Hyperpolarized [1-13C]-ascorbic and dehydroascorbic acid: Vitamin C as a probe for imaging redox status in vivo

Article abstract of DOI:10.1021/ja2045925

Dynamic nuclear polarization (DNP) of ¹³C-labeled metabolic substrates in vitro and their subsequent intravenous administration allow both the location of the hyperpolarized substrate and the dynamics of its subsequent conversion into other metabolic products to be detected in vivo. We report here the hyperpolarization of [1-¹³C]-ascorbic acid (AA) and [1- ¹³C]-dehydroascorbic acid (DHA), the reduced and oxidized forms of vitamin C, respectively, and evaluate their performance as probes of tumor redox state. Solution-state polarization of 10.5 ± 1.3% was achieved for both forms at pH 3.2, whereas at pH 7.0, [1-¹³C]-AA retained polarization of 5.1 ± 0.6% and [1-¹³C]-DHA retained 8.2 ± 1.1%. The spin-lattice relaxation times (T₁'s) for these labeled nuclei are long at 9.4 T: 15.9 ± 0.7 s for AA and 20.5 ± 0.9 s for DHA. Extracellular oxidation of [1-¹³C]-AA and intracellular reduction of [1-¹³C]-DHA were observed in suspensions of murine lymphoma cells. The spontaneous reaction of DHA with the cellular antioxidant glutathione was monitored in vitro and was approximately 100-fold lower than the rate observed in cell suspensions, indicating enzymatic involvement in the intracellular reduction. [1-¹³C]-DHA reduction was also detected in lymphoma tumors in vivo. In contrast, no detectable oxidation of [1-¹³C]-AA was measured in the same tumors, consistent with the notion that tumors maintain a reduced microenvironment. This study demonstrates that hyperpolarized ¹³C-labeled vitamin C could be used as a noninvasive biomarker of redox status in vivo, which has the potential to translate to the clinic.

Full text of DOI:10.1021/ja2045925

ARTICLE
pubs.acs.org/JACS
13
Hyperpolarized [1- C]-Ascorbic and Dehydroascorbic Acid: Vitamin C
as a Probe for Imaging Redox Status in Vivo
Sarah E. Bohndiek, Mikko I. Kettunen, De-en Hu, Brett W. C. Kennedy, Joan Boren, Ferdia A. Gallagher,
and Kevin M. Brindle*
Department of Biochemistry, University of Cambridge, 80 Tennis Court Road, Cambridge CB2 1GA, U.K., and Cancer Research UK
Cambridge Research Institute, Li-Ka Shing Centre, Robinson Way, Cambridge CB2 0RE, U.K.
S Supporting Information
b
13
ABSTRACT: Dynamic nuclear polarization (DNP) of C-labeled
metabolic substrates in vitro and their subsequent intravenous admin-
istration allow both the location of the hyperpolarized substrate and the
dynamics of its subsequent conversion into other metabolic products to
13
be detected in vivo. We report here the hyperpolarization of [1- C]-
13
ascorbic acid (AA) and [1- C]-dehydroascorbic acid (DHA), the
reduced and oxidized forms of vitamin C, respectively, and evaluate
their performance as probes of tumor redox state. Solution-state
13
polarization of 10.5 ( 1.3% was achieved for both forms at pH 3.2, whereas at pH 7.0, [1- C]-AA retained polarization of 5.1
1
3
(
1
0.6% and [1- C]-DHA retained 8.2 ( 1.1%. The spinꢀlattice relaxation times (T 's) for these labeled nuclei are long at 9.4 T:
1
13
13
5.9 ( 0.7 s for AA and 20.5 ( 0.9 s for DHA. Extracellular oxidation of [1- C]-AA and intracellular reduction of [1- C]-DHA
were observed in suspensions of murine lymphoma cells. The spontaneous reaction of DHA with the cellular antioxidant glutathione
was monitored in vitro and was approximately 100-fold lower than the rate observed in cell suspensions, indicating enzymatic
13
involvement in the intracellular reduction. [1- C]-DHA reduction was also detected in lymphoma tumors in vivo. In contrast, no
13
detectable oxidation of [1- C]-AA was measured in the same tumors, consistent with the notion that tumors maintain a reduced
13
microenvironment. This study demonstrates that hyperpolarized C-labeled vitamin C could be used as a noninvasive biomarker of
redox status in vivo, which has the potential to translate to the clinic.
’
INTRODUCTION
Dynamic nuclear polarization (DNP) of C-labeled cell
substrates enhances their sensitivity to detection in vivo by over
0,000-fold. DNP has shown great promise for metabolic
imaging of cancer in preclinical studies, with a wide range of
in vitro, but its application in vivo is likely to be limited by a lack
12 13
13
of sensitivity. To be used in vivo, C-labeled DNP substrates
must exhibit long spinꢀlattice relaxation times (T 's), be rapidly
1
transported (or react in the extracellular space), and produce an
observable chemical shift change between the injected substrate
and its product, to allow spectroscopic imaging of both metabo-
lites. The chemical and biological properties of vitamin C make it
a prime candidate for in vivo detection of redox status: the C1
1
1
13
substrates now available including hyperpolarized [1- C]-pyr-
uvate, which enables early detection of tumor response to drug
4
2
,3
treatment and noninvasive assessment of tumor grade;
1
3
carbonyl group has a long T , its oxidation produces a chemical
1
[2- C]-fructose, phosphorylation of which is influenced by both
5
13
shift difference of ∼5 ppm, the oxidized form is transported
glycolytic and pentose phosphate pathway activity; [1,4- C ]-
fumarate, which is emerging as a marker of cell death following
chemotherapy; and [ C]-bicarbonate, which allows assess-
ment of tumor pH. Although these hyperpolarized substrates
probe a wide range of biological processes, there are currently no
hyperpolarized substrates that report directly on tumor redox
status. Rapid cancer cell proliferation promotes intracellular
accumulation of reducing equivalents such as glutathione and
2
13
rapidly by the same membrane transporters utilized by glucose,
2
,6
13
and its redox state is coupled to two primary cellular antioxidants,
14
7
glutathione and NADPH.
The reduced form of vitamin C, ascorbic acid (AA), is an
essential water-soluble antioxidant and cofactor of numerous
14
enzymes involved in biosynthetic reactions. AA is reversibly
oxidized (Figure 1), forming the stable ascorbic free radical
(AFR) intermediate with the loss of one electron or dehydroas-
corbic acid (DHA) with the loss of two electrons. AFR decays by
disproportionation to AA and DHA, whereas DHA sponta-
neously hydrolyses under physiological conditions, with the open-
8
NADPH to counteract the effects of reactive oxygen species
9
(
ROS) and to maintain redox balance. The ability of certain
cancer cells to maintain a highly reduced intracellular environ-
ment is strongly correlated with aggressiveness and drug
1
5
1
0,11
ing of the lactone ring to form diketogulonic acid (DKG).
resistance.
Noninvasive imaging of tumor redox status may
therefore indicate prognosis and give an early readout of treat-
1
3
ment response. Recently, C-benzoylformic acid (BFA) was
Received: May 19, 2011
Published: June 21, 2011
shown to report specifically on hydrogen peroxide concentration
r 2011 American Chemical Society
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|J. Am. Chem. Soc. 2011, 133, 11795–11801

Journal of the American Chemical Society
ARTICLE
Figure 1. Primary intracellular oxidation, reduction, and degradation pathways for vitamin C. Enzymes involved in GSH- and NADPH-dependent
1
4
reduction of dehydroascorbic acid (DHA) are highlighted, with the primary cytosolic enzymes in bold. Abbreviations: AA, ascorbic acid; AFR,
ascorbate free radical (also known as semidehydroascorbic acid); DHA, dehydroascorbic acid; DKG, diketogulonic acid; GSH, reduced glutathione;
GSSG, oxidized glutathione.
through NADPH-dependent reactions, primarily involving the
14
enzyme thioredoxin reductase (K = 0.7 mM). Glutathione-
m
dependent enzymes include the thiol-disulfide oxidoreductases,
glutaredoxin and protein disulfide isomerase, while the predomi-
nant NADPH-dependent enzyme is the selenoprotein, thiore-
doxin reductase (Figure 1). In both cases, rapid intracellular
reduction of DHA will maintain a favorable gradient for
19
transport.
The upregulation of the glutathione and thioredoxin antiox-
idant systems in tumors has been implicated in both chemore-
10,24ꢀ26
sistance and increased metastatic potential.
While the
dominant mechanism of DHA reduction remains unresolved in
cancer cells, NADPH provides the reducing power for both of
these systems, and therefore the rate of intracellular reduction of
DHA is determined ultimately by the availability of NADPH and,
in turn, pentose phosphate pathway activity, reflecting oxidative
2
7,28
Figure 2. Mechanism of vitamin C transport, in both reduced (AA) and
oxidized (DHA) forms, by sodium-dependent vitamin C transporters 1
and 2 (SVCT 1, 2) and glucose transporters (primarily GLUT1, 3 and
also GLUT 4), respectively. This is dominated in cancer cells by GLUT1
transport of DHA, with SVCTs either not expressed or having low
stress.
Furthermore, oxidative stress is associated with almost
all major diseases and in addition to cancer has been implicated in
numerous neurodegenerative and cardiovascular conditions.
Oxidative stress has been assessed in the clinic, at a systemic level,
from measurements of plasma ubiquinol and ubiquinone and
from measurements of serum antioxidants and the presence of
oxidized protein and lipid. Although imaging regions of
oxidative stress in tissues should provide a more sensitive readout
in a range of diseases, this has been limited to date by the lack of
available probes and there are currently no methods available to
image the spatial distribution of redox state in the clinic. Imaging
1
1
2
9
19,21
activity due to the acidic extracellular pH in tumors.
30
While epithelial cells of the intestine, liver, and kidney can
transport AA directly through sodium-dependent vitamin C
16
transporters (SVCT1, 2 ), the primary means by which most
tissues acquire vitamin C is through transport of DHA, which
forms a stable bicyclic hydrated hemiacetal structure in aqueous
measurements of nitroxide reduction, either directly using EPR
imaging
1
7,18
31,32
solution.
Studies in tumor cell lines have found that they
or indirectly from T -weighted MRI measurements,
1
33
either lack the capacity to transport AA, or if they do have
the capability, the rate of uptake of DHA is at least an order
have shown great promise in preclinical studies, but these
techniques have yet to translate to the clinic. The rapid inter-
conversion of the reduced and oxidized forms of vitamin C
in vivo makes them ideal candidates for hyperpolarized redox
imaging, with AA primarily an extracellular probe and DHA an
intracellular redox probe. The goal of this study was to evaluate
19
of magnitude faster. DHA is taken up by the glucose
1
3,20
transporters
.7 mM) (Figure 2), which display a similar affinity for both
glucose and DHA and are overexpressed in most tumors
GLUT1 (K = 1.1 mM) and GLUT3 (K =
m m
1
21
13
13
compared to normal tissue. This phenomenon is widely
exploited in detection and staging of tumors by [ F]-fluoro-
deoxyglucose positron emission tomography (FDG PET).
the performance of [1- C]-AA and [1- C]-DHA as hyperpo-
larized imaging probes using cancer models in vitro and in vivo.
18
9,22
Once inside the cell, DHA must be reduced rapidly to avoid
hydrolysis and maintain antiscorbutic activity. This can occur
spontaneously or enzymatically, by reaction with glutathione
’
METHODS
2
3
13
13
Hyperpolarization of [1- C]-Ascorbic Acid and [1- C]-
Dehydroascorbic Acid. A 1.5 M solution of [1- C]-AA (Omicron
1
3
(GSH) or GSH-dependent enzymes (Km = 2.2 mM) and
1
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Journal of the American Chemical Society
Biochemicals, Inc., South Bend, IN) in 60:40 H O/glycerol or a 1.8 M
ARTICLE
2
supplemented with 10% fetal calf serum and 2 mM L-glutamine
(culture media from Invitrogen). Cell number and viability were
assessed using trypan blue dye staining. Cells (1 ꢁ 10 ) were suspended
13
solution of [1- C]-DHA (Supporting Information) in DMSO-d ,
6
8
containing 14.8 mM trityl radical (OX063; GE Healthcare, Amersham,
UK) and 1.4 mM of gadolinium chelate (Dotarem; Guerbet, Roissy,
in RPMI 1640 medium (2 mL) in a 10-mm NMR tube. For the
experiments with AA the dissolution was performed within 2.5 min of
resuspension of the cells to allow for generation of extracellular ROS and
for the DHA experiments within 1.5 min in order to minimize ROS
generation. The hyperpolarized substrate under study (2 mL, 14 mM)
was injected into the cell suspension at approximately 10 s post
dissolution. RPMI 1640 medium was found to be sufficient to buffer
the hyperpolarized sample dissolved in water to pH 7.0. Spectra (180)
were recorded using an 8° flip angle pulse (nt = 1, 16 kHz spectral
window, TR = 0.5 s, total acquisition time of 3 min) starting at the
beginning of hyperpolarized substrate injection.
3
France), were polarized as described previously. Briefly, the sample was
rapidly cooled under liquid helium in a GE Healthcare DNP prototype
hyperpolarizer at a pressure of ∼1 mBar to ∼1.2 K. Polarization of the
13
electron spins on the trityl radical were transferred to the C label using
microwave irradiation at 94.010 GHz (100 mW) over 1 and 1.5 h for
13
13
[
1- C]-AA and [1- C]-DHA, respectively. The frozen samples were
dissolved in 5 mL of water (pH ∼3.2). For in vivo administration,
samples were neutralized by addition of a further 1 mL of phosphate
buffer (200 mM; pH 7.8) containing EDTA (1.8 mM) and NaCl
(
(
400 mM) to yield a final pH and osmolality in the physiological range
pH 7.0ꢀ7.2; >310 mOsm/kg). This procedure prevented significant
1
3
Hyperpolarized C Spectroscopy (MRS) in Vivo at 9.4 T.
C
1
3
sample degradation and loss of polarization.
Hyperpolarized C NMR Spectroscopy in Vitro at 9.4 T.
For experiments in vivo, a 24-mm diameter surface coil tuned to
13
1
(100 MHz) was used in combination with a quadrature H-tuned
volume coil (Varian NMR Instruments, Palo Alto, CA), as described
NMR studies of reaction kinetics in vitro were performed on a 9.4 T
13
2
1
vertical wide-bore spectrometer (100 MHz C, Oxford Instruments,
previously. Transverse H spinꢀecho images were acquired to localize
13
Abingdon, U.K.) using a 10 mm C broadband probe (Varian NMR
the tumor and plan voxels for MRS (T = 500 ms, TE = 10 ms, field-of-
R
2
Instruments, Palo Alto, CA) and temperature-controlled at 37 °C.
view = 35 ꢁ 35 mm in a data matrix of 256 ꢁ 256; 21 slices of 2 mm
13
13
13
Samples of the [1- C]-AA (46 μL; 14 mM on dissolution) or
thickness). After administration of [1- C]-AA or [1- C]-DHA
(0.2 mL; 30 mM, 1.1 mg/kg), 200 spectra were acquired from a 6
mm tumor slice using a nominal flip angle of 10° (every fifth spectrum
was non-slice selective), with TR = 1 and 0.25 s for AA and DHA,
13
[
1- C]-DHA (39 μL; 14 mM on dissolution) preparations were
hyperpolarized, and measurements of the percentage polarization for
both substrates were made with a liquid-state polarimeter (Amersham
Health R & D, Malm €o , Sweden) approximately 8 s after the dissolution,
respectively. The T of each substrate was estimated by fitting the peak
integrals of the injected substrate in the non-slice selective spectra to
eq 1. Over 250 mg/kg intravenous AA, a 250-fold excess compared to
the dose here, is administered in pharmacological studies without side
1
13
at both pH 3.2 and 7.0. For [1- C]-AA, the reading was confirmed by
13
acquisition of 25 pulse and acquire C NMR spectra at approximately
0 s post dissolution (number of transients (nt) = 1, 32 kHz spectral
1
3
4
window, 5 s pulse repetition time (T ), 65° flip angle pulse, total
effects. High dose infusions of DHA (up to 60 mg/kg) can cause
R
35
acquisition time of 105 s) followed by acquisition of spectra at thermal
equilibrium from the same solution (nt = 16, 32 kHz spectral window,
elevated blood pressure and CNS responses in healthy rats, but this
level is 60-fold in excess of that used in this study, and adverse effects on
blood pressure were not observed here.
R
T = 200 s, 65° flip angle pulse, total acquisition time of 50 min). The
polarization was calculated by comparing the first hyperpolarized
spectrum with the spectrum at thermal equilibrium, correcting for
differences in the number of transients. This measurement could not
’ RESULTS AND DISCUSSION
13
be performed with [1- C]-DHA due to the rapid hydrolysis of the
The measured polarizations and T 's for the labeled C
1
2
3
1
sample, with a half-life of approximately 10 min at neutral pH. The T1
position in both ascorbic acid and dehydroascorbic acid at
1
3
of the hyperpolarized C label was determined in a separate experiment
13
13
9
.4 T are given in Table 1. [1- C]-DHA and [1- C]-AA exhibit
at pH 7.0 using 180 spectra acquired with an 8° flip angle pulse (nt = 1,
a similar level of polarization at pH 3.2, but at neutral pH,
1
6 kHz spectral window, T = 1 s, total acquisition time of 3 min) and
13
R
[
1- C]-DHA retains a degree of polarization significantly higher
13 13
fitting the decay of the integrated peak intensity to eq 1:
than that of [1- C]-AA (p = 0.05; n = 6). The loss of [1- C]-AA
polarization at physiological pH (5.1 ( 0.6% vs 10.5 ( 1.3% at
pH 3.2, p = 0.008; n = 6) can be attributed to the fact that the
hydroxyl group at the C position has a pK of 4.2 and so is
ꢀ
ꢁ
nTR
T1
n
S ¼ S0 exp ꢀ
cosðRÞ
ð1Þ
3
a
where S is the initial integrated peak intensity, T is the repetition time
13
0
R
completely dissociated at pH 7.0. The polarization of [1- C]-
DHA is also slightly lower at neutral pH (p = 0.12; n = 8), relating
in part to hydrolysis of the sample prior to measurement.
The polarization values reported here are lower than those
in seconds, T
angle, and n is the number of preceding RF pulses.
To determine the reaction kinetics of [1- C]-DHA with glutathione
1
is the spinꢀlattice relaxation time in seconds, R is the flip
13
13
(GSH), hyperpolarized [1- C]-DHA (2 mL) was injected into a 10 mm
13
commonly reported for [1- C]-pyruvate, which are in the range
NMR tube containing a solution of GSH (2 mL, 100 mM) in phosphate
of 20ꢀ35%, but are within the range of other molecules imaged
buffer (100 mM, pH 7.4) with EDTA (0.3 mM), to yield final
13
13
13
in vivo including [ C]-bicarbonate and [2- C]-fructose.
13
concentrations of 5 mM [1- C]-DHA and 50 mM GSH at pH 7.3;
40 spectra were then recorded using an 8° pulse (nt = 1, 24 kHz spectral
window, T = 1 s, total acquisition time of 4 min). This was repeated for
As expected, the C position in [1- C]-DHA displays a longer
1
2
13
T than that of [1- C]-AA in vitro (p = 0.003; n = 3), but
1
R
1
3
there was no significant difference in the estimated T values of
the two species in vivo, which are in agreement with those
1
final GSH concentrations of 1, 5, 10, and 25 mM. The rate of [1- C]-AA
production, kDHA, was determined by fitting the peak integrals to the
modified Bloch equations for two-site exchange
36
2,3,6
measured in erythrocyte suspensions and of a similar order to
with the back-flux of
13
13
those found for hyperpolarized [ C]-bicarbonate and [2- C]-
the reaction, kAA, set to zero. The relationship between the forward flux
fructose.
(
[DHA] ꢁ kDHA) and the rate of production of AA determined
spectrophotometrically (see Supporting Information) was established
using linear regression in Prism (Graphpad, San Diego, CA).
The intracellular reduction of DHA depends on NADPH,
both through glutathione-dependent reactions, which require
NADPH to replenish the reduced glutathione pool, and directly
through NADPH-dependent reactions. The observed reduction
For spectroscopic studies of vitamin C turnover in cancer cells,
EL-4 murine lymphoma cells were grown in RPMI 1640 medium
1
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Journal of the American Chemical Society
ARTICLE
Table 1. T Relaxation Times Measured in Vitro and in Vivo at 9.4 T and Percent Polarization Measured at Acidic and Neutral pH
1
Approximately 8 s Post Dissolution
polarization (%)
1
T (s)
vitamin C form
pH 3.2
pH 7.0
in vitro (pH 7.0)
in vivo
1
3
[
[
1- C]-ascorbic acid
10.5 ( 1.3
10.7 ( 1.3
5.1 ( 0.6
8.2 ( 1.1
15.9 ( 0.7
20.5 ( 0.9
8.6 ( 0.7
1
3
1- C]-dehydroascorbic acid
11.9 ( 0.1
measured spectrophotometrically under equivalent experimental
conditions (Figure 3c, see also Supporting Information; y = (1.00
2
(
0.04)x; r = 0.992). The second order rate constant for the
reaction derived from the spectrophotometric data, 0.34 ( 0.01
ꢀ
1
ꢀ1
M
data.
s , was in excellent agreement with previously reported
3
7
13
13
Oxidation of [1- C]-AA to [1- C]-DHA was observed at a
rate of 327 ( 41 μmol/s ([AA] x k ) in suspensions of EL4
AA
lymphoma cells at 2.5 min after resuspension of the cells in fresh
RPMI medium (Figure 4b, e). No oxidation was observed when
13
[
1- C]-AA was added to either RPMI medium alone (Figure 4a)
or to cells at 1 min after resuspension, although a low level
became apparent at 2 min after cell resuspension (data not
shown). Cell culture imposes a state of oxidative stress on
cells, with antioxidants provided primarily in supplemented
3
8,39
serum.
Cancer cells in vitro constitutively release hydrogen
38
peroxide, which readily penetrates the cell membrane and is
relatively stable in most culture media compared to reactive
oxygen species. Following resuspension, cells are not only
40
1
3
13
Figure 3. (a) C NMR spectra of hyperpolarized [1- C]-DHA
deprived of serum antioxidants, leading to an increased rate of
intracellular superoxide generation and in turn, a higher release
13
(
175 ppm) at the peak of [1- C]-AA (179 ppm) production following
reaction of 5 mM [1- C]-DHA with 1, 5, 10, 25, and 50 mM
glutathione. Peaks at 173.5 and 175.4 ppm are from diketogulonic acid
and an unknown product, respectively. (b) Time course of [1- C]-AA
peak intensity at various glutathione concentrations, normalized to the
maximum [1- C]-DHA intensity. (c) Correlation of the rate of [1- C]-
AA production, [DHA] ꢁ kDHA, with the rate of AA production
measured spectrophotometrically at equivalent concentrations of
DHA and glutathione.
38
13
of hydrogen peroxide, but will also rapidly deplete the medium of
oxygen and nutrients, leading to an acutely hypoxic state.
Furthermore, the pharmacological effects of high concentrations
of AA (>0.3 mM) have been attributed to the generation of
hydrogen peroxide through the reduction of catalytic metals such
1
3
13
13
4
1
as Cu(II), which are present in cell culture media, by AA. AA is
42
known to be oxidized by hydrogen peroxide, so the observation
13
of [1- C]-AA oxidation at increasing time after resuspension is
likely due to accumulation of hydrogen peroxide and is consistent
with the notion that this is an extracellular reaction.
13
of hyperpolarized [1- C]-DHA in vitro and in vivo is likely to
depend in large part on the reaction with reduced glutathione,
proceeding via both spontaneous and enzymatic reactions. The
spontaneous reaction of DHA with glutathione is thermodyna-
14
13
Conversely, the reduction of [1- C]-DHA (Figure 4d, f) was
observed only at early time points (<1 min) after cell resuspen-
5
ꢀ1
13
mically favorable at physiological pH (Keq = 2.5 ꢁ 10 M at
sion and returned to baseline by 2.5 min. Reduction of [1- C]-
pH 7) but proceeds slowly at endogenous concentrations of
DHA was observed in RPMI medium alone, which contains
1
0,11
43
DHA (micromolar) and glutathione (few millimolar).
fore, we sought to establish the rate at which higher concentrations
of [1- C]-DHA react spontaneously with glutathione to deter-
mine whether this could contribute to the [1- C]-DHA reduction
observed in cancer cell suspensions in vitro and tumor xenografts
in vivo. The reaction of [1- C]-DHA with glutathione yielded
There-
0.0033 mM glutathione. The medium also contains other
factors that have been documented to both reduce and hydrolyze
1
3
44
45
DHA, including sodium bicarbonate and cystine, as well as
11 mM D-glucose, which may competitively inhibit DHA trans-
port. The final concentrations of glutathione, cystine, bicarbo-
nate and glucose in these experiments were 0.0017, 0.104, 11.9,
and 5.5 mM respectively, with a final concentration of 7 mM
13
13
13
[
1- C]-AA at 179 ppm (Figure 3a) immediately following
13
13
addition of [1- C]-DHA to the glutathione-containing buffer.
[1- C]-DHA. Based on the measurements made above, the rate
1
3
13
The [1- C]-AA signal reached maximal intensity within 7ꢀ16 s
depending on the concentration of glutathione (Figure 3b). The
ratio of the peak integrals of AA and DHA in the spectrum
containing the maximum AA signal was calculated and found to be
linear with both glutathione concentration and the rate of AA
production (data not shown). The rate of [1- C]-AA production
was established by multiplying the amount of DHA (micromoles)
of [1- C]-AA production due to spontaneous reaction with the
glutathione in RPMI would be 0.15 nmol/s. The rates observed
in RPMI medium and EL4 lymphoma cell suspensions were
99 ( 6 nmol/s (n = 2) and 223 ( 18 nmol/s (n = 3), respectively.
The contribution of the spontaneous reaction with glutathione
may therefore be assumed to be negligible in EL4 cell suspen-
sions, indicating that other constituents of RPMI contribute to
the reduction of DHA in vitro and that a further reaction,
enzymatic or otherwise, occurs within EL4 cells. The shorter
13
ꢀ
1
with the apparent rate constant of the reaction kDHA (s ), and
this showed a linear correlation with the rate of AA production
1
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ARTICLE
1
3
13
Figure 4. Oxidation of [1- C]-AA (a, b, e) and reduction of [1- C]-DHA (c, d, f) in EL4 lymphoma cell suspensions. Hyperpolarized substrates were
8
added to either RPMI medium alone (a, c) or a suspension of 10 EL4 lymphoma cells in RPMI medium. (b) Spectrum obtained at 2.5 min post cell
resuspension and (d) at 1.5 min post cell resuspension. Only every fourth spectrum is displayed in the interests of clarity (e, f; time course over 46 s).
5
0
Peaks at 172.6 and 176.4 ppm are from diketogulonic acid and the C3 carbon of AA, respectively.
T1 of DHA in cell suspensions compared to RPMI medium,
0.8 ( 0.7 s (n = 5) versus 17.9 ( 1.7 s (n = 3), may be due to the
intracellular transport of this substrate.
indicate that upregulation of antioxidant systems allows tumors
48
2
to maintain a reduced microenvironment.
13
13
The in vivo metabolism of [1- C]-AA and [1- C]-DHA is
demonstrated in Figure 5. Intravenous administration of 30 mM
’ CONCLUSIONS
1
3
13
[
1- C]-AA yielded no observable [1- C]-DHA resonance in
EL4 lymphoma tumors (Figure 5a; n = 4), whereas administra-
In this study, we have demonstrated hyperpolarization of
13
13
13
[1- C]-ascorbic acid and [1- C]-dehydroascorbic acid, the
reduced and oxidized forms of vitamin C, to relatively high
levels. Both compounds have been administered previously in
high doses in clinical studies with little or no observed toxicity
tion of 30 mM [1- C]-DHA resulted in readily detectable
1
3
production of [1- C]-AA (Figure 5b; n = 4) and hydrolysis to
1
3
[
1- C]-DKG. The apparent rate constant for the in vivo reduc-
1
3
ꢀ1
tion of [1- C]-DHA was k
= 0.020 ( 0.004 s . The ratio of
DHA
13
13
and were shown here to exhibit sufficiently long T
1
's to allow
the peak integrals of [1- C]-AA and [1- C]-DHA calculated
within the first 1 s of data acquisition (summation of the first 4
FIDs) was 0.35 ( 0.08, with an average signal-to-noise ratio for
the [1- C]-DHA peak of 67 ( 20 over the same period. By
comparison, the peak ratio in the non-slice selective spectrum
taken at 1.25 s was 0.17 ( 0.03, indicating that a significant
portion of the overall reduction of [1- C]-DHA occurs in the
tumor. The EL4 tumor concentration of glutathione was deter-
observation of their signals in vitro and in vivo. The extracellular
13
pool of [1- C]-AA was oxidized rapidly in hypoxic EL4 cell
suspensions but not in vivo, as might be expected given that
1
3
13
tumors maintain a reduced microenvironment. [1- C]-AA may
find application in diseases that lead to a high level of extracellular
ROS, for example, for imaging of inflammation, where high levels
of superoxide and hydrogen peroxide are generated by immune
13
38
13
46
cells. Rapid intracellular reduction of [1- C]-DHA was ob-
served, both in EL4 tumor cell suspensions in vitro and in EL4
tumor xenografts in vivo. The rate of reduction could not be
mined by the method of Tietze et al. to be 1.18 ( 0.03 μmol/g
wet wt, corresponding to approximately 1.2 mM in the tumor
tissue.
4
6,47
Given the results in Figure 3, at this concentration the
1
3
accounted for by the direct reaction with glutathione. [1- C]-
DHA is transported by the glucose transporters GLUT1 and 3,
which are often overexpressed in tumor cells but are also
abundant at the bloodꢀbrain barrier. Given the association of
oxidative stress with neurodegenerative diseases and the avid
utilization of vitamin C in the brain, [1- C]-DHA may provide
a hyperpolarized probe for clinical imaging of redox status in vivo
not only in cancer but in a wide range of pathologies and could
spontaneous reaction of DHA with glutathione would be ex-
ꢀ
1
pected to proceed at a rate of only 0.0004 s and produce a
1
3
maximum relative signal intensity of 0.013 for [1- C]-AA. These
data provide further evidence that an enzymatic reaction is
required to account for both the rate and level of [1- C]-AA
13
49
13
produced in EL4 cells. The lack of observable oxidation of
1
3
[
1- C]-AA in vivo, despite the high level seen in cell suspen-
13
sions, together with the rapid reduction of [1- C]-DHA,
1
1799
dx.doi.org/10.1021/ja2045925 |J. Am. Chem. Soc. 2011, 133, 11795–11801

Journal of the American Chemical Society
ARTICLE
F.A.G. received a Cancer Research UK & Royal College of
Radiologists (UK) clinical research training fellowship, and
B.W.C.K. received a Cancer Research UK studentship.
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