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

The overall goal of this procedure is to isolate cervical mononuclear cells from an endocervical scraping using a cyto brush. Once isolated, these cells can be used in multiple assays, including flow cytometry, where you can determine the phenotype and function of immune cells at the female genital tract. The collection of cyto brush and cervical lavage samples is a non-invasive procedure that must be performed by a gynecologist or trained physician.

The cervical vaginal lavage is performed first where the endocervix is washed with two mills of PBS, which is then collected from the posterior fornix following the lavage. The concave end of a cervical scraper is placed against the cervix and rotated to sample the cervical opening or cervical os. Next, the cyto brush is inserted into the Endocervical OS and rotated 360 degrees.

The use of gentle pressure will help to ensure no blood contamination of the sample. The cyto brush in the scraper are collected into a tube containing five mils of PBS. After being transported on ice, the sample is rapidly vortex to dissociate the cells from the cyto brush.

Once you've washed the samples with media, they're then filtered through a hundred micron nylon filter. This allows for the collection of the lymphocytes, monocytes, and epithelial cells from the sample. The cells are now ready to be stained with antibodies against surface proteins, and once they have been fixed, they can be run on the flow cytometer.

We will show you how we analyze the resulting data to assess the phenotype of both the CD four and CD eight T-cell populations present at the cervix. Hi, I'm a PhD student in Dr.Keith folk's lab. Our research focuses on the link between immune activation and HIV susceptibility as well as disease progression.

Because the majority of HIV infections now occur through heterosexual intercourse, we're also interested in determining the correlates of protection to HIV transmission to female genital tract. Unfortunately, the collection of samples from the genital tract can be very challenging as well as fairly invasive for study participants. The use of cyto brushes to collect CMCs or cervical mononuclear cells is a faster and easier method compared to other protocols such as biopsies.

In this video, we will show you how to isolate and prepare CMCs for use in flow cytometry or other downstream applications. After collection, the cyto brush and scraper are transported in five mils of cold PBS. It is important to transport the samples on ice until the time of processing, which should be no more than two hours after sample collection.

Any samples that are contaminated with traces of blood at this point in time should be excluded from further analysis to dissociate the cells from the cyto brush. Vortex the tube on high for 45 seconds because cells will still remain on the cyto brush and scraper. They must be manually removed.

Put on a fresh pair of gloves and use the cyto brush to scrape any remaining material off of the scraper. After discarding into a bleach solution, use your thumb and forefinger to remove all remaining cells from the cyto brush by squeezing up and down over the falcon tube. After discarding the cyto brush, change your gloves before preparing another sample, use a transfer pipette to wash the sides of the tube with PBS ensuring all sample is collected.

Fit a hundred micron nylon. Filter over a fresh Falcon tube and use the pipette to filter the entire volume of PBS solution. Add five mils of RPMI media lacking FBS to the original tube.

Use the pipette to thoroughly wash the sides of the tube in order to remove any adherent mucus or debris. Then transfer the media through the filter. Use some of the filtered media to wash the bottom of the filter before discarding into bleach.

To pellet the isolated cells. Centrifuge the samples for 10 minutes at 1500 r pm to prevent the pellet from dislodging. Remember to turn off the centrifuge brake if the cells will be used for flow cytometry.

Prepare the blocking solution required to prevent non-specific antibody binding. During the centrifugation step, use 100 microliters of blocking solution per sample. The blocking solution contains 93.2 microliters of fax, wash or PBS plus 2%FBS five microliters of FBS and 1.8 microliters of mouse IgG.

After the spin, most samples with viable cells will have a visible pellet. In some cases, the pellet will be quite large. While in other samples it may be barely visible.

Pellet size is not always related to cell yield. Carefully to cant the supernatant ensuring that you do not disturb the cells resuspend the pellet by gentle agitation. Wash the cells with five mils of PBS and repeat the centrifugation for 10 minutes At 1500.

RRP M with the break off, once again, decant the supernatant and resuspend the pellet as before. At this point, the cells are ready to be either cryopreserved or surface stained prior to staining With fluorescent antibodies, it is beneficial to block non-specific binding by incubating cells with a blocking solution prepared during an earlier centrifugation. After the cell pellet is resuspended, add a hundred microliters of blocking solution to the sample and mix thoroughly.

The remaining protocol steps can be performed in either a 96 well plate or fax tube. Depending on the size of the CMC pellet. Transfer the blocking solution and cell mixture to a fax tube.

Incubate the sample for 10 minutes at four degrees Celsius. During the 10 minute incubation, prepare the viability dye that will be used to discriminate live and dead cells. During data analysis, prepare the viability dye according to manufacturer's and pre titrate as required.

Be sure to keep the dye mixture covered from light until it is ready to be used. After the 10 minute incubation, wash the cells with 500 microliters of fax, wash or PBS with 2%FBS, and then centrifuge for 10 minutes at 1600 RPM. After spinning, decant the supernatant from the fax tube.

Re suspend the cells by agitation. Now add the appropriate volume of the viability dye that was previously prepared. Incubate cells for 30 minutes at four degrees Celsius ensure that samples remain covered from the light by wrapping in tinfoil.

During this incubation, you can prepare the master mix of fluorescent antibodies that will be used to label the cells. Be sure to pre titrate and validate your antibodies on peripheral blood control samples prior to using them on CMCs. Mix the titrated antibodies and bring to a final volume of a hundred microliters per sample.

With fax, wash, protect all antibodies from the light during these steps. After the 30 minute incubation, once again, wash the sample with 500 microliters of fax, wash, centrifuge, decant, and resuspend the cells as before. It is now time to incubate the cells with 100 microliters of the antibody cocktail prepared during the previous incubation.

Once again, incubate the samples for 30 minutes at four degrees Celsius, ensuring that they're protected from the light. The incubating CMCs are now ready to be washed with 500 microliters of PBS for the final time. Centrifuge the sample at 1600 RPM for 10 minutes.

After decanting and resus suspending the pellet dilute the cells in a final volume of 750 microliters in a 1%paraldehyde solution. To fix them, it is important to resuspend the cells in this large volume in order to prevent clogging of the flow cytometer. Even at this volume, the sample tube may appear quite cloudy.

Keep the cells in the dark at four degrees Celsius until they're acquired on the flow cytometer. In addition to the cyto brush sample, we routinely collect two mils of PBS lavage from the cervix and vagina. This sample is centrifuge at 1400 RPM for seven minutes to remove cellular debris from the lavage.

The remaining supernatant can then be allotted and frozen at minus 80 degrees Celsius for the future analysis of soluble protein concentration. This data can be particularly useful in combination with cellular phenotype data obtained by flow cytometry. Before running your CMC sample.

It is helpful to prepare PBMC controls to assist with gating run compensation tubes, using beads or single stain cells in order to adjust for spectral overlap. Acquire the entire CMC tube to collect all events compared to PBMC samples. CMCs will exhibit more doublets on an FSC area versus height plot and may not exhibit as distinctive a lymphocyte population as pbmc.

After acquiring your samples, export the files for analysis in a program such as FlowJo. Here we will show the major components of a typical gating strategy for CMC derived CD four positive and CD eight positive T cells. Use an FSC area versus height plot to G on the singlets in your sample.

In this matched comparison, you can clearly see the increased number of doublets in a CMC sample compared to p BMCs. Next, plotting a fluorescent marker or FSC versus time serves as a quality control step. If any changes in flow rate introduced artifacts into the data, they can be removed at this step.

At the top, you can see a consistent fluorescence over time profile indicative of a high quality sample. Below you can see a sample with multiple interruptions and flow rate, which can be excluded from further analysis. Identification of the lymphocyte population in CMC samples can be challenging.

Using A-P-B-M-C control will assist with gait placement. As you can see, some samples exhibit large clear populations while other samples lack lymphocyte populations. Altogether, these samples should be excluded.

Although we will focus on lymphocyte populations. It is also possible to g on monocytes by gating on cells that lack the expression of lineage markers. It's possible to identify cells that express combinations of dendritic cell and monocyte markers such as CD 16 and CD 11 C.The inclusion of a viability marker or dye is crucial for CMC analysis even on fresh cells.

As you can see, the majority of CMC lymphocytes are viable, but some samples contain high frequencies of dead cells that must be excluded from analysis. Dead cells may non-specifically take up antibody and confound data analysis within the lymphocyte gait. CD three staining should be robust.

The proportion of CD three positive T cells is lower and more variable than in PBMC samples, which is why it is crucial to acquire the entire CMC sample and maximize the number of events collected. The CD four positive and CD eight positive T-cell populations can be identified just as in TBMC samples. In some CMC samples.

The CD four to CD eight ratio is lower than observed in peripheral blood and some individuals exhibit a large double negative T-cell population. After identifying the CD four positive and CD eight positive T cell populations, it is possible to quantify the expression of numerous phenotypic markers. Although CD 1 61 expression can be detected in CMCs, the CD 1 61 bright population that is evident in blood is absent at the genital tract.

Similarly, expression of activation markers such as CD 69 and H-L-A-D-R, as well as markers like CCR five and PD one can be reliably measured among CMC samples and quantified. The use of cyto brush samples allows us to characterize both the phenotype and function of mucosal immune cells, especially in the context of infectious diseases. Not only does this give us greater insight into the mechanisms regulating mucosal and reproductive immunology, and also provides the basis for the development of novel measures of mucosal vaccine efficacy.