Chlorophyll a Biosynthetic Pathway

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VI. The Chl a Carboxylic Biosynthetic Routes: (Photo)Conversion of Protochlorophyllides (Pchlides) a to Chlorophyllide (Chlide) a

The reactions that will be discussed are shown in more details in Fig. 1. .
DV Chlide a
MV Chlide a
DV and MV Chlides a are the main immediate precursors of Chl a. They are formed via multiple light-dependent and light-independent biosynthetic routes from Pchlide a (Fig. 1). In all cases, the reaction involves reduction of the double bond at position 7-8 of the macrocycle by addition of two trans-hydrogens. Most of the investigations of the photoreduction of Pchlide a have dealt with t-LW-Pchlide a H (E650 F657) ( see section VI for terminology). The latter is a ternary complex of Pchlide a, NADPH and Pchlide a oxidoreductase, a shuttling photoenzyme.

The notion that the t-LW-Pchl(ide) a H apoprotein acts as a shuttling photoenzyme that catalyzes the conversion of Pchlide a to Chlide a was first proposed by Sironval et al (1967). In this work the authors report that Pchl(ide) a (E647 F 657), with a red excitation maximum at 647 nm and a red emission maximum at 657 nm, is photoconverted to Chlide a (E676 F690). The latter shifts in darkness to a Chlide a (E682 F697) species. At this stage, the authors suggest that the apoprotein discharges the newly formed Chlide a and picks up another Pchlide a which may be photoconverted to Chlide a via a similar cycle. The spectral shifts described by Sironval et al were confirmed by Gassman et al (1968), and Bonner (1969). The concept of shuttling photoenzyme was also compatible with the photoconversion of photoinactive Pchlide a 633 to phototransformable Pchlide a 650 reported by Gassman (1973).

Also discussed will be the light-independent conversion of Pchlide a to Chlide a. Although light-independent Pchlide a reduction has been known to occur in gymnosperms its occurrence in angiosperms is novel (Adamson and Packer, 1984).

A. Kinetics of the photoconversion of the Pchlide a Chromophore of Pchlide a-H (E650 F657) to Chlide a

1. Action Spectrum of the Photoconversion

Pchl(ide) a H (E650 F657) is the photoreceptor for its own photoconversion to Chlide a (Koski and Smith, 1951). In an albino corn mutant lacking carotenoids, the action spectrum exhibited two prominent peaks, one at 650 nm and one at 445 nm that corresponded to the absorption spectrum of LW t-Pchlide a H of the mutant.

2. Effect of Temperature on the Photoconversion

The phototransformation of LW t-Pchlide a H to Chlide a was completely inhibited at -195 C (Smith and Benitez, 1954). Partial photoconversion took place at -70 C. At temperatures beyond 50 C, photoconversion was progressively inhibited due to apoprotein denaturation. Dependency of the photoconversion upon temperature indicates that the phototransformation is not a purely photochemical reaction but also involves a thermochemical component.

3. Quantum Yield of the Photoconversion

The average quantum yield of the photoconversion at 642 nm amounted to about 0.6 (Smith and Benitez, 1954). Therefore it was not clear from this work whether one or two quanta of light are required for the photoconversion. On the other hands, Thorne (1971) proposed a two quantum process for the photoconversion.

4. Effect of Environment on the Photoconversion

The rate of photoconversion expressed as a percentage of the photoconvertible protochlorophyllide was found to be independent of the initial concentration of the holochrome and was not influenced bu the viscosity of the medium (Boardman, 1962). This led to the proposal that the photoconversion did not involve a collision process between independent protein molecules or between a protein molecule and a hydrogen donor molecule. Instead, Boardman (1962) proposed a restricted collision process between the photo activated Pchl molecule and the hydrogen donor. However since the rate of phototransformation was temperature-dependent, it seemed likely that the hydrogenation involved some vibrational or rotational movement of that part of the protein molecule in close proximity to the Pchlide a chromophore. 5.

5. Photoconversion Kinetics

While Smith and Benitez (1954), opted for a bimolecular reaction with respect to Pchlide a, Thorne and Boardman (1972) suggested that by allowing for energy transfer within molecular groups, the true kinetics of the photoconversion was first order, which is still compatible with the restricted collision hypothesis (Boardman, 1962).

B. Biosynthetic Heterogeneity of Chlorophyllide a Formation

In Fig. 1, two pools of DV Chlide a formed via routes 1, and 13, four pools of MV Chlide a formed via light-dependent routes 2, 3, 10 and 12, three MV Chlide a pools formed by 4-vinyl reduction of DV Chlide a via routes 4, 8, and 13, and two MV Chlide a pools formed via light-independent routes 3D and 15D are depicted. In other words, Chlide a biosynthetic heterogeneity involves formation of Chlide a via photoreduction of DV Pchlide a (routes 1, 8 and 13), via photoreduction of MV Pchlide a (routes 2, 3, 10 and 12), via 4-vinyl reduction of DV Chlide a (routes 4, 8 and 13), and via light-independent conversion of Pchlide a to Chlide a (routes 3D and 15D). These various biosynthetic routes and their relationships to LW and SW Pchlide a H photoconversion (see Section V for terminology) are discussed below.

1.Formation of Chlide a via Light-Independent Pchlide a Reductase(s) via Routes 3D and 15D
Algae, ferns, mosses, and the cotyledons of most gymnosperms, all with a DDV-LDV-LDDV greening group affiliation (Ioannides et al, 1994) (also see section on botanical fall-outs) are capable of converting Pchlide a to Chlide a in the absence of light (Kirk and Tilney Basset, 1967; Ryberg and Sundqvist, 1991; Shulz and Senger, 1993), via a reaction catalyzed by a light-independent Pchlide a reductase (Fig. 1, route 3D). Most probably light-independent Chlide a formation is via a nt-SW-Pchlide a species (Schoefs and Franck, 1998).

Although in angiosperms light is required for the formation of photosynthetic pigment-protein complexes and the accumulation of massive amounts of Chl, Adamson and coworkers pioneered the notion that in this phylum, a certain amount of Chl a biosynthesis can also take place in darkness via a light-independent Pchlide a reductase (Adamson et al, 1997). Dark-incorporation of 14C-glutamate and 14C-ALA into 14C-Chl a in barley leaves and barley etiochloroplasts appear to confirm Adamsonís contention (Tripathy and Rebeiz, 1987). It is therefore proposed that biosynthetic routes 3D and 14 D (Fig. 1) are also functional in angiosperms such as cucumber and barley. It should be emphasized however that the amount of Chl a formed via Chlide a in darkness is very small, and its biological significance is unknown.

Genetic and sequence analysis have indicated that in R. capsulatus, three genes, bchL, bchN, and bchB appear to be involved in Pchlide a reduction in darkness (Suzuki et al, 1997). The three open frames exhibited significant sequence similarity to the three subunits of nitrogenase, which led to the proposal that light-independent Pchlide a reductase and nitrogenase share a common evolutionary ancestor. Expression of the bchL, bchN, and bchB genes has been however unsuccessful. Very recently, Yuichi and Bauer reported demonstration of dark-Pchlide a reductase activity in reconstituted systems from R. capsulatus, a purple nonsulfur photosynthetic bacterium (Yuichi and Bauer, 2000). Two of the putative three subunits, BchL and BchN were expressed in R. capsulatus as S tag fusion proteins. The third subunit, BchB, copurified with the BchN protein, thus indicating that the BchN and BchB proteins form a tight complex. Dark Pchlide a reductase activity was shown to be dependent on the presence of all three subunits, on ATP, and on the reductant dithionite. In angiosperms, the corresponding gene products ChlL, ChlN, and ChlB, also appear to be evolutionarily related to the subunits of the eubacterial nitrogenase enzyme complex (Armstrong , 1998).

2. Formation of Chlide a via Light-Dependent Pchlide a Oxidoreductases (PORs)
It has been proposed that at least four different POR isozymes may be present in plants (Ikeuchi and Murakami, 1982; Dehesh et al, 1986). In Arabidopsis thaliana and Barley, two different genes with about 75 % homology, PorA and PorB, have been shown to code for two different Pchlide a oxidoreductases, namely PORA and PORB (Armstrong et al, 1995; Holtorf et al, 1995). PORA is synthesized in the dark and constitutes the bulk of the crystalline prolamellar body of etioplasts. However the transcription of its gene is turned off in the light and the enzyme is rapidly degraded by a light-induced protease (Reinbothe et al, 1999; Reinbothe et al, 1995). On the other hand, the PorB gene is transcribed in darkness and in the light, and the transcripts are translated continuously into the enzyme which is responsible for the bulk of Chl a biosynthesis and accumulation in daylight. More recently, a gene that encodes a third POR in Arabidopsis thaliana has been reported (Oosawa et al, 2000). the Protein has been named PORC. PORC is expressed only during the light phase of the photoperiod. Since PORA, B and C respond to light differently (see below), it has been suggested tht the function of the three PORs of Arabidopsis are not redundant, but may allow the plant to adapt its needs for Chl biosynthesis according to the prevailing light regime (Su et al, 2001). In our opinion, adaptation of Chl biosynthesis to different light conditions, proceeds via multiple and different biosynthetic routes.

a. NADPH-Protochlorophyllide a (Photo)oxidoreductase A (PORA, or PCR)

As pointed out above, most of the early investigations of the photoreduction of Pchlide a dealt with t-LW-Pchlide H (E650 F657). The notion that the Pchlide a H apoprotein of t-LW-Pchlide a H (E650 F657) (i.e. PORA) acts as a shuttling photoenzyme that catalyzes the conversion of Pchlide a to Chlide a was first proposed by Sironval et al (Sironval et al, 1967). In this work the authors reported that t-LW-Pchlide a (E647 F 657), with a red excitation maximum at 647 nm and a red emission maximum at 657 nm, is photoconverted by light to Chlide a (E676 F690). The latter is converted in darkness to Chlide a (E682 F697). At this stage of the reaction, the authors suggest that the apoprotein discharges the newly formed Chlide a (E682 F697) and picks up another Pchlide a chromophore that may be photoconverted to Chlide a via a similar cycle. The spectral shifts described by Sironval et al were confirmed by Gassman et al (Gassman et al, 1968), and Bonner (Bonner, 1969). The concept of a shuttling Pchlide a reductase photoenzyme was also compatible with the reported conversion of nt-SW-Pchlide a (F633) to t-LW-Pchlide a (F650) during the photoreduction process (Gassman, 1973).

A significant step in the understanding of Pchlide a photoreduction was achieved with the realization that NADPH is the hydrogen donor for the reaction ( Griffiths, 1974). This was followed by the proposal that the shuttling photoenzyme (POR, now called PORA), NADPH, and Pchlide a formed a photoactive ternary Pchlide a-NADPH-enzyme complex with a red absorption maximum at 652 nm (Apel et al, 1980). Equally important was the purification of PORA from etiolated barley (Apel, 1981). The purified enzyme consisted of one polypeptide (Mr 36000) with two to three bound Pchlide a chromophores. It is synthesized in the cytoplasm as a precursor protein of about 44 kDa. The transit sequence of about 8 kDa, is hydrolyzed when the enzyme is transported into the plastid (Apel, 1981). The size of PORA reported by various authors depends on the plant species and varies from 33 to 38 kDa (Shulz and Senger, 1993). More recently, pigment-free PORA was purified from barley etioplasts by solubilization with n-octyl-B-D-glucoside and chromatography on DEAE-cellulose (Klement et al, 1999). Using pigment and protein analysis it was shown that barley etioplasts contained a one-to-one PORA and Pchlide a. The enzyme was twice as active towards MV than toward DV Pchlide a Klement et al, 1999.

It has also been demonstrated that during the greening of etiolated tissues a rapid decline of PORA is observed. For example, after six hours of continuous illumination, when the rate of Chl a accumulation is at its peak, only traces of the PORA protein are detected (Santel and Apel, 1981). The disappearance of PORA from etiolated tissues during greening was confirmed by Kay and Griffiths (Kay and Griffiths, 1983). These observation and further experimentation have led to the proposal that in etiolated tissues, although PORA functions only for a short period of time after the onset illumination, it is required for normal greening (Rung et al, 1996).

b. Protochlorophyllide a oxidoreductase B (PORB)

It has been proposed that PORB provides the means to sustain light-dependent Chl biosynthesis in fully greened mature plants, in the absence of PORA and t-LW-Pchlide a H (E450 F655) ( Runge et al, 1996). In other words, it was suggested that in some t-SW-Pchlide a H species, the apoprotein consists of PORB.

The photoreduction of Pchlide a by purified PORB overexpressed heterologously in E. coli has recently been described (Lebedev and Timko, 1999). The PORB reaction is described as consisting of two steps. In a first photochemical step, a single quantum mechanism leads to the formation of an unstable tetrapyrrole intermediate with a putative free electron. In a second step, the free radical intermediate is spontaneously converted to Chlide a. Both steps appear to proceed at subzero temperatures. At room temperature, the rate of the reaction depends linearly on enzyme and substrate concentrations, and the reduction kinetics are consistent with one mole of substrate bound per active PORB monomer.

c. Protochlorophyllide a Photooxidoreductase C (PORC)

Like PORA and PORB, PORC is light-and-NADPH-dependent. In contrast to the PORA and PORB mRNAs, the PORC mRNA accumulates only after the beginning of illumination (Osawa et al, 2000). In light-dadpted mature plants only PORB and PORC mRNAs were detectable, and the amounts of both mRNAs exhibited pronounced diurnal rhytmic fluctuations (Su et al, 2001). However, differences were observed between PORB and PORC. The differences can be summarized as follows: (a) While the oscillations of PORB mRNA are under the control of the circadian clock, that of PORC is not, (b) Upon transfering to darkness seedlings grown under continuous white light, the concentration of PORC mRNA rapidly declined and became undetectable, while PORB mRNA did not, (c) When seedlings were exposed to different light intensities, the amounts of PORB mRNA remained the same, while the mRNAs of PORA and PORC were modulated in an inverse way by light intensity changes.

3. Contribution of t-LW-Pchlide a (PORA) and t-SW-Pchlide a (PORB) to Photoperiodic Greening

Under natural photoperiodic greening conditions, Pchlide a accumulates during the dark cycles of the photoperiod and contributes to Chl a biosynthesis and accumulation at the onset of light (i. e. at dawn) (Cohen et al, 1977). Furthermore, Pchlide a is always present in green tissues during the light phases of the photoperiod (Carey and Rebeiz, 1985; Abd-El-Mageed et al, 1997; Cohen et al, 1977). During photoperiodic greening Pchlide a (E650 F655), also known as t-LW-Pchlide a H [(E450 F657), and its apoprotein, PORA, are transient species that peak during the 7th dark cycle and become undetectable by the 11th dark cycle (Cohen and Rebeiz, 1978). On the other hand SW Pchl(ide) a H species and their apoproteins, PORB and/or PORC are present throughout the photoperiodic greening process (Cohen and Rebeiz, 1978). In other words although t-LW-Pchlide a H (E450 F657) contributes to prolamellar body reformation and Chl a formation during the first few dark cycles, it is SW t-Pchlides a Hs and PORA/PORC that prevail during the light cycles of photoperiodic greening, and contribute the bulk of Chl a accumulation under normal field conditions (Cohen and Rebeiz, 1978).

The significance of the accumulation patterns of t-SW Pchl(ide) a Hs , and PORB/C during photoperiodic greening, to the Chl a biosynthetic process, rests upon the direct photoconvertibility of t-SW Pchlide a Hs to Chlide a without prior conversion to t-LW-Pchl(ide) a (E450 F657) and subsequent photoreduction by PORA. This was reported to be the case by Cohen and Rebeiz (Cohen and Rebeiz, 1978). However, the photoconversion of t-SW-Pchlide a H, was slower than that of t-LW-Pchlide a H (E450 F657) which is catalyzed by PORA.

In conclusion, since etiolation and prolamellar body formation are not abnormal phenomena, but are part of the natural succession of the dark (night) and light (daylight) cycles during photoperiodic greening (Cohen and Rebeiz, 1978; Rebeiz and Rebeiz, 1986), it is very plausible for PORA and PORB/C, to play definite but distinct roles during Chl a biosynthesis and accumulation under natural field conditions. If it is assumed that routes 1, 2, 3, 12 and 13 prevail in darkness while routes 8, 10 prevail in the light, it becomes logical to propose that PORA prevails in routes 1, 2, 3, 12 and 13, while PORB/C prevail in routes 8 and 10 (Fig. 1).

C. Photoreduction Intermediates and Spectral Shifts During Photoreduction of Protochlorophyll(ide) a H (E550 F655)


Fig. 6. Spectral Shifts of the Ternary Pchlide a-NADPH-Apoprotein Complex During the Photoconversion of Pchlide a to Chl(ide) a. "E" and "F" refer to the red fluorescence excitation and emission maxima respectively at 77 K.

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In etiolated tissues, the photoreduction of t-LW-Pchlide a H (E550 F655) by PORA is accompanied by complex spectral shifts (Fig. 6) of intermediates and end products that eventually result in the conversion of the crystalline prolamellar body and prothylakoids to thylakoid membranes. It appears that some of the spectral shifts may be regulated by protein phosphorylation (Wiktorsson et al, 1996).At the present stage, the significance of the postillumination spectral shifts to Chl biosynthetic heterogeneity is unclear.

1. Spectral Shift I
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Spectral shift I is light-dependent. It was reported by Thorne (1971) in etiolated bean leaves. It occurs as a result of fractional photoconversion of LW t-Pchlide a H (E650 F655), to a dark-stable pigment-apoprotein complex (E668 F674), with 77 K red excitation and maxima at 668 and 674 nm , respectively. This intermediate yields a mixture of Pchlide and Chlide a after dark-ethanol extraction. The photoconversion rate for Chlide a H (E668 F674) was twice the rate for the photoconversion of the next intermediate, thus suggesting that, in vivo, photoconversion of Pchlide a to Chlide a is a two step two photon process.

2. Spectral Shift II
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Light-dependent spectral shift II was first described by Sironval et al (1967) as a photoconversion of t-LW-Pchlide a (E650 F655) to a Chlide a (E676 F690)-apoprotein complex. Later on, that Pchlide a-protein complex was referred to as Chlide a (E676 F688) by Sironval and Kuyper (1972)]. Then in 1971, Thorne reported that the photoprecursor of Chlide a (E676 F690) was Chlide a (E668 F674) instead of t-LW-Pchlide a (E650 F655). The chemical nature of the chromophore in Chlide a (E668 F674) is not clear however. Sironval et al (1967), and Sironval and Kuyper (1972) initially proposed that it was some kind of Pchlide a-Chlide a intermediate of an ill defined nature. Thorne (1971), proposed however, that the chromophore consisted exclusively of Chlide a. Shift II was also confirmed by Gassman et al (1968), and by Bonner (1969).

3. Spectral Shift III
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Shift III, which converts Chlide a (E676 F690) to Chlide a (E682 F697) takes place very rapidly in darkness. It was considered by Sironval et al (1967) to lead to the formation of a mature Chlide a-apoprotein complex that releases Chlide a from the Pchlide a oxidoreductase complex. This in turn allows PORA to pick up another Pchlide a chromophore to yield t-LW-Pchlide a-H (E650 F655).

4. Spectral Shift IV
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The conversion of Chlide a (E682 F697) to Chl(ide) a (E672 F680) species was the first spectral shift to be described during the conversion of Pchlide a to Chl(ide) a. It was reported by Shibata in 1957, as a spectral shift that took place in darkness or in the light in about 10 to 20 minutes after the onset of illumination, depending on the age of the etiolated tissue, and the plant species. During this shift Chlide a is esterified with geranylgeraniol, which is reduced stepwise to phytol (see section dealing with the reactions between Chlide a and Chl a).

5. Spectral Shift V
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The fifth shift was also described by Shibata (1957). It takes place either in the light or in darkness, and corresponds to the final integration of Chl a into various pigment proteins of the thylakoid membranes. On the basis of energy transfer from Pchlide a (E650 F655) to Chlide a (E682 F697) at fractional or partial photoconversions, Thorne (1971) concluded that diffusion of the chromophore from the apoprotein occurs at the level of Chl(ide) a (E674 F683) (shift V) instead of Chlide a (E682 F697) as proposed by Sironval et al (1967). From the maximal appearance of the (E668 F6740) photointermediate and the Pchlide a and Chlide a content Thorne (1971) proposed that the Pchlide a aggregate comprised about 20 molecules of Pchlide a.

D. Formation of DV Chlide a by Photoconversion of DV Pchlide a to DV Chlide a

In higher plants, DV Chlide a is a transient yet important metabolic intermediate. It gives rise to MV Chlide a via routes 4, 8 and 13; to DV Chl a via route 1; and to DV Chlide b via route 6.

1. Formation of DV Chlide a by Photoconversion of DV Pchlide a via Route 1 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Greening DDV-LDV-LDDV Plants during the Initial Dark Phase of the Photoperiod

In DDV-LDV-LDDV etiolated tissues, a precursor-product relationship between DV Pchlide a and DV Chlide a was established by demonstrating the photoconversion of DV Pchlide a to DV Chlide a in etiolated cucumber cotyledons induced to accumulate DV Pchlide a exclusively (Duggan and Rebeiz, 1982). The photoreduction is most probably catalyzed by PORA. Biosynthesis of DV Chlide a via biosynthetic route 1 also takes place in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when t-LW-Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977). In the Nec 7 corn mutant (Brereton et al, 1983), and in the picoplankton of subtropical waters which accumulate DV Chl a and b (Chisholm et al, 1988; Chisholm et al, 1992; Goereke and Repeta, 1992; Veldhuis, et al, 1990), DV Chlide a accumulates in substantial amounts and is most probably formed via route 1 at the beginning of the light phases of the photoperiods.

2. Formation of DV Chlide a by Photoconversion of DV Pchlide a via Route 8 in DDV-LDV-LDDV Plants during the Light and Dark Phases of the Photoperiod

As mentioned above, light-grown DDV-LDV-LDDV plants form most of their Chl via DV Pchlide a as evidenced by the accumulation of regenerated DV Pchlide a upon interrupting the light cycle by a brief dark treatment (Abd-El-Mageed et al, 1997). We now propose that in the light, nascent t-SW-DV Pchlide a H is photoconverted to DV Chlide a by the action of PORB, and the transient DV Chlide a is converted to MV Chlide a by 4VCR as depicted in route 8. Biosynthetic route 8 is also most probably involved in the formation of DV Chlide a in the Nec 7 corn mutant (Brereton et al, 1983), and in the picoplankton of subtropical waters which accumulate DV Chl a and b (Chisholm et al, 1988; Chisholm et al, 1992; Goereke and Repeta, 1992; Veldhuis, et al, 1990) in the light. It should be stressed that although the Nec 7 corn mutant is a lethal mutant it does accumulate substantial amounts of DV Chlide a and b and DV Chl a and b in the light.

3. Formation of DV Chlide a by Photoconversion of DV Pchlide a via Route 13 in Etiolated DMV-LDV-LDMV Plants in Darkness, and in Greening DMV-LDV-LDMV Plants during the Initial Dark Phase of the Photoperiod

In DMV-LDV-LDMV etiolated tissues, a precursor-product relationship between DV Pchlide a and DV Chlide a was established by demonstrating the photoconversion of DV Pchlide a to DV Chlide a in etiolated corn leaves induced to accumulate significant amounts of DV Pchlide a in addition to MV Pchlide (Belanger and Rebeiz, 1982). The photoreduction is most probably catalyzed by PORA. Biosynthesis of DV Chlide a via biosynthetic route 13 probably also takes place in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when t-LW-Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977).

E. Formation of MV Chlide a by Photoconversion of the MV Pchlide a Chromophore to MV Chlide a

MV Chlide a can be formed from MV Pchlide a by photoreduction, via four distinct routes namely via routes 2,3, 10 and 12 (Fig. 1). These four biosynthetic routes are discussed below.

1. Formation of MV Chlide a by Photoconversion of MV Pchlide a via Routes 2 and 3 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Greening DDV-LDV-LDDV Plants during Initial the Dark Phases of the Photoperiod

In DDV-LDV-LDDV etiolated tissues, or in greening DDV-LDV-LDDV tissues during the first few dark-cycles of the photoperiod small amounts of MV Pchlide a are formed from MV Mpe via route 2 (Tripathy and Rebeiz, 1986), or by 4-vinyl reduction of DV Pchlide a during prolonged dark-incubations, via route 3 (Tripathy and Rebeiz, 1988). Such t-LW-MV Pchlides a are readily photoconvertible to MV Chlide a by PORA (Belanger et al, 1982; Belanger and Rebeiz, 1980). Biosynthesis of MV Chlide a via biosynthetic routes 2, and 3 also takes place in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when t-LW-Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977).

2. Formation of MV Chlide a by Photoconversion of MV Pchlide a via Route 10 in Greening DMV-LDV-LDMV Plants during the Light Phases of the Photoperiod

During the light phases of the photoperiod, in DMV-LDV-LDMV plant species such as corn wheat and barley, MV Pchlide a can be formed from DV Pchlide a via route 10, which involves reduction of the vinyl group of DV Pchlide a at position 4 of the macrocycle to ethyl, a reaction catalyzed by 4VPideR (Tripathy and Rebeiz, 1988). The nascent MV Pchlide a can then be rapidly photoconverted to MV Chlide a most probably by PORB which predominates in the light. The latter is active in green tissues, and is assigned to route 10 on the basis of the continuous detection of DV Pchlide a in the light, which in DMV-LDV-LDMV plant species is rapidly converted to MV Pchlide a and MV Chlide a (Abd-El-Mageed et al, 1997).

3. Formation of MV Chlide a by Photoconversion of MV Pchlide a via Route 12 in Etiolated DMV-LDV-LDMV Plants in Darkness, and in Greening DMV-LDV-LDMV Plants during the Initial Dark Phases of the Photoperiod

Etiolated DMV-LDV-LDMV tissues accumulate massive amounts of MV Pchlide a in darkness and form most of their Chl via regenerated MV Pchlide a during the dark and light cycles of the photoperiod (Carey and Rebeiz, 1985; Abd-El-Mageed et al, 1997; Tripathy and Rebeiz, 1986). In etiolated barley for example, the resulting MV Pchlide a pool can be rapidly photoconverted to MV Chlide a ( Belanger et al, 1982). In etiolated tissues and during the initial dark phases of the photoperiod when t-LW-Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977), the photoreduction of MV Pchlide a is most probably catalyzed by PORA which predominates in etiolated tissues.

F. Formation of MV Chlide a by Vinyl Reduction of DV Chlide a via Routes 4 and 8

In DDV-LDV-LDDV etiolated tissues, or in green DDV-LDV-LDDV tissues during the initial dark-cycles of the photoperiod, MV Chlide a is formed by reduction of the vinyl group of DV Chlide a at position 4 of the macrocycle to ethyl via routes 4 and 8. The reduction of the vinyl group of DV Chlide a to ethyl is catalyzed in darkness and in the light by a very potent enzyme, 4VCR (Parham and Rebeiz, 1992; Duggan and Rebeiz, 1982). 4VCR is a fast acting membrane-bound, NADPH-dependent enzyme ( Parham and Rebeiz, 1992; 1995). It is a stable enzyme that has been solubilized and purified about 20 fold ( Kolossov and Rebeiz, 2001).

1. Formation of MV Chlide a by Vinyl Reduction of DV Chlide a via Route 4 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Greening DDV-LDV-LDDV Plants during the Initial Dark Phases of the Photoperiod

The most spectacular manifestation of MV Chlide a formation via route 4 is in DDV-LDV-LDDV etiolated tissues induced to accumulate DV Pchlide a exclusively, by successive dark-light treatments (Duggan and Rebeiz, 1982). The accumulated DV Pchlide a is photoconverted by a short light flash to DV Chlide a and the latter is very rapidly converted to MV Chlide a in vivo and in organello (Duggan and Rebeiz (1982b). The photoreduction is most probably catalyzed by PORA which predominates in etiolated tissues. Biosynthesis of DV Chlide a via biosynthetic route 4 is likely to take place also in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when t-LW-Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977).

2. Formation of MV Chlide a by Vinyl Reduction of DV Chlide a via Route 8 in Greening DDV-LDV-LDDV Plants during the Light Phases of the Photoperiod

Formation of MV Chlide a by rapid vinyl reduction of transient DV Chlide a via route 8, depends upon the detection of 4VCR activity in photoperiodically grown green tissues, which has been reently documented by Kolossov and Rebeiz (in Preparatrion).the occurrence of 4VCR activity in greening cucumber cotyledons after 4 h of illumination has also been documented (Abd-El-Mageed et al, 1997).

3. Formation of MV Chlide a by Vinyl Reduction of DV Chlide a via Route 13 in Greening DMV-LDV-LDMV Plants during the Light Phases of the Photoperiod

Formation of MV Chlide a by rapid vinyl reduction of transient DV Chlide a via route 13, depends upon the detection of 4VCR activity in photoperiodically grown green tissues, which has been reently documented by Kolossov and Rebeiz (in Preparatrion). Its occurrence in barley etiochloroplasts after 4 h of illumination has been documented (Kolossovv and Rebeiz, 2001).

References

  1. I. M. Ioannides, I. M., Fasoula, D. M., Robertson. K. R., and Rebeiz, C. A. (1994) An evolutionary study of chlorophyll biosynthetic heterogeneity in green plants Biochem. Sys. Ecol. 22, 211-220.
  2. Kirk, J. T. O and Tilney-Basset R. A. E.(1967) The Plastids: Their Chemistry, Structure, Growth, and Inheritance. Freeman, London., pp 504-506.
  3. Ryberg M. R.and Sundqvist, C. (1991) Structural and functional significance of pigment-protein complexes of chlorophyll precursors. In Chlorophylls, edited by H. Cheer (CRC press, Boca Raton, Florida), pp. 587-612.
  4. Shulz R. and Senger, H. (1993) Protochlorophyllide Reductase: A key enzyme in the greening process. In: Pigment-Protein Complexes in Plastids: Synthesis and assembly., (edited by C. Sundqvist and M. Ryberg) Academic Press, New York., pp. 179-218.
  5. Schoefs B. and Franck, F. (1998) Chlorophyll synthesis in dark-grown pine primary needles. Plant Physiol. 118, 1159-1168.
  6. Adamson, H. Y., Hiller, H. J., and Walmsley, J. (1997) J. Photochem. Photophys. 41, 201-221.
  7. Tripathy B. C. and C. A. Rebeiz, C. A. (1987) Non-equivalence of glutamic acid and delta-aminolevulinic acid as substrates for protochlorophyllide and chlorophyll biosynthesis in darkness. In: Progress in Photosynthesis Research; Vol. IV, (J. Biggins, Ed.) Martinus Nijhoff, Amsterdam, pp. 439-443.
  8. Suzuki, J. Y. Bollivar, D. W.and Bauer, C. E. (1997) Genetic analysis of chlorophyll biosynthesis. Ann. Rev. Genetics 31, 61-89 .
  9. F. Yuichi F. and Bauer, C. E. (2000) Reconstitution of light-independent protochlorophyllide reductase from purified BchL and BchB subunits. In vitro confirmation of nitogenase-like features of a bacteriochlorophyll biosynthesis enzyme.J. Biol. Chem. 275, 23583-23588.
  10. Armstrong, G. A., (1998) Greening in the dark: light-independent chlorophyll biosynthesis from anoxygenic photosynthetic bacteria to gymnosperms. J. Photochem. Photobiol. B: Biol. 43, 87-100.
  11. M. Ikeuchi M. and S. Murakami, S. (1982) Measurement and identification of NADPH:protochlorophyllide oxidoreductase solubilized with Triton-X-100 from etioplast membranes of squash cotyledonsPlant Cell. Physiol. 23, 1089-1099.
  12. Dehesh, K. M. Hauser, M. and Apel, K. (1986) Light-induced changes in the distribution of the 36,000 Mr polypeptide of NADPH:protochlorophyllide oxidoreductase within different cellular compartments of barley (Hordeum vulgare, L.). I. Localization by immunoblotting in isolated plastids and total extracts.Planta 169, 162-171.
  13. Reinbothe, C., Lebedev, N., and Reinbothe, S. (199) A protochlorophyllide light-harvesting complex involved in de-etiolation of higher plants Nature 397, 80-84.
  14. Oosawa, N., Masuda, t., Awai, K., Fusada, N., Shimada, H., Ohta, H., and Takmiya, K. (2000) Identification and light-induced expression of a novel gene of NADPH-protochlorophyllide oxidoreductase isoform in Arabidopsis thaliana. FEBS Lett. 474: 133-136.
  15. Klement, H., Helfrich, M., Oster, U., Schoch, S., and Rudiger, W. (1999) Pigment-free NADPH:protochlorophyllide oxidoreductase from Avena sativa L: purification and substrate specificityEur. J. Biochem. 265, 862-874.
  16. Runge, S., Ulrich, S., Frick, J., Apel, K., and Armstrong, G. A. (1996) Distinct roles for light-dependent NADP:ptotochlorophyllide oxidorectase (POR) A and B during greening in higher plants The Plant J. 9, 513-523.
  17. N. Lebedev N., and M. P. Timko, M. P. (1999) Protochlorophyllide oxidoreductase B-catalyzed protochlorophyllide photoreduction in vitro Proc. Nat Acad. Sci. USA. 96, 9954-9954
  18. Su, Q., Frick, G., Armstrong G., and Apel, K. (2001) POR C of Arabidopsis thaliana: a third light-and NADPH-dependent protochlorophyllide oxidoreductase that is differentially regulated by light. Plant Mol. Biol. 47, 805-2001.
  19. Carey, E. E. and Rebeiz, C. A. (1985). Chloroplast Biogenesis 49. Difference among angiosperms in the biosynthesis and accumulation of monovinyl and divinyl protochlorophyllide during photoperiodic greening. Plant Physiol. 79, 1-6.
  20. Abd-El-Mageed, H. A. El Sahhar, K. F., Robertson, K. R., Parham, R., and Rebeiz, C. A. (1977). Chloroplast Biogenesis 77. Two novel monovinyl and divinyl light-dark greening groups of plants and their relationship to the chlorophyll a biosynthetic heterogeneity of green plants. Photochem. Photobiol. 66, 89-96.
  21. C. E. Cohen, C. E., Bazzaz, M. B., Fullet, S. E. and Rebeiz, C. A. (1977) Chloroplast Biogenesis XX. Accumulation of porphyrin and phorbin pigments in cucumber cotyledons during photoperiodic greening Plant Physiol. 60, 743-746.
  22. Wiktorsson, B. Ryberg, M.Sundqvist, C. (1996). Aggregation of NADPH-protochlorophyllide oxidoreductase-pigment complexes is favoured by protein phosphorylation. Plant Physiol. Biochem. 34, 23-34.
  23. Brereton, R. G., Bazzaz, M. B., Santikarn, S., Williams, D. H. (1983) Positive and Negative ion Fast Atom Bombardment Mass Spectrometric Studies on Chlorophylls: Structure of 4-vinyl-4-desethyl chlorophyll b. Tetrahedron Lett. 24: 5775-5778.
  24. Chisholm, S., Olson, R. J., Zettler, E. R., Goerike, R., Waterbury, J. B. (1988). A novel free-living prochlorophyte abundant in the oceanic euphotic zone. Nature. 334: 340-343.
  25. Chisholm, S. W., Frankel, S., Goerike, R., Olson, R., Palenic, R., Urbach, B., Waterbury, J. B., Zettler, E. R. (1992) Prochlorococcus marinus nov. gen. sp.: an oxyphototrophic marine prokaryote containing divinyl chlorophyll a and b. Arch. Mikrobiol. 157: 297-300.
  26. Goerike, R., Repeta, D. (1992). limnol. Oceanogr. 27: 425-433.
  27. Veldhuis, M. J. W., Kraay, G. W. (1990). Mr. Ecol. Progr. Ser. 68: 121-127.
  28. Belanger, F. C., Dugan, J. X., Rebeiz, C. A. (1982) Chloroplast Biogenesis: Identification of chlorophyllide a (E458F674) as a divinyl chlorophyllide a. J. Biol. Chem. 257: 4849-4858.
  29. Tripathy, B. C., Rebeiz, C. A. (1986) Chloroplast Biogenesis. Demonstration of the monovinyl and divinyl monocarboxylic routes of chlorophyll biosynthesis in higher plants. J. Biol. Chem. 261: 13556-13564.
  30. Tripathy, B. C., Rebeiz, C. A. (1988) Chloroplast Biogenesis 60. Conversion of divinyl protochlorophyllide to monovinyl protochlorophyllide in green(ing) barley, a dark monovinyl/light divinyl plant species. Plant Physiol. 87: 89-94.
  31. Parham, R. Rebeiz, C. A. (1992). Chloroplast Biogenesis: [4-vinyl] chlorophyllide a reductase is a divinyl chlorophyllide a-specific NADPH-dependent enzyme. Biochem. 31:8460-8464.
  32. Parham, R. Rebeiz, C. A. (1992). Chloroplast Biogenesis: [4-vinyl] chlorophyllide a reductase is a divinyl chlorophyllide a- specific NADPH-dependent enzyme. Biochem. 31: 8460-8464.
  33. Parham, R. Rebeiz, C. A. (1995) Chloroplast Biogenesis 72: A [4-vinyl] chlorophyllide a reductase assay using divinyl chlorophyllide a as an ecogenous substrate. Anal. Biochem. 231:164-169.
  34. Kolossov, V. L. Rebeiz, C. A. (2001) Chloroplast Biogenesis 84. Solubilization and partial purification of membrane-bound [4-vinyl] chlorophyllide a reductase from etiolated barley leaves. Anal. Biochem. 295: 214-219.
  35. Duggan, J. X. and Rebeiz, C. A. (1982a). Chloroplast Biogenesis 37. Induction of chlorophyllide a (E459 F675) accumulation in higher plants. Plant Sci. Lett. 24: 27-37.
  36. Duggan, J. X. Rebeiz, C. A (1982b). Chloroplast Biogenesis 42. Conversion of DV chlorophyllide a to monovinyl chlorophyllide a in vivo and in vitro. Plant Sci. Lett. 27: 137-145.
  37. Rebeiz, C. A., S. M. Wu, M. Kuhadja, H. Daniell, and E. J. Perkins. Chlorophyll biosynthetic routes and chlorophyll a chemical heterogeneity in plants. Mol. Cell. Biochem. 58:97-125.
  38. Rebeiz, C. A., Parham R., Fasoula D.A. and Ioannides I. M. (1994). Chlorophyll a Biosynthetic Heterogeneity. In: Biosynthesis of Tetrapyrroles. Pigments Ciba Symposium 180. pp 177-193, John Wiley, New York.
  39. Sironval, C., Kuyper, Y., Michel, J. M., and Brouers, M. (1967). The primary photoact in the conversion of protochlorphyll ide into chlorophyllide. Stud. Biophys. 5: 43-50.
  40. Gassman, M., Granick, S., and Mauzerall, D. (1968). A rapid spectral shift change in etiolated red kidney leaves following phototransformation of protochlorophyllide. Biochem. Biophys. Res. Comm. 32: 295-300.
  41. Bonner, B. (1969). A short-lived intermediate form in the in vivo conversion of protochlorphyll ide 650 to chlorophyllide 684. Plant Physiol. 44: 739-747.
  42. Gassman, M. (1973). The conversion of photoinactive protochlorophyllide 633 to phototransformable protochlorophyll ide 650 in etiolated bean leaves treated with delta-aminolevulinic acid. Plant Physiol. 52: 590-594.
  43. Adamson, H., and Packer, N. (1984). Dark-synthesis of chlorophyll in vivo and dark reduction of protochlorophyll ide in vitro by pea chloroplasts. In: Protochlorophyllide Reduction and Greening. Sironval, C. and Brouers, M. (eds.). pp. 353-363. Martinus Nijhoff, Boston.
  44. Griffiths, W. T. (1974). Source of reducing equivalents for the in vitro synthesis of chlorophyll from protochlorophyll. FEBS Lett. 46: 301-304.
  45. Griffiths, W. T. (1978). Reconstitution of chlorophyllide formation by isolated etioplast membranes. Biochem. J. 174: 681-692.
  46. Apel, K., Santel, H. J., Redlinger, T. E. and Falk, H. (1980). The protochlorophyll ide holochrome of barley (Hordeum vulgare L.). isolation and characterization of the NADPH:protochlorophyll ide oxidoreductase. Eur. J. Biochem. 111: 251-258.
  47. Apel, K. (1981). The protochlorophyllide holochrome of barley (Hordeum vulgare L.). Phytochrome-induced decrease of the translatable mRNA coding for the NADPH:protochlorophyllide:oxidoreductase. Eur. J. Biochem. 120: 89-93.
  48. Shulz R., and Senger, H. (1993). protochlorophyll ide Reductase: A keto enzyme in the greening process. In: Pigment-Protein Complexes in Plastids: Synthesis and assembly. Sundqvist, C., and Ryberg, M. (eds.). pp: 179-218. Academic Press, New York.
  49. Ikeuchi, M., and Murakami, S. (1982). Measurement and identification of NADPH:protochlorophyll ide oxidoreductase solubilized with Triton-X-100 from etioplast membranes of squash cotyledons. Plant Cell. Physiol. 23: 1089-1099.
  50. Dehesh, K., Hauser, M., and Apel, K. (1986). Light-induced changes in the distribution of the 36,000 Mr polypeptide of NADPH:protochlorophyllide oxidoreductase within different cellular compartments of barley (Hordeum vulgare, L.). I. Localization by immunoblotting in isolated plastids and total extracts. Planta 169: 162-171.
  51. Santel, H. J., and Apel, K. (1981). The protochlorophyll ide Holochrome of Barley (Hordeum vulgare L.). The effect of light on the NADPH:protochlorophyllide oxidoreductase. Eur. J. Biochem. 120: 95-103
  52. Kay, S. A., and Griffiths, W. T. (1983). Light-induced breakdown of NADPH-protochlorophyllide oxidoreductase in vitro. Plant Physiol . 72: 229-236.
  53. Armstrong, G. A., Runge, S., Frick, G., Sperlink, U., and Apel K. (1995). Identification of NADPH:protochlorophyll ide oxidoreductases A and B: A branched pathway for light-dependent chlorophyll biosynthesis in Arabidopsis thaliana. Plant Physiol. 108: 1505-1517.
  54. Holtorf, R., Reinbothe, S., Reinbothe, C., Bereza, B. and Apel, K. (1995). Two routes of chlorophyllide synthesis that are differentially regulated by light in barley (Hordeum vulgare L.). Proc. Natl. Acad. Sci. USA 92: 3254-3258.
  55. Reinbothe C, Apel, K. and Reinbothe S. (1995a). A light-induced protease from barley plastids degrades NADPH:protochlorophyllide oxidoreductase complexed with chlorophyllide. Mol. Cell. Biol. 15: 6206-6212.
  56. Reinbothe S., Reinbothe C., Holtorf H., and Apel, K. (1995b). Two NADPH:protochlorophyllide oxidoreductases in barley: evidence for the selective disappearance of POR A during the light-induced greening of etiolated seedlings. Plant Cell 7: 1933-1940.
  57. Cohen, C. E. and Rebeiz, C. A. (1978). Chloroplast Biogenesis XXII. Contribution of short wavelength and long wavelength protochlorophyll species to the greening of higher plants. Plant Physiol. 61:824-829.
  58. Rebeiz, C. C., and Rebeiz, C. A. (1986). Chloroplast Biogenesis 53: Ultrastructural study of chloroplast development during photoperiodic greening. In: Regulation of Chloroplast Differentiation. Akoyunoglou, G., and Senger, H. (eds.) pp: 389-396. Alan Liss, New York.
  59. Rebeiz, C. A. (1987). Ever Green. The Sciences.
  60. Kirk, J. T. O., and Tilney-Basset, R. A. E. (1967). The Plastids: Their Chemistry, Structure, Growth, and Inheritance. pp. 504-506. W. H. Freeman, London.
  61. Ryberg, M. R., and Sundqvist, C. (1991). Structural and functional significance of pigment-protein complexes of chlorophyll precursors. In: Chlorophylls. Cheer, H. (ed.). pp: 587-612. CRC press, , Florida.
  62. Tripathy, B. C. and Rebeiz, C. A. Non-equivalence of glutamic acid and delta-aminolevulinic acid as substrates for protochlorophyllide and chlorophyll biosynthesis in darkness. In: Progress in Photosynthesis Research Vol. IV. Biggins, J. (ed.) pp. 439-443. Martinus Nijhoff, Amsterdam.
  63. Thorne, S. W. (1971). The greening of etiolated bean leaves I. The initial photoconversion process. Biochim. Biophys. Acta. 226: 113-127.
  64. Sironval, C. and Kuyper, Y. (1972). The reduction of protochlorophyllide into chlorophyllide. Photosynthetica. 6: 254-275.
  65. Shibata, K. (1957). Spectroscopic studies on chlorophyll formation in intact leaves. J. Biochem. 44: 147-172.
  66. Cohen, C. E., Bazzaz, M. B., Fullett, S. A., and Rebeiz, C. A. (1977). Chloroplast Biogenesis XX. Accumulation of porphyrin and phorbin pigments in cucumber cotyledons during photoperiodic greening. Plant Physiol. 60: 743-746.
  67. Koski, V. M., French, C. S., and Smith, J. H. C. (1951). The action spectrum for the transformation of protochlorophyll to chlorophyll in normal and albino corn seedlings. Arch. Biochem. Biophys. 31: 1-17.
  68. Smith, J. H. C. and Benitez, A. (1954). The effect of temperature on the conversion of protochlorophyll to chlorophyll a in etiolated barley leaves. Plant Physiol. 29: 135-143.
  69. Boardman, N. K. (1962). Biochim. Biophys. Acta. 64: 279-288.
  70. Thorne, S. W. and Boardman, N. K. (1972). Biochim. Biophys. Acta. 267: 104-110.
  71. Ioannides, I., Fasoula D. A., Robertson, K. R. and Rebeiz, C. A. (1994). An evolutionary study of chlorophyll biosynthetic heterogeneity in green plants. Biochem. Syst. Ecol. 22: 211-220.
  72. Belanger, Faith C. and Rebeiz C. A. (1980b). Chloroplast Biogenesis 30. Chlorophyll(ide) (E459 F675) and Chlorophyll(ide) (E449 F675) the first detectable products of divinyl and monovinyl protochlorophyll photoreduction. Plant Sci. Lett. 18:343-350

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