Spodoptera spp., Tribolium castaneum, Nilaparvata lugens, Helicoverpa armigera, and Apis mellifera are models that have been widely used to learn how RNAi works within particular taxa of insects. Musca domestica has two Ago2 genes and Glossina morsitans three, as found by Lewis et al. 2016 and Hain et al. 2010.^[179]^[180] In the case of the miRNA pathway Diuraphis noxia has two Ago1s, M. domestica two Dcr1s, Acyrthosiphon pisum two each of Ago1 and Loqs and Dcr1 and four of Pasha. While in the piRNA, G. morsitans and A. pisum have two or three Ago3s each.^[180] This has led to identification of future insecticide development targets, and the modes of action and reasons for insecticide resistance of other insecticides.^[180]

Transgenic plants

Transgenic crops have been made to express dsRNA, carefully chosen to silence crucial genes in target pests. These dsRNAs are designed to affect only insects that express specific gene sequences. As a proof of principle, in 2009 a study showed RNAs that could kill any one of four fruit fly species while not harming the other three.^[4]

Topical

Alternatively dsRNA can be supplied without genetic engineering. One approach is to add them to irrigation water. The molecules are absorbed into the plants' vascular system and poison insects feeding on them. Another approach involves spraying dsRNA like a conventional pesticide. This would allow faster adaptation to resistance. Such approaches would require low cost sources of dsRNAs that do not currently exist.^[4]

Functional genomics

Approaches to the design of genome-wide RNAi libraries can require more sophistication than the design of a single siRNA for a defined set of experimental conditions. Artificial neural networks are frequently used to design siRNA libraries^[181] and to predict their likely efficiency at gene knockdown.^[182] Mass genomic screening is widely seen as a promising method for genome annotation and has triggered the development of high-throughput screening methods based on microarrays.^[183]^[184]

Genome-scale screening

Genome-scale RNAi research relies on high-throughput screening (HTS) technology. RNAi HTS technology allows genome-wide loss-of-function screening and is broadly used in the identification of genes associated with specific phenotypes. This technology has been hailed as a potential second genomics wave, following the first genomics wave of gene expression microarray and single nucleotide polymorphism discovery platforms.^[185] One major advantage of genome-scale RNAi screening is its ability to simultaneously interrogate thousands of genes. With the ability to generate a large amount of data per experiment, genome-scale RNAi screening has led to an explosion of data generation rates. Exploiting such large data sets is a fundamental challenge, requiring suitable statistics/bioinformatics methods. The basic process of cell-based RNAi screening includes the choice of an RNAi library, robust and stable cell types, transfection with RNAi agents, treatment/incubation, signal detection, analysis and identification of important genes or therapeutical targets.^[186]

History

RNAi discovery

The process of RNAi was referred to as "co-suppression" and "quelling" when observed prior to the knowledge of an RNA-related mechanism. The discovery of RNAi was preceded first by observations of transcriptional inhibition by antisense RNA expressed in transgenic plants,^[188] and more directly by reports of unexpected outcomes in experiments performed by plant scientists in the United States and the Netherlands in the early 1990s.^[189] In an attempt to alter flower colors in petunias, researchers introduced additional copies of a gene encoding chalcone synthase, a key enzyme for flower pigmentation into petunia plants of normally pink or violet flower color. The overexpressed gene was expected to result in darker flowers, but instead caused some flowers to have less visible purple pigment, sometimes in variegated patterns, indicating that the activity of chalcone synthase had been substantially decreased or became suppressed in a context-specific manner. This would later be explained as the result of the transgene being inserted adjacent to promoters in the opposite direction in various positions throughout the genomes of some transformants, thus leading to expression of antisense transcripts and gene silencing when these promoters are active.^{[citation needed]} Another early observation of RNAi came from a study of the fungus Neurospora crassa,^[190] although it was not immediately recognized as related. Further investigation of the phenomenon in plants indicated that the downregulation was due to post-transcriptional inhibition of gene expression via an increased rate of mRNA degradation.^[191] This phenomenon was called co-suppression of gene expression, but the molecular mechanism remained unknown.^[192]

Not long after, plant virologists working on improving plant resistance to viral diseases observed a similar unexpected phenomenon. While it was known that plants expressing virus-specific proteins showed enhanced tolerance or resistance to viral infection, it was not expected that plants carrying only short, non-coding regions of viral RNA sequences would show similar levels of protection. Researchers believed that viral RNA produced by transgenes could also inhibit viral replication.^[193] The reverse experiment, in which short sequences of plant genes were introduced into viruses, showed that the targeted gene was suppressed in an infected plant.^[194] This phenomenon was labeled "virus-induced gene silencing" (VIGS),^[174] and the set of such phenomena were collectively called post transcriptional gene silencing.^[195]

After these initial observations in plants, laboratories searched for this phenomenon in other organisms.^[196]^[197] The first instance of RNA silencing in animals was documented in 1996, when Guo and Kemphues observed that, by introducing sense and antisense RNA to par-1 mRNA in Caenorhabditis elegans caused degradation of the par-1 message.^[198] It was thought that this degradation was triggered by single-stranded RNA (ssRNA), but two years later, in 1998, Fire and Mello discovered that this ability to silence the par-1 gene expression was actually triggered by double-stranded RNA (dsRNA).^[198] Craig C. Mello and Andrew Fire's 1998 Nature paper reported a potent gene silencing effect after injecting double stranded RNA into C. elegans.^[199] In investigating the regulation of muscle protein production, they observed that neither mRNA nor antisense RNA injections had an effect on protein production, but double-stranded RNA successfully silenced the targeted gene. As a result of this work, they coined the term RNAi. This discovery represented the first identification of the causative agent for the phenomenon. Fire and Mello were awarded the 2006 Nobel Prize in Physiology or Medicine.^[6]^[200]

RNAi therapeutics

Just after Fire and Mello's ground-breaking discovery, Elbashir et al. discovered, by using synthetically made small interfering RNA (siRNA), it was possible to target the silencing of specific sequences in a gene, rather than silencing the entire gene.^[201] Only a year later, McCaffrey and colleagues demonstrated that this sequence-specific silencing had therapeutic applications by targeting a sequence from the Hepatitis C virus in transgenic mice.^[202] Since then, multiple researchers have been attempting to expand the therapeutic applications of RNAi, specifically looking to target genes that cause various types of cancer.^[203]^[204] By 2006, the first applications to reach clinical trials were in the treatment of macular degeneration and respiratory syncytial virus.^[205] Four years later the first-in-human Phase I clinical trial was started, using a nanoparticle delivery system to target solid tumors.^[206]

The FDA approved the first siRNA-based drug (patisiran) in 2018. Givosiran and lumasiran later won FDA approval for the treatment of AHP and PH1 in 2019 and 2020, respectively.^[112] Inclisiran received EMA approval in 2020 for the treatment of high cholesterol and is currently under review by the FDA.^[207]

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External links

Overview of the RNAi process, from Cambridge University's The Naked Scientists
Animation of the RNAi process, from Nature
NOVA scienceNOW explains RNAi – A 15-minute video of the Nova broadcast that aired on PBS, 26 July 2005
Silencing Genomes Archived 10 August 2019 at the Wayback Machine RNA interference (RNAi) experiments and bioinformatics in C. elegans for education. From the Dolan DNA Learning Center of Cold Spring Harbor Laboratory.
RNAi screens in C. elegans in a 96-well liquid format and their application to the systematic identification of genetic interactions (a protocol)
2 American ‘Worm People’ Win Nobel for RNA Work, from NY Times
Molecular Therapy web focus: "The development of RNAi as a therapeutic strategy", a collection of free articles about RNAi as a therapeutic strategy.
GenomeRNAi: a database of phenotypes from RNA interference screening experiments in Drosophila melanogaster and Homo sapiens
RNAi tools Archived 19 June 2018 at the Wayback Machine Pre-designed and custom RNA Interference tools

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+{{cs1 config|name-list-style=vanc|display-authors=6}}
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-[[File:ShRNA Lentivirus.svg|thumb|upright=1.5|right|[[Lentiviral]] delivery of designed shRNAs and the mechanism of RNA interference in mammalian cells.]]
+[[File:ShRNA Lentivirus.svg|thumb|upright=1.5|right|[[Lentiviral]] delivery of designed shRNAs and the mechanism of RNA interference in mammalian cells]]
-'''RNA interference''' ('''RNAi''') is a biological process in which [[RNA]] molecules are involved in sequence-specific suppression of [[gene expression]] by double-stranded RNA, through translational or transcriptional repression. Historically, RNAi was known by other names, including ''co-suppression'', ''post-transcriptional gene silencing'' (PTGS), and ''quelling''. The detailed study of each of these seemingly different processes elucidated that the identity of these phenomena were all actually RNAi. [[Andrew Fire]] and [[Craig C. Mello]] shared the 2006 [[Nobel Prize in Physiology or Medicine]] for their work on RNAi in the [[nematode]] worm ''[[Caenorhabditis elegans]]'', which they published in 1998. Since the discovery of RNAi and its regulatory potentials, it has become evident that RNAi has immense potential in suppression of desired genes. RNAi is now known as precise, efficient, stable and better than [[antisense therapy]] for gene suppression.<ref name="Saurabh Satyajit, Vidyarthi AS, Prasad D 2014 543–564">{{cite journal | vauthors = Saurabh S, Vidyarthi AS, Prasad D |title=RNA interference: concept to reality in crop improvement |journal=[[Planta (journal)|Planta]] |volume=239 | issue = 3 | pages = 543–64 | date = March 2014 | pmid = 24402564 | doi = 10.1007/s00425-013-2019-5 | doi-access = free|bibcode=2014Plant.239..543S }}</ref> [[Antisense RNA]] produced intracellularly by an expression vector may be developed and find utility as novel therapeutic agents.<ref>{{cite journal | vauthors = Weiss B, Davidkova G, Zhou LW | s2cid = 9448271 | title = Antisense RNA gene therapy for studying and modulating biological processes | journal = Cellular and Molecular Life Sciences | volume = 55 | issue = 3 | pages = 334–58 | date = March 1999 | pmid = 10228554 | doi = 10.1007/s000180050296}}</ref>
+'''RNA interference''' ('''RNAi''') is a biological process in which [[RNA]] molecules are involved in sequence-specific suppression of [[gene expression]] by [[double-stranded RNA]], through translational or transcriptional repression. Historically, RNAi was known by other names, including ''co-suppression'', ''post-transcriptional gene silencing'' (PTGS), and ''quelling''. The detailed study of each of these seemingly different processes elucidated that the identity of these phenomena were all actually RNAi. [[Andrew Fire]] and [[Craig C. Mello]] shared the 2006 [[Nobel Prize in Physiology or Medicine]] for their work on RNAi in the [[nematode]] worm ''[[Caenorhabditis elegans]]'', which they published in 1998. Since the discovery of RNAi and its regulatory potentials, it has become evident that RNAi has immense potential in suppression of desired genes. RNAi is now known as precise, efficient, stable and better than [[antisense therapy]] for gene suppression.<ref name="Saurabh Satyajit, Vidyarthi AS, Prasad D 2014 543–564">{{cite journal | vauthors = Saurabh S, Vidyarthi AS, Prasad D |title=RNA interference: concept to reality in crop improvement |journal=[[Planta (journal)|Planta]] |volume=239 | issue = 3 | pages = 543–64 | date = March 2014 | pmid = 24402564 | doi = 10.1007/s00425-013-2019-5 | doi-access = free|bibcode=2014Plant.239..543S }}</ref> [[Antisense RNA]] produced intracellularly by an expression vector may be developed and find utility as novel therapeutic agents.<ref>{{cite journal | vauthors = Weiss B, Davidkova G, Zhou LW | s2cid = 9448271 | title = Antisense RNA gene therapy for studying and modulating biological processes | journal = Cellular and Molecular Life Sciences | volume = 55 | issue = 3 | pages = 334–58 | date = March 1999 | pmid = 10228554 | doi = 10.1007/s000180050296| pmc = 11146801 }}</ref>
-Two types of small [[ribonucleic acid]] (RNA) molecules, [[microRNA]] (miRNA) and [[small interfering RNA]] ([[Small interfering RNA|siRNA]]), are central to components to the RNAi pathway. Once mRNA is degraded, post-transcriptional silencing occurs as protein translation is prevented. Transcription can be inhibited via the pre-transcriptional silencing mechanism of RNAi, through which an enzyme complex catalyzes DNA methylation at genomic positions complementary to complexed siRNA or miRNA. RNAi has an important role in defending cells against parasitic [[nucleotide]] sequences (e.g., [[virus|viruses]] or [[transposon|transposons]]) and also influences development of organisms.
+Two types of small [[ribonucleic acid]] (RNA) molecules, [[microRNA]] (miRNA) and [[small interfering RNA]] ([[Small interfering RNA|siRNA]]), are central to components to the RNAi pathway. Once mRNA is degraded, post-transcriptional silencing occurs as protein translation is prevented. Transcription can be inhibited via the pre-transcriptional silencing mechanism of RNAi, through which an enzyme complex catalyzes DNA methylation at genomic positions complementary to complexed siRNA or miRNA. RNAi has an important role in defending cells against parasitic [[nucleotide]] sequences (e.g., [[virus]]es or [[transposon]]s) and also influences development of organisms.
 The RNAi pathway is a naturally occurring process found in many [[eukaryotes]] and animal cells. It is initiated by the enzyme [[Dicer]], which cleaves long [[double-stranded RNA]] (dsRNA) [[molecules]] into short double-stranded fragments of approximately 21 to 23 [[nucleotide]] siRNAs. Each siRNA is unwound into two single-stranded RNAs (ssRNAs), the passenger (sense) strand and the guide (antisense) strand. The passenger strand is then cleaved by the protein [[Argonaute|Argonaute 2]] (Ago2). The passenger strand is degraded and the guide strand is incorporated into the [[RNA-induced silencing complex]] (RISC). The RISC assembly then binds and degrades the target mRNA. Specifically, this is accomplished when the guide strand pairs with a complementary sequence in a mRNA molecule and induces cleavage by Ago2, a catalytic component of the RISC. In some organisms, this process spreads systemically, despite the initially limited molar concentrations of siRNA.<ref name=Matranga/>
@@ Line 12: / Line 13: @@
 == Cellular mechanism ==
-[[File:2ffl-by-domain.png|thumb|right|The [[Dicer]] protein from ''[[Giardia intestinalis]]'', which catalyzes the cleavage of dsRNA to siRNAs. The [[RNase]] domains are colored green, the PAZ domain yellow, the platform domain red, and the connector helix blue.<ref name="Macrae">{{cite journal | vauthors = Macrae IJ, Zhou K, Li F, Repic A, Brooks AN, Cande WZ, Adams PD, Doudna JA | title = Structural basis for double-stranded RNA processing by Dicer | journal = Science | volume = 311 | issue = 5758 | pages = 195–8 | date = January 2006 | pmid = 16410517 | doi = 10.1126/science.1121638 | bibcode = 2006Sci...311..195M | s2cid = 23785494 }}</ref>]]
+[[File:2ffl-by-domain.png|thumb|right|The [[Dicer]] protein from ''[[Giardia]]'', which catalyzes the cleavage of dsRNA to siRNAs. The [[RNase]] domains are colored green, the PAZ domain yellow, the platform domain red, and the connector helix blue.<ref name="Macrae">{{cite journal | vauthors = Macrae IJ, Zhou K, Li F, Repic A, Brooks AN, Cande WZ, Adams PD, Doudna JA | title = Structural basis for double-stranded RNA processing by Dicer | journal = Science | volume = 311 | issue = 5758 | pages = 195–8 | date = January 2006 | pmid = 16410517 | doi = 10.1126/science.1121638 | bibcode = 2006Sci...311..195M | s2cid = 23785494 }}</ref>]]
 RNAi is an RNA-dependent [[gene silencing]] process that is controlled by [[RNA-induced silencing complex|RISC]] and is initiated by short double-stranded RNA molecules in a cell's cytoplasm, where they interact with the catalytic RISC component [[Argonaute]].<ref name="Daneholt2006">{{cite web | title=Advanced Information: RNA interference |  vauthors = Daneholt B | website=The Nobel Prize in Physiology or Medicine 2006 | url=http://nobelprize.org/nobel_prizes/medicine/laureates/2006/adv.html | access-date=25 January 2007 | archive-url=https://web.archive.org/web/20070120113455/http://nobelprize.org/nobel_prizes/medicine/laureates/2006/adv.html <!-- Added by H3llBot --> | archive-date=20 January 2007}}</ref> When the dsRNA is exogenous (coming from infection by a virus with an RNA genome or laboratory manipulations), the RNA is imported directly into the [[cytoplasm]] and cleaved to short fragments by Dicer. The initiating dsRNA can also be endogenous (originating in the cell), as in pre-microRNAs expressed from [[non-coding RNA|RNA-coding gene]]s in the genome. The primary transcripts from such genes are first processed to form the characteristic [[stem-loop]] structure of pre-miRNA in the [[cell nucleus|nucleus]], then exported to the cytoplasm. Thus, the two dsRNA pathways, exogenous and endogenous, converge at the RISC.<ref name="pmid15614608">{{cite journal | vauthors = Bagasra O, Prilliman KR | s2cid = 2966105 | title = RNA interference: the molecular immune system | journal = Journal of Molecular Histology | volume = 35 | issue = 6 | pages = 545–53 | date = August 2004 | pmid = 15614608 | doi = 10.1007/s10735-004-2192-8 | citeseerx = 10.1.1.456.1701}}</ref>
@@ Line 22: / Line 23: @@
 Exogenous dsRNA is detected and bound by an effector protein, known as RDE-4 in ''[[Caenorhabditis elegans|C. elegans]]'' and R2D2 in ''[[Drosophila]]'', that stimulates Dicer activity.<ref name="Parker">{{cite journal |vauthors=Parker G, Eckert D, Bass B |title=RDE-4 preferentially binds long dsRNA and its dimerization is necessary for cleavage of dsRNA to siRNA |journal=RNA |volume=12 |issue=5 |pages=807–18 |year=2006 |pmid=16603715 |doi=10.1261/rna.2338706 |pmc=1440910}}</ref> The mechanism producing this length specificity is unknown and this protein only binds long dsRNAs.<ref name="Parker"/>
-In ''C. elegans'' this initiation response is amplified through the synthesis of a population of 'secondary' [[Small interfering RNA|siRNAs]] during which the Dicer-produced initiating or 'primary' [[Small interfering RNA|siRNAs]] are used as templates.<ref name="Baulcombe D 2007 199–200">{{cite journal | vauthors = Baulcombe DC | s2cid = 46285020 | title = Molecular biology. Amplified silencing | journal = Science | volume = 315 | issue = 5809 | pages = 199–200 | date = January 2007 | pmid = 17218517 | doi = 10.1126/science.1138030}}</ref> These 'secondary' [[Small interfering RNA|siRNAs]] are structurally distinct from Dicer-produced [[Small interfering RNA|siRNAs]] and appear to be produced by an [[RNA-dependent RNA polymerase]] (RdRP).<ref name="Pak">{{cite journal | vauthors = Pak J, Fire A | s2cid = 46620298 | title = Distinct populations of primary and secondary effectors during RNAi in C. elegans | journal = Science | volume = 315 | issue = 5809 | pages = 241–4 | date = January 2007 | pmid = 17124291 | doi = 10.1126/science.1132839 | bibcode = 2007Sci...315..241P }}</ref><ref name="Sijen">{{cite journal | vauthors = Sijen T, Steiner FA, Thijssen KL, Plasterk RH | title = Secondary siRNAs result from unprimed RNA synthesis and form a distinct class | journal = Science | volume = 315 | issue = 5809 | pages = 244–7 | date = January 2007 | pmid = 17158288 | doi = 10.1126/science.1136699 | bibcode = 2007Sci...315..244S | s2cid = 9483460}}</ref>
+In ''C. elegans'' this initiation response is amplified through the synthesis of a population of 'secondary' [[Small interfering RNA|siRNAs]] during which the Dicer-produced initiating or 'primary' [[Small interfering RNA|siRNAs]] are used as templates.<ref name="Baulcombe D 2007 199–200">{{cite journal | vauthors = Baulcombe DC | s2cid = 46285020 | title = Molecular biology. Amplified silencing | journal = Science | volume = 315 | issue = 5809 | pages = 199–200 | date = January 2007 | pmid = 17218517 | doi = 10.1126/science.1138030}}</ref> These 'secondary' [[Small interfering RNA|siRNAs]] are structurally distinct from Dicer-produced [[Small interfering RNA|siRNAs]] and appear to be produced by an [[RNA-dependent RNA polymerase]] (RdRP).<ref name="Pak">{{cite journal | vauthors = Pak J, Fire A | s2cid = 46620298 | title = Distinct populations of primary and secondary effectors during RNAi in C. elegans | journal = Science | volume = 315 | issue = 5809 | pages = 241–4 | date = January 2007 | pmid = 17124291 | doi = 10.1126/science.1132839 | bibcode = 2007Sci...315..241P }}</ref>
 === MicroRNA ===
-[[File:Microrna secondary structure.png|thumb|right|The [[stem-loop]] [[secondary structure]] of a pre-[[microRNA]] from ''[[Brassica oleracea]]''.]]
+[[File:Microrna secondary structure.png|thumb|right|The [[stem-loop]] [[secondary structure]] of a pre-[[microRNA]] from ''[[Brassica oleracea]]'']]
 {{main|MicroRNA}}
 [[MicroRNA]]s (miRNAs) are [[genome|genomically]] encoded [[non-coding RNA]]s that help regulate [[gene expression]], particularly during [[developmental biology|development]].<ref name="Zhao Y 2007">{{cite journal | vauthors = Wang QL, Li ZH | s2cid = 23014413 | title = The functions of microRNAs in plants | journal = Frontiers in Bioscience | volume = 12 | pages = 3975–82 | date = May 2007 | pmid = 17485351 | doi = 10.2741/2364 | pmc = 2851543}}<br/>{{cite journal | vauthors = Zhao Y, Srivastava D | title = A developmental view of microRNA function | journal = Trends in Biochemical Sciences | volume = 32 | issue = 4 | pages = 189–97 | date = April 2007 | pmid = 17350266 | doi = 10.1016/j.tibs.2007.02.006}}</ref> The phenomenon of RNAi, broadly defined, includes the endogenously induced gene silencing effects of miRNAs as well as silencing triggered by foreign dsRNA. Mature miRNAs are structurally similar to [[Small interfering RNA|siRNAs]] produced from exogenous dsRNA, but before reaching maturity, miRNAs must first undergo extensive [[post-transcriptional modification]]. A miRNA is expressed from a much longer RNA-coding gene as a [[primary transcript]] known as a ''pri-miRNA'' which is processed, in the [[cell nucleus]], to a 70-nucleotide [[stem-loop]] structure called a ''pre-miRNA'' by the [[microprocessor complex]]. This complex consists of an [[RNase III]] enzyme called [[Drosha]] and a dsRNA-binding protein [[DGCR8]]. The dsRNA portion of this pre-miRNA is bound and cleaved by Dicer to produce the mature miRNA molecule that can be integrated into the RISC complex; thus, miRNA and [[Small interfering RNA|siRNA]] share the same downstream cellular machinery.<ref name="Denli">{{cite book | vauthors = Gregory RI, Chendrimada TP, Shiekhattar R | title = MicroRNA Protocols | chapter = MicroRNA biogenesis: isolation and characterization of the microprocessor complex | series = Methods in Molecular Biology | volume = 342 | pages = 33–47 | year = 2006 | pmid = 16957365 | doi = 10.1385/1-59745-123-1:33 | isbn = 978-1-59745-123-9}}</ref> First, viral encoded miRNA was described in [[Epstein–Barr virus]] (EBV).<ref name="Pfeffer 734–736">{{cite journal | vauthors = Pfeffer S, Zavolan M, Grässer FA, Chien M, Russo JJ, Ju J, John B, Enright AJ, Marks D, Sander C, Tuschl T | s2cid = 25287167 | title = Identification of virus-encoded microRNAs | journal = Science | volume = 304 | issue = 5671 | pages = 734–6 | date = April 2004 | pmid = 15118162 | doi = 10.1126/science.1096781 | bibcode = 2004Sci...304..734P }}</ref> Thereafter, an increasing number of microRNAs have been described in viruses. VIRmiRNA is a comprehensive catalogue covering viral microRNA, their targets and anti-viral miRNAs<ref name="Qureshi bau103">{{cite journal | vauthors = Qureshi A, Thakur N, Monga I, Thakur A, Kumar M | title = VIRmiRNA: a comprehensive resource for experimentally validated viral miRNAs and their targets | journal = Database | volume = 2014 | pages = bau103 | date = 1 January 2014 | pmid = 25380780 | pmc = 4224276 | doi = 10.1093/database/bau103}}</ref> (see also VIRmiRNA resource: [http://crdd.osdd.net/servers/virmirna/ http://crdd.osdd.net/servers/virmirna/).]
@@ Line 49: / Line 50: @@
 Exogenous dsRNA is detected and bound by an effector protein, known as RDE-4 in ''[[Caenorhabditis elegans|C. elegans]]'' and R2D2 in ''[[Drosophila]]'', that stimulates Dicer activity.<ref name="Parker"/> This protein only binds long dsRNAs, but the mechanism producing this length specificity is unknown.<ref name="Parker"/> This RNA-binding protein then facilitates the transfer of cleaved [[Small interfering RNA|siRNAs]] to the RISC complex.<ref name="Liu2003">{{cite journal | vauthors = Liu Q, Rand TA, Kalidas S, Du F, Kim HE, Smith DP, Wang X | title = R2D2, a bridge between the initiation and effector steps of the Drosophila RNAi pathway | journal = Science | volume = 301 | issue = 5641 | pages = 1921–5 | date = September 2003 | pmid = 14512631 | doi = 10.1126/science.1088710 | bibcode = 2003Sci...301.1921L | s2cid = 41436233}}</ref>
-In ''C. elegans'' this initiation response is amplified through the synthesis of a population of 'secondary' [[Small interfering RNA|siRNAs]] during which the Dicer-produced initiating or 'primary' [[Small interfering RNA|siRNAs]] are used as templates.<ref name="Baulcombe D 2007 199–200"/> These 'secondary' [[Small interfering RNA|siRNAs]] are structurally distinct from Dicer-produced [[Small interfering RNA|siRNAs]] and appear to be produced by an [[RNA-dependent RNA polymerase]] (RdRP).<ref name="Pak"/><ref name="Sijen"/>
+In ''C. elegans'' this initiation response is amplified through the synthesis of a population of 'secondary' [[Small interfering RNA|siRNAs]] during which the Dicer-produced initiating or 'primary' [[Small interfering RNA|siRNAs]] are used as templates.<ref name="Baulcombe D 2007 199–200"/> These 'secondary' [[Small interfering RNA|siRNAs]] are structurally distinct from Dicer-produced [[Small interfering RNA|siRNAs]] and appear to be produced by an [[RNA-dependent RNA polymerase]] (RdRP).<ref name="Pak"/>
-[[File:RNAi overview.png|thumb|400x400px|[[small RNA]] Biogenesis: primary miRNAs (pri-miRNAs) are transcribed in the nucleus and fold back onto themselves as hairpins that are then trimmed in the nucleus by a [[microprocessor complex]] to form a ~60-70nt hairpin pre-RNA. This pre-miRNA is transported through the [[Nuclear pore|nuclear pore complex]] (NPC) into the cytoplasm, where [[Dicer]] further trims it to a ~20nt miRNA duplex (pre-siRNAs also enter the pathway at this step). This duplex is then loaded into Ago to form the “pre-RISC(RNA induced silencing complex)” and the passenger strand is released to form active [[RNA-induced silencing complex|RISC]].{{imagefact|date=December 2022}}]]
+[[File:RNAi overview.png|thumb|400x400px|[[small RNA]] Biogenesis: primary miRNAs (pri-miRNAs) are transcribed in the nucleus and fold back onto themselves as hairpins that are then trimmed in the nucleus by a [[microprocessor complex]] to form a ~60-70nt hairpin pre-RNA. This pre-miRNA is transported through the [[Nuclear pore|nuclear pore complex]] (NPC) into the cytoplasm, where [[Dicer]] further trims it to a ~20nt miRNA duplex (pre-siRNAs also enter the pathway at this step). This duplex is then loaded into Ago to form the “pre-RISC(RNA induced silencing complex)” and the passenger strand is released to form active [[RNA-induced silencing complex|RISC]].{{image reference needed|date=December 2022}}]]
-<nowiki/><nowiki/><nowiki/>[[File:Argonaute 1u04 1ytu composite.png|thumb|left|300px|''Left:'' A full-length [[Argonaute]] protein from the [[archaea]] species ''[[Pyrococcus furiosus]]''. ''Right:'' The [[PIWI domain]] of an [[Argonaute]] protein in complex with [[double-stranded RNA]].]]
+[[File:Argonaute 1u04 1ytu composite.png|thumb|left|300px|''Left:'' A full-length [[Argonaute]] protein from the [[archaea]] species ''[[Pyrococcus furiosus]]''. ''Right:'' The [[PIWI domain]] of an [[Argonaute]] protein in complex with [[double-stranded RNA]].]]
-The active components of an RNA-induced silencing complex (RISC) are [[endonuclease]]s called Argonaute proteins, which cleave the target mRNA strand [[complementarity (molecular biology)|complementary]] to their bound [[Small interfering RNA|siRNA]].<ref name="Daneholt2006"/> As the fragments produced by Dicer are double-stranded, they could each in theory produce a functional [[Small interfering RNA|siRNA]]. However, only one of the two strands, which is known as the ''guide strand'', binds Argonaute and directs gene silencing. The other ''anti-guide strand'' or ''passenger strand'' is degraded during RISC activation.<ref name="Gregory">{{cite journal | vauthors = Gregory RI, Chendrimada TP, Cooch N, Shiekhattar R | title = Human RISC couples microRNA biogenesis and posttranscriptional gene silencing | journal = Cell | volume = 123 | issue = 4 | pages = 631–40 | date = November 2005 | pmid = 16271387 | doi = 10.1016/j.cell.2005.10.022 | doi-access = free}}</ref> Although it was first believed that an [[adenosine triphosphate|ATP]]-dependent [[helicase]] separated these two strands,<ref name=Lodish>{{cite book | title = Molecular Cell Biology | edition = 5th | publisher = WH Freeman: New York, NY | year = 2004 | url = https://archive.org/details/molecularcellbio00harv | isbn = 978-0-7167-4366-8 | vauthors = Lodish H, Berk A, Matsudaira P, Kaiser CA, Krieger M, Scott MP, Zipurksy SL, Darnell J | url-access = registration}}</ref> the process proved to be ATP-independent and performed directly by the protein components of RISC.<ref name="Matranga">{{cite journal | vauthors = Matranga C, Tomari Y, Shin C, Bartel DP, Zamore PD | title = Passenger-strand cleavage facilitates assembly of siRNA into Ago2-containing RNAi enzyme complexes | journal = Cell | volume = 123 | issue = 4 | pages = 607–20 | date = November 2005 | pmid = 16271386 | doi = 10.1016/j.cell.2005.08.044 | doi-access = free}}</ref><ref name="Leuschner">{{cite journal | vauthors = Leuschner PJ, Ameres SL, Kueng S, Martinez J | title = Cleavage of the siRNA passenger strand during RISC assembly in human cells | journal = EMBO Reports | volume = 7 | issue = 3 | pages = 314–20 | date = March 2006 | pmid = 16439995 | pmc = 1456892 | doi = 10.1038/sj.embor.7400637}}</ref> However, an ''in vitro'' kinetic analysis of RNAi in the presence and absence of ATP showed that ATP may be required to unwind and remove the cleaved mRNA strand from the RISC complex after catalysis.<ref name="Haley 2004 599–606">{{cite journal | vauthors = Haley B, Zamore PD | title = Kinetic analysis of the RNAi enzyme complex | journal = Nature Structural & Molecular Biology | volume = 11 | issue = 7 | pages = 599–606 | date = July 2004 | pmid = 15170178 | doi = 10.1038/nsmb780 | s2cid = 12400060}}</ref> The guide strand tends to be the one whose [[5' end|5′ end]] is less stably paired to its complement,<ref>{{cite journal | vauthors = Schwarz DS, Hutvágner G, Du T, Xu Z, Aronin N, Zamore PD | title = Asymmetry in the assembly of the RNAi enzyme complex | journal = Cell | volume = 115 | issue = 2 | pages = 199–208 | date = October 2003 | pmid = 14567917 | doi = 10.1016/S0092-8674(03)00759-1 | doi-access = free}}</ref> but strand selection is unaffected by the direction in which Dicer cleaves the dsRNA before RISC incorporation.<ref name="Preall">{{cite journal | vauthors = Preall JB, He Z, Gorra JM, Sontheimer EJ | title = Short interfering RNA strand selection is independent of dsRNA processing polarity during RNAi in Drosophila | journal = Current Biology | volume = 16 | issue = 5 | pages = 530–5 | date = March 2006 | pmid = 16527750 | doi = 10.1016/j.cub.2006.01.061 | doi-access = free}}</ref> Instead, the R2D2 protein may serve as the differentiating factor by binding the more-stable 5′ end of the passenger strand.<ref name="Tomari">{{cite journal | vauthors = Tomari Y, Matranga C, Haley B, Martinez N, Zamore PD | title = A protein sensor for siRNA asymmetry | journal = Science | volume = 306 | issue = 5700 | pages = 1377–80 | date = November 2004 | pmid = 15550672 | doi = 10.1126/science.1102755 | bibcode = 2004Sci...306.1377T | s2cid = 31558409}}</ref>
+The active components of an RNA-induced silencing complex (RISC) are [[endonuclease]]s called Argonaute proteins, which cleave the target mRNA strand [[complementarity (molecular biology)|complementary]] to their bound [[Small interfering RNA|siRNA]].<ref name="Daneholt2006"/> As the fragments produced by Dicer are double-stranded, they could each in theory produce a functional [[Small interfering RNA|siRNA]]. However, only one of the two strands, which is known as the ''guide strand'', binds Argonaute and directs gene silencing. The other ''anti-guide strand'' or ''passenger strand'' is degraded during RISC activation.<ref name="Gregory">{{cite journal | vauthors = Gregory RI, Chendrimada TP, Cooch N, Shiekhattar R | title = Human RISC couples microRNA biogenesis and posttranscriptional gene silencing | journal = Cell | volume = 123 | issue = 4 | pages = 631–40 | date = November 2005 | pmid = 16271387 | doi = 10.1016/j.cell.2005.10.022 | doi-access = free}}</ref> Although it was first believed that an [[adenosine triphosphate|ATP]]-dependent [[helicase]] separated these two strands,<ref name=Lodish>{{cite book | title = Molecular Cell Biology | edition = 5th | publisher = WH Freeman: New York, NY | year = 2004 | url = https://archive.org/details/molecularcellbio00harv | isbn = 978-0-7167-4366-8 | vauthors = Lodish H, Berk A, Matsudaira P, Kaiser CA, Krieger M, Scott MP, Zipurksy SL, Darnell J | url-access = registration}}</ref> the process proved to be ATP-independent and performed directly by the protein components of RISC.<ref name="Matranga">{{cite journal | vauthors = Matranga C, Tomari Y, Shin C, Bartel DP, Zamore PD | title = Passenger-strand cleavage facilitates assembly of siRNA into Ago2-containing RNAi enzyme complexes | journal = Cell | volume = 123 | issue = 4 | pages = 607–20 | date = November 2005 | pmid = 16271386 | doi = 10.1016/j.cell.2005.08.044 | doi-access = free}}</ref><ref name="Leuschner">{{cite journal | vauthors = Leuschner PJ, Ameres SL, Kueng S, Martinez J | title = Cleavage of the siRNA passenger strand during RISC assembly in human cells | journal = EMBO Reports | volume = 7 | issue = 3 | pages = 314–20 | date = March 2006 | pmid = 16439995 | pmc = 1456892 | doi = 10.1038/sj.embor.7400637}}</ref> However, an ''in vitro'' kinetic analysis of RNAi in the presence and absence of ATP showed that ATP may be required to unwind and remove the cleaved mRNA strand from the RISC complex after catalysis.<ref name="Haley 2004 599–606">{{cite journal | vauthors = Haley B, Zamore PD | title = Kinetic analysis of the RNAi enzyme complex | journal = Nature Structural & Molecular Biology | volume = 11 | issue = 7 | pages = 599–606 | date = July 2004 | pmid = 15170178 | doi = 10.1038/nsmb780 | s2cid = 12400060}}</ref> The guide strand tends to be the one whose [[5' end|5′ end]] is less stably paired to its complement,<ref>{{cite journal | vauthors = Schwarz DS, Hutvágner G, Du T, Xu Z, Aronin N, Zamore PD | title = Asymmetry in the assembly of the RNAi enzyme complex | journal = Cell | volume = 115 | issue = 2 | pages = 199–208 | date = October 2003 | pmid = 14567917 | doi = 10.1016/S0092-8674(03)00759-1 | doi-access = free}}</ref> but strand selection is unaffected by the direction in which Dicer cleaves the dsRNA before RISC incorporation.<ref name="Preall">{{cite journal | vauthors = Preall JB, He Z, Gorra JM, Sontheimer EJ | title = Short interfering RNA strand selection is independent of dsRNA processing polarity during RNAi in Drosophila | journal = Current Biology | volume = 16 | issue = 5 | pages = 530–5 | date = March 2006 | pmid = 16527750 | doi = 10.1016/j.cub.2006.01.061 | doi-access = free| bibcode = 2006CBio...16..530P }}</ref> Instead, the R2D2 protein may serve as the differentiating factor by binding the more-stable 5′ end of the passenger strand.<ref name="Tomari">{{cite journal | vauthors = Tomari Y, Matranga C, Haley B, Martinez N, Zamore PD | title = A protein sensor for siRNA asymmetry | journal = Science | volume = 306 | issue = 5700 | pages = 1377–80 | date = November 2004 | pmid = 15550672 | doi = 10.1126/science.1102755 | bibcode = 2004Sci...306.1377T | s2cid = 31558409}}</ref>
 The structural basis for binding of RNA to the Argonaute protein was examined by [[X-ray crystallography]] of the binding [[structural domain|domain]] of an RNA-bound Argonaute. Here, the [[phosphorylation|phosphorylated]] 5′ end of the RNA strand enters a [[Conserved sequence|conserved]] [[basicity|basic]] surface [[binding site|pocket]] and makes contacts through a [[divalent]] [[cation]] (an atom with two positive charges) such as [[magnesium]] and by [[aromaticity|aromatic]] [[stacking (chemistry)|stacking]] (a process that allows more than one atom to share an electron by passing it back and forth) between the 5′ nucleotide in the [[Small interfering RNA|siRNA]] and a conserved [[tyrosine]] residue. This site is thought to form a nucleation site for the binding of the [[Small interfering RNA|siRNA]] to its mRNA target.<ref name="Ma">{{cite journal | vauthors = Ma JB, Yuan YR, Meister G, Pei Y, Tuschl T, Patel DJ | title = Structural basis for 5'-end-specific recognition of guide RNA by the A. fulgidus Piwi protein | journal = Nature | volume = 434 | issue = 7033 | pages = 666–70 | date = March 2005 | pmid = 15800629 | pmc = 4694588 | doi = 10.1038/nature03514 | bibcode = 2005Natur.434..666M}}</ref> Analysis of the inhibitory effect of mismatches in either the 5’ or 3’ end of the guide strand has demonstrated that the 5’ end of the guide strand is likely responsible for matching and binding the target mRNA, while the 3’ end is responsible for physically arranging target mRNA into a cleavage-favorable RISC region.<ref name="Haley 2004 599–606"/>
@@ Line 68: / Line 69: @@
 The type of [[RNA editing]] that is most prevalent in higher eukaryotes converts [[adenosine]] nucleotides into [[inosine]] in dsRNAs via the enzyme [[adenosine deaminase]] (ADAR).<ref name=Bass_2002>{{cite journal | vauthors = Bass BL | title = RNA editing by adenosine deaminases that act on RNA | journal = Annual Review of Biochemistry | volume = 71 | pages = 817–46 | year = 2002 | pmid = 12045112 | pmc = 1823043 | doi = 10.1146/annurev.biochem.71.110601.135501}}</ref> It was originally proposed in 2000 that the RNAi and A→I RNA editing pathways might compete for a common dsRNA substrate.<ref name=Bass_2000>{{cite journal | vauthors = Bass BL | title = Double-stranded RNA as a template for gene silencing | journal = Cell | volume = 101 | issue = 3 | pages = 235–8 | date = April 2000 | pmid = 10847677 | doi = 10.1016/S0092-8674(02)71133-1 | doi-access = free}}</ref> Some pre-miRNAs do undergo A→I RNA editing<ref name=Luciano_2004>{{cite journal | vauthors = Luciano DJ, Mirsky H, Vendetti NJ, Maas S | title = RNA editing of a miRNA precursor | journal = RNA | volume = 10 | issue = 8 | pages = 1174–7 | date = August 2004 | pmid = 15272117 | pmc = 1370607 | doi = 10.1261/rna.7350304}}</ref><ref name="yang2006">{{cite journal | vauthors = Yang W, Chendrimada TP, Wang Q, Higuchi M, Seeburg PH, Shiekhattar R, Nishikura K | title = Modulation of microRNA processing and expression through RNA editing by ADAR deaminases | journal = Nature Structural & Molecular Biology | volume = 13 | issue = 1 | pages = 13–21 | date = January 2006 | pmid = 16369484 | pmc = 2950615 | doi = 10.1038/nsmb1041}}</ref> and this mechanism may regulate the processing and expression of mature miRNAs.<ref name="yang2006" /> Furthermore, at least one mammalian ADAR can sequester [[Small interfering RNA|siRNAs]] from RNAi pathway components.<ref name=Yang_2005>{{cite journal | vauthors = Yang W, Wang Q, Howell KL, Lee JT, Cho DS, Murray JM, Nishikura K | title = ADAR1 RNA deaminase limits short interfering RNA efficacy in mammalian cells | journal = The Journal of Biological Chemistry | volume = 280 | issue = 5 | pages = 3946–53 | date = February 2005 | pmid = 15556947 | pmc = 2947832 | doi = 10.1074/jbc.M407876200| doi-access = free }}</ref> Further support for this model comes from studies on ADAR-null ''C. elegans'' strains indicating that A→I RNA editing may counteract RNAi silencing of endogenous genes and transgenes.<ref name=Nishikura_2006>{{cite journal | vauthors = Nishikura K | title = Editor meets silencer: crosstalk between RNA editing and RNA interference | journal = Nature Reviews Molecular Cell Biology | volume = 7 | issue = 12 | pages = 919–31 | date = December 2006 | pmid = 17139332 | pmc = 2953463 | doi = 10.1038/nrm2061}}</ref>
-[[File:Rnai diagram retrovirology.png|thumb|300px|right|Illustration of the major differences between plant and animal gene silencing. Natively expressed [[microRNA]] or exogenous [[small interfering RNA]] is processed by [[Dicer]] and integrated into the [[RISC]] complex, which mediates gene silencing.<ref name="Saumet">{{cite journal |vauthors=Saumet A, Lecellier CH |year=2006 |title= Anti-viral RNA silencing: do we look like plants ?|journal=Retrovirology |volume=3 |issue=1 |pmid=16409629 |doi= 10.1186/1742-4690-3-3 |page= 3 |pmc=1363733 |doi-access=free }}</ref>]]
+[[File:Rnai diagram retrovirology.png|thumb|300px|right|Illustration of the major differences between plant and animal gene silencing. Natively expressed [[microRNA]] or exogenous [[small interfering RNA]] is processed by [[Dicer]] and integrated into the [[RNA-induced silencing complex|RISC]] complex, which mediates gene silencing.<ref name="Saumet">{{cite journal |vauthors=Saumet A, Lecellier CH |year=2006 |title= Anti-viral RNA silencing: do we look like plants ?|journal=Retrovirology |volume=3 |issue=1 |pmid=16409629 |doi= 10.1186/1742-4690-3-3 |page= 3 |pmc=1363733 |doi-access=free }}</ref>]]
 === Variation among organisms ===
-Organisms vary in their ability to take up foreign dsRNA and use it in the RNAi pathway. The effects of RNAi can be both systemic and heritable in plants and ''C. elegans'', although not in ''Drosophila'' or mammals. In plants, RNAi is thought to propagate by the transfer of [[Small interfering RNA|siRNAs]] between cells through [[plasmodesmata]] (channels in the cell walls that enable communication and transport).<ref name=Lodish /> Heritability comes from [[DNA methylation|methylation]] of promoters targeted by RNAi; the new methylation pattern is copied in each new generation of the cell.<ref>{{cite journal | vauthors = Jones L, Ratcliff F, Baulcombe DC | s2cid = 16789197 | title = RNA-directed transcriptional gene silencing in plants can be inherited independently of the RNA trigger and requires Met1 for maintenance | journal = Current Biology | volume = 11 | issue = 10 | pages = 747–57 | date = May 2001 | pmid = 11378384 | doi = 10.1016/S0960-9822(01)00226-3| doi-access = free }}</ref> A broad general distinction between plants and animals lies in the targeting of endogenously produced miRNAs; in plants, miRNAs are usually perfectly or nearly perfectly complementary to their target genes and induce direct mRNA cleavage by RISC, while animals' miRNAs tend to be more divergent in sequence and induce translational repression.<ref name="Saumet" /> This translational effect may be produced by inhibiting the interactions of translation [[eukaryotic initiation factor|initiation factor]]s with the mRNA's [[polyadenylation|polyadenine tail]].<ref name="Humphreys">{{cite journal | vauthors = Humphreys DT, Westman BJ, Martin DI, Preiss T | title = MicroRNAs control translation initiation by inhibiting eukaryotic initiation factor 4E/cap and poly(A) tail function | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 102 | issue = 47 | pages = 16961–6 | date = November 2005 | pmid = 16287976 | pmc = 1287990 | doi = 10.1073/pnas.0506482102 | bibcode = 2005PNAS..10216961H| doi-access = free }}</ref>
+Organisms vary in their ability to take up foreign dsRNA and use it in the RNAi pathway. The effects of RNAi can be both systemic and heritable in plants and ''C. elegans'', although not in ''Drosophila'' or mammals. In plants, RNAi is thought to propagate by the transfer of [[Small interfering RNA|siRNAs]] between cells through [[plasmodesmata]] (channels in the cell walls that enable communication and transport).<ref name=Lodish /> Heritability comes from [[DNA methylation|methylation]] of promoters targeted by RNAi; the new methylation pattern is copied in each new generation of the cell.<ref>{{cite journal | vauthors = Jones L, Ratcliff F, Baulcombe DC | s2cid = 16789197 | title = RNA-directed transcriptional gene silencing in plants can be inherited independently of the RNA trigger and requires Met1 for maintenance | journal = Current Biology | volume = 11 | issue = 10 | pages = 747–57 | date = May 2001 | pmid = 11378384 | doi = 10.1016/S0960-9822(01)00226-3| doi-access = free | bibcode = 2001CBio...11..747J }}</ref> A broad general distinction between plants and animals lies in the targeting of endogenously produced miRNAs; in plants, miRNAs are usually perfectly or nearly perfectly complementary to their target genes and induce direct mRNA cleavage by RISC, while animals' miRNAs tend to be more divergent in sequence and induce translational repression.<ref name="Saumet" /> This translational effect may be produced by inhibiting the interactions of translation [[eukaryotic initiation factor|initiation factor]]s with the mRNA's [[polyadenylation|polyadenine tail]].<ref name="Humphreys">{{cite journal | vauthors = Humphreys DT, Westman BJ, Martin DI, Preiss T | title = MicroRNAs control translation initiation by inhibiting eukaryotic initiation factor 4E/cap and poly(A) tail function | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 102 | issue = 47 | pages = 16961–6 | date = November 2005 | pmid = 16287976 | pmc = 1287990 | doi = 10.1073/pnas.0506482102 | bibcode = 2005PNAS..10216961H| doi-access = free }}</ref>
-Some eukaryotic [[protozoa]] such as ''[[Leishmania major]]'' and ''[[Trypanosoma cruzi]]'' lack the RNAi pathway entirely.<ref name=DaRocha_2004>{{cite journal | vauthors = DaRocha WD, Otsu K, Teixeira SM, Donelson JE | title = Tests of cytoplasmic RNA interference (RNAi) and construction of a tetracycline-inducible T7 promoter system in Trypanosoma cruzi | journal = Molecular and Biochemical Parasitology | volume = 133 | issue = 2 | pages = 175–86 | date = February 2004 | pmid = 14698430 | doi = 10.1016/j.molbiopara.2003.10.005}}</ref><ref name=Robinson_2003>{{cite journal | vauthors = Robinson KA, Beverley SM | title = Improvements in transfection efficiency and tests of RNA interference (RNAi) approaches in the protozoan parasite Leishmania | journal = Molecular and Biochemical Parasitology | volume = 128 | issue = 2 | pages = 217–28 | date = May 2003 | pmid = 12742588 | doi = 10.1016/S0166-6851(03)00079-3}}</ref> Most or all of the components are also missing in some [[fungi]], most notably the [[model organism]] ''[[Saccharomyces cerevisiae]]''.<ref name="Aravind">{{cite journal | vauthors = Aravind L, Watanabe H, Lipman DJ, Koonin EV | title = Lineage-specific loss and divergence of functionally linked genes in eukaryotes | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 97 | issue = 21 | pages = 11319–24 | date = October 2000 | pmid = 11016957 | pmc = 17198 | doi = 10.1073/pnas.200346997 | bibcode = 2000PNAS...9711319A| doi-access = free }}</ref> The presence of RNAi in other budding yeast species such as ''[[Saccharomyces castellii]]'' and ''[[Candida albicans]]'', further demonstrates that inducing two RNAi-related proteins from ''S. castellii'' facilitates RNAi in ''S. cerevisiae''.<ref>{{cite journal | vauthors = Drinnenberg IA, Weinberg DE, Xie KT, Mower JP, Wolfe KH, Fink GR, Bartel DP | title = RNAi in budding yeast | journal = Science | volume = 326 | issue = 5952 | pages = 544–550 | date = October 2009 | pmid = 19745116 | pmc = 3786161 | doi = 10.1126/science.1176945 | bibcode = 2009Sci...326..544D}}</ref> That certain [[ascomycete]]s and [[basidiomycete]]s are missing RNAi pathways indicates that proteins required for RNA silencing have been lost independently from many fungal [[Lineage (evolution)|lineages]], possibly due to the evolution of a novel pathway with similar function, or to the lack of selective advantage in certain [[Ecological niche|niche]]s.<ref name="Nakayashiki">{{cite journal | vauthors = Nakayashiki H, Kadotani N, Mayama S | s2cid = 22639035 | title = Evolution and diversification of RNA silencing proteins in fungi | journal = Journal of Molecular Evolution | volume = 63 | issue = 1 | pages = 127–35 | date = July 2006 | pmid = 16786437 | doi = 10.1007/s00239-005-0257-2 | bibcode = 2006JMolE..63..127N | url = http://www.lib.kobe-u.ac.jp/repository/90000024.pdf}}</ref>
+Some eukaryotic [[protozoa]] such as ''[[Leishmania major]]'' and ''[[Trypanosoma cruzi]]'' lack the RNAi pathway entirely.<ref name=DaRocha_2004>{{cite journal | vauthors = DaRocha WD, Otsu K, Teixeira SM, Donelson JE | title = Tests of cytoplasmic RNA interference (RNAi) and construction of a tetracycline-inducible T7 promoter system in Trypanosoma cruzi | journal = Molecular and Biochemical Parasitology | volume = 133 | issue = 2 | pages = 175–86 | date = February 2004 | pmid = 14698430 | doi = 10.1016/j.molbiopara.2003.10.005}}</ref><ref name=Robinson_2003>{{cite journal | vauthors = Robinson KA, Beverley SM | title = Improvements in transfection efficiency and tests of RNA interference (RNAi) approaches in the protozoan parasite Leishmania | journal = Molecular and Biochemical Parasitology | volume = 128 | issue = 2 | pages = 217–28 | date = May 2003 | pmid = 12742588 | doi = 10.1016/S0166-6851(03)00079-3}}</ref> Most or all of the components are also missing in some [[fungi]], most notably the [[model organism]] ''[[Saccharomyces cerevisiae]]''.<ref name="Aravind">{{cite journal | vauthors = Aravind L, Watanabe H, Lipman DJ, Koonin EV | title = Lineage-specific loss and divergence of functionally linked genes in eukaryotes | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 97 | issue = 21 | pages = 11319–24 | date = October 2000 | pmid = 11016957 | pmc = 17198 | doi = 10.1073/pnas.200346997 | bibcode = 2000PNAS...9711319A| doi-access = free }}</ref> The presence of RNAi in other budding yeast species such as ''[[Saccharomyces castellii]]'' and ''[[Candida albicans]]'', further demonstrates that inducing two RNAi-related proteins from ''S. castellii'' facilitates RNAi in ''S. cerevisiae''.<ref>{{cite journal | vauthors = Drinnenberg IA, Weinberg DE, Xie KT, Mower JP, Wolfe KH, Fink GR, Bartel DP | title = RNAi in budding yeast | journal = Science | volume = 326 | issue = 5952 | pages = 544–550 | date = October 2009 | pmid = 19745116 | pmc = 3786161 | doi = 10.1126/science.1176945 | bibcode = 2009Sci...326..544D}}</ref> That certain [[ascomycete]]s and [[basidiomycete]]s are missing RNAi pathways indicates that proteins required for RNA silencing have been lost independently from many fungal [[Lineage (evolution)|lineages]], possibly due to the evolution of a novel pathway with similar function, or to the lack of selective advantage in certain [[Ecological niche|niche]]s.<ref name="Nakayashiki">{{cite journal | vauthors = Nakayashiki H, Kadotani N, Mayama S | s2cid = 22639035 | title = Evolution and diversification of RNA silencing proteins in fungi | journal = Journal of Molecular Evolution | volume = 63 | issue = 1 | pages = 127–35 | date = July 2006 | pmid = 16786437 | doi = 10.1007/s00239-005-0257-2 | bibcode = 2006JMolE..63..127N | url = http://www.lib.kobe-u.ac.jp/repository/90000024.pdf | access-date = 4 December 2019 | archive-date = 6 August 2020 | archive-url = https://web.archive.org/web/20200806175438/http://www.lib.kobe-u.ac.jp/repository/90000024.pdf | url-status = dead }}</ref>
 === Related prokaryotic systems ===
@@ Line 81: / Line 82: @@
 == Biological functions ==
-{{outdated section|date=May 2020}}
+{{update section|date=May 2020}}
 === Immunity ===
@@ Line 89: / Line 90: @@
 Although animals generally express fewer variants of the Dicer enzyme than plants, RNAi in some animals produces an antiviral response. In both juvenile and adult ''Drosophila'', RNAi is important in antiviral [[innate immunity]] and is active against pathogens such as [[Drosophila X virus]].<ref name="Zambon">{{cite journal | vauthors = Zambon RA, Vakharia VN, Wu LP | title = RNAi is an antiviral immune response against a dsRNA virus in Drosophila melanogaster | journal = Cellular Microbiology | volume = 8 | issue = 5 | pages = 880–9 | date = May 2006 | pmid = 16611236 | doi = 10.1111/j.1462-5822.2006.00688.x| s2cid = 32439482 | doi-access = free }}</ref><ref name="Wang">{{cite journal | vauthors = Wang XH, Aliyari R, Li WX, Li HW, Kim K, Carthew R, Atkinson P, Ding SW | title = RNA interference directs innate immunity against viruses in adult Drosophila | journal = Science | volume = 312 | issue = 5772 | pages = 452–4 | date = April 2006 | pmid = 16556799 | pmc = 1509097 | doi = 10.1126/science.1125694 | bibcode = 2006Sci...312..452W}}</ref> A similar role in immunity may operate in ''C. elegans'', as Argonaute proteins are upregulated in response to viruses and worms that overexpress components of the RNAi pathway are resistant to viral infection.<ref name="Lu">{{cite journal | vauthors = Lu R, Maduro M, Li F, Li HW, Broitman-Maduro G, Li WX, Ding SW | title = Animal virus replication and RNAi-mediated antiviral silencing in Caenorhabditis elegans | journal = Nature | volume = 436 | issue = 7053 | pages = 1040–1043 | date = August 2005 | pmid = 16107851 | pmc = 1388260 | doi = 10.1038/nature03870 | bibcode = 2005Natur.436.1040L}}</ref><ref name="Wilkins">{{cite journal | vauthors = Wilkins C, Dishongh R, Moore SC, Whitt MA, Chow M, Machaca K | title = RNA interference is an antiviral defence mechanism in Caenorhabditis elegans | journal = Nature | volume = 436 | issue = 7053 | pages = 1044–7 | date = August 2005 | pmid = 16107852 | doi = 10.1038/nature03957 | bibcode = 2005Natur.436.1044W | s2cid = 4431035}}</ref>
-The role of RNAi in mammalian innate immunity is poorly understood, and relatively little data is available. However, the existence of viruses that encode genes able to suppress the RNAi response in mammalian cells may be evidence in favour of an RNAi-dependent mammalian immune response,<ref name="Berkhout">{{cite journal | vauthors = Berkhout B, Haasnoot J | title = The interplay between virus infection and the cellular RNA interference machinery | journal = FEBS Letters | volume = 580 | issue = 12 | pages = 2896–902 | date = May 2006 | pmid = 16563388 | doi = 10.1016/j.febslet.2006.02.070 | pmc = 7094296}}</ref><ref name="Schutz">{{cite journal | vauthors = Schütz S, Sarnow P | title = Interaction of viruses with the mammalian RNA interference pathway | journal = Virology | volume = 344 | issue = 1 | pages = 151–7 | date = January 2006 | pmid = 16364746 | doi = 10.1016/j.virol.2005.09.034 | doi-access = free}}</ref> although this hypothesis has been challenged as poorly substantiated.<ref name="Cullen">{{cite journal | vauthors = Cullen BR | s2cid = 23467688 | title = Is RNA interference involved in intrinsic antiviral immunity in mammals? | journal = Nature Immunology | volume = 7 | issue = 6 | pages = 563–7 | date = June 2006 | pmid = 16715068 | doi = 10.1038/ni1352}}</ref>
+The role of RNAi in mammalian innate immunity is poorly understood, and relatively little data is available. However, the existence of viruses that encode genes able to suppress the RNAi response in mammalian cells may be evidence in favour of an RNAi-dependent mammalian immune response,<ref name="Berkhout">{{cite journal | vauthors = Berkhout B, Haasnoot J | title = The interplay between virus infection and the cellular RNA interference machinery | journal = FEBS Letters | volume = 580 | issue = 12 | pages = 2896–902 | date = May 2006 | pmid = 16563388 | doi = 10.1016/j.febslet.2006.02.070 | pmc = 7094296| bibcode = 2006FEBSL.580.2896B }}</ref><ref name="Schutz">{{cite journal | vauthors = Schütz S, Sarnow P | title = Interaction of viruses with the mammalian RNA interference pathway | journal = Virology | volume = 344 | issue = 1 | pages = 151–7 | date = January 2006 | pmid = 16364746 | doi = 10.1016/j.virol.2005.09.034 | doi-access = free}}</ref> although this hypothesis has been challenged as poorly substantiated.<ref name="Cullen">{{cite journal | vauthors = Cullen BR | s2cid = 23467688 | title = Is RNA interference involved in intrinsic antiviral immunity in mammals? | journal = Nature Immunology | volume = 7 | issue = 6 | pages = 563–7 | date = June 2006 | pmid = 16715068 | doi = 10.1038/ni1352}}</ref>
 Evidence for the existence of a functional antiviral RNAi pathway in mammalian cells has been presented.<ref>{{cite journal | vauthors = Maillard PV, Ciaudo C, Marchais A, Li Y, Jay F, Ding SW, Voinnet O | title = Antiviral RNA interference in mammalian cells | journal = Science | volume = 342 | issue = 6155 | pages = 235–8 | date = October 2013 | pmid = 24115438 | pmc = 3853215 | doi = 10.1126/science.1241930 | bibcode = 2013Sci...342..235M}}</ref><ref>{{cite journal | vauthors = Li Y, Lu J, Han Y, Fan X, Ding SW | title = RNA interference functions as an antiviral immunity mechanism in mammals | journal = Science | volume = 342 | issue = 6155 | pages = 231–4 | date = October 2013 | pmid = 24115437 | pmc = 3875315 | doi = 10.1126/science.1241911 | bibcode = 2013Sci...342..231L}}</ref>
-Other functions for RNAi in mammalian viruses also exist, such as miRNAs expressed by the [[Herpes simplex virus|herpes virus]] that may act as [[heterochromatin]] organization triggers to mediate viral latency.<ref name="Li3">{{cite journal | vauthors = Li HW, Ding SW | title = Antiviral silencing in animals | journal = FEBS Letters | volume = 579 | issue = 26 | pages = 5965–73 | date = October 2005 | pmid = 16154568 | pmc = 1350842 | doi = 10.1016/j.febslet.2005.08.034}}</ref>
+Other functions for RNAi in mammalian viruses also exist, such as miRNAs expressed by the [[Herpes simplex virus|herpes virus]] that may act as [[heterochromatin]] organization triggers to mediate viral latency.<ref name="Li3">{{cite journal | vauthors = Li HW, Ding SW | title = Antiviral silencing in animals | journal = FEBS Letters | volume = 579 | issue = 26 | pages = 5965–73 | date = October 2005 | pmid = 16154568 | pmc = 1350842 | doi = 10.1016/j.febslet.2005.08.034| bibcode = 2005FEBSL.579.5965L }}</ref>
 === Downregulation of genes ===
@@ Line 115: / Line 116: @@
 === Medications ===
-[[File:Drosophila melanogaster - side (aka).jpg|thumb|left|250px|A normal adult ''[[Drosophila melanogaster|Drosophila]]'' fly, a common model organism used in RNAi experiments.]]
+[[File:Drosophila melanogaster - side (aka).jpg|thumb|left|250px|A normal adult ''[[Drosophila melanogaster|Drosophila]]'' fly, a common model organism used in RNAi experiments]]
 [[File:Vertical Timeline of RNAi's Use in Medicine.pdf|thumb|upright=1.5|Timeline of the use of RNAi in medicine between 1996 and 2017]]
-The technique of knocking down genes using RNAi therapeutics has demonstrated success in randomized controlled clinical studies. These medications are a growing class of siRNA-based drugs that decrease the expression of proteins encoded by certain genes. To date, five RNAi medications have been approved by regulatory authorities in the US and Europe: [[patisiran]] (2018), [[givosiran]] (2019), [[lumasiran]] (2020), [[inclisiran]] (2020 in Europe with anticipated US approval in 2021), and [[vutrisiran]] (2022).<ref name=Patisiran>{{cite journal |title=Patisiran, an RNAi Therapeutic, for Hereditary Transthyretin Amyloidosis |journal=New England Journal of Medicine|date=July 5, 2018 |doi=10.1056/NEJMoa1716153 |pmid=29972753|last1=Adams |first1=David |last2=Gonzalez-Duarte |first2=Alejandra |last3=O'Riordan |first3=William D. |last4=Yang |first4=Chih-Chao |last5=Ueda |first5=Mitsuharu |last6=Kristen |first6=Arnt V. |last7=Tournev |first7=Ivailo |last8=Schmidt |first8=Hartmut H. |last9=Coelho |first9=Teresa |last10=Berk |first10=John L. |last11=Lin |first11=Kon-Ping |last12=Vita |first12=Giuseppe |last13=Attarian |first13=Shahram |last14=Planté-Bordeneuve |first14=Violaine |last15=Mezei |first15=Michelle M. |last16=Campistol |first16=Josep M. |last17=Buades |first17=Juan |last18=Brannagan |first18=Thomas H. |last19=Kim |first19=Byoung J. |last20=Oh |first20=Jeeyoung |last21=Parman |first21=Yesim |last22=Sekijima |first22=Yoshiki |last23=Hawkins |first23=Philip N. |last24=Solomon |first24=Scott D. |last25=Polydefkis |first25=Michael |last26=Dyck |first26=Peter J. |last27=Gandhi |first27=Pritesh J. |last28=Goyal |first28=Sunita |last29=Chen |first29=Jihong |last30=Strahs |first30=Andrew L. |volume=379 |issue=1 |pages=11–21 |s2cid=205102839 |display-authors=1 |doi-access=free |hdl=2445/138257 |hdl-access=free }}</ref><ref>{{Cite journal |last1=Balwani |first1=Manisha |last2=Sardh |first2=Eliane |last3=Ventura |first3=Paolo |last4=Peiró |first4=Paula Aguilera |last5=Rees |first5=David C. |last6=Stölzel |first6=Ulrich |last7=Bissell |first7=D. Montgomery |last8=Bonkovsky |first8=Herbert L. |last9=Windyga |first9=Jerzy |last10=Anderson |first10=Karl E. |last11=Parker |first11=Charles |last12=Silver |first12=Samuel M. |last13=Keel |first13=Siobán B. |last14=Wang |first14=Jiaan-Der |last15=Stein |first15=Penelope E. |date=2020-06-11 |title=Phase 3 Trial of RNAi Therapeutic Givosiran for Acute Intermittent Porphyria |url=http://www.nejm.org/doi/10.1056/NEJMoa1913147 |journal=New England Journal of Medicine |language=en |volume=382 |issue=24 |pages=2289–2301 |doi=10.1056/NEJMoa1913147 |pmid=32521132 |issn=0028-4793}}</ref><ref>{{Cite journal |last1=Garrelfs |first1=Sander F. |last2=Frishberg |first2=Yaacov |last3=Hulton |first3=Sally A. |last4=Koren |first4=Michael J. |last5=O'Riordan |first5=William D. |last6=Cochat |first6=Pierre |last7=Deschênes |first7=Georges |last8=Shasha-Lavsky |first8=Hadas |last9=Saland |first9=Jeffrey M. |last10=Van't Hoff |first10=William G. |last11=Fuster |first11=Daniel G. |last12=Magen |first12=Daniella |last13=Moochhala |first13=Shabbir H. |last14=Schalk |first14=Gesa |last15=Simkova |first15=Eva |date=2021-04-01 |title=Lumasiran, an RNAi Therapeutic for Primary Hyperoxaluria Type 1 |url=https://pubmed.ncbi.nlm.nih.gov/33789010/ |journal=The New England Journal of Medicine |volume=384 |issue=13 |pages=1216–1226 |doi=10.1056/NEJMoa2021712 |issn=1533-4406 |pmid=33789010}}</ref><ref>{{Cite journal |last1=Adams |first1=David |last2=Tournev |first2=Ivailo L. |last3=Taylor |first3=Mark S. |last4=Coelho |first4=Teresa |last5=Planté-Bordeneuve |first5=Violaine |last6=Berk |first6=John L. |last7=González-Duarte |first7=Alejandra |last8=Gillmore |first8=Julian D. |last9=Low |first9=Soon-Chai |last10=Sekijima |first10=Yoshiki |last11=Obici |first11=Laura |last12=Chen |first12=Chongshu |last13=Badri |first13=Prajakta |last14=Arum |first14=Seth M. |last15=Vest |first15=John |date=March 2023 |title=Efficacy and safety of vutrisiran for patients with hereditary transthyretin-mediated amyloidosis with polyneuropathy: a randomized clinical trial |url=https://pubmed.ncbi.nlm.nih.gov/35875890/ |journal=Amyloid: The International Journal of Experimental and Clinical Investigation: The Official Journal of the International Society of Amyloidosis |volume=30 |issue=1 |pages=18–26 |doi=10.1080/13506129.2022.2091985 |issn=1744-2818 |pmid=35875890}}</ref>
+The technique of knocking down genes using RNAi therapeutics has demonstrated success in randomized controlled clinical studies. These medications are a growing class of siRNA-based drugs that decrease the expression of proteins encoded by certain genes. To date, five RNAi medications have been approved by regulatory authorities in the US and Europe: [[patisiran]] (2018), [[givosiran]] (2019), [[lumasiran]] (2020), [[inclisiran]] (2020 in Europe with anticipated US approval in 2021), and [[vutrisiran]] (2022).<ref name=Patisiran>{{cite journal |title=Patisiran, an RNAi Therapeutic, for Hereditary Transthyretin Amyloidosis |journal=New England Journal of Medicine|date=July 5, 2018 |doi=10.1056/NEJMoa1716153 |pmid=29972753|last1=Adams |first1=David |last2=Gonzalez-Duarte |first2=Alejandra |last3=O'Riordan |first3=William D. |last4=Yang |first4=Chih-Chao |last5=Ueda |first5=Mitsuharu |last6=Kristen |first6=Arnt V. |last7=Tournev |first7=Ivailo |last8=Schmidt |first8=Hartmut H. |last9=Coelho |first9=Teresa |last10=Berk |first10=John L. |last11=Lin |first11=Kon-Ping |last12=Vita |first12=Giuseppe |last13=Attarian |first13=Shahram |last14=Planté-Bordeneuve |first14=Violaine |last15=Mezei |first15=Michelle M. |last16=Campistol |first16=Josep M. |last17=Buades |first17=Juan |last18=Brannagan |first18=Thomas H. |last19=Kim |first19=Byoung J. |last20=Oh |first20=Jeeyoung |last21=Parman |first21=Yesim |last22=Sekijima |first22=Yoshiki |last23=Hawkins |first23=Philip N. |last24=Solomon |first24=Scott D. |last25=Polydefkis |first25=Michael |last26=Dyck |first26=Peter J. |last27=Gandhi |first27=Pritesh J. |last28=Goyal |first28=Sunita |last29=Chen |first29=Jihong |last30=Strahs |first30=Andrew L. |volume=379 |issue=1 |pages=11–21 |s2cid=205102839 |doi-access=free |hdl=2445/138257 |hdl-access=free }}</ref><ref>{{Cite journal |last1=Balwani |first1=Manisha |last2=Sardh |first2=Eliane |last3=Ventura |first3=Paolo |last4=Peiró |first4=Paula Aguilera |last5=Rees |first5=David C. |last6=Stölzel |first6=Ulrich |last7=Bissell |first7=D. Montgomery |last8=Bonkovsky |first8=Herbert L. |last9=Windyga |first9=Jerzy |last10=Anderson |first10=Karl E. |last11=Parker |first11=Charles |last12=Silver |first12=Samuel M. |last13=Keel |first13=Siobán B. |last14=Wang |first14=Jiaan-Der |last15=Stein |first15=Penelope E. |date=2020-06-11 |title=Phase 3 Trial of RNAi Therapeutic Givosiran for Acute Intermittent Porphyria |url=http://www.nejm.org/doi/10.1056/NEJMoa1913147 |journal=New England Journal of Medicine |language=en |volume=382 |issue=24 |pages=2289–2301 |doi=10.1056/NEJMoa1913147 |pmid=32521132 |issn=0028-4793}}</ref><ref>{{Cite journal |last1=Garrelfs |first1=Sander F. |last2=Frishberg |first2=Yaacov |last3=Hulton |first3=Sally A. |last4=Koren |first4=Michael J. |last5=O'Riordan |first5=William D. |last6=Cochat |first6=Pierre |last7=Deschênes |first7=Georges |last8=Shasha-Lavsky |first8=Hadas |last9=Saland |first9=Jeffrey M. |last10=Van't Hoff |first10=William G. |last11=Fuster |first11=Daniel G. |last12=Magen |first12=Daniella |last13=Moochhala |first13=Shabbir H. |last14=Schalk |first14=Gesa |last15=Simkova |first15=Eva |date=2021-04-01 |title=Lumasiran, an RNAi Therapeutic for Primary Hyperoxaluria Type 1 |url=https://pubmed.ncbi.nlm.nih.gov/33789010/ |journal=The New England Journal of Medicine |volume=384 |issue=13 |pages=1216–1226 |doi=10.1056/NEJMoa2021712 |issn=1533-4406 |pmid=33789010}}</ref><ref>{{Cite journal |last1=Adams |first1=David |last2=Tournev |first2=Ivailo L. |last3=Taylor |first3=Mark S. |last4=Coelho |first4=Teresa |last5=Planté-Bordeneuve |first5=Violaine |last6=Berk |first6=John L. |last7=González-Duarte |first7=Alejandra |last8=Gillmore |first8=Julian D. |last9=Low |first9=Soon-Chai |last10=Sekijima |first10=Yoshiki |last11=Obici |first11=Laura |last12=Chen |first12=Chongshu |last13=Badri |first13=Prajakta |last14=Arum |first14=Seth M. |last15=Vest |first15=John |date=March 2023 |title=Efficacy and safety of vutrisiran for patients with hereditary transthyretin-mediated amyloidosis with polyneuropathy: a randomized clinical trial |url=https://pubmed.ncbi.nlm.nih.gov/35875890/ |journal=Amyloid|volume=30 |issue=1 |pages=18–26 |doi=10.1080/13506129.2022.2091985 |issn=1744-2818 |pmid=35875890|hdl=1959.4/unsworks_80922 |hdl-access=free }}</ref>
 While all of the current regulatory body approved RNAi therapeutics focus on diseases that originate in the liver, additional medications under investigation target a host of disease areas including cardiovascular diseases, bleeding disorders, alcohol use disorders, cystic fibrosis, gout, carcinoma, and eye disorders.
-[[Patisiran]] is the first double stranded siRNA-based medication approved in 2018 and developed by [[Alnylam Pharmaceuticals]]. Patisiran uses the RNAi cascade to suppress the gene that codes for TTR (transthryetin). Mutations in this gene may cause the misfolding of a protein responsible for [[Transthyretin amyloidosis|hereditary ATTR amyloidosis]]. To achieve therapeutic response, patirisan is encased by a [[Solid lipid nanoparticle|lipid nanoparticle]] membrane that facilitates crossover into the cytoplasm. Once inside the cell, the siRNA begins processing by the enzyme Dicer. Patirisan is administered by a healthcare professional through an intravenous infusion with dosing based on body weight. Warnings and precautions include risk of infusion-related reactions and reduced vitamin A levels (serum).<ref>{{cite news|url=https://dailymed.nlm.nih.gov/dailymed/drugInfo.cfm?setid=e87ec36f-b4b4-49d4-aea4-d4ffb09b0970&audience=consumer|date=May 10, 2021|title=LABEL:ONPATTRO- patisiran injection, lipid complex|work=Daily Med}}</ref>
+[[Patisiran]] is the first double stranded siRNA-based medication approved in 2018 and developed by [[Alnylam Pharmaceuticals]]. Patisiran uses the RNAi cascade to suppress the gene that codes for TTR (transthryetin). Mutations in this gene may cause the misfolding of a protein responsible for [[Transthyretin amyloidosis|hereditary ATTR amyloidosis]]. To achieve therapeutic response, patisiran is encased by a [[Solid lipid nanoparticle|lipid nanoparticle]] membrane that facilitates crossover into the cytoplasm. Once inside the cell, the siRNA begins processing by the enzyme Dicer. Patisiran is administered by a healthcare professional through an intravenous infusion with dosing based on body weight. Warnings and precautions include risk of infusion-related reactions and reduced vitamin A levels (serum).<ref>{{cite news|url=https://dailymed.nlm.nih.gov/dailymed/drugInfo.cfm?setid=e87ec36f-b4b4-49d4-aea4-d4ffb09b0970&audience=consumer|date=May 10, 2021|title=LABEL:ONPATTRO- patisiran injection, lipid complex|work=Daily Med}}</ref>
 In 2019, the FDA and EMA approved [[givosiran]] for the treatment of adults with [[Hepatic porphyria|acute hepatic porphyria]] (AHP).<ref>{{cite news|url=https://www.accessdata.fda.gov/drugsatfda_docs/nda/2019/212194Orig1s000TOC.cfm|title=Drug Approval Package: GIVLAARI (givosiran)Injection|work=FDA}}</ref> The FDA also granted [[givosiran]] a [[breakthrough therapy]] designation, [[priority review]] designation, and [[orphan drug]] designation for the treatment of acute hepatic porphyria (AHP) in November 2019.<ref>{{cite journal |last1=Gonzalez-Aseguinolaza|first1=Gloria|title=Givosiran — Running RNA Interference to Fight Porphyria Attacks |journal=New England Journal of Medicine|date=June 11, 2020|volume=382 |issue=28–4793|pages=2366–2367 |doi=10.1056/NEJMe2010986|pmid=32521139 |s2cid=219592948 }}</ref> By 2020, givosiran received EMA approval.<ref>{{cite news|url=https://www.ema.europa.eu/en/medicines/human/EPAR/givlaari|title=Givlaari|work=European Medicines Agency}}</ref> Givosiran is an siRNA that breaks down [[aminolevulinic acid synthase]] 1 (ALAS1) mRNA in the liver. Breaking down ALAS1 mRNA prevents toxins (responsible for neurovisceral attacks and AHP disease) such as [[aminolevulinic acid]] (ALA) and [[porphobilinogen]] (PBG) from accumulating.<ref>{{cite journal |last1=Puy |first1=H |last2=Gouya |first2=L |last3=Deybach |first3=JC |title=Porphyrias |journal=Lancet|date=2010 |volume=375 |issue=9718 |pages=924–937 |doi=10.1016/S0140-6736(09)61925-5 |pmid=20226990|s2cid=208791867 }}</ref><ref>{{cite journal |last1=Simon |first1=A |last2=Pompilus |first2=F |last3=Querbes |first3=W |last4=Wei |first4=A |last5=Strzok |first5=S |last6=Penz |first6=C |last7=Howe |first7=DL |last8=Hungate |first8=JR |last9=Kim |first9=JB |last10=Agarwal |first10=S |last11=Marquis |first11=P |title=Patient Perspective on Acute Intermittent Porphyria with Frequent Attacks: A Disease with Intermittent and Chronic Manifestations|journal=Patient |date=2018|volume=11 |issue=5 |pages=527–537 | doi=10.1007/s40271-018-0319-3|pmid=29915990|pmc=6132435 }}</ref><ref>{{cite journal |last1=Pischik |first1=E |last2=Kauppinen |first2=R |title=An update of clinical management of acute intermittent porphyria |journal=Application of Clinical Genetics |date=2015 |volume=8 |pages=201–214 |doi=10.2147/TACG.S48605 |pmid=26366103|pmc=4562648 |doi-access=free }}</ref><ref>{{cite journal |last1=Bissell |first1=DM |last2=Lai |first2=JC |last3=Meister |first3=RK |last4=Blanc |first4=PD |title=Role of delta-aminolevulinic acid in the symptoms of acute porphyria |journal=The American Journal of Medicine |date=2015 |volume=128 |issue=3 |pages=313–317 |doi=10.1016/j.amjmed.2014.10.026 |pmid=25446301|pmc=4339446 }}</ref> To facilitate entry into the cytoplasm, givosiran uses [[GalNAc|GalNAc ligands]] and enters into liver cells. The medication is administered subcutaneously by a healthcare professional with dosing based on body weight. Warnings and precautions include risk of anaphylactic reactions, hepatic toxicity, renal toxicity and injection site reactions.<ref>{{cite news|url=https://dailymed.nlm.nih.gov/dailymed/drugInfo.cfm?setid=167e663c-11e1-497b-a3fc-951d65d58eaa|date=April 5, 2022|title=LABEL:GIVLAARI- givosiran sodium injection, solution|work=Daily Med}}</ref>
 [[Lumasiran]] was approved as a siRNA-based medication in 2020 for use in both the European Union and the United States.<ref>{{cite news|url=https://www.accessdata.fda.gov/drugsatfda_docs/nda/2020/214103Orig1s000TOC.cfm|title=Drug Approval Package: OXLUMO|work=FDA}}</ref><ref>{{cite news|url=https://www.ema.europa.eu/en/medicines/human/EPAR/oxlumo|title=Oxlumo|work=European Medicines Agency}}</ref> This medication is used for the treatment of [[Primary hyperoxaluria|primary hyperoxaluria type 1]] (PH1) in pediatric and adult populations. The drug is designed to reduce hepatic oxalate production and urinary oxalate levels through RNAi by targeting [[Hydroxyacid oxidase (glycolate oxidase) 1|hydroxyacid oxidase 1]] (HAO1) mRNA for breakdown. Lowering HAO1 enzyme levels reduces the oxidation of glycolate to glyoxylate (which is a substrate for oxalate). Lumasiran is administered subcutaneously by a healthcare professional with dosing based on body weight.<ref>{{cite news|url=https://dailymed.nlm.nih.gov/dailymed/drugInfo.cfm?setid=16985a31-f5e4-4557-9266-fc78d4bc5055|date=December 2, 2021|title=LABEL: OXLUMO- lumasiran injection, solution|work=Daily Med}}</ref> Data from randomized controlled clinical trials indicate that the most common adverse reaction that was reported was injection site reactions. These reactions were mild and were present in 38 percent of patients treated with lumasiran.<ref>{{cite journal |title=Lumasiran, an RNAi Therapeutic for Primary Hyperoxaluria Type 1 |journal=New England Journal of Medicine|date=April 1, 2021 |doi=10.1056/NEJMoa2021712 |pmid=33789010|last1=Garrelfs |first1=Sander F. |last2=Frishberg |first2=Yaacov |last3=Hulton |first3=Sally A. |last4=Koren |first4=Michael J. |last5=O'Riordan |first5=William D. |last6=Cochat |first6=Pierre |last7=Deschênes |first7=Georges |last8=Shasha-Lavsky |first8=Hadas |last9=Saland |first9=Jeffrey M. |last10=Van't Hoff |first10=William G. |last11=Fuster |first11=Daniel G. |last12=Magen |first12=Daniella |last13=Moochhala |first13=Shabbir H. |last14=Schalk |first14=Gesa |last15=Simkova |first15=Eva |last16=Groothoff |first16=Jaap W. |last17=Sas |first17=David J. |last18=Meliambro |first18=Kristin A. |last19=Lu |first19=Jiandong |last20=Sweetser |first20=Marianne T. |last21=Garg |first21=Pushkal P. |last22=Vaishnaw |first22=Akshay K. |last23=Gansner |first23=John M. |last24=McGregor |first24=Tracy L. |last25=Lieske |first25=John C. |author26=((ILLUMINATE-A Collaborators)) |volume=384 |issue=13 |pages=1216–1226 |s2cid=232482623 |doi-access=free }}</ref>
-In 2022, the FDA and EMA approved [[vutrisiran]] for the treatment of adults with hereditary [[Familial amyloid polyneuropathy|transthyretin mediated amyloidosis with polyneuropathy]] stage 1 or 2.<ref>{{Cite journal |last=Research |first=Center for Drug Evaluation and |date=2023-11-27 |title=Drug Trial Snapshots: AMVUTTRA |url=https://www.fda.gov/drugs/drug-approvals-and-databases/drug-trial-snapshots-amvuttra |journal=FDA |language=en}}</ref><ref>{{Cite web |title=Amvuttra {{!}} European Medicines Agency |url=https://www.ema.europa.eu/en/medicines/human/EPAR/amvuttra |access-date=2024-01-11 |website=www.ema.europa.eu}}</ref> Vutrisiran is designed to break down the mRNA that codes for [[transthyretin]].
+In 2022, the FDA and EMA approved [[vutrisiran]] for the treatment of adults with hereditary [[Familial amyloid polyneuropathy|transthyretin mediated amyloidosis with polyneuropathy]] stage 1 or 2.<ref>{{Cite journal |last=Research |first=Center for Drug Evaluation and |date=2023-11-27 |title=Drug Trial Snapshots: AMVUTTRA |url=https://www.fda.gov/drugs/drug-approvals-and-databases/drug-trial-snapshots-amvuttra |journal=FDA |language=en}}</ref><ref>{{Cite web |title=Amvuttra {{!}} European Medicines Agency |url=https://www.ema.europa.eu/en/medicines/human/EPAR/amvuttra |access-date=2024-01-11 |website=www.ema.europa.eu|date=12 October 2022 }}</ref> Vutrisiran is designed to break down the mRNA that codes for [[transthyretin]].
 Other investigational drugs using RNAi that are being developed by pharmaceutical companies such as [[Arrowhead Pharmaceuticals]], Dicerna, [[Alnylam Pharmaceuticals]], [[Amgen]], and Sylentis. These medications cover a variety of targets via RNAi and diseases.
@@ Line 134: / Line 135: @@
 Investigational RNAi therapeutics in development:
 {| class="wikitable"
-|'''Drug'''
+! Drug
-|'''Target'''
+! Target
-|'''Delivery System'''
+! Delivery System
-|'''Disease'''
+! Disease
-|'''Phase'''
+! Phase
-|'''Status'''
+! Status
-|'''Company'''
+! Company
-|'''Identifier'''
+! Identifier
 |-
 |ALN–VSP02
@@ Line 362: / Line 363: @@
 ==== Legal categorization and legal issues in a near future ====
 Currently, both miRNA and SiRNA are currently chemically synthesized and so, are legally categorized inside EU and in USA as "simple" medicinal products. But as bioengineered siRNA (BERAs) are in development, these would be classified as biological medicinal products, at least in EU. The development of the BERAs technology raises the question of the categorization of drugs having the same mechanism of action but being produced chemically or biologically. This lack of consistency should be addressed.<ref>{{Cite journal |last1=Guerriaud |first1=Mathieu |last2=Kohli |first2=Evelyne |date=2022 |title=RNA-based drugs and regulation: Toward a necessary evolution of the definitions issued from the European union legislation |journal=Frontiers in Medicine |volume=9 |doi=10.3389/fmed.2022.1012497 |pmid=36325384 |pmc=9618588 |issn=2296-858X|doi-access=free }}</ref>
 === Delivery mechanisms ===
-To achieve the clinical potential of RNAi, siRNA must be efficiently transported to the cells of target tissues. However, there are various barriers that must be fixed before it can be used clinically. For example, "naked" [[small interfering RNA|siRNA]] is susceptible to several obstacles that reduce its therapeutic efficacy.<ref name="Kanasty nmat3765">{{cite journal | vauthors = Kanasty R, Dorkin JR, Vegas A, Anderson D | title = Delivery materials for siRNA therapeutics | journal = Nature Materials | volume = 12 | issue = 11 | pages = 967–77 | date = November 2013 | pmid = 24150415 | doi = 10.1038/nmat3765 | bibcode = 2013NatMa..12..967K}}</ref> Additionally, once siRNA has entered the bloodstream, naked RNA can be degraded by serum nucleases and can stimulate the innate immune system.<ref name="Kanasty nmat3765"/> Due to its size and highly polyanionic (containing negative charges at several sites) nature, unmodified siRNA molecules cannot readily enter the cells through the cell membrane. Therefore, artificial or [[RNAi nanoparticles to target cancer|nanoparticle encapsulated siRNA]] must be used. If siRNA is transferred across the cell membrane, unintended toxicities can occur if therapeutic doses are not optimized, and siRNAs can exhibit off-target effects (e.g. unintended downregulation of genes with [[complementarity (molecular biology)|partial sequence complementarity]]).<ref name="ReferenceA">{{cite journal | vauthors = Wittrup A, Lieberman J | title = Knocking down disease: a progress report on siRNA therapeutics | journal = Nature Reviews Genetics | volume = 16 | issue = 9 | pages = 543–52 | date = September 2015 | pmid = 26281785 | pmc = 4756474 | doi = 10.1038/nrg3978}}</ref> Even after entering the cells, repeated dosing is required since their effects are diluted at each cell division. In response to these potential issues and barriers, two approaches help facilitate siRNA delivery to target cells: lipid nanoparticles and conjugates.<ref>{{cite journal | vauthors = De-Souza EA, Camara H, Salgueiro WG, Moro RP, Knittel TL, Tonon G, Pinto S, Pinca AP, Antebi A, Pasquinelli AE, Massirer KB, Mori MA | display-authors = 6 | title = RNA interference may result in unexpected phenotypes in Caenorhabditis elegans | journal = Nucleic Acids Research | volume = 47 | issue = 8 | pages = 3957–3969 | date = May 2019 | pmid = 30838421 | pmc = 6486631 | doi = 10.1093/nar/gkz154}}</ref>
+To achieve the clinical potential of RNAi, siRNA must be efficiently transported to the cells of target tissues. However, there are various barriers that must be fixed before it can be used clinically. For example, "naked" [[small interfering RNA|siRNA]] is susceptible to several obstacles that reduce its therapeutic efficacy.<ref name="Kanasty nmat3765">{{cite journal | vauthors = Kanasty R, Dorkin JR, Vegas A, Anderson D | title = Delivery materials for siRNA therapeutics | journal = Nature Materials | volume = 12 | issue = 11 | pages = 967–77 | date = November 2013 | pmid = 24150415 | doi = 10.1038/nmat3765 | bibcode = 2013NatMa..12..967K}}</ref> Additionally, once siRNA has entered the bloodstream, naked RNA can be degraded by serum nucleases and can stimulate the innate immune system.<ref name="Kanasty nmat3765"/> Due to its size and highly polyanionic (containing negative charges at several sites) nature, unmodified siRNA molecules cannot readily enter the cells through the cell membrane. Therefore, artificial or [[RNAi nanoparticles to target cancer|nanoparticle encapsulated siRNA]] must be used. If siRNA is transferred across the cell membrane, unintended toxicities can occur if therapeutic doses are not optimized, and siRNAs can exhibit off-target effects (e.g. unintended downregulation of genes with [[complementarity (molecular biology)|partial sequence complementarity]]).<ref name="ReferenceA">{{cite journal | vauthors = Wittrup A, Lieberman J | title = Knocking down disease: a progress report on siRNA therapeutics | journal = Nature Reviews Genetics | volume = 16 | issue = 9 | pages = 543–52 | date = September 2015 | pmid = 26281785 | pmc = 4756474 | doi = 10.1038/nrg3978}}</ref> Even after entering the cells, repeated dosing is required since their effects are diluted at each cell division. In response to these potential issues and barriers, two approaches help facilitate siRNA delivery to target cells: lipid nanoparticles and conjugates.<ref>{{cite journal | vauthors = De-Souza EA, Camara H, Salgueiro WG, Moro RP, Knittel TL, Tonon G, Pinto S, Pinca AP, Antebi A, Pasquinelli AE, Massirer KB, Mori MA | title = RNA interference may result in unexpected phenotypes in Caenorhabditis elegans | journal = Nucleic Acids Research | volume = 47 | issue = 8 | pages = 3957–3969 | date = May 2019 | pmid = 30838421 | pmc = 6486631 | doi = 10.1093/nar/gkz154}}</ref>
 ==== Lipid nanoparticles ====
-Lipid nanoparticles (LNPs) are based on liposome-like structures that are typically made of an aqueous center surrounded by a lipid shell.<ref>{{cite journal|last1=Li|first1=W|last2=Szoka|first2=FC|title=Lipid-based nanoparticles for nucleic acid delivery|journal=Pharm Res|date=March 24, 2007|volume=24|issue=3|pages=438–449|doi=10.1007/s11095-006-9180-5|pmid=17252188|s2cid=9995555}}</ref> A subset of liposomal structures used for delivery drugs to tissues rest in large unilamellar vesicles (LUVs) which may be 100 nm in size. LNP delivery mechanisms have become an increasing source of encasing nucleic acids and may include [[Plasmid|plasmids]], [[CRISPR]] and [[Messenger RNA|mRNA]].<ref>{{cite journal|last1=Leung|first1=AK|last2=Tam|first2=YY|last3=Cullis|first3=PR|title=Lipid nanoparticles for short interfering RNA delivery|journal=Adv Genet|series=Advances in Genetics |date=2014|volume=88 |pages=71–110 |doi=10.1016/B978-0-12-800148-6.00004-3|pmid=25409604|pmc=7149983 |isbn=9780128001486 }}</ref>
+Lipid nanoparticles (LNPs) are based on liposome-like structures that are typically made of an aqueous center surrounded by a lipid shell.<ref>{{cite journal|last1=Li|first1=W|last2=Szoka|first2=FC|title=Lipid-based nanoparticles for nucleic acid delivery|journal=Pharm Res|date=March 24, 2007|volume=24|issue=3|pages=438–449|doi=10.1007/s11095-006-9180-5|pmid=17252188|s2cid=9995555}}</ref> A subset of liposomal structures used for delivery drugs to tissues rest in large unilamellar vesicles (LUVs) which may be 100&nbsp;nm in size. LNP delivery mechanisms have become an increasing source of encasing nucleic acids and may include [[plasmid]]s, [[CRISPR]] and [[Messenger RNA|mRNA]].<ref>{{cite book|last1=Leung|first1=AK|last2=Tam|first2=YY|last3=Cullis|first3=PR|chapter=Lipid Nanoparticles for Short Interfering RNA Delivery |title=Nonviral Vectors for Gene Therapy - Lipid- and Polymer-based Gene Transfer|series=Advances in Genetics |date=2014|volume=88 |pages=71–110 |doi=10.1016/B978-0-12-800148-6.00004-3|pmid=25409604|pmc=7149983 |isbn=978-0-12-800148-6 }}</ref>
 The first approved use of [[Solid lipid nanoparticle|lipid nanoparticle]]s as a drug delivery mechanism began in 2018 with the siRNA drug patisiran, developed by Alnylam Pharmaceuticals. Dicerna Pharmaceuticals, [[Persomics]], [[Sanofi]] and [[Sirna Therapeutics]] also worked to bring RNAi therapies to market.<ref>{{cite news|url=https://www.forbes.com/sites/johnlamattina/2014/04/15/big-pharmas-turn-on-rnai-shows-that-new-technologies-dont-guarantee-rd-success/?sh=17b5334478fc|author=John LaMattina|date=April 15, 2014|title=Big Pharma's Turn On RNAi Shows That New Technologies Don't Guarantee R&D Success|work=Forbes}}</ref><ref>{{cite news|url=https://www.the-scientist.com/features/the-second-coming-of-rnai-36936|author=Eric Bender|date=September 1, 2014|title=The Second Coming of RNAi|work=The Scientist}}</ref>
 Other recent applications include two FDA approved COVID-19 vaccines: mRNA-1273, developed by [[Moderna]]. and [[Pfizer–BioNTech COVID-19 vaccine|BNT162b]], developed by a collaboration between [[Pfizer]] and [[BioNtech]].<ref>{{cite journal|last=Baden|first=LR|title=Efficacy and safety of the mRNA-1273 SARS-CoV-2 vaccine|journal=New England Journal of Medicine|date=2021|volume=384 |issue=5 |pages=403–416 |doi=10.1056/NEJMoa2035389|pmid=33378609 |pmc=7787219 }}</ref> These two vaccines use lipid nanoparticles to deliver antigen mRNA. Encapsulating the mRNA molecule in lipid nanoparticles was a critical breakthrough for producing viable mRNA vaccines, solving a number of key technical barriers in delivering the mRNA molecule into the host cell as distributed through [[apolipoprotein E]] (apoE) in the low-density lipoprotein receptor (LDLR). In December 2020, Novartis announced that positive results from phase III efficacy studies deemed [[inclisiran]] was a treatment for [[heterozygous]] [[familial hypercholesterolemia]] (HeFH) and [[atherosclerotic cardiovascular disease]] (ASCVD).<ref>{{cite journal|last1=Fitzgerald|first1=K|last2=White|first2=S|last3=Borodovsky|first3=A|last4=Bettencourt|first4=BR |last5=Strahs|first5=A|last6=Clausen|first6=V|title=A Highly Durable RNAi Therapeutic Inhibitor of PCSK9|journal=New England Journal of Medicine|date=January 2017|volume=376|issue=1|pages=41–51|doi=10.1056/NEJMoa1609243|pmid=27959715|pmc=5778873}}</ref>
 ==== Conjugates ====
-In addition to LNPs, RNAi therapeutics have targeted delivery through siRNA conjugates (e.g., GalNAc, carbohydrates, peptides, aptamers, antibodies).<ref>{{cite journal|last1=Kruspe|first1=S|last2=Giangrande|first2=P|title=Aptamer-siRNA Chimeras: Discovery, Progress, and Future Prospects|journal=Biomedicines|year=2017|volume=5|issue=4|page=45|doi=10.3390/biomedicines5030045|pmid=28792479|pmc=5618303|doi-access=free}}</ref> Therapeutics using siRNA conjugates have been developed for rare or genetic diseases such as acute hepatic porphyria (AHP), [[Haemophilia|hemophilia]], [[primary hyperoxaluria]] (PH) and hereditary ATTR [[amyloidosis]] as well as other cardiometabolic diseases such as [[hypertension]] and non-alcoholic [[steatohepatitis]] (NASH).<ref>{{cite journal|title=Phase 3 Trial of RNAi Therapeutic Givosiran for Acute Intermittent Porphyria|journal=New England Journal of Medicine|date=June 11, 2020|doi=10.1056/NEJMoa1913147|pmid=32521132|last1=Balwani |first1=Manisha |last2=Sardh |first2=Eliane |last3=Ventura |first3=Paolo |last4=Peiró |first4=Paula Aguilera |last5=Rees |first5=David C. |last6=Stölzel |first6=Ulrich |last7=Bissell |first7=D. Montgomery |last8=Bonkovsky |first8=Herbert L. |last9=Windyga |first9=Jerzy |last10=Anderson |first10=Karl E. |last11=Parker |first11=Charles |last12=Silver |first12=Samuel M. |last13=Keel |first13=Siobán B. |last14=Wang |first14=Jiaan-Der |last15=Stein |first15=Penelope E. |last16=Harper |first16=Pauline |last17=Vassiliou |first17=Daphne |last18=Wang |first18=Bruce |last19=Phillips |first19=John |last20=Ivanova |first20=Aneta |last21=Langendonk |first21=Janneke G. |last22=Kauppinen |first22=Raili |last23=Minder |first23=Elisabeth |last24=Horie |first24=Yutaka |last25=Penz |first25=Craig |last26=Chen |first26=Jihong |last27=Liu |first27=Shangbin |last28=Ko |first28=John J. |last29=Sweetser |first29=Marianne T. |last30=Garg |first30=Pushkal |volume=382 |issue=24 |pages=2289–2301 |s2cid=219586624 |display-authors=1 |doi-access=free }}</ref>
+In addition to LNPs, RNAi therapeutics have targeted delivery through siRNA conjugates (e.g., GalNAc, carbohydrates, peptides, aptamers, antibodies).<ref>{{cite journal|last1=Kruspe|first1=S|last2=Giangrande|first2=P|title=Aptamer-siRNA Chimeras: Discovery, Progress, and Future Prospects|journal=Biomedicines|year=2017|volume=5|issue=4|page=45|doi=10.3390/biomedicines5030045|pmid=28792479|pmc=5618303|doi-access=free}}</ref> Therapeutics using siRNA conjugates have been developed for rare or genetic diseases such as acute hepatic porphyria (AHP), [[Haemophilia|hemophilia]], [[primary hyperoxaluria]] (PH) and hereditary ATTR [[amyloidosis]] as well as other cardiometabolic diseases such as [[hypertension]] and non-alcoholic [[steatohepatitis]] (NASH).<ref>{{cite journal|title=Phase 3 Trial of RNAi Therapeutic Givosiran for Acute Intermittent Porphyria|journal=New England Journal of Medicine|date=June 11, 2020|doi=10.1056/NEJMoa1913147|pmid=32521132|last1=Balwani |first1=Manisha |last2=Sardh |first2=Eliane |last3=Ventura |first3=Paolo |last4=Peiró |first4=Paula Aguilera |last5=Rees |first5=David C. |last6=Stölzel |first6=Ulrich |last7=Bissell |first7=D. Montgomery |last8=Bonkovsky |first8=Herbert L. |last9=Windyga |first9=Jerzy |last10=Anderson |first10=Karl E. |last11=Parker |first11=Charles |last12=Silver |first12=Samuel M. |last13=Keel |first13=Siobán B. |last14=Wang |first14=Jiaan-Der |last15=Stein |first15=Penelope E. |last16=Harper |first16=Pauline |last17=Vassiliou |first17=Daphne |last18=Wang |first18=Bruce |last19=Phillips |first19=John |last20=Ivanova |first20=Aneta |last21=Langendonk |first21=Janneke G. |last22=Kauppinen |first22=Raili |last23=Minder |first23=Elisabeth |last24=Horie |first24=Yutaka |last25=Penz |first25=Craig |last26=Chen |first26=Jihong |last27=Liu |first27=Shangbin |last28=Ko |first28=John J. |last29=Sweetser |first29=Marianne T. |last30=Garg |first30=Pushkal |volume=382 |issue=24 |pages=2289–2301 |s2cid=219586624 |doi-access=free }}</ref>
 === Biotechnology ===
@@ Line 405: / Line 405: @@
 RNAi has been used to genetically engineer plants to produce lower levels of natural plant toxins. Such techniques take advantage of the stable and heritable RNAi phenotype in plant stocks. [[Cotton]] seeds are rich in [[protein in nutrition|dietary protein]] but naturally contain the toxic [[terpenoid]] product [[gossypol]], making them unsuitable for human consumption. RNAi has been used to produce cotton stocks whose seeds contain reduced levels of [[delta-cadinene synthase]], a key enzyme in gossypol production, without affecting the enzyme's production in other parts of the plant, where gossypol is itself important in preventing damage from plant pests.<ref name="Sunilkumar">{{cite journal | vauthors = Sunilkumar G, Campbell LM, Puckhaber L, Stipanovic RD, Rathore KS | title = Engineering cottonseed for use in human nutrition by tissue-specific reduction of toxic gossypol | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 103 | issue = 48 | pages = 18054–9 | date = November 2006 | pmid = 17110445 | pmc = 1838705 | doi = 10.1073/pnas.0605389103 | doi-access = free }}</ref>
-Development efforts have successfully reduced the levels of [[allergen]]s in [[tomato]] plants<ref name="Le_tomato">{{cite journal | vauthors = Le LQ, Lorenz Y, Scheurer S, Fötisch K, Enrique E, Bartra J, Biemelt S, Vieths S, Sonnewald U | title = Design of tomato fruits with reduced allergenicity by dsRNAi-mediated inhibition of ns-LTP (Lyc e 3) expression | journal = Plant Biotechnology Journal | volume = 4 | issue = 2 | pages = 231–42 | date = March 2006 | pmid = 17177799 | doi = 10.1111/j.1467-7652.2005.00175.x}}</ref> and fortification of plants such as tomatoes with dietary [[antioxidant]]s.<ref name="Niggeweg">{{cite journal | vauthors = Niggeweg R, Michael AJ, Martin C | title = Engineering plants with increased levels of the antioxidant chlorogenic acid | journal = Nature Biotechnology | volume = 22 | issue = 6 | pages = 746–54 | date = June 2004 | pmid = 15107863 | doi = 10.1038/nbt966 | s2cid = 21588259}}</ref> RNAi silencing of [[alpha-amylase]] have also been used to decrease [[Aspergillus flavus]] fungal growth in maize which would have otherwise contaminated the kernels with dangerous [[aflatoxin]]s.<ref>{{cite journal | vauthors = Gilbert MK, Majumdar R, Rajasekaran K, Chen ZY, Wei Q, Sickler CM, Lebar MD, Cary JW, Frame BR, Wang K | title = RNA interference-based silencing of the alpha-amylase (amy1) gene in Aspergillus flavus decreases fungal growth and aflatoxin production in maize kernels | journal = Planta | volume = 247 | issue = 6 | pages = 1465–1473 | date = June 2018 | pmid = 29541880 | doi = 10.1007/s00425-018-2875-0 | bibcode = 2018Plant.247.1465G | s2cid = 3918937}}</ref> Silencing lachrymatory factor synthase in [[onion]]s have produced tearless onions and RNAi has been used in BP1 genes in [[rapeseed]]s to improve photosynthesis.<ref>{{cite journal | vauthors = Katoch R, Thakur N | s2cid = 45212581 | title = RNA interference: a promising technique for the improvement of traditional crops | journal = International Journal of Food Sciences and Nutrition | volume = 64 | issue = 2 | pages = 248–59 | date = March 2013 | pmid = 22861122 | doi = 10.3109/09637486.2012.713918}}</ref> SBEIIa and SBEIIb genes in wheat have been targeted in wheat in order to produce higher levels of [[amylose]] in order to improve bowel function,<ref>{{cite journal | vauthors = Katoch R, Thakur N | s2cid = 23733295 | title = Advances in RNA interference technology and its impact on nutritional improvement, disease and insect control in plants | journal = Applied Biochemistry and Biotechnology | volume = 169 | issue = 5 | pages = 1579–605 | date = March 2013 | pmid = 23322250 | doi = 10.1007/s12010-012-0046-5}}</ref> and Travella et al. 2006 employed RNAi for [[functional genomics]] an investigation of [[hexaploid bread wheat|hexaploid bread races]], while virus-induced gene silencing (VIGS, a subtype of RNAi) was used by Scofield et al. 2005 to investigate the mechanism of [[plant disease resistance|resistance]] provided by ''[[Lr21]]'' against [[wheat leaf rust]] in [[hexaploid wheat]].<ref name="Downie-et-al-2021">{{cite journal | last1=Downie | first1=Rowena C. | last2=Lin | first2=Min | last3=Corsi | first3=Beatrice | last4=Ficke | first4=Andrea | last5=Lillemo | first5=Morten | last6=Oliver | first6=Richard P. | last7=Phan | first7=Huyen T. T. | last8=Tan | first8=Kar-Chun | last9=Cockram | first9=James | title=Septoria Nodorum Blotch of Wheat: Disease Management and Resistance Breeding in the Face of Shifting Disease Dynamics and a Changing Environment | journal=[[Phytopathology (journal)|Phytopathology]] | publisher=[[American Phytopathological Society]] | date=2021-07-27 | volume=111 | issue=6 | issn=0031-949X | doi=10.1094/phyto-07-20-0280-rvw | pages=PHYTO–07–20–028| pmid=33245254 | hdl=20.500.11937/83208 | s2cid=227181536 | hdl-access=free }}</ref>
+Development efforts have successfully reduced the levels of [[allergen]]s in [[tomato]] plants<ref name="Le_tomato">{{cite journal | vauthors = Le LQ, Lorenz Y, Scheurer S, Fötisch K, Enrique E, Bartra J, Biemelt S, Vieths S, Sonnewald U | title = Design of tomato fruits with reduced allergenicity by dsRNAi-mediated inhibition of ns-LTP (Lyc e 3) expression | journal = Plant Biotechnology Journal | volume = 4 | issue = 2 | pages = 231–42 | date = March 2006 | pmid = 17177799 | doi = 10.1111/j.1467-7652.2005.00175.x| doi-access = free }}</ref> and fortification of plants such as tomatoes with dietary [[antioxidant]]s.<ref name="Niggeweg">{{cite journal | vauthors = Niggeweg R, Michael AJ, Martin C | title = Engineering plants with increased levels of the antioxidant chlorogenic acid | journal = Nature Biotechnology | volume = 22 | issue = 6 | pages = 746–54 | date = June 2004 | pmid = 15107863 | doi = 10.1038/nbt966 | s2cid = 21588259}}</ref> RNAi silencing of [[alpha-amylase]] have also been used to decrease [[Aspergillus flavus]] fungal growth in maize which would have otherwise contaminated the kernels with dangerous [[aflatoxin]]s.<ref>{{cite journal | vauthors = Gilbert MK, Majumdar R, Rajasekaran K, Chen ZY, Wei Q, Sickler CM, Lebar MD, Cary JW, Frame BR, Wang K | title = RNA interference-based silencing of the alpha-amylase (amy1) gene in Aspergillus flavus decreases fungal growth and aflatoxin production in maize kernels | journal = Planta | volume = 247 | issue = 6 | pages = 1465–1473 | date = June 2018 | pmid = 29541880 | doi = 10.1007/s00425-018-2875-0 | bibcode = 2018Plant.247.1465G | s2cid = 3918937}}</ref> Silencing lachrymatory factor synthase in [[onion]]s have produced tearless onions and RNAi has been used in BP1 genes in [[rapeseed]]s to improve photosynthesis.<ref>{{cite journal | vauthors = Katoch R, Thakur N | s2cid = 45212581 | title = RNA interference: a promising technique for the improvement of traditional crops | journal = International Journal of Food Sciences and Nutrition | volume = 64 | issue = 2 | pages = 248–59 | date = March 2013 | pmid = 22861122 | doi = 10.3109/09637486.2012.713918}}</ref> SBEIIa and SBEIIb genes in wheat have been targeted in wheat in order to produce higher levels of [[amylose]] in order to improve bowel function,<ref>{{cite journal | vauthors = Katoch R, Thakur N | s2cid = 23733295 | title = Advances in RNA interference technology and its impact on nutritional improvement, disease and insect control in plants | journal = Applied Biochemistry and Biotechnology | volume = 169 | issue = 5 | pages = 1579–605 | date = March 2013 | pmid = 23322250 | doi = 10.1007/s12010-012-0046-5}}</ref> and Travella et al. 2006 employed RNAi for [[functional genomics]] an investigation of [[hexaploid bread wheat|hexaploid bread races]], while virus-induced gene silencing (VIGS, a subtype of RNAi) was used by Scofield et al. 2005 to investigate the mechanism of [[plant disease resistance|resistance]] provided by ''[[Lr21]]'' against [[wheat leaf rust]] in [[hexaploid wheat]].<ref name="Downie-et-al-2021">{{cite journal | last1=Downie | first1=Rowena C. | last2=Lin | first2=Min | last3=Corsi | first3=Beatrice | last4=Ficke | first4=Andrea | last5=Lillemo | first5=Morten | last6=Oliver | first6=Richard P. | last7=Phan | first7=Huyen T. T. | last8=Tan | first8=Kar-Chun | last9=Cockram | first9=James | title=Septoria Nodorum Blotch of Wheat: Disease Management and Resistance Breeding in the Face of Shifting Disease Dynamics and a Changing Environment | journal=[[Phytopathology (journal)|Phytopathology]] | publisher=[[American Phytopathological Society]] | date=2021-07-27 | volume=111 | issue=6 | issn=0031-949X | doi=10.1094/phyto-07-20-0280-rvw | pages=PHYTO–07–20–028| pmid=33245254 | hdl=20.500.11937/83208 | s2cid=227181536 | hdl-access=free }}</ref>
 ==== Insecticides ====
@@ Line 411: / Line 411: @@
 RNAi is under development as an [[insecticide]], employing multiple approaches, including genetic engineering and topical application.<ref name=s1308/> Cells in the midgut of some insects take up the dsRNA molecules in the process referred to as environmental RNAi.<ref>{{cite journal | vauthors = Ivashuta S, Zhang Y, Wiggins BE, Ramaseshadri P, Segers GC, Johnson S, Meyer SE, Kerstetter RA, McNulty BC, Bolognesi R, Heck GR | title = Environmental RNAi in herbivorous insects | journal = RNA | volume = 21 | issue = 5 | pages = 840–50 | date = May 2015 | pmid = 25802407 | pmc = 4408792 | doi = 10.1261/rna.048116.114}}</ref> In some insects the effect is systemic as the signal spreads throughout the insect's body (referred to as systemic RNAi).<ref>{{cite journal | vauthors = Miller SC, Miyata K, Brown SJ, Tomoyasu Y | title = Dissecting systemic RNA interference in the red flour beetle ''Tribolium castaneum'': parameters affecting the efficiency of RNAi | journal = PLOS ONE | volume = 7 | issue = 10 | pages = e47431 | year = 2012 | pmid = 23133513 | pmc = 3484993 | doi = 10.1371/journal.pone.0047431 | bibcode = 2012PLoSO...747431M| doi-access = free }}</ref>
-Animals exposed to RNAi at doses millions of times higher than anticipated human exposure levels show no adverse effects.<ref>{{cite journal | vauthors = Petrick JS, Friedrich GE, Carleton SM, Kessenich CR, Silvanovich A, Zhang Y, Koch MS | title = Corn rootworm-active RNA DvSnf7: Repeat dose oral toxicology assessment in support of human and mammalian safety | journal = Regulatory Toxicology and Pharmacology | volume = 81 | pages = 57–68 | date = November 2016 | pmid = 27436086 | doi = 10.1016/j.yrtph.2016.07.009 | doi-access = free}}</ref> RNAi has varying effects in different species of [[Lepidoptera]] (butterflies and moths).<ref>{{cite journal | vauthors = Terenius O, Papanicolaou A, Garbutt JS, Eleftherianos I, Huvenne H, Kanginakudru S, Albrechtsen M, An C, Aymeric JL, Barthel A, Bebas P, Bitra K, Bravo A, Chevalier F, Collinge DP, Crava CM, de Maagd RA, Duvic B, Erlandson M, Faye I, Felföldi G, Fujiwara H, Futahashi R, Gandhe AS, Gatehouse HS, Gatehouse LN, Giebultowicz JM, Gómez I, Grimmelikhuijzen CJ, Groot AT, Hauser F, Heckel DG, Hegedus DD, Hrycaj S, Huang L, Hull JJ, Iatrou K, Iga M, Kanost MR, Kotwica J, Li C, Li J, Liu J, Lundmark M, Matsumoto S, Meyering-Vos M, Millichap PJ, Monteiro A, Mrinal N, Niimi T, Nowara D, Ohnishi A, Oostra V, Ozaki K, Papakonstantinou M, Popadic A, Rajam MV, Saenko S, Simpson RM, Soberón M, Strand MR, Tomita S, Toprak U, Wang P, Wee CW, Whyard S, Zhang W, Nagaraju J, Ffrench-Constant RH, Herrero S, Gordon K, Swevers L, Smagghe G | display-authors = 6 | title = RNA interference in Lepidoptera: an overview of successful and unsuccessful studies and implications for experimental design | journal = Journal of Insect Physiology | volume = 57 | issue = 2 | pages = 231–45 | date = February 2011 | pmid = 21078327 | doi = 10.1016/j.jinsphys.2010.11.006 | url = https://biblio.ugent.be/publication/1101411 | hdl = 1854/LU-1101411 | hdl-access = free}}</ref>
+Animals exposed to RNAi at doses millions of times higher than anticipated human exposure levels show no adverse effects.<ref>{{cite journal | vauthors = Petrick JS, Friedrich GE, Carleton SM, Kessenich CR, Silvanovich A, Zhang Y, Koch MS | title = Corn rootworm-active RNA DvSnf7: Repeat dose oral toxicology assessment in support of human and mammalian safety | journal = Regulatory Toxicology and Pharmacology | volume = 81 | pages = 57–68 | date = November 2016 | pmid = 27436086 | doi = 10.1016/j.yrtph.2016.07.009 | doi-access = free}}</ref> RNAi has varying effects in different species of [[Lepidoptera]] (butterflies and moths).<ref>{{cite journal | vauthors = Terenius O, Papanicolaou A, Garbutt JS, Eleftherianos I, Huvenne H, Kanginakudru S, Albrechtsen M, An C, Aymeric JL, Barthel A, Bebas P, Bitra K, Bravo A, Chevalier F, Collinge DP, Crava CM, de Maagd RA, Duvic B, Erlandson M, Faye I, Felföldi G, Fujiwara H, Futahashi R, Gandhe AS, Gatehouse HS, Gatehouse LN, Giebultowicz JM, Gómez I, Grimmelikhuijzen CJ, Groot AT, Hauser F, Heckel DG, Hegedus DD, Hrycaj S, Huang L, Hull JJ, Iatrou K, Iga M, Kanost MR, Kotwica J, Li C, Li J, Liu J, Lundmark M, Matsumoto S, Meyering-Vos M, Millichap PJ, Monteiro A, Mrinal N, Niimi T, Nowara D, Ohnishi A, Oostra V, Ozaki K, Papakonstantinou M, Popadic A, Rajam MV, Saenko S, Simpson RM, Soberón M, Strand MR, Tomita S, Toprak U, Wang P, Wee CW, Whyard S, Zhang W, Nagaraju J, Ffrench-Constant RH, Herrero S, Gordon K, Swevers L, Smagghe G | title = RNA interference in Lepidoptera: an overview of successful and unsuccessful studies and implications for experimental design | journal = Journal of Insect Physiology | volume = 57 | issue = 2 | pages = 231–45 | date = February 2011 | pmid = 21078327 | doi = 10.1016/j.jinsphys.2010.11.006 | bibcode = 2011JInsP..57..231T | url = https://biblio.ugent.be/publication/1101411 | hdl = 1854/LU-1101411 | hdl-access = free}}</ref>
 ''[[Drosophila]]'' spp., ''[[Bombyx mori]]'', ''[[Locusta]]'' spp., ''[[Spodoptera]]'' spp., ''[[Tribolium castaneum]]'', ''[[Nilaparvata lugens]]'', ''[[Helicoverpa armigera]]'', and ''[[Apis mellifera]]'' are models that have been widely used to learn how RNAi works within particular [[taxon|taxa]] of insects. ''[[Musca domestica]]'' has two ''[[Ago2]]'' genes and ''[[Glossina morsitans]]'' three, as found by Lewis et al. 2016 and Hain et al. 2010.<ref name="Mongelli-Saleh-2016">{{cite journal | last1=Mongelli | first1=Vanesa | last2=Saleh | first2=Maria-Carla | title=Bugs Are Not to Be Silenced: Small RNA Pathways and Antiviral Responses in Insects | journal=[[Annual Review of Virology]] | publisher=[[Annual Reviews (publisher)|Annual Reviews]] | volume=3 | issue=1 | date=2016-09-29 | issn=2327-056X | doi=10.1146/annurev-virology-110615-042447 | pages=573–589 | s2cid=38499958 | pmid=27741406| url=https://hal-pasteur.archives-ouvertes.fr/pasteur-01957180/file/Mongelli%20and%20Saleh.pdf }}</ref><ref name="Zhu-Palli-2020" /> In the case of the [[miRNA]] pathway ''[[Diuraphis noxia]]'' has two ''[[Ago1]]''s, ''M. domestica'' two ''[[Dcr1]]''s, ''[[Acyrthosiphon pisum]]'' two each of ''[[Ago1]]'' and ''[[Loqs]]'' and ''Dcr1'' and four of ''[[Pasha (protein)|Pasha]]''. While in the [[piRNA]], ''G. morsitans'' and ''A. pisum'' have two or three ''[[Ago3]]''s each.<ref name="Zhu-Palli-2020" /> This has led to identification of future [[insecticide development]] targets, and the [[mode of action|modes of action]] and reasons for [[insecticide resistance]] of other insecticides.<ref name="Zhu-Palli-2020">{{cite journal | last1=Zhu | first1=Kun Yan | last2=Palli | first2=Subba Reddy | title=Mechanisms, Applications, and Challenges of Insect RNA Interference | journal=[[Annual Review of Entomology]] | publisher=[[Annual Reviews (publisher)|Annual Reviews]] | volume=65 | issue=1 | date=2020-01-07 | issn=0066-4170 | doi=10.1146/annurev-ento-011019-025224 | pages=293–311| pmid=31610134 | pmc=9939233 | s2cid=204702574 }}</ref>
@@ Line 445: / Line 445: @@
 Not long after, plant [[virologist]]s working on improving plant resistance to viral diseases observed a similar unexpected phenomenon. While it was known that plants expressing virus-specific proteins showed enhanced tolerance or resistance to viral infection, it was not expected that plants carrying only short, non-coding regions of viral RNA sequences would show similar levels of protection. Researchers believed that viral RNA produced by transgenes could also inhibit viral replication.<ref name=Covey_1997>{{cite journal | vauthors = Covey S, Al-Kaff N, Lángara A, Turner D | title = Plants combat infection by gene silencing | journal = Nature | year = 1997 | volume = 385 | pages = 781–2 | doi= 10.1038/385781a0 | issue = 6619 |bibcode = 1997Natur.385..781C | s2cid = 43229760}}</ref> The reverse experiment, in which short sequences of plant genes were introduced into viruses, showed that the targeted gene was suppressed in an infected plant.<ref name=Kumagai_1995>{{cite journal | vauthors = Kumagai MH, Donson J, della-Cioppa G, Harvey D, Hanley K, Grill LK | title = Cytoplasmic inhibition of carotenoid biosynthesis with virus-derived RNA | journal = Proceedings of the National Academy of Sciences of the United States of America | volume = 92 | issue = 5 | pages = 1679–83 | date = February 1995 | pmid = 7878039 | pmc = 42583 | doi = 10.1073/pnas.92.5.1679 | bibcode = 1995PNAS...92.1679K| doi-access = free }}</ref> This phenomenon was labeled "virus-induced gene silencing" (VIGS),<ref name="Downie-et-al-2021" /> and the set of such phenomena were collectively called ''post transcriptional gene silencing.''<ref name=Ratcliff_1997>{{cite journal | vauthors = Ratcliff F, Harrison BD, Baulcombe DC | title = A similarity between viral defense and gene silencing in plants | journal = Science | volume = 276 | issue = 5318 | pages = 1558–60 | date = June 1997 | pmid = 18610513 | doi = 10.1126/science.276.5318.1558}}</ref>
-After these initial observations in plants, laboratories searched for this phenomenon in other organisms.<ref name="Guo">{{cite journal | vauthors = Guo S, Kemphues KJ | title = par-1, a gene required for establishing polarity in C. elegans embryos, encodes a putative Ser/Thr kinase that is asymmetrically distributed | journal = Cell | volume = 81 | issue = 4 | pages = 611–20 | date = May 1995 | pmid = 7758115 | doi = 10.1016/0092-8674(95)90082-9 | doi-access = free}}</ref><ref name="Pal-Bhadra">{{cite journal | vauthors = Pal-Bhadra M, Bhadra U, Birchler JA | title = Cosuppression in Drosophila: gene silencing of Alcohol dehydrogenase by white-Adh transgenes is Polycomb dependent | journal = Cell | volume = 90 | issue = 3 | pages = 479–90 | date = August 1997 | pmid = 9267028 | doi = 10.1016/S0092-8674(00)80508-5 | doi-access = free}}</ref> The first instance of [[RNA]] silencing in animals was documented in 1996, when Guo and Kemphues observed that, by introducing [[Sense strand|sense]] and [[antisense RNA]] to par-1 mRNA in ''[[Caenorhabditis elegans]]'' caused degradation of the par-1 message.<ref name="Sen">{{cite journal | vauthors = Sen GL, Blau HM | s2cid = 12917676 | title = A brief history of RNAi: the silence of the genes | journal = FASEB Journal | volume = 20 | issue = 9 | pages = 1293–9 | date = July 2006 | pmid = 16816104 | doi = 10.1096/fj.06-6014rev}}</ref> It was thought that this degradation was triggered by single-stranded RNA (ssRNA), but two years later, in 1998, Fire and Mello discovered that this ability to silence the par-1 gene expression was actually triggered by double-stranded RNA (dsRNA).<ref name="Sen" /> [[Craig C. Mello]] and [[Andrew Fire]]'s 1998 ''Nature'' paper reported a potent gene silencing effect after injecting double stranded RNA into ''C. elegans''.<ref name="Fire">{{cite journal | vauthors = Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC | title = Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans | journal = Nature | volume = 391 | issue = 6669 | pages = 806–11 | date = February 1998 | pmid = 9486653 | doi = 10.1038/35888 | bibcode = 1998Natur.391..806F | s2cid = 4355692| url = http://www.dspace.cam.ac.uk/handle/1810/238264 }}</ref> In investigating the regulation of muscle protein production, they observed that neither mRNA nor [[antisense RNA]] injections had an effect on protein production, but double-stranded RNA successfully silenced the targeted gene. As a result of this work, they coined the term ''RNAi''. This discovery represented the first identification of the causative agent for the phenomenon. Fire and Mello were awarded the 2006 [[Nobel Prize in Physiology or Medicine]].<ref name="Daneholt2006" /><ref>{{Cite web|url=https://www.nobelprize.org/nobel_prizes/medicine/laureates/2006/advanced.html|title=The Nobel Prize in Physiology or Medicine 2006| vauthors = Daneholt B |date=2 October 2006|website=nobelprize.org |access-date=30 October 2017}}</ref>
+After these initial observations in plants, laboratories searched for this phenomenon in other organisms.<ref name="Guo">{{cite journal | vauthors = Guo S, Kemphues KJ | title = par-1, a gene required for establishing polarity in C. elegans embryos, encodes a putative Ser/Thr kinase that is asymmetrically distributed | journal = Cell | volume = 81 | issue = 4 | pages = 611–20 | date = May 1995 | pmid = 7758115 | doi = 10.1016/0092-8674(95)90082-9 | doi-access = free}}</ref><ref name="Pal-Bhadra">{{cite journal | vauthors = Pal-Bhadra M, Bhadra U, Birchler JA | title = Cosuppression in Drosophila: gene silencing of Alcohol dehydrogenase by white-Adh transgenes is Polycomb dependent | journal = Cell | volume = 90 | issue = 3 | pages = 479–90 | date = August 1997 | pmid = 9267028 | doi = 10.1016/S0092-8674(00)80508-5 | doi-access = free}}</ref> The first instance of [[RNA]] silencing in animals was documented in 1996, when Guo and Kemphues observed that, by introducing [[Sense strand|sense]] and [[antisense RNA]] to par-1 mRNA in ''[[Caenorhabditis elegans]]'' caused degradation of the par-1 message.<ref name="Sen">{{cite journal | vauthors = Sen GL, Blau HM | s2cid = 12917676 | title = A brief history of RNAi: the silence of the genes | journal = FASEB Journal | volume = 20 | issue = 9 | pages = 1293–9 | date = July 2006 | pmid = 16816104 | doi = 10.1096/fj.06-6014rev| doi-access = free }}</ref> It was thought that this degradation was triggered by single-stranded RNA (ssRNA), but two years later, in 1998, Fire and Mello discovered that this ability to silence the par-1 gene expression was actually triggered by double-stranded RNA (dsRNA).<ref name="Sen" /> [[Craig C. Mello]] and [[Andrew Fire]]'s 1998 ''Nature'' paper reported a potent gene silencing effect after injecting double stranded RNA into ''C. elegans''.<ref name="Fire">{{cite journal | vauthors = Fire A, Xu S, Montgomery MK, Kostas SA, Driver SE, Mello CC | title = Potent and specific genetic interference by double-stranded RNA in Caenorhabditis elegans | journal = Nature | volume = 391 | issue = 6669 | pages = 806–11 | date = February 1998 | pmid = 9486653 | doi = 10.1038/35888 | bibcode = 1998Natur.391..806F | s2cid = 4355692| url = http://www.dspace.cam.ac.uk/handle/1810/238264 }}</ref> In investigating the regulation of muscle protein production, they observed that neither mRNA nor [[antisense RNA]] injections had an effect on protein production, but double-stranded RNA successfully silenced the targeted gene. As a result of this work, they coined the term ''RNAi''. This discovery represented the first identification of the causative agent for the phenomenon. Fire and Mello were awarded the 2006 [[Nobel Prize in Physiology or Medicine]].<ref name="Daneholt2006" /><ref>{{Cite web|url=https://www.nobelprize.org/nobel_prizes/medicine/laureates/2006/advanced.html|title=The Nobel Prize in Physiology or Medicine 2006| vauthors = Daneholt B |date=2 October 2006|website=nobelprize.org |access-date=30 October 2017}}</ref>
 ===RNAi therapeutics===
 Just after Fire and Mello's ground-breaking discovery, Elbashir et al. discovered, by using synthetically made [[small interfering RNA]] (siRNA), it was possible to target the silencing of specific sequences in a gene, rather than silencing the entire gene.<ref>{{cite journal|vauthors=Elbashir S, Harborth J, Lendeckel W et al.|title=Duplexes of 21-nucleotide RNAs mediate RNA interference in cultured mammalian cells|journal=Nature|volume=411|pages=494–498|year=2001|issue=6836|doi=10.1038/35078107|pmid=11373684|bibcode=2001Natur.411..494E|s2cid=710341}}</ref> Only a year later, McCaffrey and colleagues demonstrated that this sequence-specific silencing had therapeutic applications by targeting a sequence from the [[Hepatitis C]] virus in [[Transgenesis|transgenic]] mice.<ref>{{cite journal | vauthors = McCaffrey AP, Meuse L, Pham TT, Conklin DS, Hannon GJ, Kay MA | s2cid = 4361399 | title = RNA interference in adult mice | journal = Nature | volume = 418 | issue = 6893 | pages = 38–9 | date = July 2002 | pmid = 12097900 | doi = 10.1038/418038a | bibcode = 2002Natur.418...38M | author-link5 = Gregory Hannon}} {{closed access}}</ref> Since then, multiple researchers have been attempting to expand the therapeutic applications of RNAi, specifically looking to target genes that cause various types of [[cancer]].<ref>{{cite journal | vauthors = Devi GR | title = siRNA-based approaches in cancer therapy | journal = Cancer Gene Therapy | volume = 13 | issue = 9 | pages = 819–29 | date = September 2006 | pmid = 16424918 | doi = 10.1038/sj.cgt.7700931 | doi-access = free}}</ref><ref>{{cite journal | vauthors = Wall NR, Shi Y | title = Small RNA: can RNA interference be exploited for therapy? | journal = Lancet | volume = 362 | issue = 9393 | pages = 1401–3 | date = October 2003 | pmid = 14585643 | doi = 10.1016/s0140-6736(03)14637-5 | s2cid = 25034627}}</ref> By 2006, the first applications to reach [[clinical trial]]s were in the treatment of [[macular degeneration]] and [[respiratory syncytial virus]].<ref>{{cite journal |author=Sah D |title=Therapeutic potential of RNA interference for neurological disorders |journal=Life Sci |volume=79 |issue=19 |pages=1773–80 |year=2006 |pmid=16815477 |doi=10.1016/j.lfs.2006.06.011}}</ref> Four years later the first-in-human Phase I clinical trial was started, using a [[Targeted drug delivery|nanoparticle delivery system]] to target [[Neoplasm|solid tumors]].<ref>{{cite journal | vauthors = [[Mark E. Davis|Davis ME]], Zuckerman JE, Choi CH, Seligson D, Tolcher A, [[Christopher Alabi|Alabi CA]], Yen Y, Heidel JD, Ribas A | title = Evidence of RNAi in humans from systemically administered siRNA via targeted nanoparticles | journal = Nature | volume = 464 | issue = 7291 | pages = 1067–70 | date = April 2010 | pmid = 20305636 | pmc = 2855406 | doi = 10.1038/nature08956 | bibcode = 2010Natur.464.1067D}}</ref>
 The FDA approved the first siRNA-based drug ([[patisiran]]) in 2018. [[Givosiran]] and [[lumasiran]] later won FDA approval for the treatment of AHP and PH1 in 2019 and 2020, respectively.<ref name=Patisiran/> [[Inclisiran]] received EMA approval in 2020 for the treatment of high cholesterol and is currently under review by the FDA.<ref>{{cite news|url=https://www.novartis.com/news/media-releases/novartis-receives-eu-approval-leqvio-inclisiran-first-class-sirna-lower-cholesterol-two-doses-year|date=December 11, 2020|title=Novartis receives EU approval for Leqvio(inclisiran), a first-in-class siRNA to lower cholesterol with two doses a year|work=Novartis}}</ref>