The microbial parasite Blastocystis colonizes the large intestines of numerous animal species and increasing evidence has linked Blastocystis infection to enteric diseases with signs and symptoms including abdominal pain, constipation, diarrhea, nausea, vomiting, and flatulence. It has also recently been reported to be an important member of the host intestinal microbiota. Despite significant advances in our understanding of Blastocystis cell biology and host-parasite interactions, a genetic modification tool is absent. In this study, we successfully established a robust gene delivery protocol for Blastocystis subtype 7 (ST7) and ectopic protein expression was further tested using a high sensitivity nano-luciferase (Nluc) reporter system, with promoter regions from several genes. Among them, a strong promoter encompassing a region upstream of the legumain 5′ UTR was identified. Using this promoter combined with the legumain 3′ UTR, which contains a conserved, precise polyadenylation signal, a robust transient transfection technique was established for the first time in Blastocystis. This system was validated by ectopic expression of proteins harbouring specific localization signals. The establishment of a robust, reproducible gene modification system for Blastocystis is a significant advance for Blastocystis research both in vitro and in vivo. This technique will spearhead further research to understand the parasite’s biology, its role in health and disease, along with novel ways to combat the parasite.
Blastocystis is a common enteric microbial eukaryote belonging to the Stramenopiles, infecting more than 1 billion humans along with a large variety of non-human hosts, such as farm animals, rodents, birds, reptiles and others1,2. Clinically, increasingly evidence has shown that its presence is associated with abdominal pain, constipation, diarrhea, nausea, vomiting, flatulence, irritable bowel syndrome-like symptoms (IBS)3,4,5,6, and even skin disorders7. Studies into the global causes of severe diarrhea, estimated to be responsible for 10.5% of overall child mortality8 in young children, have identified Blastocystis as an important pathogen, just after Cryptosporidium and Giardia9,10. Blastocystis is also an opportunistic pathogen in the context of human immunodeficiency virus (HIV)- AIDS infections11.
In vitro studies reveal that both parasite and parasite lysates exert damaging effects on intestinal epithelial cells, cause apoptosis and degradation of tight junction proteins (occludin and ZO1), resulting in increased intestinal permeability12,13. Adhesion of trophic forms to the intestinal epithelium and release of cysteine proteases appear to be the major triggers leading to pathogenesis, although axenic co-culture conditions and varying ratios of infection may not reflect the adhesion14,15 or pathogenesis in vivo16,17. Putative virulence factors identified include cysteine proteases legumain18,19 and cathepsin B20. Blastocystis spp. also possess immuno-modulatory effects including degradation of IgA21, inhibition of iNOS22, and upregulation of proinflammatory cytokines IL8 and GM-CSF in intestinal epithelial cells23, and IL1β, IL6 and TNFα in murine macrophages24,25. Blastocystis spp. may also dampen response to LPS in intestinal epithelial cells and monocytes26. Studies in rodent models and naturally infected pigs indicate that the parasite causes mucosal sloughing, increases goblet cell mucin, enhances intestinal permeability17,27,28, and induces a pro-inflammatory cytokine response with upregulation of TNFα, IFNγ and IL1217.
The development of suitable genetic tools including a transfection system for Blastocystis spp. to facilitate molecular studies will be an invaluable contribution in delineating the roles of virulence factors. Biologically, Blastocystis is a strict anaerobe. Although numerous intracellular organelles resembling mitochondria (mitochondrial-related organelles; MROs) exist, they are completely devoid of cytochrome enzymes29. These MROs have the property of both mitochondria of aerobes and hydrogenosomes of anaerobes, and are involved in various metabolic pathways including amino acid metabolism, iron-sulfur cluster biogenesis, oxygen stress response and a partial tricarboxylic acid cycle30,31. The organism is also capable of synthesizing various essential cellular phospholipids, and accumulates these within storage vacuoles32,33. Genetic modifications in which specific parasite molecules or organelles are rendered visible via reporter systems or epitope tags may lead to a better understanding of the cell biology of this poorly understood but widespread parasite.
Results and Discussion
Optimization of DNA delivery and transfection method
A genetic tool for manipulating gene expression in Blastocystis is hitherto lacking, although transient or stable transfection approaches have been successfully achieved in several parasitic protists, including trypanosomatid parasites Trypanosoma brucei34,35, T. cruzi36 and Leishmania spp.36; apicomplexan parasites Plasmodium spp.37, Cryptosporidium spp38. and Toxoplasma gondii39; parasitic excavates such as Trichomonas vaginalis40 and Giardia intestinalis41,42 and parasitic amoeba Entamoeba histolytica43. Nucleic acids were introduced by using Bio-Rad Gene Pulser Xcell or BTX ECM 630 Electroporation systems with commonly used cytomix buffer44 (occasionally supplemented with 2 mM ATP and 5 mM glutathione), or by using Amaxa Nucleofector device and human T-cell transfection reagent. Although the latter approach offers higher transfection efficiency in many kinds of cells, the high cost and “blackboxing” transfection programs and reagents restricted its usage in many labs.
A reliable gene delivery approach should introduce electropores in as many cells as possible and the pores should be sealed in sufficient number of cells to maintain the viability of cultures after transfection. Based on these criteria, we evaluated different transfection programs using the Bio-Rad Gene Pulser and cytomix supplemented with 2 mM ATP and 5 mM glutathione via an approach shown in Fig. 1A, in Blastocystis. To determine the electropore-forming rate, cells were washed with and resuspended in cytomix, then stained with 5 μg/ml of the cell-impermeable dye propidium iodide (PI) before transfection. The PI-positive cells were enumerated using a hemocytometer under fluorescence microscopy, and the percentage of PI-positive cells post-transfection was calculated (Fig. 1A,B). To determine the survival rate, similar batches of cells without PI staining were also subjected to electroporation and stained with PI after 12 hours’ incubation to allow membrane sealing. The PI-negative cells were counted and the survival rates were calculated (Fig. 1A,B). Two protocols (“Exponential” and “Time Constant”) and several programs with different voltage (“Exponential protocol”) or different voltage and electroporation time (“Time Constant” protocol) were evaluated. A low-voltage program was selected due to the large size of Blastocystis cells, which resulted in arcs when the voltage was set too high or when the electroporation time was too long (Fig. 1B). Amongst all the tested programs, one pulse with 370 V, 30 ms under the “Time Constant” protocol achieved the best result, with a 94.3% electropore-forming rate and 9.4% survival rate (Fig. 1B).
It was noteworthy that during these evaluations, only 2 × 107 cells were used for each transfection. When more cells were used, the probability of arcing was higher, and shorter electroporation times should therefore be employed. In the case of 108 cells per transfection, pulsing with 370 V and 20 ms resulted in excellent transfection efficiency (Ref. the followed section).
Oscillating electroporation instead of standard exponential decay electroporation was used for the slime mold D. discoidum45 and mammalian NIH 3T3 cells46 to increase the transformation efficiency. In this case, the Bio-Rad Gene Pulser II system is equipped with a radiofrequency module that can convert a direct electric field into an oscillatory field. This produces several oscillating pulses rather than a single decaying pulse and improves transformation efficiency by 20-fold in D. discoidum45. Since the Gene Pulser in our laboratory lacks this radiofrequency module, this approach could not be tested.
Evaluation of NanoLuc luciferase (NLuc) expression in transfected Blastocystis
To drive efficient ectopic gene expression, a strong promoter and a polyadenylation signal in the 3′ UTR region are required. A crucial element of successful genetic manipulation in various protistan parasites involves the usage of species-specific, endogenous promoters that regulated a variety of housekeeping genes including actin, tubulin and GAPDH, or conserved essential genes such as histone, myosin, and enolase38,47,48,49,50. To test promoter activity, Photinus pyralis (firefly) luciferase assays are commonly used due to the extremely rapid reaction, low cost, high sensitivity and the utilization of commercially available non-radioactive substrates51. A new smaller and ATP-independent luciferase, deep-sea shrimp Oplophorus gracilirostris NanoLuc luciferase (Nluc) was developed recently by Promega. Together with its synthetic imidazopyrazinone substrate furimazine, it produces signals averaging over 150 times brighter than firefly luciferase52. The high stability and sensitivity of this enzyme render it an ideal tool for testing promoter activity in Blastocystis cells.
We replaced the YFP-tag with Nluc in the pXS2 vector that was used for ectopic expression of target genes in T. brucei53. The Nluc cassette was flanked by putative promoter sequences from the 5′ UTR regions of four different Blastocystis genes: two house-keeping genes GAPDH (GAPDHP) and actin (ActP), enolase (EnoP) involved in energy metabolism, and the membrane enzyme legumain that is highly expressed in Blastocystis19, together with a 230-bp potential polyadenylation signal-containing 3′ UTR downstream of the legumain coding region (Fig. 2A,B). For the potential legumain promoter, three fragments of 630, 1140 and 1450 bp upstream of the legumain (without intron) open reading frame (ORF) (designated LeguP2, P1 and P3, respectively) were tested (Fig. 2A,B). All the constructs (25 μg per transfection) were transfected into Blastocystis ST7 B cells (2 × 107 each) using the program indicated above, and the relative Nluc luminescence was assayed using a HiDex multimodel microplate reader. Among the promoters tested, only the Nluc driven by LeguP1 produced strong luninescence, but not those by LeguP2, LeguP3, GAPDHP, EnoP and ActP (Fig. 2C). The supplementation of 2 mM ATP and 5 mM glutathione augmented the transfection efficiency about 3-fold (Fig. 2C).
The Nluc luminescence correlated with the amount of DNA (Fig. 3A) and the number of parasites (Fig. 3B) used for transfection. High transfection efficiency was achieved by using 100 μg of plasmid DNA and 108 cells (Fig. 3C), and this condition (370 V, 20 ms under Time Constant protocol) was used for further transfections. To test the best detection time point, the Nluc luminescence assays were conducted in transfectants with 6, 9, 12, 15, 18, 21 or 24 hours incubation post transfection. After 6 hours incubation, the expression was detectable (Fig. 3D). A relative higher expression was achieved after 15 hours’ incubation and slightly increasing at 24 hours (Fig. 3D). Nluc luminescence was also shown to be dependent upon the presence of parasite-specific promoters: the T. brucei PARP promoter only drives Nluc expression in trypanosomes, while the LeguP1 only drives the expression in Blastocystis (Fig. 3E).
In Blastocystis, an intriguing phenomenon was documented by Klimes et al.54; i.e. polyadenylation-mediated creation of termination codons occurs in about 15% of all nuclear-coding genes. In the 3′ UTR of these genes, a conserved motif TGTTTGTT is located 4 nucleotides downstream of the polyadenylation site54. This motif was found in legumain 3′ UTR, and a potential polyadenylation site (nucleotide T) exists 3 or 5 nucleotides upstream of the motif (Fig. 4A). To test whether this motif at the 3′ UTR region contributes to Nluc expression, two approaches were adopted. Firstly, we mutated the conserved motif to TCAAAGTT and the upstream two thymines (+3 and +5) to adenines (Fig. 4A). Secondly, we replaced the legumain 3′ UTR in the LeguP1 vector with a 150-bp GAPDH 3′ UTR fragment that lacks the conserved motif (Figs 2A and 4A). The Nluc expression level diminished 324 and 148-fold in cells transfected with vectors containing mutated legumain 3′ UTR or GAPDH 3′ UTR, respectively (Fig. 4B), indicating that the existence of a precise polyadenylation signal54 contributes significantly to efficient gene expression, although it is yet unknown how the expression was regulated in genes without the conserved motif in their 3′ UTRs.
There is a scarcity of information on how gene expression is controlled and regulated in Blastocystis. This limited our ability to select suitable promoters for ectopic gene expression. Thus far, only one promoter (LeguP1) worked well in our experience. A larger number of strong promoters need to be identified in order to expand the functionality of the existing protocol into possibilities of stable or regulatable transfection.
Determination of the efficiency of transient transfection
To evaluate the efficiency of transient transfection in Blastocystis, 108 cells were transfected with 100 μg of pXS2-PLegumain vector. After transfection, cells were aliquoted at 2-fold series dilution, and the luminescence was determined for each aliquot. After 14 dilution steps, the reading of Nluc luminescence decreased to background level (Fig. 5). Thus, the transfection efficacy is approximately 10−4 (214 ÷ 108).
The expression level in individual cells (reflecting the promoter strength) was not determined. However, since it was comparable with T. brucei, which possesses high ectopic expression level (Fig. S1), this suggested that the legumain promoter is sufficiently strong for further optimization towards stable transfection.
Evaluation the expression of the eGFP and small epitope Ty-tagged proteins
To test whether the vector can be used to track protein expression by microscopy, we replaced Nluc with eGFP together with a small Ty tag55 (Figs 6A and S2, the full sequence of the vector can be found in Supplement Data 1). The transfectants were stained with anti-Ty antibody or MitoTracker and detected by microscopy using the eGFP channel (eGFP tag) and dsRed channel (Ty tag). Less than 1% of cells with normal cell morphology exhibited strong Ty signal (red) distributed in the whole cell, but only background level of eGFP signal was displayed (Fig. 6B). Based on previous studies, oxygen is essential for the post-translational folding of eGFP into the fluorescent chromophore56, which may have hindered our detection of GFP fluorescence in the anaerobic conditions of our assays. To address this limitation, cells were transfected with proteins involved in the mitochondrial iron-sulfur cluster biosynthesis, IscU and IscS, fused to eGFP and incubated in oxygen for approximately 1 h to allow for proper GFP folding. We observed punctuated concentration of the signal in intracellular structures and co-localisation with MitoTracker, suggesting a localization in Blastocystis MROs (Fig. 7). Both proteins were previously shown to localize in the Blastocystis MROs31, and thus our results confirm successful tagging of these proteins and pathway in the organelles. In future studies, we will employ the anaerobic cyan-green fluorescent protein evoglow Pp1 (GFPana or aFP)57,58 instead of eGFP.
Towards stable transfection and potential applications
To achieve a constant expression of a target protein, several issues need to be addressed: two strong promoters to drive target and drug resistance genes; sensitive drugs for transfectant selection and incorporation of the expression cassette into the host chromosome. Till now, only one strong promoter was identified and a more comprehensive study needs to be performed to identify more robust promoters. Blastocystis genomes apparently do not encode components of the nonhomologous end-joining machinery, suggesting that homologous recombination is the principal mechanism for the repair of double-stranded DNA breaks59. Whether the recombination rate is sufficient for stable transfection selection requires further exploration.
In the post-genomic era, the generation of ectopic expression, knockdown or knockout cells using genetic tools is encouraging and will be crucial to understand the function of Blastocystis genes. Here we successfully developed a transient transfection system that facilitates the study of Blastocystis to advance in numerous new directions. To further optimize and exploit this technique, the limited knowledge of homologous recombination in Blastocystis is a critical area that needs to be addressed. The optimized genetic tools will be essential in studies to understand the function and localization of novel Blastocystis proteins as well as the orthologs of known eukaryotic and/or prokaryotic proteins that have been identified in the Blastocystis genome project. Notably, the proteins involved in host-microbiota-Blastocystis interactions, metabolic and biochemical pathways can be further characterized as these are of interest from both biomedical and evolutionary perspectives. Furthermore, such reliable expression of homologous or heterologous proteins and antigens in Blastocystis may facilitate potentially important in vivo studies into understanding the various adaptations of the parasite31,59 and the modulation of relevant immune responses in the gastrointestinal tract60.
Cell strain and culture
An axenized Blastocystis isolate ST7 B61 was used in this study. Cells were cultured in pre-reduced Iscove’s modified Dulbecco’s medium (IMDM) (Thermo Scientific) supplemented with 10% horse serum (Gibco) at 37 °C62. The culture tubes were maintained inside 2.5-liter sealed anaerobic jars with an anaerobic gas pack (Oxoid).
Procyclic T. brucei YTat1.1 cells were maintained in Cunningham’s medium supplemented with 15% fetal bovine serum (FBS) at 27 °C63.
Nluc fragment was amplified from pNL1.1[Nluc] vector (#N1001, Promega) using primers Nluc-f (5′-ctagctagcatggtcttcacactcgaagatttc-3′) and Nluc-r (5′-cgggatccttacgccagaatgcgttcg-3′) and cloned into NheI/BamHI-digested pXS2 vector to generate pXS2-Nluc plasmid.
The legumain and GAPDH 3′ UTRs were amplified from Blastocystis genome DNA using the respective primer pairs: Legu-3utr-f (5′-cgggatccgcattgagtgtatatgtttgttataaaac-3′) and Legu-3utr-r (5′-ggaattcccatcgtctgctttatccac-3′); GAPDH-utr-f (5′-cgggatccatccgtcagtgaccaaacgagt-3′) and GAPDH-utr-r (5′-ggaattcgtagtaatgaatgcttctgaagaaaggtt-3′). The 3′ UTRs were then digested by BamHI/EcoRI and cloned into the BamHI/EcoRI-digested pXS2-Nluc vector, resulting in pXS2-Nluc-GAPDH3UTR and pXS2-Nluc-Legu3UTR plasmids.
The LeguP1, LeguP2 and LeguP3 promoters were amplified from Blastocystis ST7 B genome DNA using forward primers LeguP1-f (5′-cccaagctt cgcacgtagtcagccgtt-3′), LeguP2-f (5′-cccaagcttgatcagtcggcacgttgtg-3′) and LeguP3-f (5′-cccaagcttcttgtacggattaacccatgtaaatg-3′), paired with the same reverse primer LeguP-r (5′-ctagctagcttacaaattttttatggtattatttctact-3′). Since a HindIII restriction site exists in the GAPDH promoter region, the GAPDHP with a mutated HindIII site was amplified by overlap-extension PCR using these 4 primers: GAPDHP-f1 (5′-cccaagcttgtaggcacgctctctgaatattcc-3′), GAPDHP-r1 (5′-acagaaaactgaagcatgatagagcga aaca-3′), GAPDHP-f2 (5′-tgtttcgctctatcatgcttcagttttctgt-3′) and GAPDHP-r2 (5′-ctagct agcatggaattgatatgataaagaaatcaattc-3′). EnoP and ActP promoters were amplified using the corresponding primer pairs: EnoP-f (5′-cccaagcttctactatagcagaatgtggtactgcata-3′) and EnoP-r (5′-ctagctagctctatagaaaaattttgaggatgagg-3′); ActP-f (5′-cccaagcttgcccgaccttgacttagcc-3′) and ActP-r (5′-ctagctagcatcgatgagttctttgcgtctg-3′). The amplified promoters were digested with HindIII/NheI and cloned into HindIII/NheI digested pXS2-Nluc-Legu3UTR or pXS2-Nluc-GAPDH3UTR to obtain the final vectors used for transfection (Fig. 2B).
To express the fluorescent reporter protein, an eGFP and small isotope Ty fusion fragment was amplified from pDEX-777 vector using primers eGFP-f (5′-ctagctagcagcaagggcgaggagct-3′) and eGFP-r (5′-cgggatccttagtcaagtggatcttggttagtatgg acc-3′), and digested by NheI/BamHI. The digested fragment was cloned into digested pXS2 vector to generate pXS2-eGFP-Ty plasmid.
To express IscU and IscS proteins along with eGFP tag, the IscU and IscS genes were amplified from previously published Blastocystis clones31 using primers IscU-F (5′-gctagcatgtatgcattaaccagatcg-3′), IscU-R (5′-gctagcctttgacttcttttcgctctt-3′), IscS-F (5′-gctagcatgctctcccgatttagcagt-3′) and IscS-R (5′-gctagcatgggtgctcctcttgatcgc-3′) and digested by NheI. The digested fragment was cloned into the digested pXS2-LeguP-eGFP-Ty-LeguUTR plasmid.
RNA extraction, first-strand cDNA synthesis and reverse-transcription (RT)-PCR
Total RNA was extracted from 5 × 107 Blastocystis ST7 B cells with TRIzol Reagent (Life Technologies, 15596-026) following the manufacturer’s protocol. After treaent with RNase-free DNase I (Roche, 04 716 728 001), the first-strand cDNA was synthesized with M-MLV Reverse Transcriptase (Invitrogen, 28025-013) using the standard protocol and PCR was performed with the primers indicated above.
For transfection of Blastocystis, cells were cultured to log-phase and harvested by centrifugation at 1,000 g for 10 min at room temperature. Cells were then washed once with pre-reduced incomplete cytomix buffer (10 mM K2HPO4/KH2PO4, pH 7.6, 120 mM KCl, 0.15 mM CaCl2, 25 mM HEPES, 2 mM EGTA, and 5 mM MgCl2), and then resuspended in pre-reduced complete cytomix (incomplete cytomix supplemented with 2 mM ATP and 5 mM glutathione). These cells were ready for transfection.
Blastocystis cells were mixed with an appropriate amount of plasmid DNA in a 0.4-cm transfection cuvette (Bio-Rad) and subjected to 1 pulse using the Bio-Rad Gene Pulser electroporation system under the protocol and programs shown in Fig. 1B. After electroporation, cells were transferred into fresh pre-reduced culture medium and maintained at 37 °C anaerobically for 12–16 hrs.
For stable transfection in Trypanosoma brucei, the constructs were linearized with MluI and transfected into YTat1.1 cells using Bio-Rad Gene Pulser. The stable transformants were selected with 10 μg/ml blasticidin (Invitrogen, R210-01).
Nluc luciferase assay
The Nluc luminescence was monitored using Nano-Glo Luciferase Assay System (Promega) following the protocol provided. Briefly, the cells were collected and lysed using 1 × Luciferase Cell Culture Lysis reagent (25 mM Tris-phosphate (pH 7.8), 2 mM DTT, 2 mM 1,2-diaminocyclohexane-N,N,N′,N′-tetraacetic acid, 10% glycerol, 1% Triton X-100). Cell lysate was mixed with equal volume of Nano-Glo Luciferase Assay reagents and the relative luminescence units (RLU) was measured using Hidex Sense multimode microplate reader (Hidex).
Blastocystis ST7 B cells were washed with and resuspended in PBS. Cells were fixed with 4% paraformaldehyde (PFA, Sigma, P6148) in PBS at room temperature for 30 min, permeabilized with 1% NP-40 (Igepal CA-630, Sigma, I8896) and blocked with 3% BSA (Sigma, A7906) before antibody staining. Monoclonal anti-Ty55 (a gift from Prof. Keith Gull, University of Oxford) was used at 1:100 to label eGFP-Ty, followed by staining with goat-anti-mouse IgG H&L (Alexa Fluor 594) secondary antibody at 1:2000. Cells were then mounted onto slides and observed using an Axioplan2 inverted microscope (Carl Zeiss MicroImaging, Germany) equipped with a CoolSNAP HQ2 camera (Photometrics, USA) and a Plan-Apochromat 63×/1.40 oil DIC objective.
For nanoluciferase (NLuc) assay, three independent transfections were performed and relative luminescence unit (RLU) readings were shown as mean ± SD. Statistical analyses were performed using the 2-tailed, equal variance Student t test.
Yoshikawa, H., Morimoto, K., Wu, Z., Singh, M. & Hashimoto, T. Problems in speciation in the genus Blastocystis. Trends Parasitol 20, 251–5 (2004).
Clark, C. G., van der Giezen, M., Alfellani, M. A. & Stensvold, C. R. Recent developments in Blastocystis research. Adv Parasitol 82, 1–32 (2013).
Dogruman-Al, F. et al. Blastocystis subtypes in irritable bowel syndrome and inflammatory bowel disease in Ankara, Turkey. Mem Inst Oswaldo Cruz 104, 724–7 (2009).
Stensvold, C. R. et al. Subtype distribution of Blastocystis isolates from synanthropic and zoo animals and identification of a new subtype. Int J Parasitol 39, 473–9 (2009).
Tan, K. S. New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin Microbiol Rev 21, 639–65 (2008).
Ajjampur, S. S. & Tan, K. S. Pathogenic mechanisms in Blastocystis spp. - Interpreting results from in vitro and in vivo studies. Parasitol Int 65, 772–779 (2016).
Casero, R. D., Mongi, F., Sanchez, A. & Ramirez, J. D. Blastocystis and urticaria: Examination of subtypes and morphotypes in an unusual clinical manifestation. Acta Trop 148, 156–61 (2015).
Liu L. et al. Child Health Epidemiology Reference Group of WHO, Unicef. 2012. Global, regional, and national causes of child mortality: an updated systematic analysis for 2010 with time trends since 2000. Lancet 379, 2151–61 (2013).
Ramirez, J. D., Florez, C., Olivera, M., Bernal, M. C. & Giraldo, J. C. Blastocystis subtyping and its association with intestinal parasites in children from different geographical regions of Colombia. PLoS One 12, e0172586 (2017).
Rostami, A. et al. The role of Blastocystis sp. and Dientamoeba fragilis in irritable bowel syndrome: a systematic review and meta-analysis. Parasitol Res 116, 2361–2371 (2017).
Brites, C., Barberino, M. G., Bastos, M. A., Sampaio, S. M. & Silva, N. Blastocystis hominis as a Potential Cause of Diarrhea in AIDS Patients: a Report of Six Cases in Bahia, Brazil. Braz J Infect Dis 1, 91–94 (1997).
Mirza, H., Wu, Z., Teo, J. D. & Tan, K. S. Statin pleiotropy prevents rho kinase-mediated intestinal epithelial barrier compromise induced by Blastocystis cysteine proteases. Cell Microbiol 14, 1474–84 (2012).
Puthia, M. K., Sio, S. W., Lu, J. & Tan, K. S. Blastocystis ratti induces contact-independent apoptosis, F-actin rearrangement, and barrier function disruption in IEC-6 cells. Infect Immun 74, 4114–23 (2006).
Fayer, R. et al. Blastocystis tropism in the pig intestine. Parasitol Res 113, 1465–72 (2014).
Wang, W., Bielefeldt-Ohmann, H., Traub, R. J., Cuttell, L. & Owen, H. Location and pathogenic potential of Blastocystis in the porcine intestine. PLoS One 9, e103962 (2014).
Moe, K. T. et al. Experimental Blastocystis hominis infection in laboratory mice. Parasitol Res 83, 319–25 (1997).
Iguchi, A., Yoshikawa, H., Yamada, M., Kimata, I. & Arizono, N. Expression of interferon gamma and proinflammatory cytokines in the cecal mucosa of rats experimentally infected with Blastocystis sp. strain RN94-9. Parasitol Res 105, 135–40 (2009).
Tan, S. W. et al. Production and characterization of murine monoclonal antibodies to Blastocystis hominis. Int J Parasitol 26, 375–81 (1996).
Wu, B., Yin, J., Texier, C., Roussel, M. & Tan, K. S. Blastocystis legumain is localized on the cell surface, and specific inhibition of its activity implicates a pro-survival role for the enzyme. J Biol Chem 285, 1790–8 (2010).
Wawrzyniak, I. et al. Characterization of two cysteine proteases secreted by Blastocystis ST7, a human intestinal parasite. Parasitol Int 61, 437–42 (2012).
Puthia, M. K., Vaithilingam, A., Lu, J. & Tan, K. S. Degradation of human secretory immunoglobulin A by Blastocystis. Parasitol Res 97, 386–9 (2005).
Mirza, H., Wu, Z., Kidwai, F. & Tan, K. S. A metronidazole-resistant isolate of Blastocystis spp. is susceptible to nitric oxide and downregulates intestinal epithelial inducible nitric oxide synthase by a novel parasite survival mechanism. Infect Immun 79, 5019–26 (2011).
Puthia, M. K., Lu, J. & Tan, K. S. Blastocystis ratti contains cysteine proteases that mediate interleukin-8 response from human intestinal epithelial cells in an NF-kappaB-dependent manner. Eukaryot Cell 7, 435–43 (2008).
Lim, M. X. et al. Differential regulation of proinflammatory cytokine expression by mitogen-activated protein kinases in macrophages in response to intestinal parasite infection. Infect Immun 82, 4789–801 (2014).
Long, H. Y., Handschack, A., Konig, W. & Ambrosch, A. Blastocystis hominis modulates immune responses and cytokine release in colonic epithelial cells. Parasitol Res 87, 1029–30 (2001).
Teo, J. D., Macary, P. A. & Tan, K. S. Pleiotropic effects of Blastocystis spp. Subtypes 4 and 7 on ligand-specific toll-like receptor signaling and NF-kappaB activation in a human monocyte cell line. PLoS One 9, e89036 (2014).
Zuckerman, M. J., Watts, M. T., Ho, H. & Meriano, F. V. Blastocystis hominis infection and intestinal injury. Am J Med Sci 308, 96–101 (1994).
Zhang, H. W., Li, W., Yan, Q. Y., He, L. J. & Su, Y. P. [Impact of blastocystis hominis infection on ultrastructure of intestinal mucosa in mice]. Zhongguo Ji Sheng Chong Xue Yu Ji Sheng Chong Bing Za Zhi 24, 187–91 (2006).
Zierdt, C. H. Cytochrome-free mitochondria of an anaerobic protozoan–Blastocystis hominis. J Protozool 33, 67–9 (1986).
Stechmann, A. et al. Organelles in Blastocystis that blur the distinction between mitochondria and hydrogenosomes. Curr Biol 18, 580–5 (2008).
Tsaousis, A. D. et al. Evolution of Fe/S cluster biogenesis in the anaerobic parasite Blastocystis. Proc Natl Acad Sci USA 109, 10426–31 (2012).
Keenan, T. W. & Zierdt, C. H. Lipid biosynthesis by axenic strains of Blastocystis hominis. Comp Biochem Physiol Biochem Mol Biol 107, 525–31 (1994).
Zierdt, C. H. Blastocystis hominis, a long-misunderstood intestinal parasite. Parasitol Today 4, 15–7 (1988).
Zomerdijk, J. C. et al. The promoter for a variant surface glycoprotein gene expression site in Trypanosoma brucei. EMBO J 9, 2791–801 (1990).
MacGregor, P., Rojas, F., Dean, S. & Matthews, K. R. Stable transformation of pleomorphic bloodstream form Trypanosoma brucei. Mol Biochem Parasitol 190, 60–2 (2013).
Coburn, C. M., Otteman, K. M., McNeely, T., Turco, S. J. & Beverley, S. M. Stable DNA transfection of a wide range of trypanosomatids. Mol Biochem Parasitol 46, 169–79 (1991).
Rug, M. & Maier, A. G. Transfection of Plasmodium falciparum. Methods Mol Biol 923, 75–98 (2013).
Vinayak, S. et al. Genetic modification of the diarrhoeal pathogen Cryptosporidium parvum. Nature 523, 477–80 (2015).
Soldati, D. & Boothroyd, J. C. Transient transfection and expression in the obligate intracellular parasite Toxoplasma gondii. Science 260, 349–52 (1993).
Tsai, C. D., Liu, H. W. & Tai, J. H. Characterization of an iron-responsive promoter in the protozoan pathogen Trichomonas vaginalis. J Biol Chem 277, 5153–62 (2002).
Yu, D. C., Wang, A. L. & Wang, C. C. Stable coexpression of a drug-resistance gene and a heterologous gene in an ancient parasitic protozoan Giardia lamblia. Mol Biochem Parasitol 83, 81–91 (1996).
Yee, J. & Nash, T. E. Transient transfection and expression of firefly luciferase in Giardia lamblia. Proc Natl Acad Sci USA 92, 5615–9 (1995).
Baxt, L. A., Rastew, E., Bracha, R., Mirelman, D. & Singh, U. Downregulation of an Entamoeba histolytica rhomboid protease reveals roles in regulating parasite adhesion and phagocytosis. Eukaryot Cell 9, 1283–93 (2010).
van den Hoff, M. J., Moorman, A. F. & Lamers, W. H. Electroporation in ‘intracellular’ buffer increases cell survival. Nucleic Acids Res 20, 2902 (1992).
Alibaud, L., Cosson, P. & Benghezal, M. Dictyostelium discoideum transformation by oscillating electric field electroporation. Biotechniques 35(78–80), 82–3 (2003).
Tekle, E., Astumian, R. D. & Chock, P. B. Electroporation by using bipolar oscillating electric field: an improved method for DNA transfection of NIH 3T3 cells. Proc Natl Acad Sci USA 88, 4230–4 (1991).
Clark, J. D. et al. A toolbox facilitating stable transfection of Eimeria species. Mol Biochem Parasitol 162, 77–86 (2008).
Balu, B. & Adams, J. H. Advancements in transfection technologies for Plasmodium. Int J Parasitol 37, 1–10 (2007).
Kim, K. & Weiss, L. M. Toxoplasma gondii: the model apicomplexan. Int J Parasitol 34, 423–32 (2004).
Davis-Hayman, S. R. & Nash, T. E. Genetic manipulation of Giardia lamblia. Mol Biochem Parasitol 122, 1–7 (2002).
DiLella, A. G. et al. Utility of firefly luciferase as a reporter gene for promoter activity in transgenic mice. Nucleic Acids Res 16, 4159 (1988).
Hall, M. P. et al. Engineered luciferase reporter from a deep sea shrimp utilizing a novel imidazopyrazinone substrate. ACS Chem Biol 7, 1848–57 (2012).
Bangs, J. D., Brouch, E. M., Ransom, D. M. & Roggy, J. L. A soluble secretory reporter system in Trypanosoma brucei. Studies on endoplasmic reticulum targeting. J Biol Chem 271, 18387–93 (1996).
Klimes, V., Gentekaki, E., Roger, A. J. & Elias, M. A large number of nuclear genes in the human parasite blastocystis require mRNA polyadenylation to create functional termination codons. Genome Biol Evol 6, 1956–61 (2014).
Bastin, P., Bagherzadeh, Z., Matthews, K. R. & Gull, K. A novel epitope tag system to study protein targeting and organelle biogenesis in Trypanosoma brucei. Mol Biochem Parasitol 77, 235–9 (1996).
Cubitt, A. B. et al. Understanding, improving and using green fluorescent proteins. Trends Biochem Sci 20, 448–55 (1995).
Landete, J. M. et al. Anaerobic green fluorescent protein as a marker of Bifidobacterium strains. Int J Food Microbiol 175, 6–13 (2014).
Landete, J. M. et al. Use of anaerobic green fluorescent protein versus green fluorescent protein as reporter in lactic acid bacteria. Appl Microbiol Biotechnol 99, 6865–77 (2015).
Gentekaki, E. et al. Extreme genome diversity in the hyper-prevalent parasitic eukaryote Blastocystis. PLoS Biol 15, e2003769 (2017).
Turkeltaub, J. A., McCarty, T. R. III & Hotez, P. J. The intestinal protozoa: emerging impact on global health and development. Curr Opin Gastroenterol 31, 38–44 (2015).
Wong, K. H. et al. Predominance of subtype 3 among Blastocystis isolates from a major hospital in Singapore. Parasitol Res 102, 663–70 (2008).
Ho, L. C., Singh, M., Suresh, G., Ng, G. C. & Yap, E. H. Axenic culture of Blastocystis hominis in Iscove’s modified Dulbecco’s medium. Parasitol Res 79, 614–6 (1993).
Wirtz, E., Leal, S., Ochatt, C. & Cross, G. A. A tightly regulated inducible expression system for conditional gene knock-outs and dominant-negative genetics in Trypanosoma brucei. Mol Biochem Parasitol 99, 89–101 (1999).
This project was supported by a generous grant awarded to Kevin S.W. Tan from the Ministry of Education (MOE), Singapore (R-571-000-037-114) and to Cynthia Y. He from MOE, Singapore (R-154-000-A74-114). This research was also supported by BBSRC research grant (BB/M009971/1) to Anastasios D. Tsaousis. We would also like to thank Diego M. Cantoni from the University of Kent for his assistance in using the confocal microscope.
The authors declare no competing interests.
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Li, FJ., Tsaousis, A.D., Purton, T. et al. Successful Genetic Transfection of the Colonic Protistan Parasite Blastocystis for Reliable Expression of Ectopic Genes. Sci Rep 9, 3159 (2019). https://doi.org/10.1038/s41598-019-39094-5