Name: determining the phagocytic activity of clinical antibody samples
Uploaded: 2011-11-30T13:00:00
Duration: 9 min 5 s
Description: Watch this Scientific Journal Video about Determining the Phagocytic Activity of Clinical Antibody Samples at JoVE.com

The authors wish to acknowledge funding from NIH 3R01AI080289-02S1, and a Harvard University Center for AIDS Research Scholar Fellowship under NIH/NIAID 2P30AI060354-07.

1. Culture Phagocytic Cells
<ol>
<li>Culture THP-1 cells10 in RPMI 1640 supplemented with 10% fetal bovine serum as a stationary suspension in T flasks. Cell density should be maintained below 0.5x106/mL in order to maintain consistent levels of Fc&gamma;R expression and assay performance.</li>
</ol>
2. Prepare Biotinylated Antigen
<ol>
<li>Calculate the amount of sulfo-NHS LC biotin reagent kit necessary to biotinylate the target antigen of interest according to the manufacturer&rsquo;s instructions.</li>
<li>Dissolve the biotinylation reagent in water and immediately add the calculated amount to antigen in a buffer that does not contain primary amines. Allow the reaction to proceed for 1 hour at room temperature, mixing occasionally.</li>
<li>Remove excess, unconjugated biotin by buffer exchange in an Amicon centrifugal concentration unit of an appropriate molecular weight cutoff to retain the target antigen. &nbsp;Add sample to the top chamber of the concentration unit and add PBS to bring the total volume up to 15 mL fill line. Centrifuge at 4000 x g to bring volume down to approximately 1.5 mls. Repeating this process twice will remove 99% of the free biotin to ensure maximal coating of antigen to beads.</li>
</ol>
3. Prepare Antigen Saturated Beads
<ol>
<li>Wash 100 &mu;l of 1 mm fluorescent neutravidin beads twice in 1 ml 0.1% PBS-BSA after spinning down in a microcentrifuge at high speed. Resuspend washed beads in 100 &mu;l PBS-BSA, and aliquot into 10 tubes.</li>
<li>To determine antigen coating conditions which saturate the beads, combine 10 ul of the washed bead suspension with varying concentrations of biotinylated antigen in two-fold steps. Incubate overnight at 4&deg;C in a microcentrifuge tube on a rotator. </li>
</ol>
NOTE: Saturation of beads must be determined experimentally. This can be accomplished by identifying bead-coating conditions that yield maximal phagocytosis when beads are subsequently opsonized with a control monoclonal antibody. 
<ol start="3">
<li>Remove unbound antigen by washing with 1 mL PBS-BSA and centrifuge at high speed until beads are pelleted.&nbsp; Remove PBS-BSA and repeat. </li>
<li>Resuspend antigen-coated beads in a final volume of 1 ml in PBS-BSA. Beads can be stored for up to a week at 4&deg;C prior to use.</li>
</ol>
4. Prepare Antibody Samples
<ol>
<li>Clinical antibody samples can be purified from plasma using a Melon gel IgG purification kit according to manufacturers instructions. Purified IgG can be stored at 4&deg;C until ready for use.</li>
</ol>
NOTE: Proper precautions regarding handling human samples must always been taken.
<ol start="2">
<li>Determine the concentration of purified antibodies by absorbance at A280, and dilute samples to 1 mg/ml in PBS.</li>
<li>Prepare positive and negative monoclonal control antibodies by dilution to 1 mg/ml in PBS.</li>
</ol>
5. Plating the Experiment
<ol>
<li>Resuspend the washed, antigen saturated bead solution prepared above by vortexing, and transfer 10 &mu;l into each well of a round-bottom 96 well plate. Care must be taken to continually agitate the bead suspension to ensure equal numbers of fluorescent beads are added to each well.</li>
<li>Add varying concentrations of the antibodies of interest to each well, creating a dose-response curve for each antibody. Optimal concentrations will differ between samples depending on the titer of antibodies present, but a range of 0.01- 100 &mu;g/ml final concentration will provide good coverage and allow identification of the concentration range of interest. Ensure antibodies are added in volumes no larger than 20 &mu;l.</li>
<li>Incubate beads and antibody samples for 2 hours at 37&deg;C to allow the antibodies to opsonize the beads.</li>
<li>Prepare a suspension of THP-1 cells at 2.5 x 105 cells/ml, and add 200 &mu;l of this suspension to each well, for a total of 5 x 104 THP-1 cells in each well.</li>
<li>Incubate overnight at 37&deg;C, 5% CO2, in a stationary incubator, allowing cells and beads to pellet via gravity.</li>
</ol>
NOTE: Signal to noise may be improved by determining the optimal bead:antibody:THP-1 cell ratios for a given antigen and antibody source.
6. Flow Cytometric Analysis
<ol>
<li>Remove 100 &mu;l of supernatant from each well being careful not to disturb the cell pellet. Supernatant can be saved if cytokine secretion determinations or other analyses are desired.</li>
<li>For fixation, add 100 &mu;l of 4% paraformaldehyde to each well and pipet to resuspend and mix cells.</li>
<li>High throughput flow cytometric analysis can be performed using a BD LSR II equipped with an HTS plate reader.&nbsp; </li>
<li>Program software to mix each well three times (100 &mu;l mix volume) and analyze 30 &mu;l, or at least 2000 cell events, of each sample. </li>
<li>Data collected can be analyzed in FlowJo or equivalent software.&nbsp; Useful metrics include the percent of bead+, or fluorescent cells, which provides a measure of the number of phagocytic cells present, as well as the fluorescence intensity of the phagocytic cells, which provides a measure of the number of phagocytosed beads.&nbsp; Multiplying these values generates an integrated mean fluorescent intensity, or iMFI.</li>
<li>The average number of beads phagocytosed by each phagocytic cell can be calculated by dividing the iMFI by the mean fluorescence of 1 bead.</li>
</ol>
7. Representative Results
There should be clear differentiation of antibody samples from affected and unaffected subjects. Figure 1A presents the FACS histograms of an antibody sample from an HIV negative (black trace) and an HIV positive (gray trace) subject, and demonstrates the increased phagocytosis driven by the presence of antigen-specific antibodies.
Optimal sensitivity of the assay is dependent on saturation of the beads with biotinylated antigen. Figure 1B presents the phagocytosis observed when beads coated with differing amounts of antigen were opsonized with 3 control monoclonal antibodies (including a non-binding antibody, triangles), and establishes 2&mu;g antigen/&mu;l beads as a saturating concentration for this antigen.
A dose-response curve is presented in Figure 2, and demonstrates the differential capacity of subject antibody samples to induce phagocytosis over an antibody concentration range of 0.05-5 &mu;g/ml. This differential phagocytosis may be driven by either differences in titer or Fc domain properties such as IgG subclass and glycosylation state.
When THP-1 cells are imaged by fluorescent microscopy, there is clear evidence of bead phagocytosis. Figure 3 presents two 63x still images of THP-1 cells after incubation with antibody opsonized (green) and non-opsonized (red) beads, demonstrating the lack of phagocytic uptake in the absence of antibody. When time lapse microscopy is performed, the antibody-specific phagocytic uptake of fluorescent beads is even more striking (Movie 1, 20x magnification).
Previous work has confirmed internalization of the beads associated with the cells, and experiments with primary monocytes have agreed well with phagocytic scores (iMFI values) in this high throughput assay (data not shown).
<img src="/files/ftp_upload/3588/3588fig1.jpg" alt="Figure 1"/> Figure 1.&nbsp;
Assay Quality Control. 1A, Flow cytometry histograms of phagocytosis for an antibody sample from an HIV negative subject (black trace) and an HIV positive subject (gray trace). 1B: Experimental determination of the optimal bead coating conditions for a sample antigen. Antigen-specific monoclonal antibodies (circle, square) demonstrate maximal phagocytosis of beads coated with &gt;2 &mu;g antigen/&mu;l of beads, while a control antibody (triangle) demonstrates no phagocytic activity.
<img src="/files/ftp_upload/3588/3588fig2.jpg" alt="Figure 2"/> Figure 2.&nbsp;
Phagocytosis Dose-Response Curve. Clinical antibody samples from HIV positive (treated, untreated, and exhibiting control of viral replication in the absence of anti-retroviral therapy) and HIV negative subjects drive phagocytosis of gp120 (HIV envelope) coated beads differentially.
<img src="/files/ftp_upload/3588/3588fig3.jpg" alt="Figure 3"/> Figure 3.&nbsp;
Efficient Internalization. Microscopy confirms the internalization of antibody-opsonized beads (green), while non-opsonized beads remain in solution (63x magnification).
Movie 1. Time-lapse microscopy of antibody-driven phagocytosis was performed over the course of 14 hours and allows visualization of the phagocytic activity of the THP-1 cells utilized in the high-throughput assay. Green fluorescent beads are antibody opsonized, red fluorescent beads provide a negative control. <a href="https://www.jove.com/files/ftp_upload/3588/3588movie1.mov">Click here to watch the movie.</a>

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No conflicts of interest declared.

The assay described here allows high-throughput analysis of the phagocytic activity of clinical antibody samples by utilizing a monocytic cell line long utilized to study phagocytic processes10 and plate based automated flow cytometry. This assay is advantageous over others in its ability to precisely characterize antigen-specific antibody subsets, allowing not only for study of differences in phagocytosis driven by disease-specific antibodies from different subjects, but also study of multiple antigen specificities within the same subject. Because antibody recruitment of innate effector cells, including monocytes and other antigen presenting cells strongly impacts disease outcome11-13, and is biologically variable depending antibody geometry, IgG subclass, and glycosylation at Asparagine297 of the Fc domain14-16, this method provides for evaluation of a critical antibody mechanism of action. Furthermore, because antibody-mediated phagocytosis is exploited by some pathogens during the course of infection7,17,18, this assay holds promise in evaluation of the process of antibody-dependent enhancement, and highlights the dual nature of phagocytosis as a potentially protective, as well as potentially detrimental antibody activity.
The assay described can be utilized to analyze antibodies from clinical samples of patient populations, but also of vaccinees, and may be adapted to utilize serum rather than purified IgG. Additionally, cytokine secretion in response to phagocytosis can be analysed from the culture supernatant, and the influence of specific FcR can be determined by using FcR-blocking antibodies, allowing not only differences in phagocytic potency to be determined, but downstream signaling events, with insight into mechanism, and may help to resolve and refine understanding of this immune mechanism.

<table border="1">
<tbody>
<tr>
<td>Name of the reagent</td>
<td>Company</td>
<td>Catalogue number</td>
<td>Comments</td>
</tr>
<tr>
<td>EZ-Link Micro Sulfo-NHS-LC-Biotinylation Kit</td>
<td>Thermo Scientfic</td>
<td>21935</td>
<td>&nbsp;</td>
</tr>
<tr>
<td>Amicon Ultra-15 Centrifugal Filter Unit with Ultracel-50 membrane</td>
<td>Millipore</td>
<td>UFC905024</td>
<td>Filter unit size may vary according to antigen size</td>
</tr>
<tr>
<td>FluoSpheres NeutrAvidin labeled microspheres, 1.0 &mu;m, yellow-green fluorescent (505/515) </td>
<td>Invitrogen</td>
<td>F-8776</td>
<td>Manufacturers produce this product in various colors</td>
</tr>
<tr>
<td>Melon Gel IgG Purification Kit</td>
<td>Thermo</td>
<td>45212</td>
<td>Care should be taken to verify the purity of Ab samples by SDS-page.</td>
</tr>
</tbody>
</table>

determining the phagocytic activity of clinical antibody samples

Antibody-driven phagocytosis is induced via the engagement of Fc receptors on professional phagocytes, and can contribute to both clearance as well as pathology of disease. While the properties of the variable domains of antibodies have long been considered critical to in vivo function, the ability of antibodies to recruit innate immune cells via their Fc domains has become increasingly appreciated as a major factor in their efficacy, both in the setting of recombinant monoclonal antibody therapy, as well as in the course of natural infection or vaccination1-3.
Importantly, despite its nomenclature as a constant domain, the antibody Fc domain does not have constant function, and is strongly modulated by IgG subclass (IgG1-4) and glycosylation at Asparagine 2974-6. Thus, this method to study functional differences of antigen-specific antibodies in clinical samples will facilitate correlation of the phagocytic potential of antibodies to disease state, susceptibility to infection, progression, or clinical outcome.
Furthermore, this effector function is particularly important in light of the documented ability of antibodies to enhance infection by providing pathogens access into host cells via Fc receptor-driven phagocytosis7. Additionally, there is some evidence that phagocytic uptake of immune complexes can impact the Th1/Th2 polarization of the immune response8.
Here, we describe an assay designed to detect differences in antibody-induced phagocytosis, which may be caused by differential IgG subclass, glycan structure at Asn297, as well as the ability to form immune complexes of antigen-specific antibodies in a high-throughput fashion. To this end, 1 &mu;m fluorescent beads are coated with antigen, then incubated with clinical antibody samples, generating fluorescent antigen specific immune complexes. These antibody-opsonized beads are then incubated with a monocytic cell line expressing multiple Fc&gamma;Rs, including both inhibitory and activating. Assay output can include phagocytic activity, cytokine secretion, and patterns of Fc&gamma;Rs usage, and are determined in a standardized manner, making this a highly useful system for parsing differences in this antibody-dependent effector function in both infection and vaccine-mediated protection9.

Watch this Scientific Journal Video about Determining the Phagocytic Activity of Clinical Antibody Samples at JoVE.com

Determining the Phagocytic Activity of Clinical Antibody Samples

We present a high-throughput flow cytometric assay to determine the phagocytic activity of antigen-specific antibodies from clinical samples, utilizing fluorescent antigen-coated beads and a monocytic cell line expressing multiple Fc receptors&#x2014;providing receptor usage and phagocytic activity determinations in a standardized and reproducible fashion for any antigen of interest.

Antibody-driven phagocytosis is induced via the engagement of Fc receptors on professional phagocytes, and can contribute to both clearance as well as ...

determining-the-phagocytic-activity-of-clinical-antibody-samples

Research

JoVE Journal

Immunology and Infection

Determining the Reactivity and Titre of Serum using a Haemagglutination Assay

Haemagglutination is a specific form of agglutination and is used when antibodies bind to red blood cells, which act as a particulate antigen. Red blood cells are particularly useful targets as they are readily available and agglutination is observable using the naked eye. This technique is commonly used to determine the titre of an antibody (Ab), for blood grouping and viral quantification. In this video, the steps involved in preparing and performing a haemagglutination assay is demonstrated using antibodies specific to blood group A-antigens added to red blood cells (Revercells). The antiserum is serially diluted in a 96 well U-bottom microtitre tray, to which is added a suspension of Revercells. The samples are mixed and then incubated at 37&deg;C for 60 minutes. After this time, the samples can then be easily scored for  ve, +ve and intermediate (-/+) haemagglutination reactions. This approach allows for the reactivity and titre of a serum sample to be assessed using a rapid and simple technique. The video will cover the theory behind the assay, how the results are read and interpreted, how the titre is determined, how the assay can be modified and any issues associated with the use of this technique.

Haemagglutination is a form of agglutination where antibodies bind to red blood cells. Red blood cells are both readily available and the results are readily observable using the naked eye. This video demonstrates the steps involved in a haemagglutination assay, interpreting the results and determining the titre.

Haemagglutination is a specific form of agglutination and is used when antibodies bind to red blood cells, which act as a particulate antigen. Red blood ...

Using a Pan-Viral Microarray Assay (Virochip) to Screen Clinical Samples for Viral Pathogens

The diagnosis of viral causes of many infectious diseases is difficult due to the inherent sequence diversity of viruses as well as the ongoing emergence of novel viral pathogens, such as SARS coronavirus and 2009 pandemic H1N1 influenza virus, that are not detectable by traditional methods. To address these challenges, we have previously developed and validated a pan-viral microarray platform called the Virochip with the capacity to detect all known viruses as well as novel variants on the basis of conserved sequence homology1. Using the Virochip, we have identified the full spectrum of viruses associated with respiratory infections, including cases of unexplained critical illness in hospitalized patients, with a sensitivity equivalent to or superior to conventional clinical testing2-5. The Virochip has also been used to identify novel viruses, including the SARS coronavirus6,7, a novel rhinovirus clade5, XMRV (a retrovirus linked to prostate cancer)8, avian bornavirus (the cause of a wasting disease in parrots)9, and a novel cardiovirus in children with respiratory and diarrheal illness10. The current version of the Virochip has been ported to an Agilent microarray platform and consists of ~36,000 probes derived from over ~1,500 viruses in GenBank as of December of 2009. Here we demonstrate the steps involved in processing a Virochip assay from start to finish (~24 hour turnaround time), including sample nucleic acid extraction, PCR amplification using random primers, fluorescent dye incorporation, and microarray hybridization, scanning, and analysis.

Using a Pan-Viral Microarray Assay Virochip to Screen Clinical Samples for Viral Pathogens

The Virochip is a pan-viral microarray designed to simultaneously detect all known viruses as well as novel viruses on the basis of conserved sequence homology. Here we demonstrate how to run a Virochip assay to analyze clinical samples for the presence of both known and unknown viruses.

The diagnosis of viral causes of many infectious diseases is difficult due to the inherent sequence diversity of viruses as well as the ongoing emergence ...

Multiplex Detection of Bacteria in Complex Clinical and Environmental Samples using Oligonucleotide-coupled Fluorescent Microspheres

Bacterial vaginosis (BV) is a recurring polymicrobial syndrome that is characterized by a change in the "normal" microbiota from Lactobacillus-dominated to a microbiota dominated by a number of bacterial species, including Gardnerella vaginalis, Atopobium vaginae, and others1-3. This condition is associated with a range of negative health outcomes, including HIV acquisition4, and it can be difficult to manage clinically5. Furthermore, diagnosis of BV has relied on the use of Gram stains of vaginal swab smears that are scored on various numerical criteria6,7. While this diagnostic is simple, inexpensive, and well suited to resource-limited settings, it can suffer from problems related to subjective interpretations and it does not give a detailed profile of the composition of the vaginal microbiota8. Recent deep sequencing efforts have revealed a rich, diverse vaginal microbiota with clear differences between samples taken from individuals that are diagnosed with BV compared to those individuals that are considered normal9,10, which has resulted in the identification of a number of potential targets for molecular diagnosis of BV11,12. These studies have provided a wealth of useful information, but deep sequencing is not yet practical as a diagnostic method in a clinical setting. We have recently described a method for rapidly profiling the vaginal microbiota in a multiplex format using oligonucleotide-coupled fluorescent beads with detection on a Luminex platform13. This method, like current Gram stain-based methods, is rapid and simple but adds the additional advantage of exploiting molecular knowledge arising from sequencing studies in probe design. This method therefore provides a way to profile the major microorganisms that are present in a vaginal swab that can be used to diagnose BV with high specificity and sensitivity compared to Gram stain while providing additional information on species presence and abundance in a semi-quantitative and rapid manner. This multiplex method is expandable well beyond the range of current quantitative PCR assays for particular organisms, which is currently limited to 5 or 6 different assays in a single sample14. Importantly, the method is not limited to the detection of bacteria in vaginal swabs and can be easily adapted to rapidly profile nearly any microbial community of interest. For example, we have recently begun to apply this methodology to the development of diagnostic tools for use in wastewater treatment plants.

We describe a multiplex method for the detection of microorganisms within a sample using oligonucleotide-coupled fluorescent beads. Amplicon from all organisms within a sample is hybridized to a panel of probe-coupled beads. A Luminex or Bio-Plex instrument is used to query each bead for bead type and hybridization signal.

Bacterial vaginosis (BV) is a recurring polymicrobial syndrome that is characterized by a change in the "normal" microbiota from Lactobacillus-dominated ...

Determining Optimal Cytotoxic Activity of Human Her2neu Specific CD8 T cells by Comparing the Cr51 Release Assay to the xCELLigence System

Cytotoxic CD8 T cells constitute a subgroup of T cells that are capable of inducing the death of infected or malignant host cells1. These cells express a specialized receptor, called the T cell receptor (TCR), which can recognize a specific antigenic peptide bound to HLA class I molecules2. Engagement of infected cells or tumor cells through their HLA class I molecule results in production of lytic molecules such as granzymes and perforin resulting in target cell death. While it is useful to determine frequencies of antigen-specific CD8 T cells using assays such as the ELIspot or flow cytometry, it is also helpful to ascertain the strength of CD8 T cell responses using cytotoxicity assays3. The most recognizable assay for assessing cytotoxic function is the Chromium Release Assay (CRA), which is considered a standard assay 4. The CRA has several limitations, including exposure of cells to gamma radiation, lack of reproducibility, and a requirement for large numbers of cells. Over the past decade, there has been interest in adopting new strategies to overcome these limitations. Newer approaches include those that measure caspase 
release 4, BLT esterase activity 5 and surface expression of CD107 6. The impedance-based assay, using the Roche xCelligence system, was examined in the present paper for its potential as an alternative to the CRA. Impedance or opposition to an electric current occurs when adherent tumor cells bind to electrode plates. Tumor cells detach following killing and electrical impedance is reduced which can be measured by the xCelligence system. The ability to adapt the impedance-based approach to assess cell-mediated killing rests on the observation that T cells do not adhere tightly to most surfaces and do not appear to have much impact on impedance thus diminishing any concern of direct interference of the T cells with the measurement. Results show that the impedance-based assay can detect changes in the levels of antigen-specific cytotoxic CD8 T cells with increased sensitivity relative to the standard CRA. Based on these results, impedance-based approaches may be good alternatives to CRAs or other approaches that aim to measure cytotoxic CD8 T cell functionality.

The chromium release assay, a common assay for detecting cytotoxic T cell activity, has several limitations. Using antigen-specific CD8 T cells and the human breast cancer tumor line, SKBR3, in the present article, an impedance-based approach was examined for the capability of detecting cell killing.

Cytotoxic CD8 T cells constitute a subgroup of T  cells that are capable of inducing the death of infected or malignant host  cells1. These cells express ...

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne relapsing fever (Borrelia spp.), Rocky Mountain Spotted fever (Rickettsia rickettsii), ehrlichiosis (Ehrlichia chaffeensis and E. equi), anaplasmosis (Anaplasma phagocytophilum), encephalitis (tick-borne encephalitis virus), babesiosis (Babesia spp.), Colorado tick fever (Coltivirus), and tularemia (Francisella tularensis) 1-8. To be properly transmitted into the host these infectious agents differentially regulate gene expression, interact with tick proteins, and migrate through the tick 3,9-13. For example, the Lyme disease agent, Borrelia burgdorferi, adapts through differential gene expression to the feast and famine stages of the tick's enzootic cycle 14,15. Furthermore, as an Ixodes tick consumes a bloodmeal Borrelia replicate and migrate from the midgut into the hemocoel, where they travel to the salivary glands and are transmitted into the host with the expelled saliva 9,16-19.
As a tick feeds the host typically responds with a strong hemostatic and innate immune response 11,13,20-22. Despite these host responses, I. scapularis can feed for several days because tick saliva contains proteins that are immunomodulatory, lytic agents, anticoagulants, and fibrinolysins to aid the tick feeding 3,11,20,21,23. The immunomodulatory activities possessed by tick saliva or salivary gland extract (SGE) facilitate transmission, proliferation, and dissemination of numerous tick-borne pathogens 3,20,24-27. To further understand how tick-borne infectious agents cause disease it is essential to dissect actively feeding ticks and collect tick saliva. This video protocol demonstrates dissection techniques for the collection of hemolymph and the removal of salivary glands from actively feeding I. scapularis nymphs after 48 and 72 hours post mouse placement. We also demonstrate saliva collection from an adult female I. scapularis tick.

Saliva, Salivary Gland, and Hemolymph Collection from Ixodes scapularis Ticks

The collection of infected tick hemolymph, salivary glands, and saliva is important to study how tick-borne pathogens cause disease. In this protocol we demonstrate how to collect hemolymph and salivary glands from feeding Ixodes scapularis nymphs. We also demonstrate saliva collection from female I. scapularis adults.

Ticks are found worldwide and afflict humans with many tick-borne illnesses. Ticks are vectors for pathogens that cause Lyme disease and tick-borne ...

Fluorescence Microscopy Methods for Determining the Viability of Bacteria in Association with Mammalian Cells

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols to address these questions include colony count assays, gentamicin protection assays, and electron microscopy. Colony count and gentamicin protection assays only assess the viability of the entire bacterial population and are unable to determine individual bacterial viability. Electron microscopy can be used to determine the viability of individual bacteria and provide information regarding their localization in host cells. However, bacteria often display a range of electron densities, making assessment of viability difficult. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells. These assays were developed originally to assess survival of Neisseria gonorrhoeae in primary human neutrophils, but should be applicable to any bacterium-host cell interaction. These protocols combine membrane-permeable fluorescent dyes (SYTO9 and 4&#39;,6-diamidino-2-phenylindole [DAPI]), which stain all bacteria, with membrane-impermeable fluorescent dyes (propidium iodide and SYTOX Green), which are only accessible to nonviable bacteria. Prior to eukaryotic cell permeabilization, an antibody or fluorescent reagent is added to identify extracellular bacteria. Thus these assays discriminate the viability of bacteria adherent to and inside eukaryotic cells. A protocol is also provided for using the viability dyes in combination with fluorescent antibodies to eukaryotic cell markers, in order to determine the subcellular localization of individual bacteria. The bacterial viability dyes discussed in this article are a sensitive complement and/or alternative to traditional microbiology techniques to evaluate the viability of individual bacteria and provide information regarding where bacteria survive in host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. This article outlines protocols for the use of fluorescent dyes that reveal the viability of individual bacteria inside and associated with host cells.

Central to the field of bacterial pathogenesis is the ability to define if and how microbes survive after exposure to eukaryotic cells. Current protocols ...

Scalable High Throughput Selection From  Phage-displayed Synthetic Antibody Libraries

The demand for antibodies that fulfill the needs of both basic and clinical research applications is high and will dramatically increase in the future. However, it is apparent that traditional monoclonal technologies are not alone up to this task. This has led to the development of alternate methods to satisfy the demand for high quality and renewable affinity reagents to all accessible elements of the proteome. Toward this end, high throughput methods for conducting selections from phage-displayed synthetic antibody libraries have been devised for applications involving diverse antigens and optimized for rapid throughput and success. Herein, a protocol is described in detail that illustrates with video demonstration the parallel selection of Fab-phage clones from high diversity libraries against hundreds of targets using either a manual 96 channel liquid handler or automated robotics system. Using this protocol, a single user can generate hundreds of antigens, select antibodies to them in parallel and validate antibody binding within 6-8 weeks. Highlighted are: i) a viable antigen format, ii) pre-selection antigen characterization, iii) critical steps that influence the selection of specific and high affinity clones, and iv) ways of monitoring selection effectiveness and early stage antibody clone characterization. With this approach, we have obtained synthetic antibody fragments (Fabs) to many target classes including single-pass membrane receptors, secreted protein hormones, and multi-domain intracellular proteins. These fragments are readily converted to full-length antibodies and have been validated to exhibit high affinity and specificity. Further, they have been demonstrated to be functional in a variety of standard immunoassays including Western blotting, ELISA, cellular immunofluorescence, immunoprecipitation and related assays. This methodology will accelerate antibody discovery and ultimately bring us closer to realizing the goal of generating renewable, high quality antibodies to the proteome.

A method is described with visual accompaniment for conducting scalable, high throughput selections from phage-displayed combinatorial synthetic antibody libraries against hundreds of antigens simultaneously. Using this parallel approach, we have isolated antibody fragments that exhibit high affinity and specificity for diverse antigens that are functional in standard immunoassays.

The demand for antibodies that fulfill the needs of both basic and clinical research applications is high and will dramatically increase in the future. ...

A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

The isolation of giant viruses is of great interest in this new era of virology, especially since these giant viruses are related to protists. Giant viruses may be potentially pathogenic for many species of protists. They belong to the recently described order of Megavirales. The new lineage Faustovirus that has been isolated from sewage samples is distantly related to the mammalian pathogen African swine fever virus. This virus is also specific to its amoebal host, Vermamoeba vermiformis, a protist common in health care water systems. It is crucial to continue isolating new Faustovirus genotypes in order to enlarge its genotype collection and study its pan-genome. We developed new strategies for the isolation of additional strains by improving the use of antibiotic and antifungal combinations in order to avoid bacterial and fungal contaminations of the amoeba co-culture and favoring the virus multiplication. We also implemented a new starvation medium to maintain V. vermiformis in optimal conditions for viruses co-culture. Finally, we used flow cytometry rather than microscopic observation, which is time-consuming, to detect the cytopathogenic effect. We obtained two isolates from sewage samples, proving the efficiency of this method and thus widening the collection of Faustoviruses, to better understand their environment, host specificity and genetic content.

A Rapid Strategy for the Isolation of New Faustoviruses from Environmental Samples Using Vermamoeba vermiformis

We describe here the latest advances in viral isolation for the characterization of new genotypes of Faustovirus, a new asfarvirus-related lineage of giant viruses. This protocol can be applied to the high throughput isolation of viruses, especially giant viruses infecting amoeba.

The isolation of giant viruses is of great interest in this new era of virology, especially since these giant viruses are related to protists. Giant ...

Fast and Specific Assessment of the Halogenating Peroxidase Activity in Leukocyte-enriched Blood Samples

In this paper a protocol for the quick and standardized enrichment of leukocytes from small whole blood samples is described. This procedure is based on the hypotonic lysis of erythrocytes and can be applied to human samples as well as to blood of non-human origin. The small initial sample volume of about 50 to 100 &#181;l makes this method applicable to recurrent blood sampling from small laboratory animals. Moreover, leukocyte enrichment is achieved within minutes and with low material efforts regarding chemicals and instrumentation, making this method applicable in multiple laboratory environments.
Standardized purification of leukocytes is combined with a highly selective staining method to evaluate halogenating peroxidase activity of the heme peroxidases, myeloperoxidase (MPO) and eosinophil peroxidase (EPO), i.e., the formation of hypochlorous and hypobromous acid (HOCl and HOBr). While MPO is strongly expressed in neutrophils, the most abundant immune cell type in human blood as well as in monocytes, the related enzyme EPO is exclusively expressed in eosinophils. The halogenating activity of these enzymes is addressed by using the almost HOCl- and HOBr-specific dye aminophenyl fluorescein (APF) and the primary peroxidase substrate hydrogen peroxide. Upon subsequent flow cytometry analysis all peroxidase-positive cells (neutrophils, monocytes, eosinophils) are distinguishable and their halogenating peroxidase activity can be quantified. Since APF staining may be combined with the application of cell surface markers, this protocol can be extended to specifically address leukocyte sub-fractions. The method is applicable to detect HOCl and HOBr production both in human and in rodent leukocytes.
Given the widely and diversely discussed immunological role of these enzymatic products in chronic inflammatory diseases, this protocol may contribute to a better understanding of the immunological relevance of leukocyte-derived heme peroxidases.

This protocol describes the quick enrichment of leukocytes from small blood samples for a subsequent specific determination of the halogenating peroxidase activity within the cells. The method can be applied to human and non-human material and may contribute to the evaluation of new inflammatory markers.

In this paper a protocol for the quick and standardized enrichment of leukocytes from small whole blood samples is described. This procedure is based on ...

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, including phagocytosis of damaged synapses or cells, debris, and/or invading pathogens. Impaired phagocytic function has been implicated in the pathogenesis of diseases such as Alzheimer's and age-related macular degeneration, where amyloid-&#946; plaque and drusen accumulate, respectively. Despite its importance, microglial phagocytosis has been challenging to assess in vivo. Here, we describe a simple, yet robust, technique for precisely monitoring and quantifying the in vivo phagocytic potential of retinal microglia. Previous methods have relied on immunohistochemical staining and imaging techniques. Our method uses flow cytometry to measure microglial uptake of fluorescently labeled particles after intravitreal delivery to the eye in live rodents. This method replaces conventional practices that involve laborious tissue sectioning, immunostaining, and imaging, allowing for more precise quantification of microglia phagocytic function in just under six hours. This procedure can also be adapted to test how various compounds alter microglial phagocytosis in physiological settings. While this technique was developed in the eye, its use is not limited to vision research.

Assessing Retinal Microglial Phagocytic Function In Vivo Using a Flow Cytometry-based Assay

Microglial phagocytosis is critical for the maintenance of tissue homeostasis and inadequate phagocytic function has been implicated in pathology. However, assessing microglia function in vivo is technically challenging. We have developed a simple but robust technique for precisely monitoring and quantifying the phagocytic potential of microglia in a physiological setting.

Microglia are the tissue resident macrophages of the central nervous system (CNS) and they perform a variety of functions that support CNS homeostasis, ...