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* Division of Parasitology, National Institute for Medical Research, London, United Kingdom; and
Medical Research Council Center for Immune Regulation, University of Birmingham, Edgbaston, Birmingham, United Kingdom
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Abstract |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Although Ab appears to be important in the blood stages of human malaria, immunity only develops after repeated infections and is lost without continued exposure to infection. Longitudinal studies in malaria-exposed human populations show that Ab levels to several important malarial proteins drop rapidly at the end of the malaria transmission season (6, 7, 8, 9, 10). This drop in Ab levels is also seen after people have left the transmission area (11). These data contrast with Ab responses of noninfected mice to nonrenewable protein Ags that can be sustained for years (12, 13). In mice, splenic B cells able to respond to infected erythrocytes in vitro can still be found up to 1 year after infection with P. chabaudi chabaudi, but their number is significantly reduced compared with the response 3 mo after infection (14). Together, these data indicate that the generation or maintenance of memory B cells and long-lived plasma cells could be impaired in malaria infection.
One way of examining B cell memory generation is to assess the balance between the follicular and extrafollicular B cell response. T-independent Ab responses, which typically induce extrafollicular Ab responses, are not associated with functional germinal centers or memory formation (15). Under exceptional conditions, T cell-independent germinal centers can form but do not complete their development (16). By contrast, T-dependent Ags, which produce both extrafollicular responses and germinal centers, are associated with affinity maturation and sustained Ab responses (17, 18). The differentiation between the follicular and extrafollicular pathways is performed by studying the four major B cell populations of the spleen: naive recirculating B cells, which are mainly located in the follicular mantle of the white pulp; germinal center B cells as the source of memory B cells and long-lived plasma cells; plasma cells and their plasmablast precursors; and marginal zone B cells, which include both naive and memory cells and give rise to rapid extrafollicular thymus-dependent and thymus-independent Ab responses (19, 20, 21).
In the present study, C57BL/6 mice were infected with P. chabaudi chabaudi (AS), and the resulting changes in the splenic architecture as well as the four major B cell populations of the spleen were studied by immunohistology and flow cytometry. We conclude that there are major structural changes in the spleen, but these reflect an ordered response to the parasite rather than a salvage response to indiscriminate damage.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Female C57BL/6 mice bred in the specific pathogen-free unit at the National Institute for Medical Research (London, U.K.) were used at 6–12 wk of age. They were conventionally housed on sterile bedding, food, and water.
Parasites and infection
P. chabaudi chabaudi clone AS was routinely injected from frozen stocks. Further infections were initiated by i.p. injection of 105 parasitized RBCs obtained from infected mice before the peak of parasitemia, and the infection monitored by Giemsa-stained thin blood films as previously described (22).
Immunization with chicken 
-globulin (CGG)
3
CGG (1 mg/ml; The Jackson Laboratory, Bar Harbor, ME) was mixed 1:1 with 9% potassium-aluminum-sulfate and precipitated with 1 M sodium hydroxide. Mice were immunized i.p. with 25 µg alum-precipitated Ag in 100-µl saline solution.
Abs and other reagents for immunohistology and flow cytometry
The following Abs and reagents were used in immunohistology: rat anti-mouse CD3 (KT3) and rat anti-mouse IgM H chain (LO-MM-9) (Serotec, Oxford, U.K.); rat anti-mouse syndecan-1 (CD138, 09341D; BD PharMingen, Oxford, U.K.); biotinylated peanut agglutinin (PNA; Vector Laboratories, Burlingame, CA); sheep anti-mouse IgD and HRP-labeled donkey anti-sheep/goat Ig (The Binding Site, Birmingham, U.K.); biotinylated rabbit anti-rat Ig and alkaline phosphatase (AP)-labeled StreptABC complex (DAKO, Cambridgeshire, U.K.).
The Abs for flow cytometry were obtained from BD PharMingen: FITC-labeled anti-mouse CD21/CD35 (7G6), anti-mouse IgD (11-26c.2a), and anti-mouse GL7 (GL7); PE-labeled anti-mouse CD23 (B3B4), anti-mouse CD19 (1D3), and anti-mouse syndecan-1 (CD138; 281-2); biotinylated anti-mouse CD19 (1D3). Streptavidin-Tricolor was from Caltag Laboratories (Burlingame, CA). Rat anti-mouse B220 (RA3 3A1) (23) and rat anti-mouse Fc receptor (2.4G2) Abs (24) were purified from hybridoma supernatants.
Tissue preparation for immunohistology
Spleens were snap frozen by repeated dipping in liquid nitrogen and stored at -70°C in gripseal plastic bags. The enlarged spleens from later stages of infection were halved before freezing. The spleens were embedded in cold OCT compound (Sakura, Zoeterwede, The Netherlands), and serial 5-µm cryosections were mounted on four spot-glass slides (Hendley Essex, Loughton, U.K.). Slides were allowed to air dry at room temperature for at least 1 h. They were fixed in 90% acetone at 4°C for 20 min, air dried, and stored in gripseal bags at -20°C until use.
Immunohistology
Immunohistology was performed as previously described (25). In brief, primary Abs were added to thawed sections for 1 h at room temperature, followed by washing and HRP- or biotin-labeled secondary Abs for 45 min. After another wash, AP-labeled StreptABC complex was added. HRP activity was detected with diaminobenzidine tetrahydrochloride solution containing hydrogen peroxide to inhibit endogenous peroxidases. AP activity was detected using the substrate Naphthol AS-MX phosphate (0.4 mg/ml) and the chromogen Fast Blue BB salt (1 mg/ml, Sigma-Aldrich, Poole, U.K.) in 50 mM TBS (pH 9.2), containing 0.8 mg/ml levamisole (Sigma-Aldrich) to inhibit endogenous phosphatases, and 3.8% v/v N,N-dimethylformamide (Sigma-Aldrich). The slides were then mounted in Immuno-Mount (Thermo Shandon, Pittsburgh, PA).
Size measurements in histological sections
The size of the germinal centers (PNA+ areas in the follicular mantle) was determined by systematic perusal of the entire spleen section through the grid of the microscope ocular while counting the number of grid intersections that fell within the germinal center. This measurement was converted into millimeters squared by comparison with the grid of a Neubauer counting chamber at the same magnification. Where spleens had to be halved for cryosectioning, germinal center numbers were extrapolated for whole spleens.
H&E staining of splenic sections
Spleens were fixed for 24 h in 10% neutral buffered formalin, embedded in fibrowax (BDH, Poole, U.K.), sectioned, and then stained with H&E, dehydrated in graded alcohols, cleared with Histoclear (National Diagnostics, Hull, U.K.), and mounted in DPX (BDH).
Flow cytometry
Single-cell suspensions of spleens were prepared, the erythrocytes lysed with 0.16 M NH4Cl and the cells resuspended in FACS buffer (1% weight to volume ratio BSA, 5 mM EDTA, 0.01% sodium azide in PBS, or FACS buffer without sodium azide for syndecan-1 staining). Cells (5 x 105) were incubated with appropriately diluted Abs in the presence of anti-Fc receptor Ab to prevent Ab binding via the Fc receptor. Samples were acquired on a FACSCalibur and analyzed with CellQuest software (BD Biosciences, San José, CA).
Cell death
7-Aminoactinomycin D (7-AAD, Sigma-Aldrich) was used to detect apoptotic cells, with the necrotic cell marker propidium iodide used in parallel to ascertain that the majority of the 7-AAD+ cells were actually apoptotic. Splenocytes were stained as described in the previous section. In the final step, either 7-AAD (1 µg/2.5 x 105 cells) or propidium iodide (1 µg/2.5 x 105 cells; Sigma-Aldrich) was added for 20 or 5 min, respectively. Cells were then analyzed on the FACSCalibur without further washes.
ELISA
Total IgG and malaria-specific IgG were measured using affinity-purified goat anti-mouse IgG (Southern Biotechnology Associates, Birmingham, AL) and crude malaria extract (22) as coating reagents. IgG was revealed with AP-labeled goat anti-mouse IgG Abs (Southern Biotechnology Associates) and p-nitrophenyl phosphate as described (1, 22). Normal plasma was used as a negative control, and plasma from P. chabaudi chabaudi immune mice was used as a standard (22). Data are expressed as milligrams per milliliter and arbitrary units (AU), based on a immune plasma value of 100, respectively, for total IgG and malaria-specific Abs.
Statistical analysis
Data were analyzed by a two-tailed Student t test using GraphPad Instat software (San Diego, CA). Values of p < 0.05 were considered to be significantly different.
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Results |
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P. chabaudi chabaudi (AS) infection with 105 parasitized RBCs becomes patent in C57BL/6 mice on day 3 of infection, following which parasitemia rises exponentially, peaking on day 9 or day 10 of infection with up to 30% of erythrocytes infected. Parasitemia then drops rapidly, followed by a much lower recrudescence of 
1% at day 20–25 and becomes undetectable around day 30–35 (Fig. 1a). The infection is accompanied by splenomegaly with spleen weights increasing more than 10-fold (e.g., Ref.26) and nucleated splenocyte numbers increasing 8-fold (Fig. 1b). There is a 3-fold increase in total plasma IgG peaking at 14 days of infection (6.14 ± 0.53 mg/ml compared with 2.32 ± 0.10 mg/ml in normal plasma, p < 0.01, n = 5) and levels remain elevated at 28 days (5.558 ± 0.22 mg/ml compared with normal plasma, p < 0.01, n = 5). Malaria-specific IgG was detectable only after 20 days of infection (day 20, 3 ± 0.7 AU; day 28, 30 ± 10 AU).
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The four major B cell populations of the spleen were examined during a course of infection with P. chabaudi chabaudi. Immunohistology was performed to determine: 1) the distribution of T cells and naive recirculating B cells (also referred to as follicular B cells) by staining for CD3 and IgD (Fig. 1c); 2) the extrafollicular Ab response by staining cells committed to plasma cell differentiation with syndecan-1 (CD138) (Fig. 1d); 3) the development and persistence of germinal centers by staining for IgD and CD3, where germinal centers appear as unstained areas in the follicles (Fig. 1c), or positive staining of germinal centers for PNA binding (Fig. 2a); and 4) the involvement of marginal zone B cells by IgM staining in the marginal zones (Fig. 2c). Parallel studies were conducted using flow cytometry.
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Red pulp changes
Before infection, the red pulp contains many widely spaced IgD+ and CD3+ cells (day 0, Fig. 1c). On days 8 and 10 there has been a massive increase in the number of Ab-containing cells in the red pulp (days 8 and 10, Fig. 1d). Some of these contain IgM, but many have switched to IgG (data not shown). Between days 10 and 20 of infection, the red pulp appears devoid of IgD+ and CD3+ lymphocytes (day 10 and 20, Fig. 1c). Flow cytometric analyses show that the numbers of naive recirculating B cells (Fig. 3b) and of T cells (27) are not decreased at the peak of infection, suggesting that the lymphocytes are just more dispersed in the enlarged spleens. To assess the type of cells occupying the expanded red pulp, H&E staining was performed. There is a massive but transient increase in the level of hematopoiesis in the red pulp in agreement with previous observations in P. yoelli infections (28). Large islands of erythroid precursors and megakaryocytes were observed at day 10 (Fig. 2f). Macrophage densities were not increased according to F4/80 or nonspecific esterase staining (data not shown).
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The location of the four major splenic B cell populations (naive recirculating B cells, germinal center B cells, Ab-containing cells, marginal zone B cells) during infection was determined by immunohistology. Small numbers of PNA-positive germinal centers are present in the spleens of uninfected control mice (Table I). Slightly increased numbers of small germinal centers are seen on days 7–9, and by day 10 the germinal center response is firmly established (Table I, Figs. 1c and 2a), representing the most constant element in the changing white pulp structure. The germinal centers continue to increase in size and number up to day 20. Normal germinal centers are present up to day 40 and still visible on day 60 in a more weakly stained form. Germinal centers are slightly larger and more frequent on day 10 of a P. chabaudi chabaudi infection than after immunization with a conventional Ag, alum-precipitated CGG, and at day 20 of infection, the number and average size of the germinal centers are significantly higher those seen 10 days after CGG immunization (Table I, p < 0.05).
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At day 10 of infection, the majority of the syndecan-1+ cells are located in the T cell zone of the white pulp (day 10, Fig. 1d) rather than the red pulp. Between day 12 and at least day 20 of infection (day 20, Fig. 2, d and e, and Fig. 1d), as the total number of syndecan-1+ cells begins to decrease and more T cells are seen in the T zone, T cells and Ab-containing cells occupy nonoverlapping areas of the T zone. By day 30, the Ab-containing cells are again found in the red pulp (data not shown) with numerous foci still visible at day 60 (day 60, Fig. 1d).
Marginal zone B cells (identified in immunohistology as IgM+IgD-) are normally located at the edge of the white pulp, as can be seen in the spleens of uninfected mice (day 0, Fig. 2c). After infection, the marginal zone B cells decrease in number, and after day 10 are no longer detectable in the marginal zones (day 10, Fig. 2c). The cells may have migrated to other parts of the spleen, but immunohistology cannot distinguish between IgM+ plasma cells and IgM+ B cells of marginal zone origin without the additional locational indicator. By day 40, the marginal zones contain IgM+ B cells again, but even at day 60 they form a less continuous structure than before the disruption (data not shown).
Quantification of splenic B cell populations
Due to the large increase in spleen size during infection, which involves unequal expansion of the red and white pulp, an increase in a cell population may be masked by the overall increase in spleen size. We therefore analyzed the four major splenic B cell populations quantitatively by flow cytometry. The Ab combinations used to define the B cell populations are shown for two representative time points of infection and compared with uninfected control mice in Fig. 3. Abs against CD19 (Fig. 3a) were used to determine total B cell numbers. The naive follicular B cells are defined as CD21intCD23+ (Fig. 3b, labeled FO), while marginal zone B cells are CD21+CD23- (Fig. 3c, labeled MZ) (20). Germinal center B cells were identified by positive staining with Abs against GL7 and CD19 (Fig. 3d, labeled GC), as there was a considerable level of nongerminal center binding of PNA in the infected spleens. Ab-containing cells (plasma cells and plasmablasts) are defined as syndecan-1+CD19int (Fig. 3e, labeled PC).
The graphs in Fig. 3 show how the relative population sizes, as determined by flow cytometry, translate into absolute numbers for the different B cell populations. The total number of CD19+ B cells is greatest at day 8 of infection when there are roughly twice as many B cells as in uninfected mice (Fig. 3a, p = 0.0012). After that, numbers drop slightly and fluctuate around a value that is still significantly higher than that in uninfected mice (p < 0.01 for days 10, 20, 30, and 40 postinfection). The naive recirculating B cell population (CD21intCD23+) grows in the early and late stages of infection, but even at the peak of infection, numbers are not significantly lower than in naive mice (Fig. 3b). Thus, the marked transient reduction of these cells in the follicular mantles observed by immunohistology between days 8 and 12 is not reflected in the total population size.
The total numbers of germinal center B cells and Ab-containing cells agree well with the immunohistological observations. The numbers of germinal center B cells (GL7+CD19+) rise between days 3 and 5 postinfection, with significant increases above uninfected controls by day 6 (Fig. 3d, p < 0.01). Numbers are maximal on day 20 and remain elevated relative to uninfected controls up to day 40 of infection (p < 0.01 comparing day 0 with days 10–40) and even at this late stage, levels are five times higher than at day 10 after CGG immunization (p < 0.01 comparing day 40 postinfection with CGG-immunized mice).
Ab-containing (syndecan-1+) cell numbers increase even more sharply (Fig. 3e) with the peak levels at day 10 of infection 300-fold higher than uninfected levels (p < 0.01 comparing day 0 with days 8–12 postinfection) and over 100-fold higher than on day 10 after CGG immunization (p < 0.0001). After that, levels drop, but are still over four times higher than uninfected levels 60 days after infection (p < 0.01).
The marginal zone B cell population exhibits a strong peak at days 5 and 6 of infection, with significantly higher numbers than those observed in uninfected mice (p < 0.01) followed by a sharp decline at day 7 and a gradual decrease up to day 16 (Fig. 3c). After that, the population regains and maintains a level significantly higher than the uninfected level for days 16, 30, and 40 (p < 0.05, 0.01, and 0.05, respectively). This contrasts with the apparent complete loss of marginal zone B cells seen in immunohistology, suggesting that the marginal zone B cells have migrated into other parts of the spleen.
Apoptosis
Previous studies have shown that malarial infection causes large numbers of host cells to undergo apoptosis (26, 27). To determine whether apoptosis preferentially affects any particular population of B cells, CD19+ B cells, germinal center B cells, and marginal zone B cells were labeled with 7-AAD. Splenocytes from days 8, 9, and 10 of infection were studied, as the parasite density and splenic disruption are highest during this period.
Although the total number of apoptotic CD19+, germinal center, and marginal zone B cells increases 5-fold, 24-fold, and 2-fold, respectively, by day 8 or day 9 of infection (Table II), there were no significant differences in the percentages of apoptotic cells in any of the B cell populations when compared with control uninfected mice (p > 0.05 for all B cell populations). This suggests that the increase in total numbers of apoptotic B cells is entirely explained by the size increase in these populations during infection (see Fig. 3). Examination of the H&E-stained sections from days 8 and 10 of infection supports this notion, as clusters of cells with apoptotic nuclei (condensed chromatin) were only observed in the germinal centers, where apoptosis is an integral part of affinity-based B cell selection (Fig. 2b).
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Discussion |
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The disturbance of the B and T cell areas does not appear to affect either generation of germinal centers or the production of Ab-containing cells (plasma cells and plasmablasts) in the spleen. This finding suggests that organized primary B cell follicles and T cell areas may not be necessary for the generation of germinal centers and supports previous findings that B cells can colocalize with the follicular dendritic cell network and develop fully functional germinal centers in mice incapable of forming primary splenic B cell follicles (32).
The kinetics of germinal center appearance after infection are similar to those observed after immunization with classical hapten Ags (e.g., (4-hydroxy-3-nitrophenyl)acetyl nitropropene), where PNA+ germinal centers are visible within 8 days of immunization (33). The lifespan of germinal centers depends on the duration of Ag exposure, and hapten-specific germinal centers are present in the spleen for 3 wk after immunization (e.g., Ref.19). However, germinal centers in the spleens of P. chabaudi-infected mice can persist for at least 60 days, which is reminiscent of the persistence of germinal centers at least 4 mo after murine mammary virus infection (25). In visceral leishmaniasis on the other hand, which resembles malaria in causing splenomegaly and long parasite persistence, germinal centers diminish after 4 wk of infection in parallel with destruction of follicular dendritic cells (34). This shows that closer examination of Ag-processing cells and APC during malaria infection must still be performed to understand better the functioning of the B cells.
These results indicate that if B cell memory is impaired it is not through a deficiency in germinal center formation or maintenance. Other possible causes for the reported short-lived Ab responses or poor memory responses (6, 7, 8, 9, 10), such as ablation of specific B cells by soluble Ag (35, 36) or generation of short-lived rather than long-lived plasma cells are currently being investigated. It has been shown previously that large amounts of soluble Ag can induce apoptosis of specific B cells in germinal centers and thus affect affinity maturation and B cell memory (35, 36). Because malaria infections generate a lot of circulating Ag (37), this would be an attractive mechanism of interference with memory development in malaria. However, we found no evidence that enhanced apoptosis or reduction in germinal center size takes place in germinal centers during the acute infection.
We have also shown that plasmablast and plasma cell numbers expand greatly up to the peak of the infection and then rapidly disappear, which fits with the spleen’s limited capacity to support continued plasma cell survival (38). Most plasma cells generated during an immune response are short lived, with only a small proportion of long-lived cells remaining in the spleen or migrating to the bone marrow (39, 40, 41). Increased plasma cell numbers were still found in the red pulp 60 days after the mice were infected. It is unclear whether these persistent plasma cells are long-lived cells derived from the extrafollicular response or are recent emigrants from germinal centers. All of these sources contribute to persistent red pulp plasma cells in responses of noninfected mice to hapten protein (38). It would be valuable to determine the balance between the acute short-lived plasma cells in the spleen and the long-lived plasma cells in the bone marrow because this could provide important information on the longevity of the Ab response in malaria.
Normally, plasma cells move from the edges of the T zone through the bridging channels into the extrafollicular foci of the red pulp (33). In P. chabaudi infection, large numbers of Ab-containing cells were detected within the T zone for at least 10 days after the peak of infection. In conventional histology, this atypical location of plasma cells has been previously noted for isolated time points after infection with Plasmodium berghei, P. yoelii (42), and P. chabaudi adami (43). The T zone location of plasma cells in malaria is similar to old observations in neonatally thymectomized mice, where the splenic white pulp develops an area corresponding to the T zone, but in place of T cells, most of the cells within are immature plasma cells (44). Thymic involution, which can occur in response to stress or infection and result in reduced T cell numbers in peripheral lymphoid organs, may have contributed to the lack of T cells in the T zone. Additional experiments on thymic subpopulations in P. chabaudi infection would clarify this. These data suggest that plasma cells do not migrate into the red pulp if T cells are absent or diminished in number in the periarteriolar region. Stromally produced CCL-13 is thought to play a role in plasma cell migration (45), but it is unknown whether T cells supply further migration signals via chemokines or direct cellular interaction.
This study shows that B cells disappear from the marginal zones at the peak of infection. Flow cytometry reveals that the cells are still present in the spleen and do not increase their level of apoptosis, suggesting that they migrate to other parts of the spleen. LPS causes transient loss of B cells from marginal zones (46) and immune complexes can induce transient migration of B cells from marginal zones to the follicles (47). In addition, LPS injection stimulates IgD+ blast formation reminiscent of that observed in our model (48), but malaria-infected spleens do not exhibit the rapid repopulation of the marginal zone seen after injection of nonreplicating Ags (47). The long-lasting changes in the marginal zone B cells suggest that these cells may play a role during the early acute malarial infection. Therefore, it would be interesting to see whether this phase of infection is altered in a mouse lacking marginal zone B cells, such as the Pyk-2-deficient mouse (21).
A possible function of the marginal zone B cells may lie in their capability to develop into Ab-secreting plasma cells more quickly than naive follicular B cells, particularly in response to particulate T-independent Ags (49, 50). This possibility is supported by our observation that marginal zone B cell numbers roughly double on days 5 and 6 of infection, with a sizable fraction (18% as compared with 3% in naive animals, data not shown) of the marginal zone B cell population observed on day 6 of plasmablast size.
In summary, our findings reveal no obvious deficiencies in the follicular or extrafollicular B cell pathways. On the contrary, the B cell response is much stronger than that seen in classical immunization experiments despite the gross alterations of the splenic architecture at the peak of infection. Although only P. chabaudi (AS) was studied, the observation of splenic disruption and abnormal plasma cell distribution in infections with other species of Plasmodium support the assumption that other clones of P. chabaudi and other rodent malaria species will elicit a similar B cell response.
The overall strength of the B cell response could be misleading, because malaria is known to generate a large polyclonal B cell response (51, 52). The kinetics of this response correspond well with the peak prevalence of Ab-containing cells in the infected spleens. However, the level of specific malarial IgG does not rise above the levels in normal serum until 3 wk of infection. This result suggests that a large fraction of the early B cell response may not be specific for malarial Ags. Therefore, it is essential to study how the B cell response specific for a malarial Ag develops in the spleen and the bone marrow.
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Acknowledgments |
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Footnotes |
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2 Address correspondence and reprint requests to Dr. Jean Langhorne, Division of Parasitology, National Institute for Medical Research, The Ridgeway, Mill Hill, NW7 1AA London, U.K. E-mail address: jlangho{at}nimr.mrc.ac.uk
3 Abbreviations used in this paper: CGG, chicken -globulin; AP, alkaline phosphatase; 7-AAD, 7-aminoactinomycin D; AU, arbitrary units; PNA, peanut agglutinin.
Received for publication December 24, 2002. Accepted for publication April 29, 2003.
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