The authors would like to thank Joshua Kimani, clinical director of the research program at the University of Nairobi, for his assistance with mucosal immunology studies related to this protocol. The authors would like to acknowledge funding from CHVI grant MOP 86721.

Ethics statement: The research ethics boards of both the University of Manitoba and Kenyatta National Hospital/University of Nairobi approved this study and written informed consent was obtained from all study participants.
1. Preparation of Media and CMC Collection Tubes
<ol>
	<li>Prepare phosphate buffered saline (PBS) solution (137.93 mM NaCl, 2.67 mM KCl, 8.1 mM Na2HPO4, 1.47 mM KH2PO4). Autoclave for sterility. This can be stored at 4 &#186;C for several months.</li>
	<li>Pre-aliquot 5 ml&#160;PBS into 50 ml&#160;falcon tube (one per sample to be collected) and store at 4 &#186;C until sample collection time.</li>
	<li>Label 2-4 cryovials or 1.5 ml&#160;microcentrifuge tubes per sample to collect lavage for storage.</li>
	<li>Prepare cell culture media: RPMI 1640 + 1% penicillin/streptomycin/amphotericin B (final concentration 100 units/ml, 100 &#956;g/ml&#160;and 250 ng/ml, respectively, with no FCS/FBS added.</li>
</ol>
2. Collection of Cytobrush Samples
Collection of cytobrush samples is a non-invasive procedure that must be performed by a trained MD or gynecologist.
<ol>
	<li>Collect cervical mononuclear cells (CMC) using both a cytobrush and a wooden or plastic scraper. Collect CMC from the participant under speculum examination by inserting the scraper and rotating around the cervical os (Figure 1). Insert the cytobrush into the endocervical os, rotate 360&#176;, and immediately place both the cytobrush and scraper into the 50 ml falcon tube containing 5 ml of PBS.</li>
	<li>Keep samples on ice/at 4 &#186;C until processing. To maintain cell viability, process samples within 2 hr&#160;of collection.</li>
	<li>If possible, collect matched blood samples in green top heparin vacutainers for peripheral blood mononuclear cell (PBMC) isolation to serve as a comparator/control for CMC analysis.</li>
</ol>
3. Isolation of CMCs
Perform sample processing in a biosafety level 2 laboratory, in a sterile biosafety cabinet with double gloves.
<ol>
	<li>During collection, CMC samples can become contaminated with blood. Exclude samples with visible blood contamination from the study in order to prevent confounding inclusion of PBMC in the final sample. Due to the low number of lymphocytes isolated from CMC samples, minor blood contamination can easily overwhelm the CMC lymphocyte component.</li>
	<li>Vortex falcon tubes containing both the cytobrush and scraper for 45 sec&#160;to dissociate cells from the brushes. Samples may become foamy during this process; this is normal.</li>
	<li>Use the cytobrush to scrape any remaining material off the scraper, and discard the scraper into a bleach solution. To collect any cells remaining attached to the brush/scraper, use a fresh glove to squeeze the scraper between the thumb and forefinger. Slide down the brush, allowing the cells and PBS to be collected in the same 50 ml tube. Discard the cytobrush into a bleach solution, and discard the glove. 
	IMPORTANT: Use a new glove for each sample.</li>
	<li>Fit a 100 &#956;m&#160;nylon cell strainer to a fresh 50 ml&#160;falcon tube. Using a transfer pipette, collect the PBS-cell suspension from the sample tube and filter through the strainer into the fresh tube.</li>
	<li>Add 5 ml&#160;of RPMI 1640 to the original sample tube. Using the transfer pipette, wash the sides of the sample tube with the RPMI, and transfer through the filter to the new collection tube. Wash the bottom of the nylon filter with the RPMI as well.</li>
	<li>Centrifuge the samples for 10 min at 514 x g. It is crucial to leave the centrifuge brake off during this step, in order to prevent dislodging of the CMC pellet.</li>
	<li>Gently remove the supernatant from the tube, taking care to not disturb the pellet. Pellet size will be highly variable; in some samples, the pellet is quite large due to the presence of large numbers of epithelial cells, while in other cases, the pellet is barely visible and highly transparent.</li>
	<li>Resuspend the pellet by gentle agitation of the falcon tube. Add 5 ml&#160;of PBS to wash the sample.</li>
	<li>Centrifuge again for 10 min at 514 x g with no centrifuge brake.</li>
	<li>Carefully discard the supernatant and re-suspend the pellet as above. The cells are ready for either cryopreservation (using a PBMC cryopreservation protocol), stimulation or flow cytometry phenotyping.</li>
</ol>
4. CMC Surface Staining and Flow Cytometry
<ol>
	<li>Resuspend the pellet from step 3.10 in 100 &#956;l of blocking solution and transfer either into a 96-well plate or FACS tube. The blocking solution (to block Fc&#947;&#160;receptors) recipe is as follows: 1.8 &#956;l&#160;mouse IgG (final concentration 0.2 &#956;g/&#956;l), 5 &#956;l&#160;FBS and 93.2 &#956;l&#160;FACS wash (PBS+2%FBS). Staining can be performed in a 96 well plate if pellet sizes are small enough, but may need to be performed in FACS tubes if pellets are routinely large. It is important to titrate antibodies accordingly.</li>
	<li>Block the samples for 10 min&#160;on ice/at 4 &#176;C.</li>
	<li>Wash the cells with 100 &#956;l of FACS Wash (PBS+2% FBS) in 96-well plates, or 500 &#956;l&#160;in FACS tubes. Centrifuge at 600 x g for 10 min at 4 &#176;C with the brake on low (or off, if a low brake setting is unavailable).</li>
	<li>Remove the supernatant and re-suspend the cell pellet by agitation. Add viability Stain (Live Dead fixable viability dye). Prepare viability dye according to manufacturer&#8217;s instructions. It may need to be titrated for use in each lab/cytometer setup. Incubate for 30 min&#160;in the dark at 4 &#176;C.</li>
	<li>Wash the cells 100&#160;&#956;l/500 &#956;l&#160;of PBS (for 96-well plate/FACS tubes). Centrifuge at 600 x g for 10 min&#160;with low/no brake. Remove supernatant and resuspend cells.</li>
	<li>Incubate the cells with the cocktail of surface marker antibodies, in a total volume of 100 &#956;l. IMPORTANT: Optimize and titrate each flow cytometry panel prior to sample staining. Incubate cells for 30 min&#160;in the dark at 4 &#176;C.</li>
	<li>Repeat step 4.5. if staining in a 96-well plate, transfer cells to a FACS tube and dilute to a final volume of 750 &#181;l of FACS Wash containing 1% paraformaldehyde (PFA) (or CytoFix, diluted 1 in 4). Proceed to data acquisition on a flow cytometer.</li>
</ol>
5. Collection and Preparation of Cervical Vaginal Lavage (CVL)
<ol>
	<li>Prior to cytobrush sampling, use a syringe to wash the endocervix with 2 ml of sterile PBS and aspirate the lavage from the posterior fornix.</li>
	<li>Collect the aspirated lavage sample into a 15 ml falcon tube and keep on ice/at 4 &#186;C until processing.</li>
	<li>Centrifuge sample at 400 x g for 7 min&#160;to remove cellular debris.</li>
	<li>Collect the supernatant, aliquot the sample into 1 ml&#160;or 500 &#956;l&#160;aliquots, and store at -70 &#176;C. Aliquots can be thawed for ELISA or bead array analysis of CVL proteins, but avoid multiple freeze/thaw cycles.</li>
	<li>If desired, resuspend the pellet of cellular debris in RNA Later to measure RNA expression by following the manufacturer&#8217;s protocol.</li>
</ol>
6. Data Acquisition
<ol>
	<li>Prepare a set of compensation tubes that matches the antibody panel. Run the compensation tubes to adjust spectral overlap. Run the samples on any multicolor flow cytometer that is configured for the fluorochrome conjugated antibodies used in the panel.</li>
	<li>Acquire a PBMC sample first to adjust Forward Scatter (FSC) and Side Scatter (SSC) voltage in order to detect the lymphocyte population. Try to keep them at 100 on each axis on a linear scale. Running a PBMC control will make it significantly easier to find the CMC lymphocyte population.</li>
	<li>The FSC threshold for CMC samples may need to be increased relative to PBMC samples due to smear up the SSC axis at low FSC values.</li>
	<li>Vortex each tube before acquiring. Acquire the entire tube to maximize the number of lymphocyte gate events collected. Monitor the machine carefully to avoid clogging.</li>
	<li>Export the data into a flow cytometric analysis program such as FlowJo.</li>
</ol>
7. Gating Strategy
<ol>
	<li>Select Forward side scatter high (FSC-H)/Forward side scatter area (FSC-A) to determine the singlet population.</li>
	<li>Select Time versus Fluorescent marker (as example FITC) to control for the quality of the acquisition. Change in flow rate may introduce artifacts in the data. Exclude any area that shows discrepancy in flow rate.</li>
	<li>Identify the lymphocyte population on SSC-A/FSC-A plot. Use a PBMC control to facilitate the identification. Note that the CMC lymphocyte population might not be as clear as the PBMC control.</li>
	<li>Gate on SSC-A/viability dye to exclude dead cells from live cells. Dead cells can non-specifically incorporate the antibodies, which will introduce artifacts.</li>
	<li>Gate on the SSC-A/CD3 to identify the T cell population. From this gate, identify the CD4+ and CD8+ populations.</li>
</ol>

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M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Elevated+T+cell+counts+and+RANTES+expression+in+the+genital+mucosa+of+HIV-1-resistant+Kenyan+commercial+sex+workers.">Elevated T cell counts and RANTES expression in the genital mucosa of HIV-1-resistant Kenyan commercial sex workers.</a> The Journal of infectious diseases. 192, 728-738 (2005).</li><li>Hirbod, T., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/entrez/query.fcgi?db=PubMed&cmd=Search&doptcmdl=Citation&defaultField=Title+Word&term=Stable+CD4+expression+and+local+immune+activation+in+the+ectocervical+mucosa+of+HIV-infected+women.">Stable CD4 expression and local immune activation in the ectocervical mucosa of HIV-infected women.</a> Journal of immunology. 191, 3948-3954 (2013).</li><li>Horton, R. 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The authors have nothing to disclose.

Given the large gaps in knowledge with respect to immunity at the female genital tract (FGT), phenotypic analysis of CMCs can provide a wide array of insights into multiple lymphocyte populations at the cervix. Coupled with proteomic analysis and viral load measurements in cervical lavage, immunity to sexually transmitted infections (STI)s and other pathogens can be dissected in various populations.
Technical considerations - CMCs: The isolation and successful staining of CMC ...

The majority of new HIV infections worldwide arise through heterosexual transmission, with women representing 47% of new infections in 2011 (UNAIDS1). Understanding the female genital tract (FGT), one of the main entry portals for HIV and other sexually transmitted pathogens, is of high importance on the path to finding efficient strategies to prevent infection. Immune responses at the genital mucosa are clearly unique and differ from those measured in peripheral blood2. However, current knowledge of the immune dynamics at the FGT is limited at best. To date, studies of the mucosal immune environment have largely focused on the gut-associated lymphoid tissues (GALT), where it has become clear that the early events in mucosal tissues following infection have a strong impact on subsequent disease progression3,4. Collecting samples from the genital mucosa represents a great challenge and is at least partially responsible for the lack of understanding of the immunology of the FGT. Solving the puzzle of the immune dynamic between host and pathogen in the context of the distinct environment that is the FGT necessitates efficient methods for collecting and analyzing samples from this locale.
The FGT is divided into two sections: the upper reproductive tract that includes the fallopian tubes, endometrium and endocervix, and the lower tract which contains the ectocervix and the vagina (reviewed by Kaushic et al5). It is still unclear what the relative contribution of these different sites is to HIV infection, but it is believed that both sites could contribute to HIV entry6. T cells represent 40-50% of the leucocytes in the upper and lower reproductive tracts, while macrophages comprise approximately 10% (reviewed in Rodriguez-Garcia et al2). T cells can be detected in the vagina, cervix, and endometrium. Macrophages are more strongly localized in the endometrium and myometrial connective tissue than the cervix, although they can be detected in both tissues. Finally, plasmacytoid dendritic cells (pDCs) and Langerhans cells can also be detected in FGT tissues. The phenotype and proportions of immune populations and their susceptibility to HIV infection may vary importantly according to hormonal cycles, the use of hormonal contraceptives, bacterial vaginosis or sexual activities5,7-9.
Diverse methods have been developed to study the immune populations and environment of the FGT. Cervical biopsy, cervical cytobrushes and cervicovaginal lavages (CVL)10-12 are the most commonly used across the literature. CVL collection by PBS lavage is the simplest method and allows the study of immune modulatory proteins but results in extremely low cell yield, and is therefore not suitable for studying the immune cell populations of the FGT13. CVL samples are, on the other hand, very useful for evaluating the immune environment of the FGT by measuring the expression of various cytokines, chemokines or antimicrobial factors using methods such as ELISA, cytokine bead array14 or mass spectrometry15,16. Characterization of immune cell frequencies, phenotypes and functions can be achieved by collecting cervical mononuclear cells (CMC) by cervical cytobrush or by cervical biopsy sampling.
Cervical biopsy sampling is an invasive method that increases the discomfort and risk of bleeding and takes 2 to 11 days to heal following the procedure depending on the immune status of the woman12. On the other hand, cervical cytobrushes, despite the lower yield of cells collected, is a less invasive and more convenient method to collect immune cells from the FGT. Both methods can reach the same yield of CD45+ leucocytes, but two sequential cervical cytobrushes are necessary to obtain the same amount of cells contained in one biopsy13. Nonetheless, cytobrush sampling still provides an acceptable number of cells (about 5,000 CD45+ cells/cytobrush) for further ex vivo phenotyping by flow cytometry14. Also, functional characterization can be carried out on these samples, as stimulation and intracellular flow cytometry or qPCR have been performed using cytobrush-derived CMCs to identify HIV-specific immune responses17 or Th cell polarization18. Expansion of the T cells population may also facilitate functional studies with CMCs19.
It is important to note that biopsies and cytobrushes sample distinct portions of the FGT. Biopsies are derived from the superior portion of the epithelium and stroma of the ectocervix12,13, while cervical cytobrushes sample the cervical os, collecting cells derived from the epithelium of endocervix and presumably the transformation zone. Cytobrush samples therefore sample a region composed of a single layer of columnar epithelium, while biopsies, include a region lined by a squamous stratified epithelium5. As a result, the nature of the leucocyte populations collected by cervical biopsy and cytobrush differs. Biopsies collect a higher proportion of CD3+ T cells, whereas cytobrushes result in collection of a higher proportion of CD14+ monocytes/macrophages13.
Studying the immunology of the FGT has been an interest for many years20-22 and we have accumulated a great deal of expertise with the study of cytobrush-derived CMCs. Our studies focus mostly on the study of HIV-infected, uninfected and HIV-exposed seronegative (HESN) female sex workers from Nairobi, Kenya. HIV preferentially replicates in activated T cells23 and lower numbers of activated cells that can be targeted by HIV in the FGT could contribute to protection against HIV acquisition. In line with this hypothesis, several studies have described lower immune activation among HESN sex workers who are highly exposed to HIV yet remain uninfected24,25, and this quiescent phenotype is also observed in the FGT14. Here, we describe methodology for processing and assessing T cells activation in CMC samples derived from cervical cytobrushes by ex vivo flow cytometry.

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			<td style="width:133px;">352360</td>
			<td>CMC processing</td>
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			<td height="21" style="height:21px;width:295px;">RPMI 1640</td>
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			<td style="width:133px;">SH30027.01</td>
			<td>CMC processing</td>
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			<td height="21" style="height:21px;width:295px;">Fetal Bovine serum&#160;</td>
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			<td style="width:136px;">Fisher</td>
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			<td>CVL storage</td>
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			<td height="21" style="height:21px;">LIVE/DEAD Fixable Cell Stain Kit</td>
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collection, isolation, and flow cytometric analysis of human endocervical samples

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female genital tract (FGT). Because heterosexual transmission now represents the dominant mechanism of HIV transmission, and given the continual spread of sexually transmitted infections (STIs), it is critical to understand the interplay between host and pathogen at the genital mucosa. The substantial gaps in knowledge around FGT immunity are partially due to the difficulty in successfully collecting and processing mucosal samples. In order to facilitate studies with sufficient sample size, collection techniques must be minimally invasive and efficient. To this end, a protocol for the collection of cervical cytobrush samples and subsequent isolation of cervical mononuclear cells (CMC) has been optimized. Using ex vivo flow cytometry-based immunophenotyping, it is possible to accurately and reliably quantify CMC lymphocyte/monocyte population frequencies and phenotypes. This technique can be coupled with the collection of cervical-vaginal lavage (CVL), which contains soluble immune mediators including cytokines, chemokines and anti-proteases, all of which can be used to determine the anti- or pro-inflammatory environment in the vagina.

Multiparameter flow cytometry is a powerful tool to dissect the phenotypes and functions of cell subsets in previously uncharacterized tissues. Analysis of CMC samples can yield information on both lymphocyte and monocyte populations with appropriate gating strategies.
A representative CMC gating strategy, compared to a matched PBMC profile, is shown in Figure 2. The FSC-A versus FSC-H plot allows for the exclusion of cell doublets, which are highly prevalent in CMC samples co...

Watch this Scientific Journal Video about Collection, Isolation, and Flow Cytometric Analysis of Human Endocervical Samples at JoVE.com

Collection, Isolation, and Flow Cytometric Analysis of Human Endocervical Samples

The use of cytobrush sampling to collect lymphocytes and monocytes from the endocervix is a minimally invasive technique that provides samples for analysis of female genital tract immunity. In this protocol, we describe the collection of cytobrush samples and immune cell isolation for flow cytometry assays.

Despite the public health importance of mucosal pathogens (including HIV), relatively little is known about mucosal immunity, particularly at the female ...

collection-isolation-flow-cytometric-analysis-human-endocervical

Research

JoVE Journal

Medicine

19.0K 次观看.  University of Manitoba. The use of cytobrush sampling to collect lymphocytes and monocytes from the endocervix is a minimally invasive technique that provides samples for analysis of female genital tract immunity. In this protocol, we describe the collection of cytobrush samples and immune cell isolation for flow cytometry assays. 

视频: Collection, Isolation, and Flow Cytometric Analysis of Human Endocervical Samples

Isolation of Human Islets from Partially Pancreatectomized Patients

Investigations into the pathogenesis of type 2 diabetes and islets of Langerhans malfunction 1 have been hampered by the limited availability of type 2 diabetic islets from organ donors2. Here we share our protocol for isolating islets from human pancreatic tissue obtained from type 2 diabetic and non-diabetic patients who have undergone partial pancreatectomy due to different pancreatic diseases (benign or malignant pancreatic tumors, chronic pancreatitis, and common bile duct or duodenal tumors). All patients involved gave their consent to this study, which had also been approved by the local ethics committee. The surgical specimens were immediately delivered to the pathologist who selected soft and healthy appearing pancreatic tissue for islet isolation, retaining the damaged tissue for diagnostic purposes. We found that to isolate more than 1,000 islets, we had to begin with at least 2 g of pancreatic tissue. Also essential to our protocol was to visibly distend the tissue when injecting the enzyme-containing media and subsequently mince it to aid digestion by increasing the surface area.
To extend the applicability of our protocol to include the occasional case in which a large amount (&gt;15g) of human pancreatic tissue is available , we used a Ricordi chamber (50 ml) to digest the tissue. During digestion, we manually shook the Ricordi chamber3 at an intensity that varied by specimen according to its level of tissue fibrosis. A discontinous Ficoll gradient was then used to separate the islets from acinar tissue. We noted that the tissue pellet should be small enough to be homogenously resuspended in Ficoll medium with a density of 1.125 g/ml. After isolation, we cultured the islets under stress free conditions (no shaking or rotation) with 5% CO2 at 37 &deg;C for at least 48 h in order to facilitate their functional recovery. Widespread application of our protocol and its future improvement could enable the timely harvesting of large quantities of human islets from diabetic and clinically matched non-diabetic subjects, greatly advancing type 2 diabetes research.

The supply of type 2 diabetic islets for research is insufficient. Here we share our protocol for isolating islets from patients undergoing partial pancreatectomy. This approach represents a unique venue for obtaining islets from type 2 diabetic and clinically matched non-diabetic subjects in adequate numbers for basic and clinical studies.

Investigations into the pathogenesis of type 2 diabetes and islets of Langerhans malfunction 1 have been hampered by the limited availability of type 2 ...

Collection Protocol for Human Pancreas

This dissection and sampling procedure was developed for the Network for Pancreatic Organ Donors with Diabetes (nPOD) program to standardize preparation of pancreas recovered from cadaveric organ donors. The pancreas is divided into 3 main regions (head, body, tail) followed by serial transverse sections throughout the medial to lateral axis. Alternating sections are used for fixed paraffin and fresh frozen blocks and remnant samples are minced for snap frozen sample preparations, either with or without RNAse inhibitors, for DNA, RNA, or protein isolation. The overall goal of the pancreas dissection procedure is to sample the entire pancreas while maintaining anatomical orientation.
Endocrine cell heterogeneity in terms of islet composition, size, and numbers is reported for human islets compared to rodent islets 1. The majority of human islets from the pancreas head, body and tail regions are composed of insulin-containing &beta;-cells followed by lower proportions of glucagon-containing &alpha;-cells and somatostatin-containing &delta;-cells. Pancreatic polypeptide-containing PP cells and ghrelin-containing epsilon cells are also present but in small numbers. In contrast, the uncinate region contains islets that are primarily composed of pancreatic polypeptide-containing PP cells 2. These regional islet variations arise from developmental differences. The pancreas develops from the ventral and dorsal pancreatic buds in the foregut and after rotation of the stomach and duodenum, the ventral lobe moves and fuses with the dorsal 3. The ventral lobe forms the posterior portion of the head including the uncinate process while the dorsal lobe gives rise to the rest of the organ. Regional pancreatic variation is also reported with the tail region having higher islet density compared to other regions and the dorsal lobe-derived components undergoing selective atrophy in type 1 diabetes4,5.
Additional organs and tissues are often recovered from the organ donors and include pancreatic lymph nodes, spleen and non-pancreatic lymph nodes. These samples are recovered with similar formats as for the pancreas with the addition of isolation of cryopreserved cells. When the proximal duodenum is included with the pancreas, duodenal mucosa may be collected for paraffin and frozen blocks and minced snap frozen preparations.

This video demonstrates a dissection procedure for processing human pancreas into multiple storage formats. Anatomical orientation is maintained throughout the pancreatic regions to allow definition of regional islet composition and density.

This dissection and sampling procedure was developed for the Network for Pancreatic Organ Donors with Diabetes (nPOD) program to standardize preparation ...

Isolation, Culture, and Imaging of Human Fetal Pancreatic Cell Clusters

For almost 30 years, scientists have demonstrated that human fetal ICCs transplanted under the kidney capsule of nude mice matured into functioning endocrine cells, as evidenced by a significant increase in circulating human C-peptide following glucose stimulation1-9. However in vitro, genesis of insulin producing cells from human fetal ICCs is low10; results reminiscent of recent experiments performed with human embryonic stem cells (hESC), a renewable source of cells that hold great promise as a potential therapeutic treatment for type 1 diabetes. Like ICCs, transplantation of partially differentiated hESC generate glucose responsive, insulin producing cells, but in vitro genesis of insulin producing cells from hESC is much less robust11-17. A complete understanding of the factors that influence the growth and differentiation of endocrine precursor cells will likely require data generated from both ICCs and hESC. While a number of protocols exist to generate insulin producing cells from hESC in vitro11-22, far fewer exist for ICCs10,23,24. Part of that discrepancy likely comes from the difficulty of working with human fetal pancreas. Towards that end, we have continued to build upon existing methods to isolate fetal islets from human pancreases with gestational ages ranging from 12 to 23 weeks, grow the cells as a monolayer or in suspension, and image for cell proliferation, pancreatic markers and human hormones including glucagon and C-peptide. ICCs generated by the protocol described below result in C-peptide release after transplantation under the kidney capsule of nude mice that are similar to C-peptide levels obtained by transplantation of fresh tissue6. Although the examples presented here focus upon the pancreatic endoderm proliferation and &#946;&#160;cell genesis, the protocol can be employed to study other aspects of pancreatic development, including exocrine, ductal, and other hormone producing cells.

A protocol to isolate, culture, and image islet cell clusters (ICCs) derived from human fetal pancreatic cells is described.&#160; The method details the steps necessary to generate ICCs from tissue, culture as monolayers or in suspension as aggregates, and image for markers of proliferation and pancreatic cell fate decisions.

For almost 30 years, scientists have demonstrated that human fetal ICCs transplanted under the kidney capsule of nude mice matured into functioning ...

High-throughput Flow Cytometry Cell-based Assay to Detect Antibodies to N-Methyl-D-aspartate Receptor or Dopamine-2 Receptor in Human Serum

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases in children and adults. These antibodies have been used to guide diagnosis and treatment. Cell-based assays have improved the detection of antibodies in patient serum. They are based on the surface expression of brain antigens on eukaryotic cells, which are then incubated with diluted patient sera followed by fluorochrome-conjugated secondary antibodies. After washing, secondary antibody binding is then analyzed by flow cytometry. Our group has developed a high-throughput flow cytometry live cell-based assay to reliably detect antibodies against specific neurotransmitter receptors. This flow cytometry method is straight forward, quantitative, efficient, and the use of a high-throughput sampler system allows for large patient cohorts to be easily assayed in a short space of time. Additionally, this cell-based assay can be easily adapted to detect antibodies to many different antigenic targets, both from the central nervous system and periphery. Discovering additional novel antibody biomarkers will enable prompt and accurate diagnosis and improve treatment of immune-mediated disorders.

Over the recent years, live cell-based assays have been used successfully to detect antibodies against surface and conformational antigens. Here, we describe a method using high-throughput flow cytometry enabling the analysis of large cohorts of patients. Detection of novel antibodies will improve diagnosis and treatment of immune-mediated disorders.

Over the recent years, antibodies against surface and conformational proteins involved in neurotransmission have been detected in autoimmune CNS diseases ...

Isolation and Functional Characterization of Human Ventricular Cardiomyocytes from Fresh Surgical Samples

Cardiomyocytes from diseased hearts are subjected to complex remodeling processes involving changes in cell structure, excitation contraction coupling and membrane ion currents. Those changes are likely&#160;to be responsible for the increased arrhythmogenic risk and the contractile alterations leading to systolic and diastolic dysfunction in cardiac patients. However, most information on the alterations of myocyte function in cardiac diseases has come from animal models.
Here we describe and validate a protocol to isolate viable myocytes from small surgical samples of ventricular myocardium from patients undergoing cardiac surgery operations. The protocol is described in detail. Electrophysiological and intracellular calcium measurements are reported to demonstrate the feasibility of a number of single cell measurements in human ventricular cardiomyocytes obtained with this method.
The protocol reported here can be useful for future investigations of the cellular and molecular basis of functional alterations of the human heart in the presence of different cardiac diseases. Further, this method can be used to identify novel therapeutic targets at cellular level and to test the effectiveness of new compounds on human cardiomyocytes, with direct translational value.

Current knowledge on the cellular basis of cardiac diseases mostly relies on studies on animal models. Here we describe and validate a novel method to obtain single viable cardiomyocytes from small surgical samples of human ventricular myocardium. Human ventricular myocytes can be used for electrophysiological studies and drug testing.

Cardiomyocytes from diseased hearts are subjected to complex remodeling processes involving changes in cell structure, excitation contraction coupling and ...

Isolation of Cancer Stem Cells From Human Prostate Cancer Samples

The cancer stem cell (CSC) model has been considerably revisited over the last two decades. During this time CSCs have been identified and directly isolated from human tissues and serially propagated in immunodeficient mice, typically through antibody labeling of subpopulations of cells and fractionation by flow cytometry. However, the unique clinical features of prostate cancer have considerably limited the study of prostate CSCs from fresh human tumor samples. We recently reported the isolation of prostate CSCs directly from human tissues by virtue of their HLA class I (HLAI)-negative phenotype. Prostate cancer cells are harvested from surgical specimens and mechanically dissociated. A cell suspension is generated and labeled with fluorescently conjugated HLAI and stromal antibodies. Subpopulations of HLAI-negative cells are finally isolated using a flow cytometer. The principal limitation of this protocol is the frequently microscopic and multifocal nature of primary cancer in prostatectomy specimens. Nonetheless, isolated live prostate CSCs are suitable for molecular characterization and functional validation by transplantation in immunodeficient mice.

The isolation of cancer stem cells (CSCs) directly from human tissues is requisite for their biological characterization. This manuscript describes a methodology for the isolation of prostate CSCs from human tissues, while also providing tips on troubleshooting difficult steps.

The cancer stem cell (CSC) model has been considerably revisited over the last two decades. During this time CSCs have been identified and directly ...

Rapid Fractionation and Isolation of Whole Blood Components in Samples Obtained from a Community-based Setting

Collection and processing of whole blood samples in a non-clinical setting offers a unique opportunity to evaluate community-dwelling individuals both with and without preexisting conditions. Rapid processing of these samples is essential to avoid degradation of key cellular components. Included here are methods for simultaneous peripheral blood mononuclear cell (PBMC), DNA, RNA and serum isolation from a single blood draw performed in the homes of consenting participants across a metropolitan area, with processing initiated within 2 hr of collection. We have used these techniques to process over 1,600 blood specimens yielding consistent, high quality material, which has subsequently been used in successful DNA methylation, genotyping, gene expression and flow cytometry analyses. Some of the methods employed are standard; however, when combined in the described manner, they enable efficient processing of samples from participants of population- and/or community-based studies who would not normally be evaluated in a clinical setting. Therefore, this protocol has the potential to obtain samples (and subsequently data) that are more representative of the general population.

We outline a methodology for the processing of whole blood to obtain a variety of components for further analysis. We have optimized a streamlined protocol that enables rapid, high-throughput simultaneous processing of whole blood samples in a non-clinical setting.

Collection and processing of whole blood samples in a non-clinical setting offers a unique opportunity to evaluate community-dwelling individuals both ...

Isolation and Profiling of MicroRNA-containing Exosomes from Human Bile

Exosome research in the last three years has greatly extended the scope towards identification and characterization of biomarkers and their therapeutic uses.
Exosomes have recently been shown to contain microRNAs (miRs). MiRs themselves have arisen as valuable biomarkers for diagnostic purposes. As specimen collection in clinics and hospitals is quite variable, miRNA isolation from whole bile varies substantially. To achieve robust, accurate and reproducible miRNA profiles from collected bile samples in a simple manner required the development of a high-quality protocol to isolate and characterize exosomes from bile. The method requires several centrifugations and a filtration step with a final ultracentrifugation step to pellet the isolated exosomes. Electron microscopy, Western blots, flow cytometry and multi-parameter nanoparticle optical analysis, where available, are crucial characterization steps to validate the quality of the exosomes. For the isolation of miRNA from these exosomes, spiking the lysate with a non-specific, synthetic miRNA from a species like Caenorhabditis elegans, i.e., Cel-miR-39, is important for normalization of RNA extraction efficiency. The isolation of exosome from bile fluid following this method allows the successful miRNA profiling from bile samples stored for several years at -80 &#176;C.

Bile fluid is a valuable source of extracellular vesicles/exosomes that contain potentially important biomarkers. This protocol represents a robust method to isolate exosomes from human bile for further analyses including miRNA profiling.

Exosome research in the last three years has greatly extended the scope towards identification and characterization of biomarkers and their therapeutic ...

Simultaneous Flow Cytometric Characterization of Multiple Cell Types Retrieved from Mouse Brain/Spinal Cord Through Different Homogenization Methods

Recent advances in viral vector and nanomaterial sciences have opened the way for new cutting-edge approaches to investigate or manipulate the central nervous system (CNS). However, further optimization of these technologies would benefit from methods allowing rapid and streamline determination of the extent of CNS and cell-specific targeting upon administration of viral vectors or nanoparticles in the body. Here, we present a protocol that takes advantage of the high throughput and multiplexing capabilities of flow cytometry to allow a straightforward quantification of different cell subtypes isolated from mouse brain or spinal cord, namely microglia/macrophages, lymphocytes, astrocytes, oligodendrocytes, neurons and endothelial cells. We apply this approach to highlight critical differences between two tissue homogenization methods in terms of cell yield, viability and composition. This could instruct the user to choose the best method depending on the cell type(s) of interest and the specific application. This method is not suited for analysis of anatomical distribution, since the tissue is homogenized to generate a single-cell suspension. However, it allows to work with viable cells and it can be combined with cell-sorting, opening the way for several applications that could expand the repertoire of tools in the hands of the neuroscientist, ranging from establishment of primary cultures derived from pure cell populations, to gene-expression analyses and biochemical or functional assays on well-defined cell subtypes in the context of neurodegenerative diseases, upon pharmacological treatment or gene therapy.

We present a flow cytometry method to identify simultaneously different cell types retrieved from mouse brain or spinal cord. This method could be exploited to isolate or characterize pure cell populations in neurodegenerative diseases or to quantify the extent of cell targeting upon in vivo administration of viral vectors or nanoparticles.

Recent advances in viral vector and nanomaterial sciences have opened the way for new cutting-edge approaches to investigate or manipulate the central ...

Hemocompatibility Testing of Blood-Contacting Implants in a Flow Loop Model Mimicking Human Blood Flow

The growing use of medical devices (e.g., vascular grafts, stents, and cardiac catheters) for temporary or permanent purposes that remain in the body&#39;s circulatory system demands a reliable and multiparametric approach that evaluates the possible hematologic complications caused by these devices (i.e., activation and destruction of blood components). Comprehensive in vitro hemocompatibility testing of blood-contacting implants is the first step towards successful in vivo implementation. Therefore, extensive analysis according to the International Organization for Standardization 10993-4 (ISO 10993-4) is mandatory prior to clinical application. The presented flow loop describes a sensitive model to analyze the hemostatic performance of stents (in this case, neurovascular) and reveal adverse effects. The use of fresh human whole blood and gentle blood sampling are essential to avoid the preactivation of blood. The blood is perfused through a heparinized tubing containing the test specimen by using a peristaltic pump at a rate of 150 mL/min at 37 &#176;C for 60 min. Before and after perfusion, hematologic markers (i.e., blood cell count, hemoglobin, hematocrit, and plasmatic markers) indicating the activation of leukocytes (polymorphonuclear [PMN]-elastase), platelets (&#946;-thromboglobulin [&#946;-TG]), the coagulation system (thombin-antithrombin III [TAT]), and the complement cascade (SC5b-9) are analyzed. In conclusion, we present an essential and reliable model for extensive hemocompatibility testing of stents and other blood-contacting devices prior to clinical application.

This protocol describes a comprehensive hemocompatibility evaluation of blood-contacting devices using laser-cut neurovascular implants. A flow loop model with fresh, heparinized human blood is applied to mimic blood flow. After perfusion, various hematologic markers are analyzed and compared to the values gained directly after blood collection for hemocompatibility evaluation of the tested devices.