biochemistry of plasmodium (malarial parasites)cytes by p. vivax (142, 147). indeed, the...

MICROBIOLoGICAL REVIEWs, Dec. 1979, p. 453-495 Vol. 43, No.401460749/79/04-0453/43$02.00/0

Biochemistry of Plasmodium (Malarial Parasites)IRWIN W. SHERMAN

Department ofBiology, University of California, Riverside, California 92521

INTRODUCTION AND LIFE CYCLE .......................................... 453BIOCHEMICAL DETERMINANTS OF PARASITE SPECIFICITY FOR HOST

CEILS...................... 454MORPHOLOGY AND GROWTH OF THE BLOOD STAGES. 456MORPHOLOGICAL ALIERATIONS OF INFECTED CELS. 456MEMBRANE STRUCTURE AND FUNCTION IN MALARIA .................. . 457Membrane Proteins of Infected Erythrocytes ................................ 457Plasmodial Membrane Proteins ............................................. 458Lipids of Malaria-Infected Erythrocytes .....459................ ...... .. 459Plasmodial Membrane lipids ............................................... 460ETrABOLIC PATHWAYS ................................... 462Carbohydrate Transport and Metabolism .................................. 462

Glucose uptake ........... ................ ............. 462Products of glucose catabolism............. 463Enzymes of carbohydrate metabolism ................... ......... 464

(i) Glycolytic enzymes .......................................... 464(ii) Citric acid cycle enzymes ........................................... 465(iii) COrfuiing enzymes ............................... 465(iv) Pentose phosphate pathway enzymes .............................. 466

Oxygen utilization and electron transport .......... 467Nucleic Acids.469

Deoxyribonucleic acid: characteristics and synthesis.469Ribonucleic acid: characteristics and synthesis.470Pyrimidine biosynthesis.471Purine transport and salvage pathway mechanisms.472

Protein Synthesis.475Biosynthesis of amino acids.475Uptake of free amino acids.475Proteolysis of hemoglobin and the formation of malarial pigment. 476Mechanisms Qf protein synthesis.477Histidine-rich protein.477

Lipid Biosynthesis.478Vitamins and Cofactors ................... 479Vitamin A.479Vitamin B1 (thiamine).479Vitamin B2 (riboflavin).479Biotin.479Nicotinic acid (niacin), nicotinamide, and pyridine nucleotides.479Ascorbic acid .......................................... 480Vitamin E .................................................. 480Vitamin B, group ......................................................... 480Pantothenate .................... ........................................ 480Vitamin K and ubiquinones.480Folates ..................................................... 481

(i) Biosynthesis of dihydrofolate... 481(ii) Dihydrofolate reductase (tetrahydrofolate dehydrogenase).481(iii) Tetrahydrofolate metabolism ...................................... 482

CationAlterations.483Precautions, Prospects, and Possible Conclusions ........................... 483

LITERATURE CITED ............................................4...........486

INTRODUCTION AND LIFE CYCLE tebrate host begins with the bite of a mosquitoand the injection of threadlike sporozoites. The

The disease malaria is caused by protozoan introduced sporozoites are carried by the bloodparasites belonging to the genus Plasmodium. to various organs and tissues of the body; theAll species require two hosts, a vertebrate and parenchymal cells of the liver (in mammals) oran invertebrate. Commonly, infection of the ver- the endothelial cells (in birds) are invaded by

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these sporozoites, where they divide asexually toform daughter progeny called merozoites. Ulti-mately, the merozoites are released from thetissues to enter the circulation, where they in-vade the erythrocytes. Within an erythrocyteeach parasite grows and divides asexually, pro-ducing a substantial number of merozoites. Fi-nally, the erythrocyte lyses, merozoites are re-leased, and reinvasion of erythrocytes can takeplace. The synchronous asexual developmentand rupturing of erythrocytes by the parasite ismarked by periodic fever-chill cycles which arethe hallmark of malarial infections. Some mer-ozoites continue to reinvade erythrocytes andmultiply asexually, but others enter erythrocytesand differentiate into sexual stages called ga-metocytes. When a mosquito feeds on a verte-brate host and ingests these gametocytes, a sex-ual cycle is initiated. In the mosquito stomachthe gametocytes are transformed into gametes,fertilization takes place, and the resultant worm-like zygotes penetrate the stomach wall andcome to lie on the outer surface of the stomach.Here each zygote forms a cystlike body, theoocyst, and within it thousands of sporozoitesare produced by asexual multiplication. Uponsporozoite maturation the oocyst bursts, andsporozoites are released, which in time enter thesalivary glands. When the mosquito next takesa blood meal, sporozoites are injected and thelife cycle is completed.The microscopic size and inaccessibility of the

tissue forms of malaria have limited our knowl-edge about the biochemistry of these stages;similarly, the minuteness of the mosquito stageshas also prevented biochemists from studyingthese forms. Consequently, most of the infor-mation on the malaria-host relationship is de-rived from studies of parasites growing within orremoved from erythrocytes since such stages areboth accessible and abundant. Therefore, of ne-cessity, our knowledge of the biochemistry ofPlasmodium is limited almost exclusively to theinteractions of malarial parasites with one hostcell type, the erythrocyte.BIOCHEMICAL DETERMINANTS OF

PARASITE SPECIFICITY FORHOST CELLS

The invasive stage of malaria (the merozoite)measures about 1 Lm across, is surrounded by acomplex of membranes (pellicle), and containsa nucleus, mitochondria, endoplasmic reticulum,a cytostome, and apical (polar) organelles calledrhoptries and micronemes (1, 2, 8). The lengthof the asexual cycle for merozoite production,the number of merozoites produced, and thekinds of hosts that may be infected are charac-

teristic for each species of Plasmodium. Suchspecificity does not depend on a particular spe-cies of mosquito since it is possible to initiate amalaria infection in a susceptible host simply bytransferring infected blood from one host toanother.McGhee began a series of experiments in the

1950s to determine why different host speciesvaried in susceptibility to a species of malaria(135, 136). To avoid complications by the hostimmune response, McGhee used chicken em-bryos to study the invasion process. The avianmalaria Plasmodium lophurae, a highly virulentand synchronously developing parasite, was se-lected for study. Washed uninfected ducklingerythrocytes were introduced into the circula-tion of a P. lophurae-infected chicken embryo;periodically blood samples were taken, and thedegree of infection was determined. By this tech-nique McGhee could easily calculate the per-centage of introduced cells that had been in-vaded by the P. lophurae merozoites. He foundthat although the number of chick cells wasgreater than the number of introduced ducklingcells, the rate of infection of the latter was con-siderably higher. Clearly, the P. lophurae mer-ozoites recognized different kinds of cells andpreferentially invaded a particular host cell type.What permitted such merozoite discrimination?Some years later studies of merozoite penetra-

tion were undertaken, in which an in vitro cellsuspension culture system was used. Normalduckling erythrocytes were treated with a vari-ety of enzymes to remove possible erythrocytereceptors for the parasite. These cells werewashed and introduced into a culture vessel witha small inoculum of P. lophurae-infected cells.It was reasoned that if receptors were removedby the treatment, then invasion of these cellswould be blocked, and the total parasite countsin these culture vessels would be less than thosefound in untreated samples. None of the treat-ments, however, affected parasite penetration(210). Subsequently, Butcher et al. and Miller etal. (reviewed in reference 142) showed that thein vivo infectivity of the monkey malaria P.knowlesi for a variety of hosts was directly re-lated to the degree of in vitro penetration of theerythrocytes of these species. Thus, resistantspecies, such as rats and chickens, were notinvaded in vitro, and the reduced virulence of P.knowlesi merozoites for human erythrocyteswas reflected by an in vitro rate of infection thatwas one-quarter that of rhesus monkey eryth-rocytes, a host in which the infection is ordinar-ily fatal.

Miller and his colleagues (143, 145, 146) at-tempted to clarify the nature of the erythrocyte

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receptor for P. knowlesi. Human erythrocyteswere treated with a variety of enzymes and thensubjected to invasion by P. knowlesi merozoitesin vitro. Chymotrypsin and pronase blocked in-vasion, whereas trypsin and neuraminidase werewithout effect. The results suggested that thereceptor for P. knowlesi on human erythrocyteswas a glycoprotein. In addition, chymotryptictreatment of rhesus monkey erythrocytes, thepreferred host cell, did not block parasite inva-sion. These authors suggested that the monkeyerythrocyte receptor may be a glycolipid or thereceptor may be resistant to enzyme treatment,and consequently a strong block against parasiteinvasion was not produced. Perhaps a similarexplanation can be offered for the failure toblock the P. Iophurae invasion of duck eryth-rocytes by pretreatment with enzymes (210).What does the receptor do? Light and electron

microscopic studies have shown that the inva-sion of an erythrocyte by a malarial merozoitefollows a definite sequence: merozoite attach-ment, widespread deformation of the erythro-cyte, junction formation between the merozoiteand erythrocyte, movement of the junction, in-vagination of the erythrocyte membrane to in-ternalize the merozoite, and resealing of theerythrocyte membrane after the parasite hasbeen completely surrounded by the vacuolarmembrane (5, 14, 15, 60, 122).

Initial studies by Miller et al. (132, 142)showed that Duffy blood group-negative humanerythrocytes (Fy Fy) were refractory to invasionby P. knowlesi merozoites. Duffy-positive eryth-rocytes have the determinants Fya and Fiy aswell as a third determinant, Fy, which is presenton both Fya and Fyb cells. Duffy-negative eryth-rocytes lack all three determinants. If Duffy-positive erythrocytes are treated with chymo-trypsin, the Fy8 and Fyb determinants are re-moved, but Fy3 remains. Such cells resist inva-sion. Miller has suggested that there may be twoP. knowlesi receptors on human erythrocytes.One is Duffy unrelated and chymotrypsin sen-sitive, whereas the other (Fy39) is involved injunction formation. If P. knowlesi merozoiteswere treated with cytochalasin B, attachmentdid occur with Duffy-positive cells, but no inva-sion took place; although cytochalasin B-treatedmerozoites did attach to Duffy-negative cells, nojunction was formed, and therefore parasite en-try did not occur (145). The Duffy-associateddeterminants apparently are not involved in theinitial attachment of the merozoites, but mayplay a role in junction formation.

Miller postulated that the Duffy factor mightbe involved in the invasion of human erythro-cytes by P. vivax (142, 147). Indeed, the inci-

BIOCHEMISTRY OF PLASMODIUM 455

dence of Duffy-negative cells is extremely highin West Africa, where resistance to P. vivax isalso high. To test this postulate, 11 Americanblacks who had previously been exposed to P.vivax-infected mosquitos were typed. Five Duffynegatives were resistant, whereas six Duffy pos-itives became infected. A subsequent study in-volving American blacks infected in Vietnamwith P. vivax malaria revealed that all wereDuffy positive. Thus, there is ample evidence tosupport the hypothesis that the Duffy-negativephenotype is a basis for P. vivax resistance (147).The nature of the Duffy blood group receptor

has not been characterized chemically. More-over, we do not know how it specifically interactswith the merozoites of P. vivax and P. knowlesi.What we do suspect, based on susceptibilitystudies of various erythrocytes, is that the recep-tors for the two species are similar but notidentical.

Since West Africans who are refractory to P.vivax malaria are susceptible to other humanmalarias, Miller suspected that these other ma-laria species have different erythrocyte recep-tors. The recent advance in the cultivation of P.falciparum in human blood cells (260) has madea direct test of this possible. Human erythro-cytes lacking Duffy and other blood group de-terminants were all invaded by P. falciparummerozoites. Trypsin treatment of erythrocytesreduced their susceptibility to P. falciparum,but not P. knowlesi merozoites; by contrast,chymotrypsin blocked P. knowlesi invasion butdid not influence susceptibility to P. falciparum(143). Thus, the current view is that the P.knowlesi (and probably the P. vivax) and P.falciparum receptors on the surfaces of humanerythrocytes are different.

Miller speculates that for infection to occurtwo steps must take place: the merozoite mustreact with a specific peptide or a carbohydratemoiety, which serves for attachment and con-tributes to cell specificity, and then there mustbe a reaction with another peptide, which trig-gers endocytosis (142). Once the receptors canbe characterized chemically, we may better un-derstand their role in the invasion process. Thatthe host cells cooperate in the invasion processwas demonstrated by studies in which erythro-cyte ghosts were used. Although P. knowlesimerozoites attach to susceptible human eryth-rocyte ghosts, they rapidly detach from these,and parasite invasion does not occur (142). Thestudies of erythrocyte receptors for malaria sug-gest that the merozoites contain ligand-like sub-stances on their surfaces, which are complemen-tary to the specific receptors on the erythrocytesurface. Such ligands must be exceedingly labile


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since merozoites do not attach to susceptibleerythrocytes 15 min after they have been re-

leased from an erythrocyte.The widespread deformation of the erythro-

cyte, radiating from the point at which the mer-

ozoite contacts it, suggests that the merozoitereleases a material which acts on the host cellmembrane. This substance is probably releasedfrom organelles at the apical end of the mero-

zoite. Such membrane-active substances couldalso induce endocytosis (see below).The end result of merozoite invasion is that

each parasite is bounded by two very closelyapposed membranes, an outer parasitophorousvacuolar membrane (PVM) and the parasiteplasma membrane. Both membranes are re-

tained through the entire intraerythrocytic de-velopmental cycle, and if parasites are removedfrom the cell by a potent hemolytic antiserum(prepared in rabbits by injection of washed nor-

mal erythrocytes), they are still enveloped byboth of these membranes (261). Furthermore,studies by Langreth and Trager have shown thatduring extracellular cultivation of P. lophuraeboth membranes remain (123).The nature of the PVM has received consid-

erable attention, primarily through the use ofelectron microscopic techniques (15, 120, 139,140, 200, 241, 242). Although its origins are eryth-rocytic, it appears to come under the control ofthe growing parasite. This is not unexpected; thePVM increases in size as the volume of theparasite increases, and consequently it musteither stretch or be added to. Also, althoughsmall parasites showed a localization of adeno-sine triphosphatase in the PVM that was typicalof an invaginated erythrocyte membrane, as the,parasite grew there was a reorientation of aden-osine triphosphatase in the membrane. Simi-larly, reduced nicotinamide adenine dinucleotide(NAD) oxidase, an enzyme localized on the outersurface of the erythrocyte, was not found on theinner surface of the PVM (as it would be if itwere simply an invaginated erythrocyte mem-

brane), but it was found on the outer surface ofthe PVM (120). Also, studies with cationizedferritin (241) and with freeze-fracture (139, 140)techniques show that the PVM is dynamicallyaltered by the parasite. How such alterations areinduced and by what mechanisms they occur

remain unknown.

MORPHOLOGY AND GROWTH OF THEBLOOD STAGES

Upon invasion of an erythrocyte, a merozoitededifferentiates, losing its rhoptries, micro-nemes, and pellicular membranes (1, 8, 47). Theresult is a uninucleate, amoeboid trophozoite

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which lies within the PVM and begins to ingesthost cell cytoplasm via a specialized organelle,the cytostome. Vacuoles are pinched off fromthe base of the cytostome, and within thesedigestion of hemoglobin and formation of he-mozoin (malaria pigment) take place. The tro-phozoite cytoplasm contains limited amounts ofendoplasmic reticulum, numerous ribosomes,mitochondria (cristate or in the form of concen-tric membranes), Golgi bodies, primary lyso-somes, and in some species specialized mem-brane-bound vesicles. The trophozoite feeds andgrows in size, and its nucleus enlarges. In time,the trophozoite transforms into a schizont,which undergoes nuclear division (schizogony)to yield merozoites. Schizogony involves the for-mation of intranuclear mitotic spindles, divisionof the nucleus, laying down of pellicular struc-tures, development ofrhoptrymicronemes at theperiphery, and incorporation of nuclei and mi-tochondria into the divided cytoplasm. Thoseorganelles excluded from the merozoite, such asthe pigment-containing vacuoles, membranousvesicles, and endoplasmic reticulum remnants,form a structure called the residual body. Withinthe confines of the PVM, the merozoites acquirean amorphous surface coat (1, 8, 121, 144). Mer-ozoites are released, and the remnant of the hostcell contains the residual body and the PVM.The exact mechanism whereby merozoite re-lease is accomplished is not understood.

MORPHOLOGICAL ALTERATIONS OFINFECTED CELLS

When certain species of malaria parasites in-vade erythrocytes, they alter the host cells mor-phologically. In these instances the presence ofthe parasite leaves a telltale surface scar, onewhich serves to characterize the particular kindof malaria. Erythrocytes infected with Plasmo-dium ovale- and P. vivax-type malarias, afterstaining with Giemsa stain, show a fine stipplingpattern that is easily seen with a light micro-scope. The dots in this pattern are called Schiiff-ner's dots. When these parasitized erythrocytesare examined by electron microscopy, they showmany small invaginations, which are called cav-eolae; the base of a caveola is flattened, andwithin its lumen is a fuzzy material (4, 6). Cav-eolae continuous with the erythrocyte mem-brane are surrounded by small vesicles. Thenumber of invaginations tends to depend on thesize of the parasite; cells with larger parasitesusually have more caveolae. There is now strongevidence to show that the caveolae are Schiuff-ner's dots (2, 6). Both appear at the same time,and they are of similar size; the caveolae bindhorseradish peroxidase-labeled antibody from


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monkeys chronically infected with P. vivax, andthey accumulate fine electron-dense granulesafter staining with Giemsa stain. Two functionshave been proposed for the caveola-vesicle com-plexes; one suggestion is that these vesicles areengaged in micropinocytosis, and the other isthat membrane-lined vesicles, called clefts, formin the erythrocyte stroma and that these fusewith the surface to promote expansion of thecell surface, thus contributing to the enlarge-ment ofthe P. ovale-P. vivax-infected cell. Bothviews may be correct.

Cytoplasmic clefts surrounded by a single unitmembrane have been observed in erythrocytesinfected with parasites other than P. ovale- andP. vivax-type malarias, including P. cynomolgi,P. coatneyi, P. fakciparum, P. gonderi, and P.lophurae (2, 261). Trager et al. (262) suggestedthat in the case of P. falciparum, these cleftsmay represent the coarse stippling patternswhich are seen by light microscopy after Giemsastaining and are known as Maurer's clefts. Theclefts are believed to be of parasitic origin. Thesignificance of clefts in malaria species otherthan the P. vivax and P. ovale types remainsunclear, however.

In addition to clefts and caveola-vesicle com-plexes, some malaria species induce electron-dense elevations or excrescences on the surfacesof the infected erythrocytes (7, 8, 121, 262).These knobs, as they are commonly called, areparticularly evident in the P. falciparum- andP. malariae-type malarias. The knobs appear tobe underneath the erythrocyte membrane introphozoites. The number of erythrocytic knobstends to increase as the parasite grows, and thereare indications that some P. fakciparum strainsare better knob inducers than are others andthat knobs are lost during prolonged in vitroculture (S. Langreth, personal communication).As a parasite grows, the knobs obscure the eryth-rocyte membrane, bizarre extensions may form,and some of these appear to be sloughed fromthe surface of the erythrocyte. The knobs on P.falciparum- and P. coatneyi-infected erythro-cytes form focal junctions with the membranesof endothelial cells, suggesting that they play arole in the sequestration of infected cells whereasexual multiplication takes place in the deeporgans (7, 8). Knobs are found with both asexualand sexual parasites in the P. maiariae-typeparasites, but in the P. fakciparum-type malariasonly erythrocytes bearing asexual parasites showknobs. Since both asexual and sexual P. ma-lariae-type parasites circulate in the peripheralblood, the knobs probably are unrelated to deepvascular sequestration in these species. How-ever, during the acute stage of infection of mon-


keys with P. brasilianum (a P. malariae-typemalaria), these infected cells become seques-tered, which leads one to suspect that in somespecies the knobs may be involved in the imn-mune response.The knobs on the surfaces of erythrocytes

infected with P. falciparum-type malarias con-tain parasite materials. Ferritin-labeled anti-body prepared against parasites (109) or anti-body derived from immune monkeys (Langreth,personal communication) reacts with the surfacemembrane where the protuberances are present;no binding is evident where knobs are absent.How such parasite materials become incorpo-rated into the knobs has still not been deter-mined.Very recently, freeze-fracture techniques have

been applied to studies of the erythrocytic mem-branes of normal and P. knowlesi-infected mon-key erythrocytes (139, 140). The number of in-tramembranous particles in infected erythro-cytes bearing small parasites was not signifi-cantly different from the number in uninfectederythrocytes; however, in schizont-infectederythrocytes there was both a reduction and aclumping of the remaining intramembranousparticles.

Clearly, the erythrocyte is not an inert mem-branous vessel harboring the parasite from theimmune mechanisms of the host, nor is it a staticmembrane system that serves merely to envelopthe growing and reproducing plasmodium. In-stead, sometimes in obvious morphological fea-tures, and perhaps more frequently in subtlemolecular ways, the parasite so influences thehost cell that exploitation of host resources be-comes possible.

MEMBRANE STRUCTURE ANDFUNCTION IN MALARIA

Membrane Proteins of InfectedErythrocytes

In the rodent malarias P. berghei and P. cha-baudi Weidekamm et aL (282) and Konigk andMirtsch (115) demonstrated a decrease in thestaining of spectrin (bands 1 and 2), band 4, andband PAS-1; concomitant with such reductions,the intensities of bands 7, 8, and PAS-2 in-creased, and some new bands appeared (bands2a and PASJ). The unique band PASi with amolecular weight of -165,000 has also been re-ported for P. berghei, P. yoelii YM, P. vinckei,and P. chabaudi (288). However, in some relatedstrains of P. yoelii (17x and 33x) a distinct banddid not appear, but instead there were severalweak bands in the molecular weight range of120,000 to 200,000. Weidekamm et al. (282) sug-


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gested that the decreased amounts of bands 1and 2 could be due to degradation by proteolysis,and this would contribute to band 2a and theincreased intensities of bands 7 and 8. Addition-ally, the breakdown of band 4 could also inten-sify bands 7 and 8. Yuthavong et al. (288) testedthis hypothesis by incubating normal mouse

erythrocyte membranes with lyophilized prepa-

rations of P. berghei, and although there was

degradation of spectrin, no new bands of~150,000 molecular weight developed. Such re-

sults could be explained by the action of plas-modial enzymes on the inner surface of theerythrocytes in vivo and the proteolysis of boththe inner and outer surfaces in vitro.

Eisen (62) did not find a decrease in the totalamount of spectrin in P. chabaudi-infectederythrocytes and could not detect glycophorinin such cells. Such an observation correlates wellwith the report of Trigg et al. (265), who showeda decrease in the labeling (by galactose oxidase-tritiated borohydride after pretreatment of cellswith neuraminidase) of band 3 in P. knowlesi-infected monkey erythrocytes. Wallach andConley (270) found a decrease in band PAS-1and an iodinatable component of band 5 (50,000daltons) in P. knowlesi-infected cells, new Coo-massie brilliant blue-staining bands (120,000 to200,000 daltons), and a parasite-specific bandPAS-lp (125,000 daltons); band 2 decreased,whereas band 4 increased. Recently, Schmidt-Ullrich and co-workers (196, 197) reported thatat least three proteins (-55,000 to 90,000 dal-tons) occur in the membranes of erythrocytesbearing P. knowlesi, but these proteins were notfound in parasites or normal erythrocytes. Bycontrast, Deans et al. (55) found relatively fewP. knowlesi antigens in infected erythrocytemembranes, but all of these were present in theintracellular parasites.There is one caveat that should be mentioned:

in most of the reported studies of membraneproteins of rodent and simian malaria-infectederythrocytes, there has not been a separation ofhost cell from parasite membranes nor has therebeen an attempt to control protease activity.

Modifications in the membranes of duckerythrocytes during infection with P. lophuraehave been studied by Sherman and co-workers(219, 241, 242). Plasma membranes from theerythrocytes of malaria-infected ducklings wereobtained by nitrogen cavitation; sodium tetra-thionate was used as a protease inhibitor. Theresultant membrane vesicles were purified by asucrose step gradient, and the membranes werecharacterized by the following techniques: (i)electron microscopy, (ii) polyacrylamide gelelectrophoresis, (iii) sialic acid content and (iv)

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iodinatability. Normal duckling erythrocytemembranes contained 79 nmol of sialic acid permg of membrane protein, whereas that derivedfrom infected cells had approximately one-halfthis amount (219). Furthermore, the iodinata-bility of infected erythrocyte membranes wasalso reduced. Finally, when the Coomassie bril-liant blue-stained gel patterns of infected eryth-rocyte membranes from duckling erythrocytesbearing small parasites (day 4 of infection) orlarger parasites (day 5 of infection) were exam-ined, reductions in bands 1, 2, and 3 and inten-sifications of bands 4 through 7 were observed.No new bands were found.What do these studies of host cell membranes

indicate? Changes in band intensity, reductionin iodinatability, and reduction in sialic acidcontent could be due to plasmodial enzymes aswell as to changes in the lipid milieu of themembrane. Thus, by the action of plasnodialproteases and phospholipases liberated intracel-lularly, some of the inner surface proteins, suchas spectrin and actin, as well as proteins thatspan the entire erythrocyte membrane, may becleaved, modifying these proteins and alteringthe cytoskeletal framework. Such alterationscould also serve to modify the accessibility ofouter surface proteins to reagents commonlyemployed for identiffying the external surface ofcells. Also, in some malarias, such as P. knowlesi,P. falciparum and several species of rodent ma-laria, parasite-specific proteins may be intro-duced in addition to the degradation of existingerythrocyte membrane proteins. These so-calledneoproteins could modify the antigenicity of theinfected erythrocytes and could affect the func-tion of the erythrocytes. How such proteinsmove to the erythrocyte surface and how theyare intercalated remain unknown.

Plasmodial Membrane ProteinsP. lophurae, which is easily maintained in the

laboratory by intravenous inoculation, growssynchronously in ducklings. By using such asystem, it is possible to obtain parasites at twostages of development: trophozoites (day 4 ofinfection) and schizonts (day 5 of infection).Furthermore, parasites can be removed from theerythrocytes by treating infected erythrocyteswith a hemolytic antiserum prepared in rabbits,followed by trypsin and deoxyribonuclease treat-ments. Such free parasites are surrounded by aPVM (261). Since the PVM is developmentallyrelated to the erythrocyte membrane, its char-acteristics have been compared with those of thehost cell membrane. The surface of the eryth-rocyte bears negative charges, mostly due to thepresence of sialic acid residues. By using posi-


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tively charged cationic ferritin, which can bevisualized by electron microscopy, the erythro-cyte membranes bound the stain at 0.18 mg/ml,but the PVM and parasite plasma membranesdid not. However, at a very high concentrationof stain, the three kinds of membranes could beclearly distinguished from one another (241). Aferritin-labeled antibody prepared against nor-mal duckling erythrocytes did not react with thePVM or parasite plasma membranes.

Lectins are proteins that show considerablespecificity for cell surface carbohydrates; conse-quently, they can be used as highly sensitiveprobes for the characterization of cell membranesurfaces. Free parasites were not agglutinatedby lectins, nor was there evidence for the bindingof ferritin-conjugated lectins to the surface ofthe PVM or parasite plasma membranes. Anexception to this was the weak and occasionalbinding of concanavalin A (242). Seed and Kreier(202) reported a stage-independent binding ofconcanavalin A-ferritin to erythrocyte-free P.berghei, and Bannister et al. (15) were unable toagglutinate merozoites with phytohemaggluti-nin, wheat-germ agglutinin, ricin, and concana-valin A, but occasionally some concanavalin A-ferritin binding was observed. Such results showthat the PVM, although derived from the eryth-rocytes by endocytosis, is changed. This is cor-roborated by the cytochemical and freeze-frac-ture observations of Langreth (120) and Mc-Laren et al. (139, 140) referred to above.Membrane vesicles can be produced easily by

Dounce homogenization ofP. Iophurae liberatedfrom host cells by the hemolytic antiserum tech-nique. In a linear sucrose gradient two bandswere present, a heavy one with a density of 1.158g/cm and a lighter one with a density of 1.110g/cm3. The heterodispersed heavy band had he-mozoin granules. When such membranes wereanalyzed for sialic acid, they contained -8 nmolof sialic acid per mg of protein, about one-tenththe amount of the erythrocyte cell membranes.(Reduced amounts of sialic acid were also re-ported for P. berghei [200].) The heavy and lightmembrane fractions, when analyzed by sodiumdodecyl sulfate-polyacrylamide gel electropho-resis, were qualitatively similar to one another,but were distinctly different from the membranefractions of the host cells. Plasmodial mem-branes separated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis could not bestained with periodic acid-Schiff stain and con-tained no spectrin bands (219). Wallach andConley (270) reported similar findings for eryth-rocyte-free P. knowlesi. The P. lophurae mem-branes were also deficient in their ability toincorporate radioactive iodine when the lacto-


peroxidase-glucose oxidase method was used(219).

lipids of Malaria-Infected ErythrocytesAnalyses of the lipid composition of malaria-

infected erythrocytes are complicated by thefact that the experimental material consists of amulticompartmental membrane system, whichincludes the plasma membranes of the host andparasite, the PVM, the nuclear membrane of theplasmodium (and in avians that of the host cellnucleus as well), and, depending on the species,the membranes of various organelles.

Generally speaking, the total lipid content ofmalaria-infected erythrocytes, as well as theamounts of lipids in the various fractions, ishigher than the lipid content found in normalerythrocytes. In P. knowlesi-infected erythro-cytes the total lipid content is three to five timesgreater than that in uninfected erythrocytes (10,13, 87, 137); similar increases, but of smallermagnitude, have been reported for P. berghei-infected rat erythrocytes (87, 124, 178) and P.lophurae-infected duckling cells (18) (Table 1).Lawrence and Cenedella (124) calculated that"free" P. berghei might contain as much as 5times more lipid than rat reticulocytes, but thevalue could also be as low as 1.5 times. Othersreport that free plasmodia of P. knowlesi and P.lophurae (10, 18) contained an amount of lipidabout equal to that in uninfected erythrocytes.Indeed, although Rao et al. (178) stated that P.berghei contributed -6% to the total lipid con-tent of infected rat reticulocytes, examination oftheir data (Table 1) indicates that the contri-bution was greater than 60%. Most of these lipidincreases in the infected cells can be attributedto the plasmodial membrane phospholipids. Al-though the total cholesterol content of an in-fected erythrocyte tends to increase, the ratio ofcholesterol to phospholipid decreases becausethe parasite lipids contain proportionally morephospholipid (Table 1), and the infected eryth-rocyte membranes tend to lose cholesterol (201).(How this occurs is not understood.)The major phospholipids of uninfected eryth-

rocytes are phosphatidylethanolamine andphosphatidylcholine, with lesser amounts ofsphingomyelin and phosphatidylinositol (Table2). Upon infection, phosphatidylethanolamineand phosphatidylinositol increase, and sphingo-myelin decreases.The most common fatty acids ofnormal eryth-

rocytes are palmitic (16:0), stearic (18:0), oleic(18:1), linoleic (18:2), and arachidonic (20:4);upon infection, striking increases are observedin the saturated fatty acids palmitic acid and


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TABLE 1. Lipid contents ofnormal and infected erythrocytes and malaria parasitesa

Cell type Total lipid content Phospholipid content Cholesterol content Ratio of phospholipidto cholesterol

Monkey erythrocytesb 575 283 2.15P. knowlesi-infected 625 136 2.6

erythrocytesbP. knowlesib 750 136 6.3

Monkey erythrocytesc 3.5 2.1 1.1 2P. knowlesi-infected 17.0 13.2 2.4 5.5

erythrocytescP. knowlesic 4-5 2.5 0.7 3.5

Duck erythrocytesd 650 481 130 3.6P. lophurae-infected 1,320 1,082 158 6.0

erythrocytesdP. lophurae 86% 8% 10.7

Rat erythrocytesc 5.3 4 1.26 3P. berghei-infected 8.06-10.6 6.5 2.11 3

erythrocytescP. bergheic 8-28

a Data are from references 18, 56, 87, 178, and 187. Different values for the same thing are from differentstudies.

b Expressed as micrograms per millgram of protein.'Expressed as miligrams per 1010 cels.d Expressed as miligrams per 100 ml of cells.

oleic acid (Table 3). Linoleic acid and arachi-donic acid decline in both avian and mammalianmalarias; stearic acid declines in P. knowlesi(74) and P. berghei (18) malarias, but increasesin P. lophurae-infected cells (18, 88). The en-hanced amount of 18:1 fatty acid in P. lophurae-infected erythrocytes cannot be attributed solelyto the presence of the plasmodium, but it isprobably associated with the erythrocyte mem-brane itself. (This is derived from the fact thatthe alkoxy form of phosphatidylethanolamine isnot present in the lipids of P. lophurae but doesoccur in substantial amounts in erythrocyteswhere the content of 18:1 fatty acids was in-creased.)The increased amounts of 18:1 fatty acids in

the erythrocytes of infected animals could influ-ence the properties of the infected erythrocytemembrane by altering the fluidity of the innerand outer leaflets of the phospholipid bilayer; aconsequence of this could be conformationalchanges in membrane proteins, with an effect ontransport and other membrane functions. Byimposing restraints on the movement of thephospholipid hydrocarbon chains, cholesteroltends to stabilize membrane fluidity. A loss ofcholesterol with a reduction in the membranecholesterol/phospholipid ratio of an erythrocyte(Table 1) could be reflected in reduced activetransport and increased osmotic fragility andpassive permeability of the cell, which are allcharacteristics of malaria-infected erythrocytes.

Plasmodial Membrane Lipids

Relatively few studies of the lipids of malarialparasites can be considered free of error becauseof the presence of undetermined amounts ofcontaminating erythrocytic membranes. Indeed,even in those reports describing the lipid com-position of truly free parasites, the isolated par-asites remain enclosed within the PVM, itself aderivative of the erythrocyte membrane. (How-ever, the composition of the PVM may not sub-stantially confound the analytic results sincethere is now evidence to show that shortly afterformation the PVM assumes plasmodial char-acteristics [see above].)These limitations aside, the membranes of

malaria parasites have been found to be richerin unesterified fatty acids, triacylglycerols, po-lyglycerol phosphatides, 1,2-diacylglycerols, dia-cylphosphatidylethanolamine, and phosphati-dylinositol and to contain less cholesterol, phos-phatidylserine, and sphingomyelin than themembranes of erythrocytes (Table 2). Theunique qualities of plasmodial membranes canbe determined from several bits of evidence:phosphatidylinositol, a significant phospholipidof P. lophurae and P. knowlesi (-4 to 8%)membranes, was not present to a significantdegree in erythrocytic membranes, and phos-phatidylserine, a component of mammalianmembranes, was virtually absent from plasmo-dial membranes. Also, in P. Iophurae mem-

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TABLE 2. Percent distribution ofmajorphospholipid classes in normal and malaria-

infected erythrocytes andplasmodiaa% Distributionb

Cell typePC PS SM PI PE

Rat erythrocytes 45 9.6 16.9 1.7 28Rat erythrocytesc 58 17 25P. berghei-infected 34 12 9.7 3.4 44

erythrocytesP. berghei-infected 58 9.0 32

erythrocytesd

Monkey erythrocytes 36 11 15 3 31Monkey erytbrocytes 37.1 9 18 1.2 31.1P. knowlesi-infected 44.7 1.7 3.2 8.1 40.9

erythrocytesP. knowlesi-infected 42.3 9.7 16.1 1.8 26.5

erythrocytesP. knowksi 45 2 3 8 41P. knowlesi 45 2 2 4 39

Duck erythrocytes 40 <1 1 1 20P. lophurae-infected 36 <1 3 3 33

erlythrocytesP. lophurae 40 <1 4 4 36

a Data are from references 18, 56, 87, 133, 178, and 186.Different values for the same thing are from different studies.

bAbbreviations: PC, phosphatidylcholine; PS, phosphati-dylserine; SM, sphingomyelin; PI, phosphatidylinositol; PE,phosphatidylethanolamine.

'In this study, the phosphatidylcholine plus phosphatidyl-serine content for rat erythrocytes was 58%.

d In this study, the phosphatidy1choline plus phosphatidyl-serine content for P. berghei-infected erythrocytes was 58%.

branes less of the phosphatidylethanolaminewas in the alkyl-1-enyl and alkyl-acyl forms com-pared with erythrocyte membranes (4 and 3%versus 35 and 9%, respectively), whereas forphosphatidylinositol the reverse was true (10and 15% versus 7 and 4%, respectively).Although the principal fatty acids of the

erythrocyte and Plasmodium phospholipidswere similar, the proportion of 18:1 fatty acids(oleic, cis-vaccenic) and saturated fatty acids(principally 16:0 [palmitic acid] and 18:0 [stearicacid]) was greater in the plasmodial membranes,whereas the proportion of polyunsaturates (18:2[linoleic acid], 20:4 [arachidonic acid], and 22:6[docosahexaenoic acid]) was less (Table 3).According to Beach et al. (18), the bias for

18:1 fatty acids and the relative paucity ofpolyunsaturates in the malarial parasite phos-pholipids suggest that the plasmodia are defi-cient in desaturases and chain elongation en-zyme systems. Since the parasites do differ fromtheir host cells in the proportions of neutral andpolar lipids, in the content of 1,2-diacylglycerolsin the neutral lipids, and in the content of thediphosphatidylglycerol and alkoxy lipids in thephospholipids, as well as in the amount of fattyacids in all lipids, it is apparent that despite theinability of the parasites to synthesize lipids


TABLE 3. Percent distribution ofprincipal fattyacids in the total lipids ofnormal and malaria-

infected erythrocytes and parasitesa% Distribution of the following fatty acidab:

Cell type16:0 18:0 18:1 18:2 20:4 22:6

Monkey 22-39 14-18 13-18 10-15 4-17 Trace-2erythro-cytes

P. knowlesi- 37 14 21 14 2infectederythro-cytes

P. knowlesi 33 9-14 35 15 2 Trace

Rat erythro- 24 17 8 11 31cytes

P. berghei 42 15 21 7 5P. vinckei 40 13 17 8 0.8

Duck erythro- 24 10 18 22 10 7cytes

P. lophurae- 28 14 28 12 4 3infectederythro-cytes

P. Iophurae 26 16 33 12 3 3a Data are from references 18, 17, 87, and 187.b 16:0, Palmitic acid; 18:0, stearic acid; 18:1, oleic and cia-

vaccenic acids; 18:2, linoleic acid; 20:4, arachidonic acid; 22:6,docoshexaenoic acid.

from acetate (see below), they can regulate theiruse of host cell lipids and lipid precursors.How does membrane biosynthesis by intracel-

lular plasmodia take place, and how is the asym-metry of the PVM changed? Transmembranetumbling (ffip-flop) of phosphatidylethanola-mine and cholesterol does not occur, and phos-phatidylcholine migration is very slow from oneleaflet of the lipid bilayer to the other. There-fore, it seems plausible to assume that duringintraerythrocytic feeding by the parasites muchof the PVM is recycled and that this mechanismprovides an opportunity for intercalation of newparasite-derived membrane materials into thePVM. The result is a topographic organizationof the PVM lipid bilayer similar to that of theerythrocyte and plasmodial plasma membranes.The lipid composition of the parasite membraneis regulated by the lipids and lipid precursors ofthe erythrocytes and the plasma, as well as bythe limited metabolic capabilities ofthe parasite.Exchanges of phospholipids, lysophospholipids,and cholesterol among the blood plasma, theerythrocytes, the PVM, and the parasite mem-branes obviously take place during intraerythro-cytic growth of the plasmodia. In addition, thelipids of the endocytosed cytoplasm and mem-branes of a host cell, as well as the plasma fattyacids, phosphate, inositol, choline, glycerol, andethanolamine, can be utilized for parasite mem-brane synthesis. Since it is incapable of de novo


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fatty acid biosynthesis, the parasite acts as ametabolic sink for host fatty acids, particularlyoleic acid. As a consequence, there is a netmovement of this fatty acid from the plasmainto the infected erythrocytes, and membranelipid enrichment in octadecenoic fatty acids re-sults.There are no data on the transmembrane dis-

tribution of phospholipids in the membranessurrounding the parasites; such informationcould be of significance in understanding para-site membrane function.

METABOLIC PATHWAYSCarbohydrate Transport and MetabolismGlucose uptake. The beginning biochemical

investigations with malarial parasites involvedstudies of carbohydrates, largely because thiswas an early focal point of the biochemistry ofyeasts, muscle, and liver. Today some of thesepioneering efforts may be criticized on severalgrounds: the techniques used were insensitiveand imprecise, contaminants such as platelets,leukocytes, and cellular debris were not removedor evaluated, incubation times were prolonged,the media were frequently unphysiological, im-mature erythrocytes were present, and substratelevels were often unlike those in vivo. Yet de-spite these limitations, it seems clear that ingeneral malarial parasites use mechanisms for

the breakdown of glucose which are very similarto those used by a variety of other eucaryoticcells.The intraerythrocytic stages of malaria do not

store glycogen or other reserve polysaccharides(43, 53, 90, 192); consequently, simple sugars

(notably glucose) are essential for the continuedgrowth and reproduction of the plasmodia. Thisfact was appreciated as early as 1912, when Bassand Johns (17) reported that suspensions of P.falciparum- and P. vivax-infected erythrocytesrequired the addition of glucose for parasitemaintenance in vitro; of the variety of othersugars they tested, only maltose was an adequatesubstitute for glucose. Johns (102) also foundthat by adding glucose to P. vivax-infected cellsthe survival of the parasite was prolonged at0°C. Also, Hegner and MacDougall (84) ob-served that P. gallinaceum infections in chick-ens were more severe if the animals receivedintravenous injections of glucose. Clearly, glu-cose was needed by the malarial parasites.

Systematic biochemical studies of carbohy-drate utilization in a variety of malaria specieswere undertaken during the 1930s and 1940s. Itwas found very early on that, although normalerythrocytes tended to use very little of theadded metabolite and consumed only smallamounts of oxygen in vitro, glucose and severalother sugars (Table 4), as well as glycerol, dis-appeared rapidly when added to suspensions of

TABLE 4. Substrates utilized and products formed by PlasmodiumInfected cells Free parasites

Compound P. lo- P. galli- P. P. know- p P- lo- P. galli- P. P. know-phurae naceum berghei lesi (137, arum (17) phurae naceum berghei lesi (138,(222)a (137) (89,90) 192, 193) arum (24) (152) (26,159) 173)

Substrates utilizedGlucose ++++ ++++ +++++ ++++ ++++ +++ +++ +++ ++++Mannose ++++ +Fructose ++++Maltose + ++++ +Glycerol ++++ ++++ ++++Pyruvate ++++ ++++ +++ +++ + +Oxaloacetate ++cis-Aconitatea-KetoglutarateSuccinate + + + + +?Fumarate + +Malate + +CitrateLactate ++++ ++++ +++ +++

Products formedPyruvate + + + +Acetate + + +Lactate + +++ ++++ +++ +++ ++ + +++ +Succinate + +C02 ++ +++ +Aspartic acid + + + + + +Glutamic acid + + + + + +Alanine + + + + + +

a The numbers in parentheses are the references from which the data were taken.

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infected erythrocytes, and concomitantly oxygenuptake increased (89, 90, 95, 137, 138). Theamount of substrate utilized was found to de-pend on a complex set of variables, includingcharacteristics of the substrate itself, incubationmedium, species of malarial parasite, size of theintracellular plasmodium, degree of parasitiza-tion, pH, and the kinds and amounts of contam-inants. Yet despite such overwhelming uncer-tainties, it became clear that the parasite-pre-ferred substrate for energy was glucose and thatparasitized erythrocytes consumed up to 75times more sugar than did their uninfected coun-terparts (89, 90, 137, 226).The dramatic increase in glucose consumption

by erythrocytes infected with a malarial parasitecan only be sustained by an alteration in thepermeability barrier of the erythrocyte mem-branes since some bird (chicken and duckling)and rodent (mouse) erythrocytes demonstrate alow sugar uptake. Indeed, this suggestion, firstproposed by Hermnan et al. (85), was confirmedby the studies of Sherman and Tanigoshi (226)and Neame and Homewood (93, 159). In theformer case, the non-metabolizable sugar 3-0-methylglucose was used to separate the eventsof transport and catabolism from one another,the permeability of P. lophurae-infected eryth-rocytes was markedly enhanced for 3-0-meth-ylglucose, and the accelerated entry was due toan increase in the simple diffusion component,as well as a modification-in the carrier-mediatedportion of the entry process. Furthernore, theleakiness toward 3-0-methylglucose was great-est for those erythrocytes bearing larger para-sites. Also, although nornal mouse erythrocyteswere imperneable to L-glucose (also non-metab-olizable), P. berghei-infected cells readily tookit up (159). Of some interest was the finding thatL-glucose entered P. berghei-parasitized eryth-rocytes, but not uninfected cells taken from aninfected animal.Thus, not only does a malaria-infected eryth-

rocyte consume more glucose than a normalerythrocyte, but the presence of the parasite"encourages" the entry of sugar by changing thepermeability characteristics of the host cell.How this altered permeability occurs requiresfurther clarification.Products of glucose catabolism. The end

products of glucose metabolism vary with thespecies of Plasmodium. In general, mammalianmalarias incompletely oxidize glucose, yieldingorganic acids, whereas in bird malarias a signif-icantly higher degree of oxidation occurs, withthe production of CO2 as well as organic acids(Table 4).An early report claimed that under aerobic


conditions P. knowlesi-infected erythrocytesproduced substantial amounts of lactic acid andpyruvic acid; anaerobiosis prevented pyruvicacid formation, but aerobically infected cells ox-idized some of the lactic acid (137, 283). How-ever, Scheibel and Pfiaum (193) seriously ques-tioned this. Using free P. knowlesi, they foundthat little added radioactive glucose was con-verted to "'CO2 and that all of the glucose couldbe accounted for as lactate or volatile com-pounds, notably acetate and formate. Glucosewas incorporated into keto acids to a minordegree (<2%) both aerobically and anaerobi-cally, whereas acid volatiles (acetate and for-mate) accounted for 22 to 53% and neutral vol-atiles represented 10 to 30% of the products.Aerobically, pyruvate was utilized by the freeplasmodia; however, acetate, not C02, was pro-duced. The exact mechanism for the formation-of acid and neutral volatiles by plasmodia isunknown, but the production of these com-pounds could result from a phosphoroclastic re-action similar to that described for several otherorganisms (see below).Studies with free parasites show an interesting

parallel between P. knowlesi (193) and P. gal-linaceum and P. lophurae (23, 24, 239, 240) withregard to carbohydrate metabolism. Free P. gal-linaceum oxidized pyruvate at a rate one-thirdthat of infected erythrocytes, and this rate couldbe increased by adding malate, adenosine tri-phosphate (ATP), nicotinamide adenine dinu-cleotide phosphate (NADP), and NAD in cata-lytic amounts (240). Furthermore, acetate wasproduced in large amounts by these free para-sites, but not by infected erythrocytes. Moulder(152) noted that there was a striking correlationbetween the cofactors needed for pyruvate oxi-dation in P. gallinaceum and the additions tothe culture medium, including coenzyme A(CoA), necessary for the successful extracellularcultivation of P. lophurae (253, 254). He specu-lated that the metabolic shift seen in free para-sites could be due to their lacking CoA. Sinceplasmodia are unable to synthesize CoA extra-cellularly (see below), its deficiency would leadto a reduced rate of pyruvate oxidation via thecitric acid cycle. (However, with free P. knowlesithe addition of CoA had no effect on glucoseutilization or lactate formation [191].) This couldbe true for the avian malarias, but it is alsopossible that the altered metabolism observedwith free parasites is a consequence of free par-asites being exceedingly leaky and that onceremoved from the host cell, the parasites losecritical.cofactors necessary for further metabo-lism of pyruvate. Such leakiness could be in-curred by the isolation procedure itself or may


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be an in vivo property of the parasite. To dis-criminate more fully between the "defect in car-bohydrate oxidation" hypothesis ofMoulder andacetate formation by a pyruvate clastic reaction,it will be necessary to identify the plasmodialenzymes involved in acetate formation. If thesewere found to be lacking in malarial parasites,then the hypothesis of Moulder of a leaky ma-larial parasite with a defect in CoA synthesiswould receive additional support. Conversely, ifsuch enzymes were present, then it would sug-gest that the plasmodia have an alternative cat-abolic route, one which may be less obviouswhen the parasites are intracellular.

In addition to the formation of lactate andpyruvate, P. knowlesi-infected cells also catab-olized glucose to the amino acids aspartic acid,glutamic acid, and alanine (173). These aminoacids, which were not formed in large quantities,most likely arose via C02 fixation reactions (seebelow).The metabolism of glucose by the rodent ma-

laria P. berghei parallels to a large degree thefindings for P. knowlesi. P. berghei-infectederythrocytes consumed 5 to 10 times more glu-cose than did uninfected erythrocytes, and morethan 70% of the added glucose was converted tolactate; smaller amounts of alanine, glutamicacid, and aspartic acid were also recovered (26,27, 29, 90, 158). The oxidation of citric acid cycleintermediates by erythrocyte-free P. berghei ap-pears to be restricted to succinate and pyruvate(156). Other workers have reported that lactatewas the primary end product of glucose catabo-lism by free P. berghei, but in comparison withthe intact host cells, free parasites had a de-creased ability to oxidize glucose or lactate (26,27).By contrast, studies with bird malarias (P.

cathemerium, P. gallinaceum, and P. lophurae)illustrate a rather different pattern of glucosecatabolism; extracts of free parasites carried outthe key reactions of the Embden-Meyerhoff gly-colytic pathway (23, 24, 89, 95, 137, 152, 239),free parasites oxidized pyruvate and other tri-carboxylic acid cycle intermediates, and aerobi-cally one of the principal end products was C02(23, 24, 89, 95, 240). Bovarnick et al. (23, 24)found that free P. lophurae had an 02 consump-tion rate with pyruvate and lactate equal to thatwith glucose, but with succinate and fumaratethe rate was only one-third of this. Related tothese observations are the findings of Trager(254) and Clarke (44, 45), who showed that py-ruvate, malate, and CoA enhanced the extracel-lular survival and development of P. lophuraeand P. gallinaceum. As with mammalian malar-ias, erythrocyte-free P. lophurae and P. galli-

naceum had a lower 02 consumption than didintact infected erythrocytes (23, 24), and in somecases the 02 consumed was only one-sixth thatexpected for the complete oxidation of glucose;this implies that either free parasites were dam-aged during the isolation procedure or they lackcofactors necessary for further oxidation of py-ruvate (see below). Both conditions may occur.Enzymes ofcarbohydrate metabolism. (i)

Glycolytic enzymes. The overall scheme ofglucose utilization presented above suggests thatall malarial parasites possess glycolytic enzymesof the Embden-Meyerhoff pathway. Indeed, al-though the reports are rather scattered, wheresuch enzymes have been looked for, they havebeen found (Table 5). Although it has beendifficult in some cases to decide unequivocallywhether the enzyme activity in a crude extractof parasites was due to the plasmodia, was a hostcell contaminant, or reflected the presence ofplatelets and leukocytes, in recent years some ofthese questions have been resolved by the use ofelectron microscope cytochemical methods, aswell as by the identification of plasnodium-spe-cific isoenzymes. (Ordinarily the latter are de-termined by histochemical staining of electro-phoretically separated extracts of parasites, hostcells, and presumed contaminants.)For example, the lactic dehydrogenase activ-

ity of P. knowlesi-infected cells was reported tobe 2.5 times greater than that of nornal cells(137), and increased lactic dehydrogenase activ-ity was measured with P. berghei-infected eryth-rocytes (28); however, the question remained asto whether these increases reflected enzyme ac-tivity of the parasites or stimulation of the hostcells. Such difficulties were resolved when Sher-man (206, 207) demonstrated the presence ofparasite-specific lactic dehydrogenase isoen-zymes in P. lophurae and P. berghei. Not onlywere these cathodally migrating enzymes dis-tinct from the anodal forms of the host cells, butthey showed rather different catalytic activitieswith NAD analogs (209) and pyruvate at differ-ent pH values. Subsequently, by employing sim-ilar techniques isozymic forms of hexokinase,glucose phosphate isomerase, and pyruvic kinasewere identified in rodent and human malariaparasites (34). Earlier, by more convential meth-ods, phosphoglyceric kinase, aldolase, and phos-phofructokinase were identified in P. gallina-ceum (152, 239).

It is regrettable that for any one malarialspecies there is not a single complete sequenceof glycolytic enzymes, nor do we know muchabout any of the kinetic properties of the en-zymes. Yet, all of the enzymes have been foundin one species or another, and it seems reasona-

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TABLE 5. Enzymes of carbohydrate metabolism in PlasmodiumP. gaUlinaceum P. berghei, P.

Enzyme and P. cathem- P. lophuraeb vinckei, and P. P. knowlesid P. falcprum'erium chabaudic

Hexokinase +f +Glucose phosphate + +

isomerasePhosphofructokinase +Aldolase +Those phosphate isomerase +Glyceraldehyde 3-phosphate +

dehydrogenasePhosphoglycerate kinase + +Pyruvic kinase + +Lactic dehydrogenase + + + + +G6PDH6-Phosphogluconate + + +

dehydrogenaseMalic dehydrogenase + + ±Succinic dehydrogenase + +Isocitric dehydrogenase +Phosphenol pyruvate + + +

carboxylase/carboxy-kinase

Cytochrome oxidase + + + + +a From references 239, 240, and 269.b From references 169, 206, 208, 212, 227, and 255 and unpublished data of Sherman.c From references 70, 134, 149, 156, and 166.d From references 67, 68, 137, 138, 173, and 191.e From references 34, 192, and 248.f +, Present; -, absent; ±, present in some strains.

ble to conclude that probably all ofthe glycolyticenzymes are present in all species of Plasmo-dium.

(ii) Citric acid cycle enzymes. The onlyenzyme of the citric acid cycle that has beenidentified with some degree of certainty in bothrodent and avian malarias is malic dehydrogen-ase (151, 212, 267).The P. lophurae malic dehydrogenase ap-

pears to be extramitochondrial (ie., soluble); itmay play a role in the reoxidation of reducedNAD for glycolysis (212) and in this way couldserve in a capacity smilar to that of lactic de-hydrogenase.

Isocitrate dehydrogenase specific for the birdmalaria P. lophurae has been identified (Sher-man, unpublished data), but isocitrate dehydro-genase specific for the rodent malaria P. bergheihas not been found (97, 98). There is, however,no evidence for the cellular localization of isocit-rate dehydrogenase in P. lophurae.

It has been claimed that P. lophurae and P.gallinaceum have succinate dehydrogenase ac-

tivity (199, 240); this enzyme, however, couldnot be identified in P. berghei (96, 156).Thus, there is suggestive evidence for enzymes

of the citric acid cycle in P. gallinaceum and P.lophurae, but such enzymes do not appear to be

present in the rodent malarias or P. knowlesi.Based on a product analysis of free P. knowlesi,P. lophurae, and P. gailinaceum with glucose(see above), it is possible that in addition to theconventional glycolytic pathway, malarial para-sites may be capable of enzymic fission of pyru-vate:

pyruvate + CoA -. C02 + acetyl-CoA + 2e-

acetyl-CoA + ADP + Pi -. acetate + CoA + ATP

This system has been described for the proto-zoans Entamoeba histolytica (180) and Tritri-chomonas foetus (154), for bacterial species suchas Clostridium acidi-urici (268) and Spiro-choeta aurantia (33), and for parasitic myco-plasmas (179). Alternatively, pyruvate could becleaved to acetate and formate, as occurs inEscherichia coli.

If such pathways exist in Plasmodium, theywould provide an additional source of energy forthose species lacking cytochromes and a citricacid cycle. Therefore, it will be of considerableinterest to determine whether the relevant en-zymes are present in plasmodia.

(iii) COfixing enzymes. Several species ofmalarial parasites appear to be capable of fixingC02. When radioactive NaHCO3 was added to

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cell suspensions of P. knowlesi-, P. berghei-, andP. lophurae-infected cells (70, 90, 95, 158, 215,228, 235), 14C was incorporated into keto andother organic acids as well as amino acids. Al-though glutamic and aspartic acids are minorC02 fixation products quantitatively, they areprobably important to P. knowlesi qualitativelysince such acids, when added exogenously, arenot incorporated into protein.

Free P. lophurae cells were also capable offixing C02 (227, 250). However, only in the caseof P. berghei have the enzymes of C02 fixation,phosphoenolypyruvate carboxylase and phos-phoenolpyruvate carboxykinase, been identifieddefinitively (70, 134). The phosphoenolpyruvatecarboxylase seems to be unique to P. bergheisince this enzyme has not been found in mam-malian cells.

(iv) Pentose phosphate pathway en-zymes. The pentose phosphate pathway oferythrocytes serves many roles; it is the principalsource of pentose sugars for nucleic acid synthe-sis, contributes to the maintenance of reducedglutathione, provides for regeneration ofreducedNADP (NADPH), and prevents the accumula-tion of methemoglobin. In addition, it has beenpostulated that the genetically determined defi-ciency of the first enzyme in the pathway, glu-cose 6-phosphate dehydrogenase (G6PDH), pro-vides human erythrocytes with protectionagainst malaria (20). Despite there being somecontradictory clinical evidence for this postulate,the growth of P. falciparum does appear to beimpaired in individuals with a G6PDH defi-ciency (72).The activity of the overall pentose pathway in

malaria-infected erythrocytes appears to be low.Thus, only 2% of [1-_4C]glucose was metabolizedto I'CO2 by free P. berghei, and less than 20% of[1-'4C]glucose was recovered as 14CO2 in P.knowlesi-infected cells (22, 29, 191, 205). Withfree P. knowlesi [1-_4C]glucose was incorporatedinto C02 to such a small degree that a pentosepathway was considered to be nonexistent (193).Also, little or no 6-phosphogluconate was formedby P. berghei from radioactive glucose (29). Sim-ilarly, Herman et al. (85) and Shernan et al.(230), using specifically radiolabeled glucose,came to the conclusion that the pentose pathwayplayed a minor role in P. gallinaceum and P.lophurae glucose catabolism.

Although the G6PDH activity of P. knowlesi-infected monkey erythrocytes increased with in-creasing parasitemia, there is no evidence for thepresence of G6PDH in the parasite itself (67-69,245, 246, 248). The increased levels of G6PDHin P. knowlesi-infected cells may be unusualsince in rodent and bird malarias the erythrocyte

MICROBIOL. REV.

level of G6PDH remains stable or declines de-spite the fact that the parasites are increasing insize (208). Both observations suggest that theG6PDH activity is not a reflection of a plasmo-dial enzyme.What is paradoxical about the pentose phos-

phate shunt and malarial parasites is that thesecond enzyme of the pathway, 6-phosphoglu-conate dehydrogenase, has been consistentlyidentified in plasmodial extracts and is electro-phoretically distinct from the host cell enzyme(34). However, no known mechanism exists forthe formation of the substrate for this enzymesince G6PDH, which catalyzes the formation of6-phosphogluconate from glucose 6-phosphate,is absent from all malarias studied. (Carter re-ported a G6PDH isoenzyme in P. berghei; thisis probably an error [69, 113], as is the earlierreport by Langer et al. [117].)

Therefore, it appears that for Plasmodiumthe pentose pathway is absent or a novel path-way exists. The parasite could utilize some ofthe pentose pathway intermnediates directly fromthe host cell, but as yet no compelling evidencefor this is available. The absence of a plasmodialG6PDH leaves the parasite without a mecha-nism for the regeneration of NADPH, a cofactorcritical to several biosynthetic schemes; it hasbeen suggested, however, that a plasmodium-specific glutamic dehydrogenase could assumethis role (221, 278).Decreased pentose phosphate pathway activ-

ity in host cells would impair the function of theerythrocytes by promoting the oxidation of re-duced glutathione and contribute to the forma-tion of methemoglobin. Eckman and Eaton (61)postulated that P. berghei utilized the NADPHof the host cells to maintain parasite-reducedglutathione. This hypothesis was based on theobservation that the reduced glutathione con-tent of the infected erythrocytes was increasedtwofold despite an absence ofreticulocytosis andthat most of the reduced glutathione was re-covered in the parasites themselves. Addition-ally, a plasmodial glutathione reductase specificfor NADP was also found. These authors rea-soned that if the parasites utilized host cellNADPH to reduce their oxidized glutathione,then the excess NADP would accelerate hostcell pentose shunt activity and contribute to itsoxidant sensitivity. Furthermore, in G6PDH de-ficiency the capacity of a cell to regenerateNADPH would impair parasite growth and pre-dispose the infected cell to premature destruc-tion. To be sure, pyridine nucleotide levels doincrease in malaria-infected cells (see below),and elevations of reduced glutathione and glu-tathione reductase do occur in the rodent ma-


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laria P. vinckei, but as yet the evidence for directutilization of host cell NADPH by a parasite ispurely circumstantial. Further, NADPH reduc-tion in the parasite could take place via theglutamic dehydrogenase (see above) identifiedin P. chabaudi (278) and P. berghei (118) aswell as P. lophurae (221), and thus there is noneed to postulate utilization of host cell stores.Indeed, in most ofthe malarial infections studiedto date the dislocation of the host cell pentosepathway appears to be minimal during intra-erythrocytic parasite growth.

If the pentose phosphate pathway of the par-asite is incomplete, where do the pentoses re-quired for plasmodial nucleic acid synthesiscome from? Possibly, pentoses arise from theaction of erythrocytic and/or plasmodial nucleo-side phosphorylases on host cell ATP catabo-lites, which in turn release a free base and ribose1-phosphate. Such a mechanisim of ribose for-mation does occur in mycoplasmas (179).Based on the forgoing considerations, carbo-

Glucose

6-phosphogluconate -lNADP

) 6-phosphogluconateNAPH dehydrogenoseNADPH

i ATribulose-5-PC4 Ar


hydrate metabolism for Plasmodium can be rep-resented as in Fig. 1. In the mammalian malarialspecies the primary end products are lactate andvolatile compounds, whereas in the avian speciespyruvate may be further oxidized via the citricacid cycle to C02 and water. The small amountsof succinate and keto acids, identified as endproducts of plasmodial catabolism of glucose,could arise from oxaloacetate by a reductivereversal of the usual pathway.Oxygen utilization and electron trans-

port. In vitro intraerythrocytic development ofP. lophurae, P. knowlesi, and P. falciparum(reviewed in reference 190) is favored by lowoxygen tensions. Also, malaria-infected eryth-rocytes show enhanced oxygen uptake with cer-tain exogenously added substrates when com-pared with uninfected erythrocytes (reviewed inreferences 89, 90, 137). This increased oxygenconsumption is demonstrable even under thosecircumstances where platelets, leukocytes, andreticulocytes have been accounted for or re-

'P-Ihexokinase

P'P-I-Glucose - 6-P04

PG

Fructose -6-P04TP

PFKDPrFructose 1,6, diPO4

aldolose

lP., ,..Glyceraldehyde-3- PC4TINAD

glyceroldehyde r3-PC4 DH n_NADH

glutamate 1,3, diphosphoglycerate

a ketoglutarate ADPs phosphoglycerate kinase

soc trate ATP

citrate artate 3-phosphoglycerote

O C02 l phosphoglycerate mutoseOAA *NADH- S ti 2-phosphoglycerateNAD * *malate enolase4

fumarate PEP alanine* ADP pyruvicsuccinate A kinase CoATh

PYRUVATE *acetyl CoA17-IACETATEacetyl CoA a ADP ATP

I NADH LDHCitric Acid Cycle

(Avian malarias) NAD LACTATE

FIG. 1. Proposed pathway of carbohydrate metabolism in Plasmodium. ADP, Adenosine diphosphate;PGI, glucose phosphate isomerase; PFK, phosphofructokinase; DHAP, dihydroxyacetone phosphate; TPI,triose phosphate isomerase; DH, dehydrogenase; PEP, phosphoenolpyruvate; LDH, lactic dehydrogenase;OAA, oxaloacetate.


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moved. (Interestingly, increased oxygen con-sumption by parasitized erythrocytes took placeafter exposure to 30,000 rads, a dose that de-stroyed infectivity [36].) Undoubtedly, some ofthe increased respiratory activity of malaria-in-fected erythrocytes is due to stimulation of themetabolism of the host cells themselves, but thestrong correlation of oxygen uptake with the sizeof the parasite and the time of schizogony (269),as well as the beneficial effects of lower oxygenlevels (<5%), strongly suggests that the parasitesthemselves are microaerophilic (190).

Indeed, when Warburg manometry was used,saponin-liberated P. lophurae showed a remark-able degree of oxygen uptake with pyruvate andlactate as well as glucose (23, 24). The respira-tory activity of such free parasite preparationswas inhibited by cyanide (CN), suggesting thatmetalloenzyme catalysis was somehow involvedwith oxygen uptake. Also, P. knowlesi-infectedcells were reported to have a heavy metal cata-lyst since oxygen uptake was inhibited by CNand CO. However, in the latter case the COinhibition could not always be reversed by stronglight (137, 138). Oxygen uptake may be difficultto correlate with the physiology of Plasmodiumsince there are many subtle factors that interactwith one another and are difficult to control. Forexample, temperature, time, serum concentra-tion, tonicity of the medium, pH, pO2, and thereduced amounts of hemoglobin in infected cellsmay affect the buffering capacity and oxygen-carrying capacity of the blood, and these in turnmay influence 02 uptake measurements. Sincethe malarial parasites utilize both hemoglobinand oxygen, such factors assume special signifi-cance. Indeed, although the parasites live in an02-rich environment and can assimilate 02, uti-lization may be substantial; e.g., P. falciparumgrows optimally in vitro at 3% 02 and survivesat 0.5%, whereas in vivo schizonts develop in themore anaerobic deep tissues (190).

It should be emphasized that in no instance,except for the presence of cytochrome oxidase,have cytochromes been isolated from plasmodia.CN sensitivity in itself does not prove the exist-ence of a cytochrome system, since the filarialworm Litomosoides carinii is CN sensitive, butlacks a cytochrome system and cytochrome ox-idase (30, 280). Also, Schistosoma mansoni, ablood fluke that lives in the bloodstream, wherethe supply of oxygen is ample, has cytochromec and cytochrome oxidase, and yet these respi-ratory enzymes account for <10% of the overall02 consumption of the worm. Schistosoma is ahomolactate fermenter, and the small amount of02 utilized is for tanning of the eggshells, notenergy production (30).

MICROBIOL. REV.

As noted above, malarial parasites do possessan enzyme for oxygen utilization. Scheibel andMiller (191, 192) identified cytochrome oxidaseactivity in platelet-free preparations of P. know-ksi, P. berghei, P. cynomolgi, and P. fakipa-rum. By electron microscope cytochemistry, thisactivity was localized in the acristate mitochon-dria ofP. knowlesi, P. berghei, and P. cynomolgi(249). Avian malarias show cristate mitochon-dria, and cytochrome oxidase activity was foundto be associated with this organelle (169, 249).The presence of cytochrome oxidase does not,

of itself, establish the existence of a functionalcytochrome-mediated electron transport system(see above). In the absence of a citric acid cycle,as appears to be the case for most m mlianmalarias, what can be the functional significanceof cytochrome oxidase? It is conceivable thatmalarial parasites contain a unique and as-yet-undiscovered electron transport chain or thatcytochrome oxidase does not serve as a compo-nent ofan aerobic energy-generating system, butfunctions in some other capacity. Gutteridge etal. (78) proposed that oxygen utilization by plas-modia may be coupled to the de novo biosyn-thesis of pyrimidines. The dehydrogenase thatcatalyzes the formation of dihydroorotate to or-otate (dihydroorotate dehydrogenase) is, at leastin mammals, particulate, mitochondrial, irre-versible, and closely linked to the cytochromechain, to which it passes electrons directly viathe ubiquinones and for which oxygen is theterminal acceptor (Fig. 2). This reaction, whichwas demonstrated with extracts from P. galli-naceum, P. berghei, and P. knowlesi, was in-hibited by CN and antimycin (78). If this findingis confirmed and extended (i.e., if superoxidesare formed or if the electrons pass to 02 with theformation of water), it would strongly supportthe notion that oxygen is used by Plasmodiumfor biosynthetic purposes, especially fabricationof nucleic acids.

C02, ATP, glutomine

Dihydrorotate Ubiquinone H2 Cyto H2 1/20

Orotate Ubiquinone Cyto H20

OMP

UMPFIG. 2. Proposed conversion of dihydroorotate to

orotate in Plasmodium (see reference 78). OMP,Orotidine monophosphate; UMP, uridine monophos-phate.


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It has been postulated, based on the indirecttechnique of measuring the effects of inhibitorson chloroquine-induced clumping of malarialpigment, that a branched electron transport sys-tem exists in plasmodia (94). It was hypothesizedthat one branch involved cytochromes, had ox-ygen as the terminal acceptor, and was CN sen-sitive, whereas the other branch was CN insen-sitive and had an unidentified electron acceptor.Although chloroquine-induced pigment clump-ing in P. berghei appears to be an energy-de-pendent process (and this has by no means beenunequivocally demonstrated), the unknownmanner by which the clumping occurs and therather high concentrations of inhibitors neededto prevent the occurrence of clumping make itdifficult to draw firm conclusions regarding theexistence of an electron transport system. Yet,despite severe limitations, a branched scheme isproposed here in the hope of stimulating furtherwork (Fig. 3). Substantiation or refutation ofsuch speculation will require additional inhibitorstudies, isolation and identification of cyto-chromes and other electron acceptors, and de-scriptions of the functional attributes of thesecompounds. (Branched oxidase pathways havebeen proposed for higher plant mitochondria[183], trypanosomes [reviewed in reference 86],and helminths [reviewed in reference 16].)

Nucleic AcidsDuring intraerythrocytic schizogony, the plas-

modia multiply rapidly to produce merozoites.Some appreciation of the magnitude of the syn-thetic capabilities ofthe parasites may be gainedby considering a developmental cycle in themost virulent species; in the course of 24 to 48 hthe parasite nearly fills an erythrocyte, and 10to 25 merozoite nuclei may be present. If an"average" malarial parasite produces about 16merozoite nuclei every 24 h, the parasite dou-bling time is approximately 6 h. What are thesources for such nucleic acid syntheses?Deoxyribonucleic acid: characteristics

and synthesis. Deane (54) and Chen (42), usingthe Feulgen staining technique, clearly showed

ATP ATP

Rotenone Cyto b-tCyto c _-Cyto a, a3--02AT Imya / Antimycin CN

NAD-Flavoprotein-1- *-.

Pyruvate Alternate ioxidose '02soci trate

a Ketoglutarate SHAMMalate Flavoprotein (possible inhibitor)t

SuccinateFIG. 3. Proposed electron transport system in

Plasmodium. SHAM, Salicyl-hydroxamic acid.


more than 30 years ago that malarial parasitescontained deoxyribonucleic acid (DNA). Despitethis fact, data on the biosynthesis of this nucleicacid remain scanty. Indeed, it took 5 years afterthe first identification of plasmodial DNA formetabolic studies to be initiated. In 1952 and1953 Whitfeld separated plasmodial DNA fromribonucleic acid (RNA) and found that 3P fromNa2H3PO4 was incorporated into both of thesenucleic acids by P. berghei-infected erythrocytes(284, 285). Similarly, Clarke (44, 45) demon-strated that P. gallinaceum growing either in-tracellularly or extracellularly synthesized DNA,and Schellenberg and Coatney (194) used 32pincorporation to evaluate the intracellulargrowth of P. berghei and P. gaUlinaceum in thepresence or absence of candidate antimalarials.However, what are the in vivo precursors of

this DNA? In those malarial species invadingnucleated erythrocytes, can the host cell nucleusbe a source of precursors? Lewert (129), usingultraviolet light absorption, suggested that P.gallinaceum degraded chicken erythrocyte nu-clei and thereby derived the raw materials fornucleic acid synthesis. However, almost simul-taneously Clarke (44) contradicted this by show-ing that this parasite could synthesize DNAextracellularly. The conflict was resolved in 1968with the demonstration that the synthesis ofDNA by P. lophurae was independent of thehost cell nucleus; that is, 'P was incorporateddirectly into plasmodial DNA and did not in-volve intermediates from the nucl"us of theerythrocyt'e (271). Finally, using a highly sensi-tive cytofluorometric technique, Bahr (11) wasunable to confirm the results of Lewert. Thus,at present there exists no evidence that theintraerythrocytic stages of malarial parasites usenucleic acid precursors derived from erythrocytenuclei.A merozoite or a ring-stage trophozoite con-

tains _ 10-13 g of DNA, and this amount mayincrease 10- to 20-fold during schizogony (11,12). When during intraerythrocytic developmentis this DNA made?

In vitro studies in which synchronous infec-tions of intraerythrocytic P. lophurae were usedshowed a lag of 'P incorporation into DNA for2 to 4 h, maximum synthesis for the following 20h, and then a leveling off of incorporation for thelast 8 to 10 h of development (271). P. lophuraehad a 36-h schizogonic cycle (243), and duringthe first 2 to 4 h in culture the predominantstages were early rings; the greatest incorpora-tion took place when the erythrocytes containedlate rings and early schizonts, whereas duringthe last 8 to 10 h schizogony was essentiallycomplete. Similar in vitro results were obtained


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with P. knowlesi, a species with a 24-h asexualcycle: a lag of incorporation for 5 to 10 h, maxi-mum synthesis for 10 to 15 h, and then an abruptcessation of incorporation when the parasitesentered schizogony (79, 172). In all of thesestudies, despite the variation in the duration ofthe asexual cycle, the synthesis of DNA ap-peared to occur continuously, and there was noevidence for periodicity. However, Gutteridgeand Trigg (81) and Jung et al. (103) claimned thatnucleic acid synthesis was discontinuous. Gut-teridge and Trigg (81) found that the S phase ofthe P. knowlesi cell cycle occurred during thering and trophozoite stages and that little or nosynthesis occurred during schizogony. There wasa very short G, phase (1 h) and little indicationof a G2 phase. These authors also were of theopinion that adenosine incorporation was a morevalid measure ofDNA synthesis than orotic acidincorporation (used in reference 172) for tworeasons: (i) entry of orotic acid into the eryth-rocytes could be limited during the early growthof the parasites, since the membranes of theerythrocytes and parasites are relatively im-permeable to pyrimidines; and (ii) because oroticacid has to be added at a high specific activityto see any incorporation, its incorporation rep-resents a minimal amount of DNA synthesis.Others (48) have questioned these contentions,stating that DNA synthesis occurred primarilyin late trophozoites and early schizonts regard-less ofwhether orotic acid or adenosine was usedas the tra9er. Although there has been no com-pletely satisfactory resolution ofthese contradic-tory findings, it is entirely possible that thedifferences are more apparent than real. Thedivision of nuclei is not always synchronous inall parasites, and therefore it is highly unlikelythat one would see a pattern of incorporationtypical of the cell cycle in a uninucleate eucar-yote. Furthermore, as Cooper (52) has observed,the GI period may be an artificial construct thatis only present in slowly growing cells, and it ismore reasonable to perceive G, as the time fromthe initiation of one round of DNA synthesis tothe next. Indeed, he notes, "If eukaryotic cellscould produce two 'interphase' nuclei followingthe S period (for example, if the nuclei under-went an immediate mitosis without cell division)then it would be possible for DNA synthesis tooccur in these nuclei while the cells were un-dergoing division." Such appears to be the casein Plasmodium and explains, in part, the diffi-culty in defining a G, period in these organisms.The DNA isolated from P. knowlesi- and P.

falciparum-infected cells consists of two com-ponents, a major component with a buoyantdensity of 1.697 and a minor component with a

MICROBIOL. REV.

density of 1.679. Although no unusual bases werefound, the base composition of the minor com-ponent was distinctive in its low guanine pluscytosine (G+C) content (-19%); the major (nu-clear) component was similar in host and para-site in buoyant density and G+C content(-37%). The rodent malarias P. berghei, P. cha-baudi, P. vinckei, and P. yoelii showed a singlemajor component with a buoyant density of1.683, corresponding to 24% G+C; minor satellitebands of buoyant densities 1.671 and 1.677 werefound in P. berghei and P. chabaudi (41, 82, 83).The validity of designating four distinct speciesof rodent malaria has been supported by DNA-DNA hybridization studies (41). The avian ma-larias P. gallinaceum and P. lophurae had amajor component with a density of 1.678 to1.680, corresponding to 18 to 20% G+C. It isclear that according to nuclear DNA base com-position, malarial parasites can be separated intothree discrete categories: primate malarias with37% G+C, rodent malarias with 24% G+C, andavian malarias with 18 to 20% G+C.

In P. lophurae a mitochondrial DNA with abuoyant density of 1.679 (identical to nuclearDNA density) was isolated and shown to havecircular contours of 10 ,um (105). The minorsatellite bands observed in DNA preparationsfrom P. knowlesi, P. berghei, P. chabaudi, andP. falciparum may also represent mitochondrialDNA, but DNA has not been isolated frommitochondria or mitochondrion-like bodies inthese species.Ribonucleic acid: characteristics and

synthesis. The RNA content of the malarialparasites has been estimated to be five timesgreater than the DNA content (11, 79, 284, 285).Most of the plasmodial RNA is localized inabundant cytoplasmic ribosomes which are pres-ent in the intraerythrocytic parasites. The ribo-somalRNA (rRNA) ofP. lophurae was typicallyprotozoan in its sedimentation values of 25S and17S (in contrast to the 28S and 18S rRNA's ofother eucaryotes [218]). Similarly, P. knowlesirRNA had sedimentation values of 24.2S and17.4S (216, 281), and the sedimentation valuesof P. berghei rRNA were 25S and 14.9S (251).The nucleotide base compositions of the rRNA'sof these three species were also found to bedissimilar from those of other eucaryotes, butakin to those of other protozoans, in having alow G+C content (-35%). (Typically, the rRNAof the host cells was found to be -64% G+C.)Some years ago it was postulated that the

large rRNA of P. berghei ribosomes was pro-vided by the host cell ribosomes (251); however,this could not be confirmed with P. knowlesi orP. lophurae. Thus, free P. lophurae cells, after


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being pulse-labeled in vitro with [3H]adenosine,were shown to have radioactivity in both thelarge (60S) and small (40S) ribosomal subparti-cles, and P. knowlesi-infected cells incubated inthe presence of radioactive adenosine had majorpeaks of radioactivity which corresponded torRNA's of 24.2S and 16.6S (218, 226). Based on

these findings, a reinvestigation of the charac-teristics of P. berghei rRNA was carried out.This new study (141) has shown that the longdelay in the labeling of the 25S component ofthe plamodia after cells were incubated withNaH2'PO4 or [14C]orotic acid (and which was

taken to be evidence for an absence of plasmo-dial biosynthesis of the large ribosomal subpar-ticle) was probably the result of uncontrolledribonuclease activity by the plasmodia, whichcontributed to the degradation of the 25S rRNA.(However, since these workers obtained a 3plabeled 25S RNA in some cases [251] but couldnot recover label in the 60S subparticle of theparasite, there was already a strong indicationthat the poor labeling of the plamodial RNAwas artifactual and could not be ascribed toincorporation of host materials.) Although thelarge rRNA of P. knowlesi and P. lophurae hasbeen recovered intact, in P. berghei it was al-ways isolated in a partially degraded form. Ithas been suggested that this could be due to a"nicking" of the RNA or the in situ action ofnucleases on an exposed portion of the RNA, or

it might represent a natural occurrence duringthe maturation of this rRNA. Although ribonu-clease was demonstrated in P. gaUlinaceummany decades ago (148), the ribonuclease of P.berghei remains of considerable interest in thatits activity appears to be undiminished by con-

ventional inhibitors, and its action is restrictedto the large rRNA.The pattern of rRNA synthesis in Plasmo-

dium is not completely understood. The occur-

rence of45S 37.2S, and 31.7S RNA species in P.knowlesi suggests that the processing of rRNAmay be typically eucaryotic (266). In most eu-

caryotes rRNA processing takes place in thenucleolus. A compact nucleolus is absent fromsimian and rodent malarias, whereas avian ma-

larias do have a compact nucleolus. The reasons

for the absence of a nucleolus in some malariaspecies could be due to the following factors: (i)

the number of copies of ribosomal genes is low;(ii) the rate of gene transcription is low, whereasthe processing time is rapid; (iii) the supply ofribosomal proteins synthesized in the cytoplasmis limiting; and (iv) the nucleoli are not obviousbecause they are dispersed. Which among thesealternatives (if any) is operative has not beendetermined.


The synthesis of RNA during the intra-erythrocytic development of P. knowlesi hasbeen studied in several laboratories. For exam-ple, Polet and Barr (172) followed the incorpo-ration of [14C]orotic acid into total RNA andobserved a slight lag for the first 5 h, which wasfollowed by a linear rate of incorporation. Gut-teridge and Trigg (79) found a similar in vitropattern with [3H]adenine and [3H]orotic acid asthe tracers, with a lag for 4 to 8 h, linear incor-poration for - 10 h, and then abrupt cessation asthe parasites entered schizogony. However, with[3H]adenosine no lag in incorporation was evi-dent, and maximum incorporation occurred atthe late trophozoite stage before nuclear division(81). Trigg and Gutteridge (264) found that whenP. knowlesi cells were grown in vitro in cellsuspension cultures, there was a striking defi-ciency in the synthesis of RNA during the sec-ond growth cycle, and they proposed that thiscould in part account for the inability of theparasites to multiply in culture. As noted above,periodicity of DNA and RNA syntheses by P.knowlesi is clouded by controversy (48, 81). Al-though most workers have been unable to dem-onstrate incorporation of uracil or uridine intoRNA, one laboratory (48) claimed evidence forthis. The discrepancy could be due to purity ofthe isotopes, presence of leukocytes, very highspecific activity of the tracer, and/or varied cul-ture conditions. It is likely that uridine anduracil are incorporated by malarial parasitesonly when supplied in high concentrations andthat this is responsible for some of the variedfindings. The general lack of incorporation ofuracil by Plasmodium is in contrast to the in-corporation of uracil in a related coccidian, Tox-oplasma gondii. However, in the latter caseuracil incorporation was dependent on a para-site-specific uridine phosphorylase (165), an en-zyme that is apparently lacking in malarial par-asites.Pyrimidine biosynthesis. In 1967 and 1968

Bungener and Nielsen showed that P. berghei-and P. vinckei-infected erythrocytes incorpo-rated tritiated hypoxanthine and adenosine, butnot thymidine. From these findings as well asfrom correlated studies, it became clear thatplasmodia were unable to synthesize the purinering de novo; that is, labeled formate and glycinewere not incorporated into plasmodial purinesderived from nucleic acids (114, 214, 272). There-fore, the purines necessary for the synthesis ofnucleic acids and other metabolic functions hadto be obtained by so-called salvage pathwaymechanisms. By contrast, it was found that theplasmodia did have the capacity for the de novofabrication of pyrimidines. In keeping with the


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incorporation of '4C-labeled bicarbonate intoplasmodial DNA and RNA has been the identi-fication of about six enzymes associated with denovo pyrimidine synthesis, ribonucleosidemonophosphate formation, and conversion ofthese compounds into deoxyribonucleotides(Fig. 4).

It is of considerable interest to note that inthose cases where it has been studied, only the

bicarbonate

aspartate carbamoyl P04

dihydroorotic acid

lb

orotic acid

Ic

OMP

UMP -1

UDP

Huridine

UTP dUDPT thymidine

CTP -

+ dUTP g

CDP + TdUMP -ddTMP

dCDP dCMPJ

AddCTP dTDP

dCRdTTP

Nucleic AcidsFIG. 4. Pyrimidine biosynthesis in Plasmodium.

OMP, Orotidine monophosphate; UMP, uridinemonophosphate; UDP, uridine diphosphate; UTP,uridine triphosphate; CTP, cytidine triphosphate;CDP, cytidine diphosphate; dCDP, deoxycytidine di-phosphate; dCTP, deoxycytidine triphosphate;dUDP, deoxyuridine diphosphate; dUTP, deoxyuri-dine triphosphate; dUMP, deoxyuridine monophos-phate; dCMP, deoxycytidine monophosphate; dTMP,deoxyribosylthymine monophosphate; dTDP, deoxy-ribosylthymine diphosphate; dTTP, deoxyribosylthy-mine triphosphate; dCR, deoxycytidine. The enzymesindicated on the figure are: a, aspartate transcar-bamylase (161); b, dihydroorotic acid dehydrogenase(78, 116); c, orotidine monophosphate pyrophospho-rylase (271); d, deoxycytidine kinase (114); e, deoxy-cytidylate aminohydrolase (114); f, thymidylate syn-thetase (182, 271); g, thymidine kinase (not present or

very low activity).

MICROBIOL. REV.

folate enzyme associated with pyrimidine bio-synthesis (i.e., serine hydroxymethyltransferase)has been identified in plasnodial extracts,whereas the folate enzymes involved with purinemetabolism are conspicuous by their absence(see below).Purine transport and salvage pathway

mechanisms. A clue to the requirement forexogenous sources of nucleic acid precursors bymalarial parasites was evident from some of theearliest in vitro growth studies with P. knowlesi(137); namely, omission of the "purine plus py-rimidine" component from the medium ad-versely affected the intraerythrocytic growth ofthe plasmodia. Whether both purines and py-rimidines were required could not be determinedat that time by using this complex system. Later,however, with the availability of radioisotopes itwas possible to show specific precursor require-ments more clearly. When mice and rats infectedwith the rodent malaria P. berghei or P. vinckeiwere injected with the tritiated pyrimidinesuridine and thymidine, there was no evidence ofincorporation; in vitro incubation of P. vinckei-infected mouse blood gave similar results. How-ever, P. berghei-infected cells became heavilylabeled when blood was exposed to radioactiveadenosine (32). Autoradiographic studies withhypoxanthine showed similar patterns with P.berghei- and P. vinckei-infected erythrocytes.In addition, when mouse erythrocytes labeledwith [3H]adenosine were transfused into miceinfected with P. vinckei, radioactivity was incor-porated into the malarial parasites (114, 214).Thus, Bungener and Nielsen discovered thatmalarial parasites utilized exogenous supplies ofpurines, but not pyrimidines.The data on purine uptake by rodent malarias

have been completely confirmed by primate andhuman malaria studies. Thus, intraerythrocyticP. knowlesi utilized adenine, adenosine, deoxy-adenosine, guanosine, and hypoxanthine; thy-mine, thymidine, uracil, uridine, cytidine, anddeoxycytidine were not incorporated. Only onepyrimidine, orotic acid, was incorporated intoDNA and RNA (79, 172). P. lophurae-infectederythrocytes also had a high uptake of adeno-sine, inosine, and hypoxanthine, with the degreeof accumulation being directly related to size ofthe parasite (256). In the case of adenine, thereverse was true; infected cells with multinu-cleate parasites took up less than erythrocytescontaining uninucleate forms. Adenosine mono-phosphate (AMP), ATP, and inosine monophos-phate were taken up to a very limited degree.Adenosine uptake by infected cells was inhibitedby the presence of hypoxanthine and inosine,but such mutual inhibition was not seen in nor-


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mal cells; adenine, guanine, and ATP did not actas uptake inhibitors. Tracy and Sherman (252)suggested that adenosine was deaminated toinosine shortly before or during uptake. Thus,they concluded that adenosine, inosine, and hy-poxanthine have a common 6-oxypurine trans-port site on infected erythrocyte membranes.Most workers have suggested that only pu-

rines are transported into malaria-infectederythrocytes, whereas pyrimidines (except fororotic acid) are excluded. Neame et al. (quotedin reference 214) have disputed this. They in-cubated P. berghei-infected cells with purines(adenosine and guanosine) and pyrimidines (cy-tidine and thymidine) and showed that for allsubstrates tested there was equilibration acrossthe erythrocyte membrane; however, only pu-rines were incorporated. Thus, according tothese workers, the poor utilization of pyrimi-dines was not due to their inability to permeatethe erythrocytes, but rather a deficiency of theenzymes necessary to effect phosphorylation or

incorporation was involved. It should be notedthat in the in vitro studies of Neame et al.incubation time was prolonged (1 h) and sub-strate concentrations were exceedingly high (1mmol/liter). Probably, pyrimidines (except fororotic acid) are not utilized by malaria-infectedcells for several reasons. First, they are trans-ported very slowly into the erythrocyte; second,the appropriate host cell and plasmodial en-

zymes to convert these into nucleotides are lack-ing; and third, the parasites synthesize pyrimi-dines de novo.

In 1967, it was suggested that host erythro-cytes could be an important source of purines

for the synthesis of plasmodial nucleic acids (31).Shortly thereafter, Walsh and Sherman (272)speculated, "The parasite is capable of the denovo synthesis of pyrimidines including thymi-dylate and presumably utilizes these compoundsfor the synthesis ofDNA and RNA. P. Iophuraesynthesizes purines de novo only to a limitedextent and therefore must rely on exogenous

sources of these compounds. The most obvioussource is the host erythrocyte which contains a

relatively high concentration of purines . ."

What are the purine sources in erythrocytesthat growing plasmodia require? Approximately80% of the purines of the erythrocytes are in theform of ATP, and potentially this ATP could beutilized by the parasite. Indeed, Trager (253)found that the extracellular survival of P. lo-

phurae (freed from host cells by hemolytic an-

tiserum) was favored by the addition of ATP tothe erythrocyte extract medium. Such extracel-lular plasmodia incorporated orotic acid and ad-enine but not uridine. During a 3- to 4-h incu-


bation, orotic acid incorporation was reduced ifCoA was omitted or if dilute erythrocyte extractwas used, but not if ATP and pyruvate wereabsent from the medium. Although ATP pro-moted extracellular development ofthe parasite,both AMP and ATP interfered with adenineuptake (256). This suggests that the added AMPand ATP were altered after being added to theerythrocyte extract and that the resultant nu-cleosides and nucleobases acted as competitorsfor the plasmodial adenine transport locus.There is no evidence as yet to show exactly whatrole ATP has in parasite growth. Indeed, in theview of Trager, exogenous ATP is involved inthe active transport of materials across the twoclosely apposed membranes that separate thecytoplasms of the erythrocytes and the para-sites. In support of this, Trager (257,258) showedthat bongkrekic acid (an inhibitor of mitochon-drial adenosine triphosphatase and cation trans-port) inhibited the extraceliular growth of P.lophurae and the intracellular development ofP. falciparum. Addition of ATP (2 to 12 mmol/liter) reversed the bongkrekic acid (1.4 to 35 ,ug)effects. Atractyloside, a related inhibitor, waswithout effect, however. IfATP is not the purinetransported into the parasite, what is?When "saponin-freed" P. lophurae was used,

high uptake ofhypoxanthine, adenosine, inosine,and guanine was demonstrated, whereas uptakeof guanosine, xanthine, adenine, AMP, ATP,and inosime monophosphate was low (252). Themagniitude of the uptake, the high distributionratios (>90% of the available radioactivity wasinside the cell at low substrate concentrations),the saturation kinetics at high purine levels, andthe phenomenon of mutual inhibition suggestedthat the free parasites transported adenosine,inosine, and hypoxanthine by a common carrier.Unfortunately, the transport studies were com-promised by long (5-min) incubation periods andthe fact that some substrate metabolism tookplace. P. lophurae was capable of forming bothATP and guanosine triphosphate from addedhypoxanthine, inosine, and adenosine; 90 to 95%of the added radioactivity was incorporated intoRNA, and 5 to 10% went into DNA. Tracy andSherman (252) speculated that hypoxanthinemight be the purine species available to P. lo-phurae in vivo and that the added adenosinewas probably deaminated to inosine during up-take by free parasites in vitro. It is of interest tonote that good intraerythrocytic growth of P.lophurae took place in Weymouth medium,which contains 2.5 x 10-4 mol of hypoxanthineper liter as the sole purine (271), that Gutteridgeand Trigg (79) found that one growth cycle couldbe completed in vitro by P. knowlesi without


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exogenous purines, and that the continuous cul-tivation of P. falciparum (260) was achieved ina medium devoid of purines except for thosepresent in human serum.

Studies of purine uptake by mammalian plas-modia are limited to one species, P. berghei(reviewed in reference 214). Two types of prep-aration have been used: saponin-prepared para-sites in a plasma-containing medium andsaponin-liberated parasites suspended in Krebsbuffer. The early studies with parasites in Krebsbuffer indicated that radioactivity from adeno-sine or ATP was incorporated into parasite nu-cleic acids; it was presumed (but not shown) thatATP was degraded to adenosine by adenosinetriphosphatase from contaminating erythrocytestroma. When this same scheme was used, cyti-dine and uridine were not incorporated, and theauthors suggested that "the membrane of theparasite exerts a selectivity on the penetrationof pyrimidine, but not on purine, nucleosides."When these investigators began to use parasitesin plasma, the results obtained were somewhatdifferent; the pattern of incorporation was [8-3H]AMP > [8-3H]adenosine diphosphate > [8-3H]ATP > [8-3H]adenosme, but no incorpora-tion of label was obtained with [a-32P]ATP orjfl,y-32P]ATP. To explain this paradox, they sug-gested that the 32p label was cleaved from the[32P]ATP before the nucleoside adenosine pen-etrated the plasmodia. It was suggested that thehigher efficiency of [8-3H]AMP when comparedwith [8-3H]adenosine was due to the formerbeing a protected form of the nucleoside. Inaddition, they proposed that AMP was not aseasily deaminated as adenosine and that theslow dephosphorylation of AMP gradually re-leased adenosine, which was presumably takenup by the parasite. The use of plasma as asuspending medium, coupled with the presenceof erythrocyte stroma in the reported experi-ments, makes the interpretations of substratetransport uncertain (for example, plasma con-tains adenosine deaminase so that the-addedadenosine would be converted to inosine, whichin turn could be taken up by the parasite).Although earlier Van Dyke and co-workers wereof the opinion that adenosine was phosphoryl-ated to AMP by the parasite, their recent studieswith parasites in Krebs buffer suggested thefollowing scheme: adenosine -- inosine -- hy-poxanthine in the extracellular medium, withphosphorylated compounds (inosine monophos-phate -- AMP -- adenosine diphosphate -*ATP) being present in the parasite. Moreover,compound 555 of Patterson (an inhibitor ofadenosine transport) had no effect on adenosineincorporation, whereas 10-6 to 1iO4 M purine-6-

sulfonic acid-3-N-oxide effectively blocked up-take and phosphorylation of tritiated hypoxan-thine by both parasitized and free P. berghei.Accordingly, the conclusion that they drew wasthat adenosine is probably not the immediateprecursor for phosphorylation, but that hypo-xanthine is the initial entry compound for freeparasites (131). Unfortunately, interpretation ofthis study remains ambiguous since no controlpreparations of erythrocyte ghosts (which rou-tinely contaminate saponin-freed parasites [3,50, 112]) were tested, total removal of leukocytesand platelets was not attempted, and parasitesentirely free of erythrocyte contaminants (e.g.,plasmodia freed by hemolytic antiserum) werenot employed. Furthermore, since we havefound that saponin-liberated parasites are leakyto macromolecules (286), it is difficult to deter-mine whether the added label or some interme-diate produced by plasmodial enzymes presentin the incubation medium was utilized. Despitethis, the contention that hypoxanthine is thepreferred purine for intraerythrocytic plasmodiais probably correct (see below).Support for the key role of hypoxanthine can

be found in the study of Bungener (reviewed inreference 114). He found that the addition ofallopurinol to the drinking water of rats andmice enhanced the severity of P. berghei, P.vinckei, and P. chabaudi infections. Since allo-purinol acts as an inhibitor of xanthine oxidase,it may be that allopurinol treatment of the hostincreased the blood levels of hypoxanthine. (Al-lopurinol could have other effects, however.) Ifwe assume that purines are a limiting factor inthe multiplication of the parasites, then the ad-ditional supplies of hypoxanthine in allopurinol-treated animals could contribute to parasite re-production.

If hypoxanthine were a key metabolite forparasite nucleic acid synthesis, it would be proc-essed by a plasmodial hypoxanthine phosphori-bosyltransferase. Isozymes of hypoxanthinephosphoribosyltransferase have not been iden-tified in plasmodia; the molecular weight of theenzyme from P. chabaudi was very similar tothe weight of the enzyme from erythrocytes(74,000 versus 68,000); however, the isoelectricpoints of the two enzymes did differ (276). TheKm for P. chabaudi hypoxanthine phosphori-bosyltransferase for hypoxanthine was 2.5 x 10-6M, and the enzyme was competively inhibitedby 6-mercaptopurine, 2-amino-6-mercaptopu-rine, and 3-thioguanine (276).

In deteriorating human erythrocytes the fol-lowing sequence of catabolic reactions occurs:ATP -- adenosine diphosphate -- AMP -+ ino-sine monophosphate -* inosine -- hypoxanthine

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(21). Malaria-infected erythrocytes (except per-haps for P. berghei growing in reticulocytes) canalso be considered to be deteriorating cells andthrough the action of erythrocytic purine nu-cleoside phosphorylase probably contain signif-icant amounts of hypoxanthine. Since the cir-culating levels of adenosine and adenine are low(_1O-7 M or less in human blood) and the par-asites are unable to synthesize purines de novo,the utilization of purines by salvage pathwayenzymes becomes critical to the development ofthe parasites. Although adenosine could beformed from erythrocyte AMP by a parasitemembrane-associated 5'-nucleotidase, no suchactivity is associated with plasmodia (286).Therefore, hypoxanthine assumes primary im-portance. Hypoxanthine utilization most likelyoccurs in vivo. However, in vitro the plasmodiacan also transport inosine and adenosine, be-cause of the common oxypurine transport locus.Once transported into plasmodia, the parasitesalvage pathway enzymes convert the inosinemonophosphate into adenine and guanine nu-cleotides, and ultimately these are used for thesynthesis of RNA, DNA, and nucleotide coen-zymes. Purine salvage pathway enzymes havebeen identified and characterized from P.berghei, P. chabaudi (114, 130, 164, 276), and P.Iophurae (286). The purine salvage scheme isillustrated in Fig. 5.

Protein SynthesisMalarial parasites obtain the amino acids nec-

essary for protein biosynthesis in three ways: (i)biosynthesis ofamino acids from carbon sources;(ii) uptake ofpreformed free amino acids presentin the plasma or host cells; and (iii) proteolysisof hemoglobin with the release of amino acids.Biosynthesis of amino acids. Malarial par-

asites and malaria-infected erythrocytes fix C02and thereby synthesize the amino acids alanine,aspartic acid, and glutamic acid (173, 227, 228,235). It is noteworthy that the free amino acidpool of infected erythrocytes is substantially in-creased in these same amino acids (39, 76, 220).Since very little of the amino acids formed viaC02 fixation are incorporated into plasmodialproteins, it may be that this pathway serves ananaplerotic role in the economy of the parasite.Thus, it is conceivable that-glutamic acid oxi-dized via a parasite-specific NADP-glutamic de-hydrogenase coupled to the enzymes of the citricacid cycle provides an ancillary source of energyfor the avian malarias. Indeed, free P. lophuraecells metabolize more than 60% of added ["4C]-glutamic acid to '"CO2 (223). Rickettsiae alsoobtain energy via glutamate oxidation (25). An-other functional role for glutamic dehydrogenase


glycine + formate }PRPP + glutomine o'

PARASITE RBC

ATP

AtP

AMP4IMP

Inosine'ribose-I-PO4

... L7-r --O

HT.HPRT

adenylosuccinate IMP

synthetase and lyase /AMP GMP

4 4ADP GDP

* 4ATP- GTP- folate biosynthesis

nucleic acids

FIG. 5. Salvage pathway for purine utilization inPlasmodium. PRPP, Phosphoribosylpyrophosphate;ADP, adenosine diphosphate; RBC, erythrocyte;IMP, inosine monophosphate; HPRT, hypoxanthinephosphoribosyltransferase; GMP, guanosine mono-phosphate; GDP, guanosine diphosphate; GTP, gua-nosine triphosphate.

in the rodent malarias (34, 118, 221, 278), whichappear to lack a complete citric acid cycle, aswell as in the avian malarias, may be the reduc-tion of NADP; in the absence of a plasmodialG6PDH (see above), the conversion of gluta-mate to a-ketoglutarate may be a primary mech-anism for regenerating the NADPH needed forreductive synthesis.Uptake of free amino acids. Malarial par-

asites and malaria-infected erythrocytes havethe capacity to take up free amino acids fromthe external environment. P. knowlesi-infectederythrocytes accumulate isoleucine, methionine,leucine, cystine, and histidine (reviewed in ref-erence 214), and this supports the earlier in vitroand in vivo growth studies which showed thatthe intraerythrocytic stages of P. knowlesi re-quired extracellular sources of methionine andisoleucine (73, 174). It was contended that thereason for the specific requirement of P. know-lesi for exogenous methionine (and cysteine), aswell as isoleucine, was that primate hemoglobinwas particularly deficient in these amino acids.Although a good correlation was claimed to existbetween the amino acid composition of hemo-globin and the need to supply amino acids extra-cellularly, the universality of this explanationappears to be invalid. It holds for isoleucine, butnot for other amino acids. For example, althoughmethionine and cysteine levels are low in pri-mate hemoglobin, these compounds are alsopoorly incorporated when supplied exogenously,whereas the converse is true for leucine. There-fore, the degree of plasmodial incorporation ofan amino acid is not simply a reflection of the


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availability of that amino acid in hemoglobin,but also involves the content of the free aminoacid pool, factors influencing the entry rate (i.e.,concentration, charge, etc.), the amino acid com-position of plasmodial proteins, and their rateand duration of synthesis (214). In no case hasthere been a complete assessment of the contri-butions of these factors.

Characteristically, malaria-infected erythro-cytes take up and incorporate exogenously sup-plied amino acids at an accelerated rate com-pared with uninfected erythrocytes. This featurehas been useful (especially with isoleucine,which is readily incorporated) in all malariasstudied (i.e., P. falciparum, P. berghei, and P.lophurae) for monitoring plasmodial growth un-der a variety of culture conditions, for evaluationof antibody and drug efficacy, and for assessingviability after cryopreservation (reviewed in ref-erence 214).

Unfortunately, no detailed studies exist on themechanism of transport of isoleucine (or anyother amino acid for that matter) into erythro-cytes infected with mammalian malarias. How-ever, studies ofamino acid uptake with the avianparasite P. lophurae may provide some clues asto the nature of the transport processes in theseother malarias. It has been reported thatduckling erythrocytes infected with P. lophuraetend to show diffusion entry of several aminoacids (alanine, leucine, histidine, methionine,cysteine) which are ordinarily transported bycarrier-mediated processes in normal cells, andeven for those amino acids that continue to bemoved across the erythrocyte surface by carrier-mediated processes (glycine, serine, threonine,lysine, and arginine) there was an altered (in-creased) transport constant. It was suggestedthat this leakiness of infected cells was due to adepletion of host cell ATP and plasmodium-induced (enzymic?) alterations in the erythro-cyte membrane (225). However, an attempt toestablish the transport characteristics of alanineobserved in the malaria-infected cells by treatingnormal duckling erythrocytes with parasite ex-tracts and reducing the ATP content was onlypartially successful.In 1962 Moulder (151) wrote, "In the course

of evolutionary adaptation to life inside the redcell, the malarial parasite may have lost manyof the active transport systems regulating thepassage ofmolecules in both directions across itscell membrane and may have become freelypermeable to all sorts of molecules which itderives from its host." Sherman and Tanigoshi(225) attempted to test this hypothesis for P.lophurae "freed" from the host cells by saponinlysis. (It has been shown that such saponin-

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prepared parasites are not entirely removed andthat many parasites remain enclosed by eryth-rocyte membranes [3, 50, 112, 261].) Of all theamino acids tested, only arginine, lysine, glu-tamic acid, aspartic acid, and cysteine enteredthe parasite by what appeared to be a carrier-mediated process; all of the others entered bysimple diffusion. These studies with free para-sites, although appearing to support the perme-ability defect hypothesis of Moulder, may notactually do so. Thus, the parasites used in theaforementioned studies were still enclosedwithin erythrocyte membranes, and this couldhave influenced the results. (However, it is note-worthy that ghosts prepared from normal cellsby saponin lysis had none of the properties ofthe suspensions of plasmodia prepared in thisway.) Also, the reported transport constants forthe parasites could be error prone because ofprotracted incubation times and sampling tech-niques. However, the most severe limitation tothese results involves the characteristics of theparasites themselves. Recently, we discoveredthat saponin-prepared parasites, and indeed par-asites liberated by the gentler hemolytic antise-rum method, lose high-molecular-weight cyto-plasmic constituents, including proteins (286).Therefore, it remains uncertain whether theleaky free parasites commonly used for meta-bolic work are highly penneable in situ (intra-cellularly) or whether this condition is provokedby the techniques used to remove the parasitesfrom the host cells. Until this dilemma can besatisfactorily resolved, it will be difficult to askcritical questions regarding the transport prop-erties of free parasites.Proteolysis of hemoglobin and the for-

mation of malarial pigment. Since de novobiosynthesis of amino acids is restricted in kindand because the free amino acids present withinerythrocytes are presumed not to be of sufficientquantity to serve for the synthesis of plasmodialproteins, the hemoglobin of the erythrocytesremains the most abundant reservoir of aminoacids available to malarial parasites.

Early malariologists recognized that thegolden brown-black pigment (hemozoin) accu-mulated during the intraerythrocytic develop-ment of the parasites was related to hemoglobindestruction by the plasmodia. Therefore, at leastin theory, characterization of hemozoin shouldprovide clues to the ability of the parasites toutilize hemoglobin. Some investigators believedthe pigment to be melanin (reviewed in reference137); later, however, it was shown that hemozoincontained ferriprotoporphyrin coupled to a de-natured polypeptide or protein (reviewed in ref-erences 90, 95). The nature of the polypeptide


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moiety has been the subject of much research,and the findings vary in their interpretation; onegroup believed that the protein was a partiallydegraded hemoglobin (229), whereas other in-vestigators suggested that the protein was syn-thesized by the plasmodia (91, 92). Recent ex-periments on the malarial pigment of P. lo-phurae provide evidence that this hemozoin con-sists of insoluble monomers and dimers of hem-atin, that some of it consists of ferriprotopor-phyrin coupled to a plasmodial protein, and thatinsoluble methemoglobin is also present (287).Hemozoin is not a partially degraded form ofhemoglobin. It is difficult to state with certaintythat the extracted pigment is truly identical tothe hemozoin formed in situ within parasite foodvacuoles, since hematin is known to bind avidlyto many proteins, and during extraction somecytoplasmic constituents may be released whichcan be bound to the pigment adventitiously.The functional significance of hemozoin is not

completely understood. However, it has beenproposed that plasnodial formation of insolubleferriprotoporphyrin polymers enables the para-sites to sequester in a benign form the excessheme derived from their digestion of hemoglo-bin, and the processing of hemoglobin in thisway may also reduce the untoward effects ofoxygen on the plasmodia. -

It has been estimated that an average intra-erythrocytic plasnodium destroys between 25and 75% of the host cell hemoglobin (76); duringthis process there is the release of amino nitro-gen and the deposition of insoluble heme (he-mozoin). When radiolabeled erythrocytes weretransfused into P. lophurae-, P. knowlesi-, andP. berghei-infected hosts and parasites were per-mitted to invade and grow in these tagged eryth-rocytes, the amino acids in the radioactive he-moglobin became incorporated into plasmodialproteins (73, 223, 247). Electron microscopyshows that the destruction of erythrocytic he-moglobin takes place within the food vacuoleswhich are pinched off from the base of thespecial ingestive organelle of the parasites, thecytostome (2, 8). Involved in the degradation ofhemoglobin are parasite proteases which areprobably secreted into these food vacuoles.Moulder and Evans (153) identified a proteolyticenzyme in extracts of P. gallinaceum with a pHoptimum of 6.5, and Cook et al. (49) and Cooket al. (50) reported that extracts of P. bergheiand P. knowlesi readily hydrolyzed denaturedhemoglobin. A major alkaline protease (pH 7 to8) and a minor acid protease (pH 4 to 5) weredetected in both of these species. More recently,Levy and co-workers (125-128) and Chan andLee (40) have identified a cathepsin D-like pro-


teinase from P. berghei, P. knowlesi, and P.falciparum. The enzyme is probably membranebound or enclosed in membranous vesicles, sinceactivity was enhanced by the presence of TritonX-100. Although claims for parasite-specific pro-teases have been made, there is still some reasonto believe that some of the measured activitiesmay be due to host cell contaminants; the sen-sitivity to inhibitors and pH optima is not strik-ingly different for plasmodial and host cell-de-rived enzymes. Furthermore, based on the re-sults of Levy et al. with inhibitors, it seemsprobable that the plsmodial extracts containednot a single enzyme, but a complex containingseveral proteases.Where are these plasmodial proteases synthe-

sized, and how are they delivered to the foodvacuoles? Little is known concerning the occur-rence of lysosomes in malarial parasites. Indeed,evidence for lysosomal bodies is restricted to themorphological study of P. falciparum by Lan-greth et al. (121) and the positive cytochemicalreaction for acid phosphatase demonstrated inP. gallinaceum and P. berghei (9). The reactionproduct for acid phosphatase was also detectedin the endoplasmic reticulum of the parasites,which suggests that lysosomal enzymes may betransferred either directly from the endoplasmicreticulum to the food vacuoles or indirectly vialysosomal vesicles.Mechanisms of protein synthesis. It is

known in considerable detail for eucaryotes andprocaryotes that amino acids, ribosomes, solublefactors (enzymes, transfer RNA, messengerRNA, etc.), and energy are required for thefabrication of proteins.The available evidence indicates that for P.

knowlesi (216), P. berghei (100, 101), and P.lophurae (213) the molecular mechanisms in-volved in plasmodial protein synthesis are typi-cally eucaryotic; protein synthesis is sensitive tocycloheximide and puromycin, but not to chlor-amphenicol, the optimum Mg2e concentration is5 to 7 mM for an endogenous cell-free protein-synthesizing system, and the parasite ribosomeshave a sedimentation constant of 80S and canbe dissociated into 60S and 40S subparticles (51,218). The plasmodial ribosomal RNA is dis-tinctly protozoan in its base composition (seeabove), and the ribosomal RNA's (218, 266, 281)are distinctive in sedimentation constant andelectrophoretic mobility.

All species of malaria investigated to dateappear to synthesize their proteins by utilizingtheir own metabolic machinery and do not de-pend upon host cell ribosomes for the biogenesisof plasmodAl ribosomes.Histidine-rich protein. The only well-char-


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acterized protein from a malarial parasite is thehistidine-rich protein (HRP) isolated from mem-brane-bound cytoplasmic granules of P. lo-

phurae (104). This acid-soluble protein with anestimated molecular weight of 35,000 to 40,000is particularly rich (-73%) in histidine residues(104). The protein is synthesized in largeamounts by P. lophurae, and there is evidenceto suggest that the HRP may be a component ofthe polar organelles (111). The P. lophurae HRPaltered the morphology of normal erythrocytesby inducing invaginations (106, 107), and Kile-jian (108) successfully protected ducklingsagainst ordinarily fatal infections of P. lophuraeby prior administration of HRP.Evidence of an indirect nature has been pre-

sented for the presence of an acid-soluble HRPin P. falciparum (110). The P. falciparum HRPhad a higher molecular weight (-55,000) bysodium dodecyl sulfate-polyacrylamide gel elec-trophoresis than did the protein obtained fromP. lophurae.

If the HRP is the material responsible fortriggering erythrocyte endocytosis of merozoites(and as yet this has not been proven), then itmay be possible to develop protection againstmalaria by vaccination with this protein as an-

tigen.

Lipid BiosynthesisLipids constitute a significant fraction of the

total solids of malaria parasites (87, 137), andthe plasmodia tend to be richer in phospholipidsthan their hosts; approximately 80% or more ofthe fatty acids of the parasite total lipids are

unsaturated (reviewed in reference 87). By whatmechanisms are the parasites able to fabricatesuch distinct classes of lipids?The synthesis of long-chain fatty acids from

acetate in the extramitochondrial compartmentof eucaryotic cells requires three enzymaticsteps: (i) acetate activation to form acetyl-CoA;(ii) carboxylation of acetyl-CoA to malonyl-CoAby the enzyme acetyl-CoA carboxylase; and (iii)serial condensation of malonyl-CoA to the elon-gating acyl-CoA chain. To date, direct evidencefor the presence (or absence) of these enzymaticreactions in Plasmodium has not been obtained.Therefore, what little is known about lipid bio-synthesis in malaria parasites is derived fromrather indirect and often seriously compromisedobservations.Although P. fallax growing in turkey eryth-

rocytes incorporated ['4C]acetate and "C-la-beled fatty acids (oleic, palmitic, and stearicacids) into phospholipids (primarily phosphati-dylethanolamine), there was no critical exami-nation as to whether this incorporation repre-

MICROBIOL. REV.

sented de novo biosynthesis (77). Indeed, Cene-della (37) found that despite the rapid incorpo-ration of carbon from radioactive glucose intothe phospholipid fraction of P. berghei-infectedcells, less than 5% of the label was in the fattyacid portion and the vast majority was recoveredin the glycerol moiety. Rock (185, 186) per-formed similar experiments with P. knowlesi.He found that 14C-labeled pahnitic, oleic, andstearic acids were readily incorporated during a2-h in vitro incubation by infected cells and freeparasites, whereas uninfected monkey erythro-cytes incorporated less than 2% of the extracel-lular label into lipids. A total of 80 to 90% of theincorporated fatty acids was in the phospholipidfraction, primarily in phosphatidylethanola-mine, phosphatidylcholine, and phosphatidyli-nositol; this was in marked contrast to the lowrates of incorporation oflabeled acetate, glucose,and glycerol in the acyl portion of the lipid.Additionally, [32P]orthophosphate incorporationinto P. knowlesi was 80 to 100 times higher thanin erythrocyte membranes; incorporation wasprimarily into phospholipids in the parasites,whereas in the host cells it was into phosphatidicacid. The high rates of fatty acid incorporationand the low 32P-labeling pattern suggested thata rapid turnover of phosphorus rather than a netsynthesis of parasite phospholipids occurred(188). When parasites were exposed to [14C]cho-line, incorporation was into phosphatidylcho-line, whereas with [14C]ethanolamine the labelwas recovered in phosphatidylethanolamine;over 95% of the "4C fr6m glucose was recoveredin the glycerol moiety of the parasite phospho-lipids. If normal and P. knowlesi-infected eryth-rocytes were incubated in vitro with labeledacetate, mevalonate, and cholesterol, only la-beled cholesterol was recovered from the para-site membranes (263). (Confirmation of thesemetabolic studies comes from work [231, 263] inwhich it was shown that stearic acid and choles-terol enhanced the in vitro growth of P. know-lesi.)Thus, it appears that although members of

the genus Plasmodium do synthesize lipids, theyare incapable of fabricating fatty acids de novo.Through the incorporation of acetate into exist-ing fatty acids and by means of limited chain-lengthening reactions, the parasite is able tomaintain a lipid fatty acid composition distinctfrom that of its host. Indeed, the parasite lipidsare particularly enriched (-90%) in unsaturatedfatty acids (those containing zero, one, or twodouble bonds), whereas in erythrocytes theseacids comprise less than 75% of the total lipids(Table 3). Since intracellular parasites are un-able to synthesize cholesterol or fatty acids and


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their capabilities for desaturation are restricted,it appears that the parasites are able to satisfya portion of their requirements for phospholip-ids, lysophospholipids, cholesterol, fatty acids,and phospholipid precursors (i.e., as glycerol,inositol, choline, and phosphorus) by participat-ing in dynamic exchanges with components ofthe blood plasma during the turnover of eryth-rocyte lipids. In support of this view are thefindings that there are alterations in the mem-brane phospholipids of nonparasitized erythro-cytes in P. knowlesi-infected animals (10) andthat changes in the composition of the erythro-cyte membranes of P. lophurae-infected duck-ling erythrocytes (18) also take place. Addition-ally, by endocytic feeding of the parasites boththe membrane lipids of the parasites as well asthose of the host cells become available forplasmodial lipid biosynthesis.

Vitamins and CofactorsVitamin A. A dietary deficiency of vitamin A

was reported to depress infections ofP. lophuraein chicks (189) and P. berghei in rats (63). How-ever, in another study vitamin A deficiency pro-duced a more acute P. berghei infection (22). Itis not clear from the available evidence whetherthe vitamin deficiency directly affected parasitegrowth or interfered with the nutrition of thehost.Vitamin B1 (thiamine). Thiamine is a pre-

cursor ofthe coenzyme thiamine pyrophosphate,which is involved in many key decarboxylationreactions. Thiamine deficiency is reported todepress P. berghei infections (176), and Singer(234) showed that P. berghei-infected erythro-cytes contained more thiamine than normalcells. It appears that plasmodia require thia-mine, but in what formn and whether the parasitesynthesizes it de novo or the host supplies ithave not been determined.Vitamin B2 (riboflavin). A host deficiency

of riboflavin decreased the severity of the courseof P. lophurae (204) and P. gallinaceum (177)infections of chickens. This vitamin, the precur-sor for the coenzymes flavin adenine dinucleo-tide and flavin mononucleotide, was reported tobe present in P. knowlesi (13, 137); characteri-zation and specific roles in other species havenot been clarified.

Biotin. The first report of the influence of aspecific vitamin on the course of development ofPlasmodium involved biotin. In birds made se-verely deficient in biotin, P. lophurae infectionswere of high intensity, suggesting that the defi-ciency impaired the immune response of thehost (reviewed in reference 259). However, inducklings made moderately deficient in biotin P.


cathermerium infections progressed slowly atfirst, but later the parasitemia was considerablyhigher than in birds maintained on a diet havingadequate amounts of biotin. It has been con-cluded that these results show that biotin isrequired by the parasite for growth, as well asby the host for developing resistance to theparasite. There are, however, conflicting reportson the in vitro effects of biotin. Siddiqui et al.(reviewed in reference 259) claimed that theaddition of biotin favored the growth ofP. know-lesi in monkey erythrocytes; however, with P.lophurae there was no effect, and blood frombiotin-deficient monkeys did not adversely affectthe in vitro growth of P. knowlesi (137).Nicotinic acid (niacin), nicotinamide, and

pyridine nucleotides. Direct synthesis of nic-otinamide from nicotinic acid does not occur inanimals. In human erythrocytes niacin (nicotinicacid) first reacts with 5-phosphoribosyl-1-pyro-phosphate to yield nicotinic acid mononucleo-tide, and this is subsequently converted to NADby reaction with glutamine and ATP. Erythro-cytes can also make nicotinamide mononucleo-tide, which can then form NAD by reaction withATP; the enzyme catalyzing these transforma-tions (NAD phosphorylase) is localized in thenucleus ofthe cell. In chicken erythrocytes, how-ever, nicotinamide and nicotinic acid wereequally effective for the fornation of NAD.NADP is formed by the enzymatic phosphoryl-ation of NAD.A deficiency of nicotinic acid in the diet of

chicks produced a depression of parasitemiawith P. lophurae (189), and in vitro P. lophuraerequired a high level of nicotinamide for goodextracellular growth (259).The presence ofNAD in P. gallinaceum (239)

and the pyridine contents of P. berghei- and P.lophurae-infected erythrocytes have been re-ported (154, 211). In both P. lophurae and P.berghei striking increases in the total pyridinenucleotides of infected cells were observed. In P.berghei the reduced forms increased ninefold,whereas the oxidized forms increased only abouttwofold. Since NAD and NADP were not indi-vidually measured and because P. berghei in-vades reticulocytes which have the capacity forpyridine nucleotide synthesis, it is possible thatsuch increases reflect host cell stimulation aswell as the synthetic capabilities of the parasiteitself. However, in P. lophurae (a parasite ofmature erythrocytes), the NAD, reduced NAD,and NADP levels were increased only twofold ininfected erythrocytes, and during infection theNADPH content remained unchanged. The re-duced NAD, NADPH, and NADP contents offree P. lophurae were equivalent in amount to


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those found in duckling erythrocytes, whereasthe NAD content.of the parasite was 1.5 timesgreater (211). It was med that the parasitesfabricated their own pyridine nucleotides sincethe total content of the P. lophurae-infectedcells could be arrived at by simply summing thecontent of the free parasites with that of theuninfected erythrocytes. Although utilization ofNADPH has been suggested for P. berghei (61),direct evidence for this is lacking (see above).Ascorbic acid. A dietary deficiency of vi-

tamin C (ascorbic acid) in rhesus monkeysgreatly inhibited the multiplication of P. know-le8i, and administration of this vitamin reversedthe effect (137). However, since absence of as-corbic acid in the in vitro culture medium hadno effect on parasite growth, it was assumed thatthis vitamin acted indirectly via the host; noevidence for a change in the ascorbic acid con-tent in the adrenal glands of chicks infected withP. gallinaceun was detected despite adrenalhypertrophy (137).Vitamin E. The effect of vitamin E on the

course of a malarial infections was tested indi-rectly-, the addition of vitamin E was found toreverse the depressive effect of a cod liver oildiet on P. berghei in mice (75) and P. gaUina-ceum in chicks (244).Vitamin B, group. Pyridoxine, pyridoxal,

and pyridoxamine are essential for cellular me-tabolism. Seeler (203) found that massve dosesof pyridoxine reversed the antimalarial activityof quinine and quinacrine against P. lophurae,and Ramakrshnan (175) noted that if rats wereplaced on a pyridoxine-deficient diet, P. bergheiinfections were low. Trager (259) suggested thatthe lower activity of pyridoxine kinase in theerythrocytes of Africans might exert a selectiveadvantage in such individuals if the parasitelacked this enzyme. However, Platzer and Kassis(171) found pyridoxine kinase in P. lophurae,suggesting that plasmodial metabolism of vi-tamin Be was independent of the host cell. Theparasite enzyme had a lower affinity for pyridox-ine than the host cell enzyme; thus, if the hostwere limited in the availability of pyridoxine,the parasite would be deprived of this essentialcofactor since the pyridonine must first cross thehost cell cytoplasm before it can enter the plas-modium.Pantothenate. Pantothenate, when added in

vitro to P. lophurae-infected erythroytes,maintained parasite infectivity and the viabilityof male gametocytes. In vivo studies with blood-induced P. gallnceum infections in chickensshowed that parasitemias were lower in panto-thenate-deficient birds (259). Later, the role ofpantothenate in the growth of the parasites be-

MICROBIOL. REy.

came clearer. Trager found that pantothenate,a precursor of CoA, had no beneficial effectswhen it was added to extracellular P. lophurae,only CoA would lengthen the survival of theparasites (254, 259). CoA could not be replacedby phosphopantothenoyl cysteine, althoughsome restoration of extracellular growth tookplace with phosphopantetheine and completegrowth was achieved when the erythrocyte ex-tract-containing medium was supplementedwith dephospho CoA.Trager hypothesized that the parasite re-

quired CoA, but was unable to synthesize thisfrom pantothenate. Because the parasite wasentirely dependent on the host to supply theintact coenzyme, Trager suggested that the plas-modia were obligately parasitic. This hypothesishas been supported by the finding that althoughall of the enzymes in the biosynthetic pathwayfrom pantothenate to CoA were present in eryth-rocytes, none could be found in the free parasites(259). The beneficial effects of phosphopunte-theine and dephospho CoA on the extracellulargrowth of P. lophurae can be ascribed to theconversion of these compounds by erythrocyteenzymes present in the erythrocyte extract ofthe culture medium and not by plsmodial ac-tivity. Indeed, the fact that the CoA activity ofthe host cell declined as parasite growth in-creased supports the argument that the parasitesdestroy rather than contribute to the host cellconstituents. Also, since the CoA content of theliver of the host declines during the infection(233), it may be that the CoA required by themalarial parasites is synthesized in other organsand transported to the erythrocytes.CoA is required for the oxidation of glucose

via the citric acid cycle and for the syntheses ofcellular constituents by acetylation reactions.Therefore, a deficiency in its availability couldimpair parasite growth. The antimetabolite ef-fects of pantothenate analog on the in vitrointracellular growth of P. fakiparum and P.coatei (259) are undoubtedly related to this.Vitamin K and ubiquinones. Coenzymes Q

(CoQ) are components of the electron transportsystem ofmitochondria and therefore are criticalto cellular energy production via aerobic means.Despite there being limited informaton con-cerning mitochondrial function in plasmodia, ithas been speculated that the antimalarial activ-ity of certain naphthoquinones could be relatedto structural similarity to vitamin K or ubiqui-none or both (279). An extensive search forvitamin K and its synthesis from shikimic acidproved to be fhitless. However, ubiquinones 8and 9 (CoQs and CoQ9) were identified in P.lophurae-infected duck blood, but only CoQ1o


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was found in P. knowlesi, P. cynomolgi, and P.berghei (184, 198,236,237). Although CoQio andCoQ9 as well as CoQ8 occur in the blood ofnormal rhesus monkeys and CoQ9 and CoQs arefound in mouse blood, the predominance ofCoQ8in the samples of infected blood suggests thatthis coenzyme is characteristic of the parasite.However, it is known that ubiquinones differentfrom those found in normal blood do arise withvirally induced cancers (35) and that leukocytesdo synthesize CoQ, so that the assignment ofCoQ8 to the plasmodia is not without equivoca-tion.Of the CoQ analogs tested for antimalarial

activity, only a few alkylmercaptoquinones werefound to be effective (279). Although these com-pounds do resemble CoQ8, the biochemical basisfor their antimalarial action is far from estab-lished; it is suspected that their activity is notsolely confined to the inhibition of CoQ8 activityor its biosynthesis. Based on inhibitor studies, itwas suggested that CoQ may be involved in abranched electron chain (see above). Furtherevidence for this scheme, however, is needed.

Folates. (i) Biosynthesis ofdihydrofolate.The observation in 1940 that sulfanilamide couldbe used in the treatment of human malarialinfections was the first piece of evidence that 4-aminobenzoic acid (pABA) played a role in thegrowth of malarial parasites (46). Shortly there-after, it was shown that the sulfanilamide effecton P. gallinaceum could be reversed by pABA(reviewed in reference 64). Nutritional studiesamply confirmed the plasmodial requirement forpABA; the in vitro growth of P. knowlesi wasimproved by adding pABA to the medium, par-asitemias in P. berghei-infected rats and P.cynomolgi-infected monkeys were suppressedwhen the hosts were maintained on a milk dietwhich was low in pABA, Aotus monkeys kepton a milk diet did not support good growth of P.falciparum, and in mice the severity of a P.berghei infection was directly related to thepABA level of the diet (64). Although somevariations in the response of malarial infectionsto a milk diet in the host have been reported, asnoted by Ferone (64) and Nowell (160), thesemay simply reflect differences in the abilities ofmalaria strains to utilize small amounts ofpABAand the abilities of certain hosts to maintain anadequate blood level of pABA despite a dietarydeficiency.The investigations on milk diet, sulfonamides,

and pABA strongly suggested that malarial par-asites synthesized their folate cofactors de novo

and did not utilize exogenously supplied, intactfolate molecules as did their hosts. Thus, sulfa-nilamide, sulfaquinaxoline, sulfadimethoxine,


and dapsone (analogs of pABA) all act as anti-malarials by preventing the synthesis of dihy-dropteroate, an intermediate in the formnation oftetrahydrofolate (THF) (Fig. 6).

Further confirmation of the folate biosyn-thetic pathway (Fig. 6) in malarial parasitescame from the identification of the enzymesinvolved in the formation of dihydrofolate andTHF. Extracts of P. chabaudi, P. berghei, P.knowlesi, P. lophurae, and P. gallinaceum con-tain dihydropteroate synthetase (64, 274, 275),which catalyzes the formation of dihydroptero-ate from pteridine pyrophosphate; also, in ex-tracts of P. chabaudi and P. berghei, 2-amino-4-hydroxy-6-hydroxymethyl-dihydropteridinepyrophosphokinase (the enzymne that directlyprecedes dihydropteroate synthetase in thepathway) was also present. Both of these en-zymes were separable by chromatographicmethods in P. berghei (64), whereas the P. cha-baudi pyrophosphokinase activity could not beisolated from the synthetase activity (275). Thedifferences may be due to technical proceduresrather than reflect a significant qualitative di-vergence.

Despite evidence for a biosynthetic pathwayfrom pABA, glutamate, and guanosine triphos-phate to dihydrofolate, gaps in our understand-ing still remain. For example, what is the sourceof the pteridine that is ultimately converted topteridine pyrophosphosphate? Is this synthe-sized by the parasite from host pABA with theaddition of glutamate and guanosine triphos-phate, or is it obtained by cleavage ofpreexistingfolates? Why has it been impossible to detectactivity of dihydrofolate synthetase (the enzymewhich catalyzes the formation of dihydrofolatefrom dihydropteroate) in plasmodial extracts?

GTP + pABA+ glutamate

pter idine

pyrophosphokinose

pteridine pyrophosphate

sulfanilamide block --- tdihydropteroate synthetase

dihydropteroatelutamate

? dihydrofolate synthetase Present only in hostFolate

NADP NADPHdihydrofolate o r

NADPH folate reductaseNADP 4 dihydrofolate reductose

tetrahydrofolate

FIG. 6. Synthesis of folate in Plasmodium and itshost. GTP, Guanosine triphosphate.


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(ii) Dihydrofolate reductase (tetrahydro-folate dehydrogenase). Dihydrofolate reduc-tase, a key enzyme in folate metabolism, was thefirst folate enzyme identified in plasmodia (65).It has been found in P. lophurae, P. berghei, P.chabaudi, and P. knowlesi (64, 168). In bothpyrimethamine-sensitive and normal strains ofvarious plasmodial species, the enzyme had amolecular weight of 100,000 to 200,000, which is5 to 10 times greater than the molecular weightsof the dihydrofolate reductases of avians, mam-mals, helminths, and bacteria, but similar to themolecular weights of the dihydrofolate reduc-tases reported for other protozoans.The substrate affinity of the plasmodial di-

hydrofolate reductase was two to three orders ofmagnitude greater than that of the host cellenzyme, and it is probably for this reason thatpyrimethamine as well as other antifols, such astrimethoprim, cycloguanil, and dihydrotriazines(which also bind to the enzyme), are effective asantimalarials. Certain pyrimethamine-resistantstrains of P. berghei and P. vinckei show in-creased amounts of reductase with a reduceddrug affinity, and it is possible that this accountsfor the induction of resistance (57). However,other mechanisms of resistance are known andcould be associated with resistant strains of P.falciparum.The activity of dihydrofolate reductase mark-

edly increased during the growth of P. chabaudi(273); maximum activity occurred during schi-zogony, and it was of interest to note that theschizont stage was the most susceptible to pyri-methamine (80).

(iii) Tetrahydrofolate metabolism. THFacts as an acceptor of one-carbon units, and theensuing THF cofactors participate in severalreactions, including the interconversion of cer-tain amino acids, the de novo synthesis of purinenucleotides, and the formation of the pyrimidinedeoxyribonucleotide thymidylic acid from deox-yuridylate. The pathway from deoxyuridylate tothymidylic acid is catalyzed by thymidylate syn-thetase, and N5,Nl0-methylene tetrahydrofolateserves as the methyl donor. In the course of thisreaction dihydrofolate is regenerated, but beforeit can participate again it must be reduced toTHF by dihydrofolate reductase.

Erythrocyte-free P. berghei, unlike host cells,cannot reduce exogenously supplied folate todihydrofolate, indicating that the plasmodiumlacks folate reductase (66). However, if P.berghei cells are supplied with dihydrofolate,they are capable of forming THFs since theyproduce substances that promote the growth ofPediococcus cerevesiae (66, 181). Platzer (167)reported that although serine hydroxymethyl-

MICROBIOL. REV.

transferase activity was increased in P. lo-phurae-infected cells, the activities of formyl-tetrahydrofolate synthetase and methylene tet-rahydrofolate dehydrogenase were decreased ininfected erythrocytes. Indeed, neither of the lat-ter two enzymes was demonstrable in extracts offree P. lophurae. The serine hydroxymethyl-transferase of P. lophurae was found to be cy-tosolic (169) and distinctly different from thehost cell enzyme in molecular weight, pH opti-mum, and thermostability (170). Based on thesefindings, as well as the presence of dihydrofolateand thymidylate synthetase in extracts of P.lophurae (271), P. chabaudi (277), and P.berghei (182), Platzer (168) proposed the exist-ence of a "thymidylate synthesis cycle" in plas-modia: dihydrofolate -- THF -* N5,N'0-meth-ylene tetrahydrofolate -- dihydrofolate. Such acycle (Fig. 7) conveniently accounts for the denovo biosynthesis of pyrimidines by the parasiteand supports the contention that malarial para-sites rely on salvage pathway enzymes for theirpurines and are incapable of synthesizing pu-rines de novo.

Recently, reactions involving the synthesis ofmethionine and participation of N5-methyl tet-rahydrofolate have been demonstrated in ma-laria-infected cells (119, 238). Thus, it is possiblethat in addition to the thymidylate synthesiscycle, another folate pathway exists. However,the enzymes involved in the fonnation of N5-methyl tetrahydrofolate have not been identi-fied, and the conversion rate of homocysteine tomethionine (Fig. 7) was low.

If malarial parasites synthesize THF de novoand require only pABA for the formation ofdihydrofolate, then how can one explain thebeneficial effects of added folic and folinic acid

homocysteine methionine

.------- --THF N methyl THF

Folate DHF SHMT ?/reductose J

T thymidylate 5 10DHF N ,N methylene THF

/ synthetase

FTHFST dTMP dUMP TMTHFDH

N10 formyl THF-- . 5,---N10N methenyl THF

FIG. 7. Folate metabolism in Plasmodium (basedon references 64, 119, 167, and 238). DHF, Dihydro-folate; SHMT, serine hydroxymethyltransferase;FTHFS, fornyltetrahydrofolate synthetase; dTMP,deoxyribosylthymine monophosphate; dUMP, deox-yuridine monophosphate; MTHFDH, methylene tet-rahydrofolate dehydrogenase.


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(N5-fornyl tetrahydrofolate) on plasmodiagrowing intracellularly and extracellularly (232)and the finding that these materials and pABAreversed the inhibition of DNA synthesis bysulfalene but not pyrimethamine? The simplestexplanation is that pteridines or 4-aminoben-zoyl glutamate was a contaminant in the samplesof folic or folinic acid or that the host itself orthe media which contained host enzymes wereable to break down the added folates into pABA,and then this was converted by the parasite toTHF (64, 167).

Thus, it appears that malarial parasites do notuse exogenous folates supplied from the host cellbecause they are impermeable to these mole-cules and because they lack the appropriateenzymes for conversion to utilizable THF cofac-tors (64).

Cation AlterationsOverman et al. (reviewed in references 135,

164) clearly showed that a profound cationicimbalance existed in the plasma and erythro-cytes of monkeys infected with P. knowlesi.Similar findings have been reported for P. gal-linaceum in chickens (163), P. lophurae in duck-lings (223), P. coatneyi in monkeys, P. falcipa-rum in chimpanzees, and P. berghei in hamsters(58, 59). The most significant increases in potas-sium occur in the plasma, whereas in erythro-cytes the most striking change is an elevation inthe level of sodium. Dunn (58) has studied so-dium alterations in some detail and found thatthe erythrocyte sodium levels increased dispro-portionately to the parasitemia, that these in-creases persisted for several days after drug ther-apy had eradicated the infections, and that non-parasitized as well as parasitized erythrocyteswere abnormal in their capacity for sodium ex-trusion. The ATP content of the erythrocytesalso increased two to three times, and this par-alleled the increases in parasitemia and intracel-lular sodium.To account for these findings, Dunn postu-

lated that the elevated sodium levels were aconsequence of a circulating toxin that dimin-ished the efflux of sodium while promoting itsinflux. However, cross-incubation experimentswith malarious plasma did not produce sodiumtransport changes identical to those found ininfected animals. Since the disruption of thecation gradient was found to be both gradualand unrelated to immunological phenomena, itseems more plausible to ascribe these events notto a toxin but to a modification of the membranelipid-protein architecture, which in turn affectsadenosine triphosphatase as well as other pro-


teins involved in the sodium pump. Membranechanges of this kind could also contribute to theosmotic fragility of nonparasitized cells in ma-laria-infected hosts.

Precautions, Prospects, and PossibleConclusions

No account of the biochemistry of Plasmo-dium would be complete without some mentionof the inherent pitfalls and problems encoun-tered when working with these organisms.At various times investigators have neglected

to include proper and extensive controls, so thatit is not always clear whether the parameterbeing examined was due to a parasite constituentor a contaminant. Work of this sort obviouslymust be regarded with a degree of skepticism.However, contaminants can be eliminated ortheir contribution can be evaluated, and thisshould be standard for all work. Future research-ers will have to use all available morphologicaland biochemical tools to ensure that the datatruly reflect the capacities of the parasite or thehost-parasite complex and not of some extra-neous materials.

Still, even if rigorous purification of parasiteswere to become routine, biochemists workingwith malarial parasites removed from the hostcells must demonstrate that the parasites in thecell suspension have remained intact. Indeed, ithas been our experience in recent years to findthat parasites liberated from erythrocytes by thehemolytic antiserum-complement method, aswell as parasites freed by saponin lysis of in-fected erythrocytes, leak macromolecules (286).Presently, we remain uncertain whether thisleaky condition is an artifact of the in vitrorelease technique or whether such a permeabil-ity property exists in vivo. Also, we are unableto ascertain whether the leakiness involves allcells to a lesser or greater degree or is restrictedto specific stages in the growth cycle. For somekinds of biochemical analyses, such difficultiesmay not pose a serious hazard in interpretingthe results, but where the studies involve themetabolism of substrates added to suspensionsof free parasites, it could prove to be a seriousliability. Now that we are aware of an inherentdeficiency in the currently employed methodsfor release of plasmodia, it is ixnperative thatnew techniques be developed, ones that giveassurances that each parasite has been removedcleanly and without damage to its limiting mem-branes. Possibly no such technique can be de-vised because the intracellular plasmodia trulyhave a "permeability defect"-a consequence oftheir evolutionary adaptation to the intracellular


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mode of life. Nevertheless, it should be standardprocedure for researchers to identify the meta-bolic alterations of added substrates which area direct result of enzymes that are released bydamaged parasites, before conclusions are drawnas to utilization of a specific substance.

It was once assumed that the biochemicalbasis for parasitism could be uncovered by grow-ing Plasmodium extracellularly in vitro underaxenic conditions or preferably in defined media.By providing or eliminating nutrients in culture,it was felt that a determination could be maderegarding the critical materials which were sup-plied by the host. This goal has been achievedto a limited degree for only one species, P.lophurae. The in vitro extracellular cultivationof P. lophurae from a uninucleate trophozoite toa multinucleate schizont has been a significantaccomplishment (253, 254, 256) and has providedmuch insight into the nature of parasitism bymalarial parasites. It has, for example, been ofextreme importance in identifying the CoA le-sion. However, it should be recognized that suchcultures are not without their limitations. Thegrowth of P. lophurae extracellularly requires acomplex medium, an essential component ofwhich is an extract of duck erythrocytes. Thiserythrocyte extract is a cornucopia of proteins,enzymes, coenzymes, substrates, and metabolicintermediates. As a consequence, the addition ofa substance and the assessment of its beneficialor detrimental effect on the growth and repro-duction of the parasite cannot be construed apriori to be simply a result of that added sub-stance. Cause (the substance) and effect(growth, etc.) may not be tightly linked underthese special conditions, and until the possibletransformations that occur to the substance areidentified, it behoves investigators to draw con-clusions carefully regarding growth-promotingroles. Thus, the benefits of adding folinic acid orATP to extracellular P. lophurae may not besolely due to these compounds, but could reflectsome product formed in the enzyme-rich me-dium. Then too, there exists the possibility thatit is not always the major ingredient listed onthe label of a particular biochemical that is atwork, but that the effects obtained are due to atrace contaminant. A case in point is the possiblepresence of contaminants in preparations of leu-covorin, which led to confusion of the role offolates in the nutrition of the parasites.Although extracellular cultivation has its at-

tendant pitfalls, suspensions of malaria-infectedcells may also cause difficulties to which inves-tigators are sometimes oblivious. Often malar-iologists have recognized how reticulocytes mayaffect the measured activities of infected cells,

MICROBIOL. REV.

but they have less frequently appreciated the invitro and in vivo fragility of the parasitized aswell as the unparasitized erythrocytes obtainedfrom heavily infected hosts. Such cells, noto-riously leaky, may release substances into themedium which affect a tagged substrate that hasbeen added to follow a particular pathway. In-deed, in the desire to obtain sufficient materialsfor analysis, very often virulent parasites main-tained in convenient but abnormal hosts areroutinely used. Results obtained by using suchmaterial may not always be representative ofwhat takes place in natural infections, althoughwe consistently favor the notion that extrapola-tions are proper.Biochemical investigations on Plasmodium

are not for the faint of heart, and an awarenessof the potential artifacts should provide a meansfor successfully reducing the gaps in our existingknowledge.This said, what may be concluded from this

review of the biochemistry of Plasmodium?Successful invasion of erythrocytes by mero-

zoites involves specific binding of the parasitesto a surface receptor and induction of endocy-tosis. The Duffy factor on human erythrocytesappears to be involved in junction formationwith merozoites of P. knowlesi and P. vivax. Aconsequence of this is that Duffy-negative eryth-rocytes are refractory to invasion by these spe-cies. Susceptibility studies show that the eryth-rocyte receptor for P. falciparum differs fromthat for P. vivax. How receptors promote mer-ozoite attachment and how merozoite bindingtriggers erythrocyte endocytosis are problems inneed of solution. Chemical characterization oferythrocyte receptors, as well as characterizationof the constituents in the polar organelles of themerozoite, would contribute to our understand-ing of the mechanism of erythrocyte invasion.

Malarial parasites may alter the erythrocytesurface by inducing morphological changes (i.e.,knobs and caveola-vesicle complexes), modify-ing existing membrane proteins and lipid com-position, and inserting neoproteins. The exactmechanisms whereby such parasite-provokedchanges come about is uncertain. Normal eryth-rocytes differ from malaria-infected cells andisolated parasites in lipid composition. In partic-ular, the lipids of Plasmodium are enriched inphospholipids and unsaturated fatty acids (es-pecially octadecenoic acid). Plasmodium is in-capable of de novo synthesis of fatty acids andcholesterol, and its distinct lipid characteristicsare maintained by the parasite acting as a met-abolic sink, as erythrocytes participate in dy-namic exchanges with the blood. The alteredlipid composition of infected erythrocytes could


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profoundly affect their osmotic and metabolicproperties.

Intraerythrocytically, bird and mammalianmalarias appear to derive energy by metaboliz-ing glucose to lactic acid via a conventionalpathway of anaerobic glycolysis. If supplied withoxygen, avian parasites may oxidize a portion ofthe pyruvate to C02 and water by means of thecitric acid cycle, whereas rodent and primatemalarias, which lack a functional citric acid cy-cle, are unable to do so and have lactate as theirprimary end product.

Erythrocyte-free P. gallinaceum cells pro-duce appreciable quantities of acetate from glu-cose and pyruvate. Since plasmodia are unableto synthesize CoA de novo, it is possible that inP. gallinaceum acetate formation is due to alack of host-supplied CoA. In P. knowlesi it maybe that volatile acids are formed by a pyruvateclastic reaction, but the enzymes involved (ifthey exist) have not been looked for. However,in view of the fragile nature of free parasites,acetate and formate production may reflect de-ranged metabolism due to in situ leakiness ofplasmodia or may be a consequence of insultduring the isolation procedure.There is no evidence for a pentose phosphate

shunt in malarial parasites since the first enzymein the pathway (G6PDH) is absent. Indeed, theonly enzyme in this pathway that has been iden-tified consistently is 6-phosphogluconate dehy-drogenase. Lacking a pentose shunt, the para-sites must have other means for obtaining riboseand reducing NADP. It has been suggested thataction by phosphorylases supplies the pentosesand that glutamic dehydrogenase provides forthe reduction of NADP.Evidence for an energy-yielding electron

transport chain is at best circumstantial. In allof the malarias studied, the only enzyme foundto be associated with this system is cytochromeoxidase. It is conceivable that in the acristaterodent malarias and in P. knowlesi, and perhapseven in those malarias having cristate mitochon-dria (avians and P. falciparum), cytochromeoxidase is involved in the de novo pyrimidinebiosynthetic pathway and not in energy-yieldingreactions.

Malarial parasites are incapable of de novo

purine biosynthesis; however, pyrimidines are

synthesized de novo. Exogenously supplied pu-

rines and orotic acid are transported and incor-porated by infected erythrocytes and plasmodia,whereas pyrimidines (uracil and thymidine) are

not. There is evidence to support the contentionthat hypoxanthine is the preferred purine of theparasites in vivo and that it is derived from thecatabolism of erythrocytic ATP. Plasmodia have


a distinctive DNA and rRNA base composition.Malarial parasite ribosomes are not provided forby host cell ribosomal subparticles, and themechanism of protein synthesis by the parasitesis typically eucaryotic.The capacity of the parasites for de novo

amino acid biosynthesis is limited, and it appearsthat host cell hemoglobin provides most of theamino acids. For some species, isoleucine andmethionine must be supplied exogenously forgood plasmodial growth. The degradation oferythrocyte hemoglobin by parasite proteasesleaves a golden brown-black residue called he-mozoin (malarial pigment). Hemozoin consistsof insoluble monomers and dimers of hematin,methemoglobin, and ferriprotoporphyrin cou-pled to plasmodial protein. The functional sig-nificance of hemozoin is not completely under-stood.The only well-characterized plasmodial pro-

tein is the HRP of P. lophurae. It is possiblethat HRP is localized in the polar organelles ofthe merozoites and is involved in the process ofinvasion.

Information regarding the vitamin require-ments of malarial parasites is scanty. Plasmodiaare incapable of synthesizing CoA from panto-thenate and rely on the host cell for this cofactor;this may be one reason for their being obligateintracellular parasites. By contrast, Plasmo-dium can synthesize folate from pABA.

Characteristically, malaria-infected erythro-cytes show an elevated sodium content due toan inability of the cells to extrude this cation.Although it has been postulated that this alter-ation is provoked by a circulating toxin, it is alsopossible to ascribe such a change to modifica-tions in membrane lipid-protein architecture.A fundamental goal for studying the biochem-

istry of Plasmodium is to uncover metabolicdifferences between host and parasite that couldbe exploited for designing drugs that would ex-terminate the parasites without injury to thehost. Several such examples have been alludedto and some cases clearly identified above. Plas-modium synthesizes folates de novo, whereashosts do not, and as a consequence antifols areeffective antimalarials. Particularly striking inthis regard is the finding that the moleculardifferences between the dihydrofolate reduc-tases of hosts and malarial parasites contributeto the exquisite sensitivity of the parasites topyrimethamine. Because Plasmodiwm is lackingin certain specific enzymes, the parasites areunable to synthesize CoA from pantothenate;however, the host can, and this difference couldprove to be of significance in the developmentof chemotherapeutic agents. Plasmodia fabri-


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cate a unique HRP that may be involved in hostcell invasion, and almost all of the enzymes ofthe parasite differ structurally from those of thehost cell. Here too, the differences provide forthe possibility of developing very specific par-asiticidal drugs. Indeed, the fact that the activesite of parasite (P. lophurae) lactic dehydrogen-ase is dissimilar from that of erythrocytes pro-vides encouragement that such uniqueness couldbe exploited in an antimalarial. Because theterminal stages of energy metabolism in theparasites appear to be distinct, possibly thiscould be used to inhibit parasite growth withoutdamage to the host. Furthermore, since the basecompositions of the rRNA's (and in some casesthe DNAs) of Plasmodium species are so strik-ingly different from those of the hosts whichthey parasitize, it is conceivable that base ana-logs might serve as effective antimalarials. Nodoubt other subtle, and as yet undiscovered,metabolic and structural differences exist be-tween host and parasite. It is to be hoped thatincreased research activity into the biochemicalbasis of parasitism will provide suitable "magicbullets" to effectively control this "O million-murdering Death."

ACKNOWLEDGMENTSI am indebted to W. E. Gutteridge, G. G. Holz, L.

H. Miller, E. G. Platzer, and L. W. Scheibel, who gavetheir time and expertise to review critically an earlydraft of this work. The literature on malaria is abun-dant, and in some cases I have had to resort to citingreviews or secondary sources. This was done solely torestrict the number of citations, and the absence of a

citation in no way implies diminished importance ofthe contribution of a researcher. The literature reviewwas completed in September 1979.

Research in my laboratory has been supported byPublic Health Service grants AI-05226 and AI-13422from the National Institute of Allergy and InfectiousDiseases.

LITERATURE CMD1. Aikawa, M. 1971. Plasmodium: the fine struc-

ture of malarial parasites. Exp. Parasitol. 30:284-320.

2. Aikawa, M. 1977. Variations in structure andfunction during the life cycle of malarial para-sites. Bull. W.H.O. 55:139-156.

3. Aikawa, M., and R. T. Cook. 1972. Plasmo-dium: electron microscopy of antigen prepara-tions. Exp. Parasitol. 31:67-74.

4. Aikawa, M., C. L. Hsieh, and L. Miller. 1977.Ultrastructural changes of erythrocyte mem-

brane in ovale-type malarial parasites. J. Par-asitol. 63:152-154.

5. Aikawa, M., L. H. Miller, J. Johnson, and J.Rabbege. 1978. Erythrocyte entry by malarialparasites. A moving junction between eryth-rocyte and parasite. J. Cell Biol. 77:72-82.

6. Aikawa, M., L. Miller, and J. Rabbege. 1975.

MICROBIOL. REV.

Cavelola-vesicle complexes in the plasma-lemma of erythrocytes infected by Plasmo-dium vivax and P. cynomolgi. Am. J. Pathol.79:285-294.

7. Aikawa, M., J. R. Rabbege, and B. T. Welide.1972. Junctional apparatus in erythrocytes in-fected with malarial parasites. Z. Zellforsch.Mikrosk. Anat. 124:72-75.

8. Aikawa, M., and C. Sterling. 1974. Intracellu-lar parasitic protozoa. Academic Press Inc.,New York.

9. Aikawa, M., and P. E. Thompson. 1971. Lo-calization of acid phosphatase activity in Plas-modium berghei and P. gallinaceum: an elec-tron microscopic observation. J. Parasitol. 57:603-610.

10. Angus, M. G. N., K. A. Fletcher, and B. G.Maegraith. 1972. Studies on the lipids ofPlas-modium knowlesi-infected rhesus monkeys(Macaca mulatta). IV. Changes in erythrocytelipids. Ann. Trop. Med. Parasitol. 65:429-439.

11. Bahr, G. F. 1966. Quantitative cytochemicalstudy of erythrocytic stages of Plasmodiumlophurae and Plasmodium bergehi. Mil. Med.131:1064-1070.

12. Bahr, G. F., and U. Mikel. 1972. The arrange-ment of DNA in the nucleus of rodent malariaparasites. Proc. Helminthol. Soc. Wash. 39:361-372.

13. Ball, E. G., R. W. McKee, C. B. Anfinsen, W.0. Cruz, and Q. M. Geiman. 1948. Studies onmalarial parasites. IX. Chemical and metabolicchanges during growth and multiplication invivo and in vitro. J. Biol. Chem. 175:547-571.

14. Bannister, L. H. 1977. The invasion of red cellsby Plasmodium. Symp. Br. Soc. Parasitol. 15:27-55.

15. Bannister, L., G. Butcher, and G. Mitchell.1977. Recent advances in understanding theinvasion of erythrocytes by merozoites of Plas-modium knowlesi. Bull. W.H.O. 55:163-170.

16. Barrett, J. 1976. Bioenergetics in helminths, p.67-80. In H. Van den Bossche (ed.), Biochem-istry of parasites and host-parasite relation-ships. Elsevier/North Holland BiomedicalPress, Amsterdam.

17. Bass, C. C., and F. M. Johns. 1912. The culti-vation of malarial plasmodia (Plasmodium vi-vax and Plasmodium falciparum) in vitro. J.Exp. Med. 16:567-579.

18. Beach, D. H., I. W. Sherman, and G. G. Holz,Jr. 1977. Lipids of Plasmodium lophurae, andof erythrocytes and plasmas of normal and P.lophurae-infected Pekin ducklings. J. Parasi-tol. 63:62-75.

19. Becker, Y. 1978. The chlamydia: molecular bi-ology of procaryotic obligate parasites of eu-caryotes. Microbiol. Rev. 42:274-306.

20. Bienzle, U., I. Guggenmoos-Holzmann, andL. Luzatto. 1979. Malaria and erythrocyte glu-cose-6-phosphate dehydrogenase variants inWest Africa. Am. J. Trop. Med. Hyg. 28:619-621.

21. Bishop, C. 1960. Purine metabolism in humanand chicken blood in vitro. J. Biol. Chem. 235:


mbr.asm

.org/D

ownloaded from


VOL. 43, 1979

3228-3232.22. Bouliset, L., and J. Ruffie. 1958. Course of P.

berghei malaria in white rats deficient in vi-tamin A. Ann. Parasitol. Hum. Comp. 33:209-217.

23. Bovarnick, M. R., A. Lindsay, and L. Heller-man. 1946. Metabolism of the malarial para-site, with reference particularly to the action ofantimalarial agents. I. Preparation and prop-erties of Plasmodium lophurae separated fromthe red cells ofduck blood by means ofsaponin.J. BioL Chem. 163:523-533.

24. Bovarnick, M. R., A. Lindsay, and L. Heiler-man. 1946. Metabolism of the malrial para-site, with reference particularly to the action ofantimaral agents. II. Atabrine (Quinacrine)inhibition of glucose oxidation in parasites ini-tially depleted of substrate. Reversal by aden-ylic acid. J. Biol. Chem. 163:535-551.

25. Bovarnick, M. R., and J. C. Miller. 1950. Oxi-dation and trnsamination of glutamate by ty-phus rickettsiae. J. Biol. Chem. 184:661-676.

26. Bowman, L. B. R., P. T. Grant, and W. 0.

Kermack. 1960. The metabolism of Plasmo-dium berghei, the malaria parasite of rodents.I. The preparation of the erythrocytic form ofP. berghei separated from the host cell. Exp.Parasitol. 9:131-136.

27. Bowman, L. B. R., P. T. Grant, W. 0. Ker-mack, and D. Ogston. 1961. The metabolismof Plasmodium berghei, the malaria parasiteof rodents. H. An effect of mepacrine on themetabolism of glucose by the parasite sepa-rated from its host cell. Biochem. J. 78:472-478.

28. Broun, G. 1961. Enzymes erythrocytaires et in-festation a Plasmodium berghei chez la souris.Rev. Fr. Etud. Clin. Biol. 6:695-699.

29. Bryant, C., A. Voller, and M. J. H. Smith.1964. The incorporation of radioactivity from[14C]glucose into the soluble metabolic inter-mediates of malaria parasites. Am. J. Trop.Med. Hyg. 13:515-519.

30. Bueding, E., and B. Charms. 1952. Cytochromec and cytochrome oxidase, and succinoxidaseactivities of helminths. J. Biol. Chem. 196:615-627.

31. Bungener, W. 1967. Adenosindeaminase andNucleosidphosphorylase bei Malariaparasiten.Z. Tropenmed. Parasitol. 18:48-52.

32. Bungener, W., and G. Nielsen. 1969. Nuklein-saurenstoffwechsel bei experimenteller Ma-laria. III. Einbau von Adenin aus dem Aden-innukleotidpool der Erythrozyten in die Nu-k1einsauren von Malariaparasiten. Z. Tro-penmed. Parasitol. 20:66-73.

33. Canale-Parola, E. 1977. Physiology and evolu-tion of spirochaetes. Bacteriol. Rev. 41:181-204.

34. Carter, R., and D. Walliker. 1977. Biochemicalmarkers for strain differentiation in malarialparasites. Bull. W.H.O. 55:339-345.

35. Casey, A. C., and E. G. Bliznakov. 1973. Co-enzyme Q levels in liver, spleen, and blood ofmice with Friend leukemia virus infection.


Cancer Res. 33:1183-1186.36. Ceithamn, J., and E. A. Evans, Jr. 1946. The

biochemistry of the malaria parasite. IV. Thein vitro effects of X-rays upon Plasmodiumgallinaceum. J. Infect. Dis. 78:190-197.

37. Cenedella, R. J. 1968. Lipid synthesis from glu-cose carbon by Plasmodium berghei in vitro.Am. J. Trop. Med. Hyg. 17:680-684.

38. Cenedella, R. J., J. J. Jarrell, and L H. Saxe.1969. Lipid synthesis in vivo from 1_14C-oleicacid and 6-3H-glucose by intraerythrocyticPlasmodium berghei. Mil. Med. 134:1045-1055.

39. Cenedella, R. J., H. Rosen, C. R. Angel, andL. H. Saxe. 1968. Free amino-acid productionin vitro by Plasmodium berghei. Am. J. Trop.Med. Hyg. 17:800-8.

40. Chan, V. L, and P. Y. Lee. 1974. Host-cellspecific proteolytic enzymes in Plasmodiumberghei infected erythrocytes. Southeast AsianJ. Trop. Med. Public Health 5:447-449.

41. Chance, M. L., H. Momen, D. C. Warhurst,and W. Peters. 1978. The chemotherapy ofrodent marlaria. XXIX. DNA relationshipswithin the subgenus Plasmodium (Vinckeia).Ann. Trop. Med. Parasitol. 72:13-22.

42. Chen, T. T. 1944. The nuclei in avian malariaparasites. I. The structure of nuclei in Plas-modium elongatum with some considerationson technique. Am. J. Hyg. 40:26-34.

43. Ciuca, M., A. G. Ciplea, C. Bona, N. Poszgi,T. Isfan, and G. luga. 1963. Etudes cytochi-miques sanguines dans l'infection experimen-tale avec Plasmodium berghei de la sourisblanche. I. Structure cytochimique du parasite,des globules rouges et observations effectueesau microscope a contraste de phase. Arch.Roum. Pathol. Exp. Microbiol. 22:503-515.

44. Clarke, D. H. 1952. The use of phosphorus 32 instudies on Plasmodium gallinaceum. I. Thedevelopment of a method for the quantitativedeternination of parasite growth and devel-opment in vitro. J. Exp. Med. 96:439-450.

45. Clarke, D. H. 1952. The use of phosphorus 32 instudies on Plasmodium gallinaceum. II. Stud-ies on conditions affecting parasite growth inintact cells and in lysates. J. Exp. Med. 96:451-463.

46. Coggeshall, L. T. 1940. The selective action ofsulfanilamide on the parasites of experimentalmalaria in monkeys in vivo and in vitro. J.Exp. Med. 71:13-20.

47. Collins, W. E., and H. Aikawa. 1977. Plasmo-dia of nonhuman primates, p. 467-492. In J. P.Kreier (ed.), Parasitic protozoa, vol. 3. Aca-demic Press Inc., London.

48. Conklin, K. A., S. C. Chou, W. A. Siddiqui,and J. V. Schnell. 1973. DNA and RNAsyntheses by intraerythrocytic stages of Plas-modium knowlesi. J. Protozool. 20:683-688.

49. Cook, L., P. T. Grant, and W. 0. Kermack.1961. Proteolytic enzymes of the erythrocyticforms of rodent and simian species of malarialplasmodia. Exp. Parasitol. 11:372-379.

50. Cook, R. T., M. Aikawa, R. C. Rock, W. Little,


mbr.asm

.org/D

ownloaded from


488 SHERMAN

and H. Sprinz. 1969. The isolation and frac-tionation of Plasmodium knowlesi. Mil. Med.134:866-883.

51. Cook, R. T., R. C. Rock, M. Aikawa, and M.J. Fournier, Jr. 1971. Ribosomes of the ma-larial parasite, Plasmodium knowlesi. I. Isola-tion, activity and sedimentation velocity.Comp. Biochem. Physiol. 39B:897-911.

52. Cooper, S. 1979. A unifying model for the GIperiod in prokaryotes and eukaryotes. Nature(London) 280:17-19.

53. Dasgupta, B. 1960. Polysaccharides in the dif-ferent stages of the life-cycles of certain spo-rozoa. Parasitology 60:509-514.

54. Deane, H. W. 1945. Studies on malarial para-sites. II. The staining of two primate parasitesby the Feulgen technique. J. Cell. Comp. Phys-iol. 26:139-145.

55. Deans, J. A., E. D. Dennis, and S. Cohen.1978. Antigenic analysis of sequential erythro-cytic stages of Plasmodium knowlesi. Parasi-tology 77:333-344.

56. De Zeeuw, R. A., J. Wijsbeek, R. C. Rock,and G. J. McCormick. 1972. Composition ofphospholipids in Plasmodium knowlesi mem-branes and in host rhesus erythrocyte mem-branes. Proc. Helninthol Soc. Wash. 39:412-418.

57. Diggens, S. M., W. E. Gutteridge, and P. I.Trigg. 1970. Altered dihydrofolate reductaseassociated with a pyrimethamine-resistantPlasmodium berghei berghei produced in asingle step. Nature (London) 228:579-580.

58. Dunn, M. J. 1960. Alterations of red blood cellsodium transport during malarial infection. J.Clin. Invest. 48:674-684.

59. Dunn, M. J. 1969. Alterations of red blood cellmetabolism in simian malaria: evidence for ab-normalities of nonparasitized cells. Mil. Med.134:1100-1105.

60. Dvorak, J. A., L. H. Miller, W. Whitehouse,and T. Shiroishi. 1975. Invasion of erythro-cytes by malaria merozoites. Science 187:748-750.

61. Eckman, J. R., and J. W. Eaton. 1979. De-pendence of plasmodial glutathione metabo-lism on the host cell. Nature (London) 278:754-756.

62. Eisen, H. 1977. Purification of intracellular formsof Plasmodium chabaudi and their interac-tions with the erythrocyte membrane and se-rum albumin. Bull. W.H.O. 55:333-338.

63. Fabiani, G., and P. Grellet. 1951. Etude chez lerat blanc des rapports entre la carence en vi-tamine A et le paludisme experimental a Plas-modium berghei. C. R. Soc. Biol. 146:441-444.

64. Ferone, R. 1977. Folate metabolism in malaria.Bull. W.H.O. 55:291-298.

65. Ferone, R., J. J. Burchall, and G. H. Hitch-ings. 1969. Plasmodium berghei dihydrofolatereductase. Isolation, properties, and inhibitionby antifolates. Mol. Pharmacol. 5:49-59.

66. Ferone, R., and G. H. Hitchings. 1966. Folatecofactor biosynthesis by Plasmodium berghei.Comparison of folate and dihydrofolate as sub-strates. J. Protozool. 13:504-506.

MICROBIOL. REV.

67. Fletcher, K. A., M. V. Canning, and R. D. G.Theakston. 1977. Electrophoresis of glucose-6-phosphate and 6-phosphogluconate dehydro-genases in erythrocytes from malaria-infectedanimals. Ann. Trop. Med. Parasitol. 71:125-130.

68. Fletcher, K. A., and B. G. Maegraith. 1962.Glucose-6-phosphate and 6-phosphogluconatedehydrogenase activities in erythrocytes ofmonkeys infected with Plasmodium knowlesi.Nature (London) 196:1316-1318.

69. Fletcher, K. A., and B. G. Maegraith. 1972.The metabolism of the malaria parasite and itshost. Adv. Parasitol. 10:31-48.

70. Forrester, L. J., and P. M. L. Siu. 1971. P-enolpyruvate carboxylase from Plasmodiumberghei. Comp. Biochem. Physiol. 38B:7345.

71. Fraser, D. M., and W. 0. Kermack. 1957. Theinhibitory action of some antimalarial drugsand related compounds on the hexokinase ofyeasts and Plasmodium berghei. Br. J. Phar-macol. Chemother. 12:16-23.

72. Friedman, M. J. 1979. Oxidant damage mediatesvariant red cell resistance to malaria. Nature(London) 280:245-247.

73. Fulton, J. D., and P. T. Grant. 1956. Thesulphur requirements of the erythrocytic formof Plasmodium knowlesi. Biochem. J. 63:274-282.

74. Ginger, C. D. 1967. Preliminary studies on thelipid constitution of the malaria parasite.Trans. R. Soc. Trop. Med. 61:2-3.

75. Godfrey, D. 1957. Antiparasitic action of dietarycod liver oil upon Plasmodium berghei and itsreversal by vitamin E. Exp. Parasitol. 6:555-565.

76. Groman, N. B. 1951. Dynamic aspects of thenitrogen metabolism of Plasmodium gallina-ceum in vivo and in vitro. J. Infect. Dis. 88:126-150.

77. Gutierrez, J. 1966. Effect of the antimalarialchloroquine on the phospholipid metabolismof avian malaria and heart tissue. Am. J. Trop.Med. Hyg. 15:818-822.

78. Gutteridge, W. E., D. Dave, and W. H. G.Richards. 1979. Conversion of dihydroorotateto orotate in parasitic protozoa. Biochim. Bio-phys. Acta 582:390-401.

79. Gutteridge, W. E., and P. L. Trigg. 1970. In-corporation of radioactive precursors intoDNA and RNA of Plasmodium knowlesi invitro. J. Protozool. 17:89-96.

80. Gutteridge, W. E., and P. I. Trigg. 1971. Actionof pyrimethamine and related drugs againstPlasmodium knowlesi in vitro. Parasitology62:431 444.

81. Gutteridge, W. E., and P. I. Trigg. 1972. Peri-odicity of nuclear DNA synthesis in the in-traerythrocytic cycle ofPlasmodium knowlesi.J. Protozool. 19:378-381.

82. Gutteridge, W. E., and P. I. Trigg. 1972. Somestudies on the DNA of Plasmodium knowlesi,p. 199-218. In H. Van den Bossche (ed.), Com-parative biochemistry of parasites. AcademicPress Inc., London.

83. Gutteridge, W. E., P. I. Trigg, and D. H.


mbr.asm

.org/D

ownloaded from


VOL. 43, 1979

Williamson. 1971. Properties of DNA fromsome malarial parasites. Parasitology 62:209-219.

84. Hegner, RI W., and M. S. MacDougall. 1926.Modifying the course of infections with birdmalaria by changing the sugar content of theblood. Am. J. Hyg. 6:602-608.

85. Herman, Y. F., R. A. Ward, and R. H. Her-man. 1966. Stimulation of the utili;ation of 1-'4C-glucose in chicken red blood cells infectedwith Plasmodium gallinaceum. Am. J. Trop.Med. Hyg. 15:276-280.

86. Hill, G. C. 1976. Characterization of the electrontransport systems present during the life cycleof African trypanosomes, p. 31-50. In H. Vanden Bossche (ed.), Biochemistry of parasitesand host-parasite relationships. Elsevier/North Holland Biomedical Press, Amsterdam.

87. Holz, G. G., Jr. 1977. Lipids and the malarialparasite. Bull. W.H.O. 55:237-248.

88. Holz, G. G., Jr., D. H. Beach, and L. W. Sher-man. 1977. Octadecenoic fatty acids and theirassociation with hemolysis in malaria. J. Pro-tozool. 24:566-574.

89. Homewood, C. A. 1977. Carbohydrate metabo-lism of malarial parasites. Bull. W.H.O. 55:229-235.

90. Homewood, C. A. 1978. Biochemistry, p. 170-211. In R. Killick-Kendrick and W. Peters (ed.),Rodent malaria. Academic Press Inc., NewYork.

91. Homewood, C. A., J. M. Jewsbury, and M. LChance. 1972. The pigment formed duringhaemoglobin digestion by malarial and schis-tosomal parasites. Comp. Biochem. Physiol.43B:517-523.

92. Homewood, C. A., G. A. Moore, D. C. War-hurst, and E. M. Atkinson. 1975. Purificationand some properties of malarial pigment. Ann.Trop. Med. Parasitol. 69:283-287.

93. Homewood, C. A., and K. D. Neame. 1974.Malaia and the permeability of the host eryth-rocyte. Nature (London) 252:718-719.

94. Homewood, C. A., D. C. Warhurst, W. Peters,and V. C. Baggaley. 1972. Electron transportin intraerythrocytic Plasmodium berghei.Proc. Helminthol Soc. Wash. 39:382-386.

95. Honigberg, B. M. 1967. Chemistry of parasitismamong some protozoa, p. 695-814. In G. W.Kidder (ed.), Chemical zoology, vol. 1, Proto-zoa. Academic Press Inc., New York.

96. Howells, R. E. 1970. Mitochondrial changes dur-ing the life cycle of Plasmodium berghei. Ann.Trop. Med. Parasitol. 64:181-187.

97. Howeils, R. E., and L. MaxwelL 1973. Furtherstudies on the mitochondrial changes duringthe life cycle of Plasmodium berghei: electro-phoretic studies on isocitrate dehydrogenases.Ann. Trop. Med. Parasitol. 67:279-283.

98. Howells, RI E., and L Maxwell. 1973. Citricacid cycle activity and chloroquine resistancein rodent malaria parasites: the role of reticu-locyte. Ann. Trop. Med. Parasitol. 67:285-300.

99. Howells, R. E., W. Peters, and J. Fullard.1969. Cytochrome oxidase activity in a normal

and some drug-resistant strains ofPlasmodium


berghei-a cytochemical study. I. Asexualerythrocytic stages. Mil. Med. 134:893-915.

100. Ban, J., D. R. Pierce, and F. W. Miller. 1977.Influence of 9-(B-D-arabinofuranosyladenine ontotal protein synthesis and on differential geneexpression of unique proteins in the rodentmalarial parasite Plasmodium berghei. Proc.Natl. Acad. Sci. U.S.A. 74:3386-3390.

101. Ilan, J., and K. Tokuyasu. 1969. Amino acidactivation for protein synthesis in Plasmodiumberghei. Mil. Med. 134:1026-1031.

102. Johns, F. M. 1930. Influence of dextrose and oflow temperatures on preservation, transporta-tion and viability of malaria parasites. Proc.Soc. Exp. Biol. Med. 28:743-745.

103. Jung, A., R. Jackisch, A. Picard-Maureau,and R. HeischkeiL 1975. DNA-, RNA- undLipidsynthese sowie die spezifische Aktivitatvon Glucose-6-phosphatdehydrogenase undGlucose-6-phosphatase in den verschiedenenmorphologischen Stadien von Plasmodiumvinckei. Tropenmed. Parasitol. 26:27-34.

104. Kilejian, A. 1974. A unique histidine-rich poly-peptide from the malaria parasite, Plasmo-dium lophurae. J. Biol. Chem. 249:4650-4655.

105. Kilejian, A. 1975. Circular mitochondrial DNAfrom the avian malarial parasite Plasmodiumlophurae. Biochim. Biophys. Acta 390:276-284.

106. Kilejian, A. 1976. Does a histidine-rich proteinfrom Plasmodium lophurae have a function inmerozoite penetration? J. Protozool. 23:272-277.

107. Kilejian, A. 1976. Studies on a histidine-richprotein from Plasmodium lophurae, p. 441-448. In H. Van den Bossche (ed.), Biochemistryof parasites and host-parasite relationships.Elsevier/North Holland Biomedical Press,Amsterdam.

108. Kilejian, A. 1978. Histidine-rich protein as amodel malaria vaccine. Science 201:922-924.

109. Kilejian, A., A. Abati, and W. Trager. 1977.Plasmodium fakciparum and Plasmodiumcoatneyi: immunogenicity of "knoblike protru-sions" on infected erythrocyte membranes.Exp. Parasitol. 42:157-164.

110. Kilejian, A., and J. B. Jensen. 1977. A histi-dine-rich protein from Plasmodium falcipa-rum and its interaction with membranes. Bull.W.H.O. 55:191-198.

111. Kilejian, A., T. H. Liao, and W. Trager. 1975.Studies on the primary structure and biosyn-thesis of a histidine-rich polypeptide from themalaria parasite, Plasmodium lophurae. Proc.Natl. Acad. Sci. U.S.A. 72:3057-3059.

112. Killby, V. A. A., and P. H. Silverman. 1969.Isolated erythrocytic forms of Plasmodiumberghei. An electron-microscopical study. Am.J. Trop. Med. Hyg. 18:836-859.

113. Killick-Kendrick, R. 1974. Parasitic protozoa ofthe blood of rodents: a revision of Plasmodiumberghei. Parasitology 69:225-237.

114. Konigk, E. 1977. Salvage syntheses and theirrelationship to nucleic acid metabolism. Bull.W.H.O. 55:249-252.

115. Konigk, E., and S. Mirtsch. 1977. Plasmodium


mbr.asm

.org/D

ownloaded from


490 SHERMAN

chabaudi-infection of mice: specific activitiesof erythrocyte membrane-associated enzymes

and patterns of proteins and glycoproteins oferythrocyte membrane preparations. Tro-penmed. Parasitol. 28:17-22.

116. Krooth, R. S., K. D. Wuu, and R. Ma. 1969.Dihydroorotic acid dehydrogenase: introduc-tion into erythrocyte by the malaria parasite.Science 164:1073-1075.

117. Langer, B. W., Jr., P. Phisphumvidhi, and Y.Friedlander. 1967. Malarial parasite metabo-lism: the pentose cycle in Plasmodium berghei.Exp. Parasitol. 20:68-76.

118. Langer, B. W., Jr., P. Phisphumvidhi, and D.Jampermpoon. 1970. Malarial parasite me-

tabolism: the glutamic acid dehydrogenase ofPlasmodium berghei. Exp. Parasitol. 28:298-303.

119. Langer, B. W., Jr., P. Phisphumvidhi, D.Jampermpoon, and R. P. Weidhorn. 1969.Malarial parasite metabolism: the metabolismof methionine by Plasmodium berghei. Mil.Med. 134:1039-1044.

120. Langreth, S. 1977. Electron microscope cyto-chemistry of host-parasite membrane interac-tions in malaria. Bull. W.H.O. 55:171-178.

121. Langreth, S. G., J. B. Jensen, R. T. Reese,and W. Trager. 1978. Fine structure ofhumanmalaria in vitro. J. Protozool. 25:443-452.

122. Langreth, S. G., P. Nguyen-Dinh, and W.Trager. 1978. Plasmodium falciparum: mer-

ozoite invasion in vitro in the presence of chlo-roquine. Exp. Parasitol. 46:235-238.

123. Langreth, S. G., and W. Trager. 1973. Finestructure of the malaria parasite Plasmodiumlophurae developing extracellularly in vivo. J.Protozool. 20:606-613.

124. Lawrence, C. W., and R. J. Cenedela. 1969.Lipid content of Plasmodium berghei-infectedrat red blood cells. Exp. Parasitol. 26:181-186.

125. Levy, M. R., and S. C. Chou. 1973. Activity andsome properties of an acid proteinase fromnormal and Plasmodium berghei-infected redcells. J. Parasitol. 59:1064-1070.

126. Levy, M. R., and S. C. Chou. 1974. Some prop-erties and susceptibility to inhibitors of par-tially purified acid proteases from Plasmodiumberghei and from ghosts of mouse red cells.Biochim. Biophys. Acta 334:423-430.

127. Levy, M. R., and S. C. Chou. 1975. Inhibitionof macromolecular synthesis in the malarialparasites by inhibitors of proteolytic enzymes.Exp. Parasitol. 31:52-54.

128. Levy, M. R., W. A. Siddiqui, and S. C. Chou.1974. Acid protease activity in Plasmodiumfalciparum and P. knowlesi and ghosts of theirrespective host red cells. Nature (London) 247:546-549.

129. Lewert, R. M. 1952. Changes in nucleic acidsand protein in nucleated erythrocytes infectedwith Plasmodium gaUlinaceum as shown byultraviolet absorption measurements. J. Infect.Dis. 91:180-183.

130. Lukow, I., C. Schmidt, R. D. Walter, and E.Konigk. 1973. Adenosinmonophosphat-Sal-

MICROBIOL. REV.

vage-Synthese bei Plasmodium chabaudi. Z.Tropenmed. Parasitol. 24:500-504.

131. Manandhar, M. S. P., and K. Van Dyke. 1975.Detailed purine salvage metabolism in and out-side the free malarial parasite. Exp. Parasitol.37:138-146.

132. Mason, S. J., L. H. Miller, T. Shiroishi, J. A.Dvorak, and M. H. McGinnis. 1977. TheDuffy blood group determinants: their role inthe susceptibility of human and animal eryth-rocytes to Plasmodium knowlesi malaria. Br.J. Haematol. 36:327-336.

133. McClean, S., W. C. Purdy, A. Kabat, J. Sam-pugna, and R. De Zeeuw. 1976. Analysis ofthe phospholipid composition of Plasmodiumknowlesi and rhesus erythrocyte membranes.Anal. Chim. Acta 82:175-185.

134. McDanieL H. G., and P. M. L. Siu. 1972. Puri-fication and characterization of phosphoenol-pyruvate carboxylase from Plasmodiumberghei. J. Bacteriol. 109:385-390.

135. McGhee, R. B. 1953. Factors affecting the sus-ceptibility of erythrocytes to an intracellularparasite. Ann. N. Y. Acad. Sci. 56:1070-1073.

136. McGhee, R. B. 1953. The infection by Plasmo-dium lophurae of duck erythrocytes in thechick embryo. J. Exp. Med. 97:773-782.

137. McKee, R. W. 1951. Biochemistry of Plasmo-dium and the influence of antimalarials, p. 251-322. In S. H. Hutner and A. Lwoff (ed.), Bio-chemistry and physiology of protozoa, vol. 1.Academic Press Inc., New York.

138. McKee, R. W., R. A. Ormsbee, C. B. Anfinsen,Q. M. Geiman, and E. G. BalL 1946. Studieson malarial parasites. VI. The chemistry andmetabolism of normal and parasitized (P.knowlesi) monkey blood. J. Exp. Med. 84:569-582.

139. McLaren, D. J., L. H. Bannister, P. L. Trigg,and G. A. Butcher. 1977. A freeze-fracturestudy on the parasite-erythrocyte interrela-tionship in Plasmodium knowlesi infections.Bull. W.H.O. 56:199-204.

140. MeLaren, D. J., L. H. Bannister, P. L. Trigg,and G. A. Butcher. 1979. Freeze fracture stud-ies on the interaction between the malaria par-asite and host erythrocyte in Plasmodiumknowlesi infections. Parasitology 79:125-139.

141. Miller, F. W., and J. ilan. 1978. The ribosomesof Plasmodium berghei: isolation and ribo-somal ribonucleic acid analysis. Parasitology77:345-365.

142. Miller,L H. 1977. Hypothesis on the mechanismof invasion of erythrocytes by malaria mero-zoites. Bull. W.H.O. 55:157-162.

143. Miller, L. H. 1977. Evidence for differences inerythrocyte surface receptors for the malarialparasites Plasmodium falciparum and P.knowlesi. J. Exp. Med. 146:277-281.

144. Miller, L. H., M. Aikawa, and J. A. Dvorak.1975. Malaria (Plasmodium knowlesi) mero-zoites: immunity and the surface coat. J. Im-munol. 114:1237-1242.

145. Miller, L. H., M. Aikawa, J. G. Johns, and T.Shiroishi. 1979. Interaction between cytocha-


mbr.asm

.org/D

ownloaded from


VOL. 43, 1979

lasin-B treated malarial parastes and eryth-rocytes. Attachment and junction formation. J.Exp. Med. 149:172-184.

146. Miller, L H., J. A. Dvorak, T. Shiroishi, andJ.R Durocher. 1973. Influence of erythrocytemembrane components on malaria merozoiteinvasion. J. Exp. Med. 138:1597-1561.

147. Miller, L H., KL H. McGinnis, P. V. Holland,and P. Sigmon. 1978. The Duffy blood groupphenotype in American blacks infected withPlasmodium vivax in Vietnam. Am. J. Trop.Med. Hyg. 27:1069-1072.

148. Miller, Z. B., and L M. Koziloff. 1947. Theribonuclease activity of normal and parasitizedchick erythrocytes. J. Biol. Chem. 170:105-120.

149. Momen, H. 1979. Biochemistry of intraerythro-cytic parasites. I. Identification of enzymes ofparasite origin by starch-gel electrophoresis.Ann. Trop. Med. ParasitoL 73:109-116.

150. Momen, H. 1979. Biochemistry of intraerythro-cytic parasites. II. Comparative studies in car-

bohydrate metabolism. Ann. Trop. Med. Par-asitoL 73:117-122.

151. Momen, H, E M. Atinson, and C. A. Home-wood. 1975. An electrophoretic investigationof the malate dehydrogenase of mouse eryth-rocytes infected with Pkasodium berghei. Int.J. Biochem 6:533-535.

152. Moulder, J. 1962. The biochemistry of intracel-lular parasitism. University of Chicago Press,Chicago.

153. Moulder, J. W., and E. A. Evans, Jr. 1946.The biochemistry of the malaria parasite. VI.Studies on the nitrogen metabolism of the ma-laria parasite. J. Biol. Chem. 164:135-157.

154. Miiller, 1975. Biochemistry of protozoan mi-crobodies: peroxisumes, glycerophosphate oxi-dase bodies, hydrogenosomes. Annu. Rev. Mi-crobioL 29:467-483.

155. Nagarajan, K. 1964. Pyruvate and lactate levelsin relationship to the nicotinamide adeninedinucleotide levels in malarial parasites (Plas-modium berghe/). Biochim. Biophys. Acta 93:176-179.

156. Nagarajan, K. 1968. Metabolism of Plasmo-dium berghei. I. Krebs cycle. Exp. Parasitol.22:19-26.

157. Nagarajan, K. 1968. Metabolism of Plasmo-dium berghei. II. 32P, incorporation into high-energy phosphates. Exp. ParasitoL 22:27-32.

158. Nagarajan, K. 1968. Metabolism of Plasmo-dium berghei. Ill. Carbon dioxide fixation androle of pyruvate and dicarboxylic acids. Exp.ParasitoL 22:33-42.

159. Neame, K. D., and C. A. Homewood. 1975.Alterations in the permeability ofmouse eryth-rocytes infected with the malaria parasite,Plasmodium berghei. Int. J. ParasitoL 5:537-540.

160. Nowell, F. 1970. The effect of a milk diet uponPlasmodium berghei, Nuttalia (- Babesia)rodhaini and Trypanosoma brucei infectionsin mice. Parasitology 61:425-433.

161. Oelshlegel, F. J., Jr., and G. J. Brewer. 1975.


Parasitism and the red blood cell, p. 1263-1302.In D. M. N. Surgenor (ed.), The red blood cell,2nd ed., vol. 2. Academic Press Inc., New York.

162. OelshlegeL F. J., Jr., B. J. Sander, and G. J.Brewer. 1975. Pyruvate kinase in malariahost-parasite interaction. Nature (London)255:345-347.

163. Overman, R. R., A. C. Bass, and T. H. Tom-linson, Jr. 1950. Ionic alterations in chickensinfected with Pkasmodium gallinaceum. Fed.Proc. 9:96-97.

164. Overman, R. R., T. S. Hil, and U. T. Wong.1949. Physiological studies in the human nor-mal host. I. Blood, plasma, "extracellular" fluidvolumes and ionic balance in therapeutic P.vivax and P. fakciparum. J. Natl. Malar. Soc.8:14-31.

165. Pfefferkorn, E. R., and L. C. Pfefferkorn.1977. Specific labeling of intracellular Toxo-plasna gondii with uracil. J. Protozool. 24:449-453.

166. Phisphumvidhi, P., and B. W. Langer, Jr.1969. Malarial parasite metabolism: the lacticacid dehydrogenase of Plasmodium berghei.Exp. Parasitol. 24:37-41.

167. Platzer, E. G. 1972. Metabolism of tetrahydro-folate in Plasmodium lophusrae and ducklingerytbrocytes. Trans. N. Y. Acad. Sci. Ser. 2 34:200-208.

168. Platzer, E. G. 1974. Dihydrofolate reductase inPlasmodium lophurae and duckling erythro-cytes. J. Protozool. 21:400405.

169. Platzer, E. G. 1977. Subcellular distribution ofserine hydroxymethyltranserase in Plasmo-dium lophusrae. Life Sci. 20:1417-1424.

170. Platzer, E. G., and H. C. Campuzano. 1976.The serine hydroxymethyltransferase of Plas-modium lophurae. J. ProtozooL 23:282-286.

171. Platzer, E. G., and J. A. Kassis. 1978. Pyridox-ine kinase in Plasmodium lophurae andduckling erythrocytes. J. Protozool. 25:556-559.

172. Polet, H., and C. F. Barr. 1968. DNA, RNA,and protein synthesis in erythrotic forms ofPlasmodium knowlesi Am. J. Trop. Med.Hyg. 17:672-679.

173. Polet, H., N. D. Brown, and C. R. AngeL 1969.Biosynthesis of amino acids from '4C-U-glu-cose, pyruvate, and acetate by erythrocyticforms of P. knowkesi, in vitro. Proc. Soc. Exp.BioL Med. 131:1215-1218.

174. Polet, H., and ML E. Conrad. 1968. Malaria:extracellular amino acid requirements for invitro growth of erythrocytic forms of Plasmo-dium knowlesi Proc. Soc. Exp. Biol. Med. 127:251-253.

175. Ramakrislhan, S. P. 1954. Studies on Plas-modium berghei Vincke and Lips 1948. XIX.The course of blood induced infections in pyr-idoxine or vitamin B6 deficient rats. Indian J.MalarioL 8:107-113.

176. Rama Rao, R., and M. Sirsi. 1956. Avian ma-laria and B complex vitamins. I. Thiamine. J.Indian Inst. Sci. 38:108-114.

177. Rama Rao, R., and M. Sirsi. 1956. Avian ma-


mbr.asm

.org/D

ownloaded from


492 SHERMAN

laria and B complex vitamins. II. Riboflavin. J.Indian Inst. Sci. 38:186-189.

178. Rao, K. N., D. Subrahmanyam, and S. Prak-ash. 1970. Plasmodium berghei: lipids of ratred blood cells. Exp. Parasitol. 27:22-27.

179. Razin, S. 1978. The mycoplasmas. Microbiol.Rev. 42:414-470.

180. Reeves, R., L. Warren, B. Susskind, and H.Lo. 1977. An energy conserving pyruvate-to-acetate pathway in Entamoeba histolytica. J.Biol. Chem. 252:726-731.

181. Reid, V. E., and M. Friedkin. 1973. Plasmo-dium berghei: folic acid levels in mouse eryth-rocytes. Exp. Parasitol. 33:424-428.

182. Reid, V. E., and M. Friedkin. 1973. Thymidyl-ate synthetase in mouse erythrocytes infectedwith Plasmodium berghei. Mol. Pharmacol. 9:74-80.

183. Rich, P. R., and A. L Moore. 1976. The involve-ment of the protonmotive ubiquinone cycle inthe respiratory chain of higher plants and itsrelation to the branchpoint of the alternatepathway. FEBS Lett. 65:334-344.

184. Rietz, P. J., F. S. Skelton, and K. Folkers.1967. Occurrence of ubiquinones-8 and -9 inPlasmodium lophurae. Int. J. Vitam. Nutr.Res. 37:405411.

185. Rock, R. C. 1971. Incorporation of "4C-labellednon-lipid precursors into lipids of Plasmodiumknowlesi in vitro. Comp. Biochem. Physiol.40B:657-669.

186. Rock, R. C. 1971. Incorporation of '4C-labelledfatty acids into lipids of rhesus erythrocytesand Plasmodium knowlesi in vitro. Comp. Bio-chem. Physiol. 40B:893-906.

187. Rock, R. C., J. C. Standefer, R. T. Cook, W.little, and H. Sprinz. 1971. Lipid compositionof Plasmodium knowlesi membranes: compar-ison of parasites and microsomal subfractionswith host rhesus erythrocyte membranes.Comp. Biochem. Physiol. 38B:425-437.

188. Rock, R. C., J. Standefer, and W. little. 1971.Incorporation of 3'P-orthophosphate intomembrane phospholipids of Plasmodiumknowlesi and host erythrocytes ofMacaca mu-latta. Comp. Biochem. Physiol. 40B:543-561.

189. Roos, A., D. Hegsted, and F. Stare. 1946.Nutritional studies with the duck. IV. Theeffect of vitamin deficiencies on the course ofPlasmodium lophurae infection in the duckand the chick. J. Nutr. 32:473-484.

190. Scheibel, L. W., S. H. Ashton, and W. Trager.1979. Plasmodium falciparum: microaero-philic requirements in human red blood cells.Exp. Parasitol. 47:410-418.

191. Scheibel, L. W., and J. Miller. 1969. Cyto-chrome oxidase activity in platelet-free prepa-rations of Plasmodium knowlesi. J. Parasitol.55:825-829.

192. Scheibel, L W., and J. Miller. 1969. Glycolyticand cytochrome oxidase activity in plasmodia.Mil. Med. 134:1074-1080.

193. Scheibel, L. W., and W. K. Pflaum. 1970. Car-bohydrate metabolism in Plasmodium know-lesi. Comp. Biochem. Physiol. 37:543-553.

MICROBIOL. REV.

194. Schellenberg, K. A., and G. R. Coatney. 1961.The influence of anti-malaril drugs on thenucleic acid synthesis in Plasmodium galli-naceum and Plasmodium berghei. Biochem.Pharmacol. 6:143-152.

195. Schmidt, G., R. D. Walter, and E. Konigk.1974. Adenosine kinase from normal mouseerythrocytes and from Plasmodium chabaudi:partial purification and characterization. Tro-penmed. Parasitol. 25:301-308.

196. Schmidt-Ullrich, R., and D. F. H. Wallach.1978. Plasmodium knowlesi-induced antigensin membranes of parasitized rhesus monkeyerythrocytes. Proc. Natl. Acad. Sci. U.S.A. 75:4949-4953.

197. Schmidt-Ullrich, R., D. F. H. Wallach, and J.Lightholder. 1979. Two Plasmodium know-lesi-specific antigens on the surface of schizont-infected rhesus monkey erythrocytes induceantibody production in immune hosts. J. Exp.Med. 150:86-99.

198. Schnell, J. V., W. A. Siddiqui, and Q. M.Geiman. 1971. Biosynthesis of coenzymes Qby malarial parasites. II. Coenzyme Q synthesisin blood cultures of monkeys infected withmalarial parasites (Plasmodium fakciparumand P. knowlesi). J. Med. Chem. 14:1026-1029.

199. Seaman, G. R. 1953. Inhibition of the succinicdehydrogenase of parasitic protozoans by anarsono and a phosphono analog of succinicacid. Exp. Parasitol. 2:366-373.

200. Seed, T. M., M. Aikawa, and C. R. Sterling.1973. An electron microscope-cytochemicalmethod for differentiating membranes of hostred cells and malaria parasites. J. Protozool.20:603-605.

201. Seed, T. M., and J. P. Kreier. 1972. Plasmo-dium gallinaceum: erythrocytic membrane al-terations and associated plasma changes in-duced by experimental infections. Proc. Hel-minthol. Soc. Wash. 39:387-411.

202. Seed, T. M., and J. P. Kreier. 1976. Surfaceproperties of extracellular malaria parasites:electrophoretic and lectin-binding characteris-tics. Infect. Immun. 14:1339-1347.

203. Seeler, A. 0. 1945. The inhibitory effect of pyr-idoxine on the activity of quinine and atabrineagainst avian malaria. J. Natl. Malar. Soc. 4:13-19.

204. Seeler, A., and W. Ott. 1944. Effect of riboflavindeficiency on the course of Plasmodium to-phurae infections in chicks. J. Infect. Dis. 75:175-178.

205. Shakespeare, P. G., and P. L. Trigg. 1973.Glucose catabolism by the simian malaria par-asite Plasmodium knowlesi. Nature (London)241:538-540.

206. Sherman, L. W. 1961. Molecular heterogeneityof lactic dehydrogenase in avian malaria (Plas-modium lophurae). J. Exp. Med. 114:1049-1062.

207. Sherman, I. W. 1962. Heterogeneity of lacticdehydrogenase in intraerythrocytic parasites.Trans. N. Y. Acad. Sci. Ser. 2 24:944-953.

208. Sherman, L. W. 1965. Glucose-6-phosphate de-


mbr.asm

.org/D

ownloaded from


VOL. 43, 1979

hydrogenase and reduced glutathione in ma-laria-infected erythrocytes (Plasmodium lo-phurae and P. berghei). J. ProtozooL 12:394-396.

209. Sherman, L W. 1966. Heterogeneity of lacticdehydrogenase in avian malaria demonstratedby the use of coenzyme analogs, p. 73, A2. InA. Coradetti (ed.), Proceedings of the FirstInternational Congress of Parasitology. Perga-mon Press, Oxford.

210. Sherman, L. W. 1966. In vitro studies of factorsaffecting penetration of duck erythrocytes byavian malaria (Plasmodium lophurae). J. Par-asitol. 52:17-22.

211. Sherman, L. W. 1966. Levels of oxidized andreduced pyridine nucleotides in avian malaria(Plasmodium lophurae). Am. J. Trop. Med.Hyg. 15:814-817.

212. Sherman, L. W. 1966. Malic dehydrogenase het-erogeneity in malaria (Plasmodium lophuraeand P. berghei). J. Protozool. 13:344-349.

213. Sherman, L. W. 1976. The ribosomes of thesimian malaria Plasmodium knowlesi. II. Acell-free protein synthesizing system. Comp.Biochem. PhysioL 53B:447-450.

214. Sherman, L W. 1977. Transport of amino acidsand nucleic acid precursors in malarial para-sites. Bull. W.H.O. 55:211-225.

215. Sherman, L W. 1977. Amino acid metabolismand protein synthesis in malarial parasites.Bull. W.H.O. 55:265-276.

216. Sherman, . W., R. A. Cox, B. EHgginsn, D.J. MeLaren, and J. Williamson. 1975. Theribosomes of the simian mlaria, Plasmodiumknowlesi. I. Isolation and characterization. J.ProtozooL 22:568-572.

217. Sherman, L. W., andL A. Jones. 1976. Proteinsynthesis by a cell-free preparation from thebird malaria, Plasmodium lophurae. J. Proto-zooL 23:277-281.

218. Sherman, L. W., and L A. Jones. 1977. ThePlasmodium lophurae (avian malaria) ribo-some. J. Protozool. 24:331-334.

219. Sherman, I. W., and L A. Jones. 1979. Plas-modium lophurae: Membrane proteins oferythrocyte-free plasmodia and malaria-in-fected red cells. J. Protozool. 26:489-501.

220. Sherman, L W., and J. B. Mudd. 1966. Malariainfection (P. lophurae): changes in free aminoacids. Science 154:287-289.

221. Sherman, L W., L. Peterson, L. Tagoshi,and L P. Ting. 1971. The glutamate dehydro-genase of Plasmodium lophurae (avian ma-laria). Exp. ParasitoL 29:433-439.

222. Sherman, L W., J. A. Ruble, and L P. Ting.1969. Piasmodium lophUrae: [U-_4C]-_gucosecatabolism by free plasmodia and duckling hosterythrocytes. Exp. ParasitoL 25:181-192.

223. Sherman, L. W., and L. Tanigoshi. 1970. In-corporation of '4C-amino acids by malaria(Plasmodium lophurae). IV. In vivo utilizationof host cell haemoglobin. Int. J. Biochem. 1:635-637.

224. Sherman, I. W., and L Tanigoshi. 1971. Al-terations in sodium and potassium in the red


blood cells and plasma during the malaria in-fection (Plasmodium lophurae). Comp. Bio-chem. Physiol. 40A:543-546.

225. Sherman, I. W., and L. Tanigoshi. 1974. In-corporation of '4C-amino acids by malarialplasmodia (Pklmodium lophurae). VI.Changes in the kinetic constants of amino acidtransport during infection. Exp. Parasitol. 35:369-373.

226. Sherman, I. W., and L Tanigoshi. 1974. Glu-cose transport in the malarial (Plasmodiumlophurae) infected erythrocyte. J. Protozool.21:603-607.

227. Sherman, L W., and L. P. Ting. 1966. Carbondioxide fixation in malaria (Plasmodium lo-phurae). Nature (London) 212:1387-1389.

228. Sherman, L. W., and L P. Ting. 1968. Carbondioxide fixation in malaria. II. Plasmodiumknowlesi (monkey malaria). Comp. Biochem.Physiol. 24:639-642.

229. Sherman, L. W., L. P. Ting, and J. A. Ruble.1968. Characterization of the malaria pigment(hemozoin) from the avian malaria parasitePlasmodium lophurae. J. Protozool. 15:158-164.

230. Sherman, L. W., L. P. Ting, and L. Tanigoshi.1970. Plasmodium lophurae: glucose-1-14C andglucose-6-14C catabolism by free plasmodia andduckling host erythrocytes. Comp. Biochem.Physiol. 34:625-639.

231. Siddiqui, W. A., J. V. Schell, and Q. M. Gei-man. 1967. Stearic acid as plasma replacementfor intracellular in vitro culture ofPlasmodiumknowlesi. Science 156:1623-1625.

232. Siddiqui, W. A., and W. Trager. 1966. Folicand folinic acids in relation to the developmentofPlasmodium lophurae. J. Parasitol. 52:556-558.

233. Singer, I. 1956. Coenzyme A changes in liver,spleen and kidneys of rats with infections ofPlasmodium berghei. Proc. Soc. Exp. Biol.Med. 91:315-318.

234. Singer, L. 1961. Tissue thiamine changes in ratswith experimental trypanosomiasis or malaria.Exp. Parasitol. 11:391-401.

235. Siu, P. M. L 1967. Carbon dioxide fixation inplasmodia and the effect of some antimalarialdrugs on the enzyme. Comp. Biochem. Physiol.23:785-795.

236. Skelton, F. S., K. D. Lunan, K. Folkers, J. V.Schnell, W. A. Siddiqui, and Q. M. Geiman.1969. Biosynthesis of ubiquinones by malarialparasites. I. Isolation of ['4C]ubiquinones fromcultures of rhesus monkey blood infected withPlamsodium knowlesi. Biochemistry 8:1284-1287.

237. Skelton, F. S., P. J. Rietz, and K. Folkers.1970. Coenzyme Q. CXXII. Identification ofubiquinone-8 biosynthesized by Plasmodiumknowlesi, P. cynomolgi and P. berghei. J. Med.Chem. 13:602-606.

238. Smith, C. C., G. J. McCormick, and C. J.Canfield. 1976. Plasmodium knowlesi: in vitrobiosynthesis of methionine. Exp. Parasitol. 40:432-437.


mbr.asm

.org/D

ownloaded from


494 SHERMAN

239. Speck, J. F., and E. A. Evans, Jr. 1945. Thebiochemistry of the malaria parasite. II. Gly-colysis in cell-free preparations of the malariaparasite. J. Biol. Chem. 159:71-81.

240. Speck, J. F., J. W. Moulder, and E. A. Evans,Jr. 1946. The biochemistry of the malaria par-asite. V. Mechanism of pyruvate oxidation inthe malaria parasite. J. Biol. Chem. 164:119-144.

241. Takahashi, Y., and LW. Sherman. 1978. Pics-modium lophurae: cationized feritin staining,an electron microscope cytochemical methodfor differentiating malarial parasite and hostcell membranes. Exp. Parasitol. 44:145-154.

242. Takahashi, Y., and L.W. Sherman. 1979. Plas-modium lophurae: lectin mediated agglutina-tion ofmalaria-infected red cells and fine-struc-ture cytochemical detection of lectin bindingsites on plasmodial and host cell membranes.Exp. Parasitol. (in press).

243. Taliaferro, W. H., and L. G. Taliaferro. 1940.Active and passive immunity in chickensagainst Plasmodium lophurae. J. Infect. Dis.66:153-165.

244. Taylor, A. E. R. 1958. Effects of cod liver oil andvitamin E on infections in Plasmodium galli-naceum and Treponema duttoni. Ann. Trop.Med. Parasitol. 52:139-144.

245. Theakston, R. D. G., and K. A. Fletcher. 1973.A technique for the cytochemical demonstra-tion in the electron microscope of glucose-6-phosphate dehydrogenase activity in erythro-cytes of malaria-infected animals. J. Microsc.(Oxford) 97:315-320.

246. Theakston, R. D. G., and K. A. Fletcher. 1973.An electron cytochemical study of 6-phospho-gluconate dehydrogenase activity in infectederythrocytes during malaria. Life Sci. 13:405-410.

247. Theakston, R. D. G., K. A. Fletcher, and B.G. MaeGraith. 1970. The use of electron mi-croscope autoradiography for examining theuptake and degradation of haemoglobin byPlasmodium berghei. Ann. Trop. Med. Paras-itol. 64:63-71.

248. Theakston, R. D. G., K. A. Fletcher, and G.A. Moore. 1976. Glucose-6-phosphate and 6-phosphogluconate dehydrogenase activities inhuman erythrocytes infected with Plasmo-dium falciparum. Ann. Trop. Med. Parasitol.70:125-127.

249. Theakston, R. D. G., R. E. Howells, K. A.Fletcher, W. Peters, J. Fuliard, and G. A.Moore. 1969. The ultrastructural distributionof cytochrome oxidase activity in Plasmodiumberghei and P. gallinaceum. Life Sci. 8:521-529.

250. Ting, L. P., and L. W. Sherman. 1966. Carbondioxide fixation in malaria. I. Kinetic studies inPlasmodium lophurae. Comp. Biochem. Phys-iol. 19:855-869.

251. Tokuyasu, K., J. Ilan, and J. Ilan 1969. Bio-genesis of ribosomes in Plasmodium berghei.Mil. Med. 134:1032-1038.

252. Tracy, S. M., and L. W. Sherman. 1972. Purine

uptake and utilization by the avian malariaparasite Plasmodium lophurae. J. Protozool.19:541-549.

253. Trager, W. 1950. Studies on the extracellularcultivation of an intracellular parasite (avianmalaria). I. Development of the organisms inerythrocyte extracts, and the favoring effect ofadenosine triphosphate. J. Exp. Med. 92:349-366.

254. Trager, W. 1952. Studies on the extracellularcultivation of an intracellular parasite (avianmalaria). II. The effects of malate and of co-enzyme A concentrates. J. Exp. Med. 96:465-476.

255. Trager, W. 1967. Adenosine triphosphate andthe pyruvic and phosphoglyceric kinases of themalaria parasite Plasmodium lophurae. J.Protozool. 14:110-114.

256. Trager, W. 1971. Malaria parasites (Plasmo-dium lophurae) developing extracellularly invitro: incorporation of labeled precursors. J.Protozool. 18:392-399.

257. Trager, W. 1972. Effects of bongkrekic acid onmalaria parasites (Plasmodium lophurae) de-veloping extracellularly in vitro, p. 343-350. InH. Van den Bossche (ed.), Comparative bio-chemistry of parasites. Academic Press Inc.,London.

258. Trager, W. 1973. Bongkrekic acid and the aden-osine triphosphate requirement of malaria par-asites. Exp. Parasitiol. 34:412416.

259. Trager, W. 1977. Cofactors and vitamins in themetabolism of malaria parasites. Bull. W.H.O.55:285-289.

260. Trager, W., and J. B. Jensen. 1976. Humanmalaria parasites in continuous culture. Sci-ence 193:673-675.

261. Trager, W., S. G. Langreth, and E. G. Platzer.1972. Viability and fine structure of extracel-lular Plasmodium lophurae prepared by dif-ferent methods. Proc. Hehminthol. Soc. Whsh.39:220-230.

262. Trager, W., M. A. Rudzinska, and P. Brad-bury. 1966. The fine structure of Plasmodiumfalkparum and its host erythrocytes in naturalmalarial infections in man. Bull. W.H.O. 35:883-86.

263. Trigg, P. L. 1968. Sterol metabolism of Plasmo-dium knowlesi in vitro. Ann. Trop. Med. Par-asitol. 62:481487.

264. Trigg, P. I., and W. E. Gutteridge. 1972. Arational approach to the serial culture of ma-laria parasites. Evidence for a deficiency inRNA synthesis during the first cycle in vitro.Parasitology 65:265-271.

265. Trigg, P. L., S. Hirst, P. Shakespeare, and LTappenden. 1977. Labelling of membrane gly-coprotein in erytFrocytes infected with Plas-modium knowlesi. Bull. W.H.O. 55:205-210.

266. Trigg, P. I., P. G. Shakespeare, S. J. Burt,and S. L. Kyd. 1975. Ribonucleic acid synthesisin Plasmodium knowlesi maintained both invivo and in vitro. Parasitology 71:199-209.

267. Tsukamoto, M. 1974. Differential detection ofsoluble enzymes specific to a rodent malaria

MICROBIOL. REV.


mbr.asm

.org/D

ownloaded from


VOL. 43, 1979

parasite, Piasmodium berghei, by electropho-resis on polyacrylamide gels. Trop. Med. 16:55-69.

268. Uyeda, K., and J. C. Rabinowitz. 1971. Pyru-vate-ferredoxin oxidoreductase. m. Purifica-tion and properties of the enzyme. J. BioLChem. 246:3111-3119.

269. Velick, S. F. 1942. The respiratory metabolismofthe malaria parasite, P. cathemerium, duringits developmental cycle. Am. J. Hyg. 35:152-161.

270. Wallach, D. F. H., and M. Conley. 1977. Al-tered membrane proteins of monkey erythro-cytes infected with simian malaria. J. Mol.Med. 2:119-136.

271. Walsh, C. J., and L W. Sherman. 1968. Isola-tion, characterization and synthesis of DNAfrom a malaria parasite. J. Protozool. 15:503-508.

272. Walsh, C. J., and L. W. Sherman. 1968. Purineand pyrimidine synthesis by the avian malariaparasite, Plasmodium lophurae. J. Protozool.15:763-770.

273. Walter, R. D., and E. Konigk. 1971. Syntheseder Desoxythymidylat-Synthetase und derDihydrofolat-Reduktase bei synchroner Schi-zogonie von Plasmodium chabaudi. Z. Tro-penmed. ParasitoL 22:250-255.

274. Walter, R. D., and E. Komigk. 1971. Plasmo-dium chabaudi: die enzymatische Synthesevon Dihydropteroat und ihre Hemmung durchSulfonamide. Z. Tropenmed. ParasitoL 22:256-259.

275. Walter, R. D., and E. Konigk. 1974. Biosyn-thesis of folic acid compounds in plasmodia.Purification and properties of the 7,8-dihydro-pteroate-synthesizing enzyme from Plasmo-dium chabaudi. Hoppe-Seyler's Z. Physiol.Chem. 355:431-437.

276. Walter, R. D., and E. Konigk. 1974. Hypoxan-thine-guanine phosphoribosyltransferase andadenine phosphoribosyltransferase from Plas-modium chabaudi, purification and properties.Tropenmed. Parasitol. 25:227-235.

277. Walter, R. D., H. Muhlpfordt, and E. Komgk1970. Vergleichende Untersuchungen der De-soxythymidylatsynthese bei Plasmodium cha-baudi, Trpanosoma gambiense und Trypa-nosoma lewisi. Z. Tropenmed. ParasitoL 21:


347-357.278. Walter, R. D., J. P. Nordmeyer, and E. Kon-

igk. 1974. NADP-specific glutamate dehydro-genase from Plasmodium chabaudi. Hoppe-Seyler's Z. Physiol. Chem. 355:495-500.

279. Wan, Y. P., T. H. Porter, and K. Folkers.1974. Antimalarial quinones for prophylaxisbased on a rationale of inhibition of electrontransfer in Plasmodium. Proc. Natl. Acad. Sci.U.S.A. 71:952-956.

280. Wang, E. J., and H. J. Saz. 1974. Comparativebiochemical studies of Litomosoides carinii,Dipetalonema viteae and Brugia pahangiadults. J. Parasitol. 60:316-319.

281. Warhurst, D. C., and J. Williamson. 1970.Ribonucleic acid from Plasmodium knowlesibefore and after chloroquine treatment. Chem.Biol. Interact. 2:89-106.

282. Weidekamm, E., D. F. H. Wallach, P. S. -in,and J. Hendricks. 1973. Erythrocyte mem-brane alterations due to infection with Plas-modium berghei. Biochim. Biophys. Acta 323:539-546.

283. Wendel, W. B., and S. KimbalL 1942. Forma-tion of lactic acid and pyruvic acid in bloodcontaining Plasmodium knowlesi. J. Biol.Chem. 145:343-344.

284. Whitfeld, P. R. 1952. Nucleic acids in erythro-cytic stages of a malaria parasite. Nature (Lon-don) 169:751-752.

285. Whitfeld, P. R. 1953. Studies on the nucleicacids of the malaria parasite, Plasmodiumberghei (Vincke and Lips). Aust. J. Biol. Sci.6:234-243.

286. Yamada, K., and L. W. Sherman. 1977. Regu-lation of red cell purine banwport and metab-olism by malaria parasites. J. Protozool. (inpress).

287. Yamada, K. A., and L W. Sherman. 1979.Plasmodium lop/urae: composition and prop-erties of hemozoin, the malarial pigment. Exp.ParasitoL 48:61-74.

288. Yuthavong, Y., P. Wilairat, B. Panijpan, C.Potiwan, and G. Beale. 1979. Alterations inmembrane proteins of mouse erythrocytes in-fected with different strains and species of ma-laria parasites. Comp. Biochem. Physiol. 63B:83-85.


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