We thank John Kelly and Martin Taylor (London School of Hygiene &#38; Tropical Medicine) for providing T. b. brucei GVR35-VSL-2 and Dr. Andrea Zelmer (LSHTM) for advice on in vivo imaging. This work was supported by the Bill and Melinda Gates Foundation Global Health Program (grant number OPPGH5337).

Ethics 
All work was carried out under the approval of the UK Home Office Animals (Scientific Procedures) Act 1986 and the London School of Hygiene &#38; Tropical Medicine Animal Welfare and Ethics Review Board. ARRIVE guideline are followed in this report.
1. In Vivo Passage of Bioluminescent Trypanosoma brucei brucei 
<ol>
	<li>Remove a cryopreserved stock (termed stabilate) of T. b. brucei strain GVR35-VSL-2 (available from Professor John Kelly at the London School of Hygiene &#38; Tropical Medicine through MTA)9 from liquid nitrogen and allow to equilibrate to room temperature. Dilute stabilate to 2 x 104 trypanosomes/ml in sterile phosphate buffered glucose saline (PBS-G) pH 8.0 (1 L PBS + 15 g glucose)10. 
	Note: T. b. brucei GVR35-VSL-2 is non-infectious to humans and requires SAPO licensed facilities to carry out experiments.</li>
	<li>Inject a 20-25 g female CD1 mouse with 0.2 ml diluted trypanosomes via the intraperitoneal route (i.p.) with a 25 G needle. This is the &#34;donor mouse&#34;. Place the infected mouse into a Specific Pathogen Free (SPF)-cage and feed ad libitum. 
	Note: CD1 mice are an outbred strain and T. b. brucei GVR35 infections are well-characterized in this strain. However, this protocol is also applicable to inbred or genetically-modified strains, but it is important to note that fur color may impact on the sensitivity of imaging6,9.</li>
	<li>Monitor peripheral parasitemia by blood sampling. Place the mouse into a plastic restrainer and take 5 &#181;l of blood from the tail by venepuncture with 21 G needle or venesection and mix 1/10 with 0.85% sterile ammonium chloride to lyse red blood cells and allow easier counting.</li>
	<li>Count the Diluted Blood in a Neubauer Hemocytometer.
	<ol>
		<li>Load the diluted blood into both chambers. Count the entire central grid of one chamber to give n trypanosomes. Concentration of trypanosomes in blood = n x 104 trypanosomes/ml, adjust for any dilution factor and calculate the average of the two chambers. 
		Note: After approximately 5-7 days post-infection, peripheral parasitemia should rise sufficiently to carry out an infection at a concentration of 2 x 104 trypanosomes/ml.</li>
	</ol>
	</li>
	<li>When trypanosome levels are high enough for the number of mice required, move on to step 1.6. A parasitemia level of 6 x 104 trypanosomes/ml in one donor mouse is sufficient for infection of 10 mice. 
	Note: For statistically significant group sizes, n = 6 for each treatment group of mice are needed8. For other models of infection, power calculations would need to be applied to determine appropriate group sizes.</li>
	<li>Place mice into a heat box (at 28-30 &#176;C for approximately 5-10 min) to allow the tail veins to dilate. To induce terminal anaesthesia, administer 20 mg/kg pentobarbital i.v with a 25 G needle. Confirm mouse is anesthetized by gently pressing the footpads to ensure there is no withdrawal reflex.</li>
	<li>Heparinize a 21 G needle by aliquoting 5 ml heparin at 5,000 USP units/ml into a Bijou tube and sucking twice in and out of the needle attached to a 1 ml syringe.</li>
	<li>Take the heparinized 21 G needle and 1 ml syringe, and perform cardiac puncture on the anesthetized passage mouse by inserting the needle centrally underneath the rib cage (a bead of blood will pool in the base of the needle) and gently pull back on the plunger so as not to collapse the chamber of the heart. Collect a minimum of 0.7 ml of blood.</li>
	<li>Dilute infected blood to produce an inoculum of 2 x 104 trypanosomes/ml in PBS-G pH 8.0 as step 1.1 above. 
	Note: At this point, the remaining blood can be cryopreserved to produce stabilate by mixing with 8% glycerol, 20% heat-inactivated foetal calf serum (hi-FCS) and 72% infected blood. Slowly freeze at -80 &#176;C using a slow-freeze freezing container to achieve a cooling rate of -1 &#176;C/min, before transferring to liquid nitrogen.</li>
</ol>
2. Infection of Experimental Mice
<ol>
	<li>Place CD1 female mice (~21-25 g) into a heating box to gently warm to dilate veins. Place the warmed mice into a plastic restraint and inject 0.2 ml diluted inoculum intravenously with a 25 G needle into the tail vein, equivalent to 4,000 trypanosomes per mouse. Place pressure at the injection site afterwards to stem the bleeding. 
	Note: Intraperitoneal infection is a valid route of infection and has been used in previous studies10, but we have found this less reproducible than the intravenous route.</li>
	<li>Randomize infected mice and cage in groups of 3 in SPF-vented cages. As a control, inject CD1 female mice with wild type non-bioluminescent parasite, T. b. brucei GVR35 (n = 3).</li>
	<li>Monitor the Infection Through Blood Film Microscopy
	<ol>
		<li>Spot 5 &#181;l blood from venepuncture or venesection onto a glass slide and perform a blood smear (using a second glass slide gently push the drop of blood across the slide to produce a thin film). Allow slides to air dry.</li>
		<li>Fix the blood smear with 100% methanol and stain with Giemsa for 10 min before rinsing with dH2O and allow to air dry. Under 100X objective count the number of trypanosomes in 10 fields of view (f.o.v.). 
		Note: Blood filming is carried out alongside whole animal noninvasive imaging as early as one day post-infection. When processing a number of animals, this method of trypanosome quantitation is preferred over hemocytometer counting as the samples can be stored at room temperature and counted later.</li>
	</ol>
	</li>
</ol>
3. Bioluminescence Imaging to Track Infection
Note: To monitor the infection, whole animal non-invasive imaging can be used.
<ol>
	<li>Prepare a stock solution of D-luciferin, the firefly luciferase substrate, in Dulbecco&#39;s Phosphate Buffered Saline (DPBS) at 30 mg/ml. Filter sterilize and freeze in aliquots at -20 &#176;C. 
	Note: D-Luciferin is light sensitive, therefore protect it from light by wrapping aliquots in foil. Do not repeatedly freeze-thaw luciferin as it can affect activity11.</li>
	<li>Warm an aliquot of D-luciferin to room temperature. Remove each mouse from the cage (including control mice infected with nonbioluminescent wild type parasite strain) and inject 150 mg/kg of the warmed D-luciferin (diluted in DPBS) intraperitoneally with a 25 G needle. Place the mice into an isofluorane chamber for approximately 5 min to induce anesthesia with 2% isofluorane/O2 mix at 2.5 L/min. 
	Note: Mice become immobile when they are under anesthesia. For the imager used here, 3 mice can be processed at a time. If mice are anesthetized for longer than 30 min, ophthalmic artificial tear ointment can be applied to prevent the eyes of the mice from drying out.</li>
	<li>For the imager described here, open the software and initialize the machine using the software as per manufacturer&#39;s instructions. Acquire images once camera and heated plate have reached temperature. 
	Note: This instrument is not usually switched off, enabling the machine to perform background calibrations overnight. In the event that it needs to restart, it must run a calibration check first.</li>
	<li>Place the anesthetized mice into the imager (see 3.5). Use large black separators between the mice to minimize bleed through of any bright bioluminescent signal. Image mice using a set of exposure times; 1, 3, 10, 30, 60 and 180 secs, with medium binning, 1 f/stop and an open filter, and field of view E (12.5 x 12.5 cm2). 
	Note: The control mice infected with the wild type nonbioluminescent GVR35 parasites provide a background bioluminescence value. From a luciferin kinetics experiment (Figure 6) with this model, we have found that bioluminescence signal peaks at 10 minutes post administration and imaging takes approximately 5 min for 3 mice. Take this timescale into consideration when anesthetizing and imaging the mice.</li>
	<li>To ensure thorough imaging of the mice, rotate to obtain a ventral, dorsal and lateral view. Maintain consistency in the order of orientation for imaging between sequential animals.</li>
	<li>After imaging, allow mice to recover from anesthesia under observation before returning to SPF-cage.</li>
	<li>Continue imaging mice every 7 days until day 21 post-infection. There should be a steady increase in bioluminescence signal over the entire animal.</li>
	<li>At D21, image and blood film the mice as described above, and dose mice with 40 mg/kg diminazene aceturate intraperitoneally. This drug will clear the peripheral parasitemia but will not cross the blood-brain barrier, and therefore enables better visualization of the CNS infection. 
	Note: Diminazene aceturate is a well-established drug used to treat early-stage animal trypanosomiasis.</li>
	<li>Continue imaging and monitoring at D28 and D35.</li>
</ol>
4. Confirming CNS Infection
<ol>
	<li>At D35, image mice as detailed above (steps 3.1-3.4).</li>
	<li>After imaging, allow mice to recover from anesthesia and exsanguinate mice as detailed in step 1.6-1.8, collecting blood for further analysis.</li>
	<li>Following cardiac puncture, perfuse mice with 20 ml PBS (optional: add 3 mg/ml of luciferin to perfusion solution) via the heart to clear blood from the organs and to reintroduce luciferin label that might have cleared from the brain region over the time interval from step 4.1.</li>
	<li>Gently remove the brain, by using curved scissors, cutting from the base around the circumference of the skull, and lifting off the top part of the skull, to allow the brain to be gently lifted out with curved forceps.</li>
	<li>Place the excised brain onto a section of black plastic and pipette 50 &#181;l of 15 mg/ml luciferin over the organ prior to imaging, to ensure adequate luciferin has been added to the excised brain. Image using the settings as above. This will demonstrate localization of the infection to the brain hemispheres. 
	Note: Downstream quantification of parasitemia on the excised brain can be carried out by qPCR as previously described8. Other alternative downstream applications might include histology or immunohistochemistry to identify parasites in the brain tissue following fixation.</li>
</ol>
5. Quantitation of Bioluminescence Imaging
Note: Bioluminescence can be quantified using the region of interest (ROI) with the imaging software and corrected for background bioluminescence.
<ol>
	<li>Using the ROI tool, create a rectangular ROI for whole animal quantitation and position it over the mouse image to cover tip of the nose to base of the tail of the mouse. Use the same size box for every animal and every time point. When looking at the ROI for the brain, use a circular ROI in the same way, positioning it over the mouse head region in the image.</li>
	<li>If desired, adjust images to appear on the same spectrum for a visual representation of infection. 
	Note: The software used here generates a color map for each image automatically adjusted for minimum and maximum photon counts. To enable visual comparison between a series of images, the color scale must be set to the same values for all images.
	<ol>
		<li>Using the image with the highest bioluminescence and the &#39;Image adjust&#39; tab on the tool palette, select a logarithmic scale and an appropriate color scale minimum and maximum (to be determined empirically). Apply this scale to all images throughout the experiment.</li>
	</ol>
	</li>
</ol>

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S. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Bioluminescence+imaging.">Bioluminescence imaging.</a> Proc. Am. Thorac. Soc. 2 (6), 537-540 (2005).</li><li>Burrell-Saward, H., Rodgers, J., Bradley, B., Croft, S. L., Ward, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+sensitive+and+reproducible+in+vivo+imaging+mouse+model+for+evaluation+of+drugs+against+late-stage+human+African+trypanosomiasis.">A sensitive and reproducible in vivo imaging mouse model for evaluation of drugs against late-stage human African trypanosomiasis.</a> J. Antimicrob. Chemother. 70 (2), 510-517 (2015).</li><li>McLatchie, A. 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The authors have nothing to disclose.

The development of a bioluminescent T. b. brucei GVR35 strain allows the visualization of a trypanosome infection from the early to the late stage. Previous infection models were unable to detect the late stage, when parasites are in the brain, in real time from blood film microscopy, and required the culling and removal of brains from the infected mice to determine parasite burden12. The bioluminescence reduces inter-mouse variability as a single mouse can be tracked throughout the entirety of the in...

Human African trypanosomiasis (HAT), or sleeping sickness, is caused by the vector-borne protozoan parasites of the Trypanosoma brucei spp1. Estimated numbers of current cases is fewer than 7 thousand every year with almost 70 million people exposed to the risk of the parasite infection within the African continent. The disease, which is most often lethal if left untreated, comprises an early hemolymphatic stage where parasites are present in the blood, progressing to the late stage when parasites invade the central nervous system (CNS) and are no longer susceptible to treatment by early stage trypanosomal drugs2. The current drug therapies for late-stage HAT have both complex, prolonged, treatment regimens and severe adverse effects as well as reported resistance, therefore research into new drug therapies is imperative3,4. 
The study of late-stage human African trypanosomiasis (HAT) within traditional mouse models is lengthy and complex, with the removal of brain tissue being required to monitor parasitic burden5. The animal infective strain T. b. brucei is used as the study model of trypanosomiasis with the late stage appearing 21 days post infection (dpi). To monitor the wild type nonbioluminescent parasite infection in the mouse model, peripheral blood films or quantitative PCR are the only methods to determine parasite burden. For parasite burden in the brain, the mouse needs to be culled, brain excised and qPCR carried out on tissues, making it impossible to track parasites through multiple time points in the late stage infection. This results in the inability to follow real-time infections within the central nervous system (CNS). 
In vivo bioluminescence imaging (BLI) can provide highly sensitive, non-invasive detection of parasite dissemination and disease progression in a mouse model that can be followed in a single animal for the entirety of the experiment6. BLI is based on the emission of light in the visible spectrum produced by a luciferase-catalyzed reaction. The emitted photons are then detected by a charge coupled device (CCD) camera7. For this purpose, the pathogen is genetically modified to express a luciferase protein and the substrate, luciferin, is introduced at time points of interest by injection. The main advantage of this method is the ability to carry out longitudinal studies, in which the same animal can be imaged several times with minimal adverse effects. The acquired bioluminescence signal can be quantified, thus indicating the pathogen burden.
The optimization and validation of a red-shifted bioluminescent T. b. brucei has enabled the investigation of the late stage infection through non-invasive procedures, detecting parasites earlier than blood film microscopy and greatly reducing the time, cost and numbers of animals needed to study CNS infection and drug screening in late-stage trypanosomiasis8,9. In this protocol we demonstrate infection of mice with bioluminescent trypanosomes and how to then visualize the parasites in vivo for quantification of disease progression and CNS penetration.

<table><tbody><tr><td>PBS</td><td>Sigma, UK</td><td>P4417</td><td>tablets pH 7.4</td></tr><tr><td>Glucose</td><td>Sigma, UK</td><td>G8270</td><td>99.5% (molecular) grade</td></tr><tr><td>Ammonium chloride</td><td>Sigma, UK</td><td>A9434</td><td>99.5% (molecular) grade</td></tr><tr><td>Heparin (lithium salt)</td><td>Sigma, UK</td><td>H0878</td><td></td></tr><tr><td>Hi-FCS</td><td>Gibco, Life Technologies, UK</td><td>10500-064</td><td>500 ml</td></tr><tr><td>DPBS</td><td>Sigma, UK</td><td>D4031</td><td>Sterile filtered</td></tr><tr><td>Mr. Frosty</td><td>Nalgene, UK</td><td></td><td></td></tr><tr><td>Giemsa</td><td>Sigma, UK</td><td>G5637</td><td></td></tr><tr><td>D-Luciferin</td><td>Perkin Elmer, UK</td><td></td><td></td></tr><tr><td>Sigma, UK</td><td>115144-35-9</td><td></td><td></td></tr><tr><td>Diminazene aceturate</td><td>Sigma, UK</td><td>D7770</td><td>Analytical grade</td></tr><tr><td>IVIS Lumina II</td><td>Perkin Elmer, UK</td><td></td><td>other bioimagers available &lt;em&gt;e.g.&lt;/em&gt; from Bruker, Kodak</td></tr><tr><td>Living Image v. 4.2</td><td>Perkin Elmer, UK</td><td></td><td>proprietary software for Perkin Elmer IVIS instruments; other instruments may have their own</td></tr><tr><td>1 ml Syringe</td><td>Fisher Scientific, UK</td><td>10142104</td><td></td></tr><tr><td>20 ml Syringe</td><td>Fisher Scientific, UK</td><td>10743785</td><td></td></tr><tr><td>25 G Needles</td><td>Greiner Bio-one</td><td>N2516</td><td></td></tr><tr><td>21 G Needles</td><td>Greiner Bio-one</td><td>N2138</td><td></td></tr><tr><td>Twin-frosted microscope slide</td><td>VWR, UK</td><td>631-0117</td><td></td></tr><tr><td>1.5 ml Microcentrifuge tube</td><td>StarLab, UK</td><td>I1415-1000</td><td></td></tr><tr><td>7 ml Bijou tube</td><td>StarLab, UK</td><td>E1412-0710</td><td></td></tr><tr><td>Mouse restrainer</td><td>Sigma, UK</td><td>Z756903</td><td>our restrainer was made in-house, this is a similar model</td></tr></tbody></table>

bioluminescence imaging to detect late stage infection of african trypanosomiasis

Human African trypanosomiasis (HAT) is a multi-stage disease that manifests in two stages; an early blood stage and a late stage when the parasite invades the central nervous system (CNS). In vivo study of the late stage has been limited as traditional methodologies require the removal of the brain to determine the presence of the parasites.
Bioluminescence imaging is a non-invasive, highly sensitive form of optical imaging that enables the visualization of a luciferase-transfected pathogen in real-time. By using a transfected trypanosome strain that has the ability to produce late stage disease in mice we are able to study the kinetics of a CNS infection in a single animal throughout the course of infection, as well as observe the movement and dissemination of a systemic infection.
Here we describe a robust protocol to study CNS infections using a bioluminescence model of African trypanosomiasis, providing real time non-invasive observations which can be further analyzed with optional downstream approaches.

This protocol demonstrates how to follow disease progression following infection of mice with T. b. brucei, a model for human African trypanosomiasis. Figure 1 shows the timeline of the experimental protocol, demonstrating the timetable for treatment and imaging steps. Figure 2 demonstrates a typical field of view in a fixed Giemsa-stained blood smear used to quantitate peripheral parasitemia, with trypanosomes and red blood cells present. Develo...

Watch this Scientific Journal Video about Bioluminescence Imaging to Detect Late Stage Infection of African Trypanosomiasis at JoVE.com

Bioluminescence Imaging to Detect Late Stage Infection of African Trypanosomiasis

This manuscript describes the use of a bioluminescent strain of African trypanosomes to enable the tracking of late stage infection and demonstrates how in vivo live imaging can be used to visualize infections within the central nervous system in real-time.

Human African trypanosomiasis (HAT) is a multi-stage disease that manifests in two stages; an early blood stage and a late stage when the parasite invades ...

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Immunology and Infection

7.7K Views.  London School of Hygiene & Tropical Diseases. This manuscript describes the use of a bioluminescent strain of African trypanosomes to enable the tracking of late stage infection and demonstrates how in vivo live imaging can be used to visualize infections within the central nervous system in real-time.

Video: Bioluminescence Imaging to Detect Late Stage Infection of African Trypanosomiasis

In vivo Imaging of Transgenic Leishmania Parasites in a Live Host

Distinct species of Leishmania, a protozoan parasite of the family Trypanosomatidae, typically cause different human disease manifestations. The most common forms of disease are visceral leishmaniasis (VL) and cutaneous leishmaniasis (CL). Mouse models of leishmaniasis are widely used, but quantification of parasite burdens during murine disease requires mice to be euthanized at various times after infection. Parasite loads are then measured either by microscopy, limiting dilution assay, or qPCR amplification of parasite DNA. The in vivo imaging system (IVIS) has an integrated software package that allows the detection of a bioluminescent signal associated with cells in living organisms. Both to minimize animal usage and to follow infection longitudinally in individuals, in vivo models for imaging Leishmania spp. causing VL or CL were established. Parasites were engineered to express luciferase, and these were introduced into mice either intradermally or intravenously. Quantitative measurements of the luciferase driving bioluminescence of the transgenic Leishmania parasites within the mouse were made using IVIS. Individual mice can be imaged multiple times during longitudinal studies, allowing us to assess the inter-animal variation in the initial experimental parasite inocula, and to assess the multiplication of parasites in mouse tissues. Parasites are detected with high sensitivity in cutaneous locations. Although it is very likely that the signal (photons/second/parasite) is lower in deeper visceral organs than the skin, but quantitative comparisons of signals in superficial versus deep sites have not been done. It is possible that parasite numbers between body sites cannot be directly compared, although parasite loads in the same tissues can be compared between mice. Examples of one visceralizing species (L. infantum chagasi) and one species causing cutaneous leishmaniasis (L. mexicana) are shown. The IVIS procedure can be used for monitoring and analyzing small animal models of a wide variety of Leishmania species causing the different forms of human leishmaniasis.

In vivo Imaging of Transgenic Leishmania Parasites in a Live Host

An in vivo imaging system is used to generate quantitative measurements of murine infection with the Trypanosomatid protozoan Leishmania. This is a non-invasive and non-lethal method for detecting parasites expressing luciferase within many tissues throughout the course of chronic Leishmania spp. infection.

Distinct species of Leishmania, a protozoan parasite of the family Trypanosomatidae, typically cause different human disease manifestations. The most ...

Comprehensive &amp; Cost Effective Laboratory Monitoring of HIV/AIDS: an African Role Model

We present the video about assisting anti-retroviral therapy (ART) by an apt laboratory service - representing a South-African role model for economical large scale diagnostic testing. In the low-income countries inexpensive ART has transformed the prospects for the survival of HIV seropositive patients but there are doubts whether there is a need for the laboratory monitoring of ART and at what costs - in situations when the overall quality of pathology services can still be very low. The appropriate answer is to establish economically sound services with better coordination and stricter internal quality assessment than seen in western countries. This video, photographed at location in the National Health Laboratory Services (NHLS-SA) at the Witwatersrand University, Johannesburg, South Africa, provides such a coordinated scheme expanding the original 2-color CD4-CD45 PanLeucoGating strategy (PLG). Thus the six modules of the video presentation reveal the simplicity of a 4-color flow cytometric assay to combine haematological, immunological and virology-related tests in a single tube. These video modules are: (i) the set-up of instruments; (ii) sample preparations; (iii) testing absolute counts and monitoring quality for each sample by bead-count-rate; (iv) the heamatological CD45 test for white cell counts and differentials; (v) the CD4 counts, and (vi) the activation of CD8+ T cells measured by CD38 display, a viral load related parameter. The potential cost-savings are remarkable. This arrangement is a prime example for the feasibility of performing &gt; 800-1000 tests per day with a stricter quality control than that applied in western laboratories, and also with a transfer of technology to other laboratories within a NHLS-SA network. Expert advisors, laboratory managers and policy makers who carry the duty of making decisions about introducing modern medical technology are frequently not in a position to see the latest technical details as carried out in the large regional laboratories with huge burdens of workload. Hence this video shows details of these new developments.

Comprehensive & Cost Effective Laboratory Monitoring of HIV/AIDS: an African Role Model

Anti-retroviral therapy to treat HIV/AIDS is monitored in South Africa on a large scale. Flow cytometry is combined for haematology (CD45), immunology (CD4) and viral-load linked CD38 assay. Recorded at NHLS-SA laboratories, Johannesburg, these modern methods are cost-efficient with heightened local internal quality control, serving as role-models for resource-limited diagnostics.

We present the video about assisting anti-retroviral therapy (ART) by an apt laboratory service - representing a South-African role model for economical ...

Bioluminescence Imaging of NADPH Oxidase Activity in Different Animal Models

NADPH oxidase is a critical enzyme that mediates antibacterial and antifungal host defense. In addition to its role in antimicrobial host defense, NADPH oxidase has critical signaling functions that modulate the inflammatory response 1. Thus, the development of a method to measure in "real-time" the kinetics of NADPH oxidase-derived ROS generation is expected to be a valuable research tool to understand mechanisms relevant to host defense, inflammation, and injury.
Chronic granulomatous disease (CGD) is an inherited disorder of the NADPH oxidase characterized by severe infections and excessive inflammation. Activation of the phagocyte NADPH oxidase requires translocation of its cytosolic subunits (p47phox, p67phox, and p40phox) and Rac to a membrane-bound flavocytochrome (composed of a gp91phox and p22phox heterodimer). Loss of function mutations in any of these NADPH oxidase components result in CGD. Similar to patients with CGD, gp91phox -deficient mice and p47phox-deficient mice have defective phagocyte NADPH oxidase activity and impaired host defense 2, 13. In addition to phagocytes, which contain the NADPH oxidase components described above, a variety of other cell types express different isoforms of NADPH oxidase.
Here, we describe a method to quantify ROS production in living mice and to delineate the contribution of NADPH oxidase to ROS generation in models of inflammation and injury. This method is based on ROS reacting with L-012 (an analogue of luminol) to emit luminescence that is recorded by a charge-coupled device (CCD). In the original description of the L-012 probe, L-012-dependent chemiluminescence was completely abolished by superoxide dismutase, indicating that the main ROS detected in this reaction was superoxide anion 14. Subsequent studies have shown that L-012 can detect other free radicals, including reactive nitrogen species 15, 16. Kielland et al. 16 showed that topical application of phorbol myristate acetate, a potent activator of NADPH oxidase, led to NADPH oxidase-dependent ROS generation that could be detected in mice using the luminescent probe L-012. In this model, they showed that L-012-dependent luminescence was abolished in p47phox-deficient mice.
We compared ROS generation in wildtype mice and NADPH oxidase-deficient p47phox-/- mice 2 in the following three models: 1) intratracheal administration of zymosan, a pro-inflammatory fungal cell wall-derived product that can activate NADPH oxidase; 2) cecal ligation and puncture (CLP), a model of intra-abdominal sepsis with secondary acute lung inflammation and injury; and 3) oral carbon tetrachloride (CCl4), a model of ROS-dependent hepatic injury. These models were specifically selected to evaluate NADPH oxidase-dependent ROS generation in the context of non-infectious inflammation, polymicrobial sepsis, and toxin-induced organ injury, respectively. Comparing bioluminescence in wildtype mice to p47phox-/- mice enables us to delineate the specific contribution of ROS generated by p47phox-containing NADPH oxidase to the bioluminescent signal in these models.
Bioluminescence imaging results that demonstrated increased ROS levels in wildtype mice compared to p47phox-/- mice indicated that NADPH oxidase is the major source of ROS generation in response to inflammatory stimuli. This method provides a minimally invasive approach for "real-time" monitoring of ROS generation during inflammation in vivo.

NADPH oxidase is the major source of reactive oxygen species (ROS) in phagocytes. Because of the ephemeral nature of ROS, it is difficult to measure and monitor ROS levels in living animals. A minimally invasive method for serial quantification of ROS in living mice is described.

NADPH oxidase is a critical enzyme that mediates antibacterial and antifungal host defense. In addition to its role in antimicrobial host defense, NADPH ...

In Vivo Imaging Systems (IVIS) Detection of a Neuro-Invasive Encephalitic Virus

Modern advancements in imaging technology encourage further development and refinement in the way viral research is accomplished. Initially proposed by Russel and Burch in Hume's 3Rs (replacement, reduction, refinement), the utilization of animal models in scientific research is under constant pressure to identify new methodologies to reduce animal usage while improving scientific accuracy and speed. A major challenge to Hume's principals however, is how to ensure the studies are statistically accurate while reducing animal disease morbidity and overall numbers. Vaccine efficacy studies currently require a large number of animals in order to be considered statistically significant and often result in high morbidity and mortality endpoints for identification of immune protection. We utilized in vivo imaging systems (IVIS) in conjunction with a firefly bioluminescent enzyme to progressively track the invasion of the central nervous system (CNS) by an encephalitic virus in a murine model. Typically, the disease progresses relatively slowly, however virus replication is rapid, especially within the CNS, and can lead to an often, lethal outcome. Following intranasal infection of the mice with TC83-Luc, an attenuated Venezuelan equine encephalitis virus strain modified to expresses a luciferase gene; we are able to visualize virus replication within the brain at least three days before the development of clinical disease symptoms. Utilizing CNS invasion as a key encephalitic disease development endpoint we are able to quickly identify therapeutic and vaccine protection against TC83-Luc infection before clinical symptoms develop. With IVIS technology we are able to demonstrate the rapid and accurate testing of drug therapeutics and vaccines while reducing animal numbers and morbidity.

In Vivo Imaging Systems IVIS Detection of a Neuro-Invasive Encephalitic Virus

Utilizing luciferase and in vivo imaging systems (IVIS) as a novel means to identify disease endpoints before clinical developments occur. IVIS has allowed us to visualize in real time the invasion of encephalitic viruses over multiple days, providing a more accurate disease model for future study. It has also allowed us to identify the potential protective features of antivirals and vaccines faster than currently utilized animal models. The capability to utilize individual animals over multiple time points ensures reduced animal requirements, costs, and overall morbidity to the animals utilized ensuring a more humane and more scientific means of disease study.

Modern advancements in imaging technology encourage further development and refinement in the way viral research is accomplished. Initially proposed by ...

Using Fluorescent Proteins to Monitor Glycosome Dynamics in the African Trypanosome

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in cattle1. The parasite alternates between the bloodstream of the mammalian host and the tsetse fly vector. The composition of many cellular organelles changes in response to these different extracellular conditions2-5.
Glycosomes are highly specialized peroxisomes in which many of the enzymes involved in glycolysis are compartmentalized. Glycosome composition changes in a developmental and environmentally regulated manner4-11. Currently, the most common techniques used to study glycosome dynamics are electron and fluorescence microscopy; techniques that are expensive, time and labor intensive, and not easily adapted to high throughput analyses.
To overcome these limitations, a fluorescent-glycosome reporter system in which enhanced yellow fluorescent protein (eYFP) is fused to a peroxisome targeting sequence (PTS2), which directs the fusion protein to glycosomes12, has been established. Upon import of the PTS2eYFP fusion protein, glycosomes become fluorescent. Organelle degradation and recycling results in the loss of fluorescence that can be measured by flow cytometry. Large numbers of cells (5,000 cells/sec) can be analyzed in real-time without extensive sample preparation such as fixation and mounting. This method offers a rapid way of detecting changes in organelle composition in response to fluctuating environmental conditions.

Glycosome dynamics in African trypanosomes are difficult to study by traditional cell biology techniques such as electron and fluorescence microscopy. As a means of observing dynamic organelle behavior, a fluorescent-organelle reporter system has been used in conjunction with flow cytometry to monitor real-time glycosome dynamics in live parasites.

Trypanosoma brucei is a kinetoplastid parasite that causes human African trypanosomiasis (HAT), or sleeping sickness, and a wasting disease, nagana, in ...

High-throughput Gene Tagging in Trypanosoma brucei

Improvements in mass spectrometry, sequencing and bioinformatics have generated large datasets of potentially interesting genes. Tagging these proteins can give insights into their function by determining their localization within the cell and enabling interaction partner identification. We recently published a fast and scalable method to generate Trypanosoma brucei cell lines that express a tagged protein from the endogenous locus. The method was based on a plasmid we generated that, when coupled with long primer PCR, can be used to modify a gene to encode a protein tagged at either terminus. This allows the tagging of dozens of trypanosome proteins in parallel, facilitating the large-scale validation of candidate genes of interest. This system can be used to tag proteins for localization (using a fluorescent protein, epitope tag or electron microscopy tag) or biochemistry (using tags for purification, such as the TAP (tandem affinity purification) tag). Here, we describe a protocol to perform the long primer PCR and the electroporation in 96-well plates, with the recovery and selection of transgenic trypanosomes occurring in 24-well plates. With this workflow, hundreds of proteins can be tagged in parallel; this is an order of magnitude improvement to our previous protocol and genome scale tagging is now possible.

High-throughput Gene Tagging in Trypanosoma brucei

Addition of a tag to a protein is a powerful way of gaining insight into its function. Here, we describe a protocol to endogenously tag hundreds of Trypanosoma brucei proteins in parallel such that genome scale tagging is achievable.

Improvements in mass spectrometry, sequencing and bioinformatics have generated large datasets of potentially interesting genes. Tagging these proteins ...

AI Program for Trypanosomiasis Detection and Classification

Superior Auto-Identification of Trypanosome Parasites by Using a Hybrid Deep-Learning Model

Trypanosomiasis is a significant public health problem in several regions across the world, including South Asia and Southeast Asia. The identification of hotspot areas under active surveillance is a fundamental procedure for controlling disease transmission. Microscopic examination is a commonly used diagnostic method. It is, nevertheless, primarily reliant on skilled and experienced personnel. To address this issue, an artificial intelligence (AI) program was introduced that makes use of a hybrid deep learning technique of object identification and object classification neural network backbones on the in-house low-code AI platform (CiRA CORE). The program can identify and classify the protozoan trypanosome species, namely Trypanosoma cruzi, T. brucei, and T. evansi, from oil-immersion microscopic images. The AI program utilizes pattern recognition to observe and analyze multiple protozoa within a single blood sample and highlights the nucleus and kinetoplast of each parasite as specific characteristic features using an attention map.
To assess the AI program&#39;s performance, two unique modules are created that provide a variety of statistical measures such as accuracy, recall, specificity, precision, F1 score, misclassification rate, receiver operating characteristics (ROC) curves, and precision versus recall (PR) curves. The assessment findings show that the AI algorithm is effective at identifying and categorizing parasites. By delivering a speedy, automated, and accurate screening tool, this technology has the potential to transform disease surveillance and control. It could also assist local officials in making more informed decisions on disease transmission-blocking strategies.

Author Spotlight: AI-Driven Trypanosome Species Detection from Microscopic Images 

Worldwide medical blood parasites were automatically screened using simple steps on a low-code AI platform. The prospective diagnosis of blood films was improved by using an object detection and classification method in a hybrid deep learning model. The collaboration of active monitoring and well-trained models helps to identify hotspots of trypanosome transmission.

Trypanosomiasis is a significant public health problem in several regions across the world, including South Asia and Southeast Asia. The identification of ...

Bioengineering

Dual Bioluminescence Imaging of Tumor Progression and Angiogenesis

Angiogenesis, as a crucial process of tumor progression, has become a research hotspot and target of anti-tumor therapy. However, there is no reliable model for tracing tumor progression and angiogenesis simultaneously in a visual and sensitive manner. Bioluminescence imaging displays its unique superiority in living imaging due to its advantages of high sensitivity, strong specificity, and accurate measurement. Presented here is a protocol to establish a tumor-bearing mouse model by injecting a Renilla luciferase-labeled murine breast cancer cell line 4T1 into the transgenic mouse with angiogenesis-induced Firefly luciferase expression. This mouse model provides a valuable tool to simultaneously monitor tumor progression and angiogenesis in real-time by dual bioluminescence imaging in a single mouse. This model may be widely applied in anti-tumor drug screening and oncology research.

This protocol describes the establishment of a tumor-bearing mouse model to monitor tumor progression and angiogenesis in real-time by dual bioluminescence imaging.

Angiogenesis, as a crucial process of tumor progression, has become a research hotspot and target of anti-tumor therapy. However, there is no reliable ...

Medicine

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, ...

In Vivo Imaging of Leishmania infantum-infected Hamsters by Gingival Inoculation of Axenic Amastigotes Expressing Luciferase

American tegumentary leishmaniasis (ATL) and visceral leishmaniasis (VL) are considered neglected by the World Health Organization. VL can be lethal if not treated; the drugs used in treatment are toxic, and there are cases of resistance. Preclinical tests can represent a bottleneck in discovering new medicines for treatment, depending on the animal model, the strain used, and the inoculum route. The golden hamster stands out for its high susceptibility to subgenera Viannia and Leishmania species, displaying many of the clinical and immunopathological processes observed in human disease.
By hamster anatomy, which has a short tail and limbs, the intracardiac route is usually the choice for intravenous injection of Leishmania. However, it is an inoculum that can lead to bleeding and eventually to animal death. Thus, we standardized an alternative intravenous inoculation route for infection at the gingival vein, which is minimally invasive, allows easy venous access, and causes few local and systemic injuries to the animal. Therefore, hamsters infected by the intraperitoneal (IP) or intragingival (IG) route with Leishmania Infantum expressing luciferase (Luc) were followed up for 22 days by the bioluminescence imaging system and 50 days and 8 months post infection by PCR.
After gingival inoculation of both axenic amastigotes and promastigotes of L. infantum-Luc, bioluminescence was restricted for at least 2 weeks at the site of injection, which is an indicator of infection in the tissues around the gingival plexus. Hamsters infected intraperitoneally with L. infantum-Luc displayed bioluminescence dispersed throughout the abdomen, as expected. However, by the bioluminescence imaging system infection declined until the 50th dpi and was only detectable by PCR.&#160;Axenic amastigotes showed better infection than promastigotes, as evaluated by PCR. Indeed, 8 months after infection, parasites were detected by PCR in the liver of animals inoculated with axenic amastigotes by the intravenous route, which can be a characteristic of the reference strain of L. infantum MHOM/BR/1974/PP75, whose infection progresses slowly and display low parasite burden, below the bioluminescent imaging resolution. Thus, axenic amastigotes can be a better choice for infection and follow-up than promastigotes, and the gingival inoculum is a feasible route for intravenous injection of Leishmania and other pathogens.

Here we present a longitudinal evaluation of golden hamsters infected intraperitoneally (IP) or via intragingival (IG) with L. infantum-Luc by bioluminescent imaging and by PCR. Hamsters were evaluated 1 day post infection (1 dpi), 1 week after infection (8 dpi), and 3 weeks after infection (22 dpi) and euthanized at the 50th dpi and 8 months post infection.

American tegumentary leishmaniasis (ATL) and visceral leishmaniasis (VL) are considered neglected by the World Health Organization. VL can be lethal if ...

Bioluminescence Imaging to Detect Late Stage Infection of African Trypanosomiasis

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In vivo Imaging of Transgenic Leishmania Parasites in a Live Host

Comprehensive & Cost Effective Laboratory Monitoring of HIV/AIDS: an African Role Model

Bioluminescence Imaging of NADPH Oxidase Activity in Different Animal Models

In Vivo Imaging Systems (IVIS) Detection of a Neuro-Invasive Encephalitic Virus

Using Fluorescent Proteins to Monitor Glycosome Dynamics in the African Trypanosome

High-throughput Gene Tagging in Trypanosoma brucei

Superior Auto-Identification of Trypanosome Parasites by Using a Hybrid Deep-Learning Model

Dual Bioluminescence Imaging of Tumor Progression and Angiogenesis

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

In Vivo Imaging of Leishmania infantum-infected Hamsters by Gingival Inoculation of Axenic Amastigotes Expressing Luciferase