(Received for publication, October 20, 1995; and in revised form, January 29, 1996)
From the
The subcellular distributions of folate and folate-synthesizing
enzymes were investigated in pea leaves. It was observed that the
mitochondrial folate pool (400 µM) represented
50% of the total pool. Furthermore, all the enzymes involved in
tetrahydrofolate polyglutamate synthesis were present in the
mitochondria. In marked contrast, we failed to detect any significant
activity of these enzymes in chloroplasts, cytosol, and nuclei. The
presence of the tetrahydrofolate synthesis pathway in mitochondria is
apparently a general feature in plants since potato tuber mitochondria
also contained a high folate concentration (
200 µM)
and all the enzymes required for tetrahydrofolate polyglutamate
synthesis.
The specific activities of tetrahydrofolate-synthesizing
enzymes were rather low (1.5-15 nmol h mg
matrix protein), except for dihydrofolate
reductase (180-500 nmol h
mg
matrix protein). Dihydrofolate reductase was purified to
homogeneity. The enzyme had a native molecular mass of
140 kDa and
was constituted of two identical 62-kDa subunits. Interestingly, this
mitochondrial protein appeared to be a bifunctional enzyme, also
supporting thymidylate synthesis. The cell distribution of thymidylate
synthase was also investigated. No significant activity was observed in
cell fractions other than mitochondria, indicating that plant cell
mitochondria are also a major site for thymidylate synthesis.
In all organisms, one-carbon transfer reactions are mainly
mediated by tetrahydrofolate polyglutamate coenzymes(1) . As a
result, a number of pathways such as those involved in the metabolism
of methionine, serine, purine, or thymidylate are dependent on an
endogenous supply of these coenzymes(2) . Because animals lack
the first three steps of folate biosynthesis, folate supply in these
organisms is dependent on feeding. In contrast, plants and
microorganisms are able to synthesize tetrahydrofolate de
novo. This pathway requires the sequential operation of five
enzymes: a dihydropterin pyrophosphokinase (HPPK), ()a
dihydropteroate synthase (DHPS), a dihydrofolate synthetase (DHFS), a
dihydrofolate reductase (DHFR), and a folylpolyglutamate synthetase
(FPGS) (Fig. 1). The second step of this pathway is the target
of antimicrobial sulfonamide drugs, and considerable effort has been
focused on the molecular characterization of the bacterial enzymes
involved in the early steps of folate
synthesis(3, 4, 5) . In plants, the study of
folate metabolism is complicated by the presence of different
subcellular compartments. Indeed, there is now some evidences that, in
addition to the cytosol, discrete pools of folate are associated with
mitochondria and chloroplasts(6) . The distribution of
folate-synthesizing enzymes between the cytosol, mitochondria, and
plastids remains, however, uncertain, and it is not clear whether all
these compartments are autonomous for folate synthesis. The HPPK and
DHPS activities were reported to be carried on a single bifunctional
enzyme in plants (7) and in Plasmodium(8) .
This situation is different in bacteria, where two separate proteins
are involved(3, 9) . Fractionation studies in pea
seedlings suggested the presence of the enzyme in a
``mitochondrial'' fraction (73%) and in a
``soluble'' fraction (25%)(10) , but an eventual
cross-contamination of these fractions was not investigated. Likewise,
DHFS from pea seedlings was reported to be present in a mitochondrial
fraction (40%), a ``chloroplastic'' fraction (15%), and a
soluble fraction (22%), but the cross-contamination of the fractions
was not estimated(11) . DHFR in plants was described either as
a bifunctional enzyme, also supporting the thymidylate synthase (TS)
activity(12, 13) , or as a monofunctional polypeptide
associated with TS in a large multimeric enzyme complex(14) .
In protozoa, the DHFR and TS activities were also reported to be
carried on a single protein, but not in prokaryotes and mammals, where
two different proteins are involved (for a review, see (15) ).
The enzyme has been localized in the cytosol of animal cells (16) , but its compartmentalization in plants remains to be
established. FPGS was reported to be cytosolic and mitochondrial in
mammalian cells (17) and in fungi(18) , but its
distribution in higher plant cells is still unknown. In bacteria, this
enzyme is bifunctional, supporting also the dihydrofolate synthetase
activity(5) .
Figure 1:
HPteGlu
synthesis pathway. 1, dihydropterin
pyrophosphokinase; 2, dihydropteroate synthase; 3,
dihydrofolate synthetase; 4, dihydrofolate reductase; 5, folylpolyglutamate synthetase. pABA, p-aminobenzoic acid.
Subcellular localization of folate synthesis in higher plant cells is hampered by the difficulty of obtaining large amounts of purified organelles devoid of contamination from the other cell fractions. So far, the only reported studies were focused on the dihydropteroate synthase and the dihydrofolate synthetase(10, 11) , but the different cellular fractions were not purified, and the cross-contamination of these fractions was not measured. Furthermore, the distribution of folate (and the distribution of the enzymes involved in its synthesis) might vary from one tissue to another, depending on the specificity of the tissue. From this point of view, the presence in leaf mitochondria of the glycine cleavage system, which represents 40% of the soluble proteins and requires folate as cofactor(19) , might affect the cellular folate distribution. In this study, we investigated the presence of the tetrahydrofolate-synthesizing enzymes in the different cell compartments of pea leaves using Percoll-purified mitochondria, chloroplasts, and nuclei or a cytosol-enriched fraction. In addition, we purified the mitochondrial DHFR/TS bifunctional enzyme, and we studied some of its biochemical properties. We also determined the TS distribution in the leaf cell. Our results indicate that mitochondria are a major site for tetrahydrofolate polyglutamate and thymidylate synthesis in plants.
Mitochondria were isolated and purified as described previously (23) using a self-generating gradient of Percoll. Chloroplasts were isolated and purified on a discontinuous Percoll gradient as described by Douce and Joyard(24) . Protein extracts from these cell organelles were obtained as described previously(25) . With these experimental procedures, mitochondria and chloroplasts were usually devoid of contamination from the other compartments.
Nuclei were extracted from pea leaves and purified as described by Dunham and Bryant(26) . Purified nuclei were devoid of contamination from mitochondria. Contamination by cytosol proteins was very low (up to 1% of the total protein), but contamination by proteins from chloroplasts was up to 30% of the total protein.
To obtain the cytosol-enriched fraction, leaves (0.5 kg)
were ground with a Waring blender at low speed for 3 s in 1 liter of a
medium containing 0.3 M mannitol, 20 mM sodium
pyrophosphate, pH 7.5, 0.5% (w/v) polyvinylpyrrolidone, 10 mM -mercaptoethanol, 15 mM malate, 10% (v/v) glycerol,
and 1 mM phenylmethylsulfonyl fluoride. After filtration on
three layers of gauze, the filtrate was centrifuged at 1500
g to remove cellular debris. The supernatant was centrifuged
at 18,000
g to remove all the cell organelles, and the
remaining soluble fraction, enriched with cytosol, was concentrated to
3-5 ml using a Diaflo XM-10 membrane and an Amicon stirred
cell.
To obtain the leaf extracts, leaves (0.3 kg) were ground with
a Waring blender at high speed for 10 s in 0.5 liter of a medium
containing 100 mM potassium P, pH 7.5, 0.5% (w/v)
polyvinylpyrrolidone, 10 mM
-mercaptoethanol, and 1
mM phenylmethylsulfonyl fluoride. After filtration on three
layers of gauze, the filtrate was centrifuged at 1500
g to remove cellular debris and then at 18,000
g to
remove the membranes. The supernatant was concentrated to a final
volume of 5 ml using a Diaflo XM-10 membrane and an Amicon stirred
cell.
Folate, pteroate, and pterin were obtained from Sigma. Pterin pyrophosphate and folate polyglutamate were obtained from Schircks Laboratory (Jona, Switzerland). These products were reduced as dihydro and tetrahydro compounds as described by Scrimgeour(27) .
The HPPK activity was estimated in
association with the DHPS activity. The standard reaction medium
(medium A) contained, in a total volume of 120 µl, 20 mM Tris, 20 mM KHPO
, pH 8, 20 mM
-mercaptoethanol, 15 mM MgCl
, 10 mM ATP, and various amounts of our protein extracts. 2 µl of 2
mMp-[carboxyl-
C]aminobenzoic
acid (1.85 GBq mmol
) were added to the assay medium,
and then the reaction was started by the addition of 100 µM dihydropterin. After 20 min of incubation, the reaction was
stopped by heating the samples at 100 °C for 5 min. The samples
were centrifuged to remove the precipitated proteins, and the
[
C]dihydropteroate formed was determined with a
reverse-phase HPLC system (Nova-Pak C
column, Waters)
coupled with a Berthold LB 506D scintillation counter. The HPLC
conditions were as follows: solvent A, 0.1 M sodium acetate,
pH 6; and solvent B, acetonitrile. Solvent B was increased linearly 1%
every minute.
The DHPS activity was measured in medium A (final
volume of 120 µl) devoid of ATP. 2 µl of 2 mMp-[carboxyl-C]aminobenzoic
acid (1.85 GBq mmol
) were added in the assay medium,
and then the reaction was started by the addition of 100 µM dihydropterin pyrophosphate. After 5 min of incubation, the
reaction was stopped, and the [
C]dihydropteroate
formed was estimated as described above.
The DHFS activity was
measured in medium A (final volume of 120 µl) in the presence of 3
mM [H]glutamate (0.5 GBq
mmol
). This medium contained 20 mM K
HPO
buffer. A further addition of
K
monovalent cation, which is required for DHFS
activity (29) , did not significantly improve the reaction
rate. The reaction was started by the addition of 100 µM dihydropteroate. After 20 min of incubation, the reaction was
stopped by the addition of 0.5 ml of a solution containing 0.1 M sodium acetate, pH 5, and 50 mM glutamate. The sample was
then loaded on an ion-exchange column (DE52, Whatman) previously
equilibrated with the sodium acetate/glutamate solution. The column was
washed with 12 ml of this solution, and then the
[
H]dihydrofolate formed, which was retained on
the column, was eluted with 2 ml of 1 N HCl and counted.
The FPGS activity was determined by the same method as described above, except that the reaction was initiated with 100 µM tetrahydrofolate instead of dihydropteroate.
The DHFR activity
was monitored spectrophotometrically by measuring the oxidation of
NADPH at 340 nm ( = 12.3
cm
). The assay mixture contained 50 mM potassium P
, pH 7.2, 20 mM
-mercaptoethanol, 1 mM dithiothreitol, 0.2 mM NADPH, 0.2 mM H
FGlu
, and known
amounts of protein extract in a final volume of 0.5 ml.
The TS
activity was measured by the tritium release assay (30) in 50
mM Tris, pH 7.5, containing 30 mM -mercaptoethanol, 0.25 mM H
PteGlu
, 2.5 mM formaldehyde, 100
µM [5-
H]dUMP (0.16 GBq
mmol
), and protein extract in a final volume of 1
ml. At various times, 0.2 ml of the reaction medium was withdrawn and
added to 1 ml of a 2% trichloroacetic acid solution containing 0.2 g of
charcoal (initially washed with the 2% trichloroacetic acid solution)
to remove free dUMP. After vigorous shaking, the suspension was
centrifuged, and 0.4 ml of the clear upper phase was counted to
quantify
H
O released from
[5-
H]dUMP. Activity is expressed as nmol of dUMP
formed h
.
The SHMT activity was measured according to Taylor and Weissbach (31) as described previously(32) .
To determine the methylenetetrahydrofolate
dehydrogenase activity, known amounts of our protein extract were added
to a medium containing 50 mM potassium P, pH 7.5,
5 mM NADP, and 2 mM HCHO in a final volume of 500
µl. The reaction was started by the addition of 100 µM tetrahydrofolate, and NADPH formation was monitored
spectrophotometrically at 340 nm.
The proteins were loaded on an anionic DEAE-Sepharose
column previously equilibrated with medium B at pH 7.5 and connected to
a Pharmacia fast protein liquid chromatography system. The column was
washed with medium B, at a flow rate of 0.3 ml min,
until the A
of the effluent was <0.1.
Proteins were eluted with a linear gradient of 0-1 M KCl
in medium B. Active fractions containing DHFR activity were pooled,
dialyzed against medium B, and concentrated by ultrafiltration to
2-3 ml.
In a final step, DHFR was purified by affinity
chromatography on a methotrexate-agarose column. The sample was applied
to the column previously equilibrated with medium B and connected to
the fast protein liquid chromatography system. The flow rate was 0.2 ml
min. The column was washed until the A
of the effluent was <0.1. The column was
eluted with a linear gradient of 0-1 M KCl in medium B.
At this stage, the DHFR remained fixed on the column. After washing the
column with 12 ml of medium B containing 1 M KCl, the DHFR was
finally eluted with 10 ml of a buffer containing 50 mM Tris,
pH 8.5, 1 M KCl, 1 mM H
FGlu
,
30 mM
-mercaptoethanol, 1 mM dithiothreitol, 1
mM EDTA, and 15% (w/v) glycerol. To avoid oxidation of
H
FGlu
, this buffer was previously bubbled with
argon.
The different activities
involved in tetrahydrofolate polyglutamate synthesis in mitochondria
were then separated by gel filtration (see ``Experimental
Procedures''). As shown in Fig. 2, the HPPK and DHPS
activities coeluted with an apparent molecular mass of 300 kDa.
This value is higher that the one previously reported for the HPPK/DHPS
bifunctional enzyme isolated from pea seedlings (180 kDa) (7) .
The DHFS activity was eluted with an apparent molecular mass of 54 kDa,
in good agreement with a previous report(29) . The FPGS
activity was eluted with an apparent molecular mass of 70 kDa, a value
comparable to those previously reported for plant and mammal
FPGS(39, 40) . The DHFR activity was eluted with a
molecular mass corresponding to
140 kDa. Similar molecular masses
were previously reported for DHFR originating from soybean seedling (41) and carrot cell suspension cultures(42) . In
protozoa, the apparent molecular mass of the enzyme was also estimated
at 150 kDa by gel filtration(43) . In this last case, however,
DHFR appeared to be a bifunctional protein associated with
TS(43) . Interestingly, the TS activity in plant mitochondria
also coeluted with the DHFR activity (Fig. 2). These results
suggest that the DHFR and TS activities from pea leaf mitochondria are
also supported by the same enzyme. To verify this, experiments were
undertaken to purify mitochondrial DHFR.
Figure 2:
Separation by gel filtration of the
different enzyme activities involved in tetrahydrofolate polyglutamate
synthesis. A, separation of the HPPK (), DHPS (
),
and DHFS (
) activities. To facilitate the graph reading, the DHPS
activity was reduced three times. B, separation of the DHFR
(
), TS (*), and FPGS (
) activities. To facilitate the graph
reading, the DHFR activity was reduced 20
times.
Figure 3: SDS-polyacrylamide gel electrophoresis analysis of DHFR/TS purified from pea leaf mitochondria. Lane A, standards; lane B, 90 µg of matrix extract; lane C, 1 µg of purified DHFR/TS.
Figure 4:
HFGlu
reduction
catalyzed by a mitochondrial enzyme extract. A, proteins from
the DEAE-Sepharose column (0.7 mg) were added to 500 µl of the
reaction medium (see ``Experimental Procedures''), and the
reaction was immediately initiated by adding
H
FGlu
. The DHFR activity was monitored at 30
°C by measuring the rate of NADPH consumption. B, the
proteins were incubated for 5 min in the presence of NADPH before the
reaction was initiated by adding H
FGlu
. Numbers along the lines are nmol of NADPH consumed
min
mg
protein. MTX,
methotrexate.
In contrast to DHFR activity, TS
activity exhibited no lag phase, but slowly declined with time, even in
the presence of saturating amounts of dUMP (100 µM) and
CH-H
FGlu
(100 µM) (Fig. 5). However, in the presence of 1 mM NADPH, the
reaction appeared linear for at least 90 min. These results strongly
suggest that the time-dependent inhibition was the result of
H
FGlu
accumulation. Indeed, in the presence of
NADPH, H
FGlu
produced by TS activity could be
recycled into H
FGlu
owing to the associated
DHFR activity. The addition of 1 µM FdUMP, a potent
inhibitor of monofunctional thymidylate synthase(45) ,
completely blocked the reaction (Fig. 5). The maximal TS
activity was obtained for temperatures close to 40 °C, and the
optimum pH was between 7 and 7.5 (data not shown). The affinity
constants for dUMP and CH
-H
FGlu
(assuming that only half of the
CH
-H
FGlu
, the pro-R form,
interacted with the enzyme) were 1.5 ± 0.3 and 22 ± 2.5
µM, respectively. In contrast to DHFR, increasing the
polyglutamate chain length resulted in a considerable decrease in the K
of CH
-H
FGlu
for TS (Table 4). The V
of the
reaction was also affected by the length of the polyglutamate chain and
increased
3 times when the number of glutamates increased from one
to five (Table 4).
Figure 5:
Thymidylate synthesis catalyzed by a
mitochondrial enzyme extract. Proteins from the DEAE-Sepharose column
(0.65 mg) were added to 2.5 ml of the reaction medium containing 100
µM (6R,6S)-CH-H
FGlu
.
At each time point, 200 µl were withdrawn to determine the amount
of dTMP formed (see ``Experimental Procedures''). In the
presence of NADPH, the reaction was maintained linear for at least 50
min.
The activity ratio of DHFR to TS was
generally between 20 and 30. However, we observed that, during storage,
this ratio could increase considerably because of instability of the
catalytic domain responsible for TS activity. In this connection, it is
interesting to note that the domain of the enzyme responsible for TS
activity was much more sensitive to protease action than the DHFR
domain. Indeed, as shown in Fig. 6, when partially purified
enzyme was incubated at 5 °C in the presence of trypsin (20 µg
ml), TS activity rapidly declined and dropped to 0
after 30 min. In contrast, during the same period, DHFR activity was
unaffected and remained constant even after 2 h of incubation. This
last observation strongly supports the idea that the two domains
catalyzing the DHFR and TS activities are spatially distinct. This
assumption is also reinforced by the observation that FdUMP, a potent
inhibitor of TS activity, did not affect DHFR activity (data not
shown).
Figure 6:
Effect
of trypsin on the DHFR and TS activities. The DHFR and TS activities
were measured at 30 °C under the optimal conditions shown in Fig. 4and Fig. 5. At t = 0, trypsin
(final concentration, 17 µg ml) was added to the
reaction medium. Rates are expressed as percent of the initial maximal
velocities.
The results presented here indicate, for the first time, that
higher plant mitochondria are a major site for tetrahydrofolate
synthesis. Indeed, we observed that the largest part of the cellular
folate and all the enzymes required for tetrahydrofolate synthesis were
located in this compartment. Surprisingly, we were not able to detect
these enzymes in chloroplasts, nuclei, or cytosol. Although it is
difficult to ascertain negative results, it is clear that these
activities, if present, were much lower in these last three
compartments than in mitochondria. These observations are in agreement
with previous reports indicating that a large part of the DHPS and DHFS
activities is localized in pea seedling
mitochondria(10, 11) . Some minor DHPS and DHFS
activities in chloroplastic and cytosolic fractions were also observed,
but these locations were subject to discussion because of contamination
from mitochondria(11) . In marked contrast with the results
presented here, the DHFR and FPGS activities in mammalian cells were
believed to be primarily localized in the cytosol (1, 16, 46) . In Neurospora crassa,
however, 50% of the FPGS activity was cytosolic and 50%
mitochondrial(18) . These two FPGS isoenzymes differed in their
substrate specificity, the mitochondrial isoform catalyzing glutamate
addition only with H
PteGlu
as substrate and not
with H
PteGlu
. Thus, it appears from these data
that the cell distribution of folate-synthesizing enzymes may greatly
vary from one species to another. Yet, it must be pointed out that
these localization studies were mainly based on cell fractionation
experiments, where inevitable cross-contamination makes the results
difficult to interpret. From this point of view, the localization of
the enzyme activities in purified cell fractions such as
Percoll-purified mitochondria or chloroplasts would permit more
definite conclusions. In this context, it has been recently
demonstrated that FPGS, with a high specific activity, was also present
in mammalian mitochondria purified on a Percoll density gradient (17) .
In animal cells, folate is supplied from the external medium, and the mitochondrial pool is obtained from cytosolic folate uptake(47) . In plants, the situation is totally different because plant cells can synthesize folate de novo within mitochondria. However, tetrahydrofolate derivatives are an absolute requirement for the synthesis of numerous compounds such as purine or thymidylate and must therefore also be present in chloroplasts and cytosol. Thus, taking into account our results, we are forced to imagine that, in marked contrast with the situation encountered in animal cells, polyglutamyltetrahydrofolates required in chloroplasts and cytosol are, at least partly, supplied from mitochondria. Little is known about folate transport through biological membranes, but the cell membrane permeability to polyglutamylfolate is apparently considerably less than to monoglutamate species(2) . Analysis of mitochondrial folate in pea leaves revealed a pool of polyglutamates dominated by tetraglutamate (25%) and pentaglutamate (55%)(32) , a situation that therefore will limit the mitochondrial folate ability to be transported across the membranes. Furthermore, tetrahydrofolate polyglutamate is a very labile molecule that undergoes rapid oxidation in the presence of oxygen(48) . This oxidative degradation is, however, prevented when tetrahydrofolate is bound to folate-dependent proteins, a situation greatly favored by long polyglutamate chains(48) . These observations strongly suggest that tetrahydrofolate cannot diffuse freely within the cell. Unfortunately, there is at present no information available regarding the cellular traffic of folate, and the questions concerning the mode of transport of this cofactor between the different compartments (cotransport with a binding protein, endocytosis, or potocytosis(49) ) remain to be answered.
The specific
activities of the enzymes involved in tetrahydrofolate synthesis in
plant mitochondria were rather low (1.5-15 nmol h mg
matrix protein), except for DHFR. Indeed,
DHFR had a specific activity 10-100 times higher than the other
proteins, suggesting a possible role in other metabolic functions. As a
matter of fact, DHFR in plant mitochondria is involved not only in
tetrahydrofolate synthesis, but also in thymidylate synthesis. This
situation is also found in
protozoa(43, 50, 51) . It is interesting to
note that all the experiments done with various species of protozoa (51) indicated only one DHFR/TS activity. The strong analogy
between the bifunctional plant enzyme and that from protozoa suggests
that, in these latter organisms, DHFR/TS is also localized in
mitochondria. In contrast, DHFR in mammalian cells appeared to be a
monomeric cytosolic enzyme separated from TS activity(15) .
Although the bifunctional DHFR/TS enzyme had been intensively studied
at the level of molecular biology(13, 52) , there is
little information concerning its biochemical properties. Our
preliminary studies indicate that DHFR/TS has similar properties to
monofunctional DHFR. In particular, both DHFRs exhibit a high affinity
for either the monoglutamate or the polyglutamate form of
dihydrofolate, which illustrates that both types of enzymes play a dual
role in tetrahydrofolate biosynthesis and in recycling of
H
FGlu
formed during thymidylate biosynthesis.
Finally, our results indicate that, in plants, TS was mainly localized in mitochondria. Again, the situation is very different in animal cells, where TS is a monofunctional protein localized in the cytosol and, in the case of actively dividing tissue, closely associated with nuclei(46) . Although we cannot rule out the possibility that TS cell distribution in actively dividing plant tissues is different from that reported here, mitochondria from young leaves appeared to be a major site for thymidylate synthesis. This raises the question of the transport of thymidylate toward nuclei and plastids, a point currently under investigation.