Background

A plateau is a special environment characterized by low atmospheric pressure and low partial oxygen pressure. Long-term exposure to plateau environments may lead to chronic mountain disease [1]. High-altitude polycythemia (HAPC) is a common and widespread chronic mountain sickness characterized by excessive erythrocytosis [2]. Hypobaric hypoxia (HH) is the main cause of erythrocytosis, which in turn alleviates hypoxic conditions in tissues under high-altitude (HA) exposure [3]. In a healthy organism, a balance is maintained between erythropoiesis in the bone marrow and eryptosis in the spleen [4]. Even though HA/HH exposure can disrupt the equilibrium between RBC formation and clearance, healthy individuals can swiftly establish a new physiological RBC steady state [5]. However, in patients with HAPC, RBCs homeostasis is disrupted, failing to reach a state of equilibrium, which ultimately leads to a persistent increase in RBCs [4]. It is well-established that exposure to HA/HH can induce erythropoiesis, yet the pathogenesis of eryptosis under these conditions remains poorly understood.

The spleen plays a crucial role in maintaining erythropoietic homeostasis by effectively clearing impaired and senescent RBCs from circulation [6]. Particularly, red pulp macrophages (RPMs) within the spleen, serving as primary phagocytes, are responsible for clearing senescent, damaged, and abnormal erythrocytes from circulation to recycle iron [7]. RPMs initiate the process of RBC endocytosis and lysosomal digestion into heme, following the recognition of the RBCs earmarked for removal via their cell surface signals [8, 9]. Subsequently, heme oxygenase-1 (HO-1) decomposes the heme into biliverdin, carbon monoxide, and iron [10]. Most of the iron recycled from heme is transported out of the cell through a protein called ferroportin (Fpn). This iron then binds to transferrin (Tf), a plasma protein that transports iron in the blood. The iron-transferrin complex interacts with the transferrin receptor (TfR) on the cell membrane, contributing to the formation of RBCs in the erythroid compartment [11]. A portion of the released iron is loaded into cellular ferritin (Ft; including Ft-H and Ft-L). In the presence of oxygen, the Ft-H facilitates the oxidation of Fe²⁺ (ferrous ion) to Fe³⁺ (ferric ion). Subsequently, the Ft-L stores this Fe³⁺ [12]. The Ft-L features a nucleation site, consisting of a cluster of cavity-exposed carboxyl residues, which readily bind to Fe³⁺, thereby simplifying the storage process [13]. When the body’s iron metabolism is vigorous, nuclear receptor coactivator 4 (NCOA4) can recognize Ft-H, bring Ft into the lysosomal pathway for degradation, and release iron in Ft for iron utilization in the body [14]. However, some iron is free in cells in the form of ferrous iron (Fe²⁺), which is well known to be an important initiator of free radical oxidation. Moreover, the accumulation of large amounts of Fe²⁺ ions in cells may cause ferroptosis [15].

Despite the extensive study of erythropoiesis under HA/HH conditions, the precise effects of HA/HH on erythrophagocytosis within the spleen remain largely unexplored. Considering the significant role of the spleen in RBC processing and the crucial function of RPMs in iron recycling from RBC clearance, we evaluated the impacts of HA exposure on the spleen/splenic macrophages using an HH-exposed mouse model (simulating 6000 m exposure conditions). In the current study, we sought to further investigate whether high-altitude/hypobaric hypoxia (HA/HH) can influence the erythrocyte disposal and iron recycling processes in macrophages. More specifically, we aimed to determine the potential impact of HA/HH exposure on the integrity of the erythrophagocytosis process. We discovered that exposure to HH triggered ferroptosis in the spleen, particularly in macrophages. This led to a reduction in macrophage numbers, which was subsequently followed by disruptions in eryptosis and iron recycling within the spleen. These findings may hold clinical significance, particularly in the context of continuous pathological erythrocytosis and the progression of HAPC under HA exposure.

Methods

HH mouse exposure model

Male C57BL/6 male mice (at 8 weeks of age) were obtained from the animal experimental centre of Nantong University. Mice were adapted to the facilities for 3 days before experiments. Mouse models of HH exposure were established by placing the mice in the decompression chamber (TOW-INT TECH, ProOx-810, China) for 1 d, 2 d, 3 d, 7 d, and 14 d. The parameters were set as follows: 6000 m above sea level, oxygen partial pressure of 11 kPa, lifting speed of 20 m/s, chamber pressure of 54 kPa, temperature of 25 ℃, humidity of 40%, and 12-h light/dark cycles. Tuftsin (1.5 mg/kg, MCE, HY-P0240, USA) was used to stimulate the phagocytosis of macrophages [16], and was injected at 7 d and 11 d during the 14-d period of mice exposed to HH.

Splenic macrophage culture and treatments

After isoflurane anaesthesia, mice were transcardially perfused with precooled PBS, and then splenic tissue was collected. The spleen was then dissected into minuscule fragments, each approximately 1 mm³ in volume, and separated in cold, phenol red-free Hanks’ balanced salt solution (HBSS; Gibco, Grand Island, NY, USA). The cell extracts were centrifuged at 500 rpm for 5 min at 4 ℃. The pellets were then resuspended and incubated with ammonium-chloride-potassium (ACK) lysing buffer (Thermo Fisher Scientific, USA) for 3 min at room temperature. Next, cell extracts were purified in a four-phase Percoll (Sigma‒Aldrich, St. Louis, MO, USA) gradient: 0, 30, 40, and 50% in HBSS. After centrifugation at 500 rpm for 20 min at 4°C, the macrophage-enriched fraction was collected at the interface between the 30% and 40% phases. Cells were washed in two rounds with HBSS and then resuspended in AIM V medium (Thermo Fisher Scientific). The cells in this medium were cultured at 37 ℃ and 5% CO₂ for 12 h. After the cells were treated with ferrostatin-1 (Fer-1, 2 μm; Selleck Chemicals, S7243, USA) 1 h in advance, the cells were placed in a hypoxia workstation (Invivo2, Ruskinn, UK) for 24 h.

Blood smear and hematological indices

The collected blood was gently mixed with EDTA (15 g/L) to obtain anticoagulant blood, which was detected by a hematology analyser (XE-5000, Sysmex Corporation, Japan). Blood smear preparation method: a blood sample is dropped onto a slide, and another slide is used to push the blood sample into a uniform thin layer at a constant speed. After the blood samples were dried, Wright’s stain solution was added and incubated for 1 min. Then, the slide was rinsed with water. Blood smears were photographed with a DM4000B microscope (Leica, Germany).

Lillie staining

The Lillie staining method was employed to ascertain Fe²⁺ deposition within the spleen, according to the protocol delineated by Liu et al [17]. Briefly, paraffin-embedded spleen sections underwent a process of dewaxing and rehydration via xylene and gradient alcohol. Then, the Lillie staining solution (G3320, Solarbio, China) was applied at 37°C for 50 min in a light-restricted environment, improved by a 5-min PBS wash. Subsequently, a mixture comprising 30 % hydrogen peroxide and methanol was incubated for 20 min, and then the sections were subjected to two 5-min washes with PBS. The DAB developing solution, which is not included in the Lillie bivalent iron staining kit, was applied for light-sensitive staining. The sections were stained with a nuclear fast red solution in a light-restricted setting for 10 minutes, followed by a 5-second wash with ddH2O. Lastly, the sections were dehydrated with gradient alcohol, cleared with xylene, and mounted with neutral resin.

Wright staining

Mice were anaesthetized and perfused with saline. The spleen was subsequently excised to prepare a single cell suspension, achieved by filtering through a 70 μm filter. Samples were centrifuged at 500 g for 5 min for cell separation. The cell pellet was subjected to three PBS washes. The suspension was then smeared onto a slide and air-dried. Following this, the slide was stained with Wright solution (G1040, Solarbio, China) for 3 min. An equivalent volume of pH 6.4 phosphate buffer was introduced, and the slide was gently agitated to allow mixing with the Wright’s stain solution for 5 min. After washing and drying, a microscope (DM4000B, Leica, Germany) was utilized for visualization and image acquisition.

Western blot

Tissue and cell samples were homogenized on ice after cell lysates and protease inhibitors were added. Centrifugation was performed at 12,000 rpm for 15 min at 4°C to collect the supernatant. Protein concentration was determined by the BCA detection method. A sample containing 30 μg protein was loaded and run in each well of SDS‒ PAGE gels. The membranes were incubated with the following antibodies: HO-1 (1:2000, Abcam, ab13243, USA); Ft-L (1:1000, Abcam, ab69090, USA); Ft-H (1:1000, Novus, NBP1-31944, USA); NCOA4 (1:1000, Santa Cruz, sc-373739, USA); TfR (1:1000, Thermo Fisher, 13-6800, USA); Fpn (1:1000, Novus, NBP1-21502, USA); ACSL4 (1:1000, Santa Cruz, sc-271800, USA); xCT (1:1000, Proteintech, 26864-1-AP, USA); Gpx4 (1:1000, Abcam, ab125066, USA); CD206 (1:1000, RD, AF2535, USA); CD16 (1:2000, RD, AF1960, USA); and β-actin (1:1000, Sigma‒Aldrich, A5316, USA). The secondary antibodies were as follows: goat anti-mouse (1:10000, Jackson, USA) and goat anti-rabbit (1:10000, Jackson, USA). The gray value of specific blots was scanned and analysed using ImageJ software (National Institute of Health, USA).

RT‒PCR analysis

Total RNA extraction and RT‒PCR analysis were performed essentially as described previously [18]. Primer sequences were as follows:

GEO analysis

GSE46480 samples were found in the GEO database of NCBI with the keyword “High Altitude” search. The data set was divided into two groups, one for the plain (Base Camp) and the other for the plateau (Altitude), with a sample size of 98 for each group. Blood samples were collected from the same person at McMurdo Station (48 m) and immediately transferred to Amundsen-Scott South Pole Station (2835 m) on the third day. R language was used for data analysis, and GraphPad Prism was used for statistics.

Malondialdehyde (MDA), Cysteine (Cys) and Glutathione (GSH) content detection

MDA was detected by a Micro Malondialdehyde Assay Kit (Solarbio, BC0025, China). Cys was detected by a Micro Cysteine Assay Kit (Solarbio, BC0185, China). GSH was detected by a Micro Reduced Glutathione Assay Kit (Solarbio, BC1175, China).

Immunofluorescence

Spleen sections of 9 μm thickness were flash-frozen and stored at –80°C. Before immunofluorescence staining, these sections were allowed to acclimate to room temperature for 30 minutes, then incubated in 10% BSA-PBS for 10 minutes. Next, the sections were incubated overnight with F4/80-PE (1:200, Biolegend, 123110, USA) and then washed thrice with PBS. Due to the limitations of F4/80 as a definitive macrophage marker, we concurrently conducted immunohistochemical analysis for heme oxygenase-1 (HO-1) (1:200, Abcam, ab13243, USA). Following this, sections were washed thrice with 0.05% PBST, incubated with secondary antibodies DAR-Alexa555 at room temperature for 2 hours, washed thrice with PBS again, and finally sealed with 50% glycerine-PBS. Fluorescence microscopy images were captured using a confocal laser scanning microscope (SP8, Leica Microsystems, Wetzlar, Germany). To quantify the immunostaining intensity of F4/80 and HO-1, we utilized ImageJ software. Specifically, we captured multiple fields of view for each slide, and the software was used to measure the mean intensity of the fluorescent signal in these images. These measurements were then normalized to the NN group to account for any experimental variations.

Tissue iron staining (DAB-enhanced Perls’ staining)

DAB-enhanced Perls’ staining for the spleen paraffin sections was performed as described previously [19]. Briefly, the sections were washed with PBS and cultured in freshly prepared Perls’ solution (1% potassium ferricyanide in 0.1 M hydrochloric acid buffer). The slides were then immersed and stained with DAB. All slides were counterstained with hematoxylin and visualized under a DM4000B microscope (Leica, Germany). Data were collected from three fields of view per mouse and semiquantitatively analysed with ImageJ software. Quantitative results of iron staining were finally normalized to the NN control group.

Flow cytometry

• 1) The steps of reticulocyte ratio detection were as follows: Whole blood was extracted from the mice and collected into an anticoagulant tube, which was then set aside for subsequent thiazole orange (TO) staining [20]. The experimental tube and negative control tube were prepared, 125 μL normal saline, 4 μL anticoagulant and 125 μL TO working solution (1 μg/mL, Sigma, 390062, USA) were added to each tube, and 250 μL normal saline and 4 μL anticoagulant were added to each tube of the negative control tube. After incubation at room temperature for 1 h, flow cytometry analysis was carried out by using the FL1 (488 nm/525 nm) channel.20]. The experimental tube and negative control tube were prepared, 125 μL normal saline, 4 μL anticoagulant and 125 μL TO working solution (1 μg/mL, Sigma, 390062, USA) were added to each tube, and 250 μL normal saline and 4 μL anticoagulant were added to each tube of the negative control tube. After incubation at room temperature for 1 h, flow cytometry analysis was carried out by using the FL1 (488 nm/525 nm) channel.

• 2) The steps for the determination of intracellular divalent iron content and lipid peroxidation level were as follows: Splenic tissue was procured from the mice and subsequently processed into a single-cell suspension using a 40 μm filter. The RBCs within the entire sample were subsequently lysed and eliminated, and the remaining cell suspension was resuspended in PBS in preparation for ensuing analyses. A total of 1 × 106 cells were incubated with 100 μL of BioTracker Far-red Labile Fe2+ Dye (1 mM, Sigma, SCT037, USA) for 1 h or C11-Bodipy 581/591 (10 μM, Thermo Fisher, D3861, USA) for 30 min. After the cells were washed with PBS twice, flow cytometry analysis was carried out by using the FL6 (638 nm/660 nm) channel for determination of intracellular divalent iron content or the FL1 (488 nm/525 nm) channel for determination of lipid peroxidation level.⁶ cells were incubated with 100 μL of BioTracker Far-red Labile Fe²⁺ Dye (1 mM, Sigma, SCT037, USA) for 1 h or C11-Bodipy 581/591 (10 μM, Thermo Fisher, D3861, USA) for 30 min. After the cells were washed with PBS twice, flow cytometry analysis was carried out by using the FL6 (638 nm/660 nm) channel for determination of intracellular divalent iron content or the FL1 (488 nm/525 nm) channel for determination of lipid peroxidation level.

• 3) The steps for detecting the mortality of spleen cells and peritoneal macrophages were as follows: 1 × 106 cells were incubated at room temperature for 40 min with 2 μM Calcein AM and 8 μM PI. Flow cytometry analysis was carried out by using FL1 (488 nm/525 nm, Calcein AM) and FL3 (488 nm/620 nm, PI) channels.⁶ cells were incubated at room temperature for 40 min with 2 μM Calcein AM and 8 μM PI. Flow cytometry analysis was carried out by using FL1 (488 nm/525 nm, Calcein AM) and FL3 (488 nm/620 nm, PI) channels.

• 4) The steps for detecting the number of M1/M2 macrophages in the spleen were as follows: 1 × 106 cells were incubated with 2% mouse serum-PBS for 10 min, incubated with F4/80-PE (1:200), CD86-PE/Cyanine7 (1:20, Biolegend, 105014, USA), and CD206-Alexa 647 (1:80, BD, 565250) for 30 min, and washed with 2% mouse serum-PBS twice. Flow cytometry analysis was carried out by using FL2 (488 nm/575 nm, F4/80-PE), FL5 (488 nm/755 nm, CD86-PE/Cyanine7) and FL6 (638 nm/660 nm, CD206-Alexa 647) channels.⁶ cells were incubated with 2% mouse serum-PBS for 10 min, incubated with F4/80-PE (1:200), CD86-PE/Cyanine7 (1:20, Biolegend, 105014, USA), and CD206-Alexa 647 (1:80, BD, 565250) for 30 min, and washed with 2% mouse serum-PBS twice. Flow cytometry analysis was carried out by using FL2 (488 nm/575 nm, F4/80-PE), FL5 (488 nm/755 nm, CD86-PE/Cyanine7) and FL6 (638 nm/660 nm, CD206-Alexa 647) channels.

• 5) The steps for detecting the number of monocytes in blood, spleen and bone marrow were as follows: 1 × 106 cells were incubated with 2% mouse serum-PBS for 10 min, incubated with F4/80-PE, CD11b-PE/CY7 (1:2000, BD, 552850, USA), and Ly6C-APC (1:2000, Thermo Fisher, 17-5932-82) for 30 min, and washed with 2% mouse serum-PBS twice. Flow cytometry analysis was carried out by using FL2 (488 nm/575 nm, F4/80-PE), FL5 (488 nm/755 nm, CD11b-PE/CY7) and FL6 (638 nm/660 nm, Ly6C-APC) channels.⁶ cells were incubated with 2% mouse serum-PBS for 10 min, incubated with F4/80-PE, CD11b-PE/CY7 (1:2000, BD, 552850, USA), and Ly6C-APC (1:2000, Thermo Fisher, 17-5932-82) for 30 min, and washed with 2% mouse serum-PBS twice. Flow cytometry analysis was carried out by using FL2 (488 nm/575 nm, F4/80-PE), FL5 (488 nm/755 nm, CD11b-PE/CY7) and FL6 (638 nm/660 nm, Ly6C-APC) channels.

Phagocytosis of E. coli and RBCs

E. coli was labelled with Cy5.5 (5 mg/mL), and RBCs were labelled with NHS-biotin (20 mg/mL). Macrophages (1 × 10⁶) were coincubated with E. coli-Cy5.5 or RBC-Biotin for 30 min and washed with 2% mouse serum-PBS twice. Flow cytometry analysis was carried out by using FL6 (638 nm/660 nm) to determine the phagocytosis of spleen by E. coli. Macrophages (coincubated with Biotin-RBCs) were incubated with streptavidin-FITC for 3 h and washed twice with 2% mouse serum PBS. FL1 (488 nm/525 nm) was used for flow cytometry analysis to determine the phagocytosis of RBCs in the spleen.

Single-cell RNA Sequencing

The spleen tissues were surgically removed and stored in MACS Tissue Storage Solution (Miltenyi Biotec, Bergisch Gladbach, Germany) until processing. Single-cell dissociation was performed by the experimentalists at the GENECHEM laboratory (Shanghai, China). Dissociated single cells were then stained for viability assessment using Calcein-AM (BD Biosciences, USA) and Draq7 (BD Biosciences). The BD Rhapsody system was used to capture transcriptomic information from single cells. Single-cell capture was achieved by random distribution of a single-cell suspension across >200,000 microwells through a limited dilution approach. Beads with oligonucleotide barcodes were added to saturation so that a bead was paired with a cell in a microwell. The cells were lysed in the microwell to hybridize mRNA molecules to barcoded capture oligos on the beads. Beads were collected into a single tube for reverse transcription and ExoI digestion. Upon cDNA synthesis, each cDNA molecule was tagged on the 5′ end (that is, the 3′ end of an mRNA transcript) with a unique molecular identifier (UMI) and cell barcode indicating its cell of origin. Whole transcriptome libraries were prepared using the BD Rhapsody single-cell whole-transcriptome amplification (WTA) workflow, including random priming and extension (RPE), RPE amplification PCR and WTA index PCR. The libraries were quantified using a High Sensitivity D1000 ScreenTape (Agilent) and High Sensitivity D1000 Reagents (Agilent) on a 4150 TapeStation System (Agilent, Palo Alto, CA, USA) and the Qubit High Sensitivity DNA assay (Thermo Fisher Scientific). Sequencing was performed on an Illumina sequencer (Illumina Nova Seq 6000, San Diego, CA) in a 150 bp paired-end run.

Statistical analysis

Statistical analysis was conducted using GraphPad Prism 8.0 software. All values were represented as the mean ± standard error (SEM). The homogeneity of variance was verified before the application of a parametric test. A two-tailed Student’s t-test was employed for data from two groups, while a one-way analysis of variance (ANOVA) with multiple comparisons using Tukey’s post hoc test was utilized for data from multiple groups. A p-value of less than 0.05 was considered statistically significant.

Results

HH exposure promotes erythrocytosis in mice

To mimic 6000 m HA exposure, we placed C58BL/6 mice in an animal hypobaric oxygen chamber and detected the blood indices in the blood of the mice after HH exposure for different times. The blood smear showed that the number of RBCs was increased from 3 to 14 days after HH exposure compared with the normobaric normoxia (NN) group (Figure 1A). Routine blood tests further confirmed the results in blood smears, which showed that the RBC number (Figure 1B), HGB content (Figure 1C) and HCT value (Figure 1D) were all increased significantly to varying degrees, while MCH was not changed after HH exposure (Figure 1E). We further performed flow cytometry by TO staining to detect reticulocytes after 7 and 14 days of HH treatment. The results showed that at both HH 7 and 14 d of exposure, the reticulocyte proportion was increased (Figure 1F). These results suggested that the HH-treated mouse model mimics HA exposure well, which promotes erythropoiesis and results in erythrocytosis in mice following HH exposure.

Fig. 1

Spleen inhibits the immoderate increase in RBCs under HH conditions

To determine the roles of the spleen in RBC homeostasis under HA/HH, we investigated the effects of HH on the morphology, volume and weight of the spleen as well as erythrocyte indices. As shown in Figure 2, the spleen volume and weight were decreased significantly after HH exposure for 1 day compared to NN treatment (Figure 2, A-C). However, the spleen was obviously enlarged from 2 to 14 days after HH exposure (Figure 2, A-C). The results indicated that the spleen contracted, the stored RBCs in the spleen were released into the blood at 1 day, and the RBCs were produced and/or retained in the spleen from 2 to 14 days after HH exposure. In our research, we also examined the influence of the spleen on RBCs homeostasis under HH conditions. We investigated whether the role of spleen in RBCs clearance under HH conditions could be compensated by the liver or other components of the mononuclear macrophage system. To conduct this, we performed splenectomies on mice and subsequently exposed them to HH conditions for 14 days. This allowed us to monitor RBCs counts and blood deposition. Our findings indicated that, in comparison to both the splenectomized mice under NN conditions and the sham-operated mice exposed to HH, erythrocyte deposition (Figure 2D) and counts (Figure 2E), as well as HGB (Figure 2F) and HCT (Figure 2G) levels, significantly increased 14 days post-splenectomy under HH conditions. Meanwhile, MCH levels remained stable (Figure 2H). These indices did not vary in mice, regardless of whether they had undergone a splenectomy, under NN conditions (Figure 2D and E). These results suggest that the liver and other mononuclear macrophages do not effectively compensate under HH conditions. Furthermore, our research highlights the crucial role of the spleen in maintaining RBCs homeostasis under conditions of HH. The study suggests that the spleen functions to regulate the proliferation of RBCs, preventing their overproduction in response to HH exposure.

Fig. 2

Reduction of splenic macrophages due to HH exposure

Considering macrophages as the primary cell type responsible for processing RBCs within the spleen under physiological conditions, we subsequently investigated the population and activity of these macrophages after exposure to HH for 7 or 14 days, employing flow cytometry and single-cell sequencing techniques. Calcein/PI double staining, examined via flow cytometry, revealed a significant decrease in viable spleen cells and a concurrent increase in dead cells following 7 and 14 days of HH exposure (Figure 3, A-C). Single-cell sequencing further disclosed a significant reduction in splenic macrophages after 7 days of HH exposure (Figure 3, D-F). Additionally, the expressions of CD16 (M1 macrophage marker) and CD206 (M2 macrophage marker) were observed to decrease after both 7 and 14 days of HH exposure (Figure 3, G-L). Western blot analysis corroborated the flow cytometry findings, showing a decrease in mature macrophages (F4/80⁺/CD11b⁺), including both M1-type (F4/80⁺/CD86⁺) and M2-type (F4/80⁺/CD206⁺) macrophages, in the spleens of mice after 7 days of HH exposure (Figure 3M). To further elucidate the migration and differentiation of monocytes from the bone marrow to the spleen, we analyzed the expression of chemokines CCL2, CCL7, Csf1, and Csf2 in the spleen using qPCR and determined the number of monocytes in the bone marrow (BM) and spleen via flow cytometry. Our results indicated a significant reduction in the expression of CCL2, CCL7 (Figure 3N), Csf1, and Csf2 (Figure 3O) in the spleen after 7 and 14 days of HH exposure. Furthermore, the number of monocytes (Ly6C⁺/CD11b⁺) in the bone marrow (Figure 3, P-R) and spleen (Figure 3, P and S-T) also exhibited a decline after 7 and 14 days of HH exposure. 373 We evaluated the depletion of macrophages in the spleen under HH conditions by examining the expression and distribution of HO-1 and F4/80. Figures 3U-W depict a decrease in both HO-1 and F4/80, predominantly within the spleen’s red pulp following HH exposure. Together, these findings indicate a reduction in the number of splenic macrophages after HH exposure, which could impair the spleen’s capacity to process erythrocytes.

Fig. 3

The impact of HH exposure on erythrophagocytosis and iron processing capacity of the spleen

We investigated the influence of HH exposure on erythrocyte phagocytosis and heme iron recycling within splenic macrophages. We administered NHS-biotin intravenously to the mice followed by HH treatment in vivo. The clearance of RBCs was monitored by detecting the retention of biotin-labelled RBCs using flow cytometry in both the blood and spleen of the HH-treated mice. We observed a substantial decrease in biotin-labelled RBCs in the blood and spleen cells at 7-and 14-days post HH exposure (Figure 4, A-D). Despite the natural decrease of labelled RBC over time, the decay was more pronounced in the HH exposure group compared to the normobaric normoxia (NN) group in both the blood (Figure 4, A and C) and spleen (Figure 4, B and D). The phagocytic ability of mouse spleen macrophages towards RBCs was significantly more impaired after HH exposure, particularly on the 14th day (Figure 4, A-D). Additionally, to substantiate our findings, we quantified the retention of RBC counts by Wright staining in splenic cells after 7 and 14 days of HH exposure (Figure 4, E-F). This confirmed impaired erythrophagocytosis in the spleen under HH exposure, as evidenced by an increase in deformation and retention of RBCs following 7 and 14 days of HH exposure. Furthermore, we employed Tuftsin to stimulate phagocytosis of macrophages and carried out immunofluorescence and iron staining to examine the heme iron recycling capability of the spleen under HH conditions (Figure 4G). As depicted in Figure 4H, compared with the NN group, F4/80 expression and iron deposition in the red pulp were significantly reduced after HH exposure. However, Tuftsin led to an induction of F4/80 expression and iron deposition in the red pulp of the spleen under HH (Figure 4, I-L). These results imply that the capacity for erythrocyte phagocytosis and heme iron recycling by the spleen was compromised, principally due to the reduction in macrophages in the red pulp under HH conditions.

Fig. 4

HH exposure induces iron mobilization and ferroptosis in the spleen

To elucidate the precise mechanisms underlying the macrophage reduction caused by HH, we examined the expression of proteins related to iron metabolism and ferroptosis in mice treated with HH and analyzed the corresponding gene expression in peripheral blood mononuclear cells (PBMCs) from healthy humans acutely exposed to HA conditions using GEO data (No. GSE46480). The results showed that, compared to the NN group, the expression levels of HO-1, Ft-L, Ft-H, NCOA4, and xCT were decreased, while Fpn, TfR, and ACSL4 expressions were significantly increased after 7 and 14 days of HH exposure (Figure 5, A-B, D-E; Figure S1, A-D).

Fig. 5

With the exception of NCOA4, the alterations in gene expressions related to iron metabolism and ferroptosis in PBMCs were consistent with our Western blot results (Figure 5, C and F). The GPX4 gene and protein expressions remained largely unchanged in both human PBMCs and mouse spleens. The changes in iron metabolism-and ferroptosis-related genes in PBMCs reflect the protein changes in the spleen under HH exposure. We detected Fe²⁺ and lipid ROS levels in the spleen by flow cytometry, and quantified MDA, GSH, and Cys levels using biochemical detection kits. As depicted in Figure 5, G and H; Figure S1, I-K, the Fe²⁺ (Figure 5G; Figure S1, E and F) and lipid ROS (Figure 5H; Figure S1, G and H) levels in the spleen increased significantly after HH exposure. Additionally, the MDA content (Figure 5I; Figure S1I) in the spleen increased, whereas Cys (Figure 5J; Figure S1J) and GSH (Figure 5K; Figure S1K) levels significantly decreased after HH exposure. To ascertain the increased Fe²⁺ primarily emanated from the red pulp, we detected Fe²⁺ deposition and distribution in the spleen by Lillie stain following 7 and 14 days of HH exposure (Figure 5, L-M; Figure S1, L-M). These results demonstrated increased Fe²⁺ primarily deposited in the red pulp of the spleen. Taken together, these findings suggest that iron mobilization in the spleen was enhanced and ferroptosis was induced after 7 days of HH exposure.

Hypoxia induces ferroptosis in primary splenic macrophages

To investigate whether hypoxia exposure can trigger ferroptosis in macrophages, we measured alterations in Fe²⁺ and lipid ROS levels, cell viability, phagocytosis, and ferroptosis-related protein expressions in primary splenic macrophages under hypoxia in the presence of a ferroptosis inhibitor (Fer-1). As depicted in Figure 6, compared with the normoxia control group (Nor), Fe²⁺ (Figure 6, A-C) and lipid ROS (Figure 6, D-E) levels significantly increased, whereas the number of viable cells (Figure 6G-H) and phagocytosis ability (Figure 6I) significantly decreased after 24 h of hypoxia (Hyp) treatment. In addition, prolonged hypoxia exposure led to a gradual increase in ACSL4 expression, while xCT and GPX4 expressions decreased (Figure 6F). Fer-1 mitigated the increase in Fe²⁺ content (Figure 6, J-K) and reversed the expression changes in ACSL4, xCT, and GPX4 induced by hypoxia exposure (Figure 6, L-O). Simultaneously, Fer-1 reversed the alterations in MDA, Cys, and GSH levels induced by hypoxia (Figure 6, P-R). These findings confirm that hypoxia induced ferroptosis in primary splenic macrophages.

Fig. 6

Discussion

The purpose of this study was to explore the effects and mechanisms of HA/HH exposure on erythrophagocytosis and iron circulation in mouse spleens. Here, we used an HH chamber to simulate 6000 m HA exposure and found that HH exposure induced iron mobilization and activated ferroptosis in the spleen, especially in red pulp macrophages (RPMs), which subsequently inhibited the phagocytosis and clearance of RBCs. Finally, chronic exposure to HA/HH may promote the retention of RBCs in the spleen, cause splenomegaly, advance RBC production, and promote the occurrence and development of HAPC (Figure 7).

Fig. 7

RBCs/erythrocytes are the carriers of oxygen and the most abundant cell type in the body. Erythrocytes are rapidly increased by triggering splenic contraction in acute HA/HH exposure [21] and stimulating erythropoiesis in subsequent continuous or chronic HA/HH exposure. Changes in spleen morphology and size are closely related to the spleen’s ability to recover RBCs [22], and studies have shown that the spleen is in a contraction state after short-term exposure to HA/HH, and approximately 40% of the increase in RBCs is due to the contraction of the spleen [21, 23, 24]. In the present study, mice were treated under HH to mimic HA exposure (mainly mimicking the low oxygen partial pressure environment of 6000 m HA) for different times according to other studies [25, 26]. Our results showed that HH exposure not only significantly increased the RBC and HGB contents but also significantly increased CD47⁺ cells in blood. However, short-term (1 d) HH exposure caused spleen contraction and induced splenomegaly after 3 d of HH treatment. This contraction can trigger an immediate release of RBCs into the bloodstream in instances of substantial blood loss or significant reduction of RBCs. Moreover, elevated oxygen consumption rates in certain animal species can be partially attributed to splenic contractions, which augment hematocrit levels and the overall volume of circulating blood, thereby enhancing venous return and oxygen delivery [27, 28]. Thus, we hypothesized that the body, under such conditions, is incapable of generating sufficient RBCs promptly enough to facilitate enhanced oxygen delivery. Consequently, the spleen reacts by releasing its stored RBCs through splenic constriction, leading to a measurable reduction in spleen size. These results are consistent with other research results; that is, HH or HA exposure affects spleen morphology and further 483 affects RBC counts and HGB levels [29–31]. Considering these findings, the present study postulates that hypoxia-induced inhibition of erythrophagocytosis may lead to RBC retention. However, we acknowledge that the manuscript in its current preprint form does not offer conclusive evidence to substantiate this hypothesis. To bridge this gap, we further conducted experiments where the spleen was perfused, and total cells were collected post HH exposure. These cells were then smeared onto slides and subjected to Wright staining. Our results unequivocally demonstrate an evident increase in deformation and retention of RBCs in the spleen following 7 and 14 days of HH exposure. This finding strengthens our initial hypothesis and contributes a novel perspective to the understanding of splenic responses under hypoxic conditions.

HAPC is a common chronic HA disease characterized by excessive proliferation of RBCs caused by HH conditions [32]. Under physiological conditions, RBC homeostasis is maintained by balancing erythropoiesis and erythrocyte clearance [4]. The ability of RBC disposal in the spleen for iron recycling under HA/HH exposure was important to RBC regeneration in bone marrow (BM) [33]. Thus, we hypothesized that erythrophagocytosis and iron recycling in the spleen were altered by HA/HH exposure and further disturbed RBC homeostasis and affected the progression of HAPC. We found that compared with sham group mice, HH significantly increased the contents of RBCs and HGB in the blood of splenectomy mice. This strongly verified that the spleen is a key organism that maintains RBC magnitude within a certain range of physiological statuses under HA/HH conditions.

As originally proposed by Metchnikoff in the 19th century, macrophages, especially RPMs, play a pivotal role in the regulation of RBCs and iron homeostasis [34, 35]. We detected the macrophage population in the spleen and found that HH exposure for 7 and 14 days reduced the total macrophage and M1-and M2-type macrophage proportions in the spleen. This result is supported by another study, that is, hypoxia reduces M1-type macrophages in human gastric cancer [36]. We next observed whether monocyte migration and differentiation from the BM to the spleen replenish reduced macrophages and sustain macrophage homeostasis after HH treatment. It is well known that splenic erythropoiesis is a dynamic process [4]. Upon phagocytosing erythrocytes in macrophages, spleen tissue (including macrophages and fibroblasts) produces the chemokine C–C motif chemokine ligand 2 and 7 (CCL2 and CCL7) to recruit blood monocytes to the spleen [37, 38]. Unlike most studies in tumours, which mainly suggested that macrophage accumulation in hypoxic areas was derived by enhanced chemokine expression in tumours [39, 40], CCL2, CCL7 and Csf1 expression was decreased in the spleen after HH exposure for 7 and 14 days in our study. Furthermore, we found that HH exposure inhibited Ly6C^hi monocyte migration from the BM to spleen, where these cells subsequently differentiated into RPMs. The combination of decreased splenic RPMs, along with the reduced BM-derived Ly6C^hi monocytes, suggests that the homeostasis of the RPM population of circulating monocytes was also inhibited after the RPM depletion induced by HH exposure. The observed decrease in monocytes within the BM is likely attributable to the fact that monocytes and precursor cells for RBCs both originate from the same hematopoietic stem cells within the BM [41]. As such, the differentiation to monocyte is reduced under hypoxic conditions, which may subsequently cause a decrease in migration to spleen. Furthermore, we hypothesize that an increased migration of monocytes to other tissues under HH exposure may also contribute to the decreased migration to the spleen. The liver, which partially contributes to the clearance of RBCs, may play a role in this process. Our investigations to date have indeed identified an increased monocyte migration to the liver. We were pleased to discover an elevation in CSF1 expression in the liver following HH exposure for both 7 and 14 days. This finding was corroborated through flow cytometry, which confirmed an increase in monocyte migration to the liver.

HO-1 is a major antioxidant and cytoprotective enzyme that catalyzes the degradation of heme to provide heme homeostasis and protect against free heme-induced toxicity [42]. In addition, it was supposed that mice deficient in HO-1 are largely devoid of RPMs and suggested that HO-1 is critical for RPM development and survival after erythrocyte clearance [43]. The reduced HO-1 protein of red pulp in our study also implied decreased macrophage accounts induced by HH exposure, although HO-1 was considered to be upregulated by hypoxia-inducible factor 1 (HIF-1) under hypoxia [44, 45]. Along with the decreased macrophage amount induced by HH, the phagocytic ability of macrophages in the spleen was also inhibited both in vitro and in vivo. Meanwhile, the injection of tufftsin strongly confirmed the enhancement of heme iron processing in the spleen under HH exposure. Nevertheless, our data conflict with the study of Anand et al, which reported that hypoxia causes an increase in phagocytosis by macrophages in a HIF-1α-dependent manner [46]. This may be because they performed intermittent hypoxia, while we used consistent hypoxia in the present study. Macrophages play a crucial role in maintaining erythrocyte homeostasis, partially due to their function in digesting the HGB content of cleared RBCs and recycling the iron back to erythroid progenitors for heme synthesis and HGB production [47]. In mammals, the majority of bodily iron exists in the form of heme, with the most substantial pool comprising HGB [48]. HGB contains four prosthetic hemoglobins, and each mature RBC is thought to contain approximately 1.2 × 10⁹ heme moieties [47]. As previously mentioned, the majority of iron required to sustain erythropoiesis is derived from recycled RBCs. In general, RPMs are equipped with molecular machinery capable of neutralizing the toxic effects of heme and metabolizing iron [49, 50]. Nonetheless, any disruptions in the process of erythrophagocytosis, the uptake and degradation of erythrocytes by macrophages, could potentially result in abnormal iron metabolism, potentially manifesting as conditions such as anemia or iron overload [49]. The release of heme upon processed RBCs constitutes a permanent and considerable threat for iron cytotoxicity in macrophages and may eventually result in a specific form of programmed cell death termed ferroptosis [51], which may cause decreased cell counts [52, 53].

Accordingly, we investigated the iron metabolism status and explored ferroptosis in spleen/macrophages after HH/hypoxia treatment and found that HH exposure prompted iron mobilization, especially in ferritinophagy and lipid peroxidation in the spleen. Interestingly, we found that except for NCOA4, all the gene expression changes in the PBMCs of humans after acute HA exposure were similar to the changes in the spleens of mice after HH exposure for 7 and 14 days. This is probably because NCOA4 in the spleen mediates iron release from Ft-L and facilitates iron reuse, while PBMCs do not possess this function in vivo. These results not only indicated that HH exposure induces spleen ferroptosis but also implied that the gene expression changes in PBMCs under HA/HH may reflect RBC processing functions in the spleen and further indicate the iron metabolism status in the clinic. We further found that the exact mechanism of macrophage ferroptosis induced by HH exposure was caused by increased Fe²⁺ and decreased antioxidative system expression, which finally resulted in lipid peroxidation of macrophages. It has been proposed that heme catabolism by HO-1 protects macrophages from oxidative stress [49]. The enhanced ROS and lipid peroxidation in vivo were also consistent with the decreased HO-1 expression in the spleen. In addition, 1% hypoxia treatment induced ferroptosis in vitro, which was reduced by treatment with ferrostatin-1, a ferroptosis inhibitor. However, our data were inconsistent with the study of Fuhrmann et al., which reported that hypoxia inhibits ferritinophagy and protects against ferroptosis [54]. We hypothesized that the enhanced iron demand for new RBC regeneration in BM caused by HH leads to an increase in NCOA4 and Ft-L protein expression in the spleen. On the other hand, hypoxia inhibited anti-ferroptosis system expression, including reduced Gpx4 expression and increased lipid ROS production. The results were supported by the group of Youssef LA et al, who reported that increased erythrophagocytosis induces ferroptosis in RPMs in a mouse model of transfusion [55]. However, as most studies have reported, ferroptosis is not only characterized by increased lipid ROS but also significantly shrinking mitochondria [56]. We never found shrinking mitochondria in RPMs after HH exposure (data not shown). Gao et al. proposed that mitochondria played a crucial role in cysteine deprivation-induced ferroptosis but not in that induced by inhibiting glutathione peroxidase-4 (GPX4), the most downstream component of the ferroptosis pathway [57]. Interestingly, in our in vivo study, Gpx4 expression was also not changed after HH exposure. It is still not clear whether mitochondria are involved in ferroptosis. Whether HH treatment caused mitochondrial swelling directly and the exact mechanism involved in this process still need to be further investigated.

When the splenic macrophages’ phagocytic function is compromised, the burden of managing the increased number of RBCs falls to the other cells within the spleen, as well as other clearance mechanisms within the body [58]. The endothelium, for instance, has been shown to be involved in the removal of aged and damaged RBCs [59]. The sinusoidal endothelial cells can recognize these RBCs, which often have altered surface properties, leading to their removal from circulation [59, 60]. Therefore, under HH conditions, endothelial cells may play a compensatory role in RBC clearance when macrophage function is impaired. Other macrophages, such as liver Kupffer cells and bone marrow macrophages, may also contribute to managing this buildup of RBCs [61]. Kupffer cells are the resident macrophages of the liver and have significant phagocytic ability. Similarly, macrophages in the bone marrow play a crucial role in iron recycling from aged erythrocytes [61, 62]. Therefore, these macrophages could potentially compensate for the reduction in splenic erythrophagocytosis, although their ability to effectively handle a large surge in RBC numbers remains to be determined.

It is also possible that the spleen may adapt to the increased load of RBCs by expanding the red pulp, leading to splenomegaly [63]. The red pulp is highly vascular and contains numerous sinuses lined with macrophages [64]. This expansion could potentially accommodate a greater number of RBCs, thus enhancing the spleen’s capacity for erythrophagocytosis and iron recycling. However, these compensatory mechanisms may be insufficient or become overwhelmed over time, leading to pathological conditions such as HAPC. Further research is necessary to fully understand the consequences of splenic ferroptosis on RBC homeostasis and how these effects could be mitigated or reversed. Finally, as these observations are based on experimental models, they should be corroborated in human studies to assess their clinical relevance and potential implications for managing conditions related to high altitude exposure. The potential for therapeutic interventions that target ferroptosis, erythrophagocytosis, and iron recycling should also be explored, as these could provide new avenues for treating HAPC and other conditions related to high altitude exposure.

Based on the above experiments, we first put forward the “spleen theory” of HAPC (Figure 7). It hypothesizes that HH exposure induced spleen ferroptosis, especially RPMs, which further inhibited erythrophagocytosis, RBC clearance, HGB processing and iron recycling. This theory identifies 637 a novel pathophysiological process in which HH exposure leads to the induction of spleen ferroptosis, particularly in red pulp. As a result, erythrophagocytosis, RBCs clearance, and iron recycling are impeded, causing a buildup of RBCs within the spleen, leading to splenomegaly and continued RBC production. The “spleen theory” of HAPC provides an intriguing new perspective on the physiological responses to hypoxia and highlights the role of the spleen as a vital organ in managing erythrocyte homeostasis under high altitude/hypoxic conditions. These findings may be clinically relevant to the pathological conditions of HAPC progression.

Abbreviations

BM: bone marrow; Cys: Cysteine; GSH: Glutathione; Ft-L: Ferritin light chains; Fpn: Ferroportin; Ft: Ferritin; HH: Hypobaric hypoxia; HA: High-altitude; HAPC: High-altitude polycythemia; HO-1: Heme oxygenase-1; HCT: Haematocrit; HGB: Haemoglobin; HIF-1a: Hypoxia inducible factor 1 a; MCH: Mean corpuscular haemoglobin; MDA: Malondialdehyde; NN: Normobaric normoxia; NCOA4: nuclear receptor coactivator 4; PBMCs: peripheral blood mononuclear cells; RBC: Red blood cell; RPM: Red pulp macrophage; TfR1: Transferrin receptor 1; Tf: Transferrin; TfR: Transferrin receptor.

Author contributions

QQL and GHW conceived, organized, and designed the study. YQG supervised the work. WPY, MQL, JD and JYL performed the experiments. WPY, GW and BL contributed to the analysis of data. QQL and GHW prepared, wrote and revised the manuscript. All authors contributed to read, and approved the submitted version.

Funding

This work was supported by Natural Science Foundation of China (Grants 32271228, 82171190, and 81873924), Open Cooperation Program from Key Laboratory of Extreme Environmental Medicine, Ministry of Education (KL2019GY011), China Postdoctoral Science Foundation (2020M673649), Cultivate Candidate of the Jiangsu Province “333” Project, and Nantong Municipal Science and Technology Project (MS12021020 and MS22021010).

Availability of data and materials

The datasets used and/or analyzed during the current study are available from the corresponding author on reasonable request.

Declarations

Ethics approval and consent to participate

All animal care and experimental protocols were carried out according to the Chinese Animal Management Rules of the Ministry of Health and were authorized by the Animal Ethics Committees of Nantong University research program protocol #S20190219-011. The results of the study are presented clearly, honestly, and without fabrication, falsification, or inappropriate data manipulation.

Consent for publication

Not applicable.

Competing interests

The authors have no relevant financial or non-financial interests to disclose.

Author details

¹Department of Physiology and Hypoxic Biomedicine, Institute of Special Environmental Medicine and Co-innovation Center of Neuroregeneration, Nantong University, 9 Seyuan Road, Chongchuan District, Nantong, Jiangsu 226019, China. ²College of High-Altitude Military Medicine, Institute of Medicine and Hygienic Equipment for High Altitude Region, Army Medical University, Chongqing 400038, China. ³Key Laboratory of Extreme Environmental Medicine and High-Altitude Medicine, Ministry of Education of China, Chongqing 400038, China. ⁴Department of Neurosurgery, Southwest Hospital, Army Medical University, Chongqing, Chongqing 400038, China.

Supplementary Figure S1. HH exposure promotes iron mobilization and induces ferroptosis in the spleen. The C57BL/6 mice were treated with NN and HH for 14 days, and the spleen was collected for subsequent detection. (A) Western blot detection of HO-1, Ft-L, Ft-H, NCOA4, Fpn and TfR protein expression in spleen. (B) Statistical analysis of HO-1, Ft-L, Ft-H, NCOA4, Fpn and TfR protein expression in A. (C) Western blot detection of ACSL4, GPX4 and xCT protein expression in the spleen. (D) Statistical analysis of ACSL4, GPX4 and xCT protein expression in C. (E) The content of Fe²⁺ in the spleen was detected using the FerroFarRed probe by flow cytometry. (F) Quantitative results of the mean fluorescence intensity in E. (G) The level of lipid ROS in the spleen was detected using the C11-BODIPY probe by flow cytometry. (H) Quantitative results of the mean fluorescence intensity in G. (I) MDA, (J) Cys, and (K) GSH levels in the spleen were detected using kits by biochemical methods. (L) Lillie staining of Fe²⁺ in the spleen after 14 days of HH exposure. (M) Fe²⁺ deposition in the spleen as described in (L) was quantified. Data are expressed as the means ± SEM (n = 3 per group); * P < 0.05, ** P < 0.01, *** P < 0.001 versus the NN group or the indicated group.