Approaches to Protozoan Drug Discovery ... - ACS Publications

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Approaches to Protozoan Drug Discovery: Phenotypic Screening Miniperspectives Series on Phenotypic Screening for Antiinfective Targets Melissa L. Sykes and Vicky M. Avery* Discovery Biology, Eskitis Institute for Drug Discovery, Griffith University, Nathan, Queensland, Australia ABSTRACT: Determining the activity of a compound and the potential impact on a diseased state is frequently undertaken using phenotypic or target-based approaches. Phenotypic screens have the advantage of the whole organism being exposed to the compound and thus all the targets and biological pathways associated with it. Cell penetration and access to targets in their “natural” environment are taken into account. Unless utilizing a genetically modified organism with an additional target associated indicator, elucidation of specific target(s) of active compounds is necessary. Target discovery is desirable to allow development of chemical entities based upon knowledge of the target structure. Phenotypic drug discovery has successfully identified new molecular entities for neglected protozoan disease research. In this perspective, the phenotypic approaches used to identify chemical entities for drug discovery and for use as tools against the parasites Plasmodium falciparum, Trypanosoma brucei brucei, and Trypanosoma cruzi will be outlined.

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INTRODUCTION The majority of new molecular entities approved by the FDA between 1999 and 2008 were identified by phenotypic screening (37% versus 23% discovered by target- based approaches).1 However, as target-based approaches outnumbered phenotypic screening during this time frame, the success of phenotypic screening is underrated. Phenotypic screening has the advantage of identifying new targets, unlike biochemical screens, which rely on known therapeutic pathways. Also, compounds may affect multiple proteins or pathways in the organism which would not be identified in a biochemical screen. Lack of membrane permeability can lead to inactivity being reported for a compound with demonstrated target activity when assessed in a phenotypic screen. This can be a double-edged sword, as a target specific molecule with inactivity in a phenotypic screen would be lost. However, given the costs and frequent failure to suitably optimize and progress molecules, this loss is usually considered acceptable. Phenotypic (whole cell) screening against protozoan parasites has resulted in the identification of a number of chemical entities that are currently in preclinical and clinical development. The phenotypic approaches used in drug discovery for Plasmodium falciparum, Trypanosoma cruzi, and Trypanosoma brucei brucei are outlined, and some considerations of these in vitro applications are discussed.

headache, sweating, nausea, and muscle and joint pain. In severe cases, malaria progresses to seizures, kidney failure, coma, and can be fatal. P. falciparum malaria is a major global health problem, predominantly affecting children under the age of five in endemic countries, claiming >1 million lives and resulting in some 200 million clinical cases every year.2 The life cycle of Plasmodium is reliant on a mosquito vector and mammalian host. During feeding on the human host, the Anopheles mosquito transmits the parasite to the host in the form of sporozoites, which migrate to the hepatic cells of the liver. Following extensive multiplication, merozoites are released which invade the circulating erythrocytes. The clinical manifestations of malaria are due to the asexual replication of the P. falciparum parasite in the human host. One complete asexual life cycle (Figure 1) takes approximately 48 h from the initial invasion of an erythrocyte, progression through the ring, trophozoite, and schizont stages, and subsequent invasion with new progeny. Transmission requires the sexual form of the parasite, namely the gametocytes, which are present as a minor population in the bloodstream of the human host and representing the only malaria parasite forms transmitted to, and capable of surviving in, the Anopheles mosquito. Gametocytes initiate sexual reproduction in the mosquito, resulting in a multistage development process called sporogony, ending in the formation of sporozoites, the stage transmitted back to the human host during a mosquito bite (Figure 1).

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MALARIA: PLASMODIUM The malaria parasite, Plasmodium, is a unicellular protozoan belonging to the phylum Apicomplexa. Five species of Plasmodium infect humans, namely P. falciparum, P. vivax, P. ovale, P. malariae, and P. knowlesi. Symptoms of malaria, which develop 9−14 days after being bitten by an infected mosquito, include many flu-like symptoms such as sudden fever, chills, © XXXX American Chemical Society

Special Issue: Miniperspectives Series on Phenotypic Screening for Antiinfective Targets Received: March 25, 2013

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dx.doi.org/10.1021/jm4004279 | J. Med. Chem. XXXX, XXX, XXX−XXX

Journal of Medicinal Chemistry

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erythrocytes from noninfected, which is readily identifiable through the use of mathematical algorithms. The significant advantages of the image output include the exclusion of all nonparasite related fluorescence such as fluorescent compounds, potentially reducing false negative rates, and the ability to reassess the data with alternative algorithms.11−13 Phenotypic HTS campaigns aimed at identifying potential new chemical starting points for antimalarial agents,10,12,14,15 directed at the asexual blood stages of P. falciparum, have generated a wealth of diverse chemical entities with potential for further drug development. The public availability of these molecules provides the malaria community at large with valuable tools not only for drug discovery but for furthering our understanding of malaria biology. The most well-known of these include the Genomics Institute of the Novartis Research Foundation (GNF) Malaria box,15 the GlaxoSmithKline (GSK) TCAMS library,14 and the collective known as the MMV Malaria Box, comprising representative compounds from the aforementioned libraries plus the St. Jude Children’s Research Hospital actives12 and others identified through alternative partners. During the past five years, we have performed numerous screening campaigns utilizing our image-based assay11 against various libraries of differing sizes and comprising both natural products and small molecules, and as a result have been screened at different concentrations with vastly differing hit rates (Table 1). These libraries have been sourced from both industry and academia and have included both diverse and focused libraries, with the total number of data points screened

Figure 1. Life cycle of the malaria parasite. From Epidemiology of Infectious Diseases: available at: http://ocw.jhsph.edu. Copyright Johns Hopkins Bloomberg School of Public Health. Creative Commons BY-NC-SA.

The focus of the Avery laboratory is on the asexual and sexual forms of P. falciparum found within the human host. Hence the life cycle stages within the mosquito vector will not be discussed in the context of this article. Current antimalarial therapy is primarily based on artemisinin combination therapies (ACT).3 The limited chemical diversity of aminoquinolines, antifolates, hydroxynapthoquinones, and artemisinin derivatives means resistance is an ever-present threat, thus there is a need to identify new/novel chemotypes which act upon new druggable targets. The primary objective of our laboratory is to address this issue through the identification of new selective chemical starting points which are efficacious against the malaria parasite, P. falciparum. Plasmodium falciparum Asexual HTS and Drug Discovery. Screening of compound libraries using in vitro assays in the search of new antimalarial moieties against the asexual blood stages of Plasmodium has been undertaken by numerous research laboratories over many years. The initial assay of choice was a radiometric assay measuring the incorporation of [3H]-hypoxanthine by the parasite,4 however, given the need for radio-isotopes, additional handling for filtration and cell harvesting, plus the 96-well format alternative, more efficient and cost-effective methods have emerged. Alternative approaches include flow cytometry (YOYO-1 dye),5 and fluorescence-based assays utilizing stains such as Hoescht, PicoGreen,6 SYBR green,7 and 4′,6-diamidino-2phenylindole (DAPI),8 which incorporate into the DNA. Recently, new high throughput screening (HTS)-ready luminescence approaches9 based on the measurement of the firefly luciferase reporter gene activity have also been reported. Not all of these assay formats are suitable for HTS due to a myriad of factors such as cost, assay stability, robustness, equipment availability, and data quality. The assays based on fluorescent DNA dyes are readily adaptable to high throughput,8,10 and with the advent of high throughput image-based screening technologies,8,11,12 this has led to the development of more data-rich and informative image-based assays. We have developed one such image-based assay for performing asexual HTS which utilizes the incorporation of a DNA stain to distinguish parasite infected

Table 1. Example of the Libraries Screened against the Asexual Blood Stage P. falciparuma library

NPE/NPC/SC compound no.

1 2 3 4 5 6 7

NPE: 140,000 SC: 34,654 SC: 5,279 SC: 3,215 SC: 42,880 SC: 150,352 SC: 1,996

8

SC: 3,030

9

SC: 176

10

NPC: 141

11 12 13 14 15 16 17 18 19 20

SC: 10,560 SC: 3,520 SC: 27,035 SC: 20,000 SC: 502,868 SC: 256,263 SC: 1,239 SC: 49,501 SC: 117,120 NPE: 203,000

HTS concn 0.7 μg/mL 1.82 μM 1.83 μM 11 pt CRC 1.96 μM 3.17 μM 0.784 μM 7.84 μM 0.784 μM 7.84 μM 0.784 μM 7.84 μM 0.25 μM 1.28 μM 6.4 μM 1.8 μg/mL 0.4 μg/mL 3.4 μM 1.9 μM 1.6 μM 1.92 μM 8 pt CRC 1.8 μM 1.75 μM 0.5 μge/μL

HTS actives (confirmed)

% hit rate

915 380 671 140 303 4181 321

0.65 1.09 12.70 4.30 0.70 2.78 16

154

5.00

36

20.00

99 < 1 μM

70.00

407 89 908 448 9443 1985 50 (20 < 1 μM) 508 925 2276

3.85 2.50 3.35 2.24 1.88 0.77 4.00 1.02 0.79 1.12

a

Library type, concentration provided, and size determined the screening concentrations used and reflected in the hit rates (confirmed) obtained. NPE: natural product extract or fractions; NPC: natural product compound; SC: synthetic compound.

B



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Table 2. Activities (IC50 Values) of Antimalarial Drugs and Known Inhibitors against Both the Early (I−III) and Late (IV−V) Gametocytes, Utilizing an Image-Based Assaya early (I−III) gametocyte

a

late (IV−V) gametocyte

asexual (3D7)

compound/drug

IC50 (nM) mean ± SD

n

IC50 (nM) mean ± SD

n

chloroquine pyronaridine primaquine dihydroartemisinin puromycin amodiaquine pyrimethamine mefloquine epoxymicin tafenoquine artesunate

120 ± 4.8 131.3 ± 17.3 5860 ± 193 1.80 ± 0.5 137 ± 21.0 159.0 ± 13.7 ∼40% inhibition at 120 μM 220 ± 100 ND ND ND

6 4 2 3 6 5 2 3

∼100% inhibition at 100 μM 1300 ± 900 ∼40% inhibition at 120 μM ND 46.0 ± 14.1 4490 ± 1850 ∼100% inhibition at 120 μM ND 0.7 ± 0.3 3366 ± 1290 3.43 ± 3.58

15 13 2 20 8 9 18 19 7

IC50 (nM) mean ± SD 31.0 46.0 6090 0.7 64 28.0 14.0 200 2.2 1070 2.20

± ± ± ± ± ± ± ± ± ± ±

6.4 19.0 375 1.0 16.3 0.8 2.8 110 0.9 482 0.9

Asexual activity included was determined using an image-based assay, all asexual data represented determined from n ≥ 5.11 ND = not determined.

have all been demonstrated to have gametocytocidal activity in vitro.21 Crucial for more focused screening of molecules against the gametocyte stages to identify new chemotypes with gametocidal activity, is the ability to reproducibly culture them in large scale and access to robust, biologically relevant assays. In Vitro Plasmodium falciparum Gametocyte Culture. Environmental changes, host factors, and various chemicals leading to parasite stress have consistently been shown to enhance gametocyte production in vitro. Ultimately, it is likely that this depends on a complex interplay of multiple environmental factors, some of which are difficult to control in the laboratory, hence the challenge of large scale culture of gametocytes. Consequently, this has had a direct impact on the development of gametocytocidal HTS assays. A comprehensive review of gametocyte whole cell screening has recently been published by Lucantoni and Avery (2012),21 which discusses in detail the essential requirements for gametocyte culture and available assay formats. Gametocytocidal Assays and Drug Discovery. Several reports of new techniques for screening compound libraries against gametocytes of different strains of P. falciparum have recently been published. These range from flow cytometry,22 ATP bioluminescence,23 fluorescence,24 and metabolic activity using an oxido-reduction indicator, Alamar Blue.25 Luminescence (luciferase reporter) and image-based HTS assays for screening both early (I−III) and late (IV−V) stage gametocytes developed in our laboratory have recently been used to screen the publicly accessible compound libraries comprising asexual blood stage actives, including the GSK TCAMS collection, GNF and MMV Malaria Boxes, and a considerable number of well characterized preclinical candidates and academic molecules. Exemplars of the actives identified from these libraries21 indicate that a large number of molecules are active across both the asexual and sexual stages, making them interesting candidates for either drug discovery programs or as tools to investigate the targets further. The most recent highlight being the quinolone-3-diarylethers, which are being progressed under the direction of Professor Mike Riscoe, OSHU, Portland Oregon, USA.26 Screening of the GNF malaria box (2.6 μM) identified 719 confirmed early (I−III) stage hits (19%) and 233 late (IV−V) stage confirmed hits (6%). Given that these molecules are all asexual blood stage actives, the number of early (I−III) stage gametocyte hits was lower than expected. Recent screening of a further 15,000 compounds (1.5 μM)

exceeding 3 million. This in itself is a major achievement, and a turning point in malaria drug discovery, as until this time it was unusual for large pharmaceutical companies to make their chemical libraries available to an academic institution. In total, more than 20 corporate libraries have now been screened within the Avery lab at the Eskitis Institute for Drug Discovery, Griffith University. The outcomes of HTS against the asexual blood stages of P. falciparum undertaken in our lab have resulted in a number of new chemical starting points which are currently being exploited in drug discovery programs throughout the world, in addition to those which are being used as tools to evaluate potential new targets. Some of the more recent highlights include molecules which have progressed significantly such as the 3,5-diaryl-2-aminopyridines (University of Capetown, South Africa),16 and the pyrroloiminoquinones (Eskitis Institute for Drug Discovery, Griffith University, Australia).17 The current global screening effort has afforded the malaria community a vast number of interesting and diverse chemical leads, and molecular tools, which are effective against the asexual blood stages of the parasite. However, a remaining key concern is the inability to prevent transmission of the parasite to the vector host, thus perpetuating the parasite life cycle. Transmission is reliant upon the involvement of the gametocyte, or sexual stages in the human, thus we have focused our most recent efforts in malaria drug discovery on this stage. Plasmodium falciparum Gametocyte (Sexual) HTS and Drug Discovery. Unlike the asexual blood stages, the development of P. falciparum gametocytes takes 10 or more days, involving five morphologically distinct differentiation stages. The immature stages (I−IV) are sequestered from the circulation into the bone marrow and spleen.18 Mature stage V gametocytes are released back into the bloodstream where, as circulating gametocytes, they provide the essential reservoir for reinfection of mosquitoes.19 Current therapeutics designed to be effective against the asexual blood stages have limited activity on gametocytes. The artemisinin derivatives, artesunate and artemether, and the 8aminoquinolines, for example primaquine, are the only antimalarials demonstrated to have a clinically therapeutic effect on mature gametocytes in vivo.20 Trioxaquines, 8aminoquinolines, epoxomicin, methylene blue, tipranavir, riboflavin, 9-anilinoacridines, and, more recently, spiroindolones, thiostrepton, and novel 4-aminoquinoline derivatives C



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CHAGAS: TRYPANOSOMA CRUZI Chagas disease (CD), endemic to South America, is caused by the protozoan parasite Trypanosoma cruzi. The disease is estimated to cause 14,000 deaths per year.27 There are two stages of CD, the acute and the chronic phase. The initial stage is usually asymptomatic, or may present as flu-like symptoms. Although the acute phase is not always diagnosed, there is a mortality rate of 2−10%, especially in children.28 The later chronic phase occurs when the cellular response limits parasite proliferation, leading to a sustained inflammatory response, which can ultimately result in digestive damage, with gastrointestinal complications including megaesophagus and megacolon.29 The heart is the most severely affected organ, usually associated with fatal cases of the disease.30 There are two drugs available for the treatment of CD, nifurtimox and benznidazole. Nausea, vomiting, severe weight loss, insomnia, depression, vertigo, convulsions, and disorientation are commonly reported following administration of both drugs and often force patients to stop treatment.31 The use of these drugs to treat the acute phase of the disease is widely accepted, however, the efficacy of these drugs in the chronic phase has not been demonstrated.32 Despite numerous in vitro and in vivo studies, and new targets identified, there have only been a few compounds that have advanced to clinical trials for CD. These were allopurinol, itraconazole, fluconazole, posaconazole, and K777. Although posaconazole is currently in clinical trials for the treatment of CD,33 other antifungals have failed in mouse models, including itraconazole.34 Allopurinol has shown poor efficacy in patients with chronic CD35 and K777, a protease inhibitor of the cysteine protease cruzipain, is currently undergoing preclinical safety and toxicology studies.36 Due to the high failure rates of promising compounds, and the fact that very few are in the current clinical pipeline, introduction of new compounds into the drug discovery process for the treatment of CD is required. These compounds need to display less toxicity, with activity against both the acute and chronic phases of the disease. Culture of the T. cruzi Life Cycle in Vitro. The lack of compounds currently in clinical trials may in part be due to a lack of well established procedures and in vitro screening protocols to evaluate compound activity.37 An important consideration in this process is the in vitro maintenance of the T. cruzi life cycle. During the mammalian life cycle stage of the disease, CD is transmitted by an insect vector, the domestic Triatominae bug. The nonhuman infective epimastigote form of the parasite resides in the mid gut of the bug and differentiates into metacyclic trypomastigotes. These infect mammalian hosts upon a blood meal and amastigotes form in host cells. After a few days, differentiated bloodstream trypomastigotes are released. The in vitro culture of the T. cruzi life cycle has been described in some detail for HeLa cells,38 however, information for the initiation and ongoing subculture of the T. cruzi life cycle in vitro is often with limited descriptions of methodology. There have been reports of differentiation of the noninfective epimastigote form of the parasite into the metacyclic infective form using an artificial triatomine bug urine, TAU3AAG.39 An alternative approach is by spontaneous differentiation of epimastigotes into metacyclic trypomastigotes during the stationary phase of growth, however, this is not controlled.40 Once metacyclic trypomastigotes infect mammalian tissue, they form amastigotes and differentiate into bloodstream trypomastigotes that are released

comprising 5,000 known asexual actives and 10,000 in-actives gave hit rates of 2.7% and 0.9% for the early stage and late stage gametocytes, respectively. The authors are currently preparing manuscripts detailing the active molecules identified from these screening campaigns. Table 2 illustrates the activities of well-known antimalarial drugs against both the early (I−III) and late (IV−V) stage gametocytes in our image-based assays following exposure to compound for 72 h. The data are representative of duplicate or triplicate points performed in dose response (n ≥ 2). Activity obtained with our asexual image-based assay was also included for direct comparison. HTS of late stage gametocytes at a scale feasible for screening hundreds of thousands of compounds has been made possible as a result of our optimized methodology for large scale production of pure synchronized gametocytes. Our current capacity for screening late stage gametocytes is ≥15,000 compounds per campaign, twice per week, with a greater capacity for the early stage gametocyte assays, as survival rates at the earlier stages are significantly greater. By utilizing two distinctly different technologies, i.e., luminescence and imagebased approaches, we have been able to cross validate the outcomes from our current screens, rapidly identifying molecules associated with technology interference. While a number of assay formats for screening gametocytes at both the early and late stages have emerged, to date there have been no reports of these assays being utilized for large scale screening campaigns. This has led us to believe that we are the first laboratory to undertake screening of this nature, having completed screening diverse libraries containing 15,000 and 25,000 compounds, 40,000 of a 100,000 compound library, in addition to the various Malaria Boxes. As highlighted by Lucantoni and Avery (2012),21 comparing the activities of well described drugs and/or new candidate molecules, which have been tested to date by different laboratories, is hindered by a multitude of different factors, such as the methodology used to culture, induce and purify the gametocytes, the assay format used and its inherent limitations, the compound samples themselves, and associated exposure times. With these inconsistencies between laboratories, and limited knowledge of how compound stocks were handled, comparative analysis of data in literature is difficult. This is further compounded by the potential activity differences observed for compounds dependent on their chemical class and the technology used. While highlighted here, this is a recurring theme for all drug discovery and not unique to gametocytes or parasitology per se. Understanding the benefits and limitations of the assay that is used for the identification of compound activity is important for interpretation of the results. Design of other assays to prioritize compounds may aid the decision making process. The use of imaging assays enables less interference from compounds as the technology employs specific cellular markers rather than whole fluorescent or luminescent read outs. Malaria drug discovery has come a long way in the past 10 years, and with the ongoing concerted efforts by not-for-profit organizations, such as Medicines for Malaria Venture (MMV) and the Bill & Melinda Gates Foundation (BMGF) and dedicated researchers, it is likely that we will see significant advances in the coming years, in particular in the area of transmission blocking entities. D



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blood were not able to readily incorporate 3H-thymidine.42 It was concluded that, by incorporation of 3H-thymidine, this suggested that normally nonreplicating trypomastigote forms had been modified with time in vitro and were now able to replicate. However, other data has previously suggested that incorporation of 3H-thymidine does not necessarily correlate with cell division in some cell types.43 As no correlation was observed between 3H-thymidine incorporation and increase in trypomastigote cell number, this conclusion is not definitive. We have found T. cruzi Tulahuen strain parasites maintained in vitro show some characteristic changes with the passage of time. We initiated two cultures of trypomastigotes by differentiation of epimastigotes, induced in TAU3AAG medium. One culture was maintained for 12 months and the other for 2 months by reinfection of 3T3 host cells with trypomastigotes (Figure 2). To infect a monolayer of host cells, trypomastigotes were added on day 0 and following 24 h (day 1) were washed off. By this stage, amastigotes were formed within host cells. On day 3, amastigotes began to differentiate into trypomastigotes, which were released from the host cell. On day 4, trypomastigotes released in to the supernatant were used for subculture. We found that the number of trypomastigotes and amastigotes released into the supernatant in T. cruzi infected 3T3 cultures increased in the 12 month old subcultures in comparison to the 2 month old subcultures (Figure 3). Amastigotes in the supernatant, or “extracellular amastigotes” may be intracellular amastigotes released from the host cell, or trypomastigotes may differentiate into amastigotes in the supernatant. However, this was not determined, and it would be difficult to track the source. On day 3, there was a significantly greater release of trypomastigotes in the older culture (2 × 107 cells versus 4 × 105 cells in the 2 month old culture). By day 4, although there was an increase in trypomastigote numbers in both cultures, the older subculture contained a higher percentage of amastigotes. Amastigotes comprised 3% versus 20% of the culture forms in the supernatant in the 2 and 12 month old cultures, respectively. The 2 month old culture showed 2.5 times less trypomastigotes in the supernatant on day 4. Trypomastigotes, obtained from the supernatant of the 12 month old subculture, were incubated in the absence of host cells for 48 h at a range of cell densities, and no replication of trypomastigotes was observed, as shown in Figure 4A. At all cell densities tested, there was a mixture of extracellular amastigotes and trypomastigotes observed after 48 h incubation (Figure 4B). This suggests that some trypomastigotes had differentiated into amastigotes. When relating this to infected cultures and the increase in amastigotes in the supernatant (Figure 3B), this could mean that in older cultures trypomastigotes more readily

from the host cell. Bloodstream trypomastigotes are the most clinically relevant form for human infection in comparison to the metacyclic form and thus could be considered the most appropriate form for development of assays for early drug discovery for CD. A method for the routine in vitro culture of T. cruzi human infective forms is outlined in Figure 2 for the

Figure 2. An in vitro system for maintaining infective subcultures of T. cruzi Tulahuen strain parasites using 3T3 fibroblast host cells. Trypomastigotes are added to a monolayer of host cells in a 25 cm2 tissue culture flask at a multiplicity of infection of 10:1 parasite:host cell, on day 0. Trypomastigotes invade host cells over 24 h and are then washed off. From 1 to 2 days, amastigotes can be seen developing within the host cell. After 3 days, amastigotes begin to differentiate into trypomastigotes, which are released from the host cell in to the supernatant. On day 4, trypomastigotes are collected from the supernatant and used to infect a subsequent culture of host cells. Images are not to scale for presentation purposes.

Tulahuen strain of the parasite, by infecting 3T3 host cells. To initiate infection, epimastigotes are differentiated in to metacyclic trypomastigotes in TAU3AAG, and used to infect host cells. Once sufficient trypomastigotes are produced from initial infections, a multiplicity of 10:1 trypomastigotes to host cell is used for ongoing routine culture. The age of T. cruzi life cycle forms in vitro is an important consideration in T. cruzi culture systems. The long-term maintenance of virulent EP-strain epimastigotes over several years in vitro was found to affect the ability of induced T. cruzi EP-strain metacyclic forms to infect mice. There was also an associated variation in the antigenic composition of these metacyclic trypomastigotes.41 Trypomastigotes isolated from a longer term culture (>2 months) of the Brazil strain of T. cruzi in murine PSC3H fibroblasts formed amastigotes more readily and also incorporated 3H-thymidine, although freshly isolated trypomastigotes (2−5 weeks old) of this strain from mouse

Figure 3. Enumeration of trypomastigotes and amastigotes released in to the supernatant in (A) a 2 month old subculture and (B) 12 month old subculture of T. cruzi infected 3T3 host cells. E



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Figure 4. (A) Number of host cell free trypomastigotes following incubation of a density of 2 × 105 trypomastigotes in RPMI over 48 h. (B) Percentage of trypomastigotes and amastigotes in culture following incubation of host cell free trypomastigotes in RPMI for 48 h at varying cell densities.

differentiate into amastigotes, as was found by Ashraf and coworkers.42 Although the basis for these observations was not determined, it is clear that with the use of in vitro cultured T. cruzi cells over time there are changes in the morphology of trypomastigotes in host cell free cultures. In addition, there are changes in the subcultures of infected host cells with respect to the number of extracellular amastigotes and trypomastigotes in the supernatant. Consequently, it is likely that there are associated genetic changes occurring with time. In terms of drug discovery, assay formats could be affected, as well as drug sensitivity, by these findings. We found that the use of a 12 month old subculture, compared to a 2 month old subculture, impacted on the sensitivity of an in-house image-based assay which detects amastigote infected 3T3 host cells. In this assay, intracellular amastigotes stained with Hoechst form a rounded spot shape. These are detected as spots in the host cell cytoplasm by an image-based algorithm. The stained nucleus of trypomastigotes differentiated in host cells also appear as round spots, however, they are smaller, closer to one another, overlapping, and tend to obscure the host cell nucleus. Therefore cells containing many trypomastigotes were not as accurately defined as infected. Hence the relationship of infected cells versus the multiplicity of infection (trypomastigotes:host cell ratio used to infect cultures) was effected (Figure 5). All of these factors highlight the need to closely monitor T. cruzi for temporal changes when cultured in vitro. As a result of these studies, our practice is to minimize the use of in vitro cultures of the infective form beyond 2−3 months. Phenotypic Whole Cell Assays in T. cruzi Drug Discovery. It is not clear if both life cycle stages of T. cruzi are equally important during infection, however it is suggested

that during the acute stage of the disease, extracellular and intracellular forms are the main contributors to the disease. During the chronic stage, evidence suggests that persistence of intracellular forms may be a key factor.44 Recently, a meeting “Experimental Models in Drug Screening and Development for Chagas Disease”, incorporating a variety of networks and laboratories, was held for discussion of Chagas in vivo and in vitro experimental models.37 It was agreed at this meeting that an in vitro system, which allows the effects on both amastigotes and trypomastigotes to be monitored, is required. Published assay formats for the detection of compound activity against T. cruzi employ a variety of methods. Some determine the effect of compounds on the amastigote stage, with compound addition after host cell infection, 45 while other methods have incorporated addition of compound before trypomastigote infection of host cells.46 There has also been determination of compound action on both the trypomastigote and amastigote stages in separate assays.47 Inclusion of assays that monitor the activity of compounds against both the trypomastigote and the amastigote stages may lead to more successful identification of active compounds and potentially more of an understanding of the correlation between in vitro and in vivo activity. T. cruzi Phenotypic HTS Campaigns and Outcomes. As highlighted above, the methods for determining compound activity against T. cruzi vary considerably. The technologies used to detect parasites are also different. Fluorescence-based methods include a β-galactosidase reporter gene assay with compound addition made 2 h after addition of infective trypomastigotes48 and an assay estimating the effects of amastigotes exposed to compounds utilizing T-d tomato reporter gene transfected parasites.49 Recent image-based methods using DNA staining techniques detect the effects of compound on intracellular amastigotes.45 Phenotypic HTS has enabled the activity of a large number of compounds to be evaluated against T. cruzi. A β-galactosidase reporter gene-based T. cruzi assay has been used in a 96-well format to screen 303,286 molecules (the NIH collection), and compounds were further profiled in a separate cytotoxicity assay to determine their IC50 values against host NIH-3T3 cells.46 This assay incorporated addition of compound at the same time as trypomastigotes to estimate inhibition of infection, or activity on either trypomastigotes/amastigotes, however, effects on either life cycle stage would not be able to be separated in this assay. Two of these molecules were found to reduce, but not clear, infection in an in vivo model. Further optimization of these compounds would be required due to solubility and permeability issues created by the presence of a quaternary ammonium. In more focused studies, a similar β-galactosiase reporter gene-based assay has been successful in identifying

Figure 5. Percent infection of 3T3 cells by enumeration of host cells containing T. cruzi amastigotes, over a 0−10:1 multiplicity of infection in an image-based assay. Trypomastigotes were added to a monolayer of host cells and washed off after 24 h. Percent infection was determined following a further 48 h of incubation. F



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Figure 6. T. cruzi amastigote images, captured on an Opera QEHS 2.0 at 20× magnification (A−C = infected host cell; DMSO treated for 48 h; D− F = infected host cell, treated with 12 μM nifurtimox in DMSO). (A,D) Hoechst staining of parasite and host cell nuclei (B,E) HCS CellMask Green staining of host cell cytoplasm (C,F) Analysis using Acapella Software.

division, which is necessary as this stage of the parasite is nonreplicating. The reagent, Presto Blue, is a redox indicator with improved sensitivity over Alamar Blue. The in vitro assays developed in house were used to screen a compound collection with known P. falciparum activity. Compounds with activity identified against the amastigote stage were screened against the trypomastigote stage. Compounds active on only the amastigote stage and some also active on both T. cruzi life cycle stages, with selectivity to the parasite over host 3T3 cells, were identified. These compounds are currently undergoing reconfirmation and further biological evaluation. In addition, an in-house compound library comprising known biologicals and FDA approved drugs (741 compounds) was assessed in the same manner. A number of azole antifungal compounds were identified as active in the amastigote assay, with no activity displayed in the trypomastigote assay. Some compounds that have previously been identified with other screening technologies and reported in the literature were also identified, thus further validating our approach. Cytochrome P450 14αsterol demethylase, CYP51, is recognized as a drug target in T. cruzi, against which azole antifungals have historically been shown to be active. We have shown that lanoconazole, bifonazole, and oxiconazole nitrate, previously unreported, have novel activity in our T. cruzi intracellular amastigote assay. The clinical applications of these compounds are limited as they are topical antifungals, likely with associated mammalian cytochrome P450 systemic activity; for example, bifonazole has been identified with activity against cytochrome P450 enzymes in human microsomal preparations.52 However, these results highlight the capability of the assay to identify compounds active against T. cruzi amastigotes (Table 3). Other active compounds have also been identified from this compound collection and are currently undergoing further evaluation. We are currently developing qualitative, image-based “wash off” assays to determine whether there are detectable parasites, following removal of compound/drug, and extension of incubation at the EC100 of selected compounds. Development of methods to detect the presence of amastigotes following

poscaconazole, a candidate currently in clinical trials, however, in this assay compound was added 2 h after trypomastigote addition.50 This could potentially identify activity against amastigotes and also inhibition of infections by trypomastigotes which may occur after the 2 h incubation. A compound library, comprised of 909 compounds that are mainly FDA approved, was screened in 96-well format using an image-based amastigote specific HTS assay, and the hits were used for assay validation.45 In 2012, the Institute Pasteur, Korea, screened a Pfizer library of 150,000 compounds against amastigotes using an image-based assay, in a 96-well format, with infected myoblasts.51 The outcomes of this assay have not currently been reported. It is difficult to directly compare the assays described in the literature, and their reported screening outcomes, due to the differing technologies and life cycle stages exposed to compound. However, the β-galactosidase reporter gene assay would be less costly to apply to HTS than image-based technologies and more transferrable between laboratories. A benefit of the image-based assays is that the cytotoxicity of compounds against the host cells can also be estimated in the same well. Morphological effects of compounds on the host cell can also be measured. This allows for a direct comparison of mammalian cell cytotoxicity to T. cruzi activity, removing the need to perform two independent assays which may vary in sensitivity. Image-based assays may also be able to identify fewer remaining amastigotes in a host cell than whole well fluorescence-based methods and therefore provide a more accurate determination of levels of infection. We have developed T. cruzi phenotypic, HTS ready assay formats to separately take into account both life cycle forms of the parasite. The 384-well image-based assay estimates compound activity on both the intracellular amastigote of T. cruzi Tulahuen strain and the 3T3 fibroblast host cells. Both the parasite and host nuclear material are stained with a DNA dye, and a separate cytoplasmic stain defines the host cell (Figure 6). Compound activity on the host cell-free trypomastigote is assessed using a fluorescence-based assay. This assay estimates the impact incubation of host cell-free trypomastigotes with compound for 48 h has on metabolism, rather than cell G



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Table 3. Antifungal Azoles with Activity Identified against T. cruzi Amastigotes in an Image-Based Assaya

a Values are based on two separate experiments, with the IC50 values calculated over one replicate of 15 doses of each compound. For all compounds, the IC50 value on the host cell could not be determined at these doses. The selectivity index is based on the top screening dose divided by the IC50 in the amastigote assay.

responsible for 95% of cases reported,53 some lasting several years. The disease progresses from a first, hemolymphatic stage to a second, meningoencephalitic stage in which the parasites cross the blood−brain barrier (BBB) and invade the central nervous system (CNS). Symptoms in the second stage of HAT most commonly include headache, lymphadenopathy, and sleeping disorders. The drugs used to treat HAT are limited and include those that are able to cross the BBB (melarsoprol, eflornithine, nifurtimox), for treatment of the second stage of the disease or those that are not (pentamidine, suramin), for treatment of the

removal of compound would provide a valuable tool for the drug discovery process and may enable identification of a potential suppressive action of a compound. Such a method would be beneficial for lead optimization and biological profiling.

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HUMAN AFRICAN TRYPANOSOMIASIS (HAT): TRYPANOSOMA BRUCEI BRUCEI Human African trypanosomiasis (HAT) is caused by two subspecies, Trypanosoma brucei rhodesiense and Trypanosoma brucei gambiense. T. b. gambiense is endemic to West Africa and H



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hemolymphatic stage.54 These drugs are not all able to treat both subspecies of HAT. Many of these compounds show toxicity, such as melarsoprol, which has been reported to cause the deaths of 10−70% of patients.55,56 The introduction of a combination of nifurtimox and eflornithine to treat second stage T. b. gambiense infection has been a major step forward, as this combination has been found to reduce toxicity.57 Few other compounds remain in clinical trials. Fexinidazole is in phase II/III clinical trials to treat HAT, and recent data shows promise for this compound against treating both subspecies and both stages of the disease.58 SCYX-7158, a benzoxaboral, is also in phase 1 trials for the second stage of the disease, again across subspecies.59 There have been some recent failures in the clinic, for instance, DB289, a promising candidate for oral treatment of phase 1 HAT, was discontinued due to renal toxicity.60 New compounds are therefore needed to maintain the drug discovery pipeline. Culture of T. brucei Subspecies. For the development of assays to screen compounds against the whole organism, the bloodstream form of T. brucei subspecies is the most clinically relevant. In mammalian infections, both trypanosome subspecies are transmitted by the tsetse fly. The metacyclic form of the parasite is injected into the host when the insect takes a blood meal, which then transforms into the bloodstream form. Historically, the bloodstream form that has been used for in vitro studies is the closely related nonhuman infective subspecies, T. b. brucei. The similarity between the three subspecies becomes an important consideration in the use of T. b. brucei for compound screening. While there is genetic similarity between the T. brucei subspecies, T. b. brucei does show some haplotypes not found within the other groups. T. b. gambiense consists of two subgroups, of which group 1 is most common, although it shows less genetic similarity to T. b. brucei and T. b. rhodesiense than group 2.61 Using the human infective forms could potentially increase the success of phenotypic screens due to genetic differences between subspecies. We have generally found a high confirmation rate of activity against T. b. rhodesiense for compounds identified through HTS using T. b. brucei.62 There are undoubtedly compounds which have not been identified against T. b. brucei, which would have been detected had the primary screen been against T. b. rhodesiense, but unfortunately this is not currently feasible for large compound libraries. Compounds with activity against T. b. gambiense, responsible for 95% of HAT cases, would be highly beneficial, thus a primary screening campaign undertaken with this subspecies, with compound activity reconfirmed on T. b. rhodesiense would be ideal. However, very few reports show the use of T. b. gambiense in liquid culture because of the difficulties of propagation of this subspecies both in vitro and in vivo. However, a recent report on the growth of T. b. gambiense strains over 10 consecutive subpassages in an HMI-9 medium, supplemented with methylcellulose and human serum, holds some promise.63 All seven strains of T. b. gambiense tested were able to be cultivated in this manner, and following adaption of primary cultures, displayed exponential growth. One strain showed a maximum cell number of 1.5 × 106 cells/mL with a doubling time of 12.4 ± 1.6 h when inoculated at 5 × 104 cells/ mL. It would be necessary to determine whether freezing the parasite for storage was possible and whether this, and subculturing, impacted on infectivity. If it were necessary to constantly adapt primary cultures, this would limit the application for large scale screening of compounds. Although

further work is required to determine whether this culture system would be applicable to HTS, it is a promising step forward in the culture of this infective form. Assays in T. brucei Drug Discovery. A recent review by Jones and Avery (2013) provides a comprehensive analysis of the current state of whole organism HTS for T. b. brucei.64 With the exception of eflornithine, the current pipeline of existing and potential new drugs for the treatment of HAT all resulted from phenotypic screens.65 Two assay technologies dominate phenotypic drug discovery for T. b. brucei, namely variations of the Alamar Blue assay, including 384- and 96-well formats66 and an ATP-bioluminescence based assay, which has also been developed in to 96- and 384-well assays.67 Most recently, the use of resazurin-based assays for a T. b. brucei whole cell HTS have been adopted.68,69 Once compounds have been identified from phenotypic screening and counter cytotoxicity estimation, some initial testing before progression of compounds should include determination of the static or cidal nature of a compound. Most T. b. brucei phenotypic screens rely on a low initial inoculum, and if a compound does not kill the whole population (cell division is inhibited), this may appear to be 100% active following incubation with the compound. A recent publication has focused upon this and have developed an assay that may detect anomalies in dose response compound plots (biphasic curves, which may suggest a dual static/cidal action) by increasing the concentration of the starting inoculum in the assay.70 This approach uses a 4 × 105 cell/mL inoculum, incubating the cells for 20 and 44 h (total of 24 and 48 h with 4 h incubation with resazurin) and compares the resulting dose response curves over time with an assay using 5 × 103 cells/mL, with 72 h incubation. With these higher cell densities, cultures reach stationary phase earlier, and in these studies this occurs within 48 h. This may affect calculation of the compound action, which the authors have noted. From a dose curve at 48 h at these high doses, a biphasic effect however can be seen with a compound tested that was not seen in the low inoculum, 72 h assay. This study highlights the need for static/cidal analysis, and the challenges associated with high cell numbers impacting on the sensitivity of an assay with this organism. We have used an approach with a low inoculum Alamar Blue assay to determine the EC100 and count cells over time to estimate if compounds are trypanocidal at these doses during prioritization of compound leads.62 This should be also extended to the IC50 value to determine if cells are killed over time, although even counting has limitations in terms of the ability to detect a very small population. To profile the static/cidal nature of a compound, wash off experiments would be beneficial to determine if a population of cells is able to recover from drug treatment. This has recently been used in pre-in vivo testing with a promising preclinical candidate, SCYX-7158, in combination with an ATP-based luminescence assay to determine cell viability at various time points.59 This compound showed irreversible, nM activity in these assays and in vivo activity, and is now in clinical trials. T. b. brucei HTS Screening Campaigns and Outcomes. We have recently published a number of leads identified from a T. b. brucei HTS campaign, with activity against T. b. rhodesiense,62 which are currently undergoing medicinal chemistry optimization. This includes five new scaffolds, phenylthiazol-4-ylethylamide, phenoxymethylbenzamide, 6aryl-3-aminopyrazine-2-carboxamide, pyrido-isoxazol-2-ylanilide, and aminoethyl benzoylarylguanidine. Recently, 600 I



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Table 4. Active Compounds Identified from Application of a T. b. brucei 384-Well Alamar Blue Assay to Screen a Compound Library Collection Comprising of Compounds with Known Biological Activity and FDA Approved Compounds

developed based on this compound and screened against T. b. rhodesiense whole cells, however, the technology is not mentioned. A subset was identified with nM in vitro activity, and one compound showed efficacy in a murine model of trypanosomiasis.73 A smaller (29 compounds) novel quinoline type library has also been developed and screened against T. b. brucei and T. b. rhodesiense whole cells using an Alamar Blue assay, identifying compounds with nM in vitro activity against both subspecies. One of the compounds was tested in a T. b. rhodesiense acute mouse model, however, treatment was ineffective and the authors note that solubility could be improved.72 Libraries containing compounds with known biological activity, including FDA approved drugs, have been screened against T. b. brucei using phenotypic assays. One such library, comprising primarily FDA approved drugs, bioactives, and natural products (2160), was screened in 96-well format using a

agrochemicals were screened against T. b. rhodesiense as well as T. cruzi and P. falciparum.71 A number of compounds were found to be active against T. b. rhodesiense, and zoxamide, a broad-spectrum oomyceticide, was tested in a T. b. rhodesiense acute mouse model, showing weak activity. On day 7, no parasite could be seen, however on day 10 all mice relapsed. It would be beneficial to determine whether this compound has trypanostatic activity. Quinolines have been reported in a number of publications as active against T. b. brucei, utilizing phenotypic approaches to discover compounds with demonstrated in vivo activity.72,73 One reported HTS campaign against the Specs (www.specs.net) compound library resulted in the lead identification of a series of dihydroquinolines.73 Compound 1A was tested in a T. b. brucei mouse model, and although it did not provide a cure, it did show a reduction in parasitemia. This compound served as a hit for further optimization, and 1,2-dihydroquinoline derivatives were J



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luminescence-based HTS assay, and the hits were used as a validation of the assay format.74 A resazurin-based cell viability assay was developed for in vitro phenotypic screening of the LOPAC 1280 “library of pharmacologically active compounds” against bloodstream forms of T. b. brucei.68 We have also screened a small library comprised of known biologicals and FDA approved drugs, with a number of similar compounds being identified (Table 4). Additional compounds, which previously have not been reported as trypanocidal, have also been identified and demonstrated to be selective. Biological in vitro and in vivo profiling of these compounds is currently ongoing.

and avirulent strains of the plant root pathogen, Phytophthora cinnamomi. Melissa went on to work as a research assistant at Natural Product Discovery at Griffith University in 2001. She optimized assays for application in high throughput screening (HTS) campaigns over various disease areas, including biochemical and cell-based assays. Melissa worked as a research assistant at the Eskitis Institute in 2006, whereby she developed the first published 384 well HTS assay for Trypanosoma brucei brucei whole cells. Melissa is currently studying for a Ph.D. and has developed a high throughput imaging based assay for the detection of compound activity against Trypanosoma cruzi amastigotes in 3T3 host cells.

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Vicky M. Avery obtained her Ph.D. in 1995 (Flinders University, South Australia) and was awarded an Australian NHMRC Postdoctoral Fellowship undertaken at the University of Adelaide. She gained significant industry experience at Active Biotech AB, Sweden (1998−2004). As Head of Biology for the AstraZeneca/ Griffith University collaboration, she was responsible for 50 HTS campaigns (2004 −2007), spanning many disease areas and encompassing a diverse range of technologies. Professor Avery is currently the Chief Investigator & Head of Discovery Biology, Eskitis Institute for Drug Discovery, Griffith University, Australia. Discovery Biology undertakes basic and applied research in drug discovery, primarily in the areas of cancer and neglected diseases. Her research focuses on innovative approaches for high throughput screening, with a focus on image-based technologies.

CONCLUSIONS Phenotypic screening, while informative and providing the security that the molecule does impact on the live organism, is not without its own inherent issues. In vitro maintenance of biological cultures can itself bring challenges with the need to ensure that cultures are able to produce sufficient quantities of the phenotype needed for HTS, and the health/age of the culture should also be considered. The species and strain of the organism and the life cycle stage must be clinically relevant. It should be considered which assay technologies may be applicable to the organism used, how they can be optimized, and how cytotoxicity can be estimated. Chemical lead optimization programs reliant on these assays must often contend with extended incubation times and thus turn-around time for results. Chemical optimization may not be driven by one target, there may be multiple targets, and more than one assay will be required to ensure against potential technology or off-target-related activities of compounds. If compounds identified have no previous safety testing, assays to identify absorption, distribution metabolism, elimination, and toxicity (ADMET) should be considered, as well as optimization of pharmacokinetics, bioavailability, and solubility. Once a compound with in vivo efficacy is identified from a lead optimization program, target(s) elucidation is desirable. This would ultimately allow for the development of further compounds based on the biology of the target. A balance must be sought between when molecular target/ mechanism of action studies are initiated, as ideally these should be undertaken with the best candidate molecule available. Clearly, not all molecules become drugs, and a vast number have become valuable tools for increasing our knowledge of both the biology of the disease or organism of interest and the chemical classes and properties which impact on these. Success in drug discovery relies not only on knowledge, but also on opportunity and luck.

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ACKNOWLEDGMENTS

We acknowledge and thank (DNDi) for financial support to develop the T. b. brucei assays, and we also acknowledge and thank the Medicines for Malaria Venture (MMV) for providing financial support for components of the P. falciparum asexual and gametocyte assays described here. M.L.S. is the recipient of an Australian postgraduate Scholarship Griffith University Postgraduate Scholarship (GURPS). We wish to thank Dr Achim Schnaufer, (University of Edinburgh) who whilst at the Seattle Biomedical Research Institute\ kindly supplied the trypanosome cell stock used throughout this study. We would also like to thank Professor Fred Buckner, from the University of Washington, for supplying the T. cruzi epimastigote stocks, and Dr. Victor Contreras from the Universidad de Carabobo for his aid with TAU3AAG formulation and T. cruzi differentiation protocols. We acknowledge the Australian Red Cross Blood Service for the provision of human erythrocytes. Our thanks to the members of the Discovery Biology group who have contributed to the studies referred to within the manuscript. Finally, we would like to thank Dr. Amy Jones for editorial assistance and Drs. Graeme Stevenson and Rohan Davis for assistance with redrawing chemical structures.

AUTHOR INFORMATION

Corresponding Author

*Phone: +61737356056. Fax: +61737356001. E-mail: v.avery@ griffith.edu.au.

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Author Contributions

ABBREVIATIONS USED ACT, artemisinin combination therapies; BMGF, Bill & Melinda Gates Foundation; CD, Chagas disease; DAPI, 4′,6diamidino-2-phenylindole; GNF, Genomics Institute of the Novartis Research Foundation; GSK, GlaxoSmithKline; HAT, human African trypanosomiasis; MMV, Medicines for Malaria Venture; TAU3AAG, TAU (artificial triatomine urine), supplemented with 10 mM L-proline, 50 mM L-sodium glutamate, 2 mM L-sodium aspartate, and 10 mM D-glucose

The manuscript was written through contributions of both authors. All authors have given approval to the final version of the manuscript. Notes

The authors declare no competing financial interest. Biographies Melissa L. Sykes completed an Honours degree at Murdoch University, Western Australia, on the enzyme production on virulent K



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