NMR studies of metabolism.

ANNUAL REVIEWS

Rev. Biophys. Biophys. Chern. 1990. 19:43-67 Copyright © 1990 by Annual Reviews Inc. All rights reserved

Annu.

Further

Quick links to online content

Annu. Rev. Biophys. Biophys. Chem. 1990.19:43-67. Downloaded from www.annualreviews.org by University of Chicago Libraries on 02/17/13. For personal use only.

NMR STUDIES OF METABOLISM Sebastian Cerdan I and Joachim Seelig

Biocenter of the University of Basel, Klingelbergstrasse 70, CH-4056 Basel, Switzerland KEY WORDS:

noninvasive methods, stable isotopes, metabolic regulation, in vivo NMR, localization techniques.

CONTENTS PERSPECTIVES AND OVERVIEW............................................. . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . .

43

SPATIAL LOCALIZATION METHODS FOR IN VIVO SPECTROSCOPy........................................

45

3IPNMR........................................................................................................................

47

Surface Coils.... ................... .. ........................... Image-Guided Spectroscopy...................................................................................... Metabolites Detectable by 31p NMR........................................................................ Intracellular Compartmentation.. .. ....... ................ In Situ Control of Oxidative Phosphorylation ................................ ...... ........ . ............

45 45 47 50

51

13C NMR........................................................................................................................

53

Gluconeogenesis........................................................................................................ Free Fatly Acid Metabolism and Ketogenesis ... .. ......... .............. Glycogen Synthesis and Degradation . ....... .......... . ........................................ . ............ Triglyceride and Phospholipid Metabolism .............. . ................................................

55

Tricarboxylic Acid Cycle and Related Anaplerotic Reactions....................................

IH NMR.........................................................................................................................

19F NMR ........................................................................................................................

2H NMR.........................................................................................................................

54 55

56 57

58 61

62

PERSPECTIVES AND OVERVIEW

Since its discovery by Bloch and Purcell in 1 946, nuclear magnetic reson ance (NMR) has evolved into a highly successful tool of physics, chemistry, I

Present address: Instituto de Investigaciones Biomedicas del C.S.I.C .. c/ Arzobispo

Morcillo 4, 28029 Madrid, Spain.

43 0883-9 1 82/90/06 1 0-0043$02.00


44

CERDAN & SEELIG

and biochemistry. In the last ten years new NMR applications have emerged as superconducting magnets with a horizontal bore large enough to accommodate animals or humans have been developed. For the first time it has become possible to study biological mechanisms and metabolic pathways in man and animals in a noninvasive and nondestructive manner. Typical subjects include the study of the energy metabolism of skeletal and heart muscle by 31p NMR; the study of the tricarboxylic acid cycle, of glycogen synthesis and degradation, and of gluconeogenesis and keto genesis in the liver by 13C NMR; and, most recently, the measurement of metabolites in brain by IH NMR. Modern NMR studies of in vivo metabolism probably began in 197 2 when Eakin et al (68) employed 11C NMR to monitor the metabolism of [1-13C]glucose in a eucaryotic cell system. In 1973 , Moon & Richards (121) used 31p NMR to measure inorganic phosphate and 2,3-diphospho glycerate (DPG ) in erythrocytes and to determine the intracellular pH from the separation of the phosphorus resonances. Lauterbur (106) demonstrated the use of IH NMR imaging to obtain X-ray-like images of living tissue and, almost simultaneously, the first 31P N M R spectrum of animal tissue was published (91). The study of in vivo metabolism by NMR is particularly attractive for several reasons. First, NMR experiments provide the opportunity to perform repetitive, noninvasive measurements using the same biological system. Second, unique NMR properties such as the TI and Tz relaxation times or the homonuclear and heteronuclear spin coupling patterns can be measured. These parameters contain specific information on the physio logical or pathological state of the tissue under consideration (65), on metabolic compartmentation (27), or on the flux through specific mctabolic pathways (108, Il l ) . Third, unidirectional reaction rates of enzymatic reactions can be followed in situ using magnetization transfer techniques (5). Finally, it is now possible to obtain spatially resolved metabolic infor mation yielding the distribution of metabolites within a tissue or organ (3 2, 42, 131), thus opening a new avenue to study pathology and therapy at the molecular level (150, 1 87 ). In this review we present applications of NMR to the study of different aspects of metabolism. We begin with a brief outline of localization methods that are commonly used to obtain in vivo NMR spectra. We then describe in more detail metabolic information recently obtained by NMR of perfused organs, intact animals, and humans. Previous reviews have already covered the applications of NMR to the study of metabolism in microorganisms (35), isolated or cultivated cells (3 4, 1 23, 174), and tumors (64). NMR spectroscopy of the brain (1 48), and human in vivo NMR spectroscopy (31, 1 87 ) have also been reviewed recently.

NMR

STUDIES OF METABOLISM

45

SPATIAL LOCALIZATION METHODS FOR


IN VIVO SPECTROSCOPY

A major problem associated with in vivo spectroscopy is the proper local ization of the region-of-interest (ROI). Guided by a proton MR image of the object, it should be possible to define the ROI directly on the imaging screen and to measure IH_, 31p_, or 13C-NMR spectra of the selected volume. At present, no simple method exists to solve this problem. Many different strategies have been advocated ( 1 0, 3 1 , 79, 1 79), depending, in part, on the technological state of the NMR instrument. In the following sections we summarize four methods that have proven useful in a variety of applications: (a) surface coils, (b) volume selective excitation (VSE), (c) STEAM, and (d) ISIS. Although a surface coil used alone is spatially selective, it can also be combined with one of the latter pulse sequences to further improve selectivity. Surface Coils

Surface coils are probably used in more than 80% of the published in vivo NMR studies. Although they have an inhomogeneous B1-field and a limited penetration depth, their high filling factor results in a good signal to-noise ratio. Furthermore, surface coils are easy to handle and they are the method of choice for the measurement of superficial tissue. The sensitivity and selectivity of surface coils has been investigated theoretically and experimentally (2, 69, 1 25). Recent improvements include the use of adiabatic pulses ( 1 70, 1 83, 1 84). The penetration depth of a surface coil corresponds roughly to the diameter of the sphere. Superficial tissue may be suppressed by choosing a 1 800 pulse length at the position of the surface. A useful variant of the surface coil technique is spectroscopic rotating-frame imaging (59, 1 79). The pulse angle for excitation is incremented in small steps, and for each pulse angle the corresponding free-induction decay (FID) is collected. A two-dimensional Fourier transformation will then resolve the chemical shift along one axis and the position along the other. Image-Guided Spectroscopy

Improved control of the size, shape, and position of the ROI is obtained by combining linear magnetic field gradients with frequency-selective pulses. The methods and pulse sequences employed for localization bear a close resemblance to NMR imaging routines. In volume-selective excitation spectroscopy (VSE; cf Figure IA) a fre quency-selective pulse applied in the presence of a slice gradient Gx selects

46

CERDAN & SEELIG

A

• Acq.


Gx

B

SIice1

rf

Gz

Gy

0 90

__

0 90

{} !�3- {} _

Slice3

Slice 2 - -

_

!�

0 90

{}!�!�

___

_______

� - -G-Slice1 q----------- -- - --- - - -- ---

Signal

______ ---_____________

G-Slice2--- _____ G-Slice 3

Figure 1

r:::1.

______

r=-=l

- - - - [] - - - - - - - - - r:::l. _

__

_______

- -

-

- _

-

__

Schemes for image-guided in vivo spectroscopy. A: Volume-selective excitation

(VSE) (J I, 1 25, 1 26). B: Stimulated echo acquisition mode (STEAM) (73-75).

a homogeneous slice of the sample and rotates its magnetization into the plane. A nonselective pulse is intercalated between the "soft" pulses. It rotates the magnetization of the whole sample by 90° and that of the selected slice by an additional 90°. At the end of the pulse sandwich the magnetization of the selected slice is rotated by 1 800 and oriented along the negative z axis; it decays with the corresponding Tj relaxation time. On the other hand, the magnetization outside the selected slice is rotated into the x, y plane and is rapidly dephased by the applied gradient. This sequence is repeated in the remaining two orthogonal directions and the ROI is thus further narrowed, first to a rod and then to a small cube ( 1 1 , 1 25, 1 26). The major difficulty in applying the VSE sequence to a whole body system is that the 90° hard pulse usually cannot be made short enough when using whole-body coils. This problem is avoided by the SPARS sequence, which applies hard pulses only in the absence of gradients, thus considerably reducing the bandwidth ( 11 2). Like VSE the SPARS sequence preserves the m agnetization of the selected slice along the Bo field. x, y


N MR STUDIES OF METABOLISM

47

The use of stimulated echoes appears to be the method of choice for volume-selected proton-NMR spectroscopy C H NMR) (73-75). The STEAM technique (stimulated echo acquisition mode) consists of three frequency-selective 90° pulses that are applied in the presence of ortho gonal gradients and thus select a cube-shaped region of well-defined dimen sions (cfFigure IB). STEAM avoids broad-banded radio frequency pulses, which are difficult to generate with large body coils, and it allows a relatively simple shimming of the region of interest. Such shimming is absolutely mandatory in 'H-NMR with its small chemical shift range. STEAM is extremely sensitive to eddy currents induced by gradient switch ing, since the magnetization of the region of interest dephases and is recollected in the presence of Bo-gradients. A good compensation of eddy currents is therefore required. The proton-NMR spectrum of the brain of one of the authors (Figure 3) was obtained with STEAM. The ISIS sequence (137) also provides a three-dimensional localization that combines frequency-selective 1800 pulses in the presence of Bo gradi ents with nonselective 90° acquisition pulses in the absence of gradients. The whole sequence comprises eight different experiments, and the region of-interest is generated by an add-and-subtract scheme. Since the magnet ization of interest is stored along the z-axis, ISIS is much less sensitive to eddy currents. Multivolume selective spectroscopy can be considered as a variant of ISIS (87, 124, 127). The experiment uses slice-selective inversion pulses that simultaneously excite several different slices of the same object. Thus, different regions within the same organ [e.g. the heart (127)] can be monitored at the same point in time. 31p NMR

The majority of NMR studies in living systems have been concerned with 31p NMR investigations of energy metabolism and its regulation in a variety of tissues including primarily skeletal muscle (42-48,120,143,144), hcart (96, 138), and brain (148). The following aspects are considered here: (a) metabolites that can be observed in situ by 31p NMR, (b) intracellular metabolic compartmentation and 31p NMR visibility, (c) in situ control of oxidative phosphorylation as viewed by 31p NMR. Metabolites Detectable by 31p NMR

An in vivo 31p NMR spectrum obtained from human calf muscle at 2 T in a whole body magnet routinely used for MR imaging is shown in Figure 2 (23). Five resonances, which can be assigned to the /3-, 11- and y phosphates of A TP, phosphocreatine (Per), and inorganic phosphate (Pi), respectively, are observed. Other nucleotide triphosphates arc normally

48

CERDAN & SEELIG

PCr

Figure 2

31p NMR spectrum of human

calf muscle obtained at 2T on a whole body


instrument. Resonances are assigned to

ATP, per, and Pi (23).

I

30.

20.

10.

PPM

O.

-10.

present in muscle in much smaller concentrations than A TP and are not believed to contribute significantly to the in vivo 31p NMR spectra. With few remarkable exceptions (72, 117), the a- and f)-phosphate resonances of ADP overlap in vivo with the (J.- and y-phosphate resonances of A TP. Therefore, the contribution of ADP to the 31p NMR spectrum cannot be determined directly in most cases. An indirect procedure to calculate ADP concentrations when the f)-ATPfy-ATP ratio is very close to unity is described later. In addition to resonances of nucleotides, per, and Pi, two further signals are normally observed in the 31p NMR spectra of tissues and are inter preted as phosphomonoester and phosphodiester signals. The phospho monoester resonance is a composite and contains contributions from phos phorylethanolamine, phosphorylcholine, fructose-IP, IMP, etc. The main component in brain under basal conditions is phosphorylethanolamine (86), and in liver is phosphorylcholine (66, 95), but significant contributions from inositol- l P in brain (152) or from sugar phosphates in liver or muscle (44, 93) have been detected following pilocarpine treatment, fructose infusions, or exercise-induced glycogenolysis. More recently, very low field signals (16.5 ppm) arising from inositol 1,2-cyclic phosphate have been detected in tumor cells (83). The phosphodiester resonance in vivo is also a composite signal con taining contributions mainly from glycerophosphorylcholine (GPC) and glycerophosphorylethanolamine (GPE) (36). Interestingly, in the in situ spectra of dog brain a third species, i.c. highly mobile phospholipids, also contribute to the resonance (41). This is evidenced by two facts: (a) In aqueous extracts of the brain the intensity of the phosphodiester resonance is distinctly smaller than predicted on the basis of the in vivo spectra,

NMR STUDIES OF METABOLISM

49


which indicates the presence of a diester that is hydrophobic and cannot

be extracted into the aqueous phase; (b) the phosphodiester resonance broadens selectively with increasing field strength (156), which is typical for relaxation of phospholipids via chemical shift anisotropy (28). Recently, a characteristic phosphodiester signal was found in cultured tumor adcno carcinoma cells and was shown to arise from highly mobile phospholipids (84). Quantitative information concerning the relative concentrations of the different metabolites monitored above can also be obtained. If one assumes that the volume detected is the same for all metabolites, relative variations in concentration can easily be followed if the spectra are acquired under nonsaturating conditions or are corrected for the differences in relaxation times (113, 15 1 , 176). The determination of absolute concentrations by in vivo NMR is more difficult. The most common procedure compares the resonance of interest with the area of an endogenous metabolite of known concentration, e.g. the fJ-ATP peak (180). However, other methods involv ing comparison with an exogenous standard present in an external capillary or accumulated in the tissue of interest, or with the IH NMR signal of water detected by a doubly-tuned surface coil, can be used (181). Measurement of the relative or absolute concentrations of 31p NMR observable metabolites allows the calculation of the in situ concentration of ADP and of the phosphoryl ation potential. These procedures assume that the creatine phosphokinase reaction is at equilibrium (which might not always be the case) and require knowledge of the intracellular pH, the creatine concentration (or the total creatine + phosphocreatine pool), and the value for the equilibrium constant of the creatine phosphokinase reac tion (185). The following expressions are used: [ADP] =

[ATP]· [Cr] [PCr]. KCPK· [H+]

[ATP] [ADP] [Pi]

[PCr] KCPK[H+] . [Pi] [Cr]

1. 2.

where [ATP], [PCr], [Pi], and [H+] represent the free concentrations of metabolites, which can be determined from the 31p NMR spectra, and [Cr] or KCPK are known from ancillary information. For a given condition, the ratio [Cr]/{KCPK· [H+]} is constant; therefore a qualitative index reflecting the relative changes in the concentration of ADP or the phosphorylation potential can be obtained by a simple examination of the ATP/PCr or Per/Pi ratio. In addition to detecting phosphorus-containing metabolites, 31p NMR also allows the measurement of (a) the intracellular pH and (b) the free

50

CERDAN & SEELIG

Mg2+ concentration. We do not address these questions here. Pertinent references are (12, 77,81,85,103,146).


IntracelluLar Com partmentation

Perhaps one of the most interesting results obtained by 31p NMR is the difference between the ADP and Pi concentrations measured in vivo by NMR and those measured by classical enzymatic methods in perchloric acid (peA) soluble metabolite extracts. Table 1 presents an illustrative comparison. While the concentrations of A TP and per are similar when measured by 31p NMR and classical methods, the in situ concentrations of Pi and ADP are remarkably lower with 31p NMR as compared to conventional acid extraction methods. The increase in total ADP and Pi in the peA extracts cannot be accounted for by an equivalent decrease in the A TP or per total concentrations. This indicates that the excess ADP and Pi cannot be produced by a breakdown of high energy phosphates during the extrac tion procedure and suggests the existence of pools of ADP and Pi that are not detectable by in vivo 31P NMR. Thcsc invisible pools account for approximately 50% (75%) of the total Pi and 70% (100%) of the ADP in liver (heart) (66, 80). There are several reasons to believe that only the cytosolic Pi is NMR visible. First, it is known from experiments with fast subcellular frac tionation tcchniques that approximately 50% of the total cellular Pi is mitochondrial in perfused rat liver (167, 172) and Langendorff perfused heart (100), a percentage that clearly matches the observations presented Table 1

Comparison of the concentration of phosphorylated metabo

lites in perfused heart and perfused liver as measured in situ by

31 P

NMR and in vitro by conventional enzymatic assays of perchloric acid extractsa Liver

Heart Organ technique

NMR

Extract

NMR

Extract

ATP

3.23 ±O.OSb

3.S5±0.24

2.8±0.2

2.7±0.2

ADP

O.oJ ±0.01

0.57±0.09

0.3±0.1

1.1±0.1

1.04±0.19

1.86 ±0.1O'

2.0

3.8±0.2

PCr Pi

5.98±0.10

5.43±O.17

" All concentrations are given in micromoles per gram wet weight. b Results are expressed as mean ± standard error. Values of heart and liver were taken from references (80) and (66), respectively. 'Taken from reference (102) and converted into /1mol/g wet weight using the wet/dry weight ratio of 6.37 ± 0.19 recommended in (80).



51

in Table 1 . Second, Pi visible by 31p NMR reflects the same intracellular pH as deoxyglucose-6-phosphate, an exclusively cytosolic metabolite ( 1 6). Third, mitochondria occupy only 1 0-20% of the total cellular volume and contain less water than the cytosol. High viscosity, low organelle concentration, and also a probably higher content of paramagnetic metal ions can make mitochondrial metabolites invisible to NMR in cells or tissues ( 1 7). Likewise, ADP is essentially undetectable under in vivo conditions in the heart (80), kidney ( 1 78), muscle (43), or brain (4 1 , 146), and only barely detectable in the liver (63). The reasons are similar to those described for Pi. In particular, the cytosolic ATP/ADP ratio derived from subcellular fractionation studies is 1 0 in the perfused liver ( 1 72) and between 40 and 1 22 in the Langendorf perfused heart ( 1 7 1 ). Assuming a cytosolic ATP concentration of 3-5 mM in both cases, this means that the free ADP concentration is at the edge of 31p NMR visibility in the liver and is far below the level of detection in the heart. Furthermore, ADP becomes invisible to NMR when it is bound to macromolecules like actin-myosin in muscle or heart, or to the numerous phosphoryltransferases in the liver. Recently, the visibility of ATP by 31p NMR has been investigated in perfused rat liver ( 1 29). After 1 2 min of anoxia the level of ATP visible by NMR decreased to 30% of its initial value while the total ATP, as deter mined in extracts, decayed to only 84%. A preferential decrease of the cytosolic ATP during anoxia with no alteration of the mitochondrial ATP was reported, almost simultaneously, by Aw et al ( 1 4). Taken together, these results suggest that only the cytosolic ATP is visible by NMR. This point of view is not at variance with the fact that the ATP resonances have been observed in 31p NMR spectra of isolated intact mitochondria suspensions (92). These experiments used mitochondrial concentrations that were significantly higher than those thought to occur in the intact tissue and that were prepared in the presence of chelating agents. The initial approximation that essentially all ATP is NMR visible still holds true under normoxic conditions, because the major part of cellular ATP is indeed cytosolic ( 1 67). In Situ Control of Oxidative Phosphorylation

In order to produce work, skeletal and cardiac muscles hydrolize ATP generating ADP and Pi in the exergonic reaction: 3. The MgATp2- is immediately resynthesized from ADP and Pi in the mitochondria by oxidative phosphorylation, according to the equation:

52

CERDAN & SEELIG

3MgADP- +3Pi2- +�02+NADH+H+


f'±

3MgATp 2- + NAD+ + H20.

4.

The stoichiometry of 3Pi per !Oz is only an approximation ( 1 02). The two processes (Equations 3 and 4) are tightly coupled in skeletal muscle and heart with the MgATp 2- concentration remaining essentially constant. Therefore, measurements of the work performed or of the oxygen con sumed provide quantitative information on the ATP synthesized by oxi dative phosphorylation. If the oxygen supply to the mitochondria is limited, ATP can also be generated through the creatine phosphokinase equilibrium, at the expense of PCr and MgADP: 5. The role of this reaction in cellular energetics and the measurement of its forward and reverse unidirectional fluxes using magnetization transfer techniques is not addressed here. Further information on this topic can be found in References (12, 30, 96, 123, 149, 161, 162). In the past, isolated mitochondria, cell preparations, or perfused organs were used for studies of oxidative phosphorylation; however, the advent of 31p NMR techniques made this process amenable to in situ analysis (44-48, 96, 1 20, 1 49). 3 1 p NMR has focused on the relationship between mechanical work and metabolic energy expenditure in skeletal muscle (43 , 1 1 9, 161, 162) or heart (76, 96) i n order t o understand the factors controlling oxidative phosphorylation in situ. Chance et al (46) found that the relation ship between mechanical work and the ADP concentration in the in situ skeletal muscle (denoted as transfer function), followed Michaelis-Menten kinetics, with a Km of 25 pM for ADP. These results were very similar to those previously reported for suspensions of intact isolated mitochondria (49). Under hypoxic conditions the apparent Km for ADP increased to approximately 77 ,11M, a nonregulatory value, and oxygen supply then became the limiting factor. Chance et al (46) also found that the con centrations of ADP and Pi at rest, as determined by NMR in situ, were below the K"., of respiration, which suggests a possible regulatory role not only for ADP but also for Pi in the control of muscle respiration. Similar studies based on the use of transfer functions were pcrformed in open chest (107) and closed chest dog heart preparations ( 1 38, 139) and in the in situ brain ( 1 5 7). Taken together, these studies indicated that oxidative phosphorylation could be controlled not only by the ADP concentration but by several other factors, such as the Pi concentration, the delivery rate of the substrates and of oxygen, and the phosphorylation potential. These conclusions were also supported by studies on myocardial work pro duction and free ADP concentration in Langendorf perfused hearts under



53

different substrate conditions (76). Work production was dependent on the ADP concentration only when glucose was the substrate; it became independent of ADP with glucose plus insulin or with pyruvate as substrates, indicating again that factors other than the ADP concentration might control myocardial respiration. In summary, 31p NMR has contributed significantly to our current understanding of the control of oxidative phosphorylation in situ. The in situ concentrations of free ADP and Pi were found to be significantly lower than the total values measured by conventional extraction techniques. The in vivo phosphorylation potential (see Equation 2) was thus about an order of magnitude higher than previous estimates (63, 1 1 3, 1 85). This result is inconsistent with the hypothesis of thermodynamic control of oxidative phosphorylation [see (97) for a review]. In contrast, the in vivo 31 P NMR experiments point to kinetic control and multiple possibilities of substrate limitation in which ADP, Pi, the delivery rate of substrates and oxygen, and NADH transfer to the mitochondria could become the controlling factors, depending on the physiological circumstances.

The development of in vivo 13C NMR spectroscopy has been slower than its 31P counterpart because of the low natural abundance of 1 3C, the unfavorable gyromagnetic ratio of the 13C nucleus, and the need to use double-resonance techniques to remove the strong 13C_1H couplings that complicate the undercoupled 13C spectra. These limitations can be over come, in part, by the use of selectively enriched substrates and by special double-resonance decoupling devices, such as concentrical or saddle shaped 'H coils (4, 132), doubly tuned l 3C and IH coils ( 1 53), or IH imaging probes (6 1 ). The chemical shift range of 13C NMR is about 200 ppm, and 1 3C NMR is potentially more informative than 31p NMR, since almost all metabolites contain carbon but only a few contain phosphorus. Basically, the 1 3C NMR approach is similar to the classical 14C radio active isotope experiment. However, 13C NMR offers certain advantages over the conventional radioisotope techniques. First, the kinetics of 1 3C labeling in individual carbons from a variety of metabolites can be followed simultaneously and noninvasively. Secondly, even if the analysis is per formed in extracts, the 13C NMR approach avoids, in most cases, chromatographic separations and chemical degradations, which are essen tial in conventional radioactive isotope techniques. Finally, the presence and relative proportions of different 13C isotopomers of the same meta bolite can be evaluated by the analysis of I3C_I3C homonuclear spin coupling patterns ( 1 08, Ill ), a molecular property that can only be detected

54

CERDAN & SEELIG

by NMR methods. On the other hand, 13C NMR techniques are less sensitive than conventional radioisotope techniques, and only metabolites present in the millimolar concentration range can be conveniently detected


m VIVO.

13C NMR studies have been reported for a variety of metabolic pathways, including the citrate cycle and related anaplerotic reactions, the gluconeogenesis pathway and related futile cycles, the synthesis and degradation of glycogen, the f3-oxidation of short chain and medium chain free fatty acids and ketogenesis, and the metabolism of triglycerides and phospholipids. Tricarboxylic Acid Cycle and Related Anaplerotic Reactions

13C NMR studies of the tricarboxylic acid cycle have mainly been per formed with LangendorfIperfused hearts ( 1 5, 50, 1 1 4, 1 1 5, 1 59, 1 60). The investigations have focused on the substrate preference by the heart and on the quantitation of metabolic fluxes through the oxidative and ana plerotic routes of the citrate cycle using several, specifically 13C-enriched substrates. The time course of fractional 13C enrichment in individual carbons of glutamate in heart extracts was measured for Langendorff hearts perfused with either unlabeled glucose and [ 3- 13C]pyruvate or with [2-13C]acetate (50). A sophisticated mathematical model for the tricarboxylic acid cycle allowed the calculation of (a) the total flux through the cycle, (b) the relative contributions of labeled and unlabeled forms of acetyl-CoA, and (c) the exchange fluxes through the alanine or aspartate aminotransferases. In a later study, different isotopomer populations in glutamate C4 could be detected in vivo during perfusion of the hearts with 5 mM[3- 13C]lactate and 5 mM unlabeled glucose ( 1 60). This isotopomer analysis, originally pioneered for bacteria ( 1 08), proved extremely helpful in further eluci dating the metabolism of the heart. For example, in perfusions using 5 mM [3_13C]pyruvate or 5 mM [3- 13C]lactate, it was found that approxi mately 70% of the citric acid cycle flux was derived from the [2_ 13C]_ enriched acetyl-CoA pool. Notably, when unlabeled glucose plus insulin were added to the above mentioned 13C-enriched perfusate, only a minor decrease in the percentage of 13C-Iabeled acetyl-CoA entering the cycle was measured, confirming the preference of the heart for pyruvate over glucose as substrate. The same approach was later used to estimate the relative fluxes through the combined anaplerotic reactions versus the flux through the citrate synthase in the intact heart ( 1 1 5 159). The combined flux through the anaplerotic pathways was approximately 1 0% of the citrate synthase flux with acetate as the only substrate; it increased to 20 or 40% when pyruvate or propionate were added, thus indicating significant ,

N MR STUDIES OF METABOLISM

55

contributions of pyruvate carboxylase flux and propionate carboxylase flux in situ.


Gluconeogenesis

Gluconeogenesis and its interaction with the tricarboxylic acid cycle have been analyzed in isolated rat hepatocytes (56-58), in perfused mouse or rat liver (52, 57), and in intact animals ( 1 77) by 13C NMR and by a variety of '3C-enriched gluconeogenic precursors. Most studies have focused on the in situ distribution of flux between pyruvate carboxylase and pyruvate dehydrogcnase, which provide alternative routes into the citric acid cycle, or on the in situ activity of pyruvate kinase and the phosphoenolpyruvate cycle, which are potential modulators of the net gluconeogenic flux in situ. Initial experiments with perfused mouse or rat liver (52, 56) indicated that when alanine was the only substrate, it entered the tricarboxylic acid cycle, in approximately equal proportions, through the pyruvate dehydrogenase and pyruvate carboxylase routes. However, when the liver was perfused with a mixture of alanine and ethanol (only one of which was 13C labeled), alanine entered the citrate cycle exclusively through pyruvate carboxylase while ethanol was utilized mainly for labeling the acetyl-CoA pool. These 13C NMR results were further validated by 14C tracer experiments (58). These studies showed that 13C NMR could provide a convenient alternative to the radioactive isotope method, which is nor mally used to elucidate the mechanism of gluconeogenesis [see (98) for a review]. Gluconeogenesis was also investigated with [3-13C]alanine, [2'3C]ethanol, or [2-13C]pyruvate plus ammonia in perfused livers from streptozotozin diabetic rats. The results were compared with findings from similar experiments with livers obtained from fasted or fed animals (535 5). A novel effect of insulin-the stimulation of pyruvate kinase flux in situ-was described in these studies. In contrast, no effects of insulin on the relative channeling of substrates through pyruvate carboxylase or pyruvate dehydrogenase could be detected. Free Fatty Acid Metabolism and Ketogenesis

In situ metabolism of short chain and medium chain fatty acids in the liver of intact rats has been studied by 13C NMR techniques for different nutritional or hormonal situations, including feeding, fasting, and alloxan or streptozotozin-induced diabetes ( 39, 62, 1 40, 14 1 ). Emphasis was placed on the in situ mechanism and the hormonal regulation of ketogenesis and fJ-oxidation. Two pathways for acetoacetyl-CoA formation from butyrate were considered ( 1 40): (a) complete fJ-oxidation to two molecules of [ 1 '3C]acetyl-CoA and synthesis o f acetoacetyl-CoA through the hydroxy-


56

CERDAN & SEELIG

metylglutaryl-CoA pathway or (b) direct production of acetoacetyl-CoA, followed by condensation with the mitochondrial pool of acetyl-CoA. These two mechanisms can be distinguished from one another by the relative 13C labeling in carbons Cl and C3 of fJ-hydroxybutyrate. Pathway (a) predicts equal 13C labeling in C l and C3 and pathway (b) allows asymmetric labeling because C 3 and C l are derived from different pre cursors. Although the first mechanism is usually thought to be the major pathway of acetoacetate formation, the large asymmetry found in vivo between the labeling of the C l and C 3 carbons of fJ-hydroxybutyrate provided strong evidence for a major contribution of the direct pathway (b) in situ. The mechanism and metabolic compartmentation of the fJ-oxidation pathway of I-w medium chain dicarboxylic acids in rat liver was analyzed by a combination of in vivo and in vitro 13C NMR techniques (39). It had been previously proposed that the fJ-oxidation of dicarboxylic acids in the liver followed a bidirectional fJ-oxidation mechanism. However, 13C-NMR showed that [1,2-13C2] and [ 1 ,2, 1 1 , 1 2-13C4] dodecandioic acids were degraded in vivo monodirectionally. With respect to the relative con tributions of the mitochondrial or peroxisonal fJ-oxidation systems to the overall oxidation of l-w medium chain dicarboxylic acids, no 13C label was found in metabolites of the tricarboxylic acid cycle or in the ketone bodies. Since the latter two reaction pathways are localized in the mito chondria, these results suggest that the fJ-oxidation of medium chain dicar boxylic acids proceeds primarily in the hepatic peroxisomes. 13C NMR thus provided the first in situ evidence for the activity of the peroxisomal fJ-oxidation system. Glycogen Synthesis and Degradation

13C resonances of the IX and fJ anomers of glucose (at 98.0 and 96.0 ppm, respectively) and the IX-l-4 glycosidic linkage of glycogen (Cl glycogen at 100.5 ppm) (6) made it possible to study the hormonal regulation of glycogen synthesis and degradation in situ. In spite of its high molecular weight, glycogen was shown to be completely visible by NMR and to be observable even in naturally abundant 13C spectra (169). Glycogen syn thesis and degradation have been studied in perfused or in situ liver (52 154, 158, 166, 169) and in heart (lOS, 133, 134). More recently, 13C signals of skeletal muscle glycogen have also been detected in humans (13). Initial studies analyzed the time course of[I-13C]glucose incorporation into glycogen and of glucagon-induced glycogenolysis in the perfused rat liver (169). Glycogen synthesis in the in situ liver of fed rats was found to reach steady state after 60 min of a bolus injection of[l-13C]glucose, and 50% glycogen depletion was observed after approximately 60 min of ,



57

glucagon administration ( 1 54). Similarly, glycogen synthesis from [ 1 13C]glucose was inhibited b y glucagon, whereas insulin alone or in com bination with glucagon had no stimulatory or inhibitory effect on glycogen synthesis ( 1 66). Several 13C NMR studies have recently addressed the mechanism of hepatic glycogen repletion after a glucose load in fasted rats. Previous experiments with radioactive tracers had shown that glycogen synthesis required the activity of the gluconeogenic pathway and that the indirect route glucose � trioses � glycogen could make a significant contribution to glycogen repletion relative to the conventionally accepted direct path way glucose � glucose-lP � UDP-glucose � glycogen (99, 1 35, 1 36). The contributions of the direct and indirect pathways of glycogen repletion were compared by oral administration of [ 1 - 13C]glucose or [ 1-13C]glucose plus unlabeled alanine ( 1 58, 1 6 3, 1 64). The direct and the indirect pathways accounted for 34% and 30-55%, respectively, of the glycogen production. However, the contribution of the indirect pathway, as determined in the NMR studies, was significantly smaller than previous measurements per formed with radioactive isotopes (\ 35). This discrepancy can probably be explained by the different size of the glucose loads (l04). In vivo rates of myocardial glycogen synthesis have been compared with the in vitro activity of glycogen synthase and phosphorylase, measured under conditions that preserved the original phosphorylation state and mimicked the intracellular level s of the corresponding substrates and effec tors ( 1 05). The in situ rate of glycogen synthesis closely paralleled the activity of glycogen synthase measured in vitro. These results suggest that glycogen synthase is the rate-limiting step in glycogen synthesis in vivo. In contrast, even though the activity of phosphorylase b measured in vitro was 6-7 times higher than that of glycogen synthase, no significant glyco genolysis was detected in vivo. Under in vivo conditions, phosphorylase b appears to be inhibited by an unknown factor that is not present in the in vitro assays. This result sheds doubts on the canonical view that phos phorylase b control is based on a covalent modification induced by the phosphorylation cascade ( 1 65). Triglyceride and Phospholipid Metabolism

13C NMR has been used to study the triglyceride and phospholipid com position of normal and diseased muscles (20, 2 1 , 67) of adipose tissue and isolated adipocytes ( 1 68) and of brain (19, 22). The fatty acid composition of adipose tissue in rats was shown to be modified under different dietary conditions (38). In particular, the degree of unsaturation of the fatty acid chains could be derived from the ratio of the 130.0/128.5 ppm peaks, which reflects the ratio of monounsaturated to polyunsaturated fatty acids in a

58

CERDAN & SEELIG


specific tissue. This ratio also reflects the fatty acid composition of the diet and was significantly altered in dystrophic muscles. In brain, halothane increased phospholipid mobility (as evidenced by an increase in the methylene signal) (19), and a similar effect was described for caffeine stimulated muscle (20). More recently, sufficiently mobile NMR visible phospholipid pools were characterized in brain and muscle (22). IHNMR

IH NMR has an inherently higher sensitivity than 3'p_ and 13C-NMR, and a larger number of cellular metabolites should be detectable. However, two limitations need to be considered: (a) The water signal of the tissue has to be eliminated, and (b) the complex spectral patterns that are due to overlapping proton resonances have to be resolved. Both drawbacks have been overcome, in part, by the development of water suppression tech niques (89, 90, 122, 182) and different methods of one-dimensional or two dimensional spectral editing (cf 12). Cerebral metabolism has been studied by 'H NMR under a variety of conditions (25) including hypoxia or anoxia (24, 26), hypoglycemia (24), epileptic seizures ( 1 57), genetic deficiencies (78), and traumatic shock ( 1 86). The time course of cerebral lactate production by anaerobic glyco lysis, following hypoxic insults, was monitored qualitatively by comparing the intensity of the lactate methyl resonance with that of the methyl group of N-acetylaspartic acid (NAA), assuming a complete 'H NMR visibility for both compounds in vivo (155). A quantitative interpretation of these results is difficult; recently, it has been shown that cerebral lactatc might not be completely visible by NMR (51) and that factors other than lactate concentration can also affect the intensity of the lactate methyl signal (186). In addition, different relaxation times for the methyl group of NAA have been demonstrated for white and grey matter (88), and the intensity of the in vivo NAA methyl signal may be dependent on the proportions of white and grey matter detected by the coil. A 'H NMR spectra of the human brain, as obtained in situ by the STEAM technique at 1.5 T, is shown in Figure 3 for illustrative purposes and is compared with the high resolution 'H NMR spectrum of a per chloric acid extract obtained from rat brain (at 8.4 T). Figure 3 demon strates that in spite of the large differences in field strength a significant number of resonances derived from N-acetylaspartate, creatine, and phos phocreatine, phosphorylcholine, and probably glycine can be resolved in the human brain spectrum. The high resolution spectrum shows the pres ence of up to 14 different cerebral metabolites (40) (cf 148). Other 'H NMR studies have focused on skcletal muscle metabolism. In

59


Cr/PCr

A

N-Ac-Asp CH3


Cr J PCr

Gly?

. PC/GPC

B Cr

PCr CH r

HaHa'

N-Ac -Asp

C Hr

Gly (Hal PCr

Ha Ha'

i

,

Glu Ha

Gin

Ha

N-Ac-Asp

Ha a- glucose I I

i

'

Asp Ha

I

, I' 4,0 Figure 3

l

I

l

II

GPC Tau HaHa' Tau

I

Lac

I

[H3-

! Thr CHr

I

H�H{r PC

i I

BHBA

I

Vol CHr

I!

Leu CH

II

, I' 3.0

' " PPM

'H NMR spectroscopy of the brain.

A:

, I ' 2.0

I

1.0

r

'

Localized 'H NMR spectra of human

brain in situ obtained using the STEAM technique at 1.5 T (acquisition time: 13 min, volume selected:

64

cm3) (courtesy of H. P. Hafner). B: High resolution 'H NMR spectra of a

perchloric acid extract obtained from rat brain at 8.7 T (cf 40).

CERDAN & SEELIG


60

particular, it was possible to monitor simultaneously phosphocreatine hydrolysis and lactate production in preparations of superfused frog sar torius and gastrocnemius muscles. Intracellular pH could also be deter mined from the chemical shifts of the imidazolic protons of carnosine (8). This method, initially proposed in (191),has been extended to intracellular pH determinations in human skeletal muscle (142). Two-dimensional (2D) NMR editing methods have also been used in intact cells (60, 118), tissue extracts (9,39, 71), body fluids (18, 94, 147), excised tissues (7), or even in vivo (29, 173). These methods allow a straightforward assignment of the resonances and a simultaneous editing of all metabolites (at the expense of a longer acquisition time). We illustrate this approach with the infusion of [1,2-13C Jdode 2 candicarboxylic acid into the liver of an anesthesized rat (39). The 1 3C resonances were first monitored in vivo, but at the end of the infusion period the animal was sacrificed and a perchloric acid extract of the liver was obtained. Figure 4 shows the COSY spectrum of this extract. By

1.0

2.0 I

3.0

.� t7J.�q. E .

."

QI ,Q L M

•

•

iA,,,

F�

G!

4.0

H

CilR .. .

I

"

5.0 Figure

4

til, I

Ii

i i " Iii i

4.0

II

I i I I f ii,

3.0

Ii

PPM

I iii I I Iii i i i ,

2.0

I I I Iii,

5.0 PPM

1.0

Contour plot of a 2D COSY 45 spectra obtained from a perchloric acid extract

of the liver of an alloxan diabetic rat infused with [l,2-13Czldodecandicarboxylic acid. Relevant assignments are indicated in the text. From (39).

NMR STUD IES OF METABOLISM

61

comparison with control spectra (not shown), several important differences can be detected, notably the appearance of cross peaks A, B, C, and D and an increase in the correlations I and M. These correlations were shown to be primarily derived from adipic acid (correlations A and D) and the amino acid L-Iysine (correlations B, C, I, and M) as indicated below:


D

A

D

��I� -OOC-CH2-CH2-CH2-CH2-COOI

B

C

M

nnnn

+NH3-CHz-CHz-CHz-CHz-CH-COO-

I

NHt Therefore, infusion of [1,2-13Cz]dodecandicarboxylic acid resulted in an increase of the hepatic concentrations of lysine and adipic acids. An inhibitory effect of adipic acid, the main product of the fJ-oxidation of dodecandioic acid, on the pathway of lysine degradation (at the level of a-aminoadipic acid derivatives) could be proposed from this experiment. Other applications of IH NMR have been recently reviewed (145). 19F NMR

The 19F nucleus has a slightly smaller magnetic moment than the proton, whereas its chemical shift range is larger by a factor of 20. No con tributions from tissue background signals are observed, which makes 19F an interesting magnetic label for metabolic studies. Using fluorinated indicators, 19F NMR has been extensively applied to measure the intracellular pH as well as the free Ca 2+ and free Mg2+ concentrations ( 123). Other applications of 19F NMR to the study of in situ metabolism involve (a) fluorinated anesthetics and anticancer drugs and (b) fluorinated glucose derivatives. Halothane, metoxyflurane, and isoflurane give well-resolved 19F NMR spectra in rabbit brain ( 190), and the time course of halothane elimination from rabbit brain can be followed. Further studies showed bicxponential decay curves in the cerebral halothane wash-out, which is likely associated with two different chemical environments ( 189). The metabolism of 5fluorouracyl was studied in perfused liver (37), intact animals (175), or '"


62

CERDAN & SEELIG

humans (1 88). Major metabolites in rodent liver in situ or in perfused mouse liver were ct-fluoro-fJ-alanine (FBAL) and ct-fluoro-fJ-ureidopro pionic acid (FUPA). In contrast, the only metabolite detected in man was FBAL. In no case could dihidrofluorouracyl (FUH2) be observed, even though it is a common precursor of FBAL and FUPA. The relative contributions of the aldose reductase pathway and of the pentose monophosphate shunt to the cerebral metabolism of intravenously infused 2-fluoro-2-deoxy-glucose (2FDG) were studied in rat brain ( 1 30). The in vivo 1 9F NMR spectrum showed signals of the infusate and of 2fluoro-2-deoxyglycerol (FDGL) and of 2-fluoro-2-deoxy-6-phospho gluconate (FD-6-PG), two key metabolites in the aldose reductase pathway and the pentose monophosphate shunt, respectively. Sorbinil, an inhibitor of the aldose reductase pathway, essentially eliminated 2FDG metabolism, thus indicating that this metabolic route is the main pathway of cerebral catabolism of 2- FDG. The sensitivity of 1 9F is sufficiently high to allow 1 9F MR Imaging ( 1 3 1). 1 9p images of 2-fluoro-2-deoxy-glucose-6-phosphate (2-FDG-GP), a key metabolite of the glycolytic pathway, revealed high glucose utilization in brain, spinal cord, and heart. Likewise, 1 9F images of 2-fluoro-3-deoxy sorbitol (3-FDSL), a metabolite of the aldose reductase pathway, revealed a spatial heterogeneity in the distribution of the aldose reductase; the activity of this enzyme was highest in brain and lens.

Deuterium NMR is a recent addition to the repertoire of in vivo NMR techniques. The magnetic moment of the deuterium nucleus is smaller than that of the proton by a factor of 6. 5 which reduces the sensitivity of the measurement considerably. Furthermore, the natural abundance of deuterium is low (0.02%), and 2 H N M R in vivo studies appear unrealistic at first sight. On the other hand, deuterium is a spin 1 = 1 nucleus and due to its quadrupole moment has a very short relaxation time. In biological tissues the TI relaxation time is typically of the order of 1 00 ms, which allows a very rapid repetition of the NMR measurement. In fact, water and fat resonances can be observed by in vivo 2H NM R even at the natural abundance level of deuterium. The distinct advantage of the 2H nucleus, however, is its potential as a magnetic label similar to 13C and 1 9p ( 1 1 6). In early metabolic applications 2H NMR spectra of the abdomen of a niouse were obtained at 4.7 Tesla with acquisition times of less than 2 min (33). The resonances could be assigned to HDO and to the methylene CHD-resonances of fat. The intensity of these resonances increased markedly upon addition of 1 0% D 20 to the drinking water. The turnover of water



63

and fat could be determined from the decay of the 2H NMR signal intensity after replacing the deuterated water with normal drinking water. [ 1 - 2H] or [6- 2H ]glucose have been employed to study the balance between the 2 glycolytic pathway and the polyol pathway in the eye lens (3). 2H NMR mctabolic studies have also been reported on the direct and indirect gly cogen synthesis pathways in rat liver (82) and on the in situ metabolism of L-(methyl- 2 H 3)methionine and D-(methyl- 2H 3) methionine (109, 110). D 20 has been used as a freely diffusable tracer to determine the in situ blood flow in the intact rat liver (1) and in implanted nude mouse tumors ( 1 0 1 ). 2H NMR images could be obtained from rats that for several weeks received drinking water doped with D 0 or from rats that had obtained 2 an intraperitoneal injection of D zO ( 1 28). 2H images have also been reportcd of cat brain (70). These studies suggest that D 20 at low doses may also become a useful contrast agent in medical research on humans. ACKNOWLEDGMENTS

This work was supported by Grant 4.889.8 5. 1 8 from the Swiss National Science Foundation (to 1.S.) and Grant 88/1 875 from the Spanish Fondo de Investigaciones Sanitarias de la Seguridad Social (to S.C.).

Literature Cited

I. Ackerman, J. J. H., Ewy, C S., Becker, N. N., Shalwitz, R. A. 1987. Proc. Natl. A cad. Sci. USA 84: 4099-4 102 2 . Ackerman, 1 . 1 . H., Grove, T. H . , Wong, G . C., Gadian, D. G . , Radda, G. K. 1 980. Nature 283: 1 67-70 3. Aguayo, J. B . , McLennan, I. J . , Aguiar, E., Ch eng , H. M. 1 987. Biochem. Bio phys. Res. Commun. 1 42: 359-66 4. Alger, J. R., Behar, K. L., Rothman, D. L., Shulman, R. G. 1984. J. Magn. Reson. 56: 3 34-37 5. Alger, J. R., Shulman, R. G. 1 984. Q. Rev. Biophys. 1 7 : 83-124 6. Alger, J. R . , Sillerud, L . 0., Behar, K. L . , Gillies, R. J., Shulman, R. G., et al. 1 98 1 . Science 2 1 4: 660-62 7. Alonso, 1., Arus, C, Westier, W. M . , Markley, J . L . 1 989. Magn. Reson. Med. 1 1 : 3 1 6--3 0 8 . Arus, C, Barany, M. 1 986. Biochim. Biophys. Acta 886: 4 1 1-24 9. Arus, C , Westler, W. M . , Barany, M . , Markley, J. D. 1 986. Biochemistry 2 5 : 3346--5 1 1 0 . Aue, W. P. 1 986. Rev. Magn. Reson. Med. I : 2 1 -72 I I . Aue, W. P., Muller, S., Cross, T. A.,

12.

13.

14. IS.

16.

17. 18.

19. 20.

21.

Seelig, J. 1 984. J. Magn. Reson. 56: 3 50-54 Avison, M . J . , Hetherington, H. P., Shulman, R. G. 1 986. Annu. Rev. Bio phys. Biophys. Chem. 1 5: 377�02 Avison, M. J., Rothman, D. L., Nadel, E., Shulman, R. G. 1 988. Proc. Nat!. A cad. Sci. USA 85: 1 634-36 Aw, T. Y., Andersson, B. S., Jones, D. 1 987. Am. J. Physiol. 252: C356- C361 Bailey, I. A., Gadian, D. G., Matthews, P. M . , Radda, G. K., Seeley, P. J. 1 98 1 . FEBS Lett. 1 23: 3 1 5- 1 8 Bailey, I. A., Williams, S . R . , Radda, G. K., Gadian, D. G. 198 1 . Biochem. J. 196: 1 7 1-78 Balaban, R. S. 1984. Am. J. Physiol. 246: C IO-C I9 Bales, J. R., Bell, J. D., Nicholson, J. K., Sadler, P. J., Timbrell, J . A., et al. 1 988. Magn. Reson. Med. 6: 300-6 Barany, M . , Chang, Y. C, Arus, C 1 985. Biochemistry 24: 79 1 1- 1 7 Barany, M . , Doyle, D. D., Graff, G . , WestIer, W. M . , Markley, J. L. 1 982. J. Bioi. Chem. 257: 274 1-43 Barany, M . , Doyle, D. D., Graff, G.,

64 22.

23. 24.


25.

26.

27.

28.

29.

CERDAN & SEELIG Westler, W. M . , Markley, J. L. 1984. Magn. Reson. Med. I: 30--43 Barany, M., Venkatasubramanian, P. N . 1987. Biochim. Biophys. A cta 923: 339-46 Beckman, N . , M uller, S., Seelig, J. 1 989. Magn. Reson. Med. 9: 391-94 Behar, K. L . , den Hollander, J. A., Petroff, O. A. c., Hetherington, H . P., Prichard, J. W . , Shulman, R. G. 1 985. J. Neurochem. 44: 1 045-55 Behar, K. L., den Hollander, J. A., Stromsky, M . E., Ogino, T., Shulman, R. G., et aL 1 983. Proc. Nat!. A cad. Sci. USA 80: 4945-48 Behar, K. L . , Rothman, D. L., Shulman, R. G . , Petroff, O. A. C., Prichard, J. W . 1 984. Proc. Natl. Acad. Sci. USA 8 1 : 2 5 1 7- 1 9 Belton, P . S . , Ratcliffe, R. G. 1 985. Prog. Nuc!. Magn. Reson. Spectrosc. 1 7 : 241-49 Berden, J. A . , Cullis, P. R., Hoult, D. I . , McLaughlin, A. c., Radda, G. K . , Richards, R. E. 1 974. FEBS Lell. 46: 55-58 Berkowitz, B. A . , Wolf, S. D., Balaban, R. S. 1 988. J. Magn. Reson. 79: 547-

53

30. Bessman, S . P., Carpenter, C. L . 1 985. Annu. Rev. Biochem. 54 : 83 1-62 3 \ . Bottomley, P. A. 1 989. Radiology 1 70: 1-15 3 2 . Bottomley, P. A . , Charles, H . c., Roemer, P. B . , Flamig, D . , Engeseth, H . , et aL 1 988. Magn. Reson. Med. 7: . 3 1 9-36 33. Brereton, 1. M . , Irving, M. G . , Field, J., Doddrell, D. M. 1 986. Biochem. Bio phys. Res. Commun. 1 37: 579-84 34. Brindle, K. M . , Campbell, !. D. 1 987. Q. Rev. Biophys. 19: 1 59-82 35. Burk, S. L., Shulman, R. G. 1 987. Annu. Rev. Microbiol. 4 1 : 595--6 1 6 36. Burt, C. T . 1 985. Trends Biochem. Sci. 1 0: 404-6 37. Cabanac, S . , Malet-Martino, M . c., Bon, M . , Martino, R., Nedelec, J. F., Dimicoli, J. F. 1 988. NMR Biomed. I: 1 1 3-20 38. Canioni, P. c., Alger, J. R., Shulman, R. G . 1 983. Biochemistry 22: 4974-80 39. Cerdan, S., Kunnecke, B . , Dolle, A., Seelig, J. 1 98 8 . J. Bioi. Chem. 263: 1 1 664-74 40. Cerdan, S., Parrilla, R ., Santoro, J . , Rico, M . 1985. FEBS Lett. 1 87: 1 67-72 4 1 . Cerdan, S., Subramanian. V. H .• Hil berman, M., Cone, J., Egan, J., et al. 1 986. Magn. Reson. Med. 3: 432-39 42. Cha1iss, J. R. A., Blackledge, M. J., Radda, G. K . 1 988. Am. J. Physiol. 254: C41 7-C422

43. Chaliss, J. R. A., Hayes, D. J., Radda, G. K. 1 987. Biochem. J. 246: 1 63-72 44. Chance, B . , Eleff, S., Bank, W., Leigh, J. S. Jr., Warnell, R. 1982. Proc. Natl. A cad. Sci. USA 79: 7 7 14- 1 8 45. Chance, B . , Eleff, S . , Leigh, J. S. Jf. 1 980. Proc. Natl. A cad. Sc i. USA 77: 7430-34 46. Chance, B., Eleff, S . , Leigh, J. S . Jr., Sokolow, D., Sapega, A. 1 98 1 . Proc. Natl. A cad. Sci. USA 78: 67 1 418 47. Chance, B., Leigh, J . S., Clark, B. J . , Maris, J., Kent, J . , et al. 1 985. Proc. Natl. A cad. Sci. USA 82: 8384-88 48. Chance, B . , Leigh, J. S. Jr., Kent, J., McCulli, M . K. , Nioka, S . , et al. 1 986. Proc. Natl. A cad. Sci. USA 83: 945862 49. Chance, B . , Williams, G. R. 1 956. J. Bioi. Chem. 2 1 7: 409-27 50. Chance, E. M . , Seeholtzer, S. H . , Kobayashi, K . , Williamson, J. R. 1 983. J. Bioi. Chem. 258: 1 3785-94 5 1 . Chang, L. H . , Pereira, B. M . , Wein stein, P. R . , Konig, M. A . , Murphy Boesch, J . , ct al. 1 987. Magn. Reson. Med. 4: 575-8 1 52. Cohen, S. M. 1983. J. Bioi. Chem. 258: 14294-14308 53. Cohen, S. M. 1987. B iochem istry 26: 563-72 54. Cohen, S. M. 1 987. Biochemistry 26: 573-80 55. Cohen, S. M. 1 987. Biochemistry 26: 58 1-89 56. Cohen, S . M., Glynn, P., Shulman, R. G. 1 98 1 . Proc. Natl. A cad. Sci. USA 78: 60--64 57. Cohen, S. M . , Ogawa, S., Shulman, R . G. 1 979. Proc. Natl. A cad. Sci. USA 76: 1 603-7 58. Cohen, S. M . , Rongstad, R., Shulman, R. G., Katz, J. 1 98 1 . J. Bioi. Chern. 256: 3428-32 59. Cox, K. J . , Styles, P. 1983. J. Magn. Reson. 55: 1 64-69 60. Cross, K. J. , Holmes, K. T., Mount ford, C. E., Wright, P. E. 1984. B io chemistry 23: 5895-97 6 1 . Cross, T. A., M uller, S., Aue, W. P. 1 985. J. Magn. Reson. 62: 87-98 62. Cross, T. A., Pahl, c . , Oberhansli, R . , Aue, W . P. , Keller, U. , Seelig, J. 1 984. Biochem istry 23: 6398-6402 63. Cunningham, C. c., Malloy, C. R . , Radda, G. K. 1 986. Biochim. Biophys. A cta 885: 1 2-22 64. Daly, P. F., Cohen, J. S. 1 989. Cancer Res. 49: 770-79 65. Damadian, R. 1 97 1 . Science 1 1 7: 1 1 5 153 66. Desmoulin, F., Cozzone, P. J., Cani-


67.

68.

69.


70.

71.

72. 73.

74. 75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

oni, P. 1 987. Eur. J. Biochem. 1 62: 1 5 159 Doyle, D. D . , Chalovich, J. M . , Barany, M . 1 98 1 . FEBS Lett. 1 3 1 : 1 4750 Eakin, R. T., Morgan, L. 0., Gregg, C. T., M atwiyoff, N . A. 1 972. FEBS Lett. 28: 259-64 Eveloch, J. L . , Crowley, M . G . , Acker man, J. J. H. 1 984. 1. Magn. Reson. 56: 1 1 0-24 Ewy, C. S , Ackerman, J. J. H . , Bala ban, R. S . 1 988. Magn. Reson. Med. 8: 35--44 Fan, T. W.-M., Higashi, R. M . , Lane, A. N . , Jardetzky, O. 1 986. Biochim. Biophys. Acta 882: 1 54-67 Fisher, M. J. , Dillon, P. F. 1 988. NMR Biomed. 1 : 1 2 1-26 Frahm, J., Bruhn, H., Gyngelb, M. L., Merboldt, K . D . , Hanicke, W., Sauter, R. 1 989. Magn. Reson. Med. 9: 79-93 Frahm, J. , Merboldt, K. D . , Hanicke, W . \ 987. J. Magn. Reson. 72: 502-8 Frahm, J . , Merboldt, K. D., Hanicke, W., Haase, A. 1 98 5 . J. Magn. Reson. 64: 8 1 -93 From, A. H. L. , Petein, M . A., Michur ski, S. P., Zimmer, S. D . , Ugurbil, K . 1 986. FEBS Lett. 206: 257-61 Gadian, D. G. 1 982. In NMR and its Applications to Living Systems, pp. 11 37 . Oxford: Clarendon Press Gadian, D. G., Proctor, D. L., Williams, S. R., Cady, E. B., Gardiner, M. R. 1 986. Magn. Reson. Med. 3: 1 50 56 Galloway, G. J., Brereton, 1. M . , Brooks, W . M . , Field, J . , Irving, M . , et a l . 1 987. Magn. Reson. Med. 4: 39398 Gard, J. K., Kichura, G . M., Acker man, J. J. H . , Eisenberg, J. D., Billa delio, J. J., et al. 1 985 . Biophys. J. 48: 803 1 3 Gonzalez-Mendez, R . , Hahn, G . M . , Wade-Jardetzky, N . G., Jardctzky, O . 1 988. Magn. Reson. Med. 6: 3 73-80 Goodman, M. N . , Matsuoka, L. K . , de Ropp, J. S., Jones, A. D. 1 989. Bio chem. Biophys. Res. Commun. 1 59: 52227 Graham, R. A. , Meyer, R. A., Szwer gold, B. S., Brown, T. R. 1 987. J. Bio/. Chem. 262: 35 37 Guidoni, L . , Mariutti, G., Rampelli, G . M . , Rosi, A., Viti, V. 1 987. Magn. Reson . Med. 5: 578-85 Gupta, R. K., Gupta, P., Moore, R. D . 1 984. Annu. Rev. Biophys. Bioeng. 1 3: 221--46 Gyulai, L., Bolinger, L., Leigh, J . S., Barlow, c., Chance, B . 1 984. FEBS Lett. 1 78: 1 3 7-42

65

87. Hafner, H . P. , Muller, S . , Seelig, J. 1 989. Magn. Reson. Med. I n press 88. Hanstock, C. c., Rothman, D. L., Prichard, J . W. , Jue, T., Shulman, R. G. 1 988. Proc. Natl. A cad. Sci. USA 85: 1 82 1-25 89. Hore, P. J. 1 983. J. Magn. Reson. 54: 539--42 90. Hare, P. J. 1 983. J. Magn. Reson. 55: 283-300 9 1 . Hoult, D. I . , Bushy, S. 1. W., Gadian, D. G., Radda, G. K., Richards, R. E . , Seeley, P. J. 1 974. Nature 252: 285-87 92. Hutson, S. M . , Berkik, D., Williams, G . D., La Noue, K . F., Briggs, R. W. 1 989. Biochemistry 28: 4325-32 93. Iles, R. A., Griffiths, J. R., Stevens, A. N., Gadian, D. G., Porteus, R. 1 980. Biochem. J. 1 92: 1 9 1-202 94. Iles, R. A., Jago, J. R . , Williams, S. R . , Chalmers, R. A. 1 986. FEBS Lett. 203: 49-53 95. Iles, R. A., Stevens, A. N., Griffiths, J . R ., Morris, P. G. 1 985. Biochem. J . 229: 1 4 1 -5 1 96. Ingwall, J . S . 1 982. Am. J. Physio/. 242: H729-H744 97. Jacobus, W. J. 1 98 5 . Annu. Rev. Phy sio/. 47: 707-25 98. Katz, J. 1 985. Am. J. Physio/. 248: R39 1 -R399 99. Katz, J., Kuwajima, M., Foster, D. W. , McGarry, J. D . 1 988. Trends Biochem. Sci. 1 1 : 1 36--40 100. Kauppinen, R. A., Hiltunen, J. K . , Hassinen, I . E. 1 980. FEBS Lett. 1 1 2: 273 76 1 0 1 . Kim, S. G . , Ackerman, 1. 1. H . 1 988. Cancer Res. 48: 3449-53 1 02. Kingsley-Hickman, P. B. , Sako, E. Y . , Mohanakrishnan, P., Robitaille, P. M . L., From, A. H . L . , et al. 1 987. Bio chemistry 26: 750 1 - 1 0 1 0 3 . Kushmerick, M . J. , Dillon, P. F . , Meyer, R . A . , Brown, T. R., Krisanda, J. M . , Sweeney, H. L. 1 986. J. Bio/. Chem. 26 1 : 1 4420-29 1 04. Landau, B. R., Wahren, J. 1 988. FASEB J. 2: 2368-75 1 05 . Laughlin, M. R., Petit, W. A., Dizon, J. M., Shulman, R. G. 1 988. J. Bio/. Chem. 263: 2285-91 1 06. Lauterbur, P. C. 1 973. Nature 242: 1 9091 1 07. Ligeti, L . , Osbakken, M . D. , Clark, n. J . , Schnall, M . , Bolinger, L., et a!' 1 987. Magn. Resoll. Med. 4: 1 1 2-1 9 1 08 . London, R . E . 1 988. Prog . Nucl. Magll. Reson. Spectrose. 20: 337-83 1 09. London, R . E., Gabel, S . 1 988. Bio chemistry 27: 7864-69 1 1 0. London, R. E . , Gabel, S . , Funk, A . 1 987. Biochemistry 2 7 : 7864-69


66

CERDAN & SEELIG

I l l . London, R. E., Kollman, V. H . , M atwiyoff, N. A. 1 975. 1. A m . Chern. Soc. 97: 3565-73 1 1 2 . Luyten, P. R. , Marien, A . J. Fl . , Sijtsma, B. , den Hollander, J . A. 1 986. J. Magn. Reson. 67: 1 48-55 1 1 3. Malloy, C. R., Cunningham, C. c., Radda, G. K. 1 986. Biochim. Biophys. Acta 885: 1-1 1 1 1 4. Malloy, C. R., Sherry, A. D . , Jeffrey, F. M . H. 1 98 7 . FEBS Lett. 2 1 2 : 58-62 1 1 5. Malloy, C. R., Sherry, A. D . , Jeffrey, F. M . H . 1 988. J. Bioi: Chern. 263: 6364-7 1 1 1 6. Mantsch, H. H . , Saito, H . , Smith, I . C. P. 1 977. Prog. Nuc!. Magn. Reson. Spectrosc. 1 I : 2 1 1-7 1 1 1 7. Matthews, P. M . , Shen , L. F., FoxalI, D., Mansour, T. E . 1 985. Bioehim. Bio phys. Acta 845: 1 78-88 1 1 8 . May, G. L., Wri ght, L. c., Holmes, K . T . , Williams , P. G., Smith, I . C. P., et al. 1 986. J. BioI. Chem. 26 1 : 3048-53 1 19 . Meyer, R. A . , Brown, T. R., Krilowicz, B. L., Kushmerick, M. J. 1986. Am. J. Physiol. 246: C264-C274 1 20. Meyer, R. A . , Kushmerick, M . .T., Brown, T. R. 1982. Am. J. Physiol. 242:

C I -C l l

1 2 1 . Moon, R . S . , Richards, J . H . 1 973. J. BioI. Chern. 248: 7276--7 8 1 22 . M orris, G . A. , Freeman, R. 1978. J. Magn. Reson. 29: 433-62 1 23 . Morris, P. G . 1 988. Ann. Rep. Nucl. Magn. Reson. Spec/rose. 20: 1 -60 1 24 . M uller, S. 1988. Magn. Reson. Med. 6: 364-7 1 1 25 . M uller, S., Aue, W. P., Seelig, J. 1 985. J. Magn. Reson. 63: 530-43 1 26. M uller, S., Aue, W. P., Seelig, J. 1 985. .J. Magn. Reson. 65: 332-38 1 27 . Muller, S., Sauter, R., Weber, H . , Seelig, J . 1 988. J. Magn. Reson. 76: 1 5 5-61 1 28. Muller, S., Seelig, J . 1 987. J. Magn. Reson. 72: 456--66 1 29. M urph y , E . , Gabel, S. A., Funk, A., London, R. E . 1 98 8 . Biochem istry 27: 526-28 1 30. Nakada, T., Kwee, I. L. 1 987. Magn. Resun. Med. 4: 366-7 1 1 3 1 . Nakada, T., Kwee, I. L., Card, P. J., Matwiyoff, N . A., Griffey, B . V., Griffey, R. H . 1 988. Magn. Reson. Med. 6: 307-13 1 32. Neurohr, K. J. 1 984. 1. Magn. Reson. 59: 5 1 1 - 1 4 1 33 . Neurohr, K . J., Barrett, E. J., Shulman, R. G. 1983. Proc. Nall. Acad. Sci. USA 80: 1 603-7 1 34. Neurohr, K. J., GolIin, G., Neurohr, J . M . , Rothman, D . L . , Shulman, R. G. 1 984. Biochemistry 23: 5029-35

1 3 5 . Newgard, C. B . , Hirsch, L. J . , Fo ster, D . W., McGarry, J. D . 1 983. J. BioI. Chel11. 258: 8046-52 1 36 . Newgard, C. B. , Moore, S. V . , Foster, D. W . , McGarry, J. D . 1984. J. BioI. Chem. 259: 6958-63 1 37. Ordi dge , R. J., Connelly, A., Lohman, J. A. B. 1 986. 1. Magn. Reson. 66: 28394 1 38. Osbakken, M . 1 988. In NMR Tech n iques in the Study of Cardiovascular Structure and Func tion , ed. M . Osbak ken, J. H aselgrove, pp. 207-37. Mount Kisco, NY: Futura 1 39. Os bakken, M. D., Ligeti, L., Clark, B . J . , Bolinger, L . , Subramanian, H . , e t al. 1 986. Magn. Reson. Med. 3: 801-7 140. Pahl-Wostl, c., Seelig, J. 1 986. Bio chemistry 25: 6799-6807 1 4 1 . Pahl-Wostl, c., Seelig, J. 1 987 . BioI. Chel11. Hoppe-Seyler 368: 205- 1 4 1 4 2 . Pan, J . W . , Ham, J. R . , Rothman, D . L., Shulman, R . G. 1 988. Pruc. Nail. Acad. Sci. USA 85: 7836-39 1 43 . Park , J. H., Brown, R. L., Park, C R . , Cohn , M . , Chance, B . 1 988. Proc. Natl. Acad. Sci. USA 78: 67 1 4- 1 8 144. Park, J . H . , Brown, R . L., Park, C . R . , McCully, K., Cohn, M . , et al. 1 987. Proc. Natl. A cad. Sci. USA 84: 897680 145. Petroff, O . A. C. 1988. Compo Biochem. Physiol. 908: 249-60 1 46. Petroff, O. A . c . , Prichard, 1. W., Behar, K . L., Alger, J. R., den Hollan der, J. A., Shulman, R. G. 1 98 5 . Neur ology 3 5 : 78 1 -88 1 47. Petroff, O. A . c., Yu , R. K., Ogino, T. 1 986. J. Neurochern. 47: 1 270-76 1 48 . Prichard, J. W . , Shulman, R. G. 1 986. Anna. ReI!. Neurnsci. 9: 6 1 -R5 1 49. Radda, G. K. 1 986. Biochem. Soc. Trans. 1 4: 5 I 7-25 1 50. Radda, G. K. 1 986. Science 233: 64045 1 5 1 . Remy, C, Albrand, J. P., Benabid, A . L., Decorps, M . , Jacrot, M . , et a l. 1 987. Magn. Reson. Med. 4: 1 44-52 . 1 52 . Renshaw, P. F., Schnall, M. D . , LeIgh, J. S. 1 987. Magn. Reson. Med. 4: 2 2 1 26 1 53 . Reo, N. V., Ewy, C. S . , Siegfried, B . A., Ackerman, J. J . H . 1 984. J. Magn. Reson. 58: 76-84 1 54. Reo, N. V . , Siegfried, B. A., Ackerman, J. J. H. 1 984. J. BioI. Chern. 259: 1 366467 1 55 . Richards, T. L., Kcniry, M . A., Wein stein, P. R., Pereira, B. M . , Andrews, B. T., et al. 1 987. Magn. Reson. Med. 5: 353-57 1 56. Sauter, A., Rudin, M. 1 987. Magn. Reson. Med. 4: 1 -8



1 57. Schnall, M. D . , Yoshizaki, K . , Chance, B., Leigh, 1. S. Jr. 1 988. Magn. Reson. Med. 6: 1 5-23 1 58. Shalwitz, R. A., Reo, N. V . , Becker, N . N . , Hill, A . c., Ewy, C. S., Ackerman, J. J. H. 1 989. J. Bioi. Chem. 264: 393034 1 59. Sherry, A. D., Malloy, C. R., Roby, R. E., Rajagopal, A., J effrey , P. M . H . 1 988. Biuchem. J. 254: 593-98 1 60. Sherry, A. D . , Nunnally, R. L., Peshok, R. M. 1985. J. Bioi. Chem. 260: 927279 1 6 1 . Shoubridge, E. A., Bland, J. L., Radda, G. K. 1 984. Biochim. Biophys. Acta 805: 72-78 1 62 . Shouh rid ge , E. A . , Radda, G. K. 1 984. Biochim. Biophys. Acta 805: 79-88 1 63 . Shulman, G . !" Rossetti, L . , Rothman, D. L., Blair, J. B . , Smith, D. 1987. J. Clin. Invest. 80: 387-93 1 64. Shulman, G. I., Rothman, D. L., Smith, D., Johnson, c., Blair, J. B . , et al. 1985. J. Clin. Invest. 76: 1 229-36 1 65. Shulman, R. G. 1 988. Trends Biochem. Sc i. 1 3 : 37-39 1 66. Siegfried, B. A., Reo , N. V., Ewy, C. S . , Shalwitz, R. A., Ackerman, J. J. H . , McDonald, J. M . 1 986. J . Bioi. Chem. 260: 1 6 1 37-42 1 67. Siess, E. A., Brocks, D. G . , Wieland, O. H. 1 982. In Metabolic Compartmen tation, ed. H . Sies, pp. 235 57. New York: Academic 1 68 . Sillcrud, L. 0., Han, C. H . , B itensky , M. W., Francendese, A. A. 1986. J. Biul. Chem. 26 1 : 4380-88 1 69. Sillerud, L. 0., Shulman, R . G. 1983. Biochem istry 22: 1 087-94 1 70. Silver, M . S., Joseph, R . 1., Hoult, D. I. 1 984. J. Magn. Reson. 59: 347-5 1 1 7 1 . Soboll, S., Bunger, R. 1 98 1 . Hoppe Seylers Z. Physiol. Chem. 362: 1 25-32 1 72. Soboll, S., Scholtz, R . , Heldt, H. W. 1978. Eur. J. Biochem. 87: 377-90 1 73. Sotack, C. H . , Freeman, D. M . , Hurd, D. R. 1 988. J. Magn. Reson. 78: 35561

67

1 74. Springer, C . S. 1987. Annu. Rev. Bio phys. Biophys. Chem. 1 6: 375-99 1 75. Stevens, A. N., M o rri s, P. G., Ties, R . A., Sheldon, P. W . , Griffiths, J . R . 1984. Br. J. Cancer 50: 1 1 3- 1 7 1 76. Stolk, J. A . , Olsen, J. I . , Alderman, D. W., Schweizer, M. P. 1 987. Magn. Reson Med. 4: 144-52 1 77. Stromsky, M . E., Arias-Mendoza, F . , Alger, J. R., Shulman, R. G. 1986. Magn. Reson. Med. 3: 24-32 1 78. Stubbs, M . , Freeman, D., Ross, B. D. 1984. Biochem. J. 224: 241-46 1 79. Styles, P. 1 988. In Physics of NMR

1 80. 181. 1 82. 183.

1 84. 1 85.

1 86. 1 87. 188.

1 89.

1 90.

191.

Spectroscopy in B iology and Medicine, ed. B. M araviglia, pp. 4 1 2-39. Amster dam: North-Holland Physics Tofts, P., Wray, S. 1985. J. Physiol. 359: 4 1 7-29 Tofts, P., Wray, S. 1 988. Nuc!. Magn. Reson. Biomed. 1 : 1 - 1 0 Turner, C . J . 1 984. Prog. Nuc!. Magn. Reson. Spec/rosc. 1 6: 3 1 1-70 Ugurbi1, K. , Garwood, M . , Bendall, M. R. 1 987. J. Magn. Reson. 72: 1 7785 U gurbi l , K . , Garwood, M . , Rath, A . R . 1 988. J. Magn. Reson. 80: 448-69 Veech, R. L., Randolph-Lawson, J. W., Cornell, N . W., Krebs, H . A . 1 979. J. Bioi. Chem. 254: 6538-47 Vink, R., MacIntosh, T., Faden, A. I . 1 988. Magn. Reson. Med. 7: 95-99 Weiner, M. W. 1 988. Invest. Radiol. 23: 253-61 Wolf, W., Albright, M . J., Silver, M . S . , Weber, H . , Reichardt, V . , Sauer, R . 1 987. Magn. Reson. Imag. 5: 1 65-69 Wyrwicz, A. M., Convoy, C. B., Nichols, B. G . , Ryback, K. R., Eisele, P. 1987. Biochim. Biophys. Acta 929: 27 1-77 Wyrwicz, A. M . , Pszenny, M. H . , Scho field, J. c., Tillman, P. c., Gordon, R . E., M a rt i n, P. A. 1 983. Science 232: 428-30 Yoshizaki, K . , Seo, Y., Nishikawa, H . 1 98 1 . Biochim. Biophys. Acta 678: 283 91

19F NMR studies of enflurane elimination and metabolism.

Studies on the metabolism of fluorinated xenobiotics in the rat using 19F-NMR and 1H-NMR spectroscopy.

NMR studies of the lipid metabolism of T47D human breast cancer spheroids.

Effect of glyphosate on plant cell metabolism. 31P and 13C NMR studies.

NMR studies of ligand binding.

13C NMR studies of glucose metabolism in human leukemic CEM-C7 and CEM-C1 cells.

31P-NMR studies of glucose and glutamine metabolism in cultured mammalian cells.

NMR studies of prebiotic polypertides.

NMR studies of membrane proteins.

NMR studies of retinal proteins.

Metabolic transient studies by NMR.

NMR studies of nucleic acid dynamics.

Multinuclear NMR studies of the trp-repressor.

Three-dimensional heteronuclear NMR studies of RNA.

13C NMR studies of bacterial fermentations.

NMR studies of fluorinated visual pigment analogs.

NMR studies of dynamic biomolecular conformational ensembles.

Protein-Inhibitor Interaction Studies Using NMR.

1H NMR studies on lanthanides substituted transferrins.

Biosynthesis of drug metabolites and quantitation using NMR spectroscopy for use in pharmacologic and drug metabolism studies.

Lipid metabolism in large T47D human breast cancer spheroids: 31P- and 13C-NMR studies of choline and ethanolamine uptake.

Fatty acid metabolism and contractile function in the reperfused myocardium. Multinuclear NMR studies of isolated rabbit hearts.

Lipid metabolism in T47D human breast cancer cells: 31P and 13C-NMR studies of choline and ethanolamine uptake.

13C and 31P NMR studies of glucose and 2-deoxyglucose metabolism in normal and enzyme-deficient human erythrocytes.