Research in our lab was supported by the Max von Pettenkofer Institute, Ludwig-Maximilians University Munich, the German Research Foundation (DFG, HI 1511/3-1) and the Bundesministerium f&uuml;r Bildung und Forschung (BMBF) "Medical Infection Genomics" initiative (0315834C).

1. Preparations for LCV Isolation
<ol>
	<li>Streak out Legionella pneumophila producing DsRed-Express13,14 from glycerol stocks on CYE agar plates with 5 &mu;g/mL chloramphenicol (Cam) four days before LCV isolation. Incubate the bacteria at 37 &deg;C.</li>
	<li>Seed out 1 x 107 D. discoideum producing GFP-calnexin or RAW264.7 murine macrophages in 75 cm2 tissue culture flasks one day prior infection. Use 10 ml HL5 medium with 20 &mu;g/mL G418 for D. discoideum and incubate the amoebae at 23 &deg;C. For RAW264.7 macrophages use 10 ml RPMI 1640 medium supplemented with 10% FCS (heat inactivated) and 1% glutamine, and grow the cells at 37 &deg;C and 5% CO2. Use at least three 75 cm2 flasks per infection and sample (minimum 6 x 107 cells).</li>
	<li>Inoculate an overnight culture with L. pneumophila from the CYE plate. Take a 15 mL test tube with 3 mL AYE medium and 5 &mu;g/mL Cam. Inoculate with 100 &mu;l of a bacterial suspension to yield an OD600nm of 0.1. Incubate the overnight culture on an overhead rotation wheel at 37 &deg;C for 21-22 hr.</li>
</ol>
2. LCV Isolation
<ol>
	<li>Change the medium of D. discoideum cells to remove the antibiotic, which would interfere with the following infection.</li>
	<li>Measure the OD of the overnight culture. Bacteria should have reached their peak infectivity at an OD600nm of &ge;3, which corresponds to 2 x 109 bacteria/mL.</li>
	<li>Infect the cells by adding approximately 500 &mu;l of the L. pneumophila overnight culture to the cells growing in 10 ml HL5 medium (D. discoideum) or 10 ml RPMI 1640 (macrophages). This corresponds to a multiplicity of infection (MOI) of 50. Subsequently, the infection is synchronized by centrifugation of the bacteria onto the cells at 500 &times; g for 10 min. Centrifugation is followed by an incubation of the D. discoideum cells at 25 &deg;C and the RAW264.7 macrophages at 37 &deg;C and 5% CO2, respectively. The infection time of 1 hr includes the centrifugation step.</li>
	<li>After infection remove the medium and wash the cells once to remove extracellular bacteria. Use ice-cold SorC buffer for D. discoideum and ice-cold PBS for RAW264.7 macrophages. Add 3 mL HS buffer, supplemented with protease inhibitors (Roche) to each flask and collect the cells by using a cell scraper. Pool the corresponding samples in a 15 mL test tube.</li>
	<li>For homogenization of the sample use 3 mL plastic Luer-Lock syringes and the stainless steel ball homogenizer. Make sure that you work on ice. Before starting, wash the ball homogenizer with distilled water, to avoid any detergent contamination and flush it with ice cold HS buffer to get rid of air bubbles. Use the 8 &mu;m clearance ball.</li>
	<li>Fill the first 3 mL into the syringe und mount it on the homogenizer. Press the sample back and forth nine times through the homogenizer. Exchange the syringes afterwards to avoid contamination with not-homogenized material. Collect and pool the homogenized sample in a 15 mL test tube and take a 150 &mu;l sample for microscopic analysis. Before proceeding with a different sample dismantle and wash the ball homogenizer.</li>
	<li>Block the homogenate with 2% CS or FCS for 30 min on a shaker on ice or on an overhead-spinning wheel (10-20 rpm) at 4 &deg;C.</li>
	<li>After blocking use an affinity-purified primary antibody against SidC (dilution 1:3000) or against any other bacterial marker exclusively binding to the LCV membrane. Vortex the antibody solution before adding it to the homogenate. Incubate the sample for 1 hr on a shaker on ice or on an overhead-spinning wheel (10-20 rpm) at 4 &deg;C.</li>
	<li>In the meantime, prepare the Histodenz gradient. Use a 15 mL test tube per gradient and three 75 cm2 flasks of one sample. First, add 5.75 mL of 35% Histodenz to the tube, and second, carefully add 5.75 mL of 10% Histodenz solution on top. Afterwards carefully lay the tubes down horizontally for 1 hr.</li>
	<li>After incubation of the homogenate with the blocking reagent and the primary antibody, centrifuge at 600 &times; g for 15 min at 4&deg;C to pellet the sample. Discard the supernatant and resuspend the pellet in 1.5 mL HS buffer. Transfer the samples to a fresh 15 mL test tube and take a 40 &mu;l sample for microscopic analysis later on.</li>
	<li>Incubate the pellet with the secondary antibody - MACS goat anti-rabbit IgG micro beads (dilution 1:25) - for 30 min on a shaker at 4&deg;C. In the meantime, put the columns on the MACS magnetic holder and equilibrate them with 0.5 mL HS buffer.</li>
	<li>Subsequently, apply the samples to the column. Collect 150 &mu;l of the flow-through for subsequent microscopic analysis. Wash the column three times with 0.5 mL HS buffer.</li>
	<li>Remove the columns from the magnet and eluate the LCVs with 0.5 mL HS buffer by immediate squirting. Take a 20 &mu;l sample for microscopic analysis. The purification step by immuno-magnetic separation is expected to yield per 1 &times; 107 infected D. discoideum cells approximately 1 &times; 106 intact LCVs, which are about sevenfold enriched in the eluate compared with the flow-through 13.</li>
	<li>Put the Histodenz gradient tubes back to a vertical position and carefully load the eluate on top. Centrifuge the tubes at 3350 &times; g for 1 hr at 4 &deg;C.</li>
	<li>After centrifugation take 1.5 mL fractions starting from the bottom of the test tube by using a long glass Pasteur pipette. Altogether collect eight fractions, each from the bottom of the tube. No visible layers are observed in the continuous Histodenz gradient. The highest yield of LCVs is typically expected in fraction 4 at a ratio of ~2 &times; 105 intact LCVs per 1 &times; 107 infected amoebae 13. Minor quantities of LCVs are also present in fractions 3 and 5, respectively. Thus, the overall yield of purified LCVs is ~1 &times; 106 intact LCVs per 6 &times; 107 infected D. discoideum cells 13. The recovery of LCVs can be performed at room temperature (RT), and the LCVs appear stable for several hours.</li>
</ol>
3. Microscopic Analysis of the Collected Samples
<ol>
	<li>Prepare a 24-well flat-bottom tissue culture plate with poly-L-lysine coated coverslips.</li>
	<li>Pipette every sample taken during the LCV purification plus 150 &mu;l of each density gradient fraction into the 24-well plate. Add 0.5 mL HS buffer to each well to dilute the high Histodenz concentration. Afterwards, centrifuge the plate at 600 &times; g for 10 min at 4 &deg;C. Carefully remove the supernatant and fix the samples with 4% paraformaldehyde (PFA) for 20 min at RT. After fixation wash the samples twice. Use SorC for D. discoideum and PBS for RAW264.7 macrophages.</li>
	<li>
	<ol class="lalpha">
		<li>Mount the samples from GFP-calnexin expressing D. discoideum, taken at different steps during LCV isolation, directly on glass-slides using e.g. Vectashield.</li>
		<li>Incubate the samples from RAW264.7 macrophages, taken at different steps during LCV isolation, with blocking solution (1% BSA in PBS) for 10 min at RT. Use an anti-SidC primary antibody to stain the LCVs and incubate for 1h in a wet chamber at RT (affinity-purified rabbit anti-SidC; 1:1000 in blocking buffer). Subsequently, wash the samples three times with PBS. Incubate with a secondary antibody (e.g. anti-rabbit Cy5; 1:200 in blocking solution) for 30 min in a wet and dark chamber at RT. Finally, wash the samples three times with PBS and mount them on glass-slides using e.g. Vectashield.</li>
	</ol>
	</li>
	<li>Morphology, integrity and quantity of the LCVs isolated from GFP-calnexin expressing D. discoideum or from RAW264.7 macrophages are analyzed by a fluorescence microscope equipped with the adequate filters.</li>
</ol>
4. Representative Results
The quality and yield of the LCV purification by immuno-affinity separation and density gradient centrifugation can be followed by fluorescence microscopy (Fig. 1). An overview of samples collected during the LCV isolation from D. discoideum or macrophages is shown (Fig. 2). The homogenate of L. pneumophila-infected phagocytes shows intact LCVs, but also a lot of cell debris and extracellular bacteria. After pelleting the sample, the image is similar to the homogenate, sometimes a bit denser. Since intact LCVs should stick to the column, the flow-through contains mostly extracellular bacteria and cell debris. After eluting the sample from the column, the eluate contains intact LCVs in large amounts. In our hands, the LCV purification appears to be more effective for D. discoideum than for macrophages. In D. discoideum the pathogen vacuoles appear rounder and the yield of isolated LCVs is more than 10 times higher compared with macrophages (Fig. 3). In intact macrophages LVCs stained with different antibodies also appear more irregularly shaped.
<img alt="Figure 1" src="/files/ftp_upload/4118/4118fig1.jpg" /> 
Figure 1. Schematic overview of LCV isolation. A) Isolation of LCVs by immuno-magnetic separation using an affinity-purified antibody against the bacterial effector protein SidC, localizing exclusively to the LCV membrane. A secondary MACS micro bead-coupled antibody and a magnet are used to isolate LCVs from cell debris. B) After immuno-magnetic separation LCVs are further purified by Histodenz density gradient centrifugation. LCVs are enriched in fraction 4.
<img alt="Figure 2" src="/files/ftp_upload/4118/4118fig2.jpg" /> 
Figure 2. Samples collected during LCV purification. Images from homogenate, pellet, flow-through and eluate from D. discoideum or macrophages are depicted. The samples show L. pneumophila producing DsRed-Express (red) in D. discoideum producing GFP-calnexin signal (green, upper panel) or in macrophages stained with an anti-SidC antibody (cyan, lower panel). Bars, 10 &mu;m.
<img alt="Figure 3" src="/files/ftp_upload/4118/4118fig3.jpg" /> 
Figure 3. Purified LCVs from D. discoideum or macrophages. The samples show fraction 4 after Histodenz density gradient centrifugation of L. pneumophila producing DsRed-Express (red) in (A) D. discoideum producing GFP-calnexin signal (green) or (B) in macrophages stained with an anti-SidC antibody (cyan). Bars, 10 &mu;m.

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No conflicts of interest declared.

In contrast to previously published methods, this protocol is based on two steps, first the separation of LCVs by an immuno-magnetic approach, and second, the purification of LCVs by density gradient centrifugation. The LCV isolation can be easily followed by fluorescence microscopy, when either fluorescently labeled bacteria and GFP-producing cells or antibody staining is used. The protocol described here is a simple, straight-forward method, which results in the enrichment of LCVs with high purity. Importantly, the pro...

purification of pathogen vacuoles from legionella-infected phagocytes

The opportunistic pathogen Legionella pneumophila is an amoeba-resistant bacterium, which also replicates in alveolar macrophages thus causing the severe pneumonia "Legionnaires' disease"1. In protozoan and mammalian phagocytes, L. pneumophila employs a conserved mechanism to form a specific, replication-permissive compartment, the "Legionella-containing vacuole" (LCV). LCV formation requires the bacterial Icm/Dot type IV secretion system (T4SS), which translocates as many as 275 "effector" proteins into host cells. The effectors manipulate host proteins as well as lipids and communicate with secretory, endosomal and mitochondrial organelles2-4.
The formation of LCVs represents a complex, robust and redundant process, which is difficult to grasp in a reductionist manner. An integrative approach is required to comprehensively understand LCV formation, including a global analysis of pathogen-host factor interactions and their temporal and spatial dynamics. As a first step towards this goal, intact LCVs are purified and analyzed by proteomics and lipidomics.
The composition and formation of pathogen-containing vacuoles has been investigated by proteomic analysis using liquid chromatography or 2-D gel electrophoresis coupled to mass-spectrometry. Vacuoles isolated from either the social soil amoeba Dictyostelium discoideum or mammalian phagocytes harboured Leishmania5, Listeria6, Mycobacterium7, Rhodococcus8, Salmonella9 or Legionella spp.10. However, the purification protocols employed in these studies are time-consuming and tedious, as they require e.g. electron microscopy to analyse LCV morphology, integrity and purity. Additionally, these protocols do not exploit specific features of the pathogen vacuole for enrichment.
The method presented here overcomes these limitations by employing D. discoideum producing a fluorescent LCV marker and by targeting the bacterial effector protein SidC, which selectively anchors to the LCV membrane by binding to phosphatidylinositol 4-phosphate (PtdIns(4)P)3,11 . LCVs are enriched in a first step by immuno-magnetic separation using an affinity-purified primary antibody against SidC and a secondary antibody coupled to magnetic beads, followed in a second step by a classical Histodenz density gradient centrifugation12,13 (Fig. 1).
A proteome study of isolated LCVs from D. discoideum revealed more than 560 host cell proteins, including proteins associated with phagocytic vesicles, mitochondria, ER and Golgi, as well as several GTPases, which have not been implicated in LCV formation before13. LCVs enriched and purified with the protocol outlined here can be further analyzed by microscopy (immunofluorescence, electron microscopy), biochemical methods (Western blot) and proteomic or lipidomic approaches.

Watch this Scientific Journal Video about Purification of Pathogen Vacuoles from Legionella-infected Phagocytes at JoVE.com

Purification of Pathogen Vacuoles from Legionella-infected Phagocytes

This article describes a method for the isolation and purification of intact Legionella-containing vacuoles (LCVs) from amoeba and macrophages. The two-step protocol comprises LCV enrichment by immuno-magnetic separation using an antibody against a bacterial LCV marker and further purification by density gradient centrifugation.

Purification of Pathogen Vacuoles from Legionella-infected Phagocytes

The opportunistic pathogen Legionella pneumophila is an amoeba-resistant bacterium, which also replicates in alveolar macrophages thus causing the severe ...

purification-of-pathogen-vacuoles-from-legionella-infected-phagocytes

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Immunology and Infection

12.4K vistas.  Ludwig-Maximilians-Universität. Este artículo describe un método para el aislamiento y purificación de intacta Legionella Que contienen vacuolas (LCV) de amebas y los macrófagos. El protocolo de dos pasos comprende enriquecimiento LCV por inmuno-separación magnética usando un anticuerpo contra un marcador bacteriano LCV y purificación adicional por centrifugación en gradiente de densidad.

Video: Purification of Pathogen Vacuoles from Legionella-infected Phagocytes

Invasion of Human Cells by a Bacterial Pathogen

Here we will describe how we study the invasion of human endothelial cells by bacterial pathogen Staphylococcus aureus . The general protocol can be applied to the study of cell invasion by virtually any culturable bacterium. The stages at which specific aspects of invasion can be studied, such as the role of actin rearrangement or caveolae, will be highlighted. Host cells are grown in flasks and when ready for use are seeded into 24-well plates containing Thermanox coverslips. Using coverslips allows subsequent removal of the cells from the wells to reduce interference from serum proteins deposited onto the sides of the wells (to which S. aureus would attach). Bacteria are grown to the required density and washed to remove any secreted proteins (e.g. toxins). Coverslips with confluent layers of endothelial cells are transferred to new 24-well plates containing fresh culture medium before the addition of bacteria. Bacteria and cells are then incubated together for the required amount of time in 5% CO2 at 37&deg;C. For S. aureus this is typically between 15-90 minutes. Thermanox coverslips are removed from each well and dip-washed in PBS to remove unattached bacteria. If total associated bacteria (adherent and internalised) are to be quantified, coverslips are then placed in a fresh well containing 0.5% Triton X-100 in PBS. Gentle pipetting leads to complete cell lysis and bacteria are enumerated by serial dilution and plating onto agar. If the number of bacteria that have invaded the cells is needed, coverslips are added to wells containing 500 &mu;l tissue culture medium supplemented with gentamicin and incubation continued for 1 h, which will kill all external bacteria. Coverslips can then be washed, cells lysed and bacteria enumerated by plating onto agar as described above. If the experiment requires direct visualisation, coverslips can be fixed and stained for light, fluorescence or confocal microscopy or prepared for electron microscopy.

A general protocol for the study of invasion of host cells by a bacterial pathogen, focusing on Staphylococcus aureus and human endothelial cells.

Here we will describe how we study the invasion of human endothelial cells by bacterial pathogen Staphylococcus aureus . The general protocol can be ...

One-day Workflow Scheme for Bacterial Pathogen Detection and Antimicrobial Resistance Testing from Blood Cultures

Bloodstream infections are associated with high mortality rates because of the probable manifestation of sepsis, severe sepsis and septic shock1. Therefore, rapid administration of adequate antibiotic therapy is of foremost importance in the treatment of bloodstream infections. The critical element in this process is timing, heavily dependent on the results of bacterial identification and antibiotic susceptibility testing. Both of these parameters are routinely obtained by culture-based testing, which is time-consuming and takes on average 24-48 hours2, 4. The aim of the study was to develop DNA-based assays for rapid identification of bloodstream infections, as well as rapid antimicrobial susceptibility testing. The first assay is a eubacterial 16S rDNA-based real-time PCR assay complemented with species- or genus-specific probes5. Using these probes, Gram-negative bacteria including Pseudomonas spp., Pseudomonas aeruginosa and Escherichia coli as well as Gram-positive bacteria including Staphylococcus spp., Staphylococcus aureus, Enterococcus spp., Streptococcus spp., and Streptococcus pneumoniae could be distinguished. Using this multiprobe assay, a first identification of the causative micro-organism was given after 2 h.
Secondly, we developed a semi-molecular assay for antibiotic susceptibility testing of S. aureus, Enterococcus spp. and (facultative) aerobe Gram-negative rods6. This assay was based on a study in which PCR was used to measure the growth of bacteria7. Bacteria harvested directly from blood cultures are incubated for 6 h with a selection of antibiotics, and following a Sybr Green-based real-time PCR assay determines inhibition of growth. The combination of these two methods could direct the choice of a suitable antibiotic therapy on the same day (Figure 1). In conclusion, molecular analysis of both identification and antibiotic susceptibility offers a faster alternative for pathogen detection and could improve the diagnosis of bloodstream infections.

The design of a straightforward one-day workflow scheme for bacterial pathogen diagnostics enables the rapid recognition of bloodstream infections. The inclusion of eight clinically relevant bacterial targets and their antibiotic resistance profiles offers the clinician an initial insight on the same day, which can lead to more adequate therapy.

Bloodstream infections are associated with high mortality rates because of the probable manifestation of sepsis, severe sepsis and septic shock1. ...

Purification and Visualization of Lipopolysaccharide from Gram-negative Bacteria by Hot Aqueous-phenol Extraction

Lipopolysaccharide (LPS) is a major component of Gram-negative bacterial outer membranes. It is a tripartite molecule consisting of lipid A, which is embedded in the outer membrane, a core oligosaccharide and repeating O-antigen units that extend outward from the surface of the cell1, 2. LPS is an immunodominant molecule that is important for the virulence and pathogenesis of many bacterial species, including Pseudomonas aeruginosa, Salmonella species, and Escherichia coli3-5, and differences in LPS O-antigen composition form the basis for serotyping of strains. LPS is involved in attachment to host cells at the initiation of infection and provides protection from complement-mediated killing; strains that lack LPS can be attenuated for virulence6-8. For these reasons, it is important to visualize LPS, particularly from clinical isolates. Visualizing LPS banding patterns and recognition by specific antibodies can be useful tools to identify strain lineages and to characterize various mutants. 
In this report, we describe a hot aqueous-phenol method for the isolation and purification of LPS from Gram-negative bacterial cells. This protocol allows for the extraction of LPS away from nucleic acids and proteins that can interfere with visualization of LPS that occurs with shorter, less intensive extraction methods9. LPS prepared this way can be separated by sodium dodecyl sulfate (SDS)-polyacrylamide gel electrophoresis (PAGE) and directly stained using carbohydrate/glycoprotein stains or standard silver staining methods. Many anti-sera to LPS contain antibodies that cross-react with outer membrane proteins or other antigenic targets that can hinder reactivity observed following Western immunoblot of SDS-PAGE-separated crude cell lysates. Protease treatment of crude cell lysates alone is not always an effective way of removing this background using this or other visualization methods. Further, extensive protease treatment in an attempt to remove this background can lead to poor quality LPS that is not well resolved by any of the aforementioned methods. For these reasons, we believe that the following protocol, adapted from Westpahl and Jann10, is ideal for LPS extraction.

We describe a modified hot aqueous-phenol extraction method for purifying lipopolysaccharide (LPS) from Gram-negative bacteria. Once extracted, the LPS can be subsequently analyzed by SDS-PAGE and visualized by direct staining or Western immunoblot.

Lipopolysaccharide (LPS)  is a major component of Gram-negative bacterial outer membranes. It is a tripartite molecule consisting of  lipid A, which is ...

Affinity Purification of Influenza Virus Ribonucleoprotein Complexes from the Chromatin of Infected Cells

Like all negative-strand RNA viruses, the genome of influenza viruses is packaged in the form of viral ribonucleoprotein complexes (vRNP), in which the single-stranded genome is encapsidated by the nucleoprotein (NP), and associated with the trimeric polymerase complex consisting of the PA, PB1, and PB2 subunits. However, in contrast to most RNA viruses, influenza viruses perform viral RNA synthesis in the nuclei of infected cells. Interestingly, viral mRNA synthesis uses cellular pre-mRNAs as primers, and it has been proposed that this process takes place on chromatin1. Interactions between the viral polymerase and the host RNA polymerase II, as well as between NP and host nucleosomes have also been characterized1,2.
Recently, the generation of recombinant influenza viruses encoding a One-Strep-Tag genetically fused to the C-terminus of the PB2 subunit of the viral polymerase (rWSN-PB2-Strep3) has been described. These recombinant viruses allow the purification of PB2-containing complexes, including vRNPs, from infected cells. To obtain purified vRNPs, cell cultures are infected, and vRNPs are affinity purified from lysates derived from these cells. However, the lysis procedures used to date have been based on one-step detergent lysis, which, despite the presence of a general nuclease, often extract chromatin-bound material only inefficiently. 
Our preliminary work suggested that a large portion of nuclear vRNPs were not extracted during traditional cell lysis, and therefore could not be affinity purified. To increase this extraction efficiency, and to separate chromatin-bound from non-chromatin-bound nuclear vRNPs, we adapted a step-wise subcellular extraction protocol to influenza virus-infected cells. Briefly, this procedure first separates the nuclei from the cell and then extracts soluble nuclear proteins (here termed the "nucleoplasmic" fraction). The remaining insoluble nuclear material is then digested with Benzonase, an unspecific DNA/RNA nuclease, followed by two salt extraction steps: first using 150 mM NaCl (termed "ch150"), then 500 mM NaCl ("ch500") (Fig. 1). These salt extraction steps were chosen based on our observation that 500 mM NaCl was sufficient to solubilize over 85% of nuclear vRNPs yet still allow binding of tagged vRNPs to the affinity matrix.
After subcellular fractionation of infected cells, it is possible to affinity purify PB2-tagged vRNPs from each individual fraction and analyze their protein and RNA components using Western Blot and primer extension, respectively. Recently, we utilized this method to discover that vRNP export complexes form during late points after infection on the chromatin fraction extracted with 500 mM NaCl (ch500)3.

Influenza viruses replicate their RNA genome in association with host-cell chromatin. Here, we present a method to purify intact viral ribonucleoprotein complexes from the chromatin of infected cells. Purified viral complexes can be analyzed by both Western blot and primer extension of protein and RNA content, respectively.

Like all negative-strand RNA viruses, the genome of influenza viruses is packaged in the form of viral ribonucleoprotein complexes (vRNP), in which the ...

Live-cell Video Microscopy of Fungal Pathogen Phagocytosis

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of phagocytes to the site where pathogens are located; (ii) recognition of pathogen-associated molecular patterns (PAMPs) through pattern recognition receptors (PRRs); (iii) engulfment of microorganisms bound to the phagocyte cell membrane, and (iv) processing of engulfed cells within maturing phagosomes and digestion of the ingested particle. Studies that assess phagocytosis in its entirety are informative1, 2, 3, 4, 5 but are limited in that they do not normally break the process down into migration, engulfment and phagosome maturation, which may be affected differentially. Furthermore, such studies assess uptake as a single event, rather than as a continuous dynamic process. We have recently developed advanced live-cell imaging technologies, and have combined these with genetic functional analysis of both pathogen and host cells to create a cross-disciplinary platform for the analysis of innate immune cell function and fungal pathogenesis. These studies have revealed novel aspects of phagocytosis that could only be observed using systematic temporal analysis of the molecular and cellular interactions between human phagocytes and fungal pathogens and infectious microorganisms more generally. For example, we have begun to define the following: (a) the components of the cell surface required for each stage of the process of recognition, engulfment and killing of fungal cells1, 6, 7, 8; (b) how surface geometry influences the efficiency of macrophage uptake and killing of yeast and hyphal cells7; and (c) how engulfment leads to alteration of the cell cycle and behavior of macrophages 9, 10.
In contrast to single time point snapshots, live-cell video microscopy enables a wide variety of host cells and pathogens to be studied as continuous sequences over lengthy time periods, providing spatial and temporal information on a broad range of dynamic processes, including cell migration, replication and vesicular trafficking. Here we describe in detail how to prepare host and fungal cells, and to conduct the video microscopy experiments. These methods can provide a user-guide for future studies with other phagocytes and microorganisms.

We describe methods for live-cell video microscopy of Candida albicans phagocytosis by macrophages. These methods enable stage-specific analysis of macrophage migration, recognition, engulfment and phagosome maturation and reveal novel aspects of phagocytosis.

Phagocytic clearance of fungal pathogens, and microorganisms more generally, may be considered to consist of four distinct stages: (i) migration of ...

Loop-mediated Isothermal Amplification (LAMP) Assays for the Species-specific Detection of Eimeria that Infect Chickens

Eimeria species parasites, protozoa which cause the enteric disease coccidiosis, pose a serious threat to the production and welfare of chickens. In the absence of effective control clinical coccidiosis can be devastating. Resistance to the chemoprophylactics frequently used to control Eimeria is common and sub-clinical infection is widespread, influencing feed conversion ratios and susceptibility to other pathogens such as Clostridium perfringens. Despite the availability of polymerase chain reaction (PCR)-based tools, diagnosis of Eimeria infection still relies almost entirely on traditional approaches such as lesion scoring and oocyst morphology, but neither is straightforward. Limitations of the existing molecular tools include the requirement for specialist equipment and difficulties accessing DNA as template. In response a simple field DNA preparation protocol and a panel of species-specific loop-mediated isothermal amplification (LAMP) assays have been developed for the seven Eimeria recognised to infect the chicken. We now provide a detailed protocol describing the preparation of genomic DNA from intestinal tissue collected post-mortem, followed by setup and readout of the LAMP assays. Eimeria species-specific LAMP can be used to monitor parasite occurrence, assessing the efficacy of a farm&#8217;s anticoccidial strategy, and to diagnose sub-clinical infection or clinical disease with particular value when expert surveillance is unavailable.

Loop-mediated Isothermal Amplification LAMP Assays for the Species-specific Detection of Eimeria that Infect Chickens

Diagnosis of Eimeria infection in chickens remains demanding. Parasite morphology- and host pathology-led approaches are commonly inconclusive, while molecular approaches based on PCR have proven demanding in cost and expertise. The aim of this protocol is to establish loop-mediated isothermal amplification (LAMP) as a straightforward molecular diagnostic for eimerian infection.

Eimeria species parasites, protozoa which cause the enteric disease coccidiosis, pose a serious threat to the production and welfare of chickens. In the ...

RNA Purification from Intracellularly Grown Listeria monocytogenes in Macrophage Cells

Analysis of the transcriptome of bacterial pathogens during mammalian infection is a valuable tool for studying genes and factors that mediate infection. However, isolating bacterial RNA from infected cells or tissues is a challenging task, since mammalian RNA mostly dominates the lysates of infected cells. Here we describe an optimized method for RNA isolation of Listeria monocytogenes bacteria growing within bone marrow derived macrophage cells. Upon infection, cells are mildly lysed and rapidly filtered to discard most of the host proteins and RNA, while retaining intact bacteria. Next, bacterial RNA is isolated using hot phenol-SDS extraction followed by DNase treatment. The extracted RNA is suitable for gene transcription analysis by multiple techniques. This method is successfully employed in our studies of Listeria monocytogenes gene regulation during infection of macrophage cells 1-4. The protocol can be easily modified to study other bacterial pathogens and cell types.

RNA Purification from Intracellularly Grown Listeria monocytogenes in Macrophage Cells

Here we describe a method for bacterial RNA isolation from Listeria monocytogenes bacteria growing inside murine macrophages. This technique can be used with other intracellular pathogens and mammalian host cells.

Analysis of the transcriptome of bacterial pathogens during mammalian infection is a valuable tool for studying genes and factors that mediate infection. ...

Development and Identification of a Novel Subpopulation of Human Neutrophil-derived Giant Phagocytes In Vitro

Neutrophils (PMN) are best known for their phagocytic functions against invading pathogens and microorganisms. They have the shortest half-life amongst leukocytes and in their non-activated state are constitutively committed to apoptosis. When recruited to inflammatory sites to resolve inflammation, they produce an array of cytotoxic molecules with potent antimicrobial killing. Yet, when these powerful cytotoxic molecules are released in an uncontrolled manner they can damage surrounding tissues. In recent years however, neutrophil versatility is increasingly evidenced, by demonstrating plasticity and immunoregulatory functions. We have recently identified a new neutrophil-derived subpopulation, which develops spontaneously in standard culture conditions without the addition of cytokines/growth factors such as granulocyte colony-stimulating factor (GM-CSF)/interleukin (IL)-4. Their phagocytic abilities of neutrophil remnants largely contribute to increase their size immensely; therefore they were termed giant phagocytes (G&#981;). Unlike neutrophils, G&#981; are long lived in culture. They express the cluster of differentiation (CD) neutrophil markers CD66b/CD63/CD15/CD11b/myeloperoxidase (MPO)/neutrophil elastase (NE), and are devoid of the monocytic lineage markers CD14/CD16/CD163 and the dendritic CD1c/CD141 markers. They also take-up latex and zymosan, and respond by oxidative burst to stimulation with opsonized-zymosan and PMA. G&#981; also express the scavenger receptors CD68/CD36, and unlike neutrophils, internalize oxidized-low density lipoprotein (oxLDL). Moreover, unlike fresh neutrophils, or cultured monocytes, they respond to oxLDL uptake by increased reactive oxygen species (ROS) production. Additionally, these phagocytes contain microtubule-associated protein-1 light chain 3B (LC3B) coated vacuoles, indicating the activation of autophagy. Using specific inhibitors it is evident that both phagocytosis and autophagy are prerequisites for their development and likely NADPH oxidase dependent ROS. We describe here a method for the preparation of this new subpopulation of long-lived, neutrophil-derived phagocytic cells in culture, their identification and their currently known characteristics. This protocol is essential for obtaining and characterizing G&#981; in order to further investigate their significance and functions.

Development and Identification of a Novel Subpopulation of Human Neutrophil-derived Giant Phagocytes In Vitro

We describe here a method for obtaining and identifying a newly characterized subpopulation of neutrophil-derived giant phagocytes. These cells develop in culture from fresh human blood neutrophils, and are characterized by phagocytosis, autophagy, immensely large size, and extended lifespan. This method is essential to further investigate this unique neutrophil-derived subpopulation.

Neutrophils (PMN) are best known for their phagocytic functions against invading pathogens and microorganisms. They have the shortest half-life amongst ...

Unravelling the Function of a Bacterial Effector from a Non-cultivable Plant Pathogen Using a Yeast Two-hybrid Screen

Unravelling the molecular mechanisms of disease manifestations is important to understand pathologies and symptom development in plant science. Bacteria have evolved different strategies to manipulate their host metabolism for their own benefit. This bacterial manipulation is often coupled with severe symptom development or the death of the affected plants. Determining the specific bacterial molecules responsible for the host manipulation has become an important field in microbiological research. After the identification of these bacterial molecules, called &#34;effectors,&#34; it is important to elucidate their function. A straightforward approach to determine the function of an effector is to identify its proteinaceous binding partner in its natural host via a yeast two-hybrid (Y2H) screen. Normally the host harbors numerous potential binding partners that cannot be predicted sufficiently by any in silico algorithm. It is thus the best choice to perform a screen with the hypothetical effector against a whole library of expressed host proteins. It is especially challenging if the causative agent is uncultivable like phytoplasma. This protocol provides step-by-step instructions for DNA purification from a phytoplasma-infected woody host plant, the amplification of the potential effector, and the subsequent identification of the plant&#39;s molecular interaction partner with a Y2H screen. Even though Y2H screens are commonly used, there is a trend to outsource this technique to biotech companies that offer the Y2H service at a cost. This protocol provides instructions on how to perform a Y2H in any decently equipped molecular biology laboratory using standard lab techniques.

Bacterial effector proteins are important for establishing successful infections. This protocol describes the experimental identification of proteinaceous binding partners of a bacterial effector protein in its natural plant host. Identifying these effector interactions via yeast two-hybrid screens has become an important tool in unravelling molecular pathogenicity strategies.

Unravelling the molecular mechanisms of disease manifestations is important to understand pathologies and symptom development in plant science. Bacteria ...

Isolation, Characterization, and Purification of Macrophages from Tissues Affected by Obesity-related Inflammation

Obesity promotes a chronic inflammatory state that is largely mediated by tissue-resident macrophages as well as monocyte-derived macrophages. Diet-induced obesity (DIO) is a valuable model in studying the role of macrophage heterogeneity; however, adequate macrophage isolations are difficult to acquire from inflamed tissues. In this protocol, we outline the isolation steps and necessary troubleshooting guidelines derived from our studies for obtaining a suitable population of tissue-resident macrophages from mice following 18 weeks of high-fat (HFD) or high-fat/high-cholesterol (HFHCD) diet intervention. This protocol focuses on three hallmark tissues studied in obesity and atherosclerosis including the liver, white adipose tissues (WAT), and the aorta. We highlight how dualistic usage of flow cytometry can achieve a new dimension of isolation and characterization of tissue-resident macrophages. A fundamental section of this protocol addresses the intricacies underlying tissue-specific enzymatic digestions and macrophage isolation, and subsequent cell-surface antibody staining for flow cytometric analysis. This protocol addresses existing complexities underlying fluorescent-activated cell sorting (FACS) and presents clarifications to these complexities so as to obtain broad range characterization from adequately sorted cell populations. Alternate enrichment methods are included for sorting cells, such as the dense liver, allowing for flexibility and time management when working with FACS. In brief, this protocol aids the researcher to evaluate macrophage heterogeneity from a multitude of inflamed tissues in a given study and provides insightful troubleshooting tips that have been successful for favorable cellular isolation and characterization of immune cells in DIO-mediated inflammation.

This protocol allows a researcher to isolate and characterize tissue-resident macrophages in various hallmark inflamed tissues extracted from diet-induced models of metabolic disorders.