In this protocol, fluorescently labeled T. cruzi were injected into transparent zebrafish larvae, and parasite motility was observed in vivo using light sheet fluorescence microscopy.
Chagas disease is a parasitic infection caused by Trypanosoma cruzi, whose motility is not only important for localization, but also for cellular binding and invasion. Current animal models for the study of T. cruzi allow limited observation of parasites in vivo, representing a challenge for understanding parasite behavior during the initial stages of infection in humans. This protozoan has a flagellar stage in both vector and mammalian hosts, but there are no studies describing its motility in vivo.The objective of this project was to establish a live vertebrate zebrafish model to evaluate T. cruzi motility in the vascular system. Transparent zebrafish larvae were injected with fluorescently labeled trypomastigotes and observed using light sheet fluorescence microscopy (LSFM), a noninvasive method to visualize live organisms with high optical resolution. The parasites could be visualized for extended periods of time due to this technique's relatively low risk of photodamage compared to confocal or epifluorescence microscopy. T. cruzi parasites were observed traveling in the circulatory system of live zebrafish in different-sized blood vessels and the yolk. They could also be seen attached to the yolk sac wall and to the atrioventricular valve despite the strong forces associated with heart contractions. LSFM of T. cruzi-inoculated zebrafish larvae is a valuable method that can be used to visualize circulating parasites and evaluate their tropism, migration patterns, and motility in the dynamic environment of the cardiovascular system of a live animal.
Chagas disease is caused by the protozoan parasite T. cruzi. Approximately 6 to 7 million people worldwide are infected with T. cruzi. The disease is transmitted mainly in Latin America, but has been reported in the United States, Canada, and many European as well as some Western Pacific countries, mainly due to migration of infected individuals1. Chagas is largely vector-borne and transmitted to humans by contact with the feces of hematophagic insects in the Triatominae subfamily, commonly known as "kissing bugs". However, T. cruzi can also be transmitted via blood transfusions, vertical transference from mother to child, or ingestion of food contaminated with parasites2. The acute phase the infection is mainly asymptomatic or constitutively symptomatic and lasts from 6 to 8 weeks, after which engagement of the immune system controls parasite load, but does not completely eliminate the infection3. Most individuals then enter a chronic asymptomatic phase; however, nearly 30% of patients develop a symptomatic chronic phase, in which the cardiac system and less frequently the digestive and nervous systems are compromised4. This scenario presents a challenge for disease treatment and control since there are no vaccines available, and there are only two effective drugs for Chagas: benznidazole and nifurtimox. Both treatments require prolonged administration and may have severe side effects2.
Increased understanding of the behavior of T. cruzi behavior in vivo is key to determining parasitic migration, cellular attachment, and invasion within the host; a lack of in vivo models limits the development of novel therapeutic approaches. In vitro studies of T. cruzi infection have shown that motility of trypomastigotes is important for binding to host cell membranes and subsequent cellular invasion5. Energy depletion in trypomastigotes in co-culture with a susceptible cell line has been shown to reduce cellular invasion6. Interestingly, in trypanosomatids, flagellar movement has also been characterized as an evasion mechanism against parasite-specific antibodies7.
Flagellar motility has been extensively studied in vitro in Trypanosoma brucei, a closely related parasite that causes African Trypanosomiasis8. In vitro studies of the motility of these trypanosomes showed that simulation of the conditions of blood or body fluids, including viscosity and the presence of obstacles representative of blood cells, is important for parasite forward movement9. As of yet it has not been possible to visualize the movement of parasites in the bloodstream in vivo.
Zebrafish larvae are a powerful model to study host-pathogen interactions in vivo. They are small, inexpensive, and relatively easy to raise when compared with other established vertebrate models for Chagas disease. Zebrafish have innate and adaptive immune systems similar to humans, but their adaptive immune system begins to develop at 4 days post fertilization (dpf) and is not mature for another several weeks10. During early development, when only macrophages are present, there is a large window for studying parasite behavior without immediate immune interference10. However, the greatest advantage of utilizing zebrafish larvae as a vertebrate model for studying pathogen behavior lies in their optical transparency, making them amenable for microscopic screening and imaging11. Additionally, there are multiple tools to manipulate fish genetics. For example, the Casper strain is a mutant line of zebrafish with no pigmentation, making the animal completely transparent and useful for visualization of individual organs and for real-time tracking of injected cells12.
A key limitation of longitudinal observation of swiftly moving parasites in live zebrafish using confocal or epifluorescence microscopy lies in the impossibility of imaging at high acquisition speeds and large penetration depths with good image quality and low risk of photodamage. Light sheet fluorescence micsroscopy (LSFM) is an emerging imaging technique that overcomes these limitations to permit these observations. By using one objective to detect fluorescence and a second orthogonal illumination objective that only illuminates the focal plane of the detection objective, it is possible to obtain high resolution optical sections as in a confocal microscope, but with significantly reduced photodamage, even with respect to epifluorescence microscopy13. The LSFM technique used here is called Single Plane Illumination Microscopy (SPIM), in which a thin sheet of light illuminates a single plane within the zebrafish larvae.
The objective of this methodology is to establish larval zebrafish as a viable non-infection model for understanding T. cruzi motility and related behavior in vivo. To accomplish this, we injected transparent zebrafish larvae with fluorescently labeled trypomastigotes, the cellular form responsible for infection of humans, and identified the movement of T. cruzi in the cardiovascular circulation of zebrafish using LSFM.
The following protocols were approved by the Institutional Animal Care and Use Committee of Los Andes University (CICUAL). Biosafety level 2 (BSL-2) guidelines should be strictly followed to prevent contamination with the pathogen T. cruzi.
Note: Animal care and maintenance: Casper zebrafish, a genetically modified strain of zebrafish (Danio rerio) is used in this protocol due to their valuable optical transparency in all developmental stages. Fish are manipulated using optimal care conditions for the species14, in a 14 h light-10 h dark cycle, at 28 ± 1 °C, in a pH (7.0-7.4) controlled multi-tank recirculating water system. Animals are fed twice a day with live brine shrimp (Artemia salina) and enriched with rearing food. All protocols were approved by CICUAL at Universidad de los Andes (C.FUA_14-017).
1. Preparation of Egg Water
2. Preparation of Tricaine Stock Solution
3. Preparation of 1.0% Low Melting Point Agarose
4. Mating Assay and Embryo Collection
5. Embryo Dechorionation
Note: This procedure is required if the embryos have not hatched at the time of injection. In this procedure, "larvae" are animals out of their chorion from 48 h post fertilization (hpf) onward.
6. Preparing Injection Material
7. Cell Culture for Parasite Growth
8. T. cruzi Culture and Labeling
9. Injecting Zebrafish Larvae
10. Injection of the Parasite
11. LSFM Mounting of Injected Larvae
12. LSFM Imaging of Injected Parasites
13. Image Processing and Analysis of Acquired Data
Note: Image processing was performed on a personal computer with a 2.90 GHz processor, 8.00 GB of memory, and a videocard with 1.00 GB of memory.
14. Recovering Imaged Larvae
Optimal Conditions for Injection:
Groups of zebrafish larvae were injected at 24, 48, 72, 96, and 120 hpf, at different anatomical sites, and their survival was examined every day for 5 days. After 5 days post injection, embryos injected at 24 hpf had 6.25% (2/32) survival, whereas 95% (38/40) of larvae injected at 48 hpf survived. As a control, larvae were injected with 1x PBS as a vehicle. There were no differences in survival between vehicle-injected and parasite-injected larvae, indicating no parasitic-dependent effects on survival rate of the fish (p = 0.08). Larvae injected between 72-120 hpf had comparable survival rates to 48 hpf-injected larvae at constant injection volume. For all procedures presented here, 48 hpf larvae were used due to their ease of manipulation and having developed organs and easily penetrable skin without evident damage after injection.
Larvae injected at 48 hpf were injected in the pericardial space, tail muscle, hindbrain ventricle, otic vesicle, notochord, and duct of Cuvier in the yolk sac. There were no differences in the survival of larvae injected at differing anatomical sites. However, the fastest and easiest region to inject was the duct of Cuvier located in the anterior part of yolk sac (Figure 1, Movie 1). Injections at that site allowed the introduction of higher volumes with a lower risk of injury to vital structures. Additionally, between 24-72 hpf, this region is an optimal site to directly access the developing vasculature and heart11.
Parasite Visualization Using LSFM:
Within 8-10 min following injection of T. cruzi into the duct of Cuvier, parasites were identified in zebrafish larvae using LSFM due to their CFSE fluorescent signal and the optical transparency of the larvae. After inoculation, parasites were observed either adhered to walls around the circulatory system or traveling in the direction of blood flow (Figure 2, Figure 3). When a parasite remains attached to a cardiac structure, such as the atrioventricular valve, it oscillates with heart contractions, indicating that the molecular mechanisms for the adherence of parasites might be effective in our vertebrate model ( Movie 2, Movie 3, Supplemental movie 1). T. cruzi also adhered to the walls of the larval yolk sac ( Figure 2, Movie 2), a structure which will later be reabsorbed and become part of the zebrafish intestine22. This could be similar to what happens during the chronic disease phase in infected humans, where parasites are found in cardiomyocytes and in the digestive nervous system23,24. When not attached, the parasites drifted through the blood flow in the same direction as the erythrocytes ( Figure 3, Movie 4). Parasites could be observed in different sized vessels of the fish, but were more abundant in the pericardial space and in the adjacent yolk region containing blood flow ( Figure 2, Figure 3, Supplemental movie 2).
At 10 min post injection, it was more difficult to spot single forms of the parasite due to their distribution along the vasculature, and an inability to quickly screen different anatomical sites of the fish due to a limited field of view of the LSFM (at 40X magnification). After 24 h post injection (hpi), the CFSE signal starts to accumulate in the region near the developing intestine (Supplementary figure 1).
Figure 1: Optimal injection site. (A) Image of larva 48 hpf showing the optimal injection site at the duct of Cuvier (yellow arrow) using a regular stereoscope. (B) Magnified view of box in A showing the duct of Cuvier (yellow arrow). Scale bar = 200 µm (A), 50 µm (B). Please click here to view a larger version of this figure.
Figure 2: LSFM images of a static parasite in a 48 hpf larva. The T. cruzi parasite (yellow arrow) remains adhered to the walls of the yolk sac, throughout the time-lapse sequence (7.2 s, 17.2 s, and 27.2 s), about ~15 min after parasite injection. No change in position of the parasite is observed during an acquisition period of at least 30 s. a, Atrium; v, Ventricle. Scale bar = 50 µm. Please click here to view a larger version of this figure.
Figure 3: Trajectory of a parasite traveling in the pericardial space using LSFM. The T. cruzi parasite can be tracked while drifting in the pericardial space (PS), following the direction of blood flow (track shown in red) about ~15 min after parasite injection. Scale bar = 50 µm. Please click here to view a larger version of this figure.
Movie 1: Blood circulation valley of the yolk in a larva 48 hpf. Movie of a larva 48 hpf showing the blood circulation valley or duct of Cuvier using a regular stereoscope. Different regions are focused during the video to show red blood cells circulating throughout the duct. Yellow arrow shows the optimal injection site. Movie was recorded about 10-15 min after parasite injection. Please click here to download the video.
Movie 2: T. cruzi parasites attached to walls of the yolk sac. LSFM movie of a larva 48 hpf showing that T. cruzi parasites remain adhered to the yolk sac about 10-15 min after parasite injection. No change in the position of the parasite was observed during an acquisition period of at least 30 s. a, Atrium; v, Ventricle. Please click here to download the video.
Movie 3: T. cruzi parasites attached to the walls of the heart. LSFM movie of a larva 48 hpf showing that T. cruzi parasites remain adhered to the cardiovascular wall, despite the strong heart contractions about 10-15 min after parasite injection. Erythrocytes can be observed as black rounded shadows. Please click here to download the video.
Movie 4: Parasites moving in the pericardial space. LSFM movie of a larva 48 hpf showing T. cruzi parasites drifting in the pericardial space. Two parasites can be traced at different time points (ID 1, in red circle, and ID 2 in yellow circle), following a similar trajectory. The movie was recorded about 10-15 min after parasite injection. Please click here to download the video.
Supplemental Figure 1: Accumulation of CFSE fluorescent signal in the yolk. Stereoscope images of a wildtype larva injected at 48 hpf at the duct of Cuvier. CFSE fluorescent signal progressively accumulates in the yolk, after two days post injection (48 hpi). Scale bar = 500 µm. Please click here to download the figure.
Supplemental Movie 1: Parasites attached to walls and valves of the circulatory system. Stereoscope time series images of a wild type larva injected at 48 hpf. Images are taken at 0.2 s intervals capturing T. cruzi parasites moving in synchrony with cardiac muscle contractions at the atrioventricular valve. Movie was recorded about 30 min after parasite injection. Please click here to download the video.
Supplemental Movie 2: Moving and adhered parasites in the periventricular space and yolk. LSFM movie of a larva 48 hpf showing T. cruzi parasites drifting or adhered to the pericardial space or the yolk. A transmitted light view was observed for the first 5 s. The fluorescence view was observed from 5.2-25.8 s. The movie was recorded about 10-15 min after parasite injection. Please click here to download the video.
This study highlights the advantages of zebrafish as an animal model for studying pathogen behavior in vivo. In particular, this study proposes a method to visualize the pathogen T. cruzi in its natural environment: hematic circulation. The environment of the circulatory microenvironment in fish is comparable to that of mammals, and trypanosomatids have evolved mechanisms for traveling, evading the immune system, and attaching to cells for infection in that environment25. This protocol offers a description of an optimal procedure for culture of T. cruzi in a human cell line and subsequent isolation of flagellar forms for fluorescent labeling. It then shows the appropriate settings for successful injection of the parasites into transparent zebrafish for later mounting and visualization using LSFM. Finally, this protocol offers suggestions for efficient and effective in vivo imaging of parasite location and movement in circulation using LSFM.
The flagellum of trypanosomes emerges from its posterior region, flowing from the cell body, and hangs free at the anterior part of the organism26. T. cruzi propels itself by waving the flagellum ahead of the body, which undulates the parasite's entire body. Flagellar movement is not only indispensable for parasite motility, as in the case of T. brucei27 (the causative agent of African trypanosomiasis), but it is also used for cellular infection, as has been demonstrated in T. cruzi5,28. Although zebrafish are not the natural host for T. cruzi, the parasite's motility can be studied in a developing cardiovascular circulation system using the protocols described here. Additionally, there are trypanosomes species that infect cyprinids, the class of zebrafish, such as T. carassii and T. borreli25. These parasite species could be used to study in real time the movements and attachment mechanisms of these trypanosomatids; such studies can lend insight into the mammalian cell infection process.
In this study, injected motile T. cruzi parasites were observed traveling through the cardiovascular circulation of inoculated fish, moving along with opaque erythrocytes, and adhering to structures in the cardiovascular system walls. We used a home-built LSFM with a 10X achromatic long working distance air objective (17.6 mm) for the illumination arm with a numerical aperture of 0.25. A 40X apochromatic water immersion objective with a numerical aperture of 0.8 and a working distance of 3.5 mm was used for the detection arm. The detection objective was immersed in the sample chamber, while the illumination objective was outside the chamber. A port in the chamber sealed with a coverslip and optical glue allowed for the illumination beam to enter the chamber, as depicted in Lorenzo et al.18 For illumination, we used a 50 mW DPSS laser at 488 nm whose power was modulated by an Acousto Optical Tunable Filter. The detection path used filters compatible with Green Fluorescent Protein (GFP) or FITC. A light sheet microscope equipped with a capillary sample holder (ideally with automated rotation) and temperature control of the sample chamber should be suitable for this application. The microscope should be aligned and calibrated according to the manufacturer's instructions or user's laboratory standard protocols before acquisition, if necessary. In this protocol, we controlled the microscope using the SPIM software19.
It is important to note that in the circulation of zebrafish, larvae parasitic attachment is effective. In the cardinal vein, parasites remained attached for up to several minutes; in the heart, they held on to valves and walls, oscillating with heart contractions. Further studies remain in order to elucidate whether T. cruzi interacts with the flowing erythrocytes that drift in the direction of blood flow. Previous in vitro studies have shown that the presence of solid structures (i.e., blood cells), or increased viscosity of the liquid to mimic blood in vitro, has a significant effect on the motility and velocity of the parasite9.
There are many questions regarding the course of infection of T. cruzi in humans after amastigotes escape phagocytic cells, the initially infected cell type29. For example, how do they arrive to their target organs? What are the mechanisms for tropism to the preferred organs, such as cardiac, digestive, and central nervous systems? Interestingly, in this study the parasites were initially imaged in the heart because it was the site of highest density of parasites. However, the CFSE signal subsequently accumulated in the developing intestine by 7 days post injection. Although the anatomy of fish and mammals is different, the results of this study demonstrate a form of tropism, as it was observed that parasites exhibited tropism toward known preferred target organs despite organismal differences. One significant limitation of this study concerns the temperature used in the experiments. Zebrafish larvae should be kept around 28 °C during the entire procedure. Though this temperature might be similar to the vector host (insects of the Triatominae subfamily), it is quite different from the warm-blooded mammals that comprise the final hosts (around 37 °C). T. cruzi is known to have flagellar living forms in both hosts; however, it is important to bear in mind that this factor might have an effect in the behavior of the animal in vivo.
Although the fish´s adaptive immune system is not mature until 4 weeks post fertilization, the innate immune system is active early in development10. As early as 48 or 96 hpf, phagocytic cells were observed having engulfed labeled trypanosomes (data not shown). This limits the window of time for visualization of the parasite. However, if a study was to focus on evaluating the fish´s immune response, injection at later stages may be recommended. Also, injection of parasites into transgenic fish lines with labeled macrophages or other cells of the immune system can be useful in studying parasite attachment and possible endocytosis mechanisms. It is important to note that if the parasites are labeled with CFSE, transgenic cell labels should not be GFP, and a marker in the yellow or red end of the spectrum is required.
To assess the detailed direction of parasite movement, it may be useful to follow their trajectory in 3 dimensions (3D). For 3D visualization and reconstruction of the process, a high-speed system is necessary. With the equipment used in this protocol, it is only possible to visualize the parasites in one plane. In this case, we prioritized maintaining focal plane stability during the parasite movement, and recording its trajectory in one plane.
The methodology proposed here paves the way to further investigate parasite behavior in cardiovascular circulation. In summary, the essential steps to imaging live fluorescent parasites inside zebrafish larvae are: (i) use of early hatched embryos (24-48 hpf) or larvae, or animals between 72-96 hpf with no pigmentation so that they are transparent and easy to inject; (ii) image larvae as soon as possible after injection to avoid parasite clearance by phagocytic cells; and (iii) focus the LSFM on the site of interest (e.g., pericardial region) and maintain the focus. This novel procedure allows the visualization of trypomastigotes in an environment comparable to its natural infection niche, providing for the first time the possibility to study T. cruzi in a living organism.
The authors have nothing to disclose.
This work was supported by the Convocatoria Interfacultades from Vicerrectoría de Investigaciones de la Universidad de los Andes, and the USAID Research and Innovation Fellowship program. We thank Juan Rafael Buitrago and Yeferzon Ardila for fish maintenance and assistance.
0.5-10 μL Micropipette | Fisherbrand | 21-377-815 | |
Agarose RA | Amresco | N605 | Regular |
Agarose SFR | Amresco | J234 | Low Melting point |
Aquarium salt | Instant ocean | SS15-10 | |
Cell Count chamber | Boeco | Neubauer | |
Cell culture flasks | Corning | 430639 | |
Centrifuge | Sorvall | Legend RT | |
CFSE | ThermoFisher | C34554 | |
Detection objective | Nikon 40x 0.8NA | 40x CFI APO NIR | |
DMEM medium | Sigma-Aldrich | D5648 | |
Dumont #5 fine forceps | World precision Instruments | ||
Ethyl 3-aminobenzoate methanesulfonate salt (Tricaine) | Sigma-Aldrich | A5040 | |
Fetal calf serum (FCS) | Eurobio | CVFSVF00-01 | |
Filter | Chroma | ET-525/50M | |
Glass capillaries for embryo mounting | Vitrez Medical | 160215 | |
Glass capillaries for pulling needles | World precision instruments | TW100-4 | |
Glucose | Gibco | A2494001 | |
HEPES | Gibco | 156300-80 | |
Incubator | Thermo Corporation | Revco | |
Larval microinjection mold | Adaptive Science Tools | I-34 | |
Laser | Crystalaser | DL488-050 | |
L-glutamine | Gibco | 250300-81 | |
Methylene blue | Albor Químicos | 12223 | |
Micromanipulator | Narishige | MN-153 | |
Micromanipulator system | Sutter Instrument | MP-200 | For LSFM |
Micropipette puller device | Narishige | PC-10 | |
Microscope | Olympus | CX31 | |
Microscope (inverted) | Olympus | CKX41 | |
Multipurpose microscope | Nikon | AZ100M | |
Neubauer counting chamber | Boeco Germany | ||
Penicillin-streptomycin | Gibco | 15140-163 | |
Petri dish 94×16 | Greiner bio-one | 633181 | |
Plastic pasteur pipette | Fisherbrand | 11577722 | |
Rotation stage | Newport | CONEX-PR50CC | |
RPMI-1640 medium | Sigma-Aldrich | R4130 | |
Sodium pyruvate | Gibco | 11360-070 | |
Stereoscope | Nikon | C-LEDS | |
Tricaine (MS-222) | Sigma-Aldrich | 886-86-2 | |
TRIS | Amresco | M151 | |
Trypsin-EDTA (0.25%) | Gibco | R-001-100 | |
Tubes 15 ml | Corning | 05-527-90 |