<

Chlorophyll a Biosynthetic Pathway


web page counters
Alienware Computers


The Phorbin Nucleus

Topics

    Click on a topic to go directly to it.

V. The Chl a Carboxylic Biosynthetic Routes: Protochlorophyllides (Pchlides) a

carbxlc2.gif - 10.2 K

Fig. 5: Some reactions of the DV and MV Carboxylic Chl a Biosynthetic Routes Between Mg-Proto and Chl a

The reactions between ALA and DV Proto are shown in more details in Fig. 1. DV= divinyl; MV = monovinyl; Mg-Proto = IX; Mpe = Mg-Proto monomethyl ester; Pchlide a= protochlorophyllide a; Chlide a = Chlorophyllide a; Chl a = chlorophyll a; 4VMPR = [4-vinyl] Mg-Protoporphyrin IX reductase; 4VPR = [4-vinyl] protochlorophyllide a reductase; 4VCR = [4-vinyl] chlorophyllide a reductase. Arrows joining the DV and MV branches refer to reactions catalyzed by [4-vinyl] reductases. Triple-lined arrows point to putative reactions. Adapted from Rebeiz et al, 1983, 1994.. The various routes are believed to lead to the formation of MV Chl a, at different sites of the photosynthetic membranes.

A. Protochlorophyllide a (Pchlide a)

DV Pchlide a
MV Pchlide a
The role of protochlorophyllide (Pchlide) a as an intermediate in the Chl biosynthetic pathway was based on the detection of Pchlide a in X-ray Chlorella mutants inhibited in their capacity to form Chl (Granick, 1950a). It was conjectured that since the mutants had lost the ability to form Chl but accumulated Pchlide a, the latter was a logical precursor of Chl. On the basis of chemical derivatization coupled with absorbance spectroscopic analysis, the accumulated Pchlide a was assigned a monovinyl ( MV) chemical structure. When more powerful fluorescence spectroscopic techniques were used to reinvestigate the Pchlide a pool of various plant species, it was discovered that this pool was chemically heterogeneous and consisted of DV and MV components (Belanger and Rebeiz, 1980a). Granick proposed that Pchlide a was the immediate precursor of Pchlide a phytyl ester, a fully esterified Pchlide, which was wrongly believed at that time to be the main immediate precursor of Chl a in nature (Granick, 1950; Koski, 1950). The biosynthetic function of Pchlide a as the precursor of chlorophyllide (Chlide) a, one of the immediate precursors of Chl a, was not recognized till seven years later (Wollf and Price, 1957). Then in 1963, Jones reported the detection of DV Pchlide a in R. spheroides cultures, in which the biosynthesis of bacteriochlorophyll was inhibited by incubation with 8-hydroxyquinoline (Jones, 1963). Jones proposed that DV Pchlide a was a transient immediate precursor of MV Pchlide a in all plants. He also finalized the proposal of a linear, single-branched Chl a biosynthetic pathway which started with DV Proto and ended with the formation of MV Chl a and b (Jones, 1963).

1. Formation of the Cyclopentanone Ring (Ring E) of Pchlide a


MV Mpe Acrylic
MV OH-Mpe Propionate
MV Mpe Keto-propionate
MV DVPchlide a

Formation of the cyclopentanone ring (ring E) during the proposed conversion of Mpe to Pchlide a was suggested in 1950 to involve a beta-oxidation of a putative methyl Propionate side chain to a 3-keto derivative (Granick, 1950b). Later on, detection of putative DV and MV metal-free acrylic, hydroxy and keto derivatives in ultraviolet Chlorella mutants, led to the proposal that in lower plants, the formation of DV and MV Pchlide a involves a beta-oxidation sequence of the methyl propionate of DV and MV Mpe, at position 6 of the macrocycle and the formation of acrylic, hydroxy, and keto propionate Mpe intermediates (Ellsworth and Aronoff, 1969). The authors suggested that the DV and MV keto methyl propionate species cyclized automatically to yield DV and MV Pchlide a (Ellsworth and Aronoff, 1969). This work was met with skepticism and the putative intermediates were considered to be artifacts. This feeling was reinforced by the inability of the techniques, used by Ellsworth and Aronoff as well as of other analytical techniques available at that time, to detect the proposed MV Mpe substrate (MV Mpe), the proposed DV and MV acrylic, OH, and keto intermediates, and the DV Pchlide a) end product in normal, green, lower and higher plants.

A new phase in the study of the cyclopentanone ring formation was ushered by the introduction of powerful cell-free systems capable of the massive net synthesis of Pchlide a from exogenous ALA and tetrapyrrole substrates (Rebeiz et al, 1975; Mattheis and Rebeiz, 1977a; 1977b; Daniell and Rebeiz, 192a; 1982b; Tripathy and Rebeiz, 1986), and the development of sensitive analytical fluorescence methodologies that allowed the demonstration of the DV and MV heterogeneity of the metabolic pools between Mg-Proto and Chl a (Rebeiz, et al, 1994). With the use of similar techniques, the reactions between Mpe and Pchlide a have been reinvestigated by Castelfranco and collaborators (Wong and Castelfranco, 1985; Walker et al, 1988). In a series of experiments involving the conversion of added putative tetrapyrrole intermediates to Pchlide a, in organello, it was shown that (a) both the b-OH and b-keto methyl propionate derivatives of Mpe could be converted to Pchlide a, (b) that substrates with unesterified and esterified propionic acid residues at position 7 of the macrocycle were active, (c) that both, the DV and MV b-OH and b-keto methyl propionate derivatives also served as substrates, (d) that 2-ethyl,4-vinyl analogs were inactive, (e) that the 6-methyl acrylate derivative was also inactive, (e) that only one of the two 6-b-hydroxy enantiomers was active, (f) that only one of the two MV 6-b-keto derivative was active, and (g) and that the MV 6-b-keto derivative was 4 times more active that the DV analog, whereas DV and MV Mg-Proto were equally active. It is unfortunate that in this work, no efforts were made to distinguish between the conversion of the various substrates into DV and MV Pchlide a end products.

On the basis of the above results, it has been suggested that in plants, the formation of the cyclopentanone ring involves conversion of Mpe to b-OH and b-keto methyl propionate derivatives, with stereospecificity at the level of the b-keto derivative. It should be emphasized, however, that due to the lack of absolute specificity of the putative enzymes involved in cyclopentanone ring formation, elucidation of the actual sequence of events that result in the formation of the cyclopentanone ring of Pchlide a will have to await the unambiguous identification of all the putative DV and MV intermediates between DV and MV Mpe and DV and MV Pchlide a, as well as purification of the involved enzymes.

Also at this point, it appears that the reactions between Mpe and Pchlide a require molecular oxygen and iron (Spiller et al, 1982), and are inhibited by N-ethylmaleimide, dithiothreitol, and beta-mercaptoethanol (Wong and Castelfranco). It has also been reported that the conversion of Mpe to Pchlide a requires both the membrane and stromal fractions of the plastids (Walker et al, 1991). In our hands however, excellent cyclopentanone ring synthetase activity is observed with isolated plastid membranes without the need of a stromal factor. Along with with NADPH, the stromal factor appears to be involved in the regulation of the proportions of DV and MV Pchlide a formation (Kim et al, 1997).

2. Heterogeneity of Pchlide a-Protein Complexes
Pchlides a are extremely heterogeneous. That heterogeneity is expressed chemically as DV, and MV substitutions at the level of the tetrapyrrole chromophore at position 4 of the macrocycle. It is also expressed at the level of the chromophore-protein complexes referred to as Pchlide a holochromes (Pchlide a-Hs). The overall heterogeneity is also expressed by the detection of multiple resonance excitation energy transfer bands between Pchlide a-Hs and various Chl-protein complexes (Table 1).

Discussion of the heterogeneity of Pchlide a-Hs will focus (a) on the possible nature of the chromophore-apoprotein association, (b) on the spectroscopic properties of various Pchl(ide) a-Hs in situ, and (c) on the properties of purified Pchl(ide) a-Hs.

a. Nature of the Chromophore-Protein Association of Pchl(ide) a-Hs
In etiolated tissues, Pchlide a is the most abundant tetrapyrrole (90-95%) followed by less abundant Pchlide a E (5-10%) (Rebeiz et al, 1970). Pchlide a and its ester will be referred to collectively as Pchl(ide) a. Upon binding to apoproteins, the spectroscopic properties of Pchl(ide) a chromophores change drastically (see below). It should be made clear however that the bulk of Pchl(ide) a-Hs is made up of Pchlide a-Hs.

In Pchl(ide) a-Hs, various Pchl(ide) a chromophores are bound to different apoproteins by non-covalent forces. This is evidenced by the ready extraction of the Pchl(ide) a chromophores by organic solvents such as acetone. Association of the chromophores with apoproteins, probably involve (a) axial coordination of the Pchl(ide) a central Mg-atom to nucleophyllic amino acid side chains, and (b) hydrogen bonding between the keto group of the cyclopentanone ring of the Pchl(ide) a chromophore and appropriate amino acid side chains. Pigment-pigment interaction may involve axial coordination of the keto group of the cyclopentanone ring of one Pchl(ide) a chromophore to the central Mg-atom of another Pchl(ide) a chromophore as suggested by Katz et al (Katz et al, 1966) for Chl-Chl association in hydrophobic environments, as well as P-P interactions of Pchl(ide) a chromophores (Boucher and Katz, 1967). Axial coordination of the histidine nitrogen of apoproteins to the central Mg-atom (Deisenhofer and Michel, 1991) of Pchl(ide) a has not been established for various Pchl(ide) a-Hs.

b. Spectroscopic Properties of Various Pchlide a-Hs
The existence of at least two spectroscopically different Pchl(ide) a-Hs was first reported by Hill and coworkers (Hill et al, 1953). Using a Zeiss microspectroscope they observed that in etiolated barley leaves, a band absorbing at 650 nm disappeared (was phototransformed i. e. was photoconverted to a Chl-like compound) as the light was turned on and was replaced by the appearance of two new absorbance bands: one near 670 nm which corresponded to newly formed Chl a-like compound, and one at 635 nm, which did not appear to be convertible to Chl. These results gave rise to the notion that etiolated tissues contained two spectroscopically different Pchl(ide) a-H complexes. A longer wavelength (LW), phototransformable (t) complex absorbing at 650 nm, and a shorter wavelength (SW), non-phototransformable (nt) complex, absorbing at 635 nm.

To explain the difference between the LW and SW Pchl(ide) a-Hs, Butler and Briggs proposed, on the basis of freezing and thawing treatments of plant tissues, that aggregation of pigment molecules in etioplasts shifts the absorption maximum to longer wavelengths, while disaggregation of pigment molecules shifts the absorption maximum to shorter wavelengths (Butler and Briggs, 1966). Using freezing and thawing as well as extraction, heat and acid treatments, Dujardin and Sironval (Dujardin and Sironval, 1970) suggested the presence of three universal Pchl(ide) a-Hs in plants, namely: an aggregated, phototransformable species absorbing at 647-648 nm that involves pigment-protein and pigment-pigment interactions, a second phototransformable species absorbing at 639-640 nm which involves only pigment-protein interactions, and a non-phototransformable species absorbing at 627-628 nm, which is loosely bound to proteins. They also proposed that pigment-pigment interaction is not required for phototransformation while binding to a specific protein is required. Using absorption, fluorescence emission and excitation spectroscopy at 77 K, Kahn (Kahn et al, 1970), further characterized the three Pchl(ide) a-Hs as consisting of (a) a nt-fluorescent species with a red excitation maximum at 628 nm and a red fluorescence emission maximum at 630 nm [nt-Pchl(ide) a (E628 F630) ], (b) a t, nonfluorescent species with a red excitation maximum at 639 nm [(t-Pchl(ide) a E639 F00)] which transfers its excitation energy to a Pchl(ide) a-H with a red excitation maximum at 650 nm and a red fluorescence emission maximum at 655 nm [t-(Pchl(ide) a E650 F655)]. The latter is the predominant Pchl(ide) a-H in etiolated tissues. It is now known that this Pchl(ide) a-H species is a ternary complex of Pchlide a with NADPH and Pchlide a oxidoreductase.

Using high resolution 77 K spectrofluorometry coupled to matrix analysis, Cohen and Rebeiz (Cohen and Rebeiz, 1981) carried out a detailed studies of the Pchl(ide) a-Hs that accumulate in etiolated cucumber a DDV-LDV-LDDV species, and bean a DMV-LDV-LDMV plant species. The various Pchl(ide) a-H species were assigned Soret excitation maxima (E), and fluorescence emission maxima (F). The following Pchl(ide) a-H species were detected in etiolated cucumber cotyledons: nt-SW Pchl(ide) a-H (E440 F630), t-SW Pchl(ide) a-H (E443 F633), -(E444 F636) and -(E445 F640), and t-LW Pchl(ide) a (E450 F657), which was the predominant species in etiolated cucumber. In red-kidney bean, the following Pchl(ide) a-H species were detected nt-SW Pchl(ide) a H (E440 F630), t-SW Pchl(ide) a-H (E441 F633), -(E442 F636) and -(E443 F640), and t-LW Pchl(ide) a (E447 F657), which was the predominant species in etiolated bean. It is now known that this Pchl(ide) a-H species is a ternary complex of Pchlide a with NADPH and Pchlide a oxidoreductase.

The contribution of SW and LW Pchl(ide) a-Hs to the natural greening process was assessed during photoperiodic greening, i. e. during greening under alternating light/dark photoperiods (Cohen and Rebeiz, 1978). The following observations were made (a) SW Pchl(ide) a- H species appeared within the first 24 h of germination of cucumber seedlings, (b) subsequently, LW Pchl(ide) a H species appeared then disappeared, (c) The ratio of LW/SW Pchl(ide) a H species reached a maximum of 3:1 by the end of the second dark cycle and reached a value of zero by the end of the 6th dark cycle, (d) SW Pchl(ide) a H species were continuously present during the dark and light cycles and appeared to contribute actively to the greening process, and finally (e) primary corn and bean leaves exhibited a similar pattern of Pchl(ide) a H formation.

3. Purification of Pchlide a-Hs
Early work dealing with the purification of Pchl(ide) a-Hs was described by Boardman (Boardman, 1966). The partially purified Pchl(ide) a-H (MW = 600,000) exhibited a red absorption maximum at 637.5 nm. Upon illumination, part of the Pchl(ide) a was converted into Chlide a with a red absorption maximum at 681 nm which after two minutes in darkness shifted to 675 nm. This preparation, however, did not preserve the heterogeneous spectral properties observed in vivo. A purer preparation from etiolated bean leaves (MW = 300,000) was described by Schopfer and Siegelman (Schopfer and Siegelman, 1968). The purified Pchl(ide) a-H exhibited a red absorption maximum at 639 nm, which also did not reflect the spectral heterogeneity observed in vivo. In the light the red absorption maximum shifted to a Chlide a red absorption maximum at 678 nm, which drifted to 672 nm in darkness. More purified Pchl(ide) a-Hs were prepared from etiolated barley (MW 63,000) and bean (MW 100,000) by Henningsen and Kahn (Henningsen and Kahn, 1971). Photoconversion yielded a Chl(ide) a complex with a red absorption maximum at 678 nm. In this case too, the spectral properties of the purified Pchl(ide) a-H did not reflect the spectral heterogeneity observed in vivo.

4. Biosynthetic Heterogeneity of the Pchlide a Chromophore
The Pchlide a pool of green plants consists of DV and MV Pchlide a chromophores. In Fig. 1, five DV Pchlide a, and six MV Pchlide a pools are depicted as precursors of various metabolic tetrapyrroles. The assignment of the DV or MV Pchlide a pools to different thylakoid environments is based (a) on the detection of multiple resonance excitation energy transfer bands between Pchlide a and various Chl-protein complexes (Table 1), (b) the detection of DV Pchlide a in addition to MV Pchlide a in green plants (Belanger and Rebeiz, 1980), and (c) the biosynthetic heterogeneity of DV and MV Pchlide a (vide infra). The metabolism of the various DV and MV Pchlide a pools is discussed below.

In discussing Pchlide a biosynthetic heterogeneity distinctions will be made between (a) etiolated and green plants, (b) between biosyntheses during the dark and light phases of the photoperiod in green plants, and (c) between DDV-LDV-LDDV and DMV-LDV-LDMV plant species. As will be discussed in section XIII DDV-LDV-LDDV plant species such as cucumber accumulate mainly DV Pchlide a in darkness and in the light. In the light, Chl biosynthesis proceeds mainly via regenerated DV Pchlide a. On the other hand, DMV-LDV-LDMV plant species such as corn wheat and barley accumulate mainly MV Pchlide a in darkness. In the light, some DV Pchlide a is formed, but Chl biosynthesis proceeds mainly via regenerated MV Pchlide a (Abd-El-Mageed et al, 1997).

a. Metabolism of DV Pchlide a
DV OH-Mpe Propionate
DV Mpe Keto-propionate
DV Pchlide a

dvpidea2.gif - 16.3 K

The DV nature of the DV Pchlide a component of the heterogeneous Pchlide a pool of higher plants was originally ascertained by chemical derivatization coupled with analytical fluorescence spectroscopy at 77K (Belanger and Rebeiz, 1980a). It was also confirmed by 1H nuclear magnetic resonance (NMR) and fast atom bombardment (FAB) mass spectroscopy (Wu and Rebeiz, 1984). The exact sequence, and mechanism of the reactions involved in cyclopentanone ring formation during the conversion of DV Mpe to DV Pchlide a is not completely understood and will have to await the unambiguous identification of all putative DV intermediates between DV Mpe and DV Pchlide a, as well as purification of involved enzymes.

Precursor-product relationships between DV Pchlide a and DV Chlide a (Fig. 1, routes 1, and 13) was established by demonstrating the photoreduction of DV Pchlide a to DV Chlide a in etiolated cucumber cotyledons induced to accumulate DV Pchlide a exclusively (Duggan and Rebeiz, 1982a)., and in etiolated corn seedlings exposed to brief light-dark incubations (Belanger et al, 1982).

a. Biosynthesis of DV Pchlide a via Biosynthetic Route 1 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Greening DDV-LDV-LDDV Plants during the Dark Phases of the Photoperiod

Formation of the cyclopentanone ring via biosynthetic route 1 (Fig. 1) in etiolated DDV-LDV-LDDV plants was demonstrated by conversion of exogenous DV Mpe to DV Pchlide a in isolated cucumber etiochloroplasts (Tripathy and Rebeiz, 1986). DV Mpe was converted into 83% DV Pchlide a and 17% MV Pchlide a. The formation of much smaller amounts of MV Pchlide a can be accounted by bosynthetic route 3 (Fig. 1) and suggests that biosynthetic route 1 (Fig. 1) is the predominant biosynthetic route in etiolated DDV-LDV-LDDV plant species in darkness.

Most probably, biosynthetic route 1 is also active in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977).

b. Biosynthesis of DV Pchlide a via Biosynthetic Route 8 in DDV-LDV-LDDV Plants during the Light and Dark Phases of the Photoperiod

In green DDV-LDV-LDDV plant species such as cucumber, DV Pchlide a is continuously present during the light cycles of the photoperiod and only trace amounts of MV Pchlide a are formed (Carey and Rebeiz, 1985; Ionannides et al, 1994). Interruption of the light cycle by a brief dark period (LD) indicated that such plant species form most of their Chl via regenerated DV Pchlide a (Abd-El-Mageed et al, 1997). These observation suggest very strongly that during the light cycles of the photoperiod, green DDV-LDV-LDDV plant species form most of their Chl via DV Pchlide a, DV Chlide a and MV Chlide a as depicted in route 8. DV Pchlide a is most probably formed from DV Mpe as demonstrated for route 1, and is converted to DV Chlide a by PORB which is active in the light (Runge et al, 1996). It is important to note that MV Pchlide a does not participate in this biosynthetic route. In other words biosynthetic route 8 is responsible for the formation of the bulk of MV Chl a in DDV-LDV-LDDV plant species in daytime without participation of MV Pchlide a. There is no reason to believe that route 8 is not also functional in green DDV-LDV-LDDV plants during the dark cycles of the photoperiod.

g. Biosynthesis of DV Pchlide a via Biosynthetic Routes 10 and 11 in Etiolated DMV-LDV-LDMV Plants in Darkness and in Green DMV-LDV-LDMV Plants during the Light Phases of the Photoperiod

DV Pchlide a is formed via routes 10 and 11 (Fig. 1) in dark-grown DMV-LDV-LDMV plant species and in light-grown DMV-LDV-LDMV plant species such as corn, wheat and barley, during the dark and light phases of the photoperiod. This hypothesis is based on the following experimental data.

For example, during dark incubation, of barley etiochloroplasts, exogenous DV Mpe is converted into 56% MV Pchlide a and 44% DV Pchlide a (Tripathy and Rebeiz, 1986). This in turn suggests that biosynthetic routes 10 and 11 (Fig. 1) are highly active in etiolated tissues of this greening group of plants. These results also caution against assuming that conversion of exogenous DV substrates to Pchlide a in organello is likely to yield DV Pchlide a exclusively, without specific analysis of DV and MV components as has been often assumed (Wong and Castelfranco, 1985; Walker et al, 1991). The operation of routes 10 and 11 in DMV-LD-LDMV plant species is also compatible with the detection of strong 4PideR activity that converts DV Pchlide a to MV Pchlide a in barley etiochloroplasts (Tripathy and Rebeiz, 1988).

In the light, DMV-LDV-LDMV plants form most of their Chl via regenerated MV Pchlide a (Abd-El-Mageed et al, 1997). The latter is most probably formed from DV Pchlide a, which is always detectable in the light, by the action of 4VPideR (Tripathy and Rebeiz, 1988). In summary, these results are compatible with the operations of biosynthetic routes 10 and 11 in DMV-LDV-LDMV plants in the light.

d. Biosynthesis of DV Pchlide a via Biosynthetic Route 13 in Etiolated DMV-LDV-LDMV Plants in Darkness, and in Greening DMV-LDV-LDMV Plants during the Dark Phases of the Photoperiod

The formation of the cyclopentanone ring of DV Pchlide a via biosynthetic route 13 (Fig. 1) in etiolated DMV-LDV-LDMV plants, was demonstrated by conversion of exogenous DV Mpe to DV Pchlide a in isolated barley etioplasts (Tripathy and Rebeiz, 1986). DV Mpe was converted into 44% DV Pchlide a and 56% MV Pchlide a. The formation of smaller amounts of DV Pchlide a than MV Pchlide a can be accounted for by route 13, and suggests that biosynthetic route 13 (Fig. 1) is not the predominant biosynthetic route in etiolated DMV-LDV-LDMV plant species in darkness.

Most probably, biosynthetic route 13 is also active in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977.

b. Metabolism of MV Pchlide a
MV OH-Mpe Propionate
MV Mpe Keto-propionate
MV Pchlide a

mvpidea2.gif - 45.0 K

The MV nature of the MV Pchlide a component of the heterogeneous Pchlide a pool of higher plants was determined by chemical derivatization coupled with analytical fluorescence spectroscopy at 77 K (Belanger and Rebeiz, 1980a). The MV nature was also confirmed by 1H nuclear magnetic resonance (NMR)s and fast atom bombardment (FAB) mass spectroscopy (Shedbalkar et al, 1991).

The biosynthesis of MV Pchlide a via routes 2, 3, 9, 10, 11 and 12 is discussed below. The exact sequence, and mechanism of the reactions responsible for formation of the cyclopentanone ring during conversion of MV Mpe to MV Pchlide a is not completely understood and will have to await he unambiguous identification of all putative MV intermediates between MV Mpe and MV Pchlide a, as well as purification of involved enzymes.

A precursor-product relationship between MV Pchlide a and MV Chlide a was established by demonstrating the photoconversion of MV Pchlide a to MV Chlide a in etiolated cucumber cotyledons enriched in MV Pchlide a (Belanger and Rebeiz, 1980b).

a. Biosynthesis of MV Pchlide a via Biosynthetic Routes 2 and 3 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Greening DDV-LDV-LDDV Plants during the Dark Phases of the Photoperiod

In addition to DV Pchlide a, etiolated and green DDV-LDV-LDDV plant species such as cucumber, form smaller amounts of MV Pchlide a during prolonged dark incubation (Carey and Rebeiz, 1985; Ioannides et al, 1994). This Pchlide a formation can be accounted for by routes 2 and 3.

Route 2 is initiated by reduction of the vinyl group at position 4 of the DV Mg-proto macrocycle to ethyl, and conversion of DV Mg-proto to MV Mg-proto. The reaction is catalyzed by 4VMPR (Kim and Rebeiz, 1996). The nascent MV Mg-Proto is then converted into MV Mpe and MV Pchlide a as depicted in route 2.

In route 3, MV Pchlide a formation can be accounted for by a slow conversion of DV Pchlide a to MV Pchlide a, a reaction catalyzed by 4VPideR during prolonged dark-incubations (Tripathy and Rebeiz, 1988). Probably, biosynthetic routes 2 and 3 are also active in greening DDV-LDV-LDDV plants during the first few dark phases of the photoperiod when Pchlide a accumulation is substantial (Cohen and Rebeiz, 1978; Cohen et al, 1977).

b. Biosynthesis of MV Pchlide a via Biosynthetic Route 9 in Greening DDV-LDV-LDDV Plants during the Dark and Light Phases of the Photoperiod

Route 9 branches from route 8 and is proposed to account for the formation of MV Pchlide b in green DDV-LDV-LDDV plant species in the light. In this route MV Pchlide a is considered to be formed from DV Pchlide a by the action of 4VPideR (Tripathy and Rebeiz, 1988). It should be pointed out that even during the light phases of the photoperiod DDV-LDV-LDDV plants form very small amounts of MV Pchlide a in addition to the formation of massive amounts of DV Pchlide a. It should also be stressed that in DDV-LDD-LDDV plants, formation of MV Pchlide a via route 9 is destined exclusively for the biosynthesis of MV Pchlide b in the light [153].

g. Biosynthesis of MV Pchlide a via Biosynthetic Routes 10 and 11 in Etiolated DMV-LDV-LDMV Plants in Darkness and in Green DMV-LDV-LDMV Plants during the Dark and Light Phases of the Photoperiod

As depicted in Fig. 1, MV Pchlide a can also be formed by reduction of the vinyl group of DV Pchlide a to ethyl at position 4 of the macrocycle, via routes 10 and 11. Route 11 is proposed as a second route responsible for the formation of MV Pchlide b in DMV-LDV-LDMV plant species. The operation of routes 10 and 11 in etiolated DMV-LDV-LDMV plants is based on the following experimental evidence: (a) in isolated barley etioplasts, exogenous DV Mpe is converted into 56% MV Pchlide a and 44% DV Pchlide a [97], and (b) DV Pchlide a is actively converted to MV Pchlide a by 4VPideR (Tripathy and Rebeiz, 1988).

In green DMV-LDV-LDMV plants, the operation of biosynthetic routes 10 and 11 are based on the observation that (a) in the light, in DMV-LDV-LDMV 4VPideR is very active (Tripathy and Rebeiz, 1988), and (b) DMV-LDV-LDMV plant species accumulate small amounts of residual DV Pchlide a in the light because nascent DV Pchlide a is very rapidly converted to MV Pchlide a by 4VPideR (Abd-El-Mageed et al, 1997).

It is uncertain at this stage whether vinyl reduction under various conditions is catalyzed by identical 4VPideRs or 4VPideR isozymes. It is very likely that 4VPideR is distinct from 4VMPR, which converts DV Mg-Proto to MV Mg-Proto (Kim and Rebeiz, 1996). Indeed, Rhodobacter capsulatus in which the bchJ gene which codes for 4VPideR has been deleted accumulates massive amounts of MV Mg-Proto and Mpe in addition to the accumulation of DV Pchlide a (Suzuki and Bauer, 1995). This in turn indicates that a separate [4-vinyl] reductase that acts prior to DV Pchlide a vinyl reduction is responsible for the accumulation of MV Mg-Proto in plants. Also, 4PideR is probably different from [4-vinyl] Chlide a reductase (4VCR), which converts DV Chlide to MV Chlide a (Kolossov and Rebeiz, 2001). This is evidenced by the observation that etiolated tissues containing an extremely active 4VCR, can be induced to accumulate massive amounts of DV Pchlide a in the absence of any MV Pchlide a formation. Yet when DV Chlide a is made available to the isolated plastids, it is converted to MV Chlide a by 4VCR, in less than one minute (Parham and Rebeiz, 1992). Furthermore, these results indicate that the proposal of only one [4-vinyl] reductase of wide substrate specificity, acting at all levels of DV to MV reduction as suggested by others (Whyte and Griffiths, 1993) is unfounded.

d. Biosynthesis of MV Pchlide a via Biosynthetic Route 12 in Etiolated DDV-LDV-LDDV Plants in Darkness, and in Green DDV-LDV-LDDV Plants during the Dark Phases of the Photoperiod

Biosynthetic route 12 is one of two routes that involve the participation of MV Mg-Proto in the formation of MV Mpe and MV Pchlide a. The other route tha involves MV Mg-Proto is biosynthetic route 2.

In etiolated DMV-LDV-LDMV plants the specific formation of the cyclopentanone ring in an exclusive MV sequence of reactions via route 12 (Fig. 1) was demonstrated by (a) conversion of exogenous MV Mpe to MV Pchlide a (Tripathy and Rebeiz, 1986), and (b) conversion of DV Mg-Proto to MV Mg-Proto in isolated barley etiochloroplasts (Kim and Rebeiz, 1996). In this system, the biosynthesis of MV Pchlide a occurred without formation of DV Pchlide a. The exact sequence, and mechanism of the reactions that convert MV Mpe to MV Pchlide is not completely understood and will have to await the unambiguous identification of all the putative MV intermediates between MV Mpe and MV Pchlide a, as well as purification of involved enzymes.

References

  1. 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.
  2. 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.
  3. Granick, S. (1950a). Magnesium vinyl pheoporphyrin a5, another intermediate in the biological synthesis of chlorophyll. J. Biol. Chem. 183:713-730.
  4. Belanger, F. C. and Rebeiz, C. A. (1980a) Chloroplast Biogenesis. Detection of divinyl protochlorophyllide in higher plants. J. Biol. Chem. 255:1266-1272
  5. Koski, V. M. (1950). Chlorophyll formation in seedlings of Zea mays L. Arch. Biochem. 29:339-343.
  6. Wollf, J. B., and Price, L. (1957). Terminal steps of chlorophyll a biosynthesis in higher plants. Arch, Biochem. Biophys. 72:293-301.
  7. Jones, O. T. G. (1963). Magnesium2,4-divinylphaeoporphyrin a5 monomethyl ester, a protochlorophyll-like pigment produced by Rhodopseudomonas spheroides. Biochem. J. 89:182-189.
  8. Granick, S. (1950b). The structural and functional relationships between heme and chlorophyll. Harvey Lect. 44:220-245.
  9. Ellsworth, R. K., and Aronoff, S. (1969). Investigations of the Biogenesis of Chlorophyll a. IV. Isolation and partial characterization of some biosynthetic intermediates between Mg-protoporphine IX monomethyl ester and Mg-vinylpheoporphine a5, obtained from Chlorella mutants. Arch. Biochem. Biophys. 130:374-383.
  10. Rebeiz, C. A., Smith B. B., Mattheis, J. R., Rebeiz, C. C., and Dayton, D. F. Chloroplast biogenesis. Biosynthesis and accumulation of protochlorophyll by isolated etioplasts and developing chloroplasts. Arch. Biochem. Biophys. 171:549-567.
  11. Mattheis, J. R. and Rebeiz, C. A. (1977a) Chloroplast biogenesis: New synthesis of protochlorophyllide from protoporphyrin IX by developing chloroplasts. J. Biol. Chem. 252:8347-8349.
  12. Mattheis, James R. and Rebeiz, C. A. (1977b). Chloroplast biogenesis: Net synthesis of protochlorophyllide from magnesium protoporphyrin monoester by developing chloroplasts. J. Biol. Chem. 252:4022-4024.
  13. Daniell, H. and Rebeiz, C. A. (1982a). Chloroplast Culture VIII. A new effect of kinetin in enhancing the synthesis and accumulation of protochlorophyllide in vitro. Biochem. Biophys. Res. Comm. 104:837-843.
  14. Daniell, H. and Rebeiz, C. A. (1982b). Chloroplast culture IX. Chlorophyll(ide) a biosynthesis in vitro at rates higher than in vivo. Biochem. Biophys. Res. Comm. 106:466-470
  15. Tripathy, B. C. and 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.
  16. Wong, Y. S. and Castelfranco, P. A. (1985). Properties of the Mg-protoporphyrin monomethyl ester (oxidative) cyclase system. Plant Physiol. 79:730-733.
  17. Walker, C. J. Mansfield, K. E., Rezzano, I. R., Hanamoto, C. M., Smith, K. V., and Castelfranco, P. A. (1988). The magnesium-protoporphyrin IX (oxidative) cyclase system. Studies on the mechanism and specificity of the reaction. Biochem. J. 255:685-692.
  18. Spiller, S. C., Castelfranco, A. M. and Castelfranco, P. A. (1982). Effect of iron and oxygen on chlorophyll biosynthesis. Plant physiol. 69:107-111.
  19. Walker, C. J., Castelfranco, P. A. and Whyte, B. J. (1991). Synthesis of divinyl protochlorophyllide. Enzymological properties of the Mg-protoporphyrin IX monomethyl ester oxidative cyclase system. Biochem. J. 276:691-697.
  20. Kim, J. S., Kolossov, V., and Rebeiz, C. A. (1997) Chloroplast Biogenesis 76. Regulation of 4-vinyl reduction during conversion of divinyl Mg-protoporphyrin IX to monovinyl protochlorophyllide a is controlled by plastid membranes and stromal factors. Under Review.
  21. Rebeiz, C. A., Yaghi, M., Abou-Haidar, M and Castelfranco, P. (1970) Protochlorophyll biosynthesis in cucumber (Cucumis sativus L.) cotyledons. Plant Physiol. 46: 57-63.
  22. Katz, J. J., , Dougherty. R. C., and Boucher, L. C. (1966)Infrared and nuclear magnetic resonance spectroscopy of chlorophyll. In: The Chlorophylls (Vernon L. P. and Seely, G. R., eds), PP 185-251.
  23. L. J. Boucher, L. J., and Katz, J. J. (1967) Aggregation of metalloporphyrins. J. Am. Chem. Soc. 89: 4703-4708.
  24. Deisenhofer, J. and H. Michel, H. (1991) Crystallography of chlorophyll proteins. In Chlorophylls, edited by H. Scheer (CRC Press, Boca Raton, Florida., 1991), p. 613-625.
  25. R. Hill, R., Smith, J. H. C., and French, C. S.Yearb. (1953) The absorption and fluorescence properties of natural protochlorophylls. Carneg. Inst. 52: 153-155.
  26. R. Butler R., and Briggs, W. R. (1966) The relation between structure and pigments during the first stages of proplastid greening Biochim. Biophys. Acta. 112 : 45-53.
  27. Dujardin E. and Sironval, C. (1970) The reduction of protochlorphyllide into chlorophyllide III. The phototransformation of the forms of the protochlorphyllide-lipoprotein complex found in darkness. Photosynthetica 4: 129-138.
  28. A. Kahn, A., N. K. Boardman, N. K., and Thorne, S. W. (1970) Energy transfer between protochlorophyllide molecules: Evidence for multiple chromophores in the photoactive protochlorophyllide-protein complex in vivo and in vitroJ. Mol. Biol. 48: 85-101.
  29. Cohen, C. E., and Rebeiz, C. A. (1981) Chloroplast Biogenesis 34. Spectrofluorometric characterization in situ of the protochlorophyll species in etiolated tissues of higher plantsPlant Physiol. 67, 98-103.
  30. C. E. Cohen, C. E., and Rebeiz, C. A. (1978) Chloroplast biogenesis 22. Contribution of short wavelength and long wavelength protochlorophyll species to the greening of higher plantsPlant Physiol. 61, 824-829.
  31. N. K. Boardman, N. K. (1966) In The Chlorophylls, (edited by L. P. Vernon, and G. R. Seely)academic Press, New York, p. 437-479.
  32. P. Schopfer P., and Siegelman, H. W. (1968) Purification of protochlorophyllide holochromePlant Physiol. 43, 990-996.
  33. K. W. Henningsen, K. W., and Kahn, A. (1971) Photoactive subunits of protochlorophyll(ide) holochrome Plant Physiol. 47, 685-690.
  34. Abd-El-Mageed, H. A., El Sahhar, K. F., Robertson, K. R., Parham, R., and Rebeiz, C A. (1997) 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 plantsPhotochem. Photobiol. 66, 89-96.
  35. Wu, S. M., and Rebeiz, C. A. (1984). Chloroplast Biogenesis 45: Molecular structure of protochlorophyllide (E443 F625) and of chlorophyllide a (E458 F674). Tetrahedron 40:659-664.
  36. Duggan, J. X. and Rebeiz, C. A. (1982). Chloroplast Biogenesis 37. Induction of chlorophyllide a (E459 F675) accumulation in higher plants. Plant Sci.Lett. 24:27-37.
  37. Belanger, F. C., Dugan, J. X. and Rebeiz, C. A. (1982) Chloroplast Biogenesis: Identification of chlorophyllide a (E458F674) as a divinyl chlorophyllide aJ. Biol. Chem. 257, 4849-4858 .
  38. 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.
  39. 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.
  40. 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.
  41. Abd-El-Mageed, H. A., El Sahhar, K. F., Robertson, K. R., Parham, R. and Rebeiz, C. A. (1997) 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.
  42. 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.
  43. Kolossov V. L.and 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 .
  44. Whyte, B. J., Griffiths, T. V. (1993) 8-Vinyl reduction and chlorophyll a biosynthesis in higher plants. Biochem. J. 291, 939-944.
  45. Kim J. S. and Rebeiz, C. A. (1996) Origin of the chlorophyll a biosynthetic heterogeneity in higher plants. J. Biochem. Mol. Biol. 29, 327-334.
  46. Shedbalkar V.P., Ioannides, I.M., and Rebeiz, C. A. (1991) Chloroplast Biogenesis. Detection of monovinyl protochlorophyll(ide) b in plants. J. Biol. Chem. 266 : 17151-17157
  47. Tripathy, B. C., and Rebeiz, C. A. 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.
  48. Kim, J. S., and Rebeiz, C. A. (1996). Chloroplast Biogenesis: Origin of the Chl a biosynthetic heterogeneity in higher plants. J. Biochem. Mol. Biol. 29 :327-334.
  49. Suzuki, J. Y., and Bauer, C. E. (1995). Altered monovinyl and divinyl protochlorophyllide pools in bchJ mutants of Rhodobacter capsulatus. Possible monovinyl substrate discrimination of light-independent protochlorophyllide reductase. J. Biol. Chem. 270: 3732-3740.
  50. Parham, R. and Rebeiz, C.A. (1992). Chloroplast Biogenesis: [4-Vinyl] chlorophyllide a reductase is a divinyl chlorophyllide a-specific NADPH-dependent enzyme. Biochemistry 31: 8460-8464
  51. Parham, R. And Rebeiz, C.A. (1995). Chloroplast Biogenesis 72: A [4-vinyl] chlorophyllide a reductase assay using divinyl chlorophyllide a as an exogenous substrate. Anal. Biochem. 231: 164-169.
  52. Whyte, B. J., and Griffiths, T. V. (1993). 8-Vinyl reduction and chlorophyll a biosynthesis in higher plants. Biochem. J. 291:939-944.
  53. 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.

Go Back to Main Menu

Web Analytics

Clicky