Platelet Reduction in Rats Exposed to Chronic Hypoxia Is Associated with Interaction of Glycoprotein Ib Alpha von Willebrand Factor

Abstract

Chronic high-altitude hypoxia is associated with reduced platelet count, but it is unclear whether the decrease in platelet count is due to impaired production or increased clearance. This study examines how hypoxia affects platelet production and apoptosis and elucidates the impact of glycoprotein Ibα–von Willebrand factor interaction on platelets in rats using a hypobaric hypoxia chamber. The results showed that the number of megakaryocytes increased under hypoxia; however, the levels of differentiation and polyploidy decreased, while those of apoptosis increased. Platelet production did not reduce according to the reticulated platelet percentage, while platelet apoptosis enhanced; these results suggest that increased platelet clearance was the main reason behind platelet reduction. Our previous microarray results indicated that glycoprotein Ibα (GPIbα) expression increased under hypoxia, which was a protein involved in platelet clearance; therefore, we examined the interaction of platelet GPIbα with the von Willebrand factor (vWF) both in vivo and in vitro to explore the effect of this process on platelets and whether it is related to platelet apoptosis. Under hypoxia, the stronger interaction between GPIbα and vWF promoted platelet apoptosis; inhibiting this interaction reduced platelet apoptosis and increased platelet counts. Platelet reduction is associated with apoptosis induced by the interaction between GPIbα and vWF.

Keywords

apoptosis - glycoprotein Ib alpha - hypoxia - platelet - von Willebrand factor

Introduction

Platelets not only participate in haemostasis and thrombosis but also play an important role in vascular repair, inflammation, tumour metastasis and immune regulation. Severe thrombocytopenia increases the risk of bleeding; the platelet count may reflect the outcome of the disease to a certain extent. For patients with sepsis or systemic lupus erythematosus, thrombocytopenia indicates a poor prognosis.[1] [2] [3] The number of circulating platelets is in a relatively stable state affected by platelet production and clearance. Chronic hypoxia could lead to thrombocytopenia, especially in patients with chronic altitude sickness.[4] [5] Hypoxia can inhibit the colony formation of megakaryocytes (MKs) in bone marrow.[6] Hypoxia-inducible factor-1α (HIF-1α), a nuclear transcription factor that mediates cell adaptation to hypoxia, decreases MKs in immune thrombocytopenic purpura patients,[7] indicating that hypoxia affects the production of platelets.

Research has shown that chronic hypoxia can affect platelet activity and participate in the occurrence and development of related diseases.[8] Under chronic hypoxia, the human body undergoes erythrocytosis[9] [10] [11] [12] and increases blood viscosity,[13] platelet activation and procoagulant substances, thus leading to arteriovenous thrombosis.[14] [15] [16] [17] [18] During thrombus formation, platelets are consumed, thereby lowering their counts.[19] [20] Platelets promote the inflammatory response of lung tissue under hypoxia and participate in the process of hypoxia-related pulmonary hypertension and pulmonary vascular remodelling.[21] Molecules on the surface of platelets can interact with other cells or cytokines released by activated platelets to cause inflammatory and thrombotic events.[22] Glycoprotein (GP) Ib–IX–V is the second most abundant molecule on the surface of platelets. Under shear stress, GPIb–IX–V can not only bind to VWF and collagen to induce platelet aggregation but also transmit VWF-binding signals to activate platelets. GPIb–IX–V can bind alphaMbeta2 on neutrophils, P-selectin, and activate platelets on endothelial cells, thrombin, and coagulation factors XI/XIIa in plasma and play a central role in inflammation and thrombosis.[23] [24] A previous study has reported platelet reduction accompanied by increased GPIb transcript levels in healthy individuals exposed to chronic hypoxia.[25] GPIb–IX–V plays an important role in the regulation of platelet clearance, including the apoptosis, removal of sialic acid from platelet surface glycoproteins (desialylation), glycan-mediated platelet clearance, and the clearance of VWF–platelet complexes.[26] [27]

Understanding the causes of platelet reduction under chronic hypoxia provides insights into the pathogenesis of hypoxia-related diseases. In this study, a rat model of chronic hypoxia was used to evaluate the effect of hypoxia on platelet production and apoptosis, as well as the effects of GPIb–IX–V on platelets.

Materials and Methods

The Experimental Animals

Sprague–Dawley (SD) rats (160–180 g) were purchased from Beijing HFK Bioscience Co., Ltd. (Beijing, China) and were housed in isolated, ventilated cage barrier facilities at the Animal Center of Xi'an Jiao Tong University and the Research Center for High Altitude Medicine of Qinghai University. The rats were maintained in a 12/12-hour light/dark cycle and an ambient temperature of 20 to 26 °C with sterile pellet food and water ad libitum, with one animal per cage. The minimal possible number of animals was killed and all effort was made to minimize their suffering. This study was approved by the Ethics Committee of the Affiliated Hospital of Qinghai University (P-SL-2020087) and complied with the animal management laws of the Health Commission of China.[28]

Animal Model Establishment

Male SD rats were divided randomly into three groups using a random number table. Animals were raised, respectively, at an altitude of 400 m (Xi'an, China; low-altitude (LA) group; n = 10), 2,260 m (Xining, China; mid-altitude (MA) group; n = 10), or 5,000 m (hypobaric hypoxia chamber, Dyc-3000; Guizhou Feng Lei Aviation Machinery Co., Ltd., Guangzhou, China; atmospheric pressure: 52.93 kPa, temperature: 22 °C, relative sea level oxygen concentration: 11.10%; high-altitude (HA) group; n = 20) for 30 days. In humans, males are more susceptible to hypoxia intolerance than females, which tends to cause chronic altitude sickness; therefore, male rats were selected.[29] The chronic hypoxia model was successfully established when the haemoglobin level of the rats in the HA group was ≥210 g/L.[30] [31] [32]

Preparation of Purified Platelets

SD rats were anaesthetised with isoflurane through an anaesthesia inhalation machine (R546Pro; Rayward Life Technology Co., Ltd, Shenzhen, China), and the absence of limb reflexes indicated successful anaesthesia. Whole blood (8 mL) was collected into an ACD vacutainer via pericardial puncture. Whole blood was centrifuged at 100 × g (no brake) for 20 minutes to obtain platelet-rich plasma (PRP). Subsequently, PRP supplemented with 100-nM prostaglandin E1 (PGE1; to prevent exogenous platelet activation, 13010.1; Cayman Chemical, Ann Arbor, Michigan, United States) was centrifuged at 200 × g for 20 minutes (no brake) to obtain washed platelets at the bottom. The supernatant was platelet-poor plasma, which was stored at −80 °C for follow-up enzyme-linked immunosorbent assay (ELISA) and washed platelets were resuspended in Pipes/Saline/Glucose solution (5 mM Pipes, 145 mM NaCl, 4 mM KCl, 50 M Na2HPO4, 1 mM MgCl2•6H2O, and 5.5 mM glucose) supplemented with PGE1 at 37 °C. Purified platelets were obtained via the negative selection of washed platelets supplemented with PGE1 using immunomagnetic beads (anti-CD45 microbeads to deplete leukocytes; 130-045-801; Miltenyi Biotec, Gladbach Bergisch, Germany) as previously described.[25] [33] [34] The isolated platelets were observed under a microscope to ensure >99.9% purity and subjected to flow cytometry and Western blotting.

Isolation and Culture of Megakaryocytes

MKs were cultured and isolated as previously described.[35] After blood sampling, the femurs and tibiae were dissected, and their marrow cavities were rinsed with phosphate-buffered saline (PBS, PYG0021; BOSTER, Wuhan, China). The collected fluid was centrifuged, and the bone marrow cells were obtained after red blood cell lysis was cultured in Iscove's Modified Dulbecco's Medium (IMDM). Bone marrow cells were plated at a density of 1 × 10⁶ cells/mL, and the medium was supplemented with thrombopoietin (TPO, 40 ng/mL, 400-34; PeproTech, Rocky Hill, New Jersey, United States). The plates were then placed in incubators with oxygen concentrations of 1, 5, and 21% (5% CO₂, 37 °C) for 5 days. MKs were separated via density gradient centrifugation, as follows. Cultured cells were collected and resuspended in 1 mL PBS. Next, 1.5 mL 3% bovine serum albumin (BSA) solution and 1.5 mL 1.5% BSA solution were successively added to the centrifuge tube at 37 °C, and then the cell suspension was slowly added along the wall of the tube. After allowing the suspension to stand for 40 minutes, white MKs were observed at the bottom of the tube. Next, 3 mL of the supernatant was discarded, and the remaining liquid was centrifuged (70 × g, 5 minutes) to collect the MKs.

Immunohistochemistry

After anaesthesia, the femurs of the rats were dissected off and cut into 0.5 × 0.3 × 0.2 cm pieces, soaked in 4% paraformaldehyde for 24 hours, and then decalcified using an ethylene-diamine-tetra-acetic acid (EDTA) decalcification solution. After decalcification, paraffin embedding and sectioning were performed. Immunohistochemical (IHC) staining was performed according to the instructions for the IHC kit (SA1020; BOSTER) using an anti-vWF antibody (PB9273; BOSTER) as the MK marker (1:1,000 dilution). The image was captured at 22 °C using an Olympus microscope CX 23 (40× objective lens) and APX100 software (Olympus, Tokyo, Japan). Images (no change in contrast or intensity) were analysed using ImageJ v1.8.0 (NIH, Bethesda, Maryland, United States).

Flow Cytometry

In flow cytometry experiments, the number of MKs and platelets per tube was maintained at 1 × 10⁶ cells. The flow cytometer BD FACS Aria III (Becton, Dickinson and Company, Franklin Lakes, New Jersey, United States) was used.

To detect the level of MK cell differentiation, MKs were suspended in 100 μL PBS, followed by the addition of 2 μL of anti-CD41 primary antibodies (PA5-79526; Invitrogen, Carlsbad, California, United States), incubation for 30 minutes, and washing twice with PBS. Next, 1 μL of APC-conjugated secondary antibodies (111-136-144; Jackson ImmunoResearch, Lancaster, Pennsylvania, United States), 2.5 μL of FITC-conjugated anti-CD61 antibodies (NB100-63958; NOVUS Biologicals, Littleton, Colorado, United States) were added to the cell suspension, incubated for 30 minutes, washed twice with PBS, and resuspended in 500 μL PBS to determine CD41 +/CD61+ cell percentages. Cells were incubated in the antibody solutions at 22 °C in the dark.

To detect the level of MK polyploidy, cells were added to 300 μL PBS, mixed, and fixed by adding 700 μL absolute ethanol (−20 °C) dropwise along the wall and then incubated overnight at 4 °C. The supernatant was removed via centrifugation (70 × g, 4 °C, 4 minutes) and cells were gently washed twice with PBS (4 °C) and resuspended in 1 mL PBS. Then, 10 μg RNaseA, 50 μL 1% Triton X-100 and 10 μL propidium iodide (PI, C0080; Solarbio, Beijing, China) were added to the cells, incubated for 40 minutes at 37 °C in the dark, and 400 μL PBS was added for detection. PI is a nuclear staining reagent that intercalates double-stranded DNA, emitting red fluorescence. The red fluorescence appears in the form of multiple peaks, each of which corresponds to the degree of polyploidy in MKs.

Apoptosis in MKs was detected following the instructions of an apoptosis kit (556547; BD Biosciences, Franklin Lakes). Briefly, MKs were added to 100 μL of 1× binding buffer, mixed with 5 μL FITC-Annexin V and 5 μL PI, and incubated for 15 minutes at 22 °C, followed by the addition of 400 μL 1× binding buffer; the MKs were then loaded within 1 hours to detect the percentages of Annexin V +/PI− MKs.

To measure the percentages of reticulated platelets, purified platelets were added to 1 mL thiazole orange (TO) diluent (final concentration, 0.04 μM, IT1180; Solarbio). The samples were incubated for 30 minutes at 22 °C, washed twice with PBS (containing 0.5% foetal bovine serum and 0.5% FBS), resuspended with 500 μL PBS (0.5% FBS), and loaded within 1 hour to detect TO+ cell percentages.

The platelets' mitochondrial membrane potential (MMP) was detected using the JC-1 fluorescence method. At high MMPs, the JC-1 forms complexes in the matrix known as J-aggregates, which produce red fluorescence (Ex/Em = 585/590 nm). Meanwhile, at low MMPs, the JC-1 dye is present in its monomeric form (monomers) and produces green fluorescence (Ex/Em = 510/527 nm). The positive rate of green fluorescence was calculated to determine the degree of MMP depolarisation. First, 500 μL of a working solution of the MMP detection kit (551302; BD Biosciences) was added to the purified platelets and incubated for 30 minutes at 37 °C in the dark. The platelets were washed twice with the 1× buffer provided with the kit and tested within 1 hour to detect the percentage of monomers.

The PS exposure of platelets was detected using the FITC-Annexin V (556547; BD Biosciences, Franklin Lakes). Platelets were mixed with 100 μL of 1× binding buffer and 5 μL FITC-Annexin V and incubated for 15 minutes at 22 °C, following which 400 μL 1× binding buffer was added; the platelets were then loaded within 1 hour to detect the percentages of Annexin V+ cells.

To measure the level of P-selectin (CD62P) on the platelet membrane surface, purified platelets were added to 100 μL PBS (0.5% FBS). The cell suspension was then incubated with 5 μL PE antihuman CD62P (304905; Biolegend, San Diego, California, United States) for 15 minutes at 22 °C in the dark. Finally, the cells were washed twice with Hepes-Tyrode buffer, resuspended in 500 μL PBS (0.5% FBS), and loaded to detect CD62P+ cell percentages.

To measure the level of GPIbα (CD42b) on the platelet membrane surface, purified platelets were mixed with 100 μL PBS (0.5% FBS). The cell suspension was then incubated with 5 μL anti-CD42b antibody (K11924-PLT; BaiAoLaiBo, Beijing, China) for 30 minutes and washed twice; 2.5 μL PE-conjugated secondary antibody (111-116-144; Jackson ImmunoResearch) was then added, and the suspension was incubated for an additional 30 minutes at 22 °C in the dark. Finally, the cells were washed twice, resuspended in 500 μL PBS (0.5% FBS) and loaded to detect CD42b+ cell percentages.

Western Blot

Radioimmunoprecipitation assay lysis buffer (100 μL, containing phenylmethylsulfonyl fluoride) was added to 1 × 10⁸ purified platelets and immediately stored at −80 °C. Proteins were quantified to ensure that the total protein load of each sample was 15 μg. Rat anti-caspase-3 antibody (31A1067; NOVUS Biologicals) was then added for the quantitative detection of caspase-3. Proteins were separated using 12% sodium dodecyl sulphate-polyacrylamide gel electrophoresis and transferred onto polyvinylidene fluoride membranes. The membranes were blocked with 5% BSA (A8020; Solarbio) diluted in 1× Tris-buffered saline containing 0.5% Tween 20 (TBST) for 1.5 hours. The primary antibodies were diluted at 1:1,000 with 5% BSA and incubated with the membrane overnight at 4 °C, after which the secondary antibody (goat anti-mouse IgG/HRP; E-AB-1001; Elabscience, Wuhan, diluted 1:1,000 with 1× TBST) was added and incubated at 22 °C for another 1.5 hours. The grey bands were analysed using the ImageJ software to calculate the relative expression of the target proteins.

ELISA

The platelet-poor plasma generated during the purification of platelets was thawed to 22 °C to determine vWF and GPIbα levels using ELISA kits according to the instructions of their respective manufacturers (Rat VWF ELISA Kit, E-EL-R1079; Elabscience) (Rat CD42b ELISA Kit, RA22720; Bioswamp, Wuhan, China).

Statistical Analysis

Normally distributed measurement data are expressed as mean ± standard deviation, whereas skewed measurement data are expressed as medians. Differences among groups were compared using independent t-tests, paired t-tests, one-way analysis of variance (ANOVA) or non-parametric tests. Statistical analyses were performed using the GraphPad Prism software (v8.0.2 (263); GraphPad Software, La Jolla, California, United States).

Results

Platelet Counts Decreased in Rats under Hypoxia

This blood cell and blood gas analyses of rats in the LA, MA, and HA groups were performed. As listed in [Table 1], the haemoglobin level of rats in the HA group was ≥210 g/L, indicating the successful establishment of a chronic hypoxia model.[29] [30] [31] With an increase in altitude, the partial pressure of oxygen in the three groups decreased, the number of red blood cells and haemoglobin levels increased, and the platelet counts decreased, confirming that chronic hypoxia can induce platelet reduction.

Table 1 Haematological profiles and blood gas analysis among the HA, MA and LA groups

	LA (n = 10) (400 m)	MA (n = 10) (2,260 m)	HA (n = 10) (5,000 m)	F/Z	p-Value
WBC (×109/L)	4.82 (4.48, 5.92)	6.13 (5.68, 6.21)	8.85 (7.33, 9.50)	22.893	<0.001
RBC (×1012/L)	5.96 ± 0.52[a] [b]	7.23 ± 0.81[a]	9.06 ± 0.33	69.140	<0.001
Hb (g/L)	121.6 ± 14.45[a] [b]	154.4 ± 13.41[a]	225.4 ± 9.78	174.289	<0.001
HCT (%)	37.46 ± 4.21[a]	39.43 ± 3.43[a]	59.05 ± 3.10	109.494	<0.001
PLT (×109/L)	1,014.5 ± 155.8[a] [b]	817.5 ± 145.8[a]	544.7 ± 84.5	31.697	<0.001
MPV (fL)	6.30 (6.10, 6.40)	6.30 (5.25, 7.50)	6.85 (6.18, 7.30)	1.846	0.392
PDW (fL)	15.66 ± 0.10[a]	15.70 ± 0.21[a]	15.96 ± 0.26	6.466	0.005
PaCO2 (mm Hg)	40.18 ± 3.09[a]	39.27 ± 2.55[a]	31.51 ± 3.99	21.286	<0.001
PaO2 (mm Hg)	78.51 ± 3.12[a] [b]	60.37 ± 2.45[a]	52.06 ± 4.66	146.656	<0.001
SaO2 (%)	93.60 ± 3.17[a] [b]	88.1 ± 2.81[a]	70.40 ± 4.20	124.117	<0.001

Abbreviations: HA, high altitude; HCT, haematocrit; LA, low altitude; MA, mid-altitude; MPV, mean platelet volume; PDW, platelet distribution width; PLT, platelet; RBC, red blood cell count; WBC, white blood cell count.

^a p < 0.05, compared with the HA group.

^b p < 0.05, compared with the MA group. Data represent the mean ± SD/quartile.

Under Chronic Hypoxia, the Number of MKs in the Bone Marrow and Lung Tissues Is Increased, Apoptosis Is Increased, and Maturation Is Restricted

Lefrançais et al have confirmed in mice that the lungs are another site of platelet genesis, which accounts for 50% of the total circulating platelets.[36] In this study, we observed the number of MKs in the bone marrow and lung tissues under chronic hypoxia using immunohistochemistry ([Fig. 1A]). The results showed that the number of MKs in both the bone marrow and lung tissue increased with altitude ([Fig. 1B, C]).

Fig. 1 Effect of hypoxia on the number, differentiation, polyploidy, and apoptosis of megakaryocytes. SD rats (160–180 g) were raised at an altitude of 400 m (LA group), 2,260 m (MA group) or 5,000 m (HA group) for 30 days. The number of megakaryocytes was detected using anti-von Willebrand factor (vWF) antibodies. (A) Representative immunohistochemistry image for vWF in the femurs and lungs of the rats in the LA, MA, and HA groups. Megakaryocytes (MKs) are marked with arrows. (B, C) Comparison of the average optical density (AOD) of MKs in the immunohistochemical images of the femurs and lungs of rats in the LA, MA and HA groups (n = 3 per group, 5 films per specimen). Bone marrow cells from the femur and tibia were isolated and cultured in incubators with 1, 5 or 21% oxygen. Thrombopoietin was then added (40 ng/mL) to induce cell differentiation for 5 days. Finally, MKs were isolated via density gradient centrifugation for flow cytometry analysis. (D) Comparison of the levels of MK differentiation among the three groups as calculated from the double positive rate of CD41 and CD61 expression (n = 3). (E) Comparison of the level of MK apoptosis by calculating the positivity of phosphatidylserine (PS) exposure (n = 5). (F) Comparison of the percentages of polyploids among three groups (n = 5). (G) Representative image of MK polyploidy. (H) Comparison of the average degree of polyploidy in MKs among the three groups (n = 5). Error bars represent the standard error.

In addition to the number of MKs, the platelet count is closely related to MK differentiation, polyploidy and apoptosis. The differentiation level of MKs in the 1% group was the lowest ([Fig. 1D]) and the apoptosis level was the highest ([Fig. 1E]), indicating that hypoxia inhibited the differentiation and promoted apoptosis of MKs. Polyploid results showed that the 1% group had the highest proportion in the low polyploid (4N, 8N, 16N) subpopulation and the lowest proportion in the high polyploid (32N and 64N) subpopulation ([Fig. 1F, G]). The average polyploid level in the 1% group was the lowest and the highest in the 21% group ([Fig. 1H]), indicating that hypoxia limited the polyploidization of MKs.

The Total Number of Reticulated Platelets Did Not Decrease under Hypoxia

Reticulated platelets are immature platelets newly released from the haematopoietic tissue into the peripheral blood, which can reflect the number of newly produced platelets.[37] [38] The cytoplasm of reticulated platelets contains a large amount of RNA, which can be stained with fluorescent nucleic acid-binding dyes and detected via flow cytometry.[39] We determined the percentages of reticulated platelets in the three groups and showed that reticulated platelet percentages in the HA and MA groups were higher than those in the LA group ([Fig. 2A, B]). Due to the decreased platelet counts in the HA and MA groups, the percentage of reticulated platelets could not accurately reflect the number of newly produced platelets. Therefore, the product of the percentage of reticulated platelets and the number of platelets were used to reflect the total number of new platelets. The results showed no statistical differences among the three groups ([Fig. 2C]), indicating that thrombogenesis is not reduced under chronic hypoxia and that platelet clearance may be the primary cause of thrombocytopenia.

Fig. 2 Effect of hypoxia on the number of platelets and reticulated platelets. (A) Three representative flow cytometry images of the proportions of reticulated platelets. Error bars represent the standard error. Comparison of platelet counts among the LA, MA and HA groups (n = 5 per group). (B) Comparison of the percentages of reticulated platelets among the LA, MA and HA groups (n = 5 per group). (C) Comparison of reticulated platelet counts among the three groups (n = 5 per group).

Hypoxia-Induced Increased Platelet Apoptosis in Rats

Apoptosis is the primary mechanism underlying platelet removal, in which mitochondrial apoptosis is the main pathway involved.[40] [41] [42] [43] We speculated that the decrease in platelet count under hypoxia might be related to apoptosis, and subsequently verified whether platelet apoptosis was increased under hypoxia. MMP depolarisation, caspase-3 activation and phosphatidylserine (PS) exposure are commonly used platelet apoptosis markers.[44] In the HA group, the platelets' PS exposure increased ([Fig. 3A, B]), the MMP decreased ([Fig. 3C, D]) and caspase-3 protein levels increased ([Fig. 3E, F]), confirming that hypoxia could lead to increased platelet apoptosis. It was determined that hypoxia-induced apoptosis was a reason for the reduction in platelet count.

Fig. 3 Effect of hypoxia on platelet apoptosis in rats. (A) Representative flow cytometry images of platelet apoptosis among the LA, MA and HA groups. (B) Comparison of apoptosis of purified platelets among three groups by measuring phosphatidylserine exposure (n = 4 per group). (C) Representative flow cytometry images of platelet mitochondrial membrane potential (MMP) depolarisation using JC-1 (JC-1 monomers reflect MMP depolarisation, and aggregates represent normal MMP) in the LA, MA and HA groups. (D) Comparison of MMP depolarisation in platelets among the LA, MA and HA groups (n = 5 per group). Caspase-3 levels were analysed via Western blot using anti-caspase-3 antibodies. (E) Representative image of caspase-3 protein expression in the LA, MA and HA groups. (F) Comparison of cleaved caspase-3 and procaspase-3 to beta actin ratios among the three groups (n = 5 per group, three replicates were performed).

GPIbα–vWF Interaction Induced Platelet Apoptosis under Chronic Hypoxia

Previous studies have shown that platelet clearance is most closely related to changes in GPIbα, including glycocalyx changes in the GPIbα extracellular segment, desialylation, GPIbα extracellular segment shedding, and GPIbα pathway activation.[45] [46] [47] [48] [49] [50] In the physiological state, the ligand of GPIbα on the platelet membrane surface is vWF. The other end of vWF can bind to collagen exposed in blood vessels through a process that binds platelets to the site of vascular injury and initiates platelet adhesion, simultaneously triggering intracellular signalling cascades that lead to further platelet aggregation and thrombosis.[51] The interaction of GPIbα with vWF can lead to platelet apoptosis[52] and is associated with platelet clearance.[53] [54] [55]

We have previously reported that the transcriptional levels of platelet GPIbα and vWF increased under prolonged hypoxia.[25] Next, we determined whether this was due to the increased interaction of GPIbα with vWF, which caused platelet apoptosis under hypoxia and measured the level of vWF in the plasma and the level of GPIbα on the platelet membrane in rats.

The results showed that, in the HA group, the plasma vWF level increased ([Fig. 4A]), while the expression level of GPIbα on the platelet membrane decreased ([Fig. 4B, C]). Studies have shown that the GPIbα on the platelet membrane surface can be shed.[56] [57] Therefore, we measured plasma levels of soluble GPIbα and found that soluble GPIbα levels were highest in the HA group ([Fig. 4D]), confirming that platelet GPIbα shedding increased under hypoxia. Therefore, if the binding of GPIbα to vWF results in the shedding of GPIbα, the decrease in GPIbα levels in the HA group may be the result of a GPIbα–vWF interaction.

Fig. 4 GPIbα–vWF interaction under hypoxia affects platelet apoptosis. (A) Comparison of plasma vWF levels among the LA, MA and HA groups as determined via ELISA (n = 6 per group). (B, C) Comparison of the percentages and mean fluorescence intensity (MFI) of GPIbα (CD42b) in the platelet membranes of the LA, MA and HA groups as obtained via flow cytometry (n = 5 per group). (D) Comparison of plasma soluble GPIbα levels among the LA, MA and HA groups as determined via ELISA (n = 6 per group). (E, F) Purified platelets in the HA group were divided into three groups: resting group (without activation), vWF + ristocetin group (incubated with ristocetin [0.5 mg/mL] and vWF [25 μg/mL] at 37 °C for 15 min), and the vWF + ristocetin + GlcNAc group (incubated with GlcNAc [100 mM, preventing GPIbα–vWF interaction] at 37 °C for 30 min to inhibit GPIb–IX–V signalling, followed by subsequent incubation with ristocetin (0.5 mg/mL) and vWF (25 μg/mL) at 37 °C for 15 min). (E) Representative flow cytometry images of GPIbα (CD42b) on platelet membranes in the above three groups, showing that the platelets could be divided into three subsets after vWF and ristocetin administration. Subset 1 showed a higher expression of GPIbα, whereas subset 3 exhibited a lower expression. (F) Comparison of the proportion of subset 2 (n = 5). (G) Representative flow cytometry images of platelet apoptosis among the above three groups. (H) Comparison of purified platelet apoptosis among the above three groups by measuring phosphatidylserine exposure (n = 5 per group). (I) Comparison of MMP depolarisation in platelets among three groups using JC-1 staining (n = 5 per group). (J) Representative flow cytometric images of platelet activation levels among the above three groups (n = 5 per group) (K). Comparison of platelet activation among the above three groups by measuring P-selectin expression. Error bars represent the standard error.

Ristocetin can promote the interaction of vWF with GPIbα on the platelet membrane.[58] Therefore, recombinant vWF protein (xy833Ra02; PeproTech) and ristocetin (AG004K; HYPHEN BioMed, Neuville, France) were coincubated with platelets in the HA group to detect the degree of platelet GPIbα shedding. The flow cytometry results showed that the level of GPIbα decreased after the interaction of vWF with GPIbα, which was divided into three subgroups according to their levels. Although subgroup 3, which represents complete GPIbα shedding, was not statistically different, subgroup 2, which represents partial shedding, was significantly increased following treatment with vWF and ristocetin ([Fig. 4E, F]). N-acetylglucosamine (GlcNAc) can inhibit the interaction of vWF with GPIbα by preventing the aggregation of GPIbα in the cell membrane.[26] [59] Platelets were incubated with GlcNAc (A8625; Sigma-Aldrich Inc., St. Louis, Missouri, United States) before incubation with vWF and ristocetin; thus, cells in subgroup 2 were significantly lower than those without GlcNAc ([Fig. 4E, F]). These results suggested that the GPIbα–vWF interaction could lead to the shedding of GPIbα. Ultimately, the increased GPIbα–vWF interaction was confirmed by the increased plasma vWF levels and decreased membrane GPIbα levels in the HA group.

Next, we determined whether the GPIbα–vWF interaction can lead to platelet apoptosis. First, in the HA group, in vitro results showed that platelet apoptosis increased after coincubation with vWF and ristocetin, which was specifically manifested as decreased MMP ([Fig. 4I]) and increased PS exposure ([Fig. 4G, H]). When GlcNAc was used to intervene in the above process, the MMP of platelets increased ([Fig. 4I]), and PS exposure decreased ([Fig. 4G, H]). GPIb–IX–V can cause platelet aggregation and participate in platelet activation. We found that enhancing the GPIbα–vWF interaction led to an increase in the level of platelet activation, and inhibiting the interaction alleviated the level of platelet activation ([Fig. 4J, K]). GPIbα–vWF interaction may increase the risk of thrombosis and decrease the number of platelets.

We further verified whether the GPIbα–vWF interaction is involved in the reduction of platelet count using in vivo assays. Ten SD rats in the HA group were randomly divided into control and GlcNAc groups (n = 5). On day 30 of feeding, blood was collected from the caudal vein to measure the platelet counts, and then rats in both groups were injected with 400 μL PBS or 400 μL GlcNAc through the caudal vein, respectively. GlcNAc was diluted in PBS to an injection dose of 110 μg/g.[26] The change in platelet counts in the following 5 days (the life of platelets was 3–5 days) was observed. On day 32, platelet PS exposure and MMP were detected to understand the level of platelet apoptosis.

The results showed that platelet counts increased after GlcNAc injection and peaked on the second day ([Fig. 5A]). The results showed that platelet PS exposure and MMP levels decreased after GlcNAc intervention, suggesting that inhibition of GPIbα–vWF interaction could partially alleviate platelet apoptosis caused by chronic hypoxia ([Fig. 5B, C]).

Fig. 5 Effect of GPIbα intervention on platelet counts in rats under hypoxia in vivo. Ten SD rats in the HA group were randomly divided into control and GlcNAc groups (n = 5). On day 30 of feeding, blood was collected from the caudal vein to measure the platelet counts, and then rats in both groups were injected with 400 μL PBS or 400 μL GlcNAc through the caudal vein, respectively. GlcNAc was diluted in PBS to an injection dose of 110 μg/g. (A) Changes in the platelet count before, and 5 days after, intervention were observed (n = 5). (B) Comparison of apoptosis of purified platelets among two groups by measuring phosphatidylserine exposure (n = 5 per group). (C) Comparison of apoptosis of purified platelets among two groups by measuring MMP depolarisation (n = 5 per group). Error bars represent the standard error. *p < 0.05; **p < 0.01; *****p < 0.0001.

Discussion

Chronic hypoxia can increase haemoglobin count and blood viscosity and levels of platelet activation and procoagulant factors, increasing the risk of thrombosis.[11] [12] [13] [16] [18] [25] In addition to the increased risk of blood clots, chronic hypoxia can decrease platelet count.[60] Clinicians should consider the risk of bleeding due to low platelet count when treating thrombosis. Therefore, it is important to clarify the causes of platelet reduction during chronic hypoxia.

Previous studies have speculated that, as chronic hypoxia leads to increased erythropoietin (EPO) levels, erythrocyte-MK progenitor cells undergo an erythroid transformation, resulting in a reduction in MK progenitor cell generation and, ultimately, a decrease in platelet count.[61] [62] However, Birks et al showed that EPO can lead to an increase in red blood cell count without affecting platelet numbers, suggesting that the proliferation of the erythroid lineage does not influence the expression of the MK lineage.[63] Our previous study found that the number of MK increased in patients with high-altitude polycythaemia under chronic hypoxia,[5] while this study demonstrated that the rats under chronic hypoxia also showed the same performance, which did not support the view that the thrombocytopenia was caused by the competitive inhibition of MK caused by erythroid hyperplasia. The heterogeneity among these studies may arise from differences in the time points and methodologies used to observe the MK lineage.

Furthermore, we found that the apoptosis level of MKs increased under chronic hypoxia. Apoptosis is a programmed cell death process for most cells, but for MKs, it is a sign of their maturity because the apoptosis level is higher in mature cells.[64] [65] There is a temporal correlation between MK apoptosis and platelet release; hence, MK apoptosis may play a role in thrombopoiesis.[66] An increase in the MK number and apoptosis is favourable for platelet production. Conversely, under chronic hypoxia, differentiation and polyploidy are inhibited, which is not conducive to platelet production. Therefore, the impacts of chronic hypoxia on platelet formation are multifaceted. By measuring the number of reticulated platelets, we found no difference in the number of newly produced platelets compared with that under normoxia, and the main cause of the platelet reduction may be platelet clearance.

The level of platelet apoptosis was increased in rats under chronic hypoxia, which was verified by measuring PS exposure, MMP, and caspase-3 protein expression. Platelet clearance is closely associated with GPIbα. The GPIbα–vWF interaction can lead to apoptosis or increased platelet clearance. In this study, hypoxia-promoting GPIbα–vWF interaction could lead to platelet apoptosis, and inhibiting this interaction could increase platelet counts. During physiological processes, the combination of GPIbα and vWF mainly causes platelet adhesion and aggregation, which can promote thrombosis.[67] [68] In other words, thrombosis and a reduced platelet count are closely related under chronic hypoxia. The platelet-based pathway in immune-mediated diseases can induce thrombocytopenia and is associated with thrombotic events.[69] Our results also showed that GPIbα–vWF interaction could not only induce platelet activation but also induce platelet apoptosis under chronic high-altitude hypoxia, leading to platelet reduction. Hence, for patients with high-altitude thrombosis, treatment interfering with the interaction of GPIbα and vWF can inhibit platelet adhesion and maintain platelet count by inhibiting platelet apoptosis, which may also be a beneficial target for antithrombotic therapy in patients with thrombosis and thrombocytopenia.

Hypoxia, inflammation, and thrombocytopenia are common in many diseases, such as in patients with myocardial infarct, severe COVID-19 infection, and sepsis.[8] [70] [71] Hypoxia can increase platelet reactivity and activation levels, which promote inflammation.[8] [21] [25] [72] The degree of inflammation is closely associated with the prognosis of diseases.[73] The impact of hypoxia on platelet function remains largely unresolved and still controversial. Several studies have shown that following hypoxia, the response of platelet aggregation is significantly diminished, along with a reduction in platelet aggregation and adhesion functions, as well as a decrease in platelet–monocyte complexes.[60] [74] [75] The reasons for these discrepancies may be attributed to differences in the duration of hypoxia, the distinction between pathological and physiological hypoxia, as well as variations in the technical methods used to assess platelet activity. In our study, we found that GPIbα–vWF interaction under hypoxia promoted platelet apoptosis, activation, and reduction. Previous studies have also confirmed that platelet activation is involved in thrombosis and inflammation. In clinical practice, several diseases are characterized by hypoxia and inflammation, such as chronic obstructive pulmonary disease, pulmonary hypertension, and bronchial asthma. These diseases are often accompanied by decreased platelets.[76] [77] [78] Whether the GPIbα–vWF interaction plays a role in disease onset and development disease remains to be determined.

In conclusion, this study showed that the number of MKs increased in rats under chronic hypoxia, with no effect on MK differentiation or thrombopoiesis. Platelet apoptosis was attributed to the decrease in platelet count and was closely associated with the GPIbα–vWF interaction. These two factors are involved in thrombosis formation and platelet reduction; therefore, GPIbα may be a therapeutic target for patients with chronic high-altitude thrombosis. The regulation of the GPIbα pathway has two beneficial outcomes (i.e., inhibiting thrombosis and improving platelet counts), thereby providing a feasible alternative for the clinical diagnosis and treatment of these conditions.

Limitations

The effect of chronic hypoxia on platelet production was indirectly demonstrated using the proportion of reticulocyte platelets, while new platelets were not directly observed due to the experimental conditions.

Conflict of Interest

All support for the present manuscript (e.g., funding, provision of study materials, medical writing, article processing charges, etc.): Sichuan Science and Technology Program (2023YFQ0068); The General Hospital of Western Theater Command (2021-XZYG-C30).

Acknowledgements

We thank all the members of the Tana Wuren team for their assistance. We would also like to thank Editage (www.editage.cn) for English-language editing.

Data Availability

The datasets generated for this study can be found in the figshare database (https://doi.org/10.6084/m9.figshare.23993115).

Authors' Contributions

T.W. and Z.W. conceived and designed the research; Z.W. and D.G. performed the experiments and analysed the data; Z.W. and Y.Y. prepared the figures; Y.Y. assisted in the experiment; Z.W. and D.G. drafted and revised the manuscript; and Y.W. and T.W. approved the final version of the manuscript. Z.W. and D.G. contributed equally to this work and should be considered co-first authors.

^* These authors contributed equally to this work and should be considered co-first authors.

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Eingereicht: 13. Juni 2024

Angenommen: 05. November 2024

Artikel online veröffentlicht:
28. März 2025