This protocol outlines two methods for the quantitative analysis of mitophagy in pancreatic β-cells: first, a combination of cell-permeable mitochondria-specific dyes, and second, a genetically encoded mitophagy reporter. These two techniques are complementary and can be deployed based on specific needs, allowing for flexibility and precision in quantitatively addressing mitochondrial quality control.
Mitophagy is a quality control mechanism necessary to maintain optimal mitochondrial function. Dysfunctional β-cell mitophagy results in insufficient insulin release. Advanced quantitative assessments of mitophagy often require the use of genetic reporters. The mt-Keima mouse model, which expresses a mitochondria-targeted pH-sensitive dual-excitation ratiometric probe for quantifying mitophagy via flow cytometry, has been optimized in β-cells. The ratio of acidic-to-neutral mt-Keima wavelength emissions can be used to robustly quantify mitophagy. However, using genetic mitophagy reporters can be challenging when working with complex genetic mouse models or difficult-to-transfect cells, such as primary human islets. This protocol describes a novel complementary dye-based method to quantify β-cell mitophagy in primary islets using MtPhagy. MtPhagy is a pH-sensitive, cell-permeable dye that accumulates in the mitochondria and increases its fluorescence intensity when mitochondria are in low pH environments, such as lysosomes during mitophagy. By combining the MtPhagy dye with Fluozin-3-AM, a Zn2+ indicator that selects for β-cells, and Tetramethylrhodamine, ethyl ester (TMRE) to assess mitochondrial membrane potential, mitophagy flux can be quantified specifically in β-cells via flow cytometry. These two approaches are highly complementary, allowing for flexibility and precision in assessing mitochondrial quality control in numerous β-cell models.
Pancreatic β-cells produce and secrete insulin to meet metabolic demands, and β-cell dysfunction is responsible for hyperglycemia and diabetes onset in both type 1 and type 2 diabetes. β-Cells couple glucose metabolism with insulin secretion via mitochondrial energetics and metabolic output, which depend on a reserve of functional mitochondrial mass1,2,3. To maintain optimal β-cell function, β-cells rely on mitochondrial quality control mechanisms to remove aged or damaged mitochondria and preserve functional mitochondrial mass4. Selective mitochondrial autophagy, also known as mitophagy, is a key component of the mitochondrial quality control pathway.
Assessments of mitophagy in live cells often rely on changes in mitochondrial pH that occur during mitophagy. Mitochondria have a slightly alkaline pH, and healthy mitochondria normally reside in the pH-neutral cytosol. During mitophagy, damaged or dysfunctional mitochondria are selectively incorporated into autophagosomes and eventually cleared within acidic lysosomes5. Several in vivo transgenic mitophagy reporter mouse models, such as mt-Keima6, mitoQC7, and CMMR8, as well as transfectable mitophagy probes, such as the Cox8-EGFP-mCherry plasmid9, utilize this pH change to provide quantitative assessments of mitophagy. Use of transgenic mice expressing the mt-Keima pH-sensitive dual-excitation ratiometric probe has been optimized for mitophagy assessments in islets and β-cells via flow cytometry10,11. The ratio of acidic-to-neutral mt-Keima wavelength emissions (the ratio of acidic 561 nm to neutral 480 nm excitation) can be used to robustly quantify mitophagy6,12.
This protocol describes an optimized approach to assess mitophagy flux in primary islets and β-cells isolated from mt-Keima transgenic mice10,11. While mt-Keima is a highly sensitive probe, it requires either complicated animal breeding schemes or the transfection of cells, which can often be challenging when working in combination with other genetic models or with primary human islets. Additionally, the use of multiple fluorescence lasers and detectors to identify neutral and acidic cell populations can limit the combinatorial use of other fluorescent reporters.
To overcome these challenges, this protocol also describes a complementary, single fluorescent channel, dye-based method for robust detection of mitophagy in β-cells from isolated mouse islets. This approach, referred to as the MtPhagy method, utilizes a combination of three cell-permeable dyes to select for β-cells, quantify the cell populations actively undergoing mitophagy, and assess mitochondrial membrane potential (MMP or Δψm) simultaneously.
The first of these dyes is Fluozin-3-AM, a cell-permeable Zn2+ indicator with an Ex/Em 494/516 nm13. Mouse islets comprise a heterogeneous population of functionally distinct cells, including α-, β-, δ-, and PP cells. β-Cells comprise approximately 80% of cells within the mouse islet and can be distinguished from other islet cell types due to their high Zn2+ concentration within insulin granules14,15, allowing for identification of β-cells as the Fluozin-3-AMhigh population. The MtPhagy dye, a pH-sensitive dye that is immobilized on mitochondria via a chemical bond and emits weak fluorescence, is also utilized in this protocol16. Upon mitophagy induction, damaged mitochondria are incorporated into the acidic lysosome, and the MtPhagy dye increases its fluorescence intensity within the low pH environment (Ex/Em 561/570-700 nm).
Additionally, Tetramethylrhodamine, ethyl ester (TMRE), is used to assess MMP. TMRE is a cell-permeable positively charged dye (Ex/Em 552/575 nm) that is sequestered by healthy mitochondria due to the relative negative charge upheld by their membrane potential17. Damaged or unhealthy mitochondria dissipate their membrane potential, resulting in decreased ability to sequester TMRE. Utilizing these dyes together, β-cells undergoing mitophagy can be identified as the FluozinhighMtPhagyhighTMRElow population via flow cytometry. Since mitophagy is a dynamic rather than static process, this protocol was optimized to assess mitophagy flux using valinomycin, a K+-ionophore that induces mitophagy following dissipation of MMP18. Comparison of mitophagy in the presence and absence of valinomycin allows for assessment of mitophagy flux in different sample groups.
The dye-based nature of the current approach allows it to be extrapolated to human islets and other difficult-to-transfect cell types and circumvents the need for complicated animal breeding schemes, unlike the mt-Keima protocol. The overarching goal of this protocol is to quantify mitophagy in β-cells at the single-cell level via two independent flow cytometry-based methods. Taken together, this protocol describes two powerful and complementary methods that allow for both precision and flexibility in the quantitative study of mitochondrial quality control.
The animal studies presented in this protocol were reviewed and approved by the University of Michigan Institutional Animal Care and Use Committee. Twenty-week-old male C57BL/6J mice, on either a 15-week regular fat diet (RFD) or high-fat diet (HFD), were used for this study.
1. Assessing mitophagy via the dye-based MtPhagy approach (Method 1)
2. Assessing mitophagy using the genetically encoded mt-Keima reporter (Method 2)
Assessing mitophagy via the dye-based MtPhagy approach
This dye-based approach was optimized to analyze mitophagy flux within primary mouse β-cells without the need for a genetic reporter, using Fluozin-3-AM, TMRE, and MtPhagy as well as DAPI to exclude dead cells. By pairing these dyes with valinomycin to induce mitophagy, this protocol outlines a dye-based method to selectively measure mitophagy flux in primary mouse β-cells18. For the data shown using this MtPhagy method, both basal and valinomycin-induced mitophagy were analyzed in islets isolated from either regular fat diet (RFD) or high fat diet (HFD, 60 kcal% Fat) fed mice to assess the effect of metabolic stress on mitophagy flux. To identify the population of interest, cells were gated using untreated RFD islets. FSC and SSC voltages were first adjusted to attain an even distribution of cells on a SSC-A vs. FSC-A plot (Figure 1A). To select for single cells, both FSC-H vs. FSC-W and SSC-H vs. SSC-W plots were used, where multiplets were excluded due to their higher width signal values compared to single cells (Figure 1B,C). Next, DAPI-negative cells were selected to exclude dead cells20 (Figure 1D). After establishing primary gates, single stained controls were utilized to establish fluorescence gates for Fluozin-3-AM, MtPhagy, and TMRE (Figure 1E–G) as well as compensation controls for multi-color fluorescence flow cytometry.
Once these primary and fluorescence gates were established, β-cells with high utilization of mitophagy were defined as the FluozinhighMtPhagyhighTMRElow population in quadrant 3 (Q3) using RFD without valinomycin exposure (Figure 1H). Using this gating strategy, basal and valinomycin-induced mitophagy levels were characterized in both RFD and HFD islets (Figure 2). To quantify mitophagy flux, basal vs. valinomycin-induced mitophagy levels were compared using the following ratio:
Using this ratio, mitophagy flux was quantified and compared in RFD vs. HFD β-cells to assess differences in mitophagy following the induction of obesity and peripheral insulin resistance. Quantification of mitophagy flux in RFD vs. HFD samples is shown in Figure 2E. This result highlights the feasibility of this assay to quantify mitophagy in β-cells using a straightforward dye-based approach. This method can also be employed in human islets, difficult-to-transfect cells, and islets isolated from complex genetic models where intercrossing to the mt-Keima transgenic model would be cumbersome.
Assessing mitophagy using the genetically encoded mt-Keima reporter
Mt-Keima is a dual excitation fluorescent protein fused with a Cox8-localization sequence that enables its targeting to the inner mitochondrial membrane. The bimodal fluorescent property of mt-Keima allows it to switch its excitation spectra from the neutral (405 nm) to acidic (561 nm) wavelength, depending on the pH of the intracellular compartment6. This enables a robust ratiometric fluorescence analysis of mitophagy, where an increase in acidic-to-neutral ratio indicates mitophagy induction. In this protocol, Fluozin-3-AM was also used to select for β-cells via flow cytometry. In these representative studies, mitophagy flux was assessed using islets isolated from mice fed a RFD diet10,11. FSC and SSC voltages were first adjusted to attain an even distribution of cells on a SSC-A vs. FSC-A plot (Figure 3A). To select for single cells, both FSC-H vs. FSC-W and SSC-H vs. SSC-W plots were used, where multiplets were excluded due to their higher width signal values compared to single cells (Figure 3B,C). The voltage and gating strategy for DAPI and Fluozin-3-AM were determined using single-stained islets (Figure 3D,E). Triangle gates for the acidic and neutral populations were then identified using the mt-Keima positive sample without valinomycin exposure (Figure 3F).
Once these primary and fluorescence gates were established, mitophagy flux was assessed using basal and valinomycin-induced changes in mt-Keima fluorescence (Figure 3F,G). To quantify mitophagy flux, basal mitophagy vs. valinomycin-induced levels were compared using the following ratio:
Using this ratio, mitophagy flux was quantified in RFD cells. Quantitation of this result is shown in Figure 3H. Importantly, these results are comparable to the results in RFD islets generated using the MtPhagy approach (Figure 3H).
Figure 1: Gating scheme for the MtPhagy method. (A) Flow plot displaying gating scheme to select for all cells. (B) Gating to select for singlets based on FSC-H vs. FSC-W and (C) SSC-H vs. SSC-W. (D) Gating for DAPI–negative cells to exclude dead cells. (E) Gating for Fluozin-3-AMhigh cells to select for β-cells. (F) Gating scheme for MtPhagy dye to identify MtPhagyhigh and MtPhagylow cell populations. (G) Gating scheme for TMRE to identify TMREhigh and TMRElow cell populations. (H) Quadrant gating scheme established with untreated RFD islets to identify FluozinhighMtPhagyhighTMRElow cells in quadrant 3 (Q3) as β-cells undergoing mitophagy. Please click here to view a larger version of this figure.
Figure 2: Assessing mitophagy flux differences in mouse β-cells following metabolic stress using the MtPhagy gating scheme. Representative flow cytometry plots of (A) untreated RFD β-cells, (B) untreated HFD β-cells, (C) valinomycin-exposed RFD β-cells, and (D) valinomycin-exposed HFD β-cells. (E) Quantification of mitophagy flux in β-cells, calculated using a ratio of the MtPhagyhighTMRElow cells exposed to valinomycin to the MtPhagyhighTMRElow cells not exposed to valinomycin, for both RFD and HFD samples. *p < 0.05 by Student's unpaired t-test. n = 3/group. Please click here to view a larger version of this figure.
Figure 3: Gating scheme for the mt-Keima method and comparison between both methods. (A) Flow plot displaying gating scheme to select for all cells. (B) Gating to select for singlets based on FSC-H vs. FSC-W and (C) SSC-H vs. SSC-W. (D) Gating for DAPI–negative cells to exclude dead cells. (E) Gating for Fluozin-3-AMhigh cells to select for β-cells. (F) Representative flow cytometry plots of mt-Keima/+ untreated cells and (G) mt-Keima/+ valinomycin-exposed cells. (H) Quantification of mitophagy flux in β-cells from RFD-fed mice, calculated using a ratio of the acidic/neutral cells exposed to valinomycin to ratio of the acidic/neutral cells not exposed to valinomycin using the mt-Keima method and compared to the MtPhagy method (data for MtPhagy protocol originally shown in Figure 2E). n = 3/group. Please click here to view a larger version of this figure.
This protocol described two complementary methods to quantify mitophagy flux in dissociated primary mouse islets. Using the mt-Keima method, an increase in mitophagy was quantified as an increased ratio of acidic (561 nm)/neutral (405 nm) cells, whereas in the MtPhagy method, increased mitophagy flux was quantified as an increase in the FluozinhighMtPhagyhighTMRElow cell population. These methods allow for rapid, quantitative, and β-cell-specific assessments of mitophagy flux.
Both methods are straightforward approaches. However, certain steps within this protocol are crucial for obtaining quality and reproducible results. These steps include: (1) proper islet dissociation to obtain a single cell suspension, but mild enough to ensure islet viability, (2) careful establishment of gating to correctly identify populations of interest, and (3) the objective quantification of flow cytometry data via software tools to ensure an unbiased assessment of mitophagy flux.
When selecting which method to use, the cell type and nature of the genetic models used should be considered. The mt-Keima method, either used in vivo or transfected in vitro, is a highly cited and well-regarded method for mitophagy assessment by flow cytometry or live cell imaging21. While the dye-based MtPhagy method is a newer approach compared to genetic reporters, there are instances when its use may be preferred over mt-Keima. Indeed, the MtPhagy method overcomes the need for transfection or complex breeding schemes, and MtPhagy staining takes only 30 min and is performed immediately prior to flow cytometry. The MtPhagy approach can be successfully employed in difficult-to-transfect primary human islet samples as well. This protocol, which relies on either pH-sensitive mitochondrial dyes or probes to directly measure mitophagy, is distinct from a previous approach from Mauro-Lizcano et. al that employed the membrane potential sensitive MitoTracker Deep Red dye and required use of mitophagy and lysosomal inhibitors to quantify mitophagy flux by flow cytometry22. As the Mauro-Lizcano et. al method has not been tested in islets, it is difficult to directly compare it to the efficacy of the MtPhagy or mt-Keima methods described here. Taken together, the combination of these methods provides in totality an increasing number of options to rigorously assess mitophagy flux in highly quantitative live single cell assays.
A drawback for both methods is their incompatibility with cell fixation. Although both methods are compatible with live cell imaging approaches, cell fixation interferes with the pH gradient of both MtPhagy and mt-Keima across the lysosomal membrane7,16. As an alternative, use of the mitoQC mitophagy reporter in fixed samples has been previously employed7. Additionally, a limitation for both methods is the need to dissociate islets prior to flow cytometry, which may impact cell viability. Therefore, it is critical to stain cells with DAPI to monitor cell viability following islet dissociation and ensure all samples are treated consistently. Following DAPI staining, samples had an average of 12.7% ± 7.4 dead cells (Figure 1D), indicating that >80% of cells in each sample could be used for analysis. Monitoring of cell viability at each stage (following islet isolation, culture, and then dissociation) may be additionally useful to obtain knowledge of cell survival at each step of the procedure. Variability in timing between islet isolation and single cell dissociation also may affect cell viability and results. Thus, it is recommended to remain consistent with timing across all experiments.
As mitochondrial quality control is critical to β-cell health and function, rigorous assessments of mitophagy using traditional biochemical or imaging approaches (including turnover of mitochondrial outer membrane proteins, mitochondrial localization to autophagosomes or lysosomes, and electron microscopy) can prove challenging and time consuming. Thus, the development of effective and robust mitophagy reporter systems is crucial. Both the mt-Keima and MtPhagy approach are efficient and allow for quantitative assessments of mitophagy flux. These two techniques allow for both flexibility and precision in addressing β-cell mitochondrial quality control and probing inter-organelle interactions.
The authors have nothing to disclose.
E.L-D. acknowledges support from the NIH (T32-AI007413 and T32-AG000114). SAS acknowledges support from the JDRF (COE-2019-861), the NIH (R01 DK135268, R01 DK108921, R01 DK135032, R01 DK136547, U01 DK127747), the Department of Veterans Affairs (I01 BX004444), the Brehm family, and the Anthony family.
Antibiotic-Antimycotic | Life Technologies | 15240-062 | |
Attune NxT Flow Cytometer | Thermofisher Scientific | A24858 | |
DAPI (4',6-Diamidino-2-Phenylindole, Dihydrochloride) | Thermofisher Scientific | D1306 | DAPI reconstituted in ddH2O to reach 0.2 µg/mL stock |
Dimethyl Sulfoxide | Sigma-Aldrich | 317275 | |
Fatty Acid Free heat shock BSA powder | Equitech | BAH66 | |
Fetal bovine serum | Gemini Bio | 900-108 | |
Fluozin-3AM | Thermofisher Scientific | F24195 | 100 μg Fluozin-3AM powder reconstituted in 51 μL DMSO and 51 μL Pluronic F-127 to reach 1 mM stock. |
Gibco RPMI 1640 Medium | Fisher Scientific | 11-875-093 | |
HEPES (1M) | Life Technologies | 15630-080 | |
MtPhagy dye | Dojindo | MT02-10 | 5 μg MtPhagy powder reconstituted with 50 μL DMSO to reach 100 μM stock. |
MtPhagy dye | Dojindo | MT02-10 | |
Penicillin-Streptomycin (100x) | Life Technologies | 15140-122 | 1x Solution used in procotol by diluting 1:10 in ddH2O |
Phosphate buffered saline, 10x | Fisher Scientific | BP399-20 | 1x Solution used in procotol by diluting 1:10 in ddH2O |
Sodium Pyruvate (100x) | Life Technologies | 11360-070 | 5 μg MtPhagy powder reconstituted with 50 μL DMSO to reach 100 μM stock. |
TMRE [Tetramethylrhodamine, ethyl ester, perchlorate] | Anaspec | AS-88061 | TMRE powder reconstituted in DMSO to reach 100 μM stock. |
Trypsin-EDTA (0.05%), phenol red | Thermofisher Scientific | 25300054 | |
Valinomycin | Sigma | V0627 | Valinomycin powder reconsituted in DMSO to reach 250 nM stock. |