- Email: [email protected]
A Chemical Modification Method for the Structural Analysis of RNA and RNA–Protein Complexes within Living Cells
240
NOTES & TIPS
addition of NaCl, KCl, glucose, or serum to the calibration buffer had much less effect. The difference between glass membrane and solid-state electrodes was unaffected by varying the concentration or pH of the compounds in standard calibration buffer. The solid-state electrode deviated more from the original calibration buffer pH value than the glass membrane electrode did when glucose or FBS was added to the calibration buffer and even more so with the addition of MgCl2 or CaCl2 (Table 2, columns 4 and 5). Although the solid-state system may be more sensitive to divalent cations than the glass membrane electrode, the glass membrane electrode was not completely impervious to competing ions. Although there was only a small pH difference (DpH) between measured values for glass and solid-state electrodes when NaCl or KCl was added to the standard calibration buffer, both types of electrodes exhibited a decrease in absolute pH of similar magnitude with increasing Na1 or K1 (data not shown), likely caused by pKa shifts in the acids/bases of the buffering system with increasing ionic strength. Since most physiologic buffers contain Mg21 and/or Ca21, care must be taken when selecting an electrode to measure pH, as well as the method of electrode calibration. Both types of electrodes were affected when the test solution differed greatly from the calibration buffer in salt concentration or ionic composition. The use of internal standards or matrix matching may increase accuracy of pH calibration and measurements. Nevertheless, a real discrepancy can arise when a laboratory switches to the new solid-state technology without realizing that the solid-state pH measurements in physiologic buffers are not directly comparable to the established glass membrane technology when standard calibration buffers are used. A DpH of 0.1– 0.2 between solid-state and glass membrane electrodes is significant when one considers the sensitivity of biological systems to small changes in pH within the physiologic pH range. However, this does not necessarily detract from the usefulness of the solid-state technology due to its ease of use and rapid response time. If a laboratory chooses to use a solid-state electrode for pH measurements of biological buffers, a simple conversion to glass electrode values will enable their data to be compared to the existing literature. Acknowledgments. The authors thank Ronald A. Coss, Ph.D., Dennis B. Leeper, Ph.D., Christopher Storck (Department of Radiation Oncology, Thomas Jefferson University, Philadelphia, PA), Michael D. O’Hara, Ph.D. (Cordis Institute, Johnson & Johnson, Co., Inc., Warren, NJ), and John Brekke (Sentron, Inc., Gig Harbor, WA) for their help and advice during this work. The authors also thank Andreas Woldegiorgic (Academy of Science, Stockholm, Sweden) for editorial help.
REFERENCES 1. Bergveld, P. (1972) IEEE Trans. Biomed. Eng. BME-19, 342– 351. 2. Sentron, Inc. Technical Documentation. 3. Kress-Rogers, E. (1991) Trends Food Sci. Technol., 320 –324. 4. Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985) J. Biol. Chem. 260, 3440 –3450. 5. Thomas, J., Buchsbaum, R., Zimniak, A., and Racker, E. (1979) Biochemistry USA 18, 2210 –2218.
A Chemical Modification Method for the Structural Analysis of RNA and RNA±Protein Complexes within Living Cells Monika Balzer* and Rolf Wagner1 Institut fu¨r Physikalische Biologie, Heinrich-HeineUniversita¨t Du¨sseldorf, Universita¨tsstrasse 1, Du¨sseldorf, D-40225 Germany Received August 25, 1997
Studies on the structure and function of RNA and RNA–protein complexes (RNPs)2 are of increasing interest. Chemical modification and limited enzymatic hydrolysis (structural probing) are powerful methods for obtaining detailed information on the structure and dynamics of nucleic acids in solution (1, 2). Their use, however, is widely limited to studies outside the cellular compartments. The only reagent which has gained widespread use in the in vivo modification of RNA or DNA so far is dimethyl sulfate (DMS) which methylates N7 of guanosines, the N1 position of adenosine, and N3 of cytosines. Identification of modified sites is normally carried out by the rapid primer extension procedure. However, the N7-methylguanosine cannot readily be identified by primer extension, and more tedious procedures for a chemical cleavage must be employed. To circumvent the difficulty in obtaining structural information for guanosines, and to extend the number of methods for in vivo structural probing, we developed a simple chemical method for in vivo modification taking advantage of the reagent kethoxal (a-keto-b-ethoxybutyraldehyde). Kethoxal reacts with guanosines not involved in Watson–Crick base pairs in a highly specific way and under mild conditions by forming a five-membered ring between the two carbonyl groups and the N1 and N2 positions of guanosine (3). Kethoxal-modified guanosines are not accepted as 1 To whom correspondence should be addressed. Fax: 49 211 811 5167. E-mail: [email protected]. 2 Abbreviations used: RNPs, RNA–protein complexes; DMS, dimethyl sulfate.
ANALYTICAL BIOCHEMISTRY 256, 240 –242 (1998) ARTICLE NO. AB972499
0003-2697/98 $25.00 Copyright © 1998 by Academic Press All rights of reproduction in any form reserved.
NOTES & TIPS
template for the reverse transcriptase, and can thus readily be identified by primer extension. Kethoxal has not been described for in vivo structural probing previously. We have developed a simple method which enables a rapid and universally applicable analysis of accessible guanosines within nucleic acids or protein nucleic acid complexes in vivo. Tedious additional chemical treatments for the identification of the modified guanosines are avoided. Existing in vivo modification techniques like DMS probing are thus improved. In the following we describe an in vivo modification procedure which is straightforward, consisting of three steps: (i) increasing the permeability of bacterial cell walls for the rapid uptake of kethoxal, (ii) the modification reaction which can be performed for various times at all temperatures not detrimental to the cells, and (iii) RNA (or RNP) extraction and analysis of modified nucleotides by primer extension. Suitable conditions for the modification of ribosomal RNA. HB101 cells were grown in YT medium at 30°C to an optical density of 0.5 A600. Two 100-ml aliquots of the culture were taken and the cells were harvested by centrifugation (5 min, 5000 rpm). The pellets were resuspended carefully in 1 ml 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8) and incubated at 30°C with gentle shaking. After 15 min incubation 5 ml of 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8) was added to the control culture, while the modification culture was supplemented with 5 ml 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8), 2% ethanol, presaturated with kethoxal. Incubation was continued for another 30 min at 30°C, after which the cells were harvested by centrifugation. Cell pellets were resuspended in 30 ml 25 mM Tris–HCl (pH 7.6), 60 mM KCl, 10 mM MgCl2, 20% (w/v) sucrose, 150 mg/ml lysozyme, and lysis was performed by repeated freeze–thawing (at least three times) as described (6). One hundred microliters of TKM buffer (25 mM Tris–HCl (pH 7.6), 60 mM KCl, 5 mM MgCl2), 34 ml 1% sodium– deoxycholate (w/v), 20 ml of a 5% (w/v) polyoxyethylene ether (20 cetyl ether, Brij 58), 8 ml 20 mM sodium– borate (pH 6.8), and 15 units DNase I (RNase-free) were added to the disrupted cells and incubated for 15 min on ice. The cell debris is removed by centrifugation, and the cleared supernatant can be used for RNA extraction or, as in this case, as a source for ribosome preparation. Demonstration of efficient modification of cellular RNA components. First, we established conditions under which a rapid kethoxal uptake and reaction could be obtained. As a marker we followed the guanosine modification of ribosomal 5S RNA within the cells. The kethoxal modification of 5S RNA has been carefully analyzed previously (4, 5), and G41 had been established as a position of outstanding reactivity within the complete ribosome. The kethoxal
241
FIG. 1. Analysis of 5S RNA after in vivo modification with kethoxal. The separation of 59-labeled oligonucleotides from a total RNase T1 digestion of 5S RNA extracted from in vivo kethoxalmodified cells on a 20% acrylamide/8 M urea gel is shown. (1) RNase T1 digestion after modification at G41 leads to an 11-mer (indicated by the arrow) which is not normally present in T1 digests of unmodified 5S RNA. (2) Adjustment of the oligonucleotides to pH 9 with NH4OH and incubation at 37°C for 60 min followed by a second RNase T1 digestion causes the 11-mer to disappear almost totally.
modification can be easily analyzed because modified guanosines are not recognized by RNase T1 and therefore give rise to new products after a complete RNase T1 hydrolysis (10 units, 60 min, 37°C). Because the kethoxal modification is reversible under alkaline conditions, redigestion after a mild base treatment (1 ml 25% NH4OH, 60 min, 37°C) yields the normal RNase T1 products again. Figure 1 shows an example of a pilot experiment where a clear modification of G41 within Escherichia coli cells can be seen by the appearance of an 11-mer which is not normally present in RNase T1 digests of unmodified 5S RNA. Reaction conditions can thus be optimized. Analysis of the accessibility of ribosomal 16S RNA within mutant and wild-type cells. The method has successfully been employed for a structural comparison of ribosomes within mutant and wild-type cells. Previous studies had shown that mutations within the leader region of ribosomal RNA operons cause functionally defective ribosomes (7). An analysis was performed with cells which had been transformed with plasmids containing wild-type (pSTLTO) or leader mutant (pSTLT11) rRNA operons, respectively. Modification was performed as described above. For comparison
242
NOTES & TIPS
REFERENCES 1. Christiansen, J., Egebjerg, J., Larsen, N., and Garrett, R. A. (1990). in Ribosomes and Protein Synthesis: A Practical Approach (G. Spedding, Ed.), pp. 229 –252. IRL Press/Oxford Univ. Press, London. 2. Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J.-P., and Ehresmann, B. (1987) Nucleic Acids Res. 22, 9109 –9128. 3. Shapiro, H., and Hachmann, J. (1996) Biochemistry 5, 2799 – 2807. 4. Noller, H. F., and Garrett, R. A. (1979) J. Mol. Biol. 132, 621– 636. 5. Go¨ringer, H. U., Bertram, S., and Wagner, R. (1984) J. Biol. Chem. 259, 491– 496. 6. Stark, M. J. R., Gourse, R. L., and Dahlberg, A. E. (1982) J. Mol. Biol. 159, 417– 439. 7. Theißen, G., Thelen, L., and Wagner, R. (1993) J. Mol. Biol. 233, 203–218. 8. Stern, S., Moazed, D., and Noller, H. F. (1988) Methods Enzymol. 164, 481– 489. 9. Moazed, D., Stern, S., and Noller, H. F. (1986) J. Mol. Biol. 187, 399 – 416. FIG. 2. Analysis of kethoxal reactive sites within 16S RNA in living cells. A primer extension analysis of the 59 16S RNA domain is shown. A, C, G, and T indicate sequencing lanes. Lanes showing in vivo modifications with kethoxal and DMS are indicated. Unmodified control lanes: CT0 and CT11. Analysis of RNA from cells expressing wild-type ribosomes (HB101/pSTLT0) or leader mutant ribosomes (HB101/pSTLT11) are indicated MT0 or MT11, respectively. Open arrows indicate DMS modified positions and closed arrows indicate sites of kethoxal reactivity, respectively. Modification positions coincide with previous determinations from in vitro structural probing studies (9).
a DMS modification was also performed according to (1). Ribosomal subunits were prepared from the extracted supernatant by centrifugation on 10 to 30% sucrose gradients (7) and 16S RNA was isolated by phenol extraction. Primer extension reactions were performed as described (8). An example of the analysis is shown in Fig. 2. Conclusions. The above method describes an efficient, easy, and convenient procedure for obtaining direct structural information on RNA or RNPs within living bacterial cells. It provides direct information on the accessibility of guanosines without laborious chemical cleavage of modified sites, and thereby extends and supplements existing methods for in vivo structural probing. We have successfully applied it for a structural comparison of ribosomes within wild-type and mutant cells, and it should be universally applicable without intensive adaptation to a wide variety of structural investigations. Acknowledgments. We thank Sabina Przibilla who tested the everyday lab fitness of the method. The work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie.
Elimination of Creatine Interference with the Indophenol Measurement of NH3 Produced during Nitrogenase Assays M. J. Dilworth1 and K. Fisher2 Department of Biochemistry, The Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061-0308 Received October 28, 1997
In the course of a study of azide reduction by the altered molybdenum iron (MoFe) protein in which histidine-195 in the a-subunit has been replaced by glutamine (1), we encountered a major problem with ammonia determination. This results from two factors: (i) an inherently very low activity of the altered protein towards azide, and (ii) an uncoupling by azide itself of ATP hydrolysis from substrate reduction. The combination of these two factors means that samples containing amounts of NH3 satisfactorily measurable with the indophenol method also contain large amounts of creatine (, 5 mmol) which produce almost complete inhibition of color development. Standard assays of N2 reduction by cell-free preparations (crude extracts or purified proteins) of nitrogenase require the presence of a reductant (usually so1
Permanent address: Center for Rhizobium Studies, School of Biological Sciences and Biotechnology, Division of Science, Murdoch University, Murdoch, Western Australia 6150. 2 To whom correspondence should be addressed. Fax: (540) 231 9070. E-mail: [email protected]. ANALYTICAL BIOCHEMISTRY 256, 242–244 (1998) ARTICLE NO. AB972517
0003-2697/98 $25.00 Copyright © 1998 by Academic Press All rights of reproduction in any form reserved.
NOTES & TIPS
addition of NaCl, KCl, glucose, or serum to the calibration buffer had much less effect. The difference between glass membrane and solid-state electrodes was unaffected by varying the concentration or pH of the compounds in standard calibration buffer. The solid-state electrode deviated more from the original calibration buffer pH value than the glass membrane electrode did when glucose or FBS was added to the calibration buffer and even more so with the addition of MgCl2 or CaCl2 (Table 2, columns 4 and 5). Although the solid-state system may be more sensitive to divalent cations than the glass membrane electrode, the glass membrane electrode was not completely impervious to competing ions. Although there was only a small pH difference (DpH) between measured values for glass and solid-state electrodes when NaCl or KCl was added to the standard calibration buffer, both types of electrodes exhibited a decrease in absolute pH of similar magnitude with increasing Na1 or K1 (data not shown), likely caused by pKa shifts in the acids/bases of the buffering system with increasing ionic strength. Since most physiologic buffers contain Mg21 and/or Ca21, care must be taken when selecting an electrode to measure pH, as well as the method of electrode calibration. Both types of electrodes were affected when the test solution differed greatly from the calibration buffer in salt concentration or ionic composition. The use of internal standards or matrix matching may increase accuracy of pH calibration and measurements. Nevertheless, a real discrepancy can arise when a laboratory switches to the new solid-state technology without realizing that the solid-state pH measurements in physiologic buffers are not directly comparable to the established glass membrane technology when standard calibration buffers are used. A DpH of 0.1– 0.2 between solid-state and glass membrane electrodes is significant when one considers the sensitivity of biological systems to small changes in pH within the physiologic pH range. However, this does not necessarily detract from the usefulness of the solid-state technology due to its ease of use and rapid response time. If a laboratory chooses to use a solid-state electrode for pH measurements of biological buffers, a simple conversion to glass electrode values will enable their data to be compared to the existing literature. Acknowledgments. The authors thank Ronald A. Coss, Ph.D., Dennis B. Leeper, Ph.D., Christopher Storck (Department of Radiation Oncology, Thomas Jefferson University, Philadelphia, PA), Michael D. O’Hara, Ph.D. (Cordis Institute, Johnson & Johnson, Co., Inc., Warren, NJ), and John Brekke (Sentron, Inc., Gig Harbor, WA) for their help and advice during this work. The authors also thank Andreas Woldegiorgic (Academy of Science, Stockholm, Sweden) for editorial help.
REFERENCES 1. Bergveld, P. (1972) IEEE Trans. Biomed. Eng. BME-19, 342– 351. 2. Sentron, Inc. Technical Documentation. 3. Kress-Rogers, E. (1991) Trends Food Sci. Technol., 320 –324. 4. Grynkiewicz, G., Poenie, M., and Tsien, R. Y. (1985) J. Biol. Chem. 260, 3440 –3450. 5. Thomas, J., Buchsbaum, R., Zimniak, A., and Racker, E. (1979) Biochemistry USA 18, 2210 –2218.
A Chemical Modification Method for the Structural Analysis of RNA and RNA±Protein Complexes within Living Cells Monika Balzer* and Rolf Wagner1 Institut fu¨r Physikalische Biologie, Heinrich-HeineUniversita¨t Du¨sseldorf, Universita¨tsstrasse 1, Du¨sseldorf, D-40225 Germany Received August 25, 1997
Studies on the structure and function of RNA and RNA–protein complexes (RNPs)2 are of increasing interest. Chemical modification and limited enzymatic hydrolysis (structural probing) are powerful methods for obtaining detailed information on the structure and dynamics of nucleic acids in solution (1, 2). Their use, however, is widely limited to studies outside the cellular compartments. The only reagent which has gained widespread use in the in vivo modification of RNA or DNA so far is dimethyl sulfate (DMS) which methylates N7 of guanosines, the N1 position of adenosine, and N3 of cytosines. Identification of modified sites is normally carried out by the rapid primer extension procedure. However, the N7-methylguanosine cannot readily be identified by primer extension, and more tedious procedures for a chemical cleavage must be employed. To circumvent the difficulty in obtaining structural information for guanosines, and to extend the number of methods for in vivo structural probing, we developed a simple chemical method for in vivo modification taking advantage of the reagent kethoxal (a-keto-b-ethoxybutyraldehyde). Kethoxal reacts with guanosines not involved in Watson–Crick base pairs in a highly specific way and under mild conditions by forming a five-membered ring between the two carbonyl groups and the N1 and N2 positions of guanosine (3). Kethoxal-modified guanosines are not accepted as 1 To whom correspondence should be addressed. Fax: 49 211 811 5167. E-mail: [email protected]. 2 Abbreviations used: RNPs, RNA–protein complexes; DMS, dimethyl sulfate.
ANALYTICAL BIOCHEMISTRY 256, 240 –242 (1998) ARTICLE NO. AB972499
0003-2697/98 $25.00 Copyright © 1998 by Academic Press All rights of reproduction in any form reserved.
NOTES & TIPS
template for the reverse transcriptase, and can thus readily be identified by primer extension. Kethoxal has not been described for in vivo structural probing previously. We have developed a simple method which enables a rapid and universally applicable analysis of accessible guanosines within nucleic acids or protein nucleic acid complexes in vivo. Tedious additional chemical treatments for the identification of the modified guanosines are avoided. Existing in vivo modification techniques like DMS probing are thus improved. In the following we describe an in vivo modification procedure which is straightforward, consisting of three steps: (i) increasing the permeability of bacterial cell walls for the rapid uptake of kethoxal, (ii) the modification reaction which can be performed for various times at all temperatures not detrimental to the cells, and (iii) RNA (or RNP) extraction and analysis of modified nucleotides by primer extension. Suitable conditions for the modification of ribosomal RNA. HB101 cells were grown in YT medium at 30°C to an optical density of 0.5 A600. Two 100-ml aliquots of the culture were taken and the cells were harvested by centrifugation (5 min, 5000 rpm). The pellets were resuspended carefully in 1 ml 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8) and incubated at 30°C with gentle shaking. After 15 min incubation 5 ml of 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8) was added to the control culture, while the modification culture was supplemented with 5 ml 0.1 M CaCl2, 20 mM sodium– borate (pH 6.8), 2% ethanol, presaturated with kethoxal. Incubation was continued for another 30 min at 30°C, after which the cells were harvested by centrifugation. Cell pellets were resuspended in 30 ml 25 mM Tris–HCl (pH 7.6), 60 mM KCl, 10 mM MgCl2, 20% (w/v) sucrose, 150 mg/ml lysozyme, and lysis was performed by repeated freeze–thawing (at least three times) as described (6). One hundred microliters of TKM buffer (25 mM Tris–HCl (pH 7.6), 60 mM KCl, 5 mM MgCl2), 34 ml 1% sodium– deoxycholate (w/v), 20 ml of a 5% (w/v) polyoxyethylene ether (20 cetyl ether, Brij 58), 8 ml 20 mM sodium– borate (pH 6.8), and 15 units DNase I (RNase-free) were added to the disrupted cells and incubated for 15 min on ice. The cell debris is removed by centrifugation, and the cleared supernatant can be used for RNA extraction or, as in this case, as a source for ribosome preparation. Demonstration of efficient modification of cellular RNA components. First, we established conditions under which a rapid kethoxal uptake and reaction could be obtained. As a marker we followed the guanosine modification of ribosomal 5S RNA within the cells. The kethoxal modification of 5S RNA has been carefully analyzed previously (4, 5), and G41 had been established as a position of outstanding reactivity within the complete ribosome. The kethoxal
241
FIG. 1. Analysis of 5S RNA after in vivo modification with kethoxal. The separation of 59-labeled oligonucleotides from a total RNase T1 digestion of 5S RNA extracted from in vivo kethoxalmodified cells on a 20% acrylamide/8 M urea gel is shown. (1) RNase T1 digestion after modification at G41 leads to an 11-mer (indicated by the arrow) which is not normally present in T1 digests of unmodified 5S RNA. (2) Adjustment of the oligonucleotides to pH 9 with NH4OH and incubation at 37°C for 60 min followed by a second RNase T1 digestion causes the 11-mer to disappear almost totally.
modification can be easily analyzed because modified guanosines are not recognized by RNase T1 and therefore give rise to new products after a complete RNase T1 hydrolysis (10 units, 60 min, 37°C). Because the kethoxal modification is reversible under alkaline conditions, redigestion after a mild base treatment (1 ml 25% NH4OH, 60 min, 37°C) yields the normal RNase T1 products again. Figure 1 shows an example of a pilot experiment where a clear modification of G41 within Escherichia coli cells can be seen by the appearance of an 11-mer which is not normally present in RNase T1 digests of unmodified 5S RNA. Reaction conditions can thus be optimized. Analysis of the accessibility of ribosomal 16S RNA within mutant and wild-type cells. The method has successfully been employed for a structural comparison of ribosomes within mutant and wild-type cells. Previous studies had shown that mutations within the leader region of ribosomal RNA operons cause functionally defective ribosomes (7). An analysis was performed with cells which had been transformed with plasmids containing wild-type (pSTLTO) or leader mutant (pSTLT11) rRNA operons, respectively. Modification was performed as described above. For comparison
242
NOTES & TIPS
REFERENCES 1. Christiansen, J., Egebjerg, J., Larsen, N., and Garrett, R. A. (1990). in Ribosomes and Protein Synthesis: A Practical Approach (G. Spedding, Ed.), pp. 229 –252. IRL Press/Oxford Univ. Press, London. 2. Ehresmann, C., Baudin, F., Mougel, M., Romby, P., Ebel, J.-P., and Ehresmann, B. (1987) Nucleic Acids Res. 22, 9109 –9128. 3. Shapiro, H., and Hachmann, J. (1996) Biochemistry 5, 2799 – 2807. 4. Noller, H. F., and Garrett, R. A. (1979) J. Mol. Biol. 132, 621– 636. 5. Go¨ringer, H. U., Bertram, S., and Wagner, R. (1984) J. Biol. Chem. 259, 491– 496. 6. Stark, M. J. R., Gourse, R. L., and Dahlberg, A. E. (1982) J. Mol. Biol. 159, 417– 439. 7. Theißen, G., Thelen, L., and Wagner, R. (1993) J. Mol. Biol. 233, 203–218. 8. Stern, S., Moazed, D., and Noller, H. F. (1988) Methods Enzymol. 164, 481– 489. 9. Moazed, D., Stern, S., and Noller, H. F. (1986) J. Mol. Biol. 187, 399 – 416. FIG. 2. Analysis of kethoxal reactive sites within 16S RNA in living cells. A primer extension analysis of the 59 16S RNA domain is shown. A, C, G, and T indicate sequencing lanes. Lanes showing in vivo modifications with kethoxal and DMS are indicated. Unmodified control lanes: CT0 and CT11. Analysis of RNA from cells expressing wild-type ribosomes (HB101/pSTLT0) or leader mutant ribosomes (HB101/pSTLT11) are indicated MT0 or MT11, respectively. Open arrows indicate DMS modified positions and closed arrows indicate sites of kethoxal reactivity, respectively. Modification positions coincide with previous determinations from in vitro structural probing studies (9).
a DMS modification was also performed according to (1). Ribosomal subunits were prepared from the extracted supernatant by centrifugation on 10 to 30% sucrose gradients (7) and 16S RNA was isolated by phenol extraction. Primer extension reactions were performed as described (8). An example of the analysis is shown in Fig. 2. Conclusions. The above method describes an efficient, easy, and convenient procedure for obtaining direct structural information on RNA or RNPs within living bacterial cells. It provides direct information on the accessibility of guanosines without laborious chemical cleavage of modified sites, and thereby extends and supplements existing methods for in vivo structural probing. We have successfully applied it for a structural comparison of ribosomes within wild-type and mutant cells, and it should be universally applicable without intensive adaptation to a wide variety of structural investigations. Acknowledgments. We thank Sabina Przibilla who tested the everyday lab fitness of the method. The work was supported by the Deutsche Forschungsgemeinschaft and the Fonds der Chemischen Industrie.
Elimination of Creatine Interference with the Indophenol Measurement of NH3 Produced during Nitrogenase Assays M. J. Dilworth1 and K. Fisher2 Department of Biochemistry, The Virginia Polytechnic Institute and State University, Blacksburg, Virginia 24061-0308 Received October 28, 1997
In the course of a study of azide reduction by the altered molybdenum iron (MoFe) protein in which histidine-195 in the a-subunit has been replaced by glutamine (1), we encountered a major problem with ammonia determination. This results from two factors: (i) an inherently very low activity of the altered protein towards azide, and (ii) an uncoupling by azide itself of ATP hydrolysis from substrate reduction. The combination of these two factors means that samples containing amounts of NH3 satisfactorily measurable with the indophenol method also contain large amounts of creatine (, 5 mmol) which produce almost complete inhibition of color development. Standard assays of N2 reduction by cell-free preparations (crude extracts or purified proteins) of nitrogenase require the presence of a reductant (usually so1
Permanent address: Center for Rhizobium Studies, School of Biological Sciences and Biotechnology, Division of Science, Murdoch University, Murdoch, Western Australia 6150. 2 To whom correspondence should be addressed. Fax: (540) 231 9070. E-mail: [email protected]. ANALYTICAL BIOCHEMISTRY 256, 242–244 (1998) ARTICLE NO. AB972517
0003-2697/98 $25.00 Copyright © 1998 by Academic Press All rights of reproduction in any form reserved.