Acute paranoid psychosis as sole clinical presentation of hepatic artery thrombosis after living donor liver transplantation

Armin D Goralczyk
Volker Meier
Giuliano Ramadori
Aiman Obed
Thomas Lorf
Acute paranoid psychosis as sole clinical presentation of hepatic artery thrombosis after living donor liver transplantation
C A S E R E P O R T Open Access
Background: Hepatic artery thrombosis is a devastating complication after orthotopic liver transplantation often requiring revascularization or re-transplantation. It is associated with considerably increased morbidity and mortality. Acute cognitive dysfunction such as delirium or acute psychosis may occur after major surgery and may be associated with the advent of surgical complications. Case presentation: Here we describe a case of hepatic artery thrombosis after living-donor liver transplantation which was not preceded by signs of liver failure but rather by an episode of acute psychosis. After retransplantation the patient recovered without sequelae. Conclusion: This case highlights the need to remain cautious when psychiatric disorders occur in patients after liver transplantation. The diagnostic procedures should not be restricted to medical or neurological causes of psychosis alone but should also focus vascular complications related to orthotopic liver transplantation.

Background

Hepatic artery thrombosis (HAT) is the most frequent arterial complication in orthotopic liver transplantation (OLT) occurring in 2.5-6.8% of adult transplant recipients including adult living donor liver transplant recipients [1][2][3] with an increased incidence in pediatric and ABO-incompatible liver transplantation [4,5]. The clinical presentation of HAT varies depending on the time of onset after transplantation. Severe acidosis, fever, systematic inflammatory response syndrome, or hepatic failure may ensue in early HAT (less than one month after OLT), whereas presentation of late HAT may vary from biliary complications such as recurrent biliary sepsis and stenotic lesions of the biliary tract, to an asymptomatic clinical picture with a mild elevation of serum transaminases and bilirubin levels [2,6]. HAT may thus be indicated by elevated serum transaminases, prolonged prothrombin time, or elevated total bilirubin and may be confirmed by doppler ultrasonography or angiography. Although in approximately 47% HAT requires re-OLT, cases without progressive allograft failure arterial flow may be resolved by fibrinolysis, interventional treatment, or surgical thrombectomy [1]. Mortality of HAT may vary from 55.6% (early HAT) to 15% (late HAT) [2]. The most common cause for HAT is surgical technique but hemodynamic or immunologic factors, such as reperfusion injury, hypercoagulation, and viral infections have also been recognized [1,7]. Risk factors for HAT include the number of allografts, cytomegalovirus (CMV) infection status, high recipient/donor weight ratio, biopsy proven rejection, and combination of female donor and male recipient [2].

Case presentation

A 42-year-old male patient underwent a living-donor liver transplantation (LDLT) for progressive hepatitis B, cirrhosis with 7 points on the Child-Turcotte-Pugh scale and a model for end-stage liver disease score of 10. He was placed on the waiting list for transplantation because of suspected hepatocellular carcinoma. He had no history of psychiatric disorder. The donor was his 43-year-old brother who had no major medical problems. The clinical match was appropriate (see Table 1), there were no contraindications and after further evaluation recipient and donor were found to be suitable for LDLT.

The donor underwent standard right lobectomy with preservation of the middle vein. In the recipient a standard total hepatectomy was performed, but due to successive intimal dissection of the hepatic and celiac artery complex arterial reconstruction was necessary. Due to the small diameter of the donor hepatic artery a direct anastomosis to the recipient supraceliac aorta was not feasible so an allogenic iliac artery conduit was required. The interposition graft was obtained from a heart-beating donor of a previous OLT and stored in histidinetryptophane-ketoglutarate solution at +4°C for 10 days. Anastomosis of the bile duct was performed by duct-toduct anastomosis to the donor right hepatic duct remnant.

At the time of transplantation the patient received induction immunosuppression consisting of 500 mg methylprednisolone and 20 mg basiliximab, the latter was repeated on the 4th post-operative day (POD). Immunosuppression was maintained with oral mycophenolat mofetil from the first POD at 2 × 1 g daily and oral prednisolone with an initial daily dose of 75 mg. The latter was tapered by 5 mg every two days. On the 5th POD oral tacrolimus was added to the immunosuppressive regimen starting with 0.5 mg on the first day and increasing daily dose by 1 to 2 mg to achieve trough plasma level of 4-10 μg per liter. After surgery the recipient received continuous intravenous infusion of 5000 units of unfractioned heparin per day for anticoagulation. For prevention of (re-)infection the patient also received 10 3 units of hepatitis B immunglobuline at reperfusion and intravenous infusion of piperacillin/ tazobactam at surgery until the 3rd POD but no antimycotics or prophylactic CMV therapy (both donor and recipient had experienced CMV infection).

In the postoperative course serum transaminases peaked on the first and 4th POD but then decreased continuously as prothrombin time increased (coagulation factors were not substituted). Albeit a gradual increase in total bilirubin plasma levels only a mild graft dysfunction was suspected because bile flow over the T-drain increased. Daily doppler sonography of hepatic vessels showed unaltered graft perfusion. Overall the patient had an uncomplicated postoperative without signs for infection or organ dysfunction. The clinical course and laboratory findings did show mild a graft dysfunction but neither rejection nor acute graft failure were indicated.

On the 9th POD the patient suffered from acute paranoid psychosis, being alert but agitated and physically aggressive. He was mistrustful and afraid of being harmed by people around him. The patient became unmanageable and had to be sedated with an intramuscular injection of 5 mg haloperidol and an intravenous injection of 20 mg diazepam. Due to this unusual presentation of postoperative delirium a diagnostic workup was initiated. Patent hepatic artery could not be shown in doppler sonography and subsequent angiography. Explorative laparotomy on the same day revealed a complete thrombosis of the donor hepatic artery and gross ischemic infarction in all liver segments. Because progression of infarction and subsequent graft loss was to be expected the patient was placed on the transplant waiting list for re-OLT.

An acceptable organ was offered from an 18-year-old heart-beating donor on the following day and the patient underwent re-OLT 10 days after the LDLT. The hepatic artery of the new graft was reanastomosed to the old conduit. The immunosuppressive regimen was reinitiated as described above including tacrolimus (see Figure 1). Emergence from anaesthesia was uneventful and the psychiatric symptoms did not recur in the postoperative course. Furthermore a clinically suspected episode of acute rejection indicated by rising transaminases was treated with a course of methylprednisolone (see Figure 1). Daily doppler sonography of hepatic vessels showed an unaltered graft perfusion. Examination of the explant revealed thrombotic occlusion of the hepatic artery with anaemic infarction of all liver segments.

Discussion

Here we describe a case of HAT after LDLT preceded not by signs of an liver failure but by acute paranoid psychosis. At transplantation the patient required a complex arterial reconstruction with an interposition graft which is a known risk factor for HAT [2,8]. Whether supraceliac or infrarenal placement of the interposition graft is beneficial remains equivocal [1,9]. LDLT itself has also been implied to be a risk factor for HAT, but Hatano et al could show increased incidence of HAT only in ABO-incompatible grafts [5]. In the described patient we observed a mild graft dysfunction indicated by increasing bilirubin in the first week after LDLT but accompanied by decreasing transaminases HBs-Ag neg. pos.

Anti-HBs > 1000 mlU/ml <10 mIU/ml Anti-HCV neg. neg. and an increasing synthetic liver function. Daily doppler sonography showed patent vessels until he suffered from acute psychosis on the 9th postoperative day. Psychiatric disorders are not uncommon in the postoperative course of the liver transplant patient [10]. Like other patients undergoing major surgery, transplant patients may suffer from postoperative delirium. The cause of delirium is multifactorial, including predisposing factors (age, primary disease), medical/surgical factors (length of surgery, use of a cardiopulmonary bypass, water-electrolyte imbalances, sepsis, etc), and postoperative emotional stress, like prolonged sleep deprivation, use of physical restraints, and pain [11,12]. In the described patient postoperative delirium was unlikely because after surgery delirium typically evolves immediately after emergence from general anaesthesia or, after a lucid interval, in the first 72 hours [13].

Psychiatric disorders may also be drug induced. Immunosuppressive drugs like tacrolimus and corticosteroids are known causative agents of psychiatric reactions [14][15][16]. Neurotoxic effects are dose dependent and occur in the early phase after initiation, e.g. within the first week [14,16]. In our patient induction of the described psychiatric symptoms by immunosuppressants was also unlikely because the psychosis occurred after the first week and doses were low, i. e., 5 ng/ml tacrolimus on the 9th postoperative day (see also Figure 1). After re-transplantation he received tacrolimus and prednisolone again without any sign of recurrence of the psychiatric disorder.

Because of elevated bilirubin we also suspected impaired graft function could cause acute psychosis in our patient. In chronic liver failure manifestations of encephalopathy are common and may reach from mild obtundation with dementia and movement disorders to deep coma [17]. In the early phase of acute liver failure agitation, delusional ideas, and hyperkinesis are also common until coma finally ensues [18]. Although typical laboratory findings of acute liver failure are common, only moderate alteration, as seen in our patient, is not an exclusion criterion [19]. In our patient ammonia was also not elevated but in patients with more advanced hepatocellular dysfunction the level of ammonia is only poorly correlated with the severity of hepatic encephalopathy [17]. Besides the accumulation of unmetabolized ammonia other mechanisms have been implied as causes of encephalopathy such as imbalance of neurotransmitters, endogenous benzodiazepine ligands, or elevation of circulating tumor necrosis factor (TNF) [19,20], some of them also involved in the pathogenesis of sepsis-associated delirium [12].

Besides infection and sepsis surgical complications are the most likely cause of graft dysfunction in the early post-transplant period. Therefore we initiated further diagnostic work-up. Abdominal sonography showed no flow in the hepatic artery and subsequently HAT was confirmed by angiography. During explorative laparotomy the liver was found to be with gross infarction and we decided to evaluate the patient for re-transplantation. Luckily we obtained a graft within 24 hours and after re-transplantation psychosis resolved without sequelae.

Conclusion

We propose that in the described patient HAT is the most likely cause of an acute liver failure although laboratory findings indicated only mild graft dysfunction. Acute liver failure was accompanied by hepatic encephalopathy apparent as acute delusional psychosis.

This case highlights the need to remain suspicious when psychiatric disorders occur in patients after liver transplantation. The diagnostic procedures should not be restricted to medical or neurological causes of psychosis but should also focus vascular complications of OLT.

Figure 1
1Postoperative course of the patient. Postoperative course of the patient. Upper panel depicts aspartate transaminase (AST, solid line) and alanine transaminase (ALT, dashed line) plasma levels in units per liter. In the middle panel total bilirubin (dashed line) and prothrombin time (PT, solid line) in activity percentage is shown. The lower panel displays immunosuppression: methylprednisolone, black bars; prednisolone, grey upper bars; basiliximab (Simulect), diamonds; tacrolimus trough levels, solid line and dots; tacrolimus dose, lower grey bars.

Table 1
1Characteristics of donor and recipient.Donor
Recipient

Age
43 yr.
42 yr.

Body weight
60 kg
73 kg
1.22*

BMI †
21
24
1.17*

Blood group
A Rh pos.
A Rh pos.

CMV IgG ‡
pos.
pos.

AcknowledgementsWritten informed consent was obtained from the patient for publication of this case report and accompanying images. A copy of the written consent is available for review by the Editor-in-Chief of this journal.Authors' contributions ADG and TL collected and analyzed the data and wrote the article. GR, HB and AO contributed substantially to the process of analyzing the data and writing the paper. All authors read and approved the final manuscriptCompeting interestsThe authors declare that they have no competing interests.
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Comparative performance of reference laboratory tests and in-clinic tests for Giardia in canine feces

Jennifer Mizhquiri Barbecho
Dwight D Bowman
Janice L Liotta
Comparative performance of reference laboratory tests and in-clinic tests for Giardia in canine feces
10.1186/s13071-018-2990-6R E S E A R C H Open Access
Background:We examined the performance of four in-clinic Giardia diagnostic tests by comparing results to three laboratory methods for detection of Giardia. A set of 177 fecal samples originally submitted to a commercial laboratory by veterinarians for routine ova and parasite (O&P) testing was used. Specimens were examined by direct immunofluorescence assay (DFA) for presence of Giardia cysts which served as the gold standard. Fecal samples were tested using a Giardia-specific cyst wall antigen microtiter plate format enzyme-linked immunosorbent assay (ELISA) and each of the in-clinic assays adhering to the package insert for each kit. Results: Evaluated were four in-clinic antigen test kits: VetScan® Canine Giardia Rapid Test (Abaxis), Anigen® Rapid CPV-CCV-Giardia Antigen Test (BioNote), SNAP® Giardia Test (IDEXX) and Witness® Giardia Test (Zoetis). In the comparison of the in-clinic tests to the DFA standard test sensitivity ranged between 70.0-87.1%, and specificity ranged between 71.1-93.4%. Conclusion: Of the tests evaluated here, the SNAP test had the highest sensitivity and specificity. The SNAP test had the highest percent positive and percent negative agreement when compared to the microtiter plate format ELISA and the O&P assay.

Background

Giardia are protozoan parasites capable of infecting numerous mammals including both dogs and cats and has been associated with gastrointestinal disease [1][2][3]. For dogs, diarrhea is a common clinical manifestation presented to veterinary clinics and a consideration in managing these patients is determining Giardia infection status. Giardia can be identified in feces by visualization of whole cysts by microscopy using zinc sulfate (ZnSO 4 ) fecal floatation [ova and parasite (O&P) test], by direct immunofluorescence (DFA) using Giardia-specific fluorescent antibodies or by demonstrating the presence of Giardia-specific antigen in feces using an immunoassay [4][5][6].

Fecal flotation can be performed in-clinic but identification of cysts can be difficult because of yeast and debris in the fecal sample which resemble cysts and because shedding is intermittent requiring multiday examinations [7]. Direct immunofluorescence assay has been shown to be more sensitive and specific than conventional flotation methods and herein served as the gold standard method for identifying fecal Giardia [5,8,9]. The DFA requires a fluorescent microscope limiting its usefulness in-clinic.

Giardia antigen detection tests detect a secreted soluble Giardia-specific cyst wall antigen and are available from several sources [3,6,7]. These tests are available as microtiter plate format ELISAs which can be read visually or by an instrument-based procedure in a reference laboratory setting and single use rapid tests which are read visually and are used in-clinic. A number of in-clinic tests have been described and are now available for use but studies directly comparing the performance of these commercially available tests have not been reported [6,10,11]. Using DFA as the gold standard, we examined the performance of four in-clinic rapid tests using a set of clinic samples originally submitted to a commercial laboratory by veterinarians for conventional ova and parasite (O&P) testing. Evaluated were four in-clinic antigen test kits: VetScan® Canine Giardia Rapid Test (Abaxis), Anigen® Rapid CPV-CCV-Giardia Antigen Test (BioNote), SNAP® Giardia Test (IDEXX) and Witness® Giardia Test (Zoetis).

Methods

Canine samples

Fecal samples were sourced from IDEXX Reference Laboratories (IRL) by the sole criterion that they were submitted by practicing veterinarians for general ova and parasite (O&P) testing which was performed at the reference laboratory using ZnSO 4 centrifugal floatation. No clinical information was available. Samples were stored frozen (-20°C) after the initial O&P test. A total of 87 O&P positive and 90 O&P negative samples was randomly selected so approximately an equal number of O&P positive and negative samples were obtained. Technicians performing all remaining testing had no knowledge of the O&P results.

In-clinic tests

Evaluated were four in-clinic antigen rapid test kits: VetScan® Canine Giardia Rapid Test (Abaxis, Union City, USA), Anigen® Rapid CPV-CCV-Giardia Antigen Test (BioNote, Seoul, Korea), SNAP® Giardia Test (IDEXX, Westbrook, USA) and Witness® Giardia Test (Zoetis, Parsippany, USA). The four kits were tested concurrently adhering to the package insert for each kit. Before testing samples were randomized and blind-labeled. There were no additional freeze-thaw cycles or sample handling differences in testing events between the four rapid tests. Visual interpretation of results was performed following the manufacturers' instructions supplied with each test kit.

Microtiter plate format ELISA

Samples were tested with the ProSpecT® Giardia/Cryptosporidium Microplate Assay (Thermo Fisher Scientific, Pittsburgh, USA) using instructions supplied with the kit. Plates were read spectrophotometrically at 450 nm; samples were considered positive when the net optical density reading (sample OD-negative control OD) was ≥ 0.05.

DFA test

The DFA was conducted and Giardia cysts were counted using a previously described procedure with modifications [5]. Briefly, 0.1 g feces was added to 900 μl 0.02 M sodium phosphate-buffered saline pH 7.4 (PBS) and mixed thoroughly. An aliquot of the mixed sample (100 μl) was added to a second tube containing 900 μl of PBS. The second tube was thoroughly mixed and 100 μl was added to a third tube which was mixed with 5 μl of Merifluor® detection reagent (Meridian Biosciences, Cincinnati, USA). The sample was then incubated at room temperature for 30 min in the dark and then maintained at 4°C until the quantitative cyst count was performed. A sample volume of 10.5 μl was removed and examined using a fluorescent microscope. The cysts were counted and a total cysts/gram was calculated. A positive and negative control were read prior to each test. The positive control was 85 μl PBS, 15 μl positive control from the Merifluor kit, and 5 μl Merifluor detection reagent. The negative control consisted of 100 μl PBS and 5 μl Merifluor detection reagent.

Data collection and analysis

Statistical analysis was performed in SAS© version 9.4 (SAS Institute, Inc., Cary, NC, USA). Agresti-Coull 95% confidence intervals were used for calculation of sensitivity and specificity compared to the DFA reference method, as well as percent positive and negative agreement to the microtiter plate format ELISA test. Agresti-Coull confidence intervals were calculated using the FREQ procedure in SAS 9.4. Prevalence adjusted test agreement was calculated using the following formula: agreement = prevalence × sensitivity + (1prevalence) × specificity. In this calculation, we used the prevalence (5.4%) found for client-owned symptomatic dogs presented to a veterinary hospital as determined by a monoclonal antibody-based immunofluorescent antibody assay [12].

Results

DFA, O&P and microtiter plate ELISA results

The final sample set consisted of 177 samples; all were examined by DFA and by microtiter plate format ELISA. Cysts were identified by DFA in 101 of 177 samples which were classified as positive and were not identified in 76 samples which were classified as negative. DFA results were compared to the original O&P results reported by the reference lab. In the O&P test, cysts  Table 3. Among the in-clinic rapid tests, SNAP Giardia showed the highest agreement with DFA and the microtiter plate format ELISA.

Discussion

We sought to compare the performance of four commercially available in-clinic assays designed to detect Giardia infection in dogs. Although we used the O&P test to select samples to ensure a sufficient population of negative and positives, we used direct immunofluorescence as the gold standard because several studies have demonstrated its accurate performance compared to traditional flotation methods. We chose to include the microtiter plate format test for comparison because the test detects soluble Giardia specific cell wall antigen shed in the local environment and is used for high throughput screening in reference laboratories. In comparing the microtiter plate format ELISA results to the DFA results, ELISA had a sensitivity of 94.1%, and a specificity of 97.4% (Table 1).

Results for the comparison of the in-clinic tests to the DFA and the microtiter plate format ELISA standard are shown in Tables 2 and 3, respectively. In all three measures of performance, SNAP had the highest concordance with the DFA and the ELISA standards among the tests.

We also used DFA results to retrospectively evaluate performance characteristics of the O&P method (Table 1) and compared these to those found for the 4 in-clinic tests compared to DFA shown in Table 2. We observed that the performance of SNAP Giardia was in close agreement with the O&P method while each of the 3 remaining in-clinic tests showed reduced values for percent positive agreement, percent negative agreement and prevalence adjusted agreement relative to this method.

The performance of SNAP Giardia has been studied by several groups [6,10,11]. To our knowledge, this is the first study to evaluate multiple tests currently available for detection of Giardia including several new rapid in-clinic tests. We found the performance of rapid in-clinic tests varied by comparing to different reference standards. There could be a myriad of factors attributable to the observed variability. The SNAP Giardia is an ELISA-based assay designed for in-clinic use and the methodology has been described in detail in a recent technical review [13]. In the  same journal issue dealing with "Point of Care Tests in Veterinary Medicine", the lateral flow methodology, which is the basis for the other three in-clinic tests, is also described in detail [14]. Many specific components of each individual test are proprietary. It is not clear what specific differences in the in-clinic assayed may have caused the observed performance differences between the tests.

Conclusions

We found that results of rapid in-clinic Giardia assays can vary amongst tests. Overall, of the in-clinic tests evaluated, the SNAP test most closely mirrored the results of the DFA and the microtiter plate ELISA and was the only test with a prevalence adjusted agreement greater than the O&P method. The major conclusion of this study is that among the in-clinic tests evaluated, Giardia SNAP is the most reliable method for detection of Giardia in canine stool samples.

Table 1
1Statistical comparison of ova and parasite results and reference laboratory ELISA to DFA with calculated sensitivity and specificity Percent agreement was adjusted for reported prevalence of infection determined by DFA as described in materials and methodsIn-clinic test resultsAmong the in-clinic test results, there was a single invalid result with Abaxis VetScan (positive control line did not appear) and therefore there were only 176 comparisons with this test. In all other cases, there were 177 valid results. The in-clinic tests results were comparedReference laboratory test
Sensitivity (95% CI)
Specificity (95% CI)
Prevalence adjusted % agreement a

Ova and Parasite (O&P) Test
81.2 (72.4-87.7)
93.4 (85.2-97.5)
92.7

ProSpecT Microtiter Plate ELISA
94.1 (87.4-97.5)
97.4 (90.4-99.8)
97.2

a

Table 2
2Statistical results of four in-clinic antigen tests for Giardia using DFA as the reference standard with calculated sensitivity and specificity Percent agreement was adjusted for reported prevalence of infection determined by DFA as described in materials and methodsIn-clinic test
Sensitivity (95% CI)
Specificity (95% CI)
Prevalence adjusted % agreement a

SNAP Giardia
87.1 (79.1-92.5)
93.4 (85.2-97.5)
93.1

Anigen Rapid CPV-CCV-Giardia Antigen Test
80.2 (71.3-86.9)
80.3 (69.8-87.8)
80.3

Witness Giardia Test
73.3 (63.9-81.0)
71.1 (60.0-80.1)
71.2

VetScan Canine Giardia Rapid Test
70.00 (60.4-78.1)
85.5 (75.7-91.9)
84.7

Ova and Parasite Test
81.2 (72.4-87.7)
93.4 (85.2-97.5)
92.7

a

Table 3
3Statistical results of four in-clinic antigen tests for Giardia using ProSpecT ELISA Giardia Test as the reference standard with calculated positive and negative percent agreement In-clinic test % Positive agreement (95% CI) % Negative agreement (95% CI) Prevalence adjusted % agreement a Percent agreement was adjusted for reported prevalence of infection determined by DFA as described in materials and methodsSNAP Giardia
91.8 (84.4-96.0)
95.0 (87.5-98.4)
94.8

Anigen Rapid CPV-CCV-Giardia Antigen Test 80.4 (71.3-87.2)
77.5 (67.1-85.4)
77.7

Witness Giardia Test
77.3 (68.0-84.6)
73.8 (63.1-82.2)
74.0

VetScan Canine Giardia Rapid Test
72.9 (63.2-80.8)
86.3 (76.9-92.3)
85.6

a

Abbreviations DFA: Direct immunofluorescence assay; ELISA: Enzyme-linked immunosorbent assay; IRL: IDEXX Reference Laboratories; O&P: Ova and parasite testing, zinc sulfate fecal flotation; PBS: Phosphate-buffered saline 0.02M pH 7.4; ZnSO 4 : Zinc sulfate flotation solution
AcknowledgementsThe authors thank IDEXX (Westbrook, ME) for generously providing the canine fecal samples used in this study. IDEXX had no input on the manuscript, results or interpretation of the results.FundingNot applicable.Availability of data and materialsData supporting the conclusions of this article are included within the article. The datasets used or analyzed during the present study are available from the corresponding author upon reasonable request.Authors' contributions JLL and JMB performed experiments and DDB performed analyses. All authors read and approved the final manuscript.Ethics approval and consent to participate Not applicable.Consent for publicationNot applicable.Competing interestsThe authors declare they have no competing interests.Publisher's NoteSpringer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
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PD-1 deficiency promotes TFH cells expansion in ITV-immunized mice by upregulating cytokines secretion

Taiping Liu
Xiangyun Cheng
Yan Ding
Feng Zhu
Yong Fu
Xiaohong Peng
Wenyue Xu
PD-1 deficiency promotes TFH cells expansion in ITV-immunized mice by upregulating cytokines secretion
10.1186/s13071-018-2984-4R E S E A R C H Open Access
Background: T follicular helper (TFH) cells are fundamental for the development of humoral immunity. In our previous study, we found that PD-1 deficiency substantially promoted the expansion of Plasmodium-specific TFH cells and enhanced the humoral immunity of ITV (infection treatment vaccine)-immunized mice. However, the underlying mechanism by which PD-1 signaling modulates TFH cells activation remains unclear.Methods: Mice were immunized with the ITV following the standard procedures. The activation phenotype of CD11c + CXCR5 + dendritic cells (DCs), the frequency and number of splenic follicular regulatory T cells (TFR cells), Plasmodium-specific TFH cells and germinal center (GC) B cells were analyzed by FACS. The levels of serum cytokines were quantified using the cytometric bead array (CBA) and in vivo cytokine neutralization was carried out according to a previously described protocol and verified by serum cytokine detection.Results:We found that PD-1 -/naïve and immunized mice had more TFR cells in the spleen than WT and WT immunized mice. Additionally, CXCR5 + DC, which prime TFH cells, were activated at similar levels in ITV-immunized WT and PD-1 -/mice. However, the serum levels of IL-10, IFN-γ and MCP-1 were significantly increased in ITV-immunized PD-1 -/mice, and treatment with an anti-IL-10, anti-IFN-γ or anti-MCP-1 neutralizing antibody in vivo markedly impaired the development of TFH cells and GC B cells. Conclusions: Our findings demonstrate that the modulation of TFH cells by PD-1 signaling is dependent on the cytokines IL-10, IFN-γ and MCP-1 in ITV-immunized mice. These results could facilitate the design of an effective malaria vaccine with the aim of inducing humoral immune responses.

Background

T follicular helper (TFH) cells, a particular subpopulation of CD4 + T cells, are the key cell type required for the formation of germinal centers (GCs) and the generation of long-lasting antibody responses [1,2]. These cells are characterized by high expression of the chemokine receptor CXCR5, programmed death 1 (PD-1), a master regulator transcription factor (Bcl6), IL-21, and the inducible T-cell co-stimulator (ICOS) [3]. Inducing TFH cell activation is considered to be a promising strategy for improving host immune protection induced by vaccines [4,5].

Malaria is a major global health burden, causing 212 million clinical episodes and 429,000 deaths in 2015, mostly in Africa in children under five years old [6]. The development of an effective and durable vaccine is considered to be the most cost-effective strategy for the prevention, control and eventual eradication of this disease [7]. The main approach in blood-stage malaria vaccination is to induce antibodies that can prevent merozoites from invading erythrocytes [8]. However, the need for high titers of Plasmodium-specific antibodies remains a significant clinical hurdle [9]. Therefore, understanding how Plasmodium-specific B cells are activated during vaccination could help guide the design of a more effective malaria vaccine. The infection treatment vaccine (ITV), a whole blood-stage parasite vaccine, induces protective immunity that is primarily mediated by antibodies specific for merozoites surface antigens [10,11]. By using this kind of whole blood-stage parasite vaccine mode, our previous study showed that PD-1 deficiency substantially promoted the expansion of Plasmodiumspecific TFH cells, which may enhance the Plasmodiumspecific B cell response [12]. However, it remains unknown how PD-1 signaling regulates TFH cell activation in ITVimmunized mice.

TFH cell activation is a complex and tightly regulated process. Current studies suggest that TFH cells are primed by CXCR5 + dendritic cells (DCs) and fully differentiate into mature GC TFH cells with the aid of cognate B cells [13,14]. In addition, developing TFH cells could be influenced by environmental cues, including cytokines (such as IL-6 and IL-21) and chemokines (such as CXCL13) [2,15,16]. Recently, a subset of Foxp3 + Tregs, follicular regulatory T cells (TFR cells) that inhibit TFH cell-mediated humoral immunity, has been identified [17]. Previous studies have shown that PD-1 or PD-L1 deficiency results in the presence of more TFR cells than TFH cells in lymph nodes and potently inhibits antibody production [18]. However, therapeutic in vivo blockade of PD-L1 in mice has been found to enhance the differentiation of Plasmodium-specific TFH cells and plasmablasts during malaria infection [19]. Therefore, the mechanisms by which PD-1 signaling modulates TFH cell activation remain unclear.

In this study, we found that neither CXCR5 + DCs nor TFR cells were involved in regulating Plasmodium-specific TFH cells activation in ITV-immunized PD-1 -/mice. However, elevated levels of IL-10, IFN-γ and MCP-1 substantially contribute to the expansion of Plasmodiumspecific TFH cells in ITV-immunized PD-1 -/mice. Therefore, we have revealed a novel mechanism by which PD-1 signaling regulates TFH cells activation.

Methods

Mice and malaria parasite strain

PD-1 -/mice (BALB/c background) were obtained from the Jackson Laboratory (Bar Harbor, ME, USA). Specific pathogen-free BALB/c mice were purchased from the Beijing Animal Institute. All mice ranged in age from six to eight weeks when the experiments were initiated. The lethal strain of Plasmodium yoelii 17XL was originally obtained from MR4 (Malaria Research and Reference Reagent Resource Center, Manassas, VA, USA) and maintained as cryopreserved stabilates. All animal studies were reviewed and approved by the Animal Ethics Committee of the Third Military Medical University Institute of Medical Research.

Immunization

The mice were immunized following a previously described immunization schedule [12]. Briefly, the mice were intravenously (i.v.) injected with 10 6 P. yoelii 17XL-infected red blood cells (RBCs) (Py-iRBCs) or a matching number of normal RBCs (nRBCs) (negative control). All mice were then intraperitoneally (i.p.) injected with 100 μl of 8 mg/ml chloroquine (CQ; Sigma-Aldrich, St. Louis, MO, USA) diluted in saline daily for 15 days starting the day of the iRBC injection. The absence of parasites was confirmed by Giemsa-stained blood smears in all treated mice from the beginning of CQ treatment.

Flow cytometric analysis

Spleens were collected on the appropriate days, and single-cell suspensions of splenocytes were prepared as previously described [20]. To determine the activation phenotype of CXCR5 + DCs by evaluating the expression of CD40, CD86 and MHC class II (MHC-II), 10 6 cells were pre-incubated with anti-CD16/CD32 (BioLegend, San Diego, CA, USA) to block non-specific binding to Fc receptors. Then, the cells were washed and subsequently stained with anti-mouse CXCR5 (biotin; BioLegend), streptavidin (APC; BioLegend), anti-mouse CD11c (percp/ Cy5.5; BioLegend), anti-mouse CD40 (PE; BioLegend), anti-mouse CD86 (PE/Cy7; BioLegend) and anti-mouse MHC-II (FITC; BioLegend). To analyze GC B cells, 10 6 splenocytes were blocked as described above. Then, the cells were washed and stained with anti-mouse B220 (APC; BioLegend), anti-mouse CD95 (PE; eBioscience, San Diego, CA, USA), and anti-mouse T and B cell activation marker (GL-7) (FITC; BioLegend). To analyze TFR cells, 2 × 10 6 cells were stained with anti-mouse CXCR5 (biotin; BioLegend), streptavidin (APC; BioLegend), anti-mouse CD4 (APC/Cy7; BioLegend), anti-mouse CD19 (percp/Cy5.5; BioLegend), anti-mouse ICOS (PE/Cy7; BioLegend) and anti-mouse Foxp3 (PE; eBioscience) after being permeabilized with a fixation/permeabilization agent (eBioscience). For Plasmodium-specific TFH cell analysis, cells were stained for CXCR5, CD4 and ICOS as described above. The cells were also stained with anti-mouse CD19 (Pacific Blue; BioLegend), anti-mouse CD11a (percp/Cy5.5; BioLegend), anti-mouse CD49d (FITC; BioLegend), and anti-mouse Foxp3 (PE; eBioscience) or anti-mouse Bcl6 (PE; eBioscience) and anti-mouse Foxp3 (PE/Cy7; eBioscience) after being permeabilized. The cells were analyzed on a FACSCanto II instrument (BD Biosciences, San Jose, CA, USA), and the data were analyzed with FlowJo software.

Serum cytokine detection

The levels of the proinflammatory cytokines IL-6, monocyte chemoattractant protein-1 (MCP-1), IFN-γ, TNF-α, and IL-12p70 and the anti-inflammatory cytokine IL-10 were quantified in serum samples using the cytometric bead array (CBA) Mouse Inflammation Cytokine Kit (BD Biosciences), according to the manufacturer's instructions.

Briefly, mouse inflammation standards were prepared with 2 ml of assay diluent and six mouse inflammation capture beads were mixed thoroughly. Then, 50 μl mixed capture beads were incubated with the same volume of mouse inflammation standard dilutions or each samples. Fifty microliters of the mouse inflammation PE detection reagent was added to all assay tubes and the assay tubes were incubated for 2 h at room temperature, protected from light. After being washed with washing buffer, samples were analyzed on a FACSCanto II instrument (BD Biosciences), and the data were analyzed with FCAP Array software.

In vivo cytokine neutralization

To neutralize MCP-1, IFN-γ or IL-10 in vivo, mice were respectively i.p. injected with 200 μg (per mouse) of a neutralizing mouse anti-MCP-1 mAb (BioXcell, clone: 2H5), anti-IFN-γ mAb (BioXcell, clone: XMG 1.2), anti-IL-10 mAb (BioXcell, clone: JES5-2A5) or control Ab every other day starting from one day after the last CQ injection until sacrifice, according to a previously described protocol with minor modifications [21]. Cytokine depletion was verified by serum cytokine detection as described above.

Statistical analysis

The data were analyzed using GraphPad Prism version 6 software. Nonparametric tests (Mann-Whitney test) and two-way ANOVA were used to compare groups, and P-values < 0.05 were considered statistically significant.

Results

The activation of CD11c + CXCR5 + DCs was comparable between WT and PD-1-deficient ITV-immunized mice First, the parasitemia (Fig. 1b) and survival rate (Fig. 1c) were recorded in all treated mice from the beginning of CQ treatment. As shown in Fig. 1, the parasitemia in all immunized mice appeared in day 1 and disappeared in day 5 post-infection and all mice survived during the process of CQ injection. Previous studies have found that CD11c + CXCR5 + DCs preferentially initiate TFH cell differentiation [13]. Therefore, the total number and maturation phenotype of CD11c + CXCR5 + DCs in the spleen were analyzed in ITV-immunized WT mice and PD-1 -/mice 2, 4 and 6 days after the initial immunization. As shown in Fig. 2, the total number of CD11c + CXCR5 + DCs in the spleen was comparable between WT and PD-1 -/mice, indicating no intrinsic defect in PD-1 -/mice. Although immunized mice have greater number of CD11c + CXCR5 + DCs than naïve mice in the spleen (ANOVA: F (3, 54) = 89.588, P = 0.025), there was no significant difference between the ITV-immunized WT mice and PD-1 -/mice (Fig. 2a). At resting state, the expression of CD40, CD86, and MHC-II was similar between the WT and PD-1 -/mice (Fig. 2b, c), indicating that there was no intrinsic DC defect in the absence of PD-1. Although the activation of CD11c + CXCR5 + DCs from immunized mice was significantly increased compared with DCs from naïve mice (ANOVA: F (3, 54) = 33.831, P < 0.05), no significant differences in the expression of CD40, CD86, and MHC-II on the surface of CD11c + CXCR5 + DCs were detected between WT and PD-1 -/immunized Fig. 1 The parasitemia and survival rate in all treated mice from the beginning of CQ treatment. a The procedure for the ITV immunization and experiment operation. b, c WT (n = 5) or PD-1 -/mice (n = 5) were intravenously (i.v.) injected with 10 6 P. yoelii 17XL-infected red blood cells (RBCs) (Py-iRBCs) or a matching number of normal RBCs (nRBCs) (negative control). The parasitemia and survival rate were recorded from the beginning of CQ treatment. Three independent experiments were performed. The data are presented as the mean ± SD. Data were compared with the nonparametric Mann-Whitney test. Abbreviation: n, number of samples mice (Fig. 2b, c). Thus, these results suggest that the expansion of Plasmodium-specific TFH cells in ITV-immunized PD-1 -/mice may not be attributable to enhanced DC function.

The absence of PD-1 strongly increased the number of TFR cells in ITV-immunized mice TFR cells can inhibit humoral immunity by modulating TFH cell activity [17]. To test whether the increased expansion of TFH cells in ITV-immunized PD-1-deficient mice was a result of the inhibition of TFR cell activity, the frequency and number of splenic TFR cells were compared between immunized WT and PD-1 -/mice. We defined TFR cells as CD4 + ICOS + CXCR5 + Foxp3 + CD19 − as previously reported [18] and analyzed TFR cells on days 7, 14 and 21 after the final injection of CQ. The frequency and number of TFR cells in both ITV-immunized WT mice and PD-1 -/mice gradually decreased from day 7 to 14 and then increased from day 14 to 21 after the final injection of CQ (Fig. 3b, c), suggesting that the immune-protection effect induced by ITV may be attributable to impaired TFR cell function. However, the frequency and number of TFR cells were much higher in PD-1 -/naïve and immunized mice than in WT naïve and Fig. 2 The total number and maturation phenotype of CD11c + CXCR5 + DCs were comparable between immunized WT and PD-1 -/mice. a-c Splenocytes were isolated from the immunized WT (n = 5) and PD-1 -/mice (n = 5) at 2, 4 and 6 days after the initial immunization. a Statistical analysis of the total number of CD11c + CXCR5 + DCs in the spleen from ITV-immunized WT and PD-1 -/mice on days 2, 4 and 6 after the initial immunization. b Gating strategy and FACS analysis of the expression of CD40, CD86, and MHC-II on CD11c + CXCR5 + DCs from immunized WT (thin line) and PD-1 -/-(bold line) mice. Spleen lymphocytes were gated as CD11c + CXCR5 + , and the expression levels of CD40, CD86 and MHC-II were analyzed. c A graphical representation of the geometrical mean of the immunofluorescence intensity of the expression of all three markers. Three independent experiments were performed. The data are presented as the mean ± SD. Data were compared with two-way ANOVA. Abbreviation: n, number of samples immunized mice after the final injection of CQ (ANOVA: F (3, 54) = 55.609, P < 0.05) (Fig. 3b, c), which is consistent with a previous report [18]. Therefore, these data suggest that the expansion of Plasmodium-specific TFH cells is not due to a reduction in the number or frequency of TFR cells in ITV-immunized PD-1 -/mice.

The levels of IL-10, IFN-γ and MCP-1 were significantly increased in PD-1-deficient ITV-immunized mice

Considering the important role of cytokines in supporting TFH cell differentiation [22,23], we compared the serum cytokine level between the immunized WT and PD-1 -/mice at 7, 14 and 21 days after the last CQ injection. The concentrations of IL-6, MCP-1, IFN-γ, TNF-α, IL-12p70 and IL-10 in serum were measured using a CBA. As shown in Fig. 4, the level of IL-12p70 was comparable between the two kinds of immunized mice. Although higher levels of IL-6 and TNF-α were detected in PD-1 -/immunized mice than in immunized WT mice on day 7 after the final CQ injection, these cytokines were also higher in PD-1 -/naïve mice than in WT naïve mice. Interestingly, although no significant difference was found at either day 14 or 21 after the final injection of CQ, significant increases in MCP-1 (224.89 ± 18.56 vs 20.91 ± 8.56 pg/ml; ANOVA: F (3, 54) = 9.059, P < 0.001), IFN-γ (585.64 ± 70.38 vs 48.58 ± 6.82 pg/ml; ANOVA: F (3, 54) = 9.158, P < 0.001) and IL-10 (83.45 ± 6.06 vs 18.75 ± 5.5 pg/ml; ANOVA: F (3, 54) = 10.258, P = 0.001) in PD-1 -/immunized mice were observed on day 7 after the final injection of CQ (Fig. 4), indicating that these cytokines may be involved in Plasmodium-specific TFH cell differentiation. Thus, these data suggest that increased levels of MCP-1, IFN-γ and IL-10 may promote Plasmodium-specific TFH cells expansion.

Elevated cytokine levels substantially contributed to the expansion of TFH cells and GC B cells in immunized PD-1 -/mice

To determine whether elevated levels of cytokines (MCP-1, IFN-γ and IL-10) could promote TFH cells expansion in PD-1 -/immunized mice, we performed blocking experiments using neutralizing mAbs against MCP-1, IFN-γ, or IL-10 beginning one day after the last CQ injection. The frequency and number of splenic Plasmodium-specific TFH cells were then compared between the two groups of immunized mice. According to a previous study, Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + ICOS + or Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + Bcl6 + cells are considered to be Plasmodium-specific TFH cells [12,18]. The Plasmodium-specific TFH cell frequency and number peaked on day 7 after the final CQ injection and then gradually decreased over time. Interestingly, The data are presented as the mean ± SD. Data were compared with the two-way ANOVA. Abbreviation: n, number of samples compared with normal PD-1 -/immunized mice, the frequency and number of Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + ICOS + or Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + Bcl6 + TFH cells decreased by nearly half in mice treated with blocking mAbs on day 7 after the final CQ injection (ANOVA: F (4, 44) = 19.208, P < 0.05), although no significant difference in the number of Plasmodiumspecific TFH cells was found on day 14 after the final CQ injection (Fig. 5b, c, e, f).

The most important biological function of TFH cells is to aid GC B cells in the generation of GCs and long-term protective humoral responses [1]. Thus, to further confirm the roles of MCP-1, IFN-γ and IL-10 in the regulation of TFH cell expansion, the frequency and number of GC B cells in the spleen were also detected on days 7 and 14 after the final CQ injection. As shown in Fig. 6, the GC B cells frequency and number decreased significantly in ITV-immunized PD-1 -/mice after treatment with blocking mAbs on day 14 (ANOVA: F (4, 44) = 12.790, P < 0.001), but no significant difference was found on day 7 after the final injection of CQ. Taken together, these results demonstrate that the cytokines MCP-1, IFN-γ and IL-10 substantially contribute to the expansion of TFH cells and GC B cells in immunized PD-1 -/mice.

Discussion

PD-1 and its ligands (PD-L1 and PD-L2) deliver inhibitory signals, which can regulate the balance between T cell activation and tolerance. Evidence has already confirmed that PD-1 is highly expressed on GC TFH cells and expected to provide an inhibitory signal to GC TFH cells, preventing excess CD4 + T cell proliferation in GC [16,24]. However, the precise mechanisms by which PD-1 signaling modulates TFH cell activation remain unclear. Our previous study found that PD-1 deficiency substantially promoted the expansion of Plasmodiumspecific TFH cells in ITV-immunized mice, which focused on the whole blood-stage malaria parasites [12]. However, the mechanism of Plasmodium-specific TFH cell activation in PD-1 -/immunized mice remains to be elucidated. In this study, we demonstrated that the cytokines MCP-1, IFN-γ and IL-10 participate in the induction of Plasmodium-specific TFH cell expansion in PD-1 -/immunized mice. Although recent studies have revealed that IFN-γ and IL-10 signaling can modulate TFH cell differentiation, reports regarding the function of IL-10 signaling in modulating TFH cell activation are contradictory [25][26][27][28]. Here, we found that IL-10 neutralization significantly inhibited Plasmodium-specific TFH cell proliferation in PD-1 -/immunized mice. This is consistent with a previous study that also focused on pathogen-specific TFH cells instead of total TFH cells [25]. Thus, we hypothesized that IL-10 signaling could have differential effects on the differentiation of total TFH cells and that of pathogen-specific TFH cells. Studies have found that IFN-γ can act directly on TFH cells to promote Fig. 4 MCP-1, IFN-γ and IL-10 were significantly elevated in the serum of ITV-immunized PD-1 -/mice. Serum samples were collected from immunized WT (n = 5) and PD-1 -/mice (n = 5) at the indicated times after the final immunization and the concentrations of cytokines were measured serially by a CBA. Three individual experiments were performed. The data are presented as the mean ± SD. Data were compared with the two-way ANOVA. Abbreviation: n, number of samples their expansion [28], while IL-10 can act indirectly on T cells to substantially dictate the differentiation state and function of TFH cells [25]. Surprisingly, our results also showed that MCP-1 participates in Plasmodium-specific TFH cell expansion. To the best of our knowledge, this is the first study to reveal the relationship between MCP-1 and TFH cell differentiation. Evidence has confirmed that MCP-1 is mainly produced by monocytes/macrophages [29]. Thus, in addition to eliminating Plasmodium-infected RBCs (iRBCs) [30], monocytes/macrophages may also be involved in the regulation of humoral immunity via the secretion of MCP-1.

Another important issue is the cell types that secreted the MCP-1, IFN-γ and IL-10 observed in PD-1 -/immunized mice. Although CD4 + T cells, CD8 + T cells, macrophages and DCs have been analyzed for their ability to secrete these cytokines (data not shown), we have not yet identified the major cell type producing MCP-1, IFN-γ and IL-10. It is possible that many types of cells secrete these cytokines in PD-1 -/immunized mice. A recent study Fig. 5 Frequency and number of Plasmodium-specific TFH cells in the spleens of ITV-immunized WT and PD-1 -/mice. Splenocytes were isolated from immunized WT (n = 5) and PD-1 -/mice (n = 5) on days 7 and 14 after the final immunization and the Plasmodium-specific TFH cells were analyzed by FACS. a Representative FACS analysis of Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + ICOS + Plasmodium-specific TFH cells. b, c Statistical analysis of the frequency and number of Plasmodium-specific TFH cells (Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + ICOS + ) from immunized WT and PD-1 -/mice. d Representative FACS analysis of Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + Bcl6 + Plasmodium-specific TFH cells. e, f Statistical analysis of the frequency and number of Plasmodium-specific TFH cells (Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + Bcl6 + ) from immunized WT and PD-1 -/mice. Three individual experiments were performed. The data are presented as the mean ± SD. Data were compared with two-way ANOVA. Abbreviation: n, number of samples showed that PD-1 signaling deficiency can enhance the secretion of IL-10 and IFN-γ in allogenic cultures of CD4 + T cells with various DC populations [31], which may provide clues for further experiments. Meanwhile, our results showed that the elevated levels of MCP-1, IFN-γ and IL-10 in serum were observed on day 7 after the final injection of CQ. Considering the parasitemia in immunized mice appeared on day 6 and disappeared on day 12 after the final injection of CQ (Additional file 1: Figure S1), we hypothesized that this phenomenon may correlate with the complete elimination of iRBCs in ITV-immunized mice in the early stage of CQ withdrawal.

Recent evidence has suggested that the engagement of CXCR5 on DCs may preferentially condition them to directly prime TFH cell responses [1,13]. However, unlike the maturation profiles and phenotype of DCs in mice immunized with the whole-killed P. yoelii 17XL vaccine (WKV) [20], we found that CXCR5 + DCs exhibited unique expression patterns of CD40, CD86 and MHC-II in ITV-immunized mice. The levels of CD40, CD86 and MHC-II on the CXCR5 + DCs surface were significantly higher in immunized mice than in naïve mice after the primary immunization with WKV. However, the expression level of CD40 gradually increased over time, and a significant difference between immunized mice and naïve mice only appeared on day 6 after the initial ITV immunization. In contrast to CD40, the expression of CD86 began to gradually decrease two days after the first immunization. In addition, differences in the expression of MHC-II were mainly apparent on day 2 and day 6 after the initial immunization. This is partially consistent with a recent study, which also found that each activated DC phenotype exhibited a different expression pattern during P. yoelii infection [32]. Thus, we hypothesized that pRBC lysates and live iRBCs may affect the phenotypic and functional maturation of DCs via different mechanisms. To the best of our knowledge, up to now there has not been any report concerning the relationship between the PD-1 signal and CXCR5 + DC activation. Our results showed that the total number of CD11c + CXCR5 + DCs in the spleen was comparable between WT and PD-1 -/mice, immunized WT and immunized PD-1 -/mice, which suggests that PD-1 signal may not be involved in the regulation of CXCR5 + DC activation.

TFR cells are a subset of Foxp3 + Tregs, which were found within GC and suppress the magnitude and longevity of TFH cells and GC responses [1,33]. TFR cells express many canonical TFH cell molecules, including PD-1, CXCR5, ICOS and Bcl6 and Foxp3 is currently considered to be the most important marker that distinguishes TFR cells from TFH cells [18]. Therefore, Foxp3cells were selected for further analyses of TFH cells in our study. Meanwhile, studies have found that the integrins CD49d and CD11a on antigen-experienced CD4 + T cells could be used to identify Plasmodium-specific CD4 + T cells [19], and therefore Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + ICOS + or Foxp3 -CD19 -CD4 + CD11a + CD49d + CXCR5 + Bcl6 + cells were regarded as Plasmodium-specific The data are presented as means ± SD. Data were compared with two-way ANOVA. Abbreviation: n, number of samples TFH cells in our study. Although our results also confirmed that PD-1 signaling decreased the number of TFR cells, the precise regulatory mechanism of TFR cell activation remains largely unknown. A recent study has found that PD-L1, one of the PD-1 ligands which are expressed on DC, is essential for limiting TFR cell differentiation [34]. These results may provide new ideas for further research. In addition, we found that the frequency and number of TFR cells significantly decreased from day 7 to 14 and gradually increased from day 14 to 21 after the final injection of CQ in ITV-immunized mice. Therefore, these results indicate that humoral immunity induced by ITV may be associated with the impaired function of TFR cells, which would have implications in the design of effective malaria vaccines.

Conclusions

In summary, we demonstrated that PD-1 deficiency significantly increased the levels of MCP-1, IFN-γ and IL-10 in the serum of ITV-immunized mice. We further confirmed that these elevated cytokines substantially contributed to the expansion of Plasmodium-specific TFH cells and GC B cells in PD-1 -/immunized mice. Thus, our findings not only have implications for understanding the activation mechanism of Plasmodium-specific TFH cells but also contribute to our knowledge regarding the rational design of an effective malaria vaccine.

Additional file

Additional file 1: Figure S1. The parasitemia in ITV-immunized mice after the final injection of CQ. After the last CQ injection, the parasitemia was recorded in four groups. The data are presented as the mean ± SD. Data were compared with the nonparametric Mann-Whitney test. (TIF 4538 kb)

Fig. 3
3PD-1 deficiency contributed to the expansion of TFR cells in both naïve and immunized mice. Splenocytes were isolated from immunized WT (n = 5) and PD-1 -/mice (n = 5) at the indicated times after the final immunization, and both the frequency and number of TFR cells were analyzed by FACS. a Representative FACS analysis of CD4 + ICOS + CXCR5 + Foxp3 + CD19 − cells. b, c Statistical analysis of the frequency and number of TFR cells from ITV-immunized WT and PD-1 -/mice on days 7, 14 and 21 after the final immunization. Three individual experiments were performed.

Fig. 6
6Frequency and number of GC B cells in the spleens of ITV-immunized WT and PD-1 -/mice. Splenocytes were isolated from immunized WT (n = 5) and PD-1 -/mice (n = 5) on days 7 and 14 after the final immunization and both the frequency and number of GC B cells were analyzed by FACS. a Representative FACS analysis of B220 + GL-7 + CD95 + GC B cells. b, c Statistical analysis of the frequency and number of GC B cells from ITV-immunized WT and PD-1 -/mice. Three individual experiments were performed.

Abbreviations CQ: Chloroquine; DCs: Dendritic cells; GCs: Germinal centers; ICOS: Inducible T-cell co-stimulator; iRBCs: Plasmodium-infected RBCs; ITV: Infection treatment vaccine; MCP-1: Monocyte chemoattractant protein-1; MHC-II: MHC class II; PD-1: Programmed death 1; Py-iRBCs: P. yoelii 17XL-infected red blood cells; TFH cells: T follicular helper cells; TFR cells: Follicular regulatory T cells; WKV: Whole-killed P. yoelii 17XL vaccine
AcknowledgmentsWe thank the Malaria Research and Reference Reagent Resource Center (MR4) for providing P. yoelii 17XL.FundingThis work was supported by the National Natural Science Foundation of China, nos. 81702021 and 81471976, and the Natural Science Foundation of Chongqing, nos. cstc2016jcyjA0464 and cstc2016jcyjA0485.Availability of data and materialsThe data supporting the conclusions of this article are included within the article.Authors' contributions WX conceived the experiment and wrote the paper. TL, XC, YD, FZ, YF and XP performed the research and analyzed the data. All authors read and approved the final manuscript.Consent for publicationNot applicable.Competing interestsThe authors declare that they have no competing interests.
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