Application of Flow Cytometric Analysis for Measuring Multiple Mitochondrial Parameters in 3D Brain Organoids

Summary

This protocol reports a unique method of using a streaming cytometer and multiple antibodies for simultaneous assessment of multiple mitochondrial functional parameters, including changes in mitochondrial volume, amounts of the mitochondrial respiratory chain (MRC) complex subunits, and mitochondrial DNA (mtDNA) replication.

Abstract

Mitochondrial dysfunction is a common primary or secondary contributor to many types of neurodegeneration, and changes in mitochondrial mass, mitochondrial respiratory chain (MRC) complexes, and mitochondrial DNA (mtDNA) copy number often feature in these processes. Human brain organoids derived from human induced pluripotent stem cells (iPSCs) recapitulate the brain's three-dimensional (3D) cytoarchitectural arrangement and offer the possibility to study disease mechanisms and screen new therapeutics in a complex human system. Here, we report a unique flow cytometry-based approach to measure multiple mitochondrial parameters in iPSC-derived cortical organoids. This report details a protocol for generating cortical brain organoids from iPSCs, single-cell dissociation of generated organoids, fixation, staining, and subsequent flow cytometric analysis to assess multiple mitochondrial parameters. Double staining with antibodies against the MRC complex subunit NADH: Ubiquinone Oxidoreductase Subunit B10 (NDUFB10) or mitochondrial transcription factor A (TFAM) together with voltage-dependent anion-selective channel 1 (VDAC 1) permits assessment of the amount of these proteins per mitochondrion. Since the quantity of TFAM corresponds to the amount of mtDNA, it provides an indirect estimation of the number of mtDNA copies per mitochondrial content. This entire procedure can be completed within a span of 2-3 h. Crucially, it allows for the concurrent quantification of multiple mitochondrial parameters, including both total and specific levels relative to the mitochondrial mass.

Introduction

Mitochondria are essential cellular organelles and the major site of the adenosine triphosphate (ATP) production. In addition to providing energy for cells, mitochondria also participate in multiple cellular processes, including cell information transmission, cell differentiation, and apoptosis, and have the ability to regulate cell growth and cell cycle. Changes in mitochondrial function have been identified in various neurodegenerative diseases, including Parkinson's disease (PD)^1,2, Alzheimer's disease (AD)³, and amyotrophic lateral sclerosis (ALS)⁴. Mitochondrial dysfunction plays a role in the aging process, accumulating somatic mtDNA mutations and declining respiratory chain function⁵.

Various types of mitochondrial dysfunction occur in neurodegeneration, and the ability to measure such changes is extremely useful when studying disease mechanisms and testing potential treatments. Furthermore, establishing suitable in vitro model systems that recapitulate disease in human brain cells is vital for better understanding disease mechanisms and developing new therapies. iPSCs from patients with neurodegenerative diseases have been used to generate diverse brain cells that manifest mitochondrial damage^6,7,8,9. The development of 3D brain organoids derived from iPSCs is a major step in disease modeling. These iPSC-derived brain organoids provide complexity and contain the patient´s own genetic background, thus providing a disease model that more accurately reflects pathology in the patient's brain.

While some research has been conducted on mitochondrial studies using iPSC-derived brain organoids^10,11,12, convenient and reliable techniques for determining multiple mitochondrial functional parameters in iPSC-derived brain organoids remain limited. Flow cytometry provides a powerful tool to measure mitochondrial parameters at the single-cell level, as we have demonstrated previously¹³. This study provides a detailed protocol for generating cortical organoids from iPSCs, combined with a novel flow cytometry-based approach to simultaneously measure multiple mitochondrial parameters, including mitochondrial mass, respiratory chain complex subunits, and mtDNA copy number (Figure 1). Importantly, by using mitochondrial mass as a denominator, these protocols allow one to measure both the total and specific levels per mitochondrial unit.

Protocol

1. Differentiation of iPSCs into cortical organoids

1. Preparation of matrix-coated plates
   1. Thaw the vial of commercially available basement membrane matrix on ice overnight. Dilute into 1:100 in cold Advanced Dulbecco's Modified Eagle's Medium/Ham's F12 DMEM/F12 (1% final concentration). Make aliquots and store them at -20 °C (see Table of Materials).
   2. Thaw the membrane matrix solution at 4 °C (keep it cold) and coat the required number of wells (1 mL for one well in a 6-well plate). Incubate at 37 °C for 1 h.
   3. Take the plate from the incubator and let it reach room temperature (RT) (leave it for an hour).
      NOTE: The plates can be stored for up to 2 weeks in the refrigerator (remember to take it out and warm the coated wells required for that day at RT until it is no longer cool to the touch before use). It is recommended to use the plates within 3 days. For long time storage, add 1 mL of Essential 8 Medium (see Table of Materials) to the coated plate to avoid the gel drying out.

2. Preparation Essential 8 (E8) medium for iPSC culture

1. In a sterile environment, prepare E8 Medium by combining E8 basal medium with E8 supplement (see Table of Materials). This can be stored at 4 °C for 2 weeks.
   NOTE: Thaw frozen E8 supplement at 4 °C overnight before preparing the complete medium. Thawing the frozen supplement at 37 °C is not recommended; use E8 Medium for 2 weeks. Prior to use, warm E8 Medium required for that day at RT until it is no longer cool to the touch.

3. Subculturing of iPSCs

1. Pre-warm matrix-coated plates in RT or incubator at 37 °C for 20-30 min. Pre-warm the amount of E8 Medium needed at RT.
2. Aspirate the culture medium with a pipette.
3. Rinse iPSCs (see Table of Materials) with Dulbecco's Phosphate-Buffered Saline (Ca²⁺/Mg²⁺ free) (DPBS-/-) (4 mL for one well in a 6-well plate).
4. Add Ethylenediaminetetraacetic acid (EDTA) (0.5 mM) (1 mL for one well in a 6-well plate). Incubate at 37 °C until the edges of the colonies begin to detach from the plate (usually 3-5 min).
5. Aspirate the dissociation solution.
6. Add pre-warmed E8 Medium (4 mL for one well in a 6-well plate) and use high pressure to detach iPSC colonies.
7. Transfer into two different wells in a matrix-coated 6-well plate (2 mL for one well in a 6-well plate) and incubate at 37 °C.
   NOTE: Gently shake before putting the plate into the incubator. The split ratio can be 1:2-1:4.
8. Exchange media daily till the colonies get 80% confluence with good size and connections.
   NOTE: Monitor for colonies of an appropriate size, typically characterized by a well-defined, circular shape and a diameter ranging from 0.5 to 1 mm. The colonies should exhibit dense, compact structures with clearly delineated, smooth borders.

4. Embryoid body (EB) formation and neural induction

1. Prepare neural induction medium (NIM) by combining the components listed in the Table of Materials.
2. Aspirate medium and rinse the cells with DPBS -/- (4 mL per well in a 6-well plate).
3. Add pre-warmed Accutase (1 mL per well in a 6-well plate) (see Table of Materials) in the well and incubate at 37 °C for 10 min.
4. Gently pipette the solution up and down 3 to 4 times using a 1000 µL pipette tip.
5. Neutralize with 2 mL DMEM with 10% fetal bovine serum (FBS) and centrifuge the 15 mL conical tube containing the cells at 600 x g for 5 min at RT. Aspirate the supernatant.
6. Resuspend the cell pellets by gently pipetting up and down till single-cell pallets.
7. Count and calculate live cell concentration using trypan blue¹⁴ in an automated cell counter.
8. Calculate the dilution ratio and dilute cells in NIM to make final cell concentration of 60,000 live cells/mL.
9. Add Rho-associated protein kinase (ROCK) inhibitor Y-27632 (final concentration 50 µM) (see Table of Materials) to the cell suspension and gently mix.
10. Add 150 µL of single cell suspension into each well of ultra-low attachment 96-well plate (see Table of Materials) and place in the incubator. Set up as Day 0.
11. On day 1, check the cells and EB forms in each well.
    NOTE: Evaluate successful EB formation based on their bright, spherical, and translucent appearance under a microscope, their ability to remain free-floating and separate, and the absence of dead cells, which appear as dark spots.
12. On day 2, remove 75 µL medium from each well carefully and add 150 µL NIM along with 50 µM Y-27632 to each well.
13. On days 4, 6 and 8, remove 100 µL medium from each well and add 150 µL NIM to each well.

5. Generation of cortical organoids

1. Prepare neural differentiation medium minus vitamin A (NDM-) by combining the components listed in the Table of Materials.
2. On day 10, transfer EBs from the ultra-low attachment 96-well plate to the ultra-low attachment 6-well plate using a 5 mL pipette. Transfer eight EBs to each well cautiously (Figure 2). Add 5 mL NDM- to each well.
3. Place the plate on the orbital shaker inside the incubator and start spinning at 80 rpm.
4. On days 12, 14, and 16, replace the NDM- with 4 mL after aspirating 3 mL old medium.

6. Maturation and long-term culture of cortical organoids

1. Prepare neural differentiation medium with vitamin A (NDM+) and brain-derived neurotrophic factor (BDNF). For details, see Table of Materials.
2. On day 18, remove the 3 mL medium and add 4 mL NDM+.
3. Return the plate to the spinning culture.
4. Feed every 4 days and maintain the differentiated cortical organoids in NDM+ until use. 25-40-day-old cortical organoids were used for the downstream analysis.
   NOTE: The cortical organoids can be maintained longer (up to 3-4 months); but cell viability will be dropped due to the lack of nutrition exchange.

7. Cell characterization by immunocytochemistry and immunofluorescence staining

1. Use a 1000 µL pipette to gently collect the organoid from the medium and place it gently on a medium with minimal medium on a standard microscope slide.
2. Let it dry completely at RT. Add 4% paraformaldehyde (PFA) (300 µL per sample) for 30 min. Remove PFA and then rinse twice with PBS.
   NOTE: PFA is toxic and suspected to be carcinogenic. Avoid contact with skin and eyes, and handle under a chemical fume hood.
3. Add 30% sucrose solution (300 µL per sample) to the slides and incubate overnight at 4 °C.
4. Block and permeabilize the organoids with blocking buffer (BB) (300 µL per sample) containing PBS (with Ca²⁺/Mg²⁺), 0.3% Triton X-100, and 10% normal goat serum for 2 h at RT (or overnight at 4 °C). Outline the organoids with a hydrophobic barrier pen to keep the buffer on the slide.
5. Overlay samples with primary antibody in blocking buffer (300 µL per sample), anti-SRY (sex determining region Y)-box 2 (SOX2, 1:100), and anti-neural marker tuberin beta III (Tuj1, 1:1000), and incubate overnight at 4 °C in the dark (see Table of Materials).
6. Wash in PBS for 3 h with two to three buffer changes on a gentle rocking platform.
7. Incubate with secondary antibody (300 µL per sample), anti-Alexa Flour 488 (1:800), and anti-Alexa Flour 594 (1:800), overnight at 4 °C in a humidified darkroom. Add Hoechst 33342 (1:5000) nuclear stain at the same time (see Table of Materials).
8. Aspirate the antibody and rinse quickly with PBS, then add PBS containing 0.01% NaAzide for 1-2 days at 4 °C to prevent contamination.
9. Aspirate the PBS and remove the excess solution. Mount the organoids with mounting medium (see Table of Materials), then add a coverslip. Avoid the formation of air bubbles and place in a dark room at RT for at least 12 h to allow the mounting to fully polymerize.
   NOTE: The sample is now ready to be imagined under a fluorescence microscope.

8. Dissociation of cortical organoids

1. Add 30 mL DPBS -/- in a 50 mL centrifuge tube.
2. Gently collect the brain organoids in the tube using a 10 mL pipette. At least 3 organoids are used for one staining, and they should be more than 25 days but less than 40 days old.
3. Sediment organoids for approximately 5 min. Do not centrifuge.
4. Aspirate DPBS and add 2 mL pre-warmed Accutase in the tube. Incubate the sample in the water bath for 5 min at 37 °C.
5. Triturate 15 times gently by using a 1000 µL pipette tip without generating bubbles. Incubate the sample for another 5 min at 37 °C in a water bath.
6. Dissociated gently by using 1000 µL pipette tips 15 times without generating bubbles (Figure 3A).
   NOTE: If there are still some undissociated cells, transfer the undissociated cells into a new 50 mL centrifuge tube and then incubate for another 5 min at 37 °C water bath. Then dissociate gently by using 1000 µL pipette tips 20 times.
7. Neutralize with 4 mL DMEM with 10% FBS and centrifuge the 15 mL conical tube containing the cells at 600 x g for 5 min at RT.
8. Aspirate supernatant and resuspend with 200 µL DPBS using 1000 µL pipette tips 10 times without generating bubbles.
9. Add 200 µL DPBS to make a total volume of 400 µL cell suspensions in DPBS.
10. Filter with the filter or filter tube (use 35 or 40 µm filters and rinse the filter with DPBS before using the filter).
11. Separate half of the solution in a tube and use it as unstained sample.

9. Flow cytometry measurement of MRC complex subunits and TFAM in fixed cells

1. Add Live and Dead (L/D) staining dye (1: 1000) (see Table of Materials) (either with Far Red or Near-infrared, depending on the set-up for the markers for flow cytometry).
   NOTE: If a new vial for L/D dye is opened, add 50 µL DMSO to reconstitute and make a 10 µL aliquot to store them at -20 °C.
2. Keep in RT and dark for 30 min. Add 40 mL DPBS -/-. Centrifuge at 600 x g for 5 min.
3. Resuspend into single-cell suspension after discarding the supernatant.
4. Fix and permeabilize the cells with 1 mL ice-cold 90% methanol at -20 °C for 20 min.
5. Block the samples in 1 mL block buffer containing 0.3 M glycine, 5% goat serum, and 1% bovine serum albumin (BSA) in PBS (with Ca²⁺/Mg²⁺).
6. Wash with 20 mL flow buffer containing PBS (with Ca²⁺/Mg²⁺) with 0.2% BSA once or twice.
7. Add the primary antibodies anti-NDUFB10 conjugated with Alexa Fluor 405 (1:100), anti-VDAC 1 conjugated with Alexa Fluor PE (1:100) and anti-TFAM antibody conjugated with Alexa Fluor 488 (1:200) for 30 min (see Table of Materials).
8. Wash with 20 mL flow buffer.
9. Aspirate and leave approximately 100 µL and reconstitute the cell pellets into 300 µL flow buffer.
10. Transfer the cells to 1.5 mL flow tubes. Keep in the dark and ice.
11. Analyze on the flow cytometer (see Table of Materials). Signals were detected for Anti NDUFB10-Alexa 405 using a 450/50 bandpass filter, TFAM-Alexa 488 using 530/30 bandpass filter, VDAC 1-Alexa 647 using 670/14 bandpass filter and L/D dye in 780/60 bandpass filter (Figure 3B).

10. Flow cytometry acquisition and analysis

1. Use unstained and single stained samples of the control organoid to set the voltage of cytometer settings.
2. Use the unstained control tube to set the forward scatter area (FSC-A) and side scatter area (SSC-A) scatter plots based on the size and granularity of the cell population (Figure 4A).
3. Gate out debris to select live cells from FSC-A and SSC-A plots. Gate out doublets using side scatter height (SSC-H) vs. SSC-A plot (Figure 4B).
4. Select the live cells based on the staining of L/D dye by switching to APC-cy7 vs. FSC-A plot (Figure 4C).
5. Using the unstained samples of each line, draw a gate above the main population of the events in FSC-A vs. fluorochrome channels (FITC, APC, BV 421) plots.
6. Set up compensates for the flow cytometer¹⁵.
7. Use isotype control for negative control to observe background staining.
8. Perform data acquisition.
   1. Select the live cells based on the staining of L/D dye by switching to APC-cy7 vs. the FSC-A plot (Figure 4C).
   2. Utilize the unstained as negative control. Then, while examining the FSC-A vs. fluorochrome channels (FITC, APC, BV 421) plots, establish a gate above the main population of single-cell events (Figure 4D-F).
9. Perform data analysis using FlowJo software (see Table of Materials).
   1. Duplicate the positions of the gates onto the stained cell samples. Document the number of cells that exhibit positive staining.
   2. For each targeted population, assess the median fluorescence intensity (MFI) of different channels (FITC, APC, BV 421) plotted on the x-axis of a histogram to identify the mitochondrial signal. Specific values for the complex I subunit NDUFB10 and TFAM can be determined by dividing the MFI of the complex expression or TFAM by the mitochondrial mass indicator, VDAC 1.

Results

Figure 1 provides a diagrammatic representation of the differentiation process and the strategies used for flow cytometric analysis. Human iPSCs were cultured in non-adherent 96-well plates to form EBs and then transferred to non-adherent 6-well plates to obtain fully grown cortical organoids. The cellular composition of organoids was validated using confocal microscopy after immunostaining with neuronal¹⁶ and glial markers¹⁷. Organoids were dissociated into single cells to facilitate the flow cytometry-based measurement of multiple mitochondrial parameters in individual cells. In this protocol, multiple staining strategies are used for acquiring and analyzing the parameters of mitochondrial mass, the amounts of MRC complex I subunit and TFAM, and an indirect measurement of relative mtDNA copy number. This protocol provides steps for dissociating organoids and acquiring flow cytometric data.

Using a previous protocol from Park, I. H. et al.¹⁸, cortical organoids were generated from iPSCs through inducing anterior neuroectodermal fate via dual SMA- and MAD-related protein (SMAD) inhibition and Wingless/Integrated (Wnt) inhibition. As shown in Figure 2A, human iPSCs were seeded in an iPSC culture medium on the basement membrane matrix-coated 6-well plates (Figure 2B,a). When the cells reached ~80% confluency, the colonies were dissociated into single cells and then seeded into 96-well plates with NIM. EBs were formed after 1 day of seeding (Figure 2B,b). On day 10, the EBs were transferred to 6-well plates to allow differentiation of cortical organoids (Figure 2B,c), and NIM was replaced by NDM supplemented with BDNF, N2 supplement, and B27 without vitamin A. After 18 days of differentiation, the differentiated cortical organoids were further matured in NDM, replenished with ascorbic acid, and B27 with vitamin A. At this stage, the cortical organoids displayed tissue-like morphology (Figure 2B,d) and were allowed to grow on a spinning rotator for at least 25 days and then used to study mitochondrial parameters. Cell identity was characterized during differentiation using immunofluorescence staining. In Figure 2C, immunostaining confirmed that the iPSC-derived cortical organoids expressed the specific neural progenitor marker SOX2¹⁹, and neural marker Tuj1²⁰.

Cortical organoids can be sustained for considerable durations, reaching 3-4 months. However, we have noticed a downward trend in cell viability over time. This decrease is predominantly due to inadequate nutrient exchange. As a result, for the subsequent analysis, cortical organoids in the age range of 25 to 40 days were specifically selected. At this stage, the cortical organoids were dissociated into single cells, and then cellular viability was assessed using trypan blue in a cell counter. Incubating the organoids in pre-warmed Accutase for 10 min at 37 °C with gentle trituration at 5 min intervals resulted in optimum single-cell suspension and cell viability (Figure 3A).

Flow cytometry was performed to investigate mitochondrial function in these differentiated cortical organoids. After the cells were stained, as described in Figure 3B, a flow cytometer was used for data acquisition and Flowjo Sampler for data analysis. The gating strategies are shown in Figure 4. A region for living cells was determined on the FSC vs. SSC plot to exclude dead cells and cell debris (Figure 4A). Cell doublets were excluded by making an SSC-A vs. SSC-H plot (Figure 4B). The voltage was set for each fluorophore by altering the cytometer settings using single stained samples. Background fluorescence is adequately assessed if the negative population of a specific cell type is compared with the positive population within the same cell type.

This approach compared the cortical organoids generated from human iPSCs carrying the mtDNA polymerase, POLG (W748S/W748S) mutation with disease-free samples generated from age- and gender-matched healthy control (Table 1). As demonstrated in Figure 5, decreased total and specific complex I NDUFB10 and TFAM levels were shown in POLG organoids compared with controls. However, there were no changes in terms of mitochondrial mass in POLG organoids.

These data suggest that flow cytometric analysis of different mitochondrial parameters renders a first-step approximation that would be valuable in iPSC-derived brain organoids.

Figure 1: Schematic representation of the protocol for the differentiation of cortical organoids from iPSCs and workflow of mitochondrial measurement using flow cytometry. This protocol begins with iPSCs cultured until they reach 80% confluency. These cells are then dissociated and transferred to 96-well plates, where they form EBs. EBs are transferred to 6-well plates and undergo differentiation into cortical organoids over about 18 days. The organoids are then matured for at least 25 days. Then, organoids are dissociated into single cells for flow cytometry. The cells are stained with specific markers to highlight various mitochondrial parameters, including mitochondrial mass measured by VDAC 1, MRC complex I subunit NDUFB10, and mtDNA replication measured by TFAM. Abbreviations: iPSCs: induced pluripotent stem cells; EBs: embryoid bodies; MRC: mitochondrial respiratory chain; mtDNA: mitochondrial DNA. Please click here to view a larger version of this figure.

Figure 2: Flow chart of the iPSC differentiation and representative images for the cells from different stages during the differentiation and characterization of the iPSC-derived cortical organoids. (A) A flow chart that outlines the step-by-step progression of this differentiation. This begins with cultivating iPSCs, followed by their development into EBs. These EBs then differentiate into early-stage cortical organoids and subsequently into NSCs. The NSCs mature into fully formed cortical organoids, concluding the differentiation process. (B) Representative images of cells at each of these distinct stages. Image (a) displays iPSCs, which typically exhibit a tightly packed, colony-like formation characteristic of pluripotent stem cells. Image (b) shows EBs, which are three-dimensional cell aggregates formed as the iPSCs begin to differentiate. Image (c) presents the early-stage cortical organoids, which have a more complex structure as the cells start to organize into tissue-like structures. Image (d) highlights NSCs, which are precursor cells capable of differentiating into neurons and other neural cells. Finally, image (e) shows mature cortical organoids. At this stage, the organoids exhibit a highly complex, tissue-like morphology suggestive of a well-developed neural tissue structure. The scale bar is 1 mm. (C) Confocal microscopy images of the iPSC-derived organoids. These images highlight the immunostaining of SOX2 and Tuj1. SOX2, shown in red, is a marker of neural progenitor cells, indicating the presence of cells capable of generating neural lineage cells. Tuj1, depicted in green, is a marker for neurons, implying the successful differentiation of some cells into neurons. Nuclei are stained with DAPI. The scale bar is 100 µm. Abbreviations: iPSCs: induced pluripotent stem cells; EBs: embryoid bodies; NSCs: neural stem cells. Please click here to view a larger version of this figure.

Figure 3: Single-cell dissociation and sample staining set up in flow cytometric analysis. (A) Representative image of single cells obtained after dissociation of cortical organoids. The magnification is 2.5x. This procedure involves incubating the organoids in Accutase for 10 min at 37 °C, which facilitates the breakdown of the organoids into individual cells. This process is critical as flow cytometry requires single-cell suspensions for accurate analysis. (B) The setup for sample staining to measure multiple mitochondrial parameters using flow cytometry. To evaluate mitochondrial properties, cells must be stained with specific markers or dyes highlighting these parameters. These can be non-stained samples as the negative control, single stained samples with mitochondrial mass marker VDAC 1, the complex I subunit NDUFB10, and TFAM used to indirectly measure mtDNA copy number and multiple stained samples. Abbreviations: MRC: mitochondrial respiratory chain; mtDNA: mitochondrial DNA. Please click here to view a larger version of this figure.

Figure 4: Gating strategies and data acquisition employed in the flow cytometric analysis of organoid cells. (A) A flow cytometry plot showing FSC-A vs. SSC-A. This type of plot is commonly used in flow cytometry to differentiate cell populations based on their size (FSC) and granularity or complexity (SSC). The main gate in this plot is usually drawn to include most cells, excluding debris or other non-cellular events. (B) Strategy for gating single cells by creating a plot of SSC-H vs. SSC-A. This strategy excludes cell doublets or clusters from the analysis, as these could distort the data. Single cells fall along a diagonal line in such a plot, whereas doublets or clusters of cells deviate from this line due to increased area for the same pulse height. (C) The live cells are gated based on the staining of an L/D dye. This dye differentially stains live and dead cells, allowing for their distinction in a plot of APC-cy7 (the channel where the L/D dye is detected) vs. FSC-A. (D-F) These images show gating strategies for different parameters. These plots use the FSC-A parameter vs. different fluorescence channels: in (D), the VDAC 1 is detected in the APC channel, (E) uses the BV421 channel to detect NDUFB10, which is a subunit of the MRC Complex I, and finally, (F) shows the gating for TFAM detection in the FITC channel. Abbreviations: FSC-A: Forward Scatter; SSC-A: Side Scatter; FITC = Fluorescein isothiocyanate, APC = Allophycocyanin, PE = Phycoerythrin, BV421 = Brilliant violet 421, APC-cy7 = Allophycocyanin-cyanine; MRC: mitochondrial respiratory chain; mtDNA: mitochondrial DNA. L/D: Live/Dead. Please click here to view a larger version of this figure.

Figure 5. The results from the flow cytometric analysis of cortical organoids derived from iPSCs of a patient carrying the POLG mutation and healthy control. (A) shows the measurement of total mitochondrial mass. This is assessed by staining with an antibody against VDAC 1, a protein commonly used as a marker for mitochondrial mass due to its location in the outer mitochondrial membrane. The staining intensity of VDAC 1 indicates the relative amount of mitochondria within the cells of the cortical organoids. This illustration demonstrates a similar level of VDAC 1 in organoids derived from POLG compared to those generated from the control. (B) presents the total Complex I levels, which are measured by staining for NDUFB10. Complex I is a key component of the mitochondrial respiratory chain, and its level indicates mitochondrial function. This illustration depicts a notable reduction in the total amount of NDUFB10 in organoids derived from POLG compared to those generated from the control. (C) illustrates the total amount of TFAM. TFAM is crucial for the maintenance and transcription of mtDNA, so its level can give insights into the status of mtDNA in the cells. This illustration depicts a notable reduction in the total amount of TFAM in organoids derived from POLG compared to those generated from the control. (D,E) shows the specific levels of Complex I and TFAM are presented. These are calculated as ratios of total NDUFB10 or total TFAM to the levels of VDAC 1. These ratios provide normalized measures of Complex I and TFAM, accounting for possible variations in total mitochondrial content. This illustration depicts a notable reduction in the specific amounts of NDUFB10 and TFAM in organoids derived from POLG compared to those generated from the control. The data are presented as mean values ± SEM for three independent samples (n = 6). The statistical significance of the observed differences between patient-derived and healthy control organoids was evaluated using the Mann-Whitney U test. A p-value of less than 0.05 was considered to indicate statistical significance. Abbreviations: iPSCs: induced pluripotent stem cells; SEM: standard error of the mean; ns: not significant. Please click here to view a larger version of this figure.

Line	Source	Mutation	Age (years old)	Gender
Control	AG05836 (RRID:CVCL_2B58)	No	44	Female
Patient	POLG patient	POLG homozygous for c.2243G>C; p.W748S	44	Female

Table 1: Information on mutation, age, and gender of the iPSCs used in this study.

Discussion

A protocol is presented for generating cortical brain organoids from human iPSCs and for performing the flow cytometric analysis of mitochondrial parameters in single cells isolated from these organoids. The cellular composition of the organoids was verified by confocal microscopy with immunohistochemical staining for neuronal and glial cell markers. The flow cytometry-based strategy co-staining with anti-NDUFB10, VDAC 1, and TFAM has been shown to allow the measurement of specific levels of complex I and mtDNA relative to the number of mitochondria in a cell. This protocol is suitable for measuring mitochondria in all types of brain-like cells and can be used to study diseases of mitochondrial dysfunction.

This protocol allows for the generation of brain organoids using a combination of dual SMAD and Wnt inhibition. This triple inhibition generates a more uniform cortical NSC profile compared to dual SMAD inhibition²¹. Some cell lines may require 10% heat-inactivated FBS in NIM, although FBS was not used. To ensure the successful differentiation, iPSCs should be undifferentiated prior to plating to form EBs.

A comprehensive protocol for isolating brain organoids into viable single cells is important because it is a critical step prior to procedures such as flow cytometry, and single-cell RNA sequencing. After experimenting with two different reagents (Accutase and Organoid Tissue Dissociation Kit), it was found that incubating organoids in Accutase for 10 min resulted in good cell viability and a single-cell suspension. While the duration required to dissociate them properly may be adjusted based on the size and age of the organoids, it is advisable not to exceed the 20 min incubation time. For samples that are relatively difficult to dissociate, it is recommended to use a 30-40 nm filter to remove debris and insufficiently dissociated cells.

This protocol has some key points to consider. First, the quality and size of cortical organoids are critical to ensure sufficient viable single cells for a successful flow cytometry procedure. To achieve good cell viability of isolated cells, it is recommended to use organoids differentiated for 30-40 days; organoids after 40 days and large-sized organoids should be avoided. We have developed protocols for specific stages in the differentiation process and described them above. To obtain sufficient number of cells, it is recommended to combine 3-4 organoids for a set of flow cytometry. Additionally, many other differentiation protocols and defined media^22,23,24 are available that could be effective alternatives, but these issues are not experimentally addressed in the current work. As media composition and clonal differences in iPSC lines can affect the proliferation and differentiation efficiency of the starting cell population, adaptation of this strategy to other maintenance media may require optimization.

Flow cytometry has the advantage of being able to analyze large numbers of single cells, as well as being fast and reproducible compared to other conventional microscope-based assays. Flow cytometry analysis requires less than a million cells, and sample analysis takes only a few minutes, meaning dozens of samples can be analyzed in 1-2 h. Additionally, in microscope-based analyses, researcher bias can distort results, especially with samples that vary widely in 3D and cellular composition, such as organoids. Flow cytometry analysis not only overcomes the obstacles of microscope-based analyses but can more accurately analyze the small differences between various indicators. The strategy can also be applied to a variety of cell types, including neurodegenerative disease cells and other brain organoids, and thus should help understand mechanisms and test potential treatments for different neurodegenerative diseases.

In recent years, organoids have been successfully used to model a variety of neurological diseases. They are superior to animal models due to the added advantage of being structurally similar to the human brain²⁵. This technology is expected to revolutionize high-throughput drug screening and the development of new therapies²⁶. This protocol can be used to model various mitochondrial-associated diseases and other neurodegenerative diseases, such as PD, and can be used for drug screening as well as for testing new treatments and assessing the effects of environmental toxins on the human brain.

The development of organoids, including cortical organoids, is subject to considerable variability, both within and between experimental batches²⁷. This variation is influenced by factors such as the initial cell population, minute differences in the culture environment, and random occurrences during the processes of differentiation and maturation. Consequently, different organoids may model diverse stages or features of cortical development and function. Additionally, while organoids are valuable models for understanding development and disease, they do not perfectly mirror the structure or cell diversity of the human brain. For instance, they lack key brain microenvironment components such as blood vessels and immune cells²⁷. Moreover, the proportions and types of neurons and glial cells generated may not accurately reflect those in the human brain.

Flow cytometry is a potent tool for characterizing and enumerating cell populations based on their physical and chemical properties. However, it does have constraints. One such constraint is that organoids must be dissociated into individual cells for analysis. This dissociation could modify the cell state and influence mitochondrial function, thereby affecting measurements of mitochondrial parameters²⁸. The process of dissociation can also cause cell stress and may lead to cell death, especially in more delicate cell types. Moreover, the interpretation of flow cytometry data heavily relies on proper gating strategies. Accurately distinguishing between noise and real events and setting gates demands expert knowledge²⁸. Inconsistent gating can lead to significant bias and affect results. Lastly, while flow cytometry is adept at analyzing cellular subpopulations, it may fail to capture spatial information and interactions among cells, which could be crucial in the context of organoids.

This study identified a key limitation regarding the preparation of cell suspensions for flow cytometry analysis. Our initial approach involved comparing different conditions, including no filtration, filtration with subsequent dissociation of the unfiltered sample, and direct filtration. Not using a filter was found to be unsuitable for the flow cytometer, as this equipment necessitates a single-cell suspension for accurate analysis. Moreover, a significantly low cell viability of 20% was observed while attempting to re-dissociate samples after filtration. This low viability level compromised the ability to obtain reliable flow cytometry data, effectively ruling out this approach as a viable method. As a result, direct filtration through a 35 or 40 µm filter was decided to perform. This limitation underscores the inherent difficulties in maintaining cell viability during preparation for flow cytometry. It also highlights the importance of considering equipment-specific requirements when designing an experimental methodology. These challenges are important to address in future studies to optimize cell preparation techniques to maximize cell viability and enhance flow cytometry data quality.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Antibodies using in flow cytometry
anti-DUFB10 Alexa Fluor 405	NOVUS biologicals	NBP2-72915AF405
anti-VDAC1 Alexa Fluor 647	Santa cruz technology	sc-390996
anti-TFAM Alexa Fluor 488	Abcam	ab198308
L/D fixable near-IR dead cell stain kit	Life technologies	L10119
Antibodies using in immunofluorence staining
anti-Tuj1	Abcam	ab78078
anti-SOX2	Abcam	ab97959
anti-Alexa Flour 488	Thermo Fisher Scientific	A28175
anti-Alexa Flour 594	Thermo Fisher Scientific	A-21442
Commercial cells
AG05836 (RRID:CVCL_2B58)			Provided by Gareth John Sullivan from the Institute of Basic Medical Sciences at the University of Oslo, Norway
Essential 8 Medium (iPSC culture medium)
Essential 8 Basal Medium	Thermo Fisher Scientific	A1516901
Essential 8 Supplement (50x) 2% (v/v)	Thermo Fisher Scientific	A1517101
Store at 4 °C and warm up to RT before use.
Instruments
Heracell 150i CO2 Incubators	Fisher Scientific, USA
Orbital shakers - SSM1, SSL1	Stuart Equipment, UK
CCD Microscope Camera Leica DFC3000 G	Leica Microsystems, Germany
Water Bath Jb Academy Basic Jba5 JBA5 Grant Instruments	Grant Instruments, USA
Fluid aspiration system BVC control	Vacuubrand, Germany
Leica TCS SP8 STED confocal microscope	Leica Microsystems, Germany
50 mL falcon tube	Sigma-Aldrich	CLS430828
BD LSR Fortessa	BD Biosciences, USA
Flowjo Sampler Analysis	FlowJo LLC, USA
10 mL pipette	Sigma-Aldrich	SIAL1100
1, 10, 100, 1000 mL pipette	Sigma-Aldrich
40 µm Cell stariner	Sigma-Aldrich	CLS431750
ultra-low attachment 96-well plate	S-BIO	MS-9096UZ
Countess II automated cell counter	Thermo Fisher Scientific
Neural differentiation medium  (NDM+)
DMEM/F12	Life technologies	11330032
Neurobasal medium	Life technologies	2110349
Glutamax supplement 1% (v/v)	Life technologies	35050
N2 supplement 0.5% (v/v)	Life technologies	17502-048
B27 supplement 1% (v/v)	Life technologies	17504-044
β-Mercaptoethanol 50 µM	Sigma-aldrich	M3148
BDNF 20 ng/mL	Peprotech	450-02
Ascorbic acid 200 µM	Sigma-Aldrich	A92902
Store at 4° C for upto 2 weeks
Neural differentiation medium minus viatmin A (NDM-)
DMEM/F12	Life technologies	11330032
Neurobasal medium	Life technologies	2110349
Glutamax supplement 1% (v/v)	Life technologies	35050
N2 supplement 0.5% (v/v)	Life technologies	17502-048
B27 supplement W/O vit. A 1% (v/v)	Life technologies	12587010
β-Mercaptoethanol 50 µM	Sigma-aldrich	M3148
Store at 4° C for upto 8 days
Neural Induction Medium (NIM)
DMEM/F12	Life technologies	11330032
Knockout serum replacement 15% (v/v)	Life technologies	10828028
Glutamax supplement 1% (v/v)	Life technologies	35050
β-Mercaptoethanol 100 µM	Sigma-Aldrich	M3148
LDN-193189 100 nM	Stemgent/Reprocell	04-0074
SB431542 10 µM	Tocris	1614
XAV939 2 µM	Sigma-Aldrich	X3004
Store at 4° C for upto 10 days
Neutralisation medium
IMDM	Life technologies	21980032
FBS 10%	Sigma-Aldrich	12103C
Other reagents
DPBS (Ca2+/Mg2+ free)	Thermo Fisher Scientific	14190250
Bovine Serum Albumin	Europa Bioproducts	EQBAH62-1000
Accutase	Life technologies	A11105-01
Geltrex	Life technologies	A1413302
EDTA	Life technologies	15575038
Advanced DMEM / F12	Life technologies	12634010
Neural tissue dissociation kit	Miltenyi biotec	130-092-628
Y-27632 dihydrochloride Rock Inhibitor	Biotechne Tocris	1254
Fluoromount-G™ Mounting Medium	SouthernBiotech	0100-20
PFA	Thermo Fisher Scientific	28908