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. 2025 Nov 29;45:102389. doi: 10.1016/j.bbrep.2025.102389

New insight into role of mast cells in erythrocyte homeostasis and clearance under oxidative stress conditions in vivo

Priyanka Sharma 1, Mohd Suleman 1, Niti Puri 1,⁎
PMCID: PMC12704209  PMID: 41404112

Abstract

Erythrocytes are exposed to reactive oxygen species during circulation constantly, due to normal aerobic cellular metabolism and pathology of inflammatory diseases. Genetic depletion of mast cells and systemic mastocytosis seem to affect anaemia. We attempted to reveal the participation of mast cells in erythrocytes clearance in vivo and during interaction with professional phagocytes like macrophages in vitro. Mastocytosis was induced in DBA/2 mice by injecting P815 cells and mast cell depletion was attained with compound 48/80 treatment under normal or phenylhydrazine (PHZ) induced oxidative stress conditions in mice. The mastocytosis model showed a significant decrease in circulatory erythrocytes while mast cell absence led to significantly high accumulation of erythrocytes in spleens of mast cell depleted anemic mice. The present study provides important evidences for contribution of mast cells in erythrocyte clearance during oxidative stress conditions and augmented uptake of oxydatively damaged erythrocytes (ODE) in presence of macrophages in close proximity.

Keywords: Mast cells, Erythrocytes, Mastocytosis, Compound 48/80, Phenylhydrazine, Oxidative stress

Graphical abstract

Image 1

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Highlights

  • •

    Mast cells play a role in clearance of circulatory erythrocytes.

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    Local peritoneal mast cell depletion does not affect circulatory erythrocytes.

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    Mast cell depletion reduces erythrocyte clearance in spleen in oxidative stress.

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    Mast cells show augmented uptake of ODE during direct contact with macrophages.

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    Macrophages show reduced uptake of ODE during direct contact with mast cells.

1. Introduction

The interconnection of anemia, inflammation, and oxidative stress is well known. During normal or inflammatory conditions erythrocytes are constantly exposed to oxidative stress. The interaction of mast cells with erythrocytes is of great interest but very few studies have been conducted to date as few models are available and it is very difficult to purify primary mast cells due to their tissue specific localization. Mastocytosis, the condition of variable mast cell hyperplasia has been reported to be directly linked with abnormal blood count and anemia in almost 50 % of human patients [1,2]. It has also been reported that pharmacological inhibition or genetic depletion of mast cells in mice during sickle cell anemia reduced systemic or neurogenic inflammation and also pathophysiological consequences of sickle cell anemia [3]. Mast cells, apart from their major participation during inflammatory and allergic process are now widely known as non professional phagocytes. Phagocytosis of a number of pathogens like bacteria, parasites, yeast cells and even particulate materials like latex beads, gold, ferritin, aggregated IgE and red blood cells by mast cells has been reported [[4], [5], [6], [7]]. Recently, direct uptake of ODE by mast cells in resting and activated state mimicking inflammatory conditions in vitro and by peritoneal mast cells in vivo has been reported by us [8]. Oxidatively damaged erythrocytes (ODE) are those that have been damaged due to some oxidative stress and exhibit specific hallmarks of acute stress, including hemolysis, loss of membrane integrity, and the oxidation of hemoglobin [9]. Critically, these oxidatively compromised cells present senescent markers, primarily the externalization of phosphatidylserine (PS), which acts as the ‘eat me’ signal for phagocytic clearance [8]. It is for this functional overlap - where oxidative damage directly triggers the display of the senescence marker PS that the terms oxidative stress and senescence can be employed synonymously regarding the clearance signal itself. However, it is important to recognize the biological distinction that not every senescent erythrocyte population is necessarily damaged solely by oxidative mechanisms.

Several mechanisms have been postulated for the clearance of senescent or damaged erythrocytes by macrophages [10]. Role of mast cells in erythrocyte clearance for maintaining homeostasis under normal or during oxidative stress conditions, inflammation or anemic condition in vivo is still to be elucidated. The presence of mast cells and macrophages in close proximity suggests the possibility of their interactions that may affect the release of mediators and functions of both cell types. Mast cell – macrophage interaction has been reported to be beneficial in preventing tumor metastasis [11]. The interaction between mast cells and macrophages may be a significant factor in clearance of erythrocytes by erythrophagocytosis.

In the present study, firstly we attempted to investigate the role of mast cells in erythrocyte clearance during normal and oxidative stress conditions in vivo. The effect on erythrocyte levels was observed in different mice models like mastocytoma model with high mast cell numbers and mast cell depleted model in normal and anemic/oxidative stress conditions. Secondly, the impact on the uptake of erythrocytes was observed by both cell types individually in vitro after their co-culture in direct and indirect contact with each other. The present study reveals the unique role of mast cells in anemia during inflammatory, autoimmune and oxidative stress conditions which might be useful to avoid the adverse anemic disorders.

2. Material and methods

2.1. Animals

Breeding pairs of C57BL/6 mice (inbred) were obtained from the National Institute of Nutrition, Hyderabad, India and DBA/2 mice were obtained from Indian Institute of Integrative Medicine (IIIM), Jammu, India. SWISS mice (outbred strain) were obtained from Central Laboratory Animal Resources (CLAR) Jawaharlal Nehru University (JNU), New Delhi, India. All animals (8–12 weeks old of either sex) used in the present study were bred and maintained in CLAR at JNU in positive pressure air-conditioned units (25 °C, 50 % relative humidity) under sterilized conditions at equal light and dark cycle (12 h). Both food and water were provided ad libitum. All experimental protocols were approved by the JNU Institutional Animal Ethical Committee (IAEC code 13/2013).

2.2. Culture and maintenance of cell lines

Murine P815 mastocytoma cells were obtained from National Centre for Cell Sciences (NCCS), Pune, India. Cells were maintained as suspension culture (some adherent) in Dulbecco's Modified Eagle's Medium (DMEM) (Sigma Aldrich), supplemented with, 50 μg/ml gentamicin sulfate and 10 % fetal bovine serum (FBS) (Gibco, Life technologies, Grand Island, NY, USA) in a humidified atmosphere containing 5 % CO2 at 37 °C. Murine alveolar macrophage cell line (MH-S) was procured from American type tissue culture collection (ATCC, Rockville, MD, USA) and maintained in RPMI-1640 Medium (Sigma Aldrich) supplemented with 0.05 mM 2-mercaptoethanol, 50 μg/ml gentamicin sulfate and 10 % FBS.

2.3. Induction of mastocytosis in DBA/2 mice

Mastocytosis was induced to DBA/2 mice as described elsewhere [12] with minor modifications. In brief, 1 million P815 mastocytoma cells were injected retro-orbitally to mice (male or female) and equal volume of phosphate buffer saline (PBS) was injected to control group in parallel. The detailed timeline for protocol of mastocytosis induction is depicted in Fig. 1A. After 8 days, mice were euthanized and peripheral blood was collected for analyzing blood parameters and white blood cells (WBC) staining for mast cells respectively before euthanization. Peritoneal lavage and spleen were isolated, red blood cells (RBC) were lysed and WBC pellets from peripheral blood, peritoneal lavage and spleen were stained with Phycoerythrin (PE) conjugated anti-mouse CD45, Allophcocyanin (APC) conjugated anti-mouse CD117 (c-kit) and their isotype control antibodies (Biolegend, San Diego, CA) after blocking with anti-mouse CD16/32 (Fc block antibody). Cells were washed and analyzed on BD FACSCalibur flow cytometer using CellQuest Pro software (BD Biosciences).

Fig. 1.

Fig. 1

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Mastocytosis induction and mast cell characterization in DBA/2 mice. DBA/2 mice were injected with 1 X 106 P815 mastocytoma cells suspended in PBS (treated groups) or with equal volumes of PBS (control groups) retro-orbitaly. (A) Schematic showing detailed protocol for mastocytosis induction; (B) Circulatory WBCs were analyzed from whole blood using automated cell counter. WBC levels of control and treated groups in percent relative to respective pretreatment levels are shown; (C) mice were sacrificed after 8 days of P815 injection and spleens were isolated, spleen WBCs from control and treated groups were counted using hemocytometer after hypotonic lysis of erythrocytes. (D) WBC population form peripheral blood, spleen and peritoneal lavage samples were obtained after erythrocyte lysis and stained with PE anti mouse CD45 and APC anti mouse CD117 antibodies and their respective isotype controls in parallel. Representative FACS dot plots showing percent of total double positive mast cells in peripheral blood, spleen and peritoneal cells. (E) Circulatory platelets and (F) erythrocytes (RBC) were analyzed from whole blood using automated cell counter. Values are shown relative to pretreatment levels. Data is plotted as mean ± SEM from at least 5 mice per group (∗p ≤ 0.05).

2.4. Mast cell depletion by compound 48/80 administration in vivo

Peritoneal mast cell depletion was done with continuous administration of compound 48/80 (Sigma, MO, USA) for 4 consecutive days, with minor modifications to an earlier protocol [13]. Briefly, age and weight matched SWISS, C57BL/6, or DBA/2 mice were pre-bled for analyzing blood parameters and then, treated with compound 48/80 intraperitoneally for 4 days (1.2 mg/kg, twice a day for 3 days and 2.4 mg/kg once on the day 4) or PBS alone (200 μl) to control group in parallel. The schematic is depicted with timeline in Fig. 2 (A). Mice were bled on day 3 and 5 for blood parameter analysis and sacrificed on day 6. Total peritoneal cells were isolated to assess mast cells by toluidine blue (Sigma, MO, USA) staining and Flow cytometry using anti-mouse c-kit antibody.

Fig. 2.

Fig. 2

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Mast cell depletion model in SWISS mice. SWISS mice were injected with 1.2 mg/kg of compound 48/80 twice a day for 3 days and 2.4 mg/kg of compound 48/80 (once) on day 4 intraperitoneally (treated group) or PBS alone (control group) in parallel. (A) Schematic showing detailed protocol for mast cell depletion with compound 48/80 admininstration. (B) Mice were sacrificed on the day 6, peritoneal lavage was isolated and cell recovery from peritoneal lavage (after erythrocyte lysis) of control and compound 48/80 treated mice was observed by counting live cells with trypan blue staining using hemocytometer under light microscope. (C) Peritoneal cells from both groups were stained for mast cells with APC conjugated anti mouse CD117 (c-kit) antibody and representative FACS dot plots obtained from control and compound 48/80 treated groups respective to their isotype controls were analyzed on BD FACSCalibur. (D) 1 million of peritoneal cells were adhered, fixed and stained with 0.5 % touidine blue dye as mentioned in material and methods. Cells were observed under Nikon Eclipse-Ti microscope and images were captured at 40X magnification. Arrows indicating granule filled purple/pink mast cells and other cells were observed in blue colour. Whole blood isolated from tail vein of control and compound 48/80 treated SWISS mice on day 1 (before first dose), 3 and 5 and was analyzed for (E) WBC, (F) circulatory RBC and (G) hematocrit levels on automated cell counter. Each data point is plotted as mean ± SEM and representative plots/images are from at least 6 mice per group (ns, not significant; ∗∗p ≤ 0.005; magnification 40X, scale bar 8 μm).

2.5. Phenyl hydrazine administration in vivo for anemia induction in mice

Oxidative stress related anemia was induced in mice with phenyl hydrazine (Sigma, MO, USA) as described earlier with minor modifications [14]. Briefly, age and weight matched SWISS, C57BL/6, or DBA/2 mice were injected intraperitoneally for two consecutive days with 60 mg/kg body weight of phenyl hydrazine or PBS alone (200 μl) to control group, schematic timeline depicted in Fig. 3 (A). Mice were bled and sacrificed on day 4, analyzed for blood parameters and spleen was isolated to analyze spleen morphology and number of spleen cells with or without erythrocytes. Mice body and spleen weights were observed using analytical balance and spleen size was observed using geometric scale. For reticulocyte study, 1 million erythrocytes were stained with 50 ng/ml thiazole orange (Sigma, MO, USA) for half an hour at room temperature (RT). Erythrocytes were washed with PBS supplemented with 2 % FBS at 1400 rpm at 4oC for 5 min. Percent of total thiazole orange positive reticulocytes from control and treated group were analyzed with BD FACSCalibur flow cytometer.

Fig. 3.

Fig. 3

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Induction of anemia/oxidative stress with phenylhydrazine administration and its effect on various blood parameters. SWISS mice were administered 60 mg/kg phenylhydrazine (PHZ) intraperitoneally on day 1 and 2 simultaneously to treated group and PBS alone to control group in parallel. Mice were sacrificed on the day 4 (A) Schematic showing detailed protocol for anemia induction with PHZ. Erythrocytes from control and treated groups were stained with (B) 5 μM CM-H2DCFDA for 30 min at RT to analyze ROS. (C) Circulatory RBC level was observed on day 4 of PHZ induction and compared in control and PHZ treated groups. (D) Circulatory reticulocytes were stained with 50 ng/ml thiazole orange for 30 min at RT. (E) Circulatory WBC levels in control and PHZ treated groups were observed before and after the treatment and compared. Erythrocytes were analyzed on FL-1 channel with BD FACSCalibur. Circulatory RBC and WBC were observed using automated cell counter. Data is plotted as mean ± SEM and representative FACS histograms showing mean percent of total ROS and reticulocytes with SEM values from at least 6 mice per group (∗p ≤ 0.05; ∗∗∗p ≤ 0.001).

2.6. Characterization of phenyl hydrazine induced anemia in compound 48/80 treated mast cell depleted mice in vivo

SWISS mice were co-administered with compound 48/80 and phenyl hydrazine (PHZ) intraperitoneally to deplete peritoneal mast cells followed by anemia. Mice were injected with compound 48/80 intraperitoneally for 4 days as described above. 60 mg/kg PHZ was also co-administered intraperitoneally on day 3 and 4. Control group was simultaneously injected with equal volume of PBS in parallel as shown in Fig. 5 (A). Mice were bled to analyze blood parameters before the treatment and on day 3 and 5 of treatment and sacrificed on day 6. Spleen and peritoneal cells were isolated and mast cell population was analyzed using flow cytometer using anti c-kit antibody, spleen morphology was observed and total number of spleen cells were counted using hemocytometer.

Fig. 5.

Fig. 5

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Merge model of peritoneal mast cell depletion and PHZ induced anemia. SWISS mice were administered 1.2 mg/kg of compound 48/80 (suspended in PBS) twice a day for 3 days, 2.4 mg/kg of compound 48/80 (once) on day 4 and 60 mg/kg PHZ on day 3 and 4 simultaneously to mast cell depleted anemic group, only compound 48/80 to mast cell depleted group, only PHZ to anemic group and PBS alone to control group (i.p.) in parallel. (A) Schematic showing detailed protocol for mast cell depletion followed by anemia induction. (B) Mice were sacrificed on day 6 and peritoneal cells were isolated. Peritoneal cells from each group were stained with APC conjugated anti mouse CD117 (c-kit) antibody and analyzed on BD FACSCalibur. Representative FACS dot plots obtained from total peritoneal cells from all groups respective to their isotype controls are shown. (C) Bar graph showing percent of total CD117 positive mast cells in total peritoneal cells. Data is plotted as mean ± SEM from at least 6 mice per group (ns, not significant; ∗∗∗p ≤ 0.001).

2.7. Hematological analysis

20 μl of blood samples were collected from mouse tail vein at specific time points in tip containing 15 μl of 5 mM dipotassium EDTA (Thermo fisher scientific, India). All blood parameters such as RBC, WBC, Hematocrit (HCT) and hemoglobin (Hb) levels were estimated by using an electronic hematology particle counter (MS4e, Melet Schloesing Laboratories, Chaussée Jules César, Osny, France).

2.8. Toluidine blue staining of peritoneal mast cells

1 million peritoneal cells from control and compound 48/80 treated mice were fixed on poly L lysine (Sigma, MO, USA) coated glass cover slips (200 μl of poly L lysine/per cover slip for 1 h at room temperature and washed gently with PBS) for 1 h at RT. Cells were then rinsed gently with MiliQ water (MQ) (by gently dipping cover slips) and air dried. Cells were fixed with Mota's fixative (Sigma-Aldrich) for 15 min at RT followed by a gentle rinsing in MQ. Cells were stained with toluidine blue (0.5 % in 30 % alcohol, pH < 1) for 15 min at RT, rinsed twice to remove excess stain, and left for air drying. Coverslips containing stained cells were mounted on glass slide with fluoromount-G. Images from at least 5 different fields per mouse sample were observed under Nikon Eclipse-Ti fluorescence microscope at 40X magnification (analyzed on Olympus Fluoview FV1000 software).

2.9. Intracellular reactive oxygen species (ROS) measurement in erythrocytes

1 million of normal or t-BHP treated erythrocytes were washed and resuspended in pre-warmed PBS supplemented with 2 % FBS. Erythrocytes were then incubated with 5 μM CM-H2DCFDA (chloromethyl derivative of 2′, 7′-dichlorodihydrofluorescein diacetate) (Molecular Probes, Eugene, USA). for 30 min at 37 °C protected from light as described previously [8]. ROS leads to the conversion of CMH2DCFDA to its fluorescent product as oxidative reaction. The fluorescence was measured by flow cytometry with BD FACS Calibur. Data from at least 10,000 events per sample were collected and analyzed on CellQuest Pro software (BD Biosciences).

2.10. Mast cell – macrophage interaction

0.1 million P815 mast cells and MH-S macrophages each were co-cultured together in direct contact or separated by polycarbonate membrane transwell inserts (0.4-μm pore size, 6.5-mm diameter) (Corning-Costar) in DMEM media containing 10 % FBS per well in 24 well plate. After 18 h, CFSE (Carboxyfluorescein diacetate succinimidyl ester) labeled normal or ODE (pretreated in vitro with 3 mM tert-butyl hydroperoxide (t-BHP) as described previously [8]) were co-incubated with pre-seeded mast cells-macrophages co-culture at 1:25 or 1:250 mast cells-macrophages to erythrocyte ratio for 30 min at 37oC in CO2 incubator. Mast cells-macrophages (direct contact) were then harvested, followed by hypotonic lysis of unphagocytozed erythrocytes with ACK lysis buffer (150 mM NH4Cl, 10 mM KHCO3, and 100 μM Na2EDTA, pH 7.3), washed with PBS (200×g, 5 min at room temperature) and stained with either APC conjugated anti mouse CD117 or APC conjugated anti mouse F4/80 (Affymetrix eBioscience, San Diego, CA, USA) separately (to identify mast cells or macrophages respectively from the mixed cell population). Mast cells or macrophages containing CFSE labeled erythrocytes were analyzed by flow cytometry as described above and gating strategy is described in Fig. 8 (A). For transwell experiments, mast cells cultured with macrophages or vice-versa separated by transwell were co-incubated with CFSE labeled ODE at 1:250 and 1:25 ratio respectively for 30 min at 37oC in CO2 incubator. Mast cells and macrophages co-cultured separated by transwell were harvested and analyzed for uptake of erythrocytes using BD FACSCalibur.

Fig. 8.

Fig. 8

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Uptake of normal and ODE by mast cells and macrophages in direct or indirect contact in vitro. P815 mast cells and MH-S macrophages were seeded equally and co-cultured (direct contact) or separated by transwell system (indirect contact) overnight. CFSE labeled normal or t-BHP treated ODE were added at 1: 250 cells (mast cells-macrophages) to erythrocyte ratio for 30 min. Cells were harvested and stained for APC conjugated anti mouse CD117 antibody (mast cell marker). (A) Dot plot showing P815 (mast cell) population which was gated and erythrocyte uptake was analyzed. Representative FACS histograms showing gated CD117 positive mast cells from mast cells-macrophages mixed population which were analyzed for uptake of normal and ODE are shown. (B) Dot plot showing MH-S (macrophage) population which was gated and erythrocyte uptake was analyzed. Representative FACS histograms showing gated F4/80 positive macrophages from mast cells-macrophages mixed population which were analyzed for uptake of ODE at 1:25 and 1:250 cells to erythrocyte ratio are shown. (C) Bar graph showing comparison of uptake of CFSE labeled ODE by mast cells alone and mast cells co-cultured with macrophages in direct or indirect contact. (D) Bar graph showing comparison of uptake of CFSE labeled ODE by macrophages alone and macrophages co-cultured with mast cells in direct or indirect contact. (E) Reverse transcriptase PCR analysis for RAGE and TIM3 surface receptors on RBL mast cells that help recognize and internalize damaged erythrocytes. Data is plotted with mean ± SEM values from at least three independent experiments (∗p ≤ 0.05; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001; ns, not significant).

2.11. Primer designing

Primers were designed for GAPDH, TIM3 and RAGE genes for Rattus norvegicus using Primer-BLAST software (https://www.ncbi.nlm.nih.gov/tools/primer-blast/). The parameters set for primer designing were: amplicon size between 200 and 500bp, product length 20–24 bases long, GC content 45–55 % and no self-complimentarity. Bioinformatics tools such as “Oligo Calc” (http://biotools.nubic.northwestern.edu/OligoCalc.html) and “MultAlin” (https://multialin. toulouse.infra.fr/multialin/) was used to further validate the parameters of primers designed. The primers designed are RAGE (Rat), (Sense 5′-CTACCTATTCCTGCAGCTTC-3′, anti-sense 5′-CTGATGTTGACAGGAGGGCTTTCC-3′), TIM3 (Rat) (Sense 5′-CGGCCAAGCACTCA TGTTTT-3′, anti-sense 5′-TGGGATGACTTT GGCTGGTT-3′) and GAPDH (Rat), (Sense 5′-GCGAGATCCCGCTAACATCA-3′, anti-sense 5′-CTCGTGGTTCACACCCATCA-3′).

2.12. Isolation of RNA and first strand cDNA synthesis

RNA was isolated from 5 to 10 million cells of cell lines and primary mouse or rat splenocytes using TRIZOL (TRI) reagent (Thermo Scientific) as per manufacturer instructions by Trizol-chloroform method. Briefly, 200 μl chloroform per ml TRI was added, mixed by inverting and incubated at room temperature for 15 min. After incubation, it was centrifuged at 12000×g for 15 min at 4oC. After centrifugation 3 layers was obtained and upper aqueous layer was taken out in separate eppendorf tube. 0.5 ml of isopropanol was added to precipitate RNA and RNA pellet was obtained by spinning at 7500×g and 4oC for 5 mins. The RNA pellet was washed twice with 1 ml of 75 % ethanol, supernatant was discarded followed by air drying at room temperature and finally resuspended in 40 μl (5 x 106 cells of cell lines and 10 million primary cells from mouse splenocytes) of Tris/borate/EDTA (TBE) buffer at 70oC for 10 min for proper dissolution. Concentration and purity of RNA was estimated by studying absorbance at 260/280 nm and 260/230 nm wavelength with Nanodrop (ND2000, Thermo Scientific) spectrophotometer. The integrity of RNA was analyzed by running 5 μg of RNA on 1.2 % formaldehyde agarose gel, and after confirmation of purity and integrity, the isolated RNA was used for cDNA synthesis. The first strand cDNA for the isolated RNA was synthesized by reverse transcription using M-MuLV Reverse Transcriptase enzyme (200 U/μl, NEB, UK) and oligo(dT)18 (50 μM) primer as per manufacturer instructions in a total reaction mixture of 20 μl. Amplification of cDNA was done using thermocycler (BioRad) by heating to 37oC for 60 min followed by 70oC for 10 min in order to terminate the reaction and final hold at 4oC. cDNA was confirmed by running cDNA sample in 1.8 % agarose gel. After confirmation, cDNA products were further used for amplification with PCR. The amount of RNA template per reaction mixture was standardized for the linear range for performing reverse transcriptase PCR.

2.13. Reverse transcriptase PCR

Amplification of cDNA obtained above was performed by conventional PCR using gene specific primers designed (at standardized annealing temperature for individual molecule). Taq Pol enzyme (Thermo Scientific) was used for polymerization for reverse transcriptase PCR as recommended by the manufacturer. In brief, total reaction mixture of 25 μl was prepared using 4 μl of cDNA sample, Taq Pol enzyme (5 U/μl), forward and reverse primers (61 μM each), Taq Pol buffer (1X), dNTPs mix (0.25 mM) and nuclease free water. The samples were put in thermocycler for PCR amplification with initial heating at 95oC for 5 min and cycle conditions were, denaturation for 1 min at 95oC, annealing at 60/65oC for 1 min and extension for 72oC for 1 min followed by additional extension for 5 min at 72 °C for last cycle with final hold at 4 °C. The amplified PCR products were run on 1.8 % agarose gel containing ethidium bromide (EtBr) (0.5 μg/ml) in 0.5 X TBE buffer. The agarose gel electrophoresis was carried out at 90 V for 2 h and the gel was visualized under BIO-RAD gel documentation system. The band intensity of PCR products were measured by Quantity One software version 4.6.5 (basic) from BIO-RAD. Primers of GAPDH for rat with respective template origin was amplified in parallel with each set of experiment as an endogenous control.

2.14. Statistics

Data was obtained from at least three independent experiments or at least 5 mice per group. One tailed student's t-test was used for statistical analysis using MS Excel (Microsoft office 2010). p value < 0.05 was considered for statistically significant difference between control and treated groups. Data is represented as mean ± SEM.

3. Results

3.1. Impact of mastocytosis induction on erythrocytes in DBA/2 mice in vivo

Systemic mastocytosis has been shown to be associated with anemia in at least 50 % of cases. In our experimental studies, the effect of systemic mastocytosis (SM) on erythrocytes was checked in vivo. A model designed previously for systemic mastocytosis was adapted and followed, where injection of P815 cells into DBA/2 mice induced SM [12,15]. Induction of mastocytosis in DBA/2 mice was attempted with P815 administration, retro-orbitally. The detailed timeline for protocol of mastocytosis induction is depicted in Fig. 1 A. A significant increase in circulating WBCs (2 fold increase) and slight increase in spleen WBC levels in P815 treated group compared to control group after 8 days (Fig. 1B and C) was observed. Any significant difference in body weight in control and the treated groups (data not shown) was not observed. Mast cell percentage out of total population of peripheral blood WBCs, spleen WBCs and total peritoneal cells was analyzed as CD45 and CD117 double positive population in control and P815 treated groups by flow cytometer (Fig. 1 D). A 3.6 fold increase in mast cell population in peripheral blood, almost a 1.5 fold increase in spleen WBCs and also in total peritoneal cells was observed. Hence, a small increase was observed in mast cell population in all the organs. Further, 35 % decrease in blood platelets and small but significant decrease (7 %) in circulating erythrocyte levels relative to pretreated levels was observed in P815 treated group (Fig. 1E and F). The increase in mast cell number and decrease in blood platelets in the present study correlates with the previously established mastocytosis model [12,15]. Although we were unable to obtain high increase in mast cell numbers in the mastocytosis model, but it is clear from the above model that, even a slight mastocytosis caused a significant decrease in circulating erythrocytes and platelets. This observation in the current study indicated a direct or indirect involvement of mast cells in clearance or reduction in level of circulating erythrocytes during mastocytosis conditions.

3.2. Mast cell depletion with compound 48/80 treatment

Compound 48/80 specifically activates and degranulates mast cells. It has been reported earlier that prolonged intraperitoneal administration of compound 48/80 leads to mast cell depletion [13,16]. To observe the direct impact of mast cells on erythrocytes, compound 48/80 administration to deplete mast cells in three mice strains was tried and mast cell depletion in vivo with compound 48/80 administration was standardized in SWISS mice. The schematic is depicted with timeline in Fig. 2 A. No significant difference was observed in the body weight of control and compound 48/80 treated mice after 5 days of compound 48/80 administration (Supplementary Fig. 1). Total peritoneal cells were recovered and mast cell population from control and treated groups was analyzed with flow cytometer using anti mouse CD117 (c-kit) antibody. Total peritoneal cell recovery from control and compound 48/80 treated mice were observed to be similar but 1–3 % peritoneal mast cell population (control group) decreased to less than 0.5 % (compound 48/80 treated group) and showed more than 75 % depletion of mast cell population (Fig. 2B and C). Mast cells were observed as pink/purple stained cells and other non granulated cells of peritoneal lavage were observed as blue after toluidine blue staining. No pink/purple stained mast cells were observed in compound 48/80 treated groups (Fig. 2 D). Mast cell depletion was observed in SWISS mice but not in C57BL/6 and DBA/2 mice strains, upon compound 48/80 administration, by flow cytometry (Supplementary figure 2 A). Mast cell degranulation in all mice strains was observed with toluidine blue staining (Supplementary Fig. 2B and C). Erythrocyte and hematocrit levels remained constant after compound 48/80 treatment in all mouse strains (Supplementary Fig. 3A and B). Significant increase in WBC levels was observed in C57BL/6 and DBA/2 mice but not in SWISS mice post compound 48/80 treatment (Supplementary figure 3 C). Higher level of WBCs in C57BL/6 and DBA/2 may be due to continuous mast cell degranulation but not depletion due to compound 48/80 administration, while SWISS mice were found to be depleted with peritoneal mast cell population after 5 days of treatment. In SWISS mice, we observed slight increase in WBC levels on day 3, which attained basal levels on day 5 in compound 40/80 treated group as compared to control group (Fig. 2 E). Compound 48/80 induces mast cell degranulation, which resulted in initial increase in WBC levels but prolonged administration of compound 48/80 results in mast cell depletion and WBCs attained their normal level in circulation. The effect of compound 48/80 on various blood parameters was analyzed on day 1 (before first dose of compound 48/80), day 3 (during mid-treatment) and day 5 (after all doses of compound 48/80) in control and treated groups. No significant difference in circulatory RBC and hematocrit levels in both the groups was observed throughout the treatment (Fig. 2F and G). Hence, compound 48/80 treatment resulted in mast cell depletion only in SWISS mice within 5 days. Circulating erythrocyte levels was not affected by mast cell depletion in mice in vivo.

3.3. Induction of oxidative stress/anemia with phenylhydrazine in vivo

Our previous in vitro studies had revealed a novel role for mast cells as scavenger cells involved in clearance of oxidatively damaged/stressed erythrocytes. Further, to study the involvement of mast cells in anemia in vivo, an appropriate anemic model is a prerequisite. Phenylhydrazine (PHZ) is an antipyretic drug that is well known for its ability to induce oxidative stress followed by anemia. Phenylhydrazine leads to degradation of hemoglobin and the lipid peroxidation of the erythrocyte membrane [14,17]. In the present study, oxidative stress/anemia was induced with PHZ as described in schematic timeline (Fig. 3 A). Optimum dose of PHZ causing significant anemia was standardized in SWISS mice (Supplementary Fig. 4). To confirm oxidative stress, reactive oxygen species (ROS) in erythrocytes were analyzed using CM-H2CFDA (chloromethyl derivative of H2DCFDA) dye, which binds to the intracellular glutathione and other thiols with its thiol-reactive chloromethyl group. Almost all erythrocytes from PHZ treated mice were found CM-H2CFDA positive and indicated significantly high ROS concentration in erythrocytes of PHZ treated groups (Fig. 3 B). Circulating erythrocyte levels decreased by almost 65 % in PHZ treated mice as compared to control group (Fig. 3 C). High ROS concentration along with low erythrocytes levels in PHZ treated group confirmed severe anemia under high oxidative stress in these mice. Further, circulating reticulocytes (nucleated immature erythrocytes) in whole blood were analyzed by flow cytometer using thiazole orange (1- methyl-4[(3-methyl-2(3H)-benzothiazolyli -dine) methyl]-quinolinium 4-methyl benzene sulfonate) staining, which binds both RNA and DNA. Significantly high levels of reticulocytes (20–21 %) were found in PHZ treated mice in comparison to control mice (3–4 %) (Fig. 3 D). High levels of circulating reticulocytes indicated haemopoiesis induction to compensate for low levels of erythrocytes in severe anemic conditions. High levels of circulating WBC (almost 10 times higher) were also observed after PHZ treatment (Fig. 3 E). High oxidative stress might lead to increased WBC count.

No significant decrease in body weight was observed after PHZ administration (data not shown). On day 4 after PHZ treatment, mice from control (PBS injected) and PHZ treated groups were sacrificed and spleens were isolated. Spleen morphology, in terms of spleen size and weight was analyzed and compared (Fig. 4 A, B, C). Total spleen cells were counted for both the groups before (spleen WBC + RBC) and after erythrocyte lysis (spleen WBCs). Two fold higher accumulations of erythrocytes in spleens as well as significant increase in spleen WBC (2 fold increase) was observed in PHZ treated group as compared to control group (Fig. 4 D). Various observations such as increase in spleen size, weight, WBC count in spleen cells and significant decrease in RBC count altogether indicated a significant induction of anemia with 60 mg/kg dose of PHZ and this dose was used further in the present study. To observe the strain specific differences, PHZ was administered to C57BL/6 and DBA/2 mice following similar protocol. Spleen weight was observed to be increased by 3–4 fold and 2 to 5 times higher accumulation of erythrocytes in spleens after PHZ treatment was observed within all strains with highest erythrocytes accumulation in DBA/2 mice (Supplementary Fig. 5). Spleen WBCs in all mice strains were also found in 2–3 fold high numbers as compared to their control cohorts. Circulating erythrocytes and hematocrit levels showed a decrease of 60 % in SWISS and C57BL/6, and of 40 % in DBA/2 mice (Supplementary figure 6 A). WBC levels increased up to 4–6 times in all PHZ treated groups (Supplementary figure 6 B). Reticulocyte count increased significantly (5–14 fold) in all the mouse strains to compensate for PHZ induced severe anemia under oxidative stress (Supplementary figure 6 C). SWISS mice showed higher decline in circulating erythrocytes and hematocrit levels along with a lower increase in reticulocytes as compared to other two strains during PHZ induced oxidative stress conditions. We have already observed significant mast cell depletion in only SWISS mice on compound 48/80 administration. Therefore SWISS mice were used as a model to study the mast cells role in clearance of erythrocyte during oxidative stress conditions in vivo.

Fig. 4.

Fig. 4

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Effect of PHZ administration/oxidative stress on mouse spleen. SWISS mice were administered 60 mg/kg phenylhydrazine or saline, after 4 days mice were sacrificed and spleens were isolated from control and PHZ treated groups. (A) Representative spleens, (B) spleen size, (C) spleen weight and (D) Spleen cell counts are shown before (spleen RBC + WBC) and after erythrocyte lysis (spleen WBC), observed using hemocytometer under light microscope. Data is plotted as mean ± SEM from at least 6 mice per group (∗p ≤ 0.05; ∗∗p ≤ 0.005; ∗∗∗∗p ≤ 0.0001).

3.4. Role of mast cells in erythrocyte clearance during anemia due to oxidative stress in vivo

Taking a clue from our in vitro study highlighting the ability of mast cells to phagocytose and clear oxidatively stressed erythrocytes, clearance of erythrocytes under PHZ induced oxidative stress in the absence of mast cells in vivo was studied. Peritoneal mast cells were first depleted with compound 48/80 followed by PHZ treatment to induce anemia in SWISS mice. The detailed protocol with timeline is shown in (Fig. 5 A). Mast cell population was analyzed with flow cytometer to compare mast cell depletion among mast cell depleted control (treated with compound 48/80 only) and mast cell depleted anemic (treated with compound 48/80 + PHZ) and control (PBS only) groups using anti mouse CD117 antibody (Fig. 5 B). Similar level of mast cell depletion (70 %) was observed in both mast cell depleted control and mast cell depleted anemic groups as compared to control group (Fig. 5 C). Blood parameters were further analyzed to observe anemia induction in presence or absence of peritoneal mast cells. Circulating erythrocytes and hematocrit levels showed significant and similar decrease (60 %) due to PHZ induced anemia in mice groups in presence or absence of peritoneal mast cells as compared to their pretreatment levels and control group (Fig. 6A and B). In addition, increase in circulating WBC levels (5–6 fold) was also similar in anemic control and mast cell depleted anemic groups (Fig. 6 C).

Fig. 6.

Fig. 6

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Peripheral blood parameters of merge model of peritoneal mast cell depletion and PHZ induced anemia. SWISS mice were administered PBS alone (control group), compound 48/80 alone, PHZ alone and compound 48/80 along with PHZ to get mice with mast cell depleted control, anemic control and mast cell depleted anemic models respectively. Test bleed was taken from each group from tail vein on day 1(before first dose), 5 (after all doses) and whole blood was analyzed on automated cell counter. (A) Circulatory erythrocyte levels (B) hematocrit levels and (C) WBC levels in mice from each group were observed before and after the treatment and compared. Data is plotted as mean ± SEM from at least 6 mice per group [ns, not significant; ∗p ≤ 0.05; ∗∗∗∗p ≤ 0.0001 (compared within groups); #p ≤ 0.05; ####p ≤ 0.0001, compared to pretreatment levels].

Impact of oxidative stress or anemia was analyzed on spleen in presence or absence of mast cells. Spleen morphology and cells were compared in anemic control (treated with PHZ only) and mast cell depleted anemic groups. Mice from each group were sacrificed on day 6 and spleens were isolated, analyzed for morphology (Fig. 7 A) and cell number count. Significant and similar increase in spleen size (1.5 fold) and weight (3 fold) was observed in both groups (anemic control and mast cell depleted anemic group) as compared to their control cohorts (Fig. 7B and C). Total spleen cells with and without spleen erythrocytes (depleted by hypotonic lysis) were counted and compared among these groups. Significantly higher accumulation of erythrocytes was observed in spleens of mice from both anemic control and mast cell depleted anemic groups as compared to control group. But mast cell depleted anemic group showed significant and much higher (almost twice) erythrocyte accumulation in their spleens in comparison to anemic control group, while the level of circulatory WBCs was found to be similar (Fig. 7 D). Collectively, it can be interpreted from the above observations that PHZ induced oxidative stress led to significant anemia in vivo in presence and absence of peritoneal mast cells but erythrocyte clearance was found to be reduced in spleen of mast cell depleted mice during anemia/oxidative stress condition. Hence, there was a higher accumulation of erythrocytes waiting to be scavenged or cleared in mouse spleen in vivo, in mast cell depleted mice.

Fig. 7.

Fig. 7

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Spleen characteristics in the merge model of peritoneal mast cell depletion and PHZ induced anemia.SWISS mice were administered with PBS alone (control group), PHZ alone or compound 48/80 along with PHZ to get mice with anemic control and mast cell depleted anemic models respectively. Mice were sacrificed on 6th day and spleen was isolated. (A) Representative images of spleens, (B) spleen size, (C) spleen weight and (D) Spleen cell counts are shown before (spleen RBC + WBC) and after erythrocyte lysis (spleen WBC), observed using hemocytometer under light microscope from control, anemic control (PHZ alone) and mast cell depleted anemic (compound 48/80 + PHZ) groups. Data is plotted as mean ± SEM from at least 6 mice per group (ns, not significant; ∗∗p ≤ 0.005; ∗∗∗p ≤ 0.001; ∗∗∗∗p ≤ 0.0001; ###p ≤ 0.001) [∗(comparison within WBC + RBC), # (comparison within WBC relative to control mice)].

3.5. Effect on erythrocytes uptake by mast cells and macrophages in direct or indirect contact in vitro

Direct role of mast cells in uptake of ODE as scavengers has been investigated in our previous study [8]. The present study has shown the impact of mast cells in erythrocyte clearance in vivo. In the in vivo conditions macrophages have been described as the major scavenger cells involved in uptake and clearance of damaged erythrocytes [18]. The consequence of interaction of mast cells and macrophages in erythrophagocytosis is still not known. The presence of professional phagocytes like macrophages must influence the erythrocyte clearance by mast cells to a great extent in vivo. We further tried to explore the impact of mast cell-macrophage interaction on uptake of erythrocytes by mast cells or macrophages in vitro in direct and indirect (separated by transwell) contact. P815 mast cells and MH-S macrophages were co-cultured in DMEM medium supplemented with 10 % FBS and uptake was analyzed by both cell types individually. After overnight culture CFSE labeled normal or ODE were added at 1:250 (mast cells-macrophages to erythrocytes ratio). P815 mast cells alone were also co-incubated with erythrocytes under similar conditions in parallel as control. Cells were harvested after 30 min and stained with CD117 antibody to identify P815 mast cells in the mixed cell population by flow cytometry. CD117 positive mast cells were gated and uptake of erythrocytes by mast cells from mixed population was analyzed only for gated population (Fig. 8 A). 31 ± 3.8 % of mast cells showed uptake of ODE, while only 1.2 ± 0.5 % of mast cells took up normal erythrocytes in presence of macrophages. Effect of mast cell-macrophage interaction was also observed on uptake of erythrocytes by MH-S macrophages. MH-S macrophages showed saturated uptake of ODE at macrophages to erythrocytes ratio higher than 1:100 and did not show significant uptake of normal erythrocytes in presence or absence of mast cells (data not shown). MH-S macrophages and P815 mast cells were co-cultured overnight and CFSE labeled ODE (at 1:25 and 1:250 ratio) were co-incubated for 30 min. Cells were harvested and stained with F4/80 antibody to identify MH-S macrophages in the mixed cell population by flow cytometry. F4/80 positive macrophages were gated and uptake of erythrocytes by macrophages was observed in the mixed population. 30.3 ± 4.2 % and 71.9 ± 3.9 % of total macrophages showed uptake of ODE at 1:25 and 1:250 ratio respectively in presence of mast cells (Fig. 8 B).

To confirm if erythrocyte's uptake requires direct physical contact between the two cell types or it is because of indirect interaction due to cell secreted mediators. P815 mast cells were co-cultured with MH-S macrophages separated by transwell inserts of 0.4-μm pore size, which allows free passage of media and mediators secreted by cells between both chambers preventing direct physical contact. After overnight culture, ODE were co-incubated with mast cells at 1:250 mast cells to erythrocytes ratio for 30 min. Uptake of ODE by mast cells alone, mast cells in presence of macrophages in direct physical contact and mast cells in presence of macrophages separated by transwell allowing indirect contact was observed and compared (Fig. 8 C). Uptake of ODE by mast cells increased significantly to 72 % (from 18 ± 1.8 % to 31 ± 3.8 % of total mast cells), when cultured in direct contact with macrophages, but remained unaffected due to indirect mast cell-macrophage interaction separated by transwell. Similarly, the effect of indirect interaction of both cell types on uptake of erythrocytes by macrophages was studied. Here, MH-S macrophages were co-cultured with P815 mast cells separated by transwell inserts. After overnight culture, ODE were co-incubated with macrophages at 1:25 macrophages to erythrocytes ratio for 30 min. Uptake of ODE by macrophages alone, macrophages in presence of mast cell in direct physical contact and macrophages in presence of mast cells separated by transwell allowing indirect contact was observed and compared (Fig. 8 D). Uptake of ODE by macrophages decreased significantly to 34 % (from 46 ± 4.3 % to 30.3 ± 4.2 % of total macrophages) when cultured in direct contact with mast cells but remained unaffected due to indirect mast cell-macrophage interaction separated by transwell. RT-PCR analysis confirmed the co-expression of the Receptor for Advanced Glycation End-products (RAGE) and T-cell Immunoglobulin and Mucin-domain containing-3 (TIM3) transcripts in RBL mast cells, evidenced by distinct amplification products on agarose gel electrophoresis. This finding establishes TIM3 expression in RBL cells, thereby expanding their documented receptor repertoire, as RAGE expression was previously known (Fig. 8 E). Given that macrophages utilize these same receptors (RAGE and TIM3) for the recognition and clearance of oxidatively damaged erythrocytes, this receptorial overlap suggests a potential functional convergence where RBL mast cells participate in erythrocyte interaction and uptake. Collectively, erythrophagocytic capacity of mast cells was further augmented, while it was decreased in case of macrophages when both are in direct physical contact.

4. Discussion

Anemia of inflammation or anemia of chronic disease (ACD) is the most commonly found anemia which is directly linked to inflammation [19,20]. Mast cells are well known for their crucial role in inflammatory responses, innate or adaptive immunity [21], and pathogen clearance by phagocytosis as non-professional phagocytes [4], by mediator release or by generation of extracellular traps [7]. Patients with mast cell disease, show disruptions in erythropoiesis which can lead to impaired blood cell development and function [22]. However, the direct contribution of mast cells in clearance of erythrocytes and the onset of anemia is not very well understood, but there are some previous reports, which have shown possible contribution of mast cells in the onset of anemia [1,2]. In the present study we observed significant increase in circulating WBCs, decrease in platelets and a small increase in mast cell population in peripheral blood WBCs, spleen WBCs and peritoneal lavage in our mastocytosis mouse model. Abnormal blood count such as an elevated or decreased number of WBCs or platelets, abnormal differential count, or unexplained anemia were already reported in clinical study with systemic mastocytosis patients after a bone marrow biopsy [23]. In current study, circulating erythrocytes showed significant decline relative to their pretreated levels in mastocytosis groups. We were unable to obtain higher increase in mast cell numbers in the mastocytosis model but even a little increase in mast cell numbers led to a significant decrease in circulating erythrocytes. Hence, our results are in agreement with previous reports where evidences of hematologic disorders like elevated leukocytes and unexplained anemia were correlated with systemic mastocytosis [23,24]. Correlation of iron deficiency and allergy or allergic mediators has been reported earlier [25]. Mast cell releases several pro-inflammatory and vasoactive mediators including IL-6, which is a major pro-inflammatory cytokine that stimulates hepcidin and plays important role in the iron metabolism [20,26]. Hence, mast cells may also contribute to anemia indirectly by iron sequestration through increased levels of hepcidin. The inter-connection of inflammation anemia and oxidative stress has already been reported in several previous studies [27]. Prevalence of Inflammation and oxidative stress interplay is most common in many old age disorders [28], cancer, and vascular diseases [29,30]. Oxidative stress increases exponentially during aging and variety of pathological conditions like cardiovascular diseases, diabetes and cancer [28]. In our previous study we have reported the role of mast cells as scavengers, where mast cells showed significant uptake of ODE but not of normal erythrocytes as non professional phagocytes [8].

Further, to explore the contribution of mast cells in anemia or erythrocyte clearance in vivo, we attempted to deplete mast cells with prolonged compound 48/80 treatment as reported earlier in SWISS mice and Wistar rats [13,16]. Mast cells are tissue resident cells dispersed in heart, liver, bone marrow and lungs and in low numbers in spleen and adrenals [31], but in detectable numbers in skin and peritoneum [32]. We observed the distinct effect of compound 48/80 administration in different mice strains. Significant peritoneal mast cell degranulation was observed in all strains using toluidine blue staining method, while mast cell depletion was confirmed only in SWISS mice with fluoromertic c-kit staining. Mast cell degranulation does not lead to their depletion as they have ability to regranulate by starting recovery process just after 6 h of degranulation [33]. We observed no significant difference in total peritoneal cell recoveries, circulating erythrocytes, hematocrit and hemoglobin levels in control and compound 48/80 treated groups. Mast cell degranulation followed by depletion did not affect erythrocytes levels which indicated that mast cell mediators do not affect erythrocytes and are unable to induce anemia. This was already shown in our previous study by observing erythrocyte survival in presence of activated mast cell supernatant in vitro where we did not find any decrease in normal or oxidatively damaged erythrocyte numbers due to mast cell mediators [8]. Further, we were able to induce anemia successfully with intraperitoneal PHZ administration in vivo as explained earlier [14,34]. It is well reported that narrow splenic meshwork serves as major filter for blood and erythrocytes are regularly monitored for their deformability to prevent clogging in microvessels, hence deformed aged or damaged erythrocytes are retained in spleen and cleared from circulation by professional phagocytes such as macrophages [35,36]. Elevated level of WBC count was also reported in a previous study in wistar rat due to blood loss as well as hemolysis by PHZ treatment [37]. We observed significant oxidative stress and anemia in all three mice strains, with high ROS levels in erythrocytes, splenomegaly, accumulation of erythrocytes and high levels of WBCs in spleen and peripheral blood respectively, higher reticulocytes and low levels of circulating erythrocytes and hematocrit levels upon PHZ administration which supports previous reported literature. SWISS mice showed most severe symptoms of anemia in comparison to other two strains. SWISS is an outbred mice strain and may be more comparable to humans due to genetic diversity in comparison to inbred mouse strains [38].

Induction of anemia with PHZ in compound 48/80 administered mast cell depleted SWISS model did not cause any significant difference in circulating erythrocyte but showed higher accumulation of erythrocytes in spleen with similar WBC levels in comparison to anemic control mice (only PHZ treated) in vivo. PHZ is known to induce hemolytic anemia by causing oxidative damage and cross-linking of Band 3 on erythrocytes, thereby marking them for macrophage-mediated clearance via Fc receptors. Previous studies have reported that PHZ treatment increases the total white blood cell (WBC) population [39]. Consistent with these findings, our study also observed an increase in total splenic WBC counts following PHZ administration. Since mast cells constitute only 1–2 % of the total splenic leukocyte population, the absence of a significant change in total WBC counts upon co-administration of PHZ and Compound 48/80 suggests that PHZ does not substantially alter splenic mast cell or mast cell progenitor numbers. These observations (Fig. 7D) indicate that PHZ's primary effect in our model is the induction of erythrocyte damage, rather than acting as an enhancer of the mast cell population within the spleen. Increased levels of erythrocytes in spleen result due to many reasons. First, increase in number of damaged erythrocytes; second, decreased clearance of damaged erythrocytes in spleen or third, combination of the above two reasons. Clearance of damaged or senescent erythrocytes by reticulo-endothelial system (RES) mainly in spleen, and liver through phagocytosis is well known [40,41]. In the present study, higher accumulation of erythrocytes in spleen accompanied by similar level of circulating erythrocyte and spleen WBC in comparison to anemic control mice indicates reduced clearance of erythrocytes in spleens of mast cell depleted anemic mice. Erythrocyte clearance is the crucial factor during inflammatory process due to several abnormal changes on erythrocyte surface similar to erythrocyte senescence and leads to anemia of inflammation [18]. Reduced erythrocyte clearance in spleens of mast cell depleted mice indicates the direct or indirect role of mast cells in this process. In a previous clinical study, splenic mast cell tumor and systemic mastocytosis was found to be accompanied by anemia in a domestic cat where 11 % of total mast cells showed phagocytosed erythrocytes [42]. We have already reported the significant uptake of autologous erythrocytes (oxidatively damaged in vitro) by peritoneal mast cells in vivo [8]. Our previous investigation [8] analyzed mast cell phagocytosis of oxidatively damaged, t-BHP treated, CFSE-labeled erythrocytes using both in vivo and in vitro methods, for the uptake of a damaged erythrocytes by mast cells. For the in vivo study, DBA/2 mice were injected intraperitoneally with autologous, damaged erythrocytes, 60 min later, peritoneal cells were recovered, stained with APC-conjugated anti-CD117, and analyzed by flow cytometry, quantifying phagocytosis as the percentage of CD117 positive cells that were also CFSE positive, with external RBCs lysed before analysis to ensure only internalized cells were counted [8]. The in vitro model utilized RBL-2H3 mast cells co-incubated with CFSE-labeled damaged or normal erythrocytes, with phagocytic uptake quantified by fluorescence microscopy, and confirmation of true internalization achieved through confocal microscopy z-sectioning and visualization via TEM [8]. This multi-modal approach along with evidences from older literature [43] robustly supports our conclusion that peritoneal mast cells actively phagocytose damaged erythrocytes in an extra-splenic environment, in vivo, suggesting a new functional role for these cells in localized clearance and tissue homeostasis. The present study collectively suggests the crucial contribution of mast cells in erythrocyte clearance under chronic inflammatory/oxidative stress or disease conditions in direct or indirect manner.

Mast cells share many structural and functional similarities with macrophages, as both differentiate from the common myeloid progenitors and known for their contribution in innate as well as adaptive immune response for defense against a wide range of pathogens. Both are able to perform antigen presentation and phagocytosis [44]. This is a well known fact that mast cells and macrophages are both localized throughout connective tissues in mature form at different body sites where they frequently encounter a variety of pathogens and environmental antigens and interact with each other to invoke immune response for host defense [45]. The present study reveals that direct contact of mast cells and macrophages augments mast cell uptake while inhibits uptake of erythrocytes by macrophages. It has been reported earlier that direct co-culture of both cell types led to increased MHC II expression of mast cells [46]. On the other hand, it has also been reported that mast cells impair macrophage phagocytosis by releasing prestored IL-4 in septic peritonitis and thereby aggravating severe bacterial infection [47]. Further, we did not find any difference in uptake of ODE by both cell types when co-cultured separated in a transwell system in comparison to uptake by both cell types cultured alone. Mast cells and macrophages produce many common inflammatory cytokines such as TNFα, IL-13, IL-6 and MIP-1α along with a variety of other cytokines and chemokines that could activate each other [45]. Mast cells are able to stimulate macrophages migration and recruitment at the site of inflammation [48]. On the other hand, macrophages can also induce mast cell activation and degranulation similar to Compound 48/80 or substance P [49]. Mediator release may happen on direct physical contact. Interestingly, mast cells express several receptors that overlap functionally and structurally with those found in macrophages. The detection of RAGE and TIM3 transcripts in RBL mast cells, suggests that mast cells possess the ability to recognize and internalize damaged erythrocytes. This functional convergence points to a broader immunoregulatory role for mast cells beyond their classical involvement in allergic inflammation and mediator release. It was already known that TIM3 is closely associated with FcεRI and augments IgE/antigen-dependent signaling, undergoing internalization after antigen stimulation [50]. This indicates a dual role for TIM3 in mast cells, both as an immune checkpoint receptor and as a modulator of FcεRI-driven activation. Similarly, RAGE and its ligands such as Advanced Glycation Endproducts (AGEs) and soluble RAGE are critically involved in chronic inflammation, as AGEs accumulate in numerous inflammatory and metabolic disorders [51]. Importantly, RAGE expression and its ability to induce mast cell exocytosis independent of FcεRI were previously reported in rat peritoneal mast cells [52], supporting its role in mast cell activation outside the canonical IgE pathway.

Cytoskeletal rearrangement and associated proteins also play key roles in mast cell signaling and the early stages of FcεRI stimulation [53]. Hence, during mast cell activation, receptors such as RAGE and TIM3—and their associated signaling components—may already be upregulated and spatially organized to promote rapid and efficient interactions with oxidatively damaged erythrocytes.

In literature, there are reports that direct contact interaction of mast cells and macrophages has positive as well as negative impact on immune defense. Live cell imaging of mast cell-macrophage co-culture has revealed active cellular projections by mast cells to transfer FcεRI-encompassed cellular regions directly to macrophages during F. tularensis infection. This direct interaction of mast cells with macrophages played a significant role in defense against infection by activating macrophages to upregulate caspase-1 expression [46]. On the other hand there are also reports that mast cell granules phogocytosed by macrophages decrease superoxide production by scavenging the newly generated superoxides in phagolysosomes of macrophages [54]. Effect on phagocytic activity of mast cells due to macrophages is unknown till now but induction of mast cell activation by macrophage factors, bacterial infected macrophages in direct contact and bacterial infected macrophage conditioned media has been reported earlier [48,55]. The enhanced uptake of ODE by mast cells may be due to mast cell activation induced by oxidatively damaged erythrocyte containing macropahges and these activated mast cells may in turn affect uptake by macrophages. We have already reported that mast cells show significantly enhanced uptake of ODE upon activation [8].

The present study collectively highlighted two major aspects of mast cell role in anemia. The mild mastocytosis model showed significant decrease in erythrocytes in circulation, which in a way mimics the conditions seen during aging, where mast cell numbers increase [27]. This may have implications in anemia in the aging population. On the other hand, significantly high accumulation of erythrocytes in spleen during mast cell depleted condition indicated decreased erythrocyte clearance in absence of mast cells. These in vivo studies led to the conclusion that mast cells affect erythrocyte depletion or clearance directly or indirectly. Further, we revealed that mast cell-macrophage direct physical interaction leads to augmented uptake of ODE by mast cells while decreased uptake by macrophages only in direct physical contact. The present study concludes that mast cells have an important role in erythrocyte clearance. The scavenger activity of mast cells shows a significant upgradation, when in direct interaction with macrophages. This study provides some novel insights that have major implications in anemia in various autoimmune diseases, inflammatory or oxidative stress related disorders, and anemia in the aged populations, where mast cells are in high numbers.

Funding

This work was supported by research grants from Indian Council of Medical Research (ICMR) Govt. of India (61/3/2012-BMS) and Department of Biotechnology, India (BT/PR51192/MED/29/1662/2023). We also acknowledge the facilities supported by DBT BUILDER [grant no. BT/INF/22/SP45382/2022] and DST FIST-II [grant no. SR/FST/LSII-046/2016C] and ANRF-PAIR [grant no. ANRF/PAIR/2025/000029/PAIR-A] to SLS, JNU. PS was supported by a grant from UGC, India.

CRediT authorship contribution statement

Priyanka Sharma: Data curation, Formal analysis, Investigation, Methodology, Validation, Visualization, Writing – original draft. Mohd Suleman: Formal analysis, Software, Validation, Visualization, Writing – review & editing. Niti Puri: Conceptualization, Formal analysis, Funding acquisition, Methodology, Project administration, Resources, Software, Supervision, Validation, Visualization, Writing – review & editing.

Declaration of competing interest

The authors declare no conflict of interest.

Acknowledgements

We are grateful to Prof. Shweta Saran (School of Life Sciences, JNU, New Delhi, India), and Prof. A. Selvapandiyan (Department of Molecular Medicine, Jamia Hamdard, New Delhi, India) for critical reading of the manuscript. We thank Central Instrumentation Facility (CIF, School of Life Sciences, JNU, New Delhi, India) for Flowcytometry, and CLAR (Central Laboratory Animal Resource), JNU for maintenance of animals used in the present study.

Footnotes

Appendix A

Supplementary data to this article can be found online at https://doi.org/10.1016/j.bbrep.2025.102389.

Appendix A. Supplementary data

The following is the Supplementary data to this article:

Multimedia component 1
mmc1.pdf (337.1KB, pdf)

Data availability

No data was used for the research described in the article.

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