Interaction of Cd and Zn with biologically important ligands characterized using solid-state NMR and ab initio calculations

Article abstract of DOI:10.1021/ic026287d

For the first time, coordination geometry and structure of metal binding sites in biologically relevant systems are studied using chemical shift parameters obtained from solid-state NMR experiments and quantum chemical calculations. It is also the first e

Full text of DOI:10.1021/ic026287d

Inorg. Chem. 2003, 42, 3142−3151
Interaction of Cd and Zn with Biologically Important Ligands
Characterized Using Solid-State NMR and ab Initio Calculations
,†,‡,§
Srikanth S. Kidambi,† Dong-Kuk Lee,†,‡ and A. Ramamoorthy*
Department of Chemistry, Biophysics Research DiVision, and Macromolecular Science &
Engineering, UniVersity of Michigan, Ann Arbor, Michigan 48109-1055
Received December 19, 2002
For the first time, coordination geometry and structure of metal binding sites in biologically relevant systems are
studied using chemical shift parameters obtained from solid-state NMR experiments and quantum chemical
calculations. It is also the first extensive report looking at metal−imidazole interaction in the solid state. The principal
values of the 113Cd chemical shift anisotropy (CSA) tensor in crystalline cadmium histidinate and two different
cadmium formates (hydrate and anhydrate) were experimentally measured to understand the effect of coordination
number and geometry on 113Cd CSA. Further, 13C and 15N chemical shifts have also been experimentally determined
to examine the influence of cadmium on the chemical shifts of 15N and 13C nuclei present near the metal site in
the cadmium−histidine complex. These values were then compared with the chemical shift values obtained from
the isostructural bis(histidinato)zinc(II) complex as well as from the unbound histidine. The results show that the
isotropic chemical shift values of the carboxyl carbons shift downfield and those of amino and imidazolic nitrogens
shift upfield in the metal (Zn,Cd)−histidine complexes relative to the values of the unbound histidine sample. These
shifts are in correspondence with the anticipated values based on the crystal structure. Ab initio calculations on the
cadmium histidinate molecule show good agreement with the 113Cd CSA tensors determined from solid-state NMR
experiments on powder samples. 15N chemical shifts for other model complexes, namely, zinc glycinate and zinc
hexaimidazole chloride, are also considered to comprehend the effect of zinc binding on 15N chemical shifts.
I. Introduction
ratio and I > 1/2) make it a difficult probe to perform NMR
experiments. These problems were circumvented using the
surrogate probe strategy developed by Vallee and co-
workers.1 Later, this concept of substituting insensitive metal
centers with spectroscopic-friendly nuclei was greatly ex-
ploited by several research groups2 in the field of NMR
spectroscopy. All these studies utilized cadmium ion as the
NMR probe in elucidating the structure around the metal
(mainly Zn2+ and Ca2+) centers. Cadmium ion occurs in the
same group as zinc ion and has a size comparable to both
Several metals such as Zn2+, Ca2+, Cu2+, Hg2+, Mn2+, and
Mg2+ are commonly found in living organisms. These ions
either act as a structure promoter or play an important role
in enzyme activity. A systematic study of the coordination
chemistry of metal centers in bioinorganic complexes would
be essential to obtain a clear understanding on the role of
these metals in several biological and chemical processes.
In biosystems such as photosystem-II, phosphotriesterase,
calcium, and zinc ion play an important role in the function-
ing of the protein. A thorough understanding of the properties
of these ions could provide valuable information about the
activity of these proteins. But the closed d-shell configuration
of Zn2+ and Ca2+ ions makes them unsuitable for spectro-
scopic techniques such as UV-visible or ESR spectroscopy.
Moreover, the properties of these nuclei (low gyromagnetic
(1) (a) Coleman, J. E.; Vallee, B. L. J. Biol. Chem. 1961, 236, 2244. (b)
Lindskog, S. J. Biol. Chem. 1963, 238, 945. (c) Latt, S. A.; Auld, D.
S.; Vallee, B. L. Proc. Natl. Acad. Sci. U.S.A 1970, 67, 1383.
(2) (a) Summers, M. F. Coord. Chem. ReV. 1988, 86, 43. (b) Armitage,
I. M.; Pajer, R. T.; Schoot Uiterkamp, A. J. M.; Chlebowski, J. F.;
Coleman, J. E. J. Am. Chem. Soc. 1976, 98, 5710. (c) Ellis, P. D.
Science 1983, 221, 1141. (d) Ellis, P. D. J. Biol. Chem. 1989, 264,
3108. (e) Rivera, E.; Kennedy, M. A.; Adams, R. D.; Ellis, P. D. AdV.
Magn. Reson. 1987, 13, 257. (f) Jakobsen, H. J.; Ellis, P. D.; Inners,
R. R.; Jensen, C. F. J. Am. Chem. Soc. 1982, 104, 7442. (g) McAteer,
K.; Lipton, A. S.; Ellis, P. D. Cadmium-113 NMR. A Surrogate Probe
for Zn and Calcium in Proteins. In Encyclopaedia of Nuclear Magnetic
Resonance; Grant, D. M., Harris, R. K., Eds.; John Wiley & Sons:
New York, 1996; Vol. 2, pp 1085-1091 and references cited within.
* To whom correspondence should be addressed. E-mail: ramamoor@
umich.edu. Tel: (734) 647-6572. Fax: (734) 615-3790.
† Department of Chemistry.
‡ Biophysics Research Division.
§ Macromolecular Science & Engineering.
3142 Inorganic Chemistry, Vol. 42, No. 9, 2003
10.1021/ic026287d CCC: $25.00 © 2003 American Chemical Society
Published on Web 04/09/2003

Interaction of Cd and Zn with Biologically Important Ligands
zinc and calcium ions. The nuclear spin quantum number of
1/2 for cadmium makes the 113Cd chemical shift an easily
measurable NMR parameter, and therefore it is a suitable
nucleus to replace Zn2+ or Ca2+ for NMR studies on
biological molecules. Even though Kimblin and Parkin have
reported significant differences in the structures of Cd and
Zn analogous model complexes,3 this method still seems to
be a good starting point to elucidate the structure of Zn-
containing biomolecules. Some of the most important
benefits in using 113Cd to study the coordination chemistry
of metal centers are as follows: (i) it has a large span of
113Cd chemical shift, which allows the characterization of
coordination number and type of ligands, and (ii) the
chemical shielding tensor elements of 113Cd are sensitive to
the chemical environment and provide an easy probe to
differentiate metal centers with identical coordination number
but different structure (for example: cadmium alaninate and
cadmium glycinate).4
In many of the metal-containing metalloproteins, the metal
centers form discrete units surrounded by various amino
acids. To determine the binding nature of the metal, it is
necessary to understand the coordination and structural
effects of these amino acids on the central atom. These effects
are identified either by looking at the metal center or by
looking at the ligands, namely, amino acids. It is known that
many of the amino acid residues such as glycine, glutamic
acid, histidine, cysteine, and aspartic acid in proteins bind
to a metal in metalloenzymes. Among them, the most
common amino acid that binds to a metal is histidine.
Imidazole present in histidine makes it a good complexing
agent.5 Moreover, imidazole plays an important role in
biologically active processes; it works as a proton donor and/
or acceptor.6 The ability to act as a proton donor was found
in the case of triosephosphate isomerase, where it forms a
hydrogen bond with the substrate leading to a lower energy
for the transition state.7 The imidazole moiety also enables
charge-transfer processes. One system studied extensively
was the oxygen-evolving complex in photosystem II,8 which
is responsible for the oxidation of water to dioxygen. In this
system, the imidazole moiety of the histidine appears to be
bound to the manganese cluster. The other system of
importance is phosphotriesterase (PTE).9 Cadmium-substi-
tuted PTE was found to have two cadmium centers with
hexa- and pentacoordination around the cadmium.10 Both of
these centers had two monodentate histidines chelating the
central cadmium atom. There are several other metals such
as zinc, copper, and iron that bind to the imidazole moiety
in histidine present in proteins like alkaline phosphatase5a,
plastocyanin,11 and ferredoxins.12 Thus, examining chemical
properties of imidazole, imidazole derivatives, and their metal
complexes is essential in understanding the binding nature
of imidazole. Further, zinc-containing proteins show three
different coordination numbers, 4, 5, and 6, predominantly
forming tetracoordinated species13 and are influenced by the
presence of water. Isotropic chemical shifts and scalar
coupling constants obtained from the solution NMR experi-
ments provided some insight about the structure but were
not conclusive. To get a better understanding of the structure
in rigid metal containing enzymes, NMR experiments should
be performed in the solid state. Moreover, chemical shielding
anisotropy could provide more information on the nature of
coordinating atoms and their positions around the metal. For
these reasons, solid-state NMR study of various model zinc
and cadmium complexes binding to imidazole and water
ligands will be of considerable interest in establishing a
relationship between the structure and chemical shift pa-
rameters. These NMR parameters, in turn, can be used to
characterize the structure at the metal centers in large
proteins.
Recently, with the advent of higher magnetic fields and
better radio frequency pulse techniques, it is possible to detect
zinc directly, which has a spin quantum number 5/2. Ellis et
al. in their latest work exploited cross-polarization and
quadruple echo experiments to directly detect different zinc
atoms present in zinc formate dihydrate.14 Using these
experiments, they were able to identify the two different Zn
atoms present in the crystal structure. In spite of its
advantages, direct detection of zinc is expensive and the
experimental techniques are technically demanding. Com-
pared to this method, it is relatively easier to determine the
change in the chemical shifts of 15N and 13C nuclei present
near the metal coordination center as a result of metal
coordination. There are few studies in the literature utilizing
â,â-[γ-13C] dideuteriohistidine as a nonperturbing 13C NMR
probe of the environments of the histidine residues in a zinc
(3) Kimblin, C.; Parkin, G. Inorg. Chem. 1996, 35, 6912.
(4) Barrie, P. J.; Gyani, A.; Motevalli, M.; O’Brien, P. Inorg. Chem. 1993,
32, 3862 and references cited within.
(5) (a) Otvos, J. D.; Browne, D. T. Biochemistry 1980, 19, 4011. (b) Braun,
J.; Schlabach, M.; Wehrle, B.; Kocher, M.; Vogel, E.; Limbach, H.
H. J. Am. Chem. Soc. 1994, 116, 6593. (c) Limbach, H. H.; Hennig,
J.; Kendricks, R.; Yannoni, C. S. J. Am. Chem. Soc. 1984, 106, 4059.
(d) Ellis, P. D.; Inners, R. R.; Jakobsen, H. J. J. Phys. Chem. 1982,
86, 1506.
(6) (a) Strohmeier, M.; Orendt, A.; Facelli, J. C.; Solum, M. S.; Pugmire,
R. J.; Parry, R. W.; Grant, D. M. J. Am. Chem. Soc. 1997, 119, 7114.
(b) Ramamoorthy, A.; Wu, C.; Opella, S. J. J. Am. Chem. Soc. 1997,
119, 10479. (c) Ghanotakis, D. F.; Babcock, G. T.; Yocum, C. F.
Biochim. Biophys. Acta 1984, 765, 388. (d) Nakatani, H. Y. Biochim.
Biophys. Acta 1984, 120, 299. (e) Miyao, M.; Murata, N. FEBS Lett.
1984, 168, 118. (f) Grove, G. N.; Brudvig, G. W. Biochemistry 1998,
37, 1532. (g) Debus, R. J. Biochim. Biophys. Acta 1992, 1102, 269.
(h) Chueng, W. Y. J. Biol. Chem. 1971, 246, 2589. (i) Forsen, S.;
Thulin, E.; Drakenberg, T.; Krebs, J.; Seamon, K. FEBS Lett. 1980,
117, 189.
(7) (a) Munowitz, M.; Buchovchin, W. W.; Herzfeld, J.; Dobson, C. M.;
Griffin, R. G. J. Am. Chem. Soc. 1982, 104, 1192. (b) Bachovchin,
W. W. Biochemistry 1986, 25, 7751. (c) Smith, S. O.; Farr-Jones, S.;
Griffin, R. G.; Bachovchin, W. W. Science 1989, 244, 961. (d) Yufeng,
W.; de Dios, A. C.; Mcdermott, A. E. J. Am. Chem. Soc. 1999, 121,
10389.
(8) (a) Lodi, P. J.; Knowles, J. R. Biochemistry 1991, 30, 6948. (b) Cleland,
W. W.; Kreevoy, M. M. Science 1994, 264, 1887. (c) Frey, P. A.;
Whitt, S. A.; Tobin, J. B. Science 1994, 264, 1927.
(9) Dumas, D. P.; Caldwell, S. R.; Wild, J. R.; Raushel, F. M. J. Biol.
Chem. 1989, 264, 19659.
(10) Zheng, F.; Zhan, C.-G.; Ornstein, R. L. J. Phys. Chem. B 2002, 106,
717 and references therein.
(11) (a) Dismukes, G. C. Chem. ReV. 1996, 96, 2902. (b) Ruttinger, W.;
Dismukes, G. C. Chem. ReV. 1997, 97, 1. (c) Yachandra. V. K.; Sauer,
K.; Klein, M. P. Chem. ReV. 1996, 96, 2927.
(12) (a) Kojiro, C. L.; Markley, J. L. FEBS Lett. 1983, 162, 52. (b) Chan,
T.; Markley, J. L. Biochemistry 1983, 22, 5982.
(13) Alberts, I. L.; Nadassy, K.; Wodak, S. J. Protein Sci. 1998, 7, 1700.
(14) Lipton, A. S.; Smith, M. D.; Adams, R. D.; Ellis, P. D. J. Am. Chem.
Soc. 2001, 124, 411 and references cited within.
Inorganic Chemistry, Vol. 42, No. 9, 2003 3143

Kidambi et al.
metalloenzyme alkaline phosphatase5a and 15N-enriched
strength during the data acquisition was 75 kHz. A 2 ms contact
time for CP and a recycle delay of 5 s were used. 13C and 15N
chemical shifts were referenced indirectly to adamantane (29.5 and
35.6 ppm) and ammonium sulfate powder sample (24.1 ppm),
respectively. For 113Cd NMR experiments, both static and MAS
experiments were performed on bis(histidinato)cadmium(II) com-
plex and cadmium formates in hydrous and anhydrous states.
Cadmium formate was used to set up the CP condition. 113Cd
chemical shifts are referenced relative to 0.1 M Cd(ClO4)2 (0.0
ppm). Magnitudes of the principal elements of the 113Cd CSA tensor
were determined from the powder spectrum and/or slow spinning
spectra using the Herzfeld-Berger sideband analysis program.20
The static powder spectrum was simulated using a FORTRAN-77
program. The principal elements of the chemical shift tensors are
represented according to the convention |σ33| g |σ22| g |σ11|.
Ab Initio Calculations. Isotropic and anisotropic chemical shift
values of 113Cd nuclei were calculated using Gaussian9821 and
utilized density functional theory22 quantum calculation methods.
The three-parameter hybrid functional B3PW9123 with the Kello-
Sadlej basis set23 on cadmium and 6-311+G23 on the other atoms
were used to estimate the 113Cd CSA values. We showed in our
earlier study that theoretical chemical shift values resulting from
the above-mentioned method agreed with the experimental values
to a reasonable accuracy.24 Initial positional coordinates were
obtained from neutron diffraction data.15 The values were referenced
with respect to the experimental isotropic chemical shift determined
from the powder spectrum.
imidazole ring in porphyrins to understand metal-porphyrin
bonding.5b-d These studies clearly showed the ability of 13
C
and 15N NMR to understand the structure around the metal
center.
In this paper, we use cross-polarization and magic angle
spinning (CPMAS) NMR spectroscopy to (a) determine the
chemical shift anisotropy (CSA) of the cadmium-113 nuclei
in cadmium histidinate and cadmium formates, (b) study the
effects of metal, both Zn and Cd, on natural abundance 15
N
and 13C chemical shifts in bis(histidinato)cadmium(II) hydrate
and bis(histidinato)zinc(II) hydrate complexes, and (c)
determine the 15N isotropic chemical shifts for [ZnIm6]Cl2
and bis(glycinato)zinc(II) hydrate complexes. The results
obtained from NMR experiments are correlated with well-
characterized X-ray structures.15 Experimentally determined
113Cd CSA values for cadmium histidinate are further
compared with ab initio values.
II. Experimental Section
Bis(histidinato) M(II) (M ) Cd, Zn) Hydrates. L-Histidine
was purchased from Fisher Biotech (Fairlawn, NJ), and metal-
histidine complexes were prepared using the procedure reported in
the literature.15 In brief, metal carbonate and histidine were mixed
in a 1:2 ratio in boiling water. The complexes were extracted and
recrystallized in water.
Cadmium Formates. Hydrous cadmium formate was purchased
from Aldrich (St. Louis, MO) and recrystallized in boiling water.16
The anhydrous compound was obtained from the hydrous sample
by keeping it in an oven at 140 °C for 2 days.
Gaussian calculations provide absolute shielding values, and
hence the chemical shift values of the complexes were obtained
using the equation
σ(calc) ) -σ(complex) + σ(ref)
Bis(glycinato)zinc(II) Hydrate. This sample was purchased from
TCI (Portland, OR) and used for NMR studies without further
purification.
Zinchexaimidazole Chloride Tetrahydrate. An aqueous solu-
tion of imidazole was added to zinc chloride in molecular ratio of
10:1. A trace amount of CuSO4 was added to this solution. The
sample was obtained by slow evaporation as suggested in the crystal
structure paper.17 Further elemental analysis of this complex
matched with the expected values.
III. Results and Discussion
A. 113Cd NMR Spectroscopy. (1) Bis(histidinato)cad-
mium (II) Hydrate, Cd[(C3H3N2)CH2CH(NH2)COO]2.
The structure of the bis(histidinato)cadmium(II) hydrate
complex is known from the neutron diffraction study.15 In
(20) Herzfeld, J.; Berger, E. J. Chem. Phys. 1980, 73, 6021.
(21) Frisch, M. J.; Trucks, G. W.; Schlegel, H. B.; Scuseria, G. E.; Robb,
M. A.; Cheeseman, J. R.; Zakrzewski, V. G.; Montgomery, J. A., Jr.;
Stratmann, R. E.; Burant, J. C.; Dapprich, S.; Milliam, J. M.; Daniels,
A. D.; Kudin, K. N.; Strain, M. C.; Farkas, O.; Tomasi, J.; Barone,
V.; Cossi, M.; Cammi, R.; Mennucci, B.; Pomelli, C.; Adamo, C.;
Clifford, S.; Ochterski, J.; Petersson, G. A.; Ayala, P. Y.; Cui, Q.;
Morokuma, K.; Malick, D. K.; Rabuck, A. D.; Raghavachari, K.;
Foresman, J. B.; Cioslowski, J.; Ortiz, J. V.; Stefanov, B. B.; Liu, G.;
Liashenko, A.; Piskorz, P.; Komaromi, I.; Gomperts, R.; Martin, R.
L.; Fox, D. J.; Keith, T.; Al-Laham, M. A.; Peng, C. Y.; Nanayakkara,
A.; Gonzalez, C.; Challacombe, M.; Gill, P. M. W.; Johnson, B.; Chen,
W.; Wong, M. W.; Andres, J. L.; Head-Cordon, M.; Replogle, E. S.;
Pople, J. A. Gaussian 98, revision A.5; Gaussian, Inc.: Pittsburgh,
PA, 1998.
(22) (a) Kohn, W.; Sham, L. J. Phys. ReV. A 1965, 140, 1133. (b) Becke,
A. D. J. Chem. Phys. 1988, 38, 2547. (c) Slater, J. C. Quantum theory
of Molecules and Solids. Vol. 4: The Self-Consistent Field for
Molecules and Solids; McGraw-Hill: New York, 1974.
(23) (a) Becke, A. D. J. Chem. Phys. 1988, 38, 3098. (b) Lee, C.; Yang,
W.; Par, R. G. Phys. ReV. B 1988, 37, 785. (c) Perdew, J. P.; Wang,
Y. Phys. ReV. B 1992, 45, 13244. (d) Becke, A. D. J. Chem. Phys.
1993, 98, 5648. (e) Becke, A. D. J. Chem. Phys. 1996, 104, 1040. (f)
Adamo, C.; Barone, V. Chem. Phys. Lett. 1997, 274, 242. (g) Kello,
V.; Sadlej, A. J. Theor. Chim. Acta 1995, 91, 353. (h) Clark, T.;
Chandrasekhar, J.; Spitznagel, G. W.; Schleyer, P. v. R. J. Comput.
Chem. 1983, 4, 294. (i) Stewart, J. J. P. J. Comput. Chem. 1989, 10,
209. (j) Raghavachari, K.; Pople, J. A.; Replogle, E. S.; Head-Gordon,
M. J. Phys. Chem. 1990, 94, 5579.
Solid-State NMR Spectroscopy. All NMR experiments were
carried out on a Chemagnetics/Varian Infinity 400 MHz spectrom-
eter operating at resonance frequencies of 400.1, 100.6, 40.5, and
88.7 MHz for 1H, 13C, 15N, and 113Cd, respectively at room
temperature. A 5 mm double-resonance Chemagnetics MAS probe
was used at different spinning speeds ranging from 1.0 to 9 kHz.
Cross-polarization (CP)18 and the TPPM decoupling method19 were
used to improve the sensitivity and resolution of the spectra. Typical
90° pulse widths were 3.5, 4.0, 5, and 3.5 µs for 1H, 13C, 15N, and
113Cd, respectively. The rf field strength used for 13C/1H and 15N/
1
1H cross-polarization was 35 kHz, and the H decoupling field
(15) (a) Huess, H.; Bartunik, H. Acta Crystallogr. 1976, B32, 2803. (b)
Kretsinger, R. H.; Cotton, F. A. Acta Crystallogr. 1963, 16, 651.
(16) (a) Post, M. L.; Trotter, J. Acta Crystallogr. 1979, B35, 2215. (b)
Weber, G. Acta Crystallogr. 1980, B36, 1947 (c) Harrison, W.; Trotter,
J. J. Chem. Soc., Dalton Trans. 1972, 956.
(17) (a) Garrett, T. P. J.; Guss, J. M.; Freeman, H. C. Acta Crystallogr.
1983, C39, 1031. (b) Lundberg, B. K. S. Acta Crystallogr. 1966, 21,
901.
(18) Schmidt-Rohr, K.; Spiess, H. W. Multidimensional Solid-state NMR
and Polymers; Academic Press: New York, 1994; Chapter 3. Shekar,
S. C.; Lee, D. K.; Ramamoorthy, A. J. Magn. Reson. 2002, 157, 223.
(19) Bennett, A. E.; Rienstra, C. M.; Anger, M.; Lakshmi, K. V.; Griffin,
R. G. J. Chem. Phys. 1995, 103, 6951.
(24) Kidambi, K. S.; Ramamoorthy, A. J. Phys. Chem. A 2002, 106, 10363.
3144 Inorganic Chemistry, Vol. 42, No. 9, 2003

Interaction of Cd and Zn with Biologically Important Ligands
Table 1. Cadmium-113 Isotropic and Anisotropic Chemical Shift
Values (ppm) for Cadmium Complexes
compound
σiso
σ33
σ22
σ11
∆σ
cadmium histidinate
theory
static
286
409
348
354
-42
102
88
-120
184
195
79
286a 416
cadmium formate (anhydrous) -55
-2
cadmium formateb dihydrate
Cd1
Cd2
14.4 45.6
19.9 85.1
17.4 -18.8 46.3
30.3 -55.8 97.9
cadmium acetateb dihydrate
cadmium glycinatec
cadmium alaninatec
-53.6 33.5 -69.5 -118.7 127.6
177
177
445
321
277
266
-189
-56
401
216
a σiso ) (σ11 + σ22 + σ33)/3; ∆σ ) [σ33 - (σ11 + σ22)/2]. b Reference
29. c Reference 4.
cinate consists of a distorted octahedral coordination with
two glycine ligands acting as bidentates chelating the central
cadmium atom through the nitrogen and oxygen atoms, and
the other two coordination sites are occupied by the oxygens
of two other glycine ligands, leading to a polymeric structure.
In the cadmium-alanine complex, two alanines act as
bidentates coordinating the central cadmium atom through
oxygen and nitrogen atoms. The two water oxygens also bind
to cadmium, leading to an octahedral geometry. Unlike the
cadmium glycinate structure, cadmium alaninate forms
discrete units with two bidentate alanines arranged in a cis
configuration.
Cadmium histidinate discussed in this paper does not form
a polymeric structure but forms discrete units, which are
linked to the neighboring molecules by hydrogen bondings
similar to those in cadmium alaninate. At the same time,
the two water molecules present in the cadmium histidinate
are not directly bonded to the metal, making it different from
the cadmium alaninate. Instead, cadmium histidinate forms
very weak intermolecular hydrogen bonds, making each of
these units highly distinct.
The 113Cd solid-state NMR spectrum of the cadmium
histidinate powder sample was obtained as shown in Figure
1B. The isotropic chemical shift was determined to be 286
( 3 ppm from the CPMAS spectrum (Figure 1C) of the 6
kHz sample spinning speed and further confirmed by running
the experiment at a different spinning speed (data not shown).
113Cd CSA values of cadmium histidinate were determined
from simulated static spectrum to be σ33 ) 416 ( 3 ppm,
σ22 ) 354 ( 3 ppm, and σ11 ) 88 ( 3 ppm as given in
Table 1. The simulated static spectrum is shown as a dotted
line superimposed onto the powder spectrum in Figure 1B.
The deviation between the simulated (dotted lines) and
experimental (solid line) powder patterns in Figure 1B could
be attributed to the following sources of experimental errors
that were not taken into account in the simulations. Pulse
imperfections such as resonance offset and rf field inhomo-
geneity could reduce the cross-polarization efficiency and
hence the S/N of the experimental powder pattern. Since the
span of the 113Cd chemical shift powder pattern is large (328
ppm), relative to the strength of the rf field used in the CP
experiment, for the bis(histidinato)cadmium(II) hydrate
complex, the above-mentioned pulse imperfections might
have been the main cause for the mismatch between the
simulated and experimental spectra in Figure 1. In addition,
Figure 1. (A) Neutron diffraction structure of cadmium histidinate used
in ab initio calculations and also to interpret chemical shifts measured from
solid-state NMR experiments. Cadmium-113 chemical shift spectra of bis-
(histidinato)cadmium(II) hydrate obtained at static (B) and 6 kHz (C)
spinning speeds. The CSA values are determined to be σ33 ) 416 ( 3
ppm, σ33 ) 354 ( 3 ppm, and σ33 ) 88 ( 3 ppm from computer simulation.
The simulated spectrum is superimposed onto the static powder spectrum
(see dotted line in part B). In part C, asterisks (*) denote spinning sidebands.
this complex, the cadmium center has a distorted octahedral
coordination with two histidine ligands chelating cadmium
through the imidazolic nitrogen (N2), amino nitrogen (N1),
and the carboxylic oxygens (O1) (See Figure 1A.) The
histidine complex presented in this work is structurally
distinct compared to previously studied cadmium alaninate
and cadmium glycinate complexes.4 Before starting the
discussion on cadmium histidinate, it is useful to present a
brief structural comparison between histidine and the other
two metal complexes in order to relate the structural
dependence of 113Cd chemical shift values. Cadmium gly-
Inorganic Chemistry, Vol. 42, No. 9, 2003 3145

Kidambi et al.
an inefficient decoupling of protons during the signal
acquisition could also lead to errors in the measurement of
the principal components of the 113Cd CSA tensor. The
isotropic chemical shift parameter calculated using the
magnitudes of the principal components of CSA, σiso ) (σ11
+ σ22 + σ33)/3, agreed with the value obtained from MAS
experiments. This isotropic chemical shift value differs by
approximately 110 ppm compared to the values of the two
other amino acid complexes mentioned above and as listed
in Table 1. Another complex that has a distorted octahedral
CdII environment with 4N, 2O arranged in a trans configu-
ration is the bis(2-aminomethylpyridine)dinitrato cadmium-
(II) complex. 113Cd NMR of this complex showed an
isotropic chemical shift peak at 222 ppm.25 The difference
of 64 ppm between these two cadmium complexes may be
attributed to the presence of different aromatic rings (imi-
dazole ring in the former case and pyridine in the latter
complex) and hydrogen bonds. However, the isotropic
chemical shift values of these two cadmium complexes are
much higher (or shifted toward downfield) than the one found
in the cadmium complex having similar coordination chem-
istry but with a cis configuration (52 ppm).26 The exact
reason for the effect of configurational differences on
chemical shifts is not well established.
respectively, and therefore the most shielded element, σ11,
should be oriented perpendicular to the carboxylate oxygen-
cadmium bond. Predicting the orientation of the least shielded
component, σ33, is difficult without single-crystal studies,
but we can conclude that it will be dependent on the plane
containing the four nitrogen atoms (two imidazolic nitrogens
and two amino nitrogens) on the basis of the empirical rules.
The magnitudes of the principal elements of the 113Cd CSA
tensor were also theoretically calculated on this molecule
using density functional theory and found to be σ33 ) 414
ppm, σ22 ) 343 ppm, and σ11 ) 102 ppm. These are in good
agreement with the experimental values (see Table 1). The
largest difference in the CSA values is seen for σ11
(calculated value is 14 ppm downfield to the experimental
value). However, this difference is relatively small compared
to the cadmium chemical shift span of 900 ppm for various
cadmium complexes.
(2) Cadmium Formates. Structures of both hydrous and
anhydrous cadmium formates have been determined using
X-ray crystallography.16 Hydrous cadmium formate has an
octahedral geometry with two structurally different cadmium
sites whereas anhydrous cadmium formate has a hepta-
coordinated pseudo-octahedral geometry. Another known
molecule, which has a seven-coordinate geometry, is cad-
mium acetate dihydrate.16 Figures 2A and 2B show the
crystal structures of anhydrous cadmium formate and cad-
mium acetate dihydrate, respectively. In anhydrous cadmium
formate, there are five monodendate formate oxygens [O(1),
O(2), O(4), O(5), and O(6)] and one bidentate formate
oxygen O(3) and O(7) encompass the central cadmium atom,
whereas in cadmium acetate dihydrate, two water oxygens,
O(1) and O(2), two pairs of bidentate acetate oxygens, O(3)
and O(4) and O(5) and O(7), and a monodendate bridging
oxygen, O(6), encompass the central cadmium atom. The
biggest difference between the two complexes is the occur-
rence of directly bonded water oxygens in cadmium acetate
dihydrate. This change in structure led us to explore the 113Cd
CSA tensor in anhydrous cadmium formate.
The full width at half-maximum (fwhm) of the isotropic
peak (Figure 1C) is almost 6 ppm and appears to be a
multiplet. This resembles the fine structure seen in the spectra
of other amino acids (glycine, alanine)4 and imidazole
cadmium complexes,27 which were related to the indirect
coupling (J-coupling) between 113Cd and two equivalent 14
N
nuclei. O’Brien and co-workers4 rationalized the multiplet
pattern for both the amino acid complexes using the
expression developed by Olivieri.28 Even though the spectrum
of the bis(histidinato)cadmium(II) hydrate sample shows
shoulders in Figure 1C, the multiplet pattern is not clearly
resolved; this could be due to the presence of two pairs of
similar 14N nuclei. A previous study of 113Cd CSA has shown
that the anisotropy of the shielding tensor can be correlated
with the differences in bond length around the metal center.
Also the presence of a water molecule resulted in a large
deshielding of the metal.29 Honkonen and Ellis29 have
suggested empirical rules to account for the orientations of
the three principal shielding tensors. According to this rule,
(i) the least shielded tensor element is aligned nearly
perpendicular to the plane containing water oxygens and for
molecules devoid of oxygens, (ii) the most shielded element
is perpendicular to the longest Cd-O bond, and (iii) the least
shielded element orients to maximize the shortest Cd-O or
Cd-N shielding contributions. In the case of cadmium
histidinate (Figure 1A), the bond distances for Cd-N(1),
Cd-N(2), and Cd-O(1) are 2.287, 2.290, and 2.480 Å,
Single-crystal solid-state NMR study on hydrous cadmium
formate by Honkonen and Ellis29 identified the presence of
two distinguishable cadmium sites as well as the orientation
of the tensor elements in the two sites. Hence, we will
emphasize the results for the anhydrous complex in this
article. TGA analysis on cadmium formates showed that
hydrous form loses its water and becomes anhydrous above
110 °C.30 For the present solid-state NMR study, the hydrous
sample was placed in the oven at 140 °C for 2 days to
completely remove water.
In order to corroborate and differentiate the presence of
two different forms of cadmium formates, 113Cd CPMAS
spectra for both hydrous and anhydrous forms were collected
at the sample spinning speed of 6.5 and 3.5 kHz as shown
in Figures 2C and 2E, respectively. The spectra resulted in
one sharp isotropic peak at -55 ( 3 ppm from the anhydrous
sample (minor peak at -75 ppm may be due to some other
(25) Griffith, E. A. H.; Li, H.; Amma, E. L. Inorg. Chim. Acta 1988, 148,
203.
(26) Rodeseiler, P. F.; Charles, N. G.; Griffith, E. A. H.; Lewinski, K.;
Amma, E. L. Acta Crystallogr. 1986, C42, 538.
(27) Rivera, E.; Kennedy, M. A.; Adams, R. D.; Ellis, P. D. J. Am. Chem.
Soc. 1990, 112, 1400.
(28) (a) Olivieri, A. C. J. Magn. Reson. 1989, 81, 201. (b) Olivieri, A. C.;
Hatfield, G. R. J. Magn. Reson. 1991, 94, 535.
(29) Honkonen, R. S.; Ellis, P. D. J. Am. Chem. Soc. 1984, 106, 5488.
(30) Abousekkina, M. M.; Morsi, S.; Elgeassy, A. A. Thermochim. Acta
1985, 91, 1.
3146 Inorganic Chemistry, Vol. 42, No. 9, 2003

Interaction of Cd and Zn with Biologically Important Ligands
Figure 2. Crystal structures of (A) anhydrous cadmium formate and (B) cadmium acetate dihydrate. Cadmium-113 CPMAS spectra of anhydrous cadmium
formate spun at 3.5 kHz (C) and 1.3 kHz (D) and hydrous cadmium formate spun at 6.5 kHz (E). One thousand transients obtained from ≈50 mg of powder
sample were coadded for each spectrum. Asterisks (*) denote the spinning sidebands. Plus signs (+) designate peaks of an impurity or a different form in
the sample and its spinning sidebands.
form of anhydrous cadmium formate) contrasting to two
peaks at 14.4 and 19 ppm seen in the hydrous form. Although
it is difficult to generalize coordination chemistry with
chemical shifts, Rodeseiler and Amma31 have proposed that
cadmium chemical shifts between -30 and -100 ppm
indicate a coordination number greater than 6. On the basis
of this rule, we can deduce the presence of seven-coordinated
cadmium center in the anhydrous cadmium formate. The
magnitudes of the three shielding tensor elements were found
components, σ22 and σ33, differ by 28 ppm (upfield) and 34
ppm (downfield), respectively, from that of cadmium acetate
dihydrate values (σ33 ) 33.5 ppm; σ22 ) -69.5 ppm)29,32 as
given in Table 1. The difference in the chemical shielding
values between these two complexes is small compared to
the cadmium chemical shift span of 900 ppm as mentioned
earlier. This correlation in NMR parameters can be linked
to the bond distances for the seven Cd-O bonds as listed in
Table 2. In the case of cadmium acetate dihydrate, bond
distances range between 2.294 and 2.597 Å and between
2.259 and 2.599 Å in anhydrous cadmium formate. Two of
the longest bonds seen in both the structures are from
bidentate formate (O3 and O7 in Figure 2A) and acetate
ligands (O3 and O7 in Figure 2B). Also, two oxygens from
to be σ33 ) -2 ( 3 ppm, σ22 ) -42 ( 3 ppm, and σ11
)
-120 ( 3 ppm using Herzfeld-Berger spinning sideband
analysis of the 1.3 kHz spectrum (Figure 2D). Magnitude
of the most shielded element, σ11, for the anhydrous cadmium
formate is similar to that of cadmium acetate. The other two
(31) Rodeseiler, P. F.; Amma, E. L. J. Chem. Soc., Chem. Commun. 1982,
(32) Ganapathy, S.; Chacko, V. P.; Bryant, R. G. J. Chem. Phys. 1984,
81, 661.
182.
Inorganic Chemistry, Vol. 42, No. 9, 2003 3147

Kidambi et al.
Table 3. Nitrogen-15 Isotropic Chemical Shift Values (ppm) of
Histidine and Imidazole for Free and Metal-Bound (Cd, Zn) Complexes
Obtained Using CPMAS Experimentsa
compound
N1
N2
N3
Hisb
41.2
249.5
249.8
248.0
226.3
210.4
232.5
170.8
171.6
172.2
170.8
173.5
176.8
Hisc
imidazoled
Cd-His
Zn-His
ZnIm6Cl2
33.0
31.5
a All values are indirectly referenced relative to NH3 (0.0 ppm) by setting
ammonium sulfate at 24.1 ppm. b Present result. c Literature data.7d d Ref-
erences 36 and 37.
The effect of metal-histidine interaction on 15N chemical
shifts is studied by performing CPMAS experiments on both
zinc histidinate complexes and cadmium histidinate. In the
zinc histidinate spectrum (Figure 3B), the influence of
coordinate bonds on isotropic chemical shifts is prominent
with an upfield shift of 40 ppm for the imidazolic nitrogens
(N2) and 9.7 ppm for the amino nitrogens. The upfield shift
is related to the shielding effect caused by the presence of
the metal atom. The (N3) nitrogen of the zinc complex shifts
downfield from 170.8 to 173.5 ppm. Similarly, in the
cadmium histidinate spectrum (Figure 3C), the isotropic
chemical shift value of the imidazolic nitrogen (N2) changed
from 249.5 to 226.3 ppm, an upfield shift of 23 ppm, whereas
the amino nitrogen value shifted from 41.2 to 33 ppm, an 8
ppm upfield shift. The isotropic chemical shift of the other
imidazole nitrogen (N3) remained unaffected by the cadmium
histidinate complex formation. Unlike the zinc histidine
spectrum, the isotropic chemical shifts of unbound histidine
molecules are seen in the cadmium histidine spectrum, and
this is confirmed by the 13C CPMAS spectrum shown later.
All these values are given in Table 3. The line width of the
metal complexes increases compared to the unbound histidine
molecule and may be related to longer T2 relaxation. (See
the amino nitrogen peak at 33 ppm in cadmium histidine
complex in Figure 3.) Other studies in the literature reported
the influence of pH on the chemical shifts of imidazole and
histidine in solution.6 These reports demonstrated that a
lowering of pH resulted in an upfield diamagnetic shift for
the N1 nitrogen caused by protonation of the imidazolic
nitrogens and removal of the hydrogen from the imidazole
group at a higher pH led to downfield paramagnetic shifts
for both of the nitrogens in the imidazole group, but the effect
was less prominent on the N3 nitrogen.33 On the other hand,
15N chemical shifts of doubly labeled [15N]imidazole in metal
complexes permitted evaluation of Zn(II) complex formation
in homogeneous solution. In solution of Zn2+ and imidazole,
the 15N resonance in [Zn-imidazole]2+ complexes is dia-
magnetically (upfield) shifted by 10-20 ppm relative to
neutral aqueous imidazole.34 In the case of Cd-imidazole
complexes, coordination to Cd2+ produced an observed 8-12
ppm diamagnetic shift in the 15N resonance of imidazole
relative to neutral aqueous imidazole.35 Our solid-state NMR
Figure 3. Natural abundance CPMAS 15N chemical shift spectra of (A)
histidine, (B) zinc histidinate, and (C) cadmium histidinate. The spinning
speed was 7 kHz. Ten thousand transients obtained from ≈50 mg of powder
sample were coadded for each spectrum.
Table 2. Cd-O Bond Distances (Å) from Cadmium Formate
(Anhydrous) and Cadmium Acetate Compounds
O1
O2
O3
O4
O5
O6
O7
cadmium acetate 2.299 2.325 2.546 2.303 2.294 2.296 2.597
cadmium formate 2.285 2.323 2.586 2.303 2.259 2.275 2.599
(anhydrous)
each water molecule (O1 and O2 in Figure 2B) in cadmium
acetate dihydrate have distances of 2.299 and 2.325 Å similar
to that of the two formate oxygens (2.285 and 2.323 Å).
Therefore, the minor difference in the shielding values may
be due to the presence of directly coordinated water
molecules in cadmium acetate dihydrate and the absence of
water molecules in cadmium formate. Now coming to the
analysis of the orientation of the shielding elements, σ11
should be orthogonal to the longest Cd-O(4) bond based
on the previously mentioned empirical rules. But the predic-
tion process for the other shielding components is more
difficult due to the absence of water molecules and the
presence of five monodendate formates in anhydrous cad-
mium formate.
B. 15N NMR of Metal-Histidine and Metal-Imidazole
Complexes. A commercial L-histidine sample was used for
preparing the metal (Zn, Cd) complexes as mentioned in the
Experimental Section. 15N chemical shifts of the three
nitrogens in histidine are highly dependent on the protonation
state and tautomerism,6 whereas in metal complexes the
presence of coordinate bonds between imidazolic nitrogen
and the metal prevents the formation of tautomers avoiding
complications during interpretation of NMR spectra. Natural
abundance solid-state 15N CPMAS spectrum of the histidine
sample resulted in three peaks at 41.2, 249.5, and 170.8 ppm
(Figure 3A) accounting for the three nitrogens, namely, (N1)
amino nitrogen, (N2) imidazole nitrogen, and (N3) imidazole
nitrogen, respectively. These values match with those avail-
able in the literature.7d
(33) Blomberg, F.; Ru¨terjans, H. Nucl. Magn. Reson. Spectrosc. Mol. Biol.
1978, 231.
(34) Mohammed, A., Jr.; Morgan, L. O.; Wageman, W. E. Inorg. Chem.
1978, 17, 2288.
3148 Inorganic Chemistry, Vol. 42, No. 9, 2003

Interaction of Cd and Zn with Biologically Important Ligands
Figure 5. Natural abundance CPMAS 13C chemical shift spectra of (A)
histidine, (B) zinc histidinate, and (C) cadmium histidinate. The spinning
speed was 7 kHz. One thousand transients obtained from ≈50 mg of powder
sample were coadded for each spectrum.
value of the solid imidazole sample. Moreover two peaks
are seen for the N3 nitrogens. The first peak is shifted
downfield to 176.8 ppm from 172.2 ppm relative to free
imidazole (values given in Table 3),36,37 resembling the trend
seen in the zinc histidinate, whereas the second peak occurs
at 172.6 ppm, which may be due to some unbound imidazole.
To identify this peak, we ran an experiment on a solid
imidazole sample doped with 1% CuSO4 and observed two
peaks for the two imidazolic nitrogens (N2, N3) at chemical
shift values similar to previously reported data.36,37 The S/N
ratio for this sample was found to be much lower than that
for the hexaimidazole complex, and this is due to a very
long T1 for the imidazole sample as known in the litera-
ture.36,37 The recycle delay used for the imidazole sample
was 60 s even after it was doped with 1% CuSO4 whereas
the recycle delay for the zinc-imidazole complex was 5 s.
On the basis of these two spectra, we guess that the second
N3 peak seen in the metal-imidazole complex is due to
unbound imidazole. Another perplexing aspect was the
absence of the N2 nitrogen peak arising from unbound
imidazole in this spectrum. This could be related to the longer
T1 for unbound imidazole as mentioned above, and also the
absence of hydrogen near the N2 nitrogen may lead to
Figure 4. (A) Crystal structure of hexaimidazolezinc(II) chloride. Natural
abundance CPMAS 15N chemical shift spectra of (B) hexaimidazolezinc-
(II) chloride and (C) imidazole. The samples were spun at 7 kHz. Ten
thousandtransients obtained from ≈50 mg of powder sample were coadded
for each spectrum. A recycle delay of 60 s was used for imidazole doped
with 1% CuSO4.
results agree with those obtained earlier in solution.33-35 From
the current and earlier studies, we can infer that attachment
of metal (Zn, Cd) to imidazolic nitrogen leads to an upfield
diamagnetic shift irrespective of the physical state. The
greater shift of imidazolic nitrogen (N2) and amino nitrogens
(N1) in the zinc complex can be accounted for by shorter
bond distance between metal and nitrogen (Zn-N1, 2.041
Å; Cd-N1, 2.290 Å; Zn-N3, 2.049 Å; Cd-N3, 2.287 Å).
Zinc has the ability to form both tetrahedral and octahedral
complexes17 with imidazole. The complex considered in this
study is ZnIm6Cl2, and the structure is shown in Figure 4A.
In ZnIm6Cl2, the central Zn atom is bonded to six imidazolic
nitrogens leading to an octahedral symmetry around the
metal. Natural abundance 15N CPMAS spectrum (see Figure
4B) of the hexaimidazole complex led to an upfield shift of
15 ppm for the N2 nitrogen relative to the chemical shift
1
inefficient transfer of magnetization from H to 15N in the
CPMAS experiment. As we are concerned more about the
effect of metal interaction on the chemical shift of the zinc-
bound N2 imidazolic nitrogen, which was clearly identified
by a distinct upfield change in the chemical shift from 248
ppm for unbound imidazole to 232.5 ppm using our
experiment, we did not analyze the reason for the absence
of the unbound imidazolic N2 nitrogen peak.
C. 13C NMR of Metal-Histidine Complexes. Figure 5
shows the natural abundance 13C CPMAS spectra of histidine
(36) Solum, M. S.; Altmann, K. L.; Strohmeier, M.; Berges, D. A.; Zhang,
Y.; Facelli, J. C.; Pugmire, R. J.; Grant, D. M. J. Am. Chem. Soc.
1997, 119, 9804.
(35) Mohammed, A., Jr.; Wageman, W. E.; Morgan, L. O. Inorg. Chem.
1978, 17, 3314.
(37) Hickman, B. S.; Mascal, M.; Titman, J. J.; Wood, I. G. J. Am. Chem.
Soc. 1999, 121, 11486.
Inorganic Chemistry, Vol. 42, No. 9, 2003 3149

Kidambi et al.
Table 4. Carbon-13 Isotropic Chemical Shift Values of Histidine
(ppm) for Free and Metal-Bound (Cd, Zn) Histidine Complexes
Obtained Using CPMAS Experimentsa
compound
C1
C2
C3
Câ
CR
C0
His
Cd-His
Zn-His
137.8
140.5
139.0
135.2
135.7
137.1
114.3
116.8
116.4
27.3
29.5
28.2
57.6
56.3
55.5
175.4
180.5
178.6
a All values are indirectly referenced relative to TMS by setting
admantane at 29.5 and 35.6 ppm.
(Figure 5A) and its metal complexes (zinc, cadmium
histidinates in Figures 5B and 5C, respectively). A histidine
molecule has six chemically distinct carbons, and the 13C
CPMAS spectrum obtained at 7 kHz spinning speed shows
clearly resolved isotropic peaks for all the carbons. The 13
C
CPMAS spectra of histidines bound to zinc ions also show
highly resolved isotropic chemical shift peaks with altered
chemical shift values due to the metal ion interactions. The
cadmium histidinate spectrum shows highly resolved 13C
peaks for both bound and unbound histidines (Figure 5C).
This is consistent with the 15N CPMAS spectrum seen in
Figure 3C. The presence of peaks from both bound and
unbound histidines was helpful in understanding the effect
of cadmium on the 13C chemical shifts. Two isotropic
chemical shift peaks for the carboxylate carbon in the
cadmium histidinate sample occurred at 175.4 and 180.5 ppm
(Figure 5C), where the former is for unbound histidine and
the latter in the downfield region is due to coordination
between cadmium and carboxylate oxygen. It is also seen
from Figure 5C that isotropic chemical shifts of C1, C2, C3,
and Câ carbons move downfield due to the interaction of
the metal with histidine, whereas the chemical shift of CR
carbons lying adjacent to the carboxylate carbons shifts
upfield. These effects are also seen in the zinc histidinate
with the downfield-shifted carboxylate peak occurring at
178.5 ppm. 13C chemical shift values for both unbound and
metal-bound histidine are given in Table 4. The results match
with the previous studies reported by Couves et al. on zinc-
histidine complexes in solution.38 Using titration curves, they
showed that 13C signals of carboxylate carbon shifted
downfield from 175 to 183 ppm upon coordination of varying
ratios of histidine to the zinc center.
D. 15N NMR of Zinc Glycinate. To further see the effect
of metal and ligand interaction in biologically relevant
molecules, 15N CPMAS experiments were performed on
glycine and zinc glycinate powder samples. X-ray crystal-
lography on Zn-glycine complex39 showed the presence of
coordinate bonding between amino nitrogen and the zinc
atom. On the basis of the metal-histidine study, it is
expected that the binding of metal to amino nitrogen of
glycine should lead to an upfield shift relative to that of the
free glycine chemical shifts. As anticipated, spectra shown
in Figures 6A & 6B for glycine and zinc glycinate samples,
respectively, clearly show the change in the chemical shift
Figure 6. Natural abundance CPMAS 15N chemical shift spectra of (A)
glycine and (B) zinc glycinate. The samples were spun at 7 kHz. One
thousand transients obtained from ≈50 mg of powder sample were coadded
for each spectrum.
from 30.1 ppm for the amino nitrogen of unbound glycine
to 15.1 ppm for the zinc-glycine complex.
IV. Conclusions
This paper illustrates the importance of cadmium chemical
shift tensors in model biological complexes. The 113Cd solid-
state NMR experimental data show an upfield shift of 286
ppm for bis(histidinato)cadmium(II) hydrate and a downfield
shift of -55 ppm for anhydrous cadmium formate relative
to cadmium perchlorate. Also, cadmium histidinate has a
CSA span of 330 ppm, which is smaller than the cadmium
glycinate and cadmium alaninate complexes. Moreover, the
calculated chemical shift values using density functional
theory matched well with the experimental data. The marked
difference in the chemical shift values for the cadmium
histidinate and bis(2-aminomethylpyridine) dinitrato cad-
mium(II) complexes shows that cadmium chemical shifts are
influenced not only by the neighboring ligand atoms but also
by bond distances and other secondary effects. In the case
of cadmium formates, the effect of different coordination
number and the presence of water on the cadmium chemical
shift values was clearly seen by the upfield cadmium
chemical shift values for anhydrous cadmium formate
compare to dihydrate (14.4 and 19 ppm for cadmium formate
dihydrate; -55 ppm for anhydrous cadmium formate). Along
with the chemical shift of cadmium nuclei, 13C and 15N
chemical shifts also provided useful information on the
histidine and imidazole complexes. Bonding of nitrogens N1
and N3 and the carboxylate oxygen shifted the chemical shift
values upfield and downfield, respectively, relative to
unbound histidine. Further, 15N NMR data on zinc glycinate
clearly reflected diamagnetic upfield shifts similar to that of
histidine nitrogens. These data indicate that 113Cd, 15N, and
13C NMR are all useful nuclei to probe metal centers in
intermediates or in biologically potent molecules. Availability
of ultrahigh magnetic fields (ca. 900 MHz) and a plethora
of higher resolution multidimensional solid-state NMR
(38) Couves, L. D.; Hagne, D. N.; Moreton, A. D. J. Chem. Soc., Dalton
Trans. 1992, 217.
(39) (a) Newman, J. M.; Bear, C. A.; Hambley, T. W.; Freeman, H. C.
Acta Crystallogr. 1990, C46, 44. (b) Low, B. W.; Hirshfeld, F. L.;
Richards, F. M. J. Am. Chem. Soc. 1959, 81, 4412.
3150 Inorganic Chemistry, Vol. 42, No. 9, 2003

Interaction of Cd and Zn with Biologically Important Ligands
techniques would enable experimental analysis of 113Cd CSA
tensors from a variety of biological systems such as metal-
loproteins and metal-bound RNA. In addition, recovery of
CSA tensors under fast magic angle spinning conditions40
provides 113Cd CSA tensors from multiple sites of biological
complexes. One specific system to be addressed in the future
is the coordination around the active sites in phosphotri-
esterase. Solution NMR has shown two cadmium chemical
shifts at 116 and 212 ppm, but the exact structure is uncertain.
As an extension of this work, we are currently working on
model complexes to find out the exact structure around these
metal sites using both experiments and ab initio calculations.
Acknowledgment. This study was supported by the
research funds from the National Science Foundation (CA-
REER development award to A.R.). We thank Dr. Colaneri
for his suggestions on the preparation of cadmium histidinate
complex, Dr. Rajendiran for useful comments on the
manuscript, and Manolis from Dr. Pecoraro’s lab for help
with the figures.
(40) Wei, Y.; Lee, D. K.; Mcdermott, A. E.; Ramamoorthy, A. J. Magn.
Reson. 2002, 158, 23.
IC026287D
Inorganic Chemistry, Vol. 42, No. 9, 2003 3151