Protoporphyrinogen oxidation, an enzymatic step in heme and chlorophyll synthesis: Partial characterization of the reaction in plant organelles and comparison with mammalian and bacterial systems

ARCHIVES OF BIOCHEMISTRY AND BIOPHYSICS Vol. 229, No. 1, February 15, pp. 312-319, 1984

Protoporphyrinogen Oxidation, an Enzymatic Step in Heme and Chlorophyll Synthesis: Partial Characterization of the Reaction in Plant Organelles and Comparison with Mammalian and Bacterial Systems’ J. M. JACOBS AND N. J. JACOBS2 Department

of Microbiology, Received August

Dartmouth

Medical

School, Hawver,

New Hampshire

03756

12, 1983, and in revised form October 17, 1983

High rates of oxidation of protoporphyrinogen to protoporphyrin were demonstrable in etioplasts, chloroplasts, and mitochondria from young barley shoots. Much lower rates were observed in chloroplasts from older barley or mature spinach, in mitochondria from potatoes or rat liver, and in membranes from the bacteria Escherichia coli and Rhodopseudomonas spheroides. The presence of high activity in cells capable of rapid synthesis of large amounts of chlorophyll suggests a role for this activity in chlorophyll synthesis. Characteristics of the plant protoporphyrinogen-oxidizing activity were compared to the activity in rat liver mitochondria. The activity in spinach chloroplasts exhibited a pH optimum of 7, which was lower than that of the mammalian enzyme. The plant activity was more sensitive to inhibition by glutathione or excess detergent, and was more readily inactivated at room temperature. The plant activity exhibited less specificity toward porphyrinogen substrates, oxidizing mesoporphyrinogen as rapidly as protoporphyrinogen. The mammalian enzyme oxidized mesoporphyrinogen slowly, and neither system oxidized coproporphyrinogen or uroporphyrinogen. Both the plant and the mammalian activity were bound to organelle membranes, but could be extracted with detergents. In contrast, activity from membranes of the bacteria E. coli and R. spheroides was inactivated by detergent treatment. The plant extracts could be fractionated with ammonium sulfate and retained activity after dialysis or Sephadex G25 treatment, suggesting no readily dissociable cofactor. The activity extracted from spinach chloroplasts was mostly inactivated by trypsin digestion, which was additional evidence for the protein nature of the plant activity.

The oxidation of protoporphyrinogen to protoporphyrin is a step common to the heme and chlorophyll synthesis pathways (Fig. 1). Although the oxidation can occur nonenzymatically, an enzyme has been demonstrated (l-6) and partially purified and characterized (3, 4) in yeast and liver mitochondria. Evidence for an enzymatic reaction has also been found in bacterial cells (7-13) and other mammalian tissue (14, 15). We previously demonstrated a 1 Supported by National Science Foundation Grant PCM 79 19520. *To whom all correspondence should be addressed. 0003-9861/84 $3.00 Copyright Al1 nyhts

C 1984 by Academic Press, Inc of reproduction in any form reserved

quinone involvement in anaerobic protoporphyrinogen oxidation in Escherichia coli (12). In plants, the presence of an enzyme was suggested by chemical findings (16) and our initial enzymatic studies with purified barley etioplasts and mitochondria (1’7). However, the enzyme from chlorophyll-containing cells has not been characterized, and many questions remain about the mechanism of this reaction in mammalian and especially bacterial and plant systems. Two important physiological questions are whether this enzyme is important for chlorophyll as well as heme synthesis and whether this enzyme exhib312

ENZYMATIC 8-Amlnolevullnlc

PROTOPORPHYRINOGEN

Acld-tPorphoblllnogen-Uroporphyrlnoqen

I

-4co2

Profoporphyrlnaqen

-2 co* -4H

-CCoproporphyrInoqen

i

Protoporphyrln

of heme and chlorophyll

synthesis.

its any unique properties in tissues capable of rapidly synthesizing large amounts of chlorophyll as compared to tissues which can only synthesize heme. This enzyme could play a regulatory role in chlorophyll synthesis since it is located near the branch point of the chlorophyll and heme synthesis pathway (Fig. 1). MATERIALS

AND

phyrinogen (11, 17, 19). Standard solutions of each porphyrin were analyzed spectrophotometrically USing appropriate extinction values (20) before use to construct standard curves for fluorometric determination (17, 19). Superoxide dismutase (type I from bovine blood) and catalase (from bovine liver) were from Sigma Chemical Company, St. Louis, Missouri. RESULTS

Mq-Proloporphyrln---Chlorophyll

FIG. 1. Pathway

313

OXIDATION

METHODS

Plant organelles, liver mitochondria, and bacterial membranes were prepared as previously described (g-12,17,18). Protoporphyrinogen oxidation was followed fluorometrically at room temperature in the standard assay previously described (17, 19), except that the volume of the assay mixture, which did not routinely contain added detergent, was 0.5 ml. To measure enzyme activity at pH 7,50 pmol of potassium phosphate, pH 5.4, was added as buffer before substrate addition, and the addition of the basic protoporphyrinogen solution brought the final assay pH to 7. Tris buffer was used to measure activity at pH 8.6. Glutathione (5 mM) was present to suppress protoporphyrinogen autooxidation unless indicated otherwise. To follow protoporphyrin formation, aliquots were removed into the detergent-containing measuring mixture (17,19) at recorded intervals of less than 10 min. A unit of enzyme activity was calculated as 1 nmol protoporphyrin formed/h. The nonenzymatic rate, determined with an identical amount of heatinactivated tissue (85°C for 15 min), was subtracted from the enzymatic rate (17, 19). As indicated, some tissues exhibited a more rapid rate of oxidation in the initial time interval, and this was used to calculate the activity. In surveying activity in various organelles, at least two levels of enzyme were assayed to insure an approximate linearity between activity and amount of enzyme. The oxidation of mesoporphyrinogen IX, coproporphyrinogen I, deuteroporphyrinogen, IX, and uroporphyrinogen I were measured by procedures described previously for the oxidation of protopor-

Efect of Assay Conditim on Protopwphyrinogen Oxidation Plant Organelles

by

Although our previous study demonstrated this activity in plant organelles, the optimal assay conditions were not defined (1’7). Figure 2 illustrates that barley etioplasts exhibit high rates of protoporphyrinogen oxidation. Oxidation is most rapid in the initial time interval, and is significantly inhibited by glutathione. This rapid, initial rate was not observed in our previous investigation (17), since most of those studies were conducted at higher pH values, where this phenomenon is much less apparent. For comparison, Fig. 2 also shows protoporphyrinogen oxidation by spinach chloroplasts and liver mitochondria, assayed under conditions optimal for these organelles. The barley etioplasts were tested at lower protein concentrations and are clearly more active than the organelles from spinach or liver. There are also differences in the kinetics of oxidation (Fig. 2). Some effects of glutathione and pH on plant organelles are shown in Table I. Etioplasts were considerably inhibited by glutathione and were more active at pH 7.3 than at 8.6. These effects of glutathione and pH were also observed with mitochondria from etiolated barley (data not shown). Spinach chloroplasts were also more active at pH 7 than at 8.6, but were less sensitive to glutathione inhibition (Table I). In contrast to barley organelles, the enzyme from liver mitochondria shows optimal activity at pH 8.5 and is not markedly inhibited by glutathione (4). These findings indicate that the optimal assay conditions are different for the enzyme

314

JACOBS

AND

JACOBS

Healed Controls

I

I

I

/

I

IO

20

30

50

70

Time (min) FIG. 2. Protoporphyrinogen oxidation by plant organelles. The assay conditions for barley etioplasts were at pH 7.3 and were as described under Materials and Methods. The solid line indicates assay in the absence of glutathione, and the dashed line assay in the presence of 5 mM glutathione. The circles indicate unheated barley etioplasts (415 pg) and the triangles indicate etioplasts (415 rg) heated at 85°C for 15 min. The broken line with square symbols (0) shows protoporphyrinogen oxidation by rat liver mitochondria (1.2 mg protein) obtained under assay conditions routinely used for this system (4, 19) at pH 8.6 in the presence of 5 mM glutathione. The dotted line with crosses (X) denotes oxidation by spinach chloroplasts (1.2 mg protein) at room temperature and pH 7.0 under the standard assay conditions with glutathione present. The heated controls for liver and spinach organelles were the same as shown for heated barley etioplasts in the presence of glutathione.

from various sources. We usually include glutathione in the assay mixture to minimize autooxidation, although this underestimates activity, especially in organelles from young barley. Approximate linearity between the amount of barley etioplasts added and the rate of oxidation was observed both in the TABLE

I

EFFECT OF pH AND GLUTATHIONE ON ENZYMATIC OXIDATION BY PLANT ORGANELLES

Enzyme source Barley etioplasts Barley etioplasts Spinach chloroplasts Spinach chloroplasts

Assay PH

Activity with glutathione”

Activity without glutathione”

8.6 7.3

32 58

154 223

8.6

3

6

7.0

5

8

DActivity expressed as nmol protoporphyrin formed/h/mg organelle protein. Assays conducted as described under Materials and Methods.

absence (Fig. 3) or the presence of glutathione (data not shown). The low rate of autooxidation, determined in the presence of heat-inactivated etioplasts, did not increase with increasing amounts of heated enzyme (Fig. 3). Levels of Activity in Young and Mature Plant Organelles and in Mammalian and Bacterial Systems Using these improved assay conditions, we determined whether there was a correlation between levels of this activity and capacity for rapid chlorophyll synthesis, such as shown by greening etioplasts from young plants. We found high activity in organelles from etiolated or young barley (Table II). Much lower activity was found in older barley chloroplasts, mature spinach chloroplasts, and in systems which do not synthesize chlorophyll such as liver mitochondria E. coli (Table II). We have tested several batches of spinach chloroplasts, and some have shown specific activities slightly higher than shown in Table

ENZYMATIC

PROTOPORPHYRINOGEN

315

OXIDATION

ugation (lOO,OOOg, 1 h), there was a significant solubilization of the activity from various plant organelles with both cholate and Triton X-100 (Table III). The specific activities of the detergent-extracted enzyme are slightly higher than in the original organelle (compare Tables II and III).

Preliminary Characterization of the Detergent-Extracted Enzyme mg protein in may FIG. 3. Effect of barley etioplast concentration on rate of protoporphyrinogen oxidation at neutral pH in the absence of glutathione. Assay conditions were as in Fig. 2, except that the assay pH was 6.8 rather than 7.3. The rate of activity was calculated from the nanomoles protoporphyrin formed in the initial time interval (within 10 min from the time of substrate addition) since the rate was most rapid at this time. The solid line indicates unheated enzyme, and the dashed line enzyme heated for 15 min at 85’C.

II, especially plants.

if

prepared

from

young

Extraction of the Plant Enzyme from the Membrane With Detergents Like the mammalian and bacterial enzyme (3,4,8-ll), the plant activity is bound to organelle membranes. For instance, after sonication of barley mitochondria, more than 90% of the enzyme units were sedimented with the membrane fraction following ultracentrifugation (lOO,OOOg,1 h). The enzyme can be solubilized from rat liver and yeast mitochondria with detergents (3,4). Although our earlier attempts were unsuccessful (17), we have now found that the plant enzyme can also be solubilized using lower levels of detergent than we previously employed. To achieve solubilization, we suspended barley etioplasts or mitochondria (7 mg protein/ml) or spinach chloroplasts (30 mg protein/ml) in 50 mM Tris, pH 7.6, containing 20% glycerol, followed by brief sonication and addition of a solution containing 10% detergent, 8% KCl, and 10 mM of the protease inhibitor phenylmethylsulfonyl fluoride to give a final detergent to protein ratio of 0.7. After stirring (3 h, 0°C) and centrif-

The extracted barley enzyme retained its sensitivity to glutathione, which caused a 60% inhibition of activity. The cholate-extracted barley enzyme could be fractionated with solid ammonium sulfate, with more than half the units recovered in the 40 to 80%-saturated fraction. Overnight dialysis (4°C) of the cholate-extracted spinach enzyme against 0.05 M Tris, pH 7.6, resulted in only a 20% loss of enzyme units. Passage of the extracted enzyme over a Sephadex G-25 column at 4°C resulted TABLE

II

SPECIFIC ACTIVITIES OF PROTOPORPHYRINOGEN OXIDATION IN VARIOUS TISSUES

Activity” Barley etioplasts Mitochondria from etiolated barleyb Chloroplasts from 6-day-old lightgrown barleyC Chloroplasts from ll-day-old lightgrown barley’ Chloroplasts from mature spinach Potato mitochondria Rat liver mitochondriad E. coli membranesd R. sphemides membranes”

62 40 40 18 8 14 10 9 3

’ The assay mixture was as described (17) at pH 7 with added glutathione. Rat liver mitochondria and the bacterial systems were assayed at 37°C. Activity is expressed as nmol protoporphyrin formed/h/mg/ protein. b Purified by sucrose gradient centrifugation (17). ‘Light-grown barley was grown for the days indicated in alternating day-night cycles (6 h/day in darkness). d Assayed at pH 8.5. “The membranes from R. spheroides were assayed as previously described by a spectrophotometric assay, in the absence of glutathione, at pH 7.6 (11).

316

JACOBS TABLE

AND

JACOBS

III

SOLUBILIZATION OF ACTIVITY FROM PLANT ORGANELLESWITHDETERGENTS

Enzyme source Barley etioplasts Barley etioplasts Spinach chloroplasts Spinach chloroplasts Chloroplasts from ll-day-old barley*

Detergent

Specific activity of soluble fraction”

Cholate Triton Cholate Triton

50 82 10 9

Triton

16

“Activity is expressed as nmol protoporphyrin formed/h/mg protein and is the average of at least two determinations. ‘Barley grown for 11 days in a day-night cycle.

in complete recovery of all enzyme units for the cholate-extracted enzyme from spinach chloroplasts, although a 17% loss of specific activity was noted for the Tritonextracted enzyme from barley etiopasts. This small loss was most likely due to the lability of the barley activity (see below). These results suggest a protein nature for the plant activity and the lack of a readily dissociable cofactor. The pH-activity curve of the cholate-extracted enzyme from spinach chloroplasts indicated a fairly sharp optimum at pH 7 (Fig. 4). The effect of assay temperature on the Triton extract from barley etioplasts and spinach chloroplasts is shown in Table IV. There is little activity at O”C, but activity at 25 and 37°C is nearly equal. The plant activity was labile, with the extracted etioplasts losing considerable activity when held at 37°C for 2 h (Table V). Similar treatment of the enzyme from rat liver mitochondria resulted in no loss of activity. The activity in spinach chloroplasts is more stable than in barley etioplasts (Table V). Digestion (1 h, 25°C) with trypsin (25 cLg/ mg extract protein) of the Triton extract of spinach chloroplasts resulted in a 90% loss of enzyme activity. No loss of activity occurred if the trypsin was inactivated by boiling. However, when the Triton extract of barley etioplasts was similarly treated with trypsin, a variable loss of approximately 50% of the activity was noted, but

I-

2

3

4

5

6

7

8

9

PH

FIG. 4. Effect of pH on activity of cholate-extracted spinach chloroplasts. The standard assay, with 1.5 mg of enzyme extracted with cholate from spinach chloroplasts, was used, with the following buffers used to obtain the desired final pH: 0.2 M glycine buffer, pH 2.8; 0.1 M glycine buffer, pH 3.9; 0.2 M acetate buffer, pH 5.1; 0.2 M acetate buffer, pH 6.3; 0.1 M phosphate buffer, pH 6.9; 0.1 M phosphate buffer, pH 7.3; and 0.1 M Tris buffer, pH 8.8. Each buffer was adjusted to a lower pH than indicated, but gave the final pH values shown when the basic protoporphyrinogen solution was added to the assay mixture to start the reaction. Glutathione (5 mM) was added to the assay mixture where indicated.

almost as much activity was lost due simply to enzyme lability. Further studies are needed to determine the significance of these differences between barley and spinach. TABLE

IV

EFFECTOFASSAY TEMPERATUREONACTIVITYOF EXTRACXED PLANTENZYME~ Protoporphyrin formed (nmol/h/mg) Enzyme source Barley etioplasts Spinach chloroplasts

37OC

25°C

0°C

152 9

131 7

28 2

a Assayed at pH 7.1 in the presence of 5 ITIM glutathione. Either 0.16 mg of Triton-extracted barley etioplasts, or 0.64 mg of Triton-extracted spinach chloroplast enzyme was present in the standard assay mixture.

ENZYMATIC TABLE

PROTOPORPHYRINOGEN

V

10 mg/l mg etioplast protein, caused approximately 50% inhibition of enzyme activity. Higher ratios of Tween 20 to protein caused increased inhibition. This observation is of importance since the mammalian protoporphyrinogen-oxidizing enzyme is not inhibited by these concentrations of Tween 20, and an assay for this enzyme in mammalian tissue utilizes these concentrations of Tween in the assay mixture to solubilize protoporphyrin (21).

LABILITY OF THE PLANT ACTIVITY

Treatment

Specific activity (nmol/h/mg)

Barley etioplasts Barley etioplasts Barley etioplasts

NOM! 1 h at 25°C 2 h at 37°C

62 49 25

Spinach chloroplasts Spinach chloroplasts Spinach chloroplasts

NOW 1 h at 25°C 2 h at 37°C

Fraction”

4.3 4.3 2.3

“All fractions are Triton extracts. In the assay mixture, etioplast extract was present at 0.18 mg and spinach chloroplast extract was present at 0.98 mg. The extracts were diluted with 0.1 ml of assay buffer and held at the temperatures and times indicated. Then the remaining components of the assay mixture were added and the standard assay was performed.

P-Hydroxymercuribenzoate (1 mM), a sulfhydryl reagent which is an inhibitor of the rat liver protoporphyrinogen oxidase (4), did not inhibit the activity in barley etioplasts or spinach chloroplasts. Neither salicylhydroxamic acid (4 PM), a respiratory inhibitor in some plants, nor cyanide (13) caused inhibition of protoporphyrinogen oxidation in barley mitochondria or etioplasts. Cyanide does inhibit this oxidation in the bacterium Rhodopseudomonas spheroides (11). No effect on the activity of barley mitochondria was observed when superoxide dismutase (20 pg/ml) or catalase (2.8 mg/ml) was added to the assay mixture. Tween 20, at a concentration of

Substrate Speci&ity for the Plant as Compared to the Rat Liver Enzyme The unphysiological dicarboxylic porphyrin mesoporphyrinogen is oxidized as rapidly as protoporphyrinogen by spinach chloroplasts and barley etioplasts (Table VI). Another dicarboxylic porphyrin, deuteroporphyrinogen, is oxidized less rapidly (Table VI). In contrast, liver mitochondria oxidize mesoporphyrinogen at only 20% of the rate of protoporphyrinogen (Table VI and Ref. (4)). The eight-carboxyl compound, uroporphyrinogen, and the fourcarboxyl compound coproporphyrinogen, are not oxidized at appreciable rates by either the plant or the liver enzymes (Table VI and Ref. (4)). Thus, the plant system exhibits less substrate specificity than the mammalian enzyme. of Some Properties of the Plant Enzyme to the Enzyme in E. coli

Comparison

In contrast to the plant enzyme, the E. coli enzyme shows approximately the same

TABLE SPECIFICITY OF THE PLANT AND MAMMALIAN

ENZYME

VI FOR VARIOUS

PORPHYRINOGEN

Specific activity (nmol porphyrin formed/h/mg

Spinach chloroplasts Liver mitochondria Barley etioplasts

317

OXIDATION

Proto

Meso

14 17 142

18 3 147

Deutero 4 3 82

“Enzymes were extracted with Triton. All assays contained 5 mM glutathione conditions optimal for protoporphyrinogen oxidation for each tissue. The plant 7 and 25”C, while the mammalian enzyme was measured at pH 8.7 and 37°C. * N.D., This was not determined for liver mitochondria.

SIJBSTRATES"

protein) Uro

Copro

0 N.D.* 2

1 0 14

and were conducted extracts were assayed

under at pH

318

JACOBS

AND

activity when assayed at pH 8.5 and 7.0, and is not markedly inhibited by glutathione (data not shown). All attempts to extract the E. coli enzyme with cholate resulted in no enzyme activity in either the solubilized fraction or the membrane residue. The activity in R. spheroides is also destroyed by contact with detergent (11). DISCUSSION

These findings extend and confirm the suggestion of our previous report (17), that chloroplasts and purified plant mitochondria oxidize protoporphyrinogen by a reaction which exhibits many characteristics of an enzyme. These characteristics include lability to heat, storage, and trypsin digestion; a linear relationship between activity and amount of organelle added; a pH optimum near neutrality; and extractability from the membrane by detergent. Activity was not lost following dialysis or passage over Sephadex G-25 and could be fractionated with ammonium sulfate. Although these characteristics suggest a membranebound enzyme without a readily dissociable cofactor, the exact nature of this reaction in plants must await attempts at purification and identification with a protein fraction, as has been achieved for the enzyme from liver and yeast mitochondria (3, 4). Furthermore, our present findings suggest that this oxidation may be specifically important in chlorophyll, as well as heme biosynthesis, since we found high activity in organelles such as etioplasts or chloroplasts from young barley. Much lower activity was found in chloroplasts from older barley, mature spinach, and in cells with only the capacity to synthesize heme such as liver mitochondria or E. coli. Etioplasts or developing chloroplasts from young plants are known to have the capacity for rapid chlorophyll synthesis from early precursors (22-24). Although suggestive, this finding does not prove a role for protoporphyrinogen oxidase in chlorophyll synthesis since etioplasts also contain small amounts of heme. An important question about our findings is whether the very high activity in young barley organelles represents a higher level

JACOBS

of the same activity that is found at a lower level in spinach chloroplasts. We did find some qualitative differences, in that the barley activity was more sensitive to inhibition by glutathione and was only partially destroyed by trypsin digestion. There are some interesting differences between the protoporphyrinogen-oxidizing enzyme from liver mitochondria and the plant activity. Although both are extractable from organelle membranes with detergents, the plant activity is more sensitive to glutathione inhibition and has a lower pH optimum. The kinetics of oxidation also differ in that the plant shows greatest activity in the initial time interval after substrate addition (Fig. 2). The significance of this is unknown. An interesting difference in substrate specificity was also found, with the plant using mesoporphyrinogen as rapidly as protoporphyrinogen. Mesoporphyrin, which is presumably not physiological, is a dicarboxylic porphyrin, like protoporphyrin, but differs from it in having ethyl rather than vinyl groups at positions 2 and 4 of the porphyrin ring. The liver enzyme utilized mesoporphyrinogen poorly. Further studies will be needed to determine if this difference in substrate specificity is a general property of the enzyme from green plant sources. It is interesting that mesoporphyrin is also a good substrate for the next enzyme in the heme biosynthetic pathway, ferrochelatase, which utilizes all dicarboxylic porphyrins as substrates for iron incorporation whether the enzyme is studied in animal, plant, or bacterial sources (25-29). Furthermore, our findings on substrate specificity suggest that the plant activity is not merely a reflection of a nonspecific chemical oxidant, since neither uroporphyrinogen, an eight-carboxyl porphyrin, nor coproporphyrinogen, a fourcarboxyl porphyrin, were oxidized by the plant extract. Since the plant enzyme has not been purified, we have not determined if mesoporphyrinogen and protoporphyrinogen are oxidized by the same enzyme. We have not yet determined if there are significant differences in the behavior of the enzyme activity in barley etioplasts as compared to barley mitochondria. In preliminary studies, the activity in both barley

ENZYMATIC

PROTOPORPHYRINOGEN

organelles behaved similarly, exhibiting marked inhibition by glutathione and greater activity at pH 7.0 than at pH 8.5. By these two criteria, barley mitochondria behaved more like barley etioplasts than like liver mitochondria, but this question has not been investigated carefully. The barley etioplasts used in these studies retained activity after purification on sucrose gradients (17), and were relatively free of cytochrome oxidase, a mitochondrial marker enzyme. The barley mitochondria, which exhibited high cytochrome oxidase activity, were also purified on sucrose gradients (17) to minimize contamination with etioplasts. The bacterial systems for oxidizing protoporphyrinogen have been less well characterized. Glutathione causes complete inhibition of this oxidation in extracts of photosynthetically grown R. spheroides (ll), but causes only partial inhibition of the plant and liver enzyme. Cyanide inhibits the oxidation in R. spheroides (11) but not in barley organelles. In the two bacteria studied, E. coli and R. spheroides, treatment of membranes with detergent to solubilize the enzyme has led only to irreversible inactivation. Since detergents can solubilize activity in mammalian and plant organelles, this observation could suggest that the bacterial systems require an intact electron transport system for activity, while the eucaryotic enzyme can be solubilized and still retain activity. Our previous studies suggested involvement of components of the electron transport chain in these bacteria (11, 12). We found menaquinone was absolutely required for the anaerobic oxidation of protoporphyrinogen with fumarate as electron acceptor in E. coli (12). Further studies on bacterial systems are needed to answer these questions. ACKNOWLEDGMENT We are grateful to Professor guidance concerning preparation organelles.

A. E. DeMaggio for and analysis of plant

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OXIDATION

319

3. POULSON, R., AND POLGLASE, W. J. (1975) J. BioL Chem. 250, 1269-1274. 4. POULSON, R. (1976) J. BioL Chem 251,3730-3733. 5. JACKSON, A. H., GAMES, D. E., COUCH, P., JACKSON, J. R., BELCHER, R. E., AND SMITH, S. (1975) Enzyme 17, 81-87. 6. CAMADRO, J. M., URBAN-GRIMAL, D., AND LABBE, P. (1982) Biochem. Biophgs. Res. Commun. 106, 724-730. 7. JACOBS, N. J., AND JACOBS, J. M. (1975) B&hem. Biophys. Res. Commun 65, 435-441. 8. POULSON, R., WHITLOW, K. J., AND POLGLASE, W. J. (1976) FEBS Z&t. 62, 351-353. 9. JACOBS, N. J., AND JACOBS, J. M. (1976) Biochim. Biophys. Acta 449, l-9. 10. JACOBS, N. J., AND JACOBS, J. M. (1979) Arch B&hem. Biophys. 197, 396-403. 11. JACOBS, N. J., AND JACOBS, J. M. (1981) Arch. Biochem. Biophys. 211,305-311. 12. JACOBS, N. J., AND JACOBS, J. M. (1978) B&him Biophys. Acta 544, 540-546. 13. SASARMAN, A., CHARTRAND, P., LAVOIE, M., TARDIF, D., PROSCHEK, R., AND LAPOINTE, C. (1979) J. Gem. MicrokoL 113, 397-403. 14. BRENNER, D. A., AND BLOOMER, J. R. (1980) N. Engl J. Med. 302, 765-769. 15. DEYBACH, J. C., DE VERNEUIL, H., ANDNORDMANN, Y. (1981) Human Genetics 58, 425-428. 16. BATTERSBY, A. R., MCDONALD, E., REDFERN, J. R., STAUNTON, J., AND WHIGHTMAN, R. H. (1976) J. Chem. Sot. Perkin Trans. 266-273. 17. JACOBS, J. M., AND JACOBS, N. J. (1982) Arch. Biochem. Biophys. 218,233-239. 18. LATIES, G. (1974) in Methods in Enzymology (Fleischer, S., and Packer, L., eds.), Vol. 31, pp. 589-600, Academic Press, New York. 19. JACOBS, N. J., AND JACOBS, J. M. (1982) Enzyme 28, 206-217. 20. FALK, J. E. (1964) Porphyrins and Metalloporphyrins, p. 236, Elsevier, Amsterdam/New York. 21. BRENNER, D. A., AND BLOOMER, J. R. (1980) Clin Chim. Acta 100, 259-266. 22. REIBEIZ, C. A., AND CASTELFRANCO, P. A. (1971) Plant PhysioL 47, 24-32. 23. GRANICK, S., AND BEALE, S. I. (1978). Advan EnzymoL ReL Areas MoL Biol 46, 33-203. 24. CASTELFRANCO, P. A., AND BEALE, S. I. (1981) in The Biochemistry of Plants (Stumpf, P. F., and Conn, E. E., eds.), Vol. 8, p. 375, Academic Press, New York. 25. LITTLE, H. N., ANDJONES, 0. T. G. (1976) B&hem. J. 156, 309-314. 26. PORRA, R. J., AND LASCELLES, J. (1968) B&hem. J. 108, 343-348. 27. DAILEY, H. A., AND LASCELLES, J. (1974) Arch Biochem. Biophgs. 160,523-529. 28. TAKETANE, S., AND TOKUNAGA, R. (1981) J BioL Chem. 256, 12748-12753. 29. DAILEY, H. A. (1982) .I Bid. Chem. 257, 1471414718.