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. Author manuscript; available in PMC: 2021 May 1.
Published in final edited form as: Alcohol Clin Exp Res. 2020 Apr 21;44(5):1088–1098. doi: 10.1111/acer.14328

Daily Ethanol Drinking Followed by an Abstinence Period Impairs Bone Marrow Niche and Mitochondrial Function of Hematopoietic Stem/Progenitor Cells in Rhesus Macaques

Oleg Varlamov 1,*, Matthew Bucher 2, Leslie Myatt 2, Natali Newman 3, Kathleen A Grant 3,*
PMCID: PMC7430207  NIHMSID: NIHMS1584160  PMID: 32220015

Abstract

Background:

Unhealthy consumption of alcohol is a major public health crisis with strong associations with immunological dysfunctions, high vulnerability to infectious disease, anemia, and an increase in the risk of hematological malignancies. However, there is a lack of studies addressing alcohol induced changes in bone marrow and hematopoiesis as fundamental aspects of immune system function.

Methods:

To address the effect of chronic alcohol consumption on hematopoietic stem and progenitor cells (HSPCs) and the bone marrow niche, we used an established rhesus macaque model of voluntary alcohol drinking. A cohort of young adult, male rhesus macaques underwent a standard ethanol self-administration protocol that allowed a choice of drinking alcohol or water 22 hours/day with periods of forced abstinence that elevated subsequent intakes when alcohol availability resumed. Following the last month of forced abstinence, the monkeys were euthanized. HSPCs and bone samples were collected and analyzed in functional assays and by confocal microscopy.

Results:

HSPCs from alcohol animals exhibited reduced ability to proliferate and differentiate into granulocyte-monocyte and erythroid cells. HSPCs also displayed a decrease in mitochondrial oxygen consumption linked to ATP production and basal respiratory capacity. Chronic alcohol use led to vascular remodeling of the bone marrow niche, a reduction in the number of primitive HSPCs, and a shift in localization of HSPCs from an adipose to a perivascular niche.

Conclusions:

Our study demonstrates, for the first time, that chronic voluntary alcohol drinking in rhesus macaque monkeys leads to the long-term impairment of HSPC function, a reduction in mitochondrial respiratory activity, and alterations in the bone marrow microenvironment. Further studies are needed to determine whether these changes in hematopoiesis are persistent or adaptive during the abstinent period and whether an initial imprinting to alcohol primes bone marrow to become more vulnerable to future exposure to alcohol.

Keywords: alcohol, bone marrow, bone marrow adipose tissue, hematopoiesis, hematopoietic stem and progenitor cells, nonhuman primate

Introduction

Unhealthy alcohol (ethanol) consumption is a major public health crisis (Poznyak et al., 2013) and a leading risk factor for global disease burden (Collaborators, 2018), including mortality from cancer, diabetes, liver disease, infections, and bone marrow (BM) failure (Di Rocco et al., 2019). Ethanol and its byproducts are detrimental to cellular function and metabolism, as demonstrated in different tissues and organs, including, but not limited, to the liver (Ivester et al., 2007, French, 2016, Khambu et al., 2017), skeletal (Duplanty et al., 2017) and cardiac muscle (Varga et al., 2015), alveolar macrophages (Liang et al., 2013), and the brain (Perez et al., 2018, Chen et al., 2016). Furthermore, an alcohol-induced increase in reactive oxygen species (ROS) levels has been shown to alter mitochondrial function in peripheral tissues (Ivester et al., 2007, French, 2016, Duplanty et al., 2017, Liang et al., 2013, Khambu et al., 2017).

Clinical studies show that chronic alcohol users present multiple hematological abnormalities, including altered granulopoiesis and anemia (Shi et al., 2019, Latvala et al., 2004, Yeung et al., 1988). Defects in granulocytes are associated with a high susceptibility of alcoholics to infections (Shi et al., 2019). While the detrimental effects of alcohol and its byproducts on hematopoiesis is generally accepted, only few studies to date have addressed the effect of alcohol on hematopoietic stem and progenitor cells (HSPCs). HSPCs exist in a quiescent state but can proliferate and differentiate in mature blood cells in response to injury and infection (Wilson et al., 2008, Essers et al., 2009, Baldridge et al., 2010). The localization and proliferation state of HSPCs in the BM can change in response to stress (Crane et al., 2017), irradiation-induced damage (Lo Celso et al., 2009, Xie et al., 2009), obesity (Adler et al., 2014), and aging (Mohrin and Chen, 2016). Thus, alcohol-mediate impairment of hematopoiesis may lead to a compromised response to infection and other forms of hematopoietic stress. However, the effects of chronic alcohol consumption on the functional properties and localization of HSPCs in the BM remains largely unknown.

Studies using rodent models of alcohol exposure have shown that ethanol and/or its metabolite acetaldehyde and other ethanol-generated ROS can impair HSPC function and cause gene mutations (Garaycoechea et al., 2012, Pontel et al., 2015, Garaycoechea et al., 2018). However, the effects of alcohol on human HSPCs are unknown due to difficulties in obtaining BM from alcoholic subjects. Furthermore, no studies to date have addressed the effects of chronic alcohol use on BM microenvironment and vascular endothelium that provides HSPCs with local paracrine signals essential for their maintenance and differentiation (Pinho and Frenette, 2019, Calvi and Link, 2015, Crane et al., 2017).

To bridge the gap between mouse and human studies, we took advantage of a robust preclinical nonhuman primate (NHP; rhesus macaque) model for studying the effects of daily alcohol consumption on HSPCs at a timescale similar to the developmental trajectories observed in human chronic drinkers. Furthermore, the hematopoietic system and the immunophenotypic identity of HSPCs of NHPs shares significant similarity with that of humans (Kim et al., 2014, Radtke et al., 2017, Wu et al., 2018). Previous studies with this NHP model have characterized the effects of chronic alcohol use on the peripheral immune system (Ivester et al., 2007, Allen et al., 2018, Sureshchandra et al., 2019, Balbo et al., 2016, Baker et al., 2014, Sureshchandra et al., 2016, Barr et al., 2016), however the effects of alcohol on BM microenvironment and HSPCs remain mainly unknown. We have recently demonstrated that in healthy adult male rhesus macaques, CD34+ HSPCs reside in close proximity to BM adipocytes, while only a minor fraction of CD34+ cells was associated with the CD3+/CD34+ BM endothelium (Robino et al., 2020). Here, we report, for the first time, that chronic daily alcohol drinking followed by an extended abstinence results in the long-term impairment of HSPC function, decreased mitochondrial respiratory capacity in HSPCs, and remodeling of the vascular BM niche in rhesus macaques, suggesting that this monkey model can address mechanisms of alcohol-induced alterations in hematopoiesis.

Materials and methods

Animal characteristics and study design.

Twelve experimentally naïve young adult male rhesus macaques (Macaca mullatta; 4.0–5.5 years at assignment) were assigned from a pedigreed population at the Oregon National Primate Research Center (Beaverton, OR). Housing and environmental descriptions along with detailed protocols can be found in (Allen et al., 2018). All monkeys were induced to drink a volume of ethanol or water using a schedule-induced polydipsia (SIP) procedure, equivalent to 0.5, 1.0, and 1.5 g/kg ethanol, for at least 30 sessions as described (Grant et al., 2008, Baker et al., 2014). After 183 consecutive sessions, induction conditions were discontinued and an “open-access” daily schedule of both ethanol and water were available for 22 h/day, with three meals at 0 (session onset), 2, and 4 h into the session (Baker et al., 2014). Open-access conditions continued 7 days/week for 14 consecutive months and then a repeated abstinence protocol began with no alcohol availability for 28 days and approximately 3 months of ethanol self-administration between the first two abstinence periods. The protocol ended following the third and final 28–35 days of forced abstinence (Figure 1A and B). The duration of the final abstinence reflects the throughput of necropsy schedule with 2 animals/day.

Figure 1. Study design and procedures.

Figure 1.

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A) A cohort of twelve young adult male rhesus macaques underwent a schedule-induced polydipsia procedure to establish ethanol (or water for controls) self-administration, and then the ethanol drinkers were given open access ethanol and water for 12 months, followed by repeated forced abstinence periods and 3-month relapse ethanol open-access periods. The protocol ended following the last month of forced abstinence. Femurs and BM were collected at the end of the study. Additional procedures are indicated in the legend. B) 12-month average blood ethanol concentrations in alcohol drinkers. C) Animals were subjected to several experimental procedure. Proximal femurs were fixed and analyzed by immunofluorescent confocal microscopy. BM collected from the distal and medial parts of the femur was used for isolation of CD34+ HSPCs. HSPC differentiation efficiency was measured using a colony-forming unit (CFU) assay. HSPCs were immunophenotyped by flow cytometry and analyzed by in vitro respirometry, using the Seahorse mitochondrial stress tests.

At necropsy, the left femurs were collected and cross-sectioned into two longitudinal portions. The proximal part (metaphysis) was fixed and processed for immunostaining to assess the structural organization of the BM niche by multiplexed confocal microscopy (Figure 1C). The middle portion of the femur (diaphysis) and the distal metaphysis were used for isolation of the BM mononuclear cells and antibody-based purification of CD34+ HSPCs. This experimental design allowed us to simultaneously assess the architecture of the BM niche and characterize the phenotypic and functional properties of HSPCs, including their colony-forming ability and mitochondrial respiration in vitro (Figure 1C). Body composition was determined by Dual-energy X-ray absorptiometry (DEXA), as previously described (Cameron et al., 2016).

Isolation and characterization of CD34+ cells.

BM was collected at necropsy, placed in 25 ml of ice-cold X-Vivo™ 10 media (Lonza, Basel, Switzerland) and processed within 1 hour. BM was gently disrupted using a 25-ml syringe and a blunt needle and cell suspension filtered through a 100-μm cell strainer (Life Sciences, Durham, NC). The cell suspension was layered on a Ficoll density gradient followed by centrifugation for 30 min at 1500 rpm at room temperature. Mononuclear cells were collected from the Ficoll gradient interface and HSPCs cells were isolated using rhesus-specific antibodies to CD34 (BD Pharmingen, clone 563) and the Miltenyi Biotec (Bergisch Gladbach, Germany) anti-PE MACS magnetic bead system using supplied buffers and reagents. Specifically, 1–2×108 BM mononuclear cells derived from a single femur were resuspended in 1 ml wash buffer and mixed with 200 μl FcR blocking reagent (Miltenyi Biotec) and 100 μl PE-conjugated antibodies to CD34 (BD Pharmingen, clone 563). Cells were incubated on a rocking platform for 20 min at 12–15°C, washed by centrifugation at 1000 rpm twice with 20 ml wash buffer, resuspended in 0.5–1 ml wash buffer supplied with 200 μl anti-PE MicroBeads (Miltenyi Biotec) and incubated for an additional 15 min. Cells were washed with 20 ml of wash buffer, resuspended in 1 ml wash buffer and purified on the MS columns (Miltenyi Biotec) according to the manufacturer’s instructions. To increase cell viability, CD34+ cells were eluted in 1 ml StemSpan SFEMII media (Stem Cells Technologies, Vancouver BC, Canada). We typically obtain ~1–2×106 CD34+ cells from 2×108 BM mononuclear cells derived from a single emur. 2000 cells were plated onto MethoCult™ (Stem Cells Technologies, H4435 Enriched) supplemented with supplied with human cytokines (100 ng/ml stem cell factor (SCF), thrombopoietin (TPO) and Fms-related tyrosine kinase 3 (Flt3); (Stem Cell Technologies) (Radtke et al., 2017), and the growth of hematopoietic colonies was quantified on day 10.

CD34+ cells were cryopreserved using CryoStor CS10 (Stem Cells Technologies). For flow cytometry analysis, frozen CD34+ cells were thawed, washed twice in PBS, and stained with the viability dye Zombie Aqua (BioLegend, San Diego, CA). Samples were then blocked for 45 min in PBS containing 5% each human and mouse serum, sodium azide and EDTA (FACS blocking buffer) at 4°C. Samples were stained in FACS blocking buffer using anti-CD34-PE (BD Pharmingen, clone 563), anti-CD38-APC (Caprico Biotechnologies, clone OKT10), anti-CD90-PECy7 (BD Pharmingen, clone 5E10), anti-CD45RA-APC-H7 (BD Pharmingen, clone 5H9), anti-CD49f-PECy5 (BD Pharmingen, clone GoH3), anti-Ki67-FITC (BD Pharmingen) for an additional 45 min at 4°C. Finally, samples were fixed in 2% neutral buffered formalin, washed in FACS buffer and analyzed using an LSRII flow cytometer. Data were acquired using BD FACS Diva (BD Biosciences) and post-analysis performed using FlowJo v10 (Tree Star, Ashland, OR) and Prism v6 (GraphPad Software, San Diego, CA).

Preparation of bone sections, immunohistochemistry, and optical clearing

Cross-sections of the left femur were cut at the boundary between the proximal metaphysis and the diaphysis (approximately one inch from the femoral head). The proximal metaphysis was cleared of soft tissue and fixed for 48 hours at 4°C in 4% paraformaldehyde/PBS. Following fixation, the tubular segment of the proximal metaphysis was sectioned into several 2–3 mm adjacent slices using the diamond-coated saw (Mar-med, model 80 mini-band bone saw). Each bone section was washed sequentially (4 ml, polypropylene tubes, 20 minutes each at room temperature) with the following solutions: PBS, 50% methanol/PBS, 80% methanol/DI water, 100% methanol, 20% DMSO/methanol twice, 80% methanol/DI water, 50% methanol/PBS, PBS twice, PBS/1% Triton X-100 twice before further staining procedures. Immunohistochemistry procedures were conducted at 40°C on a rocking platform. Sections were blocked in 0.5 ml 10% donkey serum, 0.25% Triton-X100/PBS for 4 hours and incubated in the following primary antibodies: (1:100 dilution) in 0.5 ml 10% donkey serum, 0.25% Triton X-100/PBS for 24 hours; mouse anti-human CD34 (BD Pharmingen, San Jose, CA, clone 581) and sheep anti-human CD31 (R&D Systems, Minneapolis, MN, clone AF806). Sections were washed 6 times x 15 minutes with 0.25% Triton X-100/PBS, incubated with the fluorescent secondary antibodies (donkey) (Invitrogen, 1:1000 dilution; 1:1000 dilution) for an additional 24 hours, then washed extensively with 0.25% Triton-X 100/PBS. For tissue clearing, stained bone sections were washed with following solutions (20 min per wash): 10 ml 50% methanol /PBS, 10 ml 70% methanol/PBS; 10 ml 100% methanol. Excess methanol was absorbed with a paper towel and the samples were incubated in 2 ml Visikol HISTO-1 at room temperature for 2 hours followed by 2 ml Visikol HISTO-2 (Visikol, Inc., Whitehouse Station, NJ) for additional 2 hours.

Image acquisition and analysis

Bone sections were transferred into a 35-mm glass-bottom culture dishes (MatTek, Ashland, MA) containing 300 μl of Visikol HISTO-2. Confocal microscopy was performed using a Leica SP5 AOBS spectral confocal system. Images were collected in a sequential mode, at 2-μm, using a ×20 PL APO NA 0.70 dry objectives. For each bone section, 3–4 random fields of z-stacks were analyzed. Z-stacks of confocal images were opened with the LOCI plug-in Fiji data browser and analyzed as follows. HSPC cell boundaries were detected using the CD34 channel. The image representing a single confocal slice was processed using median filter (rolling ball value, 2.0) and subjected to background subtraction. Individual CD34+ cells above threshold were detected using the “Analyze Particles” function. The CD34+/CD31+ endothelium vessels were traced manually and the total length of endothelium per area of the BM was determined. Megakaryocytes were identified as large multinucleated CD31+ cells.

Respirometry measurements

Mitochondrial function was assessed using the Seahorse XF Cell Mitochondrial Stress Test (Agilent Technologies, Santa Clara, CA) which measures oxygen consumption rates (OCR) using an XFe96 analyzer. Basal OCR, ATP linked OCR, maximal OCR, spare capacity, proton leak, and non-mitochondrial OCR were determined by sequential injection of Oligomycin, Carbonyl cyanide-4 (trifluoromethoxy) phenylhydrazone (FCCP), and a combination of Rotenone and Antimycin. Before inhibitors were injected, basal OCR was measured to determine baseline mitochondrial respiration. The complex V (ATP synthase) inhibitor, Oligomycin, was injected first and is used to determine OCR that is linked to ATP production. The second injection, FCCP uncouples the mitochondrial electron transport chain by collapsing the proton gradient and disrupting mitochondrial membrane potential yielding maximal OCR. Spare capacity is the mitochondria’s ability to increase ATP production to overcome stress or increased energy demand and is calculated as the difference between maximal and basal OCR. The final injection of Rotenone and Antimycin inhibits complex one and complex three of the ETC chain, respectively and completely shuts down mitochondrial respiration. Proton leak is defined as the mitochondria’s ability to respire in the presence of the ATP synthase inhibitor Oligomycin and is calculated as the difference between OCR after the Oligomycin injection and OCR after Rotenone/Animycin injection.

0.5–2 million cryopreserved CD34+ HSPCs were thawed at 37°C, washed by centrifugation twice with pre-warmed X-Vivo™ 10 media (Lonza, Basel, Switzerland), resuspended in 1 ml StemSpan SFEMII media supplemented with 100 ng/ml SCF, TPO and Flt3, and allowed to recover overnight at 37°C. Approximately 24 hrs prior to the mitochondrial respiration assay, a seahorse sensor cartridge was hydrated with seahorse calibrant solution (Agilent) and incubated in a 37°C non-CO2 incubator overnight. The following morning cells were plated at 100,000 cells/well in a 96 well seahorse culture plate coated with RetroNectin, according to the manufacturer’s instruction (Takara Shuzo Co., LTD, Kyoto, Japan). Cells were allowed to adhere for 2 hours. After cells were adherent, media were exchanged with seahorse mito-stress basal media (Agilent) supplemented with 25 mM glucose, 4 mM glutamine, and 1 mM pyruvate, and placed in a 37°C non-CO2 incubator for 1 hr. After a 1-hour incubation, the mito-stress test assay was performed according to the manufacturer’s protocol. The mitochondrial ETC drug inhibitor concentrations: Oligomycin (1 μM), FCCP (1.5 μM), and Rotenone/Antimycin (1 μM) were optimal for HSPCs. After the assay was complete DNA content was measured in each well to allow normalization of OCR. Hoechst-33342 was added to each well at a 1:1000 dilution and incubated for 30 minutes at room temperature with constant shaking. Fluorescence was measured (excitation 350nm/emission 461nm) using a plate reader. Comparisons between controls and drinkers were performed using the Student’s t-test in Excel. P < 0.05 was considered as statistically significant.

Results

Chronic Alcohol consumption impairs HSPC function.

To address the functional effects of alcohol on HSPC differentiation, purified CD34+ cells were plated in cytokine-containing methylcellulose (human cytokines, SCF, TPO, and Flt3L (Radtke et al., 2017)) for 10 days and the growth of the colony-forming units were quantified. CD34+ cells derived from alcohol drinkers produced a significantly lower number of granulocyte-monocyte (GM) and erythroid (E) colonies compared to control animals (Figure 2A and B). These results suggest that alcohol impairs HSPC differentiation, resulting in inefficient monogranulopoiesis and erythropoiesis in vitro. Furthermore, the analysis of peripheral blood revealed that alcohol consumption is associated with an increase in red blood cell distribution width (RDW), suggesting the development of macrocytosis (the enlargement of red blood cells) and a significant reduction in mean corpuscular hemoglobin (MCH) content (Figure 2C and D; Supplementary Table S1). Flow cytometry analysis of isolated CD34+ HSPCs showed that a percent of CD34+CD45RA-CD90+CD49f+ HSPCs was significantly decreased in the BM of alcohol animals compared to controls (Figure 2E and F). These studies demonstrate a long lasting decrease in the number of hematopoietic progenitors and the erythro-myeloid differentiation of HSPCs following chronic alcohol consumption in this NHP model.

Figure 2. Chronic alcohol use alters myelo-erythropoiesis.

Figure 2.

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A) Colony-forming unit (CFU) assay; representative examples of granulocyte-monocyte (GM), common myeloid progenitor (GEMM), and erythroid (E) colonies recorded on day 10; BFU, burst-forming unit-erythroid colony. B) 2000 CD34+ HSPCs were plated onto MethoCult™ in triplicates supplemented with 100 ng/ml SCF, TPO, and Flt3L, and the growth of hematopoietic colonies was quantified. C-D) Changes in the properties of red blood cells; bars are means ± SEM, n=3 controls and 9 alcohol drinkers. T-test *p<0.05. E) Flow cytometry gating strategy for phenotypic characterization of HSPC populations. F) Alcohol-induced changes in fraction of CD34+CD38+CD45RA-CD90+CD49f+ HSPCs determined by flow cytometry; 6 controls (additional controls were added) and 9 alcohol drinkers; T-test *p<0.05.

Chronic alcohol consumption impairs the respiratory capacity of CD34+ cells.

To test the hypothesis that chronic alcohol intake alters mitochondrial function, CD34+ cells isolated from the BM of control and alcohol drinkers were subjected to the mitochondrial stress test using in vitro Seahorse respirometry. The results of this analysis indicate that alcohol consumption resulted in mitochondrial dysfunction in CD34+ cells. This is evidenced by significantly reduced basal OCR (p=0.015) and OCR linked to ATP production (p=0.014) (Figure 3A and B). Importantly, there was an inverse correlation between the average blood ethanol levels during the first 12 months of drinking and basal OCR (R2=0.38; p=0.032) and ATP production rates (R2=0.43; p=0.021), suggesting that ethanol impairs mitochondrial respiratory function in a dose-dependent manner in vivo (Figure 3C and D).

Figure 3. Chronic alcohol use alters mitochondrial respiration in isolated CD34+ cells.

Figure 3.

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A) Real-time O2 consumption rates (OCR) in CD34+ cells isolated from controls and alcohol drinkers were measured by Seahorse under basal conditions as well as in response to the mitochondrial electron transport chain inhibitors Oligomycin, FCCP, and Rotenone/Antimycin, as described in “Materials and Methods.” For each animal, each assay was performed in triplicates, using 100,000 CD34+ cells per well. B) Bar graph are means± SEM of indicated parameters for n=3 controls and n=9 drinkers. T-test *p<0.05. C-D) Correlations between basal OCR, OCR coupled to ATP production, and a 12-month average ethanol concentrations.

Chronic alcohol consumption changes the composition of the BM niche.

To elucidate the effects of chronic ethanol use and then abstinence on the BM, femurs of control and alcohol-drinking animals were studied by multicolor confocal microscopy. The proximal metaphysis of the femurs were sectioned into adjacent circular segments and subjected to indirect immunofluorescence analysis (Figure 4A–D). To reduce the light-scattering effect due to the presence of the lipid-rich BM adipocytes, the bone slices containing the transverse segments of the BM were optically cleared with Visikol. To visualize vascular endothelium, required for the support and regeneration of HSPCs in both mice and humans (Sasine et al., 2017), the BM was stained with antibody to CD31. The quantitative analysis revealed prominent changes in the vascular organization of the BM in response to alcohol drinking, as evidenced by a significant increase in the length of CD31+/CD34+ arterial endothelial capillaries observed in the BM of alcohol drinkers compared to controls (Figure 4E). Furthermore, the CD31+/CD34+ endothelium in the BM of alcohol drinkers formed the capillary networks characterized by multiple branching points (Figure 4C and D). The relative expression of CD31 and CD34 antigens varied significantly between different branches of the CD31+/CD34+ endothelium (Figure 4D). The total length of CD31+/CD34+ endothelium correlated positively with the average alcohol intake (R2=0.45; p=0.034) (Figure 4F). Furthermore, the densities of megakaryocytes and CD34+ HSPCs per area of the BM were reduced in alcohol drinkers compared to controls (Figure 4G and H). In the BM of control animals, CD34+ HSPCs were found next to adipocytes (Figure 4A, right inset arrow). In contrast, in the BM of alcohol drinkers, HSPCs were associated with the CD31+/CD34+ endothelium (Figure 4B, right inset, arrows). Collectively, the analysis of the BM architecture suggests long-lasting effects of chronic ethanol drinking, even after 28–35 days of alcohol abstinence on the cellularity of the BM compartment, leading to vascular remodeling of the BM niche and the shift in localization of HSPCs from an adipose to a perivascular niche.

Figure 4. Chronic alcohol use induces bone marrow vascularity.

Figure 4.

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A-E) BM microscopy of proximal femoral cross-sections from the representative control (A) and the alcohol drinkers (B-D). The region of interest in (C) is shown enlarged in (D). A and B) Lower left insets show unstained bone sections demarcating the cortical bone (outer ring), BM adipose tissue (MAT, arrows), and red marrow (brown); HSPC, CD34+ cells; CS, central sinus. A) Right inset, two HSPCs (arrow) adjacent to the BM adipocyte (a), bar=10 μm. B) Right inset, HSPCs (arrows) adjacent to the CD34+/CD31+ capillaries; m, megakaryocyte; bar=10 μm. E) CD31+/CD34+ capillary length per BM area. F) Correlation between average alcohol intakes (grams of ethanol per kg body weight) and CD31+/CD34+ endothelial vessel length. G) Megakaryocyte and (H) HSPC density in the BM of controls (n=3) and alcohol drinkers (n=9). Bar graphs are means± SEM; T-test **p<0.01; #p<0.1. For each animal, the average BM parameters were determined using 3–4 confocal slices.

Discussion

This study provides novel data on the effect of chronic heavy alcohol consumption followed by a one-month abstinence period on reduced functionality of primate HSPCs. In human populations self-report of drinking levels and malnutrition associated with alcoholism are confounding factors in attributing ethanol dose and duration with the loss of HSPC’s ability to efficiently mature. In the present study, the monkey model of voluntary ethanol drinking can explicitly address the effects of ingested alcohol on the functional properties of HSPCs. Using this animal model we found that chronic ethanol consumption led to functional impartments in HSPCs and the structural alterations in BM niche that persisted after one month of ethanol abstinence.

The main finding of the present study is that chronic ethanol drinking leads to a reduction in the number of primitive HSPCs in the BM of rhesus macaques. This finding is consistent with an observed function decline in HSPCs of alcohol drinkers, as evidenced by reduced HSPC’s ability to form granulocyte-monocyte and erythroid colonies in vitro. Importantly, an apparent defect in erythropoiesis correlated with the development of macrocytosis, which is commonly observed in the blood of alcohol drinkers (Yokoyama et al., 2014, Latvala et al., 2004), suggesting that erythroid progenitors are particularly sensitive to the effects of chronic alcohol consumption. Indeed, cytoplasmic vacuolization of the erythroid and myeloid progenitors is commonly observed in the BM of patients with acute alcoholism (Yeung et al., 1988). Furthermore, alcoholics frequently have defective erythroid cells that are destroyed prematurely, resulting in the development of anemia (Ballard, 1997) and exhibit diverse patterns of hematological abnormalities, including cytopenia and vacuolization of BM megakaryocytes, especially in individuals with recent intoxication (Latvala et al., 2004).

Excessive alcohol consumption is also associated with impaired granulopoiesis, resulting in the compromised immune defense function, high susceptibility to infectious diseases and sepsis (Shi et al., 2019). These clinical observations are consistent with the present study, suggesting that alcohol-induced impaired granulo-erythropoiesis persists even after the abstinence period. Previous studies demonstrated that primitive HSPCs are particularly sensitive to exogenous stress, resulting in DNA damage response (Mohrin et al., 2010) and selective apoptosis of damaged HSPCs (Yamashita et al., 2016). Collectively, these mechanism may contribute to alcohol-induced functional decline in HSPCs that persists even after the termination of alcohol drinking. The latter suggests that exposure to alcohol induces long-lasting alterations in the nuclear or mitochondrial genome of HSPCs and/or damages the BM niche cells involved in HSPC maintenance, which might predispose alcohol drinkers to become more vulnerable to a future hit of alcohol exposure. Thus, even abstinent drinkers might be at risk when they drink in the future, though this is yet to be elucidated.

The present study demonstrates, for the first time, that alcohol consumption exerts a negative effect on mitochondrial function in HSPCs. Mechanistically, it has been shown that a short exposure to alcohol and its metabolites acetaldehyde and ROS exerts highly genotoxic effects on HSPCs in the transgenic models of aldehyde dehydrogenase-2 (Aldh2(−/−)) and Fanconi anemia-2 (Fancd2(−/−)) deficiencies of the DNA repair pathways (Pontel et al., 2015, Garaycoechea et al., 2018, Garaycoechea et al., 2012). Consistent with these reports, the present study shows that chronic alcohol use induces persistent alterations in mitochondrial respiratory function and ATP production, implying that alcohol may induce long-lasting genomic and/or mitochondrial DNA damage, leading to hematopoietic dysfunction. An additional mechanism that may contribute to HSPC dysfunction in our NHP model is the negative impact of alcohol on intracellular NAD+ levels (Di Rocco et al., 2019), which is required for HSPC maintenance and regeneration (Vannini et al., 2019, Luo et al., 2019, Anso et al., 2017), or the alcohol-induced damage of the supporting BM cells.

The BM niches consist of vascular endothelial and stromal cells that regulate homeostatic and regenerative aspects of HSPC function, including self-renewal, proliferation, and differentiation in response to pathological conditions such as infection or injury (Calvi and Link, 2015, Crane et al., 2017). Multiple lines of evidence suggest that vascular endothelium plays a paracrine role, supporting HSPC regeneration and maintenance in both mice and humans (Sasine et al., 2017). While arteriolar endothelial cells have been suggested to support HSPC maintenance and quiescence (Poulos et al., 2015, Xu et al., 2018), highly permeable sinusoidal endothelial cells promote HSPC maturation, proliferation, and mobilization from the BM in mice (Itkin et al., 2016, Kwak et al., 2016). Our studies demonstrate that in intact rhesus macaques, a small fraction of HSPCs is associated with the CD31+/CD34+ arterial endothelium (Robino et al., 2020). Thus, an increase in the densities of the CD31+/CD34+ arterial endothelium in the BM of alcohol drinkers may represent the adaptive hyperproliferation of supporting endothelium occurring in response to HSPC damage. Alternatively, alcohol consumption may promote the hypervascularization of the BM endothelium. Consistent with this idea, ethanol can enhance endothelial vascularization via the exosome-mediated mechanism (Lamichhane et al., 2017). Furthermore, at lower doses, ethanol can induce angiogenic proliferation and stimulates the production of vascular endothelial growth factor (Morrow et al., 2008, Gu et al., 2001).

Our study has limitations. (1) Although the data presented here offer new insight into alcohol induced HSPC dysfunction, more studies are needed to understand the precise molecular mechanism underlying alcohol-induced BM angiogenesis, its implication for hematopoiesis and the permanency of the effect from ethanol on hematopoiesis. It is unclear whether this change in angiogenesis is persistent or adaptive during the abstinent period and hence requires further studies. (2) The reduced mitochondrial respiration of CD34+ cells in response to alcohol drinking likely involves other BM cells in addition to HSPCs. For example, alcohol drinking increases the content of CD34+/CD31+ arterial endothelium and reduces the proportion of primitive CD34+CD45RA-CD90+CD49f+ HSPCs, which may collectively contribute to reduced mitochondrial respiration observed in isolated CD34+ cells of the alcohol drinkers. (3) Alcohol may affect HSPCs by indirect mechanisms through the pleotropic effect on systemic metabolism. Further studies in this NHP model are needed to discriminate the direct (HSPC and BM microenvironment damage) vs indirect (malabsorption of micro- and macro-nutrients) effects of chronic alcohol drinking on hematopoiesis and cellular metabolism in general.

Supplementary Material

Supp TableS1

Acknowledge:

we would like to thank Lindsey Crawford for help with the CFU colony assay and flow cytometry and the ONPRC pathology team for help with tissue collection.

Funding: AA019431, AA013510 and OD011092

Footnotes

Conflict of Interest: Authors declare no conflict of interest.

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