SD was supported by a Sir Henry Wellcome Fellowship [092201/Z/10/Z]. This work was funded by Wellcome Trust grants [WT066839MA][104627/Z/14/Z][108445/Z/15/Z] to Professor Keith Gull. We would like to thank Professor Keith Gull for helpful conversations and insights.

1. 96-well Long Primer PCR
<ol>
	<li>Pre-freeze 96-well PCR cooler in -80 &#176;C freezer.</li>
	<li>In a 10 ml tube, prepare Master Mix A: 2,100 &#181;l ddH2O, 105 &#181;l PCR grade DMSO, 105 &#181;l 10 mM dNTPs, 105 &#181;l pPOT template (25 ng/&#181;l) 9, for a total volume of 2,415 &#181;l per plate.</li>
	<li>Aliquot 23 &#181;l into each well of a 96-well PCR plate.</li>
	<li>Add 2 &#181;l of 100 &#181;M pooled primers to each loaded well using a P20 12 channel multichannel pipette. Seal with film, place plate on the pre-chilled PCR cooler and allow to freeze for a minimum of 20 min at -80 &#176;C.</li>
	<li>In a 10 ml tube, prepare Master Mix B: 2,048 &#181;l ddH2O, 525 &#181;l 10x buffer, 52.5 &#181;l polymerase for a total volume of 2,626 &#181;l per plate.</li>
	<li>Remove plate and PCR cooler from the -80 &#176;C freezer.</li>
	<li>With the plate still on the PCR cooler, add 25 &#181;l Master Mix B on top of the frozen Master Mix A for a total reaction volume of 50 &#181;l. This is time critical, and should be completed before Mix A can thaw. Seal plate with film.</li>
	<li>Set thermal cycler as follows: 94 &#176;C for 10 min, then 30 cycles of 94 &#176;C for 15 sec, 65 &#176;C for 30 sec, 72&#176;C for 2 min followed by a final extension period of 72 &#176;C for 7 min.</li>
	<li>Load the PCR plate after the block has reached 94 &#176;C.</li>
</ol>
2. Validation of 96-well Long Primer PCR
<ol>
	<li>Cast a large 1% (w/v) agarose gel in Tris Acetate EDTA (TAE) running buffer with 4 x 28-well combs to give a gel with 4 rows of wells. Align alternate teeth of the comb with the tips of a 12 channel multichannel pipette. Carefully submerge the gel in TAE running buffer after the lanes are loaded.</li>
	<li>Load 1 kb DNA ladder into the 1st and 28th well of each row.</li>
	<li>Using a P20 12 channel multichannel pipette, transfer 2 &#181;l of PCR product directly to each lane of the gel (Figure 1A). Do this quickly to reduce the possibility of contaminating the amplicons.
	<ol>
		<li>Load the PCR products from row A of the PCR plate into the odd numbered lanes of the 1st row of the gel (A1 - lane 3, A2 - lane 5 &#8230;&#8230; A11 - lane 23, A12 - lane 25) and load the PCR products from row B of the PCR plate into the even numbered lanes of the 1st row of the gel (B1 - lane 4, A2 - lane 6 &#8230;&#8230; B11 - lane 24, B12 - lane 26).</li>
		<li>Load the PCR products from row C and D of the PCR plate using the same pattern as described above, with row C into the odd numbered lanes and row D into the even numbered lanes. Continue this loading pattern until each well of the PCR plate is loaded onto the gel. DNA loading buffer is not required to load this gel.</li>
	</ol>
	</li>
	<li>Place the gel into the running tank and fill the tank with TAE running buffer until the gel is submerged. Run at 100 V for 30 min and visualize using a UV transilluminator. See Figure 1B for an example gel. 
	Note: PCR products can be stored at -20 &#176;C for several weeks prior to transfection</li>
</ol>
3. 96-well Transfection
<ol>
	<li>Use the T. brucei procyclic form cell line SMOXP9 10 for this procedure and grow in SDM-79 media containing 10% FCS 10.</li>
	<li>Use 1 x 107 cells per transfection (total of 1.1 x 109 cells) when tagging on the N terminus of the protein and 2 x 107 cells per transfection (total of 2.2 x 109 cells) when tagging on the C terminus of the protein. Maintain the cells in mid-log (1.2 x 106-1 x 107 cells/ml) for several days prior to transfection and harvest the cells for transfection at a density of 5-8 x 106 cells/ml.</li>
	<li>Allow frozen PCR products for transfection to thaw at room temperature.</li>
	<li>Count cells using a haemocytometer or automatic cell counter.</li>
	<li>Pellet required number of cells in multiple 50 ml tubes at 800 x g for 10 min.</li>
	<li>After centrifugation discard supernatant and resuspend the cells in 10 ml modified cytomix (0.8 mM EGTA, 24 mM KCl, 0.15 mM CaCl2, 10 mM potassium phosphate buffer pH7.6, 25 mM HEPES-KOH pH7.6, 2.6 mM MgCl2, 0.5% (w/v) glucose, 100 &#181;g/ml BSA, 1 mM hypoxanthine, 144 mM sucrose) per tube. Transfer all cell solutions to a single tube and spin again at 800 x g for 10 min.</li>
	<li>After centrifugation, discard supernatant and resuspend the cells in 23 ml of modified cytomix for a final concentration of 5 x 107 cells/ml.</li>
	<li>While cells are spinning, add 1 ml of SDM-79 media to each well of 4 x 24-well tissue culture plates. Label the plates A-D and draw a ring around well A1 on each plate in permanent marker to help plate orientation.</li>
	<li>Connect the plate handler to the electroporator. On the electroporator unit set the voltage to 1,500 V, the pulse length to 100 &#181;sec and the number of pulses to 12 and the pulse interval to 500 msec. On the plate handler, set the pulse count to 1. This ensures that that electroporator will apply a single pulse to each of the 12 columns of the electroporation plate.</li>
	<li>Using a P200 12 channel multichannel pipette, transfer the PCR reactions from the PCR plate to the 96-well disposable electroporation plates, 4 mm gap.</li>
	<li>When the cells have been spun and are resuspended to the final concentration of 5 x 107 cells/ml (Step 3.7), transfer to a reagent reservoir. Pipette 200 &#181;l of the cell solution into each well of the electroporation plate using a P200 12 channel multichannel pipette, mix with PCR product by pipetting.</li>
	<li>Using tissue remove any droplets from the top of the plate to avoid short-circuits.</li>
	<li>Apply the sealing film provided in the plate packaging to the top of the electroporation plate, positioning it so as to leave the holes for the electrodes uncovered at the top and bottom of each column. Avoid covering the raised points used to guide the film.</li>
	<li>Load the electroporation plate into the plate handler and close the lid. Press &#39;Pulse&#39; on the electroporator unit.</li>
	<li>After electroporation, quickly transfer the cells from the 96-well electroporation plate to the 4 x 24-well tissue culture plates. To transfer the cells, use a P200 12 channel multichannel pipette with every other tip missing such that the remaining 6 tips line up with the 6 rows on a 24-well tissue culture plate. Transfer the electroporated cells in the 96-well electroporations plates in pre-defined pattern to the 24-well tissue culture plates (Figure 2A).</li>
	<li>Prepare the P200 pipette tips - for each 96-well transfection prepare 2 boxes of 96 tips with every other column removed.
	<ol>
		<li>Transfer wells A1/A3/A5/A7/A9/A11 on the 96-well electroporation plate to plate A row A of the 24-well tissue culture plates, B1/B3/B5/B7/B9/B11 into row B, C1/C3/C5/C7/C9/C11 into row C and D1/D3/D5/D7/D9/D11 into row D. After transfer of each set of 6 wells from the 96-well plate to the 24-well plate, the wells in the 96-well plate are then washed with media from the corresponding wells in the 24-well plate and the wash is then transferred back to the wells in the 24-well plate.</li>
		<li>Transfer wells E1/E3/E5/E7/E9/E11 on the 96-well electroporation plate to 24-well plate B row A, F1/F3/F5/F7/F9/F11 into row B, G1/G3/G5/G7/G9/G11 into row C and H1/H3/H5/H7/H9/H11 into row D. After transfer of each set of 6 wells from the 96-well plate to the 24-well plate, the wells in the 96-well plate are then washed with media from the corresponding wells in the 24-well plate and the wash is then transferred back to the wells in the 24-well plate.</li>
		<li>Transfer wells A2/A4/A6/A8/A10/A12 on the 96-well electroporation plate to 24-well plate C row A, B2/B4/B6/B8/B10/B12 into row B, C2/C4/C6/C8/C10/C12 into row C and D2/D4/D6/D8/D10/D12 into row D. After transfer of each set of 6 wells from the 96-well plate to the 24-well plate, the wells in the 96-well plate are then washed with media from the corresponding wells in the 24-well plate and the wash is then transferred back to the wells in the 24-well plate.</li>
		<li>Transfer wells E2/E4/E6/E8/E10/E12 on the 96-well electroporation plate to 24-well plate D row A, F2/F4/F6/F8/F10/F12 into row B, G2/G4/G6/G8/G10/G12 into row C and H2/H4/H6/H8/H10/H12 into row D. After transfer of each set of 6 wells from the 96-well plate to the 24-well plate, the wells in the 96-well plate are then washed with media from the corresponding wells in the 24-well plate and the wash is then transferred back to the wells in the 24-well plate.</li>
	</ol>
	</li>
	<li>Using an inverted phase contrast microscope, examine 1 well from each column to check the cells. There will be a mixture of live and dead cells with a substantial amount of cell debris. Live cells can be distinguished from dead cells because they will be bright under phase contrast microscopy and will be moving.</li>
	<li>Place the 24-well plates in a clean, plastic box with a lid that forms a tight seal. Put the box in a 28 &#176;C incubator.</li>
	<li>Between 6-8 hr later, add 1 ml of SDM-79 with 10% FCS containing 2x selective antibiotic to each well. In our experience, cell lines where the genes are tagged on the N terminus are resistant to higher concentrations of selective drug. Therefore, using blasticidin as an example, tagging on the N terminus requires a final concentration of 20 &#181;g/ml blasticidin and tagging on the C terminus requires a final concentration of 10 &#181;g/ml blasticidin. 
	Note: Transgenic cell lines will become visible 9-10 days after transfection.</li>
	<li>Passage at least twice in fresh drug and media prior to analysis. Figure 2B shows the results from a typical 96-well transfection for tagging 96 different proteins on the N terminus. Assess each well by phase contrast microscopy to determine if it contains healthy, drug resistant cells. A healthy well contains predominantly (&#60;95%) highly active cells that are bright in phase contract microscopy. Subsequent analysis by epifluorescence microscopy or western blot is required to determine if the protein has been successfully tagged.</li>
</ol>

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M. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+tightly+regulated+inducible+expression+system+for+conditional+gene+knock-outs+and+dominant-negative+genetics+in+Trypanosoma+brucei.">A tightly regulated inducible expression system for conditional gene knock-outs and dominant-negative genetics in Trypanosoma brucei.</a> Mol Biochem Parasitol. 99 (1), 89-101 (1999).</li><li>Shi, H., Djikeng, A., Mark, T., Wirtz, L. E., Tschudi, C., Ullu, E. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Genetic+interference+in+Trypanosoma+brucei+by+heritable+and+inducible+double-stranded+RNA.">Genetic interference in Trypanosoma brucei by heritable and inducible double-stranded RNA.</a> RNA. 6 (7), 1069-1076 (2000).</li><li>Alsford, S., Turner, D. J., 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=High-throughput+phenotyping+using+parallel+sequencing+of+RNA+interference+targets+in+the+African+trypanosome.">High-throughput phenotyping using parallel sequencing of RNA interference targets in the African trypanosome.</a> Genome Res. 21 (6), 915-924 (2011).</li><li>Alsford, S., Eckert, S., 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=High-throughput+decoding+of+antitrypanosomal+drug+efficacy+and+resistance.">High-throughput decoding of antitrypanosomal drug efficacy and resistance.</a> Nature. 482 (7384), 232-236 (2012).</li><li>Mony, B. M., MacGregor, P., 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=Genome-wide+dissection+of+the+quorum+sensing+signalling+pathway+in+Trypanosoma+brucei.">Genome-wide dissection of the quorum sensing signalling pathway in Trypanosoma brucei.</a> Nature. 505 (7485), 681-685 (2014).</li><li>Rigaut, G., Shevchenko, A., Rutz, B., Wilm, M., Mann, M., S&#233;raphin, B. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=A+generic+protein+purification+method+for+protein+complex+characterization+and+proteome+exploration+:+Abstract+Nature+Biotechnology.">A generic protein purification method for protein complex characterization and proteome exploration : Abstract Nature Biotechnology.</a> Nature Biotechnol. 17 (10), 1030-1032 (1999).</li><li>Los, G. V., Encell, L. 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The authors have nothing to disclose.

Dramatic improvements in the sensitivity of proteomics and transcriptomics methods in the last 5-10 years has provided valuable data on thousands of genes and their products. However, the tools to address the function of these proteins have not kept pace.
Tagging a protein facilitates numerous experiments to determine its function. For example, a protein can be fused to a fluorescent protein in a variety of different colors to facilitate localization and co-localization studies. Tags developed for electron microscopy, such as APEX2 or miniSOG 11,12, allow ultrastructural localization of the tagged protein. Tags for biochemistry, such as the TAP tag, and ProtC-TEV-ProtA (PTP) tag 7,13 allow purification of complexes associated with the protein for identification of binding partners or in vitro biochemical assays.
The specific steps that are critical to the success of the protocol are: the incorporation of DMSO into the PCR Master Mix 1, freezing of the PCR Master Mix 1 prior to the addition of the Master Mix 2, the use of the commercial polymerase in Master Mix 2 and the modification of the cytomix electroporation buffer. In our experience, it is necessary to use double the number of cells for C terminal tagging transfections as for N terminal tagging transfections in order to achieve a similar proportion of positive wells. Therefore, all steps should be performed as described.
This technique is only likely to be successful when transfecting the insect procyclic form trypanosome. Bloodstream forms have a lower transfection efficiency 14, moreover they are likely to die during selection due to density-dependent toxicity that is unrelated to the selective drug. Therefore, our previous protocol represents the current best technology for tagging of bloodstream form trypanosomes 9. It is also likely that the transfection efficiency will vary dependent upon the specific trypanosome isolate. This protocol was optimized using 927 SMOX procyclic forms&#160;&#8211; other strains may require additional optimization. Measures that may increase the probability of success include: increasing the amount of PCR amplicon, increasing the number of cells included in the transfection.
We present a method where hundreds of proteins can be tagged in parallel. This will facilitate large scale studies on localization and interaction, complementing existing large datasets and providing invaluable information to the community. Primer templates are available upon request and a list of plasmids templates is available from: http://www.sdeanresearch.com/

Trypanosoma brucei is a protozoan parasite that causes human African trypanosomasis and nagana in cattle. T. brucei is an ideal organism for the analysis of protein function due to the combination of a high quality genome, numerous proteomics and transcriptomics datasets and well developed molecular tools 1-3. Advances in proteomics and sequencing have resulted in large datasets that highlight potentially interesting genes 4-6; however, many genes have minimal information associated with them in the existing databases. There is therefore a need for a high-throughput method to aid protein functional characterization.

Expression of a tagged protein can give a multiplicity of insights into a protein&#39;s function. For example, a protein tagged with a fluorescent protein or epitope can be localized by fluorescence microscopy, which gives information about where the protein might be exerting its biological effect. Alternatively, a protein tagged with a TAP 7, HaloTag 8 or His tag can be purified for biochemical assays and identification of its interaction partners.

We recently developed a robust tagging methodology for T. brucei&#160; 9. This used long primer PCR to generate the DNA for transfection and allowed the tagging of dozens of proteins in parallel&#160;&#8211; a major improvement to existing protocols. We have now improved the scalability of this protocol in procyclic forms by an order of magnitude. Here, we present our method where we perform the PCR and transfection into 96-well plates, with the recovery and selection occurring in 24-well plates. As hundreds of proteins can now be tagged in parallel this method provides a cost-effective and feasible method for tagging the entire trypanosome genome.

<table><tbody><tr><td>Oligonucleotide</td><td>Life technologies</td><td>na</td><td>desalt only (do not use additional purification)</td></tr><tr><td>DMSO</td><td>Roche</td><td>Included with the Expand HiFi pack</td><td>Must be PCR grade</td></tr><tr><td>dNTP</td><td>any reputable</td><td></td><td></td></tr><tr><td>Expand HiFi polymerase</td><td>Roche</td><td>11759078001</td><td></td></tr><tr><td>BTX ECM830</td><td>Harvard Apparatus</td><td></td><td>Electroporator</td></tr><tr><td>HT-200 plate handler</td><td>Harvard Apparatus</td><td>HT-200</td><td>Handles the 96-well eletroplates</td></tr><tr><td>MOS 96 ELECTROPLATE 4 mm</td><td>Harvard Apparatus</td><td>45-0452</td><td>Electroplate for the 96-well electroporation</td></tr><tr><td>Blasticidin S Hydrochloride</td><td>Melford</td><td>3513-03-9</td><td></td></tr><tr><td>Hygromycin b Gold</td><td>Invivogen</td><td>ant-hg-1</td><td></td></tr><tr><td>Phleomycin</td><td>Melford</td><td>P0187</td><td></td></tr><tr><td>225 cm TC Flask, canted neck, phenolic cap</td><td>Appletonwoods</td><td>BC006</td><td></td></tr><tr><td>24 well culture plates</td><td>Appletonwoods</td><td>BC017</td><td></td></tr><tr><td>Eppendorf&amp;reg; PCR Cooler, iceless cold storage system for 96 well plates and PCR tubes</td><td>Sigma Aldrich</td><td>Z606634-1EA</td><td></td></tr><tr><td>96-well Multiply&amp;reg; PCR plate with lateral skirt</td><td>Sarstedt</td><td>72.1979.203</td><td></td></tr></tbody></table>

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.

In this representative transfection, the primers were designed using the TagIt perl script 9 and synthesized commercially. The 96-well PCR was performed and validated as described (Figure 1A); in this example, 95/96 PCRs were successful (Figure 1B). In our experience, repeating failed reactions either with the original primers or re-synthesized primers does not result in a successful PCR.
The amplicons were transferred into 96-well electroporation plates for electroporation (Figure 2A) and then transferred to 24-well culture plates for recovery and selection. After sufficient time to allow the selection to occur, the wells were scored for survival prior to further analysis. In this example, 88/96 wells (92%) were positive after 15 days selection.
<img alt="Figure 1" src="/files/ftp_upload/54342/54342fig1.jpg" /> 
Figure 1: Validation of long primer PCR. (A) Pattern for transferring PCR products from 96-well PCR plate to the agarose gel for validation of PCR. (B) A representative 96-well PCR validation gel for tagging 95 genes on the N terminus, and 1 amplicon containing no 5&#39; targeting homology to act as a negative control in the transfection. The samples are loaded onto a 1% (w/v) agarose gel and resolved at 100 V for 30 min. <a href="https://www.jove.com/files/ftp_upload/54342/54342fig1large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" src="/files/ftp_upload/54342/54342fig2.jpg" /> 
Figure 2: 96-well electroporation and selection. (A) Pattern for transferring electroporated cells from 96-well electroporation plate to 24-well tissue culture plates. (B) A schematic showing a typical live/dead scoring after 15 days of selection in 24-well plates. Green represents a successful transfection, grey represents a well with only dead cells. In this example, 88/96 (92%) of wells contained successfully transfected parasites. <a href="https://www.jove.com/files/ftp_upload/54342/54342fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Watch this Scientific Journal Video about High-throughput Gene Tagging in Trypanosoma brucei at JoVE.com

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.

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 ...

high-throughput-gene-tagging-in-trypanosoma-brucei

Nanyang Technological University

Research

JoVE Journal

Immunology and Infection

7.9K Views.  University of Oxford. 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.

Video: High-throughput Gene Tagging in Trypanosoma brucei

Examination of the Telomere G-overhang Structure in Trypanosoma brucei

The telomere G-overhang structure has been identified in many eukaryotes including yeast, vertebrates, and Trypanosoma brucei. It serves as the substrate for telomerase for de novo telomere DNA synthesis and is therefore important for telomere maintenance. T. brucei is a protozoan parasite that causes sleeping sickness in humans and nagana in cattle. Once infected mammalian host, T. brucei cell regularly switches its surface antigen to evade the host's immune attack. We have recently demonstrated that the T. brucei telomere structure plays an essential role in regulation of surface antigen gene expression, which is critical for T. brucei pathogenesis. However, T. brucei telomere structure has not been extensively studied due to the limitation of methods for analysis of this specialized structure. We have now successfully adopted the native in-gel hybridization and ligation-mediated primer extension methods for examination of the telomere G-overhang structure and an adaptor ligation method for determination of the telomere terminal nucleotide in T. brucei cells. Here, we will describe the protocols in detail and compare their different advantages and limitations.

Examination of the Telomere G-overhang Structure in Trypanosoma brucei

Telomeres are essential for chromosome stability and the telomere G-overhang structure is essential for telomerase-mediated telomere maintenance. We have recently adopted two methods for detecting the telomere G-overhang structure in Trypanosoma brucei, which are native in-gel hybridization and ligation-mediated primer extension, which will be described.

The telomere G-overhang structure has been identified in many eukaryotes including yeast, vertebrates, and Trypanosoma brucei. It serves as the substrate ...

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

Genetic Manipulation in &Delta;ku80 Strains for Functional Genomic Analysis of Toxoplasma gondii

Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene function and phenotype(s). The development of mutant strains with targeted gene deletions, targeted mutations, complemented gene function, and/or tagged genes provides powerful strategies to address gene function, particularly if these genetic manipulations can be efficiently targeted to the gene locus of interest using integration mediated by double cross over homologous recombination.
Due to very high rates of nonhomologous recombination, functional genomic analysis of Toxoplasma gondii has been previously limited by the absence of efficient methods for targeting gene deletions and gene replacements to specific genetic loci. Recently, we abolished the major pathway of nonhomologous recombination in type I and type II strains of T. gondii by deleting the gene encoding the KU80 protein1,2. The &Delta;ku80 strains behave normally during tachyzoite (acute) and bradyzoite (chronic) stages in vitro and in vivo and exhibit essentially a 100% frequency of homologous recombination. The &Delta;ku80 strains make functional genomic studies feasible on the single gene as well as on the genome scale1-4.
Here, we report methods for using type I and type II &Delta;ku80&Delta;hxgprt strains to advance gene targeting approaches in T. gondii. We outline efficient methods for generating gene deletions, gene replacements, and tagged genes by targeted insertion or deletion of the hypoxanthine-xanthine-guanine phosphoribosyltransferase (HXGPRT) selectable marker. The described gene targeting protocol can be used in a variety of ways in &Delta;ku80 strains to advance functional analysis of the parasite genome and to develop single strains that carry multiple targeted genetic manipulations. The application of this genetic method and subsequent phenotypic assays will reveal fundamental and unique aspects of the biology of T. gondii and related significant human pathogens that cause malaria (Plasmodium sp.) and cryptosporidiosis (Cryptosporidium).

Here we report a method for using type I and type II &Delta;ku80 strains of Toxoplasma gondii to efficiently generate targeted gene deletions and gene replacements for functional genomic analysis.

Targeted genetic manipulation using homologous recombination is the method of choice for functional genomic analysis to obtain a detailed view of gene ...

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 ...

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

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 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 ...

Detection of Trypanosoma brucei Variant Surface Glycoprotein Switching by Magnetic Activated Cell Sorting and Flow Cytometry

Trypanosoma brucei, a protozoan parasite that causes both Human and Animal African Trypanosomiasis (known as sleeping sickness and nagana, respectively) cycles between a tsetse vector and a mammalian host. It evades the mammalian host immune system by periodically switching the dense, variant surface glycoprotein (VSG) that covers its surface. The detection of antigenic variation in Trypanosoma brucei can be both cumbersome and labor intensive. Here, we present a method for quantifying the number of parasites that have 'switched' to express a new VSG in a given population. The parasites are first stained with an antibody against the starting VSG, and then stained with a secondary antibody attached to a magnetic bead. Parasites expressing the starting VSG are then separated from the rest of the population by running the parasites over a column attached to a magnet. Parasites expressing the dominant, starting VSG are retained on the column, while the flow-through contains parasites that express a new VSG as well as some contaminants expressing the starting VSG. This flow-through population is stained again with a fluorescently labeled antibody against the starting VSG to label contaminants, and propidium iodide (PI), which labels dead cells. A known number of absolute counting beads that are visible by flow cytometry are added to the flow-through population. The ratio of beads to number of cells collected can then be used to extrapolate the number of cells in the entire sample. Flow cytometry is used to quantify the population of switchers by counting the number of PI negative cells that do not stain positively for the starting, dominant VSG. The proportion of switchers in the population can then be calculated using the flow cytometry data.

Detection of Trypanosoma brucei Variant Surface Glycoprotein Switching by Magnetic Activated Cell Sorting and Flow Cytometry

African trypanosomes grown in vitro undergo antigenic variation at a low rate, such that populations are made up of parasites expressing a dominant variant surface glycoprotein (VSG) type and a small population of "switched" variants. This protocol describes a fast method for detecting and quantifying these populations.

Trypanosoma brucei, a protozoan parasite that causes both Human and Animal African Trypanosomiasis (known as sleeping sickness and nagana, respectively) ...

Purification of Extracellular Trypanosomes, Including African, from Blood by Anion-Exchangers (Diethylaminoethyl-cellulose Columns)

This method allows the separation of trypanosomes, parasites responsible for animal and human African trypanosomiasis (HAT), from infected blood. This is the best method for diagnosis of first stage HAT and furthermore this parasite purification method permits serological and research investigations.
HAT is caused by Tsetse fly transmitted Trypanosoma brucei gambiense and T. b. rhodesiense. Related trypanosomes are the causative agents of animal trypanosomiasis. Trypanosome detection is essential for HAT diagnosis, treatment and follow-up. The technique described here is the most sensitive parasite detection technique, adapted to field conditions for the diagnosis of T. b. gambiense HAT and can be completed within one hour. Blood is layered onto an anion-exchanger column (DEAE cellulose) previously adjusted to pH 8, and elution buffer is added. Highly negatively charged blood cells are adsorbed onto the column whereas the less negatively charged trypanosomes pass through. Collected trypanosomes are pelleted by centrifugation and observed by microscopy. Moreover, parasites are prepared without cellular damage whilst maintaining their infectivity.
Purified trypanosomes are required for immunological testing; they are used in the trypanolysis assay, the gold standard in HAT serology. Stained parasites are utilized in the card agglutination test (CATT) for field serology. Antigens from purified trypanosomes, such as variant surface glycoprotein, exoantigens, are also used in various immunoassays. The procedure described here is designed for African trypanosomes; consequently, chromatography conditions have to be adapted to each trypanosome strain, and more generally, to the blood of each species of host mammal.
These fascinating pathogens are easily purified and available to use in biochemical, molecular and cell biology studies including co-culture with host cells to investigate host-parasite relationships at the level of membrane receptors, signaling, and gene expression; drug testing in vitro; investigation of gene deletion, mutation, or overexpression on metabolic processes, cytoskeletal biogenesis and parasite survival.

Purification of Extracellular Trypanosomes, Including African, from Blood by Anion-Exchangers Diethylaminoethyl-cellulose Columns

This method of trypanosome separation from blood depends on their surface charge being less negative than mammalian blood cells. Infected blood is placed and treated on an anion-exchanger column. This method, the most fitting diagnostic for African trypanosomiasis, provides purified parasites for immunological, biological, biochemical, pharmaceutical and molecular biology investigations.

This method allows the separation of trypanosomes, parasites responsible for animal and human African trypanosomiasis (HAT), from infected blood. This is ...

Development of pHluorin2-Based Sensor for Glycosomal pH in Trypanosoma brucei

Measuring Dynamic Glycosomal pH Changes in Living Trypanosoma brucei

Glucose metabolism is critical for the African trypanosome, Trypanosoma brucei, as an essential metabolic process and regulator of parasite development. Little is known about the cellular responses generated when environmental glucose levels change. In both bloodstream and procyclic form (insect stage) parasites, glycosomes house most of glycolysis. These organelles are rapidly acidified in response to glucose deprivation, which likely results in the allosteric regulation of glycolytic enzymes such as hexokinase. In previous work, localizing the chemical probe used to make pH measurements was challenging, limiting its utility in other applications.
This paper describes the development and use of parasites that express glycosomally localized pHluorin2, a heritable protein pH biosensor. pHluorin2 is a ratiometric pHluorin variant that displays a pH (acid)-dependent decrease in excitation at 395 nm while simultaneously yielding an increase in excitation at 475 nm. Transgenic parasites were generated by cloning the pHluorin2 open reading frame into the trypanosome expression vector pLEW100v5, enabling inducible protein expression in either lifecycle stage. Immunofluorescence was used to confirm the glycosomal localization of the pHluorin2 biosensor, comparing the localization of the biosensor to the glycosomal resident protein aldolase. The sensor responsiveness was calibrated at differing pH levels by incubating cells in a series of buffers that ranged in pH from 4 to 8, an approach we have previously used to calibrate a fluorescein-based pH sensor. We then measured pHluorin2 fluorescence at 405 nm and 488 nm using flow cytometry to determine glycosomal pH. We validated the performance of the live transgenic pHluorin2-expressing parasites, monitoring pH over time in response to glucose deprivation, a known trigger of glycosomal acidification in PF parasites. This tool has a range of potential applications, including potentially being used in high-throughput drug screening. Beyond glycosomal pH, the sensor could be adapted to other organelles or used in other trypanosomatids to understand pH dynamics&#160;in the live cell setting.

Author Spotlight: Advancements in Glycosomal pH Monitoring in Trypanosoma brucei Using pHluorin2 Biosensor

We describe a method to study how pH responds to environmental cues in the glycosomes of the bloodstream form of African trypanosomes. This approach involves a pH-sensitive heritable protein sensor in combination with flow cytometry to measure pH dynamics, both as a time-course assay and in a high-throughput screen format.

Glucose metabolism is critical for the African trypanosome, Trypanosoma brucei, as an essential metabolic process and regulator of parasite development. ...

RNA Catalyst as a Reporter for Screening Drugs against RNA Editing in Trypanosomes

Substantial progress has been made in determining the mechanism of mitochondrial RNA editing in trypanosomes. Similarly, considerable progress has been made in identifying the components of the editosome complex that catalyze RNA editing. However, it is still not clear how those proteins work together. Chemical compounds obtained from a high-throughput screen against the editosome may block or affect one or more steps in the editing cycle. Therefore, the identification of new chemical compounds will generate valuable molecular probes for dissecting the editosome function and assembly. In previous studies, in vitro editing assays were carried out using radio-labeled RNA. These assays are time consuming, inefficient and unsuitable for high-throughput purposes. Here, a homogenous fluorescence-based &#8220;mix and measure&#8221; hammerhead ribozyme in vitro reporter assay to monitor RNA editing, is presented. Only as a consequence of RNA editing of the hammerhead ribozyme a fluorescence resonance energy transfer (FRET) oligoribonucleotide substrate undergoes cleavage. This in turn results in separation of the fluorophore from the quencher thereby producing a signal. In contrast, when the editosome function is inhibited, the fluorescence signal will be quenched. This is a highly sensitive and simple assay that should be generally applicable to monitor in vitro RNA editing or high throughput screening of chemicals that can inhibit the editosome function.

A highly sensitive ribozyme-based assay, applicable to high-throughput screening of chemicals targeting the unique process of RNA editing in trypanosomatid pathogens, is described in this paper. Inhibitors can be used as tools for hypothesis-driven analysis of the RNA editing process and ultimately as therapeutics.

Substantial progress has been made in determining the mechanism of mitochondrial RNA editing in trypanosomes. Similarly, considerable progress has been ...

High-throughput Gene Tagging in Trypanosoma brucei

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