414
TRANSACTIONS OF THE ROYAL SOCIETY
Cell mediated
OF
TROPICAL MEDICINE AND HYGIENE, VOL. 71, No. 6, 1977.
and humoral immunity in experimental Plasmodium berghei infection
ARUNA PARASHAR, B. K. AIKAT, S. SEHGAL AND S. NAIK Department
of Pathology,
Postgraduate Institute of Medical Chandigarh, India
Summary
Adoptive passive transfer of immunity to Plasmodium berghei infection has been investigated in an inbred strain of Swiss mice. The mice were made hyperimmune by repeated passage of 103 parasites and subsequent therapy with an antimalarial drug. Immune sera and cells obtained from thymus, spleen and peritoneal exudate were transferred to normal animals which were subsequently challenged with standard doses of P. berghei. It was observed that: (a) immune serum in high doses (0.5 ml/mouse) enhanced parasitaemia; when used in smaller doses (0.1 ml/mouse), it afforded a considerable degree of protection; (b) viable immune lymphocytes obtained from thymus and lymph node afforded protection; (c) the mixed population of cells obtained from spleens of immunized mice, as well as peritoneal exudate, protected mice against challenge inoculum; (d) glutaraldehyde-treated spleen cells and material obtained after freezing and thawing the same number of spleen cells, macrophages and lymph node also afforded protection. These findings confirm that, under these experimental conditions, immunity against P. berghei is mediated through (i) specific antibody which is dose-dependent, (ii) cell-mediated immunity and (iii) effective response to processed antigen. Introduction
Immunity to malaria has been extensively investigated. Both humoral, antiplasmodial antibodies and cellmediated immunity have been implicated. The importance of immunoglobulins in conferring immunity in malaria has been suggested repeatedly by passive transfer of immune sera, in humans and in experimental animals (BRUCE-CHWATT & GIBSON, 1956; BRIGGS et al., 1966; DIGGS & OLSER, 1969; COHEN & BUTCHER, 1971; 1974). Protective antiplasmodial GOLENSER et al., globulins act on extracellular merozoites and on mature schizonts. More recent in vitro experiments have shown that antibodies prevent merozoite penetration into the erythrocytes, either by neutralization (COHEN &BUTCHER, 1970) or by agglutination (MILLER et al., 1975). It has also been shown by these workers that transfer of a larger dose of antibody is required to inhibit transiently parasite multiplication. JERUSALEM et al. (1971) and HAMBURGER & KREIER (1976) administered immune serum to rats and observed that there was partial or complete protection against parasitaemia in these animals. The coating of free extraerythrocytic parasites by the antibody has been suggested as the possible mode of action. However, the immune
Education
and Research,
serum does not kill the intracellular parasite, because transfer of infected blood from treated rats into mice, results in multiplication. Thus in vivo control of parasitaemia requires factors other than antibody and the possible role of macrophages has been suggested (BROWN & PHILLIPS, 1974); GOLENSER et al., 1974). Transfer of lymphoid cells from immune animals also protects against subsequent infection (STECHSHULTE, 1969 ; PHILLIPS, 1970; CABRERA & ALGER, 1971). It has been shown that cells from the spleen are more effective in conferring protection (TODORONIC et al., 1967; PHILLIPS. 1970): CABRERA & ALGER. 1970). Thoracic duct lymphocytes, bone marrow cells, thymus cells and peritoneal exudate cells were ineffective when transferred separately (ROBERT & TRACEYPATTE, 1968 ; STECHSHULTE, 1969 ). PHILLIPS (1969) was able to demonstrate protection with immune macrophages but STECHSHULTE (1969 ) could not confirm these findings. T-cell depletion by neonatal thymectomy and using antithymocyte serum made animals more susceptible to infection than were controls (BROWN et al., 1968; STECHSHULTE, 1969 ; SPIRA et al., 1970). Adoptive transfer of immunity by lymphoid cells also prompted workers to propose that immunity against P. berghei infection is dependent on effective CMI. However, BROWN et al. (1976) have shown by adoptive transfer that unfractionated spleen cells and T-cells alone could transfer protection to syngeneic recipients as early as 11 days after infection of the cell donors. They postulated that immune T-cells may act as helper cells and presumably get sensitized to variant specific antigens. Recently GRAVELY & KREIER (1976) have shown that immunity to malarial infection can also be transferred adoptively by the “Null Cells” or antibody forming “B” cells. In view of the controversial data, the present experiments were designed to study the comparative role of passively transferred immune lymphoid cells from different soumes: macrophages and immune sera in experimental P. berghei infection in Swiss inbred strain of mice. Materials
and methods
Animals
Four to six-week-old syngeneic Swiss mice/ICRC weighing 20-25 g were taken and fed ad libitum. Malarial
parasite
Strain of P. berghei obtained from N.I.C.D., New Delhi was used in this experiment. The strain was main-
ARUNA
PAUASHAR
et al.
tained by weekly serial intraperitoneal passage in Swiss inbred mice. The animals were inoculated with erythrocytes from Swiss mice infected with P. berghei. Chronic
malaria
Malaria was induced experimentally in 24 Swiss inbred (male) mice by using 1 x 103 malaria-infected red blood cells as the standard inoculum, chronic&y was maintained by giving three repeated doses of the same number of infected erythrocytes at 17-day intervals and simultaneous therapy with chloroquine (5 mg/kg body-weight). Daily blood smears were made from the tail vein from each animal in duplicate and 500 red cells were counted to determine the percentage of parasitaemia. When parasitaemia reached its peak (nearly 175 infected/500 red cells) on the tenth day, the mice were treated with chloroquine. (Two intraperitoneal injections at a two-day interval) until the parasitaemia came down to 10 to 20 infected cells/500 red cells on day 17. The animals were challenged a second time on the 17th day with the same number of parasites from the same stabilate. The parasitaemia reached its peak on the 27th day-up to 55 to 60 parasitized cells/500 red cells; the animals were treated again with the same dose of chloroquine at the same interval until parasitaemia came down to 10 parasitized cells/ 500 red cells. The third infection was given on day 34 and parasitaemia again recorded daily. It reached its peak of 23 to 25 parasitized cells/500 red cells. The animals were treated with chloroquine until the parasitaemia was completely controlled up to one week. When the animals became aparasitaemic, as confirmed from the peripheral blood smears, they were used as donors in the transfer experiments (Fig. 1). Age and sex matched normal animals were taken as donors for the control groups. Preparation
of lymphoid
cells and collection
of serum
Strictly sterile precautions were taken throughout these experiments. Before killing the animals, blood was collected from each animal, pooled and allowed to clot at 37°C for 30 min. and then centrifuged at 4°C for 30 min. at 1,500 r.p.m. The serum was separated and kept at 4°C until use. Harvesting
of macrophages
After collecting the serum the animals were lightly anaesthetized with ether. The skin was dissected with sterilized forceps and the peritoneum exposed. 5 to 7 ml of Hank’s Balanced Salt Solution (HBSS) were introduced into the peritoneal cavity by means of a syringe and the peritoneum gently massaged. The fluid was carefully withdrawn into the syringe and the cells were washed thrice in cold HBSS and resuspended in the same medium. Viability was checked in trypan blue (NAYSMITH & JAMES, 1968); cell counts were made in a haemacytometer and adjusted to the desired concentration for injection (5.5 x 106 cell/mouse). The percentage of nonviable cells was usually between five and ten. Preparation
of cell suspension
Spleens, lymph nodes and thymi were taken out under sterile conditions. teased in cold HBSS and then oassed through a 60 mesh gauze. The cells were centrifuged at 1,500 r.p.m. at 4°C for 10 min. and washed four times. The viability was tested with trypan blue; cells were counted and adjusted at 70 x 106 cells/05 ml/mouse. Cells were collected similarly from normal animals.
Fig. 1. Graph donors.
Freeze
showing
mean blood parasitaemia
in
thawing
Spleens, lymph nodes, thymi and peritoneal exudate from immune and normal animals were collected in cold HBSS under sterile conditions: teased in cold HBSS and then passed through a 60 mesh gauze. The cells were centrifuged at 1,500 r.p.m. at 4°C for 10 min. and washed four times with cold HBSS. The viability was tested with trypan blue. Cells were counted and adjusted to 70 x 106 cells/O.5 ml/mouse, kept at -70°C for one hour and thawed at 37°C in a water bath. Freezing and thawing the cells was repeated three times. After the final thawing the tubes were centrifuged at 1,500 r.p.m. for 10 min. and the supernatant was used for inoculation. A similar procedure was used with macrophages but the number of cells was adjusted to 5.5 x 106 cells/mouse. Glutaraldehyde
fixation
The method of SANDERSON & FROST (1974) was used for glutaraldehyde fixation. Spleens from normal and immune donors were removed under sterile conditions in HBSS containing antibiotics. The tissue was homogenized and the cell suspension was sieved through a fine mesh. Cells were washed twice in HBSS and the final count was adjusted to 1 x 108 cells/ml in MEM. An equal volume of 0.25 “/, glutaraldehyde (obtained from Me&h-Schuchardt M& then) in MEM was added. The cells were kept for 15 min. at 22°C. They were subsequently washed three times and centrifuged at 1,000 r.p.m. for 10 min. in a cold MSE centrifuge. Each mouse received 70 x 106 glutaraldehyde treated cells. After 24 hours the animals were challenged with parasites. Control group of age and sex matched recipients received normal cells treated with glutaraldehyde as mentioned. Transfer
experiment
Serum was transferred into three different groups each group having six test and four control animals. In the test animals, 0.5, 0.25 and 0.1 ml of the hyperimmune serum was injected intraperitoneally into the three groups on day 0. After 24 hours the animals were challenged with 1 x 103 malaria-infected red cells and parasitaemia was recorded daily in Giemsa-stained smears; the control animals were given the same dose of normal serum and challenged with the same number of parasites after 24 hours. No drug therapy was given to either the control
476
CELLMEDIATED
AND
P.berghei
HUMORALIMMUNITYINEXPERIMENTAL
or the test group after transferring immune or normal serum. Lymphoid
cells transfer
Lymphoid cells from immune lymph nodes were given to two groups. One group of six mice received viable cells (70 x 106 cells/mouse) and the other group (also of six mice) received material from the same number of frozen and thawed cells. They were challenged with 1 x 103 malaria-infected red cells and parasitaemia was recorded daily. Thymocytes were also transferred in the same dose schedule as lymph node cells. In the case of the spleen, three groups (each of six animals) were used. Viable cells were administered to the first, frozen and thawed material from an equal number of cells to the second and glutaraldehyde-treated cells to the third group. With macrophages, since the number of cells obtained in peritoneal exudate was inadequate only 5.5 x 106 cells per mouse were given. The same number of cells were disrupted by freeze thawing and the material obtained was injected into the second group of six animals. These groups were challenged with parasites. The control received the same number of cells from. normal animals.
Fig. 2. Comparative effect on parasitaemia of 0.5 ml of hyperimmune serum and serum obtained from normal animals.
Results Protective
activity
in serum
Normal serum recipients and those to which no serum was given when challenged with parasites showed parasitaemia in one or two days. They died of fulminant infection within two to three weeks. The results of injecting hyperimmune serum in different doses are presented in Figs 2 and 3. It was apparent that when 0.5 ml of hyperimmune serum was used, there was marked enhancement of parasitaemia from the fourth day onwards which reached 38% on day 14. All the animals given 0.5 ml of hyperimmune serum died within 25 days. However, when 0.1 ml of hyperimmune-serum was used the results were inconsistent and three animals died while the parasitaemia declined in the other three. There was peak parasitaemia on day six (20%) and a subsequent fall in the level of parasitaemia (approximately 25% on the 14th day). Parasitaemia was significantly lower in the group given 0.25 ml of hyperimmune serum than in the control group. of thymus cells (Fig. 4) Thymocytes afforded considerable protection and parasitaemia remained at very low levels all throughout the experiment. All the animals survived up to 45 days and the maximum parasitaemia of 5% was seen on the eighth day. Normal thymocytes did not afford protection when the mice were subsequently challenged with parasites.
Fig. 3. Comparative effect on parasitaemia of 0.1 ml, 0.25 ml of hyperimmune serum and serum obtained from normal animals.
Effect
of immune lymph node cells (Fig. 5) When viable lymph node cells were inoculated, the maximum parasitaemia of 6 to 7 % was observed on day 10. All the animals were able to clear the infection within 20 days and no mortality was observed up to 45 days. Similarly, when the same number of lymphoid cells was disrupted by freezing and thawing and the material thus obtained was injected into the host, it did afford protection in four of six animals subsequently challenged but significantly less than did viable cells. The maximum parasitaemia seen in these mice was approximately 15 % on day seven compared to 7 ‘A on day 10 in those given
EFFECT
OF
THYMOCYTES
ON
PARASITAEMIA.
70X106MO~s~
Effect
Fig. 4. Comparative effect on parasitaemia after injecting viable thymocytes from immune and control animals.
ARUNA
EFFECT
OF
LYMPH
NODE
--
CELLS 70 Xl&
PARASHAR
ON PARASITAEMIA CELLS/MOUSE
OAY-
Fig. 5. Comparative effect of viable lymph node cells, material obtained after freeze-thawing of equal number of cells from hyperimmune animals and lymph node cells from normal animals.
Fig. 6. Comparative effect of normal viable splenic cells with viable spleen cells of hyperimmune animals.
Fig. 7. Comparative effect of normal freeze thawed splenic cells with freeze thawed spleen cells of hyperimmune animals.
477
et al.
ERECT
OF
IMMUNE
SPLEEN
CELLS
(GL~ARALDEHYDE 70
TREATED) I&CELLS/MOVSE
--DAYS-
Fig. 8. Comparative effect of normal treated spleen cells with glutaraldehyde cells of hyperimmune animal.
glutaraldehyde treated spleen
-DA”l-----
Fig. 9. Comparative effect of normal peritoneal exudate cells with viable peritoneal exudate and freeze thawed peritoneal exudate obtained from immunized animals.
Fig. 10. A diagrammatic presentation of suggested mechanism of the blocking effect of antibodies.
478
CELLMEDIATED
AND HUMORALIMMUNITYINEXPERIMENTAL
viable cells. Two of the animals in this group also died at the time of peak parasitaemia, giving a high mean value for this group on the seventh day. of spleen and macrophages (Figs 6, 7, 8 and 9) Viable spleen cells, frozen and thawed material and glutaraldehyde-treated immune spleen cells, afforded a considerable degree of protection. The maximum parasitaemia observed in all these groups was 5 to I % on day 10. All the animals given glutaraldehydc treated spleen cells and frozen and thawed material from spleen cells survived up to 45 days and were able to clear the infection within 20 to 25 days. When thcsc experiments were extended with hyperimmune macrophages, both viable and frozen and thawed macrophages afforded considerable protection. The peak parasitaemia was observed on day eight in the group given viable macrophages and only one animal died on the 28th day. Similarly in the frozen and thawed macrophage group, peak parasitaemia (7 to 8 %) was observed on day 12. All the animals were able to clear the infection within 20 to 25 days. Effect
Discussion The results presented in this paper are an attempt to elucidate the role of hyperimmune serum and of sensitized lymphoid cells and macrophages in eliminating a fixed dose of parasite inoculum. The results with the serum transfer experiments indicate that whereas hyperimmune serum in doses of 0.1 ml and 0.25 ml per mouse affords a reasonable degree of protection, 0.5 ml per mouse or normal serum caused an enhancement of parasitaemia. When 0.5 ml of hyperimmune serum was used, the results were inconsistent and three animals died although the parasitaemia declined in the other three animals. There was a peak parasitaemia on day six (100 parasitized cells per 500 red cells) and a subsequent fall in the level of parasitaemia (10 to 12 parasitzed cells per 500 red cells on the 14th day) and there was a lag period of three days before onset of parasitaemia when compared to the control group. This initial lag period can be explained on the transfer of humoral antibodies. The subsequent peak on the sixth day and fall on the 14th day appear to be consequences of the transfer of processed antigen. Thus transfer of immunity via hyperimmune serum appears to be a dosedependent phenomenon. There are two possible explanations of this enhancement: (i) the relationship of the antigen and antibody must be very critical and subtle; (ii) blocking of the effective mechanism can be either due to excess of antigen or antibody. The possible pathway of such a blocking action is suggested in Fig. 10. The explanation for the dose-dependent variation in the levels of parasitaemia will have to be further worked out by estimating the actual level of protective antibody in the serum transferred and the presence of antigenantibody complex. Serum soluble plasmoidal antigens were described in the sera of monkeys infected with P. knowlesi as early as 1939 by EATON. Later such antigens were found in a variety of parasite/host combinations by MCGREGOR et al. (1968) in-man-P.‘jLzlcigavum, by Cox et al. (1968) in duck-P. lophurae and in rat-P. ber.hei and SEITZ (1972) in NMRI-mice-P. beughei. It should also be appreciated that only a fraction of antibody formed has a protective function (COHEN & BUTCHER, 1971). It has also been pointed out by COLLINS et al. (1964) that in human P. falcipavum infection, in spite of high antibody
P. berghei
titre, parasites are able to multiply to levels which make chemotherapy imperative. ROBERTS (1968) found it impossible to transfer protection with hyperimmune sera in Babesia rodhaini infections and he concluded that enhancement of protozoal blood infections might bc. due to suppression of active antibody synthesis (UHR & BANWANN 1961, quoted by ROBERTS, 1968) or to protection of parasites by covering antigenic determinants such as may occur in homograft enhancement (MOLLER & MOLLER, 1962, quoted by ROBERTS, 1968). Roberts also suggested that there are two or more types of antibody molecules which function differently, are antagonistic and which may be present in different proportions. The results of transfer of various immune cells, have shown that spleen cells are the most efficient in eliminating the parasite load (TODORONIC et al., 1967; BROWN & PHILLIPS, 1974). Thus adoptive transfer might have been successful by virtue of cells capable of producing antibody of a particular type. The transferred macrophages together with lymphoid cells help in opsonization. Thus immunity may be analogous to homograft rejection and would have been transferred by small lymphacytes together with other transferred spleen cells (GOWANS, MCGREGOR & COWEN, 1963). However, in addition to spleen cells, thymocytes, lymph node cells and peritoneal exudate cells all effectively protected the animals. These results clearly implicate the role of CM1 or helper “T” cells in eliminating infection. PHILLIPS (1969) published similar reports in which he was able to transfer immunity in rats, with viable spleen cells. However, he claimed that viability was essential for transfer of this immunitv. Similarly BROWN (1976) recently has shown no protection with freeze thawed immune spleen cells. In the present study it has been clearly shown that transfer of immunity can be afforded with frozen-thawed lymphnode, spleen and macrophages and glutaraldehyde-treated splenic cells. These results indicate that the frozen thawed material must contain some material which stimulates protective antibodies in the recipients. The significantly better results obtained with viable lymph node cells indicate the additional and more effective protective role of CM1 and co-operative role of “T” cells in affording protection. RICHARDS & KNOWLES 11968) showed that glutaraldehyde solution contains polymeric alpha and beta unsaturated aldehydes. These react with amino groups of proteins and cross link the protein molecules. The modification is not reversible. It is possible that chemical modification of protein enhances the cellular immunity. It has been shown by GRANGER & KOLB (1968) and RU~DEL & WAKSMAN (1969) that immune lymphoid cells may release a toxic factor when they come in contact with specific antigens and there is destruction of target cells. ZEMBALA et al. (1973) have shown that the presence of macrophages as helper cells is essential for lymphocyte mediated cytotoxicity. ALEXANDER et al. (1966) postulated that immune lymphoid cells involve transfer of genetic information through a subcellular component (messenger) which initiates an-immune response. -Recently COLEMAN et al. (1975) have shown the snlenic cell-effected specific lysis of SrCr-labelled erythrocytes from parasitized animals. They concluded that there is more than one cellular cytolytic effector system apparently operative in the mouse. One effector system involved splenic macrophages from normal or immune animals which were increasingly cytotoxic to target cells in the presence of antibody. A second effector system involved nylon-
ARUNA
PARASHAR
purified immune spleen cells which were significantly more cytotoxic than similarly prepared normal spleen cells in the presence of immune serum. Thus it strengthens the concept that immune spleen cells and antibody can interact in a co-operative fashion to mediate cytotoxic reactions in malaria. While it is apparent that viable cells do transfer CMI, there is an additional agent, most likely “processed antigen” produced by sensitized cells-possibly macrophages, which effectively induces immunity and is able to protect the animals. Total population of cells from spleen, lymph node and peritoneal exudate have been used in these transfer experiments. The protective role of thymocytes, indicates that “T” cells must be playing a significant role either directly or indirectly. It will be necessary to isolate the different cells populations like the lymphocytes and macrophages from the spleen, lymph nodes and peritoneal exudate before ascribing a definite independent role to any particular cell type. There appears to be an intricate interplay of multiple factors in affording protection in Plasmodium berghei infection. References Alexander, P., Connell, D. I. & Mikuska, Z. B. (1966). Treatment of murine leukemia with spleen cells or sera from allogenic mice immunized against the tumor. Cancer Research, 26, 1.508. Briggs, B. T., Wellde, B. T. & Sadun, E. H. (1966). Effects of rat antiserum on the course of P. berghei infection in mice. Military Medicine, 131, 1243-1244. Brown, 1. N., Allison, A. C. & Taylor, E. B. (1968). Plasmodium berghei infections in thymectomized rats. Nature,
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I. N. & Phillips, R. S. (1974). Immunity to P. berghei in rats: passive serum transfer and role of the spleen. Infection and Immunity. 10, 1213-1218. Brown,-K. N.,“Jarra, W. & Hills,- i,. A. (1976). T-cell in malaria immunity. Infection and Immunity, 14, 184. Bruce-Chwatt, L. J. & Gibson, F. D. (1956). Transplacental passage of P. berghei and passive transfer of immunity in rats and mice. Transactions of the Royal of Tropical
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Cabrera, E. J. & Alger, N. E. (1971). Transfer of adoptive immunity to P. berghei: a comparison of routes of injection. Journal of Protozoology, 18, 596-597. Cohen, S. & Butcher, G. A. (1970). Properties of protective malarial antibody. Immunolopv, 14, 127-138. Cohen, S. & Butcher, G.-A. (1971). Ser’um’antibody in acquired malarial immunity. Transactions of the Royal Society of Tropical Medicine and Hygiene, 65, 125-135. Cohen, S., McGregor, I. & Carrington, S. (1961). Gamma globulins and acquired immunity in human malaria. Nature,London,
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Coleman, R. M., Rencricca, N. J., Stout, J. P., Brissette, W. H. & Smith, D. M. (1975). Splenic-mediated erythrocyte cytotoxicity in malaria, Ikmunology, 29, 49-54. Collins, W. E., Jeffrey, G. M. & Skinnes, J. C. (1964). Fluorescent antibody studies in human malarial infection. III. Development of antibodies to Plasmodium falciparum in semi-immune patients. American Journal 777-782.
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Diggs, C. L. & Osler, A. (1969). Humoral immunity in rodent malaria. II. Inhibition of parasitaemia by serum antibody. Journal of Immunology, 102,298-305. Eaton, M. D. (1939). The soluble malaria antigen in the serum of mouse. Journal of Experimental Medicine, 69, 517-532. Golenser, J., Spira, D. T. & Zuckerman, A. (1974). Neutralizing antibody in rodent malaria. Tramactions of the Royal 69,251-258.
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Gowans, McGregor, I. A. & Cowen (1963). CIBA Foundation Study Groups, 16, 20. Granger, G. A. & Kolb, W. P. (1968). Lymphocyte in vitro cytotoxicity mechanism of immune and nonimmune small lymphocyte mediated target’ ‘GL” cell destruction. Journal of Immunology, 101, 11 l-120. Gravely, S. M. & Kreier, J. P. (1976). Adoptive transfer of immunity to malaria with T and B lymphocytes. Infection
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Zembala, M., Ptak, W. & Henezakowske, M. (1973). Macrophage and lymphocyte cooperation in target cell destruction in vitro. Clinical and Experimental Immunology, 15, 461. Accepted
for publication
12th May,
1977.
TRANSACTIONS OF THE ROYAL SOCIETY
Cell mediated
OF
TROPICAL MEDICINE AND HYGIENE, VOL. 71, No. 6, 1977.
and humoral immunity in experimental Plasmodium berghei infection
ARUNA PARASHAR, B. K. AIKAT, S. SEHGAL AND S. NAIK Department
of Pathology,
Postgraduate Institute of Medical Chandigarh, India
Summary
Adoptive passive transfer of immunity to Plasmodium berghei infection has been investigated in an inbred strain of Swiss mice. The mice were made hyperimmune by repeated passage of 103 parasites and subsequent therapy with an antimalarial drug. Immune sera and cells obtained from thymus, spleen and peritoneal exudate were transferred to normal animals which were subsequently challenged with standard doses of P. berghei. It was observed that: (a) immune serum in high doses (0.5 ml/mouse) enhanced parasitaemia; when used in smaller doses (0.1 ml/mouse), it afforded a considerable degree of protection; (b) viable immune lymphocytes obtained from thymus and lymph node afforded protection; (c) the mixed population of cells obtained from spleens of immunized mice, as well as peritoneal exudate, protected mice against challenge inoculum; (d) glutaraldehyde-treated spleen cells and material obtained after freezing and thawing the same number of spleen cells, macrophages and lymph node also afforded protection. These findings confirm that, under these experimental conditions, immunity against P. berghei is mediated through (i) specific antibody which is dose-dependent, (ii) cell-mediated immunity and (iii) effective response to processed antigen. Introduction
Immunity to malaria has been extensively investigated. Both humoral, antiplasmodial antibodies and cellmediated immunity have been implicated. The importance of immunoglobulins in conferring immunity in malaria has been suggested repeatedly by passive transfer of immune sera, in humans and in experimental animals (BRUCE-CHWATT & GIBSON, 1956; BRIGGS et al., 1966; DIGGS & OLSER, 1969; COHEN & BUTCHER, 1971; 1974). Protective antiplasmodial GOLENSER et al., globulins act on extracellular merozoites and on mature schizonts. More recent in vitro experiments have shown that antibodies prevent merozoite penetration into the erythrocytes, either by neutralization (COHEN &BUTCHER, 1970) or by agglutination (MILLER et al., 1975). It has also been shown by these workers that transfer of a larger dose of antibody is required to inhibit transiently parasite multiplication. JERUSALEM et al. (1971) and HAMBURGER & KREIER (1976) administered immune serum to rats and observed that there was partial or complete protection against parasitaemia in these animals. The coating of free extraerythrocytic parasites by the antibody has been suggested as the possible mode of action. However, the immune
Education
and Research,
serum does not kill the intracellular parasite, because transfer of infected blood from treated rats into mice, results in multiplication. Thus in vivo control of parasitaemia requires factors other than antibody and the possible role of macrophages has been suggested (BROWN & PHILLIPS, 1974); GOLENSER et al., 1974). Transfer of lymphoid cells from immune animals also protects against subsequent infection (STECHSHULTE, 1969 ; PHILLIPS, 1970; CABRERA & ALGER, 1971). It has been shown that cells from the spleen are more effective in conferring protection (TODORONIC et al., 1967; PHILLIPS. 1970): CABRERA & ALGER. 1970). Thoracic duct lymphocytes, bone marrow cells, thymus cells and peritoneal exudate cells were ineffective when transferred separately (ROBERT & TRACEYPATTE, 1968 ; STECHSHULTE, 1969 ). PHILLIPS (1969) was able to demonstrate protection with immune macrophages but STECHSHULTE (1969 ) could not confirm these findings. T-cell depletion by neonatal thymectomy and using antithymocyte serum made animals more susceptible to infection than were controls (BROWN et al., 1968; STECHSHULTE, 1969 ; SPIRA et al., 1970). Adoptive transfer of immunity by lymphoid cells also prompted workers to propose that immunity against P. berghei infection is dependent on effective CMI. However, BROWN et al. (1976) have shown by adoptive transfer that unfractionated spleen cells and T-cells alone could transfer protection to syngeneic recipients as early as 11 days after infection of the cell donors. They postulated that immune T-cells may act as helper cells and presumably get sensitized to variant specific antigens. Recently GRAVELY & KREIER (1976) have shown that immunity to malarial infection can also be transferred adoptively by the “Null Cells” or antibody forming “B” cells. In view of the controversial data, the present experiments were designed to study the comparative role of passively transferred immune lymphoid cells from different soumes: macrophages and immune sera in experimental P. berghei infection in Swiss inbred strain of mice. Materials
and methods
Animals
Four to six-week-old syngeneic Swiss mice/ICRC weighing 20-25 g were taken and fed ad libitum. Malarial
parasite
Strain of P. berghei obtained from N.I.C.D., New Delhi was used in this experiment. The strain was main-
ARUNA
PAUASHAR
et al.
tained by weekly serial intraperitoneal passage in Swiss inbred mice. The animals were inoculated with erythrocytes from Swiss mice infected with P. berghei. Chronic
malaria
Malaria was induced experimentally in 24 Swiss inbred (male) mice by using 1 x 103 malaria-infected red blood cells as the standard inoculum, chronic&y was maintained by giving three repeated doses of the same number of infected erythrocytes at 17-day intervals and simultaneous therapy with chloroquine (5 mg/kg body-weight). Daily blood smears were made from the tail vein from each animal in duplicate and 500 red cells were counted to determine the percentage of parasitaemia. When parasitaemia reached its peak (nearly 175 infected/500 red cells) on the tenth day, the mice were treated with chloroquine. (Two intraperitoneal injections at a two-day interval) until the parasitaemia came down to 10 to 20 infected cells/500 red cells on day 17. The animals were challenged a second time on the 17th day with the same number of parasites from the same stabilate. The parasitaemia reached its peak on the 27th day-up to 55 to 60 parasitized cells/500 red cells; the animals were treated again with the same dose of chloroquine at the same interval until parasitaemia came down to 10 parasitized cells/ 500 red cells. The third infection was given on day 34 and parasitaemia again recorded daily. It reached its peak of 23 to 25 parasitized cells/500 red cells. The animals were treated with chloroquine until the parasitaemia was completely controlled up to one week. When the animals became aparasitaemic, as confirmed from the peripheral blood smears, they were used as donors in the transfer experiments (Fig. 1). Age and sex matched normal animals were taken as donors for the control groups. Preparation
of lymphoid
cells and collection
of serum
Strictly sterile precautions were taken throughout these experiments. Before killing the animals, blood was collected from each animal, pooled and allowed to clot at 37°C for 30 min. and then centrifuged at 4°C for 30 min. at 1,500 r.p.m. The serum was separated and kept at 4°C until use. Harvesting
of macrophages
After collecting the serum the animals were lightly anaesthetized with ether. The skin was dissected with sterilized forceps and the peritoneum exposed. 5 to 7 ml of Hank’s Balanced Salt Solution (HBSS) were introduced into the peritoneal cavity by means of a syringe and the peritoneum gently massaged. The fluid was carefully withdrawn into the syringe and the cells were washed thrice in cold HBSS and resuspended in the same medium. Viability was checked in trypan blue (NAYSMITH & JAMES, 1968); cell counts were made in a haemacytometer and adjusted to the desired concentration for injection (5.5 x 106 cell/mouse). The percentage of nonviable cells was usually between five and ten. Preparation
of cell suspension
Spleens, lymph nodes and thymi were taken out under sterile conditions. teased in cold HBSS and then oassed through a 60 mesh gauze. The cells were centrifuged at 1,500 r.p.m. at 4°C for 10 min. and washed four times. The viability was tested with trypan blue; cells were counted and adjusted at 70 x 106 cells/05 ml/mouse. Cells were collected similarly from normal animals.
Fig. 1. Graph donors.
Freeze
showing
mean blood parasitaemia
in
thawing
Spleens, lymph nodes, thymi and peritoneal exudate from immune and normal animals were collected in cold HBSS under sterile conditions: teased in cold HBSS and then passed through a 60 mesh gauze. The cells were centrifuged at 1,500 r.p.m. at 4°C for 10 min. and washed four times with cold HBSS. The viability was tested with trypan blue. Cells were counted and adjusted to 70 x 106 cells/O.5 ml/mouse, kept at -70°C for one hour and thawed at 37°C in a water bath. Freezing and thawing the cells was repeated three times. After the final thawing the tubes were centrifuged at 1,500 r.p.m. for 10 min. and the supernatant was used for inoculation. A similar procedure was used with macrophages but the number of cells was adjusted to 5.5 x 106 cells/mouse. Glutaraldehyde
fixation
The method of SANDERSON & FROST (1974) was used for glutaraldehyde fixation. Spleens from normal and immune donors were removed under sterile conditions in HBSS containing antibiotics. The tissue was homogenized and the cell suspension was sieved through a fine mesh. Cells were washed twice in HBSS and the final count was adjusted to 1 x 108 cells/ml in MEM. An equal volume of 0.25 “/, glutaraldehyde (obtained from Me&h-Schuchardt M& then) in MEM was added. The cells were kept for 15 min. at 22°C. They were subsequently washed three times and centrifuged at 1,000 r.p.m. for 10 min. in a cold MSE centrifuge. Each mouse received 70 x 106 glutaraldehyde treated cells. After 24 hours the animals were challenged with parasites. Control group of age and sex matched recipients received normal cells treated with glutaraldehyde as mentioned. Transfer
experiment
Serum was transferred into three different groups each group having six test and four control animals. In the test animals, 0.5, 0.25 and 0.1 ml of the hyperimmune serum was injected intraperitoneally into the three groups on day 0. After 24 hours the animals were challenged with 1 x 103 malaria-infected red cells and parasitaemia was recorded daily in Giemsa-stained smears; the control animals were given the same dose of normal serum and challenged with the same number of parasites after 24 hours. No drug therapy was given to either the control
476
CELLMEDIATED
AND
P.berghei
HUMORALIMMUNITYINEXPERIMENTAL
or the test group after transferring immune or normal serum. Lymphoid
cells transfer
Lymphoid cells from immune lymph nodes were given to two groups. One group of six mice received viable cells (70 x 106 cells/mouse) and the other group (also of six mice) received material from the same number of frozen and thawed cells. They were challenged with 1 x 103 malaria-infected red cells and parasitaemia was recorded daily. Thymocytes were also transferred in the same dose schedule as lymph node cells. In the case of the spleen, three groups (each of six animals) were used. Viable cells were administered to the first, frozen and thawed material from an equal number of cells to the second and glutaraldehyde-treated cells to the third group. With macrophages, since the number of cells obtained in peritoneal exudate was inadequate only 5.5 x 106 cells per mouse were given. The same number of cells were disrupted by freeze thawing and the material obtained was injected into the second group of six animals. These groups were challenged with parasites. The control received the same number of cells from. normal animals.
Fig. 2. Comparative effect on parasitaemia of 0.5 ml of hyperimmune serum and serum obtained from normal animals.
Results Protective
activity
in serum
Normal serum recipients and those to which no serum was given when challenged with parasites showed parasitaemia in one or two days. They died of fulminant infection within two to three weeks. The results of injecting hyperimmune serum in different doses are presented in Figs 2 and 3. It was apparent that when 0.5 ml of hyperimmune serum was used, there was marked enhancement of parasitaemia from the fourth day onwards which reached 38% on day 14. All the animals given 0.5 ml of hyperimmune serum died within 25 days. However, when 0.1 ml of hyperimmune-serum was used the results were inconsistent and three animals died while the parasitaemia declined in the other three. There was peak parasitaemia on day six (20%) and a subsequent fall in the level of parasitaemia (approximately 25% on the 14th day). Parasitaemia was significantly lower in the group given 0.25 ml of hyperimmune serum than in the control group. of thymus cells (Fig. 4) Thymocytes afforded considerable protection and parasitaemia remained at very low levels all throughout the experiment. All the animals survived up to 45 days and the maximum parasitaemia of 5% was seen on the eighth day. Normal thymocytes did not afford protection when the mice were subsequently challenged with parasites.
Fig. 3. Comparative effect on parasitaemia of 0.1 ml, 0.25 ml of hyperimmune serum and serum obtained from normal animals.
Effect
of immune lymph node cells (Fig. 5) When viable lymph node cells were inoculated, the maximum parasitaemia of 6 to 7 % was observed on day 10. All the animals were able to clear the infection within 20 days and no mortality was observed up to 45 days. Similarly, when the same number of lymphoid cells was disrupted by freezing and thawing and the material thus obtained was injected into the host, it did afford protection in four of six animals subsequently challenged but significantly less than did viable cells. The maximum parasitaemia seen in these mice was approximately 15 % on day seven compared to 7 ‘A on day 10 in those given
EFFECT
OF
THYMOCYTES
ON
PARASITAEMIA.
70X106MO~s~
Effect
Fig. 4. Comparative effect on parasitaemia after injecting viable thymocytes from immune and control animals.
ARUNA
EFFECT
OF
LYMPH
NODE
--
CELLS 70 Xl&
PARASHAR
ON PARASITAEMIA CELLS/MOUSE
OAY-
Fig. 5. Comparative effect of viable lymph node cells, material obtained after freeze-thawing of equal number of cells from hyperimmune animals and lymph node cells from normal animals.
Fig. 6. Comparative effect of normal viable splenic cells with viable spleen cells of hyperimmune animals.
Fig. 7. Comparative effect of normal freeze thawed splenic cells with freeze thawed spleen cells of hyperimmune animals.
477
et al.
ERECT
OF
IMMUNE
SPLEEN
CELLS
(GL~ARALDEHYDE 70
TREATED) I&CELLS/MOVSE
--DAYS-
Fig. 8. Comparative effect of normal treated spleen cells with glutaraldehyde cells of hyperimmune animal.
glutaraldehyde treated spleen
-DA”l-----
Fig. 9. Comparative effect of normal peritoneal exudate cells with viable peritoneal exudate and freeze thawed peritoneal exudate obtained from immunized animals.
Fig. 10. A diagrammatic presentation of suggested mechanism of the blocking effect of antibodies.
478
CELLMEDIATED
AND HUMORALIMMUNITYINEXPERIMENTAL
viable cells. Two of the animals in this group also died at the time of peak parasitaemia, giving a high mean value for this group on the seventh day. of spleen and macrophages (Figs 6, 7, 8 and 9) Viable spleen cells, frozen and thawed material and glutaraldehyde-treated immune spleen cells, afforded a considerable degree of protection. The maximum parasitaemia observed in all these groups was 5 to I % on day 10. All the animals given glutaraldehydc treated spleen cells and frozen and thawed material from spleen cells survived up to 45 days and were able to clear the infection within 20 to 25 days. When thcsc experiments were extended with hyperimmune macrophages, both viable and frozen and thawed macrophages afforded considerable protection. The peak parasitaemia was observed on day eight in the group given viable macrophages and only one animal died on the 28th day. Similarly in the frozen and thawed macrophage group, peak parasitaemia (7 to 8 %) was observed on day 12. All the animals were able to clear the infection within 20 to 25 days. Effect
Discussion The results presented in this paper are an attempt to elucidate the role of hyperimmune serum and of sensitized lymphoid cells and macrophages in eliminating a fixed dose of parasite inoculum. The results with the serum transfer experiments indicate that whereas hyperimmune serum in doses of 0.1 ml and 0.25 ml per mouse affords a reasonable degree of protection, 0.5 ml per mouse or normal serum caused an enhancement of parasitaemia. When 0.5 ml of hyperimmune serum was used, the results were inconsistent and three animals died although the parasitaemia declined in the other three animals. There was a peak parasitaemia on day six (100 parasitized cells per 500 red cells) and a subsequent fall in the level of parasitaemia (10 to 12 parasitzed cells per 500 red cells on the 14th day) and there was a lag period of three days before onset of parasitaemia when compared to the control group. This initial lag period can be explained on the transfer of humoral antibodies. The subsequent peak on the sixth day and fall on the 14th day appear to be consequences of the transfer of processed antigen. Thus transfer of immunity via hyperimmune serum appears to be a dosedependent phenomenon. There are two possible explanations of this enhancement: (i) the relationship of the antigen and antibody must be very critical and subtle; (ii) blocking of the effective mechanism can be either due to excess of antigen or antibody. The possible pathway of such a blocking action is suggested in Fig. 10. The explanation for the dose-dependent variation in the levels of parasitaemia will have to be further worked out by estimating the actual level of protective antibody in the serum transferred and the presence of antigenantibody complex. Serum soluble plasmoidal antigens were described in the sera of monkeys infected with P. knowlesi as early as 1939 by EATON. Later such antigens were found in a variety of parasite/host combinations by MCGREGOR et al. (1968) in-man-P.‘jLzlcigavum, by Cox et al. (1968) in duck-P. lophurae and in rat-P. ber.hei and SEITZ (1972) in NMRI-mice-P. beughei. It should also be appreciated that only a fraction of antibody formed has a protective function (COHEN & BUTCHER, 1971). It has also been pointed out by COLLINS et al. (1964) that in human P. falcipavum infection, in spite of high antibody
P. berghei
titre, parasites are able to multiply to levels which make chemotherapy imperative. ROBERTS (1968) found it impossible to transfer protection with hyperimmune sera in Babesia rodhaini infections and he concluded that enhancement of protozoal blood infections might bc. due to suppression of active antibody synthesis (UHR & BANWANN 1961, quoted by ROBERTS, 1968) or to protection of parasites by covering antigenic determinants such as may occur in homograft enhancement (MOLLER & MOLLER, 1962, quoted by ROBERTS, 1968). Roberts also suggested that there are two or more types of antibody molecules which function differently, are antagonistic and which may be present in different proportions. The results of transfer of various immune cells, have shown that spleen cells are the most efficient in eliminating the parasite load (TODORONIC et al., 1967; BROWN & PHILLIPS, 1974). Thus adoptive transfer might have been successful by virtue of cells capable of producing antibody of a particular type. The transferred macrophages together with lymphoid cells help in opsonization. Thus immunity may be analogous to homograft rejection and would have been transferred by small lymphacytes together with other transferred spleen cells (GOWANS, MCGREGOR & COWEN, 1963). However, in addition to spleen cells, thymocytes, lymph node cells and peritoneal exudate cells all effectively protected the animals. These results clearly implicate the role of CM1 or helper “T” cells in eliminating infection. PHILLIPS (1969) published similar reports in which he was able to transfer immunity in rats, with viable spleen cells. However, he claimed that viability was essential for transfer of this immunitv. Similarly BROWN (1976) recently has shown no protection with freeze thawed immune spleen cells. In the present study it has been clearly shown that transfer of immunity can be afforded with frozen-thawed lymphnode, spleen and macrophages and glutaraldehyde-treated splenic cells. These results indicate that the frozen thawed material must contain some material which stimulates protective antibodies in the recipients. The significantly better results obtained with viable lymph node cells indicate the additional and more effective protective role of CM1 and co-operative role of “T” cells in affording protection. RICHARDS & KNOWLES 11968) showed that glutaraldehyde solution contains polymeric alpha and beta unsaturated aldehydes. These react with amino groups of proteins and cross link the protein molecules. The modification is not reversible. It is possible that chemical modification of protein enhances the cellular immunity. It has been shown by GRANGER & KOLB (1968) and RU~DEL & WAKSMAN (1969) that immune lymphoid cells may release a toxic factor when they come in contact with specific antigens and there is destruction of target cells. ZEMBALA et al. (1973) have shown that the presence of macrophages as helper cells is essential for lymphocyte mediated cytotoxicity. ALEXANDER et al. (1966) postulated that immune lymphoid cells involve transfer of genetic information through a subcellular component (messenger) which initiates an-immune response. -Recently COLEMAN et al. (1975) have shown the snlenic cell-effected specific lysis of SrCr-labelled erythrocytes from parasitized animals. They concluded that there is more than one cellular cytolytic effector system apparently operative in the mouse. One effector system involved splenic macrophages from normal or immune animals which were increasingly cytotoxic to target cells in the presence of antibody. A second effector system involved nylon-
ARUNA
PARASHAR
purified immune spleen cells which were significantly more cytotoxic than similarly prepared normal spleen cells in the presence of immune serum. Thus it strengthens the concept that immune spleen cells and antibody can interact in a co-operative fashion to mediate cytotoxic reactions in malaria. While it is apparent that viable cells do transfer CMI, there is an additional agent, most likely “processed antigen” produced by sensitized cells-possibly macrophages, which effectively induces immunity and is able to protect the animals. Total population of cells from spleen, lymph node and peritoneal exudate have been used in these transfer experiments. The protective role of thymocytes, indicates that “T” cells must be playing a significant role either directly or indirectly. It will be necessary to isolate the different cells populations like the lymphocytes and macrophages from the spleen, lymph nodes and peritoneal exudate before ascribing a definite independent role to any particular cell type. There appears to be an intricate interplay of multiple factors in affording protection in Plasmodium berghei infection. References Alexander, P., Connell, D. I. & Mikuska, Z. B. (1966). Treatment of murine leukemia with spleen cells or sera from allogenic mice immunized against the tumor. Cancer Research, 26, 1.508. Briggs, B. T., Wellde, B. T. & Sadun, E. H. (1966). Effects of rat antiserum on the course of P. berghei infection in mice. Military Medicine, 131, 1243-1244. Brown, 1. N., Allison, A. C. & Taylor, E. B. (1968). Plasmodium berghei infections in thymectomized rats. Nature,
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I. N. & Phillips, R. S. (1974). Immunity to P. berghei in rats: passive serum transfer and role of the spleen. Infection and Immunity. 10, 1213-1218. Brown,-K. N.,“Jarra, W. & Hills,- i,. A. (1976). T-cell in malaria immunity. Infection and Immunity, 14, 184. Bruce-Chwatt, L. J. & Gibson, F. D. (1956). Transplacental passage of P. berghei and passive transfer of immunity in rats and mice. Transactions of the Royal of Tropical
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Cabrera, E. J. & Alger, N. E. (1971). Transfer of adoptive immunity to P. berghei: a comparison of routes of injection. Journal of Protozoology, 18, 596-597. Cohen, S. & Butcher, G. A. (1970). Properties of protective malarial antibody. Immunolopv, 14, 127-138. Cohen, S. & Butcher, G.-A. (1971). Ser’um’antibody in acquired malarial immunity. Transactions of the Royal Society of Tropical Medicine and Hygiene, 65, 125-135. Cohen, S., McGregor, I. & Carrington, S. (1961). Gamma globulins and acquired immunity in human malaria. Nature,London,
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Coleman, R. M., Rencricca, N. J., Stout, J. P., Brissette, W. H. & Smith, D. M. (1975). Splenic-mediated erythrocyte cytotoxicity in malaria, Ikmunology, 29, 49-54. Collins, W. E., Jeffrey, G. M. & Skinnes, J. C. (1964). Fluorescent antibody studies in human malarial infection. III. Development of antibodies to Plasmodium falciparum in semi-immune patients. American Journal 777-782.
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Diggs, C. L. & Osler, A. (1969). Humoral immunity in rodent malaria. II. Inhibition of parasitaemia by serum antibody. Journal of Immunology, 102,298-305. Eaton, M. D. (1939). The soluble malaria antigen in the serum of mouse. Journal of Experimental Medicine, 69, 517-532. Golenser, J., Spira, D. T. & Zuckerman, A. (1974). Neutralizing antibody in rodent malaria. Tramactions of the Royal 69,251-258.
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Ruddel, N. M. & Waksman, B. H. (1967). Cytotoxic effect of lymphocyte antigen interaction in delayed hypersensitivity. Science, 157, 1060. Sanderson, C. J. & Frost, P. (1974). Induction of tumour immunity using glutaraldehyde treated tumour cells. Nature,
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Stechschulte, D. J. (1969b). Cell-mediated immunity in rats injected with Plasmodium berghei. Military Medicine, 134, 1147-l 152. Todoronic, R., Ferri, D. & Ristic, M. (1967). Role of spleen in plasmodial and babesial infections in rats. Experimental
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P. berghei
Zembala, M., Ptak, W. & Henezakowske, M. (1973). Macrophage and lymphocyte cooperation in target cell destruction in vitro. Clinical and Experimental Immunology, 15, 461. Accepted
for publication
12th May,
1977.
