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Review

Feeding Mechanisms of Pathogenic Protozoa with a Focus on Endocytosis and the Digestive Vacuole

by
Mark F. Wiser
Department of Tropical Medicine and Infectious Disease, Tulane University School of Public Health and Tropical Medicine, New Orleans, LA 70112, USA
Parasitologia 2024, 4(3), 222-237; https://doi.org/10.3390/parasitologia4030019
Submission received: 21 May 2024 / Revised: 17 June 2024 / Accepted: 26 June 2024 / Published: 1 July 2024
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Abstract

:
Endocytosis is a quintessential feature of eukaryotes, and the emergence of endocytosis played a major role in the origin and evolution of eukaryotes. During the early evolution of eukaryotes, phagocytosis and the digestion of prey (i.e., bacteria) combined with the endocytosis of macromolecules opened a new source of nutrients beyond osmotrophy. Pathogenic and commensal protozoa have retained endocytosis as a major mechanism of nutrient acquisition even though, in theory, nutrients could be obtained from the host through osmotrophy. Nearly all pathogenic protozoa exhibit endocytosis and have lysosomal-like compartments that function as digestive vacuoles, and endocytosis appears to play a major role in the acquisition of nutrients. Cryptosporidium is a possible exception that may not exhibit endocytosis. Phagotrophy, however, is only observed in parasites of the intestinal lumen and appears to have been lost in blood and tissue parasites. Overall, the basic features of endocytosis and lysosomes are similar to other eukaryotes. Nonetheless, adaptation to the host has generated some novel features that are specific to certain protozoan lineages.

1. Introduction

Endocytosis and the associated endomembrane system are prototypical features of eukaryotic cells since the ability to carry out endocytosis was a major step in the origin of eukaryotic cells [1]. In addition to functions, such as cell signaling and regulating membrane composition, endocytosis is a means to acquire nutrients that are too large to be transported by membrane channels and carriers [2]. Nutrient acquisition in prokaryotes is primarily via the uptake of small molecules called osmotrophy. In addition, prokaryotes lack an organized cytoskeletal system which is also crucial for endocytosis. Saprotrophy (i.e., extracellular digestion of macromolecules followed by osmotrophy) and autotrophy (e.g., photosynthesis) are other feeding strategies of bacteria. The acquired ability to endocytose macromolecules or to phagocytose bacteria and organic particulate matter in newly evolved eukaryotes provided an abundant food supply to complement the uptake of small-molecular-weight metabolites. This increase in potential food possibly fueled the massive expansion and diversification of the eukaryotes. Furthermore, phagocytosis set the stage for endosymbiosis and the development of mitochondria and plastids [3].
Little work has been performed on feeding mechanisms in parasitic protozoa, and most of this work is focused on prevalent or medically important pathogens. Due to the importance of eating, feeding mechanisms of pathogens likely provide sensitive targets for therapeutic intervention, and a better understanding of feeding in pathogenic protozoa may reveal novel therapeutic targets for drug development. Indeed, several highly efficacious antimalarials target the digestive vacuole or feeding mechanisms of the malarial parasite [4,5]. Despite this importance of feeding in parasitic protozoa, except for the malaria parasite, relatively little research has been carried out. For example, the unique environment of the host may lead to novel pathogen adaptations to optimize its food acquisition that can be exploited as drug targets. A priori, one might expect a reduced reliance on endocytosis for food acquisition, as compared to free-living protozoa, since pathogens may be able to acquire sufficient nutrients from the host via osmotrophy. And indeed, comparison of the genomes of the free-living Bodo saltans to related parasitic trypanosomatids reveals a loss of genes associated with macromolecular degradation and an expansion of genes associated with amino acid and nucleotide transport [6,7]. A loss of diversity in endomembrane trafficking systems is also observed in the evolution of apicomplexan parasites from free-living ancestors to parasites [8]. Nonetheless, parasitic protozoa heavily rely on endocytosis as a means to acquire nutrients.
This review provides a broad overview of endocytosis as it relates to feeding by the pathogenic protozoa of humans. The primary protozoa that are covered include Entamoeba histolytica, Giardia, Trypanosoma brucei, Trypanosoma cruzi, Leishmania, Cryptosporidium, Plasmodium, and Toxoplasma. Despite this diverse mix of protozoa in a phylogenetic sense, there are numerous similarities due to the universality of endocytosis in eukaryotes. However, there are notable unique features exhibited by these various protozoan pathogens.

2. Endocytosis

Endocytosis is a general term referring to the uptake of substances by a cell that involves surrounding the material, often referred to as cargo, with the plasma membrane and the formation of vesicles that contain the cargo (Table 1). Phagocytosis, which translates to cell eating, is a form of endocytosis in which particles ≥0.5 µm are internalized [9]. This uptake of particulate matter involves the formation of pseudopodia that extend and surround the cargo [10]. Particulate matter is internalized by fusion of the pseudopodia, which results in a large membrane-bound structure called the phagosome. Fluid-phase endocytosis, historically called pinocytosis for cell drinking, is the uptake of soluble matter by enclosing the cargo into membrane-bound vesicles. There are distinct types of fluid-phase endocytosis based on the size and volume of material being taken up and the mechanisms involved in the formation of endocytic vesicles [11,12]. Pinocytosis has been divided into macro-pinocytosis and micro-pinocytosis depending on the size of the resulting vesicles [13]. Macro-pinocytosis involves the folding back of membrane ruffles onto the plasma membrane, forming large vacuoles that range in size from 0.2 μm to >10 μm. Micro-pinocytosis involves the formation of small vesicles (<0.2 μm) at the plasma membrane and is generally divided into clathrin-mediated endocytosis (CME) and clathrin-independent endocytosis (CIE).
A well-studied and major form of endocytosis is CME, which has historically been called receptor-mediated endocytosis [14,15]. CME is proposed to be an evolutionarily ancient endocytic mechanism, and in some eukaryotic lineages, CME may be the sole mechanism for the internalization of substances from the extracellular milieu. Clathrin is recruited to the site of vesicle formation by adaptor proteins (e.g., AP-2) and assembles into a lattice work that drives the formation of a vesicle. The vesicle is separated from the membrane by a dynamin-mediated scission. Subsequently, the vesicle is uncoated so that it can fuse with endosomes. CIE is less well defined in regard to molecular components and their nomenclature is imprecise with various names depending on the molecular components that facilitate the endocytosis [16,17]. CIE comprises several parallel routes that involve several distinct mechanisms to form endocytic vesicles. Lipid rafts are also involved in the formation of many of these vesicles [18]. Lipid rafts are plasma membrane microdomains that are enriched in cholesterol and sphingolipids.
In many cells, endocytosis or phagocytosis is restricted to specific regions of the plasma membrane. For example, many protozoa have a specialized organelle called the cytostome—which translates to cell mouth—where phagocytosis or endocytosis takes place. Quite often, the cytostome is a microtubule-supported groove with a funnel shape, and food is directed down this groove. Phagocytosis or endocytosis then occurs at the bottom of the groove.

3. The Digestive Vacuole

The compartment resulting from endocytosis or phagocytosis is called either an endosome or a phagosome, respectively (Figure 1). Small endocytic vesicles can fuse to form a large endosome, or the endocytic vesicles can fuse with a pre-existing endosome to deliver the cargo [19]. Endosomes often function as a crossroads and can direct cargo to various subcellular compartments. Much of the cargo taken up by endocytosis is destined for catabolism in an acidic compartment that contains hydrolytic enzymes. Hydrolytic enzymes, such as proteases, lipases, nucleases, glycosidases, and phosphatases, break down macromolecules into metabolites that can be utilized by the cell. Thus, this compartment—especially in protozoa—is called the digestive vacuole or sometimes the food vacuole. The digestive vacuole is sometimes formed via fusion of endosomes or phagosomes with lysosomes to form endolysosomes or phagolysosomes, respectively. Thus, lysosomes deliver proton pumps to acidify the compartment as well as the acid hydrolases to the digestive vacuole. Lysosomes of animal cells are the best known and are typically small (0.1–1 mm) and numerous depending on the cell type [20]. In contrast, fungi and plants tend to have a few large acidic vacuoles [21]. Mechanisms for the formation of the digestive vacuole are not well known in the protozoa.

4. Intestinal and Other Luminal Protozoa

Several protozoan species infect the human gastrointestinal tract and Trichomonas vaginalis infects the urogenital tract. A few of these lumen-dwelling protozoa can cause clinical disease, whereas most of them do not cause disease, and some of them may even be viewed as non-pathogenic commensals. A wide variety of protozoa infect the intestinal tract including amebae, flagellates, ciliates, apicomplexans, and stramenopiles [22]. Most of these pathogenic protozoa exhibit a fecal–oral life cycle involving the formation of robust cysts that are passed with the feces and that initiate a new infection when ingested by a susceptible host. In terms of disease severity and disease prevalence, the most important intestinal protozoa of humans are Entamoeba histolytica, Giardia duodenalis, and Cryptosporidium (discussed with the Apicomplexa). Little research has been performed on other intestinal protozoa.
The lumens of the gastrointestinal and urogenital tracts are inhabited by a rich and diverse microbiota [23]. The abundance and milieu of bacteria in these environments are not overtly different than what might be seen in soil and aquatic microbiota. Therefore, one can envision a simple adaptation regarding the acquisition of nutrients as free-living protozoa evolved to lumen-dwelling parasitic protozoa. Presumably, many, if not most, lumen-dwelling protozoa acquire nutrients via the phagocytosis of bacteria or other particulate matter. However, very little work on phagocytosis and digestive vacuoles has been carried out on organisms other than Entamoeba and Giardia. Lysosomes of Balantidium coli have been described at the ultrastructural level [24], and some work has been carried out on phagocytosis in Trichomonas [25]. Highly virulent strains of T. vaginalis exhibit higher levels of phagocytic activity than non-virulent strains. The proteome of the Trichomonas lysosome reveals hydrolases and other proteins typically associated with digestive vacuoles [26].

4.1. Entamoeba histolytica

Entamoeba histolytica causes amebic dysentery as well as invasive diseases, leading to significant morbidity and mortality worldwide [27]. Entamoeba species are members of the Amoebozoa, which are a sister group of Opisthokonta that includes fungi and animals [28]. As with amebae in general, phagocytosis plays a primary role in the acquisition of food and the microbiota of the colon provides ample bacteria. E. histolytica may have a preference for particular bacterial species within the gut microbiota [29]. In addition, phagocytosis has long been noted as a virulence factor [30] and E. histolytica exhibits more phagocytic activity that the non-pathogenic E. dispar [31,32]. During severe disease, E. histolytica ingests erythrocytes and other host cells and thereby contributes to the destruction of tissues.
The hydrolytic enzymes that are associated with the digestive vacuoles of E. histolytica are similar to other organisms [33]. However, E. histolytica has a unique family of lysosomal hydrolase receptors called the cysteine protease binding protein family which function to ensure the proper targeting of hydrolases to the lysosomes [34]. Proteins involved in vesicle and endosome formation are homologous to equivalent proteins of yeasts and animals [35]. These similarities are not unexpected, due to the phylogenetic relationship between amoebozoans, fungi, and metazoans. However, endocytosis may be more complex in E. histolytica as compared to yeasts and animals since the processing of cargos obtained by fluid-phase endocytosis, such as transferrin and LDL; phagosomes containing ingested erythrocytes; or phagosomes with ingested bacteria are all processed in different vacuoles [36]. This complexity is reflected by the substantially higher number of Rab GTPase paralogs in E. histolytica than other eukaryotes [37]. Rab GTPases regulate membrane trafficking in all eukaryotic cells and Rab GTPase diversity is often regarded as a portrayal of vesicular trafficking complexity [38].
Another phagocytic mode of internalization of extracellular material is trogocytosis, or cell nibbling [39]. During trogocytosis, portions of a live cell are ingested instead of the entire cell being engulfed. E. histolytica exhibits trogocytosis and this contributes to cell killing and tissue invasion [40]. Trogocytosis has also been implicated in immune evasion by E. histolytica due to the display of host membrane proteins on the ameba’s surface [41]. Specific protein kinases, not involved in the phagocytosis of dead cells, have also been implicated in the trogocytosis of living human cells [42], suggesting multiple parallel pathways of endocytosis. Free-living ameba that are capable of causing human pathology also kill cells by trogocytosis [43,44]. Trogocytosis-mediated host cell killing by Trichomonas gallinae, a trichomonad from birds, has also been demonstrated [45].

4.2. Giardia

Giardia duodenalis is a common intestinal protozoan that colonizes the small intestine and specifically the duodenum. Phylogenetically, Giardia is a member of the Metamonada—a group composed largely of anaerobic gut commensals or parasites and includes the trichomonads [28]. A unique characteristic of Giardia is the adhesive disk on the ventral surface that firmly attaches the trophozoite to the intestinal epithelium [46]. The parasite primarily acquires nutrients through fluid-phase endocytosis on the dorsal surface. However, Giardia lacks conventional endosomes and lysosomes, and instead exhibits vacuoles just beneath the cell surface primarily on the dorsal side called peripheral vacuoles [47,48]. Endocytosed material rapidly accumulates in the peripheral vacuoles and different types of material segregates into different peripheral vacuoles [49]. Invaginations of the dorsal side of the trophozoite correlate with the presence of peripheral vacuoles on the inner surface, and these invaginations are associated with clathrin, suggesting that a clathrin-mediated process may be involved in transferring cargo from the invaginations to the peripheral vacuoles [50]. Peripheral vacuoles are also acidic and contain hydrolases [51]. Thus, the peripheral vacuoles have characteristics of digestive vacuoles and are likely to be the primary site of nutrient digestion.
Recent observations suggest that Giardia is also capable of phagocytosis, in contrast to previous beliefs [52]. Phagocytosis can occur throughout the cell surface, but preferentially occurs just below the ventral disk near the exit site of the ventral flagella. The phagocytosed material is incorporated into intracellular vacuoles that are distinct from peripheral vacuoles. The extent of the contribution of phagocytosis to nutrient acquisition is not known at this time but presumed to be minor.

5. Kinetoplastids

Kinetoplastids are a monophyletic group within the Discoba [28]. Discoba is a major eukaryotic clade of primarily heterotrophic or mixotrophic flagellates that includes the Heterolobosea and Euglenozoa as two major sub-clades. The kinetoplastids are a sister group with the euglenids that make up the Euglenozoa. A defining feature of kinetoplastids is a concatenated mass of mitochondrial DNA, called the kinetoplast, that is located near the base of the flagellum. Kinetoplastids are strictly heterotrophic and most kinetoplastids are parasitic. The medically important trypanosomatids (e.g., Trypanosoma and Leishmania) arose from the free-living bacterivorous bodonids [53]. As part of their evolution to parasites, the trypanosomatids may have lost the ability to carry out phagocytosis, and the acquisition of nutrients relies heavily on fluid-phase endocytosis and membrane channels or transporters [6,7].

5.1. Medically Important Trypanosomatids

The best studied kinetoplastids are the medically important Trypanosoma brucei complex, T. cruzi, and Leishmania which cause human African trypanosomiasis, Chagas’ disease, and leishmaniasis, respectively. These medically important kinetoplastids are all transmitted by blood-feeding vectors and exhibit complex life cycles. The life cycle stages are described as four major morphological forms designated as trypomastigotes, epimastigotes, promastigotes, or amastigotes [22]. These designations are determined by the position of the kinetoplast and the emergence of the flagellum from the cell body. The flagellum emerges from the cell near the kinetoplast from an invagination of the plasma membrane called the flagellar pocket.
Endocytosis has not been extensively studied in the kinetoplastids, and furthermore, knowledge about endocytosis tends to be focused on stages most accessible to in vitro experimentation. For example, endocytosis and secretion in the trypanosomatids are best characterized in T. brucei, which is often used as a laboratory model for human African trypanosomiasis. T. brucei infects cattle and is unable to infect humans due to a trypanosome lytic factor in the serum of humans and other higher apes [54]. However, T. brucei is closely related to the human parasites T. gambiense and T. rhodesiense, and, collectively, the three species are referred to as the T. brucei complex. Most of the studies on the African trypanosomes have been carried out in the stages found in the host blood, whereas in T. cruzi and Leishmania, most of the work has been performed in the stages found in the vector.

5.2. Endocytosis and Trafficking to the Digestive Vacuole in Kinetoplastids

Kinetoplastid endocytosis is primarily clathrin-mediated and lysosome-like organelles have been described (Table 2). A subpellicular microtubule network restricts endocytosis in kinetoplastids to specialized regions of the plasma membrane called the flagellar pocket. This flagellar pocket is the primary site of the trafficking of macromolecular material into and out of kinetoplastids [55,56]. Therefore, most of the endosomal compartments are located between the nucleus and flagellar pocket. Endocytosis primarily plays a nutritional role in kinetoplastids. For example, trypanosomatids exhibit receptor-mediated endocytosis and the uptake of host transferrin and lipoproteins is particularly well documented in T. brucei [57]. In African trypanosomes, endocytosis also participates in immune avoidance by removing antibodies from the cell surface [58].
T. brucei, T. cruzi, and Leishmania have distinct transferrin receptors and the precise trafficking mechanisms differ between these parasites and according to the life cycle stage [59]. In T. brucei, the blood-stage trypomastigotes take up transferrin at the flagellar pocket via clathrin-coated vesicles [60]. However, many genes required for clathrin-dependent endocytosis in animals and fungi are absent from trypanosomatids [61]. Likewise, genes required for clathrin-independent endocytosis are also absent. However, the trypanosomatids do possess several unique clathrin-associated proteins which may play significant roles in endocytosis [62]. T. cruzi epimastigotes take up albumin via clathrin-mediated endocytosis at the flagellar pocket, while the clathrin-independent endocytosis of transferrin occurs at the cytostome–cytopharynx complex [63]. A receptor-mediated endocytosis of hemoglobin is exhibited by Leishmania [64]. Leishmania is unable to synthesize heme and acquires heme from host hemoglobin via a specific hemoglobin receptor in a clathrin-dependent process. T. cruzi [65] and Leishmania [66,67] also take up lipoproteins by receptor-mediated endocytosis.
In T. brucei, the endocytosed vesicles are rapidly delivered to an endosomal compartment and the cargo is ultimately delivered to a single lysosome-like compartment [68,69]. Overall, the endocytic pathway and structure of the lysosome in T. brucei appear similar to other eukaryotes [70,71]. A unique lysosomal-like compartment, called the reservosome, has been identified in T. cruzi [72]. Ingested nutrients are delivered to reservosomes, which are typically located at the posterior ends of epimastigotes, and these reservosomes presumably store proteins and lipids for future use [73]. Although substantially less endocytosis is observed in the trypomastigote and amastigote stages of T. cruzi than the epimastigotes stages [65], lysosome-like organelles have been described in these life-cycle stages of T. cruzi [74]. The Leishmania lysosome has an atypical structure, consisting of an elongated vesicle-filled tubule running along the anterior–posterior axis of the promastigote [75,76]. Amastigotes have one or a few large vesicle-filled lysosomes called megasomes [77,78].

5.3. A Cytostome-Like Structure Is Found in Some Kinetoplastids

A cytostome-like organelle, called the cytostome–cytopharynx complex, is found in T. cruzi, free-living bodonids, and the parasitic Paratrypanosoma confusum [79]. P. confusum is an early branching monoxenous trypanosomatid of mosquitoes [80]. The cytostome–cytopharynx complex is a feeding structure that begins as a stable opening adjacent to the flagellar pocket and extends internally as a tubular invagination that carries endocytosed material to the posterior end of the cell for digestion. Notably, the cytostome–cytopharynx complex has been retained in monoxenous trypanosomatids that are transmitted via feces, as well as the dixenous T. cruzi which is also transmitted via feces of the vector (i.e., stercorarian transmission). Notably, the cytostome–cytopharynx complex of T. cruzi is only expressed in the epimastigote stage which inhabits the insect gut, and the cytostome–cytopharynx complex is not expressed in the trypomastigote stage which inhabits the blood and tissues of the vertebrate host [81]. Likewise, the cytostome–cytopharynx complex is neither found in Trypanosoma species nor Leishmania species that exhibit salivarian transmission. This implies that the cytostome–cytopharynx complex has been retained in free-living kinetoplastids and parasitic kinetoplastids that reside in the gut of insect hosts or vectors. The insect gut microbiota provides a potential source of nutrients via phagocytosis, thus implying that phagocytosis has been retained in some kinetoplastids. However, phagocytosis has not yet been reported in parasitic kinetoplastids.

6. Apicomplexa

The Apicomplexa are a large and diverse group of primarily parasitic protozoa [82]. Major subgroups within the Apicomplexa are the chromerids and colpodellids, gregarines including the related cryptosporidia, coccidia, and hematozoans. The chromerids and colpodellids are phylogenetically related and are the most ancestral group within the Apicomplexa, as evidenced by branching near the divergence of apicomplexans and dinoflagellates [83]. Chromerids are photosynthetic endosymbionts of corals [84] and colpodellids are free-living predators that feed on other protists [85]. The most ancestral parasitic apicomplexans are the gregarines and cryptosporidia that attach to host cells and are primarily extracellular ectoparasites. Coccidia and hematozoans are intracellular parasites that exhibit the more well-known apicomplexan features.
Apicomplexans exhibit complex life cycles involving an asexual replication called merogony, a sexual stage involving the production of gametes, and another type of asexual replication following the sexual stage called sporogony [82]. Some apicomplexan life cycles involve multiple hosts or vectors, and merogony, gametogony, and sporogony can occur in different hosts. The hematozoan Plasmodium and coccidian Toxoplasma are the best characterized among the apicomplexans due to their importance as human pathogens and robust model systems to study these organisms. Other apicomplexans that cause human disease are Cryptosporidium and intestinal coccidia. Intestinal coccidia are also important veterinary pathogens, as well as the tick-transmitted Babesia and Theileria of the hematozoans.
A defining feature of Apicomplexa—as reflected in the name—is the apical complex which consists of specialized secretory organelles called micronemes and rhoptries and a polar ring of microtubules at the apical end of the parasite. These apical organelles function in attachment to prey or host cells, cell invasion, or gliding motility [86,87,88,89]. The free-living predatory colpodellids, Cryptosporidium, and extracellular gregarines attach to prey or host cells via the apical organelles and ingest the contents of the host cell via myzocytosis [90]. The apical organelles subsequently evolved to facilitate the invasion of host cells in the coccidians and hematozoans [91]. Most intracellular apicomplexan parasites reside within an intracellular compartment called the parasitophorous vacuole. In theory, these intracellular parasites could survive via osmotrophy within this intracellular niche. However, fluid-phase endocytosis of the host cell cytoplasm is a prominent feature of Plasmodium and Toxoplasma, and is presumably widely utilized among intracellular apicomplexans. Phagotrophy, as defined by the uptake of particles greater than 0.5 μm, though, has not been observed in the apicomplexans.

6.1. Myzocytosis

Myzocytosis is a type of feeding in which a protozoan predator attaches to its prey and aspirates the cytoplasmic contents of the prey. This type of feeding is exhibited by some apicomplexans and some dinoflagellates and these two sister groups form the Myzozoa [92]. However, there are some fundamental differences in the mechanisms of myzocytosis between the apicomplexans and dinoflagellates. Some predatory dinoflagellates utilize a feeding tube, called a peduncle, to attach to the prey and suck out its cytoplasm [93], whereas in colpodellids, the apical organelles are involved in the attachment of the predator to the prey and the subsequent uptake of the prey’s cytoplasm [94]. Nuclei and mitochondria of the prey are observed to be aspirated by Colpodella possibly through a myzocytotic aperture [95], and this aperture may be analogous to a cytostome. However, no vesicular transport between this cytostome-like structure and the digestive vacuole has been reported. This suggests that this cytostome-like structure may undergo scission from the plasma membrane and develop into the digestive vacuole. In addition, detached Colpodella also exhibit endocytosis as evidenced by the uptake of nanoparticles during in vitro culture [95]. The significance of this endocytosis and its role in nutrition are unknown.
Gregarines are primarily extracellular apicomplexan parasites that mainly parasitize invertebrates by attaching to host intestinal epithelial cells and feeding via myzocytosis. A variety of structures, including mucrons, epimerites, and promerites, that are involved in these host–parasite interactions have been described [90]. The exact feeding mechanisms have not been described and likely differ between species. In some species, endocytic vesicles near the attachment site have been observed and vesicles presumably transport cargo to the digestive vacuole. In species with mucrons, an opening at the attachment site and a cytostome-like structure called the mucronal vacuole are observed [96,97]. This mucronal vacuole may be analogous to the myzocytotic aperture of the colpodellids and may develop into a digestive vacuole or essentially be the digestive vacuole.
Cryptosporidium is an intestinal parasite that is related to the gregarines and infects a wide range of vertebrates including humans [98]. Cryptosporidiosis manifests as transient diarrhea in the immunocompetent but can cause profuse and possibly life-threatening diarrhea in the immunocompromised. The parasite attaches to intestinal epithelial cells via the apical organelles and remodels actin, leading to the fusion of microvilli that ultimately encloses the parasite in an extracytoplasmic compartment [99]. A junction called the feeder organelle is formed between the parasite and host cell [100]. Nutrients are transported across the feeder organelle via an ATP-binding cassette transporter without endocytosis [101,102]. In addition, cholesterol is taken up from the host cell by the parasite by an unknown mechanism [103]. There are no reports of endocytic vesicles associated with the feeder organelle nor a digestive vacuole in Cryptosporidium. Similarly, Cryptosporidium does not have a micropore or a cytostome-like structure [104].

6.2. Plasmodium

Members of the genus Plasmodium are the causative agents of malaria and account for hundreds of thousands of deaths per year. The parasite exhibits a complex life cycle involving mosquito transmission and a transient merogony in the liver before infecting erythrocytes [22]. Merozoites released from the liver invade erythrocytes and repeated rounds of erythrocytic merogony are responsible for the clinical manifestations and pathology of the disease. Some of the merozoites develop into sexual forms called gametocytes that infect mosquitoes. During the intra-erythrocytic stage, the parasite ingests the host cell hemoglobin as a source of amino acids, and this feeding process has been extensively characterized and recently reviewed [4,105]. Endocytosis in other stages of the malaria parasite life cycle has not been studied.
Endocytosis commences shortly after invasion of the erythrocyte with the formation of small vesicles. These small vesicles initially function as individual digestive vacuoles and, as the parasite matures, the small digestive vacuoles coalesce to form large digestive vacuoles [106]. In the sexual stages, the individual digestive vacuoles do not coalesce and the small individual digestive vacuoles remain dispersed in the parasite cytoplasm [107]. The endocytic pathway in Plasmodium does not appear to involve the fusion of pre-formed lysosomes with the endosomes, and the endocytic vesicles appear to function as digestive vacuoles as soon as they are formed [105]. As the intracellular parasite matures, cytostomes develop and endocytosis primarily occurs at these structures [108].
The molecular components of the endocytic machinery of Plasmodium have not been extensively characterized and only a few of the components have been identified [105,109]. Endocytosis in Plasmodium appears to be clathrin-independent even though a clathrin adaptor protein complex, called AP-2, is likely involved [110,111]. In addition, endocytosis in Plasmodium is receptor-independent, which is consistent with CIE. Other proteins involved in endocytosis have been identified, but their precise roles are not known. Some of the Plasmodium proteins involved in endocytosis are unique to Plasmodium, suggesting some unique features of endocytosis [109].

6.3. Toxoplasma

Toxoplasmosis is a common human disease caused by Toxoplasma gondii. Most infections are quite benign and clinical disease is primarily associated with immunosuppression, congenital infections, or ocular toxoplasmosis [22]. Toxoplasma is a dixenous parasite and is categorized as a tissue-cyst-forming coccidian. The definitive hosts are felines, and the parasite exhibits a typical coccidian life cycle within the intestinal epithelium that includes asexual replication, macrogametocytes, and microgametocytes. A wide range of birds and mammals, including humans, can serve as intermediate hosts. As with most apicomplexans, Toxoplasma is an intracellular parasite. However, distinct from many Apicomplexa, Toxoplasma can replicate in a wide range of host cell types within the intermediate hosts. During the acute stage of the infection, the parasites replicate rapidly and are called tachyzoites. As immunity develops, the replication rate slows and the tachyzoites develop into bradyzoites that are readily transmitted to the definitive host [112].
Most of the work on endocytosis in Toxoplasma has been carried out on tachyzoites due to the accessibility of this stage and the ease of in vitro cultivation. Essentially nothing is known about the intestinal stages of Toxoplasma in felines. Macrogametes of the intestinal coccidian Eimeria are reported to take up nanoparticles by endocytosis [113]. There is also a report of bradyzoites endocytosing host cell cytoplasmic proteins [114]. Components of CME and CIE have been identified in Toxoplasma [115], including the molecular components that are normally involved in transporting vesicles to the lysosome. Toxoplasma also exhibits substantial endocytosis in the extracellular stages [116]. Endocytosis in the extracellular stages may not be involved in nutrient uptake, but rather may function in plasma membrane homeostasis to compensate for the exocytosis involved in gliding motility.

6.4. The Parasitophorous Vacuole

Most apicomplexans reside in a parasitophorous vacuole that is segregated from the endomembrane systems of the host cell. The membrane of this vacuole is generated during invasion of the host cell and is derived from both host membrane material and parasite membrane material excreted by the rhoptries [117,118]. Some apicomplexans such as Babesia [119,120] and Theileria [121] escape from the parasitophorous vacuole. For parasites that remain in the parasitophorous vacuole, the parasitophorous vacuolar membrane (PVM) presents a potential barrier for the acquisition of nutrients and endocytosis. Proteinaceous pores are found on the PVM of Toxoplasma [122] and Plasmodium [123]. These pores could allow for the passage of host proteins and metabolites into the parasitophorous vacuole. However, these pores may primarily function in the export of parasite proteins into the host cell.
The intracellular trophozoites of Plasmodium take up the host cell cytoplasm by endocytosis involving both the PVM and the parasite plasma membrane [105,124]. This results in the formation of double-membrane vesicles, in which the inner membrane is derived from the PVM, and this PVM-derived membrane is subsequently disintegrated. In Toxoplasma, the uptake of the host’s cytoplasm may be a two-step process in which vesicles and host cell organelles are first incorporated into the parasitophorous vacuole that are subsequently taken up by the parasite [125,126,127]. In addition, Toxoplasma may exploit the endosomal sorting complex of the host cell for uptake of host cell cytoplasm into the parasitophorous vacuole [128].

6.5. The Digestive Vacuole

The digestive vacuole of Plasmodium is well described and specializes in the ordered catabolism of hemoglobin [105,129]. In addition, the Plasmodium digestive vacuole is the site of hemozoin formation, a non-toxic biocrystal of heme [105,130]. The digestive vacuole of Toxoplasma is also relatively well characterized and contains similar proteases as the Plasmodium digestive vacuole [131], and host cytosolic proteins are digested within the vacuole [132]. The acidic pH of the vacuole is maintained by a combination of both a V-type H+-ATPase and a H+-pyrophosphatase in both Plasmodium [133] and Toxoplasma [131]. The H+-pyrophosphatase is absent in the Opisthokonta [134]. A member of the drug/metabolite transporter superfamily called the chloroquine resistance transporter is also found on the digestive vacuolar membrane in both Plasmodium [135] and Toxoplasma [136].

6.6. The Micropore of Apicomplexans

Another ultrastructural feature found in apicomplexans is the micropore [137]. The micropore usually appears as an invagination of the plasma membrane with thickened material around the neck which spans the inner membrane complex. The inner membrane complex consists of connected flattened vesicles that form a layer below the plasma membrane during some stages of apicomplexan life cycles [138]. This inner membrane complex is a barrier for endocytosis and exocytosis, and thus, the micropore may have evolved in response to the presence of the inner membrane complex. The inner membrane complex is related to the cortical alveoli that define the alveolates that includes apicomplexans, dinoflagellates, and ciliates [28,139]. Apicomplexans and dinoflagellates are sister groups and dinoflagellates also have micropores [140].
Endocytosis has been demonstrated to occur at the micropore [141,142,143]. However, the micropore is a permanent structure that does not exhibit the usual dynamics of assembly and disappearance associated with endocytosis [143]. Kelch13, a highly conserved protein in Apicomplexa [143], is localized to the electron-dense neck of the micropore in both Plasmodium and Toxoplasma [142,143]. Kelch13 is also localized to cytostomes of Plasmodium [144]. This suggests the Plasmodium cytostome may be a specialized version of the micropore. For example, the micropores of extracellular merozoites may differentiate into the cytostomes of the intracellular trophozoite. The inner-membrane complex is disassembled as merozoites differentiate into trophozoites [145] and, thus, endocytosis is no longer encumbered. Therefore, the micropore would no longer be necessary during the intracellular trophic period and may either disappear or develop into a cytostome.

7. Summary

Nutrient acquisition in protozoa relies heavily on endocytosis, which can include the phagocytosis of particles or other organisms or fluid-phase endocytosis. As protozoa evolve from free-living heterotrophs to parasitic organisms, one might expect the adaptation of endocytic processes to the distinct environments of the hosts. Overall, though, endocytosis in parasitic protozoa is similar to endocytosis in other eukaryotes. This is expected since endocytosis is a quintessential feature of eukaryotes, and the fundamental aspects of endocytosis are unlikely to exhibit major changes. Similarly, the endomembrane systems of the pathogenic protozoa are similar to eukaryotes in general with some specializations related to virulence [89].
Some lineage-specific differences in the details of endocytosis are seen in parasitic protozoa, as well as some specialized adaptations to the parasitic lifestyle (Table 3). For example, phagotrophy is common among soil, benthic, and planktonic protozoa [28] as well as E. histolytica and T. vaginalis. On the other hand, Giardia does not appear to exhibit extensive phagocytosis and relies primarily on fluid-phase endocytosis and peripheral vacuoles. Blood and tissue protozoa appear to have lost phagotrophy and rely heavily on fluid-phase endocytosis. For example, endocytosis in kinetoplastids occurs primarily at the flagellar pocket and involves clathrin-mediated endocytosis. Plasmodium and Toxoplasma also do not exhibit phagotrophy but exhibit extensive fluid-phase endocytosis of the host cell cytoplasm. Endocytosis in both species appears clathrin-independent even though a clathrin adaptor protein appears to be involved. The basal apicomplexans, including Cryptosporidium, exhibit a feeding mechanism called myzocytosis that involves attaching to a host cell and aspirating the contents of that cell.

Funding

This research received no external funding.

Conflicts of Interest

The author declares no conflicts of interest.

References

  1. Cavalier-Smith, T. Predation and eukaryote cell origins: A coevolutionary perspective. Int. J. Biochem. Cell Biol. 2009, 41, 307–322. [Google Scholar] [CrossRef]
  2. Miaczynska, M.; Stenmark, H. Mechanisms and functions of endocytosis. J. Cell Biol. 2008, 180, 7–11. [Google Scholar] [CrossRef] [PubMed]
  3. Martin, W.F.; Tielens, A.G.M.; Mentel, M.; Garg, S.G.; Gould, S.B. The physiology of phagocytosis in the context of mitochondrial origin. Microbiol. Mol. Biol. Rev. 2017, 81, e00008-17. [Google Scholar] [CrossRef]
  4. Matz, J.M. Plasmodium’s bottomless pit: Properties and functions of the malaria parasite’s digestive vacuole. Trends Parasitol. 2022, 38, 525–543. [Google Scholar] [CrossRef]
  5. Edgar, R.C.S.; Counihan, N.A.; McGowan, S.; de Koning-Ward, T.F. Methods used to investigate the Plasmodium falciparum digestive vacuole. Front. Cell. Infect. Microbiol. 2021, 11, 829823. [Google Scholar] [CrossRef]
  6. Jackson, A.P.; Otto, T.D.; Aslett, M.; Armstrong, S.D.; Bringaud, F.; Schlacht, A.; Hartley, C.; Sanders, M.; Wastling, J.M.; Dacks, J.B.; et al. Kinetoplastid phylogenomics reveals the evolutionary innovations associated with the origins of parasitism. Curr. Biol. 2016, 26, 161–172. [Google Scholar] [CrossRef]
  7. Skalický, T.; Dobáková, E.; Wheeler, R.J.; Tesařová, M.; Flegontov, P.; Jirsová, D.; Votýpka, J.; Yurchenko, V.; Ayala, F.J.; Lukeš, J. Extensive flagellar remodeling during the complex life cycle of Paratrypanosoma, an early-branching trypanosomatid. Proc. Natl. Acad. Sci. USA 2017, 114, 11757–11762. [Google Scholar] [CrossRef]
  8. Woo, Y.H.; Ansari, H.; Otto, T.D.; Klinger, C.M.; Kolisko, M.; Michálek, J.; Saxena, A.; Shanmugam, D.; Tayyrov, A.; Veluchamy, A.; et al. Chromerid genomes reveal the evolutionary path from photosynthetic algae to obligate intracellular parasites. eLife 2015, 4, e06974. [Google Scholar] [CrossRef] [PubMed]
  9. Haas, A. The phagosome: Compartment with a license to kill. Traffic 2007, 8, 311–330. [Google Scholar] [CrossRef]
  10. Uribe-Querol, E.; Rosales, C. Phagocytosis: Our current understanding of a universal biological process. Front. Immunol. 2020, 11, 1066. [Google Scholar] [CrossRef]
  11. Jeger, J.L. Endosomes, lysosomes, and the role of endosomal and lysosomal biogenesis in cancer development. Mol. Biol. Rep. 2020, 47, 9801–9810. [Google Scholar] [CrossRef]
  12. Elkin, S.R.; Lakoduk, A.M.; Schmid, S.L. Endocytic pathways and endosomal trafficking: A primer. Wien. Med. Wochenschr. 2016, 166, 196–204. [Google Scholar] [CrossRef]
  13. Doherty, G.J.; McMahon, H.T. Mechanisms of endocytosis. Annu. Rev. Biochem. 2009, 78, 857–902. [Google Scholar] [CrossRef]
  14. Kaksonen, M.; Roux, A. Mechanisms of clathrin-mediated endocytosis. Nat. Rev. Mol. Cell Biol. 2018, 19, 313–326. [Google Scholar] [CrossRef]
  15. McMahon, H.T.; Boucrot, E. Molecular mechanism and physiological functions of clathrin-mediated endocytosis. Nat. Rev. Mol. Cell Bol. 2011, 12, 517–533. [Google Scholar] [CrossRef]
  16. Ferreira, A.P.A.; Boucrot, E. Mechanisms of carrier formation during clathrin-independent endocytosis. Trends Cell Biol. 2018, 28, 188–200. [Google Scholar] [CrossRef]
  17. Mayor, S.; Parton, R.G.; Donaldson, J.G. Clathrin-independent pathways of endocytosis. Cold Spring Harb. Perspect. Biol. 2014, 6, a016758. [Google Scholar] [CrossRef]
  18. Lajoie, P.; Nabi, I.R. Lipid rafts, caveolae, and their endocytosis. Int. Rev. Cell Mol. Biol. 2010, 282, 135–163. [Google Scholar] [CrossRef]
  19. Scott, C.C.; Vacca, F.; Gruenberg, J. Endosome maturation, transport and functions. Semin. Cell Dev. Biol. 2014, 31, 2–10. [Google Scholar] [CrossRef]
  20. Luzio, J.P.; Pryor, P.R.; Bright, N.A. Lysosomes: Fusion and function. Nat. Rev. Mol. Cell Biol. 2007, 8, 622–632. [Google Scholar] [CrossRef]
  21. de Marcos Lousa, C.; Denecke, J. Lysosomal and vacuolar sorting: Not so different after all! Biochem. Soc. Trans. 2016, 44, 891–897. [Google Scholar] [CrossRef]
  22. Wiser, M.F. Protozoa and Human Disease; Garland Science: New York, NY, USA, 2011; ISBN 9780429258282. [Google Scholar]
  23. Blum, H.E. The human microbiome. Adv. Med. Sci. 2017, 62, 414–420. [Google Scholar] [CrossRef]
  24. Skotarczak, B. The formation of primary and secondary lysosomes in Balantidium coli, Ciliata. Folia Histochem. Cytobiol. 1999, 37, 261–265. [Google Scholar]
  25. Pereira-Neves, A.; Benchimol, M. Phagocytosis by Trichomonas vaginalis: New insights. Biol. Cell 2007, 99, 87–101. [Google Scholar] [CrossRef]
  26. Zimmann, N.; Rada, P.; Žárský, V.; Smutná, T.; Záhonová, K.; Dacks, J.; Harant, K.; Hrdý, I.; Tachezy, J. Proteomic analysis of Trichomonas vaginalis phagolysosome, lysosomal targeting, and unconventional secretion of cysteine peptidases. Mol. Cell. Proteomics 2022, 21, 100174. [Google Scholar] [CrossRef]
  27. Stanley, S.L.J. Amoebiasis. Lancet 2003, 361, 1025–1034. [Google Scholar] [CrossRef]
  28. Wiser, M.F. Protozoa. In Encyclopedia of Biodiversity (Volume 2); Scheiner, S.M.B.T., Ed.; Academic Press: Oxford, UK, 2024; pp. 802–817. ISBN 978-0-323-98434-8. [Google Scholar]
  29. Iyer, L.R.; Verma, A.K.; Paul, J.; Bhattacharya, A. Phagocytosis of gut bacteria by Entamoeba histolytica. Front. Cell. Infect. Microbiol. 2019, 9, 34. [Google Scholar] [CrossRef]
  30. Orozco, E.; Guarneros, G.; Martinez-Palomo, A.; Sánchez, T. Entamoeba histolytica. Phagocytosis as a virulence factor. J. Exp. Med. 1983, 158, 1511–1521. [Google Scholar] [CrossRef]
  31. Boettner, D.R.; Huston, C.D.; Sullivan, J.A.; Petri, W.A.J. Entamoeba histolytica and Entamoeba dispar utilize externalized phosphatidylserine for recognition and phagocytosis of erythrocytes. Infect. Immun. 2005, 73, 3422–3430. [Google Scholar] [CrossRef]
  32. Talamás-Lara, D.; Chávez-Munguía, B.; González-Robles, A.; Talamás-Rohana, P.; Salazar-Villatoro, L.; Durán-Díaz, Á.; Martínez-Palomo, A. Erythrophagocytosis in Entamoeba histolytica and Entamoeba dispar: A comparative study. Biomed Res. Int. 2014, 2014, 626259. [Google Scholar] [CrossRef]
  33. Watanabe, N.; Nakada-Tsukui, K.; Nozaki, T. Molecular dissection of phagocytosis by proteomic analysis in Entamoeba histolytica. Genes 2023, 14, 379. [Google Scholar] [CrossRef]
  34. Nakada-Tsukui, K.; Tsuboi, K.; Furukawa, A.; Yamada, Y.; Nozaki, T. A novel class of cysteine protease receptors that mediate lysosomal transport. Cell. Microbiol. 2012, 14, 1299–1317. [Google Scholar] [CrossRef]
  35. Bañuelos, C.; Betanzos, A.; Javier-Reyna, R.; Galindo, A.; Orozco, E. Molecular interplays of the Entamoeba histolytica endosomal sorting complexes required for transport during phagocytosis. Front. Cell. Infect. Microbiol. 2022, 12, 855797. [Google Scholar] [CrossRef]
  36. Verma, K.; Nozaki, T.; Datta, S. Role of EhRab7A in phagocytosis of type 1 fimbriated E. coli by Entamoeba histolytica. Mol. Microbiol. 2016, 102, 1043–1061. [Google Scholar] [CrossRef]
  37. Verma, K.; Srivastava, V.K.; Datta, S. Rab GTPases take centre stage in understanding Entamoeba histolytica biology. Small GTPases 2020, 11, 320–333. [Google Scholar] [CrossRef]
  38. Bonifacino, J.S.; Glick, B.S. The mechanisms of vesicle budding and fusion. Cell 2004, 116, 153–166. [Google Scholar] [CrossRef]
  39. Bettadapur, A.; Miller, H.W.; Ralston, K.S. Biting off what can be chewed: Trogocytosis in health, infection, and disease. Infect. Immun. 2020, 88. [Google Scholar] [CrossRef]
  40. Ralston, K.S.; Solga, M.D.; Mackey-Lawrence, N.M.; Somlata; Bhattacharya, A.; Petri, W.A. Trogocytosis by Entamoeba histolytica contributes to cell killing and tissue invasion. Nature 2014, 508, 526–530. [Google Scholar] [CrossRef]
  41. Miller, H.W.; Suleiman, R.L.; Ralston, K.S. Trogocytosis by Entamoeba histolytica mediates acquisition and display of human cell membrane proteins and evasion of lysis by human serum. MBio 2019, 10. [Google Scholar] [CrossRef]
  42. Somlata; Nakada-Tsukui, K.; Nozaki, T. AGC family kinase 1 participates in trogocytosis but not in phagocytosis in Entamoeba histolytica. Nat. Commun. 2017, 8, 101. [Google Scholar] [CrossRef]
  43. Kinnear, F.B. Cytopathogenicity of Acanthamoeba, Vahlkampfia and Hartmannella: Quantative & qualitative in vitro studies on keratocytes. J. Infect. 2003, 46, 228–237. [Google Scholar] [CrossRef]
  44. Marciano-Cabral, F.; Zoghby, K.L.; Bradley, S.G. Cytopathic action of Naegleria fowleri amoebae on rat neuroblastoma target cells. J. Protozool. 1990, 37, 138–144. [Google Scholar] [CrossRef]
  45. Xiang, C.; Li, Y.; Jing, S.; Han, S.; He, H. Trichomonas gallinae kills host cells using trogocytosis. Pathogens 2023, 12, 1008. [Google Scholar] [CrossRef]
  46. Holberton, D. V Fine structure of the ventral disk apparatus and the mechanism of attachment in the flagellate Giardia muris. J. Cell Sci. 1973, 13, 11–41. [Google Scholar] [CrossRef]
  47. Lanfredi-Rangel, A.; Attias, M.; de Carvalho, T.M.; Kattenbach, W.M.; De Souza, W. The peripheral vesicles of trophozoites of the primitive protozoan Giardia lamblia may correspond to early and late endosomes and to lysosomes. J. Struct. Biol. 1998, 123, 225–235. [Google Scholar] [CrossRef]
  48. Touz, M.C.; Rivero, M.R.; Miras, S.L.; Bonifacino, J.S. Lysosomal protein trafficking in Giardia lamblia: Common and distinct features. Front. Biosci. 2012, 4, 1898–1909. [Google Scholar] [CrossRef]
  49. Rivero, M.R.; Jausoro, I.; Bisbal, M.; Feliziani, C.; Lanfredi-Rangel, A.; Touz, M.C. Receptor-mediated endocytosis and trafficking between endosomal-lysosomal vacuoles in Giardia lamblia. Parasitol. Res. 2013, 112, 1813–1818. [Google Scholar] [CrossRef]
  50. Zumthor, J.P.; Cernikova, L.; Rout, S.; Kaech, A.; Faso, C.; Hehl, A.B. Static clathrin assemblies at the peripheral vacuole-plasma membrane interface of the parasitic protozoan Giardia lamblia. PLoS Pathog. 2016, 12, e1005756. [Google Scholar] [CrossRef]
  51. Abodeely, M.; DuBois, K.N.; Hehl, A.; Stefanic, S.; Sajid, M.; DeSouza, W.; Attias, M.; Engel, J.C.; Hsieh, I.; Fetter, R.D.; et al. A contiguous compartment functions as endoplasmic reticulum and endosome/lysosome in Giardia lamblia. Eukaryot. Cell 2009, 8, 1665–1676. [Google Scholar] [CrossRef]
  52. Benchimol, M. Giardia intestinalis can interact, change its shape and internalize large particles and microorganisms. Parasitology 2021, 148, 500–510. [Google Scholar] [CrossRef]
  53. Deschamps, P.; Lara, E.; Marande, W.; López-García, P.; Ekelund, F.; Moreira, D. Phylogenomic analysis of kinetoplastids supports that trypanosomatids arose from within bodonids. Mol. Biol. Evol. 2011, 28, 53–58. [Google Scholar] [CrossRef] [PubMed]
  54. Stephens, N.A.; Kieft, R.; Macleod, A.; Hajduk, S.L. Trypanosome resistance to human innate immunity: Targeting Achilles’ heel. Trends Parasitol. 2012, 28, 539–545. [Google Scholar] [CrossRef] [PubMed]
  55. Field, M.C.; Carrington, M. The trypanosome flagellar pocket. Nat. Rev. Microbiol. 2009, 7, 775–786. [Google Scholar] [CrossRef] [PubMed]
  56. Webster, P.; Russell, D.G. The flagellar pocket of trypanosomatids. Parasitol. Today 1993, 9, 201–206. [Google Scholar] [CrossRef] [PubMed]
  57. Coppens, I.; Courtoy, P.J. The adaptative mechanisms of Trypanosoma brucei for sterol homeostasis in its different life-cycle environments. Annu. Rev. Microbiol. 2000, 54, 129–156. [Google Scholar] [CrossRef] [PubMed]
  58. Engstler, M.; Pfohl, T.; Herminghaus, S.; Boshart, M.; Wiegertjes, G.; Heddergott, N.; Overath, P. Hydrodynamic flow-mediated protein sorting on the cell surface of trypanosomes. Cell 2007, 131, 505–515. [Google Scholar] [CrossRef]
  59. Reyes-López, M.; Piña-Vázquez, C.; Serrano-Luna, J. Transferrin: Endocytosis and cell signaling in parasitic protozoa. BioMed Res. Int. 2015, 2015, 641392. [Google Scholar] [CrossRef]
  60. Allen, C.L.; Goulding, D.; Field, M.C. Clathrin-mediated endocytosis is essential in Trypanosoma brucei. EMBO J. 2003, 22, 4991–5002. [Google Scholar] [CrossRef]
  61. Manna, P.T.; Obado, S.O.; Boehm, C.; Gadelha, C.; Sali, A.; Chait, B.T.; Rout, M.P.; Field, M.C. Lineage-specific proteins essential for endocytosis in trypanosomes. J. Cell Sci. 2017, 130, 1379–1392. [Google Scholar] [CrossRef]
  62. Adung’a, V.O.; Gadelha, C.; Field, M.C. Proteomic analysis of clathrin interactions in trypanosomes reveals dynamic evolution of endocytosis. Traffic 2013, 14, 440–457. [Google Scholar] [CrossRef]
  63. Kalb, L.C.; Antunes Frederico, Y.C.; Batista, C.M.; Eger, I.; Fragoso, S.P.; Soares, M.J. Clathrin expression in Trypanosoma cruzi. BMC Cell Biol. 2014, 15, 23. [Google Scholar] [CrossRef] [PubMed]
  64. Ansari, I.; Basak, R.; Mukhopadhyay, A. Hemoglobin endocytosis and intracellular trafficking: A novel way of heme acquisition by Leishmania. Pathogens 2022, 11, 585. [Google Scholar] [CrossRef] [PubMed]
  65. Soares, M.J.; de Souza, W. Endocytosis of gold-labeled proteins and LDL by Trypanosoma cruzi. Parasitol. Res. 1991, 77, 461–468. [Google Scholar] [CrossRef] [PubMed]
  66. Andrade-Neto, V.V.; Cicco, N.N.T.; Cunha-Junior, E.F.; Canto-Cavalheiro, M.M.; Atella, G.C.; Torres-Santos, E.C. The pharmacological inhibition of sterol biosynthesis in Leishmania is counteracted by enhancement of LDL endocytosis. Acta Trop. 2011, 119, 194–198. [Google Scholar] [CrossRef] [PubMed]
  67. De Cicco, N.N.T.; Pereira, M.G.; Corrêa, J.R.; Andrade-Neto, V.V.; Saraiva, F.B.; Chagas-Lima, A.C.; Gondim, K.C.; Torres-Santos, E.C.; Folly, E.; Saraiva, E.M.; et al. LDL uptake by Leishmania amazonensis: Involvement of membrane lipid microdomains. Exp. Parasitol. 2012, 130, 330–340. [Google Scholar] [CrossRef] [PubMed]
  68. Engstler, M.; Thilo, L.; Weise, F.; Grünfelder, C.G.; Schwarz, H.; Boshart, M.; Overath, P. Kinetics of endocytosis and recycling of the GPI-anchored variant surface glycoprotein in Trypanosoma brucei. J. Cell Sci. 2004, 117, 1105–1115. [Google Scholar] [CrossRef] [PubMed]
  69. Langreth, S.G.; Balber, A.E. Protein uptake and digestion in bloodstream and culture forms of Trypanosoma brucei. J. Protozool. 1975, 22, 40–53. [Google Scholar] [CrossRef]
  70. Morgan, G.W.; Hall, B.S.; Denny, P.W.; Field, M.C.; Carrington, M. The endocytic apparatus of the kinetoplastida. Part II: Machinery and components of the system. Trends Parasitol. 2002, 18, 540–546. [Google Scholar] [CrossRef] [PubMed]
  71. Link, F.; Borges, A.R.; Jones, N.G.; Engstler, M. To the surface and back: Exo- and endocytic pathways in Trypanosoma brucei. Front. Cell Dev. Biol. 2021, 9, 720521. [Google Scholar] [CrossRef]
  72. Porto-Carreiro, I.; Attias, M.; Miranda, K.; De Souza, W.; Cunha-e-Silva, N. Trypanosoma cruzi epimastigote endocytic pathway: Cargo enters the cytostome and passes through an early endosomal network before storage in reservosomes. Eur. J. Cell Biol. 2000, 79, 858–869. [Google Scholar] [CrossRef]
  73. Cunha-e-Silva, N.; Sant’Anna, C.; Pereira, M.G.; Porto-Carreiro, I.; Jeovanio, A.L.; de Souza, W. Reservosomes: Multipurpose organelles? Parasitol. Res. 2006, 99, 325–327. [Google Scholar] [CrossRef] [PubMed]
  74. Sant’Anna, C.; Parussini, F.; Lourenço, D.; de Souza, W.; Cazzulo, J.J.; Cunha-e-Silva, N.L. All Trypanosoma cruzi developmental forms present lysosome-related organelles. Histochem. Cell Biol. 2008, 130, 1187–1198. [Google Scholar] [CrossRef]
  75. Wang, Z.; Wheeler, R.J.; Sunter, J.D. Lysosome assembly and disassembly changes endocytosis rate through the Leishmania cell cycle. Microbiol. Open 2020, 9, e969. [Google Scholar] [CrossRef] [PubMed]
  76. Mullin, K.A.; Foth, B.J.; Ilgoutz, S.C.; Callaghan, J.M.; Zawadzki, J.L.; McFadden, G.I.; McConville, M.J. Regulated degradation of an endoplasmic reticulum membrane protein in a tubular lysosome in Leishmania mexicana. Mol. Biol. Cell 2001, 12, 2364–2377. [Google Scholar] [CrossRef] [PubMed]
  77. Ueda-Nakamura, T.; Attias, M.; de Souza, W. Megasome biogenesis in Leishmania amazonensis: A morphometric and cytochemical study. Parasitol. Res. 2001, 87, 89–97. [Google Scholar] [CrossRef] [PubMed]
  78. Waller, R.F.; McConville, M.J. Developmental changes in lysosome morphology and function Leishmania parasites. Int. J. Parasitol. 2002, 32, 1435–1445. [Google Scholar] [CrossRef] [PubMed]
  79. Etheridge, R.D. Protozoan phagotrophy from predators to parasites: An overview of the enigmatic cytostome-cytopharynx complex of Trypanosoma cruzi. J. Eukaryot. Microbiol. 2022, 69, e12896. [Google Scholar] [CrossRef]
  80. Flegontov, P.; Votýpka, J.; Skalický, T.; Logacheva, M.D.; Penin, A.A.; Tanifuji, G.; Onodera, N.T.; Kondrashov, A.S.; Volf, P.; Archibald, J.M.; et al. Paratrypanosoma is a novel early-branching trypanosomatid. Curr. Biol. 2013, 23, 1787–1793. [Google Scholar] [CrossRef]
  81. Alcantara, C.L.; de Souza, W.; Cunha, E.; Silva, N.L. The cytostome-cytopharynx complex of intracellular and extracellular amastigotes of Trypanosoma cruzi exhibit structural and functional differences. Cell. Microbiol. 2021, 23, e13346. [Google Scholar] [CrossRef]
  82. Votýpka, J.; Modrý, D.; Oborník, M.; Šlapeta, J.; Lukeš, J. Apicomplexa. In Handbook of the Protists; Archibald, J.M., Simpson, A.G.B., Slamovits, C.H., Margulis, L., Melkonian, M., Chapman, D.J., Corliss, J.O., Eds.; Springer International Publishing: Cham, Switzerland, 2017; pp. 1–58. ISBN 978-3-319-32669-6. [Google Scholar]
  83. Janouškovec, J.; Tikhonenkov, D.V.; Burki, F.; Howe, A.T.; Kolísko, M.; Mylnikov, A.P.; Keeling, P.J. Factors mediating plastid dependency and the origins of parasitism in apicomplexans and their close relatives. Proc. Natl. Acad. Sci. USA 2015, 112, 10200–10207. [Google Scholar] [CrossRef]
  84. Cumbo, V.R.; Baird, A.H.; Moore, R.B.; Negri, A.P.; Neilan, B.A.; Salih, A.; van Oppen, M.J.H.; Wang, Y.; Marquis, C.P. Chromera velia is endosymbiotic in larvae of the reef corals Acropora digitifera and A. tenuis. Protist 2013, 164, 237–244. [Google Scholar] [CrossRef] [PubMed]
  85. Getty, T.A.; Peterson, J.W.; Fujioka, H.; Walsh, A.M.; Sam-Yellowe, T.Y. Colpodella sp. (ATCC 50594) life cycle: Myzocytosis and possible links to the origin of intracellular parasitism. Trop. Med. Infect. Dis. 2021, 6, 127. [Google Scholar] [CrossRef] [PubMed]
  86. Frénal, K.; Dubremetz, J.-F.; Lebrun, M.; Soldati-Favre, D. Gliding motility powers invasion and egress in Apicomplexa. Nat. Rev. Microbiol. 2017, 15, 645–660. [Google Scholar] [CrossRef] [PubMed]
  87. Besteiro, S.; Dubremetz, J.-F.; Lebrun, M. The moving junction of apicomplexan parasites: A key structure for invasion. Cell. Microbiol. 2011, 13, 797–805. [Google Scholar] [CrossRef] [PubMed]
  88. Cowman, A.F.; Tonkin, C.J.; Tham, W.H.; Duraisingh, M.T. The molecular basis of erythrocyte invasion by malaria parasites. Cell Host Microbe 2017, 22, 232–245. [Google Scholar] [CrossRef]
  89. Wiser, M.F. Unique endomembrane systems and virulence in pathogenic protozoa. Life 2021, 11, 822. [Google Scholar] [CrossRef] [PubMed]
  90. Valigurová, A.; Florent, I. Nutrient acquisition and attachment strategies in basal lineages: A tough nut to crack in the evolutionary puzzle of Apicomplexa. Microorganisms 2021, 9, 1430. [Google Scholar] [CrossRef] [PubMed]
  91. Gubbels, M.-J.; Duraisingh, M.T. Evolution of apicomplexan secretory organelles. Int. J. Parasitol. 2012, 42, 1071–1081. [Google Scholar] [CrossRef]
  92. Cavalier-Smith, T.; Chao, E.E. Protalveolate phylogeny and systematics and the origins of sporozoa and dinoflagellates (phylum Myzozoa nom. nov.). Eur. J. Protistol. 2004, 40, 185–212. [Google Scholar] [CrossRef]
  93. Hansen, P.J.; Calado, A.J. Phagotrophic mechanisms and prey selection in free-living dinoflagellates. J. Eukaryot. Microbiol. 1999, 46, 382–389. [Google Scholar] [CrossRef]
  94. Brugerolle, G. Colpodella vorax: Ultrastructure, predation, life-cycle, mitosis, and phylogenetic relationships. Eur. J. Protistol. 2002, 38, 113–125. [Google Scholar] [CrossRef]
  95. Sam-Yellowe, T.Y.; Asraf, M.M.; Peterson, J.W.; Fujioka, H. Fluorescent nanoparticle uptake by myzocytosis and endocytosis in Colpodella sp. ATCC 50594. Microorganisms 2023, 11, 1945. [Google Scholar] [CrossRef] [PubMed]
  96. Simdyanov, T.G.; Kuvardina, O.N. Fine structure and putative feeding mechanism of the archigregarine Selenidium orientale (Apicomplexa: Gregarinomorpha). Eur. J. Protistol. 2007, 43, 17–25. [Google Scholar] [CrossRef]
  97. Schrével, J.; Valigurová, A.; Prensier, G.; Chambouvet, A.; Florent, I.; Guillou, L. Ultrastructure of Selenidium pendula, the type species of archigregarines, and phylogenetic relations to other marine apicomplexa. Protist 2016, 167, 339–368. [Google Scholar] [CrossRef] [PubMed]
  98. Gerace, E.; Lo Presti, V.D.M.; Biondo, C. Cryptosporidium infection: Epidemiology, pathogenesis, and differential diagnosis. Eur. J. Microbiol. Immunol. 2019, 9, 119–123. [Google Scholar] [CrossRef] [PubMed]
  99. O’Hara, S.P.; Small, A.J.; Chen, X.-M.; LaRusso, N.F. Host cell actin remodeling in response to Cryptosporidium. Subcell. Biochem. 2008, 47, 92–100. [Google Scholar] [CrossRef]
  100. Lendner, M.; Daugschies, A. Cryptosporidium infections: Molecular advances. Parasitology 2014, 141, 1511–1532. [Google Scholar] [CrossRef]
  101. Borowski, H.; Clode, P.L.; Thompson, R.C.A. Active invasion and/or encapsulation? A reappraisal of host-cell parasitism by Cryptosporidium. Trends Parasitol. 2008, 24, 509–516. [Google Scholar] [CrossRef]
  102. Perkins, M.E.; Riojas, Y.A.; Wu, T.W.; Le Blancq, S.M. CpABC, a Cryptosporidium parvum ATP-binding cassette protein at the host–parasite boundary in intracellular stages. Proc. Natl. Acad. Sci. USA 1999, 96, 5734–5739. [Google Scholar] [CrossRef]
  103. Ehrenman, K.; Wanyiri, J.W.; Bhat, N.; Ward, H.D.; Coppens, I. Cryptosporidium parvum scavenges LDL-derived cholesterol and micellar cholesterol internalized into enterocytes. Cell. Microbiol. 2013, 15, 1182–1197. [Google Scholar] [CrossRef]
  104. Beĭer, T.V.; Sidorenko, N.V.; Lakovnikova, E. V [Electron microscopic research on Cryptosporidium. I. The asexual stages in the development of Cryptosporidium parvum]. Tsitologiia 1990, 32, 462–468. [Google Scholar] [PubMed]
  105. Wiser, M.F. The digestive vacuole of the malaria parasite: A specialized lysosome. Pathogens 2024, 13, 182. [Google Scholar] [CrossRef]
  106. Wunderlich, J.; Rohrbach, P.; Dalton, J.P. The malaria digestive vacuole. Front. Biosci. 2012, 4, 1424–1448. [Google Scholar] [CrossRef]
  107. Ngwa, C.J.; Rosa, T.F.; Pradel, G. The biology of malaria gametocytes. In Current Topics in Malaria; Rodriguez-Morales, A.J., Ed.; IntechOpen: Rijeka, Croatia, 2016; Chapter 7; pp. 117–144. Available online: https://www.intechopen.com/chapters/52526 (accessed on 17 June 2024).
  108. Slomianny, C. Three-dimensional reconstruction of the feeding process of the malaria parasite. Blood Cells 1990, 16, 369–378. [Google Scholar] [PubMed]
  109. Spielmann, T.; Gras, S.; Sabitzki, R.; Meissner, M. Endocytosis in Plasmodium and Toxoplasma parasites. Trends Parasitol. 2020, 36, 520–532. [Google Scholar] [CrossRef]
  110. Henrici, R.C.; Edwards, R.L.; Zoltner, M.; van Schalkwyk, D.A.; Hart, M.N.; Mohring, F.; Moon, R.W.; Nofal, S.D.; Patel, A.; Flueck, C.; et al. The Plasmodium falciparum artemisinin susceptibility-associated AP-2 adaptin μ subunit is clathrin independent and essential for schizont maturation. MBio 2020, 11, e02918-19. [Google Scholar] [CrossRef] [PubMed]
  111. Birnbaum, J.; Scharf, S.; Schmidt, S.; Jonscher, E.; Hoeijmakers, W.A.M.; Flemming, S.; Toenhake, C.G.; Schmitt, M.; Sabitzki, R.; Bergmann, B.; et al. A Kelch13-defined endocytosis pathway mediates artemisinin resistance in malaria parasites. Science 2020, 367, 51–59. [Google Scholar] [CrossRef]
  112. Pan, M.; Ge, C.-C.; Fan, Y.-M.; Jin, Q.-W.; Shen, B.; Huang, S.-Y. The determinants regulating Toxoplasma gondii bradyzoite development. Front. Microbiol. 2022, 13, 1027073. [Google Scholar] [CrossRef] [PubMed]
  113. Frölich, S.; Wallach, M. Use of fluorescent nanoparticles to investigate nutrient acquisition by developing Eimeria maxima macrogametocytes. Sci. Rep. 2016, 6, 29030. [Google Scholar] [CrossRef]
  114. Kannan, G.; Thaprawat, P.; Schultz, T.L.; Carruthers, V.B. Acquisition of host cytosolic protein by Toxoplasma gondii bradyzoites. mSphere 2021, 6, e00934-20. [Google Scholar] [CrossRef]
  115. McGovern, O.L.; Rivera-Cuevas, Y.; Carruthers, V.B. Emerging mechanisms of endocytosis in Toxoplasma gondii. Life 2021, 11, 84. [Google Scholar] [CrossRef] [PubMed]
  116. Gras, S.; Jimenez-Ruiz, E.; Klinger, C.M.; Schneider, K.; Klingl, A.; Lemgruber, L.; Meissner, M. An endocytic-secretory cycle participates in Toxoplasma gondii in motility. PLoS Biol. 2019, 17, e3000060. [Google Scholar] [CrossRef] [PubMed]
  117. Goldberg, D.E.; Zimmerberg, J. Hardly vacuous: The parasitophorous vacuolar membrane of malaria parasites. Trends Parasitol. 2020, 36, 138–146. [Google Scholar] [CrossRef] [PubMed]
  118. Matz, J.M.; Beck, J.R.; Blackman, M.J. The parasitophorous vacuole of the blood-stage malaria parasite. Nat. Rev. Microbiol. 2020, 18, 379–391. [Google Scholar] [CrossRef] [PubMed]
  119. Gohil, S.; Kats, L.M.; Sturm, A.; Cooke, B.M. Recent insights into alteration of red blood cells by Babesia bovis: Moovin’ forward. Trends Parasitol. 2010, 26, 591–599. [Google Scholar] [CrossRef] [PubMed]
  120. Repnik, U.; Gangopadhyay, P.; Bietz, S.; Przyborski, J.M.; Griffiths, G.; Lingelbach, K. The apicomplexan parasite Babesia divergens internalizes band 3, glycophorin A and spectrin during invasion of human red blood cells. Cell. Microbiol. 2015, 17, 1052–1068. [Google Scholar] [CrossRef] [PubMed]
  121. Shaw, M.K.; Tilney, L.G.; Musoke, A.J. The entry of Theileria parva sporozoites into bovine lymphocytes: Evidence for MHC class I involvement. J. Cell Biol. 1991, 113, 87–101. [Google Scholar] [CrossRef] [PubMed]
  122. Gold, D.A.; Kaplan, A.D.; Lis, A.; Bett, G.C.L.; Rosowski, E.E.; Cirelli, K.M.; Bougdour, A.; Sidik, S.M.; Beck, J.R.; Lourido, S.; et al. The Toxoplasma dense granule proteins GRA17 and GRA23 mediate the movement of small molecules between the host and the parasitophorous vacuole. Cell Host Microbe 2015, 17, 642–652. [Google Scholar] [CrossRef]
  123. Garten, M.; Nasamu, A.S.; Niles, J.C.; Zimmerberg, J.; Goldberg, D.E.; Beck, J.R. EXP2 is a nutrient-permeable channel in the vacuolar membrane of Plasmodium and is essential for protein export via PTEX. Nat. Microbiol. 2018, 3, 1090–1098. [Google Scholar] [CrossRef]
  124. Dluzewski, A.R.; Ling, I.T.; Hopkins, J.M.; Grainger, M.; Margos, G.; Mitchell, G.H.; Holder, A.A.; Bannister, L.H. Formation of the food vacuole in Plasmodium falciparum: A potential role for the 19 kDa fragment of merozoite surface protein 1 (MSP1(19)). PLoS ONE 2008, 3, e3085. [Google Scholar] [CrossRef]
  125. Romano, J.D.; Nolan, S.J.; Porter, C.; Ehrenman, K.; Hartman, E.J.; Hsia, R.-C.; Coppens, I. The parasite Toxoplasma sequesters diverse Rab host vesicles within an intravacuolar network. J. Cell Biol. 2017, 216, 4235–4254. [Google Scholar] [CrossRef] [PubMed]
  126. Nolan, S.J.; Romano, J.D.; Coppens, I. Host lipid droplets: An important source of lipids salvaged by the intracellular parasite Toxoplasma gondii. PLoS Pathog. 2017, 13, e1006362. [Google Scholar] [CrossRef] [PubMed]
  127. Coppens, I.; Dunn, J.D.; Romano, J.D.; Pypaert, M.; Zhang, H.; Boothroyd, J.C.; Joiner, K.A. Toxoplasma gondii sequesters lysosomes from mammalian hosts in the vacuolar space. Cell 2006, 125, 261–274. [Google Scholar] [CrossRef] [PubMed]
  128. Rivera-Cuevas, Y.; Mayoral, J.; Di Cristina, M.; Lawrence, A.-L.E.; Olafsson, E.B.; Patel, R.K.; Thornhill, D.; Waldman, B.S.; Ono, A.; Sexton, J.Z.; et al. Toxoplasma gondii exploits the host ESCRT machinery for parasite uptake of host cytosolic proteins. PLOS Pathog. 2021, 17, e1010138. [Google Scholar] [CrossRef] [PubMed]
  129. Francis, S.E.; Sullivan, D.J.; Goldberg, D.E. Hemoglobin metabolism in the malaria parasite Plasmodium falciparum. Annu. Rev. Microbiol. 1997, 51, 97–123. [Google Scholar] [CrossRef] [PubMed]
  130. Egan, T.J. Haemozoin formation. Mol. Biochem. Parasitol. 2008, 157, 127–136. [Google Scholar] [CrossRef] [PubMed]
  131. Stasic, A.J.; Moreno, S.N.J.; Carruthers, V.B.; Dou, Z. The Toxoplasma plant-like vacuolar compartment (PLVAC). J. Eukaryot. Microbiol. 2022, 69, e12951. [Google Scholar] [CrossRef] [PubMed]
  132. Dou, Z.; McGovern, O.L.; Di Cristina, M.; Carruthers, V.B. Toxoplasma gondii ingests and digests host cytosolic proteins. MBio 2014, 5, e01188-14. [Google Scholar] [CrossRef] [PubMed]
  133. Saliba, K.J.; Allen, R.J.W.; Zissis, S.; Bray, P.G.; Ward, S.A.; Kirk, K. Acidification of the malaria parasite’s digestive vacuole by a H+-ATPase and a H+-pyrophosphatase. J. Biol. Chem. 2003, 278, 5605–5612. [Google Scholar] [CrossRef]
  134. Rea, P.A.; Poole, R.J. Vacuolar H+-translocating pyrophosphatase. Annu. Rev. Plant Biol. 1993, 44, 157–180. [Google Scholar] [CrossRef]
  135. Shafik, S.H.; Cobbold, S.A.; Barkat, K.; Richards, S.N.; Lancaster, N.S.; Llinás, M.; Hogg, S.J.; Summers, R.L.; McConville, M.J.; Martin, R.E. The natural function of the malaria parasite’s chloroquine resistance transporter. Nat. Commun. 2020, 11, 3922. [Google Scholar] [CrossRef] [PubMed]
  136. Warring, S.D.; Dou, Z.; Carruthers, V.B.; McFadden, G.I.; van Dooren, G.G. Characterization of the chloroquine resistance transporter homologue in Toxoplasma gondii. Eukaryot. Cell 2014, 13, 1360–1370. [Google Scholar] [CrossRef] [PubMed]
  137. Senaud, J.; Chobotar, B.; Scholtyseck, E. Role of the micropore in nutrition of the Sporozoa. Ultrastructural study of Plasmodium cathemerium, Eimeria ferrisi, E. stiedai, Besnoitia jellisoni, and Frenkelia sp. Tropenmed. Parasitol. 1976, 27, 145–159. [Google Scholar] [PubMed]
  138. Kono, M.; Prusty, D.; Parkinson, J.; Gilberger, T.W. The apicomplexan inner membrane complex. Front. Biosci. 2013, 18, 982–992. [Google Scholar] [CrossRef]
  139. Gould, S.B.; Tham, W.-H.; Cowman, A.F.; McFadden, G.I.; Waller, R.F. Alveolins, a new family of cortical proteins that define the protist infrakingdom Alveolata. Mol. Biol. Evol. 2008, 25, 1219–1230. [Google Scholar] [CrossRef] [PubMed]
  140. Appleton, P.L.; Vickerman, K. Presence of apicomplexan-type micropores in a parasitic dinoflagellate, Hematodinium sp. Parasitol. Res. 1996, 82, 279–282. [Google Scholar] [CrossRef] [PubMed]
  141. Nichols, B.A.; Chiappino, M.L.; Pavesio, C.E. Endocytosis at the micropore of Toxoplasma gondii. Parasitol. Res. 1994, 80, 91–98. [Google Scholar] [CrossRef] [PubMed]
  142. Wan, W.; Dong, H.; Lai, D.-H.; Yang, J.; He, K.; Tang, X.; Liu, Q.; Hide, G.; Zhu, X.-Q.; Sibley, L.D.; et al. The Toxoplasma micropore mediates endocytosis for selective nutrient salvage from host cell compartments. Nat. Commun. 2023, 14, 977. [Google Scholar] [CrossRef] [PubMed]
  143. Koreny, L.; Mercado-Saavedra, B.N.; Klinger, C.M.; Barylyuk, K.; Butterworth, S.; Hirst, J.; Rivera-Cuevas, Y.; Zaccai, N.R.; Holzer, V.J.C.; Klingl, A.; et al. Stable endocytic structures navigate the complex pellicle of apicomplexan parasites. Nat. Commun. 2023, 14, 2167. [Google Scholar] [CrossRef]
  144. Xie, S.C.; Ralph, S.A.; Tilley, L. K13, the cytostome, and artemisinin resistance. Trends Parasitol. 2020, 36, 533–544. [Google Scholar] [CrossRef]
  145. Ferreira, J.L.; Heincke, D.; Wichers, J.S.; Liffner, B.; Wilson, D.W.; Gilberger, T.-W. The dynamic roles of the inner membrane complex in the multiple stages of the malaria parasite. Front. Cell. Infect. Microbiol. 2020, 10, 611801. [Google Scholar] [CrossRef] [PubMed]
Parasitologia 04 00019 g001
Figure 1. Endocytosis and the digestive vacuole. Following endocytosis or phagocytosis, the digestive vacuole is formed from the endosome or phagosome. In some eukaryotes, especially animal cells, lysosomes deliver the proton (H+) pumps and acid hydrolases to the digestive vacuole. Acid hydrolases are hydrolytic enzymes, such as proteases, lipases, nucleases, glycosidases, and phosphatases, that have acidic pH optima. Complex macromolecules are broken down into small-molecular-weight metabolites.
Figure 1. Endocytosis and the digestive vacuole. Following endocytosis or phagocytosis, the digestive vacuole is formed from the endosome or phagosome. In some eukaryotes, especially animal cells, lysosomes deliver the proton (H+) pumps and acid hydrolases to the digestive vacuole. Acid hydrolases are hydrolytic enzymes, such as proteases, lipases, nucleases, glycosidases, and phosphatases, that have acidic pH optima. Complex macromolecules are broken down into small-molecular-weight metabolites.
Parasitologia 04 00019 g001
Table 1. Endocytosis mechanisms.
Table 1. Endocytosis mechanisms.
TypeMechanismSize
PhagocytosisParticles surrounded by pseudopodia>0.5 μm
Macro-pinocytosisFluid uptake by membrane ruffles0.2–10 μm
Clathrin-mediated endocytosis (CME)Clathrin in conjunction with adaptor proteins forms coated vesicles60–120 nm
Clathrin-independent endocytosis (CIE)Several distinct mechanisms involving various proteins and lipid rafts 50–80 nm
Table 2. Key features of medically important kinetoplastids.
Table 2. Key features of medically important kinetoplastids.
KinetoplastidEndocytosisDigestive Vacuole
African trypanosomesClathrin-mediated at the flagellar pocket Single large lysosome
T. cruziClathrin-mediated at the flagellar pocket and clathrin-independent at the cytostomeSeveral lysosome-like vacuoles called reservosomes
LeishmaniaClathrin-mediated at the flagellar pocketVesicle-filled tubules or vacuoles called megasomes
Table 3. Summary of feeding mechanisms by pathogenic protozoa.
Table 3. Summary of feeding mechanisms by pathogenic protozoa.
PathogenPrimary Feeding Mechanism
EntamoebaExtensive phagocytosis of bacteria and host cells including trogocytosis
GiardiaPossible clathrin-mediated process involving a unique lysosomal compartment called peripheral vacuoles
KinetoplastidsClathrin-mediated endocytosis at the flagellar pocket
PlasmodiumSpecialized digestive vacuole for hemoglobin catabolism and hemozoin formation
ToxoplasmaEndocytosis from a modified parasitophorous vacuole and a plant-like digestive vacuole
CryptosporidiumMay lack endocytosis and digestive vacuole and take up nutrients via a feeder organelle and myzocytosis
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Wiser, M.F. Feeding Mechanisms of Pathogenic Protozoa with a Focus on Endocytosis and the Digestive Vacuole. Parasitologia 2024, 4, 222-237. https://doi.org/10.3390/parasitologia4030019

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Wiser MF. Feeding Mechanisms of Pathogenic Protozoa with a Focus on Endocytosis and the Digestive Vacuole. Parasitologia. 2024; 4(3):222-237. https://doi.org/10.3390/parasitologia4030019

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Wiser, Mark F. 2024. "Feeding Mechanisms of Pathogenic Protozoa with a Focus on Endocytosis and the Digestive Vacuole" Parasitologia 4, no. 3: 222-237. https://doi.org/10.3390/parasitologia4030019

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