different parts of the world can be adapted to short- and/or long-term cultivation using the standard methods described by Trager and Jensen [109,114,198,199,335–339]. Some of the parasite populations grow readily and adapt with ease to the new environmental and nutritional conditions, while others tend to become minority populations or even disappear (do they die out because the predominant populations use up the available resources or because they are inherently incapable of adapting to in vitro conditions, or do they persist at an undetectable level?). A recent study provides further evidence that, after adaptation to in vitro cultivation, one of the parasite populations in an isolate may become predominant over others and “impose” its phenotype, such as the multiplication rate, on others, masking the phenotypes of the minority parasite populations [340]. While this is an expected result, the opposite effect may be produced in some cases. The same authors also observed that clones derived from the original isolate displayed higher multiplication rates than the original uncloned isolate, possibly due to competitive suppression within the uncloned isolate, the presence of an undetected slow-growing parasite population in the uncloned isolate, or the selection of slow-growing populations with better fitness in the uncloned isolate.

In the literature, the term “culture-adapted isolate” or “culture-adapted strain” has been used without a clear definition. An isolate that has adapted to continuous in vitro cultivation may be loosely referred to as a new “strain”. However, the definition of a “strain” has not been established in a more stringent manner [341]. Jensen and Trager have observed that isolates generally take four to six weeks of continuous cultivation to become adapted and “stabilized” and, based on a small number of isolates in their experiments, isolates that are cryopreserved during continuous cultivation can be readily recovered beyond five or six weeks [335]. Some authors have proposed that a parasite is considered to be adapted to culture conditions when a multiplication rate of >10 every 96 h is attained [114]. Others have recommended that culture-adapted malaria parasites should refer “specifically to isolates of the parasite that have been grown continuously in vitro for at least 2 months and have been frozen, thawed, and successfully recultured at least once” [342]. Based on these three criteria (i.e., duration (at least four weeks), multiplication rate, and recovery after cryopreservation), Jensen and Trager have argued that the cultivation method developed by Haynes et al. [26], terminated after 22 days, was “abandoned too soon to permit any conclusion as to whether their conditions would in fact support continuous culture” [335]. From the pragmatic viewpoint, *P. falciparum* parasites cultivated in vitro (which no longer undergo sexual reproduction) that meet the aforementioned criteria may be said to constitute a strain, defined as “a homogeneous population possessing a set of defined characters” [341]. When these defined characters are based on genetic uniformity, which is attained in the laboratory by separating a single parasite-infected erythrocyte by dilution/microscopic selection [319], limiting dilution [158,343–348], micromanipulation [349,350], or single-cell sorting [351,352] and propagating it in vitro, the strain becomes a clone. Clonality is usually evidenced by the absence of polymorphisms in well-studied antigens, such as merozoite surface antigen-1 (MSA-1) and merozoite surface antigen-2 (MSA-2).

The capacity of parasite populations to adapt to in vitro conditions may have a genetic basis. A recent study has suggested that mutations in *P. falciparum* regulator genes (*ApiAP2* transcription factor genes, *Epac* gene encoding a guanine nucleotide exchange factor, *SRPK1* (serine/threonine protein kinase) genes, and *DOC2* (double C2-like domain-containing protein) genes) may be associated with culture adaptation [353]. In another series of experiments involving bulk segregant analysis, erythrocyte-binding antigen-140 (*eba-140* (note: it is one of the merozoite ligands that plays a key role in the entry of merozoite into erythrocyte by binding to the erythrocyte receptor, glycophorin) [354]), and, to a lesser extent, aspartate transaminase (*ast*) and cysteine protease autophagy-related 4 (*atg4*) genes, were identified as the key loci that determine fitness (i.e., parasite growth rate) in parasites grown in RPMI 1640 supplemented with either human serum or Albumax® [209]. Other investigators have studied the expression patterns of clonal multigene families (or clonally variant gene families), such as *eba-140*, cytoadherence-linked asexual genes
different parts of the world can be adapted to short- and/or long-term cultivation using the standard methods described by Trager and Jensen [109,114,198,199,335–339]. Some of the parasite populations grow readily and adapt with ease to the new environmental and nutritional conditions, while others tend to become minority populations or even disappear (do they die out because the predominant populations use up the available resources or because they are inherently incapable of adapting to in vitro conditions, or do they persist at an undetectable level?). A recent study provides further evidence that, after adaptation to in vitro cultivation, one of the parasite populations in an isolate may become predominant over others and “impose” its phenotype, such as the multiplication rate, on others, masking the phenotypes of the minority parasite populations [340]. While this is an expected result, the opposite effect may be produced in some cases. The same authors also observed that clones derived from the original isolate displayed higher multiplication rates than the original uncloned isolate, possibly due to competitive suppression within the uncloned isolate, the presence of an undetected slow-growing parasite population in the uncloned isolate, or the selection of slow-growing populations with better fitness in the uncloned isolate.

In the literature, the term “culture-adapted isolate” or “culture-adapted strain” has been used without a clear definition. An isolate that has adapted to continuous in vitro cultivation may be loosely referred to as a new “strain”. However, the definition of a “strain” has not been established in a more stringent manner [341]. Jensen and Trager have observed that isolates generally take four to six weeks of continuous cultivation to become adapted and “stabilized” and, based on a small number of isolates in their experiments, isolates that are cryopreserved during continuous cultivation can be readily recovered beyond five or six weeks [335]. Some authors have proposed that a parasite is considered to be adapted to culture conditions when a multiplication rate of >10 every 96 h is attained [114]. Others have recommended that culture-adapted malaria parasites should refer “specifically to isolates of the parasite that have been grown continuously in vitro for at least 2 months and have been frozen, thawed, and successfully recultured at least once” [342]. Based on these three criteria (i.e., duration (at least four weeks), multiplication rate, and recovery after cryopreservation), Jensen and Trager have argued that the cultivation method developed by Haynes et al. [26], terminated after 22 days, was “abandoned too soon to permit any conclusion as to whether their conditions would in fact support continuous culture” [335]. From the pragmatic viewpoint, *P. falciparum* parasites cultivated in vitro (which no longer undergo sexual reproduction) that meet the aforementioned criteria may be said to constitute a strain, defined as “a homogeneous population possessing a set of defined characters” [341]. When these defined characters are based on genetic uniformity, which is attained in the laboratory by separating a single parasite-infected erythrocyte by dilution/microscopic selection [319], limiting dilution [158,343–348], micromanipulation [349,350], or single-cell sorting [351,352] and propagating it in vitro, the strain becomes a clone. Clonality is usually evidenced by the absence of polymorphisms in well-studied antigens, such as merozoite surface antigen-1 (MSA-1) and merozoite surface antigen-2 (MSA-2).

The capacity of parasite populations to adapt to in vitro conditions may have a genetic basis. A recent study has suggested that mutations in *P. falciparum* regulator genes (*ApiAP2* transcription factor genes, *Epac* gene encoding a guanine nucleotide exchange factor, *SRPK1* (serine/threonine protein kinase) genes, and *DOC2* (double C2-like domain-containing protein) genes) may be associated with culture adaptation [353]. In another series of experiments involving bulk segregant analysis, erythrocyte-binding antigen-140 (*eba-140* (note: it is one of the merozoite ligands that plays a key role in the entry of merozoite into erythrocyte by binding to the erythrocyte receptor, glycophorin) [354]), and, to a lesser extent, aspartate transaminase (*ast*) and cysteine protease autophagy-related 4 (*atg4*) genes, were identified as the key loci that determine fitness (i.e., parasite growth rate) in parasites grown in RPMI 1640 supplemented with either human serum or Albumax® [209]. Other investigators have studied the expression patterns of clonal multigene families (or clonally variant gene families), such as *eba-140*, cytoadherence-linked asexual genes
(clag), variant genes (var; encoding *P. falciparum* erythrocyte membrane protein 1, PfEMP-1), repetitive interspersed families (rif), and subtelomeric variable open reading frame (stevor) genes, which confer phenotypic diversity to genetically homogeneous parasite populations, which in turn may confer biological advantages, especially fitness, to the parasites to allow adaptation to different environments [355,356]. There is still limited experimental evidence that supports these recent findings. More studies are required to further understand whether there is a single set of genetic modifications associated with adaptation to in vitro conditions.

Continuous cultivation of mixed parasite populations usually, but not always, results in a diminution of the initial MOI and may lead to a selection of the dominant parasite population, possibly endowed with biological fitness [357,358]. For unknown reasons, several parasite populations initially present in the day 0 isolate may also continue to co-exist in vitro, and a "new parasite population" undetected by the polymerase chain reaction (PCR) in the initial isolate on day 0 may emerge within a short time period (i.e., within 9 days of continuous cultivation or four asexual intraerythrocytic cycles) [359]. In a study based on microsatellite genotyping, single nucleotide polymorphisms (SNPs) of drug resistance markers and phenotypic profiles (drug response), it was reported that three fresh Kenyan clinical isolates of *P. falciparum* exhibited various genotypic and phenotypic profiles, without any pattern or phenotype-genotype correlation, during the 2-3 month cultivation period [360]. In that study, the stabilization of the genotypic or phenotypic profile was not observed, and there was no apparent tendency toward the predominance of a distinct parasite population over the other parasite populations present in the isolate. These observations suggest that adaptation of *P. falciparum* parasites to in vitro conditions is a complex phenomenon resulting from (and/or leading to) biochemical and genetic modifications, which are in large part still unfathomed.

Once the first hurdle is cleared and the parasites become adapted to in vitro cultivation, or even during the adaptation period, the selection of parasite population(s) may or may not occur. As emphasized by Thompson and Lymbery [341], "a strain may include one or more populations". The accumulated experimental evidence has demonstrated that, during the first few days or weeks of continuous in vitro cultivation, phenotype changes occur in many *P. falciparum* isolates, including drug response, cytoadherence, and multiplication rates [336,361–363].

After or during adaptation and selection processes, further phenotypic and/or genotypic modifications may also occur in laboratory-adapted *P. falciparum* strains or even clones. Phenotypic changes that can be induced by altering in vitro culture conditions were examined in detail in earlier sections. Briefly, such changes include multiplication and invasion rates, duration of the intraerythrocytic cycle, drug response, transcriptomic profile, the number and size of knobs, the number of variant surface antigens, and cytoadhesion to endothelial receptors. The major factors that have been shown to influence the phenotype include serum supplement (human serum versus Albumax®), O₂ level, suspension culture, and hypoxanthine supplement. Some of these phenotypic changes may occur due to the cultivation conditions that the parasites encounter, i.e., they may be induced, knowingly or unknowingly, by investigators themselves. Genotypic changes likewise occur spontaneously during long-term cultivation in laboratory-adapted strains and clones, both at nucleotide and chromosome levels, which may or may not be associated with phenotype changes [353,364–366]. The "spontaneous" loss of knobs during long-term cultivation and the underlying genetic mechanisms have been studied for decades [367–370]. The underlying mechanisms are complex and still incompletely understood, and may be strain-dependent, including the selection of parasite population(s), point mutations, deletion, gene amplification, and differential transcription level. Adaptation, selection, and phenotypic and/or genotypic modifications may occur either simultaneously or in rapid succession and in a different order. There is still much work to be conducted for a better comprehension of malaria biology.
(clag), variant genes (var; encoding *P. falciparum* erythrocyte membrane protein 1, PfEMP-1), repetitive interspersed families (rif), and subtelomeric variable open reading frame (stevor) genes, which confer phenotypic diversity to genetically homogeneous parasite populations, which in turn may confer biological advantages, especially fitness, to the parasites to allow adaptation to different environments [355,356]. There is still limited experimental evidence that supports these recent findings. More studies are required to further understand whether there is a single set of genetic modifications associated with adaptation to in vitro conditions.

Continuous cultivation of mixed parasite populations usually, but not always, results in a diminution of the initial MOI and may lead to a selection of the dominant parasite population, possibly endowed with biological fitness [357,358]. For unknown reasons, several parasite populations initially present in the day 0 isolate may also continue to co-exist in vitro, and a "new parasite population" undetected by the polymerase chain reaction (PCR) in the initial isolate on day 0 may emerge within a short time period (i.e., within 9 days of continuous cultivation or four asexual intraerythrocytic cycles) [359]. In a study based on microsatellite genotyping, single nucleotide polymorphisms (SNPs) of drug resistance markers and phenotypic profiles (drug response), it was reported that three fresh Kenyan clinical isolates of *P. falciparum* exhibited various genotypic and phenotypic profiles, without any pattern or phenotype-genotype correlation, during the 2-3 month cultivation period [360]. In that study, the stabilization of the genotypic or phenotypic profile was not observed, and there was no apparent tendency toward the predominance of a distinct parasite population over the other parasite populations present in the isolate. These observations suggest that adaptation of *P. falciparum* parasites to in vitro conditions is a complex phenomenon resulting from (and/or leading to) biochemical and genetic modifications, which are in large part still unfathomed.

Once the first hurdle is cleared and the parasites become adapted to in vitro cultivation, or even during the adaptation period, the selection of parasite population(s) may or may not occur. As emphasized by Thompson and Lymbery [341], "a strain may include one or more populations". The accumulated experimental evidence has demonstrated that, during the first few days or weeks of continuous in vitro cultivation, phenotype changes occur in many *P. falciparum* isolates, including drug response, cytoadherence, and multiplication rates [336,361–363].

After or during adaptation and selection processes, further phenotypic and/or genotypic modifications may also occur in laboratory-adapted *P. falciparum* strains or even clones. Phenotypic changes that can be induced by altering in vitro culture conditions were examined in detail in earlier sections. Briefly, such changes include multiplication and invasion rates, duration of the intraerythrocytic cycle, drug response, transcriptomic profile, the number and size of knobs, the number of variant surface antigens, and cytoadhesion to endothelial receptors. The major factors that have been shown to influence the phenotype include serum supplement (human serum versus Albumax®), O₂ level, suspension culture, and hypoxanthine supplement. Some of these phenotypic changes may occur due to the cultivation conditions that the parasites encounter, i.e., they may be induced, knowingly or unknowingly, by investigators themselves. Genotypic changes likewise occur spontaneously during long-term cultivation in laboratory-adapted strains and clones, both at nucleotide and chromosome levels, which may or may not be associated with phenotype changes [353,364–366]. The "spontaneous" loss of knobs during long-term cultivation and the underlying genetic mechanisms have been studied for decades [367–370]. The underlying mechanisms are complex and still incompletely understood, and may be strain-dependent, including the selection of parasite population(s), point mutations, deletion, gene amplification, and differential transcription level. Adaptation, selection, and phenotypic and/or genotypic modifications may occur either simultaneously or in rapid succession and in a different order. There is still much work to be conducted for a better comprehension of malaria biology.
Given the large flexibility of the parasites at the individual level, the notion of clonality can be challenged. Right after cloning, when a single, isolated parasite in an infected erythrocyte multiplies "exponentially" after each intraerythrocytic cycle, some, but not all individual clones constituting the initially clonal population may undergo phenotypic and/or genotypic modifications, resulting in multiple populations, which are not genetically homogeneous. The individual clones may be subject to different cultivation conditions within the same culture vessel (e.g., in static cultures, those in the upper layer of the erythrocytes may have better access to nutrients and may have better gas exchange with the medium). Others may undergo mutations. Some of the resulting diverse populations may become more biologically fit and may be selected over time, leading to the emergence of a new population with different phenotypic and genotypic characteristics. Unless clonal populations are cloned regularly in a short interval of time, a clone may not remain a clone for a long time. One way to test this hypothesis is to compare the whole genome of the 3D7 clone used today in different laboratories around the world, which may or may not demonstrate its non-clonality.

Biological fitness is the ability of an organism to survive in its environment and reproduce and pass on its genes to the next generation of its offspring. The Darwinian concept of biological fitness and survival applied to the in vitro propagation of *P. falciparum* parasites may be juxtaposed with adaptation, selection, and phenotypic/genotypic modifications. Phenotypic features of a parasite strain related to higher parasite biomass, such as the multiplication rate, red blood cell selectivity (i.e., the number of multiply infected erythrocytes), the number of merozoites generated in each schizont, and merozoite invasion rate, are quantifiable variables that may reflect biological fitness of the asexual stage parasites [371,372]. In vivo, other phenotypic and genotypic features associated with antigenic variation and cytoadherence (the expression of *var*, *rifin*, and *stevor* multigene families; e.g., *P. falciparum* erythrocyte membrane protein 1, PfEMP1, encoded by the *var* gene family) play an important role in parasite survival and transmission [373–375]. These aforementioned parasite-associated factors that enhance biological fitness in the human host need to be counterbalanced because high virulence, defined as the ability to cause disease in the host, may lead to severe malaria and death, which is obviously unfavorable for parasite survival. A number of hypotheses have been advanced to explain these phenomena, but host–parasite interactions lie beyond the scope of this review and will not be discussed further. Suffice to say that there is evidence that biological fitness, the *P. falciparum* transmission rate, and virulence (i.e., malaria-associated morbidity and mortality) are positively correlated [376,377].

The biological fitness of drug-resistant *P. falciparum* isolates and strains has been studied extensively in the field and laboratory. The capacity to dominate or replace parasite strains during the in vitro co-cultivation of multiple parasite populations or strains has been studied, usually in the context of competition between drug-sensitive versus drug-resistant strains. Generally speaking, in the presence of selection (i.e., drug pressure), drug-resistant mutant parasites are likely to be fitter to propagate in a "hostile" environment, while in the absence of selection, the converse is more likely (i.e., drug-resistant mutant parasites are likely to be less fit than drug-sensitive wild type parasites) [378–380]. In vitro experiments have shown, however, that co-cultures initiated with equal numbers of chloroquine-sensitive and chloroquine-resistant clones in a drug-free medium lead to competition between the two clones and, after 11 days of continuous cultivation, the chloroquine-sensitive clone does not necessarily end up predominating the chloroquine-resistant clone [381]. When the same experiments were repeated with 50 nM chloroquine added to the culture medium (corresponding to > IC₅₀ for the chloroquine-sensitive clone but much below the IC₅₀ for the chloroquine-resistant clone), as expected, the chloroquine-resistant clone largely predominated over the chloroquine-sensitive clone, demonstrating the role of mutations associated with drug resistance in conferring biological fitness in an adverse environment.
Given the large flexibility of the parasites at the individual level, the notion of clonality
can be challenged. Right after cloning, when a single, isolated parasite in an infected
erythrocyte multiplies "exponentially" after each intraerythrocytic cycle, some, but not
all individual clones constituting the initially clonal population may undergo phenotypic
and/or genotypic modifications, resulting in multiple populations, which are not geneti-
cally homogeneous. The individual clones may be subject to different cultivation conditions
within the same culture vessel (e.g., in static cultures, those in the upper layer of the ery-
throcytes may have better access to nutrients and may have better gas exchange with the
medium). Others may undergo mutations. Some of the resulting diverse populations may
become more biologically fit and may be selected over time, leading to the emergence of
a new population with different phenotypic and genotypic characteristics. Unless clonal
populations are cloned regularly in a short interval of time, a clone may not remain a clone
for a long time. One way to test this hypothesis is to compare the whole genome of the
3D7 clone used today in different laboratories around the world, which may or may not
demonstrate its non-clonality.

Biological fitness is the ability of an organism to survive in its environment and repro-
duce and pass on its genes to the next generation of its offspring. The Darwinian concept of
biological fitness and survival applied to the in vitro propagation of *P. falciparum* parasites
may be juxtaposed with adaptation, selection, and phenotypic/genotypic modifications.
Phenotypic features of a parasite strain related to higher parasite biomass, such as the
multiplication rate, red blood cell selectivity (i.e., the number of multiply infected ery-
throcytes), the number of merozoites generated in each schizont, and merozoite invasion
rate, are quantifiable variables that may reflect biological fitness of the asexual stage para-
sites [371,372]. In vivo, other phenotypic and genotypic features associated with antigenic
variation and cytoadherence (the expression of *var*, *rifin*, and *stevor* multigene families;
e.g., *P. falciparum* erythrocyte membrane protein 1, PfEMP1, encoded by the *var* gene family)
play an important role in parasite survival and transmission [373–375]. These aforemen-
tioned parasite-associated factors that enhance biological fitness in the human host need to
be counterbalanced because high virulence, defined as the ability to cause disease in the
host, may lead to severe malaria and death, which is obviously unfavorable for parasite
survival. A number of hypotheses have been advanced to explain these phenomena, but
host–parasite interactions lie beyond the scope of this review and will not be discussed
further. Suffice to say that there is evidence that biological fitness, the *P. falciparum* trans-
mission rate, and virulence (i.e., malaria-associated morbidity and mortality) are positively
correlated [376,377].

The biological fitness of drug-resistant *P. falciparum* isolates and strains has been stud-
ied extensively in the field and laboratory. The capacity to dominate or replace parasite
strains during the in vitro co-cultivation of multiple parasite populations or strains has
been studied, usually in the context of competition between drug-sensitive versus drug-
resistant strains. Generally speaking, in the presence of selection (i.e., drug pressure),
drug-resistant mutant parasites are likely to be fitter to propagate in a "hostile" environ-
ment, while in the absence of selection, the converse is more likely (i.e., drug-resistant
mutant parasites are likely to be less fit than drug-sensitive wild type parasites) [378–380].
In vitro experiments have shown, however, that co-cultures initiated with equal numbers
of chloroquine-sensitive and chloroquine-resistant clones in a drug-free medium lead to
competition between the two clones and, after 11 days of continuous cultivation, the
chloroquine-sensitive clone does not necessarily end up predominating the chloroquine-
resistant clone [381]. When the same experiments were repeated with 50 nM chloroquine
added to the culture medium (corresponding to > IC₅₀ for the chloroquine-sensitive clone
but much below the IC₅₀ for the chloroquine-resistant clone), as expected, the chloroquine-
resistant clone largely predominated over the chloroquine-sensitive clone, demonstrating
the role of mutations associated with drug resistance in conferring biological fitness in an
adverse environment.
In another experiment, it was shown that when compared to a drug-sensitive strain (HB3/Honduras), a multidrug-resistant strain (Dd2/Indochina) exhibited (i) a shorter intraerythrocytic cycle (44 h versus 50 h, respectively, in a standard culture medium supplemented with 0.5% Albumax® II and cultivated in 5% CO₂, 5% O₂, and 90% N₂), (ii) a higher number of merozoites per schizont (mean number, 18 versus 16 merozoites per schizont), and (iii) a higher invasion rate (i.e., a higher number of re-infected erythrocytes) underlying its faster growth rate [236]. It is not yet clear whether *P. falciparum* has a "biological clock" to regulate its metabolic activities on its own or whether the mammalian host imposes rhythmicity on the parasites, including the duration of the intraerythrocytic cycle [382,383]. A study has suggested that the mammalian host's circadian rhythm may be controlling the synchronicity of the asexual intraerythrocytic life cycle of malaria parasites, possibly via melatonin or its precursors (e.g., serotonin) [384]. Another study has suggested that the rhythmic life cycle of each *P. falciparum* strain may be genetically programmed and driven by its endogenous "cell-cycle oscillator", independent from the human host [385]. Experiments have shown that the intrinsic duration of the intraerythrocytic cycle may vary from 36 to 54 h, depending on the *P. falciparum* strain cultivated in vitro in standard conditions (suspension cultures using RPMI 1640 supplemented with 10% human serum, 5% O₂, 5% CO₂, and 90% N₂ at 37 °C), but naturally occurring parasites adapt and align themselves with the 24 h host circadian biological clock. The precise molecular mechanisms are still under study, but similar findings were reported on *P. chabaudi* in mice [386]. The observation that the duration of the intraerythrocytic cycle of a parasite strain can be shortened or prolonged by changing the culture conditions (e.g., O₂ concentration) may suggest that it is more of a survival strategy in response to an environmental change, rather than a "fixed" phenotypic/genotypic characteristic of a given parasite strain [214,215,266,267]. Of the three main differential phenotypic factors reported by Reilly et al. [236], the number of merozoites produced by individual schizonts and invasion rates seem to be the determinant factors directly related to differential growth rates, i.e., biological fitness, in different parasite strains. The underlying genetic basis of these phenotypic features of biological fitness is still under study [387].

Many factors that influence parasite growth in vitro were comprehensively reviewed in this paper. In some areas of research, more experimental data are required for a better comprehension of malaria biology. One such area is the interplay between adaptation, selection, phenotypic and genotypic modifications, rhythmicity (or chronobiology), and biological fitness. There are reasons to believe that laboratory-adapted *P. falciparum* strains or clones may not totally behave or respond like the parasites circulating in nature. If possible, experiments conducted in the laboratory using *P. falciparum* clones and strains need to be validated using field isolates. As a final word, with all its advantages and (some) disadvantages, the in vitro cultivation of *P. falciparum* is a model system that has been, and will continue to be, an essential tool for malaria research in the coming years.

## 16. Conclusions

The development of the technique for the continuous in vitro cultivation of asexual blood stages of *P. falciparum* is one of the major advances in malariology in the 20th century. The technique, developed in parallel and independently by Haynes et al. [26] and Trager and Jensen [27], has allowed several generations of scientists to work directly with the human pathogen and reduced or eliminated their dependence on (i) short-term, suboptimal cultures of *P. falciparum*, (ii) useful but less relevant avian, rodent, and simian malaria models, and (iii) experimentally infected monkeys as sources of parasites. It has provided a large amount of parasite materials for biological, biochemical, immunological, and pharmacological studies. Most fields in malaria research have been profoundly influenced and benefited from continuous cultivation techniques [13]. The basic principles laid out by Haynes et al. [26] and Trager and Jensen [27] remain valid and are largely unchanged more than 45 years after their discovery. Still, there are some areas where improvements are possible.
In another experiment, it was shown that when compared to a drug-sensitive strain (HB3/Honduras), a multidrug-resistant strain (Dd2/Indochina) exhibited (i) a shorter intraerythrocytic cycle (44 h versus 50 h, respectively, in a standard culture medium supplemented with 0.5% Albumax® II and cultivated in 5% CO₂, 5% O₂, and 90% N₂), (ii) a higher number of merozoites per schizont (mean number, 18 versus 16 merozoites per schizont), and (iii) a higher invasion rate (i.e., a higher number of re-infected erythrocytes) underlying its faster growth rate [236]. It is not yet clear whether *P. falciparum* has a "biological clock" to regulate its metabolic activities on its own or whether the mammalian host imposes rhythmicity on the parasites, including the duration of the intraerythrocytic cycle [382,383]. A study has suggested that the mammalian host's circadian rhythm may be controlling the synchronicity of the asexual intraerythrocytic life cycle of malaria parasites, possibly via melatonin or its precursors (e.g., serotonin) [384]. Another study has suggested that the rhythmic life cycle of each *P. falciparum* strain may be genetically programmed and driven by its endogenous "cell-cycle oscillator", independent from the human host [385]. Experiments have shown that the intrinsic duration of the intraerythrocytic cycle may vary from 36 to 54 h, depending on the *P. falciparum* strain cultivated in vitro in standard conditions (suspension cultures using RPMI 1640 supplemented with 10% human serum, 5% O₂, 5% CO₂, and 90% N₂ at 37 °C), but naturally occurring parasites adapt and align themselves with the 24 h host circadian biological clock. The precise molecular mechanisms are still under study, but similar findings were reported on *P. chabaudi* in mice [386]. The observation that the duration of the intraerythrocytic cycle of a parasite strain can be shortened or prolonged by changing the culture conditions (e.g., O₂ concentration) may suggest that it is more of a survival strategy in response to an environmental change, rather than a "fixed" phenotypic/genotypic characteristic of a given parasite strain [214,215,266,267]. Of the three main differential phenotypic factors reported by Reilly et al. [236], the number of merozoites produced by individual schizonts and invasion rates seem to be the determinant factors directly related to differential growth rates, i.e., biological fitness, in different parasite strains. The underlying genetic basis of these phenotypic features of biological fitness is still under study [387].

Many factors that influence parasite growth in vitro were comprehensively reviewed in this paper. In some areas of research, more experimental data are required for a better comprehension of malaria biology. One such area is the interplay between adaptation, selection, phenotypic and genotypic modifications, rhythmicity (or chronobiology), and biological fitness. There are reasons to believe that laboratory-adapted *P. falciparum* strains or clones may not totally behave or respond like the parasites circulating in nature. If possible, experiments conducted in the laboratory using *P. falciparum* clones and strains need to be validated using field isolates. As a final word, with all its advantages and (some) disadvantages, the in vitro cultivation of *P. falciparum* is a model system that has been, and will continue to be, an essential tool for malaria research in the coming years.

**16. Conclusions**

The development of the technique for the continuous in vitro cultivation of asexual blood stages of *P. falciparum* is one of the major advances in malariology in the 20th century. The technique, developed in parallel and independently by Haynes et al. [26] and Trager and Jensen [27], has allowed several generations of scientists to work directly with the human pathogen and reduced or eliminated their dependence on (i) short-term, suboptimal cultures of *P. falciparum*, (ii) useful but less relevant avian, rodent, and simian malaria models, and (iii) experimentally infected monkeys as sources of parasites. It has provided a large amount of parasite materials for biological, biochemical, immunological, and pharmacological studies. Most fields in malaria research have been profoundly influenced and benefited from continuous cultivation techniques [13]. The basic principles laid out by Haynes et al. [26] and Trager and Jensen [27] remain valid and are largely unchanged more than 45 years after their discovery. Still, there are some areas where improvements are possible.
RPMI 1640 is the standard medium for *P. falciparum* cultivation. However, this medium was not specifically designed for malaria cultivation. Other media may support parasite growth as well as, or better, than RPMI 1640. Further studies with alternative media may be warranted. The requirement for human serum is a major disadvantage in the original technique described by Trager and Jensen, especially in developing countries. The use of Albumax® as a substitute for human serum represents a major step toward the development of a serum-free, chemically defined medium. Further studies may identify the essential fatty acids required for optimal propagation of the parasites, foregoing even the use of Albumax®. At present, human erythrocytes are required for parasite propagation.

Lastly, the unfinished work of the late Professor William Trager on axenic culture
should inspire and encourage young scientists to pursue this line of work. When some
of these modifications, notably a serum-free culture system, are successfully attained, the
cultivation of *P. falciparum* may become a routine procedure, even in developing countries
where malaria is endemic.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** The original draft of this work was initially written by the same author as a companion and introduction to the "Field application of in vitro assays for the sensitivity of human malaria parasites to antimalarial drugs", published by the World Health Organization in 2007, but has not been published before. The final work presented here is an update including peer-reviewed articles mostly indexed on PubMed up to February 2023.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** I am deeply grateful to Pascal Ringwald (Global Malaria Programme, World Health Organization, Geneva, Switzerland), Harald Noedl (Institute of Specific Prophylaxis and Tropical Medicine, Medical University of Vienna, Austria; Malaria Research Initiative Bandarban, Vienna, Austria), and the late David Charles Warhurst (Emeritus Professor of Protozoan Chemotherapy, London School of Hygiene and Tropical Medicine, London, UK) for critical reading of the early draft of this paper in 2004–2005. The present paper was partly inspired by the “representatives” of two American research teams that developed the cultivation system, whom I had the immense honor to meet during my younger years. I met the late William Trager, who had the kindness and generosity to visit and discuss with graduate students attending the summer course on “Biology of Parasitism: Modern Approaches” at Marine Biological Laboratories, Woods Hole, Massachusetts, in 1994. Later, around the year 2000, I also had the great pleasure to meet and have a face-to-face talk with J. David Haynes during my assignment to Yaoundé, Cameroon, when he came on a private visit to see his sister working in Cameroon. My gratitude also goes to Jacques Le Bras (former head of the department, Laboratoire de Parasitologie, Hôpital Bichat-Claude Bernard, Paris, France), who learned first-hand the cultivation techniques from the late William Trager at Rockefeller University (New York, NY) and who transmitted to me his know-how on in vitro cultivation as my mentor in Paris, France, in 1988–1996.

**Conflicts of Interest:** The author declares no conflict of interest.

**Abbreviations**

ACD, acid-citrate-dextrose; *ast*, gene encoding aspartate transaminase; *atg4*, gene encoding cysteine protease autophagy-related 4; ATP, adenosine triphosphate; BES, N,N'-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid; BME, basal medium Eagle; *clag*, cytoadherence linked asexual gene; CMRL-1066, the medium developed at the Connaugh Medical Research Laboratories; COI, complexity of infection; CPD, citrate-phosphate-dextrose; CPD-A, modified citrate-phosphate-dextrose with 0.2 g/L adenine; DHFR, dihydrofolate reductase; DHPS, dihydropteroate synthase; DMEM, Dulbecco's modified Eagle's medium; DOC2, double C2-like domain-containing protein; *eba-140*, gene encoding erythrocyte-binding antigen-140; G6PD deficiency, glucose-6-phosphate dehydrogenase deficiency; HDL, high-density lipoprotein; HEPES, N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid; IC50, 50% inhibitory concentration; IMDM, Iscove's modified Dulbecco's medium; LDL, low-density lipoprotein; M-199, medium 199; MCH, mean corpuscular hemoglobin; MCV, mean corpus-
RPMI 1640 is the standard medium for *P. falciparum* cultivation. However, this medium was not specifically designed for malaria cultivation. Other media may support parasite growth as well as, or better, than RPMI 1640. Further studies with alternative media may be warranted. The requirement for human serum is a major disadvantage in the original technique described by Trager and Jensen, especially in developing countries. The use of Albumax® as a substitute for human serum represents a major step toward the development of a serum-free, chemically defined medium. Further studies may identify the essential fatty acids required for optimal propagation of the parasites, foregoing even the use of Albumax®. At present, human erythrocytes are required for parasite propagation.

Lastly, the unfinished work of the late Professor William Trager on axenic culture
should inspire and encourage young scientists to pursue this line of work. When some
of these modifications, notably a serum-free culture system, are successfully attained, the
cultivation of *P. falciparum* may become a routine procedure, even in developing countries
where malaria is endemic.

**Funding:** This research received no external funding.

**Institutional Review Board Statement:** The original draft of this work was initially written by the same author as a companion and introduction to the "Field application of in vitro assays for the sensitivity of human malaria parasites to antimalarial drugs", published by the World Health Organization in 2007, but has not been published before. The final work presented here is an update including peer-reviewed articles mostly indexed on PubMed up to February 2023.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Acknowledgments:** I am deeply grateful to Pascal Ringwald (Global Malaria Programme, World Health Organization, Geneva, Switzerland), Harald Noedl (Institute of Specific Prophylaxis and Tropical Medicine, Medical University of Vienna, Austria; Malaria Research Initiative Bandarban, Vienna, Austria), and the late David Charles Warhurst (Emeritus Professor of Protozoan Chemotherapy, London School of Hygiene and Tropical Medicine, London, UK) for critical reading of the early draft of this paper in 2004–2005. The present paper was partly inspired by the “representatives” of two American research teams that developed the cultivation system, whom I had the immense honor to meet during my younger years. I met the late William Trager, who had the kindness and generosity to visit and discuss with graduate students attending the summer course on “Biology of Parasitism: Modern Approaches” at Marine Biological Laboratories, Woods Hole, Massachusetts, in 1994. Later, around the year 2000, I also had the great pleasure to meet and have a face-to-face talk with J. David Haynes during my assignment to Yaoundé, Cameroon, when he came on a private visit to see his sister working in Cameroon. My gratitude also goes to Jacques Le Bras (former head of the department, Laboratoire de Parasitologie, Hôpital Bichat-Claude Bernard, Paris, France), who learned first-hand the cultivation techniques from the late William Trager at Rockefeller University (New York, NY) and who transmitted to me his know-how on in vitro cultivation as my mentor in Paris, France, in 1988–1996.

**Conflicts of Interest:** The author declares no conflict of interest.

**Abbreviations**

ACD, acid-citrate-dextrose; *ast*, gene encoding aspartate transaminase; *atg4*, gene encoding cysteine protease autophagy-related 4; ATP, adenosine triphosphate; BES, N,N'-bis(2-hydroxyethyl)-2-aminoethanesulfonic acid; BME, basal medium Eagle; *clag*, cytoadherence linked asexual gene; CMRL-1066, the medium developed at the Connaugh Medical Research Laboratories; COI, complexity of infection; CPD, citrate-phosphate-dextrose; CPD-A, modified citrate-phosphate-dextrose with 0.2 g/L adenine; DHFR, dihydrofolate reductase; DHPS, dihydropteroate synthase; DMEM, Dulbecco's modified Eagle's medium; DOC2, double C2-like domain-containing protein; *eba-140*, gene encoding erythrocyte-binding antigen-140; G6PD deficiency, glucose-6-phosphate dehydrogenase deficiency; HDL, high-density lipoprotein; HEPES, N-2-hydroxyethylpiperazine-N'-2-ethanesulfonic acid; IC₅₀, 50% inhibitory concentration; IMDM, Iscove's modified Dulbecco's medium; LDL, low-density lipoprotein; M-199, medium 199; MCH, mean corpuscular hemoglobin; MCV, mean corpus-
cular volume; MEM, minimum essential medium; MOI, multiplicity of infection; MSA-1, merozoite surface antigen-1; MSA-2, merozoite surface antigen-2; NCTC 135, the medium formulated at the National Cancer Institute, Tissue Culture section (Bethesda, MD) for the L929 strain of mouse cells; PABA, *para*-aminobenzoic acid; PCR, polymerase chain reaction; PfEMP-1, *P. falciparum* erythrocyte membrane protein-1, expression product of *var* gene; PSAC, plasmodial surface anion channel; *rif*, gene encoding repetitive interspersed family; RIFIN, repetitive interspersed family; ROS, reactive oxygen species; rpm, rotations per minute; RPMI 1640, Roswell Park Memorial Institute cell culture medium 1640 (RPMI 1640); SAG, saline-adenine-glucose; SAGM, saline-adenine-glucose + mannitol; SNP, single nucleotide polymorphism; SRPK1, serine/threonine protein kinase 1; *stevor*, subtelomeric variable open reading frame gene family; *surf*, surface-associated interspersed gene family encoding SURFIN protein; TES, N-tris-(hydroxymethyl)methyl-2-aminoethanesulfonic acid; VLDL, very-low-density lipoprotein; WHO, World Health Organization; WRAIR, Walter Reed Army Institute of Research.

## References

1. Garnham, P.C.C. Malaria in its various vertebrate hosts. In *Malaria: Epidemiology, Chemotherapy, Morphology, and Metabolism*; Kreier, J.P., Ed.; Academic Press, Inc.: London, UK, 1980; Volume 1, pp. 95–144.

2. Wernsdorfer, W.H. The importance of malaria in the world. In *Malaria: Epidemiology, Chemotherapy, Morphology, and Metabolism*; Kreier, J.P., Ed.; Academic Press, Inc.: London, UK, 1980; Volume 1, pp. 1–93.

3. Sinden, R.E.; Gilles, H.M. The malaria parasites. In *Essential Malariology*, 4th ed.; Warrell, D.A., Gilles, H.M., Eds.; Arnold Publishers: London, UK, 2002; pp. 8–34.

4. Garnham, P.C.C. Malaria parasites of man: Life-cycles and morphology (excluding ultrastructure). In *Malaria: Principles and Practice of Malariology*; Wernsdorfer, W.H., McGregor, I., Eds.; Churchill Livingstone Inc., Longman Group UK Ltd.: Edinburgh, UK, 1988; Volume 1, pp. 61–96.

5. White, N.J.; Pukrittayakamee, S.; Hien, T.T.; Faiz, M.A.; Mokuolu, O.A.; Dondorp, A.M. Malaria. *Lancet* **2014**, *383*, 723–735.
[CrossRef] [PubMed]

6. Langreth, S.G.; Jensen, J.B.; Reese, R.T.; Trager, W. Fine structure of human malaria in vitro. *J. Protozool.* **1978**, *25*, 443–452.
[CrossRef] [PubMed]

7. Bannister, L.H.; Hopkins, J.M.; Fowler, R.E.; Krishna, S.; Mitchell, G.H. A brief illustrated guide to the ultrastructure of *Plasmodium falciparum* asexual blood stages. *Parasitol. Today* **2000**, *16*, 427–433. [CrossRef]

8. Wernsdorfer, W.H.; Wernsdorfer, M.G. The evaluation of in vitro tests for the assessment of drug response in *Plasmodium falciparum*. *Mitt. Österr. Ges. Tropenmed. Parasitol.* **1995**, *17*, 221–228.

9. Cowman, A.F.; Crabb, B.S. Invasion of red blood cells by malaria parasites. *Cell* **2006**, *124*, 755–766. [CrossRef]

10. Ngotho, P.; Soares, A.B.; Hentzschel, F.; Achcar, F.; Bertuccini, L.; Marti, M. Revisiting gametocyte biology in malaria parasites. *FEMS Microbiol. Rev.* **2019**, *43*, 401–414, Erratum in: *FEMS Microbiol. Rev.* **2019**, *43*, 684. [CrossRef] [PubMed]

11. Ferreira, M.U.; Nobrega de Sousa, T.; Rangel, G.W.; Johansen, I.C.; Corder, R.M.; Ladeia-Andrade, S.; Gil, J.P. Monitoring *Plasmodium vivax* resistance to antimalarials: Persisting challenges and future directions. *Int. J. Parasitol. Drugs Drug Resist.* **2021**, *15*, 9–24. [CrossRef]

12. Millar, S.B.; Cox-Singh, J. Human infections with *Plasmodium knowlesi*—Zoonotic malaria. *Clin. Microbiol. Infect.* **2015**, *21*, 640–648.
[CrossRef]

13. Trager, W.; Jensen, J.B. Continuous culture of *Plasmodium falciparum*: Its impact on malaria research. *Int. J. Parasitol.* **1997**, *27*, 989–1006. [CrossRef]

14. Geiman, Q.M. In vitro methods for study and cultivation of *Plasmodia*. In *Malariology. A Comprehensive Survey of All Aspects of This Group of Diseases from a Global Standpoint*; Boyd, M.F., Ed.; W.B. Saunders Company: Philadelphia, PA, USA, 1949; Volume 1, pp. 205–222.

15. Bertagna, P.; Cohen, S.; Geiman, Q.M.; Haworth, J.; König, E.; Richards, W.H.G.; Trigg, P.I. Cultivation techniques for the erythrocytic stages of malaria parasites. *Bull. World Health Organ.* **1972**, *47*, 357–373.

16. Trager, W.; Jensen, J.B. Cultivation of erythrocytic and exoerythrocytic stages of *Plasmodia*. In *Malaria. Pathology, Vector Studies, and Culture*; Kreier, J.P., Ed.; Academic Press: London, UK, 1980; Volume 2, pp. 271–319.

17. Bass, C.C. A new conception of immunity. Its application to the cultivation of protozoa and bacteria from the blood and to therapeutic measures. *J. Am. Med. Assoc.* **1911**, *57*, 1534–1535. [CrossRef]

18. Bass, C.C.; Johns, F.M. The cultivation of malarial plasmodia (*Plasmodium vivax* and *Plasmodium falciparum*) in vitro. *J. Exp. Med.* **1912**, *16*, 567–579. [CrossRef]

19. Rieckmann, K.H.; McNamara, J.V.; Frischer, H.; Stockert, T.A.; Carson, P.E.; Powell, R.D. Effects of chloroquine, quinine, and cycloguanil upon the maturation of asexual erythrocytic forms of two strains of *Plasmodium falciparum* in vitro. *Am. J. Trop. Med. Hyg.* **1968**, *17*, 661–671. [CrossRef] [PubMed]
cular volume; MEM, minimum essential medium; MOI, multiplicity of infection; MSA-1, merozoite surface antigen-1; MSA-2, merozoite surface antigen-2; NCTC 135, the medium formulated at the National Cancer Institute, Tissue Culture section (Bethesda, MD) for the L929 strain of mouse cells; PABA, para-aminobenzoic acid; PCR, polymerase chain reaction; PfEMP-1, *P. falciparum* erythrocyte membrane protein-1, expression product of var gene; PSAC, plasmodial surface anion channel; rif, gene encoding repetitive interspersed family; RIFIN, repetitive interspersed family; ROS, reactive oxygen species; rpm, rotations per minute; RPMI 1640, Roswell Park Memorial Institute cell culture medium 1640 (RPMI 1640); SAG, saline-adenine-glucose; SAGM, saline-adenine-glucose + mannitol; SNP, single nucleotide polymorphism; SRPK1, serine/threonine protein kinase 1; stevor, subtelomeric variable open reading frame gene family; surf, surface-associated interspersed gene family encoding SURFIN protein; TES, N-tris-(hydroxymethyl)methyl-2-aminoethanesulfonic acid; VLDL, very-low-density lipoprotein; WHO, World Health Organization; WRAIR, Walter Reed Army Institute of Research.

## References

1. Garnham, P.C.C. Malaria in its various vertebrate hosts. In *Malaria: Epidemiology, Chemotherapy, Morphology, and Metabolism*; Kreier, J.P., Ed.; Academic Press, Inc.: London, UK, 1980; Volume 1, pp. 95–144.

2. Wernsdorfer, W.H. The importance of malaria in the world. In *Malaria: Epidemiology, Chemotherapy, Morphology, and Metabolism*; Kreier, J.P., Ed.; Academic Press, Inc.: London, UK, 1980; Volume 1, pp. 1–93.

3. Sinden, R.E.; Gilles, H.M. The malaria parasites. In *Essential Malariology*, 4th ed.; Warrell, D.A., Gilles, H.M., Eds.; Arnold Publishers: London, UK, 2002; pp. 8–34.

4. Garnham, P.C.C. Malaria parasites of man: Life-cycles and morphology (excluding ultrastructure). In *Malaria: Principles and Practice of Malariology*; Wernsdorfer, W.H., McGregor, I., Eds.; Churchill Livingstone Inc., Longman Group UK Ltd.: Edinburgh, UK, 1988; Volume 1, pp. 61–96.

5. White, N.J.; Pukrittayakamee, S.; Hien, T.T.; Faiz, M.A.; Mokuolu, O.A.; Dondorp, A.M. Malaria. *Lancet* **2014**, 383, 723–735.
[CrossRef] [PubMed]

6. Langreth, S.G.; Jensen, J.B.; Reese, R.T.; Trager, W. Fine structure of human malaria in vitro. *J. Protozool.* **1978**, 25, 443–452.
[CrossRef] [PubMed]

7. Bannister, L.H.; Hopkins, J.M.; Fowler, R.E.; Krishna, S.; Mitchell, G.H. A brief illustrated guide to the ultrastructure of *Plasmodium falciparum* asexual blood stages. *Parasitol. Today* **2000**, 16, 427–433. [CrossRef]

8. Wernsdorfer, W.H.; Wernsdorfer, M.G. The evaluation of in vitro tests for the assessment of drug response in *Plasmodium falciparum*. *Mitt. Österr. Ges. Tropenmed. Parasitol.* **1995**, 17, 221–228.

9. Cowman, A.F.; Crabb, B.S. Invasion of red blood cells by malaria parasites. *Cell* **2006**, 124, 755–766. [CrossRef]

10. Ngotho, P.; Soares, A.B.; Hentzschel, F.; Achcar, F.; Bertuccini, L.; Marti, M. Revisiting gametocyte biology in malaria parasites. *FEMS Microbiol. Rev.* **2019**, *43*, 401–414, Erratum in: *FEMS Microbiol. Rev.* **2019**, *43*, 684. [CrossRef] [PubMed]

11. Ferreira, M.U.; Nobrega de Sousa, T.; Rangel, G.W.; Johansen, I.C.; Corder, R.M.; Ladeia-Andrade, S.; Gil, J.P. Monitoring *Plasmodium vivax* resistance to antimalarials: Persisting challenges and future directions. *Int. J. Parasitol. Drugs Drug Resist.* **2021**, *15*, 9–24. [CrossRef]

12. Millar, S.B.; Cox-Singh, J. Human infections with *Plasmodium knowlesi*—Zoonotic malaria. *Clin. Microbiol. Infect.* **2015**, *21*, 640–648.
[CrossRef]

13. Trager, W.; Jensen, J.B. Continuous culture of *Plasmodium falciparum*: Its impact on malaria research. *Int. J. Parasitol.* **1997**, *27*, 989–1006. [CrossRef]

14. Geiman, Q.M. In vitro methods for study and cultivation of *Plasmodia*. In *Malariology. A Comprehensive Survey of All Aspects of This Group of Diseases from a Global Standpoint*; Boyd, M.F., Ed.; W.B. Saunders Company: Philadelphia, PA, USA, 1949; Volume 1, pp. 205–222.

15. Bertagna, P.; Cohen, S.; Geiman, Q.M.; Haworth, J.; König, E.; Richards, W.H.G.; Trigg, P.I. Cultivation techniques for the erythrocytic stages of malaria parasites. *Bull. World Health Organ.* **1972**, *47*, 357–373.

16. Trager, W.; Jensen, J.B. Cultivation of erythrocytic and exoerythrocytic stages of *Plasmodia*. In *Malaria. Pathology, Vector Studies, and Culture*; Kreier, J.P., Ed.; Academic Press: London, UK, 1980; Volume 2, pp. 271–319.

17. Bass, C.C. A new conception of immunity. Its application to the cultivation of protozoa and bacteria from the blood and to therapeutic measures. *J. Am. Med. Assoc.* **1911**, *57*, 1534–1535. [CrossRef]

18. Bass, C.C.; Johns, F.M. The cultivation of malarial plasmodia (*Plasmodium vivax* and *Plasmodium falciparum*) in vitro. *J. Exp. Med.* **1912**, *16*, 567–579. [CrossRef]

19. Rieckmann, K.H.; McNamara, J.V.; Frischer, H.; Stockert, T.A.; Carson, P.E.; Powell, R.D. Effects of chloroquine, quinine, and cycloguanil upon the maturation of asexual erythrocytic forms of two strains of *Plasmodium falciparum* in vitro. *Am. J. Trop. Med. Hyg.* **1968**, *17*, 661–671. [CrossRef] [PubMed]
20. Ferraroni, J.J.; Speer, C.A.; Hayes, J.; Suzuki, M. Prevalence of chloroquine-resistant falciparum malaria in the Brazilian Amazon. *Am. J. Trop. Med. Hyg.* **1981**, *30*, 526–530. [CrossRef] [PubMed]

21. Stork, G.; Niu, D.Q.; Fujimoto, A.; Koft, E.R.; Balkovec, J.M.; Tata, J.R.; Dake, G.R. The first stereoselective total synthesis of quinine. *J. Am. Chem. Soc.* **2001**, *123*, 3239–3242. [CrossRef]

22. Trager, W. Studies on conditions affecting the survival in vitro of a malarial parasite (*Plasmodium lophurae*). *J. Exp. Med.* **1941**, *74*, 441–462. [CrossRef]

23. Horovitz, A.; Sautet, J. Remarques sur la culture des parasites du paludisme. *Ann. Parasitol. Hum. Comp.* **1929**, *7*, 151–160. [CrossRef]

24. Geiman, Q.M.; Anfinsen, C.B.; McKee, R.W.; Ormsbee, R.A.; Ball, E.G. Studies on malarial parasites VII. Methods and techniques for cultivation. *J. Exp. Med.* **1946**, *84*, 583–606. [CrossRef]

25. Trigg, P.I. Parasite cultivation in relation to research on the chemotherapy of malaria. *Bull. World Health Organ.* **1976**, *53*, 399–406.

26. Haynes, J.D.; Diggs, C.L.; Hines, F.A.; Desjardins, R.E. Culture of human malaria parasites (*P. falciparum*). *Nature* **1976**, *263*, 767–769. [CrossRef] [PubMed]

27. Trager, W.; Jensen, J.B. Human malaria parasites in continuous culture. *Science* **1976**, *193*, 673–675. [CrossRef]

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31. Nguyen-Dinh, P.; Campbell, C.C.; Collins, W.E. Cultivation in vitro of the quartan malaria parasite *Plasmodium inui*. *Science* **1980**, *209*, 1249–1251. [CrossRef]

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21. Stork, G.; Niu, D.Q.; Fujimoto, A.; Koft, E.R.; Balkovec, J.M.; Tata, J.R.; Dake, G.R. The first stereoselective total synthesis of quinine. *J. Am. Chem. Soc.* **2001**, *123*, 3239–3242. [CrossRef]

22. Trager, W. Studies on conditions affecting the survival in vitro of a malarial parasite (*Plasmodium lophurae*). *J. Exp. Med.* **1941**, *74*, 441–462. [CrossRef]

23. Horovitz, A.; Sautet, J. Remarques sur la culture des parasites du paludisme. *Ann. Parasitol. Hum. Comp.* **1929**, *7*, 151–160. [CrossRef]

24. Geiman, Q.M.; Anfinsen, C.B.; McKee, R.W.; Ormsbee, R.A.; Ball, E.G. Studies on malarial parasites VII. Methods and techniques for cultivation. *J. Exp. Med.* **1946**, *84*, 583–606. [CrossRef]

25. Trigg, P.I. Parasite cultivation in relation to research on the chemotherapy of malaria. *Bull. World Health Organ.* **1976**, *53*, 399–406.

26. Haynes, J.D.; Diggs, C.L.; Hines, F.A.; Desjardins, R.E. Culture of human malaria parasites (*P. falciparum*). *Nature* **1976**, *263*, 767–769. [CrossRef] [PubMed]

27. Trager, W.; Jensen, J.B. Human malaria parasites in continuous culture. *Science* **1976**, *193*, 673–675. [CrossRef]

28. Chin, W.; Moss, D.; Collins, W.E. The continuous cultivation of *Plasmodium fragile* by the method of Trager-Jensen. *Am. J. Trop. Med. Hyg.* **1979**, *28*, 591–592. [CrossRef] [PubMed]

29. Butcher, G.A. Factors affecting the in vitro culture of *Plasmodium falciparum* and *Plasmodium knowlesi*. *Bull. World Health Organ.* **1979**, *57* (Suppl. 1), 17–26.

30. Chen, Z.R.; Gao, M.X.; Li, Y.H.; Han, S.M.; Zhang, N.L. Studies on the cultivation of erythrocytic stage *Plasmodium* in vitro. *Chin. Med. J.* **1980**, *93*, 31–35.

31. Nguyen-Dinh, P.; Campbell, C.C.; Collins, W.E. Cultivation in vitro of the quartan malaria parasite *Plasmodium inui*. *Science* **1980**, *209*, 1249–1251. [CrossRef]

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270. Tonhosolo, R.; Gabriel, H.B.; Matsumura, M.Y.; Cabral, F.J.; Yamamoto, M.M.; D'Alexandri, F.L.; Sussmann, R.A.C.; Belmonte, R.; Peres, V.J.; Crick, D.C.; et al. Intraerythrocytic stages of *Plasmodium falciparum* biosynthesize menaquinone. *FEBS Lett.* **2010**, *584*, 4761–4768. [CrossRef]
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295. Pavithra, S.R.; Banumathy, G.; Joy, O.; Singh, V.; Tatu, U. Recurrent fever promotes *Plasmodium falciparum* development in human erythrocytes. *J. Biol. Chem.* **2004**, *279*, 46692–46699. [CrossRef]

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273. Mirovský, P. Continuous culture of *Plasmodium falciparum* asexual stages in "normal" air atmosphere. *Folia Parasitol.* **1989**, *36*, 107–112.

274. Southwell, B.R.; Brown, G.V.; Forsyth, K.P.; Smith, T.; Philip, G.; Anders, R. Field applications of agglutination and cytoadherence assays with *Plasmodium falciparum* from Papua New Guinea. *Trans. R. Soc. Trop. Med. Hyg.* **1989**, *83*, 464–469. [CrossRef] [PubMed]

275. Bei, A.K.; Patel, S.D.; Volkman, S.K.; Ahouidi, A.D.; Ndiaye, D.; Mboup, S.; Wirth, D.F. An adjustable gas-mixing device to increase feasibility of in vitro culture of *Plasmodium falciparum* parasites in the field. *PLoS ONE* **2014**, *9*, e09028. [CrossRef] [PubMed]

276. Rogers, M.D.; Weatherly, N.F.; Bowdre, J.H.; Desjardins, R.E. End-tidal exhaled air for the in vitro maintenance of *Plasmodium falciparum*. *Trans. R. Soc. Trop. Med. Hyg.* **1983**, *77*, 103–105. [CrossRef]

277. Taylor-Robinson, A.W. Cultivation of *Plasmodium falciparum* in vitro using microbiological gas sachets that generate microaerophilic conditions. *Trans. R. Soc. Trop. Med. Hyg.* **1998**, *92*, 357–358. [CrossRef] [PubMed]

278. Onda, T.; Lin, Q.; Kano, S.; Suzuki, M. [Continuous in vitro culture of *Plasmodium falciparum* using microaerophilic gas generators and portable incubator]. *Kansenshogaku Zasshi* **1999**, *73*, 1095–1098. [CrossRef] [PubMed]

279. Mizuno, Y.; Hatabu, T.; Kawazu, S.; Masuda, G.; Ohtomo, H.; Suzuki, M.; Kano, S. Cultivation of *Plasmodium falciparum* isolates under the Anaeropack® gas condition in a portable thermostatic incubator. *Jpn. J. Trop. Med. Hyg.* **2000**, *28*, 383–385. [CrossRef]

280. Haruki, K.; Kobayashi, F.; Fujino, T.; Matsui, T.; Tsuji, M. Evaluation of Anaeropack® (AnP) type as tools of *Plasmodium falciparum* cultivation and drug sensitivity tests. *Jpn. J. Trop. Med. Hyg.* **2001**, *29*, 365–370. [CrossRef]

281. Haruki, K.; Fujino, T.; Yokota, N.; Kobayashi, F.; Matsui, T.; Tsuji, M. Evaluation of AnaeroPack series to determine drug sensitivity of *P. falciparum*. *Jpn. J. Trop. Med.* **1997**, *25*, 87.

282. Lin, Q.; Onda, T.; Kano, S.; Masuda, G.; Suzuki, M. [In vitro drug susceptibility test of *Plasmodium falciparum* using a portable thermostat and CO₂ gas generator]. *Kansenshogaku Zasshi* **1999**, *73*, 1099–1103. [CrossRef]

283. Hatabu, T.; Kawazu, S.; Kojima, S.; Singhasivanon, P.; Krudsood, S.; Looareesuwan, S.; Kano, S. A pilot field trial of an in vitro drug susceptibility test using the Anaeropack® malaria culture system. *Trop. Med. Health* **2004**, *32*, 335–337. [CrossRef]

284. Hatabu, T.; Kawazu, S.; Kojima, S.; Sato, K.; Singhasivanon, P.; Looareesuwan, S.; Kano, S. In vitro susceptibility and genetic variations for chloroquine and mefloquine in *Plasmodium falciparum* isolates from Thai-Myanmar border. *Southeast Asian J. Trop. Med. Public Health* **2005**, *36* (Suppl. 4), 73–79.

285. Pascual, A.; Basco, L.K.; Baret, E.; Amalvict, R.; Travers, D.; Rogier, C.; Pradines, B. Use of the atmospheric generators for capnophilic bacteria Genbag-CO₂ for the evaluation of in vitro *Plasmodium falciparum* susceptibility to standard anti-malarial drugs. *Malar. J.* **2011**, *10*, 8. [CrossRef]

286. Branco, A.; Francisco, D.; Hanscheid, T. Is there a 'normal' oxygen concentration for in vitro *Plasmodium* cultures? *Trends Parasitol.* **2018**, *34*, 811–812. [CrossRef]

287. Duffy, S.; Avery, V.M. *Plasmodium falciparum* in vitro culture—The highs and lows. *Trends Parasitol.* **2018**, *34*, 812–813. [CrossRef] [PubMed]

288. Géry, A.; Basco, L.K.; Heutte, N.; Guillamin, M.; N’Guyen, H.M.; Richard, E.; Garon, D.; Eldin de Pécoulas, P. Long-term in vitro cultivation of *Plasmodium falciparum* in a novel cell culture device. *Am. J. Trop. Med. Hyg.* **2019**, *100*, 822–827. [CrossRef] [PubMed]

289. Eastham, G.M.; Rieckmann, K.H. Field incubator for measuring drug susceptibility of *Plasmodium falciparum*. *J. Trop. Med. Hyg.* **1981**, *84*, 27–28. [PubMed]

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291. Kwiatkowski, D. Febrile temperatures can synchronize the growth of *Plasmodium falciparum* in vitro. *J. Exp. Med.* **1989**, *169*, 357–361. [CrossRef]

292. Long, H.Y.; Lell, B.; Dietz, K.; Kremsner, P.G. *Plasmodium falciparum*: In vitro growth inhibition by febrile temperatures. *Parasitol. Res.* **2001**, *87*, 553–555. [CrossRef]

293. Rehman, K.; Sauerzopf, U.; Veletzky, L.; Lötsch, F.; Groger, M.; Ramharter, M. Effect of mild medical hypothermia on in vitro growth of *Plasmodium falciparum* and the activity of anti-malarial drugs. *Malar. J.* **2016**, *15*, 162. [CrossRef]

294. Singhaboot, Y.; Keayarsa, S.; Piaraksa, N.; Phumratanaprapin, W.; Kunawut, P.; Dondorp, A.; Chotivanich, K. Temperature dependence of *Plasmodium falciparum* erythrocytic stage development. *Am. J. Trop. Med. Hyg.* **2019**, *100*, 1191–1195. [CrossRef]

295. Pavithra, S.R.; Banumathy, G.; Joy, O.; Singh, V.; Tatu, U. Recurrent fever promotes *Plasmodium falciparum* development in human erythrocytes. *J. Biol. Chem.* **2004**, *279*, 46692–46699. [CrossRef]

296. Rojas, M.O.; Wasserman, M. Effect of low temperature on the in vitro growth of *Plasmodium falciparum*. *J. Eukaryot. Microbiol.* **1993**, *40*, 149–152. [CrossRef]

297. Nakazawa, S. Synchronization of cultured *Plasmodium falciparum* by cold treatment. *Kitakanto Med. J.* **1985**, *35*, 383–390. [CrossRef]

298. Yuan, L.; Hao, M.; Wu, L.; Zhao, Z.; Rosenthal, B.M.; Li, X.; He, Y.; Sun, L.; Feng, G.; Xiang, Z.; et al. Refrigeration provides a simple means to synchronize in vitro cultures of *Plasmodium falciparum*. *Exp. Parasitol.* **2014**, *140*, 18–23. [CrossRef] [PubMed]
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