J. Phys. Chem. 1995,99, 4339-4346
4339
Spectral Studies of Lanthanide Interactions with Membrane Surfaces Kerry K. Karukstis," Marie Y. Kao, Daniel A. Savin, Rachel A. Bittker, Karla J. Kaphengst, Chitoh M. Emetarom, Naomi R. Naito, and Dawn Y. Takamoto Department of Chemistry, Harvey Mudd College, Claremont, Califomia 91 711 Received: October 31, 1994; In Final Form: January 9, 1995@
We have monitored the interactions of the series of trivalent lanthanide cations with the thylakoid membrane surface of spinach chloroplasts using two complementary spectral techniques. Measurements of the fluorescence emission of the extrinsic probe 2-p-toluidinonaphthalene-6-sulfonate (TNS) and the absorbance of the intrinsic chromophore chlorophyll provide two sensitive means of characterizing the dependence of the cationmembrane interaction on the nature of the cation. In these systems, added lanthanide cations adsorb onto the membrane surface to neutralize exposed segments of membrane-embedded protein complexes. The lanthanideinduced charge neutralization increases the proximity of added TNS anion to the membrane surface as evidenced by variations in the TNS fluorescence level and wavelength of maximum emission. Our results reveal a strong dependence of TNS fluorescence parameters on both lanthanide size and total orbital angular momentum L value. Lanthanides with greater charge density (small size and/or low L value) enhance the TNS fluorescence level to a greater extent. A possible origin for the lanthanide-dependent TNS fluorescence levels is suggested in terms of a heterogeneity in the number and type of TNS binding sites. In the absence of the probe TNS, lanthanide-induced changes in the chlorophyll absorption spectrum reflect the shrinkage of chloroplasts accompanying thylakoid membrane stacking. Absorbance increases in the 500-660 nm region, attributed to increases in light scattering arising from the membrane structural reorganization, reveal a dependence on lanthanide identity. The data are consistent with the proposal that larger lanthanides with smaller enthalpies of hydration induce more significant membrane appression. These investigations illustrate a novel utilization of lanthanides in cation binding studies by employing their chemical and physical differences rather than their similarities in luminescence properties
Introduction Lanthanide cations offer an effective means of probing the role of metal cation size in the structural organization of biological membranes. The monotonic variation in effective hydrated ionic radii provided by the series of trivalent lanthanides affords a systematic manner of altering ionic size and potentially influencing the extent of cation-membrane interactions. In the thylakoid membranes of plant chloroplasts, trivalent lanthanide cations adsorb to the surface-exposed carboxyl groups of glutamic acid and aspartic acid residues of protein complexes imbedded within the thylakoid membrane.'-4 Charge neutralization of the membrane surface induces the appression or stacking of thylakoid membrane@ by reducing the thylakoid surface charge density.jq7 To monitor the interaction of lanthanides with the surface of thylakoid membranes in plant chloroplasts, we have used two techniques: measurements of the fluorescence from the molecular probe 2-ptoluidinonaphthalene-6-sulfonate(TNS) and measurements of lanthanide-induced chlorophyll absorption changes. The unique spectroscopic properties of luminescent trivalent lanthanide ions have been widely used to characrterize sites of cation binding in chemical and biochemical systems. However, previous researcher8 have demonstrated that no suitable amino acid chromophores exist on the thylakoid membrane surface to sensitize lanthanide emission indirectly via excitation transfer. Furthermore, direct lanthanide excitation is complicated by the broad and overlapping absorption band of the chlorophyll pigments in plant chloroplasts which induces lanthanide fluorescence via both the singlet and triplet states of chl~rophyll.~-"
* To whom correspondence should be addressed. @
Abstract published in Advance ACS Absrracts, March 1, 1995.
The extrinsic fluorophore TNS, however, is an ideal probe of the binding of trivalent lanthanides to the thylakoid membrane surface. TNS is a sensitive indicator of the polarity of its environment, emitting weakly in water but exhibiting intense fluorescence upon binding to a protein surface.I2-l5 A blue shift in the wavelength of maximum emission accompanies the enhancement of TNS fluorescence. As a monovalent anion, TNS fluoresces weakly in unstacked chloroplasts where negatively-charged membrane surfaces repel the anion to more aqueous regions. TNS fluorescence levels are significantly enhanced when cations are added to suppress the electrostatic repulsion between TNS and carboxylate groups on membrane surfaces, permitting the polycyclic aromatic TNS to approach the hydrophobic membrane region and bind at the polar-nonpolar interface in the lipid bilayer.I6-l8 The changes observed in TNS fluorescence intensity and emission Amax reflect the alterations in the microenvironment of the TNS probe. Early studiest9 of the interaction of TNS with chloroplast thylakoid membranes concluded that the variations in the TNS fluorescence intensity were governed principally by the cation charge (C3+ > C2+ > C+) and not by the chemical nature of the cations within a valency group. However, the data to make this assessment were severely limited, especially in the case of trivalent cations where only a single example, tris(ethy1enediamino)cobalt3+, was examined. An investigation into the effect of ionic size on lanthanide cation binding is warranted, for the affinity of trivalent lanthanide ions to selected proteins and enzymes has been shown to exhibit a strong dependence on ionic size.20-23 Conflicting observations of both higher binding constants by lanthanide ions with smaller ionic and stronger binding by the lanthanides at the start of the series23 have been reported. Accordingly, this study addresses the
0022-36541992099-4339$09.00/0 0 1995 American Chemical Society
4340 J. Phys. Chem., Vol. 99, No. 12, 1995
Karukstis et al.
sensitivity of TNS fluorescence enhancement in lanthanidethe first-reported variations of the chlorophyll absorption spectrum with cation size within a valency group. treated chloroplasts to cation size. Furthermore, linear correlations of lanthanide properties with lanthanide total orbitai angular momentum L quantum number have been observed, Experimental Section particularly for the formation constants of complexes of Sample Preparation. Chloroplasts were isolated from lanthanides with organic ligands such as acetate and EDTA.24 market spinach in a buffer medium containing 0.4 M sucrose/ A nonlinear yet periodic change in L values occurs with 50 mM HEPES-NaOH (pH 7.5)/10 mM NaC1. Centrifugation lanthanide atomic number, dividing the lanthanide series into at 6000g for 10 min at 4 OC using a Sorvall RC2-B high-speed four segments and producing an inclined-W patternz5 for L vs centrifuge was followed by resuspension of the pellet in a atomic number. The linear correlation of a property with medium of 0.1 M sucrose/lO mM HEPES-NaOH (pH 7.5)/10 lanthanide L quantum number is evidenced by the appearance mM NaCl. A second centrifugation was conducted at 6000g of such an inclined W in the plot of that property against L for 10 min, and the resulting pellet was resuspended in a final value. Thus, we will also attempt correlations of TNS fluomedium of 0.1 M sucrose/lO mM HEPES-NaOH (pH 7.5)/5 rescence parameters with lanthanide L values. mM NaCl to give approximately 1 mg of chlorophyll (Chl) per As a second means of monitoring lanthanide-chloroplast milliliter. For fluorescence and absorbance measurements the interactions, we have observed lanthanide-induced changes in chloroplasts were diluted with additional buffer and lanthanide the chlorophyll absorption spectrum of spinach chloroplasts. The stock solutions to yield [Chl] = 10 p g mL-'. All lanthanides light-harvesting pigment-protein complexes of higher plants (except promethium) were obtained (Aldrich) as the trivalent commonly contain two distinct forms of chlorophyll-chlorophyll chloride salts. Solutions were prepared to vary [Ln3+] up to a (Chl a), present in higher concentration, and chlorophyll b 1200 pM. For all lanthanides studied, concentration effects on (Chl b). Spinach chloroplasts thus exhibit an absorption fluorescence and absorbance reached a maximum level at or spectrum that is a composite of the absorption bands of Chl a before this concentration upper limit. and Chl b. The room-temperature chloroplast spectrum is TNS Fluorescence Measurements. TNS (Aldrich) was characterized by strong Soret bands which appear with maxima obtained as the potassium salt, and a stock solution of the probe at approximately 430 nm (Chl a) and 470 nm (Chl b), a was prepared in dimethyl sulfoxide (DMSO). Chloroplast minimum between 530 and 560 nm, and peaks at 650 nm for samples contained 50 pM TNS for optimal fluorescence signals. Chl b and between 670 and 700 nm for Chl a.26 The concentration of monovalent K+ counterion present in TNS is insufficient to induce any cation effects on chloroplast The in vivo absorption spectrum of chlorophyll pigments is membrane ~rganization.~ A 5-min incubation of chloroplasts significantly influenced by their environment. As Chl a and b and cation followed by a 1-min incubation of TNS occurred molecules are noncovalently bound to apoproteins in the prior to fluorescence measurement. These incubation times were thylakoid membrane, the variable spectral properties of chlorodetermined to be sufficient to produce time-independent fluophyll in vivo reflect such phenomena as pigment aggregation, rescence values, indicating that cation-induced effects on protein-chromophore interactions, and membrane conformafluorescence are complete prior to spectroscopic measurements. tional change^.^' Thus, the sensitivity of chlorophyll absorption Emission spectra were recorded at 25 "C with a Perkin-Elmer to major structural changes in chloroplast organization provides LS-5 fluorescence spectrophotometer using Aexcitatlon = 320 nm a probe of chlorophyll environment. In particular, variations and Aemission = 350-550 nm. These fluorescence parameters in the chloroplast absorption spectrum have been observed to were established using guidelines from previous studies.I9 For occur upon the addition of monovalent and divalent cations to a fixed [TNS], the TNS fluorescence intensity is dependent upon the chloroplast suspending m e d i ~ m . ~ ~ -The ~ O similar effects the chlorophyll concentration of the chloroplasts. We observed of 100 mM NaC1,28100 mM KCl,29 1 mM C a c l ~and , ~ 5~ mM (results not shown) that, both in the absence of lanthanides and MgC1z30 on the absorbance intensity in various regions of the in the presence of high [Ln3+l (Le., 1200 pM), the TNS chloroplast absorption spectrum have been attributed to two fluorescence initially increases as [Chl] increases, reaching a factors accompanying the shrinkage of chloroplasts with memmaximum at [Chl] = 8-12 p g mL-I, and then decreases at brane appression. Increases in the absorption between 500 and higher [Chl] due to the absorption of TNS emission by 660 nm suggest an increase in turbidity or light scattering chlorophyll pigments. These results match those reported for resulting from the macrostructural membrane changes upon identical studies in spinach chloroplasts with and without higher grana formation. Decreases in the Soret region between 430 concentrations of monovalent cation (Le., 100 mM K+).I9 We and 475 nm and in the peak at 675 nm indicate a mutual shading chose to examine lanthanide effects on chloroplasts with [Chl] of chlorophyll pigments within the stacked membranes. Duy= 10 p g mL-' where TNS fluorescence is maximal. sens3' interpreted this flattening effect to arise as the projected Trilinear Analyses of TNS Emission. Because of the area of the chloroplasts decreases upon grana formation. The sensitivity of the emission of TNS to its environment, we flattening effect is maximal at the wavelengths of maximum analyzed the fluorescence from TNS further, employing a absorption and increases (for fixed chlorophyll concentrations) technique of a three-mode factor analysis developed by Leurgans as the chlorophyll chromophores are concentrated in more and Ross.36 The trilinear model enables the resolution of distinct closely stacked grana and therefore smaller chloroplast^.^^^^^ This yet overlapping excitation and emission spectra for each absorption reduction arising from the mutual screening of fluorescing component in a mixture of fluorophores. The basis pigmented molecules within particles (e.g., cells or chloroplasts) for this technique is that the overall fluorescence intensity of a was altematively termed "the sieve effect" by R a b i n ~ w i t c h ~ ~ . ~mixture ~ of fluorophores is separately linear in functions of in independent studies of Chlorella suspensions. excitation wavelength, emission wavelength, and any parameter Our examination of lanthanide-induced effects on the chlothat alters fluorescence quantum yield or fluorophore concentraroplast absorption spectrum seeks to reveal lanthanide-induced tion. In mathematical terms, trilinear analysis decomposes a absorption changes that qualitatively correspond to the variations three-way array into a sum of products of parameters using observed for monovalent and divalent cations and that demonnonlinear least-squares techniques. The computational methods were used to separate the emission of bound and free TNS. strate a correlation with lanthanide size. Our results constitute
J. Phys. Chem., Vol. 99, No. 12, 1995 4341
Lanthanide Interactions with Membrane Surfaces Data for trilinear analysis were acquired by varying excitation wavelengths in 10-nm increments from 280 to 360 nm and recording emission spectra over the range from 380 to 540 nm. From the recorded emission spectra at the nine fixed excitation wavelengths, fluorescence intensities were determined at 18 wavelengths over the range from 380 to 540 nm. Data were organized into a matrix of 9 excitation wavelengths, 18 emission wavelengths, and 5 lanthanide concentrations. Nonlinear leastsquares fits to the trilinear model were performed on a VAX8600. Each data set was tested for the best fit to a system of 1,2,3, or 4 fluorescing components. Excitation and emission spectra for both bound and free TNS probe were obtained, including the relative contributions of each to the overall spectrum. Absorbance Measurements. Absorption spectra were recorded at 25 "C with a double-beam, ratio-recording PerkinElmer Lambda 6 UV-visible spectrophotometer over a range of 350-800 nm. A 5-min incubation of chloroplasts and cation occurred prior to absorption measurement. This incubation time was determined to be sufficient to produce time-independent absorbance values, indicating that cation-induced effects on absorbance are complete prior to spectroscopic measurements. For comparisons to literature data, chloroplasts were also suspended in buffer containing 5 mM MgCl2. Determination of Amount of Bound TNS. An evaluation of the amount of free and bound probe can be accomplished by separating a membrane from its suspension medium via centrif~gation.~~ To determine the amount of TNS bound to chloroplast thylakoid membranes, chloroplasts at 10 pg mL-' and [Ln3+]= 1200 pM were incubated in the dark with 50 pM TNS for 5 min. The thylakoid membranes were then pelleted by centrifugation at 32 OOOg for 1 h. Unbound TNS remained in the supematant. The volume of the decanted supematant was determined, and the absorbance spectrum was recorded. The concentration of unbound probe and the number of moles of free and bound probe were then calculated. While the percentage of bound TNS in chloroplasts with no added Ln3+ was constant for samples prepared on the same day (3~0.4%of added TNS), day-to-day variations were observed (percent TNS bound = 15.7 k 5.7%). To eliminate sample variations, the increase in the percentage of TNS bound upon Ln3+ addition was calculated. Measurement of TNS Binding Constants. The TNS binding constant was quantified by performing a titration in which [TNS] is varied (10-50 yM) for a fixed amount of chloroplast ([Chl] = 10 p g m L - I ) and Ln3+ (1200 pM). A double-reciprocal plot of TNS fluorescence as a function of [TNS] gives a straight line for a single binding site defined by the equation
where K d is the average apparent dissociation constant for the TNS-membrane interaction and F is the fluorescence intensity at A,, = 453 nm.38,39 Thus, the average apparent binding constant, K b , is the negative of the x intercept. Results TNS Fluorescence Measurements. Emission Spectra of 7TvS in Lanthanide-Treated Chloroplasts. TNS emission spectra
were recorded for spinach chloroplasts incubated with variable concentrations of all lanthanides except radioactive promethium. As an illustration of the spectra obtained, Figure 1 presents the TNS emission spectra recorded with ;lexcitation = 320 nm for spinach chloroplasts incubated with variable levels of Tm3+.
n I
380
400
420
440 460 480 500 Emlssion Wavelength / nm
520
540
Figure 1. Fluorescence emission spectra (excitation 1 = 320 nm) for TNS in spinach chloroplasts in the presence of variable levels of Tm3+. All samples contained [TNS] = 50 pM. Spectra a-g contained chloroplasts with [Chl] = 10 pg mL-'. Chloroplasts were incubated in the presence of Tm3+ at the following concentrations: (a) 1200, (b) 900, (c) 700, (d) 400, (e) 200, (0 50, and (g) 0 pM. Spectra h and i correspond to 50 pM TNS in chloroplast suspending buffer with no added chloroplasts and with either no added Tm3+ or 1200 pM Tm3+, respectively.
For all lanthanides studied, the TNS emission spectrum displayed peaks at 429 f 4, 453 f 4, and 514 k 5 nm. For comparison, Figure 1 includes the emission spectra for TNS in the presence of spinach chloroplasts without added Tm3+ and in the presence of chloroplast suspending buffer only. These latter two spectra are essentially identical with an emission A, of 520 f 2 nm. Thus, for fixed [TNS] and [Chl] levels, variations in TNS fluorescence are completely dependent upon the presence of lanthanide cations. The addition of lanthanide blue-shifts the emission maximum from 520 to 453 nm and dramatically enhances the fluorescence intensity. Both the low fluorescence intensity of TNS in untreated chloroplasts and the emission A, shift with added lanthanide enable the association of TNS with the thylakoid membrane to be reflected from increases in the fluorescence at 1 = 453 nm. Table 1 summarizes the relative fluorescence changes in these TNS emission levels. The fluorescence enhancement factor, F,,,/ Fo, is calculated with FOequal to the level in chloroplasts without added lanthanide and F,, equal to the fluorescence intensity measured at 1200 pM Ln3+. The observed fluorescence enhancement factors vary from a low of 3.7 for Pr3+to a high of 12.6 for Lu3+,but all F,,,IFo values are significantly greater than that observed for the divalent screening cation Mg2+.I9The ratio F,,/Fo generally increases with decreasing Ln3+ionic size, but periodic variations in F,,,/Fo occur with relative minima at Pr3+ and Ho3+ and maxima at La3+ % Ce3+,Tb3+,and Yb3+ x Lu3+. For a given L value, F,,,/Fo increases with decreasing ionic radius. The increase in FmaX/Fo across the lanthanide series is almost %fold for those lanthanides with L = 0, 3, and 5 (2.6, 2.7, and 2.9, respectively) but only about 1.5-fold for lanthanides with L = 6. Figure 2 illustrates both the dependence of the TNS fluorescence enhancement factor on lanthanide L value and the increase in Fmax/Foas ionic radius decreases for lanthanides of a given L value. These results are an example of an inclined-W pattern for the graph of a lanthanide property (i.e.,TNS fluorescence enhancement factor) against L Trilinear Analysis of TNS Emission. Trilinear analysis of the fluorescence emission from TNS in lanthanide-treated chloroplasts resolves the contributions of three distinct components to the overall TNS fluorescence spectrum. The predominant component, corresponding to membrane-bound TNS, has an excitation maximum of 330 f 5 nm and an emission maximum of 453 k 2 nm. Two components of minor contribution
Karukstis et al.
4342 J. Phys. Chem., Vol. 99, No. 12, 1995 TABLE 1: Observed TNS Fluorescence Enhancement Factors, Average Apparent TNS Binding Constans, and Increases in Percent TNS Bound for Lanthanide-Treated Spinach Chloroplasts lanatomic L A(% TNS thanide
no.
value
FmaJFo0
Kbb/W1
bound)'
La Ce Pr Nd
57 58 59 60 62 63 64 65 66 67 68 69 70 71
0 3 5 6 5 3 0 3 5 6 6 5 3 0
4.9 f 0.2 4.5 f 0.4 3.7 f 0.3 4.7 f 0.2 7 . 0 % 0.4 5.7 f 0.2 8.7 f 0.6 10.9 f 0.6 9.7 i 0.7 4.4 i 0.2 6.9 i 0.2 10.8 f 0.6 12.3 i 0.5 12.6 It 0.3
11.4 f 0.8 10.8 rt 2.3 14.1 rt 0.3 14.4 f 1.1 7.6 f 1.7 9.2 f 3.2 7.6 f 2.8 5.8 f 0.8 5.5 f 0.1 10.9 15.8 6.4 11.2 5.1 f 0.3 3.9 f 0.3 4.1 f 0.6
12.4 f 1.1 12.9 i 1.1 14.4 f 0.4 15.6 i 0.1 27.0 i 0.3 24.1 i 0.1 20.2 i 1.1 22.0 f 1.6 23.8 f 0.9 23.8 f 0.6 29.4 f 0.3 18.5 f 1.8 27.6 f 0.8 22.0 f 1.8
Sm Eu Gd Tb DY Ho Er Tm Yb Lu
The TNS fluorescence enhancement factor is the relative fluorescence change in the emission of TNS at 1 = 453 nm (excitation 1 = 320 nm) upon the addition of 1200 p M Ln3+ to the chloroplast suspnsion buffer. Each value reported is an average of those values obtained from six separate experiments for each Ln3+ on different days. Within a specific experiment for a given Ln3+, three measurements of F,,,,JFo were made with an average deviation of no greater than 1 0 . 2 . * The average apparent TNS binding constant describes the interaction of TNS and thylakoid membranes in spinach chloroplasts treated with 1200 p M Ln3+. Kb is calculated as the negative of the x intercept of the line described by eq 1. The increase in the percent of bound TNS was determined upon the addition of 1200 p M Ln3+ to spinach chloroplasts treated with 50 fiM TNS.
1.0 ~
z:
0.9 -
E
0.8 -
5
0.7
m
-
0.6
2
f: G .-p
-
d
.?
0.5
-
0.4 -
0.3 -
0.2 0.1 0.0
-
380
400
420
440
460
480
500
520
540
Emission Wavelength / nm
Figure 3. Normalized emission spectra for membrane-bound TNS in PP-treated (0)and Tb3+-treated (W) chloroplasts. L-3
L=5
L-6
130
a
-
20
5 y" 10
0
I
La Gd Lu Ce Eu Tb Yb
Pr
Sm Dy Tm Nd Ho Er
1
Lanthanides by L-Value
--_
Figure 4. Variation in the amount of bound TNS and in the average apparent TNS binding constant as lanthanide size decreases for those lanthanides with similar total orbital angular momentum L quantum numbers.
Tm
Er Nd
Ho 31
0
1
2
3
4
5
6
Lanthanide L Value
Figure 2. Variation in the TNS fluorescence enhancement factor at 453 nm as a function of lanthanide total angular momentum L quantum number. The appearance of an inclined-W pattem in this graph is evidence of a linear correlation of F,,,,/Fo with lanthanide L quantum number.
correspond to unbound TNS in aqueous solution (with varying orientations of the naphthalene and phenyl rings)40with Aexcltatlon = 330 nm and Aemlsslon = 520 nm and AexcltatlOn = 290 nm and AemlSSLOn= 400 nm. The shape of the emission spectrum from membrane-bound TNS varies slightly with the added lanthanide. Figure 3 illustrates the variation in the normalized emission spectrum from Pr3+- and Tb3+-treated chloroplasts, suggestive of multiple TNS environments and/or multiple TNS binding sites. Determination of Amount of Bound TNS and Measurement of TNS Binding Constants. Table 1 presents the change in the percentage of added TNS bound to the chloroplast thylakoid membrane as a consequence of the addition of 1200 pM Ln3+ (Le., percent TNS bound with Ln3+ minus percent TNS bound without Ln3+). Also listed are the average apparent TNS binding constants determined using eq 1. In general, as the effective hydrated ionic radius decreases down the series of
lanthanides, the amount of bound TNS increases and the average TNS binding constant decreases. This is consistent with the initial occupation of high-affinity binding sites and the subsequent filling of weaker binding sites as TNS is increasingly bound. No evidence of discrete binding sites (Le., multiple straight-line segments) was observed in the double-reciprocal plots of TNS fluorescence as a function of [TNS]. Figure 4 illustrates the parallel variation in the amount of bound TNS and in the average TNS binding constant as lanthanide size decreases within a lanthanide L group. Absorbance Studies. General Lanthanide Effects of Chlorophyll Absorption. To illustrate the effect of trivalent lanthanides on the absorption spectrum of spinach chloroplasts, Figure 5 presents the absorption spectra recorded in the absence and presence of 1100 pM La3+. For comparison, the absorption spectrum of chloroplasts treated with 5 mM Mg2+ is also presented. The cation-induced effects were similar for the divalent and trivalent cations, with significant increases in absorbance in the region between 450 and 650 nm and above 700 nm. The most striking difference between the two cations was in the magnitude of the absorbance increases between 525 and 650 nm-the La3+ effects were about 1.5- 1.6 times greater in magnitude than the Mg2+ increases. Figure 6 presents difference spectra determined for the cation-treated samples by subtracting the absorbance of untreated chloroplasts. To compare directly these spectra with those difference spectra previously reported for monovalent and divalent cations which were acquired using a dual-wavelength split-beam spectrophot ~ m e t e r , ~a ~reference ,~' wavelength of 492 nm should be used. Our spectra are identical in shape to previously reported
J. Phys. Chem., Vol. 99, No. 12, 1995 4343
Lanthanide Interactions with Membrane Surfaces
r
',Oo0
0.800 -
E
c
0.600 -
n 8 d
0.400
-
0200
.
300
0.000 400
500
800
700
1 0
800
200
400
800
600
1000
1200
ILas+lI pM
Wavelength / nm
Figure 5. Room-temperature absorption spectrum of spinach chloroplasts recorded in the presence of various metal cations. Spectrum a corresponds to unstacked chloroplasts suspended in a medium of 0.1 M sucrose/lO mM HEPES-NaOH (pH 7.915 mM NaCl. Spectra b and c correspond to chloroplasts suspended in the above buffer medium to which 5 mM MgC12 and 1100 p M Lac13 have been added, respectively.
Figure 7. Concentration dependence of La3+-inducedchanges in the absorption of spinach chloroplasts at 634 nm. 0.220
0.1 80 m
g
0.060 r
c
0.1 00
8 0.040
d
0.020 m
P c 8 9
-0.020 -0.020 -0.080
300
4
400
500
600
700
800
Wavelength I nm
-0.060
Figure 8. Absorbance difference spectra obtained by subtracting the absorbance spectrum of unstacked chloroplasts from the absorption spectra of chloroplasts treated with 500 p M Ln3+.
-0.1 00
300
400
500
600
700
800
Wavelength I nm
Figure 6. Absorbance difference spectra obtained by subtracting the absorbance spectrum of unstacked chloroplasts from the absorption spectra of La3+- and Mgz+-treated chloroplasts.
difference spectra for cation-induced changes, with a trough at about 430-435 nm, a shoulder at 470 nm, a peak at 525-530 nm, a second trough at 680-685 nm, and a second peak at about 700 nm. Furthermore, our difference spectra match those recorded for chloroplasts undergoing light-induced shrinkage?* Thus, the absorbance difference spectra show peaks in regions where chlorophyll absorption is generally low and indicate troughs at wavelengths where chlorophyll absorption is generally quite strong. Dependence of Lanthanide-Induced Absorption Changes at 634 nm on Lanthanide Concentration. We focused our study on the region between 500 and 660 nm where absorption changes reflect light scattering increases associated with grana f ~ r m a t i o n . ~ In ~ - particular, ~~ the cation-induced effects on chlorophyll absorption at 634 nm were examined as a function of lanthanide concentration at cation levels up to 1100pM. This wavelength corresponds to a shoulder in the chlorophyll absorption spectrum and also a region of maximal change upon cation addition. Figure 7 illustrates the concentration dependence of the La3+-induced absorbance changes at 634 nm. The concentration curve indicates a maximum increase in absorbance around 300 pM and smaller absorbance increases at higher [La3+]. We observed a similar concentration behavior for all lanthanides studied, with maximal changes occumng in the 300-500 mM region. Dependence of Lanthanide-Induced Absorption Changes on Lanthanide Identity. In light of the concentration behavior noted
TABLE 2: Absorbance Changes at 634 nm in Chloroplast Absorption Spectrum upon Addition of 500 and 1100 pM Lanthanide Ion A(absorbance) lanthanide
atomic no.
La Ce Pr Nd
57 58
Sm Eu Gd
Tb DY Ho Er Tm Yb Lu
59 60 62 63 64 65 66 67 68 69 70 71
500 p M 0.0932 f 0.0005 0.0742 f 0.0005 0.0677 f 0.0010 0.0617 f 0.0050 0.0622 f 0.0025 0.0537 f 0.0010 0.0522 f 0.0015 0.0477 f 0.0020 0.0437 f 0.0010 0.0382 f 0.0045 0.0367 f 0.0020 0.0447 f 0.0010 0.0422 f 0.0035 0.0342 f 0.0015
'
1100 p M 0.0812 f 0.0005 0.0787 f 0.0010 0.0757 f 0.0040 0.0767 f 0.0010 0.0652 f 0.0045 0.0552 f 0.0010 0.0552 f 0.0035 0.0457 f 0.0010 0.0422 f 0.0025 0.0377 f 0.0030 0.0427 f 0.0010 0.0352 f 0.0005 0.0282 f 0.0005 0.0327 f 0.0010
above, we chose to examine the dependence on lanthanide identity of cation-induced absorption changes at 634 nm for two concentration levels-500 pM (Le., at or near the concentration of maximal absorption change) and 1100 pM. The data in Table 2 correspond to the absorbance increases at 634 nm when trivalent lanthanide cations are added to the chloroplast suspending medium to yield a concentration of either 500 or 1100 p M . These data show a general decrease in the magnitude of the absorbance change as lanthanide atomic number increases. This dependence on lanthanide atomic number or essentially cation size can be illustrated graphically in a number of ways. Figure 8 presents difference spectra for spinach chloroplasts treated with 500 pM La3+, Ce3+, Pr3+, Nd3+, and Sm3+. These difference spectra illustrate a diminish-
Karukstis et al.
4344 J. Phys. Chem., Vol. 99, No. 12, 1995
events accompanying cation treatment was hypothesized. While no kinetic information is presented in our study, evidence of 0.080 both large-scale membrane conformational changes and more E localized alterations can be derived from the spectral techniques 0.070 W m selected. 5 0.060 Our results reveal a strong dependence of TNS fluorescence 0 levels on the given lanthanide. These variations correlate with 3 0.050 e both lanthanide size and L quantum number. For a given L s e ',.e 9a 0.040 value, smaller lanthanides lead to greater Fmax/Foratios. These e e, dependences reflect the fact that the effective charge density of 0.030 e'., the lanthanide is a function of both effective radius and efficiency of screening of nuclear charge. Thus, for lanthanides 56 58 60 62 64 66 68 70 72 with similar nuclear charge screening efficiencies, smaller Lanthanide Atomic Number lanthanides lead to greater charge density and higher TNS Figure 9. Absorbance change at 634 nm for chloroplasts treated with fluorescence intensities. Altematively, for lanthanides of similar 1100 pM Ln3+plotted as a function of lanthanide atomic number. size, lower L values and therefore lower nuclear charge screening lead to greater charge density on the lanthanide cation. ing effect on the chloroplast absorption spectrum as the atomic An enhanced TNS fluorescence is observed for these situations number of the lanthanide increases. Altematively, Figure 9 of low L value. presents the absorbance change at 634 nm for [Ln3+] = 1100 Our absorption studies reveal that larger lanthanide ions lead pM as a function of lanthanide number. This plot suggests a to greater increases in chlorophyll absorption in the region direct relationship between absorption changes and ionic size. between 500 and 660 nm. The lanthanide-induced absorbance Lanthanide ionic radii (for a given coordination number) increases at these wavelengths arise from light scattering decrease monotonically across the series;46 thus, absorbance increases as membranes appears. Wollmann and Dinefi4 increases in the 525-650 nm region decrease in magnitude as observed a direct proportionality between light scattering at 540 lanthanide size decreases. nm and membrane stacking as quantified directly by electron microscopy. Furthermore, a given level of light scattering Discussion corresponded to the same degree of membrane stacking, A number of photosynthetic phenomena are influenced by independent of the identity or concentration of the cation used the surface electrical properties of the thylakoid membrane.3.47 to accomplish grana formation. A similar assumption is The most direct responses to changes in membrane surface proposed in the current study-the level of lanthanide-induced charge density are variations in thylakoid membrane organizachlorophyll absorption increase in the region between 500 and tion and function. The structural differentiation of thylakoid 650 nm reflects a certain extent of membrane stacking and membranes into grana stacks interconnected by stroma-exposed chloroplast shrinkage. While a spectral selectivity of scattering thylakoids has been known for over 30 years?8 The role of can influence absorbance measurements of pigmented cell metal cations in promoting such structural differentiation has suspension^,^^-^^ our conclusions are based on changes in been surmised since the early work of Murata28949and H ~ m a n n . ~ ~ absorption spectra and on absorption differences at a specified Indeed, much is known about cation-induced electrostatic wavelength. The practice of reporting difference spectra and screening of thylakoid membra ne^^,^^ and the concomitant quantifying absorption changes at a single wavelength should For example, the extent of membrane membrane stacking. minimize the contribution of selective scattering by chloroplast stacking via electrostatic screening is predominantly dependent suspensions. Consequently, we may propose that the lanthanide on cation v a l e n ~ y , ~with ' . ~ ~divalent ions more effective in effects on chlorophyll absorption are size dependent, with larger inducing membrane appression than monovalent ions at the same lanthanides inducing a greater incidence of contact between concentration. Recent findings,61~~ however, reveal the impormembrane surfaces. tance of hydrated metal ionic size as a minor factor in This relationship of size and degree of membrane approach determining the spatial organization of thylakoid membranes suggests an influence of the strength of the enthalpies of resulting from electrostatic screening. A comparable sizehydration of the lanthanides. Adsorption of a lanthanide cation dependent phenomenon of lanthanide cation binding to thylakoid onto the thylakoid membrane surface would require the release membranes had not been investigated until this study. of water molecules, as observed upon the coordination of Our study involves complementary spectral techniques to lanthanide ions to liganding molecules or proteins.59 As the correlate cation effects with lanthanide size and L quantum enthalpies of hydration of the trivalent lanthanides increase number. Chlorophyll absorption changes, indicators of macalmost linearly across the the dehydration of trivalent roscopic structural changes, and TNS fluorescence enhancelanthanides would be thermodynamically more favorable with ments, evidence of variations in membrane microenvironments, larger size. Thus, larger lanthanides with lower enthalpies of are concurrently monitored to characterize lanthanide effects. hydration would be predicted to induce greater charge neutralA similar multivariate approach was used to examine the role ization and promote a closer approach of thylakoid membrane of monovalent (Naf) and divalent (Ca2+)cations on chloroplast surfaces. membrane structure in an earlier study.54 Parallel investigations Two possible explanations may be offered for the variation consisted of measurements of chlorophyll a fluorescence, 90" in the TNS fluorescence increases with both lanthanide size and light scattering, and extrinsic probe fluorescence using the TNS L value. A first proposal would suggest that the variations result analog 1-anilinonaphthalene-8-sulfonate (ANS). Vandermeulen from differential extents of membrane stacking, that is, variable and G ~ v i n d j e enoted ~ ~ that the kinetics of the cation-induced distances of intermembrane separation. Lanthanides with higher chlorophyll fluorescence changes were significantly slower than charge density would lead to membranes in closer proximity the time scale for the A N S probe fluorescence variations and through more extensive exclusion of water from the region 90" light-scattering changes stimulated by cation addition. A between membrane surfaces. The decrease in the polarity of mechanism involving both macroscopic and local structural O.Ogo
..,
'.\
'\
J. Phys. Chem., Vol. 99,No. 12, 1995 4345
Lanthanide Interactions with Membrane Surfaces the TNS environment following the exclusion of water and the increased orientational constraints on the membrane surface would increase the quantum yield of TNS fluorescence. Hence, the higher a lanthanide’s charge density, the greater the increase in the quantum yield of TNS fluorescence and the larger the observed enhancement of TNS fluorescence levels. However, the lanthanide-induced chlorophyll absorption changes suggest that increases in light scattering (reflecting the shrinkage of chloroplasts and decreases in the distance between membranes) correlate only with lanthanide size and not with L value. The results support the observation that lanthanides of larger size and therefore smaller enthalpies of hydration lead to smaller intermembrane distances. Thus, a simple difference in the degree of proximity between stacked membranes cannot explain TNS fluorescence variations with both lanthanide size and L value. Altematively, differences in membrane organization could potentially vary the number and type of TNS binding sites, leading to different levels of TNS fluorescence. For example, lanthanides with high charge density might give rise to greater TNS fluorescence levels as a consequence of membrane stacking that leads to additional accessible TNS binding sites. The increased binding capacity of TNS for chloroplasts treated with lanthanides of high charge density would reflect more extensive charge neutralization of the membrane surface. In essence, TNS would have access to all potential binding sites on the thylakoid membrane surface. A lowered extent of charge neutralization by lanthanides of low charge density might require the lateral separation of photosystems 1 and 2 to expel negatively charged PS 1 complexes to the stroma region. The number of binding sites for TNS would be reduced to those within the charge-neutralized region of the grana stacks. Consistent with this proposal, our measurements of the extent of TNS binding generally support an increased amount of bound TNS as lanthanide size decreases for lanthanides of similar L value. To further substantiate this hypothesis, investigations that both characterize the number and type of TNS binding sites and determine the extent of lateral segregation of the photosystems in lanthanide-treated chloroplasts would be necessary. Conclusion
Our results reveal a pronounced sensitivity of TNS fluorescence variations upon cation-membrane interaction to the nature of the cation within a valency group. The lanthanideinduced TNS fluorescence level exhibited distinct variations with two lanthanide parameters: ionic size and total orbital angular momentum L quantum number. These variations may reveal a heterogeneity in the number andor type of TNS binding sites arising from different lanthanide-induced membrane organizations. We have also observed chlorophyll absorption changes that reflect a shrinkage in chloroplast size as lanthanide-induced membrane appression occurs. These absorption changes parallel the changes previously reported for monovalent and divalent cations. Our examination of effects at 634 nm where absorption increases are sizable further reveals a lanthanide-dependent variation in the magnitude of the absorption change recorded, with larger absorbance changes as lanthanide size increases. We propose that the more thermodynamically favorable dehydration of larger lanthanides upon cation-membrane binding would yield the observed trend. These results further demonstrate a novel utility of lanthanides in cation binding studies by employing their chemical and physical differences rather than their similarities in luminescence properties. Acknowledgment. This research was supported by a National Institutes of Health Academic Research Enhancement
Award (1 R15 GM46090-Ol), by a grant from the National Science Foundation Research Experiences for Undergraduates Program (CHE-9100288), and by a grant from the Arnold and Mabel Beckman Research Fund of Harvey Mudd College. Acknowledgement is also made to the donors of the Petroleum Research Fund, administered by the American Chemical Society, for the partial support of this research. References and Notes (1) Mercer, F. V.; Hodge, A. J.; Hope, A. B.; McLean, J. D. Ausr. J. Biol. Sci. 1955, 8, 1. (2) Nakatani, H. Y.; Barber, J.; Forrester, J. A. Biochim. Biophys. Acta 1978, 504, 215. (3) Barber, J. Biochim. Biophys. Acta 1980, 594, 253. (4) Nakatani, H. Y.; Barber, J. Biochim. Biophys. Acta 1980, 591, 82. (51 Scoufflaire. C.: Lannove. R.: Barber, J. Photobiochem. Photobiophys. 1982, 4, 249. (6) Karukstis, K. K.; Gruber, S. M. Biochim. Biophys. Acta 1986,851, 722
(7) Barber, J.; Mills, J. D.; Love, A. FEBS Lett. 1977, 74, 174. ( 8 ) Mills, J. D.; Hind, G. Photochem. Photobiol. 1978, 28, 67. (9) Nieboer, E. Struct. Bonding (Berlin) 1975, 22, 1. (10) Reuben, J. Handbook on the Physics and Chemistry of Rare Earths; Gschneidner, Jr., K. A., Eyring, L., Eds.; North-Holland: Amsterdam; 1979; pp 515-552. (11) Richardson, F. S. Chem. Rev. 1982, 82, 541. (12) Weber, G.; Laurence, D. J. R. Proc. Biochem. J . 1954, 56, 31. (13) McClure, W. 0.;Edelman, G. M. Biochemistry 1966, 5, 1908. (14) McClure, W. 0.; Edelman, G. M. Biochemistry 1967, 6, 567. (15) Edelman, G. M.; McClure, W. 0. Acc. Chem. Res. 1968, 1, 65. (16) Lesslauer, W.; Cain, J. E.; Blasie, J. K. Proc. Natl. Acad. Sci. (I.S.A. 1972, 69, 1499. (17) Krishnan, K. S.; Balaram, P. Arch. Biochem. Biophys. 1976, 174, 420. (18) Cheng, S.; McQueen, H. M.; Levy, D. Arch. Biochem. Biophys. 1978, 189, 336. (19) Searle, G. F. W.; Barber, J. Biochim. Biophys. Acta 1979, 545, 508. (20) Moeller, T.; Ferms, R. Inorg. Chem. 1962, 1 , 49. (21) Epstein, M.; Levitzki, A.; Reuben, J. Biochemistry 1974, 13, 1777. (22) Gomez, J. E.; Bimbaum, E. R.; Damall, D. W. Biochemistry 1974, 13, 3745. (23) Breen, P. J.; Hild, E. K.; Horrocks, Jr., W. D. Biochemistry 1985, 24, 4991. (24) Sinha, S. P. In Structure and Bonding; Dunitz, J. D., Hemmerich, P., Ibers, J. A,, Jorgensen, C. K., Neilands, J. B., Reinen, D., Williams, R. J. P., Eds.; Springer-Verlag: Berlin, 1976; Vol. 30, pp 1-64. (25) Sinha, S. P. Helv. Chim. Acta 1975, 58, 1978. (26) Clayton, R. K. Photosynthesis: Physical Mechanisms and Chemical Patterns; Cambridge University Press: Cambridge, 1980; Chapter 2. (27) Hanson, L. K. In Chlorophylls;Scheer, H., Ed.; CRC Press: Boca Raton, FL, 1991; pp 993-1014. (28) Murata, N. Biochim. Biophys. Acta 1971, 226, 422. (29) Gross, E. L.; Libbey, J. W. Arch. Biochem. Biophys. 1972, 153, 457. (30) Henkin, B. M.; Sauer, K. Photochem. Photobiol. 1977, 26, 277. (31) Duysens, L. N. M. Biochim. Biophys. Acta 1956, 19, 1. (32) Itoh, M.; Izawa, S.; Shibata, K. Biochim. Biophys. Acta 1963, 69, 130. (33) Gross, E. L.; Packer, L. Arch. Biochem. Biophys. 1967, 121, 779. (34) Rabinowitch, E. Photosynthesis and Related Processes; Interscience Publishers: New York, 1956; Vol. 11, Part 2, Chapter 37C. (35) Das, M.; Rabinowitch, E.; Papageorgiou, G. J. Phys. Chem. 1967, 71, 3543. (36) Leurgans, S.; Ross, R. T. Star. Sci. 1992, 7, 289. (37) Azzi, A. Q. Rev. Biophys. 1975, 8, 237. (38) Brocklehurst, J. R.; Freedman, R. B.; Hancock, D. J.; Radda, G. K. Biochem. J. 1970, 116, 721. (39) Augustin, J.; Hasselbach, W. Eur. J. Biochem. 1973, 35, 114. (40) Karukstis, K. K.; Krekel, D. A,; Weinberger, D. A,; Bittker, R. A,; Naito, N. R.; Bloch, S. H. J. Phys. Chem. 1995, 99, 449. (41) Strichartz, G. R.; Chance, B. Biochim. Biophys. Acta 1972, 256, 71. (42) Yamashita, K.; Itoh, M.; Shibata, K. Biochim. Biophys. Acta 1968, 162, 610. (43) Murakami, S.; Packer, L. Arch. Biochem. Biophys. 1971, 146, 337. (44) Wollman, F.-A,; Diner, B. A. Arch. Biochem. Biophys. 1980, 202, 646. (45) Braun, G.; Malkin, S. Biochim. Biophys. Acta 1991, 1060, 303.
4346 J. Phys. Chem., Vol. 99, No. 12, 1995 (46) Greenwood, N. N.; Earnshaw, A. Chemistry of the Elements; Pergamon Press: Oxford, 1990; Chapter 30. (47) Barber, J. In The Intact Chloroplast; Barber, J., Ed.; Elsevied North-Holland: Amsterdam, 1976; pp 89- 134. (48) Menke, M. Annu. Rev. Plant Physiol. 1962, 13, 27. (49) Murata, N.; Tashiro, H.; Takamiya, A. Biochim. Biophys. Acta 1970, 197, 250. (50) Homann, P. H. Plant Physiol. 1969, 44, 932. (51) Barber, J.; Searle, G. F. W. FEBS Lett. 1978, 92, 5. (52) Mills, J. D.; Barber, J. Biophys. J . 1978, 2 1 , 257. (53) Newell, W. R.; Zara, S. J.; Barber, J. In Progress in Photosynthesis
Karukstis et al. Research; Biggins, J., Ed.; Martinus Nijhoff Publishers: Dordrecht, 1987; Vol. 11, pp 269-272. (54) Vandemeulen, D. L.; Govindjee. Biochim. Biophys. Acra 1974, 368, 6 1. (55) Latimer, P.; Rabinowitch, E. J . Chem. Phys. 1956, 24, 480. (56) Latimer, P. Science 1958, 127, 29. (57) Latimer, P.; Rabinowitch, E. Arch. Biochem. Biophys. 1959, 84, 428. (58) Latimer, P. Plant Physiol. 1959, 34, 193. (59) Kupke, D. W.; Fox,J. W. Biochemisfry 1989, 28, 4409.
JP942942U
