In Vitro 31P MR Chemical Shifts of In Vivo-Detectable Metabolites at 3T as a Basis Set for a Pilot Evaluation of Skeletal Muscle and Liver 31P Spectra with LCModel Software.

Most in vivo 31P MR studies are realized on 3T MR systems that provide sufficient signal intensity for prominent phosphorus metabolites. The identification of these metabolites in the in vivo spectra is performed by comparing their chemical shifts with the chemical shifts measured in vitro on high-field NMR spectrometers. To approach in vivo conditions at 3T, a set of phantoms with defined metabolite solutions were measured in a 3T whole-body MR system at 7.0 and 7.5 pH, at 37 °C. A free induction decay (FID) sequence with and without 1H decoupling was used. Chemical shifts were obtained of phosphoenolpyruvate (PEP), phosphatidylcholine (PtdC), phosphocholine (PC), phosphoethanolamine (PE), glycerophosphocholine (GPC), glycerophosphoetanolamine (GPE), uridine diphosphoglucose (UDPG), glucose-6-phosphate (G6P), glucose-1-phosphate (G1P), 2,3-diphosphoglycerate (2,3-DPG), nicotinamide adenine dinucleotide (NADH and NAD+), phosphocreatine (PCr), adenosine triphosphate (ATP), adenosine diphosphate (ADP), and inorganic phosphate (Pi). The measured chemical shifts were used to construct a basis set of 31P MR spectra for the evaluation of 31P in vivo spectra of muscle and the liver using LCModel software (linear combination model). Prior knowledge was successfully employed in the analysis of previously acquired in vivo data.

Origin of the 31 P MR signal at 5.3 ppm in patients with critical limb ischemia.

An unknown intense signal (Pun ) with a mean chemical shift of 5.3 ppm was observed in 31 P MR spectra from the calf muscles of patients with the diabetic foot syndrome. The aim of the study was to identify the origin of this signal and its potential as a biomarker of muscle injury. Calf muscles of 68 diabetic patients (66.3 ± 8.6 years; body mass index = 28.2 ± 4.3 kg/m2 ) and 12 age-matched healthy controls were examined by (dynamic) 31 P MRS (3 T system, 31 P/1 H coil). Phantoms (glucose-1-phosphate, Pi and PCr) were measured at pH values of 7.05 and 7.51. At rest, Pun signals with intensities higher than 50% of the Pi intensity were observed in 10 of the 68 examined diabetic subjects. We tested two hypothetical origins of the Pun signal: (1) phosphorus from phosphoesters and (2) phosphorus from extra- and intracellular alkaline phosphate pools. 2,3-diphosphoglycerate and glucose-1-phosphate are the only phosphoesters with signals in the chemical shift region close to 5.3 ppm. Both compounds can be excluded: 2,3-diphosphoglycerate due to the missing second signal component at 6.31 ppm; glucose-1-phosphate because its chemical shifts are about 0.2 ppm downfield from the Pi signal (4.9 ppm). If the Pun signal is from phosphate, it represents a pH value of 7.54 ± 0.05. Therefore, it could correspond to signals of Pi in mitochondria. However, patients with critical limb ischemia have rather few mitochondria and so the Pun signal probably originates from interstitia. Our data suggest that the increased Pun signal observed in patients with the diabetic foot syndrome is a biomarker of severe muscular damage.

Different acute effects of fructose and glucose administration on hepatic fat content.

BACKGROUND: Diets rich in fat and added sugars (especially fructose) play an important role in the pathogenesis of nonalcoholic liver disease (NAFLD), but there is only limited information on the acute effects of these nutrients on hepatic fat content (HFC). OBJECTIVES: We therefore explored how the administration of high-fat load, glucose, fructose, and combinations thereof affects HFC measured in vivo using proton magnetic resonance spectroscopy (1H-MRS) in healthy subjects. METHODS: Ten healthy nonsteatotic male volunteers (age 38.5 ± 9.6 y, body mass index [BMI, kg/m2] 26.9 ± 2.7) underwent, in random order, 6 experiments, each lasting 8 h, that included: 1) fasting; 2) a high-fat load (150 g of fat [dairy cream] at time 0); 3) glucose (3 doses of 50 g at 0, 2, and 4 h); 4) a high-fat load with glucose; 5) fructose (3 doses of 50 g at 0, 2, and 4 h); and 6) a high-fat load with fructose. HFC was measured using 1H-MRS prior to test meal administration (before time 0) and at 3 and 6 h. Plasma concentrations of triglycerides, nonesterified fatty acids, glucose, and insulin were monitored throughout each experiment. RESULTS: HFC increased to 119 ± 19% (P < 0.05) and 117 ± 17% (P < 0.01) of baseline when subjects consumed a high-fat load alone or a high-fat load with fructose, respectively, but was not affected when glucose was coadministered with a high-fat load. HFC was not affected when subjects had fasted or had consumed repeated doses of fructose. When subjects were administered 3 doses of glucose, HFC dropped to 85 ± 13% (P < 0.05) of baseline. CONCLUSIONS: Our results demonstrate that fructose and glucose have a different immediate impact on HFC in humans in vivo. Clinical trial registry: The study was registered at clinicaltrials.gov and obtained clinicaltrials.gov identifier: NCT03680248.