The GPI-Phospholipase C of Trypanosoma brucei Is Nonessential But Influences Parasitemia in Mice

In the mammalian host, the cell surface of Trypanosoma brucei is protected by a variant surface glycoprotein that is anchored in the plasma membrane through covalent attachment of the COOH terminus to a glycosylphosphatidylinositol. The trypanosome also contains a phospholipase C (GPI-PLC) that cleaves this anchor and could thus potentially enable the trypanosome to shed the surface coat of VSG. Indeed, release of the surface VSG can be observed within a few minutes on lysis of trypanosomes in vitro. To investigate whether the ability to cleave the membrane anchor of the VSG is an essential function of the enzyme in vivo, a GPI-PLC null mutant trypanosome has been generated by targeted gene deletion. The mutant trypanosomes are fully viable; they can go through an entire life cycle and maintain a persistent infection in mice. Thus the GPI-PLC is not an essential activity and is not necessary for antigenic variation. However, mice infected with the mutant trypanosomes have a reduced parasitemia and survive longer than those infected with control trypanosomes. This phenotype is partially alleviated when the null mutant is modified to express low levels of GPI-PLC

A short bifunctional element operates to positively or negatively regulate <em>ESAG9</em> expression in different developmental forms of <em>Trypanosoma brucei</em>

In their mammalian host trypanosomes generate ‘stumpy’ forms from proliferative ‘slender’ forms as an adaptation for transmission to their tsetse fly vector. This transition is characterised by the repression of many genes while quiescent stumpy forms accumulate during each wave of parasitaemia. However, a subset of genes are upregulated either as an adaptation for transmission or to sustain infection chronicity. Among this group are ESAG9 proteins, whose genes were originally identified as a component of some telomeric variant surface glycoprotein gene expression sites, although many members of this diverse family are also transcribed elsewhere in the genome. ESAG9 genes are among the most highly regulated genes in transmissible stumpy forms, encoding a group of secreted proteins of cryptic function. To understand their developmental silencing in slender forms and activation in stumpy forms, the post-transcriptional control signals for a well conserved ESAG9 gene have been mapped. This identified a precise RNA sequence element of 34 nucleotides that contributes to gene expression silencing in slender forms but also acts positively, activating gene expression in stumpy forms. We predict that this bifunctional RNA sequence element is targeted by competing negative and positive regulatory factors in distinct developmental forms of the parasite. Analysis of the 3′UTR regulatory regions flanking the highly diverse ESAG9 family reveals that the linear regulatory sequence is not highly conserved, suggesting that RNA structure is important for interactions with regulatory proteins

Murine Models for Trypanosoma brucei gambiense Disease Progression—From Silent to Chronic Infections and Early Brain Tropism

Trypanosoma brucei gambiense is responsible for more than 90% of reported cases of human African trypanosomosis (HAT). Infection can last for months or even years without major signs or symptoms of infection, but if left untreated, sleeping sickness is always fatal. In the present study, different T. b. gambiense field isolates from the cerebrospinal fluid of patients with HAT were adapted to growth in vitro. These isolates belong to the homogeneous Group 1 of T. b. gambiense, which is known to induce a chronic infection in humans. In spite of this, these isolates induced infections ranging from chronic to silent in mice, with variations in parasitaemia, mouse lifespan, their ability to invade the CNS and to elicit specific immune responses. In addition, during infection, an unexpected early tropism for the brain as well as the spleen and lungs was observed using bioluminescence analysis. The murine models presented in this work provide new insights into our understanding of HAT and allow further studies of parasite tropism during infection, which will be very useful for the treatment and the diagnosis of the disease

T. brucei Infection Reduces B Lymphopoiesis in Bone Marrow and Truncates Compensatory Splenic Lymphopoiesis through Transitional B-Cell Apoptosis

African trypanosomes of the Trypanosoma brucei species are extracellular protozoan parasites that cause the deadly disease African trypanosomiasis in humans and contribute to the animal counterpart, Nagana. Trypanosome clearance from the bloodstream is mediated by antibodies specific for their Variant Surface Glycoprotein (VSG) coat antigens. However, T. brucei infection induces polyclonal B cell activation, B cell clonal exhaustion, sustained depletion of mature splenic Marginal Zone B (MZB) and Follicular B (FoB) cells, and destruction of the B-cell memory compartment. To determine how trypanosome infection compromises the humoral immune defense system we used a C57BL/6 T. brucei AnTat 1.1 mouse model and multicolor flow cytometry to document B cell development and maturation during infection. Our results show a more than 95% reduction in B cell precursor numbers from the CLP, pre-pro-B, pro-B, pre-B and immature B cell stages in the bone marrow. In the spleen, T. brucei induces extramedullary B lymphopoiesis as evidenced by significant increases in HSC-LMPP, CLP, pre-pro-B, pro-B and pre-B cell populations. However, final B cell maturation is abrogated by infection-induced apoptosis of transitional B cells of both the T1 and T2 populations which is not uniquely dependent on TNF-, Fas-, or prostaglandin-dependent death pathways. Results obtained from ex vivo co-cultures of living bloodstream form trypanosomes and splenocytes demonstrate that trypanosome surface coat-dependent contact with T1/2 B cells triggers their deletion. We conclude that infection-induced and possibly parasite-contact dependent deletion of transitional B cells prevents replenishment of mature B cell compartments during infection thus contributing to a loss of the host's capacity to sustain antibody responses against recurring parasitemic waves

High-throughput transformation of <i>Saccharomyces cerevisiae</i> using liquid handling robots

<div><p><i>Saccharomyces cerevisiae</i> (budding yeast) is a powerful eukaryotic model organism ideally suited to high-throughput genetic analyses, which time and again has yielded insights that further our understanding of cell biology processes conserved in humans. Lithium Acetate (LiAc) transformation of yeast with DNA for the purposes of exogenous protein expression (e.g., plasmids) or genome mutation (e.g., gene mutation, deletion, epitope tagging) is a useful and long established method. However, a reliable and optimized high throughput transformation protocol that runs almost no risk of human error has not been described in the literature. Here, we describe such a method that is broadly transferable to most liquid handling high-throughput robotic platforms, which are now commonplace in academic and industry settings. Using our optimized method, we are able to comfortably transform approximately 1200 individual strains per day, allowing complete transformation of typical genomic yeast libraries within 6 days. In addition, use of our protocol for gene knockout purposes also provides a potentially quicker, easier and more cost-effective approach to generating collections of double mutants than the popular and elegant synthetic genetic array methodology. In summary, our methodology will be of significant use to anyone interested in high throughput molecular and/or genetic analysis of yeast.</p></div

Strains and plasmids used in this study.

<p>Strains and plasmids used in this study.</p

VPS38 deletion and check primers.

<p>VPS38 deletion and check primers.</p

Successful application of high-throughput transformation method to gene deletion.

<p>(A) Generation of <i>VPS38</i> knockout strain by insertion via homologous recombination of a <i>LEU2</i> selective marker at the <i>VPS38</i> locus. (B) Different volumes of 150ng/ul of the <i>LEU2</i> cassette were transformed into yeast cells. Transformants were plated on—Leucine media. (C) <i>VPS38</i> deletion was verified using check primers (<a href="http://www.plosone.org/article/info:doi/10.1371/journal.pone.0174128#pone.0174128.t002" target="_blank">Table 2</a>) from four single colonies (wild type, 1#: 30 μl spot, 2#: 60 μ spotl, 3#: 120 μl spot). Expected PCR product size of wild type: 1701 bp, <i>VPS38</i> knockout: 2810 bp. (D) Plate 1 from the non-essential yeast knockout library was cultured and transformed with 4500ng VPS38 knockout <i>LEU2</i> cassette using our methodology. Transformed cells were cultured in –Leucine SD media for 3 days and spotted on –Leucine media using prongs.</p

Plasmid and cell concentration affect transformation efficiency.

<p>(A) BY4741 cells at OD₆₀₀ 0.5 were transformed with the indicated amount of pRB1 plasmid using our automated methodology with 4-hour heat shock. After transformation, cells were spotted on a –Uracil agar plate. (B) Different OD₆₀₀ cells were transformed with 100ng pRB1 plasmid and heat shocked at 42°C for 4 hours. (C) Microscopy of BY4741 cells originally transformed either at 0.4 or 1.5 OD₆₀₀, and examined at an OD of 3.5. Numbers indicate average stress granule (Pab1-GFP, green text) or P-body (Edc3-mCh, red text) foci per cell, and the percentage of Edc3-mCh foci co-localized with Pab1-GFP (white text) (D). Quantification of stress granule (SG) and P-Body (PB) size in (C). Data is presented as mean ± standard deviation of 3 independent experiments; n.s., not significant.</p

Schematic model of yeast transformation using liquid handling robot.

<p>Step 1, Inoculate overnight culture to the deep well plate. Grow another 2 ½ -4 hours to recover cells to the mid-log phase. Step 2, Prepare plasmid and transformation mix to transformation. Normally, an OD₆₀₀ range of 0.4–1.5, ≥100ng plasmid or 4500 ng PCR product is optimal for our automated transformation method. Step 3, Heat shock of the transformants for 3–6 hours. Step 4, Transfer the transformed cells to a liquid selective media plate to grow another 2–4 days. Pin the transformed strains onto appropriate selective media to generate the new library.</p