Optimized Cardiac Leukocyte Isolation for Flow Cytometry Enumeration and Sorting

Abstract

Accurate characterization of immune cell populations within the heart is critical for understanding inflammatory mechanisms that drive cardiac injury and repair. However, isolating viable leukocytes from cardiac tissue remains technically challenging due to the dense extracellular matrix, high collagen content, and susceptibility of immune subsets to enzymatic or mechanical stress. Here, we describe an optimized protocol for cardiac leukocyte isolation that maximizes cell yield, preserves viability, and maintains the integrity of surface markers for flow cytometric analysis and cell sorting. The method integrates controlled enzymatic digestion, gentle mechanical dissociation, and precise debris removal, thereby minimizing stromal contamination and reducing cellular activation artifacts. Using this approach, we demonstrate reproducible recovery of diverse immune populations, including neutrophils, monocytes, macrophages, and lymphocyte subsets from murine hearts. Furthermore, compatibility with high-dimensional flow cytometry and fluorescence-activated cell sorting enables robust downstream applications such as transcriptomic profiling and functional assays. This optimized isolation strategy provides a reliable and versatile tool for investigating the cardiac immune landscape, offering new opportunities to delineate the immunopathology of cardiovascular disease and to develop targeted immunomodulatory therapies.

New & Noteworthy

Flow cytometry of immune cells is highly tissue-specific, requiring tailored enzymatic digestion and staining protocols. We optimized a protocol for cardiac immune cells isolation and enumeration using the following steps: 1) tissue digestion, 2) cell viability; 3) titration of live/dead dye; 4) antibody titration; 5) compensation and blocking. The resulting protocol was used both for flow cytometry and FACS analysis of neutrophils, monocytes, and macrophages, but it could easily be extended to innate immune cells.

Introduction

Precise characterization of immune cell populations within the heart is critical for understanding inflammatory mechanisms that drive cardiac injury and repair (1). Fluorescence-activated cell sorting (FACS) is vital in cardiac research as it provides high-resolution, quantitative, and functional insights into the diverse cellular players in the heart (2). It not only advances our understanding of pathophysiology but also guides the development of targeted therapies for heart disease. FACS can be used to quantify or monitor cell populations in several aspects of cardiac research - immune profiling, tissue injury, tissue regeneration, fibrosis, isolated cell studies, cultured cell studies, and clinically relevant translational studies (3,4) (Figure 1).

Figure 1.

A schematic representation of the multiple applications of FACS in cardiac research.

When isolating and enumerating inflammatory cells from the heart, protocols can vary considerably due to differences in model, tissue handling, and research focus (5). Major sources of variation include enzymatic digestion, particularly enzyme type, concentration, combination of enzymes (e.g., collagenase II, collagenase IV, DNase I), and duration and temperature of digestion, all of which affect cell yield and viability. Some protocols use Langendorff perfusion for uniform enzymatic digestion, while others rely on mechanical mincing followed by incubation with digestive enzymes, which can influence the recovery of fragile immune cells (6,7). Additionally, the use of filtration (40–100 μm mesh), red blood cell lysis, and/or Percoll/Ficoll gradients might result in distinct recovery yields and products. These steps affect purity, yield, and representation of certain immune subsets. Equally important are the steps for staining and selection of antibody panels that can bias detection and enumeration (8). Some protocols stop at enumeration (flow cytometry analysis), while others proceed to sorting, requiring stricter viability and purity. Moreover, it is also essential to consider the species and disease model due to different tissue densities and immune cell distributions. For example, protocols may differ between post-myocardial infarction (MI), myocarditis, or chronic remodeling models due to distinct inflammatory landscapes (9–11).

Antibody and reagent titration/optimization are essential when isolating and enumerating inflammatory cells in the heart because cardiac tissue is complex and prone to variability. Titrating antibodies helps enhance signal-to-noise ratio and improve fluorophore balance in multicolor panels. Careful titration and optimization of antibodies and reagents are indispensable for accurate, reproducible, and biologically meaningful results when analyzing cardiac immune cells (12,13). Here, our study provides an optimized method for cardiac leukocytes for flow cytometry and sorting in a mouse model of permanent occlusion MI. The schematic diagram showing the process involved in flow cytometry is shown in Figure 2. To produce this optimized FACS protocol for cardiac immune cells isolation and enumeration, we optimized each of the following steps: 1) tissue digestion, 2) cell viability; 3) titration of live/dead dye; 4) titration of individual antibodies; and 5) compensation and blocking. The resulting protocol was used both for flow cytometry analysis and FACS of neutrophils, monocytes, and macrophages, but it could easily be extended to innate immune cells (14–16).

Figure 2.

Depiction of the protocol workflow.

Materials and Methods

Instrument –

For protocol optimization, we used the BD FACSAria^™ Fusion Flow Cytometer, which is outfitted for multicolor analyses and cell sorting operations. The system is designed for the simultaneous sorting of up to four cell populations, with a configuration that includes four lasers and twelve detection parameters (Table 1).

Table 1-.

BD FACSAria^™ Fusion Flow Cytometer sorter parameters.

Wavelength	Operating Power	Long Pass Filter	Band Pass Filter	Fluorochromes used
405 nm	75 mW		450/50	Super Bright 436
		505	525/50	L/D Aqua
		635	670/30	BV650
488 mW	50 mW		488/10	Forward Scatter (FSC)
			488/10	Side Scatter (SSC)
		495	530/30	Alexa 488
		690	695/40	PerCP
561 nm	75 mW		586/15	PE
		595	610/20	PE-CF594
		635	670/30
		750	780/60	PE-Cy7
637 nm	100 mW		670/30	APC
		685	730/45
		750	780/60	APC-Cy7

Reagents and supplies –

Isoflurane (Isospire^™), alcohol (Pharmco-Aaper), surgical instruments for tissue harvest (forceps and scissors), 3 mL syringe with 27½G needle (BD PrecisionGlide^™ Needle, 305109), 60 ×15mm petri dish (Fisher Scientific^™, FB0875713A), 15 mL and 50 mL falcon tubes (Fisher Scientific^™, 14-959-53A and 14-432-22), collagenase II (Worthington LS004177), RPMI-1640 media (Gibco, 11875–093), phosphate buffered saline (PBS; Gibco, 10010–023), ethylenediaminetetraacetic acid (EDTA; SIGMA, E6758), bovine serum albumin (BSA; Fisher Scientific^™, BP9703–100), fetal bovine serum (FBS; Gibco 16000044), countess slides (Invitrogen, C10228), trypan blue (Invitrogen, T10282), cell strainer (FACS tube cap, Stellar Scientific, FSC-FLTCP), polystyrene round-bottom 12 × 75 mm tubes for FACS (Stellar Scientific, 352054), Live/Dead (L/D) aqua reagent (Invitrogen, L34965), Brilliant stain buffer (Bioscience BD, 563794), UltraComp eBeads (Invitrogen, 01-2222-42), Fc Block CD16/CD32 (Bioscience BD, 553142), True-Stain Monocytes Blocker (Biolegend, 426102), Alexa 488 for F4/80 (Invitrogen, 53-4801-82), PE for CD86 (BD 553692), PE-CF594 for Ly6C (Bioscience BD, 582728), PE-Cy7 for CD45R/B220 (Biolegend, 103222), APC for CD11b (Biolegend, 101212), APC-Cy7 for CD45 (Biolegend, 147718), SB 436 for Ly6G (Invitrogen, 62-9668-82), BV650 for CD3 (Invitrogen 41-60031-82), PerCp for CD163 (Invitrogen, 46-1631-82), Trizol (for sorting only; Life Technologies, 15596018).

Solutions –

Digestion media: Collagenase II A (0.5 mg/mL, 250–300 U/mg activity) prepared in RPMI-1640, prewarmed to 37°C. The digestion media was prepared fresh each day. Digestion neutralization buffer: Cold PBS with 2 mM EDTA and 4% BSA. Stain buffer: 0.2% BSA in PBS (kept cold).

Animals and Surgery –

Animal procedures were conducted in accordance with the “Guide for the Care and Use of Laboratory Animals” (17) and were approved by the Institutional Animal Care and Use Committee at East Carolina University (ECU). Both male and female C57BL/6 mice (Charles River stock #027), aged 4 to 6 months, were used in this study (n=6/sex). Animals were housed at a controlled temperature (22 ± 2 °C) on a 12 h light/dark cycle, fed standard laboratory mice chow ad libitum, and had free access to tap water. Ligation of the left anterior descending coronary artery was performed as previously described to induce permanent occlusion myocardial infarction (MI) (18).

Tissue Harvest -

Mice were anesthetized with 2–3% isoflurane in an oxygen mix. We validated the anesthetic by pinching the mouse’s toes and tail, which showed that the mouse was sleepy, and we removed the hair (Nair) from the chest. The chest was cleaned with sterile alcohol pad before harvesting the heart. Blood was collected from the carotid artery (for a separate project). Since we collect as much blood as possible, when the heart was excised, it contains a minimal amount. Mice were perfused ex vivo, through the ascending aorta, with 2–3 mL PBS, which resulted in visible blanching of the cardiac tissue, indicating effective removal of most of the circulating blood and the LV was separated from the other chambers of the heart, particularly care was taken to ensure that major arteries and veins were not present as it can skew the results. The LV was placed in a petri dish on ice for immediate cell digestion. The tissue was finely chopped utilizing cold scissors, employing only a minimal amount of cold PBS (100 μL) to preserve tissue integrity. The tissue was then transferred to a 50 mL falcon tube with 8 mL of digestion buffer and incubated at 37°C on a rotator for up to 25 minutes at 10 – 20 rpm.

Digestion and Optimization of Cellular Yield -

To maximize cellular yield and viability, we critically examined multiple protocols described in prior literature (19–21). To standardize the protocol, we weighed the tissue and started with 50–74 mg or 75–100 mg of LV tissue. Then, we systematically varied both the physical and chemical conditions of digestion, including the concentration of collagenase II (0.25 and 0.5 mg/mL) and digestion duration (5, 10, 15, and 20 minutes). Variations in the digestion neutralization buffer’s conditions and volumes were also evaluated. Specifically, we tested cold PBS enriched with 2 mM EDTA and 4% BSA and compared it to cold PBS supplemented with 2 mM EDTA and 4% FBS. After digestion, the collagenase was neutralized with 10 mL of cold neutralization buffer and kept on ice.

Optimization of Single Cell Suspension –

For optimal flow cytometry results, it is crucial to obtain a cell suspension characterized by a high proportion of viable singlets prior to antibody labeling. The cellular suspension was filtered through a 35 μm cell strainer into a 50 mL Falcon tube to remove tissue clumps and cells were then centrifuged at 400 × g for 5 minutes at 4°C. The supernatant was carefully decanted, and 3 mL of cold PBS was used to rinse cells. Gentle vortexing was used to resuspend cells prior to a second centrifugation under the same conditions. The supernatant was decanted carefully, and the leftover cell suspension was resuspended in 200 μL of cold stain buffer. Cells were then counted using Countess II (Life Technologies USA, AMQAX1000) and/or cells were counted manually using trypan blue and a hemocytometer to calculate cellular yield. We recommend performing all FACS staining steps using ice-cold reagents and maintaining samples at 4°C, as low temperatures prevent the internalization of surface antigens, thereby minimizing loss of fluorescence intensity.

Titration of Live/Dead Aqua Staining –

The goal of titrating the Live/Dead (L/D) dye differs significantly from that of antibody titration. In this context, the aim is to identify dye concentration that ensures optimal differentiation between live and dead cells, rather than achieving maximum separation. Optimal concentration will make sure the non-specific binding of the Live Dead dye is minimal to the live cells. Thus, it is critical to minimize the dye’s binding affinity to live cells. The L/D aqua stock solution was prepared as per manufacturer’s instructions before titration to identify the optimized concentration to be used with cardiac immune cells. For titration purposes, ~1 million cells were collected into a FACS tube, rinsed with 3 mL cold PBS, centrifuged at 4°C for 5 min at 400 × g, and the pellet resuspended in 1600 μL of cold PBS. The cell suspension was thoroughly mixed and distributed across seven FACS tubes. Serial dilutions of the L/D dye were prepared (1:1000, 1:2000, 1:4000, 1:8000, 1:16000, 1:32000) in PBS into a final volume of 20 μL and added individually to a tube with the cell suspension. The seventh tube served as unstained control and received 20 μL of PBS. Samples were incubated for 15 minutes at room temperature in the dark, washed with 3 mL of cold staining buffer, vortexed, and centrifuged at 1000 × g for 5 min at 4°C. After carefully discarding the supernatant, the wash step was repeated to ensure thorough removal of unbound dye. Finally, the cell pellet was then resuspended in 300 μL of cold staining buffer before being run in the flow cytometer. Once the ideal dilution was identified, the stock solution was diluted accordingly and divided into 2 μL aliquots, which were stored at −20°C, protected from light and moisture. Each aliquot was used as needed, and any remaining dye was discarded to avoid repeated freeze-thaw cycles.

Titration of Antibodies –

Our goal was to effectively enumerate neutrophils, monocytes, and macrophages, with the later possibility of differentiating between pro-inflammatory M1-type and anti-inflammatory M2-type macrophages. Thus, we performed titration of nine distinct antibodies. After LV digestion and generation of a single-cell suspension, cells were subjected to a series of centrifugations at 400 xg for 5 minutes at 4°C. After each centrifugation, the supernatant was gently removed, and cells were resuspended in cold PBS. The wash and spin cycle was performed twice to ensure thorough cleansing. Finally, the cells were resuspended in a total volume of 3850 μL of cold stain buffer and mixed well through pipetting. Cells were manually counted using trypan blue as acurate cell concentration is essential for the integrity of the titration process. The homogenous cell suspension was then equally distributed across 37 FACS tubes (4 tubes per antibody × 9 antibodies + 1 unstained control), allocating 1 million cells in 100 μL of stain buffer per tube. This allocation resulted in 36 tubes prepared for staining, with an additional tube unstained, acting as a control to benchmark the staining intensity against background fluorescence. Cells were rinsed with 3 mL of cold stain buffer, resuspended, vortexed, and centrifuged at 400 × g for 5 minutes at 4°C. The supernatant was carefully discarded and pellets were resuspended in 100 μL of cold stain buffer. To reduce non-specific binding (see Box 1), each tube received 5 μL of Fc block and 5 μL of True-Stain Monocyte Blocker.

Box 1-. Blocking:

Fc blocks inhibit non-specific binding of antibodies to cellular Fc receptors. This preemptive step is crucial for avoiding false positives and ensuring data accuracy by allowing only specific antigen recognition by the assay antibodies (19). Monocytes and macrophages are known to engage in nonspecific interactions with antibodies and fluorophores during live cell surface staining procedures (20). The utilization of True-Stain Monocyte Blocker^™, a non-antibody based blocking agent, effectively diminishes these nonspecific interactions on monocytes without impeding the desired specific staining outcomes (21).

Antibody titration was performed using four varying volumes: 5 μL, 2.5 μL, 1.25 μL, and 0.625 μL (i.e., 4 dilutions for each antibody). The total volume within each tube was brought to 200 μL with staining buffer. At this point, the cells were incubated at room temperature for 20 minutes shielded from light. Subsequently, tubes were supplemented with 3 mL of cold staining buffer and centrifuged at 400 × g for 5 minutes at 4°C. The supernatant was carefuly decanted and the pellet was washed once more before resuspension in 300 μL of staining buffer.

Subsequent to the titration of antibody dilutions, we initiated a comprehensive staining trial incorporating both L/D dye and antibodies at the optimized dilutions. Details of the reagent concentrations applied are systematically itemized in Table 2. This necessary phase corroborates the efficacy of the selected antibody concentrations, underpinning the fidelity of the staining methodology prior to scaling up for broader experimentation.

Table 2:

Antibody concentration and immune cell population targets.

Fluorochrome (Antibody details)	Marker	Clone	Catalog No.	Vendor	Antibody (μL/test) / Titer (ng/test)	Immune cell population target
Alexa 488™	F4/80	BM8	53-4801-82	Invitrogen	2.5 / 1250	Macrophages
APC	CD11b	M1/70	101212	Biolegend	2.5 / 500	Myeloid cells
APC/Cyanine7	CD45	I3/2.3	147718	Biolegend	2.5 /500	Leukocytes
BV650	CD3	RM4-5	41-60031-82	Invitrogen	2 / 400	T cells
PE	CD86	GL1	553692	BD Biosciences	2.5 / 500	M1-like macrophages
PE/Cyanine7	B220	RA3-6B2	103222	Biolegend	2 / 500	separate from CD3 to confirm B cells
PE-CF594	Ly6C	AL21	562728	BD Biosciences	0.625 / 125	Monocytes
Per-CP-eFluor™ 710	CD163	TNKUPJ	46-1631-82	Invitrogen	2.5 / 500	M2-like macrophages
Super Bright™436	Ly6G	1A8	62-9668-82	Invitrogen	2 / 400	Neutrophils

Antibody labelling, Staining, and Gating Strategy –

To analyze multiparameter flow cytometry data, it is essential to use either compensation (when the number of the instrument detectors is the same as the number of fluorochromes) or unmixing (when the number of detectors is higher than the number of fluorochromes) to rectify for spillover from fluorochromes (22). In this protocol we incorporated UltraComp eBeads^™ as these can be stained with individual fluorochrome-conjugated antibodies for use as single-color compensation controls to facilitating precise sorting and analysis (see Box 2). Each aliquot of beads presents two distinct populations: a positive population capable of capturing antibodies and a negative population that remains unreactive to antibodies. Therefore, introducing a fluorochrome-conjugated antibody to the beads results in both positive and negative populations, creating a bimodal distribution. This distribution is crucial for single-color compensation controls in complex multicolor flow cytometry experiments. On a microcentrifuge tube we added 5–6 drops of UltraComp eBeads and brought the volume up to 930 μL with cold stain buffer. This solution was later added to the multicolor testing tube that received all optimized titrated antibodies concentration and cells (Table 2). After addition of the compensation beads, the volume was brought up to 200 μL with cold stain buffer and tubes were incubated on ice for 20 minutes in the dark to preserve the integrity of the fluorescent labels and prevent photobleaching. Summary outlining the used buffers and reagents are represented in Table 3. To wash cells, 3 mL of chilled stain buffer was added to each tube, followed by a brief vortexing. Tubes were centrifuged at 400 × g for 5 minutes at 4°C, and the supernatant carefully aspirated. The washing step was repeated once more, and the pellet resuspended in 200 μL of stain buffer. This solution was then taken to the instrument to optimize the gating strategy (Figure 3). Cells were gated sequentially to exclude first debris (Figure 3A) by plotting cell size (forward scatter area, FSC-A) versus granularity (side scatter area, SSC-A); then as doublets can significantly affect analysis, we plotted FSC height (FSC-H) versus FSC-A, with the FSC-Area scaling factor to a near 45° angle distribution, as this angle produces a linear gate (cell size versus height) to select only the cells with proportional area and width; since doublets are wider this allows for doublet discrimination (Figure 3B, doublets visible as light green). Importantly, we stretched FSC-A all across its’s range to ensure that when we later gated for F4/80 we captured both F4/80⁺ with high FSC-A values, which represent resident macrophages that have matured in the tissue, expanding their cytoplasm and accumulating vacuoles/granules, and F4/80⁺ with low FSC-A values that represent infiltrating monocyte-derived macrophages. After doublet discrimination, CD45⁺ live cells were gated by plotting CD45 versus L/D viability dye (Figure 3C); lymphoid cells were then excluded by gating BV650 and CD3 (Figure 3D); this was followed by two parameter density plots to identify the myeloid subsets using CD11b, Ly6G, Ly6C, and F4/80 expression (Figure 3 E–F). Finally, we further separated the macrophage population by plotting CD86 versus CD163 (Figure 3G).

Box 2-. Compensation beads:

For our protocol we chose UltraComp eBeads^™ since these beads facilitate compensation across a range of fluorochromes, which are excited by lasers of various wavelengths including ultraviolet (355 nm), violet (405 nm), blue (488 nm), green (532 nm), yellow green (561 nm), and red (633–640 nm). The antibodies we used were conjugated to Alexa Fluor^™ 488 (blue), R-Phycoerythrin (PE, yellow-green), PE-Cyanine7 (PE-Cy7, excited by blue, green, or yellow green), allophycocyanin (APC, red), APC/Cy7 (red), Super Bright^™ 436 (violet), Brilliant Violet^™ 650 (BV650, violet), and Peridinin-Chlorophyll-Protein eFluor^™ 710 (PerCP, blue). It is critical to ensure that compensation is done for the correct spectra.

Table 3:

Composition of each buffer/reagent.

Reagents	Composition	Catalog number
Collagenase II	Collagen degrading enzymes	Worthington LS004177
RPMI 1640 media	Inorganic salts, amino acids, vitamins, glucose, phenol red	Gibco, 11875-093
Phosphate buffered saline (PBS)	Sodium chloride (NaCl) – maintains osmotic balance Potassium chloride (KCl) Disodium phosphate (Na2HPO4) Monopotassium phosphate (KH2PO4)	Gibco, 10010-023
Ethylenediaminetetraacetic acid (EDTA)	Two amine groups and two carboxyl group-use PBS as base buffer but EDTA helps to reduce cell clumping	SIGMA, E6758
Bovine serum albumin	Protein widely used in lab buffers as a stabilizer (flow cytometry-buffer (reduces nonspecific binding)	Fisher Scientific™, BP9703-100
Fetal Bovine serum	Proteins, growth factors, hormones, vitamins, lipids, carbohydrates, inorganic salts and trace elements	Gibco 16000044
Trypan blue	Azo dye of ~960 Da	Invitrogen, T10282
Digestion neutralization buffer	Cold PBS with 2 mM EDTA and 4% BSA	NIL
Stain buffer	0.2% BSA in PBS (Keep cold)	NIL

Figure 3.

Representative graphs of the gating strategy used. A: cells were gated to exclude debris by plotting cell size (forward scatter area, FSC-A) versus granularity (side scatter area, SSC-A); B: doublets were excluded (green color) and singlets chosen to move forward (FSC-A vs FSC-H); C: Live singlets were gated for CD45⁺ leukocytes; D: Lymphocytes were excluded by gating for B cells (BV220) versus T cells (CD3), non T- and B-cell populations (non TB) were chosen to move forward; E: neutrophils were identified as CD45⁺ Ly6G^high CD11⁺; F: monocytes were defined as CD45⁺ CD11b⁺ Ly6C⁺ F4/80⁺ and macrophages as CD45⁺ CD11b⁺ Ly6C⁻ F4/80⁺; G: macrophages were separated into CD86⁺ (M1-like) and CD163⁺ (M2-like).

Results

The enumeration of leukocytes in the LV was accomplished after a thorough optimization of all parameters related to the flow cytometry workflow. We started by optimizing tissue digestion, followed by titration of the L/D dye to ensure efficient recovery of viable cells. In optimal conditions the optimized protocol yielded between 2 to 2.5 million cells from 75–100 mg of LV tissue. With this method, the viability of these cells consistently ranged between 85% to 90%. On average, we obtained from each infarcted LV >23,000 live CD45⁺ cells. The average for each leukocyte subset was 5,895 monocytes; 4,284 neutrophils; and 1,253 macrophages.

To optimize tissue digestion with a high yield of viable cells, we evaluated two concentrations of collagenase II (0.5 and 1 mg/mL) and tested five incubation times for digestion (5, 10, 15, 20, and 25 minutes). Additionally, we tested two formulas for the neutralization buffer. As seen in Figure 4, higher incubation times drastically reduced the cell viability. We obtained the highest yield of live cells with 0.5 mg/mL collagenase digestion in RPMI-1640 for 5 min. Based on standard tissue digestion protocols for immune cell isolation, RPMI-1640 is preferred over HEPES as a base for collagenase II digestion of heart tissue because RPMI provides a nutrient-rich, pH-stabilized, and osmotic-compatible environment necessary for cell survival during digestion times (see Box 3), achieving a cell population with over 70% live cells (Figure 4, Condition 13). From the neutralization buffers tested, we achieved the highest cell viability when using cold PBS with 2 mM EDTA and 4% BSA.

Figure 4.

Overall viability of live cell percentile among the tested conditions. Numbers within circles refer to duration of digestion in minutes. Variables included enzyme concentration, composition of neutralization buffer, and duration of digestion. Condition 13 resulted in the highest yield of live cells and was chosen as the optimized condition. Conditions for digestion groups: #1= 1mg/mL collagenase in RPMI 25min digestion. Neutralization buffer: PBS+ 2mM EDTA + 2% FBS added after digest was strained, 10% live cells; #2 = 1mg/mL collagenase in RPMI, 20–25min digest. Neutralization buffer: PBS+ 2mM EDTA + 2% FBS added before digest was strained,12% live cells; #3 = 1mg/mL collagenase in RPMI, 25min digest. Neutralization buffer: PBS+ 2mM EDTA + 2% BSA added after digest was strained, 14% live cells; #4 = 1mg/mL collagenase in RPMI, 20–25min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA added before digest was strained, 8% live cells; #5 = 0.5mg/mL collagenase in RPMI, 20min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 8% live cells; #6 = 1mg/mL collagenase in RPMI, 20min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 12% live cells; #7 = 1mg/mL collagenase in RPMI, 20min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% FBS, 15.5% live cells; #8 = 1mL of 0.5mg/mL collagenase in RPMI, 5min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 50% live cells; #9 = 1mL of 0.5mg/mL collagenase in RPMI, 10min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 48% live cells; #10 = 1mL of 0.5mg/mL collagenase in RPMI, 15min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 61% live cells; #11 = 1mL of 0.5mg/mL collagenase in RPMI, 20min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 66% live cells; #12 = 1 mL of 0.5mg/mL collagenase in RPMI for each 10mg tissue, 5min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 73% live cells; #13 = 3 mL of 0.5mg/mL collagenase in RPMI, 5min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 75% live cells; #14 = 3 mL of 0.5mg/mL collagenase in RPMI, 10min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 44% live cells; #15 = 3 mL of 0.5mg/mL collagenase in RPMI, 15min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 58% live cells; and #16 = 3 mL of 0.5mg/mL collagenase in RPMI, 20min digest. Neutralization buffer: PBS+ 2mM EDTA + 4% BSA, 64% live cells.

Box 3-. RPMI Versus HEPES, Differences at a glance.

Feature	RPMI (Culture Medium)	HEPES (Buffering Agent)
Nature	Complete cell culture medium	Chemical buffer
Role	Supports cell growth, survival, and function	Stabilizes pH in culture media
Composition	Amino acids, vitamins, glucose, salts (+ serum optional)	Zwitterionic buffering compound
Buffering System	Bicarbonate-based (CO2-dependent)	CO2-independent buffering system
pH Control	Maintains ~7.2–7.4 (requires CO2 incubator)	Maintains ~7.2–7.6 (stable outside incubator)
CO2 Requirement	Requires ~5% CO2	No CO2 required
Primary Use	Routine cell culture and expansion	Prevents pH drift during handling
Use Outside Incubator	Limited (pH can drift)	Highly effective
Nutritional Value	Provides essential nutrients	No nutrients
Incubation time	Short	Long

Then, we evaluated multiple concentrations of L/D dye; the 1:4,000 dilution yielded the most effective results (Figure 5). This dilution effectively differentiated live from dead cells, aligning with our goal of achieving optimal separation while minimizing non-specific binding. As seen in Figure 5, the highest dilutions (1:8000, 1:16000, and 1:32000) did not yield good separation between live/dead cells. The lowest dilutions (i.e., more concentrated dye), though, presented good separation. We choose 1:4000 dilution as the optimized condition, instead of 1:1000 and 1:2000, for two reasons: first, the highest dilution allows for cost savings, and second, in the lowest dilutions (top panel left and middle) there is a fairly populated region extending from the negative population (i.e., live cells) toward the dead population, which suggests excess dye concentration and non-specific membrane binding. The 1:4000 selection guided the parameters for subsequent experiments, ensuring consistent and accurate data collection.

Figure 5.

Live/ Dead Aqua titration results demonstrate that dilution 1:4000 provided an optimal concentration.

The precise process of antibody titration is indispensable for panel optimization, ensuring unparalleled resolution by pinpointing the optimal antibody concentration. This concentration is key for achieving peak fluorescence of the target cells while concurrently suppressing non-targeted background staining, thus significantly improving the signal-to-noise ratio. Such precision not only heightens the quality of data but also economizes on reagents and costs. The third step on protocol optimization was antibody titration, as exemplified in Figure 6, and the resulting optimized flow protocol was used for leukocyte enumeration and cell sorting. The optimum antibody concentration in flow cytometry is the lowest concentration that maximizes positive–negative population separation (highest stain index) while minimizing background, spread, and fluorochrome spillover. For example, for CD86 and F4/80 antibodies we observed good separation at the highest % from parent population (Figure 6). While LY6C during titration showed continuous and similar expression across the different concentrations tested; therefore, we chose the lowest concentration to minimize non-specific binding. For CD11b all concentrations tested showed good separation and stain index, we chose one of the higher concentrations to account for experimental setting where we would encounter higher cell number. Figure 6 display examples of antibody titration and Table 2 list the optimized concentration used for each antibody.

Figure 6.

Examples of antibody titration for (A) PE-Cy7 for B220 and SB-436 for LyG; (B) PE for CD86 and APC for CD11b; and (C) Alexa 488 for F4/80 and PE-CF594 for Ly6C.

In the process of immunophenotyping leukocytes within the LV region, we performed permanent occlusion of the left descending coronary artery on both male and female mice, and harvested the heart at day 3 (D3) and day 5 (D5) post-MI. These timepoints are known for presenting peak levels of neutrophils and macrophages post-MI in rodents. Initially, there is a surge in CD45+ immune cells infiltrating the heart tissue, reflecting the inflammatory phase where neutrophils and monocytes/macrophages play a vital role in responding to the injury. First, we determined the single cell populations based on their FSC and SSC plots. The further gating details provided us the live CD45+ immune cell information, with APC-Cy fluorochrome against L/D Aqua V525 in the cell suspension (Figure 7A) revealed significant difference based on day when compared in both males and females administered saline post-MI. This was followed by gating for neutrophils and macrophages (Figure 7B–C). From our result, both males and females showed no significant difference in the number of neutrophils and macrophages at both D3 and D5.

Figure 7.

CD45+, macrophage and neutrophil populations from the heart (LV) post-MI. Unpaired t-test was used to compare the macrophages and neutrophil populations while two-way ANOVA was used for CD45+ following Tukey’s post hoc test, p ≥ 0.05.

Discussion

The enumeration of leukocytes in the heart after an MI is a cornerstone of cardiovascular immunology research, and its scientific importance extends across mechanistic understanding, therapeutic development, and translational relevance (23,24). MI initiates a highly orchestrated immune response essential for debris clearance, scar formation, and remodeling (25,26). Enumerating leukocyte subsets (e.g., neutrophils, monocytes/macrophages, T cells, B cells) reveals temporal changes in the inflammatory cascade: 0–3 days: Neutrophil dominance (acute inflammation, proteolysis); 3–7 days: Monocyte/macrophage infiltration (clearance, transition to repair); >7 days: Lymphocyte involvement (resolution, fibrosis modulation). Quantifying these cells helps define distinct phases of inflammation — a prerequisite for identifying therapeutic windows. The balance between pro-inflammatory (Ly6C⁺, CD86⁺) and pro-reparative (CD206⁺, CD163⁺) macrophages determines whether healing is adaptive (scar stabilization) or maladaptive (fibrosis and failure) (27,28). Quantification allows correlation of immune composition with LV function (Left ventricular ejection fraction, LVEF; and fraction shortening, FS), scar size and fibrosis, ventricular dilation and remodeling (29–31). Thus, immune profiling becomes a predictive biomarker of post-MI outcomes. Our results and detailed methods demonstrate adequate optimization of the flow cytometry protocol using a nine-antibody panel for leukocyte identification. Tissue digestion was accomplished with collagenase type II. Collagenase type II is generally preferred for cardiac tissue dissociation because it has a higher tryptic activity alongside its collagenolytic activity (32), making it more effective at breaking down the fibrous extracellular matrix (ECM) of well-perfused tissues like the heart, ensuring a good balance between digestion strength and cell viability. The heart is rich in type I and type III collagen arranged in a tightly cross-linked network (33); the broader protease activity of collagenase type II helps disrupt cell-matrix adhesions, yielding higher numbers of viable cells. Type I collagenase can also be used to dissociate tissues; however, collagenase type I has relatively lower protease activity making it less efficient for dense fibrous heart tissue (32,33), leading to incomplete digestion or lower cell yields. Other enzymes, such as hyaluronidase and DNase can be used as supplementary enzymes to digest hyaluronic acid or free extracellular DNA released from dying cells. Hyaluronidase can improve enzymatic access by breaking down the pericellular matrix around interstitial cells (34). If the goal of the research is to isolate cardiomyocytes, fibroblasts, and/or endothelial cells, then hyaluronidase supplementation would be beneficial. Similarly, it may need to be considered the addition of DNAse if the protocol goal is to isolate cardiomyocytes or interstitial cells. Free DNA is highly sticky and causes cell clumping, which reduces yield and skews flow cytometry or single-cell sequencing results (35). DNase I is therefore a protective additive, and it is especially valuable in cardiac protocols where prolonged digestion causes cell death and DNA release. We accomplished better immune cell isolation with shorter digestion times, compared to HEPES longer incubations, and as such did not need to include DNase. Box 4 provides a troubleshooting guide for immune cell isolation for flow cytometry.

Box 4-. Troubleshooting guide.

1). Low cell viability:

Keep cells cold and minimize total handling time, use gentler centrifugation and avoid excessive spin force, reduce harsh enzymatic or mechanical dissociation when possible, check whether fixation or permeabilization conditions are too harsh, include a viability stain so dead cells can be excluded during gating. Likely causes: prolonged processing, rough pipetting, over-digestion, delayed acquisition.

2). Cell clumping:

Filter the sample before acquisition with an appropriate cell strainer, add EDTA to the buffer if compatible with the assay, keep cells in single-cell suspension and resuspend gently, consider DNase if free DNA from dead cells is promoting aggregation, avoid overloading the cytometer; dilute dense samples if needed with strain buffer. Likely causes: sticky cells, dead-cell DNA, incomplete dissociation, or high cell density.

3). Low yield:

Pre-wet cell strainers and use appropriate pore size, reduce sample transfers and wash only as much as needed, confirm that the starting material was sufficiently dissociated, check whether cells are being lost in clumps, debris, or filter material, adjust sample concentration to an acquisition-friendly range. Likely causes: filter-related cell retention, cell loss during prep, clumping, poor recovery after dissociation, or instrument settings that suppress events.

4). High debris background:

Minimize mechanical stress during dissociation, include viability dyes and perform dead-cell exclusion gating. Likely causes: Excess tissue damage, dead cell contamination.

5). Weak fluorescence signal:

Optimize antibody titration, protect sample from prolonged light exposure, optimize digestion conditions. Likely causes: Insufficient antibody concentration, fluorophore degradation and poor antigen preservation

6). Poor population resolution:

Run single–stain compensation controls, select compactible fluorophore panels with minimal overlap, perform routine cytometer calibration and quality control. Likely causes: Incorrect compensation settings, spectra overlap between fluorophores, instrument instability.

Antibody titration was performed to determine optimal concentrations that maximized signal-to-noise ratios while minimizing nonspecific binding and fluorochrome spillover. Compensation controls were generated using single-stained compensation beads for each fluorochrome, enabling accurate correction of spectral overlaps across channels. This systematic optimization ensured reliable leukocyte discrimination, consistent gating, and reproducible quantification. Minimizing tissue digestion time, this protocol limits proteolytic cleavage and enzymatic stress that can compromise cell membranes and degrade surface antigens, which is particularly critical in inflamed post-MI myocardium. Optimization of enzyme enables efficient extracellular matrix breakdown while preventing excessive cell damage. Gentle mechanical dissociation further reduces shear stress–induced cell death and preserves fragile leukocyte populations. Collectively, these optimizations maintain high cell viability and intact antigen expression, thereby enabling accurate enumeration and reliable phenotypic characterization of diverse left ventricular leukocyte subsets by flow cytometry. Dyes such as L/D aqua are referred to as amine reactive dyes, since they bind to any available protein (36). Therefore, it is of utmost importance to use protein-free buffers in this assay to prevent nonspecific binding. While the dye does interact with the surface proteins of live cells, its binding to dead cells is exponentially greater due to their porous nature, which exposes a larger array of proteins. From the six dilutions tested, we identified 1:4000 as having the optimal results.

Antibody titration is an essential preliminary step in flow cytometry, integral to optimizing signal specificity and minimizing background noise for clear delineation of positive and negative populations. This process enhances the reproducibility and quantitative accuracy of the results. By establishing the minimal, yet effective, antibody concentration, titration mitigates non-specific binding and enhances cell population resolution, facilitating reliable and comprehensive data collection essential for advancing research findings (12,37). Optimal antibody concentration is key for achieving peak fluorescence of the target cells while concurrently suppressing non-targeted background staining, thus significantly improving the signal-to-noise ratio. Such precision not only heightens the quality of data but also economizes on reagents and costs (38). FACS as a specialized form of flow cytometry not only identifies but also physically isolates single cells based on fluorescently labeled surface or intracellular markers (3,39). Isotope controls are ideal for finding the correct amount of blocking reagent, and fluorescence minus one (FMO) controls are great for gating control; however, the use of FMO has some limitations. In our case, we have limited tissue supply (maximum of 75 g from which most of the cell population is cardiomyocytes). Including FMO controls for a panel of 10 fluorochromes would significantly reduce our source material. While we did not perform FMO or isotope controls, we included unstained controls and single-color (i.e., compensation) controls. The first measures natural cell autofluorescence, such as in the case of F4/80+ cells, and the second calculates spillover. Comparison of the unstained control with the single-color controls allowed us to check for non-specific binding (i.e., optimization of the Fc Block) and to identify degradation or uncoupling of tandem dyes (like PE-Cy7 or APC-Cy7). Nonetheless, we acknowledge that inclusion of FMO controls is recommended when necessary to resolve ambiguous or dim populations and when fluorescence spread may affect gate placement, especially for secondary expression, like CD45RA, which is not present in this protocol.

Previous protocols have described immune cell isolation and flow cytometric characterization in murine lung, retina, and aortic tissues (15,16,40). The majority of the reports that used cardiac tissues, are either focused on cardiomyocyte and/or fibroblast isolation (41,42). These methods are often tissue-specific and result in variable cell yield and viability. A landmark paper that studied monocyte populations after MI, used spleen, blood, and heart tissues; however, a detailed protocol was not provided (43). Our literature search did not identify a detailed cardiac immune cell isolation protocol that provided step by step instructions to quantify and isolate cardiac immune cells. While several manuscripts described general steps, step optimization, doses, incubation times, and antibody concentrations were often not present (44,45). The present method builds upon these approaches by optimizing enzymatic digestion and staining conditions, including an empirically determined Live/Dead Aqua dilution (1:4000) that maximizes separation index in murine heart, thereby improving cell discrimination and overall data quality. Our result also indicate that overall leukocyte recruitment to the infarcted LV is comparable between sexes following saline treatment, as reflected by similar numbers of CD45⁺ immune cells, neutrophils, and macrophages at both D3 and D5 post-MI. This suggests that early mononuclear cell infiltration and macrophage accumulation during the inflammatory and reparative phases of MI are not sex-dependent. However, an increased n number would be necessary to confirm this. Overall, this protocol demonstrated that flow cytometry enables high-resolution dissection of the immune landscape following MI/heart failure (11).

Optimized Protocol for Flow Cytometry of Cardiac Leukocytes

Following optimization of all steps mentioned above, we generated a comprehensive protocol for enumeration and isolation of immune cardiac cells as follows:

Procedure:

• 1Anesthetize the animal per AUP and harvest the heart.
• 2Perfuse the heart using retrograde perfusion via the aorta with 2–3 mL of cold PBS using a 3 mL syringe fitted with a 27½G needle.
• 3Carefully remove the connective tissues around the heart and, if needed, separate the LV from the right ventricle.
• 4Weigh the tissue(s) and record the weight. If the tissue weight is <75 mg, pool tissue from more than one animal to obtain sufficient material. This protocol is optimized for 75–100 mg LV.
• 5Transfer the tissue to a petri dish kept on ice. Add a minimal volume of cold PBS (~100 μL), just enough to keep tissue moist. Using scissors pre-chilled on ice, mince the tissue rapidly into fine pieces while keeping it cold throughout the process. Avoid over- mincing, as this increases cell death and reduces viable cell yield.
• 6Add 8 mL of digestion buffer, then transfer the minced tissue in buffer to a 50 mL falcon tube. Incubate at 37°C on a rotator for 5 minutes at 10 – 20 rpm.
• 7Once the tissue is digested, add 10 mL of cold digestion neutralization buffer.
• 8Filter the cell suspension through a 35 μm cell strainer into a new 50 mL falcon tube to remove undigested debris.
• 9
  Count cells either manually using a hemocytometer or with an automated cell counter.

  1. In a microcentrifuge tube, mix 10 μL of cell suspension with 10 μL of 0.4% trypan blue stain.
  2. Gently pipette up and down several times to mix thoroughly, avoiding bubble formation.
  3. Load the mixture onto a hemocytometer (if counting manually) and quantify live (trypan blue–negative) and dead (trypan blue–positive) cells. Record total cell number and viability.
• 10Centrifuge the cell suspension at 400 × g for 5 min at 4°C.
• 11Carefully aspirate and discard the supernatant without disturbing the cell pellet.
• 12Resuspend the pellet in 3 mL of cold PBS and briefly vortex or gently pipette to mix.
• 13Repeat the wash by centrifuging again at 400 × g for 5 minutes at 4 °C. Carefully remove and discard the supernatant. The final cell pellet should be resuspended in approximately 200 μL of PBS (or appropriate buffer) for downstream applications.
• 14
  Collect the cells in 200 μL of cold stain buffer. If needed, refilter the suspension. Mix the cell suspension (~400 μL total volume) and distribute into 3 FACS tubes.

  1. #1 FACS tube (no stain): 50 μL of cells for unstained cell tube (apply only the washing steps; steps 15 and 26–28)
  2. #2 FACS tube (L/D single stain): 50 μL of cells for L/D single stain tube (apply only washing steps and add L/D aqua at the end; steps 15 and 26–29)
  3. #3 FACS tube (all staining): 300 μL of cells for combined antibodies tube
• 15Add 3 mL of cold stain buffer (0.2% BSA/ PBS) to each tube, quickly vortex. Keep tubes #1 and #2 on ice.
• 16Tube #3: Centrifuge at 400 × g for 5 min at 4°C. Do not shake or disturb the pellet. Remove supernatant by gently decanting it and absorbing it with a Kim wipe.
• 17Resuspend in 100 μL of stain buffer and vortex.
• 18Add 20 μL Brilliant stain buffer (shake before use). Add FC block and True-Stain Monocyte Blocker (Use 5 μL of each to the 100 μL volume).
• 19Incubate for 15 min on ice protected from light.
• 20
  Prepare single-stain bead tubes while waiting for incubation. This is done only once for each run, meaning if you have 4 samples you will need tubes #1–3 for each sample (n= 12 tubes). You will only need one additional tube for each antibody tested. In this protocol we tested 9 antibodies; thus, we needed 9 single stain bead tubes (tubes #4–12).

  1. Mix the beads tube very well before using (important)
  2. Add 5–6 drops of ultracomp beads into a microtube. Measure the volume, which should be approximately 480 μL.
  3. Add cold stain buffer up to 920 μL total volume. Aliquot 100 μL into the 9 FACS tubes (keep on ice and protect from light).
• 21
  Add each Ab according to the concentrations listed in Table 2:

  1. For combined antibodies tube (#3 FACS tube) add all (total vol of 9 antibodies = 19.125 μL)
  2. For the individual single-stain bead tube (#4 to #12 FACS tubes) add the appropriate Ab

*Important - Each Ab needs to be added first to the combined tube #3 and immediately to the appropriate individual single-stained bead tube (for ex. add 2.5 μL of Alexa 488 to Tube #3 and to tube #4; then 2.5 μL of PE to tube #3 followed by 2.5 μL to tube #5 to ensure equal time of staining for each Ab.

• 22To tube #3, add 51 μL of stain buffer to make the final volume 200 μL
• 23Incubate Ab tubes (#3-#12) on ice for 20 minutes protected from light.
• 24Add 3 ml of cold stain buffer, and quickly vortex.
• 25After this step, continue the protocol with all tubes (#1 to #12)
• 26Centrifuge at 400 × g for 5 minutes at 4°C.
• 27Do not shake or disturb the pellet. Remove the supernatant by gently decanting it and absorbing with Kim wipe. Wash again and decant supernatant as described.
• 28Resuspend in 200 μL of stain buffer.
• 29This step is only for tubes #2 and #3: Add 1:4000 L/D aqua solution (i.e., 0.05 μL for a 200 μL volume). Note: We store the original L/D aqua solution as 2 μL aliquots; before use, the aliquots are diluted with 38μL of cold PBS (1:20 dilution); 1 μL of this dilution is added to the tubes to reach a 1:4000 dilution.
• 30Run in the flow cytometer and/or sort (if needed).

Conclusion

The optimized protocol for cardiac leukocyte isolation and flow cytometric analysis described here provides a reliable, reproducible, and high-yield approach to study immune cell dynamics in the injured heart. By minimizing tissue digestion time, optimizing enzyme concentration, and ensuring gentle mechanical dissociation, this method preserves both cell viability and antigen integrity, allowing accurate enumeration and phenotypic characterization of diverse leukocyte subsets. Importantly, coupling this isolation technique with FACS enables downstream applications (Figure 8) such as transcriptomic, metabolic, and functional assays on purified immune populations. This workflow thus offers a robust platform for investigating the temporal and spatial immune responses following MI, heart failure, or other cardiac injuries, and can be extended to cardiorenal syndrome models to explore systemic immune crosstalk. Overall, the optimized protocol facilitates precise quantification and mechanistic exploration of cardiac immune driven remodeling, supporting the development of targeted immunomodulatory and treatment interventions aimed at improving cardiac repair and preventing secondary organ dysfunction.

Figure 8.

Sorted cells indicating their use for downstream gene expression analysis