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

All procedures described in this protocol were submitted, approved, and conducted according to the guidelines¹⁴ preconized by the Animal Ethics Committee of Instituto de Ciências Biomédicas at Universidade de São Paulo: protocol CEUA ICB/USP no 5787250522.

1. Solutions

NOTE: Consider the preconized administrated volume of 10 mL/kg (200 µL for a mouse 20 g weight)^15,16. For instance, prepare a 15 mg/mL working solution to reach 150 mg/kg of animal dosing.

1. Hydroxypropylmethylcellulose suspension vehicle (HPMC-SV)
   NOTE: The reagents required are 0.5% (w/v) Hydroxypropylmethylcellulose (HPMC), 0.4% (v/v) Tween 80, and 0.5% (v/v) Benzyl alcohol.
   1. For 200 mL of vehicle solution, weigh 1 g of HPMC and dissolve in 64 mL of hot ultrapure water. Stir for 2 min.
   2. Add 120 mL of ice-cold ultrapure water and stir for 1 h. Add 1 mL of benzyl alcohol.
   3. Then add 0.8 mL of Tween 80 using the reverse pipetting technique and keep stirring until the solution becomes transparent.
   4. Adjust the solution for the final volume of 200 mL. Store the HPMC-SV under refrigeration (4 °C) for a maximum of 3 months.
2. Benznidazole
   1. Calculate the necessary amount of compound solution, as follows:
      volume of compound solution = (number of doses in each day x number of days x number of mice to be treated x administered volume in each mouse) + 30% extra.
   2. Weigh the necessary amount of benznidazole (BZ), considering the desired dose. For curative treatment, an oral dose of 100 mg/kg once daily for 10 days achieves a 100% cure in the CD chronic model¹⁷.
   3. Considering the desired final volume, calculate the proper amount to reach a concentration of 5% (v/v) DMSO and completely dissolve BZ in neat DMSO.
   4. Add the proper volume of the vehicle to a glass tube to reach a concentration of 95% (v/v) HPMC-SV. Transfer the BZ dissolved in DMSO to the glass tube containing HPMC-SV. Vigorously homogenize the suspension and store it at 4 °C.
      E.g.: Final volume of the BZ formulation = 10 mL (step 1.2.1)
      Amount of BZ = 100 mg (step 1.2.2)
      ​5% DMSO = 0.5 mL (step 1.2.3) and 95% HPMC-SV = 9.5 mL (step 1.2.4)
   5. Before each drug administration, the formulations are placed in an ultrasonic water bath for 5 min at 37 °C and vigorously homogenized before the animal dosing.
3. Cyclophosphamide
   1. Dissolve the necessary amount of cyclophosphamide in ultrapure water to achieve a 12.5 mg/mL solution.
   2. Sterilize the solution by filtration and store at 4 °C.
4. Giemsa stain
   1. Dissolve 0.6 mg of Giemsa powder reagent in 50 mL of methanol. Add 25 mL of glycerol and mix. Remove precipitates using filter paper and a funnel. Store the stock solution at room temperature (RT), protected from light.
   2. Prepare a working solution by diluting 10 mL of Giemsa solution in 90 mL of Mix-Phosphate Buffer (20.5 M Na₂HPO₄, 65.4 M KH₂PO₄ at pH 7.2).

2. Trypanosoma cruzi culture

1. Working in a biological safety cabinet (BSC), cultivate 4 x 10⁶ epimastigotes/mL of T. cruzi CL Brener Luc::Neon¹¹ in LIT media (68.44 mM NaCl, 5.36 mM KCl, 112.7 mM Na₂HPO₄, 5 g/L tryptone, 5 g/L liver infusion broth, 0.03 M hemin, 4.16 mM glucose), supplemented with 10% (v/v) Fetal Bovine Serum (FBS), 100 µg/mL penicillin, 100 U/mL streptomycin and 150 µg/mL hygromycin as selective drug at 28 °C. The parasites typically achieve a stationary phase in 3-4 days.
2. Induce metacyclogenesis to obtain infectious parasites by incubating 3 x 10⁸ epimastigotes in 10 mL of Grace's Insect medium with 10% (v/v) FBS in 15 mL conical tubes at 28 ˚C for 7-10 days.
3. Evaluate the differentiation rate by Giemsa-staining smears. For that, spread 20 µL of the differentiated parasites in a glass slide and let dry for 10 min.
4. In a fume hood, proceed with the sample fixation by covering the glass slide with neat methanol and let dry completely (1-2 min). Cover the glass slide with Giemsa working solution for 20 min.
5. Gently wash the glass slide with distilled water. In a microscope, count the percentage of metacyclic trypomastigotes. If a differentiation rate of at least 10 percent is achieved, proceed to the next step.
6. Centrifuge the parasites (120 x g for 10 min) and resuspend the pellet with 10 mL of DPBS. Centrifuge once more and resuspend the parasites in 5 mL of DMEM supplemented with 10% (v/v) of FBS, 100 µg/mL penicillin, and 100 U/mL streptomycin.
7. In a 25 cm² culture flask, perform the infection of 1.66 x 10⁵ LLC-MK2 cells (kidney epithelial cells from Macaca mulatta) at 37 °C in 5% CO₂, in a humidified incubator¹⁸.
8. Tissue-culture trypomastigotes (TCTs) are released in the medium after 8-9 days. Infect fresh LLC-MK2 cells monolayers with TCTs at a multiplicity of infection (MOI) of 1:40.

3. Analysis of Trypanosoma cruzi population homogeneity by flow cytometer

1. Collect TCTs of CL Brener Luc::Neon and CL Brener wild-type strains, centrifuge at 120 x g for 10 min, and resuspend the TCTs in DPBS. Adjust the parasite density to achieve 1 x 10⁶/mL.
2. Proceed with population analysis by flow cytometer based on mNeonGreen fluorescence (ex. 506/em. 517 nm)¹⁹, acquiring at least 20,000 events for each sample.
   NOTE: The fluorescent population percentage is obtained by comparing the wild type against the transfected parasite. To proceed with the mice infection, a minimum of 95% of the fluorescent parasite population of the CL Brener Luc::Neon lineage should be achieved (Supplementary Figure 1).

4. Mice experimental infection

NOTE: To increase the amount of parasites, bloodstream trypomastigotes (BT) are frequently obtained from immunodeficient mice^10,13. Here, chemical immunosuppression of BALB/C mice is used to obtain BT. For that, mild immunosuppression is achieved with four intraperitoneal injections of cyclophosphamide (CTX) at 62.5 mg/kg with 96 h intervals, which is performed concomitantly to T. cruzi infection.

1. Administer 62.5 mg/kg of sterile cyclophosphamide by intraperitoneal injection (i.p.) 1 day before the T. cruzi infection.
2. Infect each mouse with 1 x 10⁴ TCTs in 0.2 mL of DPBS through the intraperitoneal route. Attention to the high risk of incidents when handling pathogens and needles. Perform the infection in a BSC, wearing gloves and a face shield during the entire procedure.
3. Monitor the parasitemia daily by direct observation of the BT in the blood (Pizzi-Brener method)²⁰. At the parasitemia peak, around 13-17 days post-infection (dpi), conduct the blood collection. For that, anesthetize mice with 100 mg/kg ketamine and 10 mg/kg xylazine.
4. When mice are completely anesthetized²¹, proceed with the cardiac puncture using a 1 mL syringe attached to a 24 G X¾ needle containing 50 µL of 3.8% (w/v) sodium citrate. Unplug the needle and gently place the blood in a centrifuge tube.
5. In a BSC, perform a dilution of an aliquot of blood in ammonium-chloride-potassium (ACK) lysis buffer and count the parasites in the Neubauer chamber. Repeat this procedure at least twice, as small blood clots influence parasite counting.
   NOTE: To prevent counting errors, adjust the dilution factor to avoid counts lower than 30 and higher than 300 trypomastigotes in the Neubauer chamber.
6. Adjust the parasite density to 5 x 10³ BT/mL by mixing the necessary amount of pure blood in DPBS.
7. Infect non-immunosuppressed BALB/c mice with 1 x 10³ BT in 0.2 mL of DPBS per mouse (5 x 10³ BT/mL) by intraperitoneal route using a 26 G needle (narrower needles lead to parasite lysis)²². After injecting the volume intraperitoneally, hold the needle inside the mouse for 5 s to avoid backflow.
8. Place the infected mice temporarily in a box with a paper towel to identify potential leaking or bleeding and then return mice to their cages.

5. In vivo imaging

NOTE: Glossary of the terms used here are as follows:

Imaging session: Bioluminescence acquisition performed in all groups of a certain experiment on a given day. Figure 1A shows an overview of the procedure.

Imaging round: Procedure performed in subgroups of three mice, from D-luciferin injection to anesthesia recovery. It is recommended that each experimental group contain 6 mice, due to the mice´s intrinsic variability of parasite load and other factors that influence BLI quantification described below.

Imaging acquisition: Photoshooting performed by the imaging equipment to quantify bioluminescence, which results in an image overlay of a photograph and the quantified bioluminescence shown as a pseudocolor scale.

Imaging procedure: All steps discussed in this session of the protocol.

Figure 1: Acquiring BLI data. (A) Diagram of acquisition workflow applied for infectious diseases studies, using the Chagas disease bioluminescent mouse model as an example. Mice infected with genetically modified T. cruzi are analyzed by in vivo imaging on the established time points. On each imaging round, groups of up to three mice are injected with 150 mg/kg of enzyme-substrate (D-luciferin). After 5 min, anesthesia is administered at 2.5% (v/v) isoflurane in oxygen. When completely motionless, mice are placed in an imaging system, and acquisition is started following the defined settings. After imaging, mice recover from anesthesia and are returned to cages. Frequent questions and issues that the researchers might have and face during acquisition are highlighted in red. This scheme has been modified from Lewis et al. (2014)¹² (created with BioRender.com: UD26KWEVS2). (B) In vivo kinetic of D-luciferin/red-shifted firefly luciferase (PpyRE9h). Mice at the T. cruzi parasitemia peak (n = 3) were anesthetized and injected with 150 mg/kg D-luciferin. Images were acquired for 1 h (exposure time: 2 min; binning: 4). Top: ventral total flux (p/s) quantification (left Y-axis, in red) and as percentage of highest mean measurement (right Y-axis, in blue). Data are shown as mean (curves) and standard deviation (shaded area). Bottom: Acquired images of the first (4 min), highest BLI signal (30 min), and last (62 min) time points. Please click here to view a larger version of this figure.

1. Check the green bar in the Acquisition window to verify if the charge-coupled device (CCD) camera is cold (-90 °C). Click on the bar (green or red) to visualize the temperature.
   NOTE: According to the imaging equipment manual, the imaging equipment and software should be turned on all the time to keep the CCD cold.
2. Verify if the auto background adjustment was performed. To do so, click on Acquisition > Background > View available Dark Charge. A list with several setting combinations will appear.
   NOTE: This procedure reduces luminescence noise. If the chosen configuration results in a value higher than 1000, based on the formula "exposure time x (binning²)", then dark charge should not be ignored. Depending on the software configurations, binning can be shown as a number (binning factor) or a range from ‘small’ to ‘large’. The correspondence between the range and the binning factor can be found in the device manual.
3. If the Activity Window (white area at the bottom of the software) is not shown, enable it on Menu View > Activity Window.
4. Set the imaging area by clicking on Acquisition Window > Field of View. Option C (13.2 cm) acquires 3 mice, and Option D (22 cm) acquires 5 mice.
5. Select the autosave option by clicking on Acquisition > Auto-Save and select or create a new folder for each time point.
6. Lay a matte black paper on the imaging area and place the partitioning slides perpendicularly between the inlets. If the manufacturer did not provide the slides, use pieces (3 cm x 15 cm) of matte black paper.
7. Adjust the internal area of the imaging chamber to accommodate the glass nose cones to the anesthesia inlets. Use small pieces of black electrical tape to attach the anesthesia system to the platform (outside the image area).
   NOTE: In some versions of imaging devices, a laser defines the imaging area. If this tool is unavailable, acquire images through the software to check the partitioning slides within the imaging area.
8. Set the isoflurane vaporizer to 2.5% (v/v) in oxygen for the imaging and induction chambers. Check by the pressure gauge if the gas is flowing through the system²¹.
   NOTE: If the imaging device does not have an inhalation anesthesia system attached to it, other anesthesia options are possible, such as injectable xylazine and ketamine²³. In this case, mice recovery is slow, and the risk of death is high²⁴.
9. Fill syringes (31 G needles) with D-luciferin (a comprehensive protocol to prepare the D-luciferin is provided by different suppliers)^25,26, protect them from direct light, and prepare a few plastic containers or extra cages by covering the interior with a paper towel.
10. Keep a clock/watch nearby to easily track time and a lab notebook to register mice groups, luciferin injection time, and any other information during the procedure.
11. Weigh all mice and register their weight. Mark each mouse with a pen marker or tattoo on the tail. Keep mice identification well displayed during the entire experiment. This step can also be performed on the day prior to the acquisition.
    NOTE: The mark should not occupy a large body area because the ink may interfere with the bioluminescent signal. Permanent markers of different colors can also help to differentiate the groups.
12. To start the Imaging session, register a background luminescent image (settings: Exposure Time: 5 min; Binning: 'large' or '16', f/stop: 1) of the imaging chamber without mice. This procedure helps to identify auto luminescent light sources or undesired issues on the equipment.
    NOTE: Because BLI is based on enzymatic reaction, time is an essential feature of the experiment. Check all the items listed above and equipment settings before starting animal handling.
13. Inject 150 mg/kg (i.p.) of D-luciferin in the first 3 mice without reaching internal organs. Place the mice in the container box to assess whether D-luciferin leaks (bright yellow solution). Take notes if this happens and register the mice group and D-luciferin injection time.
14. Wait 5 min then transfer the mice to the anesthesia induction chamber. Turn on the system. It takes around 3 min for mice to be fully anesthetized (absence of responses like shaking legs, tail, etc.)²¹.
15. Open the imaging device's door, turn on the anesthetic flow to the imaging chamber, and, at the same time, turn off the induction chamber.
16. Place each mouse in the nose cones position: from mouse 1-3, from left to right side. Gently accommodate the mice with ventral side up inside the imaging area shown by the laser. During the accommodation, verify the mice's ID and position. Do not force the entrance of mice partially anesthetized in nose cones.
17. Place the partitioning slides between mice and close the imaging equipment's door.
18. Adjust the acquisition settings on the software Acquisition window, as follows:
    1. For non-infected mice, Exposure Time = 5 min / Binning = 'large' or '16'
       (most mathematically sensitive set of configurations to set the threshold values).
    2. For infected mice in the acute model: Exposure Time = 2 min/ Binning = 'medium' or '4'.
    3. For infected mice in the chronic model: Exposure Time = 5 min/ Binning = 'large' or '16'.
19. Verify if the D-luciferin injection time was performed at least 10 min before starting the acquisition. If not, wait the necessary period to obtain the images during the peak of the luciferase signal (Figure 1B); otherwise, click on Acquire to begin the imaging acquisition.
20. The Image Labels window will appear after the imaging starts. Fill the boxes with the appropriate information about each group, including luciferin injection time, experiment, mice identification, time point, and any other desired information that can help in keeping record. This information will be included in the measurements table file.
21. Verify whether the warning sign Saturated Image is shown at the end of the imaging acquisition. Saturated images are not acceptable. If this happens, decrease the binning and acquire a new image. Take notes to consider in the analysis.
22. Once the image is acquired, open the imaging machine door and turn the mice to the dorsal view (back side up). Perform the acquisition again and label it appropriately.
23. At the end of this imaging acquisition, turn off the anesthetic flow. Gently remove the mice from the imaging chamber and place them in a container.
24. Record the mice's body weight while observing the anesthesia recovery. Record any abnormal behavior or body abnormalities found during the procedure. Once the mice return to moving, place them in their cages.
    NOTE: Verify the BLI intensity for each acquired image. After the acquisition ends, the recorded bioluminescent image and the photograph will be displayed on the software with an automatic scale that should not be considered. Set the scale defined in data analysis (described below) on Tool Palette window.

6. Data analysis

NOTE: The above protocol is based on commercial in vivo imaging software. However, a software license-free version can perform the most basic analysis. Software details can be found in the Table of Materials.

1. Open files.
   1. Open the folder that contains the acquired data from a certain imaging time point by selecting Menu File > Browser > Select the folder (Figure 2A – Step 1).
      NOTE: A new window will open in a table format with all acquired data saved in the chosen file. It will display the Click Number sign, which is the image ID automatically given by the software (data and hour numbers), the information provided by the researcher in the Image Labels window during the acquisition, and acquisition settings. All these data help sort the images that can be used in the analysis, given that saturated images must not be used.
   2. Select multiple rows related to the chosen images, including the non-infected controls. Write down the image ID in the lab notebook along with the information registered during the imaging session.
   3. Assemble the selected images by clicking on Load as Group. This procedure creates a new sequence image. Therefore, it is possible to adjust features for all images simultaneously. Save the new sequence image in a new folder different from the one used in step 6.1 to facilitate the reanalysis of raw data if needed.
   4. In the sequence image, identify the binning used for each image by clicking on Options > Display > Select Binning Factor (Figure 2A – Step 2). The acquired binning factor will be displayed at the top of each image in the sequence.
   5. Correct all binning factors to the same number. To do so, double-click on the image to be adjusted. On the Tool Palette window > Image Adjust section, choose the appropriate binning factor (Figure 2A – Step 3) – see step 5.18 for binning factor choice. Perform this procedure in all divergent images.
      NOTE: This procedure directly influences BLI quantification (Table 1) and is further addressed in the Discussion section.
2. Set scale.
   1. Distinguish valid bioluminescent signal (above 600 counts according to the device manual) based on the non-infected control. For that, double-click on the image of the non-infected mice. Then, in the upper-left corner of the new window, select the option Counts in the Units field. Then, adjust the Color Scale for the minimum value of 600. The resulting image will be the baseline to set the minimum scale in radiance.
   2. With the sequence window active, select the option Radiance in the Units field. Adjust the Color Scale and Color Table in the Tool Palette. For that, disable the box Individual in the Color Scale Limits area.
   3. Mark the boxes Logarithmic Scale and Manual (Figure 2A – Step 4). Set the minimum scale numbers according to the non-infected control (as observed in step 6.8) and the area with the highest signal as the maximum (as shown in Figure 2A by the arrows).
      NOTE: In vivo 2D light mesurament is a relative quantification. Different scale values can be found due to differences in each device’s version and calibration. Therefore, the values demonstrated in the representative results could not fit other experiments performed on other devices and yet be equally valid according to the internal controls (non-infected and infected not treated).
   4. After setting the same scale for all images, double-click on each image, maximize the window, and export the image view in an image format (.jpg, .tiff, etc.) by using the Export Graphics button, naming the files by treatment_miceID_time point. (Figure 2A – Step 5). Perform this procedure for all images.
3. Perform measurements.
   1. Choose any image from the sequence and double-click on it. On the Tool Palette window > ROI tools section, click on the Square button (Figure 2B– Step 1) and draw a rectangle covering the whole mouse.
   2. Click on the border of the created ROI and copy and paste the same ROI for each mouse (resulting in three ROIs in the image). Save the ROIs by clicking save on the ROI Tools Section. Apply the saved ROIs for all images by selecting the box Apply to Sequence, then click on Load. The saved ROIs will be used for all mice in the entire experiment.
   3. Adjust the ROI position in each mouse to fit the animal better in the measurement area. When all mice are labeled, click on the Measure ROIs button (Figure 2B– Step 2). The ROI Measurements Table will appear. Select Measurement Types: Radiance; Image Attributes: All Possible Values; ROI Dimensions: cm. Then, click on Select All button, followed by Copy button.
   4. Paste the data directly into a table analysis software (spreadsheet). Alternatively, export a file in .csv or .txt format.
4. Work on the data.
   1. In table analysis software, organize the data by groups (treatment, control non-infected, non-treated, etc.).
   2. Parasite load is shown as whole-body bioluminescence. Therefore, arrange the data to match the same mice's ventral and dorsal Total Flux values and sum them.
   3. Calculate the mean and standard deviation of the summed values considering each group (e.g.: Supplementary Table 1). Plot the data into graphing and statistics software.
   4. Prepare a panel with bioluminescent images in an image or slide presentation software. Place each mouse in a column and each time point in the row to build a drug efficacy matrix to facilitate visualization and interpretation of the data.
      NOTE: There is an option to build the images directly into imaging software without performing step 6.2.4 (View > Image Layout Window). However, the software interface and tools limit data organization.

Figure 2: Data analysis steps from setting image scales to quantifying luminescence. (A) Living Image Software view for image data processing. Step 1: Upload the acquired data as a sequence using the Browser tool. Step 2: Identify the binning used for each acquisition (will appear on top of the individual images). Step 3: Set all images of infected mice to the same binning factor and apply binning factor 16 for the non-infected mice. Step 4: Set the color scale manually based on non-infected mice (purple arrow), which should be seen on the screen as non-bioluminescent, and infected and non-treated mice at the highest bioluminescent foci (red arrow), which should be seen as red. Step 5: To obtain an image free of any written information, double-click on each image and export the image using the Export Graphics button. (B) View of Regions of Interest (ROI) Tools. Step 1: Draw an ROI covering entirely a mouse and copy and paste the same ROI for each animal. Save the adjusted ROIs and apply them to all images in the experiment. Step 2: Click on the Measure ROIs button to generate the table to export as .csv or .txt. Please click here to view a larger version of this figure.

7. Ex vivo procedure

1. Inject 150 mg/kg D-luciferin i.p. in the mouse (one per ex vivo round) and wait for 3 min. Anesthetize the mice using 100 mg/kg ketamine and 10 mg/kg xylazine i.p.
2. In a biological safety cabinet, when the mouse is completely unresponsive to pinching of the paws, cut the mouse skin to expose the peritoneum (without opening the peritoneum cavity) and the tissue of the left-side axillary. Proceed with the exsanguination by cutting the axillary artery and vessel.
3. Collect the blood using a disposable Pasteur Pipette. Then perfuse 10 mL of 0.3 mg/mL of D-luciferin in DPBS solution through the heart´s left ventricle.
4. Lay out the selected organs on pre-established spots on a Petri dish and soak them in 0.3 mg/mL d-luciferin solution. This procedure must be performed within 30 min maximum.
5. Acquire the luminescent image of the excised organs (heart, lungs, thymus, liver, quadriceps muscle, mesentery, spleen, kidney, adrenal gland, visceral fat, esophagus, stomach, small intestine, cecum, colon, brain, subcutaneous fat, uterus, gonadal fat, bladder, and peritoneum membrane) using 5 min of exposure time and binning 16, depending on signal intensity (saturated images are not acceptable)²².
   NOTE: The same procedure must be performed in a non-infected mouse prior to the infected ones to set the threshold.