Retrospective Theses and Dissertations
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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McMahon, Robert Waring, "Relationship of porphobilinogen deaminase activity to chlorophyll content and development of 'Heinz
1350' tomato fruit " (1984). Retrospective Theses and Dissertations. Paper 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
University
iVIicrofilms
IntGrnBtiOn&l 300N.zeeb Road, Ann Arbor, Ml48106
PH.D. 1984
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
OF CHLOROPLAST PIGMENTS IN FRUITS OF
HORTICULTURAL CROPS BY HIGH PERFORMANCE
LIQUID CHROMATOGRAPHY (HPLC)
14
ABSTRACT
15
INTRODUCTION
16
MATERIALS AND METHODS
18
RESULTS AND DISCUSSION
20
LITERATURE CITED
27
SECTION II. PROPERTIES OF PORPHOBILINOGEN DEAMINASE
ISOLATED FROM 'HEINZ 1350' TOMATO FRUIT
28
ABSTRACT
29
INTRODUCTION
30
MATERIALS AND METHODS
31
RESULTS
35
DISCUSSION
44
iii
LITERATURE CITED
SECTION III. PORPHOBILINOGEN DEAMINASE ACTIVITY
IN DEVELOPING 'HEINZ 1350' TOMATO
FRUIT AND ITS RELATIONSHIP TO TOMATO
FRUIT CHLOROPHYLL CONTENT
50
53
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
GENERAL LITERATURE REVIEW
7%
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
predominately, while 3, 4-^^C glutamate labeled the
of ALA.
of ALA
to
portion
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
than glutamate-
or -
to
ALA better
, 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 enzymebound 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:
insertion of
1) the
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=yScarotene
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
Anthesis
(Days)
Fruit Fresh
Weight
(g)
Chi a
(mg/fruit)
Chi b
(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
^Mean value of 3 replicates each containing 4 fruits.
— mm
25
Table 2. Chlorophyll a and b contents of various horticultural crops
as determined using the methods developed in this report^
Chlorophyll a
UR/g, Fresh Weight
Broccoli Florets
Chlorophyll b
ttg/g Fresh Weight
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
73.4
30.8
Zucchini Squash
^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 12:808809.
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, Nethylmaleimide, 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 analytical
^G
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^ was 4.94 for 27C to 37C, 3.69 for
Q
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
R
P
0
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
0.0
0.0
-2.0
p-chloromercuribenzoate -30.0
-87.4
-94.0
HgClg
-96.0
-96.4
-97.0
+4.5
+6.9
+7.7
N-ethylmaleimide
^-aminolevulinic acid
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^ for
Q
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^ for 47C to
Q
57C was much smaller than the previous two Q^ values, and thermal
Q
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^ for 37C
Q
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 Nethylmaleimide at the concentrations tested.
For most sulfhydryl enzymes, reducing agents such as 2mercaptoethanol 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 2mercaptoethanol 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 2mercaptoethanol 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. 233:501509.
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 293:506513.
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 2aminoethyltripyrranes 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 fruitschanges 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.
6.
1981. Tomato production in bags. Benchmarks 1:4-
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.
objectives of this research were to:
Therefore, the
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
H
L
D
20
S
P
15
?
L
L
/
G
F
10
CHL a
=2,40
- XCHLb
0.05
H
10
f
1-
f
f
f
1-
f
15
20
25
30
35
40
45
50
55
f
f
- 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
LSD,
0.05
= 21.87
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
b
LS00.05=*:9
CHL
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 481510 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 437482
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 fruitschanges 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
LITERATURE CITED FOR THE GENERAL INTRODUCTION
AND THE GENERAL LITERATURE REVIEW
1. Battersby, A. R., G. L. Hodgson, E. Hunt, E. McDonald and J.
Saunders. 1976. Biosynthesis of porphyrins and related
macrocycles. Part VI. Nature of the rearrangement process
leading to the natural type III porphyrins. J. Chem. Soc.
Perkin Trans. 1:273-282.
2. Battersby, A. R. and E. McDonald. 1975. Biosynthesis of
porphyrins, chlorins and corrins. Pages 62-122 in J. E.
Smith, ed. Porphyrins and metalloporphyrins. Elsevier
Scientific Publishing Co., Amsterdam.
3. Battersby, A. R. and E. McDonald. 1976. Biosynthesis of
porphyrins and corrins. Phil. Trans. R. Soc. London, B,
273:161-180.
4. Beale, S. I. 1976. The biosynthesis of d-aminolaevulinic acid
in plants. Phil. Trans. R. Soc. London, B, 273:99-108.
5. Beale, S. I. and P. A. Castelfranco. 1973. The formation of 6arainolevulinic acid from labeled precursors by greening
cucumber cotyledons (Abstr.). Plant Physiol. 51 (Suppl.):101.
6. Beale, S. I. and P. A. Castelfranco. 1974. The biosynthesis of
(5-aminolevulinic acid in higher plants. I. Accumulation of
3-aminolevulinic acid in greening plant tissues. Plant
Physiol. 53:291-296.
7. Beale, S. I. and P. A. Castelfranco. 1974. The biosynthesis of
^^aminolevulinic acid in higher plants. II. Formation of
C-d-aminolevulinic acid from labeled precursors in greening
plant tissues. Plant Physiol. 53:297-303.
8. Beale, S. I., S. P. Gough and S. Granick. 1975. Biosynthesis of
Ô-aminolevulinic acid from the intact carbon skeleton of
glutamic acid in greening barley. Proc. Nat. Acad. Sci.
U.S.A. 72:2719-2723.
9. Bogorad, L. 1958. The enzymatic synthesis of porphyrins from
porphobilinogen. II. Uroporphyrin III. J. Biol. Chem.
233:510-515.
10. Bogorad, L. 1966. The biosynthesis of chlorophylls. Pages 481510 in L. P. Vernon and G. R. Seely, eds. The Chlorophylls.
Academic Press, London.
11. Bogorad, L. 1979. Biosynthesis of porphyrins. Pages 125-178 iji
75
D. Dolphin, ed. The Porphyrins. Volume VI. Biochemistry. Part
A. Academic Press, New York.
12. 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.
13. 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.
14. Burton, G., P. E. Fagerness, 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. Chera.
Soc. Chem. Commun. 5:202-204.
15. Davies, R. C. and A. Neuberger. 1973. Polypyrroles formed from
porphobilinogen and amines by uroporphyrinogen synthetase of
Rhodopseudomonas spheroides. Biochem. J. 133:471-492.
16. Eskin, N. A. 1979. Plant Pigments, Flavors and Textures: The
Chemistry and Biochemistry of Selected Compounds. Academic
Press, New York. 219 pages.
17. 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.
18. Frydman, B. and R. B. Frydman. 1975. Biosynthesis of
uroporphyrinogens from porphobilinogen. Mechanism and nature
of the process. Acc. Chem. Res. 8:201-208.
19. 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
20. Frydman, R. B. and B. Frydman. 1970. Purification and
properties of porphobilinogen deaminase from wheat germ.
Arch. Biochem. Biophys. 136:193-202.
21. Frydman, R. B. and B. Frydman. 1979. Disappearance of
porphobilinogen deaminase activity in leaves before the onset
of senescence. Plant Physiol. 63:1154-1157.
22. Frydman, R. B., B. Frydman and A. Valasinas. 1979.
Protoporphyrin: synthesis and biosynthesis of its metabolic
76
intermediates. Pages 1-124
D. Dolphin, ed. The
Porphyrins. Volume VI. Biochemistry. Part A. Academic Press,
New York.
23.
Frydman, R. B., B. Frydraan, A. Valasinas and E. S. Levy. 1977.
Biosynthesis of uroporphyrinogen III. Interaction among 2aminoethyltripyrranes and the enzymatic system. Pages 165-178
F. R. Longo, ed. Porphyrin Chemistry Advances. Ann Arbor
Science Publishers, Inc., Ann Arbor, Michigan.
24. Gough, S. P. 1978. Light stimulated 6-aminolevulinate
accumulation in levulinate treated barley seedlings.
Carlsberg Res. Commun. 43:497-508.
25. Harel, E. 1978. Chlorophyll biosynthesis and its control.
Prog. Phytochem. 5:127-180.
26.
Harel, E. and S. Klein. 1972. Light dependent formation of <5aminolevulinic acid in etiolated leaves of higher plants.
Biochem. Biophys. Res. Commun. 49:364-370.
27.
Harel, E., E. Meller and M. Rosenberg. 1978. Synthesis of 5aminolevulinic acidC by cell-free preparations from
greening maize leaves. Phytochemistry 17:1277-1280.
28.
Jones, 0. T. G. 1968. Biosynthesis of chlorophylls. Pages 131145 in T. W. Goodwin, ed. Porphyrins and Related Compounds.
Academic Press, New York.
29.
Jordan, P. M. and J. S. Seehra. 1979. The biosynthesis of
uroporphyrinogen III:
order of assembly of the four
porphobilinogen molecules in the formation of the tetrapyrrole
ring. FEBS Letters 104:364-366.
30.
Jordan, P. M. and D. Shemin. 1973. Purification and properties
of uroporphyrinogen I synthetase from Rhodopseudomonas
spheroides. J. Biol. Chem. 248:1019-1024.
31.
Jordan,. P. M., G. Burton, H. Nordlov, 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.
32.
Jordan, P. M., H. Nordlov, G. Burton and A. I. Scott. 1980. A
rapid direct assay for uroporphyrinogen III cosynthase. FEBS
Letters 115:269-272.
33. Khudairi, A. K. 1972. The ripening of tomatoes.
Scientist 60:696-707.
Amer.
77
34. Kipe-Nolt, J. A. and S. E. Stevens, Jr. 198O. Biosynthesis of
5-aminolevulinic acid from glutamate in Agmenellum
quadruplicatum. Plant Physiol. 65:126-128.
35. Kirk, J. T. 0. 1968. Studies on the dependence of chlorophyll
synthesis on protein synthesis in Euglena gracillis, together
with a nomogram for determination of chlorophyll
concentration. Planta 78:200-207.
36.
Krogmann, D. W. 1973» The Biochemistry of Green Plants.
Prentice-Hall, Inc., Englewood Cliffs, New Jersey. 239 pages.
37. Levin, E. Y. 1968. Uroporphyrinogen III cosynthase from mouse
spleen. Biochemistry 7:3781-3788.
38.
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.
39. 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.
40. Marriott, J. 1968. Regulation of porphyrin synthesis. Pages
61-74 in T. W. Goodwin, ed. Porphyrins and Related Compounds.
Academic Press, New York.
41. Martin, B. A., J. A. Gauger and N. E. Tolbert. 1979. Changes in
activity of ribulose-1, 5-bisphosphate carboxylase/oxygenase
and three peroxisomal enzymes during tomato fruit development
and ripening. Plant Physiol. 63:486-489.
42. Moore, M. R. 1980. The biochemistry of the porphyrins. Clin.
Haematol. 9:227-252.
43. Rebeiz, C. A. and P. A. Castelfranco. 1973. Protochlorophyll
and chlorophyll biosynthesis in cell-free systems from higher
plants. Ann. Rev. Plant Physiol. 24:129-172.
44. Rick, C. M.
1978.
The Tomato. Scientific American 239:76-87.
45. Robinson, T. 1980. The Organic Constituents pf Higher Plants.
Their Chemistry and Interrelationships. 4 edition. Cordus
Press, North Amherst, Massachusetts. 352 pages.
46. 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.
78
47. Simpson, D. J., M. R. Baqar, W. M. McGlasson and T. H. Lee.
1976. Changes in ultrastructure and pigment content during
development and senescence of fruits of normal and rin and nor
mutant tomatoes. Aust. J. Plant Physiol. 3:575-587.
48. Stevens, E. and B. Frydman. 1968. Isolation and properties of
wheat germ uroporphyrinogen III cosynthase. Biochim. Biophys.
Acta 151:429-437.
49. Warnick, G. R. and B. F. Burnham. 1971. Regulation of porphyrin
biosynthesis. Purification and characterization of 5aminolevulinic acid synthetase. J. Biol. Chem. 246:66806685.
50. Weinstein, J. D. and P. A. Castelfranco. 1977. Protoporphyrin
IX biosynthesis from glutamate in isolated greening
chloroplasts. Arch. Biochem. Biophys. 178:671-673.
51. Wider de Xifra, E. A., A. M. Stella and A. M. del C. Batlle.
1978. Porphyrin biosynthesis-immobilized enzymes and ligands.
IX. Studies on ^-aminolaevulinate synthetase from cultured
soybean cells. Plant Science Letters 11:93-98.
52. Yuan, M. and C. S. Russell. 1974. Porphobilinogen derivatives
as substrates for porphobilinogenase. FEBS Letters 46:34-38.
53.
Zelitch, I. 1971. Photosynthesis, Photorespiration and Plant
Productivity. Academic Press, New York. 347 pages.
79
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.