Biomedical applications of 133Cs NMR

NMR IN BIOMEDICINENMR Biomed. 2005;18:125–134Published online in Wiley InterScience (www.interscience.wiley.com). DOI:10.1002/nbm.909

Biomedical applications of 133Cs NMR

James Goodman,1 Jeffrey J. Neil2–4 and Joseph J. H. Ackerman1,4,5*1Department of Chemistry, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA2Department of Neurology, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA3Department of Pediatrics, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA4Department of Radiology, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA5Department of Internal Medicine, Washington University School of Medicine, 660 S. Euclid, St Louis, MO 63110, USA

Received 22 April 2004; Revised 18 June 2004; Accepted 21 June 2004

ABSTRACT: 133Cs NMR is a valuable tool for non-invasively probing biological systems. As a congener of potassium, it

accumulates in the intracellular space, primarily through the action of the Naþ–Kþ pump (ATPase). In addition, it is possible

to resolve the MR signal of 133Cs in different tissue compartments on the basis of chemical shift or MR relaxation properties.

This compartmental resolution applies not only to the intra- and extracellular spaces, but to subcellular compartments as

well. In this review, we discuss the studies defining the ion transport, chemical shift and relaxation characteristics of 133Cs in

living systems. We also review the application of 133Cs NMR to evaluation of ion transport across membranes and the kinetic/

chemical environment of the intracellular space in systems ranging from red blood cells to rat brain. Copyright # 2005 John

Wiley & Sons, Ltd.

KEYWORDS: in vivo MRS; Cs NMR; compartmental resolution; ion transport

INTRODUCTION

Over the past few decades, 133Cs NMR has proven to be avaluable tool for investigating cellular compartments andion transport in vivo. 133Cs is a spin 7/2 nucleus, whosegyromagnetic ratio is roughly 1/8 that of 1H. In its ionicform, 133Csþ relaxation is completely dominated by thequadrupolar mechanism.1 Its nuclear quadrupole momentis very small, however, resulting in long relaxation timesand narrow linewidths. 133Cs is 100% naturally abundantand 100% visible in vivo by standard, solution-state NMRmethods.2–4

The Csþ ion is one of two commonly employedphysiological analogs of the potassium ion. 133Csþ and87Rbþ, the other Kþ analog, are both much more sensitiveto the NMR experiment than is 39K, whose NMR recep-tivity is roughly 100 times less than that of the twoanalogs.5 133Csþ accumulates in the intracellular space6,7

primarily by the action of the Naþ–Kþ pump,2,3,6–13 andhas a transmembrane permeability of 0.1–0.2 of that ofKþ. Rubidium has a transmembrane permeability that is

roughly equivalent to that of Kþ,14–16 perhaps because theionic radius of Rbþ (148 pm) is more similar to Kþ

(133 pm) than is that of Csþ (167 pm).17 Interestingly,hydration numbers are roughly the same: 3.9 for Csþ, 4.0for Rbþ, and 4.6 for Kþ.18 Despite the similarity inhydration spheres for Csþ and Rbþ, the relatively largequadrupolar coupling constant of 87Rb results in a very shortT2 relaxation time constant (�0.97 ms, in vivo19) andcorrespondingly broad linewidths compared to those of133Cs. Furthermore, 133Cs is highly polarizable (its Stern-heimer antishieding factor is roughly twice that of 87Rb),which makes its chemical shift sensitive to its local envir-onment.20 Because of these attributes, a number of research-ers have employed 133Cs NMR to monitor Csþ as a probe ofthe biological activity and environment of potassium.

As Csþ is typically present only in trace amountsin vivo, it must be supplied from an exogenous source.Several researchers have introduced Csþ into cells byincubating suspensions of human erythrocytes in solu-tions containing CsCl.2,21–24 These and many othergroups have also perfused cell cultures and organs withCsþ-containing media. This method is popular becausemanipulation of perfusate composition allows the inves-tigator to easily monitor cesium influx and efflux and toapply agents that alter the biophysical process understudy. Shehan et al.4 introduced a method of dietaryloading, whereby the animal is fed a low-potassium dietand given drinking water that contains equal concentra-tions of KCl and CsCl. This method achieves higherconcentrations of cesium in some tissues (60–70mmol/g

Copyright # 2005 John Wiley & Sons, Ltd. NMR Biomed. 2005;18:125–134

*Correspondence to: J. J. H. Ackerman, Campus Box 8227, EastBuilding, 4525 Scott Avenue, St Louis, MO 63110-1093, USA.E-mail: [email protected]/grant sponsor: NIH; contract/grant numbers: R24 CA83060;R01 NS35912.

Abbreviations used: ADC, apparent diffusion coefficient; 2DG, 2-deoxy-D-glucose; DPG, 2,3-diphosphoglycerate; IBMX, 3-isobutyl-1-methylxanthine; KH, Krebs–Henseleit; L-NAME, NG-nitro-L-argininemethyl ester; NECA, 50-N-ethyl-carboxamidoadenosine; NO, nitricoxide radicals; SIN-1, 3-morpholino sydnonimine.

wet weight in liver and muscle) than can be obtainedin vitro. The 133Cs NMR signal from Csþ can be observedwithin 3 days, although the accumulation of Csþ is oftenfatal after 2 weeks of dietary loading. Intraperitonealinjection of 2 mmol CsCl per kg daily over a period of 6–12 days is a method put forth by Li et al.25 While thismethod is attractive because of its simplicity, one draw-back is that Csþ levels only reach about one-fifth of thoseattained through dietary loading.

The marked sensitivity of 133Csþ relaxation propertiesand chemical shift to local environment has been em-ployed as a powerful tool to report on compartment-specific molecular environments. Intracellular 133Csþ hasa different chemical shift than extracellular 133Csþ.Therefore, 133Cs NMR has been used to spectroscopicallyresolve Csþ in the extracellular and intracellular com-partments without the use of exogenous shift reagent. Insome cases, natural resolution of the cytosol and orga-nelles is also attainable. This compartment-specific che-mical shift allows measurements of membranetranslocation of Csþ, providing information on activeand passive compartmental exchange mechanisms inmany biological systems. 133Cs NMR has also beenused to measure local contrast agent concentration inthe intracellular space, via alterations in the T1 relaxationtime constant of Csþ in the compartment under study.26

Finally, the high intracellular compartmental specificityof Csþ in vivo has allowed measurement of the incoherentmotion (‘apparent diffusion’) of intracellular Csþ, allow-ing inferences to be made about the incoherent displace-ment of water in this compartment.27

Cesium has several properties that make it useful forprobing molecular structure. These include quadrupolarinteractions and long relaxation times. For example, therehave been 133Cs NMR reports on the quadrupolar split-tings and multiple quantum spectroscopy of Csþ inmacroscopically oriented DNA fibers.28,29 While inter-esting, these studies lie outside the scope of this review.Reports involving NMR studies of 133Csþ interactionswith biologically relevant proteins30,31 were also notincluded for the same reason.

The use of 133Cs NMR in vivo generally falls into twovery broad categories: studies of the compartment-specific properties of Csþ, and the use of Csþ as a Kþ

analog to monitor membrane transport activities in bio-logical systems. The research described below is orga-nized by these two categories, although there is somedegree of overlap between them.

133Cs NMR OF Csþ EMPLOYED AS ACOMPARTMENT-SPECIFIC PROBE

Chemical shift

One of the most interesting and useful properties of Csþ

in vivo is that its physical properties when it is present

inside cells are somewhat different from those when it ispresent outside cells. The intracellular 133Csþ signal isgenerally shifted to greater chemical shift (higher reso-nance frequency) than the extracellular signal by 1–2 ppm. Furthermore, extracellular Csþ generally has a133Cs T1 relaxation time constant that is several timeslonger than that of intracellular Csþ.

Davis et al.2 first observed the 133Csþ chemical shiftdifference between the intracellular and extracellularspace in human erythrocytes and perfused rat hearts.They proposed that the higher frequency intracellularresonance is a result of the presence of intracellularphosphates, intracellular Cl�, and, possibly, the ‘non-ideality’ of cytosolic water. The term ‘non-ideality,’ inreference to intracellular water, refers to the idea that thehydrogen bonding network among water molecules, andthus the ‘structure’ of water, inside the cell is markedlydifferent than that of ‘ideal’ water (‘ideal’ water refers towater found in the pure state or in dilute aqueoussolutions). It has been suggested that the non-ideality ofwater results from the high concentration of solute insidecells,12,32,33 such as hemoglobin in erythrocytes. It isexpected that, as the number of solvated species in-creases, so does the fraction of water that comprises thevarious spheres of hydration for these solutes. It has beenpostulated that, as this solute associated water fractionbecomes large enough, it causes disruption of the net-work of hydrogen bonds both between water moleculesand between water molecules and solutes such as Csþ.Since the Csþ ion is large and highly polarizable,20 it isespecially sensitive to these disruptions. Many authorscite ‘non-ideality’ of cytosolic water as contributing tothe 133Cs chemical shift of intracellular Csþ, and weemploy the term herein solely to indicate this usage. Toour knowledge, a quantitative measure of water ‘non-ideality’ has yet to be developed; likewise, a quantitativedescription of the effect of water non-ideality on 133Csþ

chemical shift has yet to be offered and validated.In 1992, Wittenkeller et al.33 set out to test Davis’s

proposals. In attempting to determine the source of theintracellular 133Cs chemical shift of Csþ, they examinedthe effects of the magnetic susceptibility of the medium,the non-ideality of the medium, and Csþ binding tointracellular phosphates, hemoglobin and cell mem-branes. Several buffers were made, consisting of physio-logic concentrations of individual components of theintracellular space of red blood cells. The authors ob-served a linear relationship between 133Csþ chemicalshift and 2,3-diphosphoglycerate (DPG) concentration,suggesting that interaction of Csþ with DPG plays a rolein the chemical shift of 133Cs. Through the use of abuffered red blood cell lysate solution, the authorsreported that non-ideality of water also contributes tothe difference in chemical shift. Evidence for this was theobservation that the 133Csþ chemical shift increased withincreasing concentration of proteins which were unlikelyto interact significantly with Csþ (carboxyhemoglobin

126 J. GOODMAN ET AL.


and lysozyme) but which would lead to non-ideal water.The authors concluded that binding of Csþ to intracel-lular phosphates (in particular DPG) and the non-idealityof intracellular water induced by the high concentrationof hemoglobin are primarily responsible for the differ-ence in 133Cs chemical shift of Csþ in the intra- andextracellular space of red blood cells.

Shehan et al.4 verified Wittenkeller’s conclusion con-cerning DPG using in vivo experiments. These experi-ments were performed using red blood cells taken fromCsþ-fed rats, whereas Wittenkeller’s conclusions werereached primarily through the use of buffers containing

free Csþ and DPG. When erythrocyte DPG concentrationwas varied, Shehan et al. also observed a linear relation-ship between 133Csþ chemical shift and DPG concentra-tions. However, it was apparent that DPG concentrationswere not the sole determinant of intracellular Csþ che-mical shift.

In the in vivo setting, Sakhnini et al.12 and Gilboaet al.13 studied the 133Cs chemical shift of Csþ under theconditions of extremely high salt concentrations using thehalophilic halotolerant bacterium Ba1. These cells, whichcan be found in the Dead Sea, tolerate extremely highconcentrations of salt. Ba1 cells were grown in a series of

Table 1. T1, T2 and T2* relaxation time constants for various tissues

Reference Tissue Loading Temperature Field (T) Environment Compartment T1 (s) T2 (ms) T2* (ms)method

4 Liver Diet 29 7.0 Ex vivo 1.26� 0.05 28� 6b 636 Liver Diet 29 11.75 Ex vivo IC 1.58� 0.10 62� 8a

36 Liver Diet 5 11.75 Ex vivo IC 1.04� 0.05 36� 4a

36 Liver Diet 29 7.05 Ex vivo IC 1.26� 0.11 28� 6b

36 Liver Diet 7.05 In vivo IC 1.19� 0.07 32� 3b

26 Liver Perfusion 37 4.7 Ex vivo IC 1.54 Brain Diet 29 7.0 Ex vivo 2.64� 0.12 67� 1b 1025 Brain i.p. 37 4.7 In vivo 1.9� 0.1 53� 8a 12� 325 Brain i.p. rt 7.05 Ex vivo 2.9� 0.1 87� 3a 11� 236 Brain Diet 29 11.75 Ex vivo IC 3.68� 0.16 120� 10a

36 Brain Diet 5 11.75 Ex vivo IC 2.25� 0.18 81� 10a

36 Brain Diet 29 7.05 Ex vivo IC 2.64� 0.12 67� 1b

2 Heart Perfusion 37 8.5 Ex vivo IC 2.52 Heart Perfusion 37 8.5 Ex vivo EC 123 Heart Perfusion 37 7.05 Ex vivo IC 2.1� 0.3 69� 7a 29� 725 Kidney i.p. 37 4.7 In vivo 1.6� 0.1 45� 3a 8� 125 Muscle i.p. 37 4.7 In vivo 1.8� 0.2 34� 9a 12� 225 Muscle i.p. rt 7.05 Ex vivo 2.9� 0.1 109� 6a 16� 14 Muscle Diet 29 7.0 Ex vivo 2.54� 0.24 56� 2b 1036 muscle Diet 29 11.75 Ex vivo IC 3.30� 0.30 100� 10a

36 Muscle Diet 5 11.75 Ex vivo IC 1.90� 0.16 52� 6a

36 Muscle Diet 29 7.05 Ex vivo IC 2.54� 0.24 56� 2b

36 Muscle Diet 7.05 In vivo IC 2.29� 0.11 65� 19b

4 RBC Diet 29 7.0 Ex vivo 2.02� 0.26 263� 49b 2325 RBC i.p. rt 7.05 Ex vivo 1.7� 0.2 320� 20a 19� 22 RBC Incubation 34 11.75 Ex vivo IC 4.5 330 b 332 RBC Incubation 34 11.75 Ex vivo IC 480� 40a

2 RBC Incubation 34 11.75 Ex vivo IC 610� 20a

2 RBC Incubation 34 11.75 Ex vivo BU 13.636 RBC Diet 29 11.75 Ex vivo IC 2.39� 0.14 480� 30a

36 RBC Diet 5 11.75 Ex vivo IC 1.14� 0.10 210� 20a

36 RBC Diet 29 7.05 Ex vivo IC 2.02� 0.26 260� 50b

22 RBC Incubation 9.4 Ex vivo, buffer A IC 5.1722 RBC Incubation 9.4 Ex vivo, buffer B IC 4.9621 RBC Incubation 37 7.05 or 11.75 Ex vivo IC 3.38� 0.034 Plasma Diet 29 7.0 Ex vivo 6.49� 0.19 1120� 30b 5325 Plasma i.p. rt 7.05 Ex vivo 5� 1 510� 40a 40� 4036 Plasma Diet 29 11.75 Ex vivo IC 7.06� 0.06 1190� 130a

36 Plasma Diet 5 11.75 Ex vivo IC 4.40� 0.05 380� 100a

36 Plasma Diet 29 7.05 Ex vivo IC 6.49� 0.19 1120� 30b

9 endo Perfusion 37 9.4 Ex vivo IC 2.0� 0.323 mrt Perfusion 21 9.4 In vivo CY 1.91� 0.0423 mrt Perfusion 21 9.4 In vivo VA 6.09� 0.0923 mrt Perfusion 21 9.4 In vivo ES 6.11� 0.23

RBC, red blood cells; mrt, maize root tips; i.p., intra-peritoneal injection; endo, endothelial cells; IC, intracellular; EC, extracellular; BU, buffer; CY,cytoplasm; VA, vacuole.a CPMG echo.b Hahn echo.

BIOMEDICAL APPLICATIONS OF 133Cs NMR 127


media containing 25 mM CsCl and total salt concentra-tions of 0.2–4 M. The intracellular 133Csþ resonance was2 ppm higher frequency than the extracellular resonanceat 0.2 M extracellular salt concentration and 16 ppm lowerfrequency than the extracellular resonance at 3.5 M ex-tracellular salt concentration. In addition to this increas-ing chemical shift dispersion, both resonances shifted tohigher frequencies as extracellular salt concentrationsincreased. The intracellular and extracellular resonancesshifted a total of about 8 and 25 ppm, respectively, as theextracellular salt concentration increased from 0.2 to3.5 M. These effects were attributed to the presence ofcounter ions and possibly the non-ideality of water athigher salt concentrations.

As noted above, 133Cs chemical shift is attributed notonly to non-ideality of water, but also to binding of Csþ toother intracellular molecules, particularly intracellularphosphates. Lin et al.34 quantified, through measurementof 133Cs T1 values, Csþ binding to intracellular phos-phates under physiologic conditions. They found Csþ tobind to DPG (66� 8 M

�1), erythrocyte membranes(55� 2 M

�1), ATP (25� 3 M�1), and ADP (19� 1 M

�1).Urry et al.30 used a similar analysis to determine Csþ

binding constants to the channel-forming peptide grami-cidin A (60 M

�1 tight and 4 M�1 weak).

Relaxation

In addition to compartmental differences in 133Csþ che-mical shift, there are also differences in relaxation prop-erties. Table 1 shows relaxation parameters in varioustissues in various settings. While most authors reportthese values in passing, some focus on and exploit thevariations in relaxation times in intracellular and extra-cellular compartments.

In 1977, Edzes et al.35 found the transverse relaxationof 133Csþ in halobacterium to be bi-exponential. Fromthis observation the authors speculated that Csþ is in twodifferent, non-exchanging intracellular environments,corresponding to the two different relaxation rates.Shehan et al.36 showed that this interpretation is justone of several possible explanations of this observation.Since 133Cs is a spin 7/2 nucleus, both its transverse andlongitudinal relaxation can be described by a sum of fourexponentials.36–38 The amplitudes of these exponentialsdepend on the product of the resonance frequency andthe motional correlation time. When this product ismuch less than 1, the amplitudes are such that oneexponential dominates both the transverse and longitu-dinal relaxation. When the product approaches 1, thepresence of multiple exponentials becomes strongly evi-dent in the transverse relaxation decay, yet the long-itudinal relaxation is still dominated (>99.9%) by asingle exponential. This may explain Edzes’s findings,as multiple exponentials can often be modeled well as abi-exponential function unless signal-to-noise ratio is

high and signal decay is measured at a high samplingdensity.

Shehan et al.36 measured and discussed relaxationtimes of 133Csþ in various tissues, both in vivo andex vivo. The tissues were derived from Csþ-fed rats.Measurements were made at two temperatures and twofield strengths. These different conditions change theproduct of the resonance frequency, which increaseswith increasing field strength, and the correlation time,which decreases with increasing temperature. In alltissues, longitudinal relaxation is mono-exponential,transverse relaxation is multi-exponential and much fas-ter, and both T1 and T2 relaxation time constants increasewith field strength and temperature. Shehan concludedthat �99% of Csþ ions are free, having a correlation timeon the order of 10�11 s. This free pool is in fast exchangewith �1% of the Csþ ions that are bound to othermolecules in one or more intracellular compartmentsand have correlation times on the order 10�7 or 10�8 s.

INTRACELLULAR AND EXTRACELLULARCOMPARTMENTATION

Before and during the time that the authors above weredoing work to understand the reason for the chemicalshift and relaxation differences for 133Csþ in differentcompartments in vivo, others were characterizing thisphenomenon in different systems. Davis et al.2 firstdemonstrated compartment-specific properties for Csþ

in the intracellular and extracellular space in suspendedhuman erythrocytes and perfused rat hearts. Suspendedhuman red blood cells were incubated in a buffercontaining 10 mM CsCl at 37 �C, and excised rat heartswere perfused with a buffer containing 2.5 mM CsCl at37 �C. The authors found that the intracellular 133Csþ

resonance was shifted 1–1.4 ppm to higher frequencythan the extracellular 133Csþ resonance. Intracellular133Csþ T1s (4.5 s for red blood cells and 2.5 s for cardiacmyocytes) were three to four times shorter than extra-cellular T1s (13.6 s for red blood cells and 12 s forperfusate). Both compartments gave mono-exponentialrelaxation decay curves. The authors argued that all ofthe intracellular Csþ was observable in erythrocytesbecause the summed intensity of the two signals (intra-and extracellular) was conserved during the loadingprocess, and measurement of 133Csþ T2 by a Hahn spinecho gave a mono-exponential decay. Similar observa-tions could not be made on the rat heart because of thelimitations of this preparation. Ouabain inhibited in-ward flux (into the cell) of Csþ with the rat heartpreparation, suggesting that some translocation is aresult of active transport and that Csþ substitutes for,or competes with, Kþ. This study showed that, at lowCsþ concentrations, the mechanical and electrical prop-erties of the rat heart are not seriously impaired by thepresence of Csþ in the bathing medium.



Subcellular compartmentation

In addition to exhibiting intracellular/extracellular com-partmental resolution, subcellular Csþ compartmentalresolution has been observed by 133Cs NMR in maizeroot tips and hepatocytes. In 1990, Pfeffer et al.23

demonstrated subcellular compartmentation of metalions in living plant cells. Although this study is not a‘biomedical’ one, the development of a model of sub-cellular compartmental resolution in perfused tissue isimportant for the biomedical studies that followed. Roottips from maize seedlings were perfused with Csþ con-taining buffer. While the extracellular 133Csþ resonancefrom the buffer remained constant in amplitude andfrequency, two additional resonances appeared overtime, corresponding to Csþ in the intracellular space.The two resonances developed at different rates, corre-sponding to Csþ influx in the cytosol and in the vacuole.Csþ uptake in the cytoplasm was about nine times fasterthan in the vacuole during the first 300 min of perfusion.133Csþ in the vacuolar compartment exhibited a T1

relaxation time comparable to that of the 133Csþ in theextracellular compartment, about three times longer thanthe T1 of that in the cytoplasm. This was the firstdemonstration of subcellular compartmentation of metalions in living plant tissue.

Wellard et al.39 explored the subcellular specificity ofthe chemical shift of 133Csþ in mammalian tissues.Langendorf perfused hearts, isolated hepatocytes andisolated hepatic mitochondria were obtained from ratsthat had undergone dietary loading of Csþ for 3–5 days.The perfused hearts showed one 133Cs resonance corre-sponding to the Csþ containing perfusate and twostrongly overlapping 133Cs resonances corresponding toCsþ in the intracellular space (at 7.05 T, 39.37 MHz).Packed hepatocytes were analyzed at a higher field(11.75 T, 65.60 MHz) in an attempt to better resolve thetwo intracellular resonances. When valinomycin, a po-tassium ionophore with some mitochondrial specificity,was added to the packed hepatocytes, the ratio of the twounresolved resonances changed drastically, suggestingthat one of the resonances represents Csþ in the mito-chondria. Consistent with this hypothesis, 133Cs NMR ofisolated packed mitochondria showed only a singleresonance. Wellard notes that the frequency differencebetween the two intracellular resonances places an upperlimit on the exchange rate of 32 s� 1. The relativeamplitudes and chemical shifts of the two intracellularresonances, and the redistribution on addition of valino-mycin, are evidence that the two resonances arise fromCsþ in two distinct intracellular compartments such asthe cytosol and mitochondria.

Wellard and Adam40 published another study to furtherelucidate compartmentation in the intracellular space offunctional hepatocytes. Hepatocytes were taken from thelivers of rats that had undergone dietary loading of Csþ

for 5–7 days. The cells were immobilized on agarose

threads and perfused. 31P NMR was used throughout theexperiment to confirm cell viability. As before, there werethree 133Cs resonances (Fig. 1). There was one resonanceat 0.0 ppm, which was assigned to Csþ in the extracellularspace, and two overlapping resonances centered at�1.5 ppm, which were separated by 0.86 ppm. While itis possible that there were more than two intracellularresonances present, the spectral resolution and signal-to-noise ratio were such that they were not evident. The tworesolved resonances demonstrate that there is compart-mentation in the intracellular space and that Csþ ex-change between these compartments is relatively slow.

To test whether or not the relative resonance ampli-tudes depend upon metabolic status, anoxia was inducedby perfusing the cells with a medium that had beenbubbled with 5% CO2 in N2. Glucose and glutaminewere left out of the perfusate to maximize the effects ofanoxia (ATP levels did not drop when these metabolicsubstrates were left in the perfusate). There was nochange in the relative amplitudes of the resonancescorresponding to the two intracellular compartments untilaglycemia was induced in addition to anoxia. In this state,the amplitude of the higher frequency resonance de-creased by 65� 35%, but returned to its previous ampli-tude following normoxic perfusion with a medium thatcontained glucose and glutamine. There were alsochanges in linewidth. The linewidths broadened on tran-sition from anoxia to normoxia. This was interpreted asevidence that Csþ exchange rates between compartmentsincreased.

Brief exposure of the hepatocytes to digitonin (5 mg/mlfor 45 s), which causes selective disruption of the plasmamembrane, caused a decrease in total intracellular Csþ

Figure 1. A 133Cs NMR spectrum of hepatocytes on agar-ose threads at 7.05 T (39.37MHz). Total acquisition timewas 2 h 15min. The large resonance at 0.00 ppm is from Csþ

in the perfusion medium. The two higher frequency reso-nances are from the intracellular compartment andare identified in the text. [Reproduced from Wellard RM,Adam WR. Functional hepatocyte cation compartmentationdemonstrated with 133Cs NMR. Magn. Res. Med. 2001; 46:13–17. Copyright � 2002 John Wiley & Sons, Inc. Thismaterial is used by permission of Wiley-Liss, Inc., a subsidiaryof John Wiley & Sons Inc.]40



of 19� 12%, measured as a decrease in the amplitude ofthe high frequency resonance. The lower frequency reso-nance remained constant following the disruption. Theseobservations are consistent with the assignment of the highfrequency resonance to the cytosol, which is enclosed bythe plasma membrane, and the low frequency resonanceto the spaces enclosed within mitochondrial (or otherorganelle) membranes. This is supported by the anoxic–aglycemic data. Since there are multiple organelles, how-ever, care should be taken in the assignment of subcellularCsþ populations. For example, when a Csþ-perfused intactorgan is studied from an animal having undergone dietaryloading of Csþ, even more 133Csþ resonances may bedetected. For isolated perfused rat livers obtained up to 72 hfollowing partial hepatectomy, as many as four 133Csþ

resonaces have been attributed to the intracellular space(Wellard, personal communication).

As a marker/reporter of the intracellularkinetic environment

Since Csþ accumulates in the intracellular space, Neilet al.27 used dietary loading of Csþ in rat to probeincoherent displacement motion (apparent diffusion) inthe intracellular space of healthy and injured brain inorder to make inferences about the incoherent displace-ment motion of intracellular water under both conditions.The apparent diffusion coefficient (ADC) of Csþ wasmeasured by performing a pulsed-field-gradient spin-echo experiment.41 Brain temperature, as measuredwith a fiber optic temperature probe placed in the frontallobe of the brain, was maintained near 37 �C throughoutthe entire experiment by circulating warm water throughtubes wrapped around the animal’s head.

The Csþ ADC dropped from 0.91� 0.05mm2/ms whenthe rat was alive to 0.71� 0.05mm2/ms immediately afterthe rat died (Fig. 2). The diffusion time (tdiff) in thisexperiment (27 ms) gives root mean squared displace-ments ½ð2 � ADC � tdiffÞ� of 7.0 and 6.2 mm, respectively.Relatively impermeable membranes are likely encoun-tered over this distance. These membranes can beexpected to restrict and hinder the displacement ofCsþ. Thus, this measure places a lower limit on theunrestricted diffusion coefficient within the intracellularenvironment.

The decrease in Csþ ADC after the death of the animalwas taken to indicate that injury causes changes in theintracellular environment that affect motion in that space,resulting in a change in the ADC. The authors suggestseveral possible explanations for this, such as a loss ofcytoplasmic streaming or an increase in the viscosity ofthe intracellular space following cell injury.

Sehy et al.42 performed a similar measurement onhealthy Xenopus oocytes. Because the oocytes are verylarge, single cells having volumes of �1.0ml, the in-tracellular space can be examined directly through ima-ging or volume-localized spectroscopy. The authorsemployed a series molecular species and small ions,including Csþ, and found the apparent diffusion of theintracellular milieu to be fully described by simpleBrownian motion. This suggests that cytoplasmic stream-ing or another active (i.e., ATP-dependent) process doesnot make a significant contribution to displacement mo-tion in the intracellular space of Xenopus oocytes.

. . . and chemical environment

133Cs NMR observables can also be used as reporters ofexogenous species that enter into cells. Colet et al.26 usedthe T1 sensitive amplitude of the intracellular 133Csþ

resonance observed under conditions of partial saturationas a reporter of the concentration of contrast agent, Gd-EOB-DTPA (Eovist1), which crosses into the intracel-lular space of hepatocytes.

Three livers were perfused with a Csþ containingmedium for 60 min and washed for 10 min with Csþ-free medium. Medium containing Gd-EOB-DTPA wasthen added to the perfusate and the livers perfused for30 min. 133Cs NMR data were acquired serially in 5 mintime blocks with a TR of 200 ms, which allowed onlypartial longitudinal relaxation and imparted a strong T1

dependence to the resultant signal amplitude. Aliquots of1 ml (of the 182.5 ml perfusion solution) were removedevery 5 mins for analysis of Gd-EOB-DTPA concentra-tion. After 30 min, the livers were perfused for 10 minwith fresh Krebs–Henseleit (KH) buffer to remove extra-cellular Gd-EOB-DTPA. The livers were then dried over-night for water and Gd quantification via inductivelycoupled plasma mass spectrometry (ICPMS). Forreference, relaxivity calibrations of Gd-EOB-DTPA in

Figure 2. A plot of the 133Csþ ADC (DCs) in rat brain as afunction of time after death. The error bars are standarderrors of the mean, and the asterisk denotes that the ADCfrom Csþ in live rat brain is significantly different from thevalues measured after death by ANOVA (p<0.001).27 Note:10�3mM

2/s¼ 1mm2/ms. [Reproduced from Neil J, DuongTQ, Ackerman JJH, Evaluation of intracellular diffusion innormal and globally-ischemic rat brain via 133Cs NMR.Magn. Res. Med. 1996; 35: 329–335. Copyright � 1996JohnWiley & Sons, Inc. This material is used by permission ofWiley-Liss, Inc., a subsidiary of John Wiley & Sons Inc.]27



intracellular-like buffers were made. The intracellularconcentration of Gd-EOB-DTPA was then estimatedin vivo from knowledge of the expected dependence of133Cs T1 relaxation time constant on Gd-EOB-DTPAconcentration.

After the Gd-EOB-DTPA was added to the Csþ-freemedium, the intracellular 133Csþ resonance amplitudeincreased as expected. The Csþ-free medium assuredthis was not due to further intracellular loading of Csþ.The increase in signal amplitude was interpreted asarising from the reduction in the T1 relaxation timeconstant caused by the uptake of Gd-EOB-DTPA. Theobservation of decreasing Gd-EOB-DTPA concentra-tions in the 1 ml aliquots taken out of the perfusionmedium every 5 min was consistent with this interpre-tation. Intracellular Gd-EOB-DTPA concentration de-termined by ICPMS was significantly higher thanpredicted by 133Csþ relaxivity in vivo. This could be aresult of intracellular subcompartmentation of Csþ,which would cause a fraction to be inaccessible to theGd-EOB-DTPA. It is also possible that Gd-EOB-DTPAhas a reduced relaxivity in vivo compared to thatmeasured in the intracellular-like phantoms that wereused to calibrate its relaxivity.

USES AS A Kþ ANALOG TO MONITORPUMP ACTIVITY/UPTAKE

133Cs NMR has also been used to study Naþ–Kþ pumpactivity and/or Csþ influx rates in healthy and compro-mised tissues. These studies model the total rate ofaccumulation of Csþ in the intracellular space by

d½Csþ�i=dt ¼ ðkpump þ kcÞ½Csþ�e � k�c½Csþ�i ð1Þ

where kpump is the rate constant of active transport ofCsþ into cells, kc and k�c are rate constants for passiveinflux and efflux, respectively, and [Csþ]i and [Csþ]e

are intracellular and extracellular Csþ concentrations,respectively. In perfusion studies, active transport canbe inhibited by chemical means, enabling the contribu-tion of the passive influx rate to be determined directlyby adding a pump inhibitor to the perfusate and mea-suring influx in the absence of pump activity. Studiesdesigned to measure the inhibition of energy dependenttransport have been performed in a variety of livingsystems.

In red blood cells . . .

Davis et al.2 performed the first 133Cs NMR spectroscopymeasurement of this kind in human erythrocytes. Theinvestigators suspended erythrocytes in a solution con-taining 10 mM CsCl and measured the rate of total influxto be 0.6� 10� 3 mol of Csþ per liter of cells per hour,which is about 0.3 times the uptake rate of Kþ. When

they performed the identical measurement on cells sus-pended in 10 mM CsCl and 10� 4

M ouabain (an inhibitorof the Naþ–Kþ pump), the measured rate of total influxhad decreased to 0.3� 10� 3 moles of Csþ per liter ofcells per hour.

Bramham et al.21 investigated Csþ transport in the redblood cells of individuals suffering from alcohol depen-dence. These authors were attempting to verify theobservation that Csþ levels are elevated in erythrocytesof patients with alcohol-dependence syndrome and thepostulate that there is an abnormality in the transport ofCsþ and/or distribution of Csþ in the erythrocytes ofindividuals with alcohol dependence. They found nodifference in the uptake rate of Csþ between red bloodcells from alcohol-dependent and abstinent volunteers.These data indicate that there was no transport abnorm-ality in the erythrocytes of dependent volunteers underthese conditions.

Ren et al.22 examined the effects of lanthanide ion onCsþ uptake and efflux in human erythrocytes. Theauthors used the differences in 133Cs T1s and chemicalshifts of intracellular and extracellular Csþ to estimate[Csþ]i and [Csþ]e. Erythrocytes were suspended for agiven amount of time in buffers containing constantamounts of CsCl and varying amounts of La3þ. [Csþ]i

and [Csþ]e were measured. The ratio of [Csþ]i/[Csþ]e

decreased markedly with increasing La3þ concentration,indicating uptake inhibition or competition. Decreases inCsþ uptake rates and Csþ concentration ratios at equili-brium were attributed to the inhibition of Naþ–Kþ pumpactivity in erythrocyte membranes by La3þ.

To measure the La3þ effect on efflux, Csþ-loadederythrocytes were put in buffers identical to those usedfor Csþ loading except that CsCl was replaced with anequimolar amount of NaCl. Csþ efflux was a slowprocess. No [Csþ]e was observed for 8 h when Csþ-loaded erythrocytes were put in Csþ- and La3þ-freebuffer. At high concentrations of LaCl3 (100 mmol/l),some Csþ was detected in the extracellular space, sug-gesting that La3þ at a high enough concentration candisrupt the erythrocyte membrane.

In endothelial cells . . .

Csþ translocation experiments have also been used tomonitor the efficiency of the Naþ–Kþ pump in endothe-lial cells. Gruwel et al.9 used 133Cs NMR of aorticendothelial cells to test the hypothesis that actin regulatesthe activity of the Naþ–Kþ pump. Endothelial cells frompig aortae were grown on beads and perfused alternatelywith Csþ-containing media and Csþ-free media severaltimes in order to monitor uptake and efflux. (Unlike othercell types studied, Csþ efflux from endothelial cells wasroughly equivalent to its uptake, allowing for multipleserial studies.) Different F-actin rearrangements wereinduced in the cells by perfusion with 2-deoxy-D-glucose



(2DG) and cytochalasin D, both of which also caused anincrease in G-actin. Neither agent caused inhibition ofpump activity, although there were significant effects oncell morphology due to changes in actin levels. Theauthors also investigated the effects of altered ATP levelson Naþ–Kþ pump activity, since 2DG can affect ATPlevels. To inhibit glycolytic and oxidative pathways ofATP production, 5 mM of both 2DG and NaCN wereadded to the perfusion medium. This caused a drastic, butreversible, inhibition of Naþ–Kþ pump activity, as mea-sured by Csþ influx (Fig. 3). When either 20 mM 2DG or5 mM NaCN was added alone, there was no inhibition ofpump activity, indicating that the changes in ATP levelsinduced by 2DG alone are insufficient to inhibit pumpactivity. The authors concluded that changes in actin leveldid not affect Naþ–Kþ pump activity.

Gruwel et al. also evaluated regulation of the Naþ–Kþ

pump in endothelial cells by cGMP11 and cAMP.10 Thepurpose of the cGMP study was to investigate the role ofnitric oxide radicals (NO) in regulation of the Naþ–Kþ

pump by cGMP. As before, endothelial cells taken frompig aortae were grown on microcarrier beads and per-fused with Csþ-containing media. First, cells were per-fused with Csþ-containing media for a control measure ofCsþ uptake. Once complete loading had occurred, wash-out was measured by perfusing the cells with a Csþ-freemedium. The cells were then perfused with either 0.5 mM

3-isobutyl-1-methylxanthine (IBMX), which is an inhi-

bitor of phosphodiesterases, or 0.5 mM IBMXþ 0.2 mM

8-Br-cGMP, which is a membrane-permeable cGMPderivative, or 10mM 3-morpholino sydnonimine (SIN-1), which slowly and continuously releases NO. All threeof these substances increased the intracellular cGMPconcentration, which resulted in a decrease in the amountof intracellular Csþ. IBMX reduced the pump rate of theNaþ–Kþ pump to 75% of control, and IBMXþ 8-Br-cGMP further reduced the pump rate to 59% of control.SIN-1 reduced the pump rate the most, to 52% of control.

Since increased cGMP and NO attenuated pump activ-ity, and it is known that endothelial cells maintain a basalrelease of NO, it appeared to the authors that the Naþ–Kþ

pump might be under continuous regulation by NO (i.e.NO maintains baseline cGMP levels). To test this hypoth-esis, NG-Nitro-L-arginine methyl ester (L-NAME), whichis a well-known inhibitor of NO-synthase, was added tothe perfusate. If the release of NO, which occurs endo-genously, causes an increase in cGMP, then L-NAMEshould cause a decrease in cGMP, which would in turncause an increase in Naþ–Kþ pump activity. Whenperfused with 1 mM L-NAME, the pump activity did, infact, increase 15% above the control level.

To study cAMP regulation of the Naþ–Kþ pump, theauthors used the same general strategy as in the cGMPstudy: 0.5 mM IBMX reduced the pump activity to 75% ofthe control, and 0.5 mM IBMXþ 0.2 mM 8-Br-cAMPreduced the pump activity to 48% of the control.NECA, an adenosine-A2 receptor agonist, and Iloprost,a prostaglandin receptor agonist, were both added to theperfusate at a concentration of 10mM to increase cAMPconcentration through stimulation of adenylate cyclaseactivity. These agents reduced pump efficiency to 64 and60% of control, respectively. These experiments demon-strate that increased cAMP causes a rapid, short-termdecrease in Naþ–Kþ pump activity.

. . . and in perfused organs

Schornack et al.3,43 measured the rate constants govern-ing passive influx and efflux via ion channels, and activetransport via the Naþ–Kþ pump, in perfused rat heartsderived from both healthy and septic animals. Threeexperiments were performed: (1) perfusion with Csþ

containing KH buffer to measure total Csþ influx; (2)perfusion with Csþ containing KH buffer and 0.1, 0.2or 0.4 mM ouabain to inhibit the Naþ–Kþ pump andthus measure only passive Csþ influx; and (3) perfusionof Csþ-preloaded tissue with Csþ free KH buffer tomeasure Csþ efflux. Absolute values of Csþ concentra-tion, intracellular volume and extracellular volume weremonitored or determined by other methods. (Note thatthese authors assigned only one resonance to signal fromthe intracellular space. This may be related to differencesin Cs loading conditions for this study as compared tostudies in which multiple resonances were assigned to

Figure 3. A plot of intracellular Csþ concentration in por-cine aortic endothelial cells, as measured from 133Cs NMRsignal intensity, as a function of time. The solid lines are fitsto uptake and efflux. The dotted line represents maximumefflux resulting from pump inhibition with ouabain.9 [Re-produced from Gruwel ML, Culic O, Schrader J. A 133Csnuclear magnietic resonance study of endothelial Na(+)-K(+)-ATPase activity: can actin regulate its activity? Biophys.J. 1997; 72: 2755–2782. Reprinted with permission of theBiophysical Society]9



intracellular 133Csþ.) Figure 4 shows data fromexperiments 1 and 3.

Sepsis was induced by cecal ligation and perforation24–36 h prior to removal of the heart. Control animalsunderwent surgery in which the cecum was ligated butnot perforated. Rates of Csþ influx and efflux throughpassive channels were found to be the same for bothhealthy and septic hearts, but the rate of influx via theNaþ–Kþ pump was reduced by roughly 24% in the septichearts. The authors speculated that the observed defect in

Naþ–Kþ pump rate is systemic, and that it may have arole in the disruption in the ionic concentration gradient,the reduction of which is considered to be a significantfactor in the morbidity and mortality of sepsis.

CONCLUSION

133Cs NMR has proven to be a valuable tool for exploringcompartmental structure and ion transport processes inliving systems. The compartmental specificity of Csþ

within the intracellular space has been used to probeincoherent displacement motion in the intracellular spaceas well as leakage of relaxation agent into this space.Dynamic measurements of the levels of Csþ ions inperfused tissues have provided researchers with insightinto the rates and efficiency of ion transport acrossmembranes. This has allowed exploration of the mechan-isms responsible for regulating ion transport in healthyand diseased tissue.

Acknowledgments

The authors would like to thank Drs R. Mark Wellard andWilliam R. Adam for their helpful comments on this text.

REFERENCES

1. Wehrli FW. Spin-lattice relaxation of some less common weaklyquadrupolar nuclei. J. Magn. Reson. 1978; 30: 193–209.

2. Davis DG, Murphy E, London RE. Uptake of cesium ions byhuman erythrocytes and perfused rat heart: a cesium-133 NMRstudy. Biochemistry 1988; 27: 3547–3551.

3. Schornack PA, Song S-K, Ling CS, Hotchkiss R, Ackerman JJH.Quantification of ion transport in perfused rat heart: 133Csþ as aNMR active Kþ analog. Am. J. Physiol. 1997; 272: C1618–C1634.

4. Shehan BP, Wellard RM, Adam WR, Craik DJ. The use of dietaryloading of 133Cs as a potassium substitute in NMR studies oftissues. Magn. Reson. Med. 1993; 30: 573–582.

5. Harris RK. Introduction. In NMR and the Periodic Table, HarrisRK, Mann BE (eds). Academic Press: New York: 1978; 4–7.

6. Relman AS. The physiological behavior of rubidium and cesiumin relation to that of potassium. Yale J. Biol. Med. 1956; 29: 248–262.

7. Relman AS, Lambie AT, Burrows BA, Roy AM, Allen JM,Connors HP, Dell ES. Cation accumulation by muscle tissue:the displacement of potassium by rubidium and cesium in theliving animal. J. Clin. Invest. 1957; 36: 1249–1256.

8. Wittam R, Ager ME. Vectorial aspects of adenosine-triphosphateactivity in erythrocyte membranes. Biochem. J. 1964; 93: 337–348.

9. Gruwel ML, Culic O, Schrader J. A 133Cs nuclear magneticresonance study of endothelial Na(þ )-K(þ )-ATPase activity:can actin regulate its activity? Biophys. J. 1997; 72: 2775–2782.

10. Gruwel ML, Culic O, Muhs A, Williams JP, Schrader J. Regula-tion of endothelial Na(þ )-K(þ )-ATPase activity by cAMP.Biochem. Biophys. Res. Commun. 1998; 242: 93–97.

11. Gruwel ML, Williams JP. Short-term regulation of endothelialNa(þ )-K(þ )-pump activity by cGMP: a 133Cs magnetic reso-nance study. Mol. Membr. Biol. 1998; 15: 189–192.

12. Sakhnini A, Gilboa H. Nuclear magnetic resonance studies ofcesium-133 in the halophilic halotolerant bacterium Ba1: chemi-cal shift and transport studies. NMR Biomed. 1998; 11: 80–86.

Figure 4. The top figure is a stacked plot showing theuptake of Csþ in the intracellular space of a perfused hearttaken from a healthy rat. The shift-reagent-free KH buffercontained 1.25mM Csþ. The middle figure shows plots ofthe same heart after the perfusate is switched to Csþ-freeKH buffer. The bottom plot shows the intracellular Csþ

concentration during the time periods of wash-in (solidcircles) and wash-out (open circles) from the two plotsabove. The black lines are best-fit curves to the kineticmodel, eqn (1). [Reproduced from Schornack PA, Song S-K, Ling CS, Hotchkiss R, Ackerman JJH. Quantification of iontransport in perfused rat heart 133Csþ as a NMR active Kþ

analog. Am. J. Physiol. 1997; 272: C1618–C1634. Reprintedwith permission of The American Physiological Society]3



13. Gilboa H, Sakhnini A. NMR studies of the halotolerant bacteriumHolomonas israelensis: sodium-23, cesium-133 and phosphorus-31 studies. Bull. Magn. Reson. 1999; 20: 43–46.

14. Hadley R, Hume J. Permeability of time-dependent Kþ channel inguinea pig ventricular myocytes to Csþ, Naþ, NH4

þ and Rbþ.Am. J. Physiol. 1990; 259: H1448–H1454.

15. Mitra R, Morad M. Permeance of Csþ and Rbþ through theinwardly rectifying Kþ channel in guinea pig ventricular myo-cytes. J. Membr. Biol. 1991; 122: 33–42.

16. Randle J, Oliet S, Renaud J-F. Alkali cation permeability andcaesium blockade of cromakalim-activated current in guinea-pigventricular myocytes. Br. J. Pharmac. 1991; 103: 1795–1801.

17. Berry RS, Rice SA, Ross J. Physical Chemistry. Oxford UniversityPress: New York, 2000.

18. Creekmore R, Reilley C. Nuclear magnetic resonance determina-tion of hydration numbers of electrolytes in concentrated aqueoussolutions. J. Phys. Chem. 1969; 73: 1563–1567.

19. Schornack PA. 133Csþ and 87Rbþ as NMR active Kþ transportanalogs: applications to septic shock. Doctoral thesis. WashingtonUniversity, Saint Louis, MO 1998.

20. Lindman B, Forsen S. The alkali metals. In NMR and the PeriodicTable, Harris RK, Mann BE (eds). Academic Press: New York,1978; 129–182.

21. Bramham J, Riddell FG. Cesium uptake studies on humanerythrocytes. J. Inorg. Biochem. 1994; 53: 169–176.

22. Ren J, Zhang S, Wang Y, Pei F, Zhang J, Ni J. Effects of lanthanum(III) upon cesium (I) uptake by human erythrocytes: a 133Cs NMRstudy. Chin. Sci. Bull. 1995; 40: 1617–1620.

23. Pfeffer PE, Rolin DB, Schmidt JH, Tu S-I, Kumosinski TF, DoudsDD, Jr. In vivo cesium-133 NMR a probe for studying subcellularcompartmentation and ion uptake in maize root tissue. Biochim.Biophys. Acta 1990; 1054: 169–175.

24. Pfeffer PE, Rolin DB, Schmidt JH, Tu S-I, Kumosinski TF, DoudsDD, Jr. Ion transport and subcellular compartmentation in maizeroot tissue as examined by in vivo cesium-133 NMR spectrscopy.J. Plant Nutr. 1992; 15: 913–927.

25. Li Y, Neil J, Ackerman JJH. On the use of 133Cs as an NMR activeprobe of intracellular space in vivo. NMR Biomed. 1995; 8:183–189.

26. Colet J-M, Lecomte S, Elst LV, Muller RN. Cesium-133: apotential reporter of the hepatic uptake of contrast agents.Magn. Reson. Med. 2001; 45: 711–715.

27. Neil J, Duong TQ, Ackerman JJH. Evaluation of intracellulardiffusion in normal and globally-ischemic rat brain via 133CsNMR. Magn. Reson. Med. 1996; 35: 329–335.

28. Einarsson L, Kowalewski J, Nordenskioeld L, Rupprecht A.Cesium-133 NMR of oriented CsDNA and multiple-quantum spectroscopy of I¼ 7/2. J. Magn. Reson. 1989; 85:228–234.

29. Einarsson L, Nordenskioeld L, Rupprecht A, Furo I, Tuck C.Lithium-7 and cesium-133 spin relaxation in macroscopicallyoriented in Li- and Cs-DNA fibers. J. Magn. Reson. 1991; 93:34–46.

30. Urry DW, Trapane TL, Brown RA, Venkatachalam CM, PrasadKU. Cesium-133 NMR longitudinal relaxation study of ionbinding to the gramicidin transmembrane channel. J. Magn.Reson. 1985; 65: 43–61.

31. Riddell FG, Arumugan S, Patel A. Nigericin-mediated transport ofcesium ion through phospholipid bilayers studied by a 133Csmagnetization-transfer NMR technique. Inorg. Chem. 1990; 29:2397–2398.

32. Kirk K, Kuchel PW. Physical basis of the effect of hemoglobin onthe 31P NMR chemical shifts of various phosporyl compounds.Biochemistry 1988; 27: 8803–8810.

33. Wittenkeller L, Mota de Treitas D, Geraldes CFGC, Tome AJR.Physical basis for the resolution of intra- and extracellular 133CsNMR resonances in Csþ-loaded human erythrocyte suspensions inthe presence and absence of shift reagents. Inorg. Chem. 1992; 31:1135–1144.

34. Lin W, Mota de Treitas D, Zhang Q, Olsen KW. Nuclear magneticresonance and oxygen affinity study of cesium binding in humanerythrocytes. Arch. Biochem. Biophys. 1999; 369: 78–88.

35. Edzes HT, Ginzburg M, Ginzburg BZ, Berendsen HJC. Thephysical state of alkali ions in Halobacterium: some NMR results.Experientia 1977; 33: 732–734.

36. Shehan BP, Wellard RM, Craik DJ, Adam WR. 133Cs relaxationtimes in rat tissues. J. Magn. Reson. B 1995; 107: 179–185.

37. Andersson T, Drakenberg T, Forsen S, Thulin E, Sward M. Directobservation of 43Ca NMR signals from Ca2þ ions bound toproteins. J. Am. Chem. Soc. 1982; 104: 576–580.

38. Bull TE, Forsen S, Turner DL. Nuclear magnetic relaxation ofspin 5/2 and spin 7/2 nuclei including the effects of chemicalexchange. J. Chem. Phys. 1979; 70: 3106–3111.

39. Wellard RM, Shehan BP, Craik DJ, Adam WR. Demonstration oftissue cation compartmentation using 133Cs NMR. Bull. Magn.Reson. 1996; 18: 184–185.

40. Wellard RM, Adam WR. Functional hepatocyte cation compart-mentation demonstrated with 133Cs NMR. Magn. Reson. Med.2002; 48: 810–818.

41. Stejskal EO, Tanner JE. Spin diffusion measurements: spin echoesin the presence of a time-dependent field gradient. J. Chem. Phys.1965; 42: 288–292.

42. Sehy JV, Ackerman JJH, Neil J. Apparent diffusion of water, ions,and small molecules in the Xenopus oocyte is consistent withBrownian displacement. Magn. Reson. Med. 2002; 48: 42–51.

43. Schornack PA, Song S-K, Hotchkiss R, Ackerman JJH. Inhibitionof ion transport in septic rat heart: 133Csþ as an NMR active Kþ

analog. Am. J. Physiol. 1997; 272: C1635–C1641.



Biomedical applications of 133Cs NMR

Documents

Transcript of Biomedical applications of 133Cs NMR