Advances in imaging have led to the development of powerful multispectral, quantitative imaging techniques, like histo-cytometry. The utility of this approach is limited, however, by the need for time consuming manual image analysis. We therefore developed the software Chrysalis and a group of Imaris Xtensions to automate this process. The resulting automation allowed for high-throughput histo-cytometry analysis of three-dimensional confocal microscopy and two-photon time-lapse images of T cell–dendritic cell interactions in mouse spleens. It was also applied to epi-fluorescence images to quantify T cell localization within splenic tissue by using a “signal absorption” strategy that avoids computationally intensive distance measurements. In summary, this image processing and analysis software makes histo-cytometry more useful for immunology applications by automating image analysis.

Imaging of biological samples has traditionally been used to resolve anatomic structures (1) or identify specific cells in tissues (2). Recent advances in image analysis, like histo-cytometry (3) and dynamic in situ cytometry (4) have expanded the depth of analysis by increasing characterization of cell types and objective quantification of cells in images. These new techniques combine multispectral image analysis with a quantitative workflow. The image quantification is performed by analyzing image-derived statistics in flow cytometry analysis software (3, 4). These approaches can quantify the number and location of cells throughout a tissue (5), identify cell-cell interactions (6), and correlate protein expression to cellular localization (7). Histo-cytometry and dynamic in situ cytometry have been applied to a variety of imaging systems, including confocal (810), epi-fluorescence (11, 12), and two-photon microscopy (4). However, these approaches are time consuming because of the need for extensive hands-on image processing. We addressed this issue by creating the software Chrysalis and a suite of Imaris Xtensions to batch image processing and analysis (https://histo-cytometry.github.io/Chrysalis/). This automation reduced hands-on analysis time for confocal, epi-fluorescence, and two-photon microscopy images. The broad applicability of this protocol was confirmed by quantifying cell localization and cell-cell interactions in the spleen, using multiple imaging platforms. Automation should facilitate the use of the powerful histo-cytometry technique.

Six- to eight-wk-old C57BL/6 (B6) female mice were purchased from the Jackson Laboratory or the National Cancer Institute Mouse Repository (Frederick, MD). ItgaxYFP (13) and Rag1−/− UbcGFP (14) TEa TCR transgenic (Tg) (15) female mice were a gift from B.T. Fife (University of Minnesota). Rag1−/− B3K506 TCR TG (16) and Rag1−/− B3K508 TCR Tg (16) were bred and housed in specific pathogen–free conditions in accordance with guidelines of the University Institutional Animal Care and Use Committee and National Institutes of Health. The University Institutional Animal Care and Use Committee approved all animal experiments.

Mice were injected i.v. with 1 × 107 CFUs of ActA-deficient Listeria monocytogenes expressing the P5R peptide (Lm-P5R) (17, 18).

Lymph nodes were collected from Rag1−/− B3K506 TCR Tg, Rag1−/− B3K508 TCR Tg, and Rag1−/− UbcGFP TEa TCR Tg mice, and a small sample was stained with allophycocyanin-labeled CD4 Ab (RM4-5; Tonbo Biosciences) and analyzed on an BD LSR II (BD Biosciences) flow cytometer using FlowJo software (Tree Star). The results were used to calculate the amount of the remaining sample needed to transfer 1 million CD4+ T cells. In some cases, the T cells from the Rag1−/− B3K506 and Rag1−/− B3K508 TCR Tg mice were also labeled with CellTracker Orange (Thermo Fisher Scientific) or CellTraceViolet (Thermo Fisher Scientific), respectively (19). One million TCR Tg cells were transferred into B6 mice by i.v. injection 24 h prior to infection with Lm-P5R.

Twenty-micrometer splenic sections from naive or Lm-P5R–infected mice were stained with Brilliant Violet (BV) 421–conjugated F4/80 (BM8; BioLegend), Pacific Blue–conjugated B220 (RA3-6B2; BioLegend), CF405L-conjugated CD8⍺ (53-6.7; BioLegend), AF488-conjugated phosphorylated form of S6 kinase (pS6) (2F9; Cell Signaling Technology), CF555-conjugated CD86 (GL-1; BioLegend), AF647-conjugated CD45.2 (104; BioLegend), AF700-conjugated MHC class II (MHCII) (M5/114.15.2; BioLegend), CF514-conjugated CD11c (N418; BioLegend), BV480-conjugated CD3 (17A2; BD Biosciences), and AF594-conjugated SIRP⍺ (P84; BioLegend) Abs. Certain purified Abs from BioLegend were conjugated with CF405L, CF514, or CF555 with Biotium Mix-n-Stain labeling kits. Confocal microscopy was performed with a Leica TCS SP5 confocal microscope with two HyD detectors; two PMT detectors; 405, 458, 488, 514, 543, 594, and 633 laser lines; and a 63× oil objective with a 1.4 numerical aperture. The mark-and-find feature in the Leica Application Suite was used to image 12 T cell zones in each spleen, with each image consisting of a 20-μm z-stack acquired at a 0.5-μm step size. Additionally, the Leica TCS SP5 microscope was used to image single-color–stained UltraComp eBeads (Thermo Fisher Scientific) for generating a compensation matrix.

Spleens from B6 mice infected 48 h earlier with Lm-P5R were fixed with paraformaldehyde, dehydrated with sucrose, and embedded in OCT. Seven-micrometer sections of these spleens were stained with BV421-conjugated F4/80, AF488-conjugated B220 (RA3-6B2; BioLegend), AF647-conjugated CD45.2 (104; BioLegend), and AF594-conjugated CD3 (17A2; BioLegend) Abs. The samples were imaged with a Leica DM6000 B epi-fluorescence microscope equipped with a dry 20× objective with 0.5 numerical aperture and a Leica DFC9000 camera with custom filter cubes. The tiling feature in the Leica Application Suite (Leica Microsystems) software was used to image the entire splenic section. The images were analyzed in Imaris 8.4 (Bitplane), which was used to create surfaces to identify TCR Tg cells. For quantifying T cell localization by signal absorption, statistics for the TCR Tg cell surfaces were exported with the XTStatisticsExport Xtension and imported into FlowJo v10.3 (Tree Star) for analysis. To quantify T cell localization by distance measurement, surfaces were also created for B cell follicles based on B220 staining. The Distance Transformation Xtension was then used to calculate the distance of T cells from the follicle edge toward the follicle center. Statistics for TCR Tg cells were exported with the XTStatisticsExport Xtension and imported into FlowJo v10.3 (Tree Star) for quantification. With the distance method, T cells were considered to reside in a B cell follicle if they were >0 μm into a B cell follicle. For a detailed protocol, refer to the Histo-cytometry Protocol and Documentation file available at https://histo-cytometry.github.io/Chrysalis/.

Rag1/ UbcGFP TEa TCR Tg CD4+ T cells, CMTMR-labeled B3K506 TCR Tg T cells, and CTV-labeled B3K508 TCR Tg T cells were transferred into ItgaxYFP mice that were then infected with Lm-P5R bacteria 24 h after cell transfer. Recipient spleens were immobilized on plastic coverslips, sliced longitudinally with a vibratome, and perfused with 37°C DMEM medium bubbled with 95% O2 and 5% CO2. Samples were imaged with a 4-channel Leica TCS SP8 MP microscope with a resonant scanner containing two NDD and two HyD photomultiplier tubes operating at video rate. The objective was a water dipping 25× with 0.95 numerical aperture. Samples were excited with a MaiTai TiSaphire DeepSee HP laser (15 W; Spectra-Physics) at 870 nm, and emissions at 440–480 (Cell Trace Violet), 500–520 (GFP), 520–560 (Yellow Fluorescent Protein), and 560–630 (CMTMR) nm were collected. Images acquired were 20–250 μm below the cut surface of the spleen slice, and 512 × 512 XY frames were collected at 3.0-μm steps every 30 s for 30 min.

For automated histo-cytometry analysis, a compensation matrix was created in ImageJ (National Institutes of Health) by using the GenerateCompensationMatrix script on images of single-color–stained Ultracomp eBeads. This compensation matrix was applied to three-dimensional images and movies in Chrysalis to compensate for the spillover of each fluorescent signal from its channel into other channels. Chrysalis was also used for further automated image processing as described in Figs. 1A and 5A. Imaris 8.3, 8.4, 9.0, and 9.1 (Bitplane) were used for image analysis, including surface creation to identify cells in images. The Sortomato V2.0, XTChrysalis, and XTChrysalis2phtn Xtensions were used in Imaris for identifying cellular subsets based on protein expression, quantifying cell-cell interactions, and exporting cell surface statistics. Statistics were exported from these applications and imported into FlowJo v10.3 (Tree Star) for quantitative image analysis. Details of these steps are described in the Histo-cytometry Protocol and Documentation file that is available at https://histo-cytometry.github.io/Chrysalis/.

For the traditional histo-cytometry analysis, a compensation matrix was generated and applied to the three-dimensional images with the Leica Application Suite (Leica Microsystems) software. Imaris 8.4 (Bitplane) was used to merge images from a single spleen together by stacking them in the z-plane. The dendritic cell (DC) channel was generated in Imaris 8.4 (Bitplane) using the Channel Arithmetics Xtension prior to running surface creation to identify DCs and TCR Tg cells in images. DCs were categorized as XCR1 or SIRP⍺ DCs using the Sortomato Xtension, and the distance to each DC subset was calculated with the Distance Transformation Xtension. Statistics were exported for each surface and imported into FlowJo v10.3 (Tree Star) for quantitative image analysis.

All of the code generated for image processing or analysis can be downloaded at https://histo-cytometry.github.io/Chrysalis/, including compiled versions of Chrysalis for Windows and Mac OSX, with a Linux version available upon request because of GitHub limitations on file size. Additionally, all of the Imaris Xtensions are compatible with Windows and Mac OSX. The documentation for the code as well as a detailed protocol for image acquisition and analysis is also provided at this GitHub link.

Image acquisition, processing, and analysis with histo-cytometry consists of eight steps (Fig. 1A). We developed a stand-alone software called Chrysalis for automating the three image processing steps (steps 2–4) as well as a suite of Imaris Xtensions that automate two of the image analysis steps (steps 6 and 7; Fig. 1A). For processing three-dimensional images, Chrysalis spectrally unmixes images, merges images, and generates new channels prior to image analysis in Imaris (Fig. 1A). Each of these features addresses existing issues with standard image analysis workflows and expedites image analysis. For example, spectral unmixing accounts for spectral overlap between different fluorophores and fluorescent proteins (20). To aid in this step, we wrote a script that automatically generates a compensation matrix from user-provided, single-color control images. Chrysalis uses this compensation matrix to spectrally unmix an image with a linear unmixing algorithm (Fig. 1B) (21).

FIGURE 1.

Image processing with Chrysalis. (A) Diagram of the histo-cytometry workflow on three-dimensional images when automated by Chrysalis and XTChrysalis. (B) B220 and F4/80 staining of splenic tissue before and after spectral unmixing in Chrysalis. (C) CD11c staining and histogram of DCs in 12 confocal microscopy images merged together in the z-plane. (D) Generation of a DC voxel channel with Chrysalis’ new channel feature by using the fluorescence of existing channels, including B220, CD11c, F4/80, and MHCII, which are depicted for a splenic tissue section. Scale bars, 20 μm. Data representing two to three independent experiments are shown.

FIGURE 1.

Image processing with Chrysalis. (A) Diagram of the histo-cytometry workflow on three-dimensional images when automated by Chrysalis and XTChrysalis. (B) B220 and F4/80 staining of splenic tissue before and after spectral unmixing in Chrysalis. (C) CD11c staining and histogram of DCs in 12 confocal microscopy images merged together in the z-plane. (D) Generation of a DC voxel channel with Chrysalis’ new channel feature by using the fluorescence of existing channels, including B220, CD11c, F4/80, and MHCII, which are depicted for a splenic tissue section. Scale bars, 20 μm. Data representing two to three independent experiments are shown.

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Another issue addressed by Chrysalis is the image processing required for efficiently analyzing cell-cell interactions in three-dimensional images. When analyzing cell-cell interactions, high magnification images need to be taken to observe the interaction event. Analysis of interactions in large tissues, such as spleen or lymph node, can be performed by tiling images of the entire tissue together. However, this process is extremely time intensive for image acquisition and analysis because of the high magnification and large number of images required. This approach is also inefficient in cases where the interaction event occurs only in a small percentage of the tissue. Rare interactions within three-dimensional images can instead be identified at the microscope, allowing for the acquisition of only the images that depict the relevant interactions at high magnification prior to manually merging the images together for analysis. Such a process was previously applied to analyze T regulatory cell–DC clusters (7). To make it easier to study rare interaction events, Chrysalis can automatically merge multiple images from one tissue through stacking images in the z-plane (Fig. 1C), which allows for time-efficient and consistent analysis of the relevant cell-cell interaction event.

Some cell types require identification based on expression of multiple proteins. For example, DCs are identified by their expression of CD11c and MHCII but not B220, F4/80, or CD3 (13, 22, 23). To address this issue, Chrysalis creates new channels consisting of voxels that are above a computer-generated threshold (24) for user-selected “include” channels and below a computer-generated threshold for user-selected “exclude” channels, a process called voxel gating (3). A user-selected base channel expressed by the cell type dictates the signal intensity in this new channel. For a new DC channel, CD11c and MHCII would be the include channels, whereas B220, F4/80, and CD3 would be the exclude channels, and the base channel would be CD11c (Fig. 1D). In effect, this new channel provides better DC resolution than the CD11c channel alone.

For histo-cytometry analysis, Chrysalis-processed images are imported into the image analysis software Imaris, which creates surfaces to identify cells based on the image’s channels (3, 8, 10, 25). These surfaces are created based on user-specified fluorescence intensity thresholds for the cell population of interest and the expected diameter of the cell. For example, nonproliferating adoptively transferred TCR Tg T cells can be identified based on the fluorescence intensity of a congenic marker Ab and a 6-μm diameter cell size. Once the surface creation parameters are set for one image, they can be automatically applied to other images that were acquired with the same microscope settings. However, it is important to visually inspect the quality of surface generation for each image by checking for potential issues, such as whether a group of cells is classified as a single cell. This step is necessary because differences in cell state (e.g., resting versus proliferating cells) can impact the accuracy of surface creation.

Traditionally, the steps required to analyze surfaces require extensive hands-on time. Thus, we created the Xtension XTChrysalis, which automates this process. XTChrysalis does the following: 1) it separates existing surfaces into new surfaces based on a gating scheme defined in an Xtension called Sortomato, 2) it calculates distances to each new surface, 3) it rescales signal intensities for any images, and 4) it exports statistics for any surface (Fig. 1A). The exported statistics contain each channel’s intensity mean and minimum values for each cell as well as each cell’s volume, sphericity, and position. All values have 0.1 added to them to enable logarithmic display of each parameter. These data can be directly imported into quantitative analysis software, such as FlowJo or XiT (26), for further analysis.

To demonstrate three-dimensional image analysis with Chrysalis and XTChrysalis, we analyzed images of T cells, DCs, and their interactions captured by confocal microscopy. Following infection, DCs interact with T cells by presenting MHCII-bound peptides derived from the invading pathogen, leading to TCR signaling (27, 28). To analyze this type of interaction, splenic tissue from L. monocytogenes–infected mice was analyzed by 10-color confocal microscopy. T cell responses were examined using a system involving adoptive transfer of B3K506 TCR Tg CD4+ T cells that express P5R peptide:MHCII–specific TCRs. B3K506 TCR Tg T cells were injected into B6 recipients that were then infected with Lm-P5R bacteria. Twenty-four hours postinfection, 12 T cell zones were imaged per spleen to obtain sufficient cells for analysis (29). We used Chrysalis to spectrally unmix, rescale, and merge images and generate a new channel representing DC voxels before image analysis in Imaris (Fig. 2A). TCR Tg cell surfaces were then created based on CD45.2 fluorescence (Fig. 2B). Staining for pS6, an indicator of TCR signaling (30), was examined within those surfaces to identify cells undergoing TCR signaling (Fig. 2B). DC surfaces were generated based on the DC voxel channel, thereby identifying hundreds of DCs (Fig. 2C). The Sortomato Xtension was used to identify a gating strategy to subset the DCs based on expression of CD8⍺ or SIRP⍺ (Fig. 2D) (22, 3133). XTChrysalis was then applied to the processed images, and the resulting data were analyzed in FlowJo.

FIGURE 2.

Chrysalis and XTChrysalis analysis of a three-dimensional image. (A) Confocal microscopy 10-color image before and after Chrysalis processing. (B) Identifying TCR Tg cells with CD45.2 staining and TCR signaling based on pS6 expression. (C) DC voxels (CD11c+ MHCII+ B220 CD3 F4/80) that were used to identify DCs by surface creation in Imaris. (D) Two-dimensional plot generated with Sortomato for subsetting DC surfaces into SIRP⍺+ or XCR1+ DCs based on SIRP⍺ and CD8⍺ expression. (E) Comparison of the hands-on time required for histo-cytometry analysis of a set of confocal microscopy images of a spleen using the traditional or Chrysalis-automated workflow depicted in reference to the diagram in Fig. 1A. Scale bars, 20 μm. Data representing two to three independent experiments are shown.

FIGURE 2.

Chrysalis and XTChrysalis analysis of a three-dimensional image. (A) Confocal microscopy 10-color image before and after Chrysalis processing. (B) Identifying TCR Tg cells with CD45.2 staining and TCR signaling based on pS6 expression. (C) DC voxels (CD11c+ MHCII+ B220 CD3 F4/80) that were used to identify DCs by surface creation in Imaris. (D) Two-dimensional plot generated with Sortomato for subsetting DC surfaces into SIRP⍺+ or XCR1+ DCs based on SIRP⍺ and CD8⍺ expression. (E) Comparison of the hands-on time required for histo-cytometry analysis of a set of confocal microscopy images of a spleen using the traditional or Chrysalis-automated workflow depicted in reference to the diagram in Fig. 1A. Scale bars, 20 μm. Data representing two to three independent experiments are shown.

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This automated workflow was compared with the traditional histo-cytometry protocol to determine the reduction in hands-on analysis time as a result of automation. For this experiment, 12 splenic T cell zone images were acquired by confocal microscopy as described in Fig. 2A. These images were analyzed to quantify T cell–DC interactions. The automated approach was performed as in Fig. 2A–D, whereas the traditional approach used the Leica Application Suite for image processing (steps 2–4) and Imaris for image analysis (steps 5–7; Fig. 1A). For the image processing, the traditional approach required 47 min of hands-on time, whereas Chrysalis only required 4 min, yielding a 91% reduction in hands-on time (Fig. 2E). Automation of the image analysis performed in Imaris provided a 74% reduction in hands-on time, requiring 80 min with the traditional technique and 21 min with the automated protocol (Fig. 2E). These results demonstrate that the Chrysalis-automated workflow confers a significant reduction in hands-on time required for histo-cytometry analysis of confocal images.

As expected, B3K506 TCR Tg cells in confocal images contained the CD45.2 signal, whereas DCs had CD11c and MHCII signals (Fig. 3A). Surprisingly, however, there were two populations of TCR Tg cells, one lacking CD11c and MHCII signals and one with these signals (Fig. 3B). The populations were similar in cell size but the MHCIIhigh CD11chigh population had greater TCR signaling based on pS6 expression (Fig. 3B). Because MHCII and CD11c are not expressed by T cells (34), we hypothesized that the TCR Tg cell surfaces “absorbed” MHCII and CD11c signals by being in close proximity to DCs. This hypothesis was tested by comparing the frequency of T cell–DC interactions for the MHCIIhigh CD11chigh and the MHCIIlow CD11clow T cells. The MHCIIhigh CD11chigh T cells interacted with XCR1+ and SIRP⍺+ DCs 10 times as often as the MHCIIlow CD11clow T cells, suggesting that the DC signal absorption hypothesis was correct (Fig. 3C).

FIGURE 3.

The signal absorption strategy can accurately quantify cell-cell interactions and cellular localization. (A) FlowJo analysis of CD11c, CD45.2, and MHCII expression on DCs (green) and B3K506 TCR Tg T cells (red) identified in confocal microscopy images. (B) Histogram of volume and pS6 expression for MHCIIhigh CD11chigh (red) and MHCIIlow CD11clow (blue) TCR Tg T cells. (C) Quantifying T cell–DC interactions for MHCIIhigh CD11chigh (red) and MHCIIlow CD11clow (blue) TCR Tg T cells with SIRP⍺+ and XCR1+ DCs. (D) Epi-fluorescence image of splenic tissue stained for F4/80, B220, CD4, and CD45.2, with TCR Tg cell surfaces created based on CD45.2 fluorescence. TCR Tg surfaces were subsetted into cells that absorbed B220 or F4/80, thereby allowing for the characterization of TCR Tg cell localization. Scale bar, 20 μm. (E) Representative gating scheme with 1 × 10−1 μm added to each cell for logarithmic visualization and (F) quantification of the percentage of TCR Tg cells in B cell follicles 3 d after Lm-P5R infection when analyzed by B220 absorption or T cell distance into B cell follicles (n = 7). Data representing two to three independent experiments are shown. A paired t test was used to determine significance for (F). No significant difference was detected.

FIGURE 3.

The signal absorption strategy can accurately quantify cell-cell interactions and cellular localization. (A) FlowJo analysis of CD11c, CD45.2, and MHCII expression on DCs (green) and B3K506 TCR Tg T cells (red) identified in confocal microscopy images. (B) Histogram of volume and pS6 expression for MHCIIhigh CD11chigh (red) and MHCIIlow CD11clow (blue) TCR Tg T cells. (C) Quantifying T cell–DC interactions for MHCIIhigh CD11chigh (red) and MHCIIlow CD11clow (blue) TCR Tg T cells with SIRP⍺+ and XCR1+ DCs. (D) Epi-fluorescence image of splenic tissue stained for F4/80, B220, CD4, and CD45.2, with TCR Tg cell surfaces created based on CD45.2 fluorescence. TCR Tg surfaces were subsetted into cells that absorbed B220 or F4/80, thereby allowing for the characterization of TCR Tg cell localization. Scale bar, 20 μm. (E) Representative gating scheme with 1 × 10−1 μm added to each cell for logarithmic visualization and (F) quantification of the percentage of TCR Tg cells in B cell follicles 3 d after Lm-P5R infection when analyzed by B220 absorption or T cell distance into B cell follicles (n = 7). Data representing two to three independent experiments are shown. A paired t test was used to determine significance for (F). No significant difference was detected.

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The experimental approach described above was also used to assess the locations of B3K506 TCR Tg cells by epi-fluorescence microscopy. Spleens from B6 recipients of B3K506 T cells infected three d earlier with Lm-P5R bacteria were stained for F4/80, B220, and CD4 to identify the red pulp, B cell zones, and T cell zones, respectively (Fig. 3D) (35, 36). Spleens were also stained for CD45.2 to identify the TCR Tg cells. Macrophages in the red pulp express F4/80 (36), and B cells in the B cell zone express B220 (37), but neither protein is expressed by T cells (3840). Therefore, TCR Tg surfaces that have the B220 signal should be in close proximity to B cells and reside in B cell follicles, whereas those with the F4/80 signal should be near macrophages and localize to the red pulp. Indeed, although most of the B3K506 T cells were in the T cell zones, some were in the B cell follicles and absorbed the B220 signal, whereas others were in the red pulp and absorbed the F4/80 signal (Fig. 3D). Thus, the location of a cell can be determined based on absorption of fluorescent signal from proteins expressed by nearby cells.

The signal absorption strategy was further validated by comparing this strategy to a different counting method. Epi-fluorescence microscopy images were acquired and analyzed as described in Fig. 3D and the localization of the TCR Tg cells to B cell follicles was analyzed. For the signal absorption strategy, follicular TCR Tg cells were defined based on their absorption of B220 fluorescent signal (Fig. 3E). In the other method, Imaris was used to determine the distance of each T cell from a follicle edge to the center of that follicle. A distance >0 μm indicated that a T cell resided in the follicle (Fig. 3E). There was no significant difference in the percentages of T cells found in B cell follicles based on the signal absorption or distance quantification methods (Fig. 3F). These results demonstrate that signal absorption can determine cellular localization as accurately as a more traditional counting technique.

The capacity of the signal absorption strategy to identify cell location was also employed to validate the concept that TCR signal strength influences Th cell differentiation (41). It has been shown that naive T cells with high TCR affinity for peptide:MHCII tend to differentiate into Type 1 helper (Th1) cells, whereas cells with lower affinity TCRs primarily adopt the T follicular helper (Tfh) fate (17, 42). These differences in T cell differentiation would be expected to modulate T cell localization because different Th subsets express different chemokine receptors. For example, Th1 cells express CXCR3 (43, 44), driving them toward sites of inflammation such as the splenic red pulp, whereas Tfh cells express CXCR5, allowing them to traffic into B cell follicles (45, 46). Thus, Tfh-biased low TCR affinity T cells would localize to B cell follicles at a higher frequency than Th1-biased high TCR affinity T cells.

B3K506 T cells were compared with B3K508 TCR Tg T cells, which express TCRs with lower affinity for P5R:I-Ab complexes, to test this hypothesis (16, 47). The TCR Tg populations were transferred into B6 mice, which were infected with Lm-P5R bacteria. Spleen sections were stained, imaged by epi-fluorescence microscopy, and analyzed with Chrysalis 1, 2, and 3 d postinfection. As in the previous experiment (Fig. 3D), B220 identified B cell follicles, CD4 defined T cell zones, F4/80 delineated red pulp, and CD45.2 specified TCR Tg T cells (Fig. 4A). T cell localization in the follicles or red pulp was identified based on T cell absorption of B220 or F4/80 signal, respectively (Fig. 4B). As expected, TCR Tg cells were primarily situated in T cell zones in naive mice and during the initial three d following Lm-P5R infection (Fig. 4C–E). However, the signal absorption assay revealed a greater proportion of low TCR affinity B3K508 T cells localized to B cell follicles than to high TCR affinity B3K506 T cells, in line with B3K508 T cells favoring the B cell follicle–homing Tfh cell fate (Fig. 4D) (17). This result demonstrates the ability of the improved histo-cytometry workflow to quantify cellular localization in epi-fluorescence microscopy images with a novel signal absorption strategy.

FIGURE 4.

T cells primarily reside in T cell zones following Listeria infection, and low affinity T cells traffic into B cell follicles more than high affinity T cells. (A) Representative images of B220, CD4, CD45.2, and F4/80 staining of a splenic tissue section acquired by epi-fluorescence microscopy. Scale bar, 100 μm. (B) Gating strategy for using signal absorption to identify B cell follicles (B220+) or red pulp (F4/80+) residing in TCR Tg T cells in epi-fluorescence microscopy images. (CE) Quantification of epi-fluorescence microscopy images that determine B3K506 (filled circle, n = 4) and B3K508 (empty circle, n = 4) cell localization in (C) T cell zone, (D) B cell follicle, or (E) red pulp in spleens of naive mice and mice 1, 2, or 3 d after Lm-P5R infection. Pooled data from three independent experiments are shown. One-way ANOVA was used to determine significance for (D). *p < 0.05, **p < 0.01.

FIGURE 4.

T cells primarily reside in T cell zones following Listeria infection, and low affinity T cells traffic into B cell follicles more than high affinity T cells. (A) Representative images of B220, CD4, CD45.2, and F4/80 staining of a splenic tissue section acquired by epi-fluorescence microscopy. Scale bar, 100 μm. (B) Gating strategy for using signal absorption to identify B cell follicles (B220+) or red pulp (F4/80+) residing in TCR Tg T cells in epi-fluorescence microscopy images. (CE) Quantification of epi-fluorescence microscopy images that determine B3K506 (filled circle, n = 4) and B3K508 (empty circle, n = 4) cell localization in (C) T cell zone, (D) B cell follicle, or (E) red pulp in spleens of naive mice and mice 1, 2, or 3 d after Lm-P5R infection. Pooled data from three independent experiments are shown. One-way ANOVA was used to determine significance for (D). *p < 0.05, **p < 0.01.

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Previously, histo-cytometry has been applied to three-dimensional images; however, this same methodology can be applied to two-photon time-lapse data (movies) (7, 8). Chrysalis can aid in this application because it can spectrally unmix, generate new channels, and rescale movies (Fig. 5A). Additionally, Chrysalis expedites two-photon video analysis by simplifying existing workflows. For example, two-photon movies can have variable image quality owing to poor tissue health stemming from a lack of oxygenation or low tissue temperature (48, 49). Tissue health can be assessed by examining the motility of a control population within the tissue, such as fluorescently labeled polyclonal T cells (50). By reviewing the motility of a control cell population across several movies, movies that depict healthy tissue can be identified prior to conducting in-depth analysis. To optimize this process, Chrysalis processes movies by Gaussian filtering and rescaling each channel to maximize signal intensity and video clarity. The processed movies are saved as audio video interleaved files, which can be quickly examined for tissue health prior to performing more time-consuming analysis.

FIGURE 5.

Chrysalis and XTChrysalis2phtn analysis of a two-photon microscopy video. (A) Diagram of the histo-cytometry workflow on two-photon movies when automated by Chrysalis and XTChrysalis2phtn. (B) Surface-mediated identification of B3K506, B3K508, and TEa TCR Tg cells as well as DCs in two-photon movies. Scale bar, 20 μm. (C) Quantifying cellular velocity in a two-photon video with FlowJo for B3K506 (red), B3K508 (blue), and TEa (gray) TCR Tg T cells. (D) FlowJo analysis of B3K506 and TEa TCR Tg cells in a two-photon video, with quantification of track straightness, total contact time with DCs, longest contact with a DC, and number of prolonged contacts with DCs. Data representing two to three independent experiments are shown.

FIGURE 5.

Chrysalis and XTChrysalis2phtn analysis of a two-photon microscopy video. (A) Diagram of the histo-cytometry workflow on two-photon movies when automated by Chrysalis and XTChrysalis2phtn. (B) Surface-mediated identification of B3K506, B3K508, and TEa TCR Tg cells as well as DCs in two-photon movies. Scale bar, 20 μm. (C) Quantifying cellular velocity in a two-photon video with FlowJo for B3K506 (red), B3K508 (blue), and TEa (gray) TCR Tg T cells. (D) FlowJo analysis of B3K506 and TEa TCR Tg cells in a two-photon video, with quantification of track straightness, total contact time with DCs, longest contact with a DC, and number of prolonged contacts with DCs. Data representing two to three independent experiments are shown.

Close modal

We have also written an Imaris Xtension called XTChrysalis2phtn that batches histo-cytometry analysis of two-photon movies. For each video, XTChrysalis2phtn will do the following: 1) calculate distances between cell surfaces and define cell-cell interactions at each time point, 2) rescale signal intensities, and 3) export statistics for each surface (e.g., average velocity, displacement, volume, and cell-cell interactions) (Fig. 5A). The data generated can be directly imported into FlowJo for further analysis. Thus, Chrysalis and XTChrysalis2phtn automate histo-cytometry analysis of cell-cell interactions and protein expression in two-photon movies, thereby reducing the required hands-on analysis time.

To demonstrate this improved workflow, T cell–DC interactions were quantified in two-photon microscopy movies depicting spleens from B6 recipients of B3K506, B3K508, and TEa TCR Tg cells infected 16 h earlier with Lm-P5R bacteria. The two-photon movies had four colors, which identified DCs and the three different TCR Tg populations (Fig. 5B) (19). Chrysalis spectrally unmixed and rescaled the movies, as well as generated audio video interleaved files to determine tissue health. For further analysis, the processed movies were opened in Imaris, and surfaces were generated for the DCs and TCR Tg populations (Fig. 5B). XTChrysalis2phtn then generated cell statistics for analysis in FlowJo, which provided a way to compare B3K506 and B3K508 T cells recognizing P5R:I-Ab on DCs. TEa TCR Tg cells served as control cells because they do not respond to the infection (16, 17). The B3K506 and B3K508 cells had a lower mean velocity than the TEa cells, suggesting that B3K506 and B3K508 cells interacted with DCs postinfection, whereas TEa cells did not (Fig. 5C). In line with this hypothesis, B3K506 cells had a lower confinement correlate value and greater contact time with DCs than did TEa cells (Fig. 5D). Histo-cytometry analysis of these T cell–DC interactions allowed for a more granular view of these interactions by quantifying the duration of the longest contact event as well as the number of prolonged contact events for each T cell (Fig. 5D). As expected, T cells with the longest contact events with DCs made fewer total contacts with DCs (Fig. 5D). This example demonstrates a powerful and streamlined workflow for analyzing two-photon movies.

The Chrysalis software and Imaris Xtensions described in this manuscript can be applied to a broad range of biological questions while reducing analysis time and empowering quantitative image analysis. We demonstrated the power of this workflow by quantifying T cell localization within splenic tissue in epi-fluorescence images, T cell-DC interactions in confocal microscopy images, and T cell motility and T cell–DC interactions in two-photon microscopy images. These same approaches can answer other immunological questions that require the quantification of cell localization, cell-cell interactions, or the ability to subset cells in images.

To extend the capabilities of this workflow beyond the applications described in this manuscript, we also generated separate Imaris Xtensions for each of the major steps performed by XTChrysalis, such as batched statistics export. With these additional Xtensions, users can daisy-chain Xtensions to batch image analysis in a manner that specifically addresses their research question. To further facilitate the use of this quantitative imaging approach in immunological research, we provide a step-by-step protocol that incorporates the automation steps detailed in this manuscript to streamline acquisition and analysis of confocal, epi-fluorescence, and two-photon microscopy images.

Although our protocol uses the commercial image analysis software Imaris, it can also be paired with free, publicly available software such as CellProfiler and ilastik (5154). Although these programs do not have all of the features of Imaris, these programs are able to perform cell segmentation to identify cells within images, an essential step in the histo-cytometry workflow that is performed by Imaris in our protocol. Additionally, although our protocol uses the commercial software FlowJo for comparing and quantifying image-derived statistics for each identified cell population, publicly available software such as XiT and FACSanadu (Ref. 26 and T.R. Bürglin and J. Henriksson, manuscript posted on bioRxiv) can be used within our workflow in place of FlowJo for quantifying images.

To further reduce analysis time, we developed a signal absorption technique that expedites the quantification of cellular localization. The premise of this method is that a cell near other cells will absorb the nearby cell’s fluorescence. For example, a T cell residing in a B cell follicle will absorb a B220 signal from nearby B cells. Signal absorption can then be used as a readout of cell location. This strategy is favorable over directly quantifying cell distance to a tissue structure because signal absorption only requires creating surfaces for cells and measuring their fluorescent signal. Conversely, the distance quantification approach involves creating surfaces for cells and tissue structures before quantifying the cell distance to the tissue structure. Although the distance quantification approach provides a more definitive determination of localization, the extra steps of this approach require greater hands-on analysis time and computational power. This problem is exacerbated when the distance quantification approach is applied to the analysis of large tissues, like the spleen, or to many biological samples. Therefore, the signal absorption strategy is a simpler and more time-efficient approach for quantifying cellular localization in certain cases.

Whereas we demonstrated that the signal absorption technique works with a variety of image resolutions, it might not be compatible with very high-resolution microscopy techniques, like super-resolution microscopy, because signal overlap will not occur. An additional limitation of the signal absorption technique is the fluorescence intensity of the signal being absorbed. For example, B220 is highly expressed by B cells, and they are abundant in B cell follicles. Therefore, it was possible to use signal absorption of B220 to accurately quantify T cell localization to B cell follicles. If B cells had low florescence intensity for their identifying marker or were extremely rare in the follicles, then the signal absorption method could not be used to quantify follicular T cells.

In summary, Chrysalis and the suite of Imaris Xtensions provide a high-throughput image processing workflow for confocal, epi-fluorescence, and two-photon microscopy images. This approach identifies subtle differences in cell phenotype and cell-cell interactions while also offering up to a 90% reduction in hands-on analysis time. This time-savings reduces the barrier of entry for conducting quantitative, multispectral image analysis. Accessibility to this image analysis pipeline is further enhanced by the accompanying step-by-step protocol describing how to prepare samples, acquire images, and analyze images using the novel Chrysalis software and Imaris Xtensions for confocal, epi-fluorescence, and two-photon microscopy images. An increase in the widespread adoption of these powerful, quantitative image analysis approaches will allow for novel and counterintuitive discoveries about the function and maintenance of the immune system.

We thank J. Walter and C. Ellwood for technical assistance and J. Kotov for reviewing the manuscript. We also thank P. Beemiller for creating Sortomato and M.Y. Gerner for helpful suggestions on histo-cytometry.

This work was supported by National Institutes of Health Grants T32 AI083196 and T32 AI007313 (to D.I.K.) and R01 AI039614 (to M.K.J.).

Abbreviations used in this article:

B6

C57BL/6

BV

Brilliant Violet

DC

dendritic cell

Lm-P5R

Listeria monocytogenes expressing P5R

MHCII

MHC class II

pS6

phosphorylated form of S6 kinase

Tfh

T follicular helper

Tg

transgenic.

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M.J.G. is employed by Bitplane, which produces the Imaris image analysis software that is used extensively in the image analysis pipeline described in this manuscript. The other authors have no financial conflicts of interest.