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Chloroplast biogenesis. 37. induction of chlorophyllide a (E459 F675) accumulation in higher plants
Plant Science Letters, 24 (1982) 27--37
27
Elsevier/North-HollandScientificPublishers Ltd.
CHLOROPLAST BIOGENESIS. 37. INDUCTION OF CHLOROPHYLLIDE a (E459 F675) ACCUMULATION IN HIGHER PLANTS*
JEFFREY X. DUGGANand CONSTANTINA. REBEIZ** Laboratory of Plant Pigment Biochemistry and Photobioiogy, Department of Horticulture, University of Illinois, Urbana, IL 61801 (U.S.A.) (Received May 16th, 1981) (Revision received July 14th, 1981) (Accepted July 16th, 1981)
SUMMARY
The conversion of divinyl protochlorophyllide a (DV Pchlide a) (E443 F625), where E and F refer to fluorescence excitation and emission maxima respectively in ether at 77 K, into chlorophyllide a (Chlide a) (E459 F675) is reported. This reaction was demonstrated by first inducing etiolated cucumber cotyledons to accumulate DV Pchlide a (E443 F625) devoid of monovinyl (MV) contamination by a series of light/dark treatments. The accumulated DV Pchlide a (E443 F625) was then converted into Chlide a (E459 F675) by a 2.5-ms light pulse (L). The Chlide a (E459 F675) thus generated exhibited spectrofluorometric properties which differed from those of MV Chlide a (E447 F674) at 77 K and at room temperature before and after demetaUation. The techniques described in this work permit the preparation of Chlide a (E459 F675) devoid of MV Chlide a contamination, for further chemical structural identification of this novel chlorophyll (Chl) precursor. INTRODUCTION It has recently been reported that, contrary to previous beliefs, the Chl pool of green plants is made up of several different Chl chromophoric species [1] which may have different functions in photosynthesis [2]. It has also been proposed that the different Chl a chemical species are synthesized via *This work was supported by Research Grant PCM-7811559 from the National Science Foundation and by funds from the Illinois Agricultural Experiment Station. **To whom reprint requests should be sent. Abbreviations: Chl, chlorophyll; Chlide, chlorophyllide; D, 60 min dark incubation; DV, divinyl; L, 2.5 ms exposure to actinic white light; MV, monovinyl; Pchlide, protochlorophyllide.
0304--4211/82/0000--0000/$02.75 © Elsevier/North-Holland Scientific Publishers Ltd.
28 a 4-branched Chl biosynthetic pathway instead of via a linear single chain p a t h w a y [3]. In this newly proposed pathway, most of the Chl chromophoric species are considered to be formed in parallel from two different Chlide a chemical species, i.e. Chlide a (E449 F675) and Chlide a (E459 F675), where E and F refer to the Soret excitation and red emission maxima of these tetrapyrroles in ether at 77 K [4,5]. These two Chlide a chemical species were shown in turn to be formed by photoreduction of the heterogenous MV/DV Pchlide pool of etiolated tissues [4,5]. On the basis o f the photoreduction of the heterogeneous MV/DV Pchlide a pool of etiolated tissues and the concomitant appearance of Chlide a (E449 F675) and Chlide a (E459 F675) the latter were tentatively identified as MV and DV Chlide a, respectively [4]. However, no specific precursor-product relationship between any single Pchlide a and its corresponding Chlide a was established. Neither was the chemical structure of Chlide a (E459 F675} and Chlide a (E449 F675) ascertained by a detailed chemical structural analysis of these compounds. As a preliminary step to such determinations, we describe in this work the preparation of Chlide a (E459 F675) devoid o f Chlide a (E449 F675) contamination. Furthermore, it is shown that Chlide a (E459 F675) is specifically formed from DV Pchlide a (E443 F625} by photoreduction. MATERIALS AND METHODS
Plant material. Cucumber seeds, Cucumis sativus L. cv. Beit Alpha, were sown in moist vermiculite and grown in darkness as previously described [6]. All manipulations of living plant material were performed under a green safelight. Light/dark treatments. Photoconversion of the Pchlides a into Chlides a was achieved by irradiating etiolated, excised, hookless cotyledons at room temperature with L. The light flash was generated by a Sunpack model Auto 611 photographic flash unit (Berkey Marketing Co., Woodside, NY) placed 11 cm above the tissue. The cotyledons were held either in 9-cm petri dishes or in a tea strainer (vide infra) and a mirror was placed 4 cm below the sample in order to reflect the incident light back onto the tissue. The Pchlide pool was regenerated by returning the tissue to darkness for 60 min (D). For multiple light/dark treatments this process was repeated as m a n y times as needed. When it was necessary to extract the cotyledons immediately after a light flash, the tissue was first transferred to a tea strainer, then was illuminated as described above. It was then frozen immediately by dipping the strainer into liquid N2. Pigment extraction. Pigments were extracted from the frozen tissue by homogenizing 1 g of tissue in a mortar at 0--4°C in 6.7 ml of acetone/0.1 N NH4OH (9 : 1, v/v) with a small a m o u n t of white sand. After centrifugation at 39 000 × g for 10 min, the fully esterified pigments were removed from the 80% acetone extract by extraction with hexane [7 ]. The Chlides and
29 Pchlides which remained in the hexane-extracted acetone residue were transferred to ether as described elsewhere [8]. Chromatography. Chlides were separated from Pchlides by chromatography of t h e ether extract on thin layers of polyethylene developed in 90% acetone at 4°C. This solvent system was successfully used earlier to separate MV from DV Pchlide at r o o m temperature [5]. DemetaUation of Chlides. The removal of Mg from the Chlide molecules was achieved by acid treatment as follows: 2 ml of 4.0 N HC1 were mixed with 2 ml of the ether extract containing the Chlides in a 30 ml separatory funnel. The acidic mixture was allowed to stand at room temperature for 30 rain with occasional shaking. ~olid NaHCO3 was then added to neutralize t h e acid. Alternatively the pigment was dissolved in 0.1 ml of ether which were t h e n mixed with 2 ml of 2.0 N HC1. The pigment passed immediately into the acid phase, to which 2 ml of ether were added. Solid NaHCO3 was then used to neutralize the acid. When the CO2 evolution had subsided, 3 ml of distilled H20 were added, the aqueous phase was decanted, and the ether phase containing the demetallated Chlides was washed with distilled H20 until it became acid-free. Spectrofluorometry. Fully corrected fluorescence excitation and emission spectra were recorded either with a Perkin Elmer spectrofluorometer, model MPF-3, or with an SLM spectrofluorometer, model 8000 DS interfaced with a Hewlett-Packard microcomputer model 9825S, as described elsewhere [4]. Emission spectra were recorded on the Perkin Elmer spectrofluorometer with an excitation slit width of 6 nm and an emission slit width of 3 nm, while excitation spectra were recorded with an excitation slit width of 3 nm and an emission slit width of 6 nm. Emission spectra recorded on the SLM spectrofluorometer employed an excitation slit width of 4 n m and an emission slit width of 2 nm, while excitation spectra were recorded with an excitation slit width of 2 nm and an emission slit width of 4 nm. Under these conditions the spectral accuracy of the reported maxima is + 1 nm. Low temperature spectra were recorded in ether by previously published methods
[9]. RESULTS
Experimental strategy. In order to demonstrate a precursor-product relationship between the DV Pchlide a and the Chlide a (E459 F675) pools, the experimental strategy was designed around the observation that, in contrast t o etiolated tissues, the Pchlide a pool o f greening tissues was made up mainly o f DV Pchlide a [5]. It was therefore conjectured that if, by judicious illumination of etiolated tissues, the latter were induced to accumulate only phototransformable DV Pchlide, t h e n it would be possible to obtain the immediate p h o t o p r o d u c t of this pool by a very short light treatment. This in turn would permit the unambiguous determination of the fluorescence properties of the newly formed Chlide a (E459 F675) pool.
30 An experiment was therefore designed in which etiolated cotyledons were given a series of L each followed b y D. The light flash photoconverted the Pchlide a pool into Chlide a, while D allowed the regeneration of the Pchlide a pool. Before and after each light treatment, the monocarboxylic phorbin pool containing the Pchlide a and Chlide a pools was extracted in ether and was monitored spectrofluorometrically. In this manner the time course for DV Pchlide a appearance at the end of each D could be compared with the appearance of putative DV Chlide a immediately after each subsequent phototransforming L. Thus, in the ensuing sections, the induction of DV Pchlide a accumulaL tion is first described, followed b y the conversion of the induced DV Pchlide a pool into a Chlide a (E459 F675) pool. Induction o f D V Pchlide a accumulation by successive DL treatments. The evolution of the Pchlide a pool from a mixed MV/DV pool in etiolated tissues to a DV Pchlide a pool during a sequence of three LD cycles is depicted in Fig. 1. The spectral profile of the monocarboxylic phorbin pool of etiolated cucumber cotyledons was made up exclusively of Pchlide a. As reported previously [5], this pool consisted of a MV Pchlide a (E437 F625) component and of a DV Pchlide a (E443 F625) c o m p o n e n t (Fig. l(a)). Following the first phototransforming light pulse, the regenerated monocarboxylic phorbin pool appeared to be made up mainly of DV Pchlide a (E443 F625) (Fig. l(b)). However, small amounts of MV Pchlide a (E437 F625) could still be observed as evidenced b y a small Soret excitation shoulder at 437 nm (Fig. 1B(b)). Furthermore, an u n k n o w n spectrofluorometric species, which populates the DV Pchlide a pool and which has been previously attributed to an u n k n o w n metabolic intermediate [5], also became noticeable. It exhibited a Soret excitation maximum at 451 nm and an emission maximum at 625 nm (Fig. l(b)). More recent investigations of this u n k n o w n spectrofluorometric species suggest that it m a y be attributed to the resolved Bx (o-o) band of DV Pchlide a, as suggested by Houssier and Sauer [10; Belanger and Rebeiz, unpublished]. We are n o w in the process of sorting o u t this possibility. Thus, since it appears that species (E451 F625) is some kind of a DV Pchlide a spectroscopic manifestation, its exact nature at this stage should n o t affect the o u t c o m e of the present work, and the doublet at E443 and E451 nm can be considered collectively as a marker of the DV Pchlide a pool. It was also observed that after 60 rain in the dark, during which the Pchlide a pool was regenerated following illumination, very little Chlide a remained in the monocarboxylic phorbin pool. This was evidenced b y the very small emission increment at 673--675 nm over and b e y o n d that attributed to the tail of the Pchlide a vibrational band at 675 nm (Fig. 1A(a) vs. 1A(b)). This in turn indicated that nearly all the Chlide a produced by the initial illumination was esterified to Chl a during the ensuing dark period. Finally, during the course of t w o additional LD cycles, all traces of
31 I
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Fig. 1. Fluorescence emission (A) and excitation (B) spectra in ether at 77 K of the Pchlide pools extracted from etiolated cucumber cotyledons after sequential cycles (LD) of 2.5 ms of actinic light (L) followed by 60 min of darkness (D). All emission spectra were recorded on a Perkin-Elmer, Model MPF-3 spectrofluorometer unless otherwise indicated. Excitation spectra b--d were recorded on an SLM spectrofluorometer, Model 8 0 0 0 DS operated in the ratio mode, while excitation spectrum a was recorded on the Perkin-Elmer instrument. The emission spectra were elicited by the E-wavelengths indicated. Likewise, the excitation spectra were recorded at the indicated F-wavelengths. Ordinate scale attenuation values are indicated on the spectra, 359× being the highest sensitivity used. Spectra have been vertically arranged for the purpose of clarity. Arrows point to wavelengths of interest. (a) Etiolated tissues; (b) after one LD cycle; (c) after two LD cycles; (d) after three LD cycles.
MV Pchlide a and Chlide a disappeared from the monocarboxylic phorbin pool which contained exclusively regenerated DV Pchlide a (Fig. l(c,d)). The lack of Chlide a was evidenced b y a fluorescence emission profile of the monocarboxylic phorbin pool that was identical to that of a fully etiolated tissue and which lacked the incremental fluorescence at 673--675 nm (Fig. l(a,d)). On the other hand, the absence of MV Pchlide a in this pool was obvious from its Sorer excitation profile which lacked a shoulder at 437 nm (Fig. 1B(d)). Altogether, the above results indicated that, after a third L pulse followed
32
b y a D incubation period, the monocarboxylic phorbin pool was made up o f DV Pchlide a and lacked any MV Pchlide a or Chlide a. It thus follows that the Chlide a generated by a fourth L pulse will have been formed exclusively from the regenerated DV Pchlide a pool. Photoconversion o f the induced D V Pchlide a pool into Chlide a (E459 F675). The photoconversion of the regenerated DV Pchlide a pool is contrasted with the photoconversion of the mixed MV/DV Pchlide a pools in Fig. 2. A single L pulse converted most of the mixed MV/DV Pchlide a pool of the etiolated tissue (emission maximum at 625 nm) into Chlide a, having an emission maximum at 674 nm (Figs. 1A(a) and 2A(a)). Although the newly formed Chlide a pool exhibited a single emission maximum at
F675
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Wavelength of Emission(nm] Wavelength of Excitation(nm)
Fig. 2. Fluorescence emission (A) and excitation (B) spectra at ether at 77 K of the Chlide a pools extracted from etiolated cucumber cotyledons after sequential 2.5 ms actinic light pulses (L) with intervening 60-min dark incubations. Emission spectra b--d were recorded on the Perkin-Elmer spectrofluorometer. The remaining spectra were recorded on the SLM spectrofluorometer. Arrows point to wavelengths of interest. The etiolated tissues were immediately frozen in liquid N 2 and their monocarboxylic phorbin pools extracted after (a) one actinic light pulse (L); (b) one LD cycle followed by one photoconverting flash; (c) two LD cycles followed by one photoconverting flash; (d) three LD cycles followed by one photoconverting flash. All other symbols are as in Fig. 1
33 a b o u t 674 nm, it exhibited a split Soret excitation profile with a major excitation m a x i m u m at a b o u t 448--449 nm and a shoulder at a b o u t 4 5 8 - 459 nm (Fig. 2B(a)). On the basis of the relative excitation amplitudes of the MV and DV Pchlide a Sorets at 437 and 443 nm respectively, and on the basis o f the amplitudes o f the corresponding Chlide a Sorets at 449 and 459 nm, t h e latter were previously assigned to MV and DV Chlide a respectively [ 4]. In this w o r k the precursor-product relationship o f MV Pchlide a to Chlide a (E449 F675) and that of DV Pchlide a to Chlide a (E459 F675) is demonstrated. As it m a y be recalled, the Pchlide a pool which was regenerated after the first light pulse was made up mainly of DV Pchlide a and of much smaller amounts of MV Pchlide a (Fig. 1B(b)). A L treatment of this regenerated Pchlide a pool converted most of the Pchlide a into Chlide a (Figs. 1A(b) and 2A(b)). The newly formed Chlide a pool was mainly made up of Chlide a (E459 F675) and of much smaller amounts of Chlide a (E449 F675) (Fig. 2B(b)). Actually, the latter appeared as a small excitation shoulder on t h e short wavelength tail of the Chlide a (E459 F675) Soret and its relative amplitude was commensurate with the relative amplitude o f the MV Pchlide a c o m p o n e n t that was present before photoconversion (cf. Figs. 1B(b) and Fig. 2B(b)). Likewise, the same relative relationship between the MV/DV Pchlide a components and the two Chlide a components was observed in etiolated tissues that were subjected to t w o LD cycles (Figs. 1B(c) and 2B(c)). These results strongly suggested a precursor-product relationship between MV Pchlide a (E437 F625) and Chlide a (E449 F675) and between DV Pchlide a (E443 F625) and Chlide a (E459 F675). The above relationship was further corroborated b y converting the DV Pchlide a pool which was regenerated after three LD cycles into Chlide a (E459 F675) by an L pulse (Fig. 2(d)). The absence of a MV Chlide a c o m p o n e n t is evident by the complete lack of a MV Soret excitation peak or shoulder at 448--449 nm (Fig. 2B(d)). Spectrofluorometric properties of the M V Chlide a and Chlide a (E459 F675) pools at room temperature. With the intermittent light/dark treatment o f etiolated tissues which was described above, it became possible to prepare Chlide a (E459 F675) containing very little MV Chlide a contamination and to compare its spectroscopic properties to those of MV Chlide a. A monocarboxylic phorbin pool made up mainly of MV Chlide a was extracted from etiolated cucumber cotyledons which had been illuminated for 2.5 ms with actinic white light then placed in darkness for 5 min before extraction. The L pulse converted most of the mixed MV/DV Pchlide a pool into a mixed Chlide a {E449 F675) + Chlide a (E459 F675) pool (Figs. l(a) and 2(a)). This monocarboxylic phorbin pool also contained small amounts o f non-phototransformable Pchlide a with an emission maximum at a b o u t 625 nm (Fig. 3A(a)). During the 5-rain post-illumination dark period all o f the Chlide a (E459 F675) was converted into blue-shifted (i.e. short wavelength) Chl a species of unkn own structure [ 1,4 ]. The remaining Chlide a
34
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Fig. 3. F l u o r e s c e n c e e m i s s i o n ( A ) a n d e x c i t a t i o n (B) s p e c t r a o f t h e M V Chlide a a n d Chlide a ( E 4 5 9 F 6 7 5 ) a n d o f t h e i r c o r r e s p o n d i n g p h e o p h o r b i d e s a p r e p a r e d f r o m lightt r e a t e d e t i o l a t e d c u c u m b e r c o t y l e d o n s . P i g m e n t s w h i c h w e r e m o n i t o r e d at 77 K were dissolved in e t h e r , while t h o s e m o n i t o r e d at r o o m t e m p e r a t u r e ( R T ) w e r e dissolved in h e x a n e - e x t r a e t e d 80% a c e t o n e (see Materials a n d M e t h o d s ) . S p e c t r u m f was r e c o r d e d o n t h e S L M s p e c t r o f l u o r o m e t e r , while t h e r e m a i n i n g s p e c t r a were r e c o r d e d o n t h e PerkinE l m e r f l u o r o m e t e r . A r r o w s p o i n t t o w a v e l e n g t h s o f interest. MV, refers t o t h e s t a n d a r d m o n o v i n y l Chlide a p o o l w h i l e D V refers t o t h e Chlide a ( E 4 5 9 F 6 7 5 ) p o o l b e f o r e a n d a f t e r d e m e t a l l a t i o n . All o t h e r s y m b o l s are as in Fig. 1.
35 pool (Fig. 3(a)) was then found to consist mainly of Chlide a (E446-447 F674) and of smaller amounts of blue-shifted, short wavelength Chlide a species [4]. By analogy to the MV Chl a {E446 F674) of mature green tissues [1], Chlide a {E446 F674) is here identified as MV Chlide a. A monocarboxylic phorbin pool made up mainly of Chlide a (E459 F675) was extracted from etiolated cucumber cotyledons subjected to three LD treatments, followed b y one 2.5-ms phototransforming light pulse (Fig. 3(b)). The latter converted most of the regenerated DV Pchlide a into Chlide a (E459 F675). This pool also contained some unphototransformed Pchlide a with an emission maximum at a b o u t 625 nm (Fig. 3A{b)}. The latter, did not, however, interfere with the recording of the spectrofluorometric properties of the Chlide a (E459 F675) because of the small amounts of Pchlide a present and of the insignificant contribution of its vibrational band at 673--675 nm to the red emission band of Chlide a. To insure that the spectrofluorometric differences between the standard MV Chlide a pool and Chlide a (E459 F675) were not artifacts associated with low temperature spectroscopy, a comparison was made between the fluorescence spectra of the two Chlide a species recorded at r o o m temperature in h e x a n e ~ x t r a c t e d acetone. Both pools exhibited identical emission maxima at a b o u t 675 nm (Fig. 3A(d,e}}. The much smaller emission at a b o u t 639 nm is that of the non-phototransformed Pchlide a pool (Fig. 3A(d,e}). The Soret excitation maxima of the two Chlide a pools were, however, quite different. While the standard MV Chlide a pool exhibited a ~oret excitation maximum at about 434 nm, the Chlide a (E459 F675) Soret excitation maximum was red-shifted b y about 8 nm and was observed at 442 nm (Fig. 3B(d,e)). These results indicated that the spectrofluorometric differences observed between the Soret excitation maxima of the two Chlide a pools were not due to low temperature artifacts but were most probably related to chemical differences between the two Chlide a species. Spectrofluorometric properties o f the two Chlide a species after demetallation. If the two Chlide a species do in fact differ from each other b y virtue o f chemical m o d i f i c a t i o n s ) of the porphyrin macrocycle, then removal of the central Mg atom from these two compounds should generate two spectrally different pheophorbide species. The t w o Chlide a pools were therefore demetallated and the fluorescence properties o f the two resulting pheophorbide a species were compared. The spectra were recorded at 77 K in ether. The two pheophorbide a species exhibited different fluorescence excitation maxima (Fig. 3(f,g)}. While both MV and DV pheophorbide a exhibited an emission maximum at 667 nm (Fig. 3A(f,g)}, the Soret excitation maximum of the MV pheophorbide a was observed at 415 nm, while that of the other pheophorbide a species was observed at 424 nm {Fig. 3B{f,g)). Altogether, the above results indicated that the spectrofluorometric differences b e t w e e n the t w o Chlide a species were preserved after the removal o f the central Mg atom.
36
Purification of Chlide a (E459 F675). Since further chemical characterization o f Chlide a (E459 F675) m a y require the availability o f pure quantities of this Chlide a species, it was deemed desirable to purify Chlide a (E459 F675) from the small amounts of non-phototransformed Pchlide a that were also present in the monocarboxylic phorbin pool of the lighttreated tissues. Several chromatographic systems were tested; chromatography on thin layers o f polyethylene developed in acetone/H20 (9 : 1, v/v) at 4°C gave the best results and achieved a complete separation of Chlide a (E459 F675) from Pchlide a. In this system Pchlide a migrated with an Rf-value of a b o u t 0.44, while Chlide a (E459 F675) migrated with an Rf-value of about 0.9. It was essential to carry o u t the separation at a b o u t 4°C in order to avoid demetallation o f Chlide a (E459 F675). Indeed, it was constantly observed throughout this work that this Chlide a species was much more susceptible to demetallation than MV Chlide a. Furthermore, it has been found that prepurification of the monocarboxylic phorbin pool on thin layers of silica gel H developed in toluene/ethyl acetate/ethanol (8 : 2 : 2, v/v/v) at 4°C improves the performance of the polyethylene separation, presumably by removing colorless lipids [4]; Belanger and Rebeiz, unpublished]. The fluorescence properties in ether at 77 K of purified Chlide a (E459 F675) were identical to those of the Chlide a (E459 F675) enriched monocarboxylic phorbin pool except for the absence of a Pchlide a emission at 625 nm (Fig. 3(c)). DISCUSSION Chlide a (E459 F675) was first observed by Belanger and Rebeiz [4] immediately after the photoconversion of the mixed MV Pchlide a (E437 F625)/DV Pchlide a (E443 F625) pool of etiolated tissues into a mixed Chlide a (E449 F675)/Chlide a (E459 F675) pool. On the basis of (a) the briefness of the light treatment (47 ms) and (b) assuming that the short wavelength MV Pchlide a is likely to yield a correspondingly short wavelength Chlide a, while the long wavelength DV Pchlide a is likely to yield a long wavelength Chlide a, the newly formed Chlide a (E449 F675) was tentatively identified as MV Chlide a, and Chlide a (E459 F675) was identified as DV Chlide a. The results reported in this work seem to corroborate the above assignment (vide infra). In order to minimize the occurrence of photochemical side reactions that m a y m o d i f y the structure of the reacting tetrapyrroles, the photoconversion time was reduced in this work from 47 ms to 2.5 ms. Within about 250 ms after the light treatment, all enzymatic activity was stopped b y freezing the tissue in liquid nitrogen, then the pigment was extracted in organic solvents. By assuming that during the brief period that preceded the extraction of the , pigment only photochemistry was operative while biochemical activity was ! neglible, one is led to the conclusion that following the light treatment
37 MV Pchlide a did indeed give rise to Chlide a (E449 F675), while DV Pchlide a gave rise t o Chlide a (E459 F675) as previously proposed [4]. This relationship was further supported by (a) the observation that DV Pchlide a (E443 F625) devoid of MV Pchlide a (E437 F625) contamination was converted by p h o t o r e d u c t i o n into Chlide a (E459 F675) devoid of Chlide a (E449 F675) (Figs. l ( d ) and 2(d)) and (b) by the finding that essentially the same results were obtained by freezing the cotyledons to - 1 0 ° C prior to the light treatment. The consideration of Chlide a (E459 F675) as a Chlide a species chemically distinct from MV Chlide a (E446 F674) is corroborated by its spectrofluorometric properties which differ from those of MV Chlide a (Fig. 3) (a) at 77 K and at room temperature, (b) before and after demetallation and (c) after purification. The unambiguous determination of the structure of Chlide (E459 F675) will require extensive chemical and spectroscopic studies. Such studies will require the preparation of large quantities of this c o m p o u n d in a highly purified state. The techniques described herein are presently being employed to further pursue the chemical identity of this novel Chl precursor. REFERENCES 1 C.A. Rebeiz, F.C. Belanger, G. Freyssinetand D.G. Saab, Biochim. Biophys. Acta,
590 (1980) 234. 2 G. Freyssinet, C.A. Rebeiz, J.M. Fenton, R. Khanna and Govindgee, Photobiochem. Photobiophys., 1 (1980) 203. 3 C.A. Rebeiz, F.C. Belanger, S.A. McCarthy, G. Freyssinet, J.X. Duggan, S.M. Wu and J.R. Mattheis, in: Proc. 5th Int. Cong. Photosynth., 5 (1981) 197. 4 F.C. Belanger and C.A. Rebeiz, Plant Sci. Lett., 18 (1980) 343. 5 F.C. Belanger and C.A. Rebeiz, J. Biol. Chem., 255 (1980) 1266. 6 S.I. Hardy, P.A. Castelfranco and C.A. Rebeiz, Plant Physiol., 46 (1970) 705. 7 C.A. Rebeiz, J.R. Mattheis, B.B. Smith, C.C. Rebeiz and D.F. Dayton, Arch. Biochem. Biophys., 171 (1975) 549. 8 C.A. Rebeiz, J.R. Mattheis, B.B. Smith, C.C. Rebeiz and D.F. Dayton, Arch. Biochem. Biophys., 166 (1975) 446. 9 C.E. Cohen and C.A. Rebeiz, Plant Physiol., 61 (1978) 824. 10 C. Houssier and K. Sauer, Biochim. Biophys. Acta, 172 (1969) 492.
27
Elsevier/North-HollandScientificPublishers Ltd.
CHLOROPLAST BIOGENESIS. 37. INDUCTION OF CHLOROPHYLLIDE a (E459 F675) ACCUMULATION IN HIGHER PLANTS*
JEFFREY X. DUGGANand CONSTANTINA. REBEIZ** Laboratory of Plant Pigment Biochemistry and Photobioiogy, Department of Horticulture, University of Illinois, Urbana, IL 61801 (U.S.A.) (Received May 16th, 1981) (Revision received July 14th, 1981) (Accepted July 16th, 1981)
SUMMARY
The conversion of divinyl protochlorophyllide a (DV Pchlide a) (E443 F625), where E and F refer to fluorescence excitation and emission maxima respectively in ether at 77 K, into chlorophyllide a (Chlide a) (E459 F675) is reported. This reaction was demonstrated by first inducing etiolated cucumber cotyledons to accumulate DV Pchlide a (E443 F625) devoid of monovinyl (MV) contamination by a series of light/dark treatments. The accumulated DV Pchlide a (E443 F625) was then converted into Chlide a (E459 F675) by a 2.5-ms light pulse (L). The Chlide a (E459 F675) thus generated exhibited spectrofluorometric properties which differed from those of MV Chlide a (E447 F674) at 77 K and at room temperature before and after demetaUation. The techniques described in this work permit the preparation of Chlide a (E459 F675) devoid of MV Chlide a contamination, for further chemical structural identification of this novel chlorophyll (Chl) precursor. INTRODUCTION It has recently been reported that, contrary to previous beliefs, the Chl pool of green plants is made up of several different Chl chromophoric species [1] which may have different functions in photosynthesis [2]. It has also been proposed that the different Chl a chemical species are synthesized via *This work was supported by Research Grant PCM-7811559 from the National Science Foundation and by funds from the Illinois Agricultural Experiment Station. **To whom reprint requests should be sent. Abbreviations: Chl, chlorophyll; Chlide, chlorophyllide; D, 60 min dark incubation; DV, divinyl; L, 2.5 ms exposure to actinic white light; MV, monovinyl; Pchlide, protochlorophyllide.
0304--4211/82/0000--0000/$02.75 © Elsevier/North-Holland Scientific Publishers Ltd.
28 a 4-branched Chl biosynthetic pathway instead of via a linear single chain p a t h w a y [3]. In this newly proposed pathway, most of the Chl chromophoric species are considered to be formed in parallel from two different Chlide a chemical species, i.e. Chlide a (E449 F675) and Chlide a (E459 F675), where E and F refer to the Soret excitation and red emission maxima of these tetrapyrroles in ether at 77 K [4,5]. These two Chlide a chemical species were shown in turn to be formed by photoreduction of the heterogenous MV/DV Pchlide pool of etiolated tissues [4,5]. On the basis o f the photoreduction of the heterogeneous MV/DV Pchlide a pool of etiolated tissues and the concomitant appearance of Chlide a (E449 F675) and Chlide a (E459 F675) the latter were tentatively identified as MV and DV Chlide a, respectively [4]. However, no specific precursor-product relationship between any single Pchlide a and its corresponding Chlide a was established. Neither was the chemical structure of Chlide a (E459 F675} and Chlide a (E449 F675) ascertained by a detailed chemical structural analysis of these compounds. As a preliminary step to such determinations, we describe in this work the preparation of Chlide a (E459 F675) devoid o f Chlide a (E449 F675) contamination. Furthermore, it is shown that Chlide a (E459 F675) is specifically formed from DV Pchlide a (E443 F625} by photoreduction. MATERIALS AND METHODS
Plant material. Cucumber seeds, Cucumis sativus L. cv. Beit Alpha, were sown in moist vermiculite and grown in darkness as previously described [6]. All manipulations of living plant material were performed under a green safelight. Light/dark treatments. Photoconversion of the Pchlides a into Chlides a was achieved by irradiating etiolated, excised, hookless cotyledons at room temperature with L. The light flash was generated by a Sunpack model Auto 611 photographic flash unit (Berkey Marketing Co., Woodside, NY) placed 11 cm above the tissue. The cotyledons were held either in 9-cm petri dishes or in a tea strainer (vide infra) and a mirror was placed 4 cm below the sample in order to reflect the incident light back onto the tissue. The Pchlide pool was regenerated by returning the tissue to darkness for 60 min (D). For multiple light/dark treatments this process was repeated as m a n y times as needed. When it was necessary to extract the cotyledons immediately after a light flash, the tissue was first transferred to a tea strainer, then was illuminated as described above. It was then frozen immediately by dipping the strainer into liquid N2. Pigment extraction. Pigments were extracted from the frozen tissue by homogenizing 1 g of tissue in a mortar at 0--4°C in 6.7 ml of acetone/0.1 N NH4OH (9 : 1, v/v) with a small a m o u n t of white sand. After centrifugation at 39 000 × g for 10 min, the fully esterified pigments were removed from the 80% acetone extract by extraction with hexane [7 ]. The Chlides and
29 Pchlides which remained in the hexane-extracted acetone residue were transferred to ether as described elsewhere [8]. Chromatography. Chlides were separated from Pchlides by chromatography of t h e ether extract on thin layers of polyethylene developed in 90% acetone at 4°C. This solvent system was successfully used earlier to separate MV from DV Pchlide at r o o m temperature [5]. DemetaUation of Chlides. The removal of Mg from the Chlide molecules was achieved by acid treatment as follows: 2 ml of 4.0 N HC1 were mixed with 2 ml of the ether extract containing the Chlides in a 30 ml separatory funnel. The acidic mixture was allowed to stand at room temperature for 30 rain with occasional shaking. ~olid NaHCO3 was then added to neutralize t h e acid. Alternatively the pigment was dissolved in 0.1 ml of ether which were t h e n mixed with 2 ml of 2.0 N HC1. The pigment passed immediately into the acid phase, to which 2 ml of ether were added. Solid NaHCO3 was then used to neutralize the acid. When the CO2 evolution had subsided, 3 ml of distilled H20 were added, the aqueous phase was decanted, and the ether phase containing the demetallated Chlides was washed with distilled H20 until it became acid-free. Spectrofluorometry. Fully corrected fluorescence excitation and emission spectra were recorded either with a Perkin Elmer spectrofluorometer, model MPF-3, or with an SLM spectrofluorometer, model 8000 DS interfaced with a Hewlett-Packard microcomputer model 9825S, as described elsewhere [4]. Emission spectra were recorded on the Perkin Elmer spectrofluorometer with an excitation slit width of 6 nm and an emission slit width of 3 nm, while excitation spectra were recorded with an excitation slit width of 3 nm and an emission slit width of 6 nm. Emission spectra recorded on the SLM spectrofluorometer employed an excitation slit width of 4 n m and an emission slit width of 2 nm, while excitation spectra were recorded with an excitation slit width of 2 nm and an emission slit width of 4 nm. Under these conditions the spectral accuracy of the reported maxima is + 1 nm. Low temperature spectra were recorded in ether by previously published methods
[9]. RESULTS
Experimental strategy. In order to demonstrate a precursor-product relationship between the DV Pchlide a and the Chlide a (E459 F675) pools, the experimental strategy was designed around the observation that, in contrast t o etiolated tissues, the Pchlide a pool o f greening tissues was made up mainly o f DV Pchlide a [5]. It was therefore conjectured that if, by judicious illumination of etiolated tissues, the latter were induced to accumulate only phototransformable DV Pchlide, t h e n it would be possible to obtain the immediate p h o t o p r o d u c t of this pool by a very short light treatment. This in turn would permit the unambiguous determination of the fluorescence properties of the newly formed Chlide a (E459 F675) pool.
30 An experiment was therefore designed in which etiolated cotyledons were given a series of L each followed b y D. The light flash photoconverted the Pchlide a pool into Chlide a, while D allowed the regeneration of the Pchlide a pool. Before and after each light treatment, the monocarboxylic phorbin pool containing the Pchlide a and Chlide a pools was extracted in ether and was monitored spectrofluorometrically. In this manner the time course for DV Pchlide a appearance at the end of each D could be compared with the appearance of putative DV Chlide a immediately after each subsequent phototransforming L. Thus, in the ensuing sections, the induction of DV Pchlide a accumulaL tion is first described, followed b y the conversion of the induced DV Pchlide a pool into a Chlide a (E459 F675) pool. Induction o f D V Pchlide a accumulation by successive DL treatments. The evolution of the Pchlide a pool from a mixed MV/DV pool in etiolated tissues to a DV Pchlide a pool during a sequence of three LD cycles is depicted in Fig. 1. The spectral profile of the monocarboxylic phorbin pool of etiolated cucumber cotyledons was made up exclusively of Pchlide a. As reported previously [5], this pool consisted of a MV Pchlide a (E437 F625) component and of a DV Pchlide a (E443 F625) c o m p o n e n t (Fig. l(a)). Following the first phototransforming light pulse, the regenerated monocarboxylic phorbin pool appeared to be made up mainly of DV Pchlide a (E443 F625) (Fig. l(b)). However, small amounts of MV Pchlide a (E437 F625) could still be observed as evidenced b y a small Soret excitation shoulder at 437 nm (Fig. 1B(b)). Furthermore, an u n k n o w n spectrofluorometric species, which populates the DV Pchlide a pool and which has been previously attributed to an u n k n o w n metabolic intermediate [5], also became noticeable. It exhibited a Soret excitation maximum at 451 nm and an emission maximum at 625 nm (Fig. l(b)). More recent investigations of this u n k n o w n spectrofluorometric species suggest that it m a y be attributed to the resolved Bx (o-o) band of DV Pchlide a, as suggested by Houssier and Sauer [10; Belanger and Rebeiz, unpublished]. We are n o w in the process of sorting o u t this possibility. Thus, since it appears that species (E451 F625) is some kind of a DV Pchlide a spectroscopic manifestation, its exact nature at this stage should n o t affect the o u t c o m e of the present work, and the doublet at E443 and E451 nm can be considered collectively as a marker of the DV Pchlide a pool. It was also observed that after 60 rain in the dark, during which the Pchlide a pool was regenerated following illumination, very little Chlide a remained in the monocarboxylic phorbin pool. This was evidenced b y the very small emission increment at 673--675 nm over and b e y o n d that attributed to the tail of the Pchlide a vibrational band at 675 nm (Fig. 1A(a) vs. 1A(b)). This in turn indicated that nearly all the Chlide a produced by the initial illumination was esterified to Chl a during the ensuing dark period. Finally, during the course of t w o additional LD cycles, all traces of
31 I
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Fig. 1. Fluorescence emission (A) and excitation (B) spectra in ether at 77 K of the Pchlide pools extracted from etiolated cucumber cotyledons after sequential cycles (LD) of 2.5 ms of actinic light (L) followed by 60 min of darkness (D). All emission spectra were recorded on a Perkin-Elmer, Model MPF-3 spectrofluorometer unless otherwise indicated. Excitation spectra b--d were recorded on an SLM spectrofluorometer, Model 8 0 0 0 DS operated in the ratio mode, while excitation spectrum a was recorded on the Perkin-Elmer instrument. The emission spectra were elicited by the E-wavelengths indicated. Likewise, the excitation spectra were recorded at the indicated F-wavelengths. Ordinate scale attenuation values are indicated on the spectra, 359× being the highest sensitivity used. Spectra have been vertically arranged for the purpose of clarity. Arrows point to wavelengths of interest. (a) Etiolated tissues; (b) after one LD cycle; (c) after two LD cycles; (d) after three LD cycles.
MV Pchlide a and Chlide a disappeared from the monocarboxylic phorbin pool which contained exclusively regenerated DV Pchlide a (Fig. l(c,d)). The lack of Chlide a was evidenced b y a fluorescence emission profile of the monocarboxylic phorbin pool that was identical to that of a fully etiolated tissue and which lacked the incremental fluorescence at 673--675 nm (Fig. l(a,d)). On the other hand, the absence of MV Pchlide a in this pool was obvious from its Sorer excitation profile which lacked a shoulder at 437 nm (Fig. 1B(d)). Altogether, the above results indicated that, after a third L pulse followed
32
b y a D incubation period, the monocarboxylic phorbin pool was made up o f DV Pchlide a and lacked any MV Pchlide a or Chlide a. It thus follows that the Chlide a generated by a fourth L pulse will have been formed exclusively from the regenerated DV Pchlide a pool. Photoconversion o f the induced D V Pchlide a pool into Chlide a (E459 F675). The photoconversion of the regenerated DV Pchlide a pool is contrasted with the photoconversion of the mixed MV/DV Pchlide a pools in Fig. 2. A single L pulse converted most of the mixed MV/DV Pchlide a pool of the etiolated tissue (emission maximum at 625 nm) into Chlide a, having an emission maximum at 674 nm (Figs. 1A(a) and 2A(a)). Although the newly formed Chlide a pool exhibited a single emission maximum at
F675
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Wavelength of Emission(nm] Wavelength of Excitation(nm)
Fig. 2. Fluorescence emission (A) and excitation (B) spectra at ether at 77 K of the Chlide a pools extracted from etiolated cucumber cotyledons after sequential 2.5 ms actinic light pulses (L) with intervening 60-min dark incubations. Emission spectra b--d were recorded on the Perkin-Elmer spectrofluorometer. The remaining spectra were recorded on the SLM spectrofluorometer. Arrows point to wavelengths of interest. The etiolated tissues were immediately frozen in liquid N 2 and their monocarboxylic phorbin pools extracted after (a) one actinic light pulse (L); (b) one LD cycle followed by one photoconverting flash; (c) two LD cycles followed by one photoconverting flash; (d) three LD cycles followed by one photoconverting flash. All other symbols are as in Fig. 1
33 a b o u t 674 nm, it exhibited a split Soret excitation profile with a major excitation m a x i m u m at a b o u t 448--449 nm and a shoulder at a b o u t 4 5 8 - 459 nm (Fig. 2B(a)). On the basis of the relative excitation amplitudes of the MV and DV Pchlide a Sorets at 437 and 443 nm respectively, and on the basis o f the amplitudes o f the corresponding Chlide a Sorets at 449 and 459 nm, t h e latter were previously assigned to MV and DV Chlide a respectively [ 4]. In this w o r k the precursor-product relationship o f MV Pchlide a to Chlide a (E449 F675) and that of DV Pchlide a to Chlide a (E459 F675) is demonstrated. As it m a y be recalled, the Pchlide a pool which was regenerated after the first light pulse was made up mainly of DV Pchlide a and of much smaller amounts of MV Pchlide a (Fig. 1B(b)). A L treatment of this regenerated Pchlide a pool converted most of the Pchlide a into Chlide a (Figs. 1A(b) and 2A(b)). The newly formed Chlide a pool was mainly made up of Chlide a (E459 F675) and of much smaller amounts of Chlide a (E449 F675) (Fig. 2B(b)). Actually, the latter appeared as a small excitation shoulder on t h e short wavelength tail of the Chlide a (E459 F675) Soret and its relative amplitude was commensurate with the relative amplitude o f the MV Pchlide a c o m p o n e n t that was present before photoconversion (cf. Figs. 1B(b) and Fig. 2B(b)). Likewise, the same relative relationship between the MV/DV Pchlide a components and the two Chlide a components was observed in etiolated tissues that were subjected to t w o LD cycles (Figs. 1B(c) and 2B(c)). These results strongly suggested a precursor-product relationship between MV Pchlide a (E437 F625) and Chlide a (E449 F675) and between DV Pchlide a (E443 F625) and Chlide a (E459 F675). The above relationship was further corroborated b y converting the DV Pchlide a pool which was regenerated after three LD cycles into Chlide a (E459 F675) by an L pulse (Fig. 2(d)). The absence of a MV Chlide a c o m p o n e n t is evident by the complete lack of a MV Soret excitation peak or shoulder at 448--449 nm (Fig. 2B(d)). Spectrofluorometric properties of the M V Chlide a and Chlide a (E459 F675) pools at room temperature. With the intermittent light/dark treatment o f etiolated tissues which was described above, it became possible to prepare Chlide a (E459 F675) containing very little MV Chlide a contamination and to compare its spectroscopic properties to those of MV Chlide a. A monocarboxylic phorbin pool made up mainly of MV Chlide a was extracted from etiolated cucumber cotyledons which had been illuminated for 2.5 ms with actinic white light then placed in darkness for 5 min before extraction. The L pulse converted most of the mixed MV/DV Pchlide a pool into a mixed Chlide a {E449 F675) + Chlide a (E459 F675) pool (Figs. l(a) and 2(a)). This monocarboxylic phorbin pool also contained small amounts o f non-phototransformable Pchlide a with an emission maximum at a b o u t 625 nm (Fig. 3A(a)). During the 5-rain post-illumination dark period all o f the Chlide a (E459 F675) was converted into blue-shifted (i.e. short wavelength) Chl a species of unkn own structure [ 1,4 ]. The remaining Chlide a
34
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Fig. 3. F l u o r e s c e n c e e m i s s i o n ( A ) a n d e x c i t a t i o n (B) s p e c t r a o f t h e M V Chlide a a n d Chlide a ( E 4 5 9 F 6 7 5 ) a n d o f t h e i r c o r r e s p o n d i n g p h e o p h o r b i d e s a p r e p a r e d f r o m lightt r e a t e d e t i o l a t e d c u c u m b e r c o t y l e d o n s . P i g m e n t s w h i c h w e r e m o n i t o r e d at 77 K were dissolved in e t h e r , while t h o s e m o n i t o r e d at r o o m t e m p e r a t u r e ( R T ) w e r e dissolved in h e x a n e - e x t r a e t e d 80% a c e t o n e (see Materials a n d M e t h o d s ) . S p e c t r u m f was r e c o r d e d o n t h e S L M s p e c t r o f l u o r o m e t e r , while t h e r e m a i n i n g s p e c t r a were r e c o r d e d o n t h e PerkinE l m e r f l u o r o m e t e r . A r r o w s p o i n t t o w a v e l e n g t h s o f interest. MV, refers t o t h e s t a n d a r d m o n o v i n y l Chlide a p o o l w h i l e D V refers t o t h e Chlide a ( E 4 5 9 F 6 7 5 ) p o o l b e f o r e a n d a f t e r d e m e t a l l a t i o n . All o t h e r s y m b o l s are as in Fig. 1.
35 pool (Fig. 3(a)) was then found to consist mainly of Chlide a (E446-447 F674) and of smaller amounts of blue-shifted, short wavelength Chlide a species [4]. By analogy to the MV Chl a {E446 F674) of mature green tissues [1], Chlide a {E446 F674) is here identified as MV Chlide a. A monocarboxylic phorbin pool made up mainly of Chlide a (E459 F675) was extracted from etiolated cucumber cotyledons subjected to three LD treatments, followed b y one 2.5-ms phototransforming light pulse (Fig. 3(b)). The latter converted most of the regenerated DV Pchlide a into Chlide a (E459 F675). This pool also contained some unphototransformed Pchlide a with an emission maximum at a b o u t 625 nm (Fig. 3A{b)}. The latter, did not, however, interfere with the recording of the spectrofluorometric properties of the Chlide a (E459 F675) because of the small amounts of Pchlide a present and of the insignificant contribution of its vibrational band at 673--675 nm to the red emission band of Chlide a. To insure that the spectrofluorometric differences between the standard MV Chlide a pool and Chlide a (E459 F675) were not artifacts associated with low temperature spectroscopy, a comparison was made between the fluorescence spectra of the two Chlide a species recorded at r o o m temperature in h e x a n e ~ x t r a c t e d acetone. Both pools exhibited identical emission maxima at a b o u t 675 nm (Fig. 3A(d,e}}. The much smaller emission at a b o u t 639 nm is that of the non-phototransformed Pchlide a pool (Fig. 3A(d,e}). The Soret excitation maxima of the two Chlide a pools were, however, quite different. While the standard MV Chlide a pool exhibited a ~oret excitation maximum at about 434 nm, the Chlide a (E459 F675) Soret excitation maximum was red-shifted b y about 8 nm and was observed at 442 nm (Fig. 3B(d,e)). These results indicated that the spectrofluorometric differences observed between the Soret excitation maxima of the two Chlide a pools were not due to low temperature artifacts but were most probably related to chemical differences between the two Chlide a species. Spectrofluorometric properties o f the two Chlide a species after demetallation. If the two Chlide a species do in fact differ from each other b y virtue o f chemical m o d i f i c a t i o n s ) of the porphyrin macrocycle, then removal of the central Mg atom from these two compounds should generate two spectrally different pheophorbide species. The t w o Chlide a pools were therefore demetallated and the fluorescence properties o f the two resulting pheophorbide a species were compared. The spectra were recorded at 77 K in ether. The two pheophorbide a species exhibited different fluorescence excitation maxima (Fig. 3(f,g)}. While both MV and DV pheophorbide a exhibited an emission maximum at 667 nm (Fig. 3A(f,g)}, the Soret excitation maximum of the MV pheophorbide a was observed at 415 nm, while that of the other pheophorbide a species was observed at 424 nm {Fig. 3B{f,g)). Altogether, the above results indicated that the spectrofluorometric differences b e t w e e n the t w o Chlide a species were preserved after the removal o f the central Mg atom.
36
Purification of Chlide a (E459 F675). Since further chemical characterization o f Chlide a (E459 F675) m a y require the availability o f pure quantities of this Chlide a species, it was deemed desirable to purify Chlide a (E459 F675) from the small amounts of non-phototransformed Pchlide a that were also present in the monocarboxylic phorbin pool of the lighttreated tissues. Several chromatographic systems were tested; chromatography on thin layers o f polyethylene developed in acetone/H20 (9 : 1, v/v) at 4°C gave the best results and achieved a complete separation of Chlide a (E459 F675) from Pchlide a. In this system Pchlide a migrated with an Rf-value of a b o u t 0.44, while Chlide a (E459 F675) migrated with an Rf-value of about 0.9. It was essential to carry o u t the separation at a b o u t 4°C in order to avoid demetallation o f Chlide a (E459 F675). Indeed, it was constantly observed throughout this work that this Chlide a species was much more susceptible to demetallation than MV Chlide a. Furthermore, it has been found that prepurification of the monocarboxylic phorbin pool on thin layers of silica gel H developed in toluene/ethyl acetate/ethanol (8 : 2 : 2, v/v/v) at 4°C improves the performance of the polyethylene separation, presumably by removing colorless lipids [4]; Belanger and Rebeiz, unpublished]. The fluorescence properties in ether at 77 K of purified Chlide a (E459 F675) were identical to those of the Chlide a (E459 F675) enriched monocarboxylic phorbin pool except for the absence of a Pchlide a emission at 625 nm (Fig. 3(c)). DISCUSSION Chlide a (E459 F675) was first observed by Belanger and Rebeiz [4] immediately after the photoconversion of the mixed MV Pchlide a (E437 F625)/DV Pchlide a (E443 F625) pool of etiolated tissues into a mixed Chlide a (E449 F675)/Chlide a (E459 F675) pool. On the basis of (a) the briefness of the light treatment (47 ms) and (b) assuming that the short wavelength MV Pchlide a is likely to yield a correspondingly short wavelength Chlide a, while the long wavelength DV Pchlide a is likely to yield a long wavelength Chlide a, the newly formed Chlide a (E449 F675) was tentatively identified as MV Chlide a, and Chlide a (E459 F675) was identified as DV Chlide a. The results reported in this work seem to corroborate the above assignment (vide infra). In order to minimize the occurrence of photochemical side reactions that m a y m o d i f y the structure of the reacting tetrapyrroles, the photoconversion time was reduced in this work from 47 ms to 2.5 ms. Within about 250 ms after the light treatment, all enzymatic activity was stopped b y freezing the tissue in liquid nitrogen, then the pigment was extracted in organic solvents. By assuming that during the brief period that preceded the extraction of the , pigment only photochemistry was operative while biochemical activity was ! neglible, one is led to the conclusion that following the light treatment
37 MV Pchlide a did indeed give rise to Chlide a (E449 F675), while DV Pchlide a gave rise t o Chlide a (E459 F675) as previously proposed [4]. This relationship was further supported by (a) the observation that DV Pchlide a (E443 F625) devoid of MV Pchlide a (E437 F625) contamination was converted by p h o t o r e d u c t i o n into Chlide a (E459 F675) devoid of Chlide a (E449 F675) (Figs. l ( d ) and 2(d)) and (b) by the finding that essentially the same results were obtained by freezing the cotyledons to - 1 0 ° C prior to the light treatment. The consideration of Chlide a (E459 F675) as a Chlide a species chemically distinct from MV Chlide a (E446 F674) is corroborated by its spectrofluorometric properties which differ from those of MV Chlide a (Fig. 3) (a) at 77 K and at room temperature, (b) before and after demetallation and (c) after purification. The unambiguous determination of the structure of Chlide (E459 F675) will require extensive chemical and spectroscopic studies. Such studies will require the preparation of large quantities of this c o m p o u n d in a highly purified state. The techniques described herein are presently being employed to further pursue the chemical identity of this novel Chl precursor. REFERENCES 1 C.A. Rebeiz, F.C. Belanger, G. Freyssinetand D.G. Saab, Biochim. Biophys. Acta,
590 (1980) 234. 2 G. Freyssinet, C.A. Rebeiz, J.M. Fenton, R. Khanna and Govindgee, Photobiochem. Photobiophys., 1 (1980) 203. 3 C.A. Rebeiz, F.C. Belanger, S.A. McCarthy, G. Freyssinet, J.X. Duggan, S.M. Wu and J.R. Mattheis, in: Proc. 5th Int. Cong. Photosynth., 5 (1981) 197. 4 F.C. Belanger and C.A. Rebeiz, Plant Sci. Lett., 18 (1980) 343. 5 F.C. Belanger and C.A. Rebeiz, J. Biol. Chem., 255 (1980) 1266. 6 S.I. Hardy, P.A. Castelfranco and C.A. Rebeiz, Plant Physiol., 46 (1970) 705. 7 C.A. Rebeiz, J.R. Mattheis, B.B. Smith, C.C. Rebeiz and D.F. Dayton, Arch. Biochem. Biophys., 171 (1975) 549. 8 C.A. Rebeiz, J.R. Mattheis, B.B. Smith, C.C. Rebeiz and D.F. Dayton, Arch. Biochem. Biophys., 166 (1975) 446. 9 C.E. Cohen and C.A. Rebeiz, Plant Physiol., 61 (1978) 824. 10 C. Houssier and K. Sauer, Biochim. Biophys. Acta, 172 (1969) 492.