A Quantitative Method to Detect RNA Editing Events

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vidin–HRP. The improved outcome of immunoblotting is probably a result of two separate effects: (i) improved efficacy of immuno-inert membrane blocking and (ii) the presence of Tween 20, which may aid in the removal of any residual SDS from proteins and thereby help in renaturation of proteins on the nitrocellulose (12, 13).

A Quantitative Method to Detect RNA Editing Events

SUMMARY

Received July 26, 1999

To improve the signal to noise ratio in immunoblotting with antiphosphoamino acid antibodies, an optimization study with different blocking agents was carried out. The best results were obtained using a mixture of 5% amicase, 5% BSA, and 5% MBA in TTBS over 45 min at RT. There are several advantages to our blocking formula and phosphoamino acid detection with antibodies: (i) The procedure is easy, relatively fast to perform, and widely applicable for detection with a variety of antibodies; (ii) the blocking agents used are inexpensive and available in most laboratories; (iii) the sensitivity of detection is comparable to that of autoradiography of 32 P in vivo labeled protein; (iv) immobilized proteins may be used for additional testing with different antibody after stripping (see Amersham ECL brochure for stripping protocol); and (v) handling of radioactivity is avoided. REFERENCES 1. Roffmann, E., and Meromsky, L. (1986) Biochem. Biophys. Res. Commun. 136, 80 – 85. 2. Michalewski, M. P., Kaczmarski, W., Golabek, A. A., Kida, E., Kaczmarski, A., and Wisniewski, K. E. (1998) Biochem. Biophys. Res. Commun. 253, 458 – 462. 3. http://www.novex.com/16BLOCK.htm, Technical Tips. 4. Gershoni, J. M., and Palade, G. E. (1983) Anal. Biochem. 131, 1–15. 5. Laemmli, U. K. (1970) Nature (London) 227, 680 – 685. 6. Schneider, D. L., and Chin, J. (1988) Methods Enzymol. 157, 591– 601. 7. Towbin, H., and Staehelin, T. (1979) Proc. Natl. Acad. Sci. USA 76, 4350 – 4359. 8. Spinola, S. M., and Cannon, J. G. (1985) J. Immunol. Methods 81, 161–165. 9. DenHollander, N., and Befus, D. (1989) J. Immunol. Methods 122, 129 –135. 10. Hoffman, W. L., and Jump, A. A. (1986) J. Immunol. Methods 194, 191–196. 11. Mackinley, A. G., and Wake, R. G. (1971) in Milk Proteins (McKenzie, H. A., Ed.), Vol. 2, pp. 175–215, Academic Press, New York. 12. DeBlas, A. L., and Cherwinski, H. M., (1983) Anal. Biochem. 133, 214 –219. 13. Dunn, S. D. (1986) Anal. Biochem. 157, 144 –153.

Hans H. Schiffer and Stephen F. Heinemann 1 Molecular Neurobiology Laboratory, Salk Institute for Biological Studies, 10010 North Torrey Pines Road, La Jolla, California 92037

RNA-editing mechanisms affect protein production and function in Trypanosoma, fungi, higher plants, and mammals, and are detectable as insertions, deletions, or modifications of nucleotides in RNA transcripts (1, 2). The detection and quantification of RNA editing in transcripts requires a detection method that is easy and fast to perform, sensitive, and highly reproducible. We developed an RNA-editing detection assay that improves all these features, compared to previously published detection methods. Importantly, this assay can detect all types of base modifications with equal efficiency. The methods currently in use for the quantitative detection of single nucleotide modifications are based on primer extension reactions performed on total RNA (3) or RT–PCR templates (3), or on hybridization techniques, which can discriminate single nucleotide differences in RT–PCR products (4). These methods require extended annealing times in the range of hours to obtain sufficient primer extension products or hybridized oligos, respectively. Another frequently applied method is the use of restriction enzymes, which cleave either the edited or unedited version of RT–PCR products (5). The use of restriction enzymes is limited to editing events that alter restriction enzyme recognition sequences and is further complicated by the need to standardize and control enzyme activity. We describe here a new RNA-editing assay which combines the use of the thermostable ThermoSequenase DNA polymerase (Amersham, Pharmacia, UK) and the amplification of the primer extension reaction with dideoxynucleotides in a thermocycler. Figure 1 illustrates the principle of this cycled primer extension assay, showing the steps involved in the detection of editing at the glutamine(Q)/arginine(R) site of the rat glutamate receptor subunit GluR6 transcript (3). In this case, an adenosine deaminase catalyzes the deamination of a specific adenine (A) residue to inosine (I), changing a CAG codon to CIG. During translation the inosine residue is recognized as guanosine, which causes a glutamine (Q) to arginine (R) alteration in membrane domain 2 of the 1

Supported by grants from the NIH, the McKnight Foundation, and the John Adler Foundation. Analytical Biochemistry 276, 257–260 (1999) Article ID abio.1999.4369 0003-2697/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

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FIG. 1. Schematic illustration of the cycled primer extension assay with dideoxy-terminator ddATP as used to detect Q/R site editing (A to I base modification) in rat GluR6 mRNA.

GluR6 receptor subunit. Approximately 70% of all GluR6 mRNA’s in the adult rat brain are edited at this position (2). To quantify GluR6 Q/R site editing, we first generated RT–PCR products, containing the Q/R site, from whole rat brain total RNA. The obtained RT–PCR products are a mixture of unedited and edited PCR fragments and are used as template for the cycled primer extension reaction. The reaction is catalyzed by the thermostable Thermo-Sequenase DNA polymerase (Amersham, Pharmacia, UK), which has the unique property of incorporating dideoxynucleotides as efficiently as deoxynucleotides (6, 7). During the cycling reaction, the 32P-end-labeled primer (17-mer) complementary to a sequence upstream of the GluR6 Q/R site annealed to the RT–PCR products and was extended in the presence of the dideoxynucleotide ddATP and the deoxynucleotides dTTP, dCTP, and dGTP. The primer position on the GluR6 template was selected to produce ddATP-dependent termination only at the editing site or at the next adenine residue found downstream of the editing site. Elongation of primers annealed to unedited RT–PCR products leads to a 21-mer oligonucleotide (Fig. 1). In contrast, elongation of primer annealed to edited RT–PCR products terminates at the adenine residue downstream of the editing site, leading to a 24-mer oligonucleotide (Fig. 1). Compared to previously described editing assays, the amplification of the primer extension reaction in a thermocycler reduces the reaction time from the range of hours to around 15 min, increases the total amount of extended oligonucleotide products, shows high reproducibility, and allows the simultaneous analysis of large numbers of

samples. Polyacrylamide electrophoresis in combination with phosphorimaging analysis was used to detect and quantify the amounts of extended oligonucleotides, and these data were used to calculate the percentage of editing at the Q/R site of GluR6 mRNA. An autoradiogram of an assay performed to detect the amount of GluR6 Q/R site editing in the adult rat brain is shown in Fig. 2A, lane 2. The estimated ratio of the yields of extended 21-mer and 24-mer oligonucleotides in the reaction mix indicates that ca. 72% of GluR6 mRNAs in the adult rat brain are edited at the Q/R site. Figure 2A (lane 1) shows the result of another assay detecting editing at the Q/R site of the glutamate receptor subunit GluR2 mRNA. In contrast to the GluR6 Q/R site, the GluR2 Q/R site is nearly completely edited (.99%) in adult rat brain, since only one extension product, a 24-mer oligonucleotide, was detected. The percentage of GluR2 and GluR6 Q/R site editing determined by this method is similar to the estimated percentage of editing found in several previous studies using other detection methods (2). Additional experiments were performed to determine the accuracy and sensitivity of the cycled primer extension assay. First, editing of GluR6 mRNA in the rat brain at the Q/R site was determined by performing eight independent RT–PCR reactions on the same total brain RNA, followed by eight primer extension assays. We estimated a mean of 72% GluR6 Q/R site editing with a standard deviation of 0.5%. This result demonstrates that our assay gives highly reproducible results. Second, cloned GluR6 cDNAs of the edited (R) or unedited (Q) variant were used as template for PCR reactions to obtain separate pools of edited and unedited GluR6 PCR fragments. Equal volumes of solutions containing either GluR6(Q) or GluR6(R) PCR fragments were mixed in variable ratios to produce a linear increase of GluR6(R) fragments in the mixtures. The cycled primer extension was performed three times on each mixture to determine the apparent ratio between the edited and unedited PCR products. The autoradiogram from the performed assays is shown in Fig. 2A, lanes 3–9. The estimated ratios for mixes 2– 6 were plotted as percentage editing to demonstrate the accuracy of the assay (Fig. 2B). The graph shows the expected linear relationship between the estimated values for the mixtures 2– 6 (r 5 0.994). Additional mixing experiments demonstrated that the assay can detect RNA editing at levels down to 1–2% (data not shown). When the cycled primer extension assay is performed on PCR products obtained only from unedited cloned GluR6 (Q) cDNA, no extended primer band is detected (Fig. 2A, lane 3). This result demonstrates that during the cycled primer extension assay no significant misincorporation of nucleotides occurs. We are using this assay routinely to investigate editing at eight separate sites in glutamate receptor transcripts,

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FIG. 2. Detection and quantification of RNA-editing events using the cycled primer extension assay. (A) Analysis of RNA editing (A to I base modifications) at the Q/R site of glutamate receptor subunit GluR2 and GluR6 mRNA’s in the adult rat brain (lanes 1 and 2). RT–PCR products derived from GluR2 and GluR6 mRNA were obtained by standard RT–PCR methods. Cycled primer extension assays were performed with ddATP as dideoxy-terminator. The detected 21-mer oligo-nucleotide band relates to unedited RT–PCR fragments, containing an adensosine (A) at the Q/R site. The detected 24-mer oligonucleotide band relates to edited RT–PCR fragments, containing a guanosine (G) at the Q/R site. Mixing experiments were performed to demonstrate the accuracy of the cycled primer extension assay with dideoxyterminators (lanes 3–9). Cloned GluR6 cDNA’s of the unedited (Q) or the edited (R) variant were used as template for PCR reactions to obtain separate pools of edited and unedited GluR6 PCR fragments. GluR6(Q) and GluR6(R) PCR fragments were mixed in various molar ratios, to obtain a linear increase of GluR6(R) fragments in the mixtures. Cycled primer extension assays with ddATP as dideoxy-terminator were performed on these mixes. (B) The cycled primer extension was performed three times on each mixture to determine the actual ratio between the amount GluR6(Q) and GluR6(R) fragments, expressed as percentage editing. The data are plotted 6 standard deviation. The graph shows a linear increase in detected GluR6(R) in the mixtures 2-6 (r 5 0.994).

without the need to alter the reaction conditions from those described here. RNA-editing events were detected and quantified as follows: 10 pmol primer (typically a 17-mer) was endlabeled in a volume of 10 ml using T4-polynucleotide kinase (NEB, Beverly, MA) and 5 ml [g- 32P]ATP (sp act .6000 Ci/mmol; Andotek, Irvine, CA) for 1 h at 37°C. Nonincorporated nucleotides were removed using the QIAquick nucleotide removal kit (Qiagen, Santa Clarita, CA). Purified RT–PCR products (containing the editing site of interest) were mixed together with 1 ml of the purified 32P-end-labeled primer (ca. 0.5 pmol), 100 mmol (final concentration of each nucleotide) T-Mix (dGTP, dCTP, dATP, and ddTTP) or the analogous G-MIX, A-Mix, or C-Mix (dependent on the editing event analyzed), and 1.55 ml Thermo-Sequenase reaction buffer (Amersham). Two units of 39–59 exonuclease minus Thermostable Sequenase (Thermo-Sequenase DNA polymerase, Amersham) was added to reach a final total reaction volume of 20 ml. The cycled primer extension reactions were performed without overlaying oil in a thermocycler with a heated lid (MJ Research, Inc., Watertown, MA). The amplification protocol consisted of 5 cycles with 10 s denaturation at 90°C, 30 s annealing at 50°C, and 10 s elongation at 72°C. Twenty microliters of 23 gel loading buffer (90% formamide, 0.1% bromphenol blue, 178 mM Tris– borate, 5 mM EDTA) was added to each sample after completing the

amplification reaction. The extended primers were then separated by electrophoresis in 15% denaturing polyacrylamide– 8 M urea gels (20 cm long) using 13 TBE (98 mM Tris– borate, 2.5 mM EDTA) as running buffer. All samples were heated to 70°C for 5 min and cooled on ice prior to loading. Following electrophoresis, gels were fixed by gentle shaking in 7% acidic acid, 7% methanol for 45 min, and dehydrated by gentle shaking twice in 100% methanol for 30 min. The dehydrated gels were dried on a heated (70°C) vacuum dryer for 20 min. Gels were exposed on X-ray films (X-OMAT AR, Kadak, Rochester, NY) for 10 min to 1 h to obtain the autoradiogram. Analysis of the dried gels was performed on a phosphorimager (Molecular Dynamics, Sunnyvale, CA), and the software ImageQuant NT (Molecular Dynamics) was used to determine the relative amounts of extended oligos and to quantitate the analyzed editing events. REFERENCES 1. Smith, H. C., Gott, J. M., and Hanson, M. R. (1997) RNA 3, 1105–1123. 2. Seeburg, P. H. (1996) J. Neurochem. 66, 1–5. 3. Driscoll, D. M., Wynne, J. K., Wallis, S. C., and Scott, J. (1989) Cell 58, 519 –525. 4. Sommer, R., Ko¨hler, M., Sprengel, R., and Seeburg, P. H. (1991) Cell 67, 11–19.

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5. Paschen, W., and Djuricic, B. (1994) Cell. Mol. Neurobiol. 14, 259 –270. 6. Reeve, M. A., and Fuller, C. W. (1995) Nature 376, 796 –797. 7. Vander Horn, P. B., Davis, M. C., Cunniff, J. J., Ruan, C., McArdle, B. F., Samols, S. B., Szasz, J., Hu, G., Hujer, K. M., Domke, S. T., Brummet, S. R., Moffett, R. B., and Fuller, C. W. (1997) Biotechnology 22, 764 –765.

Gas–Liquid Chromatographic Determination of Total Glycerophosphate in an Aqueous Solution Masateru Nishihara and Yosuke Koga Department of Chemistry, University of Occupational and Environmental Health, Yahatanishi-ku, Kitakyushu 807-8555, Japan Received August 2, 1999

a-Glycerophosphate (a-GP 1) is an important intermediate in glycerol metabolism and the first precursor of glycerolipid biosynthesis in most living organisms. It consists of two stereoisomers, sn-glycerol-1-phosphate (G-1-P) and sn-glycerol-3-phosphate (G-3-P). G-3-P is formed from glycerol or dihydroxyacetone phosphate (DHAP) in Eukarya, Bacteria, and in some Archaea (1), and G-3-P structure is found as a backbone of all glycerophospholipids in Eukarya and Bacteria, while G-1-P is formed by DHAP reduction in Archaea (2, 3) and also found in unacylated a-GP moiety of eukaryal and bacterial phosphatidylglycerol (4) and bacterial lipoteichoic acid (5) as well as a backbone of all glycerophospholipids in Archaea (6). A sensitive method for determination of total a-GP was developed, and is reported in the present Note. The present Note describes simplified determination method of a-GP using GLC after direct trimethylsilylation in an aqueous solution. After an in vitro a-GP-forming enzyme reaction (an a-GP dehydrogenase reaction or a glycerol kinase reaction) was completed, up to 100 ml of the aqueous solution was transferred to a test tube which had been received 30 mg icosane as an internal standard. To this solution, 0.8 ml of 1,1,1,3,3,3-hexamethyldisilazane (HMDS) and 0.4 ml of trimethylchlorosilane (TMCS) were added in this order. After bubbling ceased, 0.8 ml of HMDS and 0.4 ml of TMCS were added again and the solution was heated at 110°C for 60 min. After heating, the supernatant was transferred to an Eppen1 Abbreviations used: a-GP, a-glycerophosphate; DHAP, dihydroxyacetone phosphate; G-1-P, sn-glycerol-1-phosphate; G-3-P, snglycerol-3-phosphate; HMDS, 1,1,1,3,3,3-hexamethyldisilazane; TMCS, trimethylchlorosilane; TMS, trimethylsilyl.

Analytical Biochemistry 276, 260 –261 (1999) Article ID abio.1999.4370 0003-2697/99 $30.00 Copyright © 1999 by Academic Press All rights of reproduction in any form reserved.

FIG. 1. Standard curve of total GP determination by GLC. Up to 100 ml solution containing a-GP and 30 mg icosane (the internal standard) was treated with a large excess of HMDS and TMCS, and a portion of the concentrated solution was injected into a gas chromatograph. For experimental details, see text.

dorf tube. The test tube and the precipitates remaining in the test tube were rinsed with a small volume of hexane, and the rinse was combined to the supernatant. After a centrifugation at 10,000 rpm for 3 min, the supernatant was saved and concentrated to 10 –20 ml under N 2 at 37°C. It should be noted that the complete drying of this sample under N 2 may cause loss of trimethylsilylated (TMS)-a-GP. A small volume of the concentrated sample (2– 4 ml) was used for the determination of a-GP by a Shimadzu GC 9AM gas–liquid chromatograph equipped with flame ionization detectors and a 30-m DB-1 column (J & W Scientific, Folsom, CA) and analyzed by increasing a temperature from 125 to 285°C at 15°C/min. A peak at the retention time of 6.68 min on the chromatogram was identified as TMS–a-GP by GLC– MS (data not shown). TMS-b-glycerophosphate was eluted faster (6.01 min) than TMS-a-GP and clearly separated from TMS-a-GP. b-Glycerophosphate formation from a-GP during trimethylsilylation was not observed. The ratio of GLC signal area of a-GP to that of the known amount of the internal standard (retention time, 8.30 min) was proportional to the content of a-GP between 20 and 400 nmol in the original aqueous solution (up to 100 ml) (Fig. 1). The constituents in the aqueous enzyme reaction mixture did not disturb GLC of a-GP, although many peaks appeared on the chro-