Cryptosporidium spp.

Unlike other members of the Apicomplexan group of parasites, organisms of the genus Cryptosporidium spp. lack the apicoplast [29]. C. parvum and C. hominis are capable of causing severe diarrhea and gastrointestinal discomfort in humans, which can be life threatening in young children, the elderly, and immunocompromised individuals [30]. The disease has also major implications for life stock as it frequently causes fatal diarrhea in calves [31]. The infective oocysts when ingested establish a disease cycle within the host, which propagates itself into the surrounding environment through the shedding of oocysts in stool [32]. C. parvum is a monoxenous parasite, completing its life cycle in a single host. The asexual cycle includes multiplication of the parasite by schizogony, followed by the sexual cycle involving the production of environmentally stable thick-walled oocysts shed in feces and thin-walled oocysts causing autoinfection of host intestinal epithelium [28].

Cryptosporidium is by far the most well-researched parasite in terms of the abundance and complexity of culture models described. Previously, various human and animal cell lines have been the basis for monolayer cultures of C. parvum. However, complete development including production of sexual stages has been challenging in monolayers [32]. In recent years, various research groups have attempted to establish advanced in vitro culture models especially for C. parvum, summarized for better comparison in Table 2. The table provides at a glance the methods used as well as various advantages and disadvantages of the in vitro systems. As evident from the approaches described in the table, in spite of the large variety of in vitro models for long-term cultivation of C. parvum established so far, there is a lack of consensus among researchers regarding culture methods, as well as choice of media, and host cell lines used.

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Table 2. Recent advances in the field of in vitro culture systems for Cryptosporidium parvum.

https://doi.org/10.1371/journal.pntd.0009668.t002

Prior attempts at in vitro cultivation of C. parvum utilizing monolayers and axenic cultures have been marked by low counts of infective oocysts and incomplete life cycle with only asexual development reported [32]. The in vitro models highlighted in Table 2 offer high oocysts yields, long-term maintenance of culture and production of all parasitic stages. However, some of these models have innate disadvantages, such as the need for specialized equipment and media components as well as extensive knowhow and expertise required in implementation [33,34]. In addition, some of the in vitro models require mouse intestinal tissues [35–37] or donor biopsies [38] as basis for the model. Apart from ensuring viability of parasites for infection studies, robustness, and scalability of cultures, these advanced models are stepping-stones between cell monolayers and in vivo studies. However, there is ongoing contention in the scientific community as to whether these in vitro systems truly support uninterrupted maintenance of the life cycle due to the difficulty of direct microscopic visualization of the parasites within the various models and the fact that these results are yet to be corroborated by other research laboratories [29]. New combinations of existing technologies employing select features of current models such as simplified protocols, lower maintenance costs in terms of equipment and media, and inclusion of host immunological aspects are imperative for mass production of parasitic stages and continued infection of subsequent cultures, thereby decreasing the use of animal models in the future.

Toxoplasma gondii.

T. gondii, the causative agent of toxoplasmosis, has a worldwide distribution with most infections being mild or asymptomatic [28]. However, exposure to the parasite via consumption of raw meat and contact with contaminated soil or cat feces during pregnancy leads to a range of severe complications for the unborn child [30]. Felids are the definitive hosts for T. gondii; however, the parasite has the ability to infect all warm-blooded animals including humans [31]. The sexual cycle of T. gondii involving the production of micro- and macrogametocytes occurs exclusively in the intestinal epithelial cells of felids, while the asexual cycle involving repeated cycles of endodyogeny can take place in all mammalian cells of the indefinitive hosts such as humans [28].

In vitro cell culture methods are well established for T. gondii, with various studies published using several distinct cell lines most commonly using tachyzoite stages for infection [42–44]. Alternatively, other T. gondii stages such as oocyst-derived sporozoites or tissue cyst–derived bradyzoites may be used in experimental infection models [45]. The following features make T. gondii an ideal protozoan for infection studies: existence of several transmission stages, ease of cultivation in most vertebrate cells, availability of established animal models, and feasibility of genetic crosses. However, further insights into pathogenesis have been hindered by the absence of models simulating the architectural complexity of the host intestinal epithelium [46].

A recent study revolutionized the field of T. gondii research by successfully carrying out the sexual cycle in vitro using mouse intestinal organoids via bradyzoite infection [47]. This was achieved by making use of the fact that members of the feline family lack the enzyme delta-6-desaturase responsible for breaking down linoleic acid. Inhibiting the activity of this enzyme along with dietary supplementation of linoleic acid was conducive to the completion of the sexual cycle of T. gondii parasites in mice.

A recent study reported for the first time coinfection with T. gondii type 1 RH strain tachyzoites and Giardia duodenalis WB6 (ATCC 50803) trophozoites in murine organoid-derived monolayers [46]. By simplifying culture conditions, this study outlined protocols for the establishment and maintenance of 3D intestinal organoids and organoid-derived monolayers for 4 animal hosts with immediate relevance to the study of host–parasite interactions for gastrointestinal parasites.

Previously, Derricott and colleagues [48] developed 3D intestinal organoids using porcine and bovine jejunum as seen in Fig 1. The porcine and bovine organoids incubated with 1 × 10⁶ T. gondii RH strain tachyzoites for 1 hour, then cultured in Matrigel for 24 hours showed distinct foci of infection, thus proving their susceptibility to T. gondii infection. The use of such intestinal samples offers a viable alternative and limits the need for animal experiments. The investigators showed that the intestinal crypts thus isolated and cultured into organoids in Matrigel, could be cryopreserved and subsequently resuscitated with 75% to 80% viability. This model offers a feasible long-term solution for obtaining renewable culture material independent of the use of animal models.

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Fig 1. Schematic illustration of organoids using parasite surface antigen SAG1.

Schematic illustration showing immunofluorescent staining of bovine (a–c) and porcine (d–f) intestinal organoids using parasite surface antigen SAG1 (Alexa Fluor 488—green), F-actin (rhodamine—red), and nuclei (DAPI—blue). Organoids were incubated with 10⁶ T. gondii RH strain tachyzoites for 1 hour at 37°C, embedded in Matrigel and incubated for a further 24 hours at 37°C. SAG1 in green shows intraluminal parasites (a, d, a’, and d’), parasite rosettes indicating replication (b, e, b’, and e’), and white arrows point to intracellular parasites (c, c’, c”, f, f’, and f”). Scale bars represent 20 μm in panels a, b, d, and e and 30 μm in panels c and f. 400x magnification. Open-access image reused from [48].

https://doi.org/10.1371/journal.pntd.0009668.g001

Plasmodium spp.

A considerable amount of interest and funding has been devoted to studying malaria and its causative agent Plasmodium spp., enabling new breakthroughs toward management of the leading cause of mortality due to a parasite worldwide [30]. Malaria, transmitted by species of Anopheles mosquitoes, is characterized by relapse of the disease where the dormant stages known as hypnozoites become reactivated in liver tissues [28]. Sporozoites introduced by the mosquito into the host bloodstream undergo exoerythrocytic schizogony in liver cells, giving rise to mature schizonts that in turn release merozoites. These merozoites then undergo erythrocytic schizogony producing schizonts, which, upon maturation, release merozoites and continue further asexual multiplication. After ingestion by the mosquito vector via a blood meal, merozoites develop into male and female gametocytes and continue the sexual cycle in the insect midgut [31]. Previously, large-scale production of erythrocytic stages was achieved by 2 methods both using RPMI 1640 medium with 25mM HEPES buffer: continuous flow and petri dish–candle jar systems. Early cultivation experiments of exoerythrocytic stages utilized either cultures of infected rat liver cells or infection of embryonic fibroblasts with sporozoites [49].

A recent groundbreaking study provided unequivocal proof of the reactivation of hypnozoites in vitro by utilizing a transgenic Plasmodium cynomolgi dual fluorescent reporter line; for this purpose, the construct pCyCEN_Lisp2mCherry_hsp70_GFP (Addgene, ID 137169) was created [50]. Dormant hypnozoites were green-fluorescent protein positive and mCherry negative, while the developing forms were positive for both fluorescent reporters. Recent studies have reported parasite enrichment and short-term culture of Plasmodium vivax; however, the lack of a robust long-term culture system has hindered progress in areas such as transcriptomics for these parasites [51].

Chua and colleagues [52] published a 3D hepatic spheroid model using Cellulosponge (Invitrocue, Singapore) to study P. cynomolgi. The 3D model was successful in producing liver stages within the hepatocytes embedded in the sponge (Fig 2). This model is the first successful platform closely resembling P. cynomolgi in in vivo studies. Using this hepatic spheroid system, the complete liver stage life cycle including sporozoite invasion and merozoite release leading to new infection was achieved. The current model using P. cynomolgi parasites could also be employed to study life cycle and mode of infection in other Plasmodium species.

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Fig 2. In vitro hepatic spheroid formation.

Schematic diagram showing hepatic spheroid formation in the 3D Cellulosponge in vitro system for P. cynomolgi parasites. (A) Spheroids of uninfected hepatocytes. (B) Spheroids of hepatocytes pre-infected using sporozoites. (C) Successful reinfection and completion of life cycle in erythrocytes. (D) Potential models for drug evaluation in liver stages are shown. Open-access image reused from [52]. RBC, red blood cell.

https://doi.org/10.1371/journal.pntd.0009668.g002

Trypanosoma cruzi.

Trypanosoma cruzi is the causative organism of Chagas disease, also known as American trypanosomiasis. Originally confined to South America, this crippling disease is transmitted by direct contamination of a bite wound with feces or urine of the triatomine bug (Triatoma spp.) or via the alimentary tract by consumption of contaminated food [30]. In humans, T. cruzi exists as rounded amastigotes in muscles and nerve cells and as trypomastigotes in peripheral blood. Amastigotes undergo binary fission and transform into promastigotes, followed by epimastigote forms and are then released into the blood stream as trypomastigotes [28]. Increased international travel has led to the spread of the disease outside its endemic area [53]. Without proper treatment, the chronic phase of the disease can last a lifetime. According to WHO, it is not feasible to eradicate T. cruzi due to the extensive reservoir of the parasites in the wild animal population in endemic regions. In the future, climate change may play an increasing role in the spread of the vector to other parts of the globe.

There have been several publications on 2D in vitro cultures using monolayers for T. cruzi where both trypomastigote and amastigote stages were easily cultivated in Eagle’s Minimum Essential Medium (MEM) with 2% fetal bovine serum; however, separation of the stages to obtain pure cultures was challenging [49]. A 3D culture using cardiomyocyte spheroids was detailed by Garzoni and colleagues to study cardiomyopathy caused by T. cruzi [54]. Cardiac cells obtained from mouse embryos were plated in agarose coated 96-well plates, enabling the formation of 3D cardiac microtissues. Following infection by T. cruzi, an increase in both area and volume of the infected cardiac microtissues was observed in the spheroids.

Another recent publication sheds light on the transmigration of T. cruzi trypomastigotes by employing a 3D spheroid model mimicking infection conditions in situ [55]. Apart from the added aspects of cell-to-cell interactions and the presence of an extracellular matrix (ECM), which are both missing in monolayer cultures, a unique feature of this system is the direct visualization of host and parasite cells by differential labeling with 2 fluorescent dyes. The 3D HeLa cell spheroids were labeled with the red fluorescence protein (RFP) and the trypanomastigotes were labeled with carboxyfluorescein succinimidyl ester (CFSE) to visualize transmigration of trypomastigotes within the spheroids using confocal microscopy. A follow-up study by Rodriguez and colleagues [56] published in 2020 showed that T. cruzi trypomastigotes were less infective in HeLaR2 spheroids than in 2D monolayers and concluded that 3D spheroids provide an optimal physiological environment for studying host–parasite interactions.

Trypanosoma brucei.

Trypanosoma brucei, the causative organism of African trypanosomiasis or sleeping sickness in humans, is transmitted by the insect vector Glossina morsitans (tsetse fly) [30]. The complex life cycle between tsetse flies and their mammalian hosts involves various developmental stages. The blood stream forms known as trypomastigotes in vertebrate hosts are polymorphic, existing as long and slender, short, and stumpy as well as intermediate forms [28]. Metacyclic trypomastigotes injected into human host during a blood meal transform into bloodstream forms. In the vector midgut, bloodstream forms transform into procyclic trypomastigotes, which divide to form epimastigotes, which enter the salivary glands and replicate by binary fission. Previous attempts at retaining infectivity of the long and slender bloodstream forms to mammalian hosts in in vitro cultures were marred by their rapid transformation into noninfective short and stumpy forms resembling those seen in the vector. The long-term maintenance of the bloodstream forms has been a challenge for in vitro culture studies, which was overcome by cultivation of bloodstream trypomastigotes using mammalian feeder layer system in RPMI 1640 medium with 20% heat-inactivated fetal bovine serum and complete life cycle development of T. brucei has been achieved [49].

Among the first reports of in vitro cultivation of T. brucei testing various media, the importance of pH toward the multiplication of T. brucei was stressed. It was concluded that the best results were obtained using MEM 199 buffered with HEPES due to optimal pH conditions (7.4) provided by this medium [57].

Another early publication used bovine fibroblast-like cells in HEPES-buffered Roswell Park Memorial Institute (RPMI)-1640 medium with 20% fetal bovine serum at 37°C where the parasites were able to be maintained for more than 220 days [58]. Moreover, 18 hours after initiation of the primary bovine cell culture, the cell suspension was transferred to a new T-75 culture flask, and fresh medium was added. Sub-culturing was carried out 14 days later when fibroblasts predominated the original culture. The authors recommended to vary the time point of sub-culturing depending on the morphological stages needed as change in medium after 3 days did not lead to the transformation of the slender to stumpy trypanosomes. The parasites obtained in this study were shown to be infective to mammalian hosts.

Other studies also using RPMI-1640 on a bovine embryonic spleen (BESP) feeder layer propagated infective stages of the parasites that were shown to be infective in rats and tsetse flies [59,60]. A novel in vitro system for the cultivation of bloodstream forms of T. brucei consisted of a feeder layer of fibroblast-like cells derived from embryos of New Zealand white rabbits or the mountain vole Microtus montanus, with HEPES-buffered MEM supplemented with Earle’s salts and 15% inactivated rabbit serum [61]. The cultivated bloodstream forms thus obtained were infective to mice and were cyclically transmitted through tsetse flies.

Another approach involved cultivation of T. brucei in Cunningham’s liquid medium in the presence of head-salivary gland, alimentary tract, and abdominal body wall explants of tsetse flies for up to 66 days. It was shown that after 8 days of cultivation, some of the procyclic forms transformed into metacyclic stages, which were infective to mice [62].

In order to overcome the issue of growth inhibition due to the release of inhibitory metabolites released during culturing of T. brucei and to avoid constant medium exchanges, Ajoko and Steverding [63] developed a novel method for culturing T. brucei using transwells placed in standard tissue culture plate wells. Using this approach, a 4-fold increase in the yield of bloodstream forms of the parasites (up to 2 × 10⁷/ml) and an extension of the exponential phase by 1 day compared to previous studies [64] were achieved.

Leishmania spp.

Leishmaniasis, transmitted by infected female sandflies (Phlebotomus papatasi), refers to the disease caused by Leishmania spp. According to WHO, leishmaniasis is caused by over 20 species of Leishmania and is transmitted by over 90 species of sandflies. It is a severely debilitating neglected tropical disease with outcomes ranging from cutaneous lesions, facial disfigurement to death [30]. The life cycle involves 2 forms: spherical amastigotes found in the mammalian host and motile promastigotes occurring in phlebotomine sandflies, both forms multiply by repeated binary fission [28].

Previous studies reported the use of insect and mammalian cell culture media for large-scale production of promastigotes; however, these techniques were unsuitable for obtaining amastigotes [49]. Several studies starting from the 1980s to the turn of the century cultured Leishmania promastigotes in an axenic setting using various media, temperature, and pH conditions [65–68]. Although the promastigotes produced using cell free media were similar to in vivo–derived parasites, there was a shift in research efforts toward using cell lines for culturing Leishmania spp. due to loss of virulence of the promastigotes in long-term axenic in vitro cultures.

In spite of the recent advances in cell culture techniques, progress in the field of in vitro culture of Leishmania parasites has been slow. Petropolis and colleagues [69] developed a 3D model for studying Leishmania amazonensis promastigote migration within a Collagen I (COL I) matrix mimicking the dermal ECM in the host. This study provided valuable insights into the mode of migration within the 3D matrix not feasible in previous experimental cultures. A recent study compared for the first time the 3 most widely used media for cultivation of Leishmania infantum promastigotes: Grace’s insect cell culture medium, liver infusion tryptose, and Schneider’s insect medium, without supplementation or supplemented with fetal calf serum and bovine serum albumin [70]. This study concluded that liver infusion tryptose supplemented with fetal calf serum was best suited for growth and maintenance of L. infantum parasites.

Giardia duodenalis.

Giardiasis caused by G. duodenalis, also known as Giardia lamblia or Giardia intestinalis, is one of the most widespread gastrointestinal ailments worldwide [30]. Despite its global occurrence and severe long-term health consequences especially in children, giardiasis is a neglected tropical disease. The parasitic life cycle includes 2 forms: trophozoites and cysts. After ingestion, cysts hatch into motile trophozoites, which multiply by binary fission and colonize the human intestine [28]. A fundamental challenge for in vitro cultivation of G. duodenalis used to be the infeasibility of co-culturing parasitic trophozoites and intestinal epithelial cells. Hence, information on host–parasite interaction has been lacking. Most publications have axenically cultured trophozoites either in TYI-S-33 medium [71] or BI-S-33 medium [72] supplemented with bile.

Fisher and colleagues [73] developed a coculture model of trophozoites and Caco-2 cells mimicking the apical–basolateral direction of the small intestinal epithelium. This model using transwell inserts was able to maintain the cocultures for 21 days, thereby providing a viable system for long-term culture of G. duodenalis in the presence of host cells. This coculture model along with other in vitro cultures of G. duodenalis published so far would benefit greatly from standardization in order to implement these models across several laboratories [74]. The first report of a 3D coinfection model of G. duodenalis with T. gondii in murine organoid–derived monolayers was published recently by Holthaus and colleagues [46]. In order to ensure optimal survival of G. duodenalis trophozoites while mimicking duodenal conditions, the monolayers were incubated in a modified TYI-S-33 medium rich in cysteine with added bile salts, instead of DMEM-based media. This medium supported the growth of the parasites for over 72 hours in culture.

Entamoeba histolytica.

Amebic dysentery, also known as amebiasis, caused by the parasitic amoeba Entamoeba histolytica has a worldwide distribution [75]. After malaria, amebiasis is the second leading cause of mortality worldwide attributed to parasitic diseases [76]. The life cycle includes 2 stages: the motile trophozoites and the nonmotile, infective cysts [28]. Transmission occurs alimentarily via consumption of food and water contaminated by fecal material containing mature E. histolytica cysts.

Optimal growth conditions in vitro for E. histolytica need to mimic the anaerobic environment of the intestine. Toward this goal, Shah and colleagues [76] compared 2 cultivation methods namely: Anaerocult A (Merck, Darmstadt, Germany) and mineral oil blocking methods of in vitro cultivation of E. histolytica. The investigators cultivated strains of E. histolytica under axenic conditions by each of the 2 methods mentioned above. Minimum inhibitory concentration (MIC) of metronidazole activity against the parasites used to measure the efficacy of each culture method showed that both methods were equally reliable for the propagation of E. histolytica.

A 3D infection model for amebiasis consisting of cocultures of liver sinusoidal endothelial cells (LSECs) and hepatocytes sandwiched within a collagen-I matrix has been reported (Fig 3) [77]. Although the purpose of this model was to simulate amebic infection of the liver, it could be used as a starting point for further in vitro cultivation studies of this protozoan parasite.

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Fig 3. Illustration showing variations of a 3D liver infection model of E. histolytica infection in vitro.

Hepatic cells (red), amoebae (green), and COL-I (collagen-I, blue) are shown. (A) Virulent trophozoites added to the medium adhere to the LSEC in the 3D liver model but not in the setup without LSECs (No LSEC) seen in transversal view of reconstructed 3D images. (B) Incubation of liver model with trophozoites for 1.5 and 3 hours. (C) Incubation of the distinct setups with trophozoites for 3 hours. (B–E) Percentage of amoeba crossing the LSEC layer or COL-I matrix. Infection of the liver model (D) with various E. histolytica strains in the presence of glucose/galactose for 1.5 hours and similarly, infection of the setup without LSEC (F) for 3 hours. (F) Parasite invasion rate at 3 hours for the model and the no LSEC setup showing 0% invasion at the LSEC and 100% at the hepatocyte layer z-position, respectively. Red bars indicate mean values. (G–I) Parasite interaction with the LSEC layer of the 3D liver model at 3 hours. In panels (G) and (H), the arrows point to LSEC detaching in the presence of trophozoites. In panel (I), the asterisks point out vesicular structures localized inside the trophozoites labeled with hepatic cell staining. Open-access image reused from [77]. LSEC, liver sinusoidal endothelial cell.

https://doi.org/10.1371/journal.pntd.0009668.g003

Naegleria fowleri.

N. fowleri is the only species of the genus Naegleria (free-living amoeboflagellates) known to infect humans. It is a thermophilic amoeboflagellate commonly found in warm fresh water bodies and soils worldwide [28]. The life cycle of N. fowleri includes trophozoites, motile flagellate forms, and cysts. Infection via trophozoites occurs nasally during recreational activities in infected waters. Within 3 to 7 days after entering through the nose, the parasite crosses the blood–brain barrier leading to primary amebic meningoencephalitis (PAM) affecting the central nervous system [78]. The fatality rate for N. fowleri infections is over 97%, with only 4 survivors out of 145 known cases in the United States during the period 1962 to 2018 [79].

In one of the earliest publications on in vitro culture of N. fowleri, Wong and colleagues reported the maintenance of axenic cultures in Nelson’s medium and monkey kidney cell monolayers [80]. However, they observed a loss of virulence of N. fowleri trophozoites after 50 passages conducted weekly in Nelson’s culture medium (ATCC Medium 710). Gianinazzi and colleagues [81] described an in vitro organotypic slice culture system using rat brain tissues. In comparison to low-dose inoculum, the aforementioned N. fowleri infection model showed the multiplication of N. fowleri trophozoites as well as extensive tissue damage in the case of high-dose inoculum (10-fold). However, as pointed out by Coronado-Velazquez and colleagues [82], this model like all other in vitro models has the intrinsic disadvantage when compared to an in vivo model that the physiological conditions in the host cannot be effectively simulated yet. Another disadvantage is the lack of a representative model of a blood–brain barrier.

Burri and colleagues [83] reported in vitro proliferation and cytotoxicity of the trophozoites in various media including Nelson’s medium, PYNFH medium, and PYNFH medium (ATCC Medium 1034) supplemented with liver hydrolysate (LH) in RPMI medium in 24-well plates. To estimate cytotoxicity of N. fowleri, trophozoites were cocultured directly with the murine fibroblast cell line L929 (ATCC 2148) in a contact-dependent assay, whereas for the contact-independent assay, the trophozoites were added to cell culture inserts (Vitaris, Switzerland). Using the Cytotoxicity Detection kit ^PLUS (LDH) (Roche Pharma, Germany) to calculate cytotoxicity, the study concluded that although the addition of LH increased trophozoite proliferation in vitro, there was no correlation of cytotoxicity with the disease potential of the parasites.

Zaongo and colleagues [84] optimized growth conditions for N. fowleri in vitro by testing several media under varying temperatures and recommended Nelson’s medium supplemented with 1% peptone (N + pep) and modified PYNFH medium without yeast extract to be most suitable. These results were in keeping with previous reports of the affinity of N. fowleri toward higher temperatures for optimal growth between 30°C and 40°C. With rising global temperatures, the thermophilic nature of this free-living amoeboflagellate is an important consideration when it comes to the effect of climate change in proliferation of this life-threatening organism.