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1984 Relationship of porphobilinogen deaminase activity to chlorophyll content and development of 'Heinz 1350' tomato fruit Robert Waring McMahon Iowa State University

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Recommended Citation McMahon, Robert Waring, "Relationship of porphobilinogen deaminase activity to chlorophyll content and development of 'Heinz 1350' tomato fruit " (1984). Retrospective Theses and Dissertations. 9010. https://lib.dr.iastate.edu/rtd/9010

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Uni International 300 N. Zeeb Road Ann Arbor, Ml 48106

8423655

McMahon, Robert Waring

RELATIONSHIP OF PORPHOBILINOGEN DEAMINASE ACTIVITY TO CHLOROPHYLL CONTENT AND DEVELOPMENT OF 'HEINZ 1350' TOMATO FRUIT

Iowa State University PH.D. 1984

University iVIicrofilms IntGrnBtiOn&l 300N.zeeb Road, Ann Arbor, Ml48106

Relationship of porphobilinogen deaminase

activity to chlorophyll content and development

of 'Heinz 1350' tomato fruit

by

Robert Waring McMahon

A Dissertation Submitted to the

Graduate Faculty in Partial Fulfillment of the

Requirements for the Degree of

DOCTOR OF PHILOSOPHY

Departments: Horticulture Botany Co-majors; Horticulture Botany (Physiology) Approved:

Signature was redacted for privacy.

In Charge of Major Work

Signature was redacted for privacy.

For the Major Departments

Signature was redacted for privacy. For the Gradu College

Iowa State University Ames, Iowa

1984 ii

TABLE OF CONTENTS

Page

GENERAL INTRODUCTION 1

GENERAL LITERATURE REVIEW 3

SECTION I. METHOD FOR THE EXTRACTION AND QUANTITATION 14 OF CHLOROPLAST PIGMENTS IN FRUITS OF HORTICULTURAL CROPS BY HIGH PERFORMANCE LIQUID CHROMATOGRAPHY (HPLC)

ABSTRACT 15

INTRODUCTION 16

MATERIALS AND METHODS 18

RESULTS AND DISCUSSION 20

LITERATURE CITED 27

SECTION II. PROPERTIES OF PORPHOBILINOGEN DEAMINASE 28 ISOLATED FROM 'HEINZ 1350' TOMATO FRUIT

ABSTRACT 29

INTRODUCTION 30

MATERIALS AND METHODS 31

RESULTS 35

DISCUSSION 44 iii

LITERATURE CITED 50

SECTION III. PORPHOBILINOGEN DEAMINASE ACTIVITY 53 IN DEVELOPING 'HEINZ 1350' TOMATO FRUIT AND ITS RELATIONSHIP TO TOMATO FRUIT CHLOROPHYLL CONTENT

ABSTRACT 5%

INTRODUCTION 55

MATERIALS AND METHODS 58

RESULTS 59

DISCUSSION 67

LITERATURE CITED 72

GENERAL SUMMARY AND CONCLUSIONS 73

LITERATURE CITED FOR THE GENERAL INTRODUCTION AND THE 7% GENERAL LITERATURE REVIEW

ACKNOWLEDGMENTS 79 1

GENERAL INTRODUCTION

One phenomenon common to most ripening fruits and vegetables is

the loss of chlorophyll, followed by carotenoid biosynthesis. Loss of

chlorophyll is caused by its degradation, and, in the case of

tomatoes, results in a white, star-shaped area surrounding the blossom

scar. At this point of fruit development, the tomato is mature green.

Immediately following this stage, carotenoid biosynthesis commences,

and lycopene and >9-carotene accumulate. The fruit then changes color

from light green to red-orange, and finally to red.

Chlorophyll is biosynthesized in fruits before the ripening

process begins. However, the point at which chlorophyll production stops in relation to fruit development and ripening is not known.

Furthermore, the pattern of biosynthesis is not known. Possible

patterns include: a constant rate of synthesis up to the time fruits

begin to ripen followed by an abrupt cessation; increasing rates of chlorophyll production to a maximum rate, followed by a decline in

biosynthetic rates to zero; and an intial rapid biosynthetic rate that gradually decreases during fruit development and no longer is present at the onset of ripening. Regardless of the pattern of chlorophyll biosynthesis, the subsequent degradation of chlorophyll that follows is associated with the ripening process.

Previous findings suggest that one of the early steps of the chlorophyll biosynthetic pathway may trigger the onset of senescence in leaves. Frydman and Frydman showed that porphobilinogen deaminase activity disappeared just before the onset of senescence in pepper and 2

poinsettia leaves, and they concluded that the loss of deaminase

activity may induce senescence (21). The ripening process of fruits

and vegetables immediately precedes senescence, and, if it can be

shown that deaminase activity disappears near the onset of ripening,

then this would suggest that the deaminase enzyme also is involved in ripening and senescence in fruits.

'Heinz 1350' tomato fruits were used for the following objectives: (1) to measure the chlorophyll a and b content of fruit during development and ripening; (2) to measure porphobilinogen deaminase activity in the fruit as a function of fruit development; and (3) to correlate deaminase activity with the chlorophyll content of the tomato fruit as a function of development.

Each of these objectives are covered in separate papers written in journal form, and the author was the sole contributor to the work accomplished in each of these papers, except for the editing of the papers. 3

GENERAL LITERATURE REVIEW

General Aspects of Chlorophyll Biochemistry

Chlorophylls are magnesium-porphyrin complexes that are the chief

pigments of photosynthesis. Their structure contains four substituted

pyrrole rings, with ring IV being reduced. The four rings are

combined in a macrocyclic structure in which the four central nitrogen

atoms of the pyrrole rings are coordinated with the central Mg^"*" ion.

A phytol chain is esterified to a propionic acid residue in ring IV,

and the propionic acid side chain at position 5 on ring III has been

modified to a cyclopentanone ring (28, 45).

Higher plants contain two types of chlorophyll: chlorophyll a

and chlorophyll b. The only difference between the two is that

chlorophyll b has a formyl group instead of a methyl group at position

3 of ring II. Other types of chlorophyll exist, including chlorophyll

c, found in brown algae, and chlorophyll d, found in red algae. Not

only are there different types of chlorophyll, but six forms of

chlorophyll a have been discovered, based on absorption maxima ranging

from 663 to 700 nm (45). The ratio of chlorophyll a to chlorophyll b

usually is approximately three in most higher plants (16, 45).

Chlorophyll Biosynthesis

Biosynthesis of ^-aminolevulinic acid

^-aminolevulinic acid (ALA) is the first intermediate unique to

the tetrapyrrole pathway leading to chlorophyll, heme and vitamin B^g

(1, 34). The synthesis of ALA in plants, however, seems to differ from that of animal systems. In the latter case, ALA is formed from 4 the condensation of succinyl CoA and glycine with the elimination of the carboxyl group of glycine (50). The enzyme involved is ALA synthetase, and, in plants, it only has been detected in extracts isolated from soybean callus (51). Attempts to isolate the enzyme from other plant sources have not been successful, and many researchers believe that a different biosynthetic pathway to ALA exists in plants (4, 6, 7, 8, 26, 36, 43).

In a study with greening cucumber cotyledons, Beale fed ^'Re­ labeled glycine and succinate to the cotyledons and found that ALA was labeled only to a small extent (4). In addition, both and Cg of glycine contributed equally to the labeling, which is not consistent with the ALA synthetase reaction, in which is eliminated as COg.

Identical results were obtained with greening barley and bean leaves.

Beale, in the same study, found that the five-carbon compounds glutamate, glutaraine and ot-ketoglutarate labeled ALA to a much greater extent than glycine. glutamate labeled the of ALA predominately, while 3, 4-^^C glutamate labeled the to portion of ALA. Both labeled forms of glutamate were able to label ALA equally. Thus, it was concluded that ALA is formed from the intact carbon skeleton of glutamate by a pathway that does not involve ALA synthetase. Other researchers have obtained similar results (5, 7,

25, 34, 50).

Harel et al. showed that extracts of greening leaves of maize, pea and barley converted oe-ketoglutarate- 5-^^C to ALA better than glutamate- or - , and the incorporation of label 5 from the labeled a-ketoglutarate into ALA was not reduced by the addition of glutamate to the reaction mixture (27). Thus, Harel et al. concluded that a-ketoglutarate, rather than glutamate, is the immediate precursor of ALA in the new pathway.

In support of this observation, Beale et al. (8) found similar results in greening barley and proposed the following pathway for ALA biosynthesis: glutamate is acted upon by glutamate transaminase, which forms oe-ketoglutarate (8). This compound then is reduced, forming y,(5 dioxovalerate (DOVA), which subsequently is transaminated to ALA. As the existence of DOVA in plant tissues has not been demonstrated, it may exist ^ vivo only as an enzyme-bound intermediate.

Despite of increasing evidence against ALA synthetase functioning in plants. Wider de Xifra et al. isolated ALA synthetase from soybean callus (51). It also was demonstrated that glutamate, a-ketoglutarate and DOVA could not enhance ALA formation.

Regardless which enzyme system operates in plants, the formation of ALA is light-dependent and is one of the regulatory points of chlorophyll biosynthesis (6, 24, 25, 26, 35, 40, 49, 50, 51).

Biosynthesis of porphobilinogen

The next step in the biosynthetic pathway of chlorophyll is the conversion of ALA to porphobilinogen, and this reaction is catalyzed by the enzyme ALA dehydratase. Two molecules of ALA combine to form one molecule of porphobilinogen with the loss of two molecules of water. Porphobilinogen is the initial pyrrole synthesized in 6

chlorophyll formation. The enzyme becomes inactivated if its thiol

group(s) is (are) oxidized, and it consists of eight subunits. It is

thought that the dehydratase does not play a role in the regulation of chlorophyll biosynthesis (42).

Biosynthesis of uroporphyrinogen III

One of the most intensively studied steps of the chlorophyll biosynthetic pathway is the formation of one molecule of uroporphyrinogen III from four molecules of porphobilinogen, accompanied by the loss of four molecules of ammonia. Two enzymes are required for this step: porphobilinogen deaminase and uroporphyrinogen III cosynthase. Uroporphyrinogens are cyclic tetrapyrroles with saturated bridges between the four rings. To each ring is attached one acetic acid and one propionic acid side chain.

Four uroporphyrinogen isomers are possible, but only the type I and type III isomers have been found in living organisms (2, 11, 45). In the type I isomer, the acetic and propionic acid side chains are located in a head-to-tail fashion on all four rings, but in the type

III isomer, the positions of the two side chains are reversed in ring

IV with respect to the other three rings (1, 3, 18, 23). The mechanism by which this inversion of the two side chains occurs has not been precisely defined, although many research projects have addressed this problem.

Burton et al. used labeling in his experiments, and the results have suggested that a very short-lived intermediate was formed, and called "pre-uro'gen" (14). Jordan et al. researched this 7

subject further and believe that "pre-uro'gen" is formed by the

deaminase enzyme (31). The compound is thought to be a tetrapyrrole

because porphobilinogen was not consumed by its conversion to

uroporphyrinogen III. Rearrangement of ring IV then occurs by the

action of the cosynthase enzyme. In reaction mixtures where the

cosynthase was absent, spontaneous formation of uroporphyrinogen I

occurred. It also was shown that the type I isomer was not a

substrate for the cosynthase (31).

Frydman et al., however, believe that a tripyrrole most likely is

the intermediate in the enzymatic production of uroporphyrinogen III

(19, 23). They propose that two molecules of porphobilinogen condense

in a head to head fashion and that this condensation forms an enzyme-

bound dipyrrylmethane. Two more porphobilinogen molecules are

attached, forming first a tripyrrole, and then a bilane, which

cyclizes to uroporphyrinogen III. The tripyrrole specifically is

incorporated into the type III isomer and strongly inhibits type I

isomer formation. Several other studies have been conducted on this

topic, and more research will be required to clearly define the

mechanism that is responsible for uroporphyrinogen III biosynthesis

(1, 2, 3, 18, 29, 39).

Uroporphyrinogen III only is formed when the deaminase and

cosynthase enzymes are present together (23, 42, 43, 52). While the

deaminase by itself will consume porphobilinogen and produce

uroporphyrinogen I (14, 17, 20, 21, 30, 32), no uroporphyrinogen is formed when the cosynthase is incubated with porphobilinogen. This is 8

because porphobilinogen is not consumed by the cosynthase, and,

furthermore, it will not convert uroporphyrinogen I to the type III

isomer (9, 23, 37, 38, 46). The type I isomer is not a precursor for

chlorophyll and its only known metabolic fate is its decarboxylation

to coproporphyrinogen I. On the other hand, the type III isomer is an

intermediate in porphyrin biosynthesis in most living organisms (11).

Because the cosynthase enzyme cannot be assayed directly, the

deaminase enzyme has been characterized extensively. The

characterization of the deaminase is based on its formation of

uroporphyrinogen I. The deaminase enzyme has a very high affinity for its substrate, porphobilinogen. Km values reported for uroporphyrinogen I production include: 6.6/iM from pepper and poinsettia leaf extracts (21), 70/LlM from wheat germ (20), 13 from

Rhodopseudomonas spheroides (30), and 85 juM from Chlorella regularis

(46).

The deaminase is very heat-stable, and maximum activity has been demonstrated at 65C in extracts isolated from Chlorella regularis (46) and 67c in wheat germ extracts (20). The deaminase enzyme lost only fifteen percent of its activity when incubated for fifteen minutes at

75c (20), and similar results were reported by Shioi et al. (46). The cosynthase enzyme, however, loses most of its activity at 50C, and loses all of its activity at 50C (48). Bogorad showed that cosynthase from wheat germ was inactivated at 55C (9). The temperature at which the cosynthase showed maximum activity was 37C, which is considerably lower than that of the deaminase enzyme (48). 9

Reported pH optima for deaminase activity in phosphate and tris

buffers include: 7.2 (38), 7.6 (30), and 8.2 (20). A pH optimum of

8.2 has been shown for cosynthase (48).

When both enzymes are assayed together ^ vitro, the rate of cosynthase activity rapidly decreases to zero, while the deaminase enzyme activity continues at an undiminished rate (37). Only when the cosynthase is present in excess does uroporphyrinogen III production continue without a decrease (13, 53).

Molecular weight determinations have been performed on both enzymes. A molecular weight of 60,000 to 65,000 daltons was found for cosynthase from wheat germ (22), while molecular weights of deaminase have ranged from 35,000 to 45,000 (2, 15, 30, 39). In addition, it is believed that the deaminase enzyme is composed of one polypeptide chain (2, 30).

The deaminase enzyme is very specific for porphobilinogen.

Frydman and Frydraan incubated the enzyme with several analogs of porphobilinogen (20). The analogs differed from porphobilinogen only by the propionic acid side chain, in that an acetic acid side chain or a methyl group was substituted. The analogs inhibited the enzyme, suggesting that the propionic acid side chain of porphobilinogen is crucial for the substrate specificity of the enzyme.

Porphobilinogen deaminase is a sulfhydryl enzyme as determined by inhibitor studies because the deaminase was inhibited by the sulfhydryl reagents £-chloromercuribenzoate and N-ethylmaleimide (46).

Frydman and Frydman found that the inhibitory effects of sulfhydryl 10

reagents were more pronounced on porphyrin formation than on the

consumption of porphobilinogen (20). The inhibitory effects were

reversed by the addition of dithiothreitol and 2-mercaptoethanol,

which reduce the thiol groups of the active center. Also,

porphobilinogen-consuming activity could not be separated from

uroporphyrinogen-forming activity, and it was concluded that the

biosynthesis of uroporphyrinogen I occurs by only one enzyme that has

two active centers. The first active center polymerizes

porphobilinogen to polypyrroles, and the second active center cyclizes

them to porphyrins. N-ethylmaleimide was more inhibitory for

porphyrin formation than porphobilinogen consumption, implying the

presence of more sensitive thiol groups in the second active center.

Jordan and Shemin, however, believe that the deaminase enzyme has one

catalytic site that is involved four times during the biosynthesis of one molecule of uroporphyrinogen (30).

There is evidence that porphobilinogen deaminase may play a role in the regulation of porphyrin biosynthesis. The activity of the enzyme was very low in senescent and mature poinsettia and pepper leaves, and was high in young leaves. The cosynthase activity was the same in all three ages of the leaves, and the strong decrease in deaminase activity occurred after chlorophyll formation reached a maximum in the leaves, and it was concluded that the deaminase enzyme may regulate chlorophyll biosynthesis in leaves as a function of age

(21). Brodie et al. also believe that the deaminase plays a role in the regulation of porphyrin biosynthesis (12). Because 11

uroporphyrinogen III oosynthase activity normally is ten times greater

than that of the deaminase, the deaminase activity may be limiting and

changes in activity may provide a secondary control point in porphyrin

biosynthsis (13, 53).

Biosynthesis of coproporphyrinogen III

The next step in the biosynthesis of chlorophyll is mediated by

uroporphyrinogen III decarboxylase. This enzyme decarboxylates

uroporphyrinogen III at each of the four acetyl side chains and

coproporphyrinogen III is the end result. The decarboxylation of

uroporphyrinogen III occurs in a stepwise process (10), and it is

believed that the decarboxylation is in a clockwise fashion, starting

on ring IV and is followed by rings I, II and III (25). However,

Rebeiz and Castelfranco think that this process occurs randomly (43).

Biosynthesis of protoporphyrin IX

The propionic acid side chains on rings I and II of

coproporphyrinogen III then are decarboxylated forming vinyl groups,

and this step is mediated by coproporphyrinogen III oxidative

decarboxylase. The resulting intermediate, protoporphyrinogen IX, is oxidized by the loss of six hydrogen atoms from the ring system to form protoporphyrin IX. Whether or not the last step is enzymatic or is a spontaneous oxidation is not clear (25). The formation of protoporphyrinogen IX is the last common step of the iron- and magnesium-porphyrin biosynthetic routes, with the exception of ALA biosynthesis. Beyond this step, magnesium is inserted, leading to chlorophyll biosynthesis, or iron is inserted, leading to heme and 12 cytochrome production.

Final steps of chlorophyll biosynthesis

The remaining key steps in chlorophyll biosynthesis are: 1) the insertion of into protoporphyrin IX; 2) the conversion of magnesium protoporphyrin IX monomethyl ester to protochlorophyllide, which involves the formation of the cyclopentanone ring at position six of the macrocycle and the reduction of the vinyl group to an ethyl group at position four on ring II; 3) the photoreduction of protochlorophyllide to chlorophyllide; and 4) the esterification of the phytol tail onto chlorophyllide to form chlorophyll (2, 6, 10, 25,

36, 43, 53).

Tomato Fruit Development and Ripening

Martin et al. have classified the development of 'Heinz 1350' tomato fruit into seven stages: 1) cell division from zero to twenty days past anthesis; 2) cell enlargement from from 20 to 35 days past anthesis; 3) 36 to 40 days, mature green, which is characterized by a white, star-shaped design radiating from the blossom end scar, indicating chlorophyll degradation; 4) breaker stage from 40 to 45 days past anthesis, characterized by the development of pink color immediately around the blossom end scar; 5) light pink from 43 to 48 days past anthesis; 6) dark pink from 45 to 50 days past anthesis; and

7) red ripe from 50 to 55 days past anthesis (41).

During the early stages of tomato fruit development, rapid fruit growth occurs, accompanied by chlorophyll and protein biosynthesis.

However, as the fruit begins to ripen, several changes occur. 13

Chlorophyll degrades rapidly and carotenoid biosynthesis commences, producing>S-carotene and lycopene (33, 44, 47), which are responsible for the red color of the fruit. During this time of fruit development, the climacteric occurs. This is marked by the sudden and brief increase in the ethylene production and respiration rates that marks the beginning of senescence of the tomato fruit. 14

SECTION I. METHOD FOR THE EXTRACTION AND QUANTITATION

OF CHLOROPLAST PIGMENTS IN FRUITS OF HORTICULTURAL CROPS

BY HIGH PERFORMANCE LIQUID CHROMATOGRAPHY (HPLC) 15

ABSTRACT

Methods of chloroplast pigment extraction and sample preparation

previously established for leaves do not apply for fruits of several horticultural crops. Methods presented in this report overcome these difficulties, and results are presented that show this procedure can be used for several diverse fruits and vegetables. An HPLC run time of 25 minutes in the isocratic mode gives baseline separation of chlorophyll a, chlorophyll b, and y5-carotene while neoxanthin, violaxanthin, and lutein eluted as partially resolved peaks. A run time of 48 minutes in the linear gradient mode gives baseline separation of all 5 major chloroplast pigments. 16

INTRODUCTION

HPLC has reduced the time required for the precise separation and

quantitation of plant pigments when compared with thin layer or column

chromatography (7). Extraction and HPLC quantitation methods have

been reported for chloroplast pigments from algae (1, 2) and spinach,

bean, and soybean leaves (3, 4, 5, 8), but have not been developed for

chloroplast pigments contained in fruits and vegetables.

It was found that the previously described procedure (3, 5) for

pigment extraction works well for leaves, but that it is not effective

for fruits, especially tomato fruits. In the previously described

procedure, the tissue is ground with a glass tissue grinder in 70%

methanol and the cellular debris is washed with additional 70%

methanol to remove the remaining pigments (3, 5). The extract is filtered through a 0.5^ filter into a C^g Sep-Pal^ cartridge which retains the pigments. Stepwise elution of the Sep-Pal^ cartridge with

90% methanol removes the xanthophylls, 100% methanol removes chlorophyll a and b, and yS-carotene is eluted with acetone. The pigments are eluted into an injector vial and then are ready for HPLC.

Results of the research in this report showed that: 1) this procedure did not remove all of the pigments from the fruit tissue as it did for the leaf tissue (by visual observation); and 2) the pigments that were removed formed an emulsion in the 70% methanol. This emulsion caused only partial retention of the pigments in the Sep-Pal^ cartridges, and this led to errors in pigment quantitation. Attempts to circumvent these problems by grinding the tomato fruit in 100% methanol were not 17

successful, as the pigments still were not removed totally and

dilution of the extract to 70% methanol formed an emulsion. In

addition, it was found that the O.Sfi filter became irreversibly

blocked by cellular residue.

The speed at which many breeding and physiology research projects dealing with the chloroplast pigments of fruits and vegetables could

be increased if a technique for the rapid analysis of these pigments

by HPLC were available. The objectives of this research were to develop suitable extraction, sample preparation, and quantitation procedures for the determination of chloroplast pigments in tomato fruits by HPLC, and to test this procedure on a diverse group of other fruits and vegetables. 18

MATERIALS AND METHODS

'Heinz 1350' tomato plants were trained to a single stem, and the

pollination procedure of Lyons and Pratt was followed to obtain fruit

of the same physiological class (6). Three replicates of four fruits

of the same physiological age were harvested in 5 day increments from

15 days,past anthesis to fully ripe (50 to 55 days). Each replicate

was weighed, and, depending on fruit size, a quarter or half of each

fruit was used for extraction and sample preparation. Normally, the

sample size for extraction was 50 to 60 grams. All developmental work

was done on tomatoes produced by the above procedure; other fruits and

vegetables were purchased locally.

To overcome the difficulties cited above, samples were

homogenized for 3 minutes in 500 ml of -IOC 2:1 (v/v)

methanol:petroleum ether (38 to 54C boiling range) (9), transferred

to a 1000 ml beaker, and allowed to stand until the homogenate

separated into 2 phases. The homogenate was filtered into a 2000 ml

suction flask through a layer of glass wool over Whatman no. 1 filter

paper. The residue was washed 3 to 4 times with methanol-petroleum

ether mixture to extract the remaining pigments. The filtrate was

transferred to a séparatory funnel, an equal volume of 10^ sodium

chloride was added, and the chloroplast pigments were transferred to

the petroleum ether phase. The pigment-containing petroleum ether

phase was filtered through phase-separatory filter paper (Whatman PS

1) into an evaporation flask. Petroleum ether was removed with a rotary evaporator (bath temperature: 30C), and the pigments were 19 immediately dissolved in 75 ml of HPLC-grade methanol. Care must be exercised to not proceed to dryness for the pigments will cake onto the inside of the flask and will not be removed by the methanol. The extract was filtered through a O.SjH filter (Catalog No. SLSR025NS,

Millipore Corp., Bedford, MA 01730) into an injector vial for subsequent chromatography.

Chloroplast pigments were separated and quantitated using a

Waters Associates ALC-244 HPLC and a reverse phase /iBondapak-C^g analytical column (3.9 mm i.d. x 30 cm). In both the isocratic and linear gradient modes, a solvent flow rate of 1 ml/minute was used with detection by absorbance at 436 nm. Chlorophyll a and b concentration was determined by external standard quantitation using previously described methods to obtain the chlorophyll a and b standards (9). Research in this laboratory is related to chlorophyll a and b and only these pigments were quantified; however, standards for the other pigments can be obtained by these same methods (9). 20

RESULTS AND DISCUSSION

This research shows that chlorophyll a and b could be separated

and quantitated using an isocratic system of 50? solvent A (80%

methanol) and 50% solvent B (ethyl acetate); in addition, this system

gave baseline separation of >S-carotene (Figure 1). Baseline separation of all 5 major chloroplast pigments can be obtained using

the system previously published (5), which employs a linear gradient system starting with 100% solvent A and 0% solvent B and reaching 50% solvent A and 50% solvent B at 20 minutes, and holding to the end of the run time (Figure 2).

The extraction procedure that is described effectively removed all pigments from the fruit tissue without forming emulsions in the séparatory funnel or blockage of the 0.50^1 filter. Samples weighing

50 to 60 grams resulted in the quickest processing time, mainly due to rapid filtration; filtration took considerably longer with larger samples.

HPLC showed that 5 pigments were present in green tomato fruit: neoxanthin, violaxanthin, lutein, chlorophyll a and b, and/S-carotene

(Figures 1 and 2). Researchers interested in the baseline separation of all pigments in fruits can extract the pigments by the method in this report and use the previously described HPLC procedure (5).

Figure 2 shows that this method was very effective. However, if only chlorophyll a and b or yS-carotene are of interest, the isocratic system that is described gives baseline separation of these pigments in less than half the time of the gradient system (Figures 1 and 2). 21

8 10 12 14 16 18 20 22 TIME(MIN)

Figure 1. HPLC chromatogram of green tomato fruit pigments showing baseline separation of chlorophyll a, chlorophyll b, and )S-carotene. An isocratic solvent system of 50% solvent A (80? methanol) and 50% solvent B (ethyl acetate) was used. Other HPLC parameters in text. A=neoxanthin, B=violaxanthin, C=lutein, D=chlorophyll b, E=chlorophyll a, and F=;8-carotene 22

E C B

18 20 22 24 26 28 30 32 34 36 38 40 42 44 46 48 TIME(MIN)

Figure 2. HPLC chromatogram of green tomato fruit pigments showing baseline separation of all 6 pigments. A linear gradient solvent system starting with 100% solvent A (80$ methanol) and 0$ solvent B (ethyl acetate) and reaching 50% solvent A and 50% solvent B at 20 minutes was used, and then held until the end of the run time (48 minutes). Other HPLC parameters in text. A=neoxanthin, B=violaxanthin, C= lutein, D=chlorophyll b, E=chlorophyll a, and F=yS- carotene 23

The isocratic system eluted neoxanthin, violaxanthin, and lutein as

partially separated and identifiable peaks. Chlorophyll a and b were

eluted 10 to 12 minutes after injection, and a run time of 25 minutes

gave complete elution of all pigments (Figure 1). This is a

significant decrease in the run time when compared with previously

published HPLC methods (1, 2, 3, 4, 8).

To show the applicability of this method, the chlorophyll a and b

content of developing 'Heinz 1350' tomato fruits was determined in 5

day increments starting 15 days past anthesis and continuing until 55

days (Table 1). Data in this table show that the total amount of

either chlorophyll a or b increases during the first 35 days, reaches

a maximum at 35 days, and then proceeds downward until no chlorophyll

a or b remains at the fully ripe stage (day 55).

The chlorophyll a and b contents of several fruits and vegetables

were determined using the extraction, sample preparation, and

quantitation procedures developed in this report (Table 2). It is

evident that this method can be used on a wide range of plant organs

and, therefore, is applicable to many fruits and vegetables. As

discussed earlier, the previously published procedure (3, 5) did work

for leaves but did not work for fruits and vegetables. Cabbage was

chosen as one of the horticultural crops to be tested since it is a

leafy material. Several trials to extract and quantitate the chlorophyll in cabbage leaves by the previously published report (3,

5) resulted in the same problems encountered with tomato fruits.

Thus, the previously published report may not be applicable to all 24

Table 1. Chlorophyll a and b contents of whole 'Heinz 1350' tomato fruits as a function of days past anthesis. Results derived using methods developed in this report®

Time Past Fruit Fresh Anthesis Weight Chi a Chi b (Days) (g) (mg/fruit) (mg/fruit)

15 394 9.7 2.5

20 650 12.8 3.4

25 1,032 17.1 4.6

30 1,231 20.3 5.6

35 1,593 22.8 6.5

40 1,489 19.6 5.8

45 1,792 14.5 4.4

50 1,737 8.0 2.2

55 1,630 W — mm

^Mean value of 3 replicates each containing 4 fruits. 25

Table 2. Chlorophyll a and b contents of various horticultural crops as determined using the methods developed in this report^

Chlorophyll a Chlorophyll b UR/g, Fresh Weight ttg/g Fresh Weight

Broccoli Florets 195.3 65.5

Cabbage 75.3 25.3

Cucumber 58.1 24.3

Green Beans 54.6 17.3

Green Pepper 37.3 14.6

Green/Red Pepper 20.1 9.0

'Thompson Seedless' Grape 5.0 1.6

Lime 4.9 2.8

Zucchini Squash 73.4 30.8

^Fifty to 60 gram samples were used, and samples were intact slices, quarters, halves, or entire fruits as governed by sample size. 26 types of leaves, but may be applicable only to those having thin layers of epicuticular wax on the outer surface of the leaf.

The procedure described in this report is an easy and rapid one for the extraction, sample preparation, and quantitation of plant pigments by HPLC and should benefit researchers whose projects involve plant pigments contained in fruit and vegetable tissues. 27

LITERATURE CITED

1. Burke, S. and S. Aronoff. 1979. Separation of radiochemically pure chlorophyll a and chlorophyll b. Chromatographia 12:808- 809.

2. Davies, D. and E. S. Holdsworth. 1980. The use of high pressure liquid chromatography for the identification and preparation of pigments concerned in photosynthesis. J. Liq. Chromatogr. 3:123-132.

3. Eskins, K. and H. Button. 1979. Sample preparation for high performance liquid chromatography of higher plant pigments. Anal. Chem. 51:1885-1886.

4. Eskins, K., C. R. Scholfield and H. J. Button. 1977. High performance liquid chromatography of plant pigments. J. Chromatogr. 135:217-220.

5. Eskins, K. and L. Harris. 1981. High performance liquid chromatography of etioplast pigments in red kidney bean leaves. Photochem. Photobiol. 33:131-133.

6. Lyons, J. M. and H. K. Pratt. 1964. Effect of stage of maturity and ethylene treatment on respiration and ripening of tomato fruits. Proc. Amer. Soc. Hortic. Sci. 84:491-500.

7. Schwartz, S. J. and J. H. von Elbe. 1982. High performance liquid chromatography of plant pigments—a review. J. Liq. Chromatogr. 5 (Suppl. 1):43-73.

8. Schwartz, S. J., S. L. Woo and J. H. von Elbe. 1981. High performance liquid chromatography of chlorophylls and their derivatives in fresh and processed spinach. J. Agric. Food Chem. 29:533-535.

9. Strain, H. H. and W. A. Svec. 1966. Extraction, separation, estimation and isolation of the chlorophylls. Pages 21-66 in L. P. Vernon and G. R. Seely, eds. The Chlorophylls. Academic Press, New York. 28

SECTION II. PROPERTIES OF PORPHOBILINOGEN DEAMINASE

ISOLATED FROM 'HEINZ 1350' TOMATO FRUIT 29

ABSTRACT

Porphobilinogen deaminase was isolated from 'Heinz 1350' tomato

fruit and several properties were determined. Tomato deaminase shows

no activity below pH 6.0 and has optimum activity at pH 8.0 in tris

buffer. The enzyme was affected greatly by temperature and had

maximum activity at 57C. The Km for the deaminase was 17.5/iM,

indicating a very high affinity for porphobilinogen. HgClg and £-

chloromercuribenzoate inhibited the enzyme almost totally at a concentration of 70 /zM, suggesting that tomato fruit deaminase has

thiol groups at its active site. ^-aminolevulinic acid slightly increased deaminase activity. The reducing agents dithiothreitol and

2-mercaptoethanol did not prevent phenolic browning of the tomato fruit extracts, and 2-mercaptoethanol completely inhibited the enzyme.

The enzyme was stable for one day after extraction when stored at 4C in the dark. 30

INTRODUCTION

Porphobilinogen deaminase is one of the enzymes of the

chlorophyll and heme biosynthetic pathways. It acts in conjunction with a second enzyme, uroporphyrinogen III cosynthase, and converts four molecules of porphobilinogen into one molecule of uroporphyrinogen III (2, 5, 13, 17, 19, 21, 30, 31). When present by itself, the deaminase will consume porphobilinogen and produce uroporphyrinogen I, which is not an intermediate in chlorophyll biosynthesis. The deaminase enzyme has been characterized from a number of plant and animal sources, including: human erythrocytes (7,

12), Rhodopseudomonas spheroides (9, 10, 19, 20), leaf tissue (4, 16), wheat germ (5, 14, 15, 25), soybean callus (22) and Chlorella regularis (27). The enzyme, however, has not been isolated and characterized from fruit tissue. As a part of a project to determine the relationship of the deaminase enzyme to the development and ripening of 'Heinz 1350' tomato fruit (see section III), several properties were determined and are presented in this section. 31

MATERIALS AND.METHODS

Porphobilinogen and uroporphyrin I and III standards were

purchased from Porphyrin Products, Logan, UT.

Polyvinylpolypyrrolidone (PVPP), tris buffer, bovine serum albumin, N-

ethylmaleimide, and £-chloromercuribenzoate were obtained from Sigma

Chemical Company, St. Louis, MO. All other chemicals purchased from

commercial sources were reagent grade.

Analysis of uroporphyrin was accomplished in two ways; spectrophotometry and high performance liquid chromatography (HPLC).

After oxidation of the uroporphyrinogens to uroporphyrins, the absorbance at 405 nm was compared to a standard curve to obtain the amount of uroporphyrin formed. Absorption spectra also were determined and compared with that of the uroporphyrin standard. HPLC was used to quantitate the uroporphyrin formed and to separate uroporphyrin I and III isomers that may have formed. A Waters

Associates ALC-244 HPLC, a reverse phase //Bondapak-C^G analytical column (3.9 mm i.d. X 30 cm), and phosphate buffer were used in an isocratic system to separate the isomers. Preparation of the phosphate buffer has been described elsewhere (11, 29). A flow rate of 1 ml/min was used and uroporphyrin detection was by fluorescence.

The detector was a Schoeffel FS 970 fluorometer with the following settings; a range of 0.1/iA, sensitivity of 530 and a time constant of

6 seconds with high suppression of the signal. An excitation wavelength of 230 nm and an emission wavelength of 389 nm were used.

Uroporphyrin was identified by comparing its retention time to the 32

standards and by co-chroraatography with the standards.

'Heinz 1350' tomato plants were grown in an environmentally

controlled research greenhouse and standard cultural practices were

employed (8, 26). Plants were trained to a single stem and replicates

of fruit of the same physiological class were obtained by the

procedure of Lyons and Pratt (23). For all experiments, fruit that

were 15 to 20 days past anthesis were used, and three replicates of four tomatoes were used per experiment. Each experiment was repeated.

Immediately after harvesting, replicates of tomato fruit were

placed in ice and chilled for one hour. Each replicate was weighed, and a core of tissue (1 cm in diameter) was removed from each fruit with a stainless steel cork borer, weighed, and prepared for extraction.

The cores were diced and thoroughly ground in 4C-10 mM tris, pH

7.0. To this was added 1.25 grams of PVPP per gram fresh weight of tissue before grinding to prevent phenolic browning (18, 24). The homogenate was filtered through four layers of cheesecloth and centrifuged at 500xg for five minutes. The pellet was discarded and the supernatant was centrifuged at 100,000x£ for one hour. The pellet was discarded, and the supernatant was used as the source of enzyme.

The protein content of the extract was determined by the Bradford method (6).

For all experiments, a 1.47 mM porphobilinogen stock solution was prepared in SOmM tris, pH 8.0, immediately before each assay. Blanks omitting porphobilinogen and enzyme were run concurrently with each 33 experiment. Unless stated otherwise, the assay time was one hour in a

37C water bath with gentle shaking in the dark. The standard reaction mixture for all experiments, except when noted, consisted of: 1 ml of

30 mM tris, pH 8.15; 1 ml of extract; and 150//I of porphobilinogen stock solution in a 5 ml tube. The pH of this mixture was 8.0, and the final concentration of porphobilinogen in the reaction mixture was

102/uM and that of tris was 18.60 mM. Uroporphyrinogens biosynthesized were oxidized to uroporphyrins according to the method of Jordan and Shemin (20).

To determine the pH optimum of the deaminase enzyme from pH 5.0 to 9.0, the following mixture of buffers was prepared for the reaction mixture: 10 mM tris, 10 mM MES and 10 mM HEPES. This reaction mixture differed from the standard reaction mixture in that the 30 mM tris was replaced by the buffer mixture. The standard reaction mixture was used for the second pH study (pH range of 7.0 to 9.4).

For the time-course study (and all following experiments), the assay was started by the addition of porphobilinogen at time 0, and at

30 minute intervals the amount of uroporphyrin biosynthesized was determined. The experiment was conducted for three hours.

The assays for the temperature dependence study were conducted in

IOC increments, starting with 27C and ending at 77C. The incubation time was 30 minutes.

Km values were obtained by assaying the deaminase enzyme at increasing concentrations of porphobilinogen and measuring the amount of uroporphyrin formed per hour. A reciprocal plot of 1/ V vs 1/ S 31 was obtained and used to calculate the Km of the enzyme.

For the experiments in which the effect of various chemical reagents was determined, the chemicals were added to the standard incubation mixture just before to the start of the assay.

Concentrations of the reagents are given in the results section. 35

RESULTS

pH had a very pronounced effect on porphobilinogen deaminase

activity. The enzyme had no activity at pH 5.0 and low activity was

present at pH 6.0 (Figure 1). From pH 6.0 to 8.0, however, the

activity markedly increased to a maximum, before sharply falling at pH

9.0 (Figure 1). To more precisely determine the pH optimum of the

enzyme, another set of experiments was conducted. The enzyme was

assayed in small increments of pH between 7.0 and 9.5. The pH optimum

of porphobilinogen deaminase is approximately 8.0 (Figure 2).

The time-course study of the deaminase enzyme showed that it

biosynthesized uroporphyrin at a constant rate for at least three

hours after the assay began (Figure 3). The correlation coefficent

for the plot of uroporphyrin production vs time was 0.9942, indicating

a nearly perfect linear relationship.

Porphobilinogen deaminase activity was affected strongly by

changes in temperature (Figure 4). At the three highest assay

temperatures, chemical production of uroporphyrin occurred, and the

uroporphyrin values obtained at these three temperatures have been

corrected for chemical synthesis. The amount of uroporphyrin formed

chemically in the enzyme-free blanks was subtracted from that formed

enzymatically. Maximum enzymatic activity occurred at 57C and

abruptly diminished at 77C. The Q^Q was 4.94 for 27C to 37C, 3.69 for

37C to 47C, and 1.76 for 47C to 57C. The activity at 57C was 548% greater than that at 37C. A difference of only 5C (from 37C to 42C) doubled the uroporphyrin production rate (data not shown). Even at 36

B f

N M 0 L E U R 0 P 0 R P Ï R I N / H R

PH

Figure 1. pH profile of porphobilinogen deaminase from pH 5.0 to 9.0. Incubation mixture consisted of 1 ml of the 10 mM tris, 10 mM MES, 10 mM HEPES buffer mixture, porphobilinogen and 1 ml of enzyme extract in 2.15 ml total volume. Porphobilinogen concentration was 102/uM 37

Figure 2. pH profile of porphobilinogen deaminase from pH 7.0 to 9.5. Incubation conditions were as described for Figure 1, except that 30 mM tris was used instead of the buffer mixture 38

Figure 3. Time course of uroporphyrin formation by porphobilinogen deaminase. Incubation conditions described in text 39

TEMPERATURE (DEGREES CELSIUS)

Figure 4. Effect of assay temperature on porphobilinogen deaminase. Incubation system described in text 40

77C, the amount of uroporphyrin biosynthesized was considerably higher

than that produced at 27C.

The deaminase enzyme exhibited typical Michaelis-Menten kinetics

(Figure 5). A Km of 17.^uM and a of 5.0 nmoles uroporphyrin/hr

was calculated from a Lineweaver-Burke plot of the data.

The effect of various sulfhydryl reagents and ^-aminolevulinic

acid on deaminase activity is shown in Table 1. HgClg was the most

Table 1. Effect of chemical reagents on porphobilinogen deaminase activity

/t Change in deaminase activity from the control^

Concentration of reagent in assay mixture (^M)^

17.5 35 70

N-ethylmaleimide 0.0 0.0 -2.0 p-chloromercuribenzoate -30.0 -87.4 -94.0

HgClg -96.0 -96.4 -97.0

^-aminolevulinic acid +4.5 +6.9 +7.7

2-mercaptoethanol° -100 -100

^Assay conditions described in text.

^Activity measured as nmole uroporphyrin formed per hour.

°2-mercaptoethanol concentationss were 40 and 80 /uM. 41

5

4

3

2

1

0 0 40 80 120 160

PORPHOBILINOGEN (pM)

Figure 5. Effect of porphobilinogen concentration on uroporphyrin biosynthesis. Porphobilinogen at the following concentrations was used in the assays: 6.37 fXM; 12.75/iM; 25.5/xM; 51 /nM; 102/liM; and 204 ^iM. Other incubation conditions described in the text 42

effective inhibitor of the enzyme of the compounds tested, especially

at the two lower amounts that were added (Table 1). At a

concentration of 70 ixM, £-chloromercuribenzoate was as effective in

inhibiting the enzyme as was HgClg. N-ethylmaleimide had no effect on

enzyme activity at concentrations of 17.5 and 35/xM, and only a slight

inhibitory effect at 70/xM. ^-aminolevulinic acid increased

uroporphyrin biosynthesis to a small extent, although the difference

in effect between 35 and 70 was smaller than that between 17.5 and

35 )LtM. Curiously, 2-mercaptoethanol completely inhibited the enzyme

at Ho and 80/uM (Table 1).

The stability of porphobilinogen deaminase was determined after

24 hour intervals of storage at 4C in the dark in lOmM tris buffer, pH

7.0 (Figure 5). After 24 hours of storage, the enzyme still retained

98.7? of its original activity. However, at 48 hours, only 65.7$ of the original activity was present, and it fell to only 27% of its original activity 8 days after the enzyme was isolated. 43

98

88

70

60

58

48

38

28

10

S

DAYS AFTER EXTRACTION

Figure 6. Stability of porphobilinogen deaminase stored at 4C in the dark. The enzyme was assayed immediately after extraction and at 24 hour intervals thereafter. Activity measured in nmole uroporphyrin per hour. Incubation conditons described in text 44

DISCUSSION

The results of this research show that porphobilinogen deaminase

from 'Heinz 1350' tomato fruit has a sharp pH optimum of 8.0 in 18.60

mM tris buffer (Figure 2). This finding is in close agreement with

Davies and Neuberger (10), who found that the deaminase enzyme from R.

spheroides exhibited maximum activity at pH 7.8 to 8.0. In addition,

a pH optimum of 8.2 was determined for deaminase from wheat germ (14).

A somewhat lower pH optimum of 1.2 in tris was determined for the

enzyme from soybean callus (22). This would result in significantly

lower activity for the tomato deaminase enzyme.

Porphobilinogen deaminase biosynthesized uroporphyrin with no lag

time at a constant rate for three hours (Figure 3). This finding is in agreement with time-course studies for deaminase isolated from human erythrocytes in which a constant rate of biosynthesis occurred for at least two hours (12). Frydman and Frydman showed that the pepper leaf deaminase biosynthesized uroporphyrin at a linear rate for

30 minutes with no lag time, followed by a faster, linear rate, which was not observed in this study (16). Llambias and Batlle showed that soybean callus deaminase had a lag time of three to four hours before uroporphyrin production started (22). Because porphobilinogen consumption showed no such lag, it was concluded that an intermediate was accumulating during this time and that it was a substrate partner of porphobilinogen. However, because the tomato deaminase biosynthesized uroporphyrin at a constant rate without a lag time, it can be concluded that no prolonged accumulation of intermediates 45

occurred.

The tomato deaminase had maximum activity at 57C, and activity

was measured when assayed at 77C, showing that it is a relatively heat-stable enzyme (Figure 4). These results are similar to those found for wheat germ deaminase, except that maximum activity occurred at 67C, although the difference in activity between 57C and 67C was small (14). The fact that no uroporphyrin III was biosynthesized at

57C or higher is in agreement with other studies in which uroporphyrin

III cosynthase became inactivated around 55C (5, 28). The Q^Q for most enzymatic reactions is between 2.0 and 3.0, but the deaminase showed much larger increases in activity from 27C to 37C and from 37C to 47C. Temperature increases in this range indicate that the tomato

deaminase reaction has a high activation energy. The Q^Q for 47C to

57C was much smaller than the previous two Q^Q values, and thermal denaturation of the enzyme probably was becoming significant.

Temperatures higher than 57C resulted in more denaturation of the

enzyme, which resulted in decreased activity. While the Q^Q for 37C to 47C was still large, it was smaller than the increase over the previous IOC increase, which may indicate a small amount of thermal denaturation of the enzyme. It was shown in this study that a 5C increase in assay temperature can double the rate of uroporphyrin production, and, therefore, care must be taken to assay the enzyme at exactly the same temperature for studies that require repeated assays at a given temperature.

The Km of 17.for the tomato deaminase enzyme is very small. 46 and, thus, the deaminase enzyme has a very high affinity for porphobilinogen (Figure 5). These results are similar to Km values reported for pepper and poinsettia leaf deaminase (16) and R. spheroides (20). The enzyme for all other experiments than the Km determination was assayed with porphobilinogen at a concentration of

102 ^M, which was sufficient to keep the enzyme near maximum velocity in biosynthesizing uroporphyrin from porphobilinogen.

HgClg and £-chloromercuribenzoate effectively inhibited the deaminase enzyme (Table 1). As these chemicals are sulfhydryl-group blocking agents, their inhibitory effects show that the deaminase enzyme from tomato fruit has thiol groups at its active site. Again, this finding is similar to the results of studies on deaminase from other sources (4, 20, 25). N-ethylmaleimide at a concentration of

10 only slightly inhibited the enzyme (Table 1). However, it may be that it would have been more inhibitory at a higher concentration.

Deaminase from ^ spheroides was inhibited k5% by N-ethylmaleimide at a concentration of 1 mM (20); a considerably higher concentration than was used in this study. Evidently, the thiol groups of the enzyme are the most sensitive to HgClg and the least sensitive to N- ethylmaleimide at the concentrations tested.

For most sulfhydryl enzymes, reducing agents such as 2- mercaptoethanol would serve to promote activity and to protect such enzymes from the effects of p-chloromercuribenzoate by keeping the thiol groups at the active sites reduced. However, not only did 2- mercaptoethanol fail to reverse the effects of the inhibitors on 47

tomato deaminase, it completely inhibited the enzyme (Table 1).

Dithiothreitol and 2-mercaptoethanol have been used in extracting

media to prevent phenolic browning because of their possible

inhibitory effect on o-diphenoloxidase (1). Tomato fruit tissue has a

high phenolic content when the fruit is very young in development to

approximately half way in development to ripeness (18, 24). These two

reducing agents did not prevent phenolic browning of the extracts from

young fruit, and only when PVPP was added was browning prevented.

Thus, these reducing agents seem to be of no value both for extracting

deaminase from tomato fruit and also in enhancing deaminase activity.

Bianchi and Stegwe found that 2-mercaptoethanol also inhibited spinach leaf deaminase, but at a concentration four times greater than was used in this study (3). Frydman and Frydman showed that 2- mercaptoethanol did not improve deaminase activity when it was added to the extracting buffer (16). However, in another study, both dithiothreitol and 2-mercaptoethanol reversed the inhibition by £- chloromercuribenzoate and N-ethylmaleimide (14). It seems that deaminase from various sources is affected differently by different reducing agents. Why tomato fruit deaminase is inhibited by 2- mercaptoethanol is not known.

^-aminolevulinic acid had a small promotive effect on tomato deaminase activity (Table 1). Similar results were reported when â

-aminolevulinic acid stimulated uroporphyrin production in extracts from soybean callus (22). This suggests that ^-aminolevulinic acid dehydratase was present in the extract and that it biosynthesized 48

porphobilinogen from ^-aminolevulinic acid. The concentration of

porphobilinogen then would be increased in the assay mixture,

enhancing the rate of uroporphyrin production. As noted earlier, the

amount of porphobilinogen already in the assay mixture was enough to

keep the enzyme near maximum velocity, so that any additional

substrate would only have a small effect, which was the case in this

experiment.

Tomato fruit deaminase is stable for one day after extraction

when stored in the dark at 4C, and, after this, significant losses in

activity occur (Figure 6). This result is in contrast to findings by

Frydman and Frydman who found that the enzyme was stable at 4C for

more than a week (14). The enzyme, however, had been purified by

DEAE-cellulose chromatography, which probably removed agents from the

extract that could have resulted in less stability. ^ spheroides

deaminase still had 50$ of its original activity three weeks after

extraction when stored in 100 mM tris at pH 8.5 (20). But, this

mixture was supplemented with 2-mercaptoethanol, which would have inhibited the tomato deaminase. The loss of activity probably is

because of oxidation of the thiol groups in the active site.

Normally, this could be prevented by the addition of a reducing agent like 2-mercaptoethanol. However, because it inactivates tomato fruit deaminase, it would not be a useful additive to prolong tomato deaminase storage life.

Several properties of tomato fruit deaminase have been determined in this study. The enzyme from tomato fruit seems to be similar in 49 properties to deaminase enzymes isolated from leaf, algae and animal sources. 50

LITERATURE CITED

1. Anderson, J. W. 1968. Extraction of enzyme and subcellular organelles from plant tissues. Phytocheraistry 7:1973-1988.

2. Battersby, A. R. and E. McDonald. 1976. Biosynthesis of porphyrins and corrins. Phil. Trans. R. Soc. London, B, 273:161-180.

3. Bianchi, A. and D. Stegwee. 1979. Porphobilinogenase in greening leaves of Phaseolus vulgaris. Z. Pflanzenphysiol. 91:377-383.

4. Bogorad, L. 1958. The enzymatic synthesis of porphyrins from porphobilinogen. I. Uroporphyrin I. J. Biol. Chem. 233:501- 509.

5. Bogorad, L. 1958. The enzymatic synthesis of porphyrins from porphobilinogen. II. Uroporphyrin III. J. Biol. Chem. 233:510-515.

6. Bradford, M. M. 1976. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding. Anal. Biochem. 72:248-254.

7. Brodie, M. J., M. R. Moore, G. G. Thompson, B. C. Campbell and A. Goldberg. 1977. Is porphobilinogen deaminase activity a secondary control mechanism in heme biosynthesis in humans? Biochem. Soc. Trans. 5:1466-1468.

8. Brooks, W. M. 1973. Growing greenhouse tomatoes in Ohio. The Ohio State University Cooperative Extension Service Bulletin SB-19. 56 pages.

9. Burton, G., P. E. Eagerness, S. Hosozawa, P. M. Jordan and A. I. Scott. 1979. C-13 N. M. R. evidence for a new intermediate, pre-uroporphyrinogen, in the enzymatic transformation of porphobilinogen into uroporphyrinogens I and III. J. Chem. Soc. Chem. Commun. 5:202-204.

10. Davies, R. C. and A. Neuberger. 1973. Polypyrroles formed from porphobilinogen and amines by uroporphyrinogen synthetase of Rhodopseudomonas spheroides. Biochem. J. 133:471-492.

11. Englert, E., Jr., A. W. Wayne, E. E. Wales, Jr. and R. C. Straight. 1979. A rapid, new and direct method for isolation and measurement of porphyrins in biological samples by high performance liquid chromatography. J. High Resolution Chromatogr. Chromatogr. Commun. 2:570-574. 51

12. Ford, R. E., C. N. Ou and R. D. Ellefson. 1980. Assay for erythrocyte uroporphyrinogen I synthase activity, with porphobilinogen as substrate. Clin. Chem. 26:1182-1185.

13- Frydman, B., R. B. Frydman, A. Valasinas, S. Levy and G. Feinstein. 1975. The mechanism of uroporphyrinogen biosynthesis. The Biological Role of Porphyrins and Related Structures. Annals of the New York Academy of Sciences, New York. 244:371-395.

14. Frydman, R. B. and B. Frydman. 1970. Purification and properties of porphobilinogen deaminase from wheat germ. Arch. Biochem. Biophys. 136:193-202.

15. Frydman, R. B. and B. Frydman. 1973. The enzymatic inactivation of porphobilinogen deaminase. Biochim. Biophys. Acta 293:506- 513.

16. Frydman, R. B. and B. Frydman. 1979. Disappearance of porphobilinogen activity in leaves before the onset of senescence. Plant Physiol. 63:1154-1157.

17. Frydman, R. B., B. Frydman, A. Valasinas and E. S. Levy. 1977. Biosynthesis of uroporphyrinogen III. Interaction among 2- aminoethyltripyrranes and the enzymatic system. Pages 165-178 iji F. R. Longo, ed. Porphyrin Chemistry Advances. Ann Arbor Science Publishers, Inc., Ann Arbor, Michigan.

18. Hobson, G. E. 1967. Phenolase activity in tomato fruit in relation to growth and to various ripening disorders. J. Soi. Food Agric. 18:523-526.

19. Jordan, P. M., G. Burton, H. Mordlov, M. M. Schneider, L. Pryde and A. I. Scott. 1979. Pre-uroporphyrinogen: a substrate for uroporphyrinogen III cosynthetase. J. Chem. Soc. Chem. Commun. 5:204-205.

20. Jordan, P. M. and D. Shemin. 1973. Purification and properties of uroporphyrinogen I synthetase from Rhodopseudomonas spheroides. J. Biol. Chem. 248:1019-1024.

21. Llambias, E. B. C. and A. M. del C. Batlle. 1970. Uroporphyrinogen III cosynthase. Evidence for the existence of a polypyrrole substrate in soybean callus tissue. FEBS Letters 6:285-288.

22. Llambias, E. B. C. and A. M. del C. Batlle. 1971. Studies on the porphobilinogen deaminase-uroporphyrinogen cosynthase system of cultured soya-bean cells. Biochem. J. 121:327-340. 52

23. Lyons, J. M. and H. K. Pratt. 1964. Effect of stage of maturity and ethylene treatment on respiration and ripening of tomato fruits. Proc. Amer. Soc. Hortic. Soi. 84:491-500.

24. Mayer, A. M. and E. Harel. 1981. Polyphenol oxidases in fruits- changes during ripening. Pages I6I-I8O J. Friend and M. J. Rhodes, eds. Recent Advances in the Biochemistry of Fruits and Vegetables. Academic Press, London.

25. Russell, C. S. and P. Rockwell. 1980. The effects of sulfhydryl reagents on the activity of wheat germ uroporphyrinogen I synthase. FEES Letters 116:199-202.

26. Sheldrake, R. 1981. Tomato production in bags. Benchmarks 1:4- 6.

27. Shioi, Y., M. Nagamine, M. Kuroki and T. Sasa. 1980. Purification by affinity chromatography and properties of uroporphyrinogen I synthetase from Chlorella regularis. Biochim. Biophys. Acta 616:300-309.

28. Stevens, E. and B. Frydman. 1968. Isolation and properties of wheat germ uroporphyrinogen III cosynthase. Biochim. Biophys. Acta 151:429-437.

29. Wayne, A. W., R. C. Straight, E. E. Wales, and E. Englert, Jr. 1979. Isomers of uroporphyrin free acids separated by HPLC. J. High Resolution Chromatogr. Chromatogr. Commun. 2:621-622.

30. Yuan, M. and C. S. Russell. 1974. Porphobilinogen derivatives as substrates for porphobilinogenase. FEBS Letters 46:34-38.

31. Zelitch, I. 1971. Photosynthesis, Photorespiration and Plant Productivity. Academic Press, New York. 347 pages. 53

SECTION III. PORPHOBILINOGEN DEAMINASE ACTIVITY

IN DEVELOPING 'HEINZ 1350' TOMATO FRUIT AND ITS RELATIONSHIP

TO TOMATO FRUIT CHLOROPHYLL CONTENT 54

ABSTRACT

The chlorophyll a and b contents of 'Heinz 1350' tomato fruit

were determined in five day increments for fruit that were 15 days

past anthesis to full ripeness (55 days past anthesis). Results

showed that both the chlorophyll a and b contents were low in 15 day fruit and increased to a maximum in fruit 35 days past anthesis. The chlorophyll a and b content then fell to zero in fruit 55 days past anthesis. Similarly, prophobilinogen deaminase activity from extracts of fruit of the same stages of development showed the same changes with respect to fruit development. The stage of fruit development at which the chlorophyll content and deaminase activity were at their highest also corresponded to rapid fruit growth and maximum protein content. The chlorophyll a;b ratio did not change greatly during tomato fruit growth, and, thus, chlorophyll a and b were biosynthesized and degraded at similar rates. 55

INTRODUCTION

Chlorophyll biosynthesis is regulated by light (1, 6). One

light-regulated step is the biosynthesis of ^-aminolevulinic acid

(ALA). Etiolated plants are unable to form ALA, but, when transferred

to light, production of ALA commences. ALA is a precursor of

protochlorophyllide, and only trace amounts of protochlorophyllide

accumulate in the dark. When etiolated tissue is transferred to

light, protochlorophyllide production increases dramatically. If

etiolated leaves are fed ALA in the dark, protochlorophyllide

accumulates, showing that the enzymes are present and operative for

the biosynthesis of protochlorophyllide from ALA regardless of the effect of light. Thus, ALA formation is the limiting factor for

protochlorophyllide biosynthesis. The other light-regulated step is the photoconversion of protochlorophyllide to chlorophyllide. Once etiolated tissue is exposed to light, the accumulated protochlorophyllide rapidly is converted to chlorophyllide, and the tissue will continue to convert the protochlorophyllide to chlorophyllide for a period of time when the tissue is returned to the dark.

The light-controlled regulation of chlorophyll biosynthesis is well-documented. A study by Frydman and Frydman now suggests that there may be another control point of chlorophyll biosynthesis in relation to the age of the plant tissue (4). Porphobilinogen deaminase is one of the enzymes of the chlorophyll biosynthetic pathway that functions, with uroporphyrinogen III cosynthase, in 56

transforming four molecules of porphobilinogen into one molecule of

uroporphyrinogen III. It was found that the deaminase enzyme had high

activity in extracts from young leaves of pepper and poinsettia, low

activity in extracts from mature leaves, and almost no activity in

extracts from senescent leaves. Uroporphyrin III cosynthase, however,

showed no changes in activity in extracts from the three ages of the

leaves. Frydman and Frydman concluded that the deaminase enzyme may

control chlorophyll biosynthesis, reaching maximum activity when the

chlorophyll content of the leaf is high and then rapidly diminishing

in activity with the resultant loss of chlorophyll from the leaf

tissue. In addition, Brodie et al. believe that the deaminase also

may play a role in regulating heme biosynthesis (2).

The purpose of this project was to explore in more depth the

possibility of the regulation of chlorophyll biosynthesis by

porphobilinogen deaminase in 'Heinz 1350' tomato fruit. Considerable

research has been conducted on the ripening process of tomatoes and

the subsequent changes in pigment content. However, little, if any,

research has been done on the relationship of chlorophyll biosynthesis

to the entire development of the tomato fruit. Therefore, the

objectives of this research were to: 1) measure the chlorophyll a and

b content of 'Heinz 1350' tomato fruit ranging in age from 15 days

past anthesis to 55 days past anthesis (fully ripe); 2) measure the activity of porphobilinogen deaminase in extracts of 'Heinz 1350'

tomato fruit that were at the same stages of development as those used in the first objective; and 3) determine whether or not there was a 57 correlation between chlorophyll content and porphobilinogen deaminase activity in the tomato fruit. 58

MATERIALS AND METHODS

Procedures for the extraction of chlorophyll from tomato fruit

tissue and quantitation of chlorophyll by high performance liquid

chromatography (HPLC) are covered in the Materials and Methods section

of Section I of this dissertation. The Materials and Methods section

of Section II of this dissertation presents the cultural practices

that were used in growing the 'Heinz 1350' tomato plants. The

procedures for extracting the deaminase enzyme from tomato fruit

tissue, assaying the enzyme, and analysis of uroporphyrin by

spectrophotometry and HPLC also are covered in the Materials and

Methods section of Section II of this dissertation.

The standard incubation system for all assays in this study

consisted of: 1 ml of 30mM tris, pH 8.0, porphobilinogen and 1 ml of

extract in a total volume of 2.15 ml. The concentration of

porphobilinogen was 102/iM. 1.25 grams of PVPP per gram fresh weight

of tomato fruit tissue were added to the enzyme extracting medium

before grinding for fruits that were 15 days past anthesis in

development to full ripeness. However, it was necessary to use 5.0 grams of PVPP per gram fresh weight of 10 day old fruit tissue because of its higher phenolic content (7, 10).

To determine whether or not inhibitors were present in extracts from ripe fruit, extracts of equal volume from 15 day fruit and ripe fruit were mixed and assayed by the standard method. 59

RESULTS

Changes during in uroporphyrin production (nmoles

uroporphyrin/hr) and in chlorophyll a and b contents during fruit

development are presented on a per gram fresh weight basis (Figure 1).

Uroporphyrin biosynthesis was relatively low in 10 day fruit and significantly increased to a maximum in 15 day fruit (Figure 1A).

Fruit that was 20 days past anthesis had significantly lower uroporphyrin biosynthesis. From 25 to 50 days past anthesis uroporphyrin formation decreased overall, but not significantly from that in 20 day fruit. However, the subsequent decrease in uroporphyrin production in 55 day fruit was significant, and it was only 4$ of the peak in 15 day fruit. The uroporphyrin that was biosynthesized in 60 day fruit was not statistically different from that in 55 day fruit. Thus, porphobilinogen deaminase activity was low in 10 day fruit, peaked in 15 day fruit, gradually fell in 20 to

50 day fruit, and abrubtly diminished in 55 day fruit (Figure 1A).

The chlorophyll a and b contents were at their highest level in

15 day fruit and then decreased to zero in fruits that were 55 days past anthesis (Figure IB). Statistically, the decrease in both chlorophyll a and b contents was significant for each 5 day increment from 15 to 25 days past anthesis. However, from 25 to 40 days past anthesis, the decrease was not significant. Significant decreases in the chlorophyll a and b content occurred for each five day increment for fruit beyond 40 days in development. Generally, the decrease both in deaminase activity and chlorophyll content was similar in pattern 60

=0.58

Ï C 20 H CHL a L =2,40 D S 15 P ? 10 - XCHLb L L 0.05 / G F H f 1- f f f 1- f f f 10 15 20 25 30 35 40 45 50 55 - 60 65

FRUIT AGE (DAYS PAST ANTHESIS)

Figure 1. Effect of fruit age on porphobilinogen deaminase activity and chlorophyll content. (A) Porphobilinogen deaminase activity measured as nmole uroporphyrin/gram fresh weight/ hr. LSD (0.05)=0.58 nmole. (B) Chlorophyll a and b content measured as yug/g fresh weight. LSD (0.05) chlorophyll a=2.40 ^ig/g fresh weight, chlorophyll b=0.73 /Lig/g fresh weight. • B=chlorophyll a, A • A= chlorophyll b. Vertical bars represent the LSD, 0.05 level 61

from 15 to 55 days past anthesis.

The growth pattern of 'Heinz 1350' tomato fruit was determined

(Figure 2). The maximum growth rate occurred from 25 to 35 days past

anthesis, with lower rates before and after this time period. Growth

follows a typical sigmoid growth curve.

When the data for deaminase activity and chlorophyll content are

presented on a whole-fruit basis, both parameters show similar curves

(Figure 3A and 3B). The chlorophyll content and deaminase activity

significantly increased to a maximum in 35 day fruit. The chlorophyll

content then decreased to zero in 55 day fruit, while deaminase

activity diminished to a very low level in 60 day fruit. In addition,

the protein content of 'Heinz 1350' tomato fruit is presented on a

whole-fruit basis (Figure 4). As with the chlorophyll content and

deaminase activity curves, the protein content curve peaks in 35 day

fruit, and decreases rapidly on either side of the peak.

The chlorophyll a:b ratio of 'Heinz 1350' tomato fruit tissue was

calculated as a function of fruit development (Table 1). The ratio

was highest (3.89) in 15 day fruit and then decreased slightly to a

low of 3.28 in 45 day fruit before increasing in 50 day fruit.

Mixing extracts from 15 day fruit and ripe fruit (60 days past

anthesis) resulted in deaminase activity at a level that was additive

in relation to the activity of the two extracts (Table 2). Similar

results were obtained when extracts from 15 day fruits were mixed with

those from tomatoes at the breaker stage and pink stage of ripening

(data not shown). 62

140

120 = 21.87 LSD,0.05

100

80

60

40

20

0

FRUIT AGE (DAYS PAST ANTHESIS)

Figure 2. Growth pattern of 'Heinz 1350' tomato fruit. LSD (0.05)= 21.87 g. Vertical bar represents the LSD 63

15^0.05 = 8 70

CHL a T '•SDo.05=^®^

- X CHL b LS00.05=*:9

FRUIT AGE (DAYS PAST ANTHESISI

Figure 3. Effect of fruit age on porphobilinogen deaminase activity and chlorophyll content on a whole fruit basis. (A) Porphobilinogen deaminase activity measured as nmole uroporphyrin/fruit/hr. LSD (0.05)=8.70 nmole/fruit/hr. (B) Chlorophyll a and b content measured as mg chlorophyll/ fruit. LSD (0.05) chlorophyll a=1.07 mg/fruit, chlorophyll bzO.29 mg/fruit. —a =chlorophyll a, A A= chlorophyll b. Vertical bars represent the LSD, 0.05 level 6iJ

110

90 LSDoo5 = 1390

80

70

60

50

40

30

20

10

0

FRUIT AGE (DAYS PAST ANTHESIS)

Figure 4. Effect of fruit age on protein content. Protein content measured as mg protein/fruit. LSD (0.05)=13»90 mg. Vertical bar represents the LSD, 0.05 level 65

Table 1. Chlorophyll a:b ratio of 'Heinz 1350' tomato fruit tissue as a function of days past anthesis^

Days past anthesis Chlorophyll a:b ratio

15 3.89

20 3.75

25 3.76

30 3.55

35 3.48

40 3.44

45 3.28

50 3.64

55

^Each value is the mean of three replicates with four fruits per replicate.

Table 2. Effect of mixing extracts from 15 day past anthesis and ripe (60 days past anthesis) tomatoes on porphobilinogen deaminase activity^

Extract source Uroporphyrin formed (nmole/hr)^

Ripe tomatoes 0.15

15 day tomatoes 4.59

15 day + ripe tomatoes 2.30

®Each value is the mean of three replicates with four fruits per replicate.

^Assay conditions described in text. 66

Analysis of uroporphyrin by HPLC showed that predominantly the

type I isomer was biosynthesized in extracts from fruit at all stages

of development (Table 3). Only 10 to 20% of the total uroporphyrin

was type III, and no consistent trend in the ratio of the two isomers

was found as a function of fruit development.

Table 3- Uroporphyrin I and III isomer production by developing 'Heinz 1350' tomato fruit®

Fruit age (days past anthesis) $ Uroporphyrin I % Uroporphyrin III

10 87 13

15 90 10

20 82 18

25 86 14

30 85 15

35 90 10

40 88 12

45 81 19

50 87 13

55 80 20

60 86 14

^Each replicate represents the mean of 3 replicates, with four fruits per replicate. 67

DISCUSSION

On a per gram fresh weight basis, the decrease in chlorophyll

content over time would suggest that the rate of chlorophyll

biosynthesis is decreasing in the tomato fruit throughout its

development (Figure IB). The parallel decrease in porphobilinogen

deaminase activity could be interpreted as a cause for the decreased

chlorophyll content (Figure 1A). However, during the first 35 days of

development, 'Heinz 1350' tomato fruit were growing at a rapid rate

with large increases in volume, most of which is water (Figure 2).

Thus, a dilution effect probably was responsible for the observed

decrease in both chlorophyll content and deaminase activity of fruit

between 15 and 35 days past anthesis, and not a decrease in

chlorophyll biosynthesis per se. This observation is strengthened by

the fact that the decrease in chlorophyll content and deaminase

activity was proportionally smaller than the increase in fruit size

(Figures 1A and IB, Figure 2).

When the data are presented on a whole-fruit basis, the results are very different. The data show that chlorophyll biosynthesis is producing chlorophyll a and b in fruits up to 35 days past anthesis

(Figure 3B). During this same time, porphobilinogen deaminase activity increased in a similar fashion (Figure 3A). The decrease in chlorophyll content in 40 to 55 day fruit most likely is because the rate of chlorophyll degradation is becoming larger than the rate of chlorophyll biosynthesis. Visually, 'Heinz 1350' tomato fruit at this stage of development become a much lighter shade of green, indicating 68

chlorophyll degradation.

Because porphobilinogen deaminase still is very active in

extracts of fruit 40 to 50 days past anthesis (Figure 3A), this

supports the hypothesis that there still is chlorophyll biosynthesis

in fruit of this age. The disappearance of chlorophyll in fruit 55

days past anthesis coincided with an abrupt drop of deaminase activity

(Figure 3A and B). Thus, changes in deaminase activity correlated with changes in chlorophyll content, implying that the level of deaminase activity indeed may be a controlling point in chlorophyll biosynthesis in relation to tomato fruit development. This hypothesis is strengthened by the findings of Frydman and Frydman, who found that pepper and poinsettia leaf deaminase had the highest activity in young leaves and no activity in senescent leaves (4). This high activity corresponded to a maximum leaf chlorophyll content, after which the deaminase activity decreased. Because uroporphyrin III cosynthase activity was constant with respect to leaf age, Frydman and Frydman concluded that the deaminase regulated chlorophyll biosynthesis in leaves. Furthermore, Llambias and Batlle showed that the rate of chlorophyll biosynthesis was correlated with uroporphyin production when dark-grown soybean callus was transferred to the light (9).

A low level of activity still was present in extracts from ripe fruit, which contrasts to Frydman and Frydman's (4) finding that no deaminase activity was detected in extracts from senescent pepper and poinsettia leaves (Figure 3A). It definitely can be said that ripe tomato fruit tissue has deaminase enzyme that is capable of limited 69

uroporphyrin production, and that the deaminase may be active in ripe

fruit tissue, but at such a low level that it could be severely

limiting chlorophyll production. Furthermore, the little amount of

uroporphyrin III which is produced could be degraded immediately.

Possible reasons for the decrease in deaminase activity in older

fruit could be the presence of an inhibitor(s) or a decrease in the

actual amount of enzyme present within the fruit. The results of

mixing experiments using extracts from 15 day fruit and ripening/ripe

fruit cast doubt on the inhibitor hypothesis. Had there been

inhibitors present in extracts from ripe fruit, the amount of

uroporphyrin formed in the mixed extracts would have been less than additive, which was not the case (Table 2). Instead, the amount of

uroporphyrin was very close to the mean of the amounts biosynthesized from 15 day and ripe fruit extracts. Thus, inhibitors of chlorophyll

biosynthesis are not present, and, probably, the decreased chlorophyll content in ripening/ripe fruit is due to a lack of chlorophyll biosynthesis. This, in turn, probably is caused by decreased porphobilinogen deaminase activity or an accelerated rate of chlorophyll degradation.

To further clarify this question, protein contents of 'Heinz

1350' tomato fruit were determined. It was found that the changes in protein content closely parallel both the changes in chlorophyll content and deaminase activity. The markedly strong decrease in protein content in fruits more than 35 days past anthesis resulted in very low protein contents in 55 and 60 day fruit. As the pattern of 70

the decrease in protein content is very similar to the activity of the

deaminase, one could conclude that the decrease in deaminase activity

was caused by a decrease in the amount of the enzyme present within

the fruit tissue. However, one must keep in mind that selective

protein breakdown in the fruit tissue occurring late in the

development of the tomato may not be affecting the deaminase enzyme

and that the decrease in activity actually may be caused by another

factor.

It is interesting to note that the protein and chlorophyll

content along with deaminase activity were at a maximum in fruits that

were 35 days past anthesis. As mentioned earlier, this corresponded

to a period of maximum growth of the fruit. During this time of rapid

growth, the metabolic activities are necessarily high to sustain

growth. The maximum protein content could partly account for the many

metabolic enzymes that are required during this time of fast growth.

The fact that small amounts of uroporphyrin III were in the

extracts shows that uroporphyrin III cosynthase also was present with

the deaminase enzyme (Table 3). Unless the cosynthase is highly

purified and present in excess with respect to the deaminase,

uroporphyrin I is the major product of the reaction (5). Furthermore, cosynthase activity can decrease vitro. even when present in excess, and this leads to obvious difficulties when attempting to accurately assay the enzyme (3).

The chlorophyll a;b ratio showed no large changes in the fruit as they matured. A slight decrease in the ratio over time possibly 71

indicates that chlorophyll b was being produced at a slightly higher

rate than chlorophyll a early in fruit development and was being

degraded at a slightly slower rate in maturing fruit (Table 1).

From the results of this study, it can be concluded that the chlorophyll a and b content of 'Heinz 1350' tomato fruit peaked in 35 day fruit, and porphobilinogen deaminase isolated from the fruit showed a strong correlation in the changes of its activity to the changes in the chlorophyll content of the fruit. Increases in porphobilinogen deaminase activity were paralleled by increases in the chlorophyll a and b content. Similarly, the decrease in deaminase activity from fruits that were beyond 35 days in development was mirrored by the decrease in chlorophyll a and b contents of the fruit.

It is therefore hypothesized that porphobilinogen deaminase regulates chlorophyll biosynthesis in 'Heinz 1350' tomato fruit, and that its decrease in activity causes the rate of chlorophyll biosynthesis to become lower than that of chlorophyll degradation. This leads, ultimately, to a fruit which contains no chlorophyll as the ripe and senescent stages of tomato fruit development are approached. 72

LITERATURE CITED

1. Bogorad, L. 1965. The biosynthesis of chlorophylls. Pages 481- 510 in L. P. Vernon and G. R. Seely, Eds. The Chlorophylls. Academic Press, London.

2. Brodie, M. J., M. R. Moore, G. G. Thompson, B. C. Campbell and A. Goldberg. 1977. Is porphobilinogen deaminase activity a secondary control mechanism in heme biosynthesis in humans? Biochem. Soc. Trans. 5:1466-1468.

3. Burnham, B. F. and R. C. Bachmann. 1979. Enzymatic synthesis of porphyrins. Pages 234-257 in D. Dolphin, ed. The Porphyrins. Volume VI. Biochemistry. Part A. Academic Press, New York.

4. Frydman, R. B. and B. Frydman. 1979. Disappearance of porphobilinogen deaminase activity in leaves before the onset of senescence. Plant Physiol. 63:1154-1157.

5. Frydman, R. B., B. Frydman and A. Valasinas. 1979. Protoporphyrin: synthesis and biosynthesis of its metabolic intermediates. Pages 1-124 ^ D. Dolphin, ed. The Porphyrins. Volume VI. Biochemistry. Part A. Academic Press, New York.

6. Harel, E. 1978. Chlorophyll biosynthesis and its control. Prog. Phytochem. 5:127-180.

7. Hobson, G. E. 1967. Phenolase activity in tomato fruit in relation to growth and to various ripening disorders. J. Sci. Food Agric. 18:523-526.

8. Hobson, G. E. and J. N. Davies. 1971. The tomato. Pages 437- 482 A. C. Hulme, ed. The Biochemistry of Fruits and Their Products. Volume 2. Academic Press, New York.

9. Llambias, E. B. C. and A. M. del C. Batlle. 1971. Studies on the porphobilinogen deaminase-uroporphyrinogen cosynthase system of cultured soya-bean cells. Biochem. J. 121:327-340.

10. Mayer, A. M. and E. Harel. 1981. Polyphenol oxidases in fruits- changes during ripening. Pages 161-180 iji J. Friend and M. J. Rhodes, eds. Recent Advances in the Biochemistry of Fruits and Vegetables. Academic Press, London. 73

GENERAL SUMMARY AND CONCLUSIONS

A procedure was developed for the extraction and quantitation of chloroplast pigments in horticultural commodities by HPLC. This method was very effective for several diverse fruits and vegetables and overcame difficulties encountered when a previously published procedure for leaves was attempted on fruit and vegetable tissues.

Chlorophyll a and b and d-carotene can be extracted rapidly and quantitated by HPLC in the isocratic mode. HPLC in the linear gradient mode will resolve completely all of the six major chloroplast pigments.

Using the isocratic HPLC procedure, the chlorophyll a and b content of developing 'Heinz 1350' tomato fruit were determined in five day increments from 15 days past anthesis to full ripeness (55 days past anthesis). The chlorophyll content increased in fruit up to

35 days past anthesis and then decreased to zero in ripe fruit.

Porphobilinogen deaminase activity also was measured in extracts from developing fruit, and changes in deaminase activity correlated to changes in the chlorophyll content. Thus, it seems that porphobilinogen deaminase activity may regulate chlorophyll biosynthesis in tomato fruit. Several properties of the deaminase enzyme were determined and results showed that it is similar to deaminase enzymes isolated from leaf and animal sources. 71

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D. Dolphin, ed. The Porphyrins. Volume VI. Biochemistry. Part A. Academic Press, New York.

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ACKNOWLEDGMENTS

I wish to thank my co-major professors, Dr. Richard J. Gladon of

the horticulture department, and Dr. Cecil R. Stewart of the botany

department, for their guidance and suggestions during the course of my

Ph.D. degree program. I also thank Dr. Stewart for kindly letting me

use his Apple lie computer to plot the graphs that are in Sections II

and III. In addition, I thank Dr. Donald Nevins, Dr. Charles V. Hall and Dr. Carl L. Tipton for serving as members on my program of study committee.

Dr. William Summers of the horticulture department showed me the

basics of operating the high performance liquid chromatograph, which was vital to my research, and I am grateful to his assistance.

I also thank Dr. Kenneth Eskins of the USDA Northern Regional

Research Laboratory, Peoria, Illinois, for instructing me on various aspects of plant pigment chromatography and quantitation. This experience provided me with a basis on which I expanded that enabled me to complete the research for Section I of this dissertation.

The graduate students of the horticulture department were a special group of people. Not only did I receive a lot of good advice from my peers, but also the bonds of friendship that formed were very rewarding and will be with me always.

I wish to warmly thank my parents, who not only helped me financially all through my undergraduate and graduate schooling, but also stood behind me all the way, offering words of encouragement when needed. 80

Finally, I thank Mrs. Lena Keithahn for being a true inspiration to me. One of the best things that happened to me was the luck of having her as my biology teacher during my sophomore year in high school. Her enthusiasm for biology was contagious, and I credit her for my scientific curiosity that has brought me to this point.