Influence of the pyruvate kinase activator mitapivat (AG-348) on RBC ATP content and export and RBC adhesivity in blood from healthy subjects and sickle cell disease patients

Background

Sickle cell disease (SCD) is a significant global health burden, characterized by hemolytic anemia, vaso-occlusive crises, and end-organ damage. Sickle hemoglobin (HbS) polymerizes into rigid fibers when deoxygenated. The resulting polymers distort red blood cells (RBCs) into a rigid, sickle-like shape, leading to aggregation and obstruction of microvessels, which manifests as painful vaso-occlusive crises (VOCs) (1). Repeated sickling and unsickling, along with oxidative damage, also compromise the RBC membrane, resulting in a significantly shortened RBC lifespan and hemolytic anemia (2).

RBCs are highly dependent on glycolysis as their sole energy source due to the virtual absence of mitochondria [mature SCD RBCs can actually retain mitochondria (3)]. Pyruvate kinase (PK) catalyzes the final step of glycolysis, converting phosphoenolpyruvate (PEP) to pyruvate and generating intracellular ATP. This process generates adenosine triphosphate (ATP) and regulates levels of 2,3-diphosphoglycerate (2,3-DPG). Intracellular ATP is essential for numerous functions in RBCs, eg maintaining ion and redox homeostasis, preserving cell membrane integrity, and promoting RBC deformability (4, 5). ATP depletion can lead to the accumulation of reactive oxygen species, damage to RBC membranes, and hemolysis (6). RBCs from adult SCD patients not treated with hydroxyurea (HU) are characterized by reduced (~12%) intracellular ATP levels compared to normal RBCs (7, 8). Additionally, metabolomic profiling in SCD RBCs points to elevated 2,3-DPG and other glycolytic intermediates, consistent with a “bottleneck” that could limit ATP generation and therefore rationalizing pharmacologic PK activation (7). Acute pain in SCD has been reported to vary as a function of PKLR (encoding PK isoforms present in liver and RBCs) gene polymorphisms, suggesting a treatable trait (9). The non-selective agent mitapivat (AG-348) activates PKM2 in muscle, PKL in liver, and PKR in RBCs, and is now approved for use in PK deficiency (PKD). PK activators are also being studied in SCD patients. They increase intracellular ATP production while decreasing (consuming) upstream 2,3-DPG, consistent with the function of PKR (10). The accelerated conversion of phosphoenolpyruvate to pyruvate results in more substrate for use in ATP production, thereby lowering levels of 2,3-DPG generated via the Rapoport-Luebering shunt. The depression of DPG, levels of which are elevated in SCD, acts to disfavor O₂ offloading from HbS, in turn disfavoring polymerization and sickling. Early results from preclinical (11, 12) and early-phase clinical (6, 10, 13, 14) studies of PK or PKR activators in SCD demonstrate favorable effects such as decreased susceptibility to sickling at a given partial pressure of oxygen (pO₂) (11), sustained improvement in Hb level, and decreases in markers of hemolysis (13).

The enhanced intra-RBC ATP has both predicted and newly recognized mechanistic benefits, such as stabilizing the cytoskeleton via band 3 dephosphorylation resulting from stabilized protein tyrosine phosphatase PTP1B (15). However, the influence of PK(R) activator-induced increases in RBC ATP content on ATP export from healthy or SCD RBCs has not been determined. Exported ATP from RBCs appears to act in several ways that are favorable for efficient and adaptive blood flow, including vasodilation in proportion to the degree of hypoxia, inhibition of intercellular adhesion, and the prevention of unwanted capillary permeability (4). We aimed to investigate the effects of the PK activator (PKA) AG-348 on exported and intracellular ATP in sickle or healthy control (HC) RBCs. Given that RBCs in sickle cell anemia exhibit increased adhesivity to endothelial cells, contributing to VOCs, augmenting antiadhesive ATP and its export capacity could be a desirable result of therapy (5). The PKR activator etavopivat lowered matrix metalloproteinase 9 (MMP-9) in blood from SCD patients, consistent with attenuated neutrophil degranulation (10). Alternatively, ATP and its derivatives can, depending on concentration, microenvironment, and the presence of ATPases (e.g., ectonucleotidases), exacerbate injury during inflammatory states such as sickle cell crisis or infection (16). Therefore, determining whether PKR activators may promote ATP export from SCD (and HC) RBCs is likely to inform the safe and efficacious use of this new and emerging class of drugs.

Methods

Blood sample acquisition, treatment ex vivo, and assays

Whole blood samples from outpatients with sickle cell disease (SCD) and from healthy adult volunteers were collected per institutional review board (IRB)-approved protocols. Sickle cell patients who were admitted (did not include ER visits) in the past 30 days, received transfusions in the past 30 days, had fetal hemoglobin (HbF) >20%, or who were on Voxelotor or L-glutamine were excluded. Primary assays requiring live cells (tonometry and RBC adhesivity) were completed within 24 hours of blood acquisition. Whole blood was typically incubated with AG-348 Selleck Chemicals, S6508, (10 µM) or DMSO (vehicle) at: 4 °C for 18–24 hours; 4 °C for 1 hour, or 37 °C for 1 hour. After incubation, RBCs were separated from whole blood by centrifugation and washed twice with PBS. The washed RBCs then underwent RBC tonometry (gentle, controlled exposure to gas mixes within a rotating glass-bulb tonometer equilibrated to 37 °C). RBC tonometry was performed using a final hematocrit of 1% RBCs in Krebs buffer containing 0.5% bovine serum albumin (BSA), as previously described (17). The RBC suspensions were equilibrated for 8 minutes under normoxic (a blend of 21% O₂, 5% CO₂, balance N₂ gases) or hypoxic (1% O₂, 5% CO₂, balance N₂ gases) conditions. RBC suspensions were then removed from the bulb. Following centrifugation of the exposed sample, supernatant ATP and intracellular (RBC) ATP were assayed using a luciferase method as previously described (18). Hemoglobin (Hb) assays were also performed on samples using a spectroscopic method (FLUOstar Omega, BMG LABTECH, Ortenberg, Germany), and these values were used in determining the degree of hemolysis in the samples.

Hemolysis calculation

RBC lysis was calculated as we have described, taking into account the supernatant and suspension [Hb] and the hematocrit. To quantify the extent of hemolyzed RBCs (19), the percent hemolysis was calculated as:

% hemolysis = 100−Hct x free Hb/total Hb

where the units for the input Hb values are in moles (in the suspension). The suspension Hct (1% in this study) is expressed conventionally (%) rather than as a fraction (from 0 to 1.0). We prespecified in our study design that no samples would be excluded as a result of elevated hemolysis, because we wanted to report the entire range of consequences of the ex vivo RBC manipulation.

RBC adhesion assay

In parallel, aliquots of the AG-348-treated or vehicle-exposed SCD whole blood or RBC-only samples were diluted with PBS for adhesion assays. Ibidi microfluidic slides with 6 channels each were pre-coated with 10 µg/mL laminin (recombinant human laminin 521, Gibco A29248) by incubation at room temperature for 1 hour, then coated with 1% BSA (to block nonspecific binding) at 37 °C for 1 hour. After incubation, slides were washed with PBS and loaded with either the whole blood (WB) or washed RBC samples. The packed RBCs were suspended in PBS buffer (dilution 1:1,500) as described (20), or WB was suspended in PBS at 1:500 dilution. Using a syringe pump, the channels were exposed to increasing degrees of shear stress (0, 0.3, 1, 3, or 10 dynes/cm²) by adjusting the rate of perfusion with the RBC suspension in DPBS (Dulbecco’s phosphate buffered saline with Ca⁺⁺, Mg⁺⁺; Corning 21-030-CV) while the microfluidic slides rested on a microscope platform. Photomicrographs of the channels were obtained at baseline and five minutes after each increase in shear stress. These images were later processed using Image J software to obtain cell counts and, in turn, calculate adhesivity (defined as the % cells remaining at a given shear stress relative to baseline).

Statistical and other analyses

GraphPad Prism was used to produce figures, calculate mean and SEM values, and perform selected statistical testing.

Statistical analysis was also performed in R using lme4, lmerTest, and emmeans. For each outcome (intracellular ATP, exported ATP, % RBC lysis), we fit a prespecified linear mixed-effects model with a random intercept for donor ID to account for within-donor pairing and between-donor heterogeneity. Fixed effects included treatment (AG-348 vs vehicle), exposure (hypoxia vs normoxia), and their interaction. Outcome-specific covariates tied to our hypotheses were included as fixed effects: age for exported ATP; age, HbF, MCV (mean RBC corpuscular volume), RDW (red blood cell distribution width), and reticulocyte count for intracellular ATP. Covariates were mean-centered for interpretability, and models were fit by REML. Inference on fixed effects used Satterthwaite/Kenward–Roger degrees-of-freedom approximations (via lmerTest); we report two-sided 95% confidence intervals and the corresponding p-values.

Primary contrasts were within-exposure treatment differences, estimated from the mixed models using estimated marginal means (emmeans), and reported as the [AG – vehicle] difference within hypoxia and within normoxia (with Kenward–Roger df, two-sided 95% CIs, and p-values).

For directional covariate hypotheses, we conducted selected, prespecified one-sided tests on the relevant slopes while reporting two-sided 95% CIs for the following predictions: Exported ATP declines with age (age slope < 0); intracellular ATP declines with age (age slope < 0), declines with HbF (HbF slope < 0), and increases with reticulocyte count (reticulocyte slope > 0). Unless otherwise stated, α = 0.05, and p-values are nominal and interpreted in the context of the prespecified primary questions.

Results

We obtained whole blood samples from 23 SCD patients and 13 healthy control (HC) subjects. Samples from 15 SCD patients (8 females, ages 3 to 19 years) were incubated with standard conditions (Table 1). Hb genotype was SS in all patients, and all but one was on hydroxyurea therapy. No SCD patient had documented alpha-thalassemia. The first several experiments conducted on HC blood samples revealed that incubating with AG-348 for 24 hours at 4 °C increased intra-RBC ATP and RBC ATP export under both normoxic and hypoxic conditions. Therefore these incubation conditions were applied for the SCD patient samples as well.

Table 1. Selected demographic and clinical characteristics of the SCD patients.

Supplementary Table 1 displays the hemoglobin values in the laboratory-prepared RBC concentrates from the SCD samples, with the clinical lab-based Hgb values displayed for reference.

Intra-RBC ATP after AG-348 exposure

In HC samples, the PKRA AG-348 raised intra-RBC ATP measured after exposure to either normoxia (57.84 ± 5.64 µM (SEM) after vehicle vs. 77.59 ± 10.12 µM after AG-348; Figure 1A) or hypoxia (57.67 ± 5.69 µM for vehicle vs. 81.03 ± 9.98µM after AG-348). This ATP-augmenting effect of the PKRA is consistent with prior reports (10, 21). In SCD samples, PKRA treatment ex vivo did not increase intra-RBC ATP significantly in normoxia (71.48 ± 11.96 µM after vehicle vs. 75.17 ± 19.41 µM after AG-348) or hypoxia (Figure 1B). The SCD RBC ATP values were also expressed after normalizing to the sample-respective Hb values (Figure 2). The findings of a lack of influence of hypoxia and/or mitapivat exposure on SCD RBC ATP values are similar whether expressed after normalizing to [Hb] or not. In both HC and SCD RBCs, intra-RBC ATP values did not differ significantly as a function of exposure to hypoxia vs. exposure to normoxia. Intra-RBC ATP values in the SCD cohort did not differ significantly from those in the HC group, whether assayed after normoxic or hypoxic exposure (unpaired t-test with Welch’s correction).

Exported ATP after HC or SCD RBC exposure to AG-348

ATP exported from HC or SCD RBCs was compared as a function of the assay ‘oxia’ status (normoxia or hypoxia), AG-348 (vs. vehicle) pre-exposure, and their combination. In untreated HC samples, exported ATP was increased after exposure to hypoxia (94.28 ± 24.34 nM vs. 79.70 ± 20.58 in normoxia; Figure 1C). ATP export from HC RBCs was increased significantly by treatment with the PKRA AG-348 in normoxia (vehicle: 80.31 ± 22.27 nM vs. 109 ± 30.23 nM after AG-348) and in hypoxia (Figure 1C). ATP exported by SCD RBCs was significantly but modestly greater after exposure to hypoxic vs. normoxic conditions (Figure 1D). Exported ATP did not change significantly in SCD RBCs treated with the PKRA AG-348, whether assayed in normoxia (79.70 ± 20.58 nM vs 77.12 ± 19.91 nM; Figure 1D) or hypoxia. Some assay values for supernatant ATP (exported ATP) were negative and of a magnitude less than the measurement error. They are reported and displayed as such in order to avoid the “correction” of some values but not others. Statistical testing with these samples imputed as “zero” yielded similar results. Exported ATP values did not differ in vehicle-treated RBCs from SCD patients vs. HC subjects, whether assayed after normoxic or hypoxic exposure (unpaired t-test with Welch’s correction).

Post-tonometry RBC lysis levels were ~1-2% under all conditions (mean 1.02 to 1.48%). These low levels of lysis in the ATP export assays did not differ significantly as a function of PKRA treatment, hypoxia exposure, or between HC and SCD RBCs (Figures 1E, F).

SCD RBC ATP after AG-348 incubation: effects of varying temperature and duration

We performed additional experiments to determine whether a shorter incubation period or a higher, more physiologic, temperature enabled AG-348-induced ATP generation in SCD or HC RBCs (Figure 3), given the lack of effect of incubation of SCD RBCs with AG-348 at 4 °C for 18–24 hours. Incubation of HC RBCs with AG-348 for 1 hour at either 4 °C or 37 °C failed to substantially or significantly increase RBC ATP (Figure 3). Baseline SCD RBC ATP was significantly depressed after 24 hours incubation with vehicle (DMSO) at 4 °C. Incubation of SCD RBCs with AG-348 failed to significantly increase ATP whether incubated for one hour at 4 °C, one hour at 37 °C, or 24 hours at 4 °C. Mixed-effects model statistical testing revealed no significant effect of 1-hour incubations with AG-348 overall in either cohort. In post-hoc subgroup comparisons, SCD RBC ATP was substantially depressed in the 24 hour at 4 °C conditions as compared to either one-hour incubation at either 4 °C (p=0.0235) or 37 °C (p=0.051). Incubation of HC RBCs at 37 °C for 24 hours profoundly depressed ATP (not shown), therefore we did not pursue these conditions in SCD RBCs. In summary, this experiment demonstrates depression of SCD RBC ATP after 24 hours cold storage, and no substantial change in RBC ATP in response to AG-348, whether incubated shorter periods (1h at 4 °C or 37 °C) or longer (24h at 4 °C). The lack of changes in SCD intra-RBC ATP or ATP exported from SCD RBCs after incubation with AG-348 for 24h at 4 °C are in direct contrast to the results in HC RBCs (Figure 1).

Additional statistical analyses of effects of AG-348 on RBC ATP and post-assay lysis of HC or SCD RBCs

Test estimates, confidence intervals and additional statistical test results for the effects of AG-348 on HC or SCD intracellular (intra-RBC) ATP, exported ATP, and post-assay % RBC lysis are summarized in Table 2.

Table 2. Summary of statistical testing of AG-348 (vs. vehicle) effects on intra-RBC ATP, exported ATP, and post-assay RBC lysis in HC or SCD RBCs assayed in either normoxia or hypoxia.

Selected clinical/assay correlation and linear mixed model testing

When correlating the experimental findings with SCD patient clinical data, we found a positive and significant (p=0.046) correlation of the reticulocyte count and intra-RBC ATP in SCD samples (Table 3). HbF did not correlate significantly with intra-RBC ATP in SCD samples (Table 3). Neither age nor sex associated significantly with SCD RBC ATP content or export (Table 3) or RBC lysis findings. MCV and RDW did not correlate significantly with basal (vehicle-exposed) or post-AG-348 intra-RBC ATP values in normoxia or hypoxia in SCD patients (not shown). When MCV and RDW were included along with age, HbF, and reticulocyte count there was no significant relationship to intra-RBC ATP using a linear mixed model (not shown). There was no significant correlation between the % reticulocyte count and the basal (vehicle-exposed) or post-AG-348 intra-RBC ATP values in normoxia in SCD (not shown).

Table 3. Directional test results for selected tests of correlations between the clinical data and the lab assay results from SCD patients.

SCD RBC adhesivity after AG-348

We assayed the adhesivity of SCD RBCs to the basement membrane substrate laminin after treating either whole blood or isolated RBCs with AG-348 (or vehicle), followed by microfluidic assay of adhesion of RBCs (perfused as suspensions of RBCs or whole blood). SCD RBC treatment with AG-348 ex vivo did not significantly alter RBC adhesivity to laminin across a range of physiologically relevant shear stresses (Figure 4, upper), nor did the shear effect (p<0.0001 alone) interact with the AG-348. By contrast, there was a significant interaction (p = 0.028) between shear (again p<0.0001 alone) and AG-348 effect for the SCD RBC adhesion to laminin after WB exposure to AG-348 (vs. vehicle) (Figure 4, lower). The effect of WB exposure to AG-348 on RBC adhesivity did not reach statistical significance (p = 0.15).

Discussion

The polymerization of deoxygenated sickle hemoglobin (HbS) is just one key underlying event in sickle cell disease (SCD) (22). Mature RBCs are highly dependent on glycolysis to generate ATP (4), which is critical for maintaining cellular viability and membrane integrity (6, 21, 22). RBCs from adult SCD patients not on hydroxyurea (HU) treatment have modestly (~12-19%) decreased intracellular ATP levels and elevated 2,3-DPG when compared to healthy control RBCs (5, 6, 8, 21). Indeed, a significantly but mildly greater RBC ATP concentration was measured in SCD patients on HU therapy in another cohort (23). In the present study, RBC ATP was not depressed in children and adolescents with SCD (nearly all of whom were treated with HU), consistent with published results in adults with SCD on HU therapy (8). Notably, in that study, patients not on HU therapy had ATP values ~12% lower than those in blood from HCs (8).

Such metabolic defects are relevant to the clinical symptoms in patients with sickle cell disease and may impact underlying pathological events such as susceptibility to lysis and RBC adhesivity (5). Deficient intracellular ATP compromises cell health and deformability, contributing to the pathophysiology of SCD (22). Elevated 2,3-DPG (a negative allosteric effector of O₂-binding by Hb) reduces the oxygen affinity of HbS, stabilizing the low-affinity deoxyhemoglobin conformation and promoting polymerization (11). The increased 2,3-DPG and decreased intracellular ATP in sickle cells from non-HU-treated adults are viewed as consistent with impaired glycolytic flux, which is controlled in part by the rate-limiting enzyme pyruvate kinase (11). PKR activation is therefore a rational therapeutic strategy in SCD because it exerts multimodal effects: increasing ATP production (which could be advantageous even when basal ATP levels are normal) and simultaneously decreasing 2,3-DPG by enhancing utilization of upstream substrate (4, 12).

The PKR activator etavopivat (FT-4202) significantly increased intracellular ATP levels in nonhuman primates and dose-dependently increased RBC ATP in healthy human subjects and patients with SCD (up to 12 weeks of treatment) (6, 10, 21, 22). This increase in ATP is proposed to enhance overall RBC function and health (22). PKR activation reduced 2,3-DPG levels, leading to a leftward shift in the oxygen dissociation curve (decreased P₅₀; increased Hb O₂ affinity) in healthy subjects and in SCD RBCs treated ex vivo (6, 22). This increased oxygen affinity reduces the propensity for HbS polymerization and RBC sickling, resulting in increased hemoglobin levels.

Both RBC-specific (such as the PKRA etavopivat/FT-4202) and non-specific PK activators (exemplified by mitapivat/AG-348) are under development and clinical investigation. Tebapivat (formerly AG-946) is a newer PKRA, which activates PKR and PKM2 isoforms (24). In a non-randomized, open-label, phase 1 study in adult patients with SCD, tebapivat was noted to elicit increases from baseline hemoglobin, increases in RBC ATP from baseline, and reductions in 2,3-DPG from baseline (25). Improvements in markers of hemolysis (lactate dehydrogenase and bilirubin) and erythropoiesis (reticulocytes) were also observed.

In the present study, ex vivo treatment of healthy RBCs with a single dose of the PKR activator mitapivat (AG-348) raised RBC ATP levels and ATP export, whereas it did not alter RBC ATP content or export in SCD RBCs. To our knowledge, our finding that AG-348 increased ATP export by HC RBCs in normoxia and in hypoxia is novel. The ability of PKR activator to promote RBC export of ATP may have new and broad therapeutic and safety implications, for example in cardiovascular disease or sepsis, since ATP has vasoregulatory, anti-permeability, and antiadhesive actions in the vasculature (4, 26).

The reason for the discordant findings, namely increased ATP content in and export from HC RBCs but not SCD RBCs, after PKR activator treatment ex vivo, is uncertain. One possibility relates to the enzyme-stabilizing actions of the PKR activators on PKR itself (6, 27). Single-dose treatment schemes such as the one we used may therefore underestimate the (potential) effects of longer-term, repeated administration. In the case of healthy RBCs where PK is presumably stable, PKR activation may be robust after a single dose, as suggested by the changes in RBC ATP in the present study and in prior work (22, 23). Our samples are from patients being treated with hydroxyurea, which has been reported in the past to increase ATP generation in and release from RBCs (28). We therefore speculate that in these patients, ATP content and export may already be elevated secondary to HU treatment and thus unlikely to rise further after PK activation. The lack of effect of the PK(R)A on SCD RBC ATP ex vivo in our study also contrasts with findings that in vivo treatment with AG-348/mitapivat or etavopivat raises RBC ATP levels in SCD patients (10, 29, 30). Moreover, others have demonstrated increases in RBC ATP after single-dose treatment ex vivo, particularly in PKD patients. Differences in incubation conditions and the specific RBC disorder studied may account for the lack of increased RBC ATP in our study (in SCD RBCs ex vivo) as compared to others (in PKD RBCs ex vivo) (27, 31). We elected not to pursue PKA incubations in RBCs at 37 °C for 24 hours because this led to modest (SCD) or marked (HC) RBC ATP depletion. The reason for the lack of effect of the PKR activator on DPG levels in either HC or SCD RBCs ex vivo is also uncertain, and may reflect suboptimal incubation conditions. Given the significant association between reticulocyte count and RBC ATP in the SCD cohort, it will be interesting to investigate in future studies whether therapeutic benefits relate to effects in reticulocytes (and later perhaps, in the more mature RBCs they become). Future clinical investigation should examine or take into account both potential differences in HC vs. SCD RBCs in responses to PK activators, the potential role of WBCs in the effects of PK activation in whole blood or in vivo, and the caveats associated with ex vivo studies of novel therapeutics, particularly recognizing limits in the modeling of duration of therapy exposure and longer-term cellular turnover and maturation.

Beyond its vital role in cell-intrinsic functions, ATP is recognized as a key vasoregulatory and anti-adhesive mediator exported by RBCs, with particular importance in the responses to hypoxia or mechanical deformation of the RBC (4, 5). A relationship between basal RBC ATP levels and subsequent ATP export has been established, suggesting that the extent of ATP export depends in part on the RBC ATP content (5, 32, 33). Other determinants include shear stress, the ambient PO₂, and the position and shape of the O₂ dissociation curve (i.e. O₂ affinity, as reflected in the P₅₀) (4, 34). Allosteric effectors of Hb’s O₂ binding function are therefore also determinants of ATP export since they influence P₅₀ and therefore Hb O₂ saturation. We demonstrated that, like HC RBCs, SCD RBCs do export additional ATP in hypoxia, and the increased extracellular ATP we measured could not be accounted for by increased RBC lysis, which did not differ significantly. However, unlike with healthy RBCs, ATP export did not increase when SCD RBCs were treated with a PKR activator (Figure 1D). Since PKR activators depress DPG (which is consumed as PK generates ATP), the DPG-depressing effects of PKR activation might tend to disfavor net ATP export by raising O₂ affinity. SCD RBCs have elevated DPG levels compared to healthy AA RBCs. Thus, the DPG-depressing effects of PKRAs may be greater in SCD RBCs than in healthy RBCs, in turn disfavoring O₂ desaturation and thus ATP export (despite any PKRA-induced increased ATP content). Extracellular ATP derived from RBCs or from other cell types, including endothelial cells and leukocytes, can play injurious roles, acting as a DAMP (damage-associated molecular pattern) in inflammatory milieux from atherosclerosis to sepsis (4, 35, 36). In this context, the overall lack of increased ATP export from SCD RBCs following PKR activator treatment ex vivo could therefore be viewed as reassuring.

Exported ATP acts to limit intercellular adhesion of RBCs to endothelial cells (17). Reduced ATP export capacity has been observed in banked RBCs and corresponded with increased adhesivity (17). In SCD, RBCs can adhere excessively to endothelial cells or to basement membrane substrates such as laminin (5). We observed that the effects of the PKR activator on RBC adhesivity differed depending on whether the drug was applied to isolated RBCs or to whole blood. This differential effect suggests the possibility of a modulatory interaction between ATP derived from RBCs and another element of whole blood, e.g., plasma, platelets, or leukocytes. We point out that ex vivo, microfluidics-based models of RBC adhesivity have inherent limitations. The absolute extent of RBC adhesivity ex vivo is not believed to match that taking place in vivo, but does correlate with clinical severity in SCD (37). However, the model does have utility in predicting the direction (if any) of disease- or drug-induced changes in RBC adhesivity, which have typically matched those seen in vivo in our and others’ work (17, 20, 38). The generalizability of our findings is also limited by the fact that we examined RBC adhesivity to a relevant basement membrane substrate (laminin), but did not study RBC adhesivity to endothelial cells.

The mechanisms by which PKR activators could improve adhesivity and other cellular functions are multiple. Interventions that increase intracellular ATP content, such as PKR activation, are hypothesized to promote the capacity for increased ATP export (5). Elevated intra-RBC ATP supports membrane homeostasis, counteracting damage that could otherwise lead directly or indirectly to a pro-adhesive phenotype (22). Exported ATP is thought to limit the tendency of RBCs to adhere to endothelial cells (ECs) (5), because impaired ATP release from RBCs has been shown to promote their adhesion to ECs. PKR activation has been shown to improve the mechanical properties of sickle RBCs, including deformability and the tendency to sickle, which can be quantified as the point of sickling (PoS) (6, 22). For example, ex vivo incubation of SCD RBCs with etavopivat significantly reduced the P₅₀ and delayed the PoS (the pO₂ at which SCD RBC deformability drops sharply) in both HbSS and HbSC donors (10, 22). Since mechanical deformation is a known stimulus for ATP release, maintaining better RBC deformability and structural integrity through adequate ATP levels is essential for effective vasoregulation (5).

Findings from multiple preclinical and early clinical studies of the PKR activator drug class support the hypothesis that modulating RBC glycolysis may be a potent strategy in SCD therapeutics (21). The dual action of decreasing the sickling propensity (via lowering 2,3-DPG levels) and increasing RBC resilience and function (via ATP production) yielded benefits such as increased hemoglobin and reduced hemolysis markers (21). Increased intra-RBC ATP directly addresses the energy deficit characteristic of sickle cells (at least in the absence of HU treatment) and is predicted to restore the RBC’s capacity to export anti-adhesive ATP (4, 5), thereby mitigating the hyper-adhesivity and microvascular dysfunction seen in SCD (5).

Our data show, for the first time to our knowledge, that ATP export from HC RBCs increases after treatment with a PKRA (which also increased intra-RBC ATP, confirming prior reports). In contrast, similar one-time exposure to PKRA did not significantly increase intra-RBC ATP in SCD, and ATP export from SCD RBCs was no greater after ex vivo PKRA treatment. PKRA treatment of whole blood but not isolated RBCs decreased SS RBC adhesion to laminin. Ongoing work addresses the determinants of cellular and exported ATP in HC and SCD RBCs (basal and treated) and the basis of the antiadhesive effect of this PKRA on SS RBCs. Also of interest is future study of the ability of RBCs from SCD patients maintained on PKRA therapy (vs. treated ex vivo with a single PKRA dose) to export ATP, as well as the functional (eg antiadhesive) consequences of such changes, if any. It would also be of interest to study the effect of ex vivo PKRA treatment on samples obtained from SCD patients not on hydroxyurea to see if there is significant difference in ATP export.

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Supplementary material

The Supplementary Material for this article can be found online at: https://www.frontiersin.org/articles/10.3389/frhem.2026.1766692/full#supplementary-material

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Footnotes

Edited by: Joan-lluis Vives-Corrons, Josep Carreras Leukemia Research Institute (IJC), Spain

Reviewed by: Lars Kaestner, Saarland University, Germany

Jonathan De Wilde, Feinstein Institute for Medical Research, United States

Chiara Federici, Case Western Reserve University, United States