S.C.J. was supported by an International Society for Heart and Lung Transplantation Research Fellowship and a Royal College of Physicians and Surgeons of Canada Detweiler Traveling Fellowship.S.M. was supported in part by an International Society for Heart and Lung Transplantation Career Development Award (to S.C.J.). S.S. was supported by the National Institutes of Health Research Oxford Biomedical Research Centre.J.H. is the recipient of a Kidney Research U.K. Senior Non-Clinical Fellowship. This work was funded by the following grants to A.B. and K.W.: a Wellcome Trust Programme Grant (082519Z07Z), a British Heart Foundation Programme Grant (PG/10/62.28504), and the EU Framework Program 7 (ONE Study; BioDRIM). The authors wish to thank Michael Parsons and the Flow Cytometry Core Facility at the Lunenfeld-Tanenbaum Research Institute, Sinai Health System, Toronto for providing access to and support with the ImageStream Mark X instrument.

The authors have nothing to disclose.

Imaging flow cytometry has been used to demonstrate immune synapse formation between monoclonal T cells and APCs or in the presence of superantigens24,25,26,27,28. This method takes advantage of the fact that after a productive T cell-APC contact, the T cell rearranges its actin cytoskeleton, polarizing it toward the site of contact21. This rearrangement does not occur without TCR signaling, and it is therefore an early correlate of T-cell activation19,20,21. The method presented here adapts this approach to the measurement of alloreactive T-cell frequency in polyclonal T-cell populations. As such, it may in the future serve as the basis for the development of assays for donor reactivity in clinical transplantation.
Although direct comparisons have not yet been made, the detection of alloreactive immune synapses appears to have superior predictive power than the conventional MLR. For example, previous work has shown that, in the tolerizing protocol described above, the results of an MLR fail to reliably correlate with graft outcome2.
A number of assays have been developed for the operationally tolerant state in humans9,10,11, although these do not measure effector cell function in response to alloantigen. In contrast, IFN&#947; ELISPOT assays8 measure effector T-cell function but cannot capture the full spectrum of cytokine secretion that may be relevant to acute and chronic allograft rejection, such as IL-1738,39. The limiting dilution assay4, which is labor intensive, and the trans-vivo assay6, which requires mice, have significant practical limitations that would hinder their application in a clinical setting. Recent improvements on the analysis of proliferating cells using TCR sequence analysis of T cells responding in the MLR may be of value, but like the assay presented here, will require further validation in clinical studies18,40.
Further development of the immune synapse detection assay will require that a number of important questions be answered. First, the assay as developed only measures direct alloreactivity. The direct pathway involves the presentation of allogeneic MHC/peptide complexes on donor-derived APCs. The latter are generally eliminated quickly after transplantation, and further alloantigen presentation is carried out by recipient APCs presenting intact donor MHC (semi-direct pathway) or processed donor antigens on self MHC (indirect pathway). The indirect pathway is an important driver of chronic allograft rejection33,41.
In principle, it should be possible to detect indirect immune synapses using this assay, but indirectly alloreactive T cells have a much lower frequency than direct ones42,43, meaning that the analysis of a larger number of events will be required. A second consideration is that we have only tested this assay using CD4+ T cells, whereas CD8+ T cells are also an important component of the anti-donor response. Again, it should be possible to detect CD8+ T cell-APC synapses using this method. Another limitation is that the method requires the manual review and analysis of cell images in the final membrane contact gate, and we are currently working on the automation of this step.
Finally, the method requires testing and development in human subjects, and preliminary studies with human samples are currently being performed. Further phenotypic T-cell subset analysis (i.e., effector, memory, regulatory, etc.) in combination with the detection of immune synapses in transplant recipients would represent a powerful approach for characterizing the alloreactive T-cell repertoire and will be an important focus for future work.

Solid organ transplantation has transformed the care of patients with end-stage diseases of the kidney, liver, heart, and lungs. Owing to disparities in major and minor histocompatibility antigens, however, allografts are promptly rejected by recipient T cells if immunosuppressive drugs are not used. These agents have numerous adverse effects, including risks for cancer and organ dysfunction. A major clinical goal is therefore to lower the dose of immunosuppression to the minimum level required to prevent allograft rejection. This level is likely to vary depending on the degree of activation of the innate immune system; the degree of donor-recipient alloantigen mismatch; and inter-patient differences in immune function, pharmacokinetics, and pharmacodynamics.
Unfortunately, transplant clinicians do not have any tools for accurately assessing donor reactivity in individual patients1. The mixed leukocyte reaction (MLR) can detect donor reactivity, but it fails to reliably predict graft outcome2,3. Limiting dilution assays, cytokine ELISPOTs, and the trans-vivo assay either measure a limited range of responses or are not practical4,5,6,7,8.Gene expression profiles have revealed signatures related to operational tolerance9,10,11,12 and rejection13,14,15, but these are not always generalizable across populations16 and may ultimately have limited usefulness in individual patients. Sequence-based analyses of the T-cell receptor (TCR) of T cells in the peripheral blood17 or proliferating in the MLR18 have also been developed but require further validation.
Conceptually, it would be desirable to have an assay that detects the earliest requisite steps in recipient T-cell activation by a donor antigen. Since culturing cells over days (as in the MLR) can introduce artefacts, such a test would ideally not require the measurement of downstream events, such as proliferation or effector function. Equally, however, it would also be desirable for the test to depend on some element of T-cell function, since purely descriptive assessments (e.g., TCR sequencing) may be unable to distinguish between anergic and functional T cells.
Numerous studies have indicated that prolonged T-APC contact is required for the formation of an immune synapse, which is an essential first step in the T-cell response19,20,21,22. We recently reported that, during dynamic in vitro time lapse imaging, about 5 - 10% of mouse CD4+ T cells form long-lasting contacts with allogeneic bone marrow-derived dendritic cells (BMDCs)23. The frequency of prolonged contact was increased in animals that rejected a graft, whereas in mice previously rendered tolerant to the same antigens, it remained at levels seen in untransplanted mice23. Prolonged interactions were reduced in the presence of recipient Tregs and increased in their absence, and we observed similar phenomena using human T cells and allogeneic monocyte-derived DCs (MoDCs)23.
However, the enumeration of prolonged contacts made within a polyclonal T-cell population is time-consuming and labor-intensive. We therefore made use of imaging flow cytometry to examine allogeneic immune synapse formation. Imaging flow cytometry incorporates the multiparametric data acquisition and analysis capabilities of conventional flow cytometry with the single-cell imaging abilities of fluorescence microscopy. This technique has been used by other investigators to study immune synapse formation by monoclonal T cells24,26,27or in the presence of superantigens28. In such settings, however, the frequency of responding T cells ranges from 30-100%, whereas alloreactive T cells are generally estimated to represent 5-15% of the total T-cell repertoire29,30,31,32. Importantly, we showed that imaging flow cytometry can produce a very comparable measure of alloreactive T-cell frequency23 and that changes in synapse frequency within a polyclonal T-cell population are predictive of graft outcome23. Currently, this approach has been optimized to measure the direct alloreactivity of CD4+ T cells, but, in principle, it could also be developed to examine CD8+ T cells and the indirect pathway. Indirect alloreactivity is believed to become increasingly relevant at longer times post-transplant33. We are currently developing this method to use human cells, which will allow for testing in patients. Thus, in the future, the overall approach may be useful for the functional evaluation of T-cell responses in transplant recipients before transplant; immediately after transplant; and in the long-term, when drug minimization becomes an important goal.

<table border="0" cellpadding="0" cellspacing="0" width="706">
	<tbody>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Phosphate-buffered saline</td>
			<td align="left" style="width:133px;">Various</td>
			<td align="left" style="width:119px;">Varies</td>
			<td style="width:167px;"></td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">Ethylenediamenetetraacetic acid, 0.5M solution</td>
			<td align="left" style="width:133px;">Thermo Fisher Scientific</td>
			<td align="left" style="width:119px;">AM9260G</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Triton X-100 nonionic detergent</td>
			<td align="left" style="width:133px;">Sigma-Aldrich</td>
			<td align="left" style="width:119px;">X100</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Beta-mercaptoethanol</td>
			<td align="left" style="width:133px;">Sigma-Aldrich</td>
			<td align="left" style="width:119px;">M3148</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Dimethyl sulfoxide</td>
			<td align="left" style="width:133px;">Sigma-Aldrich</td>
			<td align="left" style="width:119px;">D8418</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Formaldehyde</td>
			<td align="left" style="width:133px;">Sigma-Aldrich</td>
			<td align="left" style="width:119px;">F1635</td>
			<td align="left">Solution is 37% formaldehyde and so must be diluted 25 times for 1.5% solution</td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Cell strainers, 70 &#956;m pore size</td>
			<td align="left" style="width:133px;">Fisher Scientific</td>
			<td align="left" style="width:119px;">08-771-2</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Phalloidin-fluorescein isothiocyanate</td>
			<td align="left" style="width:133px;">Sigma-Aldrich</td>
			<td align="left" style="width:119px;">P5282</td>
			<td></td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">7-aminoactinomycin D</td>
			<td align="left" style="width:133px;">Thermo Fisher Scientific</td>
			<td align="left" style="width:119px;">A1310</td>
			<td align="left">Reconstitute in DMSO</td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">Allophycocyanin-conjugated anti-mouse CD90.2</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">17-0902</td>
			<td></td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">Pacific blue-conjugated anti-mouse CD11b</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">48-0112</td>
			<td align="left">Pacific blue has been replaced by eFluor 450</td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Biotinylated anti-mouse CD3</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">13-0032</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Biotinylated anti-mouse MHC class II</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">13-5321</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Biotinylated anti-mouse B220</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">13-0452</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Biotinylated anti-mouse CD8</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">13-0081</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Biotinylated anti-mouse CD19</td>
			<td align="left" style="width:133px;">eBioscience</td>
			<td align="left" style="width:119px;">13-0193</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Anti-biotin microbeads</td>
			<td align="left" style="width:133px;">Miltenyi Biotec</td>
			<td align="left" style="width:119px;">130-090-485</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">LS columns</td>
			<td align="left" style="width:133px;">Miltenyi Biotec</td>
			<td align="left" style="width:119px;">130-042-401</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">MidiMACS magnetic cell separator</td>
			<td align="left" style="width:133px;">MIltenyi Biotec</td>
			<td align="left" style="width:119px;">130-042-302</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">recombinant mouse GM-CSF</td>
			<td align="left" style="width:133px;">Peprotech</td>
			<td align="left" style="width:119px;">315-03</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">recombinant human TGF&#946;1</td>
			<td align="left" style="width:133px;">Peprotech</td>
			<td align="left" style="width:119px;">100-21</td>
			<td align="left">Human TGF&#946;1 has activity on mouse cells</td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">Amnis ImageStream X Mark II</td>
			<td align="left" style="width:133px;">Amnis/EMD Millipore</td>
			<td align="left" style="width:119px;">N/A</td>
			<td align="left">Imaging flow cytometer; details available at http://www.emdmillipore.com/</td>
		</tr>
		<tr height="42">
			<td align="left" height="42" style="height:42px;width:287px;">IDEAS Software</td>
			<td align="left" style="width:133px;">Amnis/EMD Millipore</td>
			<td align="left" style="width:119px;">N/A</td>
			<td align="left">Free download (registration required): https://www.amnis.com/index.php/page/Display/login%20%20</td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Cell culture medium&#160;</td>
			<td align="left" style="width:133px;">Various</td>
			<td align="left" style="width:119px;">Varies</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Fetal bovine serum</td>
			<td align="left" style="width:133px;">Various</td>
			<td align="left" style="width:119px;">Varies</td>
			<td></td>
		</tr>
		<tr height="21">
			<td align="left" height="21" style="height:21px;width:287px;">Cell culture plates</td>
			<td align="left" style="width:133px;">Various</td>
			<td align="left" style="width:119px;">Varies</td>
			<td></td>
		</tr>
	</tbody>
</table>

measurement of t cell alloreactivity using imaging flow cytometry

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal of immunosuppression. The mixed leukocyte reaction (MLR), limiting dilution assays, and trans-vivo delayed-type hypersensitivity (DTH) assay have all been applied to this question, but these methods have limited predictive ability and/or significant practical limitations that reduce their usefulness. Imaging flow cytometry is a technique that combines the multiparametric quantitative powers of flow cytometry with the imaging capabilities of fluorescent microscopy. We recently made use of an imaging flow cytometry approach to define the proportion of recipient T cells capable of forming mature immune synapses with donor antigen-presenting cells (APCs). Using a well-characterized mouse heart transplant model, we have shown that the frequency of in vitro immune synapses among T-APC membrane contact events strongly predicted allograft outcome in rejection, tolerance, and a situation where transplant survival depends on induced regulatory T cells. The frequency of T-APC contacts increased with T cells from mice during acute rejection and decreased with T cells from mice rendered unresponsive to alloantigen. The addition of regulatory T cells to the in vitro system reduced prolonged T-APC contacts. Critically, this effect was also seen with human polyclonally expanded, naturally occurring regulatory T cells, which are known to control the rejection of human tissues in humanized mouse models. Further development of this approach may allow for a deeper characterization of the alloreactive T-cell compartment in transplant recipients. In the future, further development and evaluation of this method using human cells may form the basis for assays used to select patients for immunosuppression minimization, and it can be used to measure the impact of tolerogenic therapies in the clinic.

This method was used to investigate CD4+ T-cell alloreactivity in mice rendered tolerant to donor alloantigens before heterotopic cardiac allograft transplantation. CBA mice (H-2k) were given a tolerizing protocol consisting of a donor-specific (B6, H-2b) blood transfusion combined with a non-depleting CD4 antibody one month prior to receiving a B6 cardiac transplant. This protocol results in long-term allograft survival that is dependent on Foxp3+ regulatory T cells36,37. Seven days post-transplant, splenic CD4+ T cells were obtained from tolerized and non-tolerized recipients of B6 cardiac allografts and were co-incubated with B6 bone marrow-derived DCs according to this protocol. Figure 2 shows representative data from this experiment. The membrane contact gate is shown in Figure 2A, with green crosshairs placed on a synaptic event (left panel, 1) and on a non-synaptic event (right panel). Figure 2B shows the brightfield and fluorescence channels for this event. To reduce bias, data were analyzed by an observer blinded to treatment assignment23. As shown in several examples in Figure 3, both from non-tolerized (Figure 3A-B) and tolerized (Figure 3C-D) CBA recipients of B6 hearts, synapses are easily distinguished from non-synaptic contacts by the presence of a dense FITC-positive ridge at the T-APC interface. These results show that visual detection of immune synapses made by recipient T cells tracks with the degree of alloreactivity in the recipient.
<img alt="Figure 2" src="/files/ftp_upload/55283/55283fig2.jpg" /> 
Figure 2. Identification of T-APC Doublets with Membrane Contact and Immune Synapse Formation. Events in the final doublet gate (Figure 1F) are analyzed. A. T-cell marker fluorescence in the APC object mask is plotted against APC marker fluorescence in the DC object mask. Some doublet events have an APC and a T cell without cell-cell contact and appear in the lower left corner of the plot (images not shown). A membrane contact gate can thus be drawn that includes only doublets in which T cells and APCs are in contact. Images of each event in this gate are reviewed for evidence of actin cytoskeletal rearrangement in the phalloidin-FITC channel and can be tagged using the analysis software. The left panel indicates an immune synapse event (labeled 1 and indicated by green crosshairs), whereas the right panel indicates a membrane contact event without immune synapse formation (labeled 2 and indicated by green crosshairs). The determination of synapse formation requires manual review of these images, shown in B. B. The top row shows brightfield and fluorescence channel images for a doublet with an immune synapse (corresponds to event 1 in A); the bottom row shows a doublet with membrane contact but lacking synapse formation (corresponds to event 2 in A). The data were analyzed in a blinded fashion with respect to treatment assignment and are from a previously published experiment23. <a href="https://www.jove.com/files/ftp_upload/55283/55283fig2large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 3" src="/files/ftp_upload/55283/55283fig3.jpg" /> 
Figure 3. Examples of T-APC Synapse Formation. CBA mice received cardiac allografts from B6 donors after either no pre-treatment (A-B) or after tolerance induction with B6 whole blood under the cover of a non-depleting anti-CD4 antibody (C-D). After 7 days, splenic CD4+ T cells were tested for synapse formation with B6 DCs. A. Three examples of non-synaptic doublets with membrane contact from a non-tolerized animal. B. Three examples of immune synapses from a non-tolerized animal. C. Three examples of non-synaptic doublets with membrane contact from a tolerized animal. D. Three examples of immune synapses from a tolerized animal. Synapse formation is indicated by the presence of a bright, FITC-positive ridge at the T-APC interface (Ch03). The data were analyzed in a blinded fashion with respect to treatment assignment and are from a previously published experiment23. <a href="https://www.jove.com/files/ftp_upload/55283/55283fig3large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<table border="1" fo:keep-together.within-page="1" fo:keep-with-next.within-page="always">
	<tbody>
		<tr>
			<td>Antibody/Dye</td>
			<td>Fluorochrome</td>
			<td>Channel</td>
			<td>Concentration</td>
		</tr>
		<tr>
			<td>CD11c</td>
			<td>eF450</td>
			<td>Ch02</td>
			<td>5 &#181;g/mL, titrate empirically</td>
		</tr>
		<tr>
			<td>CD90.2</td>
			<td>APC</td>
			<td>Ch06</td>
			<td>5 &#181;g/mL, titrate empirically</td>
		</tr>
		<tr>
			<td>Phalloidin</td>
			<td>FITC</td>
			<td>Ch03</td>
			<td>0.05 - 0.5 &#181;g/mL</td>
		</tr>
		<tr>
			<td>7-AAD</td>
			<td>-</td>
			<td>Ch05</td>
			<td>25 &#181;g/mL</td>
		</tr>
	</tbody>
</table>
Table 1. Antibodies and Dyes Used in this Study. Fluorochrome-conjugated antibodies, dyes, suppliers, and recommended concentrations are presented in the table. The imaging flow cytometer channel that was used to detect each fluorochrome is also shown in the table.

Watch this Scientific Journal Video about Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry at JoVE.com

Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

This paper describes a method for measuring alloreactivity in a mixed population of T cells using imaging flow cytometry.

The measurement of immunological reactivity to donor antigens in transplant recipients is likely to be crucial for the successful reduction or withdrawal ...

measurement-of-t-cell-alloreactivity-using-imaging-flow-cytometry

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Immunology and Infection

13.4K Views.  University of Toronto and University Health Network. This paper describes a method for measuring alloreactivity in a mixed population of T cells using imaging flow cytometry.

Video: Measurement of T Cell Alloreactivity Using Imaging Flow Cytometry

Flow Cytometry Analysis of Immune Cells Within Murine Aortas

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam cells, immune cells, vascular endothelial and smooth muscle cells, platelets, extracellular matrix, and a lipid-rich core with extensive necrosis and fibrosis of surrounding tissues.1 The innate and adaptive arms of the immune response are involved in the initiation, development and persistence of atherosclerosis.2, 3 There is a significant body of evidence that different subsets of the immune cells, such as macrophages, dendritic cells, T and B lymphocytes, are present within the aortas of healthy and atherosclerosis-prone mice4. Additionally, immune cells are found in the surrounding aortic adventitia which suggests an important role of this tissue in atherogenesis.2
For some time, the quantitative detection of different types of immune cells, their activation status, and the cellular composition within the aortic wall was limited by RT-PCR and immunohistochemical methods for the study of atherosclerosis. Few attempts were made to perform flow cytometry using human aortas, and a number of problems, such as a high autofluorescence, have been reported5,6. Human atherosclerotic plaques were digested with collagenase 1, and free cells were collected and stained for CD14+/CD11c+ to highlight macrophage-derived foam cells. In this study, a "mock" channel was used to avoid false-positive staining.6 Necrotic materials accumulating during the digestion process give rise in a large amount of debris that generates a high autofluorescence in aortic samples. To resolve this problem, a panel of negative and positive controls has been proposed, but only double staining could be applied in these samples. We have developed a new flow cytometry-based method7 to analyze the immune cell composition and characterize the activation, proliferation, differentiation of immune cells in healthy and atherosclerosis-prone aorta. This method allows the investigation of the immune cell composition of the aortic wall and opens possibilities to use a broad spectrum of immunological methods for investigations of immune aspects of this disease.

This paper presents a flow cytometry-based method to investigate the immune composition of aortas. The paper also illustrates an additional technique that allows examining surrounding adventitia and vessel wall separately. This method opens possibilities to perform phenotypical analyses of aortic leukocytes and apply several immunological assays for atherosclerosis studies.

Atherosclerosis is a chronic inflammatory process of medium and large size vessels that is characterized by the formation of plaques consisting of foam ...

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly constituted by proteins and exopolysaccharides, among other factors 7-10. The microbial community encased within the biofilm often shows the differentiation of distinct subpopulation of specialized cells 11-17. These subpopulations coexist and often show spatial and temporal organization within the biofilm 18-21.
Biofilm formation in the model organism Bacillus subtilis requires the differentiation of distinct subpopulations of specialized cells. Among them, the subpopulation of matrix producers, responsible to produce and secrete the extracellular matrix of the biofilm is essential for biofilm formation 11,19. Hence, differentiation of matrix producers is a hallmark of biofilm formation in B. subtilis.
We have used fluorescent reporters to visualize and quantify the subpopulation of matrix producers in biofilms of B. subtilis 15,19,22-24. Concretely, we have observed that the subpopulation of matrix producers differentiates in response to the presence of self-produced extracellular signal surfactin 25. Interestingly, surfactin is produced by a subpopulation of specialized cells different from the subpopulation of matrix producers 15.
We have detailed in this report the technical approach necessary to visualize and quantify the subpopulation of matrix producers and surfactin producers within the biofilms of B. subtilis. To do this, fluorescent reporters of genes required for matrix production and surfactin production are inserted into the chromosome of B. subtilis. Reporters are expressed only in a subpopulation of specialized cells. Then, the subpopulations can be monitored using fluorescence microscopy and flow cytometry (See Fig 1).
The fact that different subpopulations of specialized cells coexist within multicellular communities of bacteria gives us a different perspective about the regulation of gene expression in prokaryotes. This protocol addresses this phenomenon experimentally and it can be easily adapted to any other working model, to elucidate the molecular mechanisms underlying phenotypic heterogeneity within a microbial community.

Single-cell Analysis of Bacillus subtilis Biofilms Using Fluorescence Microscopy and Flow Cytometry

Microbial biofilms are generally constituted by distinct subpopulations of specialized cells. Single-cell analysis of these subpopulations requires the use of fluorescent reporters. Here we describe a protocol to visualize and monitor several subpopulationswithin B. subtilis biofilms using fluorescence microscopy and flow cytometry.

Biofilm formation is a general attribute to almost all bacteria 1-6. When bacteria form biofilms, cells are encased in extracellular matrix that is mostly ...

Cell-based Flow Cytometry Assay to Measure Cytotoxic Activity

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells to kill CD4+ T cells loaded with their cognate peptide. Target CD4+ T cells are divided into two populations, labeled with two different concentrations of CFSE. One population is pulsed with the peptide of interest (CFSE-low) while the other remains un-pulsed (CFSE-high). Pulsed and un-pulsed CD4+ T cells are mixed at an equal ratio and incubated with an increasing number of purified CD8+ T cells. The specific killing of autologous target CD4+ T cells is analyzed by flow cytometry after coculture with CD8+ T cells containing the antigen-specific effector CD8+ T cells detected by peptide/MHCI tetramer staining. The specific lysis of target CD4+ T cells measured at different effector versus target ratios, allows for the calculation of lytic units, LU30/106 cells. This simple and straightforward assay allows for the accurate measurement of the intrinsic capacity of CD8+ T cells to kill target CD4+ T cells.

This protocol describes a sensitive, cell-based cytotoxicity assay. By enumerating the decrease in frequency of live target CD4+ T cells in the presence of an increasing number of effector CD8+ T cells, this assay allows for the direct assessment of cytolytic activity of antigen-specific CD8+ T cells.

Cytolytic activity of CD8+ T cells is rarely evaluated. We describe here a new cell-based assay to measure the capacity of antigen-specific CD8+ T cells ...

Fluorescence-activated Cell Sorting for Purification of Plasmacytoid Dendritic Cells from the Mouse Bone Marrow

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method enables researchers to better understand the characteristics of a single cell population without the influence of other cells. Compared to other methods of cell enrichment, such as magnetic-activated cell sorting (MCS), FACS is more flexible and accurate for cell separation due to the ability of phenotype detection by flow cytometry. In addition, FACS is usually capable of separating multiple cell populations simultaneously, which improves the efficiency and diversity of experiments. Although FACS has some limitations, it has been broadly used to purify cells for functional studies in both in vitro and in vivo settings. Here we report a protocol using fluorescence-activated cell sorting to isolate a very rare population of immune cells, plasmacytoid dendritic cells (pDC), with high purity from the bone marrow of lupus-prone mice for in vitro functional studies of pDC.

We report a protocol using fluorescence-activated cell sorting to isolate plasmacytoid dendritic cells (pDC) with high purity from the bone marrow of lupus-prone mice for functional studies of pDC.

Fluorescence-activated cell sorting (FACS) is a technique to purify specific cell populations based on phenotypes detected by flow cytometry. This method ...

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, ...

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and their important role in lung development and function, as well as their lung-localized responses to infection and inflammation. To date, no unified method for identification, isolation, and handling of alveolar macrophages from humans and mice exists. Such a method is needed for studies on these important innate immune cells in various experimental settings. The method described here, which can be easily adopted by any laboratory, is a simplified approach to harvesting alveolar macrophages from bronchoalveolar lavage fluid or from lung tissue and maintaining them in vitro. Because alveolar macrophages primarily occur as adherent cells in the alveoli, the focus of this method is on dislodging them prior to harvest and identification. The lung is a highly vascularized organ, and various cell types of myeloid and lymphoid origin inhabit, interact, and are influenced by the lung microenvironment. By using the set of surface markers described here, researchers can easily and unambiguously distinguish alveolar macrophages from other leukocytes, and purify them for downstream applications. The culture method developed herein supports both human and mouse alveolar macrophages for in vitro growth, and is compatible with cellular and molecular studies.

Isolation and In Vitro Culture of Murine and Human Alveolar Macrophages

This communication describes methodologies for isolation and culture of alveolar macrophages from humans and murine models for experimental purposes.

Alveolar macrophages are terminally differentiated, lung-resident macrophages of prenatal origin. Alveolar macrophages are unique in their long life and ...

Multicolor Flow Cytometry-based Quantification of Mitochondria and Lysosomes in T Cells

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular components responsible for supplying cell energy; however, excess mitochondria also produce reactive oxygen species (ROS) that could cause cell death. Therefore, the number of mitochondria must constantly be adjusted to fit the needs of the cells. This dynamic regulation is achieved in part through the function of lysosomes that remove surplus/damaged organelles and macromolecules. Hence, cellular mitochondrial and lysosomal contents are key indicators to evaluate the metabolic adjustment of cells. With the development of probes for organelles, well-characterized lysosome or mitochondria-specific dyes have become available in various formats to label cellular lysosomes and mitochondria. Multicolor flow cytometry is a common tool to profile cell phenotypes, and has the capability to be integrated with other assays. Here, we present a detailed protocol of how to combine organelle-specific dyes with surface markers staining to measure the amount of lysosomes and mitochondria in different T cell populations on a flow cytometer.

This article illustrates a powerful method to quantify mitochondria or lysosomes in living cells. The combination of lysosome- or mitochondria-specific dyes with fluorescently conjugated antibodies against surface markers allows the quantification of these organelles in mixed cell populations, like primary cells harvested from tissue samples, by using multicolor flow cytometry.

T cells utilize different metabolic programs to match their functional needs during differentiation and proliferation. Mitochondria are crucial cellular ...

Determination of Regulatory T Cell Subsets in Murine Thymus, Pancreatic Draining Lymph Node and Spleen Using Flow Cytometry

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, thymic derived Treg (tTreg) cells and peripherally induced Treg (pTreg) cells. They are present in different organs of our body and can be distinguished by specific markers, such as Helios and Neuropilin 1. It has been reported that tTreg cells are functionally more suppressive than pTreg cells. Therefore, it is important to determine the proportion of both tTreg and pTreg cells when investigating heterogeneous cell populations. Herein, we collected thymic glands, pancreatic draining lymph nodes and spleens from normoglycemic non-obese diabetic mice to distinguish tTreg cells from pTreg cells using flow cytometry. We manually prepared single cell suspensions from these organs. Fluorochrome conjugated surface CD4, CD8, CD25, and Neuropilin 1 antibodies were used to stain the cells. They were kept in the fridge overnight. On the next day, the cells were stained with fluorochrome conjugated intracellular Foxp3 and Helios antibodies. These markers were used to characterize the two subsets of Treg cells. This protocol demonstrates a simple but practical way to prepare single cells from murine thymus, pancreatic draining lymph node and spleen and use them for subsequent flow cytometric analysis.

Herein, we present a protocol to prepare single cells from murine thymus, pancreatic draining lymph node and spleen to further study these cells using flow cytometry. In addition, this protocol was used for determining the subsets of regulatory T cells using flow cytometry.

Our immune system consists of a number and variety of immune cells including regulatory T cells (Treg) cells. Treg cells can be divided into two subsets, ...

Preparation of Whole Bone Marrow for Mass Cytometry Analysis of Neutrophil-lineage Cells

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a myeloid-biased 39-antibody CyTOF panel to evaluate the hematopoietic system with a focus on the neutrophil-lineage cells by using this protocol. The CyTOF result was analyzed with an open-resource dimensional reduction algorithm, viSNE, and the data was presented to demonstrate the outcome of this protocol. We have discovered new neutrophil-lineage cell populations based on this protocol. This protocol of fresh whole BM preparation may be used for 1), CyTOF analysis to discover unidentified cell populations from whole BM, 2), investigating whole BM defects for patients with blood disorders such as leukemia, 3), assisting optimization of fluorescence-activated flow cytometry protocols that utilize fresh whole BM.

Here, we present a protocol to process fresh bone marrow (BM) isolated from mouse or human for high-dimensional mass cytometry (Cytometry by Time-Of-Flight, CyTOF) analysis of neutrophil-lineage cells.

In this article, we present a protocol that is optimized to preserve neutrophil-lineage cells in fresh BM for whole BM CyTOF analysis. We utilized a ...

Measuring Erythrocyte Complement Receptor 1 Using Flow Cytometry

CR1 (CD35, Complement Receptor type 1 for C3b/C4b) is a high molecular weight membrane glycoprotein of about 200 kDa that controls complement activation, transports immune complexes, and participates in humoral and cellular immune responses. CR1 is present on the surface of many cell types, including erythrocytes, and exhibits polymorphisms in length, structure (Knops, or KN, blood group), and density. The average density of CR1 per erythrocyte (CR1/E) is 500 molecules per erythrocyte. This density varies from one individual to another (100&#8211;1,200 CR1/E) and from one erythrocyte to another in the same individual. We present here a robust flow cytometry method to measure the density of CR1/E, including in subjects expressing a low density, with the help of an amplifying immunostaining system. This method has enabled us to show the lowering of CR1 erythrocyte expression in diseases such as Alzheimer&#39;s disease (AD), systemic lupus erythematosus (SLE), AIDS, or malaria.

The aim of this method is to determine the CR1 density in the erythrocytes of any subject by comparing with three subjects whose erythrocyte CR1 density is known. The method uses flow cytometry after immunostaining of the subjects&#39; erythrocytes by an anti-CR1 monoclonal antibody coupled to an amplified system using phycoerythrin (PE).

CR1 (CD35, Complement Receptor type 1 for C3b/C4b) is a high molecular weight membrane glycoprotein of about 200 kDa that controls complement activation, ...