Vol. 14, No. 3 Printed in U.S.A.
INFECTION AND IMMUNITY, Sept. 1976, p. 613-617 Copyright X 1976 American Society for Microbiology
Immunodepression by Rowson-Parr Virus in Mice: Lymphocyte Markers and Capping Response of Spleen and Lymph Node Cells After Infection M. BENDINELLI* AND H. FRIEDMAN Institute of Microbiology, University ofPisa, 56100 Pisa, Italy*; and Department of Microbiology, Albert Einstein Medical Center, Philadelphia, Pennsylvania 19141
Received for publication 2 April 1976
Infection with Rowson-Parr virus (RPV) induced a rapid reduction in the number of immunoglobulin-positive and theta antigen-positive cells detectable by immunofluorescence in the spleens of susceptible BALB/c mice. The changes produced by RPV infection in the lymph nodes were different, since the number of immunoglobulin-positive cells was increased and the proportion of thetapositive cells remained unchanged. However, the ability of immunoglobulinbearing cells to redistribute their receptors into caps was reduced in both types of lymphoid tissue. A similar pattern of changes was produced by infection with Friend leukemia complex, from which RPV was originally obtained. These effects of RPV and Friend leukemia complex may contribute to the immunodepressed state of infected mice. A recent study (15) has described the occurrence of profound changes in the distribution of cells exhibiting lymphocyte markers in the spleens and lymph nodes of mice infected with Friend leukemia complex (FLC). These changes are important because they may be related to the state of immunodepression induced by FLC, the exact mechanism of which remains unknown (12, 14). One of the viral entities forming FLC is Rowson-Parr virus (RPV), which was isolated from FLC by end-point dilution (21). When injected alone into BALB/c mice, RPV induces a chronic viremia and a slight hyperplasia of the spleen that peaks on the 3rd week of infection and then subsides until, after a latency of several months, splenic lymphomas develop in most infected mice (7, 9, 10, 17). The relationships between RPV and other viruses present in FLC are not clear (11, 18, 19, 22), but it has been shown that RPV can serve as helper for the component of the complex responsible for the production of spleen foci by certain strains of FLC in susceptible hosts (23). The observation that RPV can markedly depress the antibody response (2, 8, 9) has raised the question of its role in the immunological impairment induced by the entire complex (3). It was therefore of interest to study the effects of RPV infection on the distribution of lymphocyte markers in the spleens and lymph nodes of susceptible mice. The present report shows that the modifications detectable by immunofluores613
cence in the distribution of lymphocyte markers after RPV infection closely resemble those produced by FLC, except for the absence of the changes appearing in the spleens of FLC-infected mice as a consequence of the early neoplastic enlargement of this organ induced by FLC. MATERIALS AND METHODS Experimental animals. Inbred female BALB/c mice were obtained from Cumberland View Farms, Clinton, Tenn., and weighed 18 to 20 g when used. The animals were housed in groups of six to eight in plastic mouse cages and fed Purina mouse chow and water ad libitum. Viruses. The RPV used for these experiments consisted of a single preparation of a clarified 10% spleen cell extract obtained from adult BALB/c mice 8 days after infection; this preparation contained 104°0 mean infectious doses per 0.1 ml when inoculated intraperitoneally into adult BALB/c mice (7). The FLC used was prepared from 8- to 10-day infected BALB/c mice and had a titer of 103-75 mean infectious doses per 0.1 ml. Both virus preparations were free of lactic dehydrogenase virus (20). Infection was initiated by inoculation of 0.2 ml of the appropriate virus either intraperitoneally or intravenously. Antisera. The preparation and characteristics of the reagents used have been described in detail previously (15). In brief, the anti-mouse immunoglobulin serum consisted of purified immunoglobulin from New Zealand White rabbits immunized with mouse immunoglobulin in complete Freund adjuvant and was conjugated with fluorescein isothiocyanate. The anti-theta serum was prepared by re-
614
BENDINELLI AND FRIEDMAN
peated injection of AKR mice intraperitoneally with 107 thymocytes from C3H mice and was absorbed with agarose as well as with erythrocytes and bone marrow cells from C3H and BALB/c mice. Anti-lymphocyte serum (ALS) was prepared in New Zealand White rabbits by repeated subcutaneous injections of 108 BALB/c lymph node cells in complete Freund adjuvant, and was absorbed with mouse erythrocytes and directly conjugated with fluorescein isothiocyanate. The anti-virus serum was obtained from isologous BALB/c mice inoculated with a preparation of FLC inactivated by incubation with 0.1% formalin at 4°C for 3 weeks. The use of this antiserum to detect RPV antigen was appropriate since RPV is present in very high titers in FLC preparations and cross-reacts serologically with FLC (7). Immunofluorescent assays. Lymphoid cells obtained from control and infected mice were stained for membrane immunofluorescence as suspensions by the procedures described in detail previously (15). In brief, the cells were obtained by squeezing the spleens or lymph nodes (pooled mesenteric, axillary, and popliteal nodes) through nylon fibers into phosphate-buffered saline (PBS), pH 7.2. The cells were then washed three times in the cold with PBS, resuspended in the various antisera at a final concentration of 107 nucleated cells per ml, and incubated at 4°C for 30 min. For the direct assays (using ALS and anti-immunoglobulin sera), the cells, after incubation in the sera diluted 1:50 in PBS, were washed three times in the cold and smeared on triplicate glass slides. For the indirect assays (using the anti-theta and anti-virus sera), the cells were incubated first with the specific antiserum diluted 1:20; after three washings, they were resuspended in labeled anti-mouse immunoglobulin serum diluted 1:150 and then incubated for an additional 30 min at 40C. The cells were washed three times, and triplicate smears were prepared. At least 500 nucleated cells were examined for each cell suspension in triplicate. For assessment of the immunoglobulin receptor capping response, the lymphoid cells were processed exactly as for the immunoglobulin assay except that incubation was performed at 37°C and that the PBS used for the final washing contained 3 x 10-3 M sodium azide. At least 500 nucleated cells were examined for each smear, and those cells displaying polar localization of the labeled anti-immunoglobulin serum were considered capped. The results are expressed as the percent of immunoglobulin-positive fluorescent cells showing capping.
INFECT. IMMUN. Igor w
.jSO 'u ,40
r-
220
1
,=-
-11-.
T.- !1 CoNTftS' e
3
5
7
DAYS AFTER RPV
5.
=:.,
-L 10 -o -.15
-L2- io "32 20
INFECTION
FIG. 1. Surface markers and capping response of spleen cells of mice at various times after RPV infection. Percentages of total cells labeled with anti-lymphocyte (O), anti-immunoglobulin (O), anti-theta (O), or anti-viral antigen (V) sera and percentage of immunoglobulin-positive cells showing capping at 37°C (O). Each point represents the mean of four or more mice. Vertical bars represent the range.
time, the percentage of splenic cells reacting with anti-theta and anti-immunoglobulin serum, as well as the percent of immunoglobulin-positive cells that capped, decreased to less than half the values observed with spleen cells from control, noninfected mice. Subsequently there was a gradual return to normal values, but even 32 days after infection there was still a decreased percentage of positive cells. On the other hand, the percentage of splenic cells reactive with ALS was not significantly affected by RPV infection, and throughout the observation period only moderate fluctuations were noted. The anti-virus serum did not react with detectable numbers of spleen cells from normal mice. However, 1 day after infection, approximately 10 to 15% of the spleen cells reacted with the anti-virus serum, and this proportion remained stable throughout the observation period. In the lymph nodes (Fig. 2), the changes after RPV infection were markedly different. The proportion of immunoglobulin-positive cells increased, rather than decreased, as a consequence of virus injection. Peak numbers of immunoglobulin-positive cells occurred on days 1 to 3 after infection, with nearly twice as many cells exhibiting this surface marker as comRESULTS pared with the lymph node cells in control After intraperitoneal inoculation with RPV, mice. There was then a gradual decline toward the distribution pattern of cells with distinctive control values over the following weeks. In consurface markers underwent rapid alterations in trast, the percentage of immunoglobulin-posiboth the spleens and lymph nodes. In the tive cells that capped when incubated with the spleens (Fig. 1), the percentages of cells that anti-immunoglobulin serum at 3700 decreased stained with either anti-immunoglobulin or rapidly from a value of about 75% in control anti-theta serum, as well as the proportion of animals to less than 45% 1 day after infection, immunoglobulin-positive cells displaying polar and remained lower than 50% throughout the capping, were markedly reduced. Maximum next weeks. In the lymph nodes there was little changes occurred 1 day after infection. At this if any change in the percentage of ALS-positive
VOL. 14, 1976
615
LYMPHOCYTE MARKERS IN RPV INFECTION
and theta-positive cells after infection, and the cells positive for virus antigen ranged between 10 and 20% throughout the period of observation. In control experiments, it was clear that injection of extracts of normal BALB/c spleens, formalin-treated infected spleens, or a 20% suspension of washed sheep erythrocytes had no detectable effects on the distribution of the surface markers being studied. Furthermore, in other control studies it was found that there were no significant differences in the alterations of these markers when RPV was inoculated intravenously rather than intraperitoneally, or when infected plasma rather than a spleen preparation was used to infect the animals. Table 1 gives the results of one experiment in which the effects of RPV and FLC on the staining properties of spleen and lymph node cells
were compared. Mice were examined 5 and 15 days after infection. At both these times, the alterations induced by the two virus preparations on immunoglobulin-positive, theta-positive, and capping-positive cells were similar. However, a discrepancy was evident in regards to ALS- and viral antigen-positive cells in the spleens. This was especially apparent 15 days after infection, when (i) ALS-positive cells were markedly reduced in FLC-infected spleens but present in normal numbers in the spleens of RPV-infected mice, and (ii) the cells positive for virus antigen were 9% in the spleens of RPVinfected animals and 33% in the spleens of FLCinfected animals.
DISCUSSION The presence of theta antigen or of large amounts of immunoglobulin molecules on lymphocyte surfaces is considered a valuable marker for B- and T-cells, respectively (16, 24, 25). The present study, by the use of appropriate antisera in fluorescent microscopy, demon._40 strates profound changes in the proportions of cells evidencing such markers in both spleens and lymph nodes of RPV-infected mice. Since the weight and cellularity of these lymphoid organs in RPV-infected mice increased only during the 3rd week of infection (confirming earlier findings [7]), all the variations in percentages observed before this time must be considered to reflect true modifications in the abso16 1 21 32 CONotSL 1 lute number of cells exhibiting a marker. DAYS AFTER RPV INFECTION In the spleens, RPV infection was followed by FIG. 2. Surface markers and capping response of a decrease in the number of cells staining rapid lymph node cells of mice at various times after RPV infection. Symbols are as in Fig. 1. Each point repre- with anti-immunoglobulin or anti-theta sera. sents the mean of four or more mice. Vertical bars This decrease could be due to a depletion of cells represent the range. bearing these membrane markers or, alterna,,
_6
TABLE 1. Effect of infection of BALBIc mice with RPV or FLC on surface markers and on the capping response of spleen and lymph node cells
Virus infection"
Mean spleen wt
Lymphocytes
Theta antigen
Viral antigen
(mg)
None
Percentage of
Percentage of cells labeled with antiserum to
Spleen 79
LN
Spleen 29
LN 58
Spleen 0
LN 0
Immunoglobu-
cappedc
lin
Spleen 32
LN 14
Spleen 87
LN
74 >95 88 39 15 20 25 56 >95 66 9 100 78 18 87 16 17 24 49 69 >95 19 56 9 NMY 45 16 24 33 74 >95 17 56 22 151 58 ND 9 13 43 38 >95 20 41 33 365 38 a Groups of five to six mice injected intraperitoneally with the indicated virus on day indicated before the day of testing. b Average percentage of fluorescent cells for 500 or more spleen or superficial lymph node (LN) cells tested in triplicate for each group. e Average percentage of immunoglobulih (Ig)-positive cells that capped when incubated with antiimmunoglobulin serum at 37 'C. d ND, Not done.
RPV at day 5 RPV at day 15 FLC at day 5 FLC at day 15
616
BENDINELLI AND FRIEDMAN
tively, to a reduction in the concentration ofthe marker antigens on the cell surface or to impaired ability to interact with the corresponding antibody. The results of the present study do not indicate which ofthese interpretations is correct. However, previous studies have shown that in vitro RPV-infected spleen cells, after appropriate treatments, can develop antibodyforming cell responses in the normal range (6), suggesting that normal numbers of precursor cells are present and thus arguing against the first possibility, as far as immunoglobuliA-positive cells are concerned. RPV infection was also followed by a decrease in the number of spleen cells that could redistribute their immunoglobulin receptors into polar caps when incubated with anti-immunoglobulin serum at 37°C. This finding indicates that immunoglobulin molecules move less actively or freely in or on the lymphocyte membrane of RPV-infected spleens (16, 24, 25). The decrease in the ability of lymphocytes to display cap formation was very pronounced (the results are expressed as the percentage of immunoglobulin-positive cells that were also strongly decreased). Since capping is believed to precede the interiorization of antigen and the triggering of precursor cells (13, 16, 24, 25), such a decrease in capping response may be related to the impairment of antibody responsiveness exerted by RPV. A major objection to this possibility, however, is the absence of a close temporal relationship between reduction in the ability to cap and development of immunosuppression (8). The effects of RPV on the lymph nodes were different. As in the spleens, the percentage of immunoglobulin-positive cells that could form caps was reduced after infection, but in the lymph nodes this reduction was compensated for by an increase of total immunoglobulinpositive cells. If inability to cap and immunodepression are indeed related phenomena, then this compensation might explain the virtually normal response of RPV-infected mice given antigen by foodpad inoculation (4). Moreover, theta-positive cells in the lymph nodes were not significantly affected by RPV infection. This agrees with the earlier finding that contact sensitization after percutaneous immunization, a typical T-cell-mediated reaction depending on the normal functioning of superficial lymph nodes, is not impaired during RPV infection (5). The proportion of ALS-positive cells in both spleens and lymph nodes was not altered by RPV infection. This is not an unexpected finding because RPV does not cause proliferation of neoplastic cells 'that might dilute lymphoid cells early in the course of infection (9, 17). The
INFECT. IMMUN.
proportion of virus antigen-positive cells in the spleens and lymph nodes was 10 to 20% throughout the period of observation, indicating that RPV replicates in only a small proportion of cells in these organs and that this population of cells does not expand significantly during infection. As a corollary to this conclusion, it seems likely that the immunoglobulin and theta marker changes of spleen and lymph node cells are not due to viral replication in these same cells. For the reasons explained in the introduction, it was of obvious interest to compare the effects of RPV with those of FLC. In the present study, the FLC strain from which RPV was initially derived was used. Although somewhat less pronounced, its effects were generally similar to those observed in previous investigations using another strain of FLC originally derived from the American Type Culture Collection (15). The effects of RPV and FLC infections on immunoglobulin-positive cells, on theta-positive cells, and on capping in the spleen and lymph nodes were essentially the same. The only major difference concerned the proportions of ALS-positive and virus-positive cells in the spleens: as FLC infection proceeded, there was a decrease in the former cell population paralleled by an increase in the latter, whereas during RPV infection there were no effects on ALSpositive cells and the percentage of virus antigen-positive cells remained essentially steady after the early increase. This discrepancy appears to reflect the rapid proliferation of neoplastic cells releasing virus that occurs in FLCbut not RPV-infected spleens: the virus-transformed cells rapidly dilute the lymphoid cells to become the predominant cell population in the spleens of FLC-infected mice (1). The rapid onset of lymphocyte changes brought about by RPV infection is noteworthy. Most of the changes appeared, and some peaked, within 1 day after infection, raising the question of the mechanism by which they were induced. Since little RPV is detectable as infectivity in the spleen 1 day after infection, and since the virus titer does not peak in the spleen until the 3rd day (7), it appears that the alterations in surface markers require little if any virus replication. Since the inoculation of another antigen (sheep erythrocytes) and of a formalin-inactivated virus preparation had no detectable effects, it is not feasible that the lymphocyte changes noted were due to the antigenic moiety of the virus. In this regard, and to clarify the relationship between lymphocyte changes and immunodepression, it would be of obvious interest to investigate the effects on lymphocyte surface markers of infections with
VOL. 14, 1976
LYMPHOCYTE MARKERS IN RPV INFECTION
other viruses, both oncogenic and nononcogenic, having different effects on host's immune responses. Such studies are underway in our laboratories. ACKNOWLEDGMENTS This investigation was supported in part by grants from the U.S. National Science Foundation and the National Institute of Allergy and Infectious Diseases. M. B. was supported by a grant from the Italian National Research Council. We acknowledge the excellent technical assistance of John Holderbach. LITERATURE CITED 1. Bainbridge, D. R., and M. Bendinelli. 1972. Circulation of lymphoid cells in mice infected with Friend leukemia virus. J. Natl. Cancer Inst. 49:773-781. 2. Bendinelli, M. 1971. Effect of Friend leukemia virus and Rowson-Parr virus on immunological maturation of mice. Infect. Immun. 4:1-5. 3. Bendinelli, M. 1971. Immunodepression by Friend virus, p. 314. In J. L. Melnick (ed.), Proc. 2nd Int. Congr. Virol. S. Karger, Basel. 4. Bendinelli, M. 1973. Immunodepression by RowsonParr virus, p. 181-221. In W. S. Ceglowski and H. Friedman (ed.), Virus tumorigenesis and immunogenesis. Academic Press Inc., New York. 5. Bendinelli, M., M. Campa, and A. Toniolo. 1975. Immunodepression by Rowson-Parr virus: effect of Rowson-Parr virus and Friend leukemia complex infections on contact sensitivity in susceptible and resistant mice. Infect. Immun. 11:1031-1037. 6. Bendinelli, M., G. S. Kaplan, and H. Friedman. 1975. Reversal of leukemia virus-induced immunosuppression in vitro by peritoneal macrophages. J. Natl. Cancer Inst. 55:1425-1432. 7. Bendinelli, M., and L. Nardini. 1973. Immunodepression by Rowson-Parr virus in mice. I. Growth curves of Rowson-Parr virus and immunological relationships with Friend virus. Infect. Immun; 7:152-159. 8. Bendinelli, M., and L. Nardini. 1973. Immunodepression by Rowson-Parr virus in mice. II. Effect of Rowson-Parr virus infection on the antibody response to sheep red cells in vivo and in vitro. Infect. Immun. 7:160-166. 9. Carter, R. L., F. C. Chestennan, K. E. K. Rowson, M. H. Salaman, and N. Wedderburn. 1970. A new virus of minimal pathogenicity associated with Friend virus. II. Histological changes and immunodepressive effect. Int. J. Cancer. 5:103-110. 10. Carter, R. L., F. C. Chestennan, K. E. K. Rowson, M. H. Salaman, and N. Wedderburn. 1970. Induction of
617
lymphoma in BALB/c mice by Rowson-Parr virus. Int. J. Cancer 6:290-303. 11. Dawson, R. J., R. B. Tacke, and A. H. Fieldsteel. 1968. Relationship between Friend virus and an associated lymphatic leukemia virus. Br. J. Cancer 22:569-576. 12. Dent, P. B. 1972. Immunodepression by oncogenic viruses. Prog. Med. Virol. 14:1-35. 13. Dunham, E. K., E. R. Unanue, and B. Benaceraff. 1972. Antigen binding and capping by lymphocytes of genetic nonresponder mice. J. Exp. Med. 136:403-408. 14. Friedman, H., and W. S. Ceglowski. 1971. Immunosuppression by tumor viruses: effect of leukemia virus infection on immune response. Prog. Immunol. 1:815-819. 15. Kateley, J. R., J. Holderbach, and H. Friedman. 1974. Leukemia virus-induced alteration of lymphocyte Ig surface receptors and the "capping" response of mouse spleen and lymph node cells. J. Natl. Cancer Inst. 53:1135-1140. 16. Loor, F., L. Forni, and B. Pernis 1972. The dynamic state of the lymphocyte membrane. Factors affecting the distribution and turnover of surface immunoglobulins. Eur. J. Immunol. 2:203-212. 17. Michaels, L., K. E. K. Rowson, and S. E. Bird. 1972. Electron microscopical study of Rowson-Parr virus infection in BALB/c mice. Int. J. Cancer 9:162-171. 18. Pope; J. H. 1963. Detection of an avirulent virus apparently related to Friend virus. Aust. J. Exp. Biol. Med. Sci. 41:349-362. 19. Rich, M. A., and L. W. Johns, Jr. 1963. Morphology of an agent associated with a murine leukemia. Virology 20:373-376. 20. Riley, V., F. Lilly, E. Huerto, and D. Bardell. 1960. Transmissible agent associated with 26 types of experimental mouse neoplasms. Science 132:545-547. 21. Rowson, K. E. K., and I. Parr. 1970. A new virus of minimal pathogenicity associated with Friend virus. I. Isolation by end-point dilution. Int. J. Cancer 5:96102. 22. Steeves, R. A., R. J. Eckners M. Bennett, E. A. Mirand, and P. J. Trudel. Isolation and characterization of a lymphatic leukemia virus in the Friend virus complex. J. Natl. Cancer Inst. 46:1209-1217. 23. Steeves, R. A., R. J. Eckner, E. A. Mirand, and R. L. Priore. 1971. Rapid assay of murine leukemia virus helper activity for Friend spleen focus-forming virus. J. Natl. Cancer Inst. 46:1219-1228. 24. Taylor, R. B., W. P. H. Duffus, M. C. Raff, and S. De Petris. 1971. Redistribution and pinocytosis of lymphocyte surface immunoglobulin molecules induced by anti-immunoglobulin antibody. Nature (London) New Biol. 233:225-229. 25. Unanue, E. R., W. D. Perkins, and M. J. Karnovsky. 1972. Endocytosis by lymphocytes of complexes of anti-Ig with membrane-bound Ig. J. Immunol. 108:569-572.
INFECTION AND IMMUNITY, Sept. 1976, p. 613-617 Copyright X 1976 American Society for Microbiology
Immunodepression by Rowson-Parr Virus in Mice: Lymphocyte Markers and Capping Response of Spleen and Lymph Node Cells After Infection M. BENDINELLI* AND H. FRIEDMAN Institute of Microbiology, University ofPisa, 56100 Pisa, Italy*; and Department of Microbiology, Albert Einstein Medical Center, Philadelphia, Pennsylvania 19141
Received for publication 2 April 1976
Infection with Rowson-Parr virus (RPV) induced a rapid reduction in the number of immunoglobulin-positive and theta antigen-positive cells detectable by immunofluorescence in the spleens of susceptible BALB/c mice. The changes produced by RPV infection in the lymph nodes were different, since the number of immunoglobulin-positive cells was increased and the proportion of thetapositive cells remained unchanged. However, the ability of immunoglobulinbearing cells to redistribute their receptors into caps was reduced in both types of lymphoid tissue. A similar pattern of changes was produced by infection with Friend leukemia complex, from which RPV was originally obtained. These effects of RPV and Friend leukemia complex may contribute to the immunodepressed state of infected mice. A recent study (15) has described the occurrence of profound changes in the distribution of cells exhibiting lymphocyte markers in the spleens and lymph nodes of mice infected with Friend leukemia complex (FLC). These changes are important because they may be related to the state of immunodepression induced by FLC, the exact mechanism of which remains unknown (12, 14). One of the viral entities forming FLC is Rowson-Parr virus (RPV), which was isolated from FLC by end-point dilution (21). When injected alone into BALB/c mice, RPV induces a chronic viremia and a slight hyperplasia of the spleen that peaks on the 3rd week of infection and then subsides until, after a latency of several months, splenic lymphomas develop in most infected mice (7, 9, 10, 17). The relationships between RPV and other viruses present in FLC are not clear (11, 18, 19, 22), but it has been shown that RPV can serve as helper for the component of the complex responsible for the production of spleen foci by certain strains of FLC in susceptible hosts (23). The observation that RPV can markedly depress the antibody response (2, 8, 9) has raised the question of its role in the immunological impairment induced by the entire complex (3). It was therefore of interest to study the effects of RPV infection on the distribution of lymphocyte markers in the spleens and lymph nodes of susceptible mice. The present report shows that the modifications detectable by immunofluores613
cence in the distribution of lymphocyte markers after RPV infection closely resemble those produced by FLC, except for the absence of the changes appearing in the spleens of FLC-infected mice as a consequence of the early neoplastic enlargement of this organ induced by FLC. MATERIALS AND METHODS Experimental animals. Inbred female BALB/c mice were obtained from Cumberland View Farms, Clinton, Tenn., and weighed 18 to 20 g when used. The animals were housed in groups of six to eight in plastic mouse cages and fed Purina mouse chow and water ad libitum. Viruses. The RPV used for these experiments consisted of a single preparation of a clarified 10% spleen cell extract obtained from adult BALB/c mice 8 days after infection; this preparation contained 104°0 mean infectious doses per 0.1 ml when inoculated intraperitoneally into adult BALB/c mice (7). The FLC used was prepared from 8- to 10-day infected BALB/c mice and had a titer of 103-75 mean infectious doses per 0.1 ml. Both virus preparations were free of lactic dehydrogenase virus (20). Infection was initiated by inoculation of 0.2 ml of the appropriate virus either intraperitoneally or intravenously. Antisera. The preparation and characteristics of the reagents used have been described in detail previously (15). In brief, the anti-mouse immunoglobulin serum consisted of purified immunoglobulin from New Zealand White rabbits immunized with mouse immunoglobulin in complete Freund adjuvant and was conjugated with fluorescein isothiocyanate. The anti-theta serum was prepared by re-
614
BENDINELLI AND FRIEDMAN
peated injection of AKR mice intraperitoneally with 107 thymocytes from C3H mice and was absorbed with agarose as well as with erythrocytes and bone marrow cells from C3H and BALB/c mice. Anti-lymphocyte serum (ALS) was prepared in New Zealand White rabbits by repeated subcutaneous injections of 108 BALB/c lymph node cells in complete Freund adjuvant, and was absorbed with mouse erythrocytes and directly conjugated with fluorescein isothiocyanate. The anti-virus serum was obtained from isologous BALB/c mice inoculated with a preparation of FLC inactivated by incubation with 0.1% formalin at 4°C for 3 weeks. The use of this antiserum to detect RPV antigen was appropriate since RPV is present in very high titers in FLC preparations and cross-reacts serologically with FLC (7). Immunofluorescent assays. Lymphoid cells obtained from control and infected mice were stained for membrane immunofluorescence as suspensions by the procedures described in detail previously (15). In brief, the cells were obtained by squeezing the spleens or lymph nodes (pooled mesenteric, axillary, and popliteal nodes) through nylon fibers into phosphate-buffered saline (PBS), pH 7.2. The cells were then washed three times in the cold with PBS, resuspended in the various antisera at a final concentration of 107 nucleated cells per ml, and incubated at 4°C for 30 min. For the direct assays (using ALS and anti-immunoglobulin sera), the cells, after incubation in the sera diluted 1:50 in PBS, were washed three times in the cold and smeared on triplicate glass slides. For the indirect assays (using the anti-theta and anti-virus sera), the cells were incubated first with the specific antiserum diluted 1:20; after three washings, they were resuspended in labeled anti-mouse immunoglobulin serum diluted 1:150 and then incubated for an additional 30 min at 40C. The cells were washed three times, and triplicate smears were prepared. At least 500 nucleated cells were examined for each cell suspension in triplicate. For assessment of the immunoglobulin receptor capping response, the lymphoid cells were processed exactly as for the immunoglobulin assay except that incubation was performed at 37°C and that the PBS used for the final washing contained 3 x 10-3 M sodium azide. At least 500 nucleated cells were examined for each smear, and those cells displaying polar localization of the labeled anti-immunoglobulin serum were considered capped. The results are expressed as the percent of immunoglobulin-positive fluorescent cells showing capping.
INFECT. IMMUN. Igor w
.jSO 'u ,40
r-
220
1
,=-
-11-.
T.- !1 CoNTftS' e
3
5
7
DAYS AFTER RPV
5.
=:.,
-L 10 -o -.15
-L2- io "32 20
INFECTION
FIG. 1. Surface markers and capping response of spleen cells of mice at various times after RPV infection. Percentages of total cells labeled with anti-lymphocyte (O), anti-immunoglobulin (O), anti-theta (O), or anti-viral antigen (V) sera and percentage of immunoglobulin-positive cells showing capping at 37°C (O). Each point represents the mean of four or more mice. Vertical bars represent the range.
time, the percentage of splenic cells reacting with anti-theta and anti-immunoglobulin serum, as well as the percent of immunoglobulin-positive cells that capped, decreased to less than half the values observed with spleen cells from control, noninfected mice. Subsequently there was a gradual return to normal values, but even 32 days after infection there was still a decreased percentage of positive cells. On the other hand, the percentage of splenic cells reactive with ALS was not significantly affected by RPV infection, and throughout the observation period only moderate fluctuations were noted. The anti-virus serum did not react with detectable numbers of spleen cells from normal mice. However, 1 day after infection, approximately 10 to 15% of the spleen cells reacted with the anti-virus serum, and this proportion remained stable throughout the observation period. In the lymph nodes (Fig. 2), the changes after RPV infection were markedly different. The proportion of immunoglobulin-positive cells increased, rather than decreased, as a consequence of virus injection. Peak numbers of immunoglobulin-positive cells occurred on days 1 to 3 after infection, with nearly twice as many cells exhibiting this surface marker as comRESULTS pared with the lymph node cells in control After intraperitoneal inoculation with RPV, mice. There was then a gradual decline toward the distribution pattern of cells with distinctive control values over the following weeks. In consurface markers underwent rapid alterations in trast, the percentage of immunoglobulin-posiboth the spleens and lymph nodes. In the tive cells that capped when incubated with the spleens (Fig. 1), the percentages of cells that anti-immunoglobulin serum at 3700 decreased stained with either anti-immunoglobulin or rapidly from a value of about 75% in control anti-theta serum, as well as the proportion of animals to less than 45% 1 day after infection, immunoglobulin-positive cells displaying polar and remained lower than 50% throughout the capping, were markedly reduced. Maximum next weeks. In the lymph nodes there was little changes occurred 1 day after infection. At this if any change in the percentage of ALS-positive
VOL. 14, 1976
615
LYMPHOCYTE MARKERS IN RPV INFECTION
and theta-positive cells after infection, and the cells positive for virus antigen ranged between 10 and 20% throughout the period of observation. In control experiments, it was clear that injection of extracts of normal BALB/c spleens, formalin-treated infected spleens, or a 20% suspension of washed sheep erythrocytes had no detectable effects on the distribution of the surface markers being studied. Furthermore, in other control studies it was found that there were no significant differences in the alterations of these markers when RPV was inoculated intravenously rather than intraperitoneally, or when infected plasma rather than a spleen preparation was used to infect the animals. Table 1 gives the results of one experiment in which the effects of RPV and FLC on the staining properties of spleen and lymph node cells
were compared. Mice were examined 5 and 15 days after infection. At both these times, the alterations induced by the two virus preparations on immunoglobulin-positive, theta-positive, and capping-positive cells were similar. However, a discrepancy was evident in regards to ALS- and viral antigen-positive cells in the spleens. This was especially apparent 15 days after infection, when (i) ALS-positive cells were markedly reduced in FLC-infected spleens but present in normal numbers in the spleens of RPV-infected mice, and (ii) the cells positive for virus antigen were 9% in the spleens of RPVinfected animals and 33% in the spleens of FLCinfected animals.
DISCUSSION The presence of theta antigen or of large amounts of immunoglobulin molecules on lymphocyte surfaces is considered a valuable marker for B- and T-cells, respectively (16, 24, 25). The present study, by the use of appropriate antisera in fluorescent microscopy, demon._40 strates profound changes in the proportions of cells evidencing such markers in both spleens and lymph nodes of RPV-infected mice. Since the weight and cellularity of these lymphoid organs in RPV-infected mice increased only during the 3rd week of infection (confirming earlier findings [7]), all the variations in percentages observed before this time must be considered to reflect true modifications in the abso16 1 21 32 CONotSL 1 lute number of cells exhibiting a marker. DAYS AFTER RPV INFECTION In the spleens, RPV infection was followed by FIG. 2. Surface markers and capping response of a decrease in the number of cells staining rapid lymph node cells of mice at various times after RPV infection. Symbols are as in Fig. 1. Each point repre- with anti-immunoglobulin or anti-theta sera. sents the mean of four or more mice. Vertical bars This decrease could be due to a depletion of cells represent the range. bearing these membrane markers or, alterna,,
_6
TABLE 1. Effect of infection of BALBIc mice with RPV or FLC on surface markers and on the capping response of spleen and lymph node cells
Virus infection"
Mean spleen wt
Lymphocytes
Theta antigen
Viral antigen
(mg)
None
Percentage of
Percentage of cells labeled with antiserum to
Spleen 79
LN
Spleen 29
LN 58
Spleen 0
LN 0
Immunoglobu-
cappedc
lin
Spleen 32
LN 14
Spleen 87
LN
74 >95 88 39 15 20 25 56 >95 66 9 100 78 18 87 16 17 24 49 69 >95 19 56 9 NMY 45 16 24 33 74 >95 17 56 22 151 58 ND 9 13 43 38 >95 20 41 33 365 38 a Groups of five to six mice injected intraperitoneally with the indicated virus on day indicated before the day of testing. b Average percentage of fluorescent cells for 500 or more spleen or superficial lymph node (LN) cells tested in triplicate for each group. e Average percentage of immunoglobulih (Ig)-positive cells that capped when incubated with antiimmunoglobulin serum at 37 'C. d ND, Not done.
RPV at day 5 RPV at day 15 FLC at day 5 FLC at day 15
616
BENDINELLI AND FRIEDMAN
tively, to a reduction in the concentration ofthe marker antigens on the cell surface or to impaired ability to interact with the corresponding antibody. The results of the present study do not indicate which ofthese interpretations is correct. However, previous studies have shown that in vitro RPV-infected spleen cells, after appropriate treatments, can develop antibodyforming cell responses in the normal range (6), suggesting that normal numbers of precursor cells are present and thus arguing against the first possibility, as far as immunoglobuliA-positive cells are concerned. RPV infection was also followed by a decrease in the number of spleen cells that could redistribute their immunoglobulin receptors into polar caps when incubated with anti-immunoglobulin serum at 37°C. This finding indicates that immunoglobulin molecules move less actively or freely in or on the lymphocyte membrane of RPV-infected spleens (16, 24, 25). The decrease in the ability of lymphocytes to display cap formation was very pronounced (the results are expressed as the percentage of immunoglobulin-positive cells that were also strongly decreased). Since capping is believed to precede the interiorization of antigen and the triggering of precursor cells (13, 16, 24, 25), such a decrease in capping response may be related to the impairment of antibody responsiveness exerted by RPV. A major objection to this possibility, however, is the absence of a close temporal relationship between reduction in the ability to cap and development of immunosuppression (8). The effects of RPV on the lymph nodes were different. As in the spleens, the percentage of immunoglobulin-positive cells that could form caps was reduced after infection, but in the lymph nodes this reduction was compensated for by an increase of total immunoglobulinpositive cells. If inability to cap and immunodepression are indeed related phenomena, then this compensation might explain the virtually normal response of RPV-infected mice given antigen by foodpad inoculation (4). Moreover, theta-positive cells in the lymph nodes were not significantly affected by RPV infection. This agrees with the earlier finding that contact sensitization after percutaneous immunization, a typical T-cell-mediated reaction depending on the normal functioning of superficial lymph nodes, is not impaired during RPV infection (5). The proportion of ALS-positive cells in both spleens and lymph nodes was not altered by RPV infection. This is not an unexpected finding because RPV does not cause proliferation of neoplastic cells 'that might dilute lymphoid cells early in the course of infection (9, 17). The
INFECT. IMMUN.
proportion of virus antigen-positive cells in the spleens and lymph nodes was 10 to 20% throughout the period of observation, indicating that RPV replicates in only a small proportion of cells in these organs and that this population of cells does not expand significantly during infection. As a corollary to this conclusion, it seems likely that the immunoglobulin and theta marker changes of spleen and lymph node cells are not due to viral replication in these same cells. For the reasons explained in the introduction, it was of obvious interest to compare the effects of RPV with those of FLC. In the present study, the FLC strain from which RPV was initially derived was used. Although somewhat less pronounced, its effects were generally similar to those observed in previous investigations using another strain of FLC originally derived from the American Type Culture Collection (15). The effects of RPV and FLC infections on immunoglobulin-positive cells, on theta-positive cells, and on capping in the spleen and lymph nodes were essentially the same. The only major difference concerned the proportions of ALS-positive and virus-positive cells in the spleens: as FLC infection proceeded, there was a decrease in the former cell population paralleled by an increase in the latter, whereas during RPV infection there were no effects on ALSpositive cells and the percentage of virus antigen-positive cells remained essentially steady after the early increase. This discrepancy appears to reflect the rapid proliferation of neoplastic cells releasing virus that occurs in FLCbut not RPV-infected spleens: the virus-transformed cells rapidly dilute the lymphoid cells to become the predominant cell population in the spleens of FLC-infected mice (1). The rapid onset of lymphocyte changes brought about by RPV infection is noteworthy. Most of the changes appeared, and some peaked, within 1 day after infection, raising the question of the mechanism by which they were induced. Since little RPV is detectable as infectivity in the spleen 1 day after infection, and since the virus titer does not peak in the spleen until the 3rd day (7), it appears that the alterations in surface markers require little if any virus replication. Since the inoculation of another antigen (sheep erythrocytes) and of a formalin-inactivated virus preparation had no detectable effects, it is not feasible that the lymphocyte changes noted were due to the antigenic moiety of the virus. In this regard, and to clarify the relationship between lymphocyte changes and immunodepression, it would be of obvious interest to investigate the effects on lymphocyte surface markers of infections with
VOL. 14, 1976
LYMPHOCYTE MARKERS IN RPV INFECTION
other viruses, both oncogenic and nononcogenic, having different effects on host's immune responses. Such studies are underway in our laboratories. ACKNOWLEDGMENTS This investigation was supported in part by grants from the U.S. National Science Foundation and the National Institute of Allergy and Infectious Diseases. M. B. was supported by a grant from the Italian National Research Council. We acknowledge the excellent technical assistance of John Holderbach. LITERATURE CITED 1. Bainbridge, D. R., and M. Bendinelli. 1972. Circulation of lymphoid cells in mice infected with Friend leukemia virus. J. Natl. Cancer Inst. 49:773-781. 2. Bendinelli, M. 1971. Effect of Friend leukemia virus and Rowson-Parr virus on immunological maturation of mice. Infect. Immun. 4:1-5. 3. Bendinelli, M. 1971. Immunodepression by Friend virus, p. 314. In J. L. Melnick (ed.), Proc. 2nd Int. Congr. Virol. S. Karger, Basel. 4. Bendinelli, M. 1973. Immunodepression by RowsonParr virus, p. 181-221. In W. S. Ceglowski and H. Friedman (ed.), Virus tumorigenesis and immunogenesis. Academic Press Inc., New York. 5. Bendinelli, M., M. Campa, and A. Toniolo. 1975. Immunodepression by Rowson-Parr virus: effect of Rowson-Parr virus and Friend leukemia complex infections on contact sensitivity in susceptible and resistant mice. Infect. Immun. 11:1031-1037. 6. Bendinelli, M., G. S. Kaplan, and H. Friedman. 1975. Reversal of leukemia virus-induced immunosuppression in vitro by peritoneal macrophages. J. Natl. Cancer Inst. 55:1425-1432. 7. Bendinelli, M., and L. Nardini. 1973. Immunodepression by Rowson-Parr virus in mice. I. Growth curves of Rowson-Parr virus and immunological relationships with Friend virus. Infect. Immun; 7:152-159. 8. Bendinelli, M., and L. Nardini. 1973. Immunodepression by Rowson-Parr virus in mice. II. Effect of Rowson-Parr virus infection on the antibody response to sheep red cells in vivo and in vitro. Infect. Immun. 7:160-166. 9. Carter, R. L., F. C. Chestennan, K. E. K. Rowson, M. H. Salaman, and N. Wedderburn. 1970. A new virus of minimal pathogenicity associated with Friend virus. II. Histological changes and immunodepressive effect. Int. J. Cancer. 5:103-110. 10. Carter, R. L., F. C. Chestennan, K. E. K. Rowson, M. H. Salaman, and N. Wedderburn. 1970. Induction of
617
lymphoma in BALB/c mice by Rowson-Parr virus. Int. J. Cancer 6:290-303. 11. Dawson, R. J., R. B. Tacke, and A. H. Fieldsteel. 1968. Relationship between Friend virus and an associated lymphatic leukemia virus. Br. J. Cancer 22:569-576. 12. Dent, P. B. 1972. Immunodepression by oncogenic viruses. Prog. Med. Virol. 14:1-35. 13. Dunham, E. K., E. R. Unanue, and B. Benaceraff. 1972. Antigen binding and capping by lymphocytes of genetic nonresponder mice. J. Exp. Med. 136:403-408. 14. Friedman, H., and W. S. Ceglowski. 1971. Immunosuppression by tumor viruses: effect of leukemia virus infection on immune response. Prog. Immunol. 1:815-819. 15. Kateley, J. R., J. Holderbach, and H. Friedman. 1974. Leukemia virus-induced alteration of lymphocyte Ig surface receptors and the "capping" response of mouse spleen and lymph node cells. J. Natl. Cancer Inst. 53:1135-1140. 16. Loor, F., L. Forni, and B. Pernis 1972. The dynamic state of the lymphocyte membrane. Factors affecting the distribution and turnover of surface immunoglobulins. Eur. J. Immunol. 2:203-212. 17. Michaels, L., K. E. K. Rowson, and S. E. Bird. 1972. Electron microscopical study of Rowson-Parr virus infection in BALB/c mice. Int. J. Cancer 9:162-171. 18. Pope; J. H. 1963. Detection of an avirulent virus apparently related to Friend virus. Aust. J. Exp. Biol. Med. Sci. 41:349-362. 19. Rich, M. A., and L. W. Johns, Jr. 1963. Morphology of an agent associated with a murine leukemia. Virology 20:373-376. 20. Riley, V., F. Lilly, E. Huerto, and D. Bardell. 1960. Transmissible agent associated with 26 types of experimental mouse neoplasms. Science 132:545-547. 21. Rowson, K. E. K., and I. Parr. 1970. A new virus of minimal pathogenicity associated with Friend virus. I. Isolation by end-point dilution. Int. J. Cancer 5:96102. 22. Steeves, R. A., R. J. Eckners M. Bennett, E. A. Mirand, and P. J. Trudel. Isolation and characterization of a lymphatic leukemia virus in the Friend virus complex. J. Natl. Cancer Inst. 46:1209-1217. 23. Steeves, R. A., R. J. Eckner, E. A. Mirand, and R. L. Priore. 1971. Rapid assay of murine leukemia virus helper activity for Friend spleen focus-forming virus. J. Natl. Cancer Inst. 46:1219-1228. 24. Taylor, R. B., W. P. H. Duffus, M. C. Raff, and S. De Petris. 1971. Redistribution and pinocytosis of lymphocyte surface immunoglobulin molecules induced by anti-immunoglobulin antibody. Nature (London) New Biol. 233:225-229. 25. Unanue, E. R., W. D. Perkins, and M. J. Karnovsky. 1972. Endocytosis by lymphocytes of complexes of anti-Ig with membrane-bound Ig. J. Immunol. 108:569-572.
