Structural difference in the H+,K+-ATPase between the E1 and E2 conformations . An attenuated total...

Structural difference in the H+,K+-ATPase between the E1 and E2conformationsAn attenuated total reflection infrared spectroscopy, UV circular dichroism and

Raman spectroscopy study

Vincent Raussens1, M. PeÂzolet2, Jean-Marie Ruysschaert1 and Erik Goormaghtigh1

1Laboratoire de Chimie-Physique des MacromoleÂcules aux Interfaces, CP 206/2, UniversiteÂ Libre de Bruxelles, Campus Plaine, Belgium;2DeÂpartement de Chimie, Centre de Recherche en Science et IngeÂnierie des MacromoleÂcules, UniversiteÂ Laval, CiteÂ Universitaire, QueÂbec, Canada

Conformational changes taking place in the gastric H+,K+-ATPase when shifting from the K+-induced E2 form to the

E1 form upon replacing K+ ions by Na+ were investigated by different spectroscopic approaches. No significant

secondary-structure change or secondary-structure reorientation with respect to the membrane plane could be

measured by attenuated total reflection Fourier transform infrared spectroscopy of oriented films. Circular dichroism

and Raman spectra obtained on tubulovesicle suspensions indicated no significant secondary structure or tyrosine

and tryptophan side-chain environment changes in tubulovesicle suspensions. The smallest observable structural

changes are discussed in term of the number of amino-acid residues involved for each technique.

Keywords: H+, K+-ATPase; gastric; attenuated total reflection; Fourier transform infrared spectroscopy; structural

changes.

Since the sequences of many P-type ATPases were determinedin the eighties, huge efforts have been devoted to theunderstanding of the mechanism of the coupling of ATPhydrolysis to ion transport. However, this goal remainsremote because of the lack of quantitative measurements ofthe ATPase structure changes accompanying the catalyticcycle.

The gastric H+,K+-ATPase is an a,b-heterodimer [1,2]belonging to the P-type ATPase family. These ATPases, whichinclude the Na+,K+-ATPase, the Ca++-ATPases, the yeast andNeurospora plasma membrane H+-ATPases, are thought to sharea common mechanism and a common structure. According tothe hydropathy plots, the catalytic subunit contains 8±10transmembrane segments. The first four transmembrane seg-ments located in the N-terminal third of the protein are clearlydefined on the hydropathy plots and their existence has beendemonstrated by proteolysis and in-vitro expression experiments[3,4]. The other predicted transmembrane segments, located inthe C-terminal third of the protein, are more ambiguous.Predictions and experiments suggested from four to sixtransmembrane segments in this region of the protein [3,5].Recently, extensive proteolysis experiments have suggested thatmuch more of the ATPase molecule is embedded in the lipidbilayer (see [6,7] for the H+-Neurospora plasma membraneH+-ATPase and gastric H+,K+-ATPase, respectively).

Even less is known about the conformational changes takingplace in the enzyme in the course of the catalytic cycle. Until

now, structural changes in the H+,K+-ATPase have been detectedby fluorosciene isothiocyanate (bound on Lys517) [8] orchanges in trypsin proteolysis susceptibility [9,10] upon shiftingthe conformation from E1 to E2.

Yet, no quantitative determination of the size of the proteinregion affected by the changes has been made. Recentdifferential scanning calorimetry experiments show that arrange-ment of two distinct structural domains characterized by twotemperature-induced cooperative unfolding are present in the E1conformation (Na+ buffer) but that one disappears when theenzyme is forced into the E2 conformation (K+ buffer) [11]

One potential explanation for the vanishing transition in oneof the protein domains is that this domain becomes embedded orpartially embedded in the membrane [11]. The K+ binding sitehas also been suggested to be on the H6 segment [12], a part ofthe H5-H6 hairpin, accessible to trypsin cleavage in the absenceof ligands but not in the presence of K+ [13]. This region can befound either membrane embedded, when occluding cations arepresent, or in the aqueous phase in their absence in the(Na+,K+)-ATPase [14] with a 19-kDa peptide protected againsttrypsin hydrolysis [15]. Therefore, it is a possibility that thesecond cooperative domain observed by differential scanningcalorimetry contains the H5-H6 hairpin. Amide hydrogen/deuterium exchange kinetics for Neurospora plasma membraneH+-ATPase have shown that at least 175 additional amino-acidresidues are shielded from the solvent in an E2 conformation[16]. Whether the conformational change involves majorsecondary-structure changes or small amplitude rearrangementsof existing secondary structures is a major issue for theunderstanding how the catalytic process is coupled to iontransport, and it remains a matter of debate. Gresalfi andWallace [17] suggested a structural change affecting 7% of theNa+,K+-ATPase on the basis of CD measurements. Later,Chetverin and Brazhnikov [18], using infrared spectroscopy,came to the conclusion that the change affected less than 2% of

Eur. J. Biochem. 262, 176±183 (1999) q FEBS 1999

Correspondence to E. Goormaghtigh, Free University of Brussels, Campus

Plaine CP 206/2, B1050 Brussels, Belgium. Fax: +32 26505113,

Tel: + 32 26505386, E-mail: [email protected]

Abbreviations: ATR-FTIR, attenuated total reflection Fourier transform

infrared spectroscopy.

(Received 1 October 1998, revised 24 February 1999, accepted

25 February 1999)

q FEBS 1999 H+,K+-ATPase structural changes (Eur. J. Biochem. 262) 177

the residues, in agreement with a similar estimation obtained forthe H+,K+-ATPase [19].

In the present paper we used different spectroscopictechniques to obtain quantitative information about theH+,K+-ATPase structural changes experienced when theenzyme conformation shifts from E1 to E2. We also discussthe limits of the spectroscopy approaches to their ability todetect small conformational changes in the protein.

MATERIALS AND METHODS

Materials

Proteinase K, ATP and nigericin were purchased from Sigma, StLouis, MO, USA. Deuterium oxide was from Janssen Chimica(Geel, Belgium). All other reagents were of the highest puritygrade commercially available.

Tubulovesicles

Tubulovesicles were isolated from hog gastric fundus bydifferential centrifugation as described by Soumarmon [20].They were further purified from the microsomal fractions bycentrifugation on a sucrose discontinuous density gradient at100 000 g overnight. The material collected at the 8±30%sucrose interface is referred to as the tubulovesicles. ATPasepurity was checked by gel electrophoresis carried out on a 7.5%gel according to Laemmli [21]. A single band at 95 kDacorresponding to the a-subunit of the H+,K+-ATPase was visibleafter SDS/PAGE analysis of a 6-mg sample on a mini protean II(Biorad). ATPase activity of the tubulovesicles was determinedin a medium containing 40 mm Hepes, 2 mm ATP, 2 mm MgCl2at pH 7.2 in the presence or in the absence of 20 mm KCl. Thevesicles were incubated at 37 8C for 15 min in this medium. Thereaction was stopped by addition of 7% SDS (final concentra-tion 1.75%). Inorganic phosphate formation was assayedaccording to Stanton [22], except that the coloration wasrevealed with ascorbate. Sucrose was present at a concentrationof 8% (w/v) in order to keep iso-osmotic conditions andnigericin was present at a concentration 14 mm. Proteins wereassayed using the bicinchoninic acid kit from Pierce or by amodified Lowry method [23]. Bovine serum albumin was usedas standard in both cases. Lipids were assayed with theenzymatic test for the choline reaction group from Boehringerassuming that lipids carrying a choline function represent 50%of the total of the lipids present in the tubulovesicles [24].According to this, the final protein concentration was5.6 mg´mL21 and the lipid concentration was 10 mg´mL21.Sucrose from the gradient was removed for all spectroscopicmeasurements. Sucrose was removed by suspending the vesiclesin 40 mm Hepes/Tris buffer, pH 7.2, and pelleting them. Thepellet was finally resuspended at 5 mg´mL21 protein concentra-tion in the 40 mm Hepes/Tris buffer.

Fourier transform infrared spectroscopy (FTIR)

Attenuated total reflection (ATR) infrared spectra were recordedon a Bruker IFS-50 infrared spectrophotometer equipped with aliquid nitrogen cooled MCT detector. The internal reflectionelement was a germanium ATR plate (50 £ 20 £ 2 mm) with anaperture angle of 458 yielding 25 internal reflections. Twohundred and fifty-six scans were averaged for each spectrum.Spectra were recorded at a nominal resolution of 2 cm21. Thespectrophotometer was continuously purged with air dried on asilicagel column (5 £ 130 cm) at a flow rate of 7 L´min21.

Sample preparation. Tubulovesicles were placed in 2 mmHepes/Tris buffer, pH 7.2, containing either 10 mm NaCl or10 mm KCl. (referred to below as Na+ buffer and K+ buffer,respectively) by pelleting the vesicles at 100 000 g for 30 minand resuspending the in the appropriate buffer. The operationwas repeated twice. Thin films were obtained as described byFringeli and GuÈnthard [25] by slowly evaporating 40±100 mL ofthe tubulovesicles under a N2 stream on one side of agermanium plate. This resulted in the formation of orientedmultilayers at the surface of the plate. The ATR plate was thensealed in a universal sample holder.

Deconvolution. Fourier deconvolution was carried out accordingto Kauppinen et al. [26±28] with a 30 cm21 full-width at half-height Lorentzian function and a 15 cm21 full-width at half-height Gaussian function for the apodization. Secondarystructure was estimated by fitting the amide I band as previouslydescribed [29,30].

Linear polarization. The potential energy distribution of amide Iconsists of about 80% n(C = O), 10% n(C ± N) and 10%d(N ± H) (reviewed in [31]). In an a-helix, the main transitiondipole moment lies approximately parallel to the helical axis. Apreferential orientation of the a-helix with respect to themembrane plane is therefore suggested by linear polarization ofamide I. Dichroic spectra were obtained by subtraction of thespectrum recorded with parallel (') polarization from thespectrum recorded with perpendicular (//) polarization. Thedifference (dichroic) spectrum is computed as (// ± ')´c wherec is a coefficient which takes into account the difference in theevanescent field amplitude in the film. In the present experi-mental conditions, c = 1.50 (reviewed in [32]).

Raman spectroscopy

Samples for Raman spectroscopy were transferred into glasscapillary tubes (1.8 mm inside) and placed in a thermoelec-trically regulated sample holder at 10 8C. Raman spectra wereobtained with a Spex Model 1400 double monochromator. Amultichannel CCD9000 system from Photometric Ltd. (Tucson,AZ, USA) with a EEV CCD-05-X detector composed of1152 £ 298 pixels, was used as described elsewhere [33].Spectra were composed of 1152 data points covering a spectralregion of <500 cm21 recorded with diffraction gratings of 1200grooves per mm. The Raman spectra were excited with the514.5 nm line (300 mW at the sample) of an argon ion laser(Spectra Physics Model 2020). Spectra reported in this paperwere obtained from the average of 30 exposures of 50 s. Thebaseline obtained for the pure buffer was subtracted and theresulting spectra were smoothed with an 11-point Savitsky-Golay smooth.

Circular dichroism

Tubulovesicle suspension (protein 0.55 mg´mL21) was usedwithout further preparation for a `solution' CD spectrum in a0.02-cm cuvette on a Jasco-710 spectropolarimeter at roomtemperature. Twenty scans recorded in the 180±260 nm regionwith a scan speed of 50 nm´min21, a step resolution of 0.2 nmand a time constant of 0.125 s were averaged [34]. Atubulovesicle lipid extract was used for the baseline. Nosignificant intensity appeared in the spectra of the lipid extract(data not shown) ruling out differential scattering artifacts.

178 V. Raussens et al. (Eur. J. Biochem. 262) q FEBS 1999

RESULTS

ATPase activity (mean values ^ SD from four experiments)expressed in mmol´h21´mg21 protein at 37 8C was 36 ^ 7 inisotonic conditions; 111 ^ 30 after addition of 14 mm nigericinand 122 ^ 20 in hypotonic conditions (see Materials andmethods) in good agreement with previously published values[35].

The ATR-FTIR spectra of the H+,K+-ATPase in the presenceof Na+ or K+ buffer in the amide I region are presented inFig. 1A. The lipid ester n(C = O) is found at 1739 cm21, amideI at 1654 cm21, amide II at 1546 cm21 and d(CH2) at1467 cm21, mainly arising from the lipid component. ATR-FTIR spectra were recorded on films prepared from tubulove-sicles in suspension in either the Na+ or the K+ buffer. In a firststep, the reproducibility of the ATR spectra was established.Several films of the same sample in the K+ buffer were preparedand their ATR spectra recorded.

In order to evaluate the intrinsic reproducibility of theexperiment, all possible differences between six such spectrawere computed. They are reported in Fig. 2A. Before subtrac-tion, the spectra were rescaled to a constant amide I areacomputed between 1711 and 1590 cm21 and a baseline goingthrough these two points was also subtracted. Expressed as apercentage of the total amide I intensity, the large majority ofthese difference spectra have a relative intensity varyingbetween 1 and 5% of the signal intensity. The reasons for thisrather poor reproducibility are discussed below. Similarly, allpossible differences spectra were computed between spectrarecorded in the presence of Na+ and K+ ions. Spectra arereported in Fig. 2B. Comparison of Fig. 2A and Fig. 2Bindicates that no significant feature related to the E1±E2conformation change of the ATPase can be revealed within thereproducibility limits of this type of experiment.

In order to improve the sensitivity to potential secondary-structure changes which should affect amide I shape, decon-volved spectra were obtained by Fourier deconvolution (seeMaterials and methods) and are presented in Fig. 1B. Amide Ishows three distinct components: the 1655 cm21 main band canbe assigned to a-helix and the 1639 and 1680 (shoulder) cm21

to b-sheet or extended structures. In amide II, the maximum is

Fig. 1. ATR-FTIR spectra of 50 mg of

tubulovesicles. (A) Spectra performed in Na+

buffer (curve a) and K+ buffer (curve b). The

amide I intensity is 0.18 absorbance units. (B)

Same spectra after Fourier deconvolution with a

Lorentzian lineshape (FWHH = 30 cm21) and

apodization with a Gaussian lineshape

(FWHH = 15 cm21) as described by

Kauppinen et al. [26±28].

Fig. 2. All possible difference spectra obtained for six films prepared in

the presence of K+ (Panel A) and obtained by subtracting spectra of

films prepared in the presence of Na+ from spectra of films prepared in

the presence of K+ (Panel B). The averaged amide I intensity of the spectra

before rescaling was 0.2 absorbance units.


found at 1547 cm21 and the tyrosine-ring vibration at1516 cm21 is now clearly distinct. Evidently, the proteinspectral features revealed by the deconvolution process areidentical for the Na+ and K+ forms. Determination of thesecondary-structure content by curve fitting (see Materials andmethods) yielded the same a-helical content (33%) within 1%for the Na+ and K+ forms of the enzyme. The b-sheet contentwas found to be 31.5% within 1.5% for the Na+ and K+ forms, ingood agreement with our previous estimates [7]. One advantage

of ATR-FTIR is that it allows the determination of theorientation of secondary structures (reviewed in [25,32]).Dichroic difference spectra were computed for both the Na+

and K+ forms of the enzyme (Fig. 3b,c, respectively) bysubtracting the spectrum recorded with the perpendicularpolarization from the spectrum with the parallel polarization.In the absence of preferential orientation, the dichroic spectrumis expected to be flat. Figure 3b,c reports a positive deviation ofthe dichroic spectrum in the amide I region. This deviation issimilar in intensity and frequency for both the Na+ and K+

conditions, demonstrating that no major reorientation of theprotein secondary structures occurred upon exchanging the Na+

salt by a K+ salt before film formation. Interestingly, thefrequency maximum occurs at 1658 cm21 (i.e. in the a-helixfrequency range of amide I [36]), strongly suggesting thepresence of transmembrane helices. No particular dichroismseems to be associated to the 1639 cm21 shoulder visible on thespectrum (Figs 1 and 3a), suggesting no particular orientationfor the associated secondary structure (b-strand).

One concern in these experiments was the H+,K+-ATPasestability in the course of the experiments. Recovery of more than70% of the original activity after resuspending the film seems tobe satisfactory. Indeed, the possible presence of multilamellarvesicles or sealed vesicles with the ATP site inside the vesicles islikely to account for some of the lost activity.

Raman spectra of tubulovesicles were recorded in order toprobe the secondary structure and the amino-acid side-chainenvironment in solution. Intensity reproducibility from experi-ment to experiment for the Raman spectra was found to be in therange of 5%. Raman spectra of the H+, K+-ATPase in the Na+

and K+ buffer are reported in Fig. 4. Both spectra are similar inthe amide I region, which is characterized by a maximum at1658 cm21 assigned to a-helix structure and a marked shoulderat 1668 cm21 assigned to b-sheet structure [37]. These resultsconfirm those obtained by ATR-FTIR spectroscopy and indicatethat, even in solution, the structure of the protein in Na+ and K+

buffer is quite similar. When analyzed in the 1000±500 cm21

region, most features characteristic of the side-chain environ-ment were unchanged upon replacing the Na+ buffer by the K+

Fig. 3. Dichroic difference ATR-FTIR spectra in the amide I and amide

II region for a film of tubulovesicles. Curve (a) is the original, unpolarized,

spectrum recorded in the presence of the Na+ buffer; (b) difference spectrum

in Na+ buffer; and (c) difference spectrum in K+ buffer. Dichroic spectra

intensity has been expanded twofold with respect to the original spectrum.

Fig. 4. Raman spectrum in the

1350±1800 cm21 range of tubulovesicles in

(a) Na+ buffer and (b) K+ buffer.


buffer (Fig. 5), indicating that the environment of the amino-acid side chains is similar. Phe is characterized by the1003 cm21 band. The value of the I622/I642 ratio, whichcharacterizes the Phe/Tyr ratio, is 1.05 ^ 0.02 independent ofthe buffer used. The Raman doublet near 850 and 830 cm21

arises from a Fermi resonance between ring-breathing vibrationand the overtone of an out-of-plane bending vibration of thepara-substituted benzene ring of the tyrosine residues [38]. Theratio between the 849 cm21 band (exposed tyrosine) and831 cm21 (buried tyrosine) is constant at 0.94 ^ 0.02, indicat-ing that Tyr exposure to the solvent is not significantly modifiedupon shifting from E1 to E2 conformation. Ratios I849/I831 areusually below 1 for buried Tyr and above 1 for exposed Tyr [37].Using the empirical equation proposed by Yu [39] to quantifythe number of buried and exposed Tyr residues, it can becalculated that 27 out of 47 Tyr residues are exposed on thesurface of the protein. Consistent with this, the contribution of

buried Tyr at 875 cm21 is unchanged upon buffer exchange. Theenvironmental conditions experienced by the indol side chainof Trp residues can be correlated with the sharp bandpresent at 1360 cm21. Its presence indicates the presence ofburied Trp residues [39]. Clearly, the buried Trp contributionat 1360 cm21 is not affected by replacing Na+ buffer by K+

buffer (not shown). The major lipid contribution in thisspectral region appears at 717 cm21 due to ns(C ± N) in thecholine group [40].

Tubulovesicle CD spectra were recorded in the presence ofeither Na+ or K+ buffer between 180 and 260 nm. Intensityreproducibility from experiment-to-experiment for CD spectrawas <1% above 200 nm and 3% below 200 nm. Figure 6presents the UV CD spectra recorded in the presence of Na+ orK+ buffer. As for ATR-FTIR and Raman spectroscopy, nosignificant change in the spectrum is visible within the noiselevel associated with this type of experiment.

Fig. 5. Raman spectrum in the 550±1050 cm21

range of tubulovesicles in (a) Na+ buffer and

(b) K+ buffer.

Fig. 6. UV-CD spectrum in the 180±260 nm

range of tubulovesicles (0.55 mg protein´mL21)

suspended in (a) Na+ buffer and (b) K+ buffer.


DISCUSSION

ATR-FTIR on films, and solution UV CD and Raman spectrareported here indicate that conformational changes at the levelof the protein secondary structure (ATR-FTIR and UV CD) andside-chain environment (Raman) are not detectable within thelimits imposed by the experiment reproducibility (1±5% of totalsignal). Nevertheless, these experiments test a broad range ofpotential changes including secondary-structure changes, orien-tation of secondary structures and Tyr or Trp side-chainenvironment, which can not be tested with a quantitative aspectby other means. The failure to detect any significant modifi-cation upon shifting from E1 to E2 conformation indicatesclearly that no major structural change is involved. It isimportant to define here the limits from which a structuralchange can be detected, taking into account the reproducibilitylevel associated with a method. Because the protein studied islarge (1324 amino-acid residues), changes affecting only a fewresidues are not expected to be detected in these experimentalconditions.

Reproducibility was estimated by subtracting spectra obtainedwith different samples (prepared under identical conditions) andtaking the ratio of largest positive or negative deviations in thedifference spectrum to the original spectrum intensity at thesame frequency. Before subtraction, infrared spectra wererescaled to the same amide I area and to the same 1003 cm21

Phe band for Raman spectra. Rescaling was not necessary forCD spectra since concentration and optical length of the cell areeasy to control. The largest difference observed were 5% forATR-FTIR and Raman spectra and 3% for CD spectra.Assuming a reproducibility level of 5%, it could be evaluatedthat <66 amino-acid residues (5% of 1324) should change theirstructure to yield a signal above the reproducibility level.However, two antagonist effects on the sensitivity preclude anaccurate determination of the detection threshold for a structuralchange.

Firstly, suggesting that 5% reproducibility results in adetection threshold just larger than 66 amino-acid residuessuppose an all-or-none effect with no overlap of the bandspresent before and after the structural change. Such an effectwill be found for example in the infrared spectrum uponprotonation of a carboxylic acid (disappearance of the n(COO2)band near 1580 cm21 and appearance of the n(COOH) bandnear 1720 cm21). As far as secondary-structure changesare concerned, structural changes usually result in smallband shifts. The expected spectral change is then only afraction of the total intensity in the infrared, Raman orCD spectrum, thereby making the detection thresholdlarger. In order to illustrate and investigate the impor-tance of this effect, we computed the intensity of adifference spectrum obtained when an absorbance bandshifts, as expected in the amide I frequency range, uponstructural change. The bands were taken to be Lorentzianin shape in order to obtain an analytical solution to theproblem. The difference spectrum diff(n) obtained bysubtracting a band of full-width at half-height (FWHH)and shifted by Dn cm21 upon conformational change isdescribed by:

diff�n� �FWHH2

FWHH2 1 4�n 2 Dn�22

FWHH2

FWHH2 1 4n2�1�

In order to determine the frequency of the maxima inthe difference spectrum, the derivative of this expression

was set equal to 0 and solved for n, yielding tworeal solutions:

vmax;min �Dn

2^

��3

p6

£��Dn 2 2 FWHH2 1 2

��FWHH4 1 FWHH2´Dn2 1 Dn4

pq�2�

Introducing the frequency of the maximum nmax in Eqn (1)yields an expression which describes how the intensity of themaximum of the difference spectrum depends on FWHH andDn. The described band shapes for a-helices and b-sheetstructures in the amide I range have a FWHH = 16±32 cm21 andunordered structures have a FWHH of 50±60 cm21 (reviewed in[36]). The maximum amplitude of the difference spectrumrelative to the absorbance of the original band as theconformational change-induced band shift Dn increases from 0to 20 cm21 is reported in Fig. 7. Four values of the FWHHranging from 20 to 50 cm21 were tested. The data reported inFig. 7 indicate that in most instances the intensity of thedifference spectrum will be smaller than 50% of the originalsignal intensity. If the spectral intensity changes by only 50% ofthe total signal upon complete structural change, the detectionthreshold in our previous example would increase from 66 to132 amino-acid residues. In conclusion, the actual amplitude ofthe spectral change depends on the nature of the structuralchange (e.g. from random coil to b-sheet or from random coil toa-helix) and on the method considered (infrared, Raman, CD).Secondly, there is a scattering of the amino-acid contributions inthe spectra that means that the total intensity taken into accountat a given frequency represents less that 1324 amino-acidresidues. For instance, in the Raman spectra, the 849 and831 cm21 bands represent only the 47 Tyr present in the protein.Therefore, an uncertainty of 5% means an uncertainty on 2.4residues (5% of 47). This effect is more difficult to evaluate inthe infrared amide I region but can only improve the sensitivity.As both effects described above affect the sensitivity in oppositedirections, it can be concluded that the detection threshold canonly be evaluated approximately. It is important to mention herethat we do not present in this paper quantitative analyses of thesecondary structures because (a) such an information has beenpresented before [8, 41] and (b) structure determination isinherently much less accurate than the reproducibility of themeasurements.

It would be expected that modern instrumentation would yielda much better accuracy than the 1±5% of the signal discussedabove. For example, in the infrared spectrum, the maximumdeviation is typically smaller than 2 £ 1026 absorbance unitsaround 2200 cm21 and 2 £ 1025 absorbance units in the amideI band in the presence of the sample. With respect to the amide Iintensity around 0.2 absorbance units, the maximum deviationdue to the instrumentation is at most 0.01%, i.e. good enough tomonitor structural changes at the level of a single amino-acidresidue.

Spectral reproducibility attainable with the techniques usedabove is limited because they involve several manipulationsincluding sample preparation, cell filling or film casting, takingcells apart for washing followed by re-mounting. In addition, inthe case of films used for ATR-FTIR, differences in evenness ofthe formed film could also slightly affect the spectral shape. Forthe rather thick films used here, this effect is ,however, expectedto be small. In turn, the various handling resulted in areproducibility in the range of 1±3% of the total signal, i.e.well below the capabilities of the spectrophotometers used. Onepossibility to improve the reproducibility is to change theATPase conformation from E1 to E2 and vice versa repeatedly,


in situ, thereby working on the same ATPase-containing film butwithout handling of the measurement cell. Two approaches havebeen developed to improve sensitivity to structural changes asmonitored by infrared spectroscopy. In the first one, ligands arereleased in situ from a caged precursor [42±45]. This techniqueis amenable for solution FTIR studies but suffers from theformation of new chemicals produced by photolysis of the cagedligand and from rapid exhaustion of the caged compound uponrepeating the experiment although, they do allow kineticmeasurements. Alternatively, ATR-FTIR experiments underbuffer flow reach a sensitivity of at least one order of magnitudebetter than the classical techniques as previously demonstratedfor thin films by Fringeli et al. [41], for thick purple membranefilms by Marrero and Rothschild [46] and for thick acetylcholinereceptor-containing films by Baenziger [47±51] . The resultsreported here clearly indicate that one of these techniques willhave to be used to gain more insight into the gastric ATPaseconformational changes.

ACKNOWLEDGEMENTS

V. R. is Senior Research Assistant of the National Fund for Scientific

Research (Belgium). E. G. is Senior Research Associate of the National

Fund for Scientific Research (Belgium).

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Fig. 7. Intensity of a difference

spectrum obtained by subtraction of two

Lorentzian bands of bandwidth FWHH but

shifted by a frequency reported in abscissa.

The intensity reported is relative to the original

spectrum intensity. Insert, difference spectra

obtained by subtraction a 20-cm21 Lorentzian

band shifted by 5, 10, 15, 20 or 40 cm21 with

respect to the original one. The change in the

intensity of the lobes of the difference spectra

are reported for a series of conditions (FWHH and

shifts) in this figure.


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Structural difference in the H+,K+-ATPase between the E1 and E2 conformations . An attenuated total...

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