Skip Navigation

This Article
Right arrow Abstract Freely available
Right arrow Print PDF (252K) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (5)
Right arrowRequest Permissions
Right arrow Commercial Re-use Guidelines
for Open Access NAR Content
Google Scholar
Right arrow Articles by Sheehan, D.
Right arrow Articles by Dellinger, D. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sheehan, D.
Right arrow Articles by Dellinger, D. J.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?

Nucleic Acids Research, 2003, Vol. 31, No. 14 4109-4118
© 2003 Oxford University Press

Biochemical properties of phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides

David Sheehan, Benjamin Lunstad, Christina M. Yamada, Brian G. Stell, Marvin H. Caruthers* and Douglas J. Dellinger1

Department of Chemistry and Biochemistry, University of Colorado, Boulder, CO 80309, USA and 1 Agilent Laboratories, 4380 Ziegler Road, Fort Collins, CO 80525, USA

*To whom correspondence should be addressed. Tel: +1 303 492 6095; Fax: +1 303 492 6194; Email: marvin.caruthers{at}colorado.edu

Received as resubmission March 17, 2003; Revised April 14, 2003; Accepted May 2, 2003


   ABSTRACT
TOP
 ABSTRACT
INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Phosphorus-modified phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides were chemically synthesized and their biochemical properties evaluated. Under physiological pH, these DNA analogs possess negative charge and form stable, complementary A-like DNA:RNA heteroduplexes when analyzed via circular dichroism spectroscopy. Phosphonoacetate and thiophosphonoacetate oligomers were found to stimulate RNase H activity and to be completely resistant to degradation by snake venom phosphodiesterase, DNase I and HeLa cell nuclear extract. Further research has demonstrated that neutral, esterified forms of these analogs can be taken up by cells. Phosphonoacetate and thiophosphonoacetate oligomers therefore represent a new class of oligodeoxyribonucleotide analogs having phosphorus– carbon bonds with considerable potential for use in biological research.


   INTRODUCTION
TOP
 ABSTRACT
 INTRODUCTION
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 
Over the years, oligodeoxyribonucleotides (ODNs) and their analogs have become indispensable research tools for studying various biological processes, diagnosing diseases and identifying single nucleotide polymorphisms. More recently, modified ODNs have been used to modulate and control gene expression, which has broad implications from basic research to the use of these oligomers as therapeutic agents (1,2).

Most ODNs useful for modulating gene expression function by specifically inhibiting the production of a targeted gene product. Generally this process involves forming a duplex of mRNA with a complementary oligomer (hence antisense ODNs) followed by inhibition of mRNA expression or its degradation. Because natural DNA is inherently sensitive to cellular nucleases, a useful antisense ODN usually is modified so as to render it resistant to enzymatic degradation. In addition, an antisense ODN most generally activates RNase H which selectively degrades the mRNA strand of an ODN– mRNA duplex and thus eliminates expression of the targeted gene product (3). There are, however, certain nuclease-resistant antisense ODNs that effectively block gene expression by forming a stable hybrid with mRNA, which leads to inhibition of expression (4).

Among the many proposed antisense ODNs, the phosphorothioate and methylphosphonate derivatives have been the most widely studied (1,4). These analogs have either a sulfur atom or a methyl group in place of one non-bridging oxygen at each internucleotide linkage. Both are P-chiral but differ as the phosphorothioate derivative is isoelectronic with natural DNA whereas the methylphosphonate is electronically neutral. Relative to their use as antisense compounds, both have a combination of useful and undesirable features. For example, while phosphorothioate ODNs effectively stimulate RNase H activity and have increased resistance to cellular nucleases (when compared with natural DNA), they are still degraded (1,5,6). Additionally, phosphorothioate ODNs have relatively high affinity for certain proteins, which obscures the antisense mode of action and can lead to cellular toxicity (7). In contrast, methylphosphonate ODNs are stable towards nucleases but, when hybridized to RNA, they do not direct RNase H activity and therefore cannot inhibit gene expression by an RNase H-mediated mechanism (8). Loss of RNase H activity has been attributed to the lack of charge at the internucleotide linkage. Other analogs that possess RNase H activity and nuclease resistance also suffer from various limitations. These include phosphorodithioate ODNs (6,9), cyclohexene nucleic acids (10) and 2'-fluoro-2'-deoxyarabinofuranosyl nucleic acids (11).

One of the requirements for RNase H activity is that the ODN be charged, which creates a problem since polyanionic oligomers do not readily diffuse across cell membranes. In order to overcome this problem for cultured cells, cationic lipids when combined with anionic ODNs are generally used to assist uptake. Unfortunately, this complex is generally toxic to cells, which means that both the exposure time and concentration of cationic lipid must be carefully controlled to insure transfection of viable cells.

Because none of the currently popular analogs satisfies all the criteria expected of an antisense ODN, we have continued searching for a superior derivative. These studies have led us to the phosphonoacetate- and thiophosphonoacetate-modified ODNs (Fig. 1) (12). These analogs, which have an acetic acid moiety in place of oxygen at a non-bridging position of an internucleotide linkage, were expected to possess many of the desirable antisense properties observed for phosphorothioate and methylphosphonate ODNs. Here we report our initial observations regarding their biophysical and biochemical properties. We have found that these analogs as heteroduplexes with RNA form A-type helices which have reduced melting temperatures (compared with natural heteroduplexes) and that the carboxylate moiety possesses a pKa of approximately 3.8 (phosphonoacetate) and 3.9 (thiophosphonoacetate). They were found to be nuclease resistant, capable of stimulating RNase H activity and, as neutral esters, to be taken up by cells via passive diffusion. The latter observation may lead to a new prodrug approach for uptake of RNase H-active ODNs by cells in culture.



View larger version (11K):
[in this window]
[in a new window]
  		Figure 1. Chemical structure of phosphonoacetate (X = O) and thiophosphonoacetate (X = S) ODNs. B represents adenine, guanine, cytosine or thymine.

 

   MATERIALS AND METHODS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 REFERENCES
 
Phosphonoacetate and thiophosphonoacetate ODNs with the structures shown in Figure 1 were synthesized and characterized as described (12). ODNs with various combinations of phosphorothioate and phosphonoacetate/thiophosphonoacetate internucleotide linkages were synthesized via the cycle shown in Figure 2. Appropriate synthons (standard deoxynucleoside 3'-phosphoramidites and deoxynucleoside 3'-O-diisopropylaminophosphinoacetic acid dimethyl-ß- cyanoethyl esters) were introduced according to the synthetic plan in order to generate chimeric ODNs. Deoxynucleoside 3'-O-diisopropylaminophosphinoacetic acid dimethyl-ß- cyanoethyl esters were prepared as described (12) or obtained from MetaSense Technologies (Des Moines, IA). Control ODNs (DNA, phosphorothioate DNA) were synthesized by solid phase phosphoramidite chemistry on controlled pore glass (CPG) (13) using an automated synthesizer (Applied Biosystems Model 394, Foster City, CA). Oligo ribonucleotides and 2'-O-methyl oligoribonucleotides were obtained from Dharmacon Research Inc., Louisville, CO. 5'-32P-labeled oligomers were prepared by standard procedures (14) followed by purification using gel electrophoresis and ethanol precipitation.



View larger version (21K):
[in this window]
[in a new window]
  		Figure 2. Synthesis cycle for phosphonoacetate and thiophosphonoacetate ODNs. The open circle represents controlled pore glass support; B is an appropriately protected purine or pyrimidine base (12).

 
Chimeric ODNs having phosphorothioate ß-cyanoethyl esters and thiophosphonoacetate methyl and butyl esters were synthesized on 3'-phosphate-CPG (Glen Research) using procedures as published previously (12). 5'-Fluorescein phosphoramidite (Glen Research) and standard sulfurization (13) were used to generate a 5'-fluorescein label joined to these ODNs through a nuclease-stable phosphorothioate linkage. To avoid hydrolysis of the acetate esters, it was necessary to remove the ß-cyanoethyl group and cleave DNA from the support under non-nucleophilic conditions. Post-synthesis, the CPG for each ODN synthesis was transferred to an oven-dried 2.0 ml screw-cap vial. Diazabicyclo[5.4.0] undec-7-ene (DBU, 1.5%) in anhydrous acetonitrile was then added to selectively remove the ß-cyanoethyl group from phosphorothioate linkages and cleave the ODN from the support. After 30 min, the DBU was quenched with excess anhydrous acetic acid (relative to DBU) and solvents were removed under reduced pressure. The sample was then diluted with triethylammonium acetate buffer (50 mM, pH 8.0) and filtered through a 0.45 µm microcentrifuge tube (Millipore). The buffer was then removed under reduced pressure, and the ODN was dissolved in 0.5 ml of water and purified by reverse phase HPLC utilizing the 5'-fluorescein (FITC) group for hydrophobic affinity. Preparative HPLC was carried out on a 25 cm Hypersil BDS-C18pn column with a 9.4 mm ID. Gradient eluents were (A) 50 mM triethylammonium bicarbonate (pH 8.0) and (B) acetonitrile. A gradient of 8–90% B over 50 min at a flow rate of 1.2 ml/min was used to elute the ODN. Product fractions were pooled, concentrated under reduced pressure, and diluted in a buffer of 10 mM Tris–HCl pH 8.0 and 1 mM EDTA (TE buffer).

Matrix-assisted laser desorption ionization time of flight (MALDI-TOF) mass spectrometry was performed on a PerSeptive Biosystems Voyager Biospectrometry Workstation (Framingham, MA). ODNs were concentrated to dryness and dissolved in isopropanol/water (1:1) to a final concentration of 200 µM. Samples were prepared as described in the Sequazyme Oligonucleotide Sequencing Kit Manual (PerSeptive Biosystems) with the following modifications: 1 µl of ODN, 1 µl of 25mer DNA standard, 1 µl of ammonium citrate buffer and 7 µl of matrix were combined on a layer of parafilm coated with ammonium cation exchange beads. After pipetting the solution over the beads for 60 s, 5 µl was transferred to a gold-plated 100-well sample plate. The MALDI-TOF measurements were observed in the positive-ion mode.

Ionization of phosphonoacetate and thiophosphonoacetate linkages was examined by monitoring HPLC retention time as a function of HPLC buffer pH for thymidine–thymidylate dimers. Analytical HPLC utilized a 1 ml Resource RPC Reverse-Phase Column (6.4 x 30 mm). Gradient eluents were (A) 100 mM sodium formate adjusted to the appropriate pH with formic acid and (B) acetonitrile. The following gradient was used to evaluate dimer retention time: 2–22% B in 30 min at a flow rate of 1.0 ml/min followed by a 10 min wash with 100% B. Dimer retention time was monitored using HPLC buffers at pH values between 1 and 8 (one unit increments). To achieve a buffer pH of 1, HCl was added to aqueous formic acid. A 20 µl aliquot of a 100 µM dimer solution (in water) was used per sample injection. Dimer retention time in minutes at a given pH was determined from the average of three separate chromatograms. Approximate pKa values were obtained from the midpoint of the titration curve for plots of retention time versus HPLC buffer pH. A dimer containing a natural phosphodiester linkage was used as a control.

Melting points (Tms) were determined on a Varian Cary 1E UV-Visible Spectrometer. The absorbance at 260 nm was measured while the temperature of the sample was increased at a rate of 1.0°C/min. Phosphonoacetate, thiophosphonoacetate and phosphorothioate oligonucleotides were mixed separately with complementary RNA or DNA in a 1 ml cuvette and the Tm determined as the maximum of the first derivative of the melting curve. Control experiments were performed using unmodified DNA duplexes. Typical concentrations were 1 µM for each strand.

Circular dichroism (CD) spectra were recorded using a JASCO J-720 Spectropolarimeter. All CD experiments were performed at 25°C in 1 mm path length cuvettes with a buffer (pH 7.4) containing 25 mM sodium phosphate and 100 mM NaCl. The concentration was 10 µM for each strand. Prior to CD analysis, each sample was incubated briefly at 95°C, equilibrated to room temperature, chilled on ice for 30 min and finally re-equilibrated to 25°C.

Exonuclease digestion experiments were carried out using snake venom phosphodiesterase (SVP) I from Crotalus adamanteus (USB). The assays were performed using a mixture of 5'-32P-labeled ODN (100 000 c.p.m.) and unlabeled ODN (40 pmol) in a buffer (pH 8.5) containing 50 mM Tris–HCl, 72 mM NaCl and 14 mM MgCl2. Enzyme was added to a final concentration of 0.5 U/ml (40 µl total reaction volume). The reaction mixture was overlaid with 25 µl of mineral oil and incubated at 37°C. Aliquots (3.5 µl) were removed, quenched by adding 12 µl of 7 M urea in 90 mM Tris-borate pH 8.3 and 2 mM EDTA (TBE buffer), and heated to 95°C for 5 min. Samples were taken at 0, 1, 3, 7 and 18 h after the addition of enzyme and were stored at –70°C until analysis by PAGE (20% containing 7 M urea) or HPLC. Autoradiographic imaging was performed using a Molecular Dynamics Phosphorimager (Storm 820). Gel bands were quantitated using ImageQuant software (version 5.1).

Endonuclease digestion experiments were carried out using DNase I (Boehringher-Mannheim). The assays were performed using a mixture of 5'-32P-labeled ODN (100 000 c.p.m.) and unlabeled ODN (40 pmol) in a buffer (pH 7.5) containing 40 mM Tris–HCl and 6 mM MgCl2. Enzyme was added to a final concentration of 1 U/µl (40 µl total reaction volume). The reaction mixture was overlaid with 25 µl of mineral oil and incubated at 37°C. Aliquots (3.5 µl) were removed, quenched by adding 12 µl of 7 M urea in TBE buffer, and heated to 95°C for 5 min. Samples were taken at 0, 1, 3, 7 and 18 h after the addition of enzyme and were stored at –70°C until analysis by PAGE (20% containing 7 M urea). Autoradiographic imaging was performed on a Molecular Dynamics Phosphorimager (Storm 820). Gel bands were quantitated using ImageQuant software (version 5.1).

Phosphonoacetate and thiophosphonoacetate ODNs were tested for stability toward HeLa cell nuclear extracts using the following assay. HeLa cell nuclear extracts were prepared by methods described previously (6). ODNs (500 pmol) in a buffer containing 50 mM Tris–HCl pH 8.0, 20 mM KCl, 10 mM MgCl2 and 3 mM dithiothreitol (DTT) were incubated with 20 µg of HeLa cell nuclear extract (1 µl) at 37°C for 5 h (40 µl total volume). A second 20 µg aliquot of cell extract was added and the reaction proceeded for another 5 h. This step was repeated once more for a total of 15 h exposure to HeLa cell nuclear extract. The reaction was quenched by adding 3 vols of 8.3 M urea in TBE buffer and heating the samples at 95°C for 5 min. The reaction products were analyzed by ion-exchange HPLC using a 1 ml Resource Q column of 6.4 mm ID obtained from Amersham/Pharmacia. Gradient eluents were (A) 10 mM NaOH/80 mM NaBr and (B) 10 mM NaOH/1.5 M NaBr. The gradient was 0% A to 100% B in 45 min at a flow rate of 1.5 ml/min.

Escherichia coli RNase H1 kinetic experiments were done as follows. 5'-32P-labeled RNA (100 000 c.p.m./reaction), unlabeled RNA (100 pmol) and complementary ODN (100 pmol) were added to a buffer (pH 7.8) containing 20 mM HEPES-KOH, 50 mM KCl, 10 mM MgCl2 and 1 mM DTT. The duplexes were hybridized by heating briefly to 95°C and then incubated at 4°C for 30 min. Escherichia coli RNase H1 (Promega) was added (8 U) and the reactions were allowed to proceed for various times at 25°C (40 µl total reaction volume). Aliquots of the reaction mixture (3.5 µl) were quenched with 6.5 µl of 7 M urea and 20 mM EDTA and stored on ice until analysis by gel electrophoresis (20%, 19:1 cross-link). All reactions were performed in triplicate. Data were analyzed by KaleidaGraph using a linear curve fit.

Cell transfection experiments were performed as follows. SKBR-3 cells (American Type Culture Collection) were propagated in RPMI 1640 containing 10% (v/v) fetal bovine serum. Cells (100 000 per well) were plated onto 4-well chambered coverglass slides (Nalge Nunc) and grown overnight. Following removal of medium, transfections were carried out at 10 µM ODN in serum-free Opti-MEM. All cells were viewed live in culture medium or phosphate-buffered saline at various times on an Olympus IMT2-RFC reflected light fluorescence inverted microscope. Propidium iodide counterstaining was carried out in order to assess cell viability. Minimal cell death was observed with cells incubated at 10 µm ODN.


   RESULTS
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 REFERENCES
 
The chemical synthesis of phosphonoacetate and thiophosphonoacetate ODNs was carried out in the solid phase using the scheme outlined in Figure 2. Briefly, appropriately protected deoxyribonucleoside 3'-O-diisopropylaminophosphinoacetic acid dimethyl-ß-cyanoethyl esters were condensed to a deoxyribonucleoside linked to CPG (12). The phosphonite linkage was then oxidized to phosphonate using (1S)-(+)(10-camphorsulfonyl)-oxaziridine. Following capping with acetic anhydride and treatment with trichloroacetic acid to remove the 5'-O-dimethoxytrityl group, the cycle was repeated an appropriate number of times to yield the phosphonoacetate ODN. By substituting 3H-1,2-benzodithiol-3-one-1,1-dioxide (BDT) as the oxidant, a thiophosphonoacetate internucleotide linkage was generated. Using this approach, coupling yields of >97% were observed for oligomers up to 18 nucleotides in length. Isolated yields of 18mers (single peaks, ion exchange HPLC) were 32–33%. 31P-Nuclear magnetic resonance (NMR) analysis for these P-chiral oligomers gave single, broad peaks as expected at 28 and 94 p.p.m., respectively, for the phosphonoacetate and thiophosphonoacetate derivatives. The lack of any phosphate signal indicated that these oligomers were free of phosphate internucleotide linkages. Using the same synthesis cycle, chimeric ODNs containing phosphate, phosphorothioate and phosphonoacetate/thiophosphonoacetate internucleotide linkages were also prepared from standard deoxynucleoside 3'-phosphoramidites and deoxynucleoside 3'-O-diisopropylaminophosphinoacetic acid dimethyl-ß-cyanoethyl esters. Mass characterization for phosphonoacetate, thiophosphonoacetate, chimeric ODNs and control ODNs used for various studies gave the expected results (Table 1).


View this table:
[in this window]
[in a new window]
  		Table 1. MALDI-TOF mass spectrometry of ODNsa
 
For transfection experiments, oligodeoxythymidine 12mers were synthesized that had variable ratios of phosphorothioate and esterified thiophosphonoacetate internucleotide linkages. These oligomers were prepared using standard 5'-O- dimethoxytrityldeoxythymidine 3'-O-diisopropylphosphoramidites (13) and appropriately esterified (methyl or butyl) 5'-O-dimethoxytrityldeoxythymidine 3'-O-diisopropylaminophosphinoacetic acid esters. The synthesis cycle was the same as outlined in Figure 2, with BDT as the sulfurizing reagent. These oligomers were synthesized on 3'-phosphate-CPG so that they could be cleaved from the support under non-nucleophilic conditions.

To indirectly compare the charge state of a phosphonoacetate linkage with that of a natural phosphodiester, the retention times for dTT dimers containing a phosphonoacetate, thiophosphonoacetate or phosphodiester linkage were determined on a reverse-phase HPLC column using buffers at various pH values between 1 and 8. A change in retention time for the phosphonoacetate and thiophosphonoacetate dimers occurred at ~pH 4, while the retention time of the phosphodiester dimer remained unchanged until ~pH 1.5 (Fig. 3). This increase in HPLC retention time observed at lower pH for the phosphonoacetate and thiophosphonoacetate dimers indicated an increase in hydrophobic character due to ionic charge neutralization. Approximate pKa values for the phosphonoacetate and thiophosphonoacetate linkages were 3.8 and 3.9, respectively. While only estimates, these values indicated that phosphonoacetate and thiophosphonoacetate ODNs were likely to be charged near physiological pH. As expected due to the presence of hydrophobic sulfur, the thiophosphonoacetate dimer had a greater retention time than the corresponding phosphonoacetate compound. This relative enhanced hydrophobicity remained irrespective of the dimer charge state.



View larger version (10K):
[in this window]
[in a new window]
  		Figure 3. HPLC column retention times relative to pH for thymidylate dimers. Each data point represents the average value from three separate experiments. Thiophosphonoacetate (squares), phosphonoacetate (circles) and DNA (triangles).

 
Owing to the presence of a carbon–phosphorus bond, we expected the phosphonoacetate and thiophosphonoacetate ODNs to be nuclease resistant. In order to test this possibility, nuclease studies were completed using DNase I, SVP and HeLa cell nuclear extract. Attempts to monitor these studies using 5'-32P-labeled, fully modified ODNs was not possible as T4 kinase failed to phosphorylate these oligomers. With methylphosphonate DNA, similar problems were overcome by adding one phosphodiester linkage at the 5' end of an oligomer (15). This proved to be a successful strategy for the phosphonoacetate and thiophosphonoacetate oligomers as well. This type of ODN was therefore synthesized, purified and characterized by 31P NMR and mass spectra (Table 1, ACE* and S-ACE*). Phosphorylation of these modified ODNs required a 4-fold excess of [{gamma}-32P]ATP with respect to the conditions used for unmodified DNA.

When tested with SVP, these oligomers generated the profiles shown in Figure 4. Under conditions where natural DNA has undergone considerable degradation in 1 h and complete hydrolysis within 3 h (lanes 8 and 9), neither the phosphonoacetate nor the thiophosphonoacetate oligomers were degraded, even after 18 h (lanes 1–6 and 13–18, respectively). Lanes 19–24 show the hydrolysis products from incubation of phosphorothioate DNA with SVP. As expected from previous results (6), this analog degraded, but at a slower rate than natural DNA (65 and 40% full-length oligomer remains after 8 and 18 h, respectively). Chimeric ODNs when analyzed by HPLC were essentially resistant towards degradation by SVP. For example, oligomers with phosphonoacetate/phosphate (ACE EO) or thiophosphonoacetate/phosphate (S-ACE EO) at every other linkage were stable toward degradation. Similarly, phosphorothioate oligomers with the 3' and 5' ends capped with either phosphonoacetate (ACE Cap) or thiophosphonoacetate (S-ACE Cap) were also stable (see Table 1 for the sequences of these oligomers).



View larger version (64K):
[in this window]
[in a new window]
  		Figure 4. SVP analysis of radiolabeled ODNs. The SVP digestion results with various ODNs are as follows: ACE*, lanes 1–6; P, lanes 7–12; S-ACE*, lanes 13–18; PS, lanes 19–24. Control reactions without enzyme are displayed in lanes 1, 6, 7, 12, 13, 18, 19 and 24. See Table 1 for the structures defined by these abbreviations.

 
In order to characterize the stability of these ODNs towards endonucleases, the radiolabeled oligomers having one 5'-internucleotide phosphate linkage were incubated with DNase I under conditions similar to those used with SVP. Results are shown in Figure 5. Analysis of lanes containing phosphonoacetate (lanes 1–6) and thiophosphonoacetate (lanes 13–18) ODNs showed no degradation even after 18 h. Again, as expected from previous results (6), the phosphorothioate oligomer was partially degraded (78% full-length product remained after 18 h, lanes 19–24). Under these conditions, full-length natural DNA was degraded even after 1 h to a predicted series of smaller oligomers.



View larger version (61K):
[in this window]
[in a new window]
  		Figure 5. DNase I analysis of radiolabeled ODNs. The DNase I digestion results with various ODNs are as follows: ACE*, lanes 1–6; P, lanes 7–12; S-ACE*, lanes 13–18; PS, lanes 19–24. Control reactions without enzyme are displayed in lanes 1, 6, 7, 12, 13, 18, 19 and 24. See Table 1 for the structures defined by these abbreviations.

 
To further examine the stability of phosphonoacetate and thiophosphonoacetate ODNs towards nucleases, fully modified oligomers were incubated with HeLa cell nuclear extract and analyzed by ion-exchange HPLC (5'-32P-labeling could not be used for reaction product analysis because this extract was rich in phosphatases). Under conditions where the unmodified oligomer was completely degraded, both the phosphonoacetate and thiophosphonoacetate ODNs were unchanged and there were no degradation products (data not shown).

These results encouraged us to examine whether modified oligomers having phosphonoacetate and thiophosphonoacetate internucleotide linkages would form duplexes with natural, complementary DNA and RNA. This was important as our initial objective was to examine these modified oligomers with enzymes that required duplexes for biochemical activity (RNase H and DNA polymerases). Heteroduplexes were prepared using equimolar amounts of phosphonoacetate, thiophosphonoacetate and phosphorothioate ODNs in combination with an unmodified, complementary DNA or RNA strand. Natural duplexes with the same sequences were used as controls. Experiments with RNA–DNA duplexes 18 nucleotides in length were carried out with buffers of variable ionic strength and pH (Table 2). One of the buffers (buffer B) was also used for RNase H studies. As expected for oligomers with multiple P-chiral centers, the melting curves, although cooperative, were not as sharp as for natural DNA, but more similar to results observed previously with phosphorothioate DNA (3). The Tms with complementary DNA indicated that both the phosphonoacetate and thiophosphonoacetate oligomers reduced duplex stability. Further analysis confirmed that the presence of sulfur depressed the Tm for the thiophosphonoacetate-containing duplex relative to phosphonoacetate. For RNA–DNA duplexes, lower Tms were observed with buffer B, which has reduced ionic strength and higher pH than buffer A. The observation that duplexes with phosphonoacetate and thiophosphonoacetate oligomers were still formed but with reduced Tms relative to the unmodified control was encouraging. As was the case with VII and VIII, the RNA–DNA hybrid duplex having sulfur (IV) displayed a lower Tm than the phosphonoacetate duplex (III).


View this table:
[in this window]
[in a new window]
  		Table 2. Tms for modified duplexesa
 
To further characterize duplexes III and IV (Table 2), CD spectra were recorded at 25°C and compared with duplex I (Fig. 6). The CD spectra of duplex I exhibited the characteristics of A-form rather than B-form as expected for an RNA–DNA hybrid (16). For example, the positive band at 266 nm and the negative band at 211 nm are characteristic of the A-form pattern. Similarly, the phosphonoacetate–RNA (III) and thiophosphonoacetate–RNA (IV) heteroduplexes form A-like conformations. This A-like form appears to be important for the recognition and cleavage of substrates by RNase H (17).



View larger version (17K):
[in this window]
[in a new window]
  		Figure 6. CD spectra of 18mer ODN–RNA duplexes. Phosphonoacetate– RNA (circles), thiophosphonoacetate–RNA (squares) and DNA–RNA (triangles).

 
Because these analogs appear to be anionic at physiological pH, nuclease stable and form A-like duplexes with complementary RNA, their ability to stimulate E.coli RNase H1 activity was examined. In order to monitor for this activity, the RNA component (an 18mer) was 5' labeled with [32P]phosphate using T4 kinase and [{gamma}-32P]ATP. RNA substrate was then annealed with various phosphonoacetate- and thiophosphonoacetate-modified, complementary ODNs and tested for cleavage with RNase H1. Results are summarized in Figure 7 and Table 3. For each modification, three oligomers were examined (Table 1 contains the sequences, order of internucleotide modifications and appropriate abbreviations for phosphonoacetate, ACE, and thiophosphonoacetate, S-ACE, oligomers). In all cases, these oligomers stimulated degradation with RNase H1. There were, however, major differences. For example, the completely modified ODNs (ACE and S-ACE) were the least active and the capped oligomers (ACE Cap and S-ACE Cap) were even more efficient at stimulating RNase H1 than unmodified DNA. The remaining oligomers were of intermediate activity, with ACE EO and S-ACE EO having rates comparable with the phosphorothioate control ODN.



View larger version (10K):
[in this window]
[in a new window]
  		Figure 7. RNA degradation profiles for chimeric phosphonoacetate- and thiophosphonoacetate-containing heteroduplexes. Duplexes having radiolabeled RNA 18mer oligonucleotide with a sequence complementary to the 18mer deoxynucleotides defined in Table 1 were incubated with RNase H1 for the times indicated. Deoxynucleotide structures: open squares, normal DNA; open diamonds, phosphorothioate DNA; open circles, ACE Cap; filled diamonds, S-ACE Cap; filled circles, ACE EO; plus signs, S-ACE EO; filled squares, ACE; crosses, S-ACE.

 

View this table:
[in this window]
[in a new window]
  		Table 3. Initial rates for RNase H1
 
Because of encouraging results with RNase H1, experiments were carried out to determine the transfection ability of these ODNs. The approach was based upon earlier work with AZT and oligodeoxyribonucleotide derivatives where charge neutralization was shown to improve cell uptake over fully ionized species. These compounds were designed to initially diffuse across cell membranes and then be converted to biochemically active components by endogenous cellular carboxy esterases (18,19). Since the converted species were anionic, they were unable to diffuse out of the cell, which resulted in cellular enrichment. We speculated that ODNs with neutral thiophosphonoacetate esters would be hydrolyzed in a similar manner and that the ionized species would prevent diffusion out of cells.

Preliminary studies were carried out with chimeric oligomers having a variable number of phosphorothioate linkages interspersed with either the methyl or butyl thiophosphonoacetate ester linkages. The thiophosphonoacetate modification was chosen over the phosphonoacetate because of its increased hydrophobic properties (Fig. 3). Chimeric ODNs were prepared as oligodeoxythymidine 12mers having a 5'-pivaloyl-protected FITC group. Hydrolysis of pivaloyl-protected FITC generates a green fluorescent signal and served as an indicator for cellular uptake and esterase activity (19). Initial results are presented in Figure 8. As can be seen in Figure 8B, the thymidine oligomer with a charge-to-linkage ratio of 0.5 (anionic phosphate/total number of linkages) was taken up by the cells (compare Fig. 8A and B) after only 6 h. The fluorescent stain did not immediately localize in the nucleus but remained diffuse throughout the cells (not on the cell surface). This was shown by observing internal fluorescence around subcellular structures and the nucleus. Confocal microscopy (not shown) also confirmed internalization of fluorescence. After 24 h, these ODNs began to accumulate in the nucleus (Fig. 8C). When the transfection medium (6 h exposure to cells) was replaced with fresh medium lacking ODN, the diffuse stain disappeared within 2 h (Fig. 8D) and left behind a stable punctate pattern. These cells continued to grow after fluorescence had diffused away and remained viable.



View larger version (93K):
[in this window]
[in a new window]
  		Figure 8. Transfection with methyl-protected thiophosphonoacetate ODNs. (A–D) and (F) show results obtained with 5' FITC-TxTxT-TxT-TxTxT-TxT-TxTx where - represents phosphorothioate (including a phosphorothioate linkage between FITC and 5' thymidine) and x represents methyl- protected thiophosphonoacetate (including a thiophosphonoacetate at the 3' end having one negative phosphate charge). Phase contrast (A) and the fluorescent pattern (B) of the same cells after 6 h exposure with this ODN. (C) A 24 h exposure to this ODN. (D) A 6 h exposure to this ODN, removal of medium containing this ODN, and replacement with fresh medium for 2 h. (F) Lipofectin-treated ODN exposed to cells for 4 h followed by fresh medium for 24 h. (E) Results where cells were transfected for 6 h with 5' FITC-T-TxT-TxT-TxT-TxT-TxT-Tx having a phosphorothioate between FITC and 5' thymidine and a thiophosphonoacetate with one negative charge at the 3' end.

 
The residual punctate pattern (Fig. 8D) suggests that a fraction of the methyl phosphonoacetate esters were hydrolyzed (and trapped) within endosomes and/or lysosomes. To test this theory, ODNs with a charge-to-linkage ratio of 0.75 were incubated for 6 h with cells. The results (Fig. 8E) illustrate a typical punctate pattern, as has been observed many times when negatively charged oligomers, such as phosphorothioates, are exposed to cells in culture. These patterns have typically been interpreted as the irreversible accumulation of oligomers in endosomes and/or lysosomes (20). As also observed with phosphorothioate DNA (20), transfection of these ODNs with cationic lipids such as lipofectin resulted in a nuclear stain (Fig. 8F) that did not diffuse when the medium was replaced. Since lipofection goes through an endosomal pathway, this result indicates that endosomal/lysosomal enzymes are capable of deprotection (20). Results similar to those presented in Figure 8A–D and F were also obtained with the corresponding butyl derivative and with a thymidine 12mer where six phosphorothioates were flanked on each side by esterified thiophosphonoacetate linkages.


   DISCUSSION
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
REFERENCES
 
A comparison of melting temperatures ({Delta}Tm expressed as a change in °C per linkage) showed a loss of heteroduplex stability for these modified ODNs relative to natural DNA–RNA and DNA–DNA duplexes. When hybridized to RNA in high salt buffer (Table 2, buffer A), the average decreases in Tms were 1.3 and 1.8°C per linkage (18mer ODNs) for phosphonoacetate and thiophosphonoacetate duplexes (III and IV). This decrease was considerably greater than for the corresponding phosphorothioate duplex (0.5°C/linkage). In the low salt buffer used for experiments with RNase H1 (buffer B), trends were similar but with reduced Tms, as expected. With DNA in a high salt buffer (buffer A), these analogs were also less stable than the duplex with natural DNA (V). For example, with the 18mer duplexes, the decrease in stability was ~0.3 and 1.0°C per linkage, respectively, for the phosphonoacetate (VII) and thiophosphonoacetate (VIII) analogs. Interestingly, the {Delta}Tm per linkage for the phosphonoacetate ODN was less than that observed with phosphorothioate DNA (0.3 versus 0.5°C). Without more data relative to conformational analysis, hydrogen bonding of the acetate moiety and duplex hydration, it is difficult to formulate an appropriate explanation for this observation. However, one possible speculation relates to the conformational flexibility of the acetate group. To elaborate, for phosphorothioate-containing duplexes, where sulfur retains the negative charge, the Rp configuration places sulfur, and therefore the negative charge, in the major groove (B conformation). In contrast, with the Sp isomer, the phosphorothioate sulfur with negative charge is located directly on the face of the sugar–phosphate backbone and hence protrudes into solvents surrounding the duplex. Phosphorothioate duplexes with the Sp configuration can have higher Tms than those with Rp under certain sequence circumstances (21). With phosphonoacetate DNA and owing to the greater flexibility of the acetate group, perhaps the negative carboxylate for both isomers can be positioned exterior to the duplex, which should destabilize the duplex less than for phosphorothioate.

Phosphonoacetate and thiophosphonoacetate ODNs, when hybridized to complementary RNA, were substrates for E.coli RNase H1. It has been reported that increasing the affinity of an ODN for complementary RNA (by increasing G–C base pairs or through chemical modification) can result in increased RNase H-mediated RNA hydrolysis (22). However, the reduced Tm for the thiophosphonoacetate–RNA duplex (IV) relative to the phosphonoacetate–RNA duplex (III) did not lead to a subsequent reduction in RNase H1 activity. Qualitatively, the thiophosphonoacetate–RNA heteroduplex appeared to be a better substrate for RNase H1 than the phosphonoacetate–RNA heteroduplex (Fig. 7 and Table 3). Perhaps this increase in hydrolysis products might be caused by differences in the global conformation of the two heteroduplexes. However, analysis of the CD spectra of these two heteroduplexes showed that both closely resembled that of the natural DNA–RNA duplex (Fig. 6), indicating that the difference in RNase H1 activity is not likely to be a result of duplex structure. For some ribozyme-mediated reactions, duplex substrates with low Tms lead to increased product release and substrate turnover during the course of the reaction (23). Perhaps the reduced Tm for the thiophosphonoacetate– RNA duplex promoted product release and substrate turnover during the RNase H1 reaction.

Of particular interest is the unprecedented observation that the capped ODNs stimulated RNase H1 catalysis at a rate faster than with unmodified DNA (Fig. 7, Table 3). [Although capped phosphorodithioate oligomers have been shown to be more RNase H1 active than natural DNA (6), the results cannot be easily compared with those obtained with the capped chimeras. This is because the experiments with capped phosphorodithioate DNA were carried out in a background rich in nucleases (HeLa cell extracts). As a result, it is difficult to differentiate any intrinsic catalytic rate with capped phosphorodithioate ODNs from a secondary effect due to enhanced stabilization against nuclease degradation.] Earlier work involving similar chimeric, antisense ODNs routinely led to reduced catalysis rates. The rationale for these chimeras is to design antisense oligomers having a central region (phosphorothioate or phosphate internucleotide linkages) that stimulates RNase H1 activity flanked by capped analogs that enhance nuclease resistance and, owing to their ability to form A-type heteroduplexes, have increased affinity for RNase H1 (24). So far, the same properties that perhaps enhance affinity of these chimeras for RNA/RNase H1 result in loss of RNase H1 activity. This does not appear to be the case with a chimera having phosphonoacetate or thiophosphonacetate capped ends and a catalytic phosphorothioate central region (ACE Cap and S-ACE Cap). Without extensive analysis, we can only speculate on the reasons for this enhanced activity. Based upon the CD data with the phosphonoacetate ODN, this chimera probably forms an A-type duplex with RNA, which is the preferred structure recognized by RNase H1 and could lead to a more rapid initial rate. Unlike other modified bases commonly used as capping agents, such as 2'-O-methyl nucleosides, the phosphonoacetate or thiophosphonoacetate analog in complex with RNA forms a less stable duplex. Perhaps, as is the case with certain ribozymes (23), a weaker binding affinity enhanced by cleavage of RNA could destabilize the heteroduplex, which could also increase the enzyme turnover rate. Finally, the phosphonoacetate internucleotide linkage with carboxylic anionic character could form a less stable (relative to phosphate) chimeric duplex with the RNase H1 lysine cluster which is believed to define the enzyme-binding surface (25). This could lead to a more rapid initial rate. Further kinetic analysis of several chimeric constructs may lead to even more enhanced RNase H1 activity.

Because of the unique functionality of these analogs, transfection via a prodrug approach was considered. Others have shown that phosphate-protected S-pivaloylthioethyl ODNs are efficiently transfected into cells in culture via a passive diffusion mechanism (18,19). Following membrane transport, endogenous esterases cleaved the pivaloyl ester and the resulting thio-alcohol eliminated via cyclization. These ODNs were irreversibly trapped and concentrated inside cells, presumably due to an increase in charge character afforded by cell-mediated ester hydrolysis. Furthermore, this approach eliminates the need for toxic cationic lipids during transfection, and the increased hydrophobicity of esterified ODNs has been shown to significantly reduce binding to serum proteins (26).

For the work described here, thiophosphonoacetate- containing ODNs were esterified with either methyl or n-butyl groups and transfection studied as a function of the thiophosphate or anionic content of the oligomers. The results shown in Figure 8 suggest that the relative ratio of anionic charge is important. When half the charge was neutralized via hydrophobic esters, oligomers diffused reversibly into cells and accumulated after removal of the methyl esters. In contrast, oligomers with relatively higher anionic content failed to undergo transfection unless treated with lipofectin. Of particular interest was the reversible diffusion of these esterified analogs. The challenge for future research will be to identify thiophosphonoacetate esters that more clearly resemble natural substrates for esterases and lipases. Perhaps such analogs will undergo hydrolysis more rapidly and thus increase the cellular concentration of the RNase H1-active form of the corresponding ODNs.

Based upon these results, phosphonoacetate- and thiophosphonoacetate-modified ODNs merit further research as these analogs may prove useful for various diagnostic applications and for controlling gene expression via an antisense mechanism. Of particular interest were observations that these ODNs were completely resistant to the nucleases tested, exhibited RNase H1 activity and can be taken up by cells as neutral esters.


   ACKNOWLEDGEMENTS
 
We thank Dr Robert Barkley and the Central Analytical Laboratory for recording the MALDI-TOF spectra. This research was supported by NIH (GM25680 and CA83953) and the University of Colorado.


   REFERENCES
TOP
 ABSTRACT
 INTRODUCTION
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 REFERENCES
 

  1. Crooke,S.T. (1999) Molecular mechanisms of action of antisense drugs. Biochim. Biophys. Acta, 1489, 31–44.[Medline]

  2. Yuen,A., Halsey,J., Fisher,G., Advani,R., Moore,M., Saleh,M., Ritch,P., Harker,G., Ahmed,F., Jones,C. et al. Phase I/II trial of ISIS 3521, an antisense inhibitor of PKC-alpha with carboplatin and paclitaxel in non-small cell lung cancer. 37th Annual Meeting of the American Society of Clinical Oncology, San Francisco, CA, May 12, 2001.

  3. Verma,S. and Eckstein,F. (1998) Modified oligonucleotides: synthesis and strategy for users. Annu. Rev. Biochem., 67, 99–134.[CrossRef][ISI][Medline]

  4. Kean,J.M., Kipp,S.A., Miller,P.S., Kulka,M. and Aurelian,L. (1995) Inhibition of herpes simplex virus replication by antisense oligo-2'-O-methylribonucleoside methylphosphonates. Biochemistry, 34, 14617–14620.[CrossRef][Medline]

  5. Agarwal,S. and Zhang,R. (1997) Pharmacokinetics of oligonucleotides. In Chadwick,D.J. and Cardew,G. (eds), Oligonucleotides as Therapeutic Agents. Ciba Foundation Symposium 209. John Wiley and Sons, New York, pp. 60–78.

  6. Cummins,L., Graff,D., Beaton,G., Marshall,W.S. and Caruthers,M.H. (1996) Biochemical and physicochemical properties of phosphorodithioate DNA. Biochemistry, 35, 8734–8741.[CrossRef][Medline]

  7. Eckstein,F. (2000) Phosphorothioate oligodeoxyribonucleotides: what is their origin and what is unique about them? Antisense Nucleic Acid Drug Dev., 10, 117–121.[ISI][Medline]

  8. Quartin,R.S., Brakel,C.L. and Wetmur,J.G. (1989) Number and distribution of methylphosphonate linkages in oligodeoxyribonucleotides affect exo- and endonuclease sensitivity and ability to form RNase H substrates. Nucleic Acids Res., 17, 7253–7263.[Abstract/Free Full Text]

  9. Vaughn,J.P., Stekler,J., Demirdji,S., Mills,J.K., Caruthers,M.H., Iglehart,J.D. and Marks,J.R. (1996) Inhibition of the ErbB-2 tyrosine kinase receptor in breast cancer cells by phosphoromonothioate and phosphorodithioate antisense oligonucleotides. Nucleic Acids Res., 24, 4558–4564.[Abstract/Free Full Text]

  10. Verbeure,B., Lescrinier,E., Wang,J. and Herdewijn,P. (2002) RNase H mediated cleavage of RNA by cyclohexene nucleic acid (CeNA). Nucleic Acids Res., 29, 4941–4947.

  11. Damha,M.J., Wilds,C.J., Noronha,A., Brukner,I., Borkow,G., Arion,D. and Parniak,M.A. (1998) Hybrids of RNA and arabinonucleic acids (ANA and 2'F-ANA) are substrates for ribonuclease H. J. Am. Chem. Soc., 120, 12976–12977.[CrossRef]

  12. Dellinger,D.J., Sheehan,D.M., Christensen,N.K., Lindberg,J.G. and Caruthers,M.H. (2003) Solid phase chemical synthesis of phosphonoacetate and thiophosphonoacetate oligodeoxyribonucleotides. J. Am. Chem. Soc., 125, 940–950.[CrossRef][ISI][Medline]

  13. Caruthers,M.H., Barone,A.D., Beaucage,S.L., Dodds,D.R., Fisher,E.F., McBride,L.J., Matteucci,M., Stabinsky,Z. and Tang,J.-Y. (1987) Chemical synthesis of deoxyoligonucleotides by the phosphoramidite method. Methods Enzymol., 154, 287–313.[ISI][Medline]

  14. Gupta,N.K., Ohtsuka,E., Weber,H., Chang,S.H. and Khorana,H.G. (1968) The joining of short deoxyribopolynucleotides by DNA-joining enzymes. Proc. Natl Acad. Sci. USA, 60, 285–292.[Free Full Text]

  15. Miller,P.S., Reddy,M.P., Murakami,A., Blake,K.R., Lin,S.B. and Agris,C.H. (1986) Solid phase syntheses of oligodeoxyribonucleoside methylphosphonates. Biochemistry, 25, 5092–5097.[CrossRef][Medline]

  16. Cantor,C.R. and Schimmel,P.R. (1980) Biophysical Chemistry. W.H. Freeman, San Francisco, pp. 118–124.

  17. Lima,W.F., Mohan,V. and Crooke,S.T. (1997) The influence of antisense oligonucleotide-induced RNA structure on Escherichia coli RNase HI activity. J. Biol. Chem., 272, 18191–18199.[Abstract/Free Full Text]

  18. Tosquellas,G., Bologna,J.C., Morvan,F., Rayner,B. and Imbach,J.L. (1998) First synthesis of alternating SATE-phosphotriester/phosphodiester prooligonucleotides on solid support. Bioorg. Med. Chem. Lett., 8, 2913–2918.[CrossRef][Medline]

  19. Vives,E., Dell’Aquila,C., Bologna,J.C., Morvan,F., Rayner,B. and Imbach,J.L. (1999) Lipophilic pro-oligonucleotides are rapidly and efficiently internalized in HeLa cells. Nucleic Acids Res., 27, 4071–4076.[Abstract/Free Full Text]

  20. Bennett,F.C. (1998) Use of cationic lipid complexes for antisense oligonucleotide delivery. In Stein,C.A. and Krieg,A.M. (eds), Applied Antisense Oligonucleotide Technology. John Wiley and Sons, New York, pp. 129–145.

  21. Cosstick,R. and Eckstein,F. (1985) Synthesis of d(GC) and d(CG) octamers containing alternating phosphorothioate linkages: effect of the phosphorothioate group on the B–Z transition. Biochemistry, 24, 3630–3638.[CrossRef][Medline]

  22. Monia,B.P., Lesnik,E.A., Gonzalez,C., Lima,W.F., McGee,D., Guinosso,C.J., Kawasaki,A.M., Cook,P.D. and Freier,S.M. (1993) Evaluation of 2'-modified oligonucleotides containing 2'-deoxy gaps as antisense inhibitors of gene expression. J. Biol. Chem., 268, 14514–14522.[Abstract/Free Full Text]

  23. Herschlag,D. (1991) Implications of ribozyme kinetics for targeting the cleavage of specific RNA molecules in vivo: more isn’t always better. Proc. Natl Acad. Sci. USA, 88, 6921–6925.[Abstract/Free Full Text]

  24. Lima,W.F. and Crooke,S.T. (1997) Binding affinity and specificity of Escherichia coli RNase H1: impact on the kinetics of catalysis of antisense oligonucleotides–RNA hybrids. Biochemistry, 36, 390–398.[CrossRef][Medline]

  25. Kanaya,S., Katsuda-Nakai,C. and Ikehara,M. (1991) Importance of the positive charge cluster in Escherichia coli ribonuclease H1 for the effective binding of the substrate. J. Biol. Chem., 266, 11621–11627.[Abstract/Free Full Text]

  26. Tosquellas,G., Bryksin,A., Alvarez,K., Morvan,F., Vasseuer,J.J., Rayner,B., Rykova,E., Laktionov,P., Vlassov,V. and Imbach,J.L. (2000) Prooligonucleotides exhibit less serum-protein binding than phosphodiester and phosphorothioate oligonucleotides. Nucleosides, Nucleotides Nucleic Acids, 19, 995–1003.[ISI][Medline]


Add to CiteULike CiteULike   Add to Connotea Connotea   Add to Del.icio.us Del.icio.us    What's this?



This Article
Right arrow Abstract Freely available
Right arrow Print PDF (252K) Freely available
Right arrow Alert me when this article is cited
Right arrow Alert me if a correction is posted
Services
Right arrow Email this article to a friend
Right arrow Similar articles in this journal
Right arrow Similar articles in ISI Web of Science
Right arrow Similar articles in PubMed
Right arrow Alert me to new issues of the journal
Right arrow Add to My Personal Archive
Right arrow Download to citation manager
Right arrow Search for citing articles in:
ISI Web of Science (5)
Right arrowRequest Permissions
Right arrow Commercial Re-use Guidelines
for Open Access NAR Content
Google Scholar
Right arrow Articles by Sheehan, D.
Right arrow Articles by Dellinger, D. J.
Right arrow Search for Related Content
PubMed
Right arrow PubMed Citation
Right arrow Articles by Sheehan, D.
Right arrow Articles by Dellinger, D. J.
Social Bookmarking
Add to CiteULike   Add to Connotea   Add to Del.icio.us  
What's this?