On-bead combinatorial synthesis and imaging of chemical exchange saturation transfer magnetic resonance imaging agents to identify factors that influence water exchange

Article abstract of DOI:10.1021/ja201123f

The sensitivity of magnetic resonance imaging (MRI) contrast agents is highly dependent on the rate of water exchange between the inner sphere of a paramagnetic ion and bulk water. Normally, identifying a paramagnetic complex that has optimal water exchange kinetics is done by synthesizing and testing one compound at a time. We report here a rapid, economical on-bead combinatorial synthesis of a library of imaging agents. Eighty different 1,4,7,10- tetraazacyclododecan-1,4,7,10-tetraacetic acid (DOTA)-tetraamide peptoid derivatives were prepared on beads using a variety of charged, uncharged but polar, hydrophobic, and variably sized primary amines. A single chemical exchange saturation transfer image of the on-bead library easily distinguished those compounds having the most favorable water exchange kinetics. This combinatorial approach will allow rapid screening of libraries of imaging agents to identify the chemical characteristics of a ligand that yield the most sensitive imaging agents. This technique could be automated and readily adapted to other types of MRI or magnetic resonance/positron emission tomography agents as well.

Full text of DOI:10.1021/ja201123f

ARTICLE
pubs.acs.org/JACS
On-Bead Combinatorial Synthesis and Imaging of Chemical Exchange
Saturation Transfer Magnetic Resonance Imaging Agents To Identify
Factors That Influence Water Exchange
Roberta Napolitano,† Todd C. Soesbe,† Luis M. De Leꢀon-Rodríguez,†,§
||
A. Dean Sherry,†, and D. Gomika Udugamasooriya*,†,‡
†Advanced Imaging Research Center and ‡Department of Biochemistry, University of Texas Southwestern Medical Center,
5323 Harry Hines Boulevard, Dallas, Texas 75390-8568, United States
§Departamento de Química, Universidad de Guanajuato, Cerro de la Venada s/n, Guanajuato, Gto. 36040, Mexico
Department of Chemistry, University of Texas at Dallas, 800 West Campbell Road, Richardson, Texas 75083-3021, United States
S Supporting Information
b
ABSTRACT:
The sensitivity of magnetic resonance imaging (MRI) contrast agents is highly dependent on the rate of water exchange between the
inner sphere of a paramagnetic ion and bulk water. Normally, identifying a paramagnetic complex that has optimal water exchange
kinetics is done by synthesizing and testing one compound at a time. We report here a rapid, economical on-bead combinatorial
synthesis of a library of imaging agents. Eighty different 1,4,7,10-tetraazacyclododecan-1,4,7,10-tetraacetic acid (DOTA)-tetraamide
peptoid derivatives were prepared on beads using a variety of charged, uncharged but polar, hydrophobic, and variably sized primary
amines. A single chemical exchange saturation transfer image of the on-bead library easily distinguished those compounds having the
most favorable water exchange kinetics. This combinatorial approach will allow rapid screening of libraries of imaging agents to
identify the chemical characteristics of a ligand that yield the most sensitive imaging agents. This technique could be automated and
readily adapted to other types of MRI or magnetic resonance/positron emission tomography agents as well.
’ INTRODUCTION
in a decrease in the measured water signal, hence a darkening of
the image. Originally, the attention was focused on protons from
small diamagnetic molecules with ÀNH and ÀOH protons in
exchange with bulk water.8,12,13 However, the chemical shift
difference between the exchanging protons and bulk water in
such diamagnetic CEST agents is typically <5 ppm. This makes it
technically more difficult to saturate one proton pool without
inadvertently also partially affecting the other as well. One
advantage of using paramagnetic lanthanide complexes as CEST
agents (PARACEST) is that the chemical shift difference be-
tween the two exchangeable pools can be increased substantially.
Magnetic resonance imaging (MRI) is one of the most
important diagnostic imaging tools in clinical medicine today.
Exogenous paramagnetic metal complexes are often used as
contrast agents to shorten the relaxation times of the water
protons and thereby enhance tissue contrast in regions where
such agents accumulate. Over the past few years, new methods
have been explored for introducing image contrast,1 including
those based on chemical exchange saturation transfer (CEST)
mechanisms.2À11 Briefly, CEST contrast arises as a result of
selective saturation (through application of a suitable radio-
frequency field) of a proton pool that is in slow-to-intermediate
exchange with bulk water protons. Chemical transfer of those
saturated spins originally on the agent into the water pool results
Received: February 4, 2011
Published: July 27, 2011
r
2011 American Chemical Society
13023
dx.doi.org/10.1021/ja201123f J. Am. Chem. Soc. 2011, 133, 13023–13030
|

Journal of the American Chemical Society
ARTICLE
Figure 1. Schematic representation of a EuIIIDOTA-based PARACEST agent where the position of the Eu(III)-bound water molecule is shown
(arrows). The exchange kinetics of this water molecule with bulk water is modulated by the surrounding ligands on the four arms of the DOTA scaffold
(e.g., R1 and R2 represent the two peptoid residues that build diversity into the combinatorial library developed in this study).
For example, a slowly exchanging bound water resonance near 50
ppm is often seen in CEST spectra of Eu-1,4,7,10-tetraazacyclo-
dodecan-1,4,7,10-tetraacetic acid-tetraamide (EuDOTA-tretra-
amide) complexes such as those shown in Figure 1.7 This makes it
easy to selectively saturate this exchanging pool without affecting
the bulk water signal. PARACEST agents such as these have
some potential advantages over the more conventional T1 and T2
relaxation agents because image contrast is under operator
control and can be turned on and off at will.8 A second advantage
is that the CEST signal produced by EuDOTA-tetraamide
complexes is quite sensitive to the rate of water molecule
exchange between the inner sphere of the Eu(III) coordination
site and bulk water.14 This makes it easy to design responsive
agents that alter this rate by ligand electronic effects, stereo-
chemistry, and polarity of the neighboring substituents
[Figure 1(R1 and R2)].6 The primary disadvantage of current
PARACEST agents is their low sensitivity compared to that of
conventional T1 and T2 agents.5
Previous experiments have shown that a complex interplay of
chemical factors contribute to water exchange in these complexes
and that CEST sensitivity may be improved substantially by
dramatically slowing the rate of water exchange in these
complexes.6 However, it is a difficult challenge to understand
how all these possible parameters collectively influence the
CEST signal by synthesizing and studying one agent at a time,
the usual approach. Also, this is a major limitation in identifying
the optimal PARACEST agents using this time-consuming
conventional solution-phase synthesis and MRI analysis. For this
reason, we have been exploring the possibility of introducing a
large number of chemical variables into the basic DOTA scaffold
using on-bead combinatorial chemical methods in an effort to
directly identify those chemical features that yield optimal water
exchange rates for CEST imaging. The advantages of collecting a
single CEST image of a small library of polypeptides in solution
have been demonstrated previously.15,16 Here, we report (i) a
rapid, convenient on-bead synthesis of a library of 80 different
PARACEST agents that differ only in the chemical identity of
side arms on the DOTA scaffold [Figure 1(R1 and R2)] and (ii) a
method for simultaneous CEST imaging of the entire library
without removal of the agents from the beads. The resulting
images provide a direct readout of those structures that display
the best water exchange properties for CEST imaging.
200 mg/vessel), swelled in N,N-Dimethylformamide (DMF), and
treated with Fmoc-Met-OH, O-Benzotriazole-N,N,N',N'-tetramethyl-
uronium-hexafluoro-phosphate (HBTU) and 4-Methylmorpholine
(NMM) in DMF overnight. After the beads were washed with DMF,
the Fmoc group was removed (20% piperidine in DMF). The peptoid
portion (two methoxyethylamine units) was added using a two-step
(bromoacetylation and methoxyethylamine substitution) microwave-
assisted synthesis protocol [10% power (2 Â 15 s)] as previously
reported.17 Then Fmoc-β-Ala-OH was coupled as described above for
the methionine coupling. After removal of the Fmoc group, the beads
were treated with bromoacetic acid/N,N-Diisopropylcarbodiimide
(DIC) in the microwave oven at 10% power (2 Â 15 s). The beads
were then treated with 1 M 1,4,7,10-tetraazacyclododecan-1,4,7-triacetic
acid (DO3A)-tris(tert-butyl ester) solution in the microwave oven at
10% power (3 Â 15 s). After the beads were washed with DMF and
Dichloromethane (DCM), the tert-butyl groups were removed
(95% TFA, 2.5% triisopropylsilane, and 2.5% water, 4 h). After
being washed with DCM and DMF, the beads were treated with
N-Boc-1,2-diaminoethane, HBTU, and NMM for 24 h. Finally, the
Boc group was removed as described above using 95% TFA.
Synthesis of the Library. The library was synthesized starting
from the parent compound 16 (Scheme 2) and adding two peptoid
moieties onto each of the three free arms of the DOTA scaffold.
Bromoacetic acid coupling of the first peptoid residue was performed
in bulk synthesis. Then the beads were equally (∼10 mg/well)
distributed into a 96-AcroWell filter plate (PALL). The first eight
amines (Chart 1; some were protected accordingly) were added (2 M,
180 μL/well) to each row of the plate (shaking for 2 h, 250 rpm). After
DMF washing by centrifugation, bromoacetic acid/DIC was added for
30 min twice (shaking for 2 h, 250 rpm). After DMF washing, 10 amines
(Chart 1) were added to each of the first 10 columns of the plate. Beads
containing β-alanine and 1,2-diaminoethane were transferred to another
TFA-resistant 96-well plate, treated with a 95% TFA, 2.5% triisopro-
pylsilane, and 2.5% water mixture for 4 h to remove tert-butyl and the
Boc protection group. The beads were washed with DCM and DMF,
finally neutralized with 20% piperidine in DMF for 5 min, and
transferred to the previous plate. All wells were washed with water
and allowed to swell in water for 1 h. The metal complexation was
performed with a 0.2 M EuCl3 solution at pH 6.3 overnight. Finally, the
beads were washed with water (5Â), dried, and stored at 4 °C.
Cleavage, Purification, and Mass Analysis. The cyanogen
bromide cleavage18 was used to confirm the synthesis of each step up to
compound 16 and then for the 10 representative compounds of the library.
The beads were treated with the cleavage solution (0.28 M CNBr in a 5:4:1
acetonitrile/acetic acid/water mixture) overnight. CNBr and acetic acid
were removed under a nitrogen flow. The residue was redissolved in a 1:1
acetonitrile/water mixture for mass and HPLC analysis.
’ EXPERIMENTAL METHODS
On-Bead Bulk Synthesis of Parent Compound 16. TentaGel
macrobeads were placed in five peptide reaction vessels (total 1 g,
13024
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
Chart 1. Structures of the 10 Primary Amines Used To Build the Library
Scheme 1. Two Basic Peptoid Synthesis Steps Carried Out on a Bead-Attached DOTA Scaffold Adding One Peptoid Residue
NMR. CEST spectra were collected on beads placed in 3 mm NMR
tubes filled with water using a Varian 9.4 T NMR system with the sample
temperature maintained at 25 °C. A B1 of 300 Hz and 5 s duration was
applied at each frequency prior to collection of a 1H spectrum of water.
CEST Imaging of the Library. In vitro imaging was performed on
a Varian 9.4 T animal system using a 63 mm diameter quadrature volume
coil. The samples were placed at the gradient isocenter with the
temperature maintained at 20 °C using warmed air and a thermocouple.
A single coronal plane (64 Â 64 Â 2 mm) passing through the bottom
portion of each sample well was selected for imaging. Image-based
CEST spectra acquired with the fast spinÀecho sequence (TR/TE =
69.8 ms/8.4 ms, echo train 8, averages 2, 64 Â 64 Â 1 pixel matrix) were
used to measure the CEST frequency (Δω) for each well. Images were
then acquired using the fast spinÀecho sequence (TR/TE = 69.8 ms/
8.4 ms, echo train 8, averages 2, 64 Â 64 Â 1 pixel matrix) with saturation
at both +50 ppm (“on”) and À50 ppm (“off”). A 5 s long, 10 μT
saturation pulse was used for every TR. A CEST image was created by
subtracting the two images (off À on). Spatial susceptibility effects
within the samples were minimized using two methods. First, the 8 Â 11
sample wells were surrounded by a two-well thick boarder of pure water.
Second, after the first set of images was taken, the sample was rotated by
180° and imaged again. The two CEST images at 0° and 180° were then
averaged. These methods assured that the CEST variations from well to
well were due to changes in chemical configuration and not susceptibility
effects.
the remaining three carboxyl groups of DOTA to provide
terminal amines for building the library, and (iii) add two rounds
of “peptoid” monomers to the three free primary amino groups
to create the ligand library. Peptoids were selected as the diversity
component [Figure 1(R1 and R2)] because the chemistry is
simple and fast and many primary amines are available that would
introduce a variety of chemical properties into the ligand side
arms (hydrophobicity, size, charge). The well-established on-
bead peptoid synthesis is based on simple two-step chemical
reactions, an acylation step using diimide-activated bromoacetic
acid followed by nucleophilic displacement of the bromide with a
primary amine (Scheme 1).19 These reactions are not air
sensitive or particularly moisture sensitive and typically take
place in high yield at each step. Thus, the entire process can be
automated, adapted to most commercial peptide synthesizers, and
even accelerated to less than 1 min using microwave irradiation.17
The amines chosen for the library provided variable charge, polarity,
and steric bulkiness around the Eu(III)Àwater exchange site
(Chart 1). The blue-colored nitrogen atom in each amine
(Chart 1) will be incorporated into the peptoid backbone (see
Scheme 1, second step), leaving the remaining portion as the
“R” group.
The synthesis scheme is shown in Scheme 2. The initial
common synthetic steps (steps 11À16, Scheme 2) were carried
out in bulk using standard glassware, while the remaining diverse
regions (steps 17 and 18, Scheme 2) were carried out using a 96-
well synthesis filter plate [Scheme 2 (inset)]. First, a spacer
between the bead and the DOTA scaffold was introduced.
Methionine was added as the first residue of the spacer arm so
the final product could be cleaved from the resin with cyanogen
bromide for subsequent mass spectroscopic confirmation. To
obtain reasonably symmetrical DOTA-tetraamide products and
to limit possible steric hindrance between the EuDOTA complex
and the bead surface, two neutral methoxyethylamine moieties were
added next. Here, both the bromoacetic acid addition and amine
couplingreactionswereassistedbymicrowaveirradiation.17,19 Then
Eu(III) Quantitative Assay. Ten beads from each of the individual
wells selected were isolated and collected in Eppendorf tubes. A 2.0 M
solution of HCl (50 μL/tube) was added and incubated at 37 °C for 2 h.
Then the samples were neutralized using 2.0 M NaOH (55 μL/tube).
The enhancement solution was added (115 μL/tube) and incubated for
30 min at 37 °C. Fluorescence emission at 615 nm was detected using a
SpectraMax M5 spectrophotometer.
’ RESULTS AND DISCUSSION
The solid-phase synthesis strategy was to (i) attach a DOTA
scaffold to TentaGel macrobeads (300 μm diameter) through a
linker to a single carboxyl group, (ii) couple ethylenediamine to
13025
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
Scheme 2. Flow Chart of the Complete Library Synthesis
(step 15 and 16, Scheme 2) to yield 16, the starting material for
building the peptoid library.
A critical test was carried out at this point to verify that resin-
bound DOTA-tetraamide forms a complex with Eu(III) and to
test whether a CEST signal could be detected with the EuDOTA-
tetraamide complex bound to the beads. Eu(III) complexation
was performed by addition of ∼15-fold excess EuCl3 at pH 6.3
overnight, followed by removal of excess Eu(III) by washing the
resin several times with water. A few beads were then removed
and placed in a glass capillary for collection of a CEST spectrum
using a vertical bore 400 MHz NMR spectrometer. Importantly,
a large Eu(III)-bound water exchange peak was observed in the
CEST spectrum near 50 ppm (Figure 2, green data points). No
CEST was observed for ligand 16 alone [no Eu(III)] or empty
beads treated with EuCl3, confirming that the CEST signal
reflects Eu*16 attached to the beads (Figure 2, blue and black
data points). The CEST signal of a solution of Eu*16 after it was
cleaved from the beads (see section S2, Supporting Information)
displayed a chemical shift similar to that of the bead-bound
complex, but the exchange peak was slightly broader (Figure 2,
red data points). This implies that water exchange may be
somewhat slower for the complex attached to beads, although
absolute comparisons of exchange rates were not possible
because the concentrations of the two samples were unknown.
Most importantly, these data show that the CEST signal from a
covalently attached complex (Eu*16) is not strongly influenced
by the presence of the Tentagel beads. This observation allowed
us to proceed with the synthesis of a diverse chemical library of
Figure 2. Comparison of Z-spectra of Eu*16 attached to beads with
different controls [Eu*16 attached to beads, Eu*16 in solution without
the beads, beads alone (no 16, but treated with Eu(III)), or beads with
ligand 16 but no added Eu(III)].
β-alanine was added to mimic the ethylenediamine spacer
planned for the other three DOTA arms, and finally DO3A
tert-butyl ester was coupled. The three remaining carboxyl groups
of DOTA were then reacted with monoprotected 1,2-diaminoethane
13026
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
Figure 3. (A) CEST image of the 80-compound library of EuIIIDOTA-tetraamide peptoid derivatives attached to beads. The color scale displays the
percent change in bulk water intensity (1 À Ms/Mo) with a presaturation pulse set either off resonance (Mo) or on resonance (Ms). The chemical
structures of each amine used to build the library are shown along each horizontal row (R1) and each vertical column (R2). A number of control samples
were located in column 11, identified as indicated in the figure. (B) Structure of the EuIIIDOTA-tetraamide peptoid (bead-attached form) in well (7, 1)
that showed the highest CEST intensity.
CEST ligands, including hydrophobic side arms that might not
ordinarily be soluble when prepared in bulk.
etc. to yield an array of compounds having the same first peptoid
along each row (R1 = 1À8) and the same second residue along
each column (R2 = 1À10). This simple approach allowed rapid
access to 8 Â 10 = 80 different compounds, all built on the same
parental DOTA-tetraamide scaffold and differing only in the
chemical identity of R1 and R2. Finally, Eu(III) complexation was
performed as described previously. Before proceeding with the
MRI measurements, the quality of the library was checked by
analyzing the representative compounds synthesized in the
12th column using MALDI-TOF MS to confirm the complete
synthesis of all eight compounds (see section S3, Supporting
Information). Random samples were not withdrawn from the
actual 80-compound library for this purpose to have the same
number of beads in each well for imaging.
To register a single image for all 80 compounds plus controls,
the beads were first transferred from the synthesis/filtration plate
to the central 80 wells of a 145-well plate to maximize the height
of the packed beads in the 4 mm diameter well. All 145 wells were
filled with water, including those on the perimeter that did not
contain beads, to make the sample as homogeneous as possible.
The plate was then positioned in the center of a 63 mm quadrature
volume coil, and CEST imaging was performed by applying a
10 μT frequency-selective presaturation pulse for 5 s followed by
The parallel combinatorial synthesis approach was used to
couple two peptoid residues onto 16. Bromoacetic acid was
coupled (for the first peptoid residue) to 16 in bulk before the
beads were distributed equally into a 96-well synthesis plate
[Scheme 2 (inset)]. To avoid possible inhomogeneous micro-
wave irradiation in the different wells of the plate, the remaining
peptoid coupling steps were performed for an extended period of
time at room temperature.17 The first eight rows and ten columns
were used for synthesis of the library; column 11 was used for
controls, and column 12 was used for synthesis of eight additional
DOTA-tetraamide peptoids using random combinations of
amines (see section S3, Supporting Information) for cleavage
and mass spectroscopy analyses.
Eight different amines (1À8, Chart 1) were coupled to the
three remaining arms of DOTA to complete the first residue in
each row [i.e., R1 in row 1 was methoxyethylamine (1), R1 in row
2 was piperonylamine (2), R1 in row 3 was 4-methoxybenzyla-
mine (3), etc]. This was followed by a second bromoacetic acid
coupling step to prepare for the second amine. Amines 1À10
(Chart 1) were chosen as residue R2; 1 was added to each of eight
wells in column 1, 2 was added to each of eight wells in column 2,
13027
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
a spinÀecho sequence using a 9.4 T Varian scanner. A 2 mm thick
coronal slice was selected near the bottom of each well to ensure that
each image would reflect an equal number of beads (the height of
packed beads in each well was >3 mm). The resulting CEST image
(Figure3) isdefined as the difference in water intensities in an image
collected with the presaturation pulse set to À50 ppm (control)
minus a second image collected with the presaturation pulse set to
+50 ppm. This experiment was repeated on another day, and the
same image was obtained, thereby confirming the reproducibility of
this library imaging experiment.
After the plate was imaged, 10 beads were collected from each
of 11 wells selected from those that showed large variations in
CEST signal intensities that include empty beads and beads from
the well containing Eu*16. These beads were treated with HCl to
displace all Eu(III) from each chelate and analyzed for total
Eu(III) using a commercially available fluorescence enhance-
ment assay.20,21 These data (Figure 4) indicated that there was an
identical amount of total Eu(III) in each 10-bead sample and that
empty beads treated with Eu(III) did not retain a significant
amount of ion. Thus, any intensity differences in the CEST
images shown in Figure 3 could be attributed solely to con-
sequences of the chemical differences of the side arms in the 80
different EuDOTA-tetraamide complexes and not differences in
agent concentrations.
The resulting CEST image (Figure 3) illustrates the power of
the technique; the CEST signal in some wells was quite intense
(up to a 30% reduction in water intensity), while little to no
CEST was detected in other wells. To check whether this may be
due to frequency differences in the exchanging Eu(III)-bound
water resonance in each complex and to show that the spectra
were consistent with the imaging data, complete CEST spectra
were collected on several individual wells (Figure 5). For those
wells showing a CEST signal, the Eu(III)-bound water exchange
peak appeared between 50 and 51 ppm (Figure 5) in all cases.
The quantitative percent CEST signal (1 À (M(+50)/M(À50)
Â
100) from each well varied from 0 to 25% (Table 1, Supporting
Information) and was consistent with the qualitative image
intensities shown in Figure 3. The most intriguing part of the
study came from a comparison of the combined effects of various
physicochemical properties of R1 and R2 on water exchange at
each Eu(III) center.
The most intense CEST signals came from wells containing
Eu(III) complexes with small ether, polar, or charged residues
such as methoxyethyl (1), carboxyethyl (7), or the furans (6 and
8) [compounds in wells (1, 1), (1, 6), (1, 7), (7, 1), (7, 6), (7, 7),
and (7, 8), where the designation (R1, R2) refers to (row,
column), Figure 3A]. Interestingly, the position of each residue
also had consequences on the CEST signal intensity. A more
intense CEST signal was seen when a carboxyethyl group (7) was
Figure 4. Eu(III) quantification of 10 beads collected from each of 11
wells (as identified below each column).
Figure 5. On bead CEST spectra collected from six selected wells of the library. CEST spectra are shown in red and the quantitative percent CEST signal is shown in blue.
13028
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
the first residue (almost independent of which residue was
present at the second position; see row 7) compared to when
it was the second residue (column 7). For example, equally
intense CEST signals were seen for compounds containing the
carboxyethyl group (7) in the first peptoid position and either
2-methylfuran (7, 6) or 2-methylthiophene (7, 8) in the second
position, but when the sequence was inverted [(6, 7) and (8, 7)],
both resulted in a weak CEST signal. Thus, one can conclude that
the presence of a negatively charged group in the peptoid
positioned closest to the Eu(III) center has a positive influence
on the CEST signal. This is consistent with previous data
reporting that EuDOTA-tetraglycinate has one of the slowest
water exchange rates of any CEST agent reported to date.22 The
opposite trend was seen for the positively charged aminoethyl
group (4). Here, the CEST intensities did depend on the identity
of the second group, but in general, a larger CEST signal was seen
for those compounds where the positively charged group (4)
occupied the second position of the peptoid sequence, not the
first (compare the three bright signals in column 4 with the
virtually all dark signals in row 4). This was highlighted even
further for those compounds having one positively and one
negatively charged group together in one sequence; in this case,
as usual, the best CEST signal was found for those compounds
with the negatively charged carboxyethyl group (7) closest to the
metal center (7, 4). When the positively charged aminoethyl
group (4) occupied the first position, this canceled most of the
advantage of having the negatively charged carboxyethyl group in
the second position [compare the signal intensity of (4, 7) with
that of (3, 7) or (5, 7)]. This suggests that having a nonpolar
group close to the metal center is more advantageous than having
a positively charged group near the metal center, as long as the
second position is occupied by the negatively charged, carbox-
yethyl group [(1, 7) (3, 7), (5, 7), (6, 7)].
In all cases, the bulkier aromatic groups, 5-benzyl-1,3-dioxole
(2), 4-methoxybenzyl (3), or 4-chlorobenzyl (9), resulted in the
weakest CEST signals, regardless of their position in the peptoid
sequence (compare the number of dark wells in columns 2, 3, 8,
and 9 with that in any of the other columns). This is in agreement
with our prior observations that bulkier aromatic residues such as
these increase the rate of water exchange in EuDOTA-tetraamide
complexes.23 Interestingly, the smaller aliphatic, hydrophobic
isobutyl group (5) did not show a similar trend. When this group
appeared in combination with a carboxyethyl group (in either
order), the CEST intensities were similar [compare well (5, 7)
with well (7, 5)]. A similar but less obvious trend was observed
with 2-methylfuran (6) or 2-methylthiophene (8) acting as the
small nonpolar residue in combination with a charged group
(either positive or negative) as the second component [(6, 4), (8,
4), (6, 7), (8, 7)]. This suggests that having a small nonpolar
group closest to the Eu(III)Àwater exchange site may act to slow
dissociation of the inner-sphere water molecule from the Eu(III)
coordination site while the charged group further away acts to
organize second-sphere water molecules to slow their entrance
into a position where they can be in a position to exchange with
the single, inner-sphere water molecule.
peak did not vary substantially (all were near 50À51 ppm)
because the ethylenediamine spacer placed the chemically di-
verse substituents far enough away from the metal to minimize
any changes in the ligand field. Nevertheless, the most surprising
finding of this study was that the water exchange rate and hence
the CEST intensity can vary substantially even with these
chemically diverse groups positioned well away from the central
Eu(III). Thus, the differences in CEST signal intensities ob-
served here can only reflect differences in water accessibility to
the inner-sphere coordination sphere.
The use of peptoids as the diversity component has some
advantages for future potential applications. These include rapid
and inexpensive synthesis of peptoid libraries.19,25À28 In addi-
tion, peptoids are serum stable,28,29 more cell permeable,30 and
nonimmunogenic,31 important considerations for any potential
clinical application.
’ CONCLUSION
We have presented a combinatorial approach for an easy,
rapid, and cost-effective synthesis and screening of a library of
possible PARACEST contrast agents by utilizing peptoid chem-
istry and the chelating property of the DOTA structure. By
changing the amines and the basic structure of the system, many
different combinations can be prepared and studied in a single
screening assay with the agents attached to the solid support, a
much easier way than the conventional solution-phase practices.
This parallel synthesis method can, in principle, be used to create
diverse libraries of MRI or PET agents for protein and cell
targeting purposes. Thousands of different amines are commer-
cially available to bring the ligand diversity. Today, high-through-
put parallel synthesizers are also commercially available, making
these studies even easier to apply. Upon use of these automated
synthesizers, the diversity can be applied to all four arms of
DOTA scaffold equally, and thereby near symmetrical and even
much larger DOTA libraries can be synthesized in the future.
Nonetheless, the best compounds we isolated in this study can
also be synthesized with full symmetry in the solution phase for
practical applications as better CEST agents. Studies are being
continued to validate these compounds further in both in vitro
and in vivo applications.
’ ASSOCIATED CONTENT
S
Supporting Information. Detailed experimental proce-
b
dures, chemical structure of compound 16 (CNBr cleaved form),
spectral data (MALDI-MS, HPLC) of compound 16 and the
library, and complete ref 29. This material is available free of
charge via the Internet at http://pubs.acs.org.
’ AUTHOR INFORMATION
Corresponding Author
E-mail: gomika.udugama@utsouthwestern.edu.
’ ACKNOWLEDGMENT
It has been reported that the resonance frequency of the
bound water is quite sensitive to the identity of the first amide
side chain group in a series of tetrasubstituted amino acids, for
example.24 This did not hold true for the 80 complexes prepared
in this library because all four side chains were positioned well
away from the central EuDOTA structure by the four ethylene-
diamine arms. In this case, the frequency of the water exchange
We thank the National Institutes of Health (Grants
CA115531, RR02584, and EB004582), the Robert A. Welch
Foundation (Grant AT-584), and the University of Texas South-
western Medical Center for financial support. Also, we thank Dr.
Kathlynn Brown for providing access to the fluorescence
spectrophotometer.
13029
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030

Journal of the American Chemical Society
ARTICLE
’ REFERENCES
(1) Terreno, E.; Dastru, W.; Delli Castelli, D.; Gianolio, E.; Geninatti
Crich, S.; Longo, D.; Aime, S. Curr. Med. Chem. 2010, 17, 3684–3700.
(2) Hancu, I.; Dixon, W. T.; Woods, M.; Vinogradov, E.; Sherry,
A. D.; Lenkinski, R. E. Acta Radiol. 2010, 51, 910–923.
(3) Wu, Y.; Soesbe, T. C.; Kiefer, G. E.; Zhao, P.; Sherry, A. D. J. Am.
Chem. Soc. 2010, 132, 14002–14003.
(4) Wu, Y.; Zhao, P.; Kiefer, G. E.; Sherry, A. D. Macromolecules
2010, 43, 6616.
(5) Sherry, A. D.; Woods, M. Annu. Rev. Biomed. Eng. 2008,
10, 391–411.
(6) Woods, M.; Woessner, D. E.; Sherry, A. D. Chem. Soc. Rev. 2006,
35, 500–511.
(7) Zhang, S.; Merritt, M.; Woessner, D. E.; Lenkinski, R. E.; Sherry,
A. D. Acc. Chem. Res. 2003, 36, 783–790.
(8) Ward, K. M.; Aletras, A. H.; Balaban, R. S. J. Magn. Reson. 2000,
143, 79–87.
(9) Vinogradov, E.; He, H.; Lubag, A.; Balschi, J. A.; Sherry, A. D.;
Lenkinski, R. E. Magn. Reson. Med. 2007, 58, 650–655.
(10) Liu, G.; Ali, M. M.; Yoo, B.; Griswold, M. A.; Tkach, J. A.; Pagel,
M. D. Magn. Reson. Med. 2009, 61, 399–408.
(11) Zhao, J. M.; Har-el, Y. E.; McMahon, M. T.; Zhou, J.; Sherry,
A. D.; Sgouros, G.; Bulte, J. W.; van Zijl, P. C. J. Am. Chem. Soc. 2008,
130, 5178–5184.
(12) Ling, W.; Regatte, R. R.; Navon, G.; Jerschow, A. Proc. Natl.
Acad. Sci. U.S.A. 2008, 105, 2266–2270.
(13) McMahon, M. T.; Gilad, A. A.; Zhou, J.; Sun, P. Z.; Bulte, J. W.;
van Zijl, P. C. Magn. Reson. Med. 2006, 55, 836–847.
(14) Zhang, S.; Sherry, A. D. J. Solid State Chem. 2003, 171, 38–43.
(15) Liu, G.; Gilad, A. A.; Bulte, J. W.; van Zijl, P. C.; McMahon,
M. T. Contrast Media Mol. Imaging 2010, 5, 162–170.
(16) McMahon, M. T.; Gilad, A. A.; DeLiso, M. A.; Berman, S. M.;
Bulte, J. W.; van Zijl, P. C. Magn. Reson. Med. 2008, 60, 803–812.
(17) Olivos, H. J.; Alluri, P. G.; Reddy, M. M.; Salony, D.; Kodadek,
T. Org. Lett. 2002, 4, 4057–4059.
(18) Simpson, L. S.; Burdine, L.; Dutta, A. K.; Feranchak, A. P.;
Kodadek, T. J. Am. Chem. Soc. 2009, 131, 5760–5762.
(19) Zuckermann, R. N.; Kerr, J. M.; Kent, S. B. H.; Moos, W. H.
J. Am. Chem. Soc. 1992, 114, 10646–10647.
(20) De Silva, C. R.; Vagner, J.; Lynch, R.; Gillies, R. J.; Hruby, V. J.
Anal. Biochem. 2010, 398, 15–23.
(21) Handl, H. L.; Vagner, J.; Yamamura, H. I.; Hruby, V. J.; Gillies,
R. J. Anal. Biochem. 2005, 343, 299–307.
(22) Dixon, W. T.; Ren, J.; Lubag, A. J.; Ratnakar, J.; Vinogradov, E.;
Hancu, I.; Lenkinski, R. E.; Sherry, A. D. Magn. Reson. Med. 2010,
63, 625–632.
(23) Mani, T.; Tircso, G.; Togao, O.; Zhao, P.; Soesbe, T. C.;
Takahashi, M.; Sherry, A. D. Contrast Media Mol. Imaging 2009,
4, 183–191.
(24) Viswanathan, S.; Ratnakar, S. J.; Green, K. N.; Kovacs, Z.; De
Leon-Rodriguez, L. M.; Sherry, A. D. Angew. Chem., Int. Ed. 2009,
48, 9330–9333.
(25) Alluri, P. G.; Reddy, M. M.; Bachhawat-Sikder, K.; Olivos, H. J.;
Kodadek, T. J. Am. Chem. Soc. 2003, 125, 13995–14004.
(26) Udugamasooriya, D. G.; Dineen, S. P.; Brekken, R. A.;
Kodadek, T. J. Am. Chem. Soc. 2008, 130, 5744–5752.
(27) Udugamasooriya, D. G.; Dunham, G.; Ritchie, C.; Brekken,
R. A.; Kodadek, T. Bioorg. Med. Chem. Lett. 2008, 18, 5892–5894.
(28) Zuckermann, R. N.; Kodadek, T. Curr. Opin. Mol. Ther. 2009,
11, 299–307.
(29) Simon, R. J.; et al. Proc. Natl. Acad. Sci. U.S.A. 1992, 89,
9367–9371.
(30) Kwon, Y. U.; Kodadek, T. J. Am. Chem. Soc. 2007, 129,
1508–1509.
(31) Astle, J. M.; Udugamasooriya, D. G.; Smallshaw, J. E.; Kodadek,
T. Int. J. Pept. Res. Ther. 2008, 14, 223–227.
13030
dx.doi.org/10.1021/ja201123f |J. Am. Chem. Soc. 2011, 133, 13023–13030