Hexose-monophosphate shunt activity in intact Plasmodium falciparum-infected erythrocytes and in free parasites

ELSEVIER Molecular and Biochemical Parasitology 67 (1994) 79-89

MOLECULAR AND BIOCHEMICAL PARASITOLOGY

Hexose-monophosphate shunt activity in intact Plasmodium falciparum-infected erythrocytes and in free parasites

Hani Atamna a, Gianpiero Pascarmona b, Hagai Ginsburg a,, a Department of Biological Chemistry, Institute of Life Sciences, Hebrew University, Jerusalem 91904, Israel

b Dipartimento di Genetica, Biologia e Chimica Medica, University of Torino Medical Schoo~ 10126 Torino, Italy

Received 15 March 1994; accepted 21 May 1994

Abstract

The hexose monophosphate shunt (HMS) produces NADPH for reductive antioxidant protection and for metabolic regulation, as well as ribose-5-phosphate needed for the synthesis of nucleic acids. Since malaria-infected red blood cells (RBC) are under endogenous oxidant stress, it was interesting to determine HMS activity in intact infected cells, as well as in free parasites. HMS activity was determined by measuring the evolution of 14CO2 from D-[1-14C]glucose in normal RBC, in intact Plasmodiumfalciparum-infected RBC (IRBC) and in free parasites. The HMS activity of IRBC was found to be 78 times higher than that of normal RBC. This activity increased with parasite maturation from the ring stage toward the trophozoite stage, and declined at the schizont stage. The HMS activity of the parasite contributes 82% of the total observed in the intact IRBC, and that of the host cell is increased some 24-fold. The increased reducing capacity of IRBC and free parasites were also evidenced by the larger ability for reductive accumulation of methylene blue. Since the endogenous oxidative stress is produced by the parasite digestion of the host cell's hemoglobin, inhibition of this process with protease inhibitors, by alkalinization of the parasite's food vacuole, or by the application of antimalarial drugs, resulted in 20-44% inhibition of IRBC HMS activity. A similar inhibition was observed in the presence of scavengers of oxidative radicals, uric and benzoic acids. These inhibitors had only a minor effect on the HMS activity of free parasites. D-[1-14C]glucose and D-[6-t4C]glucose contributed equally to newly synthesized nucleic acids, suggesting that ribose-5-phosphate needed for this synthesis is contributed by the non-oxidative activity of HMS. These results imply that a major portion of parasite HMS activity and the activated HMS of the host cell are devoted to counteract the endogenously generated oxidative stress.

Keywords: Malaria; Plasmodium falciparum; Oxidative stress; Hexose monophosphate shunt; NADPH; Compartment analysis

1. Introduction

Abbreviations: G6PD, glucose-6-phosphate dehydrogenase; GSH, glutathione; GSSG, oxidized GSH; HMS, hexose monophosphate shunt; IRBC, infected red blood cells; RBC, red blood cells; R-5-P, ribose-5-phosphate; ROS, reactive oxygen species; TRBC, trophozoite-infected RBC.

* Corresponding author. Tel.: 972-2-585-539; Fax: 972-2-666- 804; E-mail: [email protected].

The hexose monophosphate shunt (HMS) is the principal source of pentose sugars needed for nucleotide synthesis, and for the reduction of NADP + to NADPH. The human erythrocyte has a limited need for nucleotides and for reducing power, and hence, its HMS activity is rather restricted under normal conditions [1,2]. Thus, the first enzyme glu-

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80 H. Atamna et al. / Molecular and Biochemical Parasitology 67 (1994) 79-89

cose-6-phosphate dehydrogenase (G6PD) of the HMS which is regulated by the ratio NADPH/NADP +, operates at less than 1% of its maximal capacity. Reduction of NADPH levels either with methylene blue [3] or by its consumption by glutathione reductase which reduces oxidized glutathione (GSSG) produced during the reduction of H202 by glutathione peroxidase (the glutathione cycle) [4,5], lead to substantial activation of HMS.

The malaria-infected erythrocyte is demonstrably under oxidative stress: it produces 02, H202 and OH radicals [6-9], emerging from the digestion of host cell cytosol inside the acidic food vacuole of the parasite [10]. H202 is only partially reduced by the parasite's enzymes, and demonstrably reaches the host cell compartment. Hence, both the host's and the parasite's antioxidant defence systems must be considerably activated to counteract this oxidative challenge. Such activation requires the presence of the relevant enzymes, and it should be reflected by increased HMS activity and glutathione cycling. In- deed, parasites were shown to contain glutathione peroxidase [11,12] and glutathione reductase [13], enabling them to sustain the glutathione cycle. The presence of G6PD in the parasite has been a subject of controversy for many years [14], but it now seems to be convincingly established and partially characterized [15,16]. The enzyme detected in P. falciparum was reported to be different from that of the host, its activity being modest compared to that of the host (about 10% of the host activity), but its affinity for all substrates is higher than that of the host cell. HMS activity has been measured in very few instances in intact infected red blood cells (IRBC), but not in free parasites [17-19]. Thus, it could not be established whether the measured increase in activity was due to parasite HMS or due to the activation of the host cell pathway.

Other enzymes were suggested to mediate the reduction of NADP + to NADPH, namely, glutamate and isocitrate dehydrogenases: parasites contain glutamate dehydrogenase [19-22], and both host cell and parasite contain isocitrate dehydrogenase [22,23], but their physiological roles in the reduction of NADP + have not been assessed.

Investigations on the role of HMS in supplying ribose-5-phosphate (R-5-P) for the synthesis of nucleic acids, revealed that R-5-P is mostly produced

by the non-oxidative activity of HMS, i.e., using glyceraldehyde-3-P and fructose-6-P produced by glycolysis by HMS transaldolase and transketolase [24]. In view of the possible biochemical interrela- tionship between host cell and parasite, it seems that investigations of the HMS activities in isolated parasites are warranted.

In this report we demonstrate a considerable increase of HMS activity in IRBC, which depends on parasite development and on the endogenous production of oxidative species. Most of this activity can be assigned to the parasite itself, while some of it stems from the activated HMS activity of the host cell.

2. Materials and methods

Materials. D-J1-14 C]glucose and D-[6-14 C]glucose were obtained from Amersham; leupeptin, pepstatin, uric acid, benzoic acid, hypoxanthine, chloroquine, quinine, and glucose-free RPMI-1640, were pur- chased from Sigma Chemical Co, and methylene blue from Merck. Mefloquine was kindly supplied by Hoffman-La-Roche.

Parasite cultures. The FCR3 strain of Plasmodium falciparum was cultivated in either O + or A + washed human erythrocytes at 37°C, in RPMI-1640 (GIBCO) medium supplemented with 10% (v /v ) AB + or A + heat-inactivated plasma, 25 mM Hepes (N-2-hy- droxyethyl piperazine-N-2-ethane sulfonic acid), 30 mM NaHCO 3 and 10 mM glucose. The growth medium in 150 cm 2 culture flasks was replaced daily followed by gassing with a mixture of 90% N2, 5% CO 2 and 5% 02.

Infected red blood cells (IRBC) harboring mature parasite stages (trophozoites and schizonts) were concentrated from culture to > 90% parasitemia using the gelatin flotation method [25]. After separa- tion, cells were washed twice in wash medium (i.e., growth medium without plasma) and allowed to recover for 1 hour in culture conditions prior to use for HMS activity measurement. Cells were counted in a cell counter (Analys Instrument, Stockholm, Sweden) and the percentage of infected cell was determined by microscopic inspection of Giemsa- stained thin blood smears.

H. Atamna et al. / Molecular and Biochemical Parasitology 67 (1994) 79-89 81

For synchronization of parasite stage, asyn- chronous cultures (consisting mainly of rings) were incubated in isotonic solution of alanine (300 mM) at a final hematocrit of 8% for 5 min at 37°C. The ceils were then washed twice with wash medium and returned to culture conditions.

Measurement of riMS activity. Measurement of HMS activity was performed as previously described [26] in wash medium containing 5 mM glucose, 0.1 mM hypoxanthine and 10 mM NaHCO 3. A known number of cells was resuspended into 400 /zl of wash medium which contained 0.24/~Ci D-[1-14C]glucose. The cells were incubated for 30 min at 37°C. The 14CO2 released due to HMS activity was collected into 1 N KOH solution placed in a chamber tightly connected to the reaction chamber. At the end of the incubation, 0.35 ml of 3.5 N perchloric acid was injected into the cell suspension to stop the reaction, and the KOH-containing chamber was dipped into ice for 45 min, whereas the chamber of the ceils was left at 37°C. By this procedure the transfer of 14CO2 to the KOH is maximized (the solubility of CO 2 is known to increase with decreasing temperature). The KOH solution is thereafter added to scintillation solution and radioactivity is measured in a fl-counter. Normal BRC used in these experiments were maintained for 24 h under culture conditions.

For the measurement of HMS activity of free parasites, the infected cells (trophozoite stage) were treated with Sendai virus in order to release the host cell cytosol, as previously described [27]. Briefly, infected cells (2.5% hematocrit in wash medium) were incubated with Sendai virions (20-50 /zg protein ml-1, equivalent to 400-600 hemagglutination units) for 7 min. Cells were then washed twice with the same medium and used for measuring HMS activity. The results are given in /zmoles of glucose used by the HMS per 1 h per 101° cells.

Kinetics of methylene blue reduction. Normal RBC, IRBC and free parasites were suspended in wash medium containing 0.5 mM methylene blue, and incubated at 37°C. At the desired time interval, aliquots (100 /xl) were taken, centrifuged (14000 rpm in an Eppendorf microfuge, 1 min), and the supernatant was diluted 1:25 in wash medium. Ab- sorbance of oxidized methylene blue was measured

at 666 nm, and the amount of methylene blue reduced (101°) -1 cells was plotted against time.

Incorporation of radiolabel from D-[1-14C]glucose and D-[6-14C]glucose into parasite nucleic acids. Trophozoite IRBC were isolated by gelatin flotation and one half of the cells were used for obtaining free parasites. Cells were resuspended at 2.95 × 108 cells m1-1 wash medium containing 5 mM glucose and 50 /xM hypoxanthine. Either D-[1-14C]glucose or D-[6-14 C]glucose were added at 6/xCi m1-1, and the suspensions were incubated for 1 h at 37°C. Cells were then harvested with a cell Harvester (Dynatech, Inc.). The filters were washed with distilled water, then dried for 2 h at 60°C and transferred into toluene-based scintillation fluid for counting of radioactivity. No difference in incorporated label were observed if cells were first lysed by freezing and thawing, and then treated with proteinase K, implying that no measurable label was incorporated into proteins. Results (in DPM) were converted to /xmol glucose equivalents incorporated in nucleic acids h -1 (101°) -1 parasites.

3. Results

HMS activity in normal and infected RBC, and in free parasites. Incubation of uninfected or of P. falciparum-infected RBC in presence of D-[1- 14C]glucose resulted in the release of 14CO2, indicating HMS activity. Identical rates of HMS activity (in terms of amounts of glucose consumed were obtained when total glucose concentration was increased up to 25 mM, indicating that already at 5 mM the rate-limiting enzymatic step was functioning at its maximal rate, and that depletion of glucose from the medium was minimal during the time of the assay even at 5 mM total glucose. To assess that the release of 14CO2 is a genuine reflection of HMS activity, NRBC and IRBC were incubated also in the presence of D-[6-14C]glucose. No 14CO2 could be found in the KOH solution when [14 C6 ]-glucose was used as a substrate instead of [14C1]-glucose, even after 3 h of incubation (data not shown). Labeled [laC6]-glucose will not lose its labeled carbon by passing through the HMS, unless other metabolic

82 H. Atamna et al. /Molecular and Biochemical Parasitology 67 (1994) 79-89

I

2

trl "1-

0 i i i i

0 20 40 80 80 100

Parasitemia (%)

Fig. 1. HMS activity in parasite cultures as a function of parasitemia. Trophozoite-infected RBC were isolated by gelatin flotation to 100% parasitemia. These cells were diluted with uninfected cells of the same culture, to obtain the desired parasitemia. HMS activity (in /~mol glucose consumed h -1 (101° cells) -1 ) was measured as described in Materials and methods. Line was drawn by linear regression, r = 0.996.

pa thways which can oxidize glucose up to CO 2 and H 2 0 , like the Krebs cycle, are operating. The later is absent in either N R B C or parasites [14].

The amounts o f 14CO2 released increased with increas ing parasi temia (Fig. 1) as expected if HM S activity is essent ial ly due to parasite operation. Al- ready at the r ing stage the H M S activity was 7.2-fold higher compared to NRBC, whereas at the trophozoite stage the activity was 77.8-fold more in tensive (Table 1). A cont inuous moni to r ing of the HM S activity of infected cells in t ightly synchronized cultures, was performed starting f rom the r ing stage (14 h post invasion). The HMS activity increased gradu- ally with parasite deve lopment (Fig. 2). The peak of this activity occurred at 30 h post invasion, i.e, at the trophozoite stage. Thereafter, when the parasite ma- tured to the schizont and segmenter stages, the activity of HMS decreased. Al though the absolute activity differed somehow from one culture to the other, the s tage-dependent pattern remained identical (compare the two traces of Fig. 2).

In order tO verify whether the increased HMS activity of IRBC is due to the paras i te ' s or the host ce l l ' s activity, H M S was measured in parasites which were freed from their host cells by the Sendai virus- induced lysis which does not impair parasite viabi l-

Table 1 HMS activity in different types of cells and the effect of methylene blue

Cell type Control + Methylene Activation due to activity blue methylene blue

NRBC 0.054 + 0.002 1.55 + 0.9 1.496 (n = 3) (n = 4)

Rings 0.39 + 0.04 0.9 + 0.03 0.51 (n = 3) (n = 2)

TRBC 4.2 + 0.5 6.89 + 1.5 2.69 (n = 12) (n = 7)

Free 3.4 + 0.5 5.7 + 1.2 2.3 parasites (n = 7) (n = 3)

Cells were preincubated with methylene blue for 30 min prior to the addition of D-[1-14C]glucose. n is the number of independent experiments using different cultures. NRBC, uninfected RBC; TRBC, trophozoite-infected RBC. * HMS activity: glucose consumed in /~mol h-1 (1010 cells)-1.

ity, at least for several hours [27]. Free parasites display considerable HM S activity (Table 1) that amounts to about 81% of the total activity seen in intact t rophozoi te-IRBC (TRBC). The difference between the total HM S activity in T R B C and that of the isolated parasite is 0.8 /xmol h -1 (1010 cells) -1.

W h e n this activity is normal ized to the relative vol- um e of the host compar tment (60% of the total T R B C volume) and subtracted from the activity

1.2

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0.2

0 . 0 . . . . . . . . . . . . . . . . . . . . . . . .

0 10 20 30 40 50

TIME ( h r s )

Fig. 2. HMS activity as a function of parasite development. Parasite cultures were synchronized to a time window of 4 h. When they reached 25% parasitemia, they were taken for the measurement of HMS activity. HMS activity (in /~mol glucose consumed h -1 (101° cells) -1) is shown as a function of time post-invasion. Results of two independent cultures are shown.


measured in NRBC, we get that the specific HMS activity of the host is increased 24.7-fold (compare this value to the 28.7-fold stimulation by methylene blue of HMS activity in NRBC, see below). The HMS activity in uninfected RBC isolated from the culture by gelatin flotation was 0.138 ___ 0.03 /zmol h -1 (101° cells) -1, i.e, 2.56-fold higher than in NRBC. This result suggests that some of the oxidative stress generated by IRBC is transferred to the uninfected RBC.

Effect o f methylene blue on H M S activity. The maximal capacity of HMS activity can be estimated by exposing the cells to methylene blue which oxidizes NADPH to NADP +. A 29-fold activation of HMS in NRBC was measured, resulting in an increment of 1.5 /xmol h -1 (1010 cells) -1 (Table 2). A higher increment was seen in TRBC, amounting to 2.7 /xmol h -1 (101° cells) -1. Methylene blue-induced increase in HMS activity has also been observed in P~ berghei-infected mouse RBC [28]. An increment of 2.3 /xmol h -1 (1010 cells) -1 was observed in free parasites. These results imply a) that parasites also contain NADPH-diaphorase activity, and b) that in untreated IRBC both the HMS of the host cell and that of the parasite are not maximally activated since they can be further enhanced with methylene blue.

We have also measured the kinetics of methylene blue reduction in normal RBC, in IRBC and in free parasites. As shown in Fig. 3, the depletion of methylene blue from the medium by normal RBC is rapid and reaches a steady-state. The extent of deple-

tion observed with IRBC and in the presence of free parasites are much larger.

Effect o f endogenously produced oxidative stress on H M S activity. During the digestion of ingested host cell cytosol inside the acidic food vacuole of the parasite, oxidative species are produced by the oxidation and degradation of hemoglobin in a pH- and proteolysis-dependent manner [10,29]. The protease inhibitors leupeptin and pepstatin, and the food vacuole alkalinizing agent NHaC1, are known to inhibit the digestion process [30,31] and the generation of oxidative stress [10]. To verify to what extent the endogenous generation of oxidative stress activates the HMS activity, cells were preincubated for 30 min with the protease inhibitors or with NH4CI, and then their HMS activity was measured in the presence of these agents. Results are depicted in Table 2. The HMS activity of NRBC is slightly but not signifi- cantly increased in presence of these agents. That of ring IRBC displays a similar rise. This result is expected because ring-stage parasites do not digest the host cell cytosol, and hence, there is no endogenous production of oxidative stress, as we have reported previously. Such was also the case of parasites freed from their host cell, here again because ingestion and digestion do not occur. Protease inhibitors caused a 20% reduction of HMS activity of IRBC harboring trophozoites, whereas NHaC1 caused a reduction of 55%. To assure that the inhibitory effect was not due to some non-specific effect of the antiprotease or of NHaC1, methylene blue was added

Table 2 Inhibition of HMS activity by antiproteases and NH4C1 , and its reversal by methylene blue

Cell type Control * + NH4C1 + Antiprotease MB + AP NH 4C1 + MB

NRBC 0.054 -t- 0.002 0.063 + 0.01 0.074 4- 0.02 1.75 _+ 0.2 1.52 4- 0.1 n = 3 n = 3

Ring 0.39 + 0.04 0.4 + 0.005 0.48 + 0.1 N.D. N.D. n = 2 n = 3

TRBC 4.24 4-0.3 1.9 4- 0.7 3.34 4- 0.2 5.9 4- 0.2 4.45 4- 0.2 n = 7 n = 6

Free parasites 3.4 4- 0.5 3.1 4- 0.3 3.2 4- 0.3 N.D. N.D. n = 3 n = 3

Cells were preincubated with the inhibitors for 30 min prior to the addition of D-[1-14 C]glucose. n is the number of independent experiments using different cultures. AP, antiproteases; MB, methylene blue; NRBC, uninfected RBC; TRBC, trophozoite-infected RBC. N.D.: not determined. * HMS activity: glucose consumed in p~mol h-1 (1010 cells)-1.


in presence of these agents, and HMS activity was assayed again. The results depicted in Table 2, indi- cate that methylene blue induced in these cells an increase in HMS activity that is identical to that observed in IRBC in absence of antiproteases or NH4CI.

Other modifiers of endogenous reactive oxygen species (ROS) production were also tested. As shown in Table 3, uric and benzoic acids, reduced the HMS activity in TRBC. Since deoxy-D-ribose, a demon- strable scavenger of OH radicals, had no similar effect, it is suggested that the effects of benzoic and uric acids was due to scavenging of O 2 and/or H202. Quinoline-containing antimalarials were recently shown to inhibit the spontaneous denaturation of hemoglobin and the release of iron from thus produced heme [32]. Since it has been shown that this process constitutes the biochemical mechanism of endogenous ROS production in TRBC [10], the effect of these drugs on HMS activity was tested. Therapeutically relevant concentrations of the anti-

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Fig. 3. Reduction of methylene blue by normal and infected RBC, and by free parasites. Normal ( 0 ) and trophozoite-infected ( A ) RBC, and free parasites ( • ) were incubated in presence of 0.5 mM methylene blue. Aliquots of the supematant were taken with time, and the content of reduced methylene blue was determined spectrophotometrically. Results are depicted as methylene blue reduced (in ~m01 (10 l° cells) -1) against time. Lines were drawn according to first order kinetics obtained by non-linear least square regression. The parameters of the regressions (Limit + SD ( ~ m o l (101° cells)-1), k + S D (min -1) and the initial rate = Limit * k ( ~mol min- 1 (1010 cells)- l ) were: NRBC, 2.85 + 0.24, 0.74_+0.32 and 2.10; TRBC, 21.09_+2.5, 0.18_+0.1 and 3.86; free parasites, 19.4 + 0.62, 0.154-+ 0.02 and 2.98.

Table 3 Effect of oxidative radical scavengers and antimalarial drugs on the HMS activity of different cell types

Treatment TRBC Free NRBC parasites

Chloroquine (5 /zM) 24.16 + 8.5 5.95 + 3.7 3.6 + 2.7 (n = 5) ( n = 4) ( n = 3)

Quinine (5/xM) 33 + 10.9 8 _+ 3 0 (n = 5) (n = 4) (n = 3)

Mefloquine (1 /xM) 37.8+16.5 14.8+4.9 1 .4+2 (n = 5) (n = 4) (n = 3)

Uric acid (4 mM) 23.5 _+ 15 8.5 + 12 0 (n = 3) (n = 3) (n = 3)

Benzoic acid (4 mM) 39.2+4.1 9 .6+7.2 0 (n = 3) (n = 3) (n = 3)

Cells were preincubated with the inhibitors for 30 min prior to the addition of D-[1-14C]glucose. Results are expressed as percent inhibition, n is the number of independent experiments using different cultures. NRBC, uninfected RBC; TRBC, trophozoite-infected RBC.

malarials chloroquine, quinine and mefloquine, caused substantial inhibition of HMS activity (Table 3). However, since some inhibition was also observed in non-infected RBC, it is not unlikely that these drugs may also have some ROS scavenging effect.

The role of glutamate dehydrogenase in NADP + reduction. We have verified whether the parasite's glutamate dehydrogenase could play a role in main- taining the reduced NADPH pool in the IRBC and the isolated parasite, by comparing HMS activity in these cells in absence or presence of glutamine or glutamate ethyl ester. The latter was used because glutamate uptake is rather slow, whereas the ester readily enters the cell and is hydrolysed to glutamate be resident hydrolases. We could not observe any effect of either glutamine or glutamate ethyl ester addition on HMS activity (data not shown).

The role of HMS in nucleic acid synthesis. One of the major roles of HMS is to provide ribose-5-phosphate (R-5-P) for the synthesis of nucleic acids. If R-5-P is produced exclusively by the oxidative arm of the HMS, one should not expect to find labeled nucleic acids when cells are fed with D-[1-14C]glucose. The labelling of nucleic acids from this substrate implies that R-5-P is produced by the non- oxidative activity of the HMS. Measuring the label


incorporated in nucleic acids from D-[1-14C]glucose or from D-[6-14C]glucose, showed equal incorporation (in /xmoles glucose equivalents h -1 (101° cells) -1, mean + SD, n--- 3): 0.151 ___ 0.02 and 0.1.58 + 0.02 in TRBC, and 0.183 _ 0.04 and 0.188 _ 0.04, in free parasites, respectively. The difference observed between TRBC and free parasites could be a result of glucose utilization by the host cell, or from faster uptake of glucose into free parasites than into TRBC.

The importance of HMS activity for the synthesis of nucleic acids can be tested from different angles. Since hypoxanthine is the major substrate for the synthesis of purine nucleotides, its omission from the assay system should decrease the activity of HMS if it is important for nucleotide synthesis. Only after 3 h of incubation could a reduction of HMS activity by 16% be observed in absence of hypoxanthine. This result suggests that there may be a substantial endogenous pool of purines that through the salvage pathways of host and parasite [33] could supply the necessary purine bases. This presumption is supported by the observation that in presence of the RNA synthesis inhibitor actinomycin D (6.6 /zg ml-1; [34]) the HMS activity in TRBC is inhibited by 38% (n = 3). Since RNA is overwhelmingly the major oligonucleotide synthesized by the parasite, accumulation of nucleotides in absence of RNA synthesis and the inhibition of DNA synthesis by actinomycin D, could in principle feed-back inhibit NADPH-consuming steps in pyrimidine and DNA syntheses, although one can not exclude other indirect and/or non specific effects.

4. Discussion

A long controversy has beset the substantiation of G6PD activity in malaria parasites, to the point of suggesting that the parasite depends on the host cell for the shunt pathway [14]. There was no question that the gene encoding the enzyme is present in the parasite's genome [35,36], but based on the retarded growth of P. falciparum in G6PD-deficient erythrocytes, it has been suggested that it is expressed only when the host cell enzyme is absent [37,38]. This allusion was supported by the demonstration that extracellular NADPH maintains the reduced state of

glutathione in isolated P. berghei [39]. It seems, however, that the many conflicting results, have now been reconciled: G6PD has been identified and partially characterized in P. falciparum [16] and in P. berghei [15]. It is rather surprising that in view of this controversy, only very few attempts have been made to measure HMS activity directly in intact IRBC or in free parasites.

In the present work we were able to show that the HMS activity of trophozoite IRBC is 78-fold more intense than in NRBC. A 51-fold increase, and absolute values of HMS activity in IRBC (4.4 ~mol (ml)- packed cells) h - l ) , i.e., similar to those measured in this work, were reported previously [19]. Measuring the HMS of the isolated parasite, revealed that it is much more active than that of the host cell, and contributes some 82% of the total activity of the intact IRBC. HMS activity of G6PD-deficient IRBC was 72% of that measured in normal IRBC [19], implying that most of the activity in the former cells can be attributed to the parasite, since HMS can not be activated in G6PD-deficient RBC. Furthermore, we can exclude the possibility that the parasite's activity may have been contributed by residues of the host cell cytosol entrapped within the host cell membrane which spontaneously reseals after the Sendai virus-induced lysis. Measuring the residual hemoglobin as a probe for other cytosolic compo- nents, revealed less than 5% of the original content. To account for the 78-fold higher HMS activity found in the isolated parasites, one would have to assume that the residual rate-limiting enzyme (G6PD) works 1560 times faster, a rather improbable as- sumption. We must therefore conclude that our mea- surements genuinely reflect the HMS activity of the parasite.

TRBC and isolated parasites are much more effec- tive than normal RBC in accumulating exogenous methylene blue. This observation is congruous with the increased HMS activity of TRBC and the relative contribution of the parasite to the total HMS activity of the infected cell. The ratio of steady-state levels of accumulated methylene blue (TRBC/NRBC) are almost equal to the ratio of HMS activity in these cells. Methylene blue is able to diffuse into RBC where it is reduced by NADPH-dependent diaphorase to leukomethylene. Since the latter can not diffuse across membranes, it concentrates intracellu-


larly as long as there is an adequate supply of glucose to fuel the HMS [40]. Indeed, we found that preaccumulated methylene blue diffuses out of the cells when glucose is removed from the incubation medium (data not shown). These results suggest that either the levels of NADPH in the IRBC or in free parasites are much larger than in the non infected red cells, or that the capacity to reduce NAPD + back to NADPH is greater, or both. Since free parasites display almost the same accumulative capacity as IRBC, we conclude that the capacity of the latter cells to reduce and accumulate methylene blue mostly reflects parasite activity.

The boosted HMS activity of TRBC could also explain the increased stability of GSH in infected cells [19] and their increased ability to reduce extracellular ferricyanide [41] compared to non-infected RBC. Hence, although intraerythrocytic parasites can be killed by extracellular oxidants, including those generated by macrophages [9], this is not an indica- tion of reduced antioxidant defence (compared to normal RBC), but rather of the presence of oxidant- sensitive targets in the parasite.

Comparison of the partial contributions of host cell and parasite to this activity, indicates that the host cell's activity is also considerably activated compared to that measured in normal RBC. Such activation most probably results from the oxidative stress that the parasite inflicts on its host cell. We have previously shown that H20 2 generated by the parasite reaches the host cell compartment and is reduced by catalase. The activation of the host's HMS indicates that the host's glutathione peroxidase is also involved in the reduction of H20 2, since this enzyme utilizes NADPH as a reducing cofactor. The activities of both catalase and glutathione peroxidase imply that the host cell provides some protection for the parasite against oxidative offense, by reducing H20 2 and thus increasing the gradient along which this oxidant diffuses out of the parasite.

The HMS activity of IRBC increases with parasite maturation. This rise is timed to the increased de- mand for NADPH. NADPH is required for DNA synthesis (as a cofactor for ribonucleotide reductase) which peaks during the trophozoite stage [42]. It is also probably maximally consumed by the antioxidant defence at the trophozoite stage which is most active in host cell cytosol digestion [43], and in the

endogenous production of oxidative stress [10]. It is therefore not surprising that HMS activity declines at the schizont stage when both processes virtually come to a halt. The increase in HMS activity with parasite maturation could be a simple consequence of the increased parasite biomass, but it is tempting to suggest that the synthesis of HMS enzymes is up-regulated, say by the endogenous oxidative stress, because the increase in HMS activity is much larger than the increase in biomass.

To further assess that the 24-fold activation of host cell HMS activity is due to the oxidative stress generated by the proliferating parasite, IRBC were preincubated in presence of protease inhibitors and with NHaC1 which demonstrably raises the pH of the parasite's food vacuole. The use of these agents is warranted by the recent demonstration that the biochemical origin of ROS generation is the proteolytic oxidation of host cell hemoglobin inside the acidic environment of the food vacuole [10]. Protease inhibitors and NHaC1, which inhibit this process, caused 20% and 44% inhibition at the HMS activity, respectively. That this was not due to some non- specific impairment of HMS activity, is manifested by the fact that HMS can be activated in these cells by methylene blue to the same extent as non-treated cells. These results mean that 0.84-1.8 /xmol glucose h -1 (101° cells) -1 entered the oxidative arm of the HMS to counteract the oxidative stress. These inhibitors had no effect on HMS activity at the ring or the schizont stages or in isolated parasites, when no hemoglobin digestion occurs, and hence, much less ROS are produced. Scavengers of oxidative radicals, such as uric and benzoic acids, also reduced HMS activity in TRBC (where hemoglobin digestion occurs) but much less so in free parasites that have no hemoglobin to digest.

Furthermore, therapeutic concentrations of quinoline-containing antimalarials also reduced HMS activity in TRBC. These drugs were recently shown to inhibit the proteolytic degradation of hemoglobin in IRBC [43] and in acidified red cell lysates and the release of iron from heme thus produced [32]. Other ROS generating processes may be present in the parasite, such as the one electron reduction of 0 2 by the electron transport chain of the mitochondrion [44], but the electron transport inhibitor antimycin A had no effect on HMS activity (data not shown),


indicating that this process contributes only mini- mally to ROS production, if at all. These observa- tions imply that the endogenously produced oxidative stress is a major reason for activation of HMS. We. have no way to determine whether these treat- ments affected the HMS activation in the parasite, in the host or in both. The latter is the most plausible in view of the rapid translocation of ROS in the dimen- sions of the infected cell. However, it should be emphasized that none of the inhibitors tested could completely obliterate HMS activity, either because they were not used at the maximal possible concentration (to minimize non-specific or indirect effects) or because the consumption of NADPH for anabolic processes such as DNA and tetrahydrofolic acid syntheses, presents an additional driving force for HMS activation.

Although it has been contended that parasite glutamate dehydrogenase could be instrumental in the reduction of NADP + [22], we could not observe any reduction of HMS activity with either glutamine or glutamate ethyl ester addition on HMS activity. This result raises a question mark about the significance of the glutamate dehydrogenase as potential sources for NADPH. It is important to note that glutamate dehydrogenase can work also in reverse, i.e., synthe- sizing glutamate from a-ketoglutarate and NH ~. Al- though the Km for the latter is in the millimolar range (the Vmax is much larger in the reverse reaction), the consumption of glutamate in protein synthesis can drive the reaction in the synthetic direc- tion, thus constituting an NADPH consuming process, and linking HMS directly with protein synthesis.

Despite the substantial flux of glucose through the parasite's HMS, no difference could be observed between label incorporated into nucleic acids if the substrate was D-[1-14C]glucose or D-[6-14C]glucose. We interpret this ostensible enigma as follows: the total amount of glucose consumed by TRBC is ca 27-fold larger than that engaged in HMS. The latter is some 27-fold larger than that incorporated into nucleic acids (Table 4). Since the reversible reactions of the transaldolase, transketolase, xylulose-5-P epimerase and ribose-5-P isomerase (the non-oxidative reactions of HMS) are much faster than the rate-limiting forward reactions of G6PD and 6PGD [4,5], the enormous surplus of glyceraldehyde-3-P

Table 4 Glucose consumption by glycolysis and HMS in normal and in trophozoite-infected RBC, and by synthesis of nucleic acids in infected RBC Cell type Glucose consumed a by

Oxidative Glyco- A/B * 100 Nucleotide HMS (A) lysis

NRBC 0.054 1.53 3.53 N.A. TRBC 4.2 110 3.82 0.155 Relative 27 710 1 consumption

a Units are: p.mol glucose consumed h-1 (1010 cells)-1. b Results taken from Pfaller et al. [53]. N.A.: Not applicable.

and fructose-6-P produced by glycolysis (compared to those generated by the HMS) could recycle into the HMS and provide for all the necessary R-5-P needed for the synthesis of nucleic acids. Indeed, it has been recently demonstrated that considerable recycling of D-[1-14C]glucose or D-[6-14C]glucose takes place in the HMS even in the presence of net flow of glucose through the oxidative arm of HMS [45].

Both in normal RBC and in TRBC a similar relative amount of glucose is utilized by the HMS. HMS activity in normal RBC is determined by the activity of G6PD which is controlled by the NADP+/NADPH ratio, whereas when they are ox- idatively stressed, hexokinase activity may become rate-limiting [4,5]. The ability to inhibit HMS in TRBC by reducing the oxidative stress, in conjunc- tion with the fact that most of this activity is contributed by the parasite, may suggest that it is the NADP+/NADPH ratio which controls HMS activity in the parasite. Hence, it is very likely that the major function of the oxidative arm of the HMS is to keep a high level of NADPH for the protection of the parasite from the oxidative stress generated during host cytosol digestion by keeping the reduced form of glutathione (through the action of glutathione reductase). NADPH is also needed for the activity of the rate limiting enzyme in DNA synthesis, diribonu- cleotide reductase, for the proper function of catalase [46], for the synthesis of pyrimidines, and maybe for other purposes which still need to be investigated.


Acknowledgements

This investigation received financial support from the UNDP/World Bank/WHO Special Programme for Research and Training in Tropical Diseases (TDR).

References

[1] Eaton, J.W. and Brewer, G.J. (1974) Pentose phosphate metabolism, In: The Red Blood Cell (Surgenor, MacN., ed., Academic Press, New York, Vol. I, pp. 435-471.

[2] Stern, A. (1985) Red cell oxidative damage. In: Oxidative Stress, (Sies, H., ed.), Academic Press, London, pp. 331-349.

[3] Jacob, H.S. and Jandl, J.H. (1966) Effects of sulphydryl inhibition on red blood cells. II. Glutathione in the regulation of the hexose monophosphate pathway. J. Biol. Chem. 241, 4243-4250.

[4] Thorbum, D.R. and Kuchei, P.W. (1985) Regulation of human erythrocyte hexose-monophosphate shunt under conditions of oxidative stress. Eur. J. Biochem. 150, 371-386.

[5] Schuster, R., Holzhutter, H.-G. and Jacobash, G. (1988) Interrelations between glycolysis and the hexose monophosphate shunt in erythrocytes as studied on the basis of a mathematical model. BioSystems 22, 19-36.

[6] Friedman, M.J. (1979) Oxidant damage mediates variant red cell resistance to malaria. Nature 280, 245-247.

[7] Vennerstrom, J.L. and Eaton, J.W. (1988) Oxidants, oxidant drugs, and malaria. J. Med. Chem. 31, 1269-1277.

[8] Golenser, J. and Chevion, M. (1989) Oxidant stress and malaria: Host-parasite interrelationships in normal and abnor- mal erythrocytes. Semin. Hematol. 26, 313-325.

[9] Hunt, N.H. and Stocker, R. (1990) Oxidative stress and the redox status of malaria-infected erythrocytes. Blood Cells 16, 499-526.

[10] Atamna, H. and Ginsburg, H. (1993) Origin of reactive oxygen species in erythrocytes infected with Plasmodium falciparum. Mol. Biochem. Parasitol. 61,231-242.

[11] Seth, R.K., Sahni, A.S. and Jaswal, T.S. (1985) Plasmodium berghei: oxidant defense system. Exp, Parasitol. 60, 414-416.

[12] Fairfield, A.S., Abosch, A., Ranz, A., Eaton, J.W. and Meshnick, S.R. (1988) Oxidant defense enzymes of Plas- modium falciparum. Mol. Biochem. Parasitol. 30, 77-82.

[13] Zhang, Y., Konig, I. and Schirmer, R.H. (1988) Gluathione reductase-deficient erythrocytes as host cells of malarial parasites. Biochem. Pharmacol. 37, 861-865.

[14] Scheibel, L.W. (1988) in Malaria: Principles and Practices of Malariology (Wernsdorfer, W.H. and McGregor, I., eds.), pp. 199-212, Churchill Livingston, Edinburgh.

[15] Buckwitz, D., Jacobash, G., Kuckelkom, U., Plonka, A. and Gerth, C. (1990) Glucose-6-phosphate dehydrogenase from Plasmodium berghei: kinetic and electrophoretic characterization. Exp. Parasitol. 70, 264-275,

[16] Kurdi-Haidar, B. and Luzzatto, L. (1990) Expression and

characterization of glucose-6-phosphate dehydrogenase of Plasmodium falciparum. Mol. Biochem. Parasitol. 41, 83-92.

[17] Neame, K.D. and Homewood, C.A. (1975) Alterations in the permeability of mouse erythrocytes infected with the malaria parasite Plasmodium berghei. Int. J. Parasitol. 5, 537-540.

[18] Shakespeare, P.G., Trigg, P.I., Kyd, S.I. and Tappenden, L. (1979) Glucose metabolism in the simian malaria parasite Plasmodium knowlesi: activities of the glycolytic and pentose phosphate pathways during the intraerythrocytic. Ann. Trop. Med. Parasitol. 73, 407-415.

[19] Roth, E.F., Raventos-Suarez, C., Perkins, M. and Nagel, R.L. (1982) Glutathione stability and oxidative stress in P. falciparum infection in vitro: responses of normal and G6PD deficient cells. Biochem. Biophys. Res. Commun. 109, 355- 362.

[20] Sherman, I.W., Peterson, I., Tanigoshi, L. and Ting, I.P. (1971) The glutamate dehydrogenase of Plasmodium lophu- rae (avian malaria). Exp. Parasitol. 29, 433-439.

[21] Walter, R.D., Nordmeyer, J.P. and Konigk, E. (1974) NADP-specific glutamate dehydrogenase from Plasmodium chabaudi. Hoppe-Seyler's Z. Physiol. Chem. 355, 495-500.

[22] Vander Jagt, D.L., Hunsaker, L.A., Kibrige, M. and Campos, N.M. (1989) NADPH production by the malarial parasite Plasmodium falciparum. Blood 74, 471-474.

[23] Sahni, S.K., Saxena, N., Puri, S.K., Dutta, G.P. and Pandey, V.C. (1992) NADP-specific isocitrate dehydrogenase from the simian malaria parasite Plasmodium knowlesi: partial purification and characterization. J. Protozool. 39, 338-342.

[24] Roth, E.F., Ruprecht, R.M., Schulman, S., Vanderberg, J. and Olson, J.A. (1986b) Ribose metabolism and nucleic acid synthesis in normal and glucose-6-phosphate dehydrogenase-deficient human crythrocytes infected with Plasmod- ium falciparum. J. Clin. Invest. 77, 1129-1135.

[25] Jensen, J.B. (1978) Concentration from continuous culture of erythrocytes infected with trophozoites and schizonts of Plasmodium falciparum. Am. J. Trop. Med. Hyg. 27, 1274- 1276.

[26] Pescarmona, G.P., Bosia, A., Arese, F., Sartori, M.L. and Ohigo, D. (1982) A simplified method for the pentose phosphate pathway assay in red cells. Int. J. Biochem. 14, 243- 245.

[27] Kanaani, J. and Ginsburg, H. (1989) Metabolic interconnec- tion between the human malarial parasite Plasmodium falciparum and its host erythrocyte: regulation of ATP levels by means of an adenylate translocator and adenylate kinase. J. Biol. Chem. 264, 3194-3199.

[28] Deslauriers, R., Butler, K. and Smith, I.C.P. (1987) Oxidant stress in malaria as probed by stable nitroxide radicals in erythrocytes infected with Plasmodium berghei. The effects of primaquine and chloroquine. Biochim. Biophys. Acta 931, 267-275.

[29] Gabay, T, and Ginsburg, H. (1993) Hemoglobin denaturation and iron release in acidified red blood cell lysate: a possible source of iron for intraerythrocytic malaria parasites. Exp. Parasitol. 77, 261-272.

[30] Rosenthal, P.J., McKerrow, J.H., Rasnick, D. and Leech, J.H. (1989) Plasmodium falciparum: inhibitors of lysosomal


cysteine proteinases inhibit a trophozoite proteinase and block parasite development. Mol. Biochem. Parasitol. 35, 177-184.

[31] Zhang, Y. (1987) Inhibition of hemoglobin degradation in Plasmodium falciparum by chloroquine and ammonium chlo- ride. Exp. Parasitol. 64, 322-327.

[32] Gabay, T., Krugliak, M., Shalmiev, G. and Ginsburg, H. (1993) Inhibition by antimalarial drugs of hemoglobin denaturation and iron release in acidified red blood cell lysates: a possible mechanism of their antimalarial effect? Parasitology. 108, 371-381.

[33] Gero, A.M. and O'Sullivan, W.J. (1990) Purines and pyrimidines in malarial parasites. Blood Cells 16, 467-484.

[34] Geary, T.G., Divo, A.A. and Jensen, J.B. (1989) Stage specific actions of antimalarial drugs on Plasmodium falciparum in culture. Am. J. Trop. Med. Hyg. 40, 240-244.

[35] Luzzatto, L., O'Brien, S., Usanga, E. and Wanachiwanawin, W. (1986) Origin of G6PD polymorphism: malaria and G6PD deficiency. In: Glucose-6-phosphate dehydrogenase (Yoshida, A. and Beutler, E., eds.), pp. 181-193, Acedemic Press, Orlando.

[36] Yoshida, A. and Roth, E.F. (1987) Glucose-6-phosphate dehydrogenase (G6PD) of malaria parasite (Plasmodium falciparum). Blood 69, 1528-1531.

[37] Usanga, E.A. and Luzzatto, L. (1985) Adaptation of Plas- modium falciparum to glucose-6-phosphate dehydrogenase- deficient host red cells by production of parasite-encoded enzyme. Nature 313, 197-204.

[38] Roth, E.F. Jr., Raventos-Suarez, C., Rinaldi, A. and Nagel, R.L. (1983) Glucose-6-phosphate dehydrogenase deficiency inhibits in vitro growth of Plasmodium falciparum. Proc. Natl. Acad. Sci. USA 80, 298-302.

[39] Eckman, J.R. and Eaton, J.W. (1979) Dependence of plas- modial glutathione metabolism on the host cell. Nature 278, 754-756.

[40] Sass, M.D., Caruso, C.J. and Axelrod, D.R. (1967) Accumu- lation of methylene blue by metabolizing erythrocytes. J. Lab. Clin. Med. 69, 447-455.

[41] Simoes, A.P.C.F., Vandenberg, J.J.M., Roelofsen, B. and Op den Kamp, J.A.F. (1992) Lipid peroxidation in Plasmodium falciparum-parasitized human erythrocytes. Arch. Biochem. Biophys. 298, 651-657.

[42] Inselburg, J. and Banyal, H.S. (1984) Synthesis of DNA during the asexual cycle of Plasmodium falciparum in culture. Mol. Biochem. Parasitol. 10, 79-87.

[43] Zarchin, S., Krugliak, M. and Ginsburg, H. (1986) Host cell digestion by intraerythrocytic malarial parasites is the pri- mary target for quinoline-containing antimalarials. Biochem. Pharmacol. 35, 2435-2442.

[44] Fry, M. and Beesley, J.E. (1991) Mitochondria of mam- malian Plasmodium spp. Parasitology. 102, 17-26,

[45] Schrader, M.C., Simplaceanu, V. and Ho, C. (1993) Mea- surement of Fluxes through the pentose phosphate pathway in erythrocytes from individuals with sickle cell anemia by carbon-13 nuclear magnetic resonance spectroscopy. Biochim. Biophys. Acta 1182, 179-188.

[46] Kirkman, H.N., Galiano, S. and Gaetani, G.F. (1987) The function of catalase-bound NADPH.J. Biol. Chem. 262, 660-666.

[47] Pfaller, M.A., Krogstad, D.J. and Parquette, A.R. (1982) Plasmodium falciparum: Stage-specific lactate production in synchronized cultures. Exp. Parasitol. 54, 391-396.

Documents

Hexose-monophosphate shunt activity in intact Plasmodium falciparum-infected erythrocytes and in free parasites