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. 2018 Sep 24;155(4):519–532. doi: 10.1111/imm.12997

γδ T cells modulate humoral immunity against Plasmodium berghei infection

Shin‐Ichi Inoue 1,2,✉, Mamoru Niikura 2, Hiroko Asahi 2, Yasushi Kawakami 3, Fumie Kobayashi 2,3
PMCID: PMC6231001  PMID: 30144035

Summary

It is unclear whether γδ T cells are involved in humoral immunity against Plasmodium infection. Here, we show that B‐cell‐immunodeficient mice and γδ T‐cell‐deficient mice were incapable of protecting against Plasmodium berghei XAT parasites. γδ T‐cell‐deficient mice developed reduced levels of antigen‐specific antibodies during the late phase of infection. The numbers of follicular helper T cells and germinal centre B cells in γδ T‐cell‐deficient mice were lower than in wild‐type mice during the late phase of infection. Expression profiling of humoral immunity‐related cytokines in γδ T cells showed that interleukin‐21 (IL‐21) and interferon‐γ (IFN‐γ) are increased during the early stage of infection. Furthermore, blockade of IL‐21 and IFN‐γ signalling during the early stage of infection led to reduction in follicular helper T cells and germinal centre B cells. γδ T‐cell production of IL‐21 and IFN‐γ is crucial for the development and maintenance of follicular helper T cells and germinal centre B cells during the late phase of infection. Our data suggest that γδ T cells modulate humoral immunity against Plasmodium infection.

Keywords: humoral immunity, malaria, γδ T cells

Abbreviations

GC B cell

germinal centre B cell

IFN‐γ

interferon‐γ

IL‐21

interleukin‐21

iRBCs

infected red blood cells

KO

knockout

mAb

monoclonal antibody

M‐CSF

macrophage colony‐stimulating factor

PAMP

pathogen‐associated molecular patterns

PBS

phosphate‐buffered saline

PbXAT

Plasmodium berghei XAT

PE

phycoerythrin

p.i.

post‐infection

Tfh cell

follicular helper T cell

Th1

T helper type 1

TLR

Toll‐like receptor

WT

wild‐type

Introduction

Despite great efforts toward eradication, malaria remains one of the most common infectious diseases, causing high morbidity and mortality among children in the world.1 Plasmodium parasites are malaria‐causing protozoa that develop through complex life stages.2 In the mammalian host, Plasmodium parasites first develop in hepatocytes (liver stage), followed by red blood cells (RBCs; blood stage). In contrast to asymptomatic liver‐stage malaria, blood‐stage malaria is the symptomatic phase. Hence, protective immunity against blood‐stage malaria is important for reducing the severity of the disease. An efficient malaria vaccine has not yet been developed. Therefore, immune mechanisms involved in malaria must be elucidated to advance malaria vaccine development.

Passive transfer of serum antibodies from a repeatedly immunized donor into human patients can reduce parasitaemia and protect from severe clinical symptoms.3, 4 This evidence strongly indicates that malaria‐specific antibodies play a crucial role in protective immunity against malaria. Previous studies using a rodent model of malaria showed that CD4+ T‐cell‐depleted mice fail to control Plasmodium infection,5, 6, 7, 8 as is the case with interferon‐γ (IFN‐γ)‐signalling‐deficient mice.2, 9, 10, 11, 12, 13 IFN‐γ‐signalling‐deficient mice may exhibit decreased dendritic cell activation and T helper type 1 (Th1) responses during infection.9 Therefore, both cellular and humoral immunities are crucial for protective immunity against Plasmodium infection.

As innate‐like lymphocytes, γδ T cells are the first line of defence related to host protective immunity against various infections, including malaria.5, 9, 14, 15, 16, 17, 18, 19, 20 As malaria patients and Plasmodium‐infected animals show increased γδ T‐cell numbers in peripheral blood and the spleen, γδ T cells have been suggested to have a role in malaria.5, 21 After inoculation with blood‐stage Plasmodium berghei XAT (PbXAT), a low‐virulence strain, wild‐type (WT) mice show some fluctuations in parasitaemia levels in peripheral blood and eventually controlled the parasite after 1 month. Inhibition of IFN‐γ signalling by gene knockout (KO) or injection of neutralizing anti‐IFN‐γ antibody led to worsening of the parasitaemia, resulting in loss of fluctuation of the parasitaemia.9, 22 Interferon‐γ activates phagocytosis by macrophages and monocytes. Therefore, the fluctuations in the parasitaemia of blood‐stage PbXAT parasites are strongly related to IFN‐γ. In contrast to WT mice, γδ T‐cell‐deficient T‐cell receptor‐δ (TCR‐δ) KO mice cannot control PbXAT parasites, and they develop high parasitaemia and eventually die. γδ T cells elicit dendritic cell activation via CD40 ligand expression during PbXAT infection, resulting in induction of adequate Th1 cell differentiation for protection against parasites.5, 23 A previous study showed that Vγ1+ γδ T cells modulate the development of the pre‐immune peripheral B‐cell population but not the immature B‐cell population in bone marrow in the naive condition.24 Recently, we reported that Vγ1+ γδ T cells preferentially increase during PbXAT infection.9 These observations lead to the question of whether γδ T cells are involved in humoral immunity during PbXAT infection. Furthermore, a recent study revealed that Vδ6.3+ γδ T cells begin to expand and produce macrophage colony‐stimulating factor (M‐CSF) during the late phase of Plasmodium chabaudi infection. The M‐CSF‐producing Vδ6.3+ γδ T cells function to prevent parasitaemia recrudescence.25 Therefore, it will be interesting to determine whether the Vδ6.3+ γδ T cells are the same as, or a different subset of, Vγ1+ γδ T cells in terms of their ability to produce cytokines. Previously, we found that anti‐TCR‐γδ monoclonal antiobody (mAb) ‐treated mice had reduced levels of some isotypes of malaria‐specific antibodies during the late phase of infection.19 Although administration of antibodies is useful for cell depletion, this method may influence other immune cells such as Fc receptor‐expressing macrophages. Furthermore, it remains to be determined whether γδ T cells are involved in the cellularity of humoral immunity‐related cells during PbXAT infection.

In this study, we found that PbXAT‐infected TCR‐δ KO mice transiently produce high levels of Plasmodium antigen‐specific antibodies but eventually decrease this antibody production. Levels of Plasmodium antigen‐specific antibodies in plasma and the secreting cells decrease during the late phase of infection. The numbers of follicular helper T (Tfh) cells and germinal centre (GC) B cells in TCR‐δ KO mice were lower than in WT mice during the late phase of infection. Expression profiling of humoral immunity‐related cytokines in γδ T cells revealed that interleukin‐21 (IL‐21), IFN‐γ and IL‐4 are increased during the early stage of infection. We found that IL‐21 and IFN‐γ signalling during the early stage of infection was related to the development of Tfh and GC B cells, due to blockade of IL‐21 and IFN‐γ signalling. Therefore, γδ T cells may modulate humoral immunity against Plasmodium infection.

Materials and methods

Mice

Age‐matched female C57BL/6 WT (CLEA Japan, Meguro, Tokyo, Japan), TCR‐δ KO, and Ighm KO mice (The Jackson Laboratories, Bar Harbor, ME, USA; 8–12 weeks old) were bred in the pathogen‐free unit of the animal facility of Kyorin University. The Kyorin University School of Medicine Animal Care Committee approved all animal protocols.

Parasites and infection

A low‐virulence strain of PbXAT, derived from the high‐virulence strain P. berghei NK65, was used as described previously.5, 9, 19, 23, 26 The parasites were stored as frozen stock in liquid nitrogen or a deep freezer. Freshly thawed parasites were passaged once through naive WT mice, and 104 PbXAT‐infected red blood cells (iRBCs) from the passaged mice were inoculated intravenously into experimental mice. The resulting parasitaemia was assessed by counting 250–10 000 RBCs in a Giemsa‐stained thin blood film. The percentage of parasitaemia was calculated as [(number of iRBCs)/(total number of RBCs)] × 100.

Passive transfer of serum

Wild‐type and TCR‐δ KO mice were infected with PbXAT. On days 14, 16, 18, 20, 22 and 24 post‐infection (p.i.), serum was taken from the PbXAT‐infected WT mice. PbXAT‐infected TCR‐δ KO mice were intravenously injected with the serum from PbXAT‐infected WT mice or control serum from naive WT mice (400 μl per recipient mouse). Parasitaemia was assessed during infection.

Flow cytometry and antibodies

Each day after infection, single‐cell suspensions from mouse spleens were stained using the fluorescent antibody method in cold phosphate‐buffered saline (PBS) containing 0·5% bovine serum albumin (FUJIFILM Wako Pure Chemical Corporation, Chuo‐ku, Osaka, Japan) and 0·01% sodium azide as a staining buffer for FACS analyses. Phycoerythrin (PE)‐conjugated anti‐CD138 mAb and Pacific Blue‐conjugated anti‐B220‐mAb were used for plasma cell analyses. PE‐conjugated anti‐CXCR5 mAb, PE‐Cy7‐conjugated anti‐PD‐1 mAb, allophycocyanin‐conjugated anti‐CD3ε, allophycocyanin‐Cy7‐conjugated anti‐CD4 mAb, and Pacific Blue‐conjugated anti‐TCR‐β mAb were used for Tfh‐cell analyses. PE‐conjugated anti‐GL7 mAb and Pacific Blue‐conjugated anti‐B220 mAb were used for the GC B‐cell analyses. All mAbs were purchased from BioLegend, Japan. The analyses were performed using a FACS Canto II with FACS diva software (BD Biosciences, San Jose, CA, USA). The data were analysed using flowjo software (Tree Star, Ashland, OR, USA).

ELISA and ELISpot assay

Plasmodium berghei parasite‐specific antibodies were measured in the plasma of mice using soluble malaria antigens as the capture antigens, as previously described19 with some modifications. First, 50 μl of soluble malaria antigens in carbonate–bicarbonate buffer (20 μg/ml) was coated on each well of a 96‐well plate for ELISA (ELISA plate H; Sumitomo Bakelite Co., Ltd., Shinagawa‐ku, Tokyo, Japan), and incubated for 16 hr at 4°. The plate was washed three times with washing buffer (0·05% Tween‐20 in washing buffer). Then, the plate was treated with blocking solution (3% skim milk in washing buffer) for 2 hr at room temperature. The plate was washed three times with washing buffer (0·05% Tween‐20 in washing buffer). Plasma from naive and PbXAT‐infected mice was diluted with dilution buffer (3% bovine serum albumin in washing buffer). Twofold dilution series of plasma from naive and PbXAT‐infected mice were prepared. Then, 50 μl of each solution of diluted plasma was applied on wells in the plate, and then the plate was incubated for 2 hr at room temperature. Horseradish peroxidase‐conjugated anti‐mouse IgG1, IgG2c and IgG (Southern Biotech, Birmingham, AL, USA) were diluted 2000‐fold with dilution buffer and used to detect specifically bound mouse IgG isotypes and total IgG. The reactions were visualized using the substrate 2,2′‐azino‐bis(3‐ethylbenzothiazoline‐6‐sulphonic acid) (FUJIFILM Wako Pure Chemical Corporation) during a 30‐min incubation at room temperature. The absorbance of individual wells at 414 nm was determined using Multiskan FC (Thermo Fisher Scientific, Waltham, MA, USA). For calculating the end‐point titre of Plasmodium berghei parasite‐specific antibodies, we used the mean value of buffer control +3 SD (for IgG1 and IgG2c) or mean value of 32‐fold‐diluted naive plasma control × 2 (for total IgG) as cut‐off points.

Plasmodium berghei parasite‐specific antibody‐producing cells in the spleens of mice were measured using soluble malaria antigens as the capture antigens. The antigens were coated on the wells of MultiScreen filter plates (100 μl 25 μg/ml in PBS) at room temperature for 16 hr. After antigen coating, the plates were washed with PBS and incubated with 10% fetal bovine serum‐containing RPMI‐1640 medium at 37° for 1 hr. Next, splenocytes were cultured in the wells at 37° for 16 hr. After culture, the plates were washed with PBS. Horseradish peroxidase‐conjugated anti‐mouse IgG (BioLegend Japan) antibody was diluted 2000‐fold with PBS and used to detect mouse total IgG. The reactions were visualized using the substrate 3‐amino‐9‐ethyl‐carbazole (FUJIFILM Wako Pure Chemical Corporation) at room temperature for 5–15 min.

Quantitative RT‐PCR

γδ T cells were isolated using FACS Aria II (BD Biosciences). Total RNA from the isolated γδ T cells was extracted using a ReliaPrep RNA Miniprep System (Promega, Madison, WI, USA) according to the manufacturer's instructions. Total RNA was reverse‐transcribed into cDNA using a PrimeScript RT reagent kit with gDNA Eraser (Takara Bio, Kusatsu‐shi, Shiga, Japan). Quantitative real‐time PCR was performed on a 7500 Real Time PCR System (Applied Biosystems, Foster City, CA, USA; Thermo Fisher Scientific). The primers were designed manually using primer‐BLAST (NCBI). The primer sequences used for quantitative real‐time PCR were as follows: IFN‐γ (forward 5′‐TGGAGGAACTGGCAAAAGGA‐3′ reverse 5′‐TGTTGTTGCT GATGGCCTGA‐3′), IL‐4 (forward 5′‐GTAGGGCTTCCAAGGTGCTT‐3′, reverse 5′‐AAGCCCGAAAGAGTCT CTGC‐3′), IL‐10 (forward 5′‐CAGAGAAGCATGGCCCAGAA‐3′, reverse 5′‐GCTCCACTGCCTTGCTCTTA‐3′), IL ‐21 (forward 5′‐AGGACCCTTGTCTGTCTGGT‐3′, reverse 5′‐TAATCAGGAGGCGATCTGGC‐3′), IL‐6 (forward 5′‐GGGACTGATGCTGGTGACAA‐3′, reverse 5′‐ACAGGTCTGTTGGGAGTGGT‐3′), M‐CSF (forward 5′‐GCTGGCTTGGCTTGGGATGA‐3′, reverse 5′‐TGGCAGG TACTCCTGGGTGG‐3′). The quantitative PCR results were analysed using 7500 SDS v1.2.1 (Applied Biosystems; Thermo Fisher Scientific).

Statistical methods

Survival data were plotted using the Kaplan–Meier method and analysed with the log‐rank test. γδ T‐cell numbers were analysed using the independent Student's t‐test with Welch's correction, as variances were unequal. The other data were analysed using the Mann–Whitney U‐test. All data were analysed using statcel (The Publisher OMS Ltd., Higashikurume, Tokyo, Japan).

Results

The humoral immune response is crucial for protective immunity against PbXAT infection

To determine whether humoral immunity is essential for control of PbXAT parasites, we used B‐cell‐immunodeficient mice (Ighm KO mice; Fig. 1). After inoculation of iRBCs, WT mice showed several fluctuations in parasitaemia levels in peripheral blood and eventually controlled the parasite after a month. On the other hand, Ighm KO mice showed gradually increasing parasitaemia that ultimately resulted in the death of all mice (Fig. 1a,b). These results suggest that humoral immunity is required for protective immunity against PbXAT infection.

Figure 1.

Figure 1

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Humoral immunity is required for elimination of Plasmodium berghei XAT (Pb XAT). (a and b) Wild‐type (WT) and Ighm knockout (KO) mice were infected with Pb XAT through intravenous inoculation of blood‐stage parasites [104 Pb XAT‐infected red blood cells (iRBCs)]. (a) Time–course analysis of parasitaemia (n = 5 per group). Data are representative of two experiments. (b) Kaplan–Meier survival curves (n = 10 per group). Data are representative of four experiments, and statistical significance was analysed using the log‐rank test. **P < 0·01.

γδ T cells elicit persistent humoral immunity against PbXAT infection

T‐cell receptor‐δ KO mice cannot control blood‐stage PbXAT parasites.5, 23 This evidence indicates that γδ T cells are required for protective immunity against these parasites. However, it was unclear whether γδ T cells are involved in humoral immunity against PbXAT infection. Therefore, to compare the plasma levels of Plasmodium antigen‐specific antibodies between WT and TCR‐δ KO mice during PbXAT infection, we performed titration ELISA and calculated the end‐point titres of Plasmodium antigen‐specific antibodies (Fig. 2a–f). Plasma levels of Plasmodium antigen‐specific IgG were significantly increased on day 14 p.i. in both mouse groups compared with levels in the naive condition (day 0). Plasmodium antigen‐specific IgG levels were comparable between WT and TCR‐δ KO mice on day 14 p.i. On days 21 and 28 p.i., Plasmodium antigen‐specific IgG plasma levels were significantly lower in TCR‐δ KO mice than in WT mice (Fig. 2a). The TCR‐δ KO mice showed reduced titres of Plasmodium antigen‐specific total IgG and the IgG2c isotype, but not the IgG1 isotype, during the late phase of Plasmodium infection (Fig. 2b). Furthermore, to compare the trend in the induction of IgG isotypes, we calculated the IgG2c/IgG1 ratios. The IgG2c/IgG1 ratios of WT mice increased incrementally during the late phase of infection. In contrast, the IgG2c/IgG1 ratios of TCR‐δ KO mice were comparable during the late phase of infection (Fig. 2c). These results suggest that γδ T cells regulate IgG2c isotype induction. Next, we determined the numbers of Plasmodium antigen‐specific antibody‐secreting cells in the spleen and bone marrow during infection (Fig. 2d). Both WT and TCR‐δ KO mice had markedly increased antibody‐secreting cell numbers on day 14 p.i. compared with those in the naive condition. As shown in Fig. 2(a,b), those levels were comparable between WT and TCR‐δ KO mice. On the other hand, antibody‐secreting cell numbers in TCR‐δ KO mice began to decrease from day 21 p.i. On day 28 p.i., the levels in bone marrow were slightly higher in TCR‐δ KO mice than in WT mice. These data suggest that γδ T cells elicit persistent humoral immunity against PbXAT infection.

Figure 2.

Figure 2

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γδ T‐cell deficiency impairs the persistence of humoral immunity during Plasmodium berghei XAT (Pb XAT) infection. (a) Titration curves of Plasmodium antigen‐specific antibodies in plasma: Plasmodium antigen‐specific total IgG (left panels), IgG1 (middle panels) and IgG2c (right panels) levels in plasma from wild‐type (WT) and T‐cell receptor‐δ knockout (TCR‐δ KO) mice on days 7, 14, 21 and 28 post‐infection (p.i.; n = 3 per group). Data are representative of two experiments, and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01. (b) End‐point titres of Plasmodium antigen‐specific antibodies in plasma from WT and TCR‐δ KO mice on days 7, 14, 21 and 28 p.i. (n = 3 per group). (c) IgG2c and IgG1 antibody profiles against Plasmodium antigen were analysed by calculating the IgG2c/IgG1 ratios using plasma from WT and TCR‐δ KO mice on days 7, 14, 21 and 28 p.i. (n = 3 per group). (d) Plasmodium antigen‐specific IgG‐secreting cells in the spleen (Sp) and bone marrow (BM) were detected by ELISpot. Data represent Plasmodium antigen‐specific total IgG levels in plasma from WT and TCR‐δ KO mice on days 0, 7, 14, 21 and 28 p.i. (n = 5 per group). Data are representative of four experiments, and statistical significance was determined using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

Passive transfer of serum into TCR‐δ KO mice leads to a transient reduction of the parasitaemia during the late phase of PbXAT infection

The level of Plasmodium antigen‐specific antibodies was reduced in TCR‐δ KO mice during the late phase of infection. However, it is still unclear whether the decreased antibody levels in the TCR‐δ KO mice were related to the susceptibility to PbXAT. Therefore, we passively transferred serum from PbXAT‐infected WT mice into PbXAT‐infected TCR‐δ KO mice. Because the TCR‐δ KO mice could produce comparable levels of Plasmodium antigen‐specific antibodies to those of WT mice on day 14 p.i., the passive transfer of serum into PbXAT‐infected TCR‐δ KO mice was started after day 14 p.i. The passive transfer of serum led to a transient reduction in the parasitaemia in TCR‐δ KO mice during the time of passive transfer (Fig. 3). This suggests that the decreased antibody level in TCR‐δ KO mice during infection is partially related to their susceptibility to PbXAT. Both antibodies and the Th1 immune response are affected in TCR‐δ KO mice during PbXAT infection. Therefore, the passive transfer could not cure the parasitaemia completely.

Figure 3.

Figure 3

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Passive transfer of serum from Plasmodium berghei XAT (Pb XAT)‐infected wild‐type (WT) mice led to a reduction in the parasitaemia in γδ T‐cell‐deficient mice during Pb XAT infection. WT and T‐cell receptor‐δ knockout (TCR‐δ KO) mice were infected with Pb XAT via intravenous inoculation of blood‐stage parasites (104 iRBCs). On days 14, 16, 18, 20, 22 and 24 post‐infection (p.i.), serum was taken from Pb XAT‐infected WT mice and then intravenously injected into Pb XAT‐infected TCR‐δ KO mice on each day p.i. The time–course analysis of the parasitaemia is shown (n = 6 per group). Data are representative of two experiments, and statistical significance was analysed using the Mann–Whitney U‐test. **P < 0·01.

γδ T‐cell deficiency does not reduce the numbers of plasma cells during PbXAT infection

Plasma cell differentiation may be impaired by γδ T‐cell deficiency during infection. Therefore, we compared the levels of plasma cells in the spleen between WT and TCR‐δ KO mice (Fig. 4). In both mouse groups, there were more plasma cells, both in number and as a proportion of all cell types, during PbXAT infection. Unexpectedly, cell numbers during infection were comparable between TCR‐δ KO and WT mice. These data suggest that γδ T‐cell deficiency does not impair plasma cell differentiation or survival during PbXAT infection.

Figure 4.

Figure 4

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Plasma cell numbers are maintained in γδ T‐cell‐deficient mice during Plasmodium berghei XAT (Pb XAT) infection. (a) Zebra plots represent splenic CD138+ plasma cells from wild‐type (WT) mice and T‐cell receptor‐δ knockout (TCR‐δ KO) mice on days 0, 7, 14, 21 and 28 post‐infection (p.i.). (b) Proportions and (c) absolute numbers of splenic germinal centre (GC) B cells were compared between WT and TCR‐δ KO mice on days 0, 7, 14, 21 and 28 p.i. (n = 5 per group). Data are representative of three experiments, and statistical significance was analysed using the Mann–Whitney U‐test.

γδ T cells affect the persistence of Tfh cells during PbXAT infection

After activation by antigens, B cells change from isotype IgM to IgG production in the GC of secondary lymphoid organs such as the spleen. Tfh cells are involved in the class switch of GC B cells.27 Therefore, the levels of Tfh cells in the spleen during infection were estimated and compared between WT and TCR‐δ KO mice (Fig. 5). There were more Tfh cells, both in number and as the proportion of all CD4+ T cells, in both groups starting on day 7 p.i. The proportion was comparable between WT and TCR‐δ KO mice on days 7, 14 and 21 p.i. (Fig. 5b), and cell numbers were comparable on days 7 and 14 p.i. (Fig. 5c). In WT mice, both the proportion and number increased incrementally during PbXAT infection. However, on day 21 p.i., the number of cells was significantly lower in TCR‐δ KO mice than in WT mice. Furthermore, on day 28 p.i., both the proportion and number were significantly lower in TCR‐δ KO mice. These data suggest that γδ T cells promote persistence of Tfh cells during PbXAT infection.

Figure 5.

Figure 5

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γδ T‐cell deficiency indirectly reduces follicular helper T (Tfh) ‐cell development during the late phase of Plasmodium infection. (a) Zebra plots represent splenic CXCR5+ and PD‐1+ cells (gated by CD4+, CD3+, TCR β +) from wild‐type (WT) mice and T‐cell receptor‐δ knockout (TCR‐δ KO) mice on days 0, 7, 14, 21 and 28 post‐infection (p.i.). Histograms show splenic follicular helper T (Tfh) cells as ICOS + cells in the CXCR5+ and PD‐1+ population of each upper zebra plot. (b) The proportions and (c) absolute numbers of splenic Tfh cells were compared between WT and TCR‐δ KO mice on days 0, 7, 14, 21 and 28 p.i. (n = 4 per group). Data are representative of two experiments and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

γδ T cells affect the persistence of germinal centre B cells during PbXAT infection

Next, the levels of GC B cells in the spleen during PbXAT infection were estimated and compared between WT and TCR‐δ KO mice (Fig. 6). There were more GC B cells, both in number and as a proportion of all B cells, in both groups starting on day 7 p.i. The proportion of cells was comparable between the groups on days 7, 14 and 21 p.i. (Fig. 6b). Moreover, the number of cells was comparable between the groups on days 7 and 14 p.i. (Fig. 6c). In WT mice, the number of cells increased incrementally during PbXAT infection. However, on day 21 p.i., cell numbers were significantly lower in TCR‐δ KO mice than in WT mice. Furthermore, on day 28 p.i., both the proportion and number of GC B cells were significantly lower in TCR‐δ KO mice. These data suggest that γδ T cells promote the persistence of GC B cells during PbXAT infection.

Figure 6.

Figure 6

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γδ T cells suppress germinal centre (GC) B‐cell development during Plasmodium infection but support persistence of GC B cells. (a) Zebra plots represent splenic GC B cells as Fas+ and GL7+ cells (gated by B220+) from wild‐type (WT) mice and T‐cell receptor‐δ knockout (TCR‐δ KO) mice on days 0, 7, 14, 21 and 28 p.i. (b) The proportions and (c) absolute numbers of splenic GC B cells were compared between WT and TCR‐δ KO mice on days 0, 7, 14, 21 and 28 p.i. (n = 4 per group). Data are representative of three experiments, and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

γδ T‐cell levels increase incrementally and alter cytokine production during PbXAT infection

As in our previous study, the number of splenic γδ T cells increased incrementally during PbXAT infection (Fig. 7a).5 The expansion of γδ T cells led to the repression of IFN‐γ production in γδ T cells on day 14 p.i. compared with those in the early phase of PbXAT infection.9 However, it remains to be determined whether the production of other cytokines in γδ T cells is altered during PbXAT infection. Therefore, expression levels of cytokines including IFN‐γ, IL‐4, IL‐6, IL‐10 and IL‐21, which are related to the development of humoral immunity28, 29, 30, 31, 32 in γδ T cells, were estimated during PbXAT infection and compared with those in γδ T cells from naive WT mice (Fig. 7b–f). As in our previous study, IFN‐γ expression levels in γδ T cells were significantly increased on day 7 p.i. On the other hand, IFN‐γ expression level on day 14 p.i. was comparable to that of the naive condition.9 Moreover, the expression of all cytokines other than IL‐6 was increased on day 7 p.i. Interferon‐γ expression levels in γδ T cells were also increased from day 21 p.i. Compared with IL‐4 expression levels in the naive condition, they were significantly reduced from day 14 p.i. Expression levels of IL‐10 were significantly increased during PbXAT infection. The IL‐21 expression in γδ T cells from day 14 p.i. was comparable to that of the naive condition. These data suggest that γδ T cells drastically affect cytokine production during PbXAT infection.

Figure 7.

Figure 7

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Cytokine expression profiles in γδ T cells during Plasmodium infection. (a) Absolute numbers of γδ T cells in wild‐type (WT) mice on days 0, 7, 14, 21 and 28 post‐infection (p.i.) (n = 8 per group). Data are representative of three experiments, and statistical significance was analysed using the independent Student's t‐test with Welch's correction for unequal variances. **P < 0·01. mRNA expression of (b) interferon‐γ (IFN‐γ), (c) interleukin‐4 (IL‐4), (d) IL‐6, (e) IL‐10, and (f) IL‐21 in γδ T cells from WT mice on days 0, 7, 14, 21 and 28 p.i. determined using RT‐PCR (n = 4 per group). Data are representative of two experiments, and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

Cytokine gene expression by Vγ1+ Vδ6.3+ γδ T cells was similar to that of Vγ1+ Vδ6.3− γδ T cells during PbXAT infection

A recent study showed that Vδ6.3+ γδ T cells expanded clonally and expressed M‐CSF.25 The M‐CSF‐producing Vδ6.3+ γδ T cells were necessary for preventing the parasitaemic recrudescence of P. chabaudi. Therefore, we investigated whether the cytokine production differed among the γδ T‐cell subsets (Fig. 8). We identified three main γδ T‐cell subsets during PbXAT infection: Vγ1+ Vδ6.3+, Vγ1+ Vδ6.3− and Vγ4+ Vδ6.3− γδ T cells (naive, day 7, and day 21 p.i., respectively). As previously reported, the Vγ1+ Vδ6.3+ γδ T cells expanded incrementally and became the major γδ T‐cell subset during the late phase of infection. Although M‐CSF expression was detected in all γδ T‐cell subsets on day 21 p.i., the expression in Vγ1+ Vδ6.3+ and Vγ1+ Vδ6.3− γδ T cells was markedly higher than that of Vγ4+ Vδ6.3− γδ T cells (Fig. 8a). The expression of IFN‐γ, IL‐4 and IL‐21 of all γδ T‐cell subsets was increased on day 7 p.i. (Fig. 8b,d,e). However, the IFN‐γ expression by Vγ4+ Vδ6.3− γδ T cells was lower than that of Vγ1+ Vδ6.3+ and Vγ1+ Vδ6.3− γδ T cells during infection. On the other hand, there was no difference in the IL‐4 level on day 7 p.i. among the three γδ T‐cell subsets. The IFN‐γ expression of Vγ1+ Vδ6.3+, but not Vγ1+ Vδ6.3−, γδ T cells was significantly reduced on day 21 p.i. compared with that on day 7. The IL‐6 expression of all γδ T‐cell subsets was comparable during infection (Fig. 8f). Although the IL‐10 expression was increased in both Vγ1+ Vδ6.3+ and Vγ1+ Vδ6.3− γδ T cells on day 21 p.i., it was increased in Vγ1+ Vδ6.3−, but not in Vγ1+ Vδ6.3+,γδ T cells on day 7 p.i. (Fig. 8c). Collectively, these results suggest that Vγ1+ Vδ6.3+ and Vγ1+ Vδ6.3− γδ T cells express similar cytokines.

Figure 8.

Figure 8

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Cytokine expression profiles of γδ T‐cell subsets during Plasmodium infection. (a) Three main γδ T‐cell subsets, Vγ1+ Vδ6.3+, Vγ1+ Vδ6.3−, and Vγ4+ Vδ6.3− γδ T cells were isolated from spleen of wild‐type (WT) mice on days 0 (naive), 7 and 21 post‐infection (p.i.). Expression of (a) macrophage colony‐stimulating factor (M‐CSF), (b) interferon‐γ (IFN‐γ), (c) interleukin‐10 (IL‐10), (d) IL‐4, (e) IL‐21 and (f) IL‐6 mRNA in the three γδ T‐cell subsets from WT mice on days 0, 7 and 21 p.i. was determined using RT‐PCR (n = 4 per group). Data are representative of two experiments, and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

IL‐21 and IFN‐γ production during the early phase of PbXAT infection is crucial for the development of Tfh and GC B cells

We observed that γδ T cells produced IL‐21 protein transiently in the early phase of PbXAT infection using an intracellular cytokine assay (Fig. 9a,b). Interleukin‐21 plays a crucial role in Tfh‐cell and GC B‐cell development.32 Therefore, to test whether IL‐21 during the early phase of PbXAT infection is crucial for Tfh‐cell and GC B‐cell development, we used IL‐21R‐Fc to block IL‐21 signalling.33, 34 The administration of IL‐21R‐Fc during the early phase of infection led to a reduction in Tfh and GC B cells during the late phase of infection (Fig. 9c–h). These results suggest that γδ T‐cell‐produced IL‐21 during the early phase of infection promotes the development of Tfh and GC B cells during the late phase of infection.

Figure 9.

Figure 9

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Blocking interleukin‐21 (IL‐21) and interferon‐γ (IFN‐γ) signalling leads to a reduction in follicular helper T (Tfh) and germinal centre (GC) B cells during the late phase of Plasmodium infection. (a) Zebra plots represent IL‐21+ γδ T cells (gated by CD3+, TCR‐γδ +, TCR‐β −) from the spleens of wild‐type (WT) mice on days 0, 7 and 14 p.i. (b) The proportion of IL‐21+ γδ T cells in WT mice on days 0, 7 and 14 p.i. Zebra plots represent (c) splenic CXCR5+ and PD‐1+ cells (gated by CD4+, CD3+, TCR‐β +) and (f) GC B cells as Fas+ and GL7+ cells (gated by B220+) on days 21 and 28 p.i. from WT mice that were treated with IL‐21R‐Fc to block IL‐21 signalling, and with anti‐IFN‐γ neutralizing antibody to block IFN‐γ signalling during the early phase of infection (on days 4–9). The proportions (d, g) and absolute numbers (e, h) of splenic (d, e) Tfh and (g, h) GC B cells were compared between WT mice and IL‐21R‐Fc‐treated and anti‐IFN‐γ antibody‐treated mice on days 21 and 28 p.i. (n = 4 per group). Data are representative of two experiments, and statistical significance was analysed using the Mann–Whitney U‐test. *P < 0·05, **P < 0·01.

Previously, we showed that enhancement of dendritic cell activation via CD40 ligand‐ and IFN‐γ‐expressing γδ T cells was responsible for protective immunity against Plasmodium parasites during the early stage of infection.5, 9 Next, to investigate whether IFN‐γ during the early phase of PbXAT infection is crucial for Tfh‐cell and GC B‐cell development, we used anti‐IFN‐γ neutralizing antibody to block IFN‐γ signalling. The administration of anti‐IFN‐γ antibody during the early phase of infection led to a reduction in Tfh and GC B cells during the late phase of infection (Fig. 9c–h). These results suggest that γδ T‐cell‐produced IFN‐γ during the early phase of infection is related to the development of Tfh and GC B cells during the late phase of infection.

Discussion

γδ T cells are crucial for protective immunity against PbXAT infection.5, 9, 19, 23 However, the role of γδ T cells in humoral immunity against malaria parasites is not well understood. Here, we report that γδ T cells help to maintain humoral immunity during PbXAT infection. Although the levels of Plasmodium antigen‐specific IgG and the numbers of their producing cells in the spleen were comparable between WT and TCR‐δ KO mice on day 14 p.i., they were decreased during the late phase of infection (Fig. 2). These data suggest that γδ T cells play a role in maintaining humoral immunity against Plasmodium infection.

Our previous study showed that γδ T‐cell depletion during the early phase of PbXAT infection (~day 10 p.i.) resulted in the death of mice.5 Therefore, the effector function of γδ T cells essential for protective immunity is expected to cease before day 10 p.i. We show here that γδ T‐cell deficiency leads to impaired humoral immunity during the late phase of infection. To explain this discrepancy, we hypothesized that γδ T cells might affect humoral immunity by orchestrating other immune cells, such as natural killer cells. In the late phase of infection, other immune cells may compensate for the absence of γδ T cells. Moreover, it is possible that γδ T cells have some effects on fibroblastic reticular cells, causing the formation of a GC in the spleen for the development of humoral immunity during the early phase of infection.35

Furthermore, a recent study showed that γδ T cells affect systemic IgG antibody levels in non‐immunized mice.36 In that study, a lower total IgG level was observed in TCR‐δ KO mice than in WT mice in the pre‐immune condition. On the other hand, TCR Vγ4/6 KO mice exhibit higher levels of IgM and IgG and increased spontaneous GC formation and B‐cell activation. TCR Vγ4/6 KO mice exhibit a high proportion of Vγ1+ γδ T cells in the spleen. Interleukin‐4 produced from Vγ1+ γδ T cells induces B‐cell formation from the GC and B‐cell antibody production. In our previous study, TCR‐δ KO mice exhibited lower IFN‐γ levels in plasma and a smaller population of Th1 cells in the spleen compared with WT mice during PbXAT infection.5 Moreover, we recently reported that Vγ1+ γδ T cells up‐regulated IFN‐γ production after PbXAT infection and that Vγ1+ γδ T cells were necessary for protective immunity against PbXAT infection.9 Furthermore, a recent study reported that Vδ6.3+ γδ T cells begin to expand and produce M‐CSF during the late phase of Plasmodium infection. The M‐CSF‐producing Vδ6.3+ γδ T cells prevented parasitaemia recrudescence, and most Vδ6.3+ γδ T cells expressed TCR Vγ1+.25 In comparison, in this report, we found that Vγ1+ Vδ6.3− γδ T cells are present during infection and in the naive condition. The Vγ1+ Vδ6.3− γδ T cells also express M‐CSF during the late phase of infection (Fig. 8a,b). Interleukin‐4 from Th2 and Tfh cells induces an IgG1 class switch in B cells. In contrast, IFN‐γ from Th1 cells induces an IgG2c class switch in B cells. In this report, TCR‐δ KO mice had reduced IgG2c levels, but not IgG1 levels, in plasma during the late phase of Plasmodium infection. Furthermore, a previous study showed that IFN‐γ is an important factor involved in the development of spontaneous growth of GC B cells.28 Therefore, IFN‐γ from γδ T cells or Th1 cells may elicit increases in GC B‐cell numbers and the production of antigen‐specific antibodies. Indeed, during the late phase of infection, γδ T cells expressed slightly higher levels of IFN‐γ than that in the naive condition (Fig. 6b). Furthermore, blocking IFN‐γ signalling with neutralizing antibody during the early phase of infection led to reductions in Tfh and GC B cells during the late phase of infection (Fig. 9c–h). Interleukin‐6 enhances the development of Tfh cells.30 On the other hand, IL‐21 enhances the development of both Tfh cells and GC B cells.30, 32 Although levels of IL‐21, but not IL‐6, from γδ T cells were transiently higher in the early phase of infection (day 7 p.i.), levels during the late phase of infection were comparable to those of naive γδ T cells (Fig 6b). Furthermore, blocking IL‐21 signalling with IL‐21R‐Fc during the early phase of infection led to reductions in Tfh and GC B cells during the late phase of infection (Fig. 9a,b,d,e). Therefore, IL‐21 from γδ T cells affects the maintenance of Tfh and GC B cells in the late phase of infection. A recent study showed that IL‐10 is essential for humoral immunity against malaria.31 In this paper, we found that levels of IL‐10 in γδ T cells were higher during infection. Therefore, IL‐10 from γδ T cells may be involved in maintaining Tfh and GC B cells in the late phase of infection.

A recent study reported that human Vγ9+ Vδ2+ γδ T cells, a major γδ T‐cell subset in humans, facilitated H9N2 influenza virus‐specific IgG production.37 This study showed that human γδ T cells promote Tfh‐cell differentiation, resulting in the induction of plasma cell differentiation and B‐cell immunoglobulin class switching. Moreover, they observed that human γδ T cells induced CD4 T cells to produce IL‐21 and IL‐13, resulting in enhanced plasma cell differentiation. Although they could not detect IL‐21 expression by human γδ T cells in their experiment, another study observed that IL‐21 was produced by human γδ T cells from patients with autoimmune diseases.38 Using mouse γδ T cells, a third group reported that mouse γδ T cells begin to express IL‐21 on stimulation with IL‐23.39, 40 Expression of IL‐21 in human γδ T cells may also require the presence of factors such as IL‐23. Therefore, human γδ T cells may express IL‐21 in the human body during influenza virus infection. Furthermore, it was suggested that direct contact between human γδ T cells and CD4 T cells was required for B‐cell activation. In this paper, we suggest that IL‐21 and IFN‐γ secretion from γδ T cells is crucial for Tfh and GC B‐cell development during the late phase of Plasmodium infection. However, direct contact between γδ T and Tfh cells may also play an important role in the persistence of Tfh and GC B cells during the late phase of Plasmodium infection.

Although the levels of Plasmodium antigen‐specific antibodies were significantly reduced by γδ T‐cell deficiency during the late phase of Plasmodium infection, plasma cell numbers were maintained during the infection (Fig. 3). Therefore, plasma cells in TCR‐δ KO mice may fail to produce antibodies during infection. Plasmodium parasites exhibit different varieties of pathogen‐associated molecular patterns (PAMPs).41 Plasmodium PAMPs are detected by pattern recognition receptors, such as Toll‐like receptors (TLRs), in host immune cells. Excessive TLR signalling by high parasitaemia in TCR‐δ KO mice during the late phase of infection may lead to a reduction in antigen‐specific antibody production, even though TLR signalling is crucial for B‐cell responses.42

There remains controversy about whether γδ T cells elicit or regulate protective immunity against Plasmodium infection. In patients with severe malaria, γδ T cells are a major source of cytokines including IFN‐γ and tumour necrosis factor‐α in response to P. falciparum iRBCs. The cytokines and chemokines produced by γδ T cells are associated with the severity of malaria.43 Therefore, γδ T cells may be cellular factors influencing immunopathology in malaria. On the other hand, many reports indicate that γδ T cells are cellular factors influencing protective immunity against Plasmodium infection. A P. falciparum sporozoite vaccine increases γδ T‐cell numbers in peripheral blood. The frequencies of IFN‐γ‐producing γδ T cells in immunized humans are well correlated with protection against P. falciparum infection.44 This evidence strongly indicates that γδ T cells contribute to protective immunity against Plasmodium infection. Furthermore, we previously showed that dendritic cell stimulation by γδ T cells is necessary for protective immunity against PbXAT infection.5, 23 Here, we show that γδ T cells modulate humoral immunity against Plasmodium infection. Consequently, γδ T cells may be important effector cells for modulation of not only Th1 cell‐based cellular immunity but also humoral immunity against malaria. Further investigation of γδ T cells in malaria will help to achieve new approaches for more effective malaria vaccine development.

Disclosures

There is no conflict of interest in this study.

Acknowledgements

SI performed the experiments; SI, MN, HA, YK and FK analysed the data; SI designed the study; SI and FK wrote the paper. This work is supported by Takeda Science Foundation, Kato Memorial Bioscience Foundation, and a Grant‐in‐Aid for Young Scientists (B) (No. 15K19085) from the Japan Society for the Promotion of Science (JSPS) to SI. This work was also supported in part by a Grant‐in‐Aid for Scientific Research (C) from JSPS to FK (No. 15K08451) and to MN (No. 15K08449) and by a Strategic International Research Cooperative Program (SICP) from the Japan Agency for Medical Research and Development (AMED) to FK.

[Correction added on 11 October 2018, after first online publication: Figures 2 and 4 were previously wrong and have been corrected in this version.]

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