From the Institute of Organic Chemistry, University of Fribourg, Chemin du Musée 9, CH-1700 Fribourg, Switzerland
Received for publication, October 11, 2000, and in revised form, December 11, 2000
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ABSTRACT |
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A new type of chlorophyll catabolite was isolated
from extracts of de-greened primary leaves of barley (Hordeum
vulgare cv. Lambic). Its constitution was elucidated by
one-dimensional and two-dimensional
{1H,13C}-homo- and heteronuclear NMR
spectroscopic techniques and by high resolution mass spectroscopy. The
isolated catabolite, a water-soluble, colorless, and nonfluorescent
linear tetrapyrrole, resembles urobilinogen in which one of the
propionic side chains forms a five membered isocylic ring system,
indicating its origin from the chlorophylls.
Metabolic disappearance of the chlorophylls
(Chls)1 in phototrophic
organisms indicates programmed close down of photosynthesis. Although
several linear tetrapyrrolic Chl catabolites were isolated during the
last decade from green algae and higher plants, the metabolic pathway
of tremendous amounts of the Chls is still under question. What
products follow after the familiar formybilinones? What are the
ultimate products of Chl degradation? Degradation of the Chls occurs in
light as well as in darkness. During senescence cellular components are
hydrolyzed and metabolized; liberated rare elements such as [N],
[P], [S] and metal ions are relocated (1, 2).
The first structures of Chl catabolites isolated from a green alga and
a higher plant were published in 1991 (3, 4). Several Chl catabolites
have been isolated since then (Fig. 1). The similarity of the
structures of the red Chl catabolites (3, 4) isolated from the green
alga Chlorella protothecoides with the colorless catabolites
isolated from higher plants (5-7) suggest a close relationship in the
basic skeleton. Biologists regard members of the phylum Chlorophyta as
progenitor of the higher plant cell (9, 10). This information triggered
research activities in several disciplines to elucidate the apparently
unique catabolic pathway of the Chls in the green plant lineage. The
studies range from the elucidation of chemical and enzymatic reaction
mechanisms to molecular biological research (6, 11, 12). Previous labeling experiments with oxygen isotopes and heavy water showed a high
regio- and stereoselectivity of the oxidative ring opening mechanism
and the involvement of a monooxygenase in the ring cleaving step
(13-15). The remarkable allylic pyrroline/pyrrole rearrangement was
studied in detail (11, 16). Most recent in vivo deuterium labeling has shown that in higher plants Chl b is degraded
via Chl a (17).
Chl breakdown has long been considered as a detoxification process in
which photodynamic active Chls convert to colorless tetrapyrrolic
products, all of them carry a characteristic formyl group (Fig. 1).
Those products were regarded as the final products of Chl breakdown in
senescent plants, which apparently do not cause further cleavage into
smaller fragments (6).
This work focuses on the isolation and determination of the
constitution of a new type of chlorophyll catabolite by spectroscopic methods and discusses the origin and relevance in the catabolic pathway
of the chlorophylls in plants.
General--
Chemicals were reagent grade; all solvents were
distilled before use. HPLC solvents were supplied from Fluka (Buchs,
Switzerland). HPLC columns and Nucleosil 100-7 C8 VP 250/10 were
from Macherey-Nagel (Oensingen, Switzerland), and MPLC columns
Lobar® LiChroprep® RP-18 (40-63 µm) were
purchased from Merk (Darmstadt, Germany). 35cc
Sep-Pack® Vac C-18, 10 g, were provided from Waters
(Milford, MA).
Homo- and heteromagnetic resonance experiments were performed on a
Bruker Avance DRX-500 spectrometer operating at the frequencies of
500.13 MHz for 1H and 125.75 MHz for 13C.
Chemical shifts ( Plant Matertial--
Barley seeds (Hordeum vulgare L. cv. Lambic) were a gift from Florimond Desprez. The seeds were
germinated in high density (5 seeds/cm2) in moist garden
soil and grown under natural light conditions. The primary leaves were
harvested when they reached about 10-15 cm in height. The greening and
de-greening procedure of barley leaves was essentially as described
previously (17).
Isolation of the Chlorophyll Catabolite--
150 g of de-greened
yellow leaves of H. vulgare (fresh weight) were homogenized
in a blender with 300 ml of a solution consisting of 0.1 M
potassium phosphate (KP) buffer, pH 6.8:acetone:MeOH (1:1:1).
Work-up was essentially as described previously (17). Aliquots of the
aqueous phase were injected into a Lobar®
LiChroprep® RP-18 (40-63 µm) column and eluted (4 bar,
22 ml/min) with a solution of 30 volume % MeOH in 0.01 M
KP buffer, pH 6.8. Two fractions positive in the chromic acid
degradation assay were sampled. MeOH was evaporated in
vacuo. The polar fraction contained both urobinlinogens, the less
polar fraction contained formylbilinone 5. The resulting
aqueous phases were again concentrated on a 35cc Sep-Pack®
RP-18 cartridge. Separation and purification of the urobinlinogens were
achieved on a HPLC Nucleosil 100-7 C8 VP 250/10 column eluted isochratically (flow rate: 5 ml/min) with a solution of MeOH in 0.01 M KP buffer, pH 6.8 (13 volume %). The fractions with a
retention time of 13 min and of 14 min, respectively (DAD-UV detection
at 250 nm) were collected at 0 °C and stored under argon. After
removal of the volatile solvent in vacuo, each product was
de-salted on a 35cc Sep-Pack® C-18 cartridge by first
washing with 200 ml of distilled water and afterward eluted with
aqueous acetone (50 ml of 50 volume %). The gross solvent was
eliminated in vacuo, and the remaining aqueous solution was
lyophilized giving 15 mg and 10 mg, respectively, of a slightly yellow powder.
A triad of tetrapyrrolic compounds was isolated by HPLC from
yellow cotyledons of barley. After chemical degradation all showed on
TLC plates the same characteristic maleimide fragments, namely 3-(2-hydroxyethyl)-4-methyl maleimide, 3-(2,3-dihydroxyethyl)-4-methyl maleimide and hematinic acid imide (cf. Ref.
17).
The less polar compound isolated in about 7 mg was spectroscopically
identical in all aspects with catabolite 5 (Fig. 1) previously isolated from H. vulgare cv. Gerbel (4). The second product, which was isolated in
about 15 mg, was spectroscopically analyzed. Mass spectrometric
analysis at high resolution showed two molecular ions at m/z
705.2530 atomic mass units (100%) and m/z 743.2093 atomic mass units (10%). This corresponds exactly with a molecular ion
[C34H41KN4O10 + H]+ (calculated: 705.2532; error: 2.5 × 10
INTRODUCTION
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
EXPERIMENTAL PROCEDURES
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
) are given in parts/million downfield from the
solvent used and coupling constants (J) in Hertz (Hz). Mass spectra
were obtained with a Bruker FTMS 4.7T BioAPEXII, using electrospray
ionization technique in the positive mode. UV-visible spectra were recorded on a Perkin-Elmer Lambda 40 spectrometer,
max (log
) in nanometers. CD spectra were measured on
a Jobin-Yvon Auto Dichograph Mark V,
[l
mol
1 cm
1].
RESULTS
TOP
ABSTRACT
INTRODUCTION
EXPERIMENTAL PROCEDURES
RESULTS
DISCUSSION
REFERENCES
4) and
[C34H41KN4O10 + K]+ (calculated: 743.2091; error: 1.8 × 10
4), respectively, establishing the bulk
molecular formula to be C34H41KN4O10.
View larger version (18K):
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Fig. 1.
Structures of chlorophyll a
and b (1, 2) and of the catabolites derived from
the green alga C. protothecoides (3, 4) (Ref. 5),
barley (H. vulgare) (5) (Ref. 6),
Liquidambar spec. (6) (Ref. 7), C. japonicum (6) (Ref. 8), and rape (Brassica
napus) (7) (Ref. 6). Distinct chlorophyll a
and b catabolites are excreted from green algae, whereas
higher plants degrade chlorophyll a and b to the
same catabolite (see Introduction).
Details on the constitution of this metabolite were deduced from the
analysis of the one-dimensional and two-dimensional
{1H,13C}-homo- and heteronuclear NMR
spectroscopy (Fig. 2). The compound when
dissolved in D2O shows 33 carbon-bound hydrogen atoms and 33 carbon atoms. Seven protons are bound to hetero-atoms, which rapidly
exchange for solvent. Attached proton test spectra show 5 primary, 7 secondary, 4 tertiary, and 17 quaternary carbon atoms. All protons
resonate largely as isolated systems and are well resolved, except the
signal group at about H 2.7 (Ha(5) and
Ha(15)) and
H 3.6 (H2(32) and Hb(182)) in
which two and three protons overlap, respectively (Table I). Four separated methyl groups
resonance at high field
H 1.45 H3(21),
H 1.82 H3(131),
H 1.91 H3(171), and
H 1.97 H3(71). The fifth methyl group, which absorbs
at
H 3.69 H3(85), is assigned to
a methyl ester group, because of its chemical shift, line sharpness,
and intensity. The highest resonance frequency was observed at
H 4.74 H(10); this excludes the presence of an aldehyde
group. Two-dimensional {1H,1H}- and
{1H,13C}-correlation spectra show the
presence of two ethylene groups and three AMX systems, the X
portions of which have resonances at
H 4.16 H(16),
H4.32 H(4), and
H 4.44 H(181), respectively. At
C 35.35 a weak CH
cross-peak was found in the
{1H,13C}-correlation spectrum with the
H(10) at
H 4.74 ppm.
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Although the proton part of the spectra of the compound when measured
in Me2SO-d6 as solvent showed
considerable lower resolution, the missing proton became clearly
visible at H 3.88 H(82). This proton couples
with a frequency of 3.35 Hz with a proton at
H
4.65 H(10). The {1H,13C}-correlation
spectrum showed in addition a cross-peak indicating a tertiary carbon
atom at
C 65.44 C(82), which correlates with
the H(82) proton at
H 3.88. This indicates
that this carbon-bound proton is in dynamic equilibrium (keto-enol) and
rapid exchange with deuterium from the solvent D2O. This
"blind spot" has been previously observed when the plant catabolite
5 was measured in D2O (20).
In the COSY spectrum (Fig. 2), particularly the appearance of six cross-peaks caused by long range couplings of the four methyl groups facilitated the constitutional assignment. Together with the information gathered from the heteronuclear correllation (HETCOR) spectrum, they provided the reference and indicated the starting points of the specific side chains and the bridging network for each of the two dipyrrylmethanone units.
Nuclear Overhauser effect experiments corroborated the assignment of
the fragments and showed, in addition, that both units are
interconnected (Fig. 3 and Table I).
Thus, when the proton at H 4.74 H(10) was irradiated the
signal group at
H 2.45-2.50 H2(121) was enhanced. When the latter signal
was irradiated a methyl group H3(132) and a
methylene group H2(122) responded through
space.
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The overall bonding network and chemical shift assignments of the
quaternary carbon atoms were deduced from correlation
spectroscopy via long-range coupling (COLOC)
spectra. The relaxation times used for the evolution of the long range
couplings were equivalent to 1H,13C coupling
constants of 5, 10, and 20 Hz, respectively. Although two quaternary
carbon atoms at C 191.77 and
C 160.19 showed no correlation, the former value was assigned for chemical shift reasoning to the carbonyl group C(81) and the latter
consequently to the remaining aromatic carbon atom C(9). Thus, chemical
degradation, mass, nuclear magnetic resonance, and UV spectra are
consistent for the constitution of a potassium
31,181,182-trihydroxy-85-methyl-81-oxo-82,10-cyclourobilinogen
(8) shown in Fig. 4.
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The third tetrapyrrole was isolated in about 10 mg and showed in the
mass spectrometer a molecular mass ion at m/z 705.2534 atomic mass units (100%), indicating the presence of a configurational isomer of the former. 1H,13C NMR spectroscopic
investigation corroborates a diastereomeric relationship. A 1:1 mixture
of both diastereomers shows in 1H NMR only slight
differences in chemical shifts. Two methyl groups (21) and
(171) are shifted relative 0.079 ppm to lower field and
0.0134 ppm to higher field, respectively, the rest of the spectrum
remains nearly identical. The CD spectrum shows that this compound too is optically active and that both diastereomers are chiroptically almost indistinguishable (Fig. 5).
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DISCUSSION |
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The new catabolites isolated from yellow cotyledons of barley are
colorless, nonfluorescent, and optically active. The constitution resembles Urobilinogen IX, a common metabolite of heme
catabolism in warm-blooded organisms. The similarities in the
peripheral substitution pattern of both catabolites indicate that the
new catabolite derives from catabolite 5 by an oxidative process.
The configurations of the five asymmetric centers cannot be deduced
from the spectroscopic data. Suitable crystals are not available yet.
Only one stereocenter at C(82) has its origin from the
Chls, but due to the -keto ester functional group, it is prone to
epimerization. When measured in
Me2SO-d6, a coupling of only 3.35 Hz
between H(10) and H(82) indicates anticonfiguration.
As found previously, proton H(15) in formula 5 arrives from the protic solvent during acid-catalyzed pyrroline/pyrrole rearrangement of both geometric isomers of the red chlorophyll catabolite from C. protothecoides (11, 16). Intensive mechanistic-chemical studies showed a remarkable high stereoselectivity for this rearrangement. Because of the stereochemical consequence of these experiments, this position was tentatively assigned an R configuration. The proton at this position is highly resistant, even in boiling acetic acid-d1 it does not exchange (11, 16). It is most likely that both diastereomers arise from the uniformly configured 5 previously isolated from barley leaves by a consecutive oxidative process.
Chemically, the cleavage of the formyl group of the main red chlorophyll catabolite from C. protothecoides (3) was achieved using H2O2/pyridine at 70 °C. Both possible C(4) diastereomers of the corresponding biliverdine were obtained from this reaction in a yield of 48% (16) and in a relation of 1:1. A comparable enzymatic formyl cleavage can be envisioned, for example, through a Baeyer-Villiger reaction with subsequent hydrolysis of the formiate ester group or through an oxidative decarboxylation mechanism. Nevertheless, the terminal protonation at C(4) is irreversible (vide infra) under those conditions (Fig. 4). Both diastereomers are observed in a relation of 1:0.6 as determined by HPLC. This indicates that this protonation most likely is nonenzymatically. Instead, the relation reflects the different rate constants of protonation of both diastereotopic faces of the double bond C(3)=C(4) in 9.
That the catabolite 8 is produced in the leaf tissue of barley and not during the work-up procedure was demonstrated by an experiment in which the whole work-up procedure, including cell opening and HPLC separation, was performed in the presence of heavy water (30 atom % D). 1H and 2H NMR experiments showed that all carbon bond hydrogen atoms remained unlabeled; exchange occurred, as expected, only at position C(82). Moreover, a mirror experiment in which the leaves were bleached in the presence of heavy water (80 atom % D) showed by 2H NMR that a deuterium atom resides in position C(4) of compound 8 (data not shown). These results demonstrate that the catabolite is formed entirely in the plant cell and not during the work-up procedure and further that H(4) is tautomerizationally stable under those conditions. Skeletal transformation between the two valence tautomers of urobilin/bilirhodin was reported to be reversible only under basic conditions (21).
From the Chlorophyte Bryopsis maxima a red, water-soluble tetrapyrrole was isolated (22), and a portion of structural information was published (23). The proposed constitution, a biliviolin, lacks the formyl group too and contains a phytol group and a sugar moiety. Nevertheless, the UV-visible spectrum is in disagreement with the constitution, which remains to be determined.
In Cercidiphyllum japonicum almost quantitative amounts of the chls were isolated as catabolite 6 (8), whereas in this work only about 20% of the tetrapyrrolic catabolites were recovered. As charged from a screening test of several autumnal plants using the chromic acid degradation method, (tetra)pyrroles are not always present. This raises the questions whether urobilinogenoids are peculiar or common intermediates in the catabolic pathway of the chls and whether they are finally degraded in the plant cell to nonpyrrolic, nitrogen-less compounds.
Most recently, several maleimides were isolated from senescent
cotyledones of barley (24). Therefore, it appears that phototrophic organisms are capable to degrade chls to maleimides derivatives (possibly via urobilnogenoids), a pathway that has already been discussed by Hendry et al. (25). Maleimides still contain
the nitrogen atoms of the former Chls. Can this nitrogen be re-utilized by the plant? This adaptation would certainly be of evolutionary advantage for plants, which grow in environments in which nitrogen is
in high demand. Nevertheless, the ultimate fate of the Chls is still a
matter of speculation.
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ACKNOWLEDGEMENTS |
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We appreciate the professional knowledge of F. Nydegger and F. Fehr to obtain optimal mass and NMR spectra. We are indebted to Florimond Desprez, Cappelle en Pévèle F-59242, for providing us with H. vulgare cv. Lambic.
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FOOTNOTES |
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* This work was supported by the Swiss National Science Foundation (Project No. 2000-50725.97/1).The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
To whom correspondence should be addressed. Tel.: 41-26-300-8785;
Fax: 41-26-300-9739; E-mail: norbert.engel@unifr.ch.
Published, JBC Papers in Press, December 13, 2000, DOI 10.1074/jbc.M009288200
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ABBREVIATIONS |
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The abbreviations used are: Chl(s), chlorophyll(s); MPLC and HPLC, medium and high pressure liquid chromatography, respectively.
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REFERENCES |
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1. | Stahl, E. (1909) in Zur Biologie des Chlorophylls, Laubfarbe und Himmelslicht (Stahl, E., ed) , G. Fischer Verlag, Jena, Germany |
2. | Peoples, M. B., and Dalling, M. J. (1988) in Senescence and Aging in Plants (Noodén, L. D. , and Leopold, A. C., eds) , pp. 181-217, Academic Press, New York |
3. | Engel, N., Jenny, T. A., Mooser, V., and Gossauer, A. (1991) FEBS Lett. 293, 131-133[CrossRef][Medline] [Order article via Infotrieve] |
4. | Kräutler, B., Jaun, B., Bortlik, K., Schellenberg, M., and Matile, P. (1991) Angew. Chem. Int. Ed. Engl. 30, 1315-1318 |
5. | Gossauer, A., and Engel, N. (1996) J. Photochem. Photobiol. B Biol. 32, 141-151[CrossRef] |
6. | Kräutler, B., and Matile, P. (1999) Acc. Chem. Res. 32, 35-43[CrossRef] |
7. | Iturraspe, J., Moyano, N., and Frydman, B. (1995) J. Org. Chem. 60, 6664-6665 |
8. | Curty, C., and Engel, N. (1996) Phytochemistry 42, 1531-1536[CrossRef] |
9. | Campbell, N. A. (1990) in Biology (Campbell, N. A., ed) , pp. 566-567, The Benjamin/Cummings Publishing Company, Inc., Redwood City, CA |
10. | van den Hoek, C., Mann, D. G., and Jahns, H. M. (1995) in Algae. An Introduction to Phycology (van den Hoek, C. , Mann, D. G. , and Jahns, H. M., eds) , pp. 483-512, Cambridge University Press |
11. | Engel, N., Curty, C., and Gossauer, A. (1996) Plant Physiol. Biochem. 34, 77-83 |
12. | Rosybel, D., Hörtensteiner, S., Donnison, I., Bird, C. R., and Seymour, G. B. (1999) Physiol. Plant. 107, 32-38[CrossRef] |
13. | Curty, C., Engel, N., and Gossauer, A. (1995) FEBS Lett. 364, 41-44[CrossRef][Medline] [Order article via Infotrieve] |
14. |
Hörtensteiner, S.,
Wüthrich, K. L.,
Matile, P.,
Ongania, K.-H.,
and Kräutler, B.
(1998)
J. Biol. Chem.
273,
15335-15339 |
15. | Curty, C., and Engel, N. (1997) Plant Physiol. Biochem. 35, 707-711 |
16. | Curty, C. (1996) Chlorophyll Catabolism in Plants: Chlorella protothecoides and Cercidiphyllum. Ph.D. thesis No. 1143 , University Fribourg, Switzerland |
17. |
Folly, P.,
and Engel, N.
(1999)
J. Biol. Chem.
274,
21811-21816 |
18. | Moss, G. P. (1987) Pure Appl. Chem. 59, 779-832 |
19. | Merritt, J., and Loening, K. L. (1979) Pure Appl. Chem. 51, 2251-2304 |
20. | Kräutler, B., Jaun, B., Amrein, W., Bortlik, K., Schellenberg, M., and Matile, P. (1992) Plant Physiol. Biochem. 30, 333-346 |
21. | Gossauer, A., and Plieninger, H. (1979) in The Porphyrins (Dolphin, D., ed), Vol. 6 , pp. 625-627, Academic Press, New York |
22. | Okada, M., Ohtomi, M., Nakayama, K., Sakamoto, Y., Satoh, H., Konagaya, M., and Yoshizaki, M. (1987) Plant Cell Physiol. 28, 1435-1441 |
23. | Miyake, K., Ohtomi, M., Yoshizawa, H., Sakamoto, Y., Nakayama, K., and Okada, M. (1995) Plant Cell Physiol. 36, 109-113 |
24. | Suzuki, Y., and Shioi, Y. (1999) Plant Cell Physiol. 40, 909-915 |
25. | Hendry, G. A. F., Houghton, J. D., and Brown, S. B. (1987) New Phytol. 107, 255-302 |