Controlling Gene Expression with 2-5A Antisense

METHODS: A Companion to Methods in Enzymology 18, 252–265 (1999) Article ID meth.1999.0782, available online at http://www.idealibrary.com on

Controlling Gene Expression with 2-5A Antisense Douglas W. Leaman 1 and Hagen Cramer Gemini Technologies Inc., 11,000 Cedar Avenue, Suite 140, Cleveland, Ohio 44106

Recent work has demonstrated that the activity of a ubiquitous cellular enzyme, ribonuclease L (RNase L), can be harnessed to cleave targeted RNA species. Activation of RNase L is dependent on the presence of 29,59-linked oligoadenylates (2-5A), usually produced by cells infected with viruses. By conjugating synthetic 2-5A to specific antisense compounds, it is now possible to selectively degrade RNAs in an RNase L-dependent manner, thereby providing an alternative to RNase H-dependent approaches. In this summary, we provide an updated description of the synthesis procedure for constructing these chimeric 2-5A antisense molecules. Examples of successful applications of the 2-5A antisense strategy are described, along with some of the procedures involved in those studies. Several methods are also provided for optimizing compound uptake and analyzing their effects on cells. Finally, we discuss the current body of evidence that supports the contention that RNase L is indeed the primary mediator of 2-5A antisense effects and the possible implications that this has on the future of this therapeutic approach. © 1999 Academic Press Key Words: ribonuclease L; telomerase; respiratory syncytial virus.

Many human diseases result from the inappropriate production of harmful proteins encoded by genes of cellular or pathogenic origin. Efforts to identify compounds capable of inhibiting the functions of such proteins through standard medicinal chemistry-based approaches can be very effective. Unfortunately, such approaches are also hampered by the need to screen enormous numbers of compounds and by the speculative nature of the approach. Antisense therapeutics is based on the hypothesis that complementary nucleic acids can anneal by Watson–Crick pairing within cells to “short-circuit” the production of harmful proteins encoded by targeted 1 To whom correspondence should be addressed. Fax: (216) 231– 2931. E-mail: [email protected].

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genes (1–3). By introducing short, synthetic antisense oligonucleotides (ONs) into cells, it is possible to selectively inhibit the translation of specific cellular mRNAs into protein. Unlike compounds isolated serendipitously from combinatorial chemical libraries, these “rational design” pharmaceutical compounds offer the potential to selectively target only those genetic messages that contribute to a particular disease state while leaving other cellular processes largely undisturbed. The mechanisms by which standard antisense ONs inhibit RNA expression or function can vary depending on the region of the RNA molecule to which the compound is directed. ONs that are complementary to sequences at or near the translation start site can inhibit interactions between the mRNA and components of the protein synthesis machinery (e.g., ribosomes). Appropriately targeted antisense ONs may also inhibit transcript splicing, polyadenylation, or transport out of the nucleus (4). More commonly, however, ONs are directed to single-stranded portions of the targeted mRNA where, once bound, they can attract a cellular ribonuclease (RNase), RNase H, which cleaves the RNA portion of the RNA:ON hybrid (3). In doing so, the transcript is degraded, thereby preventing its translation into protein, while the antisense molecule itself is released to hybridize with additional target sequences until it too is degraded by cellular enzymes. Antisense ONs used for therapeutic or experimental purposes were originally composed of oligodeoxynucleotides joined by phosphodiester (PO) linkages. However, the majority of antisense compounds intended for therapeutic use today have more advanced chemistries. The most widely used modification incorporates a thiol group onto the internucleotide bond, creating a phosphorothioate (PS) linkage (5, 6). Compounds with this chemical modification are more resistant to enzymatic degradation, yet still support RNase H-dependent cleavage of targeted mRNA (7). Unfortunately, PS-containing compounds also may exhibit sig1046-2023/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

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nificant non-antisense or aptameric effects that are difficult to predict or control, a major problem that has contributed to delays in getting antisense therapeutics to market (8). To avoid PS-induced problems and enhance half-life and affinity for targeted RNA, a plethora of new generation compounds with novel base compositions have been developed [see (9, 10)]. These are often more stable and less toxic than traditional antisense compounds, but the majority of the modified oligomers are no longer substrates for RNase H, necessitating use of so-called “gapmers.” Gapmers are chimeric antisense compounds in which the molecule is stabilized against nuclease degradation, usually by incorporating advanced nucleic acid modifications at the 59 and 39 termini, while the internal five to eight bases are composed of standard deoxyribonucleotide phosphorothioates (9). Alternatively, the internal bases may be modified, with the termini composed of standard nucleotides with PS linkages (11). Regardless, most of these modifications provide both increased stability and higher affinity for target sequences, and some gapmers can even provide for oral bioavailability. Dr. Paul Torrence, Dr. Robert Silverman, and their colleagues have developed a novel class of chimeric ONs for use in antisense therapeutic strategies (12– 14). These chimeras are composed of an activator moiety, 29,59-linked oligoadenylates (2-5A), attached to a standard antisense cassette. In essence, the antisense portion of the chimera functions to recruit RNase L to the targeted RNA molecule while the 2-5A moiety activates the latent enzyme (Fig. 1). The utilization of RNase L has been shown to be an effective means of degrading targeted RNAs, as described in subsequent sections of this review. Furthermore, this strategy may provide a use for the advanced oligonucleotide chemis-

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tries, mentioned earlier, that are incompatible with RNase H-dependent strategies. 2-5A and The Virus Response Pathway The physiological function of 2-5A in the cellular response to virus is summarized in Fig. 2. Doublestranded RNA (dsRNA) accumulates in cells infected by a variety of different viruses, and cells have evolved mechanisms to sense and respond to this dsRNA. dsRNA induces expression of the interferon (IFN) genes, the protein products of which prime the surrounding cells to repel potential virus infection [see (15)]. dsRNA also activates a family of latent cellular enzymes known as the 2-5A synthetases (16). Once activated by viral dsRNA, the 2-5A synthetases polymerize ATP into 2-5A, the sole function of which appears to be to activate another latent cellular factor, RNase L. RNase L is a ubiquitous 80-kDa endoribonuclease that is activated on binding of 2-5A, thereby initiating the decay of mRNA and rRNA (17). Although RNase L will target RNA indiscriminately, synthesis of 2-5A in the proximity of viral RNA may promote preferential degradation of viral transcripts, inhibiting viral replication. In addition, RNase L itself is transcriptionally upregulated by IFN (17), and so maximal protection is obtained in cells exposed first to IFN. Harnessing RNase L for Antisense Therapeutics The ubiquitous distribution and tight regulatory features of RNase L led to speculation that by coupling a 2-5A activator to an antisense molecule, RNase L could be used to selectively degrade only targeted RNAs (12–14). As described in the following sections, this approach has been put to the test in a variety of controlled experimental situations. Although some issues remain unresolved (see later),

FIG. 1. Schematic representation of 2-5A antisense targeting of mRNA. The 2-5A chimeric molecule is composed of a specific antisense component and a 2-5A activator moiety. The antisense portion targets a complementary mRNA sequence, while the 2-5A component recruits and activates RNase L. Once activated, RNase L cleaves the targeted mRNA leading to its degradation. The 2-5A antisense is then released and can bind to and selectively degrade additional mRNA molecules.

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the overall conclusion at this time is that 2-5A antisense is a viable alternative to standard, RNase H-dependent antisense strategies. The general formula for a “second-generation” 2-5A antisense molecule is shown in Fig. 3. The two features of 2-5A that contribute most significantly to RNase L activation are (1) the number of 29,59-linked adenylates that are oligomerized and (2) the obligatory presence of a 59-terminal monophosphate. Experimental evidence suggests that at least three 29,59-linked oligoadenylates are required to activate RNase L (18). Thus, all of the 2-5A-antisense chimeras currently in use contain either three or four 2-5A moieties, abbreviated pA 3 and pA 4, respectively. The lowercase “p” represents the terminal monophosphate, without which the chimera is less capable of activating RNase L. Indeed, to protect this critical residue, all second-generation 2-5A antisense compounds under evaluation have a protecting 59-phosphothioate group to inhibit phosphatidic cleavage of this critical phosphate, represented as spA 3– 4. While it is clear that the addition of an antisense component to 2-5A reduces its ability to activate RNase L (19), this reduced activity is likely what allows 2-5A antisense chimeras to specifically target only selected RNA molecules, rather than triggering a general endoribonucleolytic activity that degrades all RNA

FIG. 2. Role of 2-5A in the antiviral response. dsRNA accumulates in the cytoplasm of cells infected by a variety of different viruses. This dsRNA can activate the 2-5A synthetases, a family of enzymes whose expression can be induced transcriptionally following exposure of cells to IFN. Once activated, 2-5A synthetases catalyze the formation of oligomers of 2-5A from ATP. The only known function of 2-5A is to activate latent RNase L, resulting in a generalized degradation of RNA within the cell.

within the cell. It is important to recognize that these requirements are valid only for human RNase L (and that of at least several primate species). RNase L from rodent species requires a 59-pyrophosphate for activation, and so modified 2-5A compounds must be used (20).

SYNTHESIS OF 2-5A ANTISENSE COMPOUNDS Second-generation 2-5A antisense chimeras (Fig. 3) are synthesized by means of solid-phase synthesis using phosphoramidite methodology (14, 21, 22). The different moieties of the chimera are synthesized by applying the appropriate phosphoramidites on an automated DNA synthesizer (such as an Expedite 8909, PE Biosystems). By following the synthesis procedure described by Xiao et al. (21) and Li et al. (22), it is possible to synthesize the entire chimera without the need to reinitiate synthesis of each component part (Fig. 4). The following update of their protocol incorporates information provided by members of the Torrence laboratory (to whom we are grateful), along with modifications made in our own laboratory. To introduce the stabilizing 39-cap (39,39 internucleotide linkage at the 39 terminus), chimera synthesis is initiated with a commercially available inverted 59support using a 1 mmol 39-O-dimethoxytritylthymidine59-lcaa-CPG column (Glen Research). The antisense domain is extended in the 39-to-59 direction using DNA synthesis cycles with standard deoxynucleoside amidites (Glen Research) and concentrations recommended by the supplier. For incorporating the linker, a 2-(cyanoethyl)N,N-diisopropyl-4-O-(dimethoxytrityl)butyl phosphoramidite (ChemGenes) is coupled twice to the growing chain (coupling time 60 s). For synthesis of the 2-5A domain, RNA cycles are used with a 59-O-dimethoxytrityl-N6-benzoyl-39O-(tert-butyldimethylsilyl)adenosine 29-(N,Ndiisopropylcyanoethyl) phosphoramidite (ChemGenes) (coupling time 600 s). The 59-phosphorothioate cap is introduced by using the commercially available (2-[2-4,49-dimethoxytrityl)ethylsulfonyl]ethyl(N,N-diisopropylcyanoethyl) phosphoramidite (Glen Research) and the sulfurization reagent 3H-1,2-benzodithiol-3-one 1,1dioxide (Beaucage reagent, Glen Research) in place of iodine for the oxidation step. This sulfurization reaction is slower than the oxidation and therefore requires a longer waiting step (60 s). Deprotection After synthesis, the solid support is removed from the column and transferred into a 3-ml vial with a Teflonlined screw cap. The chimera is cleaved from the support and deprotected in a mixture of concentrated aqueous ammonia and ethanol (2 ml; 3:1, v/v) at 55°C for 8 h. After

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cooling, the ammonia solution is filtered and evaporated to dryness. Alternatively, it is possible to reduce the time of ammonia treatment (to help prevent possible damage to the oligonucleotide) by using phenoxyacetyl carbonyl (PAC) as the base-protecting group (Fig. 5). This fastdeprotecting group allows base deprotection and cleavage from the support to occur within 20 min at 55°C. To remove the tert-butyldimethylsilyl groups from the 39hydroxy of the 2-5A part, the residue from the aforementioned ammonia treatment is dissolved, with sonication, in 1 ml of a 1 M tetrabutylammonium fluoride (TBAF) solution in tetrahydrofuran (Aldrich, Milwaukee, WI) and incubated for 20 h at room temperature. Purification Chimeras are purified by means of HPLC (125P binary pump and a 166P detector module, Beckman). Material resulting from a 1-mmol synthesis is puri-

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fied with a polystyrene reverse-phase (PRP) column using an ion-pair method (21–23). Briefly, the TBAF solution from above is diluted with 2 ml of water and concentrated in a Speed-Vac at low heat to about two-thirds the volume to remove the majority of tetrahydrofuran. The remaining solution is injected onto a PRP-1 preparative column (300 3 25 mm, Hamilton) with a flow rate of 5 ml/min running the following gradient: isocratic at 30% mobile phase B in A for 2 min, then within 3 min to 70% B followed by a linear gradient from 75% to 95% B within 25 min and holding an additional 10 min at 95% B, where A is 10 mM tetrabutylammonium dihydrogen phosphate (TBAP) in water, pH 7.5, and B is 10 mM TBAP in acetonitrile/water (8:2, v/v), pH 7.5. Detection is at 260 nm. Fractions containing the chimera are pooled, evaporated to about 2 ml, and desalted by solid-phase extrac-

FIG. 3. Structure of a second-generation 2-5A antisense chimera. The 59-monophosphate is required for activation of RNase L, and the phosphorothioate modification of this residue inhibits phosphatidic cleavage. The 2-5A component binds to (e.g., recruits) and activates RNase L. The antisense portion binds to complementary RNA sequences within the cell, thereby directing the activated RNase L to cleave the targeted mRNA. This component of the chimera is typically 15–20 nucleotides in length. The 39-inverted nucleotide inhibits the action of 39-exonucleases, thereby providing additional stability.

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tion with a C 18 Sep-Pak cartridge (Waters). The chimera is loaded onto the cartridge, salts are removing by washing the cartridge with H 2O, and the chimera is eluted with MeOH/H 2 O (50:50 to 80:20, v/v). The chimera–TBA salt in 1 ml of water is converted into its sodium form by adding 1 ml of Dowex 50WX8 ionexchange wet resin (sodium form) and mixed for 30 min at 4°C, and the resin is removed using a Poly-Prep chromatography column (Bio-Rad). Finally the chimera is dialyzed against water for 48 h using a Spectra/Por DispoDialyzer with a 3500 MW cutoff cellulose ester membrane (Spectrum). After dialysis, the

chimera is sterilized with a 0.22-mm filter unit, quantitated, and evaporated to dryness. Characterization Key considerations in the characterization of 2-5A antisense chimeras include (1) that none of the 29,59phosphodiester bonds in the 2-5A portion of the chimera are isomerized to 39,59 during or after synthesis, (2) that phosphorylation and subsequent sulfurization occur at the 59 terminus in high yield, (3) that the butanediol phosphate linker connects 2-5A and antisense DNA in the anticipated position, and (4) that the

FIG. 4. Synthesis cycle for the solid phase synthesis of ONs using phosphoramidite chemistry.

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antisense part of the molecule has the right sequence (21). Points (1) and (2) are essential for the activation of RNase L, whereas (4) is needed for the specific recognition of the targeted RNA. 1 H NMR and Maxam–Gilbert chemical sequencing can be used to provide detailed information about the overall configuration of the chimera and the nucleotide

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sequence of the antisense portion, respectively (21). However, both of these procedures are laborious and require substantial amounts of starting material. 2-5A antisense chimeras are routinely characterized by using anion exchange HPLC or capillary gel electropheresis (CGE) to provide information about chimera purity (21, 22). Reversed-phase HPLC analysis following

FIG. 5. Phosphoramidite building blocks used in the synthesis of second-generation 2-5A chimeras.

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snake venom phosphodiesterase (SVPD) digestion is used to confirm that the individual components of the chimeras are present at the anticipated ratios (21, 22). Matrix-assisted laser desorption ionization mass spectrometry can then be used to determine with a high degree of accuracy the molecular weight of 2-5A antisense chimeras (21, 24). Methods Anion-exchange HPLC or CGE can distinguish between 59-phosphothioated, 59-phosphorylated, and 59unphosphorylated chimeras (25). This is helpful for determining whether a given chimera will be competent to activate RNase L (e.g., 59-phosphorylated or not) and whether its 59-phosphate is completely thioated, thereby enhancing its stability against phosphatases. For anion-exchange HPLC analyses, we use a Beckman 126 binary pump with a 168 diode array UV/vis detector. A small sample is injected onto a PA100 (250 3 4 mm) column (Dionex) with a flow rate of 1.5 ml/min running a linear gradient from 1 to 95% mobile phase B [25 mM NaOH, 1 M NaCl in acetonitrile/ water (1:1000, v/v), pH 12.4] in A [25 mM NaOH in acetonitrile/water (1:1000, v/v), pH 12.4] within 45 min while detecting from 200 to 350 nm. Figure 6A shows an HPLC chromatogram for sp59-(A29) 4-Bu 2-d(GGCGAGTCGGCTTATAAA)-39-39-dT-59. The major peak corresponds to the 59-phosphothioated chimera, and the minor peaks represent 59-phophorylated and, to a lesser extent, 59-unphosphorylated products. Digestion of Chimeras with Snake Venom Phosphodiesterase The digestion of a 2-5A antisense chimera with SVPD provides valuable information about the DNA antisense sequence, and can also be used to confirm that the butanediol phosphate linker, 29,59-linked AMP residues, and 59-phosphothioate moiety are each present in the proper amounts (21, 22). Complete digestion of a second-generation chimera yields deoxynucleotide 59phosphates (dNTPs) as well as a pdN-39-p-39-dN dinucleotide that results from the 39-cap structure (the 39,39-internucleotide bond cannot be cleaved under the conditions employed) (21, 22). Peaks corresponding to AMP and pApBupBu can be seen in the HPLC chromatogram, and these result from the 2-5A and linker moieties of the chimera, respectively. The thioated AMP (AMPS) resulting from the 59-thiophosphorylation may or may not show up on the chromatogram. When purified AMPS is digested with SVPD, the majority is hydrolyzed to AMP and, to a lesser extent, to adenosine. Nonetheless, if the chimera is not 59-phosphorylated, the adenosine peak will increase in intensity on the chromatogram, and such chimeras are not likely to activate RNase L. The procedure described

below should allow separation of the peaks corresponding to each of the above components (21, 22). The composition of each peak is confirmed by comparing the retention time with those of authentic samples, e.g., 59-dAMP, 59-dCMP, 59-dGMP, 59-dTMP, 59-AMP, 59-AMPS, adenosine (Sigma), pApBupBu (synthesized on an universal support), and the pdN-39-p-39-dN dinucleotide. Peak areas are determined by integration, and are related to the overall concentration either by dividing this value by the appropriate e value, or by the integral of a standard containing all nucleosides in the same concentration (e.g., 5 3 10 27 M). This provides a molar ratio of nucleotides present in the digest, and should match the expected ratios based on the nucleotide content of the synthesized chimera. Methods Two-tenths OD 260 of the chimera is incubated with 0.15 U of SVPD (from Crotalus adamanteus, Amersham Pharmacia Biotech) in 100 ml of 50 mM Tris-HCl (pH 8.0), 0.5 mM MgCl 2 at 37°C for 3 h. The digestion products are passed through a 10,000-MW-cutoff spin filter (Microcon-10 concentrator, Amicon, Danvers, MA) that is washed twice with 100 ml of H 2O to ensure that all nucleotides have passed through the membrane. The eluate is analyzed on a Beckman HPLC system with 126 binary pump and a 168 diode array UV/vis detector. A TLC Advantage C 18 ODS (60 Å, 5 mm, 4.6 3 250 mm) HPLC column (Thompson) is used with a flow rate of 0.5 ml/min running 1% B isocratically for 20 min, then a linear gradient of 1– 45% B in A for 30 min, followed by a linear gradient from 45 to 95% B in A for 30 min, where A is 100 mM ammonium phosphate (pH 5.5) and B is MeOH/H 2O (1:1, v/v). When a representative 39-capped chimera, sp59-(A29) 4Bu 2 -d(AGTCAGCGAGAAAAACA)-39-39-dT-59, is digested under those conditions, HPLC of digestion products reveals the peaks dCMP with a retention time of 15 min, AMPS (38 min), dTMP (39 min), dGMP (40 min), AMP (41 min), pApBupBu (43 min), dAMP (48 min), and 39-cap (pdA-39p39-dT; 62 min) in the expected ratios (except for AMPS, which is understandably low) (Fig. 6B).

TESTING AND APPLICATIONS OF THE 2-5A ANTISENSE HYPOTHESIS In the following sections, we focus on the analysis of 2-5A antisense compounds by reviewing some of the successful applications of this approach, emphasizing the types of analyses specific to these individual studies and the types of controls used to confirm the specificity of the effects. These experiments provide essential information that has aided in the development of

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new-generation compounds that are currently under evaluation. At the end of each section, a brief summary of the specific methods employed to generate the data is provided. Cell-Free Testing The effectiveness of 2-5A antisense compounds can be tested in cell-free systems using either purified RNase L or whole cell extracts. The methodology for using purified recombinant RNase L to study 2-5A antisense activity in vitro has been described in detail in a recent edition of this journal (26), and is not reiterated here. The use of cell extracts to study 2-5A antisense activity provided an essential first step in determining the specificity of the approach, as described below. A prototypical 2-5A-antisense chimera was constructed by linking pA 4 to 18 39,59-linked deoxythymi-

FIG. 6. HPLC-based analysis of 2-5A antisense chimeras. (A) Dionex PA-100 anion exchange HPLC profile of a second-generation 2-5A chimera. The major peak represents the fully 59phosphothioated chimera, and the minor peaks, 59-phosphorylated and 59-unphosphorylated products. (B) HPLC analysis of the SVPD digestion products from chimera spA 4 -Bu 2 -AGTCAGCGAGAAAAACA-39-39-dT-59, where Bu 2 represents the 2-butanediol phosphate linkers, and 39-dT-59, the stabilizing 39-inverted residue. The identities of the major peaks are indicated.

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dines (dT), yielding a compound that was abbreviated pA 4:dT 18. Like unmodified 2-5A, pA 4:dT 18 was able to bind and activate RNase L in vitro, albeit at a reduced level as compared with to the free 2-5A (see earlier). Furthermore, it was able to form a stable complex with a complementary poly(rA) RNA target at an affinity comparable to that of unmodified dT 20 (12). A synthetic target RNA was generated by interrupting a partial cDNA for the HIV vif protein with 25 dA residues (TAR:A25:vif). A control TAR:vif RNA lacking the internal poly(A) tract was also made. Both RNA species lacked 39-poly(A) tails, and were radiolabeled at their 39 termini with 32P for detection of specific cleavage products. The TAR:A25:vif and TAR:vif RNA species were added to Daudi lymphoblastoid cell extracts in the presence or absence of the pA 4:dT 18 test compound. In the presence of pA 4:dT 18, the TAR:A25:vif RNA, but not the TAR:vif RNA, was rapidly and selectively converted into a specific cleavage product (12). Three important controls were run to confirm that the cleavage was RNase L dependent and not due to RNase H activity present in the cell lysates. First, addition of excess dT 20 to the assay competed for binding of pA 4:dT 18 to the target and prevented degradation. Since this same competitor would have enhanced, rather than prevented RNase H-dependent degradation, the observed degradation could not have been due to contaminating RNase H. Second, a 59-unphosphorylated compound (A 4 :dT 18 ) that lacked the 59phosphate required for RNase L activation did not effect RNA cleavage. Finally, addition to the reaction mixture of ppp59I29p59A29p59A, a specific inhibitor of RNase L (but not RNase H), prevented pA 4:dT 18 from effecting the cleavage of the TAR:A25:vif RNA (12). These data provided clear in vitro evidence that 2-5Alinked antisense oligonucleotides could effectively recruit and activate RNase L to selectively degrade targeted RNA species. Methods Specific to These Studies (12) Target RNA species are transcribed in vitro (Promega kit) and are 39-end labeled with cytidine 39,59-[5932 P]bisphosphate using T4 RNA ligase (Pharmacia). The labeled products are purified from denaturing urea polyacrylamide gels and the specific activity is determined as described (27). Cell extracts are prepared by lysing cells in 40 mM KCl, 10 mM Hepes, pH 7.5, 2.5 mM magnesium acetate, 0.5 mM ATP, 2.5% (v/v) glycerol, 2 mM 2-mercaptoethanol. After incubation on ice, lysates are centrifuged at 100,000g, and the cleared supernatant is used directly in cleavage assays. Approximately 100 nmol of the radiolabeled target RNA is incubated with 2-5A ONs and 10 ml of the cell extract that has been adjusted to 75 mM KCl in a final volume of 20 ml. After 30 min at 30°C, the RNA is isolated by extraction with phenol/chloroform, precipitated with

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ethanol, and analyzed on an 8 M urea, 6% polyacrylamide gel, followed by autoradiography (12). Cell-Based Confirmation of the 2-5A Antisense Hypothesis As with standard antisense compounds, 2-5A antisense oligomers must pass a number of rigid tests before the observed physiological responses can be attributed to true antisense effects. These criteria include demonstration of (1) sequence and dose dependence of the effects, (2) direct cleavage of the targeted RNA species, (3) lack of effects on unrelated RNA species, and (4) lack of generalized cytotoxicity within the therapeutic dose range (28). In addition to these important standards, 2-5A antisense compounds must also be shown to act in an RNase L-dependent manner. Therefore, it is critical to synthesize non-2-5A antisense controls with wild-type (e.g., fully complementary) antisense cassettes and compounds with defective 2-5A moieties that either lack the 59-terminal phosphate residue or possess only two 2-5A residues, either of which is a very poor activator of RNase L. The first cell-based tests of the 2-5A antisense hypothesis were aimed at ablating the mRNA for the cellular serine/threonine kinase PKR (29). Those studies have been described in detail in several recent reviews (13, 14, 30), and are not covered again here. However, the 2-5A antiPKR results were fundamental in that the 2-5A anti-PKR lead and control compounds satisfied each of the above criteria. In HeLa cells, a fully functional 2-5A anti-PKR compound possessing a 29,59-tetraadenylate was able to ablate PKR mRNA and protein expression. In contrast, compounds with a defective (or no) 2-5A moiety and compounds with scrambled antisense components had little or no influence on PKR message or protein expression (29). When studies were conducted to assess the physiological consequence of ablating PKR from cells, it was found that dsRNA signaling was defective in cells treated with 2-5A anti-PKR, but that tumor necrosis factor a signaling, which does not involve PKR, was unaffected (29). Thus, these early results provided the first cellbased evidence that the 2-5A antisense hypothesis was sound. An important footnote to these studies is that subsequent work in PKR-knockout mice has revealed that dsRNA-dependent signaling is also defective in PKR–minus cells (32), thereby confirming the physiological significance of the 2-5A anti-PKR findings. In each of the following experimental situations, similar control compounds have been employed to verify the specificity of the observed effects, confirming both that an antisense mechanism is involved and that RNase L activation is critical for the response.

Telomerase Among the compounds currently under development is a 2-5A anticancer therapeutic targeting telomerase. Telomeres are TG-rich structures found at chromosomal ends that provide a protective cap to prevent loss of genetic information that would eventually occur as the chromosomes became shorter during cellular divisions (33). Telomere integrity is maintained by the ribonucleoprotein complex telomerase. The telomerase holoenzyme comprises a catalytic protein component, called TERT (telomerase reverse transcriptase) (34) and an RNA component (hTR) that provides the template for synthesizing nascent telomeric sequences (35). In cells that have a finite life span, including most somatic cell types, telomerase activity is undetectable, suggesting that normal cellular senescence results from the gradual shortening of telomeric ends. In contrast to normal cells, the majority (,80%) of human tumor cells have high levels of telomerase expression, and telomerase is thought to be essential for cellular immortality and oncogenesis [see (36, 37)]. The tumorrestricted pattern of telomerase expression makes it a strong candidate for targeted therapy. Indeed, numerous previous studies have shown that selective degradation of hTR in human tumor cell lines (38) or inhibition of its activity by using antisense compounds complementary to the template portion of the molecule (35, 39) prevents the growth of cancer cells in culture. Thus, it was of interest to determine whether an antihTR 2-5A antisense approach could be used to inhibit telomerase activity and thus prevent tumor cell growth. A test compound, spA 4-anti-hTR, was developed in Dr. John Cowell’s laboratory at the Cleveland Clinic Foundation’s Lerner Research Institute. spA 4 -antihTR was targeted to a putative stem–loop region of the hTR transcript (bases 76 –94). Two control ONs were also synthesized, spA 2-anti-hTR containing only two 2-5A moieties and spA 4-anti-(M6)hTR that contained six mismatches. The original studies were conducted with telomerase-positive malignant glioma cell lines (40). Reverse transcription polymerase chain reaction (RT-PCR) analysis demonstrated a selective degradation of the hTR transcript in glioma cells treated for 5 h with spA 4-anti-hTR. In contrast, the control ONs had little influence on hTR levels, and none of the treatments affected cellular glyceraldehyde-3-phosphate dehydrogenase (GAPDH) mRNA levels (40). Longerterm cell growth experiments demonstrated that treatment with spA 4-anti-hTR (two treatments per day, 14 days) resulted in 70 – 80% inhibition of glioma cell viability (40). Little or no inhibition was observed with the control ONs. Importantly, spA 4-anti-hTR did not affect the growth of telomerase-minus human cells such as P1N astrocytes, thereby confirming that its

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effects were specific to cells with detectable telomerase activity. In vivo tests of the antitumor effectiveness of spA 4-anti-hTR have been performed with human glioma xenografts in nude mice. spA 4-anti-hTR injection into glioma tumors resulted in a significant reduction in tumor volume as compared with injection of vehicle, the mismatch control spA 4-anti-(M6)hTR, or the 2-5Adefective control spA 2-anti-hTR (40). Together, these data suggest that spA 4-anti-hTR is a potent therapeutic compound that functions in vitro and in vivo to inhibit the growth of telomerase-positive tumor cells. Since cells treated with spA 4-anti-hTR begin to die within several days, the mechanism of death may not be due exclusively to the serial shortening of chromosomal ends with each cell division. Glioma cells treated with spA 4-anti-hTR take on the appearance of cells undergoing apoptosis (nuclear blebbing, etc.), and the TUNEL assay (Apo-Tag, Oncor Inc.), which detects broken ends of DNA, demonstrates quite clearly that U251-MG cells treated with spA 4 -anti-hTR have a higher incidence of apoptotic cells than control cultures (40). Similar results were observed when tumors treated with spA 4-anti-hTR were observed in situ (40). Although not yet proven, the rapid induction of apoptosis in spA 4-anti-hTR-treated cells (or tumors) suggests that a sensing mechanism may be used by human cells, whereby the loss of telomeric sequences from even a fraction of the cell’s chromosomes activates an apoptotic pathway. Methods Specific to These Studies RNA analysis is performed by using standard RTPCR procedures (see later) with primer sets described in Kondo et al. (40). Cell growth studies are performed by adding 2-5A anti-hTR compounds directly to the cells twice a day (12-h intervals) until the cells in control wells reach confluence. At that time, cells are dissociated in 50 ml trypsin EDTA (0.05% trypsin, 0.53 mM EDTA; Gibco BRL), 50 ml of complete medium is added, and 25 ml of the cell slurry from each well is transferred to a new well containing 75 ml of complete medium to continue the assay. From the remaining cells, 25 ml is added to an equal volume of trypan blue solution (0.4% w/v, Gibco BRL), and 10 ml of the mixture is counted on a hemacytometer to determine the number of viable cells in each well. The TRAP assay, which detects telomerase activity, is performed using the remaining 50% of the trypsinized cells. The TRAPeze kit (Oncor Inc., No. S7700-KIT) is used for these studies according to the manufacturer’s directions. The experiment continues for 10 –14 days or until one or more of the treatments has resulted in a dramatic (;90%) reduction in cell viability.

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Respiratory Syncytial Virus Respiratory syncytial virus (RSV) is a nonsegmented, negative-strand RNA virus that is a major cause of lower respiratory tract disease in infants, young children, and the elderly, particularly those who are institutionalized. It is the most common cause of viral bronchiolitis and pneumonia in children, and outbreaks in the United States frequently reach epidemic proportions during the winter months, accounting for roughly 90,000 hospitalizations and 4500 deaths per year (41). The only approved anti-RSV compound, ribavirin, has been shown to affect virus shedding, but it has little influence on patient mortality or duration of hospitalization (42). Antibody treatments RespiGam and Synagis (Med Immune) have been approved for prophylactic use in high-risk infants. The experimental utilization of 2-5A antisense ONs to inhibit RSV replication has involved two substantially different approaches. The first approach targeted several RSV transcripts for degradation with 2-5A antisense ONs. The most dramatic antiviral effects were observed with compounds directed against the RSV M2 mRNA, which encodes a transcription elongation factor essential for viral replication (43). Inhibition of RSV replication by the 2-5A anti-M2 compound was both sequence specific and 2-5A dependent (43). Recently, a more effective second-generation anti-RSV 2-5A compound has been developed that targets the viral RNA genome directly (44). The 2-5A antigenome chimera, NIH351, contains core sequences complementary to a nine-nucleotide sequence found in the intergenic regions between most RSV genes. Thus, this single antisense compound is potentially able to target multiple sequences along the RSV genome, thereby increasing the likelihood of an effective interaction with the genome. The evolution of this new anti-RSV compound followed a logical progression that provides a good example of how highly specific 2-5A antisense compounds can be developed. The original anti-RVS compound (NIH273) was an all-PS, non-2-5A ON that had significant antiviral activity. However, when the sequence of this compound was scrambled, excepting the four contiguous G residues (G-quartet), the new compound (NIH317) had slightly better antiviral activity than the wild-type NIH273. When the G-quartet was disrupted (NIH318), antiviral activity dropped to insignificant levels. Thus, it was clear that the effects of the original compound were largely nonspecific and were attributable to the PS G-quartet. When the G-quartet and flanking PS linkages were changed to PO, the activity of the resulting compound (NIH320) was negligible. However, by adding a 2-5A moiety to this compound the activity of the PS/PO 2-5A chimera NIH351 was increased by more than 100-fold. To ensure that these

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effects were sequence specific, the antisense portion of the chimera was scrambled (NIH426), resulting in a ;30-fold drop in activity. A compound lacking the 59phosphate also had little antiviral activity, confirming that RNase L activation was important for the antiviral effects (44). NIH351 was subsequently found to be effective when administered only once immediately before addition of virus (44), and studies in cultured cells have shown NIH351 to be 80- to 125-fold more potent than ribavirin (44). Importantly, NIH351 is equally effective against both A and B strains of RSV (44). Thus, these results suggest that NIH351 is an important new antiviral agent with a broad spectrum of anti-RSV activity that is both sequence and RNase L dependent. Methods Specific to These Studies The effects of 2-5A anti-RSV compounds are assessed by using a cytopathic effects (CPE) reduction assay, as described previously (45). The Player et al. (44) article described two methods to assess the effectiveness of 2-5A antisense compounds. The first was a grading system in which infected cultures were scored with a value from 1 to 4, with 1 representing cultures exhibiting no cytopathicity and 4 representing a culture in which 100% of the monolayer shows viral cytopathic effects (giant cells, syncytia, etc.). The EC 50 (effective concentration of ON resulting in a 50% decrease in virus replication) was then calculated by regression analysis using the means of the CPE ratings at each concentration of the test compounds. The second method was a neutral red assay (46), using a modification described in Player et al. (44), that provided critical quantitative data to support the visual analyses. It is also possible to obtain quantitative data by fixing the monolayers with methanol, staining them in 0.1% (w/v) crystal violet in PBS, and counting the plaques/syncytia under a microscope, or by performing an ELISA with an antibody against the RSV F-protein (Chemicon International).

GENERAL TECHNIQUES FOR OPTIMIZING AND ANALYZING 2-5A ANTISENSE COMPOUNDS Selection of Target Sequences A variety of methods have been used to try to predict, in advance, what constitutes an optimal antisense target sequence. RNA folding algorithms, such as the MFOLD program, can be used to predict the locations of putative open structures, such as stem–loop and hairpin structures. Since antisense compounds interact more efficiently with complementary sequences that are single stranded, such structures have fre-

quently been used to select target sites. However, it is unclear precisely how useful this information is when trying to identify a compound with optimal activity (47). For this reason, it is still accepted that the best approach is to make multiple antisense compounds targeting a variety of sequences along the transcript to be cleaved and to analyze each empirically to determine which is the most active. Compound Uptake in Cells Although 2-5A antisense compounds can be used without uptake enhancers (29, 43, 44), it is possible to improve their uptake into cells by adding commercially available cationic lipid transfection reagents. We have used fluorescence-tagged 2-5A compounds to optimize conditions for compound uptake and to assess the subcellular distribution of 2-5A and non-2-5A antisense compounds. Synthesis of fluorescein-tagged ONs is performed following the protocol described earlier except that a fluorescein-conjugated solid support (Glen Research) is used, and the synthesis and purification are carried out in a low-light environment to avoid blanching of the fluorescence. Oligonucleotide uptake can be examined by fluorescence-activated cell scanning (FACS) which depicts the percentage of cells that have incorporated fluorescence and also quantifies the relative fluorescence intensities within the total population(s). Treatment is performed by mixing varying amounts of transfection-enhancing reagents with serum-free culture medium and adding this to cells that have been washed once in serum free medium. Fluoresceinconjugated ONs (0.5 mM stocks in water) are then added to a final concentration of 100 –500 nM, and the cells cultured for 3 to 24 h, the same times used for short-term RNA degradation studies (see later). After treatment, cells are washed in ice-cold phosphatebuffered saline (PBS) and then fixed in 0.5% paraformaldehyde (w/v in PBS) for 30 min prior to scanning [a modification of the procedure described in Ref. (48)]. We FACScan ON-transfected cells on a Becton– Dickinson FACS machine using the CELLQuest software package (Becton–Dickinson). Results typical of a 16-h treatment period are shown in Fig. 7. By comparing the mean fluorescence intensity values, it is clear that the addition of lipofectamine can increase uptake by 10- to 30-fold as compared with unassisted uptake. Fluorescent microscopic analysis of these same cells confirmed that the fluorescence profiles represent compound that has entered the cytoplasm (data not shown). We have also found that, in general, subcellular distribution of transfected 2-5A compounds is typical of other antisense compounds (data not shown). Whether the enhanced uptake observed with transfection enhancers also improves compound activity is still under investigation.

2-5A ANTISENSE THERAPEUTICS

Treating Cells with 2-5A Antisense Compounds By combining the data obtained from fluorescent oligo uptake and RNA degradation studies, we have standardized the procedure for treating cultured cells with 2-5A antisense compounds. It is important, however, to note that cell lines can vary significantly in their abilities to take up ONs, and the nucleotide sequence and chemical makeup of an antisense compound can have subtle to dramatic effects on its ability to cross the cellular membrane. Thus, these conditions should be used as a starting point from which optimization is carried out in a cell line- and oligonucleotidespecific manner. As described throughout this report, direct analysis of the targeted and nontargeted control RNAs is the best way to assess the effectiveness of lead and control 2-5A antisense compounds. These analyses are then followed up with tests of the physiological response, such as examining protein function, cell viability, and viral replication. Since these later analyses require varied methods, we focus only on RNA-based studies. Treatment Protocol Cells are plated at 60 – 80% density and allowed to attach overnight. Six-well dishes are typically used for RNA studies, and each well will yield between 20 and 80 mg of total cellular RNA. The day following plating, monolayers are washed once in serum-free medium and then overlaid with serum-free medium containing between 2 and 10 mg/ml lipofectamine (Gibco BRL). 2-5A test and control ONs are added directly to the wells to give the desired final concentration (usually

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100 –1000 nM for RNA ablation studies). This treatment is repeated after 12 h to ensure that sufficient compound is in the cells to effect complete RNA cleavage. RNA Analysis After 24 h of treatment, cells are washed in ice-cold PBS and RNA is isolated by using Trizol reagent according to the manufacturer’s specifications. RNA that is isolated by this method can be used in RT PCR (Gibco-BRL) studies directly. We perform RT-PCR experiments by reverse transcribing RNA into cDNA by using either random hexamer ONs (for cellular mRNAs) or viral genome-specific ONs (for RSV studies) as primers in a 20-ml RT reaction. Briefly, 2 mg of total cellular RNA and 500 ng of primer (in 7 ml total volume) are denatured at 70°C for 5 min, quick-chilled on ice, and then added to 13 ml of a RT mix [4 ml 5X RT buffer, 4 ml dNTP mix (10 mM), 1 ml RNase inhibitor, 1 ml MMTV reverse transcriptase, 2 ml water). The RT reaction is carried out at 42°C for 45 min, and the reaction stopped by heating to 65°C for 10 min. Generally, 2 ml of the resulting cDNA solution is subjected to PCR amplification using specific primer sets and reaction conditions that have been optimized for the target message to be analyzed (hTR, GAPDH, PKR, etc.). PCR products can be visualized by agarose gel electrophoresis and ethidium bromide staining or by transfer to nylon membrane and hybridization with specific 32P-labeled probes.

SUMMARY

FIG. 7. Fluorescent 2-5A ON uptake in cultured human cells. Cells were left untreated (thin line) or were incubated with a fluoresceintagged 2-5A ON (500 nM) for 16 h in the absence (dashed line) or presence of transfection-enhancing reagents (lipofectin, heavy solid line; lipofectamine, dotted line). After treatment, cells were washed extensively, fixed, and subjected to FACS analysis. The horizontal axis represents relative fluorescence, and the vertical axis, cell numbers.

The 2-5A antisense approach is an effective means of selectively degrading cellular or viral RNAs. However, an issue that remains unresolved is whether the RNA ablation observed with each of these compounds is due exclusively to RNase L-dependent cleavage, or whether RNase H also contributes to the effects. Most of the convincing mechanistic studies have been performed in cell-free systems, some of which were described earlier [reviewed in detail in Ref. (30)]. In particular, the RNase L inhibitor ppp59I29p59A29p59A completely inhibits transcript cleavage when included in cell-free studies, and competitor ONs that block RNase L but not RNase H activation have been useful in dissecting the relative contribution of these two ubiquitous endoribonucleases. Although definitive cell-based elucidation of the above issues is forthcoming, much of the current evidence in favor of an RNase L-dependent mechanism is somewhat circumstantial. As mentioned, the complete removal of the 2-5A moiety decreases activity by one to three orders of magnitude, suggesting that any RNase H contribution is minor at

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best. Removing the obligatory 59-phosphate residue from 2-5A chimeras, which incapacitates the RNase L activation, also decreases or eliminates activity. It is hard to explain either of these observations if RNase L is not the primary mediator of 2-5A antisense effects. Nonetheless, conclusive demonstration of a requirement for RNase L in the effects of 2-5A antisense effects will require the use of antisense cassettes that are unable to activate RNase H. These studies must be done carefully, since not all oligonucleotide modifications are compatible with RNase L activation. All-PS antisense cassettes, for example, are incompatible with RNase L activation in vitro (P. Torrence and R. Silverman, 1998, personal communication). However, limited numbers of sulfur substitutions can be tolerated without loss of RNase L activation (such as in the anti-RSV compound NIH351). Other substitutions to the antisense cassette, including 29-O-methyl base modifications, are currently under evaluation. If these can be used in the context of a 2-5A attachment to activate RNase L, it would improve on the stability of current-generation 2-5A chimeras. More importantly, this breakthrough would expand the utility of chemical base modifications that are otherwise incompatible with RNase H activation. It is this feature of the 2-5A approach that is perhaps most attractive.

10. 11.

12.

13.

14.

15. 16. 17.

18.

19.

20.

ACKNOWLEDGMENTS We thank Dr. Paul Torrence and Dr. Robert Silverman for helpful discussions and critical reading of this manuscript. We also thank Frank Longano and James Okicki for excellent technical assistance and members of Dr. Silverman’s and Dr. Torrence’s laboratories for providing updated, unpublished information to keep these methods current.

21.

22. 23. 24. 25.

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