Abstract
Elevated sphingosine 1-phosphate (S1P) is detrimental in Sickle Cell
Disease (SCD), but the mechanistic basis remains obscure. Here, we
report that increased erythrocyte S1P binds to deoxygenated sickle Hb
(deoxyHbS), facilitates deoxyHbS anchoring to the membrane, induces
release of membrane-bound glycolytic enzymes and in turn switches
glucose flux towards glycolysis relative to the pentose phosphate
pathway (PPP). Suppressed PPP causes compromised glutathione
homeostasis and increased oxidative stress, while enhanced glycolysis
induces production of 2,3-bisphosphoglycerate (2,3-BPG) and thus
increases deoxyHbS polymerization, sickling, hemolysis and disease
progression. Functional studies revealed that S1P and 2,3-BPG work
synergistically to decrease both HbA and HbS oxygen binding affinity.
The crystal structure at 1.9 Å resolution deciphered that S1P binds to
the surface of 2,3-BPG-deoxyHbA and causes additional conformation
changes to the T-state Hb. Phosphate moiety of the surface bound S1P
engages in a highly positive region close to α1-heme while its
aliphatic chain snakes along a shallow cavity making hydrophobic
interactions in the “switch region”, as well as with α2-heme like a
molecular “sticky tape” with the last 3–4 carbon atoms sticking out
into bulk solvent. Altogether, our findings provide functional and
structural bases underlying S1P-mediated pathogenic metabolic
reprogramming in SCD and novel therapeutic avenues.
Introduction
Sickle cell disease (SCD) is a prevalent life-threatening
hemoglobinopathy characterized by a point mutation in the β-chain of
hemoglobin. The aggregation of polymers of mutated sickle hemoglobin
(HbS) under deoxygenated conditions causes sickling, a fundamental
pathogenic process of the disease^[78]1. Although SCD was discovered
more than a century ago and identified as the “first molecular disease”
in 1949^[79]2, it is extremely disappointing that hydroxyurea is
currently the only FDA-approved treatment. Notably, increased oxidative
stress is also found in sickle erythrocytes and linked with hemolysis
and disease progression^[80]3. Therefore, identifying specific factors
and signaling pathways that contribute to sickling and oxidative stress
is essential to advance our understanding of this pathogenic process
and develop novel strategies for the treatment of SCD.
Recently, through accurately measuring functional phenotypes that are
the net result of genomic, transcriptomic and proteomic changes,
metabolomics profiling have become particularly useful to study mature
erythrocytes, where gene expression profiling is not an option due to
lack of a nucleus and de novo protein synthesis machinery. It has led
to the discovery of substantial metabolic alterations in SCD
erythrocytes of humans^[81]4,[82]5 and mice^[83]6 and implicated
multiple therapeutic possibilities. For example, metabolomics screening
revealed that circulating adenosine and erythrocyte
2,3-bisphosphoglycerate (2,3-BPG), an erythroid specific glycolytic
intermediate and potent allosteric modulator of Hb, and S1P, a
bioactive signaling molecule highly enriched in erythrocytes, are
elevated in patients and mice with SCD^[84]6,[85]7. Mechanistic studies
revealed that adenosine signaling through the adenosine A2B receptor
(ADORA2B) underlies increased erythrocyte 2,3-BPG^[86]6 and S1P^[87]8
in patients and mice with SCD. Additional studies showed that
pharmacologic inhibition or shRNA knockdown of sphingosine kinase 1
(Sphk1), the major enzyme in the spingolipid metabolism pathway to
produce S1P from sphingosine in erythrocytes, significantly attenuated
sickling and other deadly complications^[88]7. Moreover, a recent study
reported that increased S1P induces oxygen (O[2]) delivery to
counteract tissue hypoxia by inducing 2,3-BPG production in healthy
individuals at high altitude and in normal mice exposed to hypoxia,
which revealed the beneficial role of elevated erythrocyte S1P in
normal individuals^[89]9. However, it is puzzling why elevated S1P is
detrimental in SCD. To solve this puzzle, here we demonstrated the
genetic, functional, metabolic and structural mechanisms underlying why
the beneficial adaptation to high altitude in healthy individuals via
induction of S1P in normal erythrocytes is detrimental in sickle
erythrocytes. In contrast to normal erythrocytes, we revealed that
genetic deletion of Sphk1 in SCD has potent anti-sickling and
anti-hemolysis effects by correcting pathogenic metabolic
reprogramming, channeling glucose to pentose phosphosphate pathway
(PPP) relative to glycolysis, lowering 2,3-BPG production and boosting
NADPH/glutathione-mediated detoxification. These findings open new
promising scenarios in the development of innovative mechanism-based
therapies for SCD.
Results
Genetic evidence for the pathogenic role of elevated Sphk1 in SCD mice
To precisely asses the detrimental role and mechanisms of elevated S1P
in SCD, we generated a strain of mice with humanized sickle Hb and
Sphk1 deficiency by crossing the SCD Berkeley mice^[90]10 with Sphk1
^−/− mice^[91]11 (Supplementary Fig. [92]1a). The SCD/Sphk1 ^−/−
offspring were viable and lived to adulthood. PCR analysis confirmed
that the Sphk1 gene was deleted, and high-performance liquid
chromatography (HPLC) analysis of Hb species reveals the presence of
only HbS in SCD/Sphk1 ^−/− mice (Supplementary Fig. [93]1b,c). In
SCD/Sphk1 ^−/−erythrocytes, Sphk1 activity is undetectable; erythrocyte
and plasma S1P levels also decreased dramatically (Supplementary
Fig. [94]1d,e). The remaining plasma S1P is presumably derived from the
sphingosine kinase 2 (Sphk2) isoform expressed in a variety of
cells^[95]12, but not in mature erythrocytes due to lack of a nucleus.
Moreover, upstream sphingolipids such as ceramides and ceramide
1-phosphates increased significantly in SCD/Sphk1 ^−/− mice
(Supplementary Fig. [96]1g,h). Together, these data demonstrate that we
have successfully deleted Sphk1 in the SCD Berkeley mice.
Next, we compared sickling in age and gender matched SCD/Sphk1 ^−/−
mice and SCD mice. Erythrocyte shape was much more uniform and
organized in SCD/Sphk1 ^−/− mice (Fig. [97]1a), and the percentage of
irreversible sickle-shaped erythrocytes was significantly reduced
(Fig. [98]1b). Because intravascular hemolysis is one of the major
complications of SCD^[99]1, we assayed erythrocyte hemolysis by
measuring plasma Hb concentration, which is significantly lower in
SCD/Sphk1 ^−/− mice (Fig. [100]1c). In agreement, we found the clear
improvement of erythrocyte life-span in SCD/Sphk1 ^−/− mice
(Fig. [101]1d). Because of severe anemia, there is a large number of
reticulocytes in SCD mice^[102]10, which were significantly reduced in
SCD/Sphk1 ^−/− mice (Fig. [103]1e). Complete blood count (CBC) analysis
revealed higher total erythrocyte number, Hb concentration and
hematocrit in SCD/Sphk1 ^−/− mice (Supplementary Table [104]1).
Moreover, the erythrocyte distribution width was also significantly
reduced (Supplementary Table [105]1). Because S1P is a potent immune
regulator^[106]13, we found that the peripheral white blood cell count
was dramatically decreased in SCD/Sphk1 ^−/− mice with both neutrophil
and lymphocyte counts back to the normal range (Supplementary
Table [107]1). Splenomegaly and multiple organ damage are the hallmarks
of SCD progression^[108]1. Consistent with the above improvements,
splenomegaly (Fig. [109]1f), congestion and damage in spleen, lungs and
liver were also significantly improved in SCD/Sphk1 ^−/− mice, as
indicated by histology analysis (Fig. [110]1g; Supplementary
Fig. [111]2). Albumin level in the bronchoalveolar lavage fluid was
also significantly reduced (Fig. [112]1h), indicating less vascular
leakage in the lungs of SCD/Sphk1 ^−/− mice. Taken together, these data
provide solid genetic evidence demonstrating that deletion of Sphk1 is
beneficial in SCD.
Figure 1.
[113]Figure 1
[114]Open in a new tab
Genetic deletion of Sphk1 improves disease conditions in SCD Berkeley
mice. (a) Representative pictures of blood smears from SCD and
SCD/Sphk1 ^−/− mice (magnification ×400). Percentage of sickle cells
(b), plasma Hb (c) and reticulocytes (e) were significantly reduced
while erythrocyte lifespan was significantly prolonged (d) by genetic
deletion of Sphk1. Spleen size (f), H&E staining of spleens, livers,
and lungs (g), and albumin concentrations in bronchial alveolar lavage
(BAL) fluid (h) collected from SCD and SCD/Sphk1 ^−/− mice. Values
shown represent the mean ± SEM (n = 5-10); *p < 0.05 versus SCD,
Student’s t-test. Scale bar: 20μm in blood smear pictures; 200μm in H&E
staining pictures. Indicates sickled RBCs.
Enhanced erythrocyte pentose phosphate pathway and anti-oxidation capacity in
SCD/Sphk1^−/− mice
Giving the solid genetic evidence of elevated Sphk1 contributing to
sickling and disease progression^[115]7, we sought to further determine
the molecular basis. Because erythrocytes lack nuclei and organelles,
metabolic adaptation has a key role in erythrocyte homeostasis^[116]14.
Therefore, we chose to exploit an unbiased high throughput metabolomic
profiling to compare global metabolic changes in the erythrocytes among
WT, Sphk1 ^−/−, SCD and SCD/Sphk1 ^−/− mice. A total of 222 named
metabolites were detected in over 9,000 features screened
(Supplementary Data [117]1). Next, we performed an unbiased
pathway-enrichment analysis using MetaboAnalyst^[118]15. First, genetic
deletion of Sphk1 in normal mice did not affect erythrocyte metabolism
other than the significantly decreased S1P levels in erythrocytes and
plasma and increased ceramides (Supplementary Fig. [119]1e–g). However,
a large portion of the 25 pathways identified were affected by Sphk1
deficiency in SCD Berkeley mice, and the top three metabolic pathways
affected by genetic deletion of Sphk1 in SCD mice were pentose
phosphate pathway (PPP), glutathione metabolism, and sphingolipid
metabolism (Fig. [120]2a). Sphingolipid metabolism alteration validates
the impact of Sphk1 deletion (Supplementary Fig. [121]1d–h). Moreover,
we found substantially increased steady state levels of multiple
intermediates of PPP including glucose 6-phosphate (G6P),
gluconate-6-phosphate (6-P-gluconate), ribose 1-phosphate (R1P),
erythrose 4-phosphate (E4P) and sedoheptulose 7-phosphate (S7P) in the
erythrocytes of SCD/Sphk1 ^−/− mice relative to SCD mice
(Fig. [122]2b,c), suggesting that the PPP is significantly enhanced in
SCD/Sphk1 ^−/− erythrocytes. In agreement with enhanced PPP, we found
increased NADPH, an important byproduct of this pathway, in SCD/Sphk1
^−/− erythrocytes (Fig. [123]2d). As such, reduced glutathione (GSH), a
key NADPH-dependent antioxidant, was substantially elevated
(Fig. [124]2e). Altogether, these data strongly suggest a decrease in
oxidative stress in SCD/Sphk1 ^−/− erythrocytes. Not surprisingly, we
detected significantly lower ROS levels, MetHb and COHb in SCD/Sphk1
^−/− erythrocytes (Fig. [125]2f). Numerous studies have indicated that
excessive oxidative stress in SCD leads to hemolysis and erythrocyte
destruction^[126]3. Thus, we determined if deletion of Sphk1 increases
resistance of SCD erythrocytes to hemolytic challenges induced by
oxidative stress. After exposure to hydrogen peroxide (H[2]O[2]),
SCD/Sphk1 ^−/− erythrocytes had a significantly lower osmotic fragility
with increased half-maximal effective concentrations (EC50)
(Fig. [127]2g), consistent with increased GSH and NADPH-dependent
antioxidant capacity.
Figure 2.
[128]Figure 2
[129]Open in a new tab
Enhanced pentose phosphate pathway and anti-oxidant capacity in
SCD/Sphk1 ^−/− mouse erythrocytes. (a) Top metabolic pathways affected
by genetic deletion of Sphk1 in SCD mice. (b) Relative abundance of
selected PPP metabolites in erythrocytes from WT, Sphk1 ^−/−, SCD and
SCD/Sphk1 ^−/− mice. (c) Intensity peak of selected PPP metabolites in
erythrocytes from WT, Sphk1 ^−/−, SCD and SCD/Sphk1 ^−/− mice detected
by metabolomics screening. Levels of NADPH (d), GSH (e) and in
erythrocytes from WT, Sphk1 ^−/−, SCD and SCD/Sphk1 ^−/− mice. (f) ROS,
MetHb and COHb levels in erythrocytes from WT, Sphk1 ^−/−, SCD and
SCD/Sphk1 ^−/− mice. (g) Resistance of WT, Sphk1 ^−/−, SCD and
SCD/Sphk1 ^−/− erythrocytes to osmolality-induced hemolysis with or
without oxidative stress challenge. Values shown represent the
mean ± SEM (n = 5); *p < 0.05 versus WT; **p < 0.05 versus SCD;
Student’s t-test. G6P: Glucose 6-phosphate; 6-P-Gluconate:
Gluconate-6-phosphate; R1P: Ribose 1-phosphate; E4P: Erythrose
4-phosphate; S7P: Sedoheptulose 7-phosphate; GSH: reduced glutathione.
Reduced glycolysis and Hb-O[2] binding affinity in SCD/Sphk1^−/− erythrocytes
Glucose in erythrocytes is metabolized through either PPP, to generate
reducing equivalents to preserve redox homeostasis, or glycolysis, to
produce ATP as an energy source^[130]16. Additionally, approximately
19~25% of the glucose is utilized to produce 2,3-BPG, a key allosteric
regulator of Hb-O[2] affinity, which derives from the
Rapoport-Luebering branch of glycolysis^[131]17. Under high O[2]
saturation conditions, oxidative stress promotes PPP to generate NADPH.
To deliver O[2] efficiently while neutralizing excessive oxidative
stress caused by a heavy load of O[2], erythrocytes rely on a
finely-tuned O[2]–dependent modulation of glucose
metabolism^[132]18–[133]20. Based on the enhanced PPP and glutathione
metabolism in the erythrocytes of SCD/Sphk1 ^−/− mice (Fig. [134]2), we
sought to test if increased steady state levels of PPP intermediates in
SCD/Sphk1 ^−/− erythrocytes correspond to a decline of metabolic flux
through glycolysis. First, we found significantly increased glycolytic
intermediates including G6P, fructose 1,6-bisphosphate (FBP),
glyceraldehyde 3-phosphate (G3P), 2/3-phosphoglyceric acid (2/3-PG),
phosphoenolpyruvate (PEP) and pyruvate in SCD mouse erythrocytes
compared to WT (Fig. [135]3a,b), confirming that glycolysis rather than
the PPP is preferentially active in SCD mouse erythrocytes
(Fig. [136]3a,b), which explains the compromised capacity to produce
reducing equivalents (Fig. [137]2d) and preserve glutathione
homeostasis in SCD (Fig. [138]2e). In addition, levels of these
metabolites were not different between Sphk1 ^−/− and WT erythrocytes.
To our surprise, the levels of three upstream metabolites of glycolysis
including G6P, FBP and G3P were increased in SCD/Sphk1 ^−/−
erythrocytes compared to SCD (Fig. [139]3a,b), suggesting a metabolic
bottleneck downstream to G3P dehydrogenase (GAPDH)^[140]21. In
contrast, the levels of three glycolytic intermediates downstream of
G3P including 2/3-PG, PEP and pyruvate were significantly reduced in
SCD/Sphk1 ^−/− erythrocytes (Fig. [141]3a,b). More importantly, the
levels of 2,3-BPG, an erythrocyte-specific metabolite contributing to
sickling^[142]6,[143]22,[144]23 and an intermediate downstream of G3P,
was increased in the SCD erythrocytes but decreased in those of
SCD/Sphk1 ^−/− mice (Fig. [145]3c).
Figure 3.
[146]Figure 3
[147]Open in a new tab
Genetic deletion of Sphk1 reduces glycolysis and O[2] release and
channels glucose flux to PPP in SCD erythrocytes. (a) Relative
abundance of selected glycolysis metabolites in erythrocytes from WT,
Sphk1 ^−/−, SCD and SCD/Sphk1 ^−/− mice (upper); glycolysis is blocked
at the step where G3P is metabolized by GAPDH in erythrocytes of
SCD/Sphk1 ^−/− mice compared to SCD mice (lower). (b) Intensity peak of
selected glycolysis metabolites in erythrocytes from WT, Sphk1 ^−/−,
SCD and SCD/Sphk1 ^−/− mice detected by metabolomics screening. 2,3-BPG
level (c) and P50 (d) in WT, Sphk1 ^−/−, SCD and SCD/Sphk1 ^−/− mouse
erythrocytes. (e) Schematic illustration of glucose metabolism flux
detection using^13C[1,2,3]-Glucose. Ratios of
^13C[1,2,3]-Lactate/^13C[1,2,3]-Glucose (f)
and^13C[2,3]-/^13C[1,2,3]-G3P (g) in WT, Sphk1 ^−/−, SCD and SCD/Sphk1
^−/− mouse erythrocytes. Values shown represent the mean ± SEM (n = 5);
*p < 0.05 versus WT; **p < 0.05 versus SCD; Student’s t-test. PPP:
pentose phosphate pathway; G6P: Glucose 6-phosphate; FBP: Fructose
1,6-bisphosphate; G3P: Glyceraldehyde 3-phosphate; 2/3-PG:
2/3-Phosphoglyceric acid; PEP: Phosphoenolpyruvate; CO[2]: carbon
dioxide.
Given the fact that 2,3-BPG decreases HbS-O[2] binding
affinity^[148]22,[149]24 and in view of our findings that elevated
Sphk1 contributes to increase 2,3-BPG production in sickle
erythrocytes, we hypothesize that elevated Sphk1 underlies sickling by
inducing 2,3-BPG, decreasing HbS-O2 binding affinity and thus
increasing deoxy-HbS polymerization. To test this hypothesis, we
measured the O[2] equilibrium curve (OEC) by calculating the partial
pressure of O[2] required to produce 50% Hb-O[2] saturation (P50), and
found increased Hb-O[2] binding affinity and thus reduced P50 in
SCD/Sphk1 ^−/− mouse erythrocytes (Fig. [150]3d). However, there was no
difference between WT and Sphk1 ^−/− erythrocytes. These findings
indicate that decreased 2,3-BPG due to deficiency of Sphk1 results in
increased Hb-O[2] binding affinity and decreased deoxyHbS level, which
support the observation of less sickling in SCD/Sphk1 ^−/− mice
(Fig. [151]1a). Altogether, the beneficial role of Sphk1 deficiency in
anti-sickling and anti-hemolysis is strongly supported by metabolic
rewiring in SCD/Sphk1 ^−/− erythrocytes.
Genetic deletion of Sphk1 channels glucose fluxes to PPP in SCD erythrocytes
Next, to provide direct mechanistic insight about intracellular glucose
flux, we first assayed glucose uptake in WT, Sphk1 ^−/−, SCD and
SCD/Sphk1 ^−/− erythrocytes. Interestingly, although glucose uptake is
significantly increased in the SCD Berkeley mouse erythrocytes compared
to normal, which agrees with previous studies done in human SCD
erythrocytes^[152]5, there was no difference between SCD and SCD/Sphk1
^−/− or WT and Sphk1 ^−/− erythrocytes (Supplementary Fig. [153]3a),
indicating that differences in PPP and glycolysis pathways in SCD and
SCD/Sphk1 ^−/− erythrocytes are not caused by variation in glucose
uptake. In addition, we used the stable ^13C[1,2,3-]glucose isotope to
trace intracellular glucose metabolism through glycolysis and PPP in
SCD and SCD/Sphk1 ^−/− erythrocytes at different time points.
Specifically, we investigated whether glycolysis or the PPP is the
major contributor to the accumulation of G3P in SCD/Sphk1 ^−/− mouse
erythrocytes by determining the ratios of the isotopologues
^13C[2,3]/^13C[1,2,3] of G3P (Fig. [154]3e). First,
^13C[1,2,3]-lactate/^13C[1,2,3]-glucose ratios were significantly
increased in SCD but not in SCD/Sphk1 ^−/− mouse erythrocytes in a
time-dependent manner, indicating significantly increased metabolic
flux through PPP in SCD mouse erythrocytes (Fig. [155]3f). In
agreement, ratios of ^13C[1,2,3]-6-P-gluconate, a PPP metabolite, to
^13C[1,2,3]-glucose were significantly higher in SCD/Sphk1 ^−/− mouse
erythrocytes (Supplementary Fig. [156]3b). As expected, ratios of
^13C[2,3]-G3P/^13C[1,2,3]-G3P isotopologues were significantly higher
in erythrocytes from SCD/Sphk1 ^−/− erythrocytes (Fig. [157]3g),
indicating that glucose flux through the PPP was enhanced.
Sphk1 regulates GAPDH localization in SCD erythrocyte
Under normoxia, erythrocyte glucose flux through glycolysis is limited
by the inhibitory sequestration of glycolytic enzymes, including GAPDH,
to the cytosolic domain of the membrane protein Band3 (cdB3)^[158]20.
However, under hypoxia, deoxygenated Hb (deoxyHb) competes with
glycolytic enzymes for binding to cdB3, which results in the release of
those enzymes, thereby promoting glycolysis^[159]18–[160]20. Recent
studies have revealed that deoxyHbS disturbs normal coupling among
erythrocyte O[2] content, glycolysis and antioxidant capacity by
increasing release of membrane anchored GAPDH to the cytosol^[161]5.
Therefore, we investigated if elevated erythrocyte Sphk1-mediated
increased S1P is involved in regulating the intracellular location of
GAPDH by facilitating the binding of deoxyHbS to cdB3 and the release
of GAPDH to the cytosol. Western blot results indicated no obvious
difference in total amount of GAPDH (Fig. [162]4a,b). However, a
significantly larger percentage of GAPDH in SCD/Sphk1 ^−/− erythrocyte
was found on the membrane (Fig. [163]4a,b). In agreement, cytosolic
GAPDH activity was significantly reduced in SCD/Sphk1 ^−/− erythrocytes
compared to SCD (Fig. [164]4c). However, no significant difference of
GAPDH localization and activity was found between WT and Sphk1 ^−/−
erythrocytes (Fig. [165]4a–c). Thus, genetic and biochemical evidence
demonstrate that Sphk1 enhances release of membrane anchored GAPDH and
increases cytosolic GAPDH activity in SCD erythrocytes. Next, to
dissect the influence of non-specifically bound HbS, we isolated
erythrocyte membrane ghosts from SCD and SCD/Sphk1 ^−/− mice and
inverted the membrane ghost on silicon beads and washed for 8 times
with low-salt buffer. Then, we detected significantly higher levels of
heme anchored on the ghost membrane of erythrocytes from SCD
(Fig. [166]4d). Thus, these findings indicate that elevated Sphk1 is
associated with enhanced HbS anchoring to membrane, and are consistent
with the release of membrane bound GAPDH and increased cytosolic GAPDH
activity.
Figure 4.
[167]Figure 4
[168]Open in a new tab
Sphk1-mediated production of S1P functions intracellularly to regulate
GAPDH and Hb localization and subsequent metabolic consequences. (a,b)
Total and membrane bound GAPDH protein levels in WT, Sphk1 ^−/−, SCD
and SCD/Sphk1 ^−/− mouse erythrocytes detected by western blot (cropped
blots displayed, whole blots see supplementary information). Cytosolic
GAPDH activity (c) and membrane bound heme (d) in WT, Sphk1 ^−/−, SCD
and SCD/Sphk1 ^−/− mouse erythrocytes. Cytosolic GAPDH activity (e),
membrane bound heme (f), 2,3-BPG levels (g), NADPH levels (h) and
percentage of sickled erythrocytes in SCD/Sphk1 ^−/− mouse erythrocytes
treated with different concentration of S1P. Values shown represent the
mean ± SEM (n = 5); *p < 0.05 versus WT or 2.5 µM; **p < 0.05 versus
SCD or 5 µM, Student’s t-test.
S1P is the ligand to five G-protein coupled receptors. To test if
extracellular or intracellular S1P is playing the major role in
regulating Hb and GAPDH localization and erythrocyte metabolism, we
treated SCD/Sphk1 ^−/− erythrocytes with different concentrations of
S1P to mimic conditions in SCD mice. Interestingly, S1P treatment up to
2.5 µM, which are sufficient to activate all of the S1P receptors, did
not affect cytosolic GAPDH, membrane heme, 2,3-BPG and NADPH in
SCD/Sphk1 ^−/− mouse erythrocytes (Fig. [169]4e–h). However, when
treated with higher concentrations of S1P that can lead to increase of
intracellular S1P levels^[170]25, all of the above parameters showed
dose-dependent increase (Fig. [171]4e–h). Moreover, we found that
sickling of SCD/Sphk1 ^−/− erythrocyte was induced not in low
concentrations of S1P at but only in 5 and 8 µM (Fig. [172]4i). Thus,
these data implicate that decreased sickling mediated by Sphk1
deficiency is independent of S1P receptors. Overall, we provide direct
evidence that S1P functions intracellularly as a modulator promoting
deoxy-HbS anchoring to the membrane and subsequently enhancing the
release of membrane bound GADPH to the cytosol, which in turn leads to
increased cytosolic GAPDH activity in SCD erythrocytes.
Co-binding of 2,3-BPG and S1P to Hb is required for S1P-induced decrease in
Hb-O[2] affinity
Since S1P directly induces HbS anchoring to membrane, we speculated
that S1P binds to HbS and HbA and reduces Hb-O[2] affinity as other
phosphates do. Indeed, S1P-conjugated beads successfully pulled down
HbA and HbS from normal and SCD patients erythrocyte lysates, while
sphingosine or lysophosphatidic acid beads cannot (Fig. [173]5a),
indicating that S1P directly and specifically binds to both human HbA
and HbS in erythrocyte lysates. Next, to determine if S1P regulates
Hb-O[2] binding affinity, we assayed HbA and HbS O[2] binding
equilibrium curves in the absence or presence of different
concentrations of S1P. To mimic the molar ratio of S1P to Hb from
1:2500 to 1:500 as seen in normal and sickle human erythrocytes, we
used HbA or HbS at 10 µM with the concentrations of S1P ranging from 0
to 10 nM. Unexpectedly, S1P alone has no effect on Hb-O[2] binding
affinity. Realizing that there is very abundant 2,3-BPG is in
erythrocytes which binds to deoxyHb, we tested if 2,3-BPG is required
for S1P-mediated reduction in Hb-O[2] affinity. P50 of purified HbA or
HbS in the presence of 2,3-BPG along with S1P revealed that S1P
decreased HbA and HbS-O[2] binding affinity in a dose-dependent manner
(Fig. [174]5b and c). Thus, we provide biochemical and functional
evidence that S1P binds directly to Hb but requires co-binding of
2,3-BPG to decrease O[2] binding affinity, presumably by further
stabilizing deoxyHb and increasing its T-state character.
Figure 5.
[175]Figure 5
[176]Open in a new tab
Functional and structural evidence for S1P binding to Hb and
stabilization of deoxyHb in T-state. (a) Pull-down of Hb by
lysophosphatidic acid (LPA), Sphingosine and S1P beads from normal
human and SCD patient RBC lysates (cropped blots displayed, for full
blots see supplementary information). S1P, at physiological and
pathological molar ratios, induces further O[2] release from HbA (b)
and HbS (c) in the presence of 2,3-BPG. (d) Crystal structure of
deoxyHbA in complex with 2,3-BPG (bound at the β-cleft), and S1P (bound
both at the central water cavity and the protein surface). (e) Close
view of 2,3-BPG binding at the β-cleft. (f–h) S1P binds to the surface
of HbA in the presence of 2,3-BPG and induces further conformational
change stabilizing the complex in T-state. (i) Working model: elevated
erythrocyte Sphk1 activity increases production of S1P, which binds to
deoxyHbS and facilitates deoxyHbS anchoring to membrane and release of
GAPDH. Increased cytosolic GAPDH accelerates glycolysis and 2,3-BPG
production while decreasing PPP and antioxidant production. Increased
2,3-BPG leads to more deoxyHbS and more sickling while decreased
antioxidant causes more oxidative stress (ROS) and more hemolysis.
Altogether, erythrocyte S1P induced by elevated Sphk1 activity leads to
impaired metabolic reprogramming and thus underlies sickling, hemolysis
and disease progression in SCD.
X-ray crystallography reveals atomic level insight into S1P-Hb binding
Given the fact that structures (both tertiary and quaternary) of
liganded or unliganded normal HbA and sickle HbS are identical even at
the pathogenic βVal6 mutation site^[177]26,[178]27 and that it is
easier to crystalize HbA, we chose to determine the crystal structures
of deoxyHbA in complex with S1P alone (deoxyHbA-S1P) or in combination
with 2,3-BPG (deoxyHbA-S1P-BPG) (subsequently solved at 2.4 Å and
1.8 Å) to gain structural insight into the above described S1P-mediated
functional/biological effects. The structures were determined by
molecular replacement using the high resolution native deoxyHbA
structure (PDB code: 2DN2)^[179]28. Expectedly, and consistent with
published studies, we observed 2,3-BPG bound in two alternate
conformations at the dyad axis of the β-cleft in the ternary
deoxyHbA-S1P-2,3-BPG complex to tie together the two β-subunits via
interactions with the residues βHis2, βLys82, βAsn139, and βHis143 from
both β-subunits (intermolecular interactions) in symmetry-related
fashion (Fig. [180]5d and e). In both the binary deoxyHbA-S1P and
ternary deoxyHbA-S1P-2,3-BPG complexes, S1P was observed bound in the
central water cavity, with the phosphate and the amide moieties located
in a pocket formed by α1Lys99, α1His103, β1Asn108, β1Tyr35 and
β1Gln131, while the flexible aliphatic long chain snaked toward the
β-cleft making hydrophobic interactions with α1 Phe36, α1Ser35,
β1Lys132, β1Gln131, β1Ala135, β1Val1 and β1His2 (Supplementary
Fig. [181]4). However, while the ternary complex showed S1P bound in a
symmetry-related fashion (Fig. [182]5d; Supplementary Fig. [183]4), the
binding of S1P in the binary deoxyHbA-S1P complex was very weak
(Supplementary Fig. [184]5), and only one S1P binding site could be
unambiguously fitted in the complex (α1β1 site). In the deoxyHbA-S1P
complex, the side-chain of α2Lys99 was in a similar position as 2DN2,
consistent with very weak S1P binding as opposed to the ternary
deoxyHbA-S1P-2,3-BPG complex. Binding of 2,3-BPG might have increased
the affinity of the protein for S1P in the central water cavity. Note
that the same concentration of S1P was used during crystallization of
both the binary and ternary complexes. Each S1P-associated interaction
in the central water cavity was essentially intramolecular in nature
(i.e. make interactions with only α1β1 or α2β2) and suggest that
central-water cavity bound S1P might not contribute significantly to
the stabilization of the T-state structure^[185]29. This observation
indicates that although the affinity of S1P binding to central water
cavity of the protein is increased by 2,3-BPG, it is unlikely to cause
significant changes to the deoxyHbA conformation.
Interestingly, besides the central water cavity bound S1P as described
above, we also found two additional S1P molecules bound in a
symmetry-related fashion at the surface of the deoxyHbA-S1P-2,3-BPG
complex but not in the deoxyHbA-S1P complex, indicating that 2,3-BPG
binding most likely is required for S1P binding at the surface of the
protein. Specifically, the phosphate moiety binds close to the α1-heme
and located in a highly positive environment formed by α1Arg92,
β2Arg40, α1His45, and α1Lys90, as well as with β2Glu43; either making
direct salt-bridge/hydrogen-bond interactions and/or water-mediated
hydrogen-bond interactions with these residues (Fig. [186]5f). The S1P
amide nitrogen makes water-mediated interaction with α1Lys90 and the
α1-heme propionate. The side-chains of both β2Glu43 and α1Lys90 have
moved from their native positions to make interactions with the S1P.
The highly flexible aliphatic chain snaked along a shallow cavity wall
making hydrophobic interactions with the so-called “switch region”
residues of β2Phe41, α1Thr41, α1Pro44, β2Leu96, β2His97, as well as
with β2-heme, like a molecular sticky tape. The last 3–4 carbon atoms
of the aliphatic chain stick out into the bulk solvent
(Fig. [187]5f,g). Similar symmetry related interactions are observed
from the β2-heme site to the β1-heme site. As previously noted, the
switch region is characterized by significant structural changes during
the T to R transition^[188]29,[189]30, and effectors that prevent these
changes are known to decrease Hb affinity for O[2] ^[190]29. These
findings raise an intriguing possibility that the surface bound S1P
which makes several inter-subunit interactions that involve residues
from the switch region serve to stabilize the T-state, and presumably
further decrease the T-state affinity for O[2].
To test this hypothesis, we compared the T-state structures
deoxyHbA-S1P, deoxyHbA-S1P-2,3-BPG, native deoxyHbA (PDB code: 2DN2)
and native R-state COHbA structure (PDB code: 2DN1)^[191]28 by
superposing their α1α1 dimers (~0.3 Å) and then obtaining the screw
rotation angles that are required to superpose the non-superposed α2β2
Hb dimers as a quaternary measure^[192]29,[193]30. Notably, we found
that deoxyHbA-S1P-BPG was further removed from the R-state (15.7°) more
than deoxyHbA-S1P (14.8°) and T-state HbA (PDB code: 2DN2) (14.2°).
Consistently, the dimer interface β2 F helix/β2FG corner at the switch
region show some significant positional differences, with that of
deoxyHbA-S1P-BPG further removed from the R-state (Fig. [194]5h). These
observations support our conclusion that 2,3-BPG is required for S1P
binding to the protein, especially to the surface of the protein which
leads to the protein becoming more tense, and presumably lower affinity
for O[2] compared to either the deoxyHbA or the binary deoxyHbA-S1P
complex structures.
Discussion
Balance of glucose flux between glycolysis and PPP is extremely
important in mature erythrocytes and therefore finely tuned. A myriad
of studies reported metabolic reprogramming in normal erythrocytes in
response to hypoxia through the binding of deoxyHb to cdB3 and
subsequently increased cytosolic glycolytic enzymes
availability^[195]18–[196]20. However, in SCD erythrocytes, glucose
metabolism is constantly programmed with glycolysis “on” and PPP “off”
even at steady normoxia condition^[197]5, leading to the over
production of 2,3-BPG and the shortage of antioxidant glutathione; yet
the regulating factors and the underlying mechanisms remain
underdetermined before our study. Our findings immediately suggest that
glucose in sickle erythrocytes was predominantly metabolized via
glycolysis rather than the PPP, as confirmed by tracing experiments.
Moreover, genetic deletion of Sphk1 in SCD reprograms glucose
metabolism by channeling glucose to PPP instead of glycolysis, which in
turn leads to increased NADPH and decreased 2,3-BPG production.
S1P is a versatile bio-active signaling lipid highly abundant in the
erythrocytes^[198]25. Although previous studies showed that S1P levels
are increased and contribute to sickling and disease progression in
SCD^[199]7, nothing is known about the underlying mechanism. It is
indeed interesting that intracellular S1P at µM concentrations can
mediate metabolic reprogramming in SCD by regulating binding of deoxyHb
to cdB3. Due to the molar ratio of about 1:300 between cdB3 and
Hb^[200]5, cdB3, not deoxyHb (present at mM concentrations), is the
rate-limiting factor in deoxyHb-cdB3 interaction. Interestingly, S1P
has an approximately 1:1 molar ratio with cdB3 in normal erythrocytes
and an even higher ratio in SCD erythrocytes. Thus, although deoxyHb is
expected to be present at a much higher concentration than S1P, it is
the concentration of S1P that controls the amount of deoxyHb that binds
to cdB3. Besides regulating the sequestration of glycolytic enzymes, it
is reasonable to speculate that increased S1P in SCD erythrocytes may
also play a role in other cdB3-mediated effects including the binding
of S-nitrosohemoglobin and spectrin to cdB3. The former is involved in
the nitric oxide (NO) metabolism^[201]31,[202]32 in erythrocytes while
the latter plays a key role in erythrocyte deformability^[203]33, both
of which are important in the pathophysiology of
SCD^[204]23,[205]34,[206]35.
Our data indicate that S1P binds to the surface of 2,3-BPG-Hb and leads
to considerable additional conformational change of deoxyHb (by making
hydrophobic interactions at the switch interface) to a more T-state
character that in part should explain the decreased Hb-O[2] affinity.
It is also notable that the surface-bound S1P could sterically impede
diffusion of diatomic ligands (O[2]) into the heme, and in part also
decreased Hb-O[2] affinity. Similar studies have been reported for
allosteric effectors that bind and block the heme access to the bulk
solvent^[207]29,[208]36. Although S1P was also observed bound in the
central water cavity, the water cavity is known to be a “sink” for
several compounds especially those with anionic groups and not all of
these compounds show an allosteric effect^[209]29. Since in the absence
of 2,3-BPG, we observed weak binding of S1P at the water cavity and no
apparent effect on the protein’s allosteric activity, it is possible
that the central water cavity S1P binding is non-specific. Another
interesting structural observation is that the last 3–4 carbon atoms of
the surface bound S1P do not make any interaction with the protein
residue but hang out in the bulk solvent, which could possibly mediate
hydrophobic interactions with other proteins, akin to the hydrophobic
βVal6 pathogenic mutation involvement in HbS polymerization^[210]1.
S1P, like other effectors of Hb, binds to multiple residues. Mutation
of one or multiple residues may result in destabilization of the Hb
tetramer. Such study is thus rarely used to ascertain the binding of an
effector, but instead structural and/or O[2] equilibrium studies
(Hb-O[2] binding studies) have been the norm. Importantly, our
structural study showing surface S1P binding only occurred in the
presence of 2,3-BPG binding in the central water cavity is highly
suggestive that the surface binding is specific. There are similar
reported studies where binding of allosteric effectors to Hb lead to
subtle but significant tertiary and/or quaternary structural changes at
the heme environment, α1α2 interface, α-cleft or
β-cleft^[211]33,[212]37,[213]38. Such changes have been used to explain
the differences in the allosteric activities of these effectors.
Notably, effectors that lead to increase in Hb affinity of O[2] show
more relaxed Hb structural features, while the opposite is true for
effectors that bind to Hb and decrease its O[2] affinity for
oxygen^[214]29.
In conclusion, we found that: S1P works collaboratively with 2,3-BPG to
cause further conformational changes and stabilize the 2,3-BPG-bound
deoxyHbS and HbA to a more enhanced T-state; deoxyHbA or deoxyHbS binds
to the membrane protein cdB3, promote release of GAPDH to cytosol and
thus channel glucose to glycolysis relative to PPP (Fig. [215]5i).
Altogether, our findings add significant new insight to erythrocyte
pathology and physiology and pave the way for novel therapeutic
interventions in SCD.
Methods
Mice
Berkeley SCD transgenic mice expressing exclusively human HbS were
purchased from The Jackson Laboratory (Bar Harbor, ME). Sphk1 ^−/− mice
were initially acquired from Dr. Richard L. Proia at the National
Institute of Diabetes and Digestive and Kidney Diseases, NIH (Bethesda,
MD) and bred in The University of Texas Health Science Center at
Houston. Eight-ten weeks old wild-type C57BL6/J mice were purchased
from The Jackson Laboratory. All protocols involving animal studies
were reviewed and approved by the Institutional Animal Welfare
Committee of The University of Texas Health Science Center at Houston.
All experiments were performed in accordance with relevant guidelines
and regulations from NIH and The University of Texas Health Science
Center at Houston.
Human subjects
Individuals with SCD were identified by hematologists on the faculty of
The University of Texas McGovern Medical School at Houston. Subjects
participating in this study had no blood transfusion for at least 6
months before blood samples were collected. Normal human subjects were
of African descent and were free of hematological disease. The research
protocol, which included informed consent from the subjects, was
approved by The University of Texas Health Science Center at Houston
Committee for the Protection of Human Subjects. Informed consent was
obtained from all participants and/or their legal guardians. All
experiments involved in human samples were performed in accordance with
relevant guidelines and regulations.
Blood collection and preparation
Mouse blood was collected with heparin or EDTA as an anti-coagulant and
centrifuged at 2,000 g for 5 min, followed by aspiration of plasma and
white interface. WT erythrocytes were washed once with 5X volume of PBS
before storing in the −80 °C freezer. Mature erythrocytes from SCD and
SCD/Sphk1 ^−/− mice were isolated using Percoll density centrifugation
media (GE Healthcare Life Sciences).
Isolation of total erythrocytes and treatment of mouse erythrocytes in vitro
Blood collected with heparin as an anti-coagulant was centrifuged at
2,000 g for 5 min at room temperature, followed by aspiration of plasma
and white interface. Packed mature RBCs were washed 3 times with
culture media (F-10 nutrients mix, Life Technologies Thermo Fisher
Scientific, Waltham, MA) and re-suspended to 4% hematocrit (HCT). One
ml of RBCs were added to each well of a 12-well plate and treated with
different concentrations of S1P (Sigma-Aldrich, St. Louis, MO) for
6 hours.
Sphk1 activity assay
Erythrocyte Sphk1 activity was measured using previously described
methods^[216]8. Briefly, RBCs were lysed in a pH7.4 Tris-HCl buffer
containing 1 mM EDTA, 1 mM β-Mercaptoethanol, 0.3% Triton X-100, 50%
glycerol and protease and phosphatase inhibitors. Then, the lysates
were assayed using 250 µM D-erythro-sphingosine in bovine serum albumin
(0.4%) and [γ-^32P]ATP (10 μCi, 20 mM) containing 200 mM MgCl[2].
Lipids were extracted and then resolved by TLC on silica gel G60 with
1-butanol/methanol/acetic acid/water (80:20:10:20, v/v). The plates
were then exposed to phosphor-imaging screening (Bio-Rad) and scanned
for radioactive signals as indications of the amount of S1^32P
synthesized.
Hemoglobin analysis by HPLC
Analysis of different hemoglobin variants was performed by HPLC using
Agilent 1100 series HPLC system (Agilent Technologies, CA) and PolyCAT
ATM weak cation-exchange column (100 × 4.6-mm, 3 μM, 1500 A; catalog
#104CT0315, PolyLC inc., Columbia, MD) as previously described^[217]39.
The chromatographic separation was achieved at 24 °C by a gradient
elution of the following mobile phases: mobile phase A contained 40 mM
Bis-Tris, 2 mM KCN, pH 6.5; mobile phase B contained 40 mM Bis-Tris,
2 mM KCN, 0.2 M NaCl, pH 6.8. Using a flow rate of 1 ml/min, HPLC
column was pre-incubated for 5 min with 18% mobile phase B before
sample application. Elution of sample was performed by increasing the
mobile phase B to 45% at 8 min, and to 100% at 12 min, then decreasing
it back to 18% at 13 min. The column was ready for next sample after
re-equilibrating with 18% mobile phase B for 5 min.
MetHb and COHb measurement
MetHb and COHb were measured using previously described
spectrophotometric methods^[218]40,[219]41. Briefly, oxyHb, COHb, and
MetHb standards were prepared from stock standard hemoglobin. Following
dilution to 1% (v/v) in distilled water, a 100% saturated solution of
oxyhemoglobin was obtained by bubbling oxygen for 10 min, excess O[2]
being removed by bubbling nitrogen for 5 min. Absorbance at wavelength
645 nm was used to calculate concentration of MetHb. For COHb, the
absorbance readings were taken at 420 and 432 nm wavelengths. The
absorbance ratio (Ar) was calculated as: Ar = A420/A432. Factors F1,
F2, and F3, were calculated by subjecting the 100% COHb and 100% O[2]Hb
samples to the same procedure. Thus, F1 = A(O[2]Hb)432/A(O[2]Hb)420,
F2 = A(COHb)432/A(O[2]Hb)420, and F3 = A(COHb)432/A(O[2]Hb)420. The %
COHb was calculated using the following equation:
[MATH: %COHb=[1−(Ar×F1)]/[Ar(F2−F1)−F3+1]. :MATH]
Glucose uptake assay
Blood was collected with heparin as an anticoagulant and centrifuged at
2,400 g for 5 min at room temperature, followed by aspiration of plasma
and buffy coat. Packed erythrocytes were purified using percoll
gradients to remove reticulocytes. The mature erythrocytes were washed
three times with PBS and re-suspended to 4% hematocrit. The uptake
assay started with transferring 54 µl of erythrocyte suspension to an
Eppendorf tube with 6 µl C[14]-glucose (PerkinElmer, Waltham, MA)
master mix (50 mM adenosine with 1 mCi∙ml^−1 C[14]-glucose in PBS) to
get a final glucose concentration of 5 mM. The uptake was performed for
up to 120 min and stopped by adding 100 ml cold stop solution (0.9%
saline), then centrifuged at 2,400 g for 5 min. The supernatant was
withdrawn and the erythrocyte pellet was lysed in 60 µl water and the
lysate was spread on a glass microfiber filter (GE Healthcare Life
Sciences, catalogue number: 1825-025), heat-dried for counting of C[14]
isotope using a scintillation counter (LKB WALLAC 1209 EACKBETA Liquid
Scintillation Counter, LKB Instruments, Victoria, Australia). Also,
54 µl of erythrocyte suspension (washed and re-suspended to 4%
hematocrit as mentioned above) was aliquoted for total protein
measurement using Pierce BCA Protein Assay kit (Thermo Scientific,
catalogue #: 23225, Rockford, IL, USA).
Metabolomics Profiling
Metabolomics extraction
RBCs (100 µl) and plasma samples (20 µl) were immediately extracted in
ice-cold lysis/extraction buffer (methanol:acetonitrile:water 5:3:2) at
1:9 and 1:25 dilutions, respectively. Samples were agitated at 4 °C for
30 min, and then centrifuged at 10,000 g for 15 min at 4 °C. Protein
pellets were discarded, and supernatants were stored at −80 °C prior to
metabolomics analyses^[220]42.
Metabolomics analysis
Ten µl of RBC extracts were injected onto a UHPLC system (Ultimate
3000, Thermo, San Jose, CA, USA) and run on a Kinetex XB-C18 column
(150 × 2.1mm, 1.7 µm particle size - Phenomenex, Torrance, CA, USA)
using a 3 min isocratic flow (5% acetonitrile, 95% water, 0.1% formic
acid) at 250 µl/min or a 9 min linear gradient (5–95% acetonitrile with
0.1% formic acid at 400 µl/min). The UHPLC system was coupled online
with a Q Exactive mass spectrometer (Thermo, Bremen, Germany), scanning
in Full MS mode (2 µscans) at 70,000 resolution in the 60–900 m/z
range, 4 kV spray voltage, 15 sheath gas and auxiliary gas, operated in
negative and then positive ion mode (separate runs). Calibration was
performed before each analysis using positive and negative ion mode
calibration mixes (Pierce, Rockford, IL, USA) to ensure sub ppm error
of the intact mass. Metabolite identifications were assigned using the
software Maven (Princeton, NJ, USA), upon conversion of raw files into
mzXML format through MassMatrix (Cleveland, OH, USA). The software
allows for peak picking, feature detection and metabolite assignment
against the KEGG pathway database. Assignments were further confirmed
against chemical formula determination (as gleaned from isotopic
patterns and accurate intact mass), and retention times against a >750
standard compound library (Sigma-Aldrich, St. Louis, MO, USA; IROA
Tech, Bolton, MA, USA)^[221]42.
Metabolic flux analysis
For the glucose flux experiment, RBCs were cultured in HEPES buffer
with 6 mM D-Glucose-1,2,3-^13C[3] (Sigma Aldrich)^[222]20,[223]43. RBCs
were extracted and processed as described above. Packed mature RBCs
were washed 3 times with HEPES buffer and re-suspended to 4% hematocrit
(HCT). One ml of RBCs was added to each well of a 12-well plate and
pretreated for 30 min before sample collection started. Flux analysis
was performed by determining the integrated peak areas of isotopologues
+2.0068 and +3.0102 Da of lactate, glucose, and G3P in negative ion
mode through the software Maven (Princeton, NJ, USA).
S1P quantification and sphingolipids analysis
Validation and quantitative analyses for sphingolipids and S1P were
performed using a Thermo Vanquish UHPLC system coupled to a Thermo Q
Exactive mass spectrometer and determined against commercial standard
compounds sphingosine 1-phosphate (>95% pure - no. S9666, Sigma
Aldrich, St. Louis, MO, USA) and sphingosine-1-phosphate-d7 (>99% pure
- no. 860659 P – Avanti Lipids Polar Inc, Alabaster, AL, USA) or
sphingosine/ceramide deuterated mixes (LM-6002 - Avanti Lipids Polar
Inc, Alabaster, AL, USA) within the linearity range, as determined
through external calibration curves across 5 orders of magnitude.
Samples were diluted 1:10 with methanol:acetonitrile:water (5:3:2)
containing 100 nM S1P-d7 or sphingosine/ceramide mixes, then agitated
and centrifuged as described above. Supernatant (10 µl per injection)
was analyzed using both a 4 and 9 minute gradient of 50–95%
acetonitrile containing 0.1% formic acid (400 µl/min) and a Kinetex C18
column (150 × 2.1 mm, 1.7 µm – Phenomenex) held at 35 °C. The mass
spectrometer was operated in positive ion mode at 70,000 resolution,
scan range of 90–1350 m/z, sheath gas 25, auxiliary gas 5.
Quantification was performed by exporting integrated peak area values
for endogenous and heavy S1P, sphingosine or ceramides. Absolute
quantitation was determined according to the formula: [light] = Peak
Area (Light)/(Heavy)*[Heavy]*dilution factor (10 for red blood cells,
25 for plasma). Results were imported into GraphPad Prism 5.0 (GraphPad
Software Inc., La Jolla, CA, USA) for statistical analysis (One way
ANOVA with Tukey multiple column comparison test; significance
threshold for p-values < 0.05).
Morphology study of erythrocytes
Blood smears were made using 1% glutaradehyde fixed cultured human or
tail blood from bone marrow transplanted mice. Blood smears were
stained by WG16-500ml kit (Sigma-Aldrich) for sickle cell. Blood smears
stained by these procedures were observed using the 40x objective of an
Olympus BX60 microscope. Areas where red blood cells do not overlap
were randomly picked, at least 10 fields were observed and 1000 red
blood cells including sickle cells were counted. The percentages of
sickle cells in red blood cells were calculated.
Hemolytic analysis
The hemoglobin in mouse plasma was quantified by ELISA kits following
instructions provided by the vendor (BioAssay Systems, Hayword, CA).
Mouse organ isolation and histological analysis
Mice were anesthetized and organs were isolated and fixed with 10%
paraformaldehyde in PBS overnight at 4 °C. Fixed tissues were rinsed in
PBS, dehydrated through graded ethanol washes, and embedded in
paraffin. 5μm sections were collected on slides and stained with
hematoxylin and eosin (H&E). The semi-quantitative analysis of
histological changes was conducted as previously described using a
computerized program^[224]7. Ten digital images were taken from each
H&E stained mouse tissue section at 20X magnification from different
areas. The congestion, necrosis or cysts on sections were identified
according to their structure and color. Briefly, the dark red color was
chosen for quantification of congestion and it was performed on 10
fields/mouse tissue sections at 20× magnification using software
analysis (Image Pro Plus 4.0; Media Cybernetics, Bethesda, MD, USA).
Additionally, the areas of necrosis in the livers and cysts in the
renal cortex were first manually marked by a magical pen tool available
in Adobe Photoshop Program. Then the quantification was conducted on 10
fields/mouse tissue sections at 20× using software analysis (Image Pro
Plus 4.0; Media Cybernetics, Bethesda, MD, USA). The whole areas of
each image were considered as 100%. The percentage of pathological
areas to whole area of image was recorded.
Measurement of life span of erythrocytes in SCD Tg mice
Erythrocytes were labeled in vivo by using N-hydroxysuccinimide (NHS)
biotin and the life span of circulating red blood cells was measured as
described^[225]7. Specifically, 50 mg/kg of NHS biotin was injected
into the retro-orbital plexus of SCD mice (prepared in 100 μl sterile
saline just prior to injection; initially dissolved at 50 mg/mL in
dimethyl sulfoxide). Blood samples (5 μl) were collected the first day
after biotin-injection from tail vein by venipuncture to determine the
percentage of erythrocytes labeled with biotin. Subsequently, 5 μl of
blood were obtained by tail vein venipuncture on day 1, 3, 5 and 7 for
measurement of biotinylated erythrocytes. The percentage of
biotinylated erythrocytes was calculated by determining the fraction of
peripheral blood cells labeled with Ter-119 (to identify erythrocytes)
that were also labeled with a streptavidin-conjugated fluorochrome by
flow cytometry
2,3-BPG analysis and erythrocyte O[2] release capacity (P50) measurement
RBC 2,3-BPG was isolated as indicated before and quantified by a
commercially available kit (Roche, Nutley, NJ)^[226]6. For P50
measurement in intact cells, 10 μl of whole blood aliquot were mixed
with 4.5 ml Hemox Buffer (TCS Scientific Corporation, PA), 10 μl
anti-foaming reagent ((TCS Scientific Corporation, PA) and 20 μl 22%
BSA in PBS; for P50 measurement of Hb with or without 2,3-BPG or S1P,
the system was prepared first as indicated above. Then, the mixture was
then injected in the Hemox Analyzer (TCS Scientific Corporation, PA)
for measurement of O[2] equilibrium curve at the temperature of 37 °C.
Measurement of NADPH
RBC NADPH was quantified by a commercially available kit
(Sigma-Aldrich). Briefly, 10 µl RBCs were used for each assay. NADPH
was extracted with 800 µl of NADP/NADPH Extraction Buffer and placed on
ice for 10 minutes, then centrifuged the samples at 10,000 g for
10 minutes to remove insoluble material. Then, 10KDa molecular weight
cut-off columns were used to filter out enzymes in the lysate. The
filtered solution was then applied to the measurement of NAPDH through
a chain colorimetric reaction and the read-outs were detected by
spectrophotometer.
Isolation of RBC cytoplasm and measurement of GAPDH activity
RBCs were lysed by freeze and thaw in 10 volume of 5 mM cold phosphate
buffer (pH 8.0) and vortexed. RBC membrane was removed by centrifuged
at 20,000 g for 20 minutes at 4°C. The supernatant was saved and used
to measure cytosolic GAPDH activity by KDalert GAPDH assay kit (Life
technologies).
Western Blot detection of GAPDH in erythrocytes
Pelleted erythrocytes were first frozen and then thawed in 20-fold
volume of 5 mM phosphate buffer containing 150 mM NaCl, protease
inhibitors (Roche) and phosphatase inhibitors (Sigma). Then, 200 µl
were isolated and used as total lysate. The rest were centrifuged with
20,000 g for 20 min. Supernatant were removed and pellets were washed
for four times before dissolving in the same buffer with 1% Triton
X-100. 50 µg of membrane protein and 150 µg total protein were loaded
for western blot detection of membrane bound and total GAPDH using
monoclonal anti-GAPDH antibody (Sigma-aldrich) (1:1000 in blocking
buffer).
Membrane bound heme measurement
Heme bound on membrane was measured previously described^[227]44 with
minimum modification. In brief, nonporous slica beads with 3.15 µm
diameter (Bangs Laboratories, IN) were pretreated as previously
described^[228]44. Human or mouse ghost cells were prepared as follow:
heparin-blood was centrifuged at 2,400 g for 5 minutes. The plasma and
buffy coat were removed. The pellet was washed twice with Phosphate
Buffered Saline (PBS). The cells were lysed in 5 mM phosphate buffer
(pH8.0), centrifuged at 18,000 g for 15 minutes. The supernatant was
removed and the pellet was washed in phosphate buffer for 7 times to
obtain ghost cells. The beads were coated by ghost cells to produce
inside-out membrane (IOM). The IOM was washed 6 times with 5 mM
phosphate buffer (PB, pH 8.0). Packed 5 × 10^9 IOM beads were added
100 μl 5 mM PB (pH7.4) with 100 μM hemoglobin, varied concentration
S1P. Beads were incubated at 37°C for 10 minutes, centrifuged at room
temperature for 1 minute at 500 g. The supernatant was transferred to
new tube for GAPDH activity assay. Pellet beads were washed 6 times
with PB (pH7.4). Beads were added to 100 µl of concentrated formic acid
(Sigma-Aldrich), vortexed for 5 minutes. The beads were centrifuged at
2000rpm for 2 minutes. 80 µl of the supernatant was transferred to a
new 1.5 ml tube and added 400 µl 5 M NaOH. The heme concentration was
determined at 398 nm wavelength as described and normalized to protein
concentration. Human Hb A was used as standards for heme assay.
S1P beads pull down assay
Two µg of total erythrocyte lysate from normal individuals was adjusted
to 100 µL using lysis buffer (20 mM PIPES, 150 mM NaCl, 1 mM EGTA, 1%
V/V Triton-X-100, 1.5 mM MgCl[2] and 1 mM Naorthovanidate, 0.1% SDS, 1X
protease inhibitors (Roche Applied) pH7.4). Approximately 100 µl of
various lipids conjugated to agarose beads including S1P-agarose beads,
lysophosphatic acid-beads or sphingosine-beads (Echelon Biosciences
Inc, Salt Lake City, UT) were washed twice with lysis buffer. The
lysates were incubated with beads overnight at 4°C with constant gentle
rotation. Protein-bound beads were washed by wash buffer (10 mM HEPES
pH 7.4, 150 mM NaCl, 0.25% NP-40) for 6 times. Washed beads were added
50 µL of 2× Laemmli buffer (Sigma-Aldrich) and heated at 100°C for
5 minutes. Beads were centrifuged at 5000 g for 5 minutes and
supernatants (eluted proteins) were separated by SDS-PAGE, transblotted
to nitrocellulose membrane. Sickle Hemoglobin on the membrane was
probed with anti-human hemoglobin antibody (Santa Cruz, CA).
Immunoreactive bands were visualized by ECL using secondary antibodies
conjugated with horseradish peroxidase and Super-signal West Pico
chemiluminesence substrate (Piere).
Crystal Structural studies
Freshly prepared solution of S1P in methanol was incubated with
deoxygenated Hb (40 mg/mL deoxyHb) with and without freshly prepared
solution of 2,3-BPG in water for 60 minutes at Hb tetramer: 2,3-BPG:S1P
molar ratio of 1:5:5 or Hb tetramer:S1P molar ratio of 1:5. The binary
(deoxyHb-S1P) and ternary (deoxyHb-S1P-2,3-BPG) complex solutions were
then crystallized with 0.2 M sodium acetate trihydrate, 0.1 M sodium
cacodylate trihydrate, pH 6.5 and 30% PEG 8000 using the batch method
as previously described^[229]45. Crystals were cryo-protected with
mother liquor and glycerol (3:1 ratio) prior to diffraction data
collection at 100 K with a Rigaku IV ++ image plate detector using a
CuKα X-rays (λ = 1.54 Å) from a MicroMax-007 source fitted with Varimax
Confocal optics (Rigaku, The Woodlands, TX). The two complexes
crystalized in orthorhombic space group P2[1]2[1]2, each with one
tetramer per asymmetric unit. The datasets were processed with the
d*trek software (Rigaku) and the CCP4 suite of programs^[230]46.
The deoxyHb-S1P structure was first determined using molecular
replacement method with Phenix v.1.8^[231]47, with the native deoxyHb
structure, deoxyHb (2DN2)^[232]28 and refined with both Phenix^[233]47
and the CNS programs^[234]38. Model building and correction were
carried out using the graphic program COOT^[235]48. The refined-S1P
structure was then used as a starting model to refine the
deoxyHb-S1P-BPG complex structure. The deoxyHb-S1P refines to
Rfactor/Rfree of 22.4/27.9% at 2.4 Å, while deoxyHb-S1P-2,3-BPG refines
to 18.2/21.1% at 1.8 Å. A significant number of the low-resolution
reflections in the ternary deoxyHb-S1P-BPG complex were characterized
by high mosaicity, which could have contributed to the large difference
in the Rfactor and Rfree. The atomic coordinate and structure factor
files have been deposited in the RCSB Protein Data Bank with accession
codes 5KSJ for deoxyHb-S1P and 5KSI for deoxyHb-S1P-2,3-BPG. Detailed
crystallographic and structural analysis parameters are reported in
Table [236]1.
Table 1.
Crystallographic data for deoxyHbA-S1P-2,3-BPG and deoxyHbA-S1P complex
structures.
deoxyHbA-S1P-2,3-BPG deoxyHbA-S1P
Data Collection Statistics
Space group P2[1]2[1]2 P2[1]2[1]2
Cell dimensions (Å) 95.94, 98.08, 65.14 97.56, 95.15, 64.98
Molecules/asymmetric unit 1 tetramer 1 tetramer
Resolution (Å) 29.42–1.80 (1.86–1.80) 29.33–2.40 (2.49–2.40)
No. of measurements 221938 (21321) 119348 (10341)
Unique reflections 54124 (5483) 24236 (2293)
I/sigma I 11.5 (3.4) 9.4 (3.1)
Completeness (%) 93.9 (96.4) 96.6 (96.8)
Rmerge (%)^a 7.6 (38.0) 12.0 (39.8)
Refinement Statistics
Resolution limit (Å) 29.42–1.80 (1.88–1.80) 29.08–2.40 (2.51–2.40)
Sigma cutoff (F) 0.0 0.0
No. of reflections 54123 (6869) 24093 (2969)
Rfactor (%) 18.1 (29.6) 22.4 (34.3)
Rfree (%)^b 21.8 (32.7) 27.9 (38.3)
Rmsd standard geometry
Bond-lengths (Å)/−angles (°) 0.010/1.5 0.000/1.6
Dihedral angles
Most favored/allowed regions 96.8/3.2 92.4/6.7
Average B-Factors
All atoms/Protein/Heme 23.5/19.3/16.9 43.2/42.6/41.6
Water/S1P/2,3-BPG 40.4/73.5/58.9 47.5/96.2
[237]Open in a new tab
Values in parentheses refer to the outermost resolution bin. ^a R
[merge] = Σ[hkl]Σ[i]|I [i](hkl) − |/Σ[hkl]Σ[i] I [i](hkl).
^bR[free] was calculated from 5% randomly selected reflection for
cross-validation. All other measured reflections were used during
refinement.
Statistical analysis
All data were presented as mean ± standard deviation and analyzed
statistically using GraphPad Prism 5 software (GraphPad Software). The
significance of differences among two groups was assessed using
Two-tailed Student’s t-test. Differences between the means of multiple
groups were compared by one-way analysis of variance (ANOVA) or two way
ANOVA, followed by a Turkey’s multiple comparisons test. A P value of
less than 0.05 was considered significant.
Electronic supplementary material
[238]SupplementaryInformation^ (1.1MB, pdf)
Acknowledgements