Microbiology

The study of host/pathogen interactions at the cellular level during Plasmodium intra-erythrocytic cycle requires differential extraction techniques aiming to analyze the different compartments of the infected cell. Various protocols have been proposed in the literature to study specific compartments and/or membranes in the infected erythrocyte. The task remains delicate despite the use of enzymes or detergents theoretically capable of degrading specific membranes inside the infected cell.

The remit of this protocol is to propose a method to isolate the erythrocyte cytosol and ghosts from the other compartments of the infected cell via a percoll gradient. Also, the lysis of the erythrocyte membrane is done using equinatoxin II, which has proven to be more effective at erythrocyte lysis regardless of the cell infection status, compared to the commonly used streptolysin. The parasitophorous vacuole (PV) content is collected after saponin lysis, before recovering membrane and parasite cytosol proteins by Triton X-100 lysis. The lysates thus obtained are analyzed by Western blot to assess the accuracy of the various extraction steps. This protocol allows the separation of the host compartment from the parasite compartments (PV and parasite), leading to potential studies of host proteins as well as parasite proteins exported to the host cell.

Cyanobacteria represent a frequently used model organism for the study of oxygenic photosynthesis. They belong to prokaryotic microorganisms but their photosynthetic apparatus is quite similar to that found in algal and plant chloroplasts. The key players in light reactions of photosynthesis are Photosystem I and Photosystem II complexes (PSI and PSII, resp.), large membrane complexes of proteins, pigments and other cofactors embedded in specialized photosynthetic membranes named thylakoids. For the study of these complexes a mild method for the isolation of the thylakoids, their subsequent solubilization and analysis is essential. The presented protocol describes such a method which utilizes breaking the cyanobacterial cells using glass beads in an optimized buffer. This is followed by their solubilization using dodecyl-maltoside and analysis using optimized clear-native gel electrophoresis which preserves the native oligomerization state of both complexes and allows the estimation of their content.

There exists a wide variety of techniques to isolate and purify viral particles from cell culture supernatants. However, these techniques vary greatly in ease of use, purity, yield and impact on viral structural integrity. Most importantly, it is becoming evident that secreted extracellular vesicles (EVs) co-purify with retroviruses using nearly all purification methods due to nearly indistinguishable biophysical characteristics such as size, buoyant density and nucleic acid content. Recently, our group has illustrated a means of isolating intact and highly enriched retroviral virions from EV-containing cell supernatants using an immunoprecipitation approach targeting the viral envelope glycoprotein of the Moloney Murine Leukemia Virus (Renner et al., 2018). This technique, that we call intact virion immunoprecipitation (IVIP), enabled us to characterize the accessibility of epitopes on the surface of these retroviruses and assess the orientation of the virus-encoded integral membrane protein Glycogag (gPr80) in the viral envelope. Proper implementation of this protocol enables fast, simple and reproducible preparations of intact and highly purified retroviral particles devoid of detectable EV contaminants.

Cyanobacteria are photosynthetic bacteria that thrive in diverse ecosystems and play major roles in the global carbon cycle. The abilities of cyanobacteria to fix atmospheric CO₂ and to allocate the fixed carbons to chemicals and biofuels have attracted growing attentions as sustainable microbial cell factories. A better understanding of activities of enzymes involved in the central carbon metabolism might lead to increased product yields. Currently, cell-free lysates are widely used for the determination of intracellular enzyme activities. However, due to thick cell walls in cyanobacteria, lysis of cyanobacterial cells is inefficient and often laborious. The present protocol describes an easy and efficient method to permeabilize cyanobacterial cells, without lysing them, and direct usage of the permeabilized cells for the determination of metabolic enzyme activities in vivo.

CRISPR-Cas (Clustered regularly interspaced short palindromic repeats-CRISPR-associated proteins) is a class of prokaryotic immune systems that degrade foreign nucleic acids in a sequence-specific manner. These systems rely upon ribonucleoprotein complexes composed of Cas nucleases and small CRISPR RNAs (crRNAs). Staphylococcus epidermidis and Staphylococcus aureus are bacterial residents on human skin that are also leading causes of antibiotic resistant infections (Lowy, 1998; National Nosocomial Infections Surveillance, 2004; Otto, 2009). Many staphylococci possess Type III-A CRISPR-Cas systems (Marraffini and Sontheimer, 2008; Cao et al., 2016), which have been shown to prevent plasmid transfer and protect against viral predators (Goldberg et al., 2014; Hatoum-Aslan et al., 2014; Samai et al., 2015) in these organisms. Thus, gaining a mechanistic understanding of these systems in the native staphylococcal background can lead to important insights into the factors that impact the evolution and survival of these pathogens. Type III-A CRISPR-Cas systems encode a five-subunit effector complex called Cas10-Csm (Hatoum-Aslan et al., 2013). Here, we describe a protocol for the expression and purification of Cas10-Csm from its native S. epidermidis background or a heterologous S. aureus background. The method consists of a two-step purification protocol involving Ni²⁺-affinity chromatography and a DNA affinity biotin pull-down, which together yield a pure preparation of the Cas10-Csm complex. This approach has been used previously to analyze the effects of mutations on Cas10-Csm complex integrity (Hatoum-Aslan et al., 2014), crRNA formation (Hatoum-Aslan et al., 2013), and to detect binding partners that directly interact with the core Cas10-Csm complex (Walker et al., 2016). Importantly, this approach can be easily adapted for use in other Staphylococcus species to probe and understand their native Type III-A CRISPR-Cas systems.

Isolation of ribosomal particles is an essential step in the study of ribosomal components as well as in the analysis of trans-acting factors that interact with the ribosome to regulate protein synthesis and modulate the expression profile of the cell in response to different environmental conditions. In this protocol, we describe a procedure for the isolation of 70S ribosomes from the unicellular cyanobacterium Synechocystis sp. PCC 6803 (hereafter Synechocystis). We have successfully used this protocol in our study of the cyanobacterial ribosomal-associated protein LrtA, which is a homologue of bacterial HPF (hibernation promoting factor) (Galmozzi et al., 2016).

Circadian rhythms are biological processes displaying an endogenous oscillation with a period of ~24 h. They allow organisms to anticipate and get prepared for the environmental changes caused mainly by the rotation of Earth. Circadian rhythms are driven by circadian clocks that consist of proteins, DNA, and/or RNA. Circadian clocks of cyanobacteria are the simplest and one of the best studied models. They contain the three clock proteins KaiA, KaiB, and KaiC which can be used for in vitro reconstitution experiments and determination of the auto-phosphatase activity of KaiC as described in this protocol.

The SLC26 or SulP proteins constitute a large family of anion transporters that are ubiquitously expressed in pro- and eukaryotes. In human, SLC26 proteins perform important roles in ion homeostasis and malfunctioning of selected members is associated with diseases. This protocol details the production and crystallization of a prokaryotic SLC26 homolog, termed SLC26Dg, from Deinococcus geothermalis. Following these instructions we obtained well-folded and homogenous material of the membrane protein SLC26Dg and the nanobody Nb5776 that enabled us to crystallize the complex and determine its structure (Geertsma et al., 2015). The procedure may be adapted to purify and crystallize other membrane protein complexes.

The direct regulation of a mycobacterial adenylate cyclase (Rv1625c) via exchange of its membrane anchor by the quorum sensing receptor CqsS (Vibrio harveyi) has recently been reported (Beltz et al., 2016). This protocol describes the expression and membrane preparation for these chimeric proteins.

The magnesium transporter A (MgtA) is a magnesium transporting P-type ATPase present in prokaryotes and plants (Subramani et al., 2016). In Salmonella typhimurium and Escherichia coli (E. coli), MgtA is expressed only in magnesium limiting conditions and plays an important role in Mg²⁺ homeostasis (Groisman et al., 2013). The transcription of mgtA is regulated by the two-component system PhoP/PhoQ (Soncini et al., 1996; Kato et al., 1999). The membrane bound histidine kinase, PhoQ, senses low Mg²⁺ concentration in the periplasmic space and phosphorylates its cognate response regulator, PhoP, which initiates mgtA transcription (Groisman et al., 2013). MgtA is targeted to the plasma membrane and facilitate the bacterial survival under low Mg²⁺ condition, by importing Mg²⁺ into the cytoplasm. The MgtA homolog in petunia (PH1) is found in the vacuolar membrane and involved with the coloration of the flower petals (Faraco et al., 2014). As a first step towards understanding the molecular details of MgtA Mg²⁺ transport, we describe a detailed protocol for the purification of E. coli MgtA that can be used for biochemical and biophysical studies. Recombinant E. coli MgtA with hexa histidine tag at the N-terminus was cloned from E. coli DH5α and over expressed in the E. coli C43(DE3) by fermentation to an OD > 6. Cell lysis was performed in a high pressure homogenizer and the membranes were isolated by ultracentrifugation. Membrane proteins were solubilized with the detergent dodecyl-β-D maltoside. MgtA was purified by affinity and size exclusion chromatography. Final yields of purified MgtA reach ~1 mg MgtA per 3 g of wet cell pellet.