We thank Dolores Campos for assisting with Vero cell culture and Romina Manarin for assisting with T. cruzi culture. This work was supported by Agencia Nacional de Promoci&#243;n Cient&#237;fica y Tecnol&#243;gica, Ministerio de Ciencia e Innovaci&#243;n Productiva from Argentina (PICT2019-0526), Consejo Nacional de Investigaciones Cient&#237;ficas y T&#233;cnicas (PIBAA 1242), and Research Council United Kingdom [MR/P027989/1].

NOTE: Figure 1 illustrates the complete experimental design.
<img alt="Trypanosomatid immunofluorescence staining; microscopy workflow diagram; fixation, gel expansion." class="xfigimg" src="/files/ftp_upload/65381/65381fig01.jpg" data-altupdatedat="1747911167000" data-alt="Figure 1"> 
Figure 1: U-ExM workflow for three in vitro life cycle stages of T. cruzi. <a href="https://www.jove.com/files/ftp_upload/65381/65381fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
1. Preparation of the poly-D-lysine-coated coverslips
<ol>
	<li>Place a 10 cm x 10 cm square of sealing film in a Petri dish. Wash the coverslips by bathing them in absolute ethanol in a 35 mm glass Petri dish.</li>
	<li>Remove the coverslips with tweezers from the ethanol bath and drain the excess liquid with tissue paper. Place the coverslips over the sealing film.</li>
	<li>Absorb the rest of the ethanol with microscopy-grade paper. Add a 0.1% v/v solution of poly-D-lysine on the center of the coverslip and spread it with the tip to cover approximately 80% of its surface. Close the Petri dish and incubate for 1 h at 37 °C. 
	NOTE: For 22 mm2 coverslips, use 200 µL of poly-D-lysine solution; for 12 mm round coverslips, use 100 µL of poly-D-lysine solution.</li>
	<li>Wash the coverslips with ultra-pure water three times. Use aspiration with a vacuum to remove the water between each wash. Keep at 4 °C for up to 1 week.</li>
</ol>
2. Solution preparation
<ol>
	<li>Prepare stock solutions of 38% (w/w) sodium acrylate (SA).
	<ol>
		<li>Slowly add 19 g of SA to 31 mL of nuclease-free water while stirring. 
		NOTE: This solution is very viscous; pay attention when pipetting.</li>
		<li>Once the SA is completely dissolved, store it in a sterile container. Keep it at 4 °C and change it every 6 months. 
		NOTE: SA sometimes shows signs of contamination with sodium polyacrylate depending on the brand, which is noticeable when preparing the stock solution as it turns yellowish and cloudy. Use it cautiously if this is the case.</li>
	</ol>
	</li>
	<li>Prepare a stock solution of 40% (w/v) acrylamide (AA) and a stock solution of 2% (w/v) N, N'-methylene bisacrylamide (BIS). Dissolve each compound in ultrapure water and filter with a 0.22 µm sterile syringe filter. Store in a sterile container at 4 °C. 
	CAUTION: AA and BIS are highly toxic substances. Work under a fume hood and use proper protection elements (gloves, protective clothing, mask, and safety glasses).</li>
	<li>Prepare the protein crosslinking prevention (CP) solution by mixing 38 µL of a 37% formaldehyde (FA) solution with 50 µL of the 40% acrylamide stock solution (step 2.2) in 912 µL of phosphate-buffered saline (PBS; Table 1) to obtain a final concentration of 1.4% formaldehyde and 2% acrylamide. 
	NOTE: Always prepare the CP solution freshly and be extremely precise when pipetting. For example, use a P1000 pipette to take 900 µL of PBS and a P20 pipette to take the remaining 12 µL of PBS.</li>
	<li>Prepare the monomeric solution 
	NOTE: Do not use the solution immediately after being prepared; store at -20 °C at least 24 h before being used.
	<ol>
		<li>Mix 500 µL of the 38% SA solution (step 2.1), 250 µL of the 40% AA solution, and 50 µL of the BIS solution (step 2.2), as well as 100 µL of 10x PBS (Table 1).</li>
		<li>Aliquot this final volume of 900 µL in 10x 1.5 mL microcentrifuge tubes with 90 µL of monomeric solution each. Store at -20 °C for up to 2 weeks. 
		NOTE: The SA solution may stay on the tip when pipetting because it is very viscous; be careful and dispense everything.</li>
	</ol>
	</li>
	<li>Prepare the denaturing solution
	<ol>
		<li>Mix 114.3 mL of a 350 mM sodium dodecyl sulfate (SDS) solution prepared in ultrapure water with 10 mL of a 4 M sodium chloride solution prepared in ultrapure water. Add 12 g of Tris while stirring in a 250 mL beaker. 
		CAUTION: SDS is highly toxic; use it under a fume hood wearing gloves, protective clothing, a mask, and safety glasses.</li>
		<li>Adjust the pH to 9 with a concentrated solution of hydrochloric acid. Make up to 200 mL with ultrapure water and store in a sterile flask at 4 °C.</li>
	</ol>
	</li>
	<li>Prepare a 10% ammonium persulfate (APS) solution and a 10% TEMED solution.
	<ol>
		<li>Dissolve 0.1 g of APS in 1 mL of ultrapure water.</li>
		<li>Prepare 1 mL of a 10% TEMED solution in ultrapure water.</li>
		<li>Prepare 100 µL aliquots of both solutions in 1.5 mL microcentrifuge sterile tubes and store them at -20 °C for up to 1 month.</li>
	</ol>
	</li>
	<li>Prepare the paraformaldehyde solution.
	<ol>
		<li>Dissolve 2 g of paraformaldehyde in 40 mL of PBS while stirring at 60 °C. Add 1 M NaOH dropwise until the solution turns from white to colorless. 
		CAUTION: Formaldehyde is highly toxic; use it under a fume hood wearing gloves, protective clothing, a mask, and safety glasses.</li>
		<li>Cool the solution at room temperature and adjust the pH with NaOH to 7.2 in a final volume of 50 mL. Filter with a 0.22 µm sterile syringe filter and store in a sterile container.</li>
		<li>Prepare aliquots of 1 mL in 1.5 mL sterile microcentrifuge tubes and store at -20 °C.</li>
	</ol>
	</li>
	<li>Prepare the paraformaldehyde/glutaraldehyde solution.
	<ol>
		<li>Add 0.2 g of paraformaldehyde in 3.5 mL of ultrapure water and 50 µL of 16 M NaOH solution. Heat the solution to 60 °C to dissolve the paraformaldehyde.</li>
		<li>Cool down and add 300 µL of 70% glutaraldehyde. Bring up the volume to 5 mL with ultrapure water and finally to 10 mL with PBS. 
		CAUTION: Paraformaldehyde and glutaraldehyde are highly toxic; use them under a fume hood wearing gloves, protective clothing, a mask, and safety glasses.</li>
		<li>Prepare aliquots of 1 mL in 1.5 mL sterile microcentrifuge tubes and store at -20 °C.</li>
	</ol>
	</li>
</ol>
3. Preparation of the parasite cultures
<ol>
	<li>Grow T. cruzi epimastigotes.
	<ol>
		<li>Use an axenic culture in a T-25 flask (25 cm2 growth area) and maintain the cultures in the logarithmic phase by sub-culturing every 48-72 h in liver infusion tryptose (LIT) medium with 10% fetal calf serum (FCS; Table 1).</li>
		<li>Ensure the cap is securely closed and keep the culture flask vertically at 28 °C for incubation. Monitor the parasite growth by cell counting in a Neubauer chamber during each subculture. 
		NOTE: For the Dm28c strain used in this study, the concentration of epimastigotes in log-phase cultures is between 1-5 x 107 parasites/mL.</li>
		<li>Prepare a suspension of 2 x 106 epimastigotes/mL from a log-phase culture in LIT medium supplemented with 10% FCS. Centrifuge the suspension at 5,000 x g for 10 min at room temperature (RT). Wash with PBS once or twice and resuspend in 200 µL of PBS.</li>
		<li>Adhere to the round 12 mm coverslip previously coated with poly-D-lysine (section 1). Incubate at RT for 15-20 min. Continue to the crosslinking prevention step (section 4). 
		NOTE: Alternatively, fix the epimastigotes with cold methanol for 7 min or paraformaldehyde/glutaraldehyde solution (step 2.8) for 10 min at RT prior to adherence of the parasites to the coverslip. It is possible to store the fixed parasites at 4 °C for up to 1 week.</li>
	</ol>
	</li>
	<li>Obtain the amastigotes from infected Vero cells.
	<ol>
		<li>Lay a sterile 12 mm round coverslip on the bottom of a 24-well tissue culture plate. Prepare a suspension of 2 x 105 Vero cells/mL in Dulbecco's modified Eagle medium (DMEM) supplemented with 2% fetal calf serum (FCS). Seed 500 µL of the suspension per well. 
		NOTE: This study uses DMEM with 2% FCS to allow the cells to grow slower.</li>
		<li>Incubate overnight (ON; 12-16 h) at 37 °C and 5% CO2 to ensure cell attachment.</li>
		<li>Following incubation, wash the cells twice using 500 µL of sterile PBS. Add T. cruzi trypomastigotes into the cells at a multiplicity of infection (MOI) of 10 in 100 µL of DMEM with 2% FCS per well, corresponding to one million trypomastigotes per well. Incubate at 37 °C and 5% CO2 for 6 h.</li>
		<li>After incubation, rinse the plates twice with PBS. Add 500 µL of DMEM supplemented with 2% FCS. At this point, continue to the crosslinking prevention step (section 4). 
		NOTE: Intracytoplasmic amastigotes will be visible through an inverted optical microscope 2 days post-infection (Supplementary Figure 1). Alternatively, fix the trypomastigotes with cold methanol for 7 min at -20 °C, or paraformaldehyde/glutaraldehyde for 10 min at RT, prior to adherence of the parasites to the coverslip.</li>
	</ol>
	</li>
	<li>Obtain trypomastigotes from infected Vero cells.
	<ol>
		<li>Collect the supernatant of a Vero cell monolayer (30%-40% confluence) infected with trypomastigotes (MOI 1:10; ON incubation) 4 days post-infection. 
		NOTE: For a T-25 flask, the initial concentration of Vero cells used is 800,000 cells, and for a T-75 flask is two million cells.</li>
		<li>Determine the trypomastigote concentration using a Neubauer chamber for cell counting. Centrifuge 4 x 106 trypomastigotes at 7,000 x g at RT for 10 min.</li>
		<li>Rinse with PBS twice and resuspend in 200 µL of PBS. Apply to a round, 12 mm coverslip coated with poly-D-lysine (section 1). Incubate for 10-15 min at RT. Proceed with the crosslinking prevention step (section 4). 
		​NOTE: Alternatively, fix the trypomastigotes with cold methanol for 7 min at -20 °C or paraformaldehyde/glutaraldehyde for 10 min at RT prior to adherence of the parasites to the coverslip.</li>
	</ol>
	</li>
</ol>
4. Performing crosslinking prevention (DAY 1)
<ol>
	<li>Submerge the 12 mm coverslip with the adhered parasites or infected cells (facing upward) in a 24-well plate with 0.5 mL of CP solution (step 2.3) in each well.</li>
	<li>Fill the empty wells with water to reduce evaporation. Seal the plate with a sealing film. Incubate for 5 h at 37 °C. This step can be extended up to an ON incubation at 4 °C. 
	​NOTE: Always submerge the coverslip in the solution; do not pipette the fixative solution over the coverslips.</li>
</ol>
5. Performing gelation of the sample
<ol>
	<li>Assemble a humid chamber in a Petri dish with a sealing film on top of tissue paper (Figure 2A). Add water to the tissue paper and incubate at -20 °C for 20 min to cool down. Thaw a TEMED and an APS aliquot on ice for 20 min (step 2.6). 
	NOTE: Do not freeze-thaw the APS more than three times.</li>
	<li>Lay the cool, humid chamber on ice (Figure 2A). Take the 24-well plate prepared in section 4 out of the incubator. Aspirate the CP solution with a 3 mL Pasteur pipette, leaving some solution, otherwise the coverslips will be hard to remove.</li>
	<li>Remove the 12 mm coverslips from the fixative solution with tweezers and lay them on tissue paper with the parasites facing up. 
	NOTE: It is helpful to use a sterile needle to lift the coverslip and then hold it with the tweezers.</li>
	<li>To an aliquot of 90 µL of monomeric solution (step 2.4), add 5 µL of TEMED and 5 µL of APS previously thawed. Mix with a vortex mixer for no more than 2-3 s; it is not necessary to close the tube with the lid. 
	NOTE: Always add TEMED first and APS last. The addition of APS first and TEMED last to the monomeric solution makes the gelation process faster, giving no time to manipulate it.</li>
	<li>Quickly make one drop of 35 µL over the sealing film of the humid chamber for each coverslip. Immediately pick up the coverslip with tweezers and lay over the drop (Figure 2B) with the parasites facing down. 
	​NOTE: Make a maximum of two coverslips at a time; it is crucial not to delay this step because the solution polymerizes very quickly.</li>
	<li>Incubate the humid chamber for 5 min on ice and then for 1 h at 37 °C. Turn on a heating block at 95 °C to ensure the correct temperature for the next step.</li>
</ol>
<img alt="Cell culture in gel: Petri dish setup, diagram of gel-layered cells on coverslip." class="xfigimg" src="/files/ftp_upload/65381/65381fig02.jpg" data-altupdatedat="1747911167000" data-alt="Figure 2"> 
Figure 2: Gelation step details. (A) Assembly of the humid chamber. (B) Dropping the coverslips onto the monomer solution with TEMED and APS for gelation. (C) Schematic representation of the gel assembled for imaging. <a href="https://www.jove.com/files/ftp_upload/65381/65381fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
6. Denaturing the gelified samples and performing the isotropic expansion
<ol>
	<li>Remove the denaturation solution (step 2.5) from 4 °C. If it is precipitated, put it in a hot water bath until it dissolves completely.</li>
	<li>Add 2 mL of the denaturing solution to each well of a 6-well plate. Transfer the coverslips from step 5.3 to the plate with the denaturation solution. Incubate for 15 min at RT with gentle shaking so the gel detaches from the coverslip.</li>
	<li>Carefully transfer the gel (removing it from the 12 mm coverslip) with a metal spatula into a 1.5 mL sterile microcentrifuge tube with 1 mL of denaturing solution. Use cap locks to secure the tubes. Incubate for 1 h and 30 min at 95 °C in a heating block. 
	NOTE: Gels start to expand during this step; be gentle when transferring the gel to the 1.5 mL microcentrifuge tube. 
	CAUTION: After the incubation, the tube containing the gel is at 95 °C, which can be dangerous. Use protective gloves and let the tubes cool down before handling them to avoid burns and projections.</li>
	<li>Perform the first round of expansion
	<ol>
		<li>Aspirate the denaturing solution from the 1.5 mL microcentrifuge tube with the gel with a P1000 pipette. Transfer the gel from the 1.5 mL microcentrifuge tube to a Petri dish with 10 mL of ultrapure water for 30 min using a small spatula.</li>
		<li>Change the ultrapure water using a disposable 3 mL Pasteur pipette. Incubate ON at RT. 
		NOTE: Be gentle with the gels because, after the first 30 min of incubation, they became fragile.</li>
		<li>Change the ultrapure water using a disposable 3 mL Pasteur pipette one more time. 
		NOTE: Three water incubations of 30 min each are sufficient, but for practicality, it is better to leave the second incubation ON.</li>
	</ol>
	</li>
</ol>
7. Performing fluorescence labeling of the target proteins (DAY 2)
<ol>
	<li>Remove the water from the Petri dish with the gel with a disposable 3 mL Pasteur pipette. Measure the diameter of the gel with a caliper to calculate the expansion (between four and five times).</li>
	<li>Wash twice with 10 mL of PBS for 15 min. Cut the gel with a razor blade into squares of approximately 10 mm x 10 mm in the center of the circular gel. Use one square per condition to be tested. 
	NOTE: The gel shrinks after the PBS incubations. Gels can be stored in PBS at 4 °C for up to 1 week.</li>
	<li>Transfer each square to a 12-well plate and incubate with 500 µL of primary antibody diluted in 2% PBS-bovine serum albumin (BSA) for 2 h and 30 min at 37 °C with shaking. 
	NOTE: Alternative antibody incubation can be performed ON at 4 °C. The minimum antibody volume that can be used is 300 µL in a 24-well plate. As a general rule, use twice the concentration of antibodies used for conventional immunolabeling.</li>
	<li>Wash three times with 2 mL of PBS with 0.1% polysorbate 20 for 10 min while shaking in a 6-well plate.</li>
	<li>Transfer the gel to a 12-well plate and incubate with 500 µL of secondary antibody in PBS with DAPI and 10 µg/mL NHS-ester conjugated to the desired fluorophore for 2 h and 30 min at 37 °C with gentle shaking. 
	NOTE: As a general rule, use twice the concentration of antibodies used for conventional immunolabeling. Alternatively, this incubation can be performed ON at 4 °C.</li>
	<li>Wash three times with 2 mL of PBS with 0.1% polysorbate 20 for 10 min while shaking in a 6-well plate.</li>
	<li>Transfer the gel to a Petri dish with ultrapure water. Incubate for 30 min. Change the water twice with a disposable 3 mL Pasteur pipette, as done in step 6.4.</li>
</ol>
8. Imaging and image processing (DAY 3)
<ol>
	<li>Remove the water from the Petri dishes with a disposable 3 mL Pasteur pipette and measure the gel diameter with a caliper to calculate the expansion factor.</li>
	<li>Cut a small piece of ~10 mm x 10 mm with a razor blade and place it on a 35 mm glass bottom dish. 
	NOTE: As an alternative, place the gel between a slide and a coverslip (without poly-D-lysine). The slide should have two smaller slide pieces cut with a diamond knife attached on the sides to form a chamber about the thickness of the gel.</li>
	<li>Check the orientation at 10x or 20x magnification. In order to focus properly, ensure that the parasites are facing the coverslip; otherwise, one has to turn the gel around and check again (Figure 2C).</li>
	<li>Once the proper orientation is found, dry the remaining gel with microscopy paper. Cover the gel with a poly-D-lysine-coated coverslip.</li>
	<li>Add a small drop of ultrapure water to the gel to visualize with a confocal microscope. 
	NOTE: To ensure proper imaging, it is crucial to maintain the gel orientation throughout the process so that the side of the gel containing cells near its surface is facing the glass-bottom dish at the end. When using 35 mm dishes, adding poly-D-lysine to the coverslip placed in the bottom reduces the gel shifting during the image acquisition.</li>
	<li>Image acquisition parameters
	<ol>
		<li>Image the samples in a confocal microscope (Table of Materials) using a 63x oil immersion 1.4 numerical aperture (NA) objective.</li>
		<li>Acquire Z stacks, the width of each z step, and an exposure time per pixel, to be determined empirically depending on the sample, the signal intensity, and optimization of acquisition times. Use the scan zoom for effective magnification if desired.</li>
		<li>Open the Z stack using image processing software (Table of materials). For each channel, group the stacked images using the Group Z-project option. Select the maximum intensity projection.</li>
		<li>To merge images, use the Merge Channels tool and select the color of each channel. Add a scale bar using the Scale Bar tool in the processing software. 
		NOTE: Alternatively, one can color each channel as desired (see examples in the representative results).</li>
	</ol>
	</li>
</ol>

Ultrastructural Expansion Microscopy in In Vitro Life Cycle of Trypanosoma cruzi

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M., Kumar, P., Conwell, A. N., Wu, K., Baskin, J. M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Lipid+expansion+microscopy.">Lipid expansion microscopy.</a> Journal of the American Chemical Society. 144 (40), 18212-18217 (2022).</li><li>Steib, E., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=TissUExM+enables+quantitative+ultrastructural+analysis+in+whole+vertebrate+embryos+by+expansion+microscopy.">TissUExM enables quantitative ultrastructural analysis in whole vertebrate embryos by expansion microscopy.</a> Cell Reports Methods. 2 (10), 100311 (2022).</li><li>Shaib, A. H., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Visualizing+proteins+by+expansion+microscopy.">Visualizing proteins by expansion microscopy.</a> bioRxiv. , (2023).</li><li>M'Saad, O., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=All-optical+visualization+of+specific+molecules+in+the+ultrastructural+context+of+brain+tissue.">All-optical visualization of specific molecules in the ultrastructural context of brain tissue.</a> bioRxiv. , (2022).</li></ol>

The authors have no conflicts of interest to disclose.

Ultrastructural expansion microscopy is a technique that allows obtaining high-resolution images of biological samples by physically expanding them to several times their original size. The U-ExM protocol involves several critical steps that must be carefully executed to achieve optimal results4. First, the sample must be fixed with a CP agent and embedded in a swellable hydrogel matrix. The formaldehyde present in the CP solution interacts with the free covalent bonds of the acrylamide to prevent...

Expansion microscopy (ExM) was described for the first time in 2015 by Boyden et al.1. It is an imaging protocol with which a conventional microscope can achieve a spatial resolution below the diffraction limit. This higher resolution is obtained because of a physical enlargement of the sample. To accomplish this, fluorescently labeled molecules are crosslinked to a hydrogel, which is subsequently expanded isotropically with water. As a result of this expansion, the signals are separated nearly isotropically in all three dimensions. This method employs low-cost chemicals and enables a spatial resolution of approximately 65 nm using conventional (confocal) microscopes, which is roughly four times better than the standard resolution of a confocal microscope (approximately 250 nm)1.
The next milestone, that has enabled the use of expansion microscopy in many biological fields, was the adaptation of immunofluorescence labeling with conventional antibodies2. Another adaptation from the initially published ExM protocol is the magnified analysis of the proteome (MAP)3. This method introduced the use of high concentrations of acrylamide and paraformaldehyde prior to sample-hydrogel immersion to prevent intra- and inter-protein crosslinking, which led to better preservation of the samples&#39; protein content and subcellular architecture. This alternative protocol was optimized to obtain enhanced conservation of the overall ultrastructure of isolated organelles by utilizing lower concentrations of the fixative agents (formaldehyde/paraformaldehyde and acrylamide); this approach was termed ultrastructure expansion microscopy (U-ExM)4.
To gain even more resolution, the combination of ExM with super-resolution microscopy techniques, including stimulated emission depletion microscopy or single-molecule localization microscopy, has also been reported in order to reach resolutions below 20 nm5.
The use of ExM has been widely reported in the fields of neuroscience and cytoskeleton research6, but only a few studies have been conducted on parasitic protists. Our laboratory was the first to report the application of U-ExM in T. cruzi7. The foundation protocol is mainly based on the previous U-ExM reports in Toxoplasma gondii, Plasmodium ssp., and Trypanosoma brucei8,9,10,11.
One of the greatest advantages of ExM is its modular nature, which allows great flexibility to adapt to different biological samples. The protocol can be divided into steps (such as fixation, crosslinking prevention, or gelation) that can be easily adjusted by the user to meet their experimental requirements. Additionally, this pipeline can be modified to enhance compatibility with the model organism or to achieve a specific resolution. As a result, ExM offers tremendous potential for both advanced and non-advanced optical systems, ensuring wider applications in the future.
Chagas disease, also called American trypanosomiasis, is an endemic disease in Latin America caused by Trypanosoma cruzi, a protozoan parasite. The parasite&#39;s life cycle is complex and involves two developmental stages in mammals and two in the insect host (members of the Triatominidae family), which is the biological vector of this disease. Chagas disease belongs to the group of neglected tropical diseases listed by the World Health Organization and represents a significant economic and social problem in Latin America. Epidemiological studies estimate 8 million people around the world live with Chagas disease and over 10,000 deaths per year. These numbers exemplify the significance of Chagas disease as a public health issue worldwide. The geographic distribution of Chagas disease has changed in recent decades, with many infected individuals now residing in large urban areas globally due to increased migrations, as opposed to the primarily rural areas of Latin America where it was originally found12.
The developmental stages of T. cruzi differ throughout its life cycle, which can be replicated completely in vitro. Epimastigotes are replicative forms in the insect vector, and they have a spherical nucleus in the central region of the cell body and a bar-shaped kinetoplast (a mitochondrial DNA-containing structure unique to kinetoplastids) in the anterior region relative to the nucleus, with a free flagellum. Trypomastigotes are the infective, non-replicative form, and have an elongated nucleus, a rounded posterior kinetoplast, and a flagellum attached to the plasma membrane along the entire length of the parasite. Amastigotes are the intracellular replicative form; they have a nucleus in the central region, a rod-shaped kinetoplast in the anterior part of the cell body, and a reduced flagellum. The parasite&#39;s adaptability to different environments is a reflection of these morphological variations. It is also worth mentioning that this life cycle involves symmetrical division and different transitional developmental stages13. During differentiation, the trypanosomatids&#39; cytoskeleton plays a critical role. This structure is formed by a corset of subpellicular microtubules arranged in an ordered array of stable microtubules below the plasma membrane. Also, a paraflagellar rod is present in these organisms, which is a lattice-like structure that runs parallel and is attached to the flagellar axoneme14. The precise cytoskeletal organization and nuclear structural changes along the cell cycle stages involve unique gene regulation mechanisms specific to trypanosomatids, making them interesting models for cell biology studies.
Given the small size of T. cruzi and other protozoan parasites, U-ExM presents an excellent tool for analyzing the structural features of these important pathogens. As mentioned earlier, the applicability of this technique on T. cruzi was validated for the first time by Dr. Alonso7. This report details a complete U-ExM protocol, with emphasis on the immunolocalization of cytoskeletal proteins during the different life cycle stages of T. cruzi. Also, we have optimized the use of N-Hydroxysuccinimide ester (NHS), a pan-proteome label that enables us to mark various parasite structures. In addition, an in vitro methodology to obtain the three stages of the parasite is described.

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Aldrich</td><td>408220-25G</td><td>SA</td></tr><tr><td>Sodium Bicarbonate</td><td>Cicarelli</td><td>929211</td><td>NaHCO&lt;sub&gt;3&lt;/sub&gt;</td></tr><tr><td>Sodium Chloride</td><td>Cicarelli</td><td>750214</td><td>NaCl</td></tr><tr><td>Sodium Dodecyl Sulfate</td><td>BioRad</td><td>1610302</td><td>SDS</td></tr><tr><td>Sodium Hidroxide</td><td>Merk</td><td>1-06498</td><td>NaOH</td></tr><tr><td>Sorvall ST 16 Centrifuge</td><td>Thermo Fisher Scientific</td><td>75004380</td><td></td></tr><tr><td>T-25 flasks</td><td>Corning</td><td>430639</td><td></td></tr><tr><td>TEMED</td><td>Invitrogen</td><td>15524-010</td><td></td></tr><tr><td>Tissue paper</td><td>Elite</td><td></td><td></td></tr><tr><td>Triptose</td><td>Merck</td><td>1106760500</td><td></td></tr><tr><td>Tris</td><td>BioRad</td><td>1610719</td><td></td></tr><tr><td>Tween-20</td><td>Biopack</td><td>2003-07</td><td>Polysorbate 20</td></tr><tr><td>Vaccum pump</td><td>Silfab</td><td>N33-A</td><td></td></tr><tr><td>Vero cells</td><td>ATCC</td><td>CRL-1587</td><td></td></tr><tr><td>Vortex MIxer</td><td>Dragon Lab</td><td>MX-S</td><td></td></tr></tbody></table>

ultrastructural expansion microscopy in three in vitro life cycle stages of trypanosoma cruzi

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial resolution of a cell or tissue for microscopic imaging. This is performed by physically expanding a sample with off-the-shelf chemicals and common lab equipment.
Chagas disease is a widespread and pressing public health concern caused by T. cruzi. The disease is prevalent in Latin America and has become a significant problem in non-endemic regions due to increased migration. The transmission of T. cruzi occurs through hematophagous insect vectors belonging to the Reduviidae and Hemiptera families. Following infection, T. cruzi amastigotes multiply within the mammalian host and differentiate into trypomastigotes, the non-replicative bloodstream form. In the insect vector, trypomastigotes transform into epimastigotes and proliferate through binary fission.The differentiation between the life cycle stages requires an extensive rearrangement of the cytoskeleton and can be recreated in the lab completely using different cell culture techniques. 
We describe here a detailed protocol for the application of U-ExM in three in vitro life cycle stages of Trypanosoma cruzi, focusing on optimization of the immunolocalization of cytoskeletal proteins. We also optimized the use of N-Hydroxysuccinimide ester (NHS), a pan-proteome label that has enabled us to mark different parasite structures.

If the protocol has been properly executed (Figure 1), samples will be visible as a planar and translucent gel that can be expanded up to a factor of 4-4.5x in water (Figure 3A). This expansion provided an effective resolution of about 70 nm, which may vary depending on the final expansion factor and imaging system employed. After the second expansion process and image acquisition in a confocal microscope, we were able to observe expansion factors of around 4.5....

Read this Scientific Article Protocol about Ultrastructural Expansion Microscopy in Three In Vitro Life Cycle Stages of Trypanosoma cruzi at JoVE.com

Ultrastructural Expansion Microscopy in Three In Vitro Life Cycle Stages of Trypanosoma cruzi

This study shows a detailed protocol to perform ultrastructure expansion microscopy in three in vitro life cycle stages of Trypanosoma cruzi, the pathogen responsible for Chagas disease. We include the optimized technique for cytoskeletal proteins and pan-proteome labeling.

We describe here the application of ultrastructure expansion microscopy (U-ExM) in Trypanosoma cruzi, a technique that allows increasing the spatial ...

ultrastructural-expansion-microscopy-three-vitro-life-cycle-stages

Research

JoVE Journal

Biology

737 Views.  CONICET. Dit protocol beschrijft de methode van ultrastructuurexpansiemicroscopie in drie in vitro levenscyclusstadia van Trypanosoma cruzi, de ziekteverwekker die verantwoordelijk is voor de ziekte van Chagas. Dit protocol biedt een alternatief voor de huidige beeldvormings- en elektronenmicroscopietechnieken met superresolutie, met het voordeel dat het compatibel is met conventionele microscopen die in de meeste biologielaboratoria en in machinekernfaciliteiten word...

Video: Ultrastructurele expansiemicroscopie in drie in vitro levenscyclusstadia van Trypanosoma cruzi 

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target against T. cruzi, it is important to validate the essentiality of the target gene in the mammalian stage of the parasite, the amastigote. Amastigotes of T. cruzi replicate inside the host cell; thus, it is difficult to conduct a knockout experiment without going through other developmental stages. Recently, our group reported a growth condition in which the amastigote can replicate axenically for up to 10 days without losing its amastigote-like properties. By using this temporal axenic amastigote culture, we successfully introduced gRNAs directly into the Cas9-expressing amastigote to cause gene knockouts and analyzed their phenotypes exclusively in the amastigote stage. In this report, we describe a detailed protocol to produce in vitro derived extracellular amastigotes, and to utilize the axenic culture in a CRISPR/Cas9-mediated knockout experiment. The growth phenotype of knockout amastigotes can be evaluated either by cell counts of the axenic culture, or by replication of intracellular amastigote after host cell invasion. This method bypasses the parasite stage differentiation normally involved in producing a transgenic or a knockout amastigote. Utilization of the temporal axenic amastigote culture has the potential to expand the experimental freedom of stage-specific studies in T. cruzi.

Introducing a Gene Knockout Directly Into the Amastigote Stage of Trypanosoma cruzi Using the CRISPR/Cas9 System

Here, we describe a protocol to introduce a gene knockout into the extracellular amastigote of Trypanosoma cruzi, using the CRISPR/Cas9 system. The growth phenotype can be followed up either by cell counting of axenic amastigote culture or by proliferation of intracellular amastigotes after host cell invasion.

Trypanosoma cruzi is a pathogenic protozoan parasite that causes Chagas&#8217; disease mainly in Latin America. In order to identify a novel drug target ...

Genetics

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

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&#39;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.

Establishment of Larval Zebrafish as an Animal Model to Investigate Trypanosoma cruzi Motility In Vivo

In this protocol, fluorescently labeled T. cruzi&#160;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 ...

Developmental Biology

Purification of the M. magneticum Strain AMB-1 Magnetosome Associated Protein MamA&Delta;41

Magnetotactic bacteria comprise a diverse group of aquatic microorganisms that are able to orientate themselves along geomagnetic fields. This behavior is believed to aid their search for suitable environments (1). This capability is conferred by the magnetosome, a subcellular organelle that consists of a linear-chain assembly of lipid vesicles each able to biomineralize and enclose a ~50-nm crystal of magnetite or greigite. A principle component of the magnetosome that was shown to be required for the formation of functional vesicles is MamA. MamA is a highly abundant magnetosome-associated protein which is one of the most characterized magnetosome-associated proteins in vivo (2-6). This article focuses on the purification of MamA, which despite being studied in vivo, no clear functional or structural details have been identified for it. Bioinformatics analysis suggested that MamA is a tetra-tricopeptide repeat (TPR) containing protein. TPR is a structural motif found as such or forming part of a bigger fold in a wide range of proteins, it serves as a template for protein-protein interactions and mediates multi-protein complexes (7). TPRs are involved in many crucial tasks in eukaryotic cell organelle processes and many bacterial pathways (8-14). In order to understand MamA, a unique TPR containing protein, highly purified protein is required as a first step. In this article, we present the purification protocol for a stable MamA deletion mutant (MamA&Delta;41) from M. magneticum AMB-1.

MamA is a unique Magnetosome associated protein which was shown to be involved in magnetosome activation. Here we present the purification protocol of MamA deletion mutant (MamA&Delta;41) from M. magneticum AMB-1.

Magnetotactic bacteria comprise a diverse group of aquatic microorganisms that are able to orientate themselves along geomagnetic fields. This behavior is ...

In Vitro Drug Screening Against All Life Cycle Stages of Trypanosoma cruzi Using Parasites Expressing &#946;-galactosidase

Trypanosoma cruzi is the causative agent of Chagas disease (ChD), an endemic disease of public health importance in Latin America that also affects many non-endemic countries due to the increase in migration. This disease affects nearly 8 million people, with new cases estimated at 50,000 per year. In the 1960s and 70s, two drugs for ChD treatment were introduced: nifurtimox and benznidazole (BZN). Both are effective in newborns and during the acute phase of the disease but not in the chronic phase, and their use is associated with important side effects. These facts underscore the urgent need to intensify the search for new drugs against T. cruzi.
T. cruzi is transmitted through hematophagous insect vectors of the Reduviidae and Hemiptera families. Once in the mammalian host, it multiplies intracellularly as the non-flagellated amastigote form and differentiates into the trypomastigote, the bloodstream non-replicative infective form. Inside the insect vector, trypomastigotes transform into the epimastigote stage and multiply through binary fission.
This paper describes an assay based on measuring the activity of the cytoplasmic &#946;-galactosidase released into the culture due to parasites lysis by using the substrate, chlorophenol red &#946;-D-galactopyranoside (CPRG). For this, the T. cruzi Dm28c strain was transfected with a &#946;-galactosidase-overexpressing plasmid and used for in vitro pharmacological screening in epimastigote, trypomastigote, and amastigote stages. This paper also describes how to measure the enzymatic activity in cultured epimastigotes, infected Vero cells with amastigotes, and trypomastigotes released from the cultured cells using the reference drug, benznidazole, as an example. This colorimetric assay is easily performed and can be scaled to a high-throughput format and applied to other T. cruzi strains.

In Vitro Drug Screening Against All Life Cycle Stages of Trypanosoma cruzi Using Parasites Expressing &#946;-galactosidase

We describe a high-throughput colorimetric assay measuring &#946;-galactosidase activity in three life cycle stages of Trypanosoma cruzi, the causative agent of Chagas disease. This assay can be used to identify trypanocidal compounds in an easy, fast, and reproducible manner.

Trypanosoma cruzi is the causative agent of Chagas disease (ChD), an endemic disease of public health importance in Latin America that also affects many ...

Biochemistry

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes. The transparency of the 
embryos, the genetic resources, and the relative ease of transformation are qualities that make C. elegans an excellent model for early embryogenesis. Laser-based 
confocal microscopy and fluorescently labeled tags allow researchers to follow specific cellular structures and proteins in the developing embryo. For example, 
one can follow specific organelles, such as lysosomes or mitochondria, using fluorescently labeled dyes. These dyes can be delivered to the early embryo by means 
of microinjection into the adult gonad. Also, the localization of specific proteins can be followed using fluorescent protein tags. Examples are presented here 
demonstrating the use of a fluorescent lysosomal dye as well as fluorescently tagged histone and ubiquitin proteins. The labeled histone is used to visualize 
the DNA and thus identify the stage of the cell cycle. GFP-tagged ubiquitin reveals the dynamics of ubiquitinated vesicles in the early embryo. Observations 
of labeled lysosomes and GFP:: ubiquitin can be used to determine if there is colocalization between ubiquitinated vesicles and lysosomes. A technique for 
the microinjection of the lysosomal dye is presented. Techniques for generating transgenenic strains are presented elsewhere (1, 2). For imaging, embryos 
are cut out of adult hermaphrodite nematodes and mounted onto 2% agarose pads followed by time-lapse microscopy on a standard laser scanning confocal 
microscope or a spinning disk confocal microscope. This methodology provides for the high resolution visualization of early embryogenesis.

Time-lapse Microscopy of Early Embryogenesis in Caenorhabditis elegans

This article describes a technique for the visualization of the early events of embryogenesis in the nematode Caenorhabditis elegans.

Caenorhabditis elegans has often been used as a model system in studies of early developmental processes.  The transparency of the 
embryos, the genetic ...

Sexual Transmission of American Trypanosomes from Males and Females to Naive Mates

American trypanosomiasis is transmitted to humans by triatomine bugs through the ingestion of contaminated food, by blood transfusions or accidently in hospitals and research laboratories. In addition, the Trypanosoma cruzi infection is transmitted congenitally from a&#160;chagasic mother to her offspring, but the male partner&#39;s contribution to in utero contamination is unknown. The findings of nests and clumps of amastigotes and of trypomastigotes in the theca cells of the ovary, in the goniablasts and in the lumen of seminiferous tubules suggest that T. cruzi infections are sexually transmitted. The research protocol herein presents the results of a family study population showing parasite nuclear DNA in the diploid blood mononuclear cells and in the haploid gametes of human subjects. Thus, three independent biological samples collected one year apart confirmed that T. cruzi infections were sexually transmitted to progeny. Interestingly, the specific T. cruzi antibody was absent in the majority of family progeny that bore immune tolerance to the parasite antigen. Immune tolerance was demonstrated in chicken refractory to T. cruzi after the first week of embryonic growth, and chicks hatched from the flagellate-inoculated eggs were unable to produce the specific antibody.&#160;Moreover, the instillation of the human semen ejaculates intraperitoneally or into the vagina of naive mice yielded T. cruzi amastigotes in the epididymis, seminiferous tubule, vas deferens and uterine tube with an absence of inflammatory reactions in the immune privileged organs of reproduction. The breeding of T. cruzi-infected male and female mice with naive mates resulted in acquisition of the infections, which were later transmitted to the progeny. Therefore, a robust education, information and communication program that involves the population and social organizations is deemed necessary to prevent Chagas disease.

The Trypanosoma cruzi agent of Chagas disease produces long-lasting asymptomatic infections that abruptly develop into clinically recognized pathology. The following research protocol describes a short-run family-based epidemiological study to unravel the T. cruzi infection transmitted sexually from parent to progeny.

American trypanosomiasis is transmitted to humans by triatomine bugs through the ingestion of contaminated food, by blood transfusions or accidently in ...

Immunology and Infection

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the messagefor review see 1. However, it's now widely accepted that synonymous codon usage is the primary cause of non-uniform translational elongation rates1. Synonymous codons are not used with identical frequency. A bias exists in the use of synonymous codons with some codons used more frequently than others2. Codon bias is organism as well as tissue specific2,3. Moreover, frequency of codon usage is directly proportional to the concentrations of cognate tRNAs4. Thus, a frequently used codon will have higher multitude of corresponding tRNAs, which further implies that a frequent codon will be translated faster than an infrequent one. Thus, regions on mRNA enriched in rare codons (potential pause sites) will as a rule slow down ribosome movement on the message and cause accumulation of nascent peptides of the respective sizes5-8. These pause sites can have functional impact on the protein expression, mRNA stability and protein foldingfor review see 9. Indeed, it was shown that alleviation of such pause sites can alter ribosome movement on mRNA and subsequently may affect the efficiency of co-translational (in vivo) protein folding1,7,10,11. To understand the process of protein folding in vivo, in the cell, that is ultimately coupled to the process of protein synthesis it is essential to gain comprehensive insights into the impact of codon usage/tRNA content on the movement of ribosomes along mRNA during translational elongation.
Here we describe a simple technique that can be used to locate major translation pause sites for a given mRNA translated in various cell-free systems6-8. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA. The rationale is that at low-frequency codons, the increase in the residence time of the ribosomes results in increased amounts of nascent peptides of the corresponding sizes. In vitro transcribed mRNA is used for in vitro translational reactions in the presence of radioactively labeled amino acids to allow the detection of the nascent chains. In order to isolate ribosome bound nascent polypeptide complexes the translation reaction is layered on top of 30% glycerol solution followed by centrifugation. Nascent polypeptides in polysomal pellet are further treated with ribonuclease A and resolved by SDS PAGE. This technique can be potentially used for any protein and allows analysis of ribosome movement along mRNA and the detection of the major pause sites. Additionally, this protocol can be adapted to study factors and conditions that can alter ribosome movement and thus potentially can also alter the function/conformation of the protein.

Isolation of Ribosome Bound Nascent Polypeptides in vitro to Identify Translational Pause Sites Along mRNA

A technique to identify translational pause sites on mRNA is described. This procedure is based on isolation of nascent polypeptides accumulating on ribosomes during in vitro translation of a target mRNA, followed by the size analysis of the nascent chains using a denaturing gel electrophoresis.

The rate of translational elongation is non-uniform. mRNA secondary structure, codon usage and mRNA associated proteins may alter ribosome movement on the ...

Quantitative 3D Imaging of Trypanosoma cruzi-Infected Cells, Dormant Amastigotes, and T Cells in Intact Clarified Organs

Chagas disease is a neglected pathology that affects millions of people worldwide, mainly in Latin America. The Chagas disease agent, Trypanosoma cruzi (T. cruzi), is an obligate intracellular parasite with a diverse biology that infects several mammalian species, including humans, causing cardiac and digestive pathologies. Reliable detection of T. cruzi in vivo infections has long been needed to understand Chagas disease&#39;s complex biology and accurately evaluate the outcome of treatment regimens. The current protocol demonstrates an integrated pipeline for automated quantification of T. cruzi-infected cells in 3D-reconstructed, cleared organs. Light-sheet fluorescent microscopy allows for accurately visualizing and quantifying of actively proliferating and dormant T. cruzi parasites and immune effector cells in whole organs or tissues. Also, the CUBIC-HistoVision pipeline to obtain uniform labeling of cleared organs with antibodies and nuclear stains was successfully adopted. Tissue clearing coupled with 3D immunostaining provides an unbiased approach to comprehensively evaluate drug treatment protocols, improve the understanding of the cellular organization of T. cruzi-infected tissues, and is expected to advance discoveries related to anti-T. cruzi immune responses, tissue damage, and repair in Chagas disease.

Quantitative 3D Imaging of Trypanosoma cruzi-Infected Cells, Dormant Amastigotes, and T Cells in Intact Clarified Organs

The present protocol describes light-sheet fluorescent microscopy and automated software-assisted methods to visualize and precisely quantify proliferating and dormant Trypanosoma cruzi parasites and T cells in intact, cleared organs and tissues. These techniques provide a reliable way to evaluate treatment outcomes and offer new insights into parasite-host interactions.

Chagas disease is a neglected pathology that affects millions of people worldwide, mainly in Latin America. The Chagas disease agent, Trypanosoma cruzi ...

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) maintain the turnover of all mature blood cell lineages. The coordination of the complex signals leading to specific HSC fates relies upon the interaction between HSCs and the intricate bone marrow microenvironment, which is still poorly understood[1-2].
We describe how by combining a newly developed specimen holder for stable animal positioning with multi-step confocal and two-photon in vivo imaging techniques, it is possible to obtain high-resolution 3D stacks containing HSPCs and their surrounding niches and to monitor them over time through multi-point time-lapse imaging. High definition imaging allows detecting ex vivo labeled hematopoietic stem and progenitor cells (HSPCs) residing within the bone marrow. Moreover, multi-point time-lapse 3D imaging, obtained with faster acquisition settings, provides accurate information about HSPC movement and the reciprocal interactions between HSPCs and stroma cells.
Tracking of HSPCs in relation to GFP positive osteoblastic cells is shown as an exemplary application of this method. This technique can be utilized to track any appropriately labeled hematopoietic or stromal cell of interest within the mouse calvarium bone marrow space.

In Vivo 4-Dimensional Tracking of Hematopoietic Stem and Progenitor Cells in Adult Mouse Calvarial Bone Marrow

The nature of the interactions between hematopoietic stem and progenitor cells (HSPCs) and bone marrow niches is poorly understood. Custom hardware modifications and a multi-step acquisition protocol allow the use of two-photon and confocal microscopy to image ex vivo labeled HSPCs homed within bone marrow areas, tracking interactions and movement.

Through a delicate balance between quiescence and proliferation, self renewal and production of differentiated progeny, hematopoietic stem cells (HSCs) ...

In Vivo Bioluminescent Model of Chagas Disease for Drug Evaluation

Demystifying In Vivo Bioluminescence Imaging of a Chagas Disease Mouse Model for Drug Efficacy Studies

To control and decrease the public health impact of human protozoan diseases such as Chagas disease, leishmaniasis, and human African trypanosomiasis, expediting the development of new drugs and vaccines is necessary. However, this process is filled with difficulties such as highly complex parasite biology and disease pathogenesis and, as typical for neglected tropical diseases, comparatively limited funding for research and development. Thus, in vitro and in vivo study models that can sufficiently reproduce infection and disease key features while providing rational use of resources are essential for progressing research for these conditions. One example is the in vivo bioluminescence imaging (BLI) mouse model for Chagas disease, which provides highly sensitive detection of long wavelength light generated by Trypanosoma cruzi parasites expressing luciferase. Despite this technique becoming the standard approach for drug efficacy in vivo studies, research groups might still struggle to implement it due to a lack of proper practical training on equipment handling and application of quality control procedures, even when suitable BLI equipment is readily available. Considering this scenario, this protocol aims to guide from planning experiments to data acquisition and analysis, with details that facilitate the implementation of protocols in research groups with little or no experience with BLI, either for Chagas disease or for other infectious disease mouse models.

Author Spotlight: Real-Time Monitoring of Parasite Burden and Host Response

The protocol provided here describes the detailed steps to perform drug efficacy studies in a bioluminescent model of Trypanosoma cruzi infection, with a focus on data acquisition, analysis, and interpretation. Troubleshooting and quality control procedures to minimize technical issues are also provided.

To control and decrease the public health impact of human protozoan diseases such as Chagas disease, leishmaniasis, and human African trypanosomiasis, ...

Ultrastructural Expansion Microscopy in Three In Vitro Life Cycle Stages of Trypanosoma cruzi

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