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33 gesichtete, geschützte Fragmente: Plagiat

[1.] Ww/Fragment 065 08 - Diskussion
Bearbeitet: 3. February 2016, 22:25 Schumann
Erstellt: 29. October 2014, 12:19 (Hindemith)
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Moreover, McIntyre and Fanning (2006) revealed that shRNA vector construction can be disturbed by high mutation rates and ensuing sequencing is often problematic. shRNA expression vectors are constructed by one of three different methods. The most common method requires the synthesis, annealing and ligation of two complementary oligonucleotides into an expression vector. The frequency of false positives determined by sequencing is high, about 20-40% high (Miyagishi et al, 2004). The unreliability of this method is in part due to the difficulty in synthesizing long oligonucleotides (> 35 bases) and this method requires two long oligonucleotides then the chance of mutation is doubled. The second strategy is a PCR approach in which a promoter sequence serves as the template. Although it is advantageous that only single long oligonucleotides is required, the strong secondary structure predicted to form within this primer can lead to the amplification of false products (Castanotto et al., 2005). The third method comprises several techniques relating to primer extension. Each is based on the principle of a polymerase extending the 3’ end of overlapping oligonucleotides. Nevertheless, this method reduces the cost of oligonucleotides and does not need purification but may cause off-set by a high rate of polymerase-induced mutations in both the initial extension and repeated cycling steps. In order to reduce mutations, conducting all reactions as single-step extensions and replacing Taq polymerase with an enzyme better able to counter the secondary structure of the hairpin template were adopted. Another reported strategy to alleviate sequencing difficulties is to include mismatched bases within the shRNA stem (Yu et al., 2003). A survey of the literature revealed that shRNA vector construction can be hindered by high mutation rates and the ensuing sequencing is often problematic. [...]

[...]

[...] In a survey of more than 100 papers applying expressed shRNA in mammalian systems we determined that shRNA expression vectors are constructed by one of three different methods (see Additional file 1).

[Seite 2]

The most common method for making shRNA constructs (74 % of surveyed studies) requires the synthesis, annealing and ligation of two complementary oligonucleotides (oligos) into an expression vector (Fig. 1b and Additional file 2). While this cloning method is quick, the oligo synthesis cost is nearly double that of other methods and the frequency of false positives determined by sequencing is high (typically 20 – 40 %) [3]. The unreliability of this method is in part due to the difficulty in the synthesis of long oligos (length > 35 bases) [4]. As this method requires two long oligos the chance of mutation due to synthesis error is doubled.

The second strategy (employed in 22 % of studies) is a PCR approach in which a promoter sequence serves as the template (Fig. 1c). [...] Although it is advantageous that only a single long oligo is required, the strong secondary structure predicted to form within this primer can lead to the amplification of false products. [...] [6]

The third method (applied in 4 % of studies) encompasses several techniques relating to primer extension. Each is based on the principle of a polymerase extending the 3' end of overlapping oligos [7]. [...] This technique is the cheapest of all the construction methods discussed as it both halves the cost of unique oligos (compared to the annealed oligo method) and does not need costly oligo purification (compared to the promoter based PCR method). However, this saving may be off-set by a high rate of polymerase-induced mutation in either the initial extension step or by repeated cycling [4].

[Seite 3]

Our first step to reduce mutations was to remove the possibility of cycling-induced errors by conducting all reactions as single-step extensions. [...]

To improve upon these results, we substituted Taq polymerase with an enzyme better able to counter the secondary structure of the hairpin template.

[Seite 4]

Another reported strategy to alleviate sequencing difficulties is to include mismatched bases within the shRNA stem [3,11].


3. Miyagishi M, Sumimoto H, Miyoshi H, Kawakami Y, Taira K: Optimization of an siRNA-expression system with an improved hairpin and its significant suppressive effects in mammalian cells. J Gene Med 2004, 6:715-723.

4. Paddison PJ, Cleary M, Silva JM, Chang K, Sheth N, Sachidanandam R, Hannon GJ: Cloning of short hairpin RNAs for gene knockdown in mammalian cells. Nat Methods 2004, 1:163-167.

6. Castanotto D, Scherer L: Targeting Cellular Genes with PCR Cassettes Expressing Short Interfering RNAs. Methods Enzymol 2005, 392:173-185.

7. Rossi JJ, Kierzek R, Huang T, Walker PA, Itakura K: An alternate method for synthesis of double-stranded DNA segments. J Biol Chem 1982, 257:9226-9229.

11. Yu JY, Taylor J, DeRuiter SL, Vojtek AB, Turner DL: Simultaneous inhibition of GSK3alpha and GSK3beta using hairpin siRNA expression vectors. Mol Ther 2003, 7:228-236.

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[2.] Ww/Fragment 064 15 - Diskussion
Bearbeitet: 3. February 2016, 22:23 Schumann
Erstellt: 26. July 2015, 09:43 (Hindemith)
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Paddison et al. (2002) showed that shRNA were somewhat less potent silencing triggers than that were siRNAs. siRNAs homologous yielded 90-95% suppression of gene expression, whereas suppression levels achieved with shRNAs ranged from 80-90% on average because of mammalian cells contain several endogenous systems that were predicted to hamper the application of RNAi. Chief among these is a dsRNA-activated protein kinase, PKR, which effects a general suppression of translation via phosphorylation of EIF-2α. However, mammalian cells contain several endogenous systems that were predicted to hamper the application of RNAi. Chief among these is a dsRNA-activated protein kinase, PKR, which effects a general suppression of translation via phosphorylation of EIF-2α (Williams 1997; Gil and Esteban 2000). [...]

[...]

[...] Overall, shRNAs were somewhat less potent silencing triggers than were siRNAs. Whereas siRNAs homologous to firefly luciferase routinely yielded ∼90%–95% suppression of gene expression, suppression levels achieved with shRNAs ranged from 80%–90% on average.

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[3.] Ww/Fragment 041 11 - Diskussion
Bearbeitet: 3. February 2016, 22:21 Schumann
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The Midi plasmid purification protocols were based on a modified alkaline lysis procedure, followed by binding of plasmid DNA to Anion-Exchange Resin under appropriate low-salt and pH conditions. RNA, proteins, dyes, and low-molecular-weight impurities were removed by a medium-salt wash. Plasmid DNA was eluted in a high salt buffer and then concentrated and desalted by isopropanol precipitation. This plasmid preparation protocol used in this embodiment is based on a modified alkaline lysis procedure, followed by binding of plasmid DNA to anion-exchange resin under appropriate low-salt and pH conditions. RNA, proteins, dyes, and low-molecular-weight impurities in the plasmid are removed by a medium-salt wash. Plasmid DNA is eluted in a high-salt buffer and then concentrated and desalted by isopropanol precipitation.
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[4.] Ww/Fragment 005 20 - Diskussion
Bearbeitet: 3. February 2016, 22:18 Schumann
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Short hairpin RNAs (shRNAs) are a sequence of RNAs that makes a tight hairpin turn and are transcribed by RNA polymerase Ⅲ. It uses a vector introduced into cells and utilizes the U6 promoter to ensure that the shRNA is always expressed and can be cleaved by the cellular machinery into siRNA, which is then bound to the RISC silencing gene expression via RNAi (Brummelkamp et al., 2002; Lee et al., 2002; Miyagishi and Tiara, 2002; [Paddison, et al. 2002; Paul et al., 2002; Sui et al., 2002; Cao et al, 2005; Harper et al., 2005; McIntyre and Fanning, 2006).] A short hairpin RNA (shRNA) is a sequence of RNA that makes a tight hairpin turn that can be used to silence gene expression via RNA interference. shRNA uses a vector introduced into cells and utilizes the U6 promoter to ensure that the shRNA is always expressed. This vector is usually passed on to daughter cells, allowing the gene silencing to be inherited. The shRNA hairpin structure is cleaved by the cellular machinery into siRNA, which is then bound to the RNA-induced silencing complex (RISC).

[...]

References

McIntyre G, Fanning G (2006). "Design and cloning strategies for constructing shRNA expression vectors". BMC Biotechnol. 6: 1. PMID 16396676.

Harper S, Staber P, He X, Eliason S, Martins I, Mao Q, Yang L, Kotin R, Paulson H, Davidson B (2005). "RNA interference improves motor and neuropathological abnormalities in a Huntington's disease mouse model". Proc. Natl. Acad. Sci. U.S.A. 102 (16): 5820–5. PMID 15811941.

Nielsen M, Pedersen F, Kjems J (2005). "Molecular strategies to inhibit HIV-1 replication". Retrovirology 2: 10. PMID 15715913.

Paddison P, Caudy A, Bernstein E, Hannon G, Conklin D (2002). "Short hairpin RNAs (shRNAs) induce sequence-specific silencing in mammalian cells". Genes Dev. 16 (8): 948–58. PMID 11959843.

Cao W, Hunter R, Strnatka D, McQueen C, Erickson R (2005). "DNA constructs designed to produce short hairpin, interfering RNAs in transgenic mice sometimes show early lethality and an interferon response". J. Appl. Genet. 46 (2): 217–25. PMID 15876690.

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[5.] Ww/Fragment 027 01 - Diskussion
Bearbeitet: 20. January 2016, 18:15 Schumann
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A more specific use of HEK cells is in the propagation of adenoviral vectors.

Viruses offer an extremely efficient means of delivering genes into cells, since this is what they have evolved to do, and are thus of great use as experimental tools. However, as pathogens, they also present a degree of danger to the experimenter. This danger can be avoided by the use of viruses which lack key genes, and which are thus unable to replicate after entering a cell. In order to propagate such viral vectors, a cell line that expresses the missing genes is required. Since HEK cells express a number of adenoviral genes, they can be used to propagate adenoviral vectors in which these genes (typically, E1 and E3) are deleted, such as AdEasy. Another application of 293, especially 293T, cells is commonly used for the production of lentiviral and retroviral vectors. Various retroviral and lentiviral packaging cell lines are based on these cells.

A more specific use of HEK cells is in the propagation of adenoviral vectors. Viruses offer an extremely efficient means of delivering genes into cells, since this is what they have evolved to do, and are thus of great use as experimental tools. However, as pathogens, they also present a degree of danger to the experimenter. This danger can be avoided by the use of viruses which lack key genes, and which are thus unable to replicate after entering a cell. In order to propagate such viral vectors, a cell line that expresses the missing genes is required. Since HEK cells express a number of adenoviral genes, they can be used to propagate adenoviral vectors in which these genes (typically, E1 and E3) are deleted, such as AdEasy (He 1998).

293, and especially 293T, cells are commonly used for the production of lentiviral and retroviral vectors. Various retroviral and lentiviral packaging cell lines are based on these cells.

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[6.] Ww/Fragment 010 01 - Diskussion
Bearbeitet: 20. January 2016, 18:13 Schumann
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[Class III enzymes are further characterized] by a helicase domain and a PAZ (Piwi/Argonaute/Zwille) domain. This last domain is also present in Argonaute family proteins, already known to be essential for RNAi, which led to the proposal that Class III enzymes are the initiator of RNAi (Bass, 2000).

The generation of a siRNA from dsRNA potentially requires four endonucleolytic reactions. It has been revealed that Dicer acts as a monomer, using two endonucleolytic reactions to generate one new terminus (Zhang et al, 2004). This would occur if Dicer binds to an existing terminus and makes a cut ~21 nucleotides from the end (Schütze, 2004). If however the enzyme can not initiate processing from the end and is forced to cut internally, the reaction becomes significantly delayed. Once binding of Dicer occurred and a single new terminus is created, further processing occurrs at normal rates, since the enzyme now has terminal ends from which to process.

MicroRNAs (miRNAs) are transcribed by RNA polymerase II as long primary transcripts (Kim, 2005). The active miRNA species, termed the mature RNA, is present in a stem–loop structure within the primary transcript. The stem–loop can be located in an exon or in an intron. For example, the miRNAs miR-106b, miR-93, and miR-25 are located within an intron of the protein coding gene mcm-7. After transcription, the miRNAs are processed from the primary transcript, and the spliced mRNA is exported and translated . Sequential processing of the primary transcript by the RNase III enzymes Drosha and Dicer liberates the mature RNA. Drosha cleavage releases the stem–loop, termed the precursor, which is exported from the nucleus in an exportin-5/RAN-GTPase-dependent manner. In the cytoplasm, the precursor is processed into a siRNA-like structure by Dicer. Drosha generates a 2 nt 3’ overhang terminus on the precursor which is recognized by the PAZ domain of Dicer, analogous to the recognition of dsRNA termini. The double stranded miRNA is incorporated into RISC in a similar manner as siRNAs.

Drosha is a Class II enzyme. This enzyme assumes a pseudo-dimer catalytic [core similar to Dicer (Han et al, 2004).]

Class III enzymes are further characterized by a helicase domain and a PAZ (Piwi/Argonaute/Zwille) domain. This last domain is also present in Argonaute family proteins, already known to be essential for RNAi, which led to the proposal that Class III enzymes are the initiator of RNAi [12,13]. [...]

The generation of an siRNA from dsRNA potentially requires four endonucleolytic reactions. How does Dicer achieve this? Early models were based on the prediction that Dicer forms a dimer on the substrate and performs four cleavage reactions [15]. Recent data, however, favors a model whereby Dicer acts as a monomer, using two endonucleolytic reactions to generate one new terminus [16,17]. This would occur if Dicer bound to an existing terminus and made a cut ~21 nucleotides from the end. This was first suggested by studies using dsRNA substrates with blocked termini [16]. If the enzyme could not initiate processing from the end and was forced to process internally, the reaction was significantly delayed. The authors' interpretation was that internal binding was less efficient and caused a lag in processing. Once binding occurred and a single new terminus was created, further processing occurred at normal rates, since the enzyme now had terminal ends from which to process.

[Seite 5824]

MicroRNAs are transcribed from RNA polymerase II as long primary transcripts (see [26] for a review). The active microRNA species, termed the mature RNA, is present in a stem–loop structure within the primary transcript. The stem–loop can be located in an exon or an intron. For example, the microRNAs miR-106b, miR-93, and miR-25 are located within an intron of the protein coding gene mcm-7. After transcription, the microRNAs are processed out of the primary transcript, and the spliced mRNA is exported and translated. Whether the microRNA is processed before, during, or after splicing is not known. Sequential processing of the primary transcript by the RNaseIII enzymes Drosha and Dicer liberates the mature RNA. Drosha cleavage releases the stem–loop, termed the precursor, which is exported from the nucleus in an Exportin-5/RAN-GTPase-dependent manner. In the cytoplasm, the precursor is processed into a siRNA-like structure by Dicer. Drosha generates a 2 nt 30 overhang terminus on the precursor which is recognized by the PAZ domain of Dicer, analogous to the recognition of dsRNA termini. The doublestranded microRNA is incorporated into RISC in a similar manner as siRNAs.

Drosha is a Class II enzyme as shown in Fig. 1. This enzyme assumes a pseudo-dimer catalytic core similar to Dicer [27].


[12] Tabara, H., Sarkissian, M., Kelly, W.G., Fleenor, J., Grishok, A., Timmons, L., Fire, A. and Mello, C.C. (1999) The rde-1 gene, RNA interference, and transposon silencing in C. elegans. Cell 99, 123–132.

[13] Bass, B.L. (2000) Double-stranded RNA as a template for gene silencing. Cell 101, 235–238.

[14] Bernstein, E., Caudy, A.A., Hammond, S.M. and Hannon, G.J. (2001) Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409, 363–366.

[15] Zamore, P.D. (2001) Thirty-three years later, a glimpse at the ribonuclease III active site. Mol. Cell 8, 1158–1160.

[16] Zhang, H., Kolb, F.A., Brondani, V., Billy, E. and Filipowicz, W. (2002) Human Dicer preferentially cleaves dsRNAs at their termini without a requirement for ATP. EMBO J. 21, 5875–5885.

[17] Zhang, H., Kolb, F.A., Jaskiewicz, L., Westhof, E. and Filipowicz, W. (2004) Single processing center models for human Dicer and bacterial RNase III. Cell 118, 57–68.

[26] Kim, V.N. (2005) MicroRNA biogenesis: coordinated cropping and dicing. Nat. Rev. Mol. Cell. Biol. 6, 376–385.

[27] Han, J., Lee, Y., Yeom, K.H., Kim, Y.K., Jin, H. and Kim, V.N. (2004) The Drosha-DGCR8 complex in primary microRNA processing. Genes Dev. 18, 3016–3027.

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[7.] Ww/Fragment 029 03 - Diskussion
Bearbeitet: 30. October 2014, 08:32 Hindemith
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Ww 29a diss.png

Fig. 2. Map for the pLKO.1 lentiviral vector. It derives from pRRLSIN.cPPT.PGK/GFP/WRPE and contains elements for efficient viral packaging and shRNA expression. These include Ψ, the lentiviral packaging site; RRE, the Rev-responsive element; cPPT. RSV/5’LTR is a hybrid of the Rous Sarcoma virus promoter and the HIV 5’LTR. SIN 3’LTR is the HIV 3’LTR with a self-inactivating U3 deletion. Expression of the shRNA is driven by the human U6 promoter (hU6). The lentiviral vector also contains the mammalian selection marker puromycin resistance gene (PAC) under the control of the PGK promoter as a mammalian selection marker. pUCori is the bacterial origin of replication of the plasmid, F1 ori is the single-stranded phage F1 origin of replication, and AmpR is the ampicillin resistance gene.

Ww 29a source.png

Figure 1 Vector map for the pLKO.1 lentiviral vector. The self-inactivating lentiviral vector backbone contains elements for efficient viral packaging and shRNA expression. These include Ψ, the lentiviral packaging site; RRE, the Rev-responsive element; cPPT. RSV/ 5′ LTR is a hybrid of the Rous Sarcoma virus promoter and the HIV 5′ LTR. SIN 3′ LTR is the HIV 3′ LTR with a self-inactivating U3 deletion. Expression of the shRNA is driven by the human U6 promoter (hU6). The lentiviral vector also contains the mammalian selection marker puromycin resistance gene (PAC) under the control of the PGK promoter as a mammalian selection marker. pUCori is the bacterial origin of replication of the plasmid, F1 ori is the single-stranded phage F1 origin of replication, and AmpR is the ampicillin resistance gene.

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[8.] Ww/Fragment 007 01 - Diskussion
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shRNAs can also be made for use in plants and other systems, and are not necessarily driven by a U6 promoter. In plants the traditional promoter for strong constitutive [sic] expression (in most plant species) is the cauliflower mosaic virus 35S promoter (CaMV35S), in which case RNA polymerase II is used to express the transcript destined to initiate RNAi. shRNAs can also be made for use in plants and other systems, and are not necessarily driven by a U6 promoter. In plants the traditional promoter for strong consitutive expression (in most plant species) is the cauliflower mosaic virus 35S promoter (CaMV35S), in which case RNA Polymerase II is used to express the transcript destined to initiate RNAi
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RNA interference (RNAi) was a term coined by Fire and coworkers to describe the inhibition of gene expression by double-stranded RNAs (dsRNAs) when introduced into nematode worms (Caenorhabditis elegans). Following on from the studies of Guo and Kemphues (1995), who had found that sense RNA was as effective as antisense RNA for suppressing gene expression in worms, Fire et al. (1998) applied single-stranded antisense RNA and double stranded RNA in their experiments. To their surprise, they found that dsRNA was more effective in producing interference than was either strand individually. After injection into adult C. elegans, single-stranded antisense RNA had a modest effect in diminishing specific gene expression, whereas double-stranded mixtures caused potent and specific interference.

Today we know that RNAi is a multistep process involving the generation of small interfering RNAs (siRNAs) in vivo through the action of the RNase III endonuclease “Dicer”. The resulting 21- to 23-nt siRNAs mediate the degradation of their complementary RNA (Shi, 2003).

RNA interference (RNAi) was a term coined by Fire and coworkers (1998) to describe the inhibition of gene expression by double-stranded RNAs (dsRNAs) when introduced into nematode worms (Caenorhabditis elegans). Following on from the studies of Guo and Kemphues (1995), who had found that sense RNA was as effective as antisense RNA for suppressing gene expression in worms, Fire et al. (1998) applied single-stranded antisense RNA and double stranded RNA in their experiments. To their surprise, they found that dsRNA was more effective at producing interference than was either strand individually. After injection into adult C. elegans, single-stranded antisense RNA had a modest effect in diminishing specific gene expression, whereas double-stranded mixtures caused potent and specific interference (Fire et al., 1998). RNAi is a multistep process involving the generation of small interfering RNAs (siRNAs) in vivo through the action of the RNase III endonuclease ‘Dicer’. The resulting 21- to 23-nt siRNAs mediate degradation of their complementary RNA (Shi, 2003).
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A major breakthrough in the elucidation of the underlying mechanism was the biochemical analysis of RNAi using Drosophila embryo or cell extracts (Tuschl et al., 1999; Hammond et al., 2000; Zamore et al., 2000), which led to the identification of the dsRNA processing enzyme Dicer (Bernstein et al., 2001a) as well as the RNA induced silencing complex, RISC (Hammond et al., 2000), which executes RNAi by using the small dsRNA species generated by Dicer as guidance molecules to target the homologous, endogenous mRNA for [degradation (Elbashir et al., 2001b,c; Zamore et al., 2000).] A major breakthrough in the elucidation of the underlying mechanism was the biochemical analysis of RNAi using Drosophila embryo or cell extracts (Hammond et al., 2000; Tuschl et al., 1999; Zamore et al., 2000), which led to the identification of the dsRNA processing enzyme Dicer (Bernstein et al., 2001a) as well as the RNA induced silencing complex, RISC (Hammond et al., 2000), which executes RNAi by using the small dsRNA species generated by Dicer as guidance molecules to target the homologous, endogenous mRNA for degradation (Elbashir et al., 2001b,c; Zamore et al., 2000).
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These discoveries led to the rapid improvement of RNAi tools, tailored to the needs of the various experimental systems, and triggered intense genetic and biochemical research into the molecular basis and regulation of RNAi (Hammond et al., 2001b; Tijsterman et al., 2002). It became clear that RNAi is a highly conserved mechanism that functions in many different cellular pathways from regulating gene expression to fighting infection and the dangers of mobile genetic elements.

1.2.1 Mechanism of RNAi

The genetic and biochemical analysis of RNAi has led to a model, in which RNAi can be divided into two distinct phases: an initiation and an execution phase. The initiation phase involves the processing of dsRNA into siRNA. In the execution phase, siRNAs are then incorporated into large ribonucleoprotein complexes. These effector complexes interfere with gene expression by using the small RNA strand to identify their complementary mRNA, which becomes cleaved and degraded. In a related pathway, short non-coding single stranded RNAs, which are derived from partially complementary dsRNA precursor molecules, are used to regulate the translation of mRNAs harbouring complementary sequences in their 3’'UTRs (Fig. 1).

These discoveries led to the rapid improvement of RNAi tools, tailored to the needs of the various experimental systems, and triggered intense genetic and biochemical

[Seite 986]

research into the molecular basis and regulation of RNAi (Hammond et al., 2001b: Tijsterman et al., 2002). It became clear that RNAi is a highly conserved mechanism that functions in many different cellular pathways from regulating gene expression to fighting infection and the dangers of mobile genetic elements.

[...]

2. The RNAi mechanism

The genetic and biochemical analysis of RNAi has led to a model, in which RNAi can be divided into two distinct phases: an initiation and an execution phase. The initiation phase involves processing of dsRNA into small RNA molecules, called small interfering RNAs (siRNA). In the execution phase, siRNAs are then incorporated into large ribonucleoprotein complexes. These effector complexes interfere with gene expression by using the small RNA strand to identify their complementary mRNA, which becomes cleaved and degraded. In a related pathway, short non-coding single stranded RNAs, which are derived from partially complementary dsRNA precursor molecules, are used to regulate the translation of mRNAs harbouring complementary sequences in their 3' UTRs.

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1.2.1.2 Execution phase: assembly of siRNA containing silencing complexes

Dicer-generated siRNAs are then incorporated into a large multiprotein complex, which is involved in various gene-silencing modes, and is called the RNA induced silencing complex, or RISC (Hammond et al, 2000; Nykanen et al, 2001). Processing of dsRNA and assembly of a functional RISC likely occurs in the cytoplasm, as Dicer is a cytosolic enzyme and RISC activity can be purified from the cytosol (Billy et al, 2001). R2D2, a Drosophila gene related to the C. elegans RNAi gene RDE-4, has been implicated in the transfer of siRNAs into the RISC (Liu et al, 2003). Generation of siRNAs from dsRNA in Drosophila embryo extracts, unwinding of the siRNA duplex, and incorporation into the RISC requires ATP (Nykanen et al, 2001). In contrast, human Dicer does not seem to rely on ATP for processing of dsRNA into siRNA molecules (Zhang et al., 2002).

2.2. The execution phase: assembly of siRNA containing silencing complexes

Dicer-generated siRNAs are then incorporated into a large multiprotein complex, which is involved in various gene-silencing modes, and is called the RNA induced silencing complex, or RISC (Hammond et al., 2000; Nykanen et al., 2001). Processing of dsRNA and assembly of a functional RISC likely occurs in the cytoplasm, as Dicer is a cytosolic enzyme and RISC activity can be purified from cytosol (Billy et al., 2001). R2D2, a Drosophila gene related to the C. elegans RNAi gene RDE-4, has been implicated in the transfer of siRNAs into the RISC (Liu et al., 2003). Generation of siRNAs from dsRNA in Drosophila embryo extracts, unwinding of the siRNA duplex, and incorporation into the RISC require ATP (Nykanen et al., 2001). In contrast, human Dicer does not seem to require ATP for processing of dsRNA into siRNA molecules (Zhang et al., 2002).

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[Therefore, a stoichiometric excess of a virus-specific siRNA, for] example, could saturate RNAi and interrupt the pathway’s normal functions in the cell. Interferons, which form part of the host’s defense against viral infection, are activated by long dsRNA (more than 500 bp). It is now apparent that siRNAs (Sledz et al, 2003) as well as shRNAs expressed from DNA vectors (Bridge et al, 2003) can trigger the activation of interferons. However, there is no evidence that the activation of interferons by short RNAs influences the degree or specificity of RNA silencing. In addition, these effects have to be reconciled with the manufacturing in cells of many thousands of copies of pre-miRNAs (Lagos et al, 2001) that do not appear to activate interferons. Therefore, a stoichiometric excess of a virus-specific siRNA, for example, could saturate RNAi and interrupt the pathway's normal functions in the cell.

Interferons, which form part of the host's defense against viral infection, are activated by long dsRNA (more than 500 bp). It is now apparent that siRNAs45 as well as short hairpin RNAs (short sequences of RNA that make tight hairpin turns and can be used to silence gene expression) expressed from DNA vectors46 can trigger the activation of interferons. However, there is no evidence that the activation of interferons by short RNAs influences the degree or specificity of RNA silencing. In addition, these effects have to be reconciled with the manufacturing in cells of many thousands of copies of pre-micro RNAs47,48,49 that do not appear to activate interferons.


45. Sledz CA, Holko M, de Veer MJ, Silverman RH, Williams BR. Activation of the interferon system by short-interfering RNAs. Nat Cell Biol 2003;5:834-9.

46. Bridge AJ, Pebernard S, Ducraux A, Nicoulaz AL, Iggo R. Induction of an interferon response by RNAi vectors in mammalian cells. Nat Genet 2003;34:263-4.

47. Lau NC, Lim LP, Weinstein EG, Bartel DP. An abundant class of tiny RNAs with probable regulatory roles in Caenorhabditis elegans. Science 2001;294:858-62.

48. Lee RC, Ambros V. An extensive class of small RNAs in Caenorhabditis elegans. Science 2001;294:862-4.

49. Lagos-Quintana M, Rauhut R, Lendeckel W, Tuschl T. Identification of novel genes coding for small expressed RNAs. Science 2001;294:853-8.

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Delivery is probably the single biggest obstacle to the development of RNAi-based therapeutic agents. Trigger RNAs (dsRNAs from which siRNAs are derived by the action of Dicer) can be expressed from vectors or delivered as artificial siRNAs. A variety of strategies to express interfering RNAs with the use of plasmid and virus vector-based cassettes have been explored (Li et al, 2002; Dykxhoorn et al, 2003). Well-documented hazards of inserting foreign vector sequences into chromosomal DNA include insertional activation and inactivation of cellular genes. Direct (intravenous) administration of siRNAs would require siRNAs that are modified to be resistant to nucleases and perhaps conjugated with a ligand to target the siRNA to specific tissues. In mice, intravenous introduction of Fas siRNAs leads to specific silencing of Fas mRNA in the liver (Song et al, 2003), so in principle, unmodified siRNAs can be taken up by the liver and perhaps other tissues. It is not clear, however, whether there are selective tissue sites for the uptake of siRNAs and whether the lymphoid system or the brain, for instance, is accessible by this route. Furthermore, the silencing effect of siRNAs is short-lived, because the siRNAs eventually decay within the cell. In addition to the danger of using vectors that integrate into the genome, the expression or injection of siRNAs may also have unwanted biologic side effects. Researchers are continually finding new cellular processes in which RNAi is involved. Therefore, a stoichiometric excess of a virus-specific siRNA, for [example, could saturate RNAi and interrupt the pathway’s normal functions in the cell. Interferons, which form part of the host’s defense against viral infection, are activated by long dsRNA (more than 500 bp).] Delivery is probably the single biggest obstacle to the development of RNAi-based therapeutic agents. Trigger RNAs (dsRNAs from which siRNAs are derived by the action of Dicer) can be expressed from vectors or delivered as artificial siRNAs. A variety of strategies to express interfering RNAs with the use of plasmid and virus vector-based cassettes have been explored.7,37 Well-documented hazards of inserting foreign vector sequences into chromosomal DNA include insertional activation and inactivation of cellular genes.

Direct (e.g., intravenous) administration of siRNAs would require siRNAs that are modified to be resistant to nucleases and perhaps conjugated with a ligand to target the siRNA to specific tissues. In mice, intravenous introduction of Fas siRNAs leads to specific silencing of Fas mRNA in the liver,25 so in principle, unmodified siRNAs can be taken up by the liver and perhaps other tissues. It is not clear, however, whether there are selective tissue sites for the uptake of siRNAs and whether the lymphoid system or the brain, for instance, is accessible by this route. Furthermore, the silencing effect of siRNAs is short-lived, because the siRNAs eventually decay within the cell.

In addition to the danger of using vectors that integrate into the genome, the expression or injection of siRNAs may also have untoward biologic effects. Researchers are continually finding new cellular processes in which RNAi is involved. Therefore, a stoichiometric excess of a virus-specific siRNA, for example, could saturate RNAi and interrupt the pathway's normal functions in the cell.

Interferons, which form part of the host's defense against viral infection, are activated by long dsRNA (more than 500 bp).


7. Dykxhoorn DM, Novina CD, Sharp PA. Killing the messenger: short RNAs that silence gene expression. Nat Rev Mol Cell Biol 2003;4:457-67.

25. Song E, Lee SK, Wang J, et al. RNA interference targeting Fas protects mice from fulminant hepatitis. Nat Med 2003;9:347- 51.

37. Li H, Li WX, Ding SW. Induction and suppression of RNA silencing by an animal virus. Science 2002;296:1319-21.

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The translocation of the Philadelphia chromosome (Ph) generates a fusion gene called BCR-ABL. The translation product of this gene creates a constitutively active protein tyrosine kinase that induces and maintains leukemic transformation in chronic myelogenous leukemia and Ph-positive acute lymphoblastic leukemia. The siRNAs specific for the BCR-ABL transcript have been shown to silence the oncogenic fusion transcripts without affecting expression levels of normal c-ABL and c-BCR transcripts (Scherr et al, 2003; Wohlbold et al, 2003).

Pancreatic and colon carcinomas, in which RAS genes are often mutated, provide another example for potential RNAi applications. In many cases, the RAS oncogenes contain point mutations that differ by a single-base mutation from their normal counterparts. The use of retroviral vectors to introduce interfering RNAs specific for an oncogenic variant of KRAS (called K-RASV12) reduced the level of K-RASV12 transcripts and resulted in a loss of anchorage-independent growth and tumorigenicity (Brummelkamp et al, 2002). Studies of this kind provided proof-of-concept data for RNAi-based strategies aiming to reverse tumorigenesis.

1.2.2.2 Infectious diseases

The translocation of the Philadelphia chromosome (Ph) generates a fusion gene called BCR-ABL. The translation product of this gene creates a constitutively active protein tyrosine kinase that induces and maintains leukemic transformation in chronic myelogenous leukemia and Ph-positive acute lymphoblastic leukemia. The siRNAs specific for the BCR-ABL transcript have been shown to silence the oncogenic fusion transcripts without affecting expression levels of normal c-ABL and c-BCR transcripts.20,21

Pancreatic and colon carcinomas, in which RAS

[Seite 1775]

genes are often mutated, provide another example. In many cases, the RAS oncogenes contain point mutations that differ by a single-base mutation from their normal counterparts. The use of retroviral vectors to introduce interfering RNAs specific for an oncogenic variant of K-RAS (called K-RASV12) reduces the level of K-RASV12 transcripts and effects a loss of anchorage-independent growth and tumorigenicity.22,23 Studies of this kind provide proof-of-concept for RNAi-based strategies aimed at reversing tumorigenesis. [...]

[...]

Infectious Diseases


20. Scherr M, Battmer K, Winkler T, Heidenreich O, Ganser A, Eder M. Specific inhibition of bcr-abl gene expression by small interfering RNA. Blood 2003;101: 1566-9.

21. Wohlbold L, van der Kuip H, Miething C, et al. Inhibition of bcr-abl gene expression by small interfering RNA sensitizes for imatinib mesylate (STI571). Blood 2003; 102:2236-9.

22. Brummelkamp TR, Bernards R, Agami R. Stable suppression of tumorigenicity by virus-mediated RNA interference. Cancer Cell 2002;2:243-7.

23. Wilda M, Fuchs U, Wossmann W, Borkhardt A. Killing of leukemic cells with a BCR/ABL fusion gene by RNA interference (RNAi). Oncogene 2002;21:5716-24.

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1.2.2 Therapeutic applications of RNAi

The therapeutic applications of RNAi are potentially enormous. The genetic etiology of many disorders has now been defined and, in some cases, has been targeted by RNAi in in vitro and in vivo model systems. Because the specificity of RNAi is governed by sequence complementarity between the siRNA and the target RNA, the most obvious application would be to treat diseases in which genetic polymorphisms within the disease-inducing gene in a particular lesion or tumor can be targeted for degradation without affecting RNA from wild-type alleles.

Therapeutic Applications

The therapeutic applications of RNAi are potentially enormous. The genetic etiology of many disorders has now been defined and, in some cases, has been targeted by RNAi in in vitro and in vivo model systems. Because the specificity of RNAi is governed by sequence complementarity between the siRNA and the target RNA, the most obvious application would be to treat diseases in which genetic polymorphisms within the disease-inducing gene in a particular lesion or tumor can be targeted for degradation without affecting RNA from wild-type alleles.

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Structurally, this protein family is characterized by two domains, the PAZ domain and the PIWI domain. Structures for both domains have been solved. Additional RISC components with unknown roles in RNAi have also been identified. These include the RNA binding protein VIG, the Drosophila homolog of the Fragile X protein, dFXR, helicase proteins, and Tudor-SN (Ishizuka et al, 2002; Caudy et al, 2003). This last protein has five staphylococcal nuclease (SNase) domains and a Tudor domain. In humans, there are four closely related Argonaute family members, named Ago1-4. All four bind siRNAs and miRNAs at similar levels, and are widely expressed. Only Ago2, however, is present in a cleavage-competent RISC-complex (Meister et al, 2004). Similarly, siRNA-mediated knockdown, or targeted knockout, of Ago2 impaired RNAi of a reporter, while knockdown of Ago1, 3, 4 had no effect. The crystal structure of an Argonaute family member from Pyrococcus furiosus has been revealed (Song et al, 2004). The structure displayed an RNaseH fold for the signature PIWI domain. The crystal structure of a second archaean argonaute, Archaeoglobus fulgidus Piwi (AfPiwi), confirmed the RNaseH fold (Parker et al, 2004). The final demonstration that Slicer activity was contained within Ago2 was the reconstitution of minimal RISC with bacterially expressed, purified Ago2 and a single-stranded siRNA (Rivas et al, 2005). Structurally, this protein family is characterized by two domains, the PAZ domain and the PIWI domain. Structures for both domains have been solved (see below). Additional RISC components with unknown roles in RNAi

[5826:]

have also been identified. These include the RNA binding protein VIG, the Drosophila homolog of the Fragile X protein, dFXR, helicase proteins, and Tudor-SN [46–48]. This last protein has five staphylococcal nuclease (SNase) domains and a Tudor domain. [...]

[...]

[...] In humans, there are four closely related Argonaute family members, named Ago1-4. All four bind siRNAs and microRNAs at similar levels, and are widely expressed. Only Ago2, however, is present in a cleavage- competent RISC [52,53]. Similarly, siRNA-mediated knockdown, or targeted knockout, of Ago2 impairs RNAi of a reporter, while knockdown of Ago1, 3, 4 had no effect. These data can be interpreted in two ways: Ago2 alone is capable of interacting with Slicer, or Ago2 itself is Slicer. The answer was provided by the crystal structure of an Argonaute family member from Pyrococcus furiosus [54]. The structure revealed an RNaseH fold for the signature PIWI domain. The crystal structure of a second archaean Argonaute, Archaeoglobus fulgidus Piwi (AfPiwi), confirmed the RNaseH fold [55]. The final demonstration that Slicer activity was contained within Ago2 was the reconstitution of minimal RISC with bacterially expressed, purified Ago2 and a single-stranded siRNA [45].


[46] Caudy, A.A., Myers, M., Hannon, G.J. and Hammond, S.M. (2002) Fragile X-related protein and VIG associate with the RNA interference machinery. Genes Dev. 16, 2491–2496.

[47] Caudy, A.A. et al. (2003) A micrococcal nuclease homologue in RNAi effector complexes. Nature 425, 411–414.

[48] Ishizuka, A., Siomi, M.C. and Siomi, H. (2002) A Drosophila fragile X protein interacts with components of RNAi and ribosomal proteins. Genes Dev. 16, 2497–2508.

[52] Meister, G., Landthaler, M., Patkaniowska, A., Dorsett, Y., Teng, G. and Tuschl, T. (2004) Human Argonaute2 mediates RNA cleavage targeted by miRNAs and siRNAs. Mol. Cell 15, 185–197.

[53] Liu, J. et al. (2004) Argonaute2 is the catalytic engine of mammalian RNAi. Science 305, 1437–1441.

[54] Song, J.J., Smith, S.K., Hannon, G.J. and Joshua-Tor, L. (2004) Crystal structure of Argonaute and its implications for RISC slicer activity. Science 305, 1434–1437.

[55] Parker, J.S., Roe, S.M. and Barford, D. (2004) Crystal structure of a PIWI protein suggests mechanisms for siRNA recognition and slicer activity. EMBO J. 23, 4727–4737.

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Chromatographic purification of RISC nuclease activity from Drosophila cells revealed several RISC components. The first identified component was Argonaute2 (Ago2) (Hammond et al, 2001). This protein is a member of a gene family conserved in most eukaryotic and several prokaryotic genomes. The C. elegans homolog, rde-1, was previously identified in a genetic screen for RNAi-deficient mutants, reinforcing its connection with RNAi (Tabara et al, [1999).] Chromatographic purification of RISC nuclease activity from Drosophila cells revealed several RISC components. The first identified component was Argonaute2 (Ago2) [43]. This protein is a member of a gene family conserved in most eukaryotic and several prokaryotic genomes. The C. elegans homolog, rde-1, was previously identified in a genetic screen for RNAi-deficient mutants, reinforcing its connection with RNAi [12].

[12] Tabara, H., Sarkissian, M., Kelly, W.G., Fleenor, J., Grishok, A., Timmons, L., Fire, A. and Mello, C.C. (1999) The rde-1 gene, RNA interference, and transposon silencing in C. elegans. Cell 99, 123–132.

[43] Hammond, S.M., Boettcher, S., Caudy, A.A., Kobayashi, R. and Hannon, G.J. (2001) Argonaute2, a link between genetic and biochemical analyses of RNAi. Science 293, 1146–1150

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[Drosha is a Class II enzyme. This enzyme assumes a pseudo-dimer catalytic] core similar to Dicer (Han et al, 2004). The substrates of Drosha, miRNA primary transcripts, are structurally distinct from Dicer substrates. Drosha does not process from a dsRNA terminus. Rather, data suggests that primarily the stem–loop structure is recognized. In particular, the loop size appears to be important for recognition (Zeng et al, 2005). In addition, unstructured sequences flanking the stem–loop are essential for processing (Chen et al, 2004; Zeng et al, 2005). It is not evident how Drosha is able to recognize these sequences, as they are outside of the dsRNA stem. Possibly other unidentified cofactors play a role. Conserved sequence elements have been found in flanking regions of C. elegans miRNAs (Ohler et al, 2004). Drosha is a Class II enzyme as shown in Fig. 1. This enzyme assumes a pseudo-dimer catalytic core similar to Dicer [27]. The substrate of Drosha, microRNA primary transcripts, is structurally distinct from Dicer substrates. Drosha does not process from a dsRNA terminus. Rather, data suggests that the stem–loop structure is recognized. In particular, the loop size appears to be important for recognition [28]. In addition, unstructured sequences flanking the stem–loop are essential for processing [29,30]. It is not evident how Drosha would recognize these sequences, as they are outside of the dsRNA stem. Possibly other, unidentified cofactors play a role. Conserved sequence elements have been found in flanking regions of C. elegans microRNAs [31].

[27] Han, J., Lee, Y., Yeom, K.H., Kim, Y.K., Jin, H. and Kim, V.N. (2004) The Drosha-DGCR8 complex in primary microRNA processing. Genes Dev. 18, 3016–3027.

[28] Zeng, Y., Yi, R. and Cullen, B.R. (2005) Recognition and cleavage of primary microRNA precursors by the nuclear processing enzyme Drosha. EMBO J. 24, 138–148.

[29] Chen, C.Z., Li, L., Lodish, H.F. and Bartel, D.P. (2004) MicroRNAs modulate hematopoietic lineage differentiation. Science 303, 83–86.

[30] Zeng, Y. and Cullen, B.R. (2005) Efficient processing of primary microRNA hairpins by Drosha requires flanking non-structured RNA sequences. J. Biol. Chem..

[31] Ohler, U., Yekta, S., Lim, L.P., Bartel, D.P. and Burge, C.B. (2004) Patterns of flanking sequence conservation and a characteristic upstream motif for microRNA gene identification. RNA 10, 1309–1322

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The goal of the initiation step of RNAi is the generation of siRNAs from long dsRNAs or mature miRNAs from their primary transcripts. This is achieved by the action of two families of RNase III-dependent genes, Dicer and Drosha. RNase III enzymes fall into three classes (Nicholson, 2003). Class I enzymes, found in bacteria and yeast, contain a single RNase III domain joined to a dsRBD (dsRNA binding domain proteins). Class II and III enzymes contain two RNase III catalytic domains. Class III enzymes are further characterized [by a helicase domain and a PAZ (Piwi/Argonaute/Zwille) domain.] The goal of the initiator step of RNAi is the generation of siRNAs from long dsRNAs, or mature microRNAs from their primary transcripts. This is achieved by the action of two families of RNase III genes, Dicer and Drosha.

[Seite 5823]

RNaseIII enzymes fall into three classes (see Fig. 1, [11] for a review). Class I enzymes, found in bacteria and yeast, contain a single RNaseIII domain joined to a dsRBD. Class II and III enzymes contain two RNaseIII catalytic domains. Class III enzymes are further characterized by a helicase domain and a PAZ (Piwi/Argonaute/Zwille) domain.


[11] Nicholson, A.W. (2003) The ribonuclease III superfamily: forms and functions in RNA maturation, decay, and gene silencing (Hannon, G.J., Ed.), RNAi: A Guide to Gene Silencing, vol. 8, pp. 149–174, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY.

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PLC-β4 function as effector enzymes for receptors belonging to the rhodopsin superfamily of transmembrane proteins that contain seven transmembrane spanning (heptahelical) segments (Ji et al, 1998). They are activated by a wide range of stimuli, from photons and tiny odorant molecules, to full-sized proteins and require specific combinations of Gɑ subunits to couple to their effectors. In the standard G protein model of PLC-β4 activation, binding of agonist triggers receptor-catalyzed exchange of GTP for bound GDP on the ɑ-component of the heterotrimer. The GTP-charged subunit then dissociates in the plane of the membrane, increasing its catalytic activity and thereby amplifying the initial receptor stimulus.

PLC-β4 was first isolated from cerebellum and retina (Min et al, 1993; Jiang et al, 1994). Its mRNA is highly concentrated in cerebellar Purkinje and granule cells, the median geniculate body, whose axons terminate in the auditory cortex, and the lateral geniculate nucleus, where most retinal axons terminate [in a visuotopic representation of each half of the visual field.]

PLC-β4 was first isolated from cerebellum (244, 245) and retina (173, 210). Its mRNA is highly concentrated in cerebellar Purkinje and granule cells (308, 362), the median geniculate body (308), whose axons terminate in the auditory cortex, and the lateral geniculate nucleus, where most retinal axons terminate in a visuotopic representation of each half of the visual field.

PLC-β isoforms function as effector enzymes for receptors belonging to the rhodopsin superfamily of trans-

[Seite 1299]

membrane proteins that contain seven transmembrane spanning (heptahelical) segments (169). They are activated by a wide range of stimuli, from photons and tiny odorant molecules, to full-sized proteins and require specific combinations of Gɑ and Gβγ subunits to couple to their effectors. In the standard G protein model of PLC activation, binding of agonist triggers receptor-catalyzed exchange of GTP for bound GDP on the ɑ-component of the heterotrimer. The GTP-charged subunit then dissociates in the plane of the membrane, and either the ɑ-subunit monomer, the βγ-heterodimer, or both bind to PLC-β, increasing its catalytic activity and thereby amplifying the initial receptor stimulus (Fig. 2).


169. JI TH, GROSSMANN M, AND JI I. G-protein coupled receptors. J Biol Chem 273: 17299–17302, 1998.

173. JIANG H, WU D, AND SIMON MI. Activation of phospholipase C beta 4 by heterotrimeric GTP-binding proteins. J Biol Chem 269: 7593– 7596, 1994

210. LEE CW, LEE KH, AND RHEE SG. Characterization of phospholipase C isozymes in bovine retina: purification of phospholipase C-beta 4. Methods Enzymol 238: 227–237, 1994

244. MIN DS, KIM DM, LEE YH, SEO J, SUH PG, AND RYU SH. Purification of a novel phospholipase C isozyme from bovine cerebellum. J Biol Chem 268: 12207–12212, 1993.

245. MIN DS, KIM Y, LEE YH, SUH PG, AND RYU SH. A G-protein-coupled 130 kDa phospholipase C isozyme, PLC-beta 4, from the particulate fraction of bovine cerebellum. FEBS Lett 331: 38–42, 1993.

308. ROUSTAN P, ABITBOL M, MENINI C, RIBEAUDEAU F, GERARD M, VEKEMANS M, MALLET J, AND DUFIER JL. The rat phospholipase C beta 4 gene is expressed at high abundance in cerebellar Purkinje cells. Neuroreport 6: 1837–1841, 1995.

362. TANAKA O AND KONDO H. Localization of mRNAs for three novel members (beta 3, beta 4 and gamma 2) of phospholipase C family in mature rat brain. Neurosci Lett 182: 17–20, 1994.

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[In such cases, the formation of the double-stranded RNA through the binding of the miRNA triggers the degradation of the mRNA transcript through a process similar to RNAi, though] in other cases it is believed that the miRNA complex blocks the protein translation machinery or otherwise prevents protein translation without causing the mRNA to be degraded. miRNAs may also target methylation of genomic sites which correspond to targeted mRNAs. miRNAs function in association with a complement of proteins collectively termed the miRNP (micro-ribonucleic protein).

This effect was first described for the worm C. elegans in 1993 (Lee et al., 1993). As of 2002, miRNAs have been confirmed in various plants and animals, including C. elegans, human and the plant Arabidopsis thaliana. Genes have been found in bacteria that are similar in the sense that they control mRNA abundance or translation by binding a mRNA by base pairing, however they are not generally considered to be miRNAs because the Dicer enzyme is not involved.

In plants, similar RNA species termed short-interfering RNAs are used to prevent the transcription of viral RNA. While this siRNA is double-stranded, the mechanism seems to be closely related to that of miRNAs, especially taking the hairpin structures into account. siRNAs are also used to regulate cellular genes, as miRNAs do.

In such cases, the formation of the double-stranded RNA through the binding of the miRNA triggers the degradation of the mRNA transcript through a process similar to RNA interference (RNAi), though in other cases it is believed that the miRNA complex blocks the protein translation machinery or otherwise prevents protein translation without causing the mRNA to be degraded. miRNAs may also target methylation of genomic sites which correspond to targeted mRNAs. miRNAs function in association with a complement of proteins collectively termed the miRNP.

This effect was first described for the worm C. elegans in 1993 by Victor Ambros and coworkers (Lee et al., 1993). As of 2002, miRNAs have been confirmed in various plants and animals, including C. elegans, human and the plant Arabidopsis thaliana. Genes have been found in bacteria that are similar in the sense that they control mRNA abundance or translation by binding an mRNA by base pairing, however they are not generally considered to be miRNAs because the Dicer enzyme is not involved.

In plants, similar RNA species termed short-interfering RNAs siRNAs are used to prevent the transcription of viral RNA. While this siRNA is double-stranded, the mechanism seems to be closely related to that of miRNA, especially taking the hairpin structures into account. siRNAs are also used to regulate cellular genes, as miRNAs do.

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[There are few possible explanations for] such selectivity. One could be that dsRNAs longer than 21 base pairs activate an interferon response and the anti-viral machinery in the cell. Another plausible explanation could be that the thermodynamical profile of

pre-miRNAs determines which strand will be incorporated into the Dicer complex. Indeed, the aforementioned study by Han et al. demonstrated very clear similarities between pri-miRNAs encoded in respective (5’- or 3’-) strands.

When Dicer cleaves the pre-miRNA stem-loop, two complementary short RNA molecules are formed, but only one becomes integrated into the RISC complex. This strand is known as the guide strand and is selected by the argonaute protein, the catalytically active RNase in the RISC complex, on the basis of the stability of the 5’ end (Preall et al, 2006). The remaining strand, known as the anti-guide or passenger strand is degraded as a RISC complex substrate (Gregory et al, 2005). After integration into the active RISC complex, miRNAs base pair with their complementary mRNA molecules and induce mRNA degradation by argonaute proteins. It is as yet unclear how the activated RISC complex locates the mRNA targets in the cell, though it has been shown that the process is not coupled to ongoing protein translation from the mRNA (Sen et al, 2005).

1.1.3.2 Cellular functions of miRNA

The miRNAs appear to be important for gene regulation. An individual miRNA is complementary to a part of one or more messenger RNAs (mRNAs). Animal miRNAs are usually complementary to a site in the 3’ UTR whereas plant miRNAs are usually complementary to coding regions of mRNAs. The annealing of the miRNA to the mRNA then inhibits protein translation, but sometimes facilitates cleavage of the mRNA. This is thought to be the primary mode of action of plant miRNAs. In such cases, the formation of the double-stranded RNA through the binding of the miRNA triggers the degradation of the mRNA transcript through a process similar to RNAi, though [in other cases it is believed that the miRNA complex blocks the protein translation machinery or otherwise prevents protein translation without causing the mRNA to be degraded.]

There are few possible explanations for such selectivity. One could be that dsRNAs longer than 21 base pairs activate interferon response and anti-viral machinery in the cell. Another plausible explanation could be that thermodynamical profile of pre-miRNA determines which strand will be incorporated into Dicer complex. Indeed, aforementioned study by Han et al. demonstrated very clear similarities between pri-miRNAs encoded in respective (5'- or 3'-) strands.

When Dicer cleaves the pre-miRNA stem-loop, two complementary short RNA molecules are formed, but only one is integrated into the RISC complex. This strand is known as the guide strand and is selected by the argonaute protein, the catalytically active RNase in the RISC complex, on the basis of the stability of the 5' end.[6] The remaining strand, known as the anti-guide or passenger strand, is degraded as a RISC complex substrate.[7] After integration into the active RISC complex, miRNAs base pair with their complementary mRNA molecules and induce mRNA degradation by argonaute proteins, the catalytically active members of the RISC complex. It is as yet unclear how the activated RISC complex locates the mRNA targets in the cell, though it has been shown that the process is not coupled to ongoing protein translation from the mRNA.[8]

Cellular functions

The function of miRNAs appears to be in gene regulation. For that purpose, a miRNA is complementary to a part of one or more messenger RNAs (mRNAs). Animal miRNAs are usually complementary to a site in the 3' UTR whereas plant miRNAs are usually complementary to coding regions of mRNAs. The annealing of the miRNA to the mRNA then inhibits protein translation, but sometimes facilitates cleavage of the mRNA. This is thought to be the primary mode of action of plant miRNAs. In such cases, the formation of the double-stranded RNA through the binding of the miRNA triggers the degradation of the mRNA transcript through a process similar to RNA interference (RNAi), though in other cases it is believed that the miRNA complex blocks the protein translation machinery or otherwise prevents protein translation without causing the mRNA to be degraded.


6. Preall JB, He Z, Gorra JM, Sontheimer EJ. (2006). Short interfering RNA strand selection is independent of dsRNA processing polarity during RNAi in Drosophila. Curr Biol 16(5):530-5.

7. Gregory RI, Chendrimada TP, Cooch N, Shiekhattar R. (2005). Human RISC couples microRNA biogenesis and posttranscriptional gene silencing. Cell 123(4):631-40.

8. Sen GL, Wehrman TS, Blau HM. (2005). mRNA translation is not a prerequisite for small interfering RNA-mediated mRNAs cleavage. Differentiation 73(6):287-93.

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[This] processing is performed in animals by a protein complex known as the Microprocessor complex, consisting of the nuclease Drosha and the double-stranded RNA binding protein Pasha (Denli et al, 2004). These pre-miRNAs are then processed to mature miRNAs in the cytoplasm by interaction with the endonuclease Dicer, which also initiates the formation of the RNA-induced silencing complex (RISC) (Bernstein et al, 2001). This complex is responsible for the gene silencing observed due to miRNA expression and RNA interference. The pathway in plants varies slightly due to the lack of Drosha homologs; instead, Dicer homologs alone affect several processing steps (Kurihara and Watanabe, 2004).

It has been shown that the efficient processing of pre-miRNAs by Drosha requires the presence of extended single-stranded RNAs on both 3’- and 5’-ends of hairpin molecules (Zeng et al, 2005). This study showed that these motifs could be of different composition while their defined length is of high importance for processing to take place. Findings were confirmed in another work by Han et al (2004). Using bioinformatics tools the folding of 321 human and 68 fly pri-miRNAs was analysed. 280 human and 55 fly pri-miRNAs were selected for further study excluding those molecules where folding showed the presence of multiple loops. All human and fly pri-miRNAs contained very similar structural regions, which the authors called ‘’basal segments’’, ‘’lower stem’’, ’’upper stem’’ and ‘’terminal loop’’. Based on the encoding position of miRNAs, in the 5’-strand (5’-donors) or 3’-strand (3’-donors), thermodynamic profiles of pri-miRNAs were determined (Zeng et al, 2005). Subsequent experiments showed that Drosha complex cleaves RNA molecules ~2 helical turns away from the terminal loop and ~1 turn away from basal segments. In most analyzed molecules this region contains unpaired nucleotides and the free energy of the duplex is relatively high compared to lower and upper stem regions (Zeng and Cullen, 2005).

Most pre-miRNAs don’t have a perfect double-stranded RNA (dsRNA) structure topped by a terminal loop. There are few possible explanations for [such selectivity.]

This processing is performed in animals by a protein complex known as the Microprocessor complex, consisting of the nuclease Drosha and the double-stranded RNA binding protein Pasha.[3] These pre-miRNAs are then processed to mature miRNAs in the cytoplasm by interaction with the endonuclease Dicer, which also initiates the formation of the RNA-induced silencing complex (RISC).[4] This complex is responsible for the gene silencing observed due to miRNA expression and RNA interference. The pathway in plants varies slightly due to their lack of Drosha homologs; instead, Dicer homologs alone effect several processing steps.[5]

Zeng et al. have shown that efficient processing of pre-miRNA by Drosha requires presence of extended single-stranded RNA on both 3'- and 5'-ends of hairpin molecule. They demonstrated that these motifs could be of different composition while their length is of high importance if processing is to take place at all. Their findings were confirmed in another work by Han et al. Using bioinformatical tools Han et al. analysed folding of 321 human and 68 fly pri-miRNAs. 280 human and 55 fly pri-miRNAs were selected for further study, excluding those molecules which folding showed presence of multiple loops. All human and fly pri-miRNA contained very similar structural regions, which authors called 'basal segments', 'lower stem', 'upper stem' and 'terminal loop'. Based on the encoding position of miRNA, i.e. in the 5'-strand (5'-donors) or 3'-strand (3'-donors), thermodynamical profiles of pri-miRNA were determined. Following experiments have shown that Drosha complex cleaves RNA molecule ~2 helical turns away from the terminal loop and ~1 turn away from basal segments. In most analysed molecules this region contains unpaired nucleotides and the free energy of the duplex is relatively high compared to lower and upper stem regions.

Most pre-miRNAs don't have a perfect double-stranded RNA (dsRNA) structure topped by a terminal loop. There are few possible explanations for such selectivity. One could be that dsRNAs longer than 21 base pairs activate interferon response and anti-viral machinery in the cell.


3. Denli AM, Tops BB, Plasterk RH, Ketting RF, Hannon GJ. (2004). Nature 432(7014):231-5.

4. Bernstein E, Caudy AA, Hammond SM, Hannon GJ. (2001). Role for a bidentate ribonuclease in the initiation step of RNA interference. Nature 409(6818):363-6.

5. Kurihara Y, Watanabe Y. (2004). Arabidopsis micro-RNA biogenesis through Dicer-like 1 protein functions. Proc Natl Acad Sci USA 101(34):12753-8.

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In genetics, miRNAs are single-stranded RNA molecules of about 21-23 nucleotides in length, which regulate gene expression. miRNAs are encoded by genes that are transcribed from DNA but not translated into protein (non-coding RNA); instead they are processed from primary transcripts known as pri-miRNAs to short stem-loop structures called pre-miRNAs and finally to function miRNAs. Mature miRNA molecules are partially complementary to one or more messenger RNA (mRNA) molecules, and their main function is to down-regulate gene expression.

1.1.3.1 Formation and processing of miRNA

The genes encoding miRNAs are much longer than the processed mature miRNA molecule; miRNAs are first transcribed as primary transcripts or pri-miRNAs with a cap and poly-A tail and processed to short, 70-nucleotide stem-loop structures known as pre-miRNAs in the cell nucleus. This [processing is performed in animals by a protein complex known as the Microprocessor complex, consisting of the nuclease Drosha and the and the double-stranded RNA binding protein Pasha (Denli et al, 2004).]

In genetics, microRNAs (miRNA) are single-stranded RNA molecules of about 21-23 nucleotides in length regulating gene expression. miRNAs are encoded by genes that are transcribed from DNA but not translated into protein (non-coding RNA); instead they are processed from primary transcripts known as pri-miRNA to short stem-loop structures called pre-miRNA and finally to functional miRNA. Mature miRNA molecules are partially complementary to one or more messenger RNA (mRNA) molecules, and their main function is to downregulate gene expression. [...]

[...]

The genes encoding miRNAs are much longer than the processed mature miRNA molecule; miRNAs are first transcribed as primary transcripts or pri-miRNA with a cap and poly-A tail and processed to short, 70-nucleotide stem-loop structures known as pre-miRNA in the cell nucleus. This processing is performed in animals by a protein complex known as the Microprocessor complex, consisting of the nuclease Drosha and the double-stranded RNA binding protein Pasha.[3]


3. Denli AM, Tops BB, Plasterk RH, Ketting RF, Hannon GJ. (2004). Nature 432(7014):231-5.

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Phospholipase C (PLC) constitutes a large family of mammalian hydrolytic phosphodiesterase enzymes that participate in phosphatidylinositol (PIP2) metabolism and lipid signaling pathways in a calcium dependent manner. Currently, the superfamily consists of six sub-families comprising a total of 13 separate isoforms that differ in their mode of activation, catalytic regulation, cellular localization, membrane binding avidity, and tissue distribution. All are capable of catalyzing the hydrolysis of PIP2 into two important second messenger molecules, which go on to alter cell responses such as proliferation, differentiation, apoptosis, cytoskeleton remodeling, vesicular trafficking, ion channel conductance, endocrine function and neurotransmission (Wu et al., 2000; Rhee, 2001). Phospholipase C (PLC) constitutes a large family of mammalian hydrolytic phosphodiesterase enzymes that participate in phosphatidylinositol (PIP2) metabolism and lipid signaling pathways in a calcium dependent manner. Currently, the superfamily consists of six sub-families comprising a total of 13 separate isoforms that differ in their mode of activation, catalytic regulation, cellular localization, membrane binding avidity, and tissue distribution. All are capable of catalyzing the hydrolysis of PIP2 into two important second messenger molecules, which go on to alter cell responses such as proliferation, differentiation, apoptosis, cytoskeleton remodeling, vesicular trafficking, ion channel conductance, endocrine function, and neurotransmission.
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[However the fact that the cells originated from] cultured kidney cells does not clearly indicate the exact cellular origin of the HEK 293, as embryonic kidney cultures may contain small numbers of almost all cell types of the body. In fact Graham and coworkers more recently provided evidence that HEK 293 cells and several other human cell lines generated by adenovirus transformation of human embryonic kidney cells have many properties of immature neurons, suggesting that the adenovirus was taken up and transformed a neuronal lineage cell in the original kidney culture (Shaw et al, 2002).

1.4.2 Applications of 293 cells

As an experimentally transformed cell line, HEK cells are not a particularly good model for normal cells, cancer cells, or any other kind of cell that is a fundamental object of research. However, they are extremely easy to work with, being straightforward to culture and to transfect, and so can be used in experiments in which the behaviour of the cell itself is not of interest. Typically, these experiments involve transfection in a gene (or combination of genes) of interest, and then analyzing the expressed protein; essentially, the cell is used simply as a test tube with a membrane. The widespread use of this cell line is due to its extreme transfectability by the calcium phosphate method, achieving efficiencies approaching 100% as determined by FACS using a 2 x PBS buffer. A lower efficiency might be achievable with an HBS buffer.

An important variant of this cell line is the 293T cell line that contains, in addition, the SV40 large T antigen, that allows for episomal replication of transfected plasmids containing the SV40 origin of replication. This allows for amplification of transfected plasmids and extended temporal expression of the desired gene products. Note that any similarly domesticated cell line can be used for this fort of work; Hela, COS and Chinese Hamster Ovary cell are common alternatives. Examples of such experiments include: A study of the effects of drug on sodium channels; testing of an inducible RNAi system; testing of an isoform-selective protein kinase C agonist; investigation of the interaction between two proteins; analysis of a nuclear export signal in a [protein (He et al, 1998).]

However the fact that the cells originated from cultured kidney cells does not say much about the exact cellular origin of the HEK 293, as embryonic kidney cultures may contain small numbers of almost all cell types of the body. In fact Graham and coworkers more recently provided evidence that HEK 293 cells and several other human cell lines generated by adenovirus transformation of human embryonic kidney cells have many properties of immature neurons, suggesting that the adenovirus was taken up and transformed a neuronal lineage cell in the original kidney culture (Shaw et al. 2002[3]).

Uses of HEK 293 Cells

As an experimentally transformed cell line, HEK cells are not a particularly good model for normal cells, cancer cells, or any other kind of cell that is a fundamental object of research. However, they are extremely easy to work with, being straightforward to culture and to transfect, and so can be used in experiments in which the behaviour of the cell itself is not of interest. Typically, these experiments involve transfecting in a gene (or combination of genes) of interest, and then analysing the expressed protein; essentially, the cell is used simply as a test tube with a membrane. The widespread use of this cell line is due to its extreme transfectability by the calcium phosphate method, achieving efficiencies approaching 100% as determined by FACS using a 2XPBS buffer. A lower efficiency might be achievable with an HBS buffer.

An important variant of this cell line is the 293T cell line that contains, in addition, the SV40 large T antigen, that allows for episomal replication of transfected plasmids containing the SV40 origin of replication. This allows for amplification of transfected plasmids and extended temporal expression of the desired gene products. Note that any similarly domesticated cell line can be used for this sort of work; HeLa, COS and Chinese Hamster Ovary cell are common alternatives.

Examples of such experiments include:

  • A study of the effects of a drug on sodium channels [4]
  • Testing of an inducible RNA interference system [5]
  • Testing of an isoform-selective protein kinase C agonist [6]
  • Investigation of the interaction between two proteins [7]
  • Analysis of a nuclear export signal in a protein [8]
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Subsequent analysis has shown that the transformation was brought about by an insert consisting of -4.5 kilo bases from the left arm of the viral genome, which became incorporated into human chromosome 19 (Louis et al, 1997).

For many years it was assumed that HEK 293 cells were generated by transformation of either a fibroblastic, endothelial or epithelial cell all of which are abundant in kidney. However the fact that the cells originated from [cultured kidney cells does not clearly indicate the exact cellular origin of the HEK 293, as embryonic kidney cultures may contain small numbers of almost all cell types of the body.]

Subsequent analysis has shown that the transformation was brought about by an insert consisting of ~4.5 kilobases from the left arm of the viral genome, which became incorporated into human chromosome 19 (Louis 1997[2]).

For many years it was assumed that HEK 293 cells were generated by transformation of either a fibroblastic, endothelial or epithelial cell all of which are abundant in kidney. However the fact that the cells originated from cultured kidney cells does not say much about the exact cellular origin of the HEK 293, as embryonic kidney cultures may contain small numbers of almost all cell types of the body.

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1.4 293FT cell lines

Human Embryonic Kidney cells, also known as HEK cells, HEK 293 or just 293 cells, are a cell line originally derived, as their name indicates, from embryonic human kidney. HEK cells are very easy to grow and transfect very readily and so are widely-used in cell biology research. They are also used by biotechnology industry to produce therapeutic proteins and viruses for gene therapy.

1.4.1 Origins of 293 cells

293 cells were generated by transformation of cultures of normal human embryonic kidney cells with sheared adenovirus 5 DNA in the laboratory of Alex Van der Eb in Leiden, Holland in the early 1970s. They are called HEK for human embryonic kidney, while the number 293 roots from numbering of experiments

Human Embryonic Kidney cells, also known as HEK cells, HEK 293 or just 293 cells, are a cell line originally derived, as their name indicates, from embryonic human kidney. HEK cells are not themselves particularly interesting, but are very easy to work with, and so are a widely-used cell line in cell biology research. [...]

[...]

HEK 293 cells were generated by transformation of cultures of normal human embryonic kidney cells with sheared adenovirus 5 DNA in the laboratory of Alex Van der Eb in Leiden, Holland in the early 70s.

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1.2.2.1 Cancer

There are two general abnormalities in cancer cells that exhibit a dysregulation of the cell cycle resulting in uncontrolled growth and are resistant to death as a result of abnormalities in one or more proteins that mediate apoptosis (Nam and Parang, 2003). The goals for RNAi approaches for cancer therapy are therefore to silence the expression of a cell cycle gene and/or an anti-apoptotic gene in the cancer cells thereby stopping tumor growth and killing the cancer cells. To selectively eliminate cancer cells without damaging normal cells, the RNAi needs to be targeted to a gene specifically involved in the growth or survival of the cancer cell, or the siRNAs would be selectively delivered into the cancer cells.


Nam NH, Parang K. Current targets for anticancer drug discovery. Curr Drug Targets. 2003 Feb;4(2):159-179.

A. Cancer

There are two general abnormalities in cancer cells — they exhibit dysregulation of the cell cycle resulting in uncontrolled growth and they are resistant to death as a result of abnormalities in one or more proteins that mediate apoptosis (Nam and Parang, 2003). The goals for RNAi approaches for cancer therapy are therefore to knock out the expression of a cell cycle gene and/or an anti-apoptotic gene in the cancer cells thereby stopping tumor growth and killing the cancer cells. To selectively eliminate cancer cells without damaging normal cells, the RNAi would be targeted to a gene specifically involved in the growth or survival of the cancer cell, or the siRNAs would be selectively delivered into the cancer cells.


Nam NH and Parang K (2003) Current targets for anticancer drug discovery. Curr Drug Targets 4:159–179.

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The ability of RNAi to inhibit the replication or cellular uptake of viruses and other infectious agents has been clearly demonstrated in cell culture studies and, therefore, holds promise for the treatment of human patients. The ability of HIV-1 to infect cells and replicate can be severely compromised by targeting viral genes using siRNAs. Examples include the suppression of HIV-1 replication in human cells transfected with siRNA directed against the tat and the rev gene (Capodici et al, 2002; Jacque et al, 2002; Lee et al, 2002a; Novina et al, 2002). Transfection of human cells with siRNAs directed against different genes in the poliovirus genome resulted in resistance of the cells to infection with poliovirus (Gitlin et al., 2002). The ability of siRNAs targeting the gene encoding the death receptor Fas to protect mice from liver failure and fibrosis in two models of autoimmune hepatitis was tested by Song and colleagues (Song et al., 2003). Intravenous injection of Fas siRNA specifically reduced Fas protein levels in the livers of mice during a 10-day period. Fas siRNA treatment abrogated hepatocyte necrosis and inflammatory infiltration and markedly reduced serum concentrations of transaminases demonstrating a clear hepatoprotective effect of the siRNA therapy. [Seite 643]

The ability of RNAi to inhibit the replication or cellular uptake of viruses and other infectious agents has been clearly demonstrated in cell culture studies and, therefore, holds promise for the treatment of human patients. The ability of HIV-1 to infect cells and replicate can be severely compromised by targeting of

[Seite 644]

viral genes using siRNAs. Examples include the suppression of HIV-1 replication in human cells transfected with siRNA directed against tat and the rev gene (Capodici et al., 2002; Jacque et al., 2002; Lee et al., 2002a; Novina et al., 2002). Transfection of human cells with siRNAs directed against different genes in the poliovirus genome resulted in resistance of the cells to infection with poliovirus (Gitlin et al., 2002). The ability of siRNAs targeting the gene encoding the death receptor Fas to protect mice from liver failure and fibrosis in two models of autoimmune hepatitis was tested by Song and colleagues (Song et al., 2003). Intravenous injection of Fas siRNA specifically reduced Fas protein levels in the livers of mice during a 10-day period. Fas siRNA treatment abrogated hepatocyte necrosis and inflammatory infiltration and markedly reduced serum concentrations of transaminases demonstrating a clear hepatoprotective effect of the siRNA therapy.

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[32.] Ww/Fragment 002 01 - Diskussion
Bearbeitet: 28. October 2014, 02:51 Hindemith
Erstellt: 4. October 2014, 09:12 (SleepyHollow02)
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[Most notably, siRNAs are involved in the RNA interference (RNAi) pathway] where the siRNA interferes with the expression of a specific gene (Tuschl et al, 2001). In addition to their role in the RNAi pathway, siRNAs also act in RNAi-related pathways, as an antiviral mechanism or in shaping the chromatin structure of a genome. siRNAs can also be exogenously (artificially) introduced into cells by various transfection methods to bring about the specific knockdown of a gene of interest. Essentially any gene of which the sequence is known can thus be targeted based on sequence complementarity with tailored siRNA. This has made siRNA an important tool for gene function and drug target validation studies in the post-genomic era. Most notably, siRNA is involved in the RNA interference (RNAi) pathway where the siRNA interferes with the expression of a specific gene. In addition to their role in the RNAi pathway, siRNAs also act in RNAi-related pathways, e.g. as an antiviral mechanism or in shaping the chromatin structure of a genome; the complexity of these pathways is only now being elucidated.

[...]

[...] SiRNAs can also be exogenously (artificially) introduced into cells by various transfection methods to bring about the specific knockdown of a gene of interest. Essentially any gene of which the sequence is known can thus be targeted based on sequence complementarity with an appropriately tailored siRNA. This has made siRNAs an important tool for gene function and drug target validation studies in the post-genomic era.

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[33.] Ww/Fragment 001 13 - Diskussion
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1.1.2 siRNA

Small interfering RNAs (siRNAs), sometimes known as short interfering RNA or silencing RNAs, represent a class of 20-25 nucleotide-long double-stranded RNA molecules that play a variety of roles in biology.

siRNAs have a well defined structure: a short (usually 21-nt) double-strand of RNA (dsRNA) with 2-nt 3' overhangs on either end. Each strand has a 5' phosphate group and a 3' hydroxyl (-OH) group. This structure is the result of processing by Dicer, an enzyme that converts either long dsRNA or hairpin RNAs into siRNAs (Baulcombe et al, 1999).

Most notably, siRNAs are involved in the RNA interference (RNAi) pathway [where the siRNA interferes with the expression of a specific gene (Tuschl et al, 2001).]

Small interfering RNA (siRNA), sometimes known as short interfering RNA or silencing RNA, are a class of 20-25 nucleotide-long double-stranded RNA molecules that play a variety of roles in biology. Most notably, siRNA is involved in the RNA interference (RNAi) pathway where the siRNA interferes with the expression of a specific gene. [...]

[...]

SiRNAs have a well defined structure: a short (usually 21-nt) double-strand of RNA (dsRNA) with 2-nt 3' overhangs on either end:

[...]

Each strand has a 5' phosphate group and a 3' hydroxyl (-OH) group. This structure is the result of processing by dicer, an enzyme that converts either long dsRNAs or small hairpin RNAs into siRNAs.[3]


3. Bernstein E, Caudy A, Hammond S, Hannon G (2001). "Role for a bidentate ribonuclease in the initiation step of RNA interference". Nature 409 (6818): 363–6. PMID 11201747.

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