Complete Spectra of the Far-red Chemiluminescence of the Oxygenase Reaction of Mn2+-activated Ribulose-bisphosphate Carboxylase/Oxygenase Establish Excited Mn2+ as the Source*

Ross McC. LilleyDagger §, XueQin Wang, Elmars Krausz, and T. John Andrews||

From the Dagger  Department of Biological Sciences, University of Wollongong, Northfields Ave., Wollongong 2522, Australia, the  Research School of Chemistry, Australian National University, Canberra, ACT 0200 Australia, and the || Research School of Biological Sciences, Australian National University, P. O. Box 475, Canberra, ACT 2601, Australia

Received for publication, December 5, 2002, and in revised form, February 24, 2003

    ABSTRACT
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Chemiluminescence emitted by Mn2+-activated ribulose-1,5-bisphosphate carboxylase/oxygenase (rubisco) while catalyzing oxygenation was analyzed to clarify the source of the emission. Using dual detectors capturing radiation over a wide range of visible and infrared wavelengths, we tested for radiation from singlet O2 decay and found it to be essentially absent (less than 0.1% of the total luminescence intensity). Spectra were determined between 647 and 885 nm with a very sensitive, charge-coupled detector-based spectrograph to detect differences in the emission spectra between rubiscos from bacterial and higher plant sources. All Mn2+-activated rubiscos emitted a broad, smooth spectrum of chemiluminescence, unchanging as the reaction progressed. The spectra from higher plant rubiscos (spinach and both the wild type and an L335V mutant from tobacco), all exhibited maxima at about 800 nm. However, Mn2+-activated rubisco from the bacterium, Rhodospirillum rubrum, emitted at shorter wavelengths (760 nm peak), demonstrating host ligand-field influences arising from aminoacyl residue differences and/or conformational changes caused by the absence of small subunits. The findings provide strong evidence that the chemiluminescence arises from an excited state of the active-site Mn2+ that is produced during oxygenation. We propose that the Mn2+ becomes excited by a one-electron exchange mechanism of oxygenation that is not available to Mg2+-activated rubisco.

    INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Weak far-red chemiluminescence is emitted when Mn2+-activated rubisco1 catalyzes its oxygenase reaction (1). Neither the physiological cofactor, Mg2+, nor any other metal ion has been found to support observable luminescence. An incomplete spectrum, based on the (then) unproven assumption that the spectrum did not change as the oxygenase reaction proceeded, exhibited characteristics typical of Mn2+ luminescence (2). Cox et al. (3) investigated the effects of potential inhibitors and enhancers and found no evidence to support the suggestion (1) that singlet O2 has a role in this luminescence. However, the Mn2+-rubisco luminescence is weak, with quantum yield estimated at 10-7 to 10-9 photons per turnover (2). This leaves open the possibility that some infrequent side reaction of the oxygenase reaction of Mn2+-activated rubisco, undetectable by other means, might generate singlet O2, which contributes to the chemiluminescence directly or indirectly by a sensitization mechanism (4).

Here we perform a direct spectroscopic test for the production of singlet O2 by this system, seeking to observe the emission at 1268 nm associated with its monomolecular transition to the triplet O2 ground state. This emission is much more intense than that arising from the dimolecular process resulting from encounters between two molecules of singlet O2. The latter generates emissions at 633 and 703 nm that are detected more readily by photomultipliers. Furthermore, the 1268 nm luminescence is independent of the concentration and lifetime factors that determine the emission from such encounters. Therefore, the presence of 1268 nm emission can reveal low levels of singlet O2 in biological systems (5). Previously, enzyme-associated emission of far-red light at wavelengths longer than 1100 nm confirmed the production of singlet O2 by lactoperoxidase (6), chloroperoxidase and catalase (7). On the other hand, chemiluminescence by acetolactate synthase detectable with a photomultiplier was attributed to the dimolecular reaction (8), but the absence of detectable IR emission did not support the involvement of singlet O2 (9).

Prompted by observations that the luminescence yield appeared to differ between rubiscos from different sources (3), we here determine a more complete and less equivocal spectrum of the chemiluminescence. The spectrum reported previously for spinach rubisco (2) was subject to spectrophotometric constraints, including the need to correct for decay of enzyme activity during scanning. Here, we use a very sensitive spectrograph to accumulate the entire spectrum in a few seconds. This enables the measurement of spectral differences in the emission between rubiscos from different sources.

    EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Enzyme and Substrate Preparation-- Rubisco was purified from spinach (Spinacea oleracea L.) leaves by anion exchange chromatography (10). Rubisco from wild type tobacco (Nicotiana tabaccum L.) and its L335V mutant (11) were purified by crystallization (12). The rbcM gene from Rhodospirillum rubrum, encoding an N-terminal hexahistidine tag, was expressed in Escherichia coli and the recombinant protein was purified by nickel-nitrilotriacetic acid-agarose chromatography using the manufacturer's procedure (Qiagen). Purified rubisco was stored at -80 °C in 50 mM NaCl, 10 mM sodium phosphate, 1 mM EDTA, 10% (v/v) glycerol, pH 7.6. The total concentration of rubisco active sites was determined by [14C]2'carboxy-D-arabitinol-1,5-bisphosphate binding (13, 14). RuBP was synthesized by the procedure of Horecker et al. (15) and purified by Dowex 1-Cl chromatography (16).

Survey of Visible and IR Emission Wavelengths-- A dual-detector apparatus with a Hamamatsu R943-02 photomultiplier and an ADC 403L germanium detector was used to measure a wide range of visible and IR wavelengths (Fig. 1). The relative responses of the two detectors were established from the manufacturer's specifications and spectral measurements in the range from 600 to 1000 nm. Data for the spectral sensitivity of the germanium detector were extended beyond the range provided by the manufacturer (1200-1800 nm) to the important 700-1200 nm region by measuring a black-body source dispersed through a monochrometer. A correction for the variation in diffraction efficiency of the grating with wavelength was taken from data provided by the grating manufacturer (Richardson Grating Laboratories).

Reaction mixtures (1 ml) in a quartz cuvette were monitored simultaneously by both detectors (Fig. 1). For experiments with rubisco using H2O as solvent, the spinach enzyme (76 µM active sites) was preactivated by incubation for 30 min in 25 mM Tris-HCl or EPPS-NaOH, pH 8.1, 0.1 mM EDTA, 10 mM NaHCO3, 2 mM MnCl2 (Aldrich). The reaction mixture contained 3.1 µM active sites of activated rubisco, 25 mM Tris (adjusted to pH 8.1 with HCl), 0.1 mM EDTA, 0.4 mM NaHCO3, and 2 mM MnCl2 and the reaction was started by addition of RuBP to 390 µM. For experiments with 2H2O as solvent, spinach rubisco (3.1 µM active sites) was preactivated for 10 min in 25 mM Tris-HCl, 0.19 mM EDTA, 1.9 mM MnCl2, 3.9 mM NaHCO3, 97.9% 2H2O. The pD was 8.2, obtained by mixing calculated weights of Tris base and Tris-HCl. The reaction was started by the addition of RuBP (to 190 µM).

Lactoperoxidase (Sigma, from bovine milk, lyophilized) was dissolved (0.1 mg ml-1) in 20 mM acetic acid-NaOH, pH 4.5. The enzyme solution (1 ml) was added first to the cuvette, followed by 0.1 ml of 200 mM NaBr. After thorough mixing, 0.1 ml of 200 mM H2O2 was pipetted onto the top of the solution in the cuvette without additional mixing.

Determination of the Chemiluminescence Spectrum-- An f4 0.5m spectrograph was constructed using a back-thinned, deep-depleted, charge-coupled detector (CCD, supplied by Roper Scientific) and a low-dispersion grating blazed at 800 nm. Each spectrum (650-890 nm) was accumulated over 20 s. The spectra were calibrated and corrected for the wavelength dependence of the sensitivity of the CCD by reference to black-body radiation from a tungsten filament of known temperature (determined by an optical pyrometer, Leeds and Northrup 8627). The light from the center of the tungsten filament was attenuated uniformly by passing it through a low-duty cycle chopper before being focused on the spectrometer slit. Details about reaction mixtures and other procedures are given in the legends.

    RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

Survey of Visible and IR Emissions-- Two detectors were required to cover the visible/IR range of wavelengths of interest to this study. The sensitivity of the photomultiplier was greatest in the visible region, declining in the IR. By contrast, the germanium detector was most sensitive in the IR beyond 1200 nm, with declining sensitivity toward the visible. Using the dual-detector system (Fig. 1) without filters, luminescence was detected by both the photomultiplier and the germanium detector while rubisco was catalyzing the oxygenation of RuBP (Table I). A filter that blocked all radiation with wavelengths shorter than 1000 nm reduced the signal from the germanium detector by over 90%; a filter that blocked all wavelengths shorter than 1200 nm eliminated it entirely. The nature of the buffer used in the experiments had no effect on the intensity of the luminescence; identical results were obtained with Tris-HCl and EPPS-NaOH.


View larger version (7K):
[in this window]
[in a new window]
 
Fig. 1.   Schematic representation of the dual-detector apparatus. Total luminescence emitted during oxygenation was monitored simultaneously with a red-sensitive, high-gain photomultiplier (sensitive to wavelengths from 200 to 900 nm) and a cooled germanium detector (most sensitive to wavelengths between 1100 and 1700 nm, but with low level sensitivity to wavelengths as short as 700 nm) positioned on opposite sides of the source. The source was a 1-cm2 quartz cuvette containing enzyme reaction mixtures. Filters were interposed between the source and the detectors to select the required wavelength ranges.


                              
View this table:
[in this window]
[in a new window]
 
Table I
Photomultiplier (PM) and germanium detector outputs for a range of experimental conditions and filter sets
Spinach rubiosco was used.

When the solvent was 98% 2H2O, the signal at both detectors doubled and a filter that blocked wavelengths shorter than 1100 nm eliminated the signal from the germanium detector. Calibration with a black-body source showed that the germanium detector was approximately one-tenth (±20%) as sensitive at 800 nm (the peak of the rubisco emission, see later) as at 1200 nm. Because the minimum observable signal of the germanium detector was about 0.01 mV, more than 99% of the signal measured for rubisco chemiluminescence in 2H2O occurred at wavelengths shorter than 1100 nm (Table I). Hence radiation emitted by the rubisco samples at wavelengths longer than 1100 nm must be <0.1% of the total chemiluminescence intensity.

As a positive control, a very strong, short-lived luminescence signal was measured at wavelengths longer than 1200 nm by the germanium detector when singlet O2 was generated by lactoperoxidase. However, at the same time, no radiation was detected by the photomultiplier at wavelengths longer than 650 nm (Table I).

Spectrum of Rubisco Luminescence-- Although both detectors of the dual-detector system had significant sensitivity over the 700-850 nm range of wavelengths, the extremely low intensity of rubisco chemiluminescence and its decay with time precluded accurate determination of the spectrum with these detectors. A spectrograph based on a CCD detector was constructed for this purpose.

The output from the CCD spectrograph required correction for the wavelength dependence of the sensitivity of the CCD detector. An example of a spectrum of the luminescence from Mn2+-activated spinach rubisco before and after correction is shown in Fig. 2. The intensity of chemiluminescence for each pixel of the detector, accumulated over 20 s, is expressed as the specific intensity per nanomole of rubisco active sites present in the cuvette. The correction was greatest at the longer wavelengths, as expected, and increased the scatter of the spectral data (Fig. 2b). Corrected spectra for spinach, tobacco, and R. rubrum rubiscos are shown in Figs. 3 and 4. When the substrate RuBP was present in excess, the chemiluminescence exhibited a slow single-phase exponential decay (Fig. 4, a and b). When the reaction conditions were such that the RuBP was fully consumed eventually (Figs. 3 and 4c), this exhaustion was accompanied by a further decay in the luminescence to zero.


View larger version (25K):
[in this window]
[in a new window]
 
Fig. 2.   Emission spectrum of chemiluminescence by spinach rubisco. The abscissa is linear in terms of wave number. Spinach rubisco (77.5 µM active sites) was activated for 30 min in 25 mM Tris-HCl, 0.1 mM EDTA, 2 mM MnCl2, 10 mM NaHCO3, pH 8.1. The activated rubisco was transferred to a reaction mixture containing a solution saturated with O2 by bubbling. The final concentrations were 3.1 µM rubisco, 24 mM Tris-HCl, 0.1 mM EDTA, 2 mM MnCl2, 0.4 mM NaHCO3 and the reaction was started by the addition of RuBP (to 390 µM). Accumulation of the spectrum by the CCD apparatus (see "Experimental Procedures") commenced 5 s later and continued for 20 s. a, uncalibrated spectral data. b, data after correction for variation in the sensitivity of the CCD with wavelength. The fitted Gaussian regression curve had a mean (dotted line) of 797 nm and r2 of 0.963.


View larger version (31K):
[in this window]
[in a new window]
 
Fig. 3.   Emission spectra and time course of peak intensity (inset) of chemiluminescence by spinach rubisco in 1H2O (a) and 96.5% (v/v) 2H2O (b). Spinach rubisco (7.6 µM sites) was preincubated for 10 min in 24 mM Tris-HCl, 0.19 mM EDTA, 1.9 mM MnCl2, 3.9 mM NaHCO3 in 1H2O or 2H2O. The pH/pD was 8.2, obtained by mixing calculated weights of Tris base and Tris-HCl. The reaction was started by the addition of RuBP (to 190 µM) and accumulation of the first spectrum commenced 5 s later. The spectra were accumulated over consecutive 20 s intervals and those (from top down) for 5-25, 85-105, 125-145, and 165-185 s are shown. The Gaussian regression curves fitted to the 5-25 s spectra had means (dotted lines) and r2 of 800 nm and 0.991 (1H2O) and 799 nm and 0.996 (2H2O), respectively. The time courses (insets) show the peak chemiluminescence intensity for all spectra collected against the midpoint time during collection of each spectrum and the line represents the regression for single-phase exponential decay.


View larger version (34K):
[in this window]
[in a new window]
 
Fig. 4.   Emission spectra and time course (inset) of chemiluminescence by rubisco. a, wild type tobacco rubisco in 1H2O. Enzyme (99 µM sites) was activated for 10 min at 50 °C, cooled to 20 °C, and then maintained at that temperature for about 30 min in 25 mM Tris-HCl, 0.2 mM EDTA, 2 mM MnCl2, 10 mM NaHCO3, pH 8.1, before transfer to the assay. The reaction mixture (1.0 ml) contained 7.9 µM rubisco, 25 mM Tris-Cl, 0.1 mM EDTA, 2 mM MnCl2, 0.8 mM NaHCO3, pH 8.1, and the reaction was started by the addition of RuBP (to 770 µM). The spectra were accumulated over consecutive 20 s and those (from top down) for 5-25, 85-105, 185-205, and 585-605 s are shown. The Gaussian regression curve fitted to the 2-25 s spectrum had a mean (dotted line) of 798 nm and r2 of 0.987. The time course (inset) shows the peak chemiluminescence intensity for all spectra collected against the midpoint time for collection of each spectrum and the line represents the regression for single-phase exponential decay. b, emission spectra and time course of chemiluminescence by the L335V mutant of tobacco in 1H2O. Enzyme (110 µM) was activated as for a in 12 mM Tris-HCl, 0.1 mM EDTA, 2 mM MnCl2, 10 mM NaHCO3, pH 8.1, before transfer to the assay. The reaction mixture (1.0 ml) contained 8.5 µM rubisco, 22 mM Tris-Cl, 0.17 mM EDTA, 2 mM MnCl2, 0.8 mM NaHCO3, pH 8.1, and the oxygenase reaction was started by the addition of RuBP (to 770 µM). The spectra were accumulated over consecutive 20 s and those (from top down) for 5-25, 65-85, and 85-305 s, are shown. The Gaussian regression curve fitted to the 5-25 s spectrum had a mean (dotted line) of 800 nm and r2 of 0.988. The time course (inset) shows the peak chemiluminescence intensity for all spectra collected against the midpoint time for collection of each spectrum. c, emission spectra and time course of chemiluminescence by R. rubrum rubisco in 1H2O. Enzyme (6.2 µM in 500 µl) was dialyzed against 25 mM Tris-HCl, 0.1 mM EDTA, 2 mM dithiothreitol, pH 8.1. Dialyzed enzyme was preincubated for 30 min in 23 mM Tris-HCl, 0.1 mM EDTA, 1.8 mM dithiothreitol, 2 mM MnCl2, 40 mM NaHCO3, pH 8.1. The reaction mixture (1.0 ml) contained 2.7 µM rubisco, 24 mM Tris-HCl, 0.1 mM EDTA, 1.1 mM dithiothreitol, 2 mM MnCl2, 22 mM NaHCO3, pH 8.1, and the oxygenase reaction was started by the addition of RuBP (to 770 µM). The spectra were accumulated over consecutive 20 s and those (from top down) for 2-25 and 45-65 s are shown. The Gaussian regression curve fitted to the 5-25-s spectrum had a mean (dotted line) of 760 nm and r2 of 0.870. The time course (inset) shows the peak chemiluminescence intensity for all spectra collected against the midpoint time for collection of each spectrum.

The luminescence spectrum is modeled closely, but not quite perfectly (r2 values in figure legends), by a Gaussian curve, symmetrical in terms of wave number. The fitted line is shown for the first and most intense corrected spectrum accumulated between 5 and 25 s after addition of RuBP to the reaction mixtures (Figs. 2-4). With time, the luminescence diminished in intensity but the spectrum was unchanged.

The wavelength of peak emission, estimated from the fitted curves for spinach rubisco in H2O, was 797 nm (Fig. 2b) or 800 nm (Fig. 3a). A similar spectrum, peaking at 799 nm, was observed for spinach rubisco in 2H2O (Fig. 3b), but the intensity approximately doubled. The chemiluminescence decayed to zero when the RuBP had been consumed completely. This took about 170 s for rubisco in H2O and about 200 s in 2H2O (Fig. 3), showing that the total rubisco activity (carboxylase plus oxygenase) was slightly lower in 2H2O. Rubisco from wild type tobacco (Fig. 4a) exhibited a chemiluminescence spectrum indistinguishable in profile from that of spinach rubisco, with peak emission at 798 nm. The peak wavelength of the emission from the L335V mutant of tobacco rubisco was similar (800 nm), but the intensity was approximately halved (Fig. 4b).

The spectrum of the chemiluminescence emitted by rubisco from R. rubrum (Fig. 4c) was blue-shifted compared with that of the enzymes from higher plants, with peak emission at 760 nm. The specific intensity per nanomole of active sites was also smaller.

Photoluminescence Could Not Be Induced by Laser Excitation-- We attempted to excite photoluminescence from Mn2+-activated rubisco by laser excitation. Some weak, variable emission was indeed seen, as well as some Raman features. The emission could not be identified clearly with the chemiluminescence measured. Although previous attempts to measure Mn2+ emission using aqueous inorganic solutions of Mn(II) have failed (17), we have observed a number of examples of laser-excited photoluminescence from aqueous Mn2+ with the same apparatus used in this study.2 However, the process is remarkably feeble, requiring very high (5 M) managanese concentrations. The Mn2+ concentrations in our rubisco samples were far lower (up to 2 mM active-site concentrations, requiring purified rubisco at up to 140 mg ml-1), precluding direct photoexcitation of measurable photoluminescence. Laser excitation of other chromophoric units in the protein does not lead to efficient electronic excitation transfer to the manganese. Consequently, the luminescence spectrum is dominated by other (weak) emitters.

    DISCUSSION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

The Spectral Data Eliminate Singlet O2 as a Possible Source of Luminescence-- The germanium detector of the two-detector, visible/IR measurement system (Fig. 1) detected intense luminescence at wavelengths longer than 1200 nm when the lactoperoxidase reaction, a known producer of singlet O2, was used (Table I). This luminescence is attributable to the monomolecular decay of singlet O2 (1268 nm) (7). On the other hand, the photomultiplier, whose sensitivity extends from visible wavelengths into the IR to ~900 nm, detected no emission from the same system. Therefore, we conclude that luminescence caused by the dimolecular dismutation of singlet O2 pairs (expected to show peaks at 634 and 703 nm (4)) is negligible in this system, even when the monomolecular reaction is readily detectable. The germanium detector detected no luminescence from manganese-rubisco at wavelengths longer than 1100 nm. Even in 98% 2H2O, which greatly enhances chemiluminescence from enzyme-generated singlet O2 (19) no emission from manganese-rubisco was detected at wavelengths longer than 1100 nm (<0.1% of that detected by the photomultiplier at shorter wavelengths). Therefore, the emission from manganese-rubisco detected by the photomultiplier cannot originate from the dimolecular singlet-O2 reaction. Consistent with this conclusion, the spectra of the Mn2+-rubisco luminescence recorded by the CCD detector showed no sign of any features at 634 and 703 nm, even in 98% 2H2O (Fig. 3).

The Luminescence Is Characteristic of Octahedral Mn2+-- The broad smooth spectra of the luminescence from Mn2+-activated rubisco resembles the emission from six-coordinate Mn2+ compounds. Manganese luminescence exhibits spectra with peak wavelengths depending strongly on the local environment of the metal ion and the resultant crystal (ligand) fields (20). Non-enzymic reactions involving an excited Mn2+ species show such broad emission spectra. In a set of inorganic reactions involving Mn3+ reduction to Mn2+, the peak emission was at either 689 or 730 nm (17), whereas luminescence from the oxidation of glyoxal by Mn(III)-lactate chelate peaked at about 710 nm (21). These systems also lacked any emission attributable to singlet O2.

An alternative conceivable source of the rubisco chemiluminescence that must be considered is the possible involvement in the oxgenase reaction mechanism of a dioxetane intermediate involving carbons 2 and 3 of the RuBP substrate. Although such a mechanism was excluded for the main, Mg2+-dependent oxygenase pathway by 18O2 labeling studies (22), the possibility that a trace of the reaction flux passes through a dioxetane intermediate cannot be excluded, particularly when Mn2+ is substituted as the active-site metal. Dioxetanes are known to luminesce during coupled O-O and C-C bond cleavage but usually at much shorter wavelengths than observed with rubisco. Dioxetane emissions at wavelengths longer than 540 nm are rare. Only in the case of chemically initiated electron exchange luminescence of phenolate-substituted dioxetanes has longer wavelength light been observed, and even then the maximum wavelengths were shorter than 630 nm (23).

Higher plant rubisco-manganese emits at substantially longer wavelengths than the laser-excited luminescence from hexa-aquo Mn(II), which has a maximum at 770 nm.2 In a precisely octahedral field with six identical ligands, the manganese emission corresponds to a 4T1g right-arrow 6A1g transition (both spin and parity forbidden) within the weak field d5 manifold of Mn(II). If the ligand field is increased, the energy of this transition decreases. Thus, in higher plant rubisco-manganese, either the ligating atoms at the manganese site are slightly stronger ligands than the oxygen of H2O or the local field has significantly lower symmetry. In the latter case, an inequivalence of the ligands, either in nature or deviation from precise octahedral position, leads to a splitting of the 4T1 state. Only the lowest excited state can emit and then the emission seen from the distorted system could thus appear as a lower energy component of the split 4T1 state. These possibilities could be distinguished by measurement of the absorption energies and characteristics of the rubisco-manganese site. Although the Mn(II) transitions are not easily seen directly by absorption spectroscopy because of their intrinsic weakness, they may be made visible in temperature-dependent magnetic circular dichroism spectroscopy, which is a particularly sensitive and selective method for observing paramagnetic metalloenzyme centers. Previous ESR studies have demonstrated varying degrees of rhombic distortion of the coordination sphere of rubisco-bound Mn2+ in enzymes from spinach and R. rubrum complexed with the six-carbon intermediate analog, 2'-carboxy-D-arabinitol-1,5-bisphosphate (24-26). Furthermore, x-ray crystallography studies showed that the bound Mg2+ ion in activated spinach rubisco is in close to octahedral coordination when the enzyme is not ligated by sugar phosphate, but that considerable distortion of the metal ion coordination sphere occurs after binding of 2'-carboxy-D-arabinitol-1,5-bisphosphate (27).

The Manganese Luminescence Spectrum of Hexadecameric Rubisco-- The intensity of successive spectra diminished exponentially with time when the RuBP concentration was far in excess of the Km (400 nM for the spinach enzyme (2)). This well known "fallover" property of rubisco results from the tight binding of inhibitors originating both from rubisco side reactions and from D-glycero-2,3-pentodiulose-1,5-bisphosphate, a contaminant of RuBP (16). No change in the spectral peak or distribution was apparent with time. This eliminates one of the potential criticisms of the spectrum reported by Lilley et al. (2). In that study, the spectra were recorded much more slowly than in the present study and required correction for the time-dependent decay of the luminescence; a correction that assumed that the spectrum did not change with time. The unchanging spectra also suggest that the presence of such tight-binding inhibitors on one or more of the active sites does not affect the orientation of amino acid residues that coordinate Mn2+ ions within the remaining unbound and catalytically active sites on the same rubisco hexadecameric holoenzyme.

The spectra for rubisco from spinach, tobacco wild type, and tobacco L335V mutant are indistinguishable in terms of general shape and maxima in the 797-800 nm range. Although spinach and tobacco rubisco differ by several amino acid residues in the large and small subunits, the residues involved in their active sites are the same and they have similar structures (28) and kinetic characteristics. L335V rubisco (Mg2+-activated) has an oxygenase Vmax 33% of the wild type and a CO2/O2 specificity ratio 25% of the wild type (11). The L335V mutation represents a shortening of the side chain at this residue, which does not participate directly in catalysis, but may contribute to positioning the mobile loop 6 as its terminal methyl groups contact the P2 phosphate of bound RuBP in the active site. This has the potential to cause a small change in the position or orientation of the epsilon  amino group of the adjacent Lys-334 that is catalytically critical (11). This group is involved in hydrogen bonding with an intermediate carboxyl group formed by the attack of CO2 on the enediol form of RuBP. Here, the L335V rubisco, when Mn2+-activated, exhibited 52% of the chemiluminescence intensity of the wild type. However, the absence of any perceptible effect on the spectrum suggests that the ligand field of Mn2+ at the active site is not disturbed by the L335V mutation.

When the reaction was performed in 96.5% 2H2O, the intensity was approximately doubled as reported previously (1, 3). However, the peak wavelength (799 nm) and distribution of the emission was unchanged.

Difference in Luminescence between Dimeric and Hexadecameric Rubiscos-- The spectrum for R. rubrum rubisco, a homodimer devoid of small subunits, has a maximum at 760 nm, clearly blue-shifted in comparison to that from the hexadecameric higher plant enzymes composed of both large and small subunits. Indeed, this peak is located at a wavelength slightly shorter than photoluminescence from hexa-aquo Mn(II).2 Although the aminoacyl side chains near the metal ion are conserved between the rubisco types, the dispositions of these critical ligand-forming groups may be slightly altered by substitutions in residues a little more removed, or by the absence in the dimeric enzyme of small subunits. The latter influence the catalytic activity of higher plant rubiscos by remote interactions, which telegraph structural alterations to the active site (29). ESR spectral differences between the Mn2+-2'-carboxy-D-arabinitol-1,5-bisphosphate complexes of the plant and R. rubrum enzymes also indicate differences in metal coordination between the two types of rubisco (24, 25).

Temperature and pH have a marked effect on the relative luminescence yield of R. rubrum rubisco, unlike the situation with hexadecameric enzyme (3). The temperature response of the dimeric enzyme was attributed to temperature sensitivity of the balance between radiationless and luminescent decay pathways and the pH effect to the presence in the dimeric, but not the hexadecameric, enzyme of an active-site group with pKa above 7.4. Clearly this group influences the Mn2+ ligand field.

The comparatively lower intensity of luminescence from R. rubrum rubisco (Fig. 4c) is partly because of competitive inhibition of the oxygenase activity by the high bicarbonate concentrations in the reaction mixtures in the present experiments. These result from bicarbonate carried over from the preactivation mixture; a higher bicarbonate concentration is required to activate the bacterial rubisco fully (30). Additionally, the relative luminescence yield of R. rubrum rubisco is lower than that of the spinach enzyme (30). The burst of luminescence in the initial seconds of catalysis (a further distinguishing feature of chemiluminescence from R. rubrum rubisco (2)) will have decayed before measurement of the first spectrum (commencing 5 s after addition of RuBP) and thus had little influence (Fig. 3c).

How Does Mn2+ Become Excited?-- The luminescence emitted by Mn2+-activated rubisco offers a unique window into the oxygenase chemistry catalyzed by this enzyme. One possible mechanism that would cause excitation of the active-site Mn2+ during oxygenation is an electron-exchange process involving a transient Mn3+ intermediate (Fig. 5). This mechanism entails a succession of single-electron transfers initiated by donation of a ground-state electron from Mn2+ to O2 and completed by acceptance of an electron from the sugar substrate into a higher energy orbital of Mn3+. This single-electron process would facilitate the spin inversion required for ground-state, triplet O2 to react with singlet RuBP while maintaining spin conservation overall. The free energy change associated with the strongly exergonic oxygenation reaction, estimated by microcalorimetry to be ~300 kJ mol-1 (31), is sufficient to achieve the excitation, even allowing for a somewhat greater energy requirement for the excitation than that indicated by the wavelength of the radiation accompanying the decay of the excited Mn2+ to ground state (151 kJ mol-1 for 800 nm photons).


View larger version (29K):
[in this window]
[in a new window]
 
Fig. 5.   One possible electron-exchange mechanism for excitation of the Mn2+ ion during addition of triplet O2 to the singlet enediolate of RuBP. R1 = -CH2OPO<UP><SUB>3</SUB><SUP>=</SUP></UP>; R2 = -CHOH-CH2OPO<UP><SUB>3</SUB><SUP>=</SUP></UP>.

This mechanism implies that excitation of the Mn2+ occurs on every oxygenation turnover, consistent with our previous observations that the chemiluminescence intensity is always proportional to the O2 consumption rate (3) and that the quantum efficiency is extremely low because of competing non-radiative decay processes (2). Single-electron exchanges are not possible with the natural metal activator, Mg2+; hence the mechanism of O2 addition catalyzed by Mg2+-activated rubisco must be different, involving less facile two-electron processes. This is consistent with the ~10-fold greater catalytic effectiveness (kcat/Km for O2) of the Mn2+-activated oxygenase, compared with its Mg2+-activated counterpart (32).

The proposed mechanism also implies that Mn2+ excitation and luminescence is linked to the catalytic step in which the 3'-keto-2'-peroxyarabinitol-P2 intermediate is formed. Therefore, the hydration and cleavage of this intermediate to form the oxygenase products, P-glycolate and P-glycerate, should not emit light. This prediction could be checked by feeding the peroxyketone intermediate as substrate.

The Physiological Importance of Activation of Rubisco by Mg2+-- Rubisco-catalyzed oxygenation is maladaptive, increasing the energy requirements of photosynthesis (33). The oxygenating capacity of Mn2+-activated rubisco is so large that photosynthesis supported by this catalyst would be incapable of sustaining a positive carbon balance in the current Earth atmosphere. Calculations (34) using a leaf-photosynthesis model (18) and the kinetic parameters of higher plant manganese-rubisco (32), estimate that, if Mn2+ replaced Mg2+ in the active site of rubisco, the CO2 compensation point of the leaf (the minimum CO2 partial pressure required for positive net CO2 assimilation) would rise 30-fold to ~1500 µbar. Natural selection of carboxylation chemistry facilitated by the ligand field of Mg2+, rather than Mn2+ (which has a vital role in other photosynthetic complexes, such as Photosystem II), may thus be viewed as one of the enabling fundamentals of autotrophic life.

    ACKNOWLEDGEMENTS

We are grateful to S. Whitney, G. Pearce, and H. Kane for provision of the purified rubisco preparations and H. Riesen for useful discussions.

    FOOTNOTES

* The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

§ To whom correspondence should be addressed: Dept. of Biological Sciences, University of Wollongong, Northfields Ave., Wollongong 2522, Australia. Tel.: 61242-213-431; Fax: 61242-214-135; E-mail: rossl@uow.edu.au.

Published, JBC Papers in Press, February 25, 2003, DOI 10.1074/jbc.M212402200

2 G. M. Moran, X-Q. Wang, and E. Krausz, personal communication.

    ABBREVIATIONS

The abbreviations used are: rubisco, ribulose-1,5-bisphosphate carboxylase/oxygenase (EC 4.1.1.39); EPPS, N-2-hydroxyethypiperazine-N'-3-propanesulfonic acid; RuBP, D-ribulose-1,5-bisphosphate; CCD, charge-coupled detector.

    REFERENCES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES

1. Mogel, S. N., and McFadden, B. A. (1990) Biochemistry 29, 8333-8337[Medline] [Order article via Infotrieve]
2. Lilley, R. McC., Riesen, H., and Andrews, T. J. (1993) J. Biol. Chem. 268, 13877-13884[Abstract/Free Full Text]
3. Cox, S. D., Lilley, R. McC., and Andrews, T. J. (1999) Aust. J. Plant Physiol 26, 475-484
4. Khan, A. U., and Kasha, M. (1970) J. Am. Chem. Soc. 92, 3293-3300
5. Oelckers, S., Ziegler, T., Michler, I., and Roder, B. (1999) J. Photochem. Photobiol. B Biol. 53, 121-127[CrossRef][Medline] [Order article via Infotrieve]
6. Kanofsky, J. R. (1983) J. Biol. Chem. 258, 5991-5993[Abstract/Free Full Text]
7. Khan, A. U. (1984) J. Photochem. 25, 327-334[CrossRef]
8. Durner, J., Gailus, V., and Böger, P. (1994) FEBS Lett. 354, 71-73[CrossRef][Medline] [Order article via Infotrieve]
9. Chipman, D., Barak, Z. A., and Schloss, J. V. (1998) Biochim. Biophys. Acta 1385, 401-419[Medline] [Order article via Infotrieve]
10. Edmondson, D. L., Badger, M. R., and Andrews, T. J. (1990) Plant Physiol. 93, 1376-1382
11. Whitney, S. M., von Caemmerer, S., Hudson, G. S., and Andrews, T. J. (1999) Plant Physiol. 121, 579-588[Abstract/Free Full Text]
12. Servaites, J. C. (1985) Arch. Biochem. Biophys. 238, 154-160[Medline] [Order article via Infotrieve]
13. Butz, N. D., and Sharkey, T. D. (1989) Plant Physiol. 89, 735-739
14. Ruuska, S., Andrews, T. J., Badger, M. R., Hudson, G. S., Laisk, A., Price, G. D., and von Caemmerer, S. (1998) Aust. J. Plant Physiol. 25, 859-870
15. Horecker, B. L., Hurwitz, J., and Weissbach, A. (1958) Biochem. Prep. 6, 83-90
16. Kane, H. J., Wilkin, J. M., Portis, A. R., and Andrews, T. J. (1998) Plant Physiol. 117, 1059-1069[Abstract/Free Full Text]
17. Barnett, N. W., Hindson, B. J., Jones, P., and Smith, T. A. (2002) Anal. Chim. Acta 451, 181-188[CrossRef]
18. Farquhar, G. D., von Caemmerer, S., and Berry, J. A. (1980) Planta 149, 78-90
19. Kanofsky, J. R. (2000) Methods Enzymol. 319, 59-67[CrossRef][Medline] [Order article via Infotrieve]
20. Blasse, G., and Grabmaier, B. C. (1994) Luminescent Materials , Springer-Verlag, Berlin
21. Watanabe, T., Shirai, N., Okada, H., Honda, Y., and Kuwahara, M. (2001) Eur. J. Biochem. 268, 6114-6122[Abstract/Free Full Text]
22. Lorimer, G. H., Andrews, T. J., and Tolbert, N. E. (1973) Biochemistry 12, 18-23[Medline] [Order article via Infotrieve]
23. Matsumoto, M., Hiroshima, T., Chiba, S., Isobe, R., Watanabe, N., and Kobayashi, H. (1999) Luminescence 14, 345-348[CrossRef][Medline] [Order article via Infotrieve]
24. Miziorko, H. M., and Sealy, R. C. (1980) Biochemistry 19, 1167-1171[Medline] [Order article via Infotrieve]
25. Miziorko, H. M., and Sealy, R. C. (1984) Biochemistry 23, 479-485
26. Gutteridge, S., Sigal, I., Thomas, B., Arentzen, R., Cordova, A., and Lorimer, G. (1984) EMBO J. 3, 2737-2743
27. Taylor, T. C., and Andersson, I. (1996) Nat. Struct. Biol 3, 95-101[Medline] [Order article via Infotrieve]
28. Schreuder, H. A., Knight, S., Curmi, P. M. G., Andersson, I., Cascio, D., Sweet, R. M., Brändén, C-I., and Eisenberg, D. (1993) Protein Sci. 2, 1136-1146[Abstract/Free Full Text]
29. Paul, K., Morell, M. K., and Andrews, T. J. (1993) Plant Physiol. 102, 1129-1137[Abstract/Free Full Text]
30. Martin, M. L., and Tabita, F. R. (1981) FEBS Lett. 129, 39-43[CrossRef]
31. Frank, J., Kositza, M. J., Vater, J., and Holzwarth, J. F. (2000) Phys. Chem. Chem. Phys. 2, 1301-1304
32. Christeller, J. T., and Laing, W. A. (1979) Biochem. J. 183, 747-750[Medline] [Order article via Infotrieve]
33. Morell, M. K., Paul, K., Kane, H. J., and Andrews, T. J. (1992) Aust. J. Bot. 40, 431-441
34. Jordan, D. B., and Ogren, W. L. (1983) Arch. Biochem. Biophys. 227, 425-433[Medline] [Order article via Infotrieve]


Copyright © 2003 by The American Society for Biochemistry and Molecular Biology, Inc.



This Article
Abstract
Full Text (PDF)
All Versions of this Article:
278/19/16488    most recent
M212402200v1
Purchase Article
View Shopping Cart
Alert me when this article is cited
Alert me if a correction is posted
Citation Map
Services
Email this article to a friend
Similar articles in this journal
Similar articles in PubMed
Alert me to new issues of the journal
Download to citation manager
Copyright Permissions
Google Scholar
Articles by Lilley, R. McC.
Articles by Andrews, T. J.
Articles citing this Article
PubMed
PubMed Citation
Articles by Lilley, R. McC.
Articles by Andrews, T. J.


HOME HELP FEEDBACK SUBSCRIPTIONS ARCHIVE SEARCH TABLE OF CONTENTS
 All ASBMB Journals   Molecular and Cellular Proteomics 
 Journal of Lipid Research   Biochemistry and Molecular Biology Education 
Copyright © 2003 by the American Society for Biochemistry and Molecular Biology.