Glutathione depletion, pentose phosphate pathway activation, and hemolysis in erythrocytes protecting cancer cells from vitamin C-induced oxidative stress

Article abstract of DOI:10.1074/jbc.C116.748848

The discovery that oxidized vitamin C, dehydroascorbate (DHA), can induce oxidative stress and cell death in cancer cells has rekindled interest in the use of high dose vitamin C (VC) as a cancer therapy. However, high dose VC has shown limited efficacy in clinical trials, possibly due to the decreased bioavailability of oral VC. Because human erythrocytes express high levels of Glut1, take up DHA, and reduce it to VC, we tested how erythrocytes might impact high dose VC therapies. Cancer cells are protected from VC-mediated cell death when co-cultured with physiologically relevant numbers of erythrocytes. Pharmacological doses of VC induce oxidative stress, GSH depletion, and increased glucose flux through the oxidative pentose phosphate pathway (PPP) in erythrocytes. Incubation of erythrocytes with VC induced hemolysis, which was exacerbated in erythrocytes from glucose-6-phosphate dehydrogenase (G6PD) patients and rescued by antioxidants. Thus, erythrocytes protect cancer cells from VC-induced oxidative stress and undergo hemolysis in vitro, despite activation of the PPP. These results have implications on the use of high dose VC in ongoing clinical trials and highlight the importance of the PPP in the response to oxidative stress.

Full text of DOI:10.1074/jbc.C116.748848

JBC Papers in Press. Published on September 22, 2016 as Manuscript C116.748848
The latest version is at http://www.jbc.org/cgi/doi/10.1074/jbc.C116.748848
Hemolysis from VC-induced oxidative stress protection
Glutathione depletion, Pentose Phosphate Pathway Activation, and Hemolysis in Erythrocytes
Protecting Cancer Cells from Vitamin C Induced Oxidative Stress
Zhuzhen Z. Zhang1, Eunice E. Lee1, Jessica Sudderth2, Yangbo Yue1, Ayesha Zia3, Donald
Glass1,4, Ralph J. Deberardinis2,3,4, Richard C. Wang1
1Department of Dermatology, The University of Texas Southwestern Medical Center, Dallas, TX,
75390; 2Children’s Medical Center Research Institute, The University of Texas Southwestern Medical
Center, Dallas, TX 75390, USA; 3Department of Pediatrics, The University of Texas Southwestern
Medical Center, Dallas, TX 75390, USA; 4Eugene McDermott Center for Human Growth and Devel-
opment, The University of Texas Southwestern Medical Center, Dallas, TX 75390, USA
Running title: Hemolysis from DHA-induced oxidative stress protection
To whom correspondence should be addressed: R. Wang, Department of Dermatology, NL08.110FB,
UT Southwestern Medical Center, 5323 Harry Hines Blvd, Dallas, TX 75390-9069, Office:
+1-214-633-1860, Fax: +1-214-648-5554; Email: richard.wang@utsouthwestern.edu
Keywords: vitamin C, erythrocyte, oxidative stress, pentose phosphate pathway (PPP), cancer, glu-
cose-6-phosphate dehydrogenase (G6PD or G6PDH), glucose metabolism, metabolomics, hydrogen
peroxide
ABSTRACT
oxidants. Thus, erythrocytes protect cancer cells
from VC induced oxidative stress and undergo
hemolysis in vitro, despite activation of the PPP.
These results have implications on the use of
high dose VC in ongoing clinical trials and high-
light the importance of the PPP in the response
to oxidative stress.
The discovery that oxidized vitamin C, dehy-
droascorbate (DHA), can induce oxidative stress
and cell death in cancer cells has rekindled in-
terest in the use of high dose vitamin C (VC) as
a cancer therapy. However, high dose VC has
shown limited efficacy in clinical trials, possibly
due to the decreased bioavailability of oral VC.
Because human erythrocytes express high levels
of Glut1, take up DHA, and reduce it to VC, we
tested how erythrocytes might impact high dose
VC therapies. Cancer cells are protected from
VC-mediated cell death when co-cultured with
Consistent with its function as a potent reducing
agent essential for numerous biological reactions,
VC can be readily oxidized to dehydroascorbate
(DHA) in aqueous solution. DHA is transported
intracellularly by Glut1 (1) after which it is re-
physiologically relevant numbers of erythrocytes. cycled to reduced VC at the expense of cellular
Pharmacological doses of VC induce oxidative
stress, glutathione (GSH) depletion, and in-
creased glucose flux through the oxidative pen-
tose phosphate pathway (PPP) in erythrocytes.
Incubation of erythrocytes with VC induced
hemolysis, which was exacerbated in erythro-
cytes from G6PD patients and rescued by anti-
antioxidants, such as GSH (2,3). This paradoxi-
cal role for VC as a source of oxidative stress
has been found to be selectively toxic to some
cancer cells, particularly to those overexpressing
Glut1 (4,5). While the benefits of VC supple-
mentation as a cancer therapy have been incon-
sistent (6), higher doses of intravenous VC have
1
Copyright 2016 by The American Society for Biochemistry and Molecular Biology, Inc.

Hemolysis from VC-induced oxidative stress protection
shown more promise, and several clinical trials
are ongoing (7).
cumulated intracellular VC much more rapidly
than HCT116 cells (Figure 1A). Consistent with
the previous studies on the accumulation of
ascorbate in erythrocytes (8), we found that ac-
cumulation of VC by RBCs peaked rapidly (~1
minute) and decreased slightly over time con-
sistent with the slow release of recycled ascor-
bate back into the media. In contrast, the kinetics
of DHA uptake by HCT116 cells was delayed
and increased slowly over 30 minutes. The rapid
uptake and reduction of DHA by erythrocytes is
consistent with the undetectable levels of DHA
in healthy human serum (12).
Like many cancer cells, human erythrocytes
(RBCs) express very high levels of Glut1. The
abundance of the Glut1 transporter in erythro-
cyte membrane allows RBCs to mediate the
transport of glucose and DHA at rates that far
exceeds their capacity to utilize it (3). There is
substantial evidence that erythrocytes participate
in VC recycling in vivo (8,9). In addition, eryth-
rocytes have also been found to protect both tis-
sues and cells from H2O2 mediated damage
through their high-capacity redox systems (10).
We hypothesized that erythrocytes might protect
cancer cells from VC-mediated toxicity through
similar mechanisms. Indeed, we find that physi-
ologically relevant numbers of erythrocytes pro-
tected HCT116, A375, and SK-MEL-28 cells
from VC mediated toxicity. Erythrocytes incu-
bated with VC showed higher levels of oxidative
stress and shifted glucose metabolism towards
the oxidative PPP, which produces NADPH for
ROS mitigation. Finally, we find that VC in-
duced hemolysis of erythrocytes, which was ex-
acerbated by chemical or genetic inhibition of
G6PD and rescued by incubation with β mer-
captoethanol. This study broadens our current
understanding of VC metabolism by erythro-
cytes and has implications on the use and moni-
toring of VC in the clinical setting.
Given the rapid kinetics of DHA uptake by
erythrocytes, we hypothesized that RBCs might
protect cancer cells from VC mediated toxicity.
First, we determined the sensitivities of HCT116
and two BRAFV600E mutant melanoma cell
lines to VC. While HCT116 cells were the most
sensitive to VC, both BRAF mutant melanoma
cell lines were also sensitive to VC-induced cell
death at physiological glucose concentrations
(Figure 1B). The intravenous administration of
high dose ascorbate has the ability to increase
serum ascorbate to >25 mM (7). Thus, all three
mutant cells showed sensitivity to VC at phar-
macologically relevant doses. Next, these cancer
cells were co-cultured with RBCs and then
treated with VC. The addition of as few as ten
thousand RBCs significantly protected HCT116,
A375, and SK-MEL-28 cells from VC-induced
toxicity (Figure 1C, 1D). Higher numbers of
erythrocytes offered near complete protection of
all three cancer cell lines from VC-mediated
toxicity. These striking in vitro findings suggest
circulating erythrocytes might exert modifying
effects on VC toxicity in the in vivo setting.
RESULTS
Erythrocytes Protect Cancer Cells from VC
and H2O2 Induced Cell Death—The selective
toxicity of VC to KRAS and BRAF mutant colo-
rectal cancer cells is due, in part, to the high ex-
pression levels of Glut1 and preferential uptake
of oxidized DHA by the facilitative glucose
transporter (5). Consistent with their very high
expression of Glut1, erythrocytes have also been
shown to transport DHA efficiently (11). We
compared the kinetics of DHA uptake by eryth-
rocytes and HCT116 at physiological concentra-
tions of glucose and found that erythrocytes ac-
The protective effects of RBCs against oxi-
dative stress were not limited to VC. Consistent
with reports that RBCs can protect tissues from
hydrogen peroxide (H2O2)-induced tissue dam-
age (10), we found that RBCs could protect
cancer cells from H2O2 mediated cell death. Af-
ter first determining the sensitivity of each of the
2

Hemolysis from VC-induced oxidative stress protection
cell lines to H2O2, we also found that as few as
ten thousand RBCs could also protect cancer
cells from H2O2 toxicity (data not shown).
Moreover, the addition of both VC and H2O2
resulted in a synergistic toxicity to the cancer
cells lines. Co-culture with erythrocytes provid-
ed protection against this synergistic oxidative
stress (data not shown).
tone phosphate (DHAP), phosphoenolpyruvate
(PEP), and pyruvate) arise from flux through the
oxidative PPP and m+2 labeled metabolites,
from glycolysis (Figure 2C). Importantly, these
measurements underestimate total flux through
the oxidative PPP as they report only carbons
flowing back into the pools of glycolytic inter-
mediates as opposed to supplying ri-
bose-phosphate pools for nucleotide biosynthesis
and other branch pathways. In unperturbed
erythrocytes, ~4-8% of lower glycolytic inter-
mediates were labeled as m+1. After treatment
with VC, the fractional abundance of m+1 la-
beled products increased significantly, as much
as 3-fold for some metabolites (Figure 2D, sup-
plemental data). Similar changes were observed
Metabolomic Flux Analysis of RBCs Treat-
ed with VC—Despite the high redox capacity of
erythrocytes, the uptake and recycling of large
amounts of DHA might present an oxidative
burden to erythrocytes. To test this, erythrocytes
were incubated with the ROS sensitive dye,
2’,7’–dichlorofluorescin diacetate (DCFDA).
We found that VC increased the intracellular
ROS of erythrocytes in a dose dependent manner. in H2O2 treated cells. Analysis of other labeled
The amount of oxidative stress induced by VC
was comparable to similar concentrations of
H2O2 (Fig 2A). Consistent with GSH’s function
as a the principal cellular antioxidant responsible
for the recycling of DHA by erythrocytes (8), we
found that erythrocyte GSH levels decreased
significantly upon incubation with VC (Figure
2B).
isotopologues revealed significant decreases in
the levels of m+2 labeled isotopologues con-
sistent with decreases of up to 20% in the gly-
colytic pathway after VC treatment (Figure 2E,
supplemental data). Interestingly, changes in
glucose flux were actually less pronounced in
RBCs treated with 1.0 mM VC or 0.1 mM H2O2
than those treated with 0.2 mM VC. High levels
of oxidative stress can overwhelm the antioxi-
dant systems of RBCs and compromise RBC
membrane integrity and metabolism (15). Thus,
we speculate that high levels of oxidant stress
may irreparably damage RBC and render them
metabolically inactive.
The regeneration of GSH from oxidized
glutathione (GSSG) requires NADPH and GSH
reductase. Keratinocytes, fibroblasts, and cancer
cells can respond rapidly to oxidative stress by
routing glucose flux through the oxidative pen-
tose phosphate pathway (PPP) to generate
NADPH (13,14). Because erythrocytes lack or-
ganelles to mount a transcriptional response to
oxidative stress, the reduction of NADP+ to
NADPH might constitute an especially im-
portant facet of the response against RBC oxida-
tive stress. DHA has previously been shown to
stimulate oxidation of glucose through the pen-
tose phosphate pathway (9). To quantitate the
impact of oxidative stress on glucose flux in
RBCs, we used [1,2-13C]glucose to monitor the
labeling of glucose-derived metabolites. In this
labeling scheme, m+1 labeled metabolites from
the lower half of glycolysis (i.e., dihydroxyace-
Hemolysis in Erythrocytes Treated with
VC—Incubation of RBCs with VC results in an
induction of oxidative stress, depletion of GSH,
and increased flux through the PPP. These find-
ings are consistent with the PPP’s key role in
producing NADPH for RBCs, and suggest that
inhibition of the PPP could damage VC-treated
RBCs. Although several cases of VC-induced
hemolysis have been reported in the literature
(16-18), the mechanism of this hemolysis has
not been described in the literature. Importantly,
all patients reported to have significant
VC-induced hemolysis were found to possess a
3

Hemolysis from VC-induced oxidative stress protection
genetic deficiency of glucose-6-phosphate dehy-
drogenase (G6PD), a key upstream enzyme of
the enzyme of the PPP. First, we tested whether
VC might induce hemolysis in normal erythro-
cytes in vitro. Indeed, we found that pharmaco-
logical concentrations of VC induced significant
hemolysis as measured by both hemoglobin re-
lease and decreased RBC number, with complete
hemolysis noted at 1.6 mM VC (Figure 3A). At
lower concentrations of VC, hemolysis was
more striking at later time points, likely reflect-
ing the depletion of glucose from the incubation
media (Figure 3B). Consistent with an important
role for the PPP in maintaining RBC redox, de-
hydroepiandrosterone (DHEA), a PPP inhibitor,
significantly exacerbated VC-induced RBC he-
molysis under all conditions (Figure 3A, 3B).
Moreover, erythrocytes from a G6PD patient
showed increased oxidative stress both at base-
line and also in response to VC and H2O2 treat-
ments (Figure 3C). G6PD erythrocytes were also
significantly more susceptible to VC induced
hemolysis (Figure 3D). The incubation of nor-
mal RBCs with β-mercaptoethanol (β-ME), an
antioxidant, partially rescued RBCs from both
VC-induced oxidative stress (Figure 3E) and
hemolysis (Figure 3F).
low levels of hemolysis might even be detectable
in otherwise healthy patients. To test this possi-
bility, clinical monitoring of hemolytic parame-
ters could be considered in future or ongoing
clinical trials. Moreover, our studies suggest that
VC alone may have limited utility in blood stor-
age where it has been tested extensively (20,21).
Rather, preservation of erythrocytes in solutions
with glucose and alternative anti-oxidants might
better prevent RBC oxidative stress and hemoly-
sis and promote longer-term erythrocyte stability
during storage.
Our results also highlight the importance of
glucose flux through the oxidative PPP as a crit-
ical component in the response to oxidative
stress. While many studies have focused on glu-
cose’s roles as an energy source and biosynthetic
intermediate, more recent studies have begun to
probe the role of glucose flux through the PPP in
the response to oxidative stress (13,22). We
speculate that the dependence of RBCs on the
PPP in the response to oxidative stress might
help explain why they have evolved to express
extraordinarily high levels of the Glut1 trans-
porter. It will be interesting to test whether other
cell types that express high levels of glucose
transporters show a similar dependence on glu-
cose in the response to oxidative stress.
DISCUSSION
Our findings that erythrocytes can protect
cancer cells from VC induced oxidative stress
extend previous studies suggesting an antioxi-
dant role for erythrocytes. While most clinical
trials involving high-dose VC include G6PD
deficiency as exclusion criteria, we suggest that
all patients at risk of hemolysis be excluded
from such trials. Consistent with this suggestion,
a recent randomized, double-blind, placebo con-
trolled trial found that oral supplementation of
VC significantly worsened hemolytic markers in
sickle cell anemia and β-thalassemia patients,
contrary to their expectations (19). While the
presence of sufficient serum glucose might mit-
igate hemolysis in ongoing high-dose VC trials,
EXPERIMENTAL PROCEDURES
Cell lines, Cell culture, RBC Co-culture,
and Treatments—A375 and SK-MEL-28 cell
lines were cultured in DMEM (Gibco 11965)
supplemented with 10% FBS and 1% Penicil-
lin-Streptomycin. HCT116 cells were cultured in
the same medium with 5% FBS. For VC and/or
H2O2 treatment, cells were seeded at a density of
5000 cells/well in a 96-well plate in 100 µl
DMEM (Gibco A14430) supplemented with 5
mM glucose and 10% heat inactivated FBS. Af-
ter 36 hours, 100 µl of fresh medium containing
VC and/or H2O2 was added to the existing cul-
ture, and cells were cultured for an additional 24
hours. For co-culture experiments, human RBCs
4

Hemolysis from VC-induced oxidative stress protection
(RBCs) were added to the culture just prior to
supplementation with VC and/or H2O2. After 24
hours, the RBCs were gently washed out before
measuring cell viability. Cell viability was de-
termined through the Dimethylthiazolyl Diphe-
(DCFDA) staining. For H2O2 treatment, 5 µM
DCFDA was first added to the cells, followed
quickly by the addition of different doses of
H2O2; samples were collected after 30 min. For
VC treatment, VC was first added to the RBCs,
nyltetrazolium Bromide MTT (ThermoScientific)
assay according to the manufacturer’s instruc-
tions. Cancer cells cultured in the absence of
RBCs and treatments were used as the reference
to calculate the relative survival of the different
experimental conditions. Cell viability was also
separately determined by counting cells that
were seeded at a density of 1×105 cells/well in a
12-well plate in 1 ml of medium. Cells were
trypsinized and counted in the presence of Try-
pan Blue. For β-mercaptoethanol (β−ME) rescue
experiments, 0.5 mM VC was added first to the
RBCs, followed quickly by the indicated amount
of β−ME. Because the toxicity of VC varied
between 96-well and 12-well plates, the concen-
tration of VC used in MTT and cell counting
experiments were independently determined.
Patient Enrollment and Preparation of
RBCs—Erythrocyte analyses were approved by
the institutional review board, and written in-
formed consent was received from participants
prior to inclusion in the study. The G6PD patient
had a documented G6PD deficiency of 0.8 IU/g
Hb (reference: 8.8 to 13.4). Venous blood was
collected from volunteers in heparinized tubes;
anticoagulated blood was centrifuged at 500g for
10 min at 4°C and the leukocytes were aspirated.
The RBC pellet was then washed three times
with DPBS under the same conditions. Washed
cells were resuspended in PBS at ~80% hemato-
crit and kept at 4°C until use. RBCs were used
on the day of collection.
after a 2 hr incubation, 5µM DCFDA was added
and cells were collected after 30 min. For collec-
tion, the samples were placed on ice, centrifuged
at 1500 rpm for 4min, gently washed once with
FACS buffer (DPBS with 2% FBS and 0.2 mM
EDTA), and resuspended in FACS buffer for
analysis. For cancer cell ROS measurements,
cells were seeded at a density of 1 × 105
cells/well in a 12 well plate in the same medium
as described for VC and H2O2 treatment. After
24 hours, 1×106 RBCs were added to indicated
wells. Next, H2O2 was added to the cells, fol-
lowed quickly by the addition of DCFDA. Can-
cer cells were collected after 30 min for FACS
analysis. For intracellular GSH measurement,
RBCs were treated with VC or H2O2 for 120 or
30 min respectively, quickly collected on ice,
gently washed once with ice-cold culture medi-
um, and then lysed in 200 µl GSH assay buffer.
GSH was measured using the GSH Colorimetric
Assay Kit (Biovison) following the manufactur-
er’s instructions.
[1,2-13C]glucose Flux Analysis—2×107
RBCs were added to 5ml of medium supple-
mented with 5 mM [1,2-13C]glucose or unla-
beled glucose and 10% dialyzed FBS with or
without vitamin C or H2O2. RBCs were carefully
dispersed after addition to the media. After 6
hours, cells were collected by centrifugation at
1500rpm for 10 min at 4°C. The media was as-
pirated and the cells frozen in liquid nitrogen,
lysed with three freeze thaw cycles in 1 mL of
50% methanol (MeOH), then centrifuged to re-
move protein debris. The lysates were dried,
methoximated and derivatized with 10mg/ml
methoxyamine (Fisher PI-45950) at 70oC for 10
min followed by 105 ml MTBSTFA (Sigma
394882) at 70oC for 1 hour. After derivitization,
1 ml of the sample was injected onto an Agilent
Intracellular ROS and GSH assay—2×107
RBCs were plated in 6 well plate in 2ml DMEM
(Gibco A14430) supplemented with 5 mM Glu-
cose and 10% heat inactivated FBS at 37°C in at
5% CO2. ROS levels in RBCs were quantified
by BD FACSCalibur flow cytometer us-
ing 2',7'-dichlorodihydrofluorescein
diacetate
5

Hemolysis from VC-induced oxidative stress protection
6970 gas chromatograph networked to an Ag-
ilent 5973 mass selective detector. The abun-
dance of the following ions were monitored: m/z
484-487 for DHAP, m/z 261-264 for lactate, m/z
453-456 for PEP and m/z 174-177 for pyruvate.
DHA uptake assay—Transport measure-
ments were performed as previously described
with modifications (11,23). Ascorbic acid,
L-[1-14C] (ARC 1569) was purchased from
American Radiolabeled Chemicals. Labeled
ascorbic acid purchased from PerkinElmer did
not perform as reported in control uptake assays.
HCT116 cells were seeded in triplicate in
12-well plates (250,000 cells/well) for 18-24 hrs,
washed twice with PBS, and incubated with se-
action. Uptake was initiated by adding 100 µl of
Ringer solution containing 0.5 µCi and 100 µM
cold AA to each sample for the indicated time
points. Transport was terminated by the addition
of ice-cold stop solution. Cells were centrifuged
at 16,000xg for 1 min followed by three addi-
tional washes. Both HCT116 and pelleted eryth-
rocytes were lysed in 0.5ml of 0.2% SDS. 375
µl of the lysate was quantitated by liquid scintil-
lation counting.
RBC hemolysis assay—Washed RBCs were
resuspended in Ringer’s solution at a cell density
of 1×107 cells/ml. RBC suspensions were prein-
cubated with 25 µM DHEA or DMSO control at
37°C with shaking for 15 min prior to treatment
rum-free DMEM containing 0.1% BSA for 2 hrs. with VC. For antioxidant rescue experiments,
Cells were then washed and incubated in a
Ringer’s solution (126 mM NaCl, 5 mM KCl, 1
mM MgSO4, 32 mM HEPES, 2.5 mM glucose, 1
mM CaCl2, 0.1% BSA, pH 6.0). Ascorbate oxi-
dase (10U/ml) was used to convert [14C]-VC to
[14C]-DHA, and transport was initiated by the
addition of 0.5 µCi of [14C]AA and 100 µM cold
AA to each well. Uptake assays were performed
for 0, 1, 3, 9, and 27 min at RT. Transport was
terminated by the removal of uptake solution
followed by 4 washes with ice-cold stop solution
containing 100 µM phloretin and 10 µM cyto-
chalasin B in PBS. For DHA uptake in RBCs, 20
µl of erythrocyte suspension containing 2.5×106
cells in Ringer’s solution was used for each re-
RBCs were preincubated with Ringer’s solution
supplemented with the indicated amounts of
β–ME. Aliquots of 2 ml RBC suspension con-
taining 1×107 cells/ml were used in each repli-
cate. RBC suspensions were incubated with var-
ious concentrations of freshly prepared VC at
37°C for indicated time points. For time course
experiments, 0.6 mM VC was used for all time
points. RBC hemolysis was determined by both
absorbance at 540 nm and cell count from the
same sample. Absorbance at 540 nm from he-
moglobin released into RBC supernatants was
normalized to the absorbance of RBCs com-
pletely hemolysed with 3 mM VC (100% he-
molysis).
Acknowledgments: We thank the volunteers for their participation and Amanda Ziegeweid for her
help in recruiting patients. This work was supported by R01 CA157996 to RD; K23 AR069728 to DG;
K23 HL132054 to AZ; and K08 CA164047, Burroughs Wellcome Fund CAMS (1010978), and a
Disease Oriented Clinical Scholar Awards to RW.
Conflict of Interest: The authors declare that they have no conflicts of interest with the contents of
this article.
Footnote: The content is solely the responsibility of the authors and does not necessarily represent the
official views of the National Institutes of Health.
6

Hemolysis from VC-induced oxidative stress protection
Author contributions: ZZ, EL, RD, and RW designed the experiments; ZZ, EL, and JS performed
experiments; ZZ, EL, JS, and YY acquired data; ZZ, EL, JS, RD, and RW analyzed data; DG and AZ
enrolled volunteers; ZZ and RW wrote the manuscript with contributions from EL, JS, and RD.
7

Hemolysis from VC-induced oxidative stress protection
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Hemolysis from VC-induced oxidative stress protection
Abbreviations: BRAF, Proto-oncogene v-Raf murine sarcoma viral oncogene homolog B; DCFDA,
Dichlorofluorescin diacetate; DHA, Dehydroascorbate; DHAP, Dihydroxyacetone phosphate; DHEA,
Dehydroepiandrosterone; Glut1, Solute carrier family 2, facilitated glucose transporter member 1;
GSSG, Glutathione disulfide; KRAS, Proto-oncogene V-Ki-ras2 Kirsten rat sarcoma viral oncogene
homolog; PEP, Phosphoenolpyruvate; PPP, Pentose Phosphate Pathway; VC, Vitamin C
FIGURE LEGENDS
FIGURE 1. Erythrocytes protect cancer cells from VC and H2O2 toxicity. (A) 14C-dehydroascorbate
(DHA) uptake assay demonstrates that RBCs accumulate DHA more rapidly than HCT116 cells. (B)
HCT116 and BRAF mutant melanoma cells (A375 and SK-MEL-28) show a dose dependent toxicity
to VC mediated toxicity as assessed by MTT. (C) Photomicrographs of A375 cells reveals that VC
treatment (0.6 mM for 24 hrs) caused cytotoxicity. Co-culture with RBCs (indicated by arrows) res-
cues cell viability. (D) The co-culture of RBCs with HCT116, A375, and SK-MEL-28 cancer cells
protects them from VC-induced toxicity as assessed by MTT assay. An equal number of cells cultured
without VC or RBCs was used as the reference (100%) to calculate relative survival (n=3, error bars =
SD; ANOVA with Dunnett’s correction; * P≤0.05, ** P≤0.01, ***P≤0.001, ****P≤0.0001). (E) The
co-culture of RBCs with HCT116, A375, and SK-MEL-28 protects them from VC-induced toxicity as
assess by cell number.
FIGURE 2. Erythrocytes show increased oxidative stress, decreased GSH, and increased glucose flux
through the oxidative PPP after treatment with VC. (A) FACS analysis of erythrocytes treated with
increasing amounts of VC (0, 0.1, 0.2, 0.4, 0.6, and 1 mM) or H2O2 (0, 0.05, 0.25, 1 mM) show in-
creased levels of ROS as indicated by DCFDA fluorescence. The grey curve represents a control
without DCFDA. (B) Intracellular GSH decreases after incubation with VC. (C) Schematic of
[1,2-13C]glucose metabolic tracing experiments. Some of the glucose proceeding through the PPP re-
sult in the generation of m+1 labeled products. However, some products of the oxidative PPP will not
be detected as m+1 labeled products (dashed arrows). Red boxes highlight the subset of monitored
products. (D) Quantitation of labeled products in metabolic tracing experiments. Incubation of eryth-
rocytes with VC or H2O2 results in significant increases of m+1 labeled products, indicating increased
flux through the PPP (top row), and significant decreases of m+2 labeled products, indicating de-
creased flux through glycolysis (bottom row). (n=3, error bars = SD; ANOVA with Dunnett’s correc-
tion; * P≤0.05, ** P≤0.01, ***P≤0.001, ****P≤0.0001 when compared to control).
FIGURE 3. VC induced hemolysis is exacerbated by G6PD inhibition. (A) Dose dependent hemoly-
sis of RBCs by VC as assessed by hemoglobin absorbance (left) and cell counts (right) after 1.5 hours
of incubation. Dehydroepiandrosterone (DHEA) significantly exacerbated hemolysis as indicated by
red asterisks (n=3, error bars = SD; t-test with Holm-Sidak correction, alpha=0.05). Data represent
replicate measurements that are representative of >3 independent experiments. (B) Incubation of
RBCs with 0.6 mM VC resulted in extensive hemolysis at 8 hrs. DHEA significantly exacerbated
hemolysis at 8 hrs (n=3, error bars = SD; t-test with Holm-Sidak correction, alpha=0.05). (C) RBCs
from a G6PD patient showed higher levels of ROS at baseline, when treated with VC (0.1 and 0.2
mM), and when treated with H2O2 (0.1, 0.2 mM) compared with control RBCs as indicated by
DCFDA fluorescence. (D) RBCs from a G6PD patient showed significantly higher levels of hemoly-
10

Hemolysis from VC-induced oxidative stress protection
sis in response to incubation with VC (1.0 mM, 1.5 hrs) compared to control RBCs (n=3, t-test,
P≤0.0001). (E) Incubation of RBCs with β-mercaptoethanol rescued VC-induced oxidative stress. (F)
Incubation of RBCs with β-ME resulted in a dose-dependent rescue of hemolysis (right) (n=3, t-test,
P≤0.0001).
11

Hemolysis from VC-induced oxidative stress protection
FIGURE 1
12

Hemolysis from VC-induced oxidative stress protection
FIGURE 2
13

Hemolysis from VC-induced oxidative stress protection
FIGURE 3
14

Glutathione Depletion, Pentose Phosphate Pathway Activation, and Hemolysis in
Erythrocytes Protecting Cancer Cells from Vitamin C Induced Oxidative Stress
Zhuzhen Z. Zhang, Eunice E. Lee, Jessica Sudderth, Yangbo Yue, Ayesha Zia, Donald
Glass, Ralph J. DeBerardinis and Richard C. Wang
J. Biol. Chem. published online September 22, 2016
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