JOURNAL OF CLINICAL MICROBIOLOGY, Apr. 1992, p. 1030-1032
Vol. 30, No. 4
0095-1137/92/041030-03$02.00/0
Copyright © 1992, American Society for Microbiology
Distinguishing between Respiratory Syncytial Virus Subgroups by Protein Profile Analysis PRAMILA WALPITA,lt* MAURICE A. MUFSON,2 RONALD J. STANEK,2 DEBORAH PFEIFER,' AND JAMES D. CONNOR' Department of Pediatrics, University of California-San Diego, La Jolla, California 92093,1 and Department of Medicine and Veterans Affairs Medical Center, Marshall University School of Medicine, Huntington, West Virginia 257012 Received 3 July 1991/Accepted 6 January 1992
We subgrouped 75 strains of respiratory syncytial virus by a protein profile method (PPM) which relies on different mobilities of the phosphoprotein in one-dimensional polyacrylamide gel electrophoresis and does not require monoclonal antibodies. When compared with enzyme immunoassay, PPM correctly subgrouped 54 of 56 subgroup A and all 19 subgroup B strains.
Infection with respiratory syncytial virus (RSV) is a leading cause of hospital admission among infants and children with bronchiolitis and pneumonia and of morbidity in the first year of life (4). Monoclonal antibodies (MAbs) generated against RSV distinguish two stable subgroups, A and B, on the basis of epitope variations of the two envelope proteins, the large glycoprotein (G) and the fusion protein (F) (2, 15). The importance of subgroup-specific immunity in RSV infection has been demonstrated in several recent reports that describe various aspects of RSV disease in children and in experimental animal models (9, 12, 14, 20). MAb-based enzyme immunoassay (EIA) and immunofluorescence remain the only methods used routinely for subgrouping of RSV strains. MAbs to RSV structural proteins are not widely available, and the results of both methods depend in part on the numbers and types of antibodies employed (2, 14). Recently, DNA probes have been used to provide an accurate identification of the two subgroups (23), but very few laboratories are currently geared to using this technology. In this study, we distinguished clinical strains of subgroups A and B of RSV by a protein profile method (PPM) which relies on differing mobilities of the phosphoproteins (P) of the two subgroups in one-dimensional sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE). This method provides a rapid means of subgrouping large numbers of RSV strains without using RSV-specific MAbs. It has an overall accuracy of 97%. Viruses. Seventy-five field strains of RSV recovered during two epidemic seasons (1986 to 1987 and 1987 to 1988) constituted the group of RSV strains for subgroup classification by two methods, PPM and EIA. Two prototype strains of RSV, Long (subgroup A) and CH18537 (subgroup B), were included in all tests. RSV strains were propagated in HEp-2 cells. The EIA procedure has been described previously (15). Each field strain was subgrouped by using 31 MAbs generated to the subgroup A prototype strain (Long) and 8 MAbs generated to the subgroup B strain (WV4843) (1, 15). PPM. The PPM for viruses has been described previously
(22). Briefly, HEp-2 cell monolayers were grown in 24-well microplates (Nunc Inc., Naperville, Ill.). Each well was inoculated with 0.2 ml of virus, and the plates were incubated at 37°C for 24 to 48 h. At the end of this period, the medium was removed and replaced with 1 ml of methioninefree medium containing 0.15 M NaCl to initiate suppression of host protein synthesis and deplete methionine in the cells. [35S]methionine (35S Translabel; ICN Radiochemicals) was added (50 LCi per well), and the plates were incubated for 4 h at 37°C. Then the medium was removed completely by suction, 150 ,ul of SDS lysis buffer was added to each well, and the plate was covered and held at room temperature for 2 min. The lysate was transferred to appropriately labeled vials, heated at 100°C for 2 min, thoroughly mixed, and stored at -70°C until used for electrophoresis. The proteins were separated by SDS-PAGE on an automated electrophoresis system (AMBIS Systems, San Diego, Calif.). Three molecular weight (MW) standard lanes and the two prototype strains were included on each gel. The dried gels were scanned by using a beta scanning system, which provided an image resembling that obtained by autoradiography. Each gel lane was delineated, and the protein fingerprint was converted to a digital lane profile (10, 22). A computer program (Dendrogram) compared protein profiles of each of the clinical and prototype strains in the data base by employing the Pearson correlation coefficient (10, 18). Correlation among strains of the same subgroup was r > 0.6; correlation between subgroups was r < 0.2. Autoradiographs of the gels were also developed for visual subgroup categorization of the field strains by matching their protein profiles with those of the prototype strains. Three [35S]methionine-labeled RSV proteins, NP, P, and M, were seen clearly at 39,000 (NP), 32,000 to 36,000 (P), and 29,000 (M) (Fig. 1). An additional band at 26,000 was seen also but not clearly and consistently. The virus-specific nature of these proteins was confirmed in our previous study (22). HEp-2 proteins did not interfere with clear visualization of viral proteins because host proteins were selectively suppressed with an optimum concentration (0.15 M) of NaCl (22). All clinical strains showed mobilities essentially similar to those of the NP (39,000) and M (29,000) proteins, irrespective of subgroup. P mobility. P-protein mobility differences among 75 clinical strains were analyzed in relation to the subgroup categorization of the strains by EIA. Fifty-six of 75 RSV strains
* Corresponding author. t Present address: Pathology Department, Children's Hospital-
San Diego, 8001 Frost St., San Diego, CA 92123.
1030
VOL. 30, 1992 1
2
NOTES 3
4
5 6 7 8 9 10 11 12 13 14
1031
TABLE 1. Seasonal distribution and correlation between RSV subgroups as determined by EIA and PPM Subgroup determined by:
No. of RSV strainsa
PPMb
EIA
1986-1987
1987-1988
A B B
A A B
27 2 2
27 0 17
-i -w+.
Ulf
a Seventy-five clinical strains were tested. hPPM subgroups were assigned by comparing P-protein mobilities of the clinical strains with those of prototype strains of subgroup A (Long) and subgroup B (CH18537). PPM accuracy = 97%. -__ ____
-__
^
FIG. 1. P-protein mobility variation between prototype strains of subgroup A (Long strain, lane 8) and subgroup B (CH18537 strain, lane 9) and within subgroup B clinical strains (lanes 4 and 12). Mobility differences within subgroup A strains were not obvious (lanes 2, 3, 5, 10, 11, and 13). Lane 7 shows uninfected HEp-2 cells. Lanes 1, 6, and 14 are identical MW marker lanes ranging from 14,000 to 200,000.
subgroup A, and 19 were subgroup B. Sixteen of the subgroup B strains were further categorized in EIA with MAbs to the Gl and G2 epitopes of subgroup B, and they all exhibited Bi specificity (1). Fifty-four of 56 subgroup A strains matched the P-protein profile of Long strain by visual examination and by computerized data analysis (10, 18). Minimal variation was observed in the mobility of the P protein among these subgroup A strains (Fig. 1). The remaining 2 of the 56 subgroup A strains had P proteins with smaller MW that were unlike those of subgroup A but similar to that of subgroup B strain CH18537. The 19 subgroup B strains were not homogeneous for P-protein mobility; only 1 had a P-protein MW close to that of the prototype strain, CH18537 (MW 32,500). However, they all exhibited P-protein mobilities distinct from that of Long strain, and all were correctly assigned to the B subgroup on that basis (Fig. 1). The relation of EIA and PPM phenotypes and their distributions during the 1987 to 1988 and 1988 to 1989 epidemic seasons were
are
shown in Table 1.
Usually, one-dimensional separation by SDS-PAGE of whole-cell lysates of virus-infected cells produces numerous host cell-specific protein bands which obscure viral protein bands. We used a simple procedure of differential chemical suppression to eliminate host proteins, so that viral proteins were clearly visualized by PAGE (5, 22). Hypertonic stress results in rapid but reversible selective inhibition of host protein synthesis (18). We used a salt concentration which effectively suppressed most of the host protein synthesis but
exerted little effect on viral protein synthesis (5, 22). Thus, PPM distinguished between subgroup A and B strains with about 97% accuracy. The marked homogeneity of P-protein mobility among subgroup A strains facilitated ready identification of 54 of 56 strains. The subgroup B strains had P-protein mobilities which were heterogeneous with respect to that of the prototype subgroup B strain CH18537. However, they all had protein profiles which were quite distinct from that of the Long strain. Heterogeneity of the P protein of subgroup B has been reported previously (1, 7, 8, 13). Subgroup B prototype CH18537 and some other RSV strains also isolated in the 1960s and early 1970s possess comparatively smaller P proteins (1, 13). These strains, although they appear infrequently, seem to be still in circulation; one of our strains from the 1986 to 1987 season had such a small MW for the P protein (32,000) and matched the P protein MW of CH18537. Variation in the mobilities of the P proteins of the subgroups of RSV has been described elsewhere, although the molecular nature of such variation remains unexplained (1, 13). A comparison of the few published P-gene sequences of RSV subgroup A strains, of the single subgroup B strain (CH18537), and of our own unpublished P-gene sequences of two recent subgroup B field strains (21a) confirms the view that sequence diversity fails to explain the variation in P-protein mobility (11). Also, the results of analysis of the phosphoprotein of other negative-strain viruses, for example, vesicular stomatitis virus, suggest that the degree of phosphorylation does not explain the variation in P-protein mobility (3). Acidic domains have been suggested as the cause of conformational changes resulting in mobility variations of some RNA viral phosphoproteins (16). However, such domains are not obvious in the published (and our unpublished) P-gene sequence data for RSV. Since envelope proteins play an important role in receptor binding, neutralization, and immunomodulation, subgrouping of RSV on the basis of antigenic variation of the G and F proteins may be more direct than subgrouping on the basis of antigenic differences of other proteins (19, 23). However, the results of our study suggest that the physical properties of the P protein may be used as proxy for identifying antigenic differences of RSV when the aim is to distinguish clinical strains of subgroups A and B. PPM should facilitate the undertaking of large-scale epidemiologic studies of RSV disease. This work was supported in part from a grant awarded to Pramila Walpita by the American Lung Association of California. We are grateful for the expert technical assistance of Charisse Davidson and Dianne Holland.
1032
NOTES
REFERENCES 1. Akerlind, B., E. Norrby, C. Orvell, and M. A. Mufson. 1988. Respiratory syncytial virus: heterogeneity of subgroup B strains. J. Gen. Virol. 69:2145-2154. 2. Anderson, L. J., J. C. Hierholzer, C. Tsou, R. M. Hendry, B. F. Fernie, Y. Stone, and K. McIntosh. 1985. Antigenic characterization of respiratory syncytial virus strains with monoclonal antibodies. J. Infect. Dis. 151:626-633. 3. Barik, S., and A. K. Banerjee. 1991. Cloning and expression of the vesicular stomatitis virus phosphoprotein gene in Escherichia coli: analysis of phosphorylation status versus transcriptional activity. J. Virol. 65:1719-1726. 4. Belshe, R. B., and M. A. Mufson. 1991. Respiratory syncytial virus, p. 388-407. In R. B. Belshe (ed.), Textbook of human virology. Mosby-Year Book, Inc., St. Louis. 5. Bernstein, J. M., and J. Hruska. 1981. Respiratory syncytial virus proteins: identification by immunoprecipitation. J. Virol. 38:278-285. 6. Cristina, J., J. A. Lopez, C. Albo, B. Garcia-Barreno, B. J. Garcia, J. A. Melero, and A. Portela. 1989. Analysis of genetic variability in human respiratory syncytial virus by the RNase A mismatch cleavage method: subtype divergence and heterogeneity. Virology 174:126-134. 7. Gimenez, H. B., P. Cash, and W. T. Melvin. 1984. Monoclonal antibodies to human respiratory syncytial virus and their use in comparison of different viral isolates. J. Gen. Virol. 65:963-971. 8. Gimenez, H. B., N. Hardman, H. M. Keir, and P. Cash. 1986. Antigenic variation between human respiratory syncytial virus isolates. J. Virol. 67:863-870. 9. Hendry, R. M., J. C. Burns, E. E. Walsh, B. S. Graham, P. F. Wright, V. G. Hemming, W. J. Rodriquez, H. W. Kim, A. P. Gregory, K. McIntosh, R. M. Chanock, and B. R. Murphy. 1988. Strain-specific serum antibody responses in infants undergoing primary infection with respiratory syncytial virus. J. Infect. Dis.
1570:640-649. 10. Hook, L. A., P. L. Block, R. W. Kohlenberger, and P. A. Kinningham. 1987. Automated microbial identification system for computer programmed analysis of radio-labeled protein banding patterns. Developments in industrial microbiology. J. Ind. Microbiol. 28(Suppl. 2):149-160. 11. Johnson, P. R., and P. Collins. 1990. Sequence comparison of phosphoprotein in RNAs of antigenic subgroups A and B of human respiratory syncytial virus identified a highly conserved domain in the predicted protein. J. Gen. Virol. 71:481-485. 12. Johnson, P. R., R. A. Olmsted, G. A. Prince, B. R. Murphy,
J. CLIN. MICROBIOL.
D. W. Ailing, E. E. Walsh, and P. L. Collins. 1987. Antigenic relatedness between glycoproteins of human respiratory syncytial virus subgroups A and B: evaluation of the contributions of F and G glycoproteins to immunity. J. Virol. 61:3163-3166. 13. Morgan, L. A., F. G. Routledge, M. M. Willcocks, A. C. Samson, R. Scott, and G. L. Toms. 1987. Strain variation of respiratory syncytial virus. J. Gen. Virol. 68:2781-2788. 14. Mufson, M. A., R. B. Belshe, C. Orvell, and E. Norrby. 1987. Subgroup characteristics of respiratory syncytial virus strains recovered from children with two consecutive infections. J. Clin. Microbiol. 25:1535-1539. 15. Mufson, M. A., C. Orvell, B. Rafnar, and E. Norrby. 1985. Two distinct subtypes of human respiratory syncytial virus. J. Gen. Virol. 66:2111-2124. 16. Pakacs, A. M., K. G. Perrine, S. Barik, and A. K. Banerjee. 1991. Alteration of specific amino acid residues in the acidic domain I of vesicular stomatitis virus phosphoprotein (P) converts GLA 4-P (I) hybrid into a transcriptional activator. New Biol. 3:581-591. 17. Sneath, H. H., and R. R. Sokal. 1973. Numerical taxonomy: the principles and practice of numerical classification. The W. H. Freeman Co., San Francisco. 18. Storch, G. A., C. S. Park, and D. E. Dohner. 1989. RNA fingerprinting of respiratory syncytial virus using ribonuclease protection: application to molecular epidemiology. J. Clin. Invest. 83:1894-1902. 19. Stott, E. J., and G. Taylor. 1985. Respiratory syncytial virus brief review. Arch. Virol. 84:1-52. 20. Stott, E. J., G. Taylor, L. A. Ball, K. Anderson, K. K. Y. Young, A. M. Q. King, and G. W. Wertz. 1987. Immune and histopathological responses in animals vaccinated with recombinant vaccinia viruses that express individual genes of human respiratory syncytial virus. J. Virol. 61:3855-3861. 21. Sullender, W. M., L. J. Anderson, K. Anderson, and G. W. Wertz. 1990. Differentiation of respiratory syncytial virus subgroups with cDNA probes in a nucleic acid hybridization assay. J. Clin. Microbiol. 28:1683-1687. 21a.Walpita, P., and S. Barik. Personal communication. 22. Walpita, P., J. D. Connor, and D. Pfeifer. 1989. Protein fingerprinting: a novel virus identification system. J. Virol. Methods 25:315-324. 23. Walsh, E. E., and J. Hruska. 1983. Monoclonal antibodies to respiratory syncytial virus proteins: identification of the fusion protein. J. Virol. 1983:171-177.
Vol. 30, No. 4
0095-1137/92/041030-03$02.00/0
Copyright © 1992, American Society for Microbiology
Distinguishing between Respiratory Syncytial Virus Subgroups by Protein Profile Analysis PRAMILA WALPITA,lt* MAURICE A. MUFSON,2 RONALD J. STANEK,2 DEBORAH PFEIFER,' AND JAMES D. CONNOR' Department of Pediatrics, University of California-San Diego, La Jolla, California 92093,1 and Department of Medicine and Veterans Affairs Medical Center, Marshall University School of Medicine, Huntington, West Virginia 257012 Received 3 July 1991/Accepted 6 January 1992
We subgrouped 75 strains of respiratory syncytial virus by a protein profile method (PPM) which relies on different mobilities of the phosphoprotein in one-dimensional polyacrylamide gel electrophoresis and does not require monoclonal antibodies. When compared with enzyme immunoassay, PPM correctly subgrouped 54 of 56 subgroup A and all 19 subgroup B strains.
Infection with respiratory syncytial virus (RSV) is a leading cause of hospital admission among infants and children with bronchiolitis and pneumonia and of morbidity in the first year of life (4). Monoclonal antibodies (MAbs) generated against RSV distinguish two stable subgroups, A and B, on the basis of epitope variations of the two envelope proteins, the large glycoprotein (G) and the fusion protein (F) (2, 15). The importance of subgroup-specific immunity in RSV infection has been demonstrated in several recent reports that describe various aspects of RSV disease in children and in experimental animal models (9, 12, 14, 20). MAb-based enzyme immunoassay (EIA) and immunofluorescence remain the only methods used routinely for subgrouping of RSV strains. MAbs to RSV structural proteins are not widely available, and the results of both methods depend in part on the numbers and types of antibodies employed (2, 14). Recently, DNA probes have been used to provide an accurate identification of the two subgroups (23), but very few laboratories are currently geared to using this technology. In this study, we distinguished clinical strains of subgroups A and B of RSV by a protein profile method (PPM) which relies on differing mobilities of the phosphoproteins (P) of the two subgroups in one-dimensional sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE). This method provides a rapid means of subgrouping large numbers of RSV strains without using RSV-specific MAbs. It has an overall accuracy of 97%. Viruses. Seventy-five field strains of RSV recovered during two epidemic seasons (1986 to 1987 and 1987 to 1988) constituted the group of RSV strains for subgroup classification by two methods, PPM and EIA. Two prototype strains of RSV, Long (subgroup A) and CH18537 (subgroup B), were included in all tests. RSV strains were propagated in HEp-2 cells. The EIA procedure has been described previously (15). Each field strain was subgrouped by using 31 MAbs generated to the subgroup A prototype strain (Long) and 8 MAbs generated to the subgroup B strain (WV4843) (1, 15). PPM. The PPM for viruses has been described previously
(22). Briefly, HEp-2 cell monolayers were grown in 24-well microplates (Nunc Inc., Naperville, Ill.). Each well was inoculated with 0.2 ml of virus, and the plates were incubated at 37°C for 24 to 48 h. At the end of this period, the medium was removed and replaced with 1 ml of methioninefree medium containing 0.15 M NaCl to initiate suppression of host protein synthesis and deplete methionine in the cells. [35S]methionine (35S Translabel; ICN Radiochemicals) was added (50 LCi per well), and the plates were incubated for 4 h at 37°C. Then the medium was removed completely by suction, 150 ,ul of SDS lysis buffer was added to each well, and the plate was covered and held at room temperature for 2 min. The lysate was transferred to appropriately labeled vials, heated at 100°C for 2 min, thoroughly mixed, and stored at -70°C until used for electrophoresis. The proteins were separated by SDS-PAGE on an automated electrophoresis system (AMBIS Systems, San Diego, Calif.). Three molecular weight (MW) standard lanes and the two prototype strains were included on each gel. The dried gels were scanned by using a beta scanning system, which provided an image resembling that obtained by autoradiography. Each gel lane was delineated, and the protein fingerprint was converted to a digital lane profile (10, 22). A computer program (Dendrogram) compared protein profiles of each of the clinical and prototype strains in the data base by employing the Pearson correlation coefficient (10, 18). Correlation among strains of the same subgroup was r > 0.6; correlation between subgroups was r < 0.2. Autoradiographs of the gels were also developed for visual subgroup categorization of the field strains by matching their protein profiles with those of the prototype strains. Three [35S]methionine-labeled RSV proteins, NP, P, and M, were seen clearly at 39,000 (NP), 32,000 to 36,000 (P), and 29,000 (M) (Fig. 1). An additional band at 26,000 was seen also but not clearly and consistently. The virus-specific nature of these proteins was confirmed in our previous study (22). HEp-2 proteins did not interfere with clear visualization of viral proteins because host proteins were selectively suppressed with an optimum concentration (0.15 M) of NaCl (22). All clinical strains showed mobilities essentially similar to those of the NP (39,000) and M (29,000) proteins, irrespective of subgroup. P mobility. P-protein mobility differences among 75 clinical strains were analyzed in relation to the subgroup categorization of the strains by EIA. Fifty-six of 75 RSV strains
* Corresponding author. t Present address: Pathology Department, Children's Hospital-
San Diego, 8001 Frost St., San Diego, CA 92123.
1030
VOL. 30, 1992 1
2
NOTES 3
4
5 6 7 8 9 10 11 12 13 14
1031
TABLE 1. Seasonal distribution and correlation between RSV subgroups as determined by EIA and PPM Subgroup determined by:
No. of RSV strainsa
PPMb
EIA
1986-1987
1987-1988
A B B
A A B
27 2 2
27 0 17
-i -w+.
Ulf
a Seventy-five clinical strains were tested. hPPM subgroups were assigned by comparing P-protein mobilities of the clinical strains with those of prototype strains of subgroup A (Long) and subgroup B (CH18537). PPM accuracy = 97%. -__ ____
-__
^
FIG. 1. P-protein mobility variation between prototype strains of subgroup A (Long strain, lane 8) and subgroup B (CH18537 strain, lane 9) and within subgroup B clinical strains (lanes 4 and 12). Mobility differences within subgroup A strains were not obvious (lanes 2, 3, 5, 10, 11, and 13). Lane 7 shows uninfected HEp-2 cells. Lanes 1, 6, and 14 are identical MW marker lanes ranging from 14,000 to 200,000.
subgroup A, and 19 were subgroup B. Sixteen of the subgroup B strains were further categorized in EIA with MAbs to the Gl and G2 epitopes of subgroup B, and they all exhibited Bi specificity (1). Fifty-four of 56 subgroup A strains matched the P-protein profile of Long strain by visual examination and by computerized data analysis (10, 18). Minimal variation was observed in the mobility of the P protein among these subgroup A strains (Fig. 1). The remaining 2 of the 56 subgroup A strains had P proteins with smaller MW that were unlike those of subgroup A but similar to that of subgroup B strain CH18537. The 19 subgroup B strains were not homogeneous for P-protein mobility; only 1 had a P-protein MW close to that of the prototype strain, CH18537 (MW 32,500). However, they all exhibited P-protein mobilities distinct from that of Long strain, and all were correctly assigned to the B subgroup on that basis (Fig. 1). The relation of EIA and PPM phenotypes and their distributions during the 1987 to 1988 and 1988 to 1989 epidemic seasons were
are
shown in Table 1.
Usually, one-dimensional separation by SDS-PAGE of whole-cell lysates of virus-infected cells produces numerous host cell-specific protein bands which obscure viral protein bands. We used a simple procedure of differential chemical suppression to eliminate host proteins, so that viral proteins were clearly visualized by PAGE (5, 22). Hypertonic stress results in rapid but reversible selective inhibition of host protein synthesis (18). We used a salt concentration which effectively suppressed most of the host protein synthesis but
exerted little effect on viral protein synthesis (5, 22). Thus, PPM distinguished between subgroup A and B strains with about 97% accuracy. The marked homogeneity of P-protein mobility among subgroup A strains facilitated ready identification of 54 of 56 strains. The subgroup B strains had P-protein mobilities which were heterogeneous with respect to that of the prototype subgroup B strain CH18537. However, they all had protein profiles which were quite distinct from that of the Long strain. Heterogeneity of the P protein of subgroup B has been reported previously (1, 7, 8, 13). Subgroup B prototype CH18537 and some other RSV strains also isolated in the 1960s and early 1970s possess comparatively smaller P proteins (1, 13). These strains, although they appear infrequently, seem to be still in circulation; one of our strains from the 1986 to 1987 season had such a small MW for the P protein (32,000) and matched the P protein MW of CH18537. Variation in the mobilities of the P proteins of the subgroups of RSV has been described elsewhere, although the molecular nature of such variation remains unexplained (1, 13). A comparison of the few published P-gene sequences of RSV subgroup A strains, of the single subgroup B strain (CH18537), and of our own unpublished P-gene sequences of two recent subgroup B field strains (21a) confirms the view that sequence diversity fails to explain the variation in P-protein mobility (11). Also, the results of analysis of the phosphoprotein of other negative-strain viruses, for example, vesicular stomatitis virus, suggest that the degree of phosphorylation does not explain the variation in P-protein mobility (3). Acidic domains have been suggested as the cause of conformational changes resulting in mobility variations of some RNA viral phosphoproteins (16). However, such domains are not obvious in the published (and our unpublished) P-gene sequence data for RSV. Since envelope proteins play an important role in receptor binding, neutralization, and immunomodulation, subgrouping of RSV on the basis of antigenic variation of the G and F proteins may be more direct than subgrouping on the basis of antigenic differences of other proteins (19, 23). However, the results of our study suggest that the physical properties of the P protein may be used as proxy for identifying antigenic differences of RSV when the aim is to distinguish clinical strains of subgroups A and B. PPM should facilitate the undertaking of large-scale epidemiologic studies of RSV disease. This work was supported in part from a grant awarded to Pramila Walpita by the American Lung Association of California. We are grateful for the expert technical assistance of Charisse Davidson and Dianne Holland.
1032
NOTES
REFERENCES 1. Akerlind, B., E. Norrby, C. Orvell, and M. A. Mufson. 1988. Respiratory syncytial virus: heterogeneity of subgroup B strains. J. Gen. Virol. 69:2145-2154. 2. Anderson, L. J., J. C. Hierholzer, C. Tsou, R. M. Hendry, B. F. Fernie, Y. Stone, and K. McIntosh. 1985. Antigenic characterization of respiratory syncytial virus strains with monoclonal antibodies. J. Infect. Dis. 151:626-633. 3. Barik, S., and A. K. Banerjee. 1991. Cloning and expression of the vesicular stomatitis virus phosphoprotein gene in Escherichia coli: analysis of phosphorylation status versus transcriptional activity. J. Virol. 65:1719-1726. 4. Belshe, R. B., and M. A. Mufson. 1991. Respiratory syncytial virus, p. 388-407. In R. B. Belshe (ed.), Textbook of human virology. Mosby-Year Book, Inc., St. Louis. 5. Bernstein, J. M., and J. Hruska. 1981. Respiratory syncytial virus proteins: identification by immunoprecipitation. J. Virol. 38:278-285. 6. Cristina, J., J. A. Lopez, C. Albo, B. Garcia-Barreno, B. J. Garcia, J. A. Melero, and A. Portela. 1989. Analysis of genetic variability in human respiratory syncytial virus by the RNase A mismatch cleavage method: subtype divergence and heterogeneity. Virology 174:126-134. 7. Gimenez, H. B., P. Cash, and W. T. Melvin. 1984. Monoclonal antibodies to human respiratory syncytial virus and their use in comparison of different viral isolates. J. Gen. Virol. 65:963-971. 8. Gimenez, H. B., N. Hardman, H. M. Keir, and P. Cash. 1986. Antigenic variation between human respiratory syncytial virus isolates. J. Virol. 67:863-870. 9. Hendry, R. M., J. C. Burns, E. E. Walsh, B. S. Graham, P. F. Wright, V. G. Hemming, W. J. Rodriquez, H. W. Kim, A. P. Gregory, K. McIntosh, R. M. Chanock, and B. R. Murphy. 1988. Strain-specific serum antibody responses in infants undergoing primary infection with respiratory syncytial virus. J. Infect. Dis.
1570:640-649. 10. Hook, L. A., P. L. Block, R. W. Kohlenberger, and P. A. Kinningham. 1987. Automated microbial identification system for computer programmed analysis of radio-labeled protein banding patterns. Developments in industrial microbiology. J. Ind. Microbiol. 28(Suppl. 2):149-160. 11. Johnson, P. R., and P. Collins. 1990. Sequence comparison of phosphoprotein in RNAs of antigenic subgroups A and B of human respiratory syncytial virus identified a highly conserved domain in the predicted protein. J. Gen. Virol. 71:481-485. 12. Johnson, P. R., R. A. Olmsted, G. A. Prince, B. R. Murphy,
J. CLIN. MICROBIOL.
D. W. Ailing, E. E. Walsh, and P. L. Collins. 1987. Antigenic relatedness between glycoproteins of human respiratory syncytial virus subgroups A and B: evaluation of the contributions of F and G glycoproteins to immunity. J. Virol. 61:3163-3166. 13. Morgan, L. A., F. G. Routledge, M. M. Willcocks, A. C. Samson, R. Scott, and G. L. Toms. 1987. Strain variation of respiratory syncytial virus. J. Gen. Virol. 68:2781-2788. 14. Mufson, M. A., R. B. Belshe, C. Orvell, and E. Norrby. 1987. Subgroup characteristics of respiratory syncytial virus strains recovered from children with two consecutive infections. J. Clin. Microbiol. 25:1535-1539. 15. Mufson, M. A., C. Orvell, B. Rafnar, and E. Norrby. 1985. Two distinct subtypes of human respiratory syncytial virus. J. Gen. Virol. 66:2111-2124. 16. Pakacs, A. M., K. G. Perrine, S. Barik, and A. K. Banerjee. 1991. Alteration of specific amino acid residues in the acidic domain I of vesicular stomatitis virus phosphoprotein (P) converts GLA 4-P (I) hybrid into a transcriptional activator. New Biol. 3:581-591. 17. Sneath, H. H., and R. R. Sokal. 1973. Numerical taxonomy: the principles and practice of numerical classification. The W. H. Freeman Co., San Francisco. 18. Storch, G. A., C. S. Park, and D. E. Dohner. 1989. RNA fingerprinting of respiratory syncytial virus using ribonuclease protection: application to molecular epidemiology. J. Clin. Invest. 83:1894-1902. 19. Stott, E. J., and G. Taylor. 1985. Respiratory syncytial virus brief review. Arch. Virol. 84:1-52. 20. Stott, E. J., G. Taylor, L. A. Ball, K. Anderson, K. K. Y. Young, A. M. Q. King, and G. W. Wertz. 1987. Immune and histopathological responses in animals vaccinated with recombinant vaccinia viruses that express individual genes of human respiratory syncytial virus. J. Virol. 61:3855-3861. 21. Sullender, W. M., L. J. Anderson, K. Anderson, and G. W. Wertz. 1990. Differentiation of respiratory syncytial virus subgroups with cDNA probes in a nucleic acid hybridization assay. J. Clin. Microbiol. 28:1683-1687. 21a.Walpita, P., and S. Barik. Personal communication. 22. Walpita, P., J. D. Connor, and D. Pfeifer. 1989. Protein fingerprinting: a novel virus identification system. J. Virol. Methods 25:315-324. 23. Walsh, E. E., and J. Hruska. 1983. Monoclonal antibodies to respiratory syncytial virus proteins: identification of the fusion protein. J. Virol. 1983:171-177.
