Live-cell imaging of mammary organoids using light sheet microscopy

Abstract

The mammary gland is a dynamic organ whose parenchyma undergoes major development during puberty and extensive remodeling with each estrous cycle. These processes can be modelled and investigated in vitro via 3D cell culture techniques that employ specialized extracellular matrices and appropriate growth factors. The resulting mammary organoid cultures faithfully represent the mammary gland with respect to cellular heterogeneity, cell-cell contacts, overall architecture as well as response to growth factor stimuli and are amendable to a variety of molecular methods as well as microscopy techniques. Among the imaging techniques, light sheet microscopy (single plane illumination microscopy; SPIM) represents a useful method for longitudinal monitoring of morphological changes and cell behavior during the establishment of mammary gland ductal systems. In contrast to other fluorescence microscopy techniques such as widefield- and confocal-microscopy, SPIM exerts minimal phototoxicity while allowing fast acquisition of different fluorophores within organoids arranged in a 3D matrix under optimized environmental conditions. Here, we provide a detailed protocol for organoid acquisition and culture and describe two sample mounting variants for use with multiview and inverted light sheet microscopes.

Introduction

The mammary gland is a unique organ whose primary function is to secrete milk to nourish and enable the survival of postnatal offspring. The epithelial ductal network is embedded in a fibroadipose stroma and represents the main functional compartment of the gland. Development of the epithelial network starts prenatally, but the complex glandular structure is achieved through the process of branching morphogenesis initiated at puberty [1, 2]. This mechanism is orchestrated by reproductive hormones and various cellular signaling molecules [3]. Altered components and/or regulation of developmental processes are often implicated in the pathogenesis of benign and malignant breast tumors.

In recent years, the use of 3D culture techniques has greatly contributed to the understanding of basic developmental processes and their association with malignant transformation. Mammary gland epithelial organoids provide reproducible and robust systems to study branching morphogenesis and associated factors [4]. Primary mammary organoids recapitulate branching morphogenesis within seven to nine days when embedded in the extracellular matrix (ECM) and provided with key signaling factors [5]. The essential tool for monitoring these dynamic changes is light microscopy. Standard brightfield microscopy allows global morphological changes to be followed at the whole organoid level, while deeper insights into the cellular behavior can be gained by using fluorescently labeled structures, such as nuclei or membranes, as well as individual proteins. In addition, the use of biosensor constructs allows the visualization of protein activity dynamics within single cells and at the tissue level.

Here, we use primary mammary organoids from the B6-Tg(EKAREV)pT2A-3905NLS (EKAREV-NLS) mouse strain [6, 7] and organoids expressing nuclear H2B-mCherry marker (stained with actin SPY650-FastAct dye) to track signaling and/or morphological changes in response to different growth stimuli. While this model enables the spatiotemporal tracking of signaling dynamics within a selected sample, the protocol presented here is broadly applicable to a range of experimental systems. It can be employed for the observation and monitoring of various MGO model types, including mammary single cell-derived organoids, mammary epithelial fragment–derived organoids, and induced pluripotent stem cell–derived organoids that either endogenously express fluorescent proteins or are labeled with live-cell dyes. These models can be used to track chemical or morphological responses using gentle light sheet microscopy setups that preserve sample integrity.

Light microscopy methods are an indispensable tool to follow dynamic morphological changes along with protein dynamics in living 3D systems. The key functional feature of light sheet fluorescence microscopy (LSFM) is the perpendicular positioning of the illumination and detection objectives [8–12]. This allows fluorophore excitation in one section plane while the signal is instantly collected through the detection objective. Therefore, phototoxicity and bleaching are minimized because fluorophores and cells are only exposed to laser light for a short time period when simultaneously recorded [13]. LSFM has several advantages when used for live time-lapse (4D) imaging. First, excitation is achieved by illumination with a laser light sheet, which significantly increases acquisition speed. Second, only the observed region of interest is illuminated, reducing photodamage and cell stress in the surrounding sample. Finally, the lateral resolution of light sheet microscopes is comparable to that of a standard fluorescence microscope, while the axial resolution depends on the adjustable thickness of the light sheet. Depending on the imaging goals and the sample type, different mounting techniques can be used in live LSFM. In the context of cell biology these are most commonly adherent cell cultures on a coverslip or special tissue culture plates, samples embedded in soft gel suspended in an imaging chamber, and cultivation in V-shaped FEP chambers.

The last two approaches are particularly useful for the 4D imaging because they allow sample mounting in an upright position and simultaneous imaging of multiple samples. Suspending samples in a larger imaging chamber is commonly used in what is known as multiview light sheet microscope (LSM) setup, which is suitable for larger samples where light penetration can be a challenge. Multiview mounting is characterized by a sample suspended from or standing on a vertical support within the imaging chamber, with illumination and detection objective being in the same plane around the sample (Fig. 1a, b). This can be achieved by embedding the sample in a cylinder of soft gel (agarose or ECMs) that is extruded from a capillary into the imaging chamber. Alternatively, gel-embedded samples can be placed on a coverslip, in FEP tubes or on custom printed holders [14]. The multiview setup allows the acquisition of multiple samples of larger volumes and fast acquisition from multiple angles, resulting in a complete image after post-processing.

Fig. 1.

Light sheet fluorescent microscopy principle. a Multiview setup with a sample suspended in the middle of an imaging chamber with exemplary sample holder schematics. b Representative brightfield and fluorescent images of mammary EKAREV-NLS organoids acquired on a multiview LSM. c Inverted setup with detection objective under a V-shaped sample holder within water-filled imaging chamber. d Representative brightfield and fluorescent images of mammary EKAREV-NLS organoids acquired on an inverted LSM

Alternatively, inverted light sheet systems can be used that allow to image deep into the sample with sufficiently high working-distance range of the setup. Here, the samples can be mounted in culture medium with or without ECM gel by simply pipetting them into the bottom of a well of a special light sheet sample holder (Fig. 1c, d). The holder has V-shaped wells made of FEP film with a total volume capacity of 400–500 µL. The holder is placed inside a larger, water-filled imaging chamber. The specific setup we used (Luxendo TruLive3D LSM) allows imaging of multiple samples in up to 6 wells, which is advantageous when testing different treatments and conditions.

In this protocol article we provide comprehensive guidelines for the isolation, cryopreservation, and long-term imaging of MGOs in two different light sheet microscopy setups. First, we describe methods for tissue processing and organoid collection. Second, we focus on the organoid culture and freezing. Finally, we describe two approaches for sample mounting in a light sheet microscope. Special emphasis is placed on the aspects of specimen mounting in multiview and inverted light sheet systems, as this is one of the most challenging factors in long-term imaging of living specimens.

Materials

Biological materials, reagents, and solutions

Biological materials, reagents, and solutions common to all procedures

1. Female mice of desired strain, age and phenotype. Optimally, use strains with an endogenously expressed fluorescent marker (see Note 1).

2. Dulbecco’s Modified Eagle Medium: Nutrient Mixture F-12 without L-glutamine with phenol red (DMEM/F12; 21331-020, Thermo Fisher Scientific).

3. Dulbecco’s Modified Eagle Medium: Nutrient Mixture F-12 with L- glutamine without phenol red (PR-free DMEM/F12; 21041-025, Thermo Fisher Scientific) (see Note 2).

4. Digestion solution 1: 2 mg/mL collagenase (C9891, Merck), 2 mg/mL trypsin (27250018, Thermo Fisher Scientific), 5% (v/v) fetal bovine serum (FBS), 5 µg/mL insulin, 50 µg/mL gentamicin in DMEM/F12. Prepare freshly for each isolation.

5. Deoxyribonuclease I (DNase I) stock solution: 2 U/µL DNase I (D4263, Merck) in 0.15 M NaCl. Add 1 mL of 0.15 M NaCl to 2000 U (1 mg) DNase I. Aliquot and store at −20 °C.

6. Deoxyribonuclease I (DNase I) working solution: 20 U/mL DNase I in DMEM/F12. Add 40 µL of 2 U/µL DNase I to 4 mL of DMEM/F12. Prepare freshly for each isolation.

7. Basal organoid medium: 2.5 mM glutamine, 1× ITS (10 µg/mL insulin, 5.5 µg/mL transferrin, 6.7 ng/mL selenium; 41400-045, Thermo Fisher Scientific), 100 U/mL penicillin, 100 µg/mL streptomycin in PR-free DMEM/F12. Store at 4 °C.

8. Phosphate buffered saline (PBS).

9. 2.5% bovine serum albumin (BSA) solution: 2.5% (w/v) BSA in PBS. Store at 4 °C.

10. Fixation solution: 4% (w/v) paraformaldehyde in PBS.

11. ECM gel, e.g. Matrigel (see Note 3).

12. Serum-free cryopreservation medium, such as Synth-A-Freeze (A1254201, Thermo Fisher Scientific) or Cellbanker 2 (11914, ZENOGEN PHARMA Co., Ltd.) (see Note 4). Store at −20 °C.

13. Digestion solution 2: 150 U/mL collagenase type 3 (LS004180, CLS-3 Worthington); 20 µg/mL Liberase Blendzyme 2 (11988425001, Roche), 25 mM HEPES in DMEM/F12 with 3.151 g/L glucose and L-glutamine. Store at 4 °C.

14. 3D ECM matrix: 15 mg/mL basement membrane extract (BME) and 1.5 mg/mL collagen I (see Notes 5 and 6).

15. Mammary epithelial growth medium (MEGM), e.g. MEGM™ Mammary Epithelial Cell Growth Medium BulletKit™ (CC-3150, Lonza) (see Note 7). Store at 4 °C for up to a month.

16. STOP medium: 10% (v/v) FBS, 25 mM HEPES, and 100 U/mL penicillin, 100 µg/mL streptomycin in DMEM/F12 with 3.151 g/L glucose and L-glutamine. Store at 4 °C.

17. Collagenase stock solution: 75000 U/mL collagenase type 3 in MEGM medium. Store aliquots at −20°C.

Instruments, equipment, and other materials

1. Centrifuge tubes, 1.5 mL, 15 mL, 50 mL.

2. Cryovials.

3. Electronic pipettor and disposable plastic serological pipettes, 5 mL, 10 mL, 25 mL.

4. Adjustable volume automatic pipettes (10 µL, 20 µL, 100 or 200 µL, 1000 µL) and pipette tips.

5. 24-well cell culture plate, flat bottom.

6. Cell culture dishes, 30 mm and 100 mm.

7. Collagen I-coated 6-well cell culture plate, flat bottom (A1142801, Thermo Fisher Scientific, or 354400, Corning).

8. Ice container.

9. Surgical scalpels, sterile.

10. Forceps, sterile.

11. Scissors, sterile.

12. Cotton buds, sterile.

13. Needles, 25 G and 26 G.

14. Insulin syringes.

15. 1 mL syringes.

16. Cell counting chamber, e.g. Neubauer type.

17. Parafilm.

18. Incubated orbital shaker (37°C) (see Note 8).

19. Centrifuge.

20. Cell culture incubator (37°C, 5% CO₂).

21. Laminar flow cell culture hood.

22. Stereoscope.

23. Light sheet microscope, such as as a multiview LSM (e.g. ZEISS Lightsheet 7) or an inverted LSM (Luxendo TruLive3D).

24. FEP tubing, ID × OD = 1.3 × 1.6 mm, wall thickness 0.15 mm (FT1.3X1.6, Adtech) (see Note 9).

25. FEP V-shaped wells, FEP foil thickness 0.025 mm (Trulive 3D dishes, Bruker/Luxendo) (see Note 10).

26. Holder for manipulation with FEP V-shaped wells (Bruker/Luxendo).

Methods

Mammary gland dissection

In this paragraph we describe the major steps of mammary gland tissue collection for isolation of primary MGOs. The reader is kindly referred to other, more detailed versions of this protocol for graphical aids [15–17].

1. After euthanasia, pin the mouse to the dissection board and wash the ventral side with 70% EtOH.

2. Using forceps lift the abdominal skin and make a small incision in the midline. Use this incision as a starting point to make sagittal cuts towards the neck and the tail, then make cuts towards the limbs.

3. Lift and slightly pull away the skin on one side of the mouse. Use a cotton bud to help separate the skin from the peritoneum. Pin the skin to the dissection board and repeat on the opposite side (see Note 11).

4. Remove the lymph node from the mammary glands number four and collect the remaining mammary tissue together with the mammary glands number three into a 2 mL centrifuge tube with cold PBS (see Note 12).

Primary mammary epithelial organoid isolation

Before starting, prepare ice in a styrofoam box and preheat the water bath and orbital shaker to 37 °C. Make sure that the centrifuge is set for the room temperature (RT). The following steps should be performed in a biosafety cell culture hood.

1. Prepare digestion solution 1 (10 mL per mouse) in a 50 mL tube, pre-warm it in a water bath, and filter-sterilize.

2. Transfer the mammary gland tissue to a sterile 100 mm Petri dish. Remove the excess of PBS and use scalpels to chop the tissue into small pieces approximately 0.5 × 0.5 × 0.5 mm³ in size and then transfer the chopped tissue to the tube containing digestion solution.

3. Place the tube containing tissue in digestion solution horizontally in the incubated orbital shaker and incubate at 110 rpm, 37 °C for 30 min.

4. After incubation, inspect the digested tissue. If no large pieces remain, proceed with the next steps. If large pieces of tissue are still visible, extend the incubation time accordingly.

5. Centrifuge the suspension at 450 × g for 10 min.

6. Meanwhile, coat 15 mL tubes with BSA solution; prepare 1 tube per each approx. 10 mL of digestion solution. Use BSA-coated tubes, pipettes, and pipette tips for all sample handling (see Note 13).

7. After centrifugation, transfer the supernatant from the 50 mL tube to the coated 15 mL tubes at a maximum volume of 10 mL per tube. Leave the pellet in the 50 mL tube.

8. Add DMEM/F12 to each tube to a total volume of 14 mL and mix well. Centrifuge the tubes at 450 × g for 5 min.

9. Meanwhile, resuspend the pellet from the 50 mL tube in 4 mL of DMEM/F12 (see Note 14).

10. After centrifugation, aspirate the supernatants from the 15 mL tubes.

11. Perform a series of pellet resuspensions by collecting the resuspended cells from the initial 50 mL tube (in 4 mL) and transferring the suspension to the pellet in the first 15 mL tube. Resuspend the pellet from this tube with the cell suspension from the 50 mL tube and use the entire volume to repeat the resuspension in the next 15 mL tube. Repeat until you reach the last 15 mL tube, resuspend the pellet and leave the entire suspension in the tube. Add 4 mL of DMEM/F12 to the 50 mL tube, wash the inside of the tube with it, and repeat the transfer and wash in the 15 mL tubes. Collect the suspension in the final 15 mL tube containing the pooled cell suspensions. Empty tubes may be discarded after this step.

12. Centrifuge the suspension at 450 × g for 5 min.

13. Aspirate the supernatant and resuspend the pellet in 4 mL of DNase I working solution (see Note 14). Incubate for 5 min at RT while shaking gently.

14. Add 6 mL of DMEM/F12 and mix well.

15. Centrifuge the suspension at 450 × g for 5 min.

16. Aspirate the supernatant and resuspend the pellet in 9 mL of DMEM/F12.

17. Perform differential centrifugation: Centrifuge at 450 × g for 10 s (see Note 15).

18. Repeat steps 17 and 18 four more times for a total of 5 times.

19. After the last round of differential centrifugation, resuspend the pellet in 1-2 mL of basal organoid medium and place on ice until further use (see Note 16).

20. Count the number of organoids. Resuspend the organoid suspension with a 1 mL automated pipette. Immediately take 20 µL of the suspension and place it in two drops (approximately 10 µL each) on a cell culture dish (see Note 17). Count the organoids in the drops using an inverted microscope.

21. Calculate the organoid concentration and total number of organoids in the 15 mL tube.

    

    

Cryopreservation of primary organoids

Primary organoids can be cryopreserved for a later use. We recommend use of serum-free cryopreservation media because in our experience they provide better viability and morphogenetic response after organoid revival (Fig. 2). Before starting, prepare your cell freezing container and bring it to RT if necessary.

Fig. 2.

Morphogenesis of revived organoids in a 3D culture. Organoids were frozen with different freezing media (indicated in the top panel), thawed after 5 weeks' storage in liquid nitrogen, seeded in ECM drops, and cultivated in the presence of 2.5 nM FGF2 for 6 days. Scale bar, 100 μm

1. Calculate the total number of organoids in the suspension (subheading 3.2, steps 21–22).

2. Centrifuge the suspension at 450 × g for 5 min.

3. Meanwhile, label the cryovials with the date, sample name, number of organoids, and any other relevant information.

4. Aspirate the supernatant. Resuspend the organoid pellet in serum-free cryopreservation medium (see Note 4). Use 1 mL of cryopreservation medium per 1000–3000 organoids (see Note 18).

5. Transfer 1 mL of suspension to each cryovial.

6. Place the cryovials in the cell freezing container and transfer to a −80 °C freezer.

7. After 24 h at −80 °C, transfer the frozen vials to a liquid nitrogen storage tank.

Cryopreserved primary organoid thawing

Before starting, prepare ice, 37 °C water bath and preheat the media.

1. Remove the cryovial from the liquid nitrogen tank and place it on ice.

2. Place the cryovial in a float in the 37 °C water bath and incubate until there is only a small amount of ice left (see Note 19).

3. Meanwhile, coat a 15 mL tube with BSA, and add 10 mL of pre-warmed DMEM/F12.

4. Add 1 mL of pre-warmed DMEM/F12 to the cryovial containing the thawed organoids and gently but thoroughly resuspend.

5. Transfer the suspension (2 mL in total) to the 15 mL tube containing 10 mL of pre-warmed DMEM/F12.

6. Centrifuge the suspension at 450 × g for 5 min.

7. Aspirate the supernatant and resuspend the pellet in 1 mL of basal organoid medium and place on ice.

8. Count the number of organoids (see paragraph 3.2, steps 21 and 22) (see Note 20).

Organoid seeding

In this paragraph, we describe procedure for seeding either fresh or revived primary mammary organoids. Before starting, prepare ice, 37 °C water bath, pre-cool PBS in a 15 mL tube, and pre-warm basal organoid medium.

1. Calculate the volume of organoid suspension required to obtain the desired number of organoids per condition and well (see Note 21):

   

• 2.Transfer the appropriate volume of organoid suspension into a 1.5 mL BSA-coated centrifuge tube (see Note 22).
• 3.Centrifuge the suspension at 450 × g for 5 min.
• 4.Place the tubes on ice and remove the supernatant (see Note 23).
• 5.Use a 10 µL automated pipette to resuspend the organoids in the remaining supernatant, taking care not to create bubbles.
• 6.Add the required volume of the ECM gel to the tube containing the resuspended organoids (50–70 µL/well), resuspend and place on ice.
• 7.Place the cold PBS and 24-well plate on ice.
• 8.Using a 200 µL pipette, create 3–6 mm wide discs (round patches) of ~ 10 µL ECM each at the bottom of the wells. Before taking up the ECM gel in the pipette tip, precool the pipette tip with the cold PBS (by pipetting the cold PBS in and out of the tip) (see Note 24).
• 9.Once all the discs have been created, incubate the plate 10–15 min at 37 °C.
• 10.Use the pre-cooled pipette tip to resuspend the organoid suspension in the ECM gel. Take the desired volume of the suspension (50–70 µL) and pipette it into the center of the previously created disc. Avoid making bubbles (see Note 25). Discard the tip.
• 11.Repeat step 10 until all of the organoid suspension is used up and all desired culture wells are populated with the organoids. Place the plate in the tissue culture incubator at 37 °C, 5% CO₂ for 30–60 min to allow the ECM gels to solidify.
• 12.Meanwhile, prepare culture medium variants according to your experimental setup (e.g., adding growth factors or drugs) and warm them up in the water bath. When using a 24-well cell culture plate, use 1 mL of medium per well.
• 13.After incubation, when the ECM gels have set, add 1 mL of preheated medium variants to each of the culture wells. Place the plate in the tissue culture incubator and incubate for several days according to the experimental plan.
• 14. Change the culture medium every 2 days, or according to your experimental design.

Organoid culture method from single cells

As an alternative to epithelial fragments, mammary organoids can be grown from single mammary epithelial cells. Here we present a protocol optimized to produce organoids that are better suited for morphogenetic studies using an inverted light sheet microscope (LSM). Since inverted LSM has a more limited focal depth compared to multiview LSM, both the placement and size of the sample significantly affect imaging quality. Therefore, smaller organoids are more appropriate for imaging on an inverted LSM system. Please also see previous versions [18–20] of this protocol, which have been adapted for a number of applications.

1. Harvest mammary glands as described in Sect. 3.1.

2. Place three mammary glands without mechanical dissociation in 5 mL of digestion solution 2 in a loosely capped 50 mL tube. Digest for 15–16 h at 37 °C.

3. Wash the resultant organoid suspension with 40 mL of PBS.

4. Centrifuge at 450 × g for 5 min at RT.

5. Remove the supernatant, and resuspend the pellet in 5 mL of trypsin-EDTA. Screw the cap loosely onto the tube, and incubate it for 20 min at 37 °C.

6. After the incubation, stop the digestion by adding 40 mL of STOP medium.

7. Centrifuge the cells at 450 × g for 5 min at RT.

8. Discard the supernatant and resuspend the pellet in 3 mL of MEGM.

9. Plate the cells onto collagen I–coated 3.5-cm dishes (6-well plates). Incubate the plates in a tissue-culture incubator at 37 °C, 5% CO₂ for 26–30 h.

10. Aspirate the medium and wash the plate with PBS. Treat the attached cells with 0.5 mL of trypsin-EDTA (prewarmed to 37 °C) and incubate at 37 °C, 5% CO₂ until the cells detach.

11. After cell detachment, inactivate the trypsin with 9 mL of STOP medium and transfer the suspension to a 15 mL tube.

12. Centrifuge the cells at 190 × g for 5 min at RT.

13. Discard the supernatant, and resuspend the cells in 1 mL of MEGM.

14. Count the number of single cells using a cell counting chamber. Exclude dead cells, erythrocytes, and cell aggregates from your count.

15. Working quickly, mix 10,000 to 12,000 cells (in 10–20 μL MEGM) with 100 μL of Matrigel on ice. Dispense 95 μL into one well of a 12-well tissue-culture plate.

16. Let the gels solidify on a level surface in a cell culture incubator at 37 °C for 30 min.

17. Add 1.5 mL of MEGM to each gel, and incubate at 37 °C in a tissue culture incubator. Replace the medium every 2 days. Organoids are typically grown till 8-10 cell stage, which corresponds to 3 days in culture.

Collection of organoids for mounting

We describe a sample mounting strategy for two different LSM setups when preselection and preservation of organoid morphological features are required for imaging. The following steps, inspired by a protocol for organoid mounting for immunostaining [21], are common to both LSM approaches and should be performed under a stereomicroscope.

1. To prepare an ECM “dispenser”, place a sterile insulin syringe with needle in the refrigerator at least 2 h before starting. When the syringe has cooled down, working in a cell culture hood, aspirate a small amount (about 100 µL) of ECM gel in it, cover the needle with a cap to keep it sterile, and place it on ice (see Note 26).

2. Use a 100 mm tissue culture dish to prepare the collection chamber. If you are collecting samples from multiple conditions, use a permanent marker to mark individual areas on the bottom of the dish. Mark each area with a sample identifier (e.g. number or name). Line the bottom of the dish with a sterile parafilm (see Note 27).

3. Prepare and preheat fresh phenol red free organoid culture medium (either basal organoid medium or MEGM) according to your experimental conditions.

4. Create 50 µL drops of the preheated medium in all areas of the collection chamber.

5. Place the 24-well plate containing the organoid culture under the stereomicroscope and locate the desired organoids.

6. Using two 24 G needles cut the ECM gel around the selected organoids (see Note 28). Gently move the organoids to the edge of the ECM drop. Repeat this step until you have collected the desired number of organoids.

7. Use a 200 µL pipette with cut sterile tips to collect the excised organoids (see Note 29). To prevent sticking of the organoids to the pipette tip, first aspirate a small amount of the medium in the tip, then continue to aspirate the organoids. Transfer the organoids to the appropriate drop in the collection chamber.

8. Repeat the collection until you have acquired desired number of organoids in all experimental conditions.

9. Once all the organoids have been collected, use the 200 µL pipette to slowly remove (almost) all of the medium from each of the drops in the collection chamber. Monitor the process under the stereomicroscope to avoid collecting the organoids.

10. Add 50 µL of the phenol red free organoid culture medium to the organoids.

11. Proceed immediately to the next steps for organoid mounting. We describe a sample mounting strategy for two different LSM setups when preselection and preservation of organoid morphological features are required for imaging.

Organoid mounting for a multiview LSM

Multiview LSMs typically have an imaging chamber and a stage that can be adapted for different sample types, holders, and mounting solutions. FEP tube fragments are the main component of our organoid sample holder and this section describes how to prepare them and load them with samples. To prepare appropriate environmental conditions for imaging, clean the imaging chamber with 70% ethanol or sterilize it by gamma irradiation and then set the temperature to 37 °C and CO₂ to 5%.

Organoid mounting in an FEP tube fragment

To prevent rapid ECM polymerization, perform the following steps at a low temperature. Keep instruments on ice, pre-cool the room using air conditioning, or work in a cold room. When handling ECM, work quickly and minimize manipulation to preserve the texture and integrity of the ECM surrounding the sample.

1. Using a surgical scalpel, pre-cut the FEP tube into approximately 1–3 mm long fragments (see Note 30).

2. Collect the fragments in a centrifuge tube and sterilize them by gamma irradiation.

3. Cut a thin (10 × 100 mm) piece of parafilm and wrap it diametrically around the bottom of a 100 mm cell culture dish (save the lid for later steps): Using about a half of the parafilm, make one layer first. Then place a sterile 1 mL syringe with a removable 26 G needle on the parafilm on the dish and use the rest of the parafilm to secure the syringe in the position (Fig. 3a, b).

Fig. 3.

Equipment and crucial mounting steps for a multiview LSM. a Equipment needed for assembling sample holder; from left 100 mm cell culture dish, 24–26 G needle, 1 mL syringe, a piece of parafilm. b An assembled holder with mounted FEP tube fragment (indicated by an arrowhead). c-j Stages of organoid mounting. c Empty FEP tube fragment. d FEP tube fragment loaded with ECM. e FEP tube fragment with ECM and mounted organoid (white arrowhead) with the drop of media. f FEP tube with a mounted organoid (white arrowhead) covered with ECM. g Higher magnification of F. h Syringe mounted in multiview stage adapter. i Holder with an organoid (white arrowhead) mounted in a multiview LSM stand in an imaging chamber (j)

• 4.Using the remaining lid of the 100 mm cell culture dish as a sterile surface, place a sterile FEP fragment on the lid. Position the fragment horizontally so that both ends of the fragment are visible and only the walls of the tube fragment touch the plastic of the dish.
• 5.Remove the needle from the syringe and pierce the FEP fragment through the center. Lift the needle with the FEP fragment and place it back on the syringe.
• 6.Using a stereomicroscope, locate and observe the needle tip containing the FEP fragment. Using a sterile 10 µL pipette tip, position the FEP fragment so that the openings are horizontal (see Note 31) (Fig. 3c).
• 7.Take the insulin needle containing the ECM gel and inject the gel into the FEP fragment from one side until it forms convex domes on both sides of the FEP fragment (Fig. 3d). Put the cap on the needle and place it back on ice.
• 8.Using a pipette with sterile-cut 10 µL pipette tips, collect the organoids from the collection chamber drop. Observe the drops under a stereomicroscope and use no more than 3 µL of medium to collect the organoids.
• 9.Locate the needle containing the ECM-filled FEP fragment and pipette the organoids to one side of the FEP fragment (see Note 32) (Fig. 3e). Use the same pipette to remove excess medium while keeping organoids on the ECM-filled FEP fragment.
• 10.Take the insulin needle with ECM gel and place small drop of ECM gel on top of the organoids (Fig. 3f, g). This will seal them between two layers of ECM gel and ensure that they remain in place during the imaging.
• 11.To image multiple organoids simultaneously, repeat the process on the other side of the FEP fragment (see Note 33).
• 12.Allow the ECM gel to polymerize for 5–10 min at RT (see Note 34).
• 13.Place the entire 100 mm dish with syringe and mounted organoids in a closed dark chamber (see Note 35) and transfer it to your LSM location.
• 14.There are two ways to place the syringe holder on the microscope, depending on the user’s dexterity level: Insertion through a stage opening (Sect. 3.8.2) or insertion through the microscope chamber door (Sect. 3.8.3). It is important to work as carefully as possible and to prevent the FEP fragment from touching any surfaces, as this would result in the loss of the sample. The first approach is faster, but the risk of losing the sample is higher. The second approach is safer but requires the ability to manipulate the microscope hardware.

Direct specimen insertion through stage opening

1. Gently remove the syringe containing the needle and the FEP fragment from the parafilm.

2. Slide the entire holder into a round adapter for multiview LSM and lock it in place (Fig. 3h).

3. Place the adapter with the syringe in the microscope in the same way as the glass capillaries (see Note 36.

4. Lower the microscope stage so that the holder and specimen are immersed in the medium (Fig. 3i, j).

5. Locate the specimen and set acquisition parameters according to your experimental design.

Inserting the specimen through the microscope chamber door

1. Gently remove the needle containing the FEP fragment from the syringe. Balance it on the sterile surface or place it on the rim of the culture dish, making sure that FEP fragment remains in the air.

2. Remove the syringe from the parafilm and place it in the adapter for multiview LSM (Fig. 3h).

3. Place the adapter with the syringe in the microscope in the same way as glass capillaries.

4. Withdraw the medium from the imaging chamber and remove the chamber from the microscope. Lower the microscope stage so that the tip of the syringe is visible.

5. Replace the needle with the FEP fragment on the syringe and raise the microscope stage to the initial position (Fig. 3i).

6. Return the imaging chamber to the microscope and refill it with medium.

7. Lower the microscope stage so that the holder and specimen are immersed in the medium (Fig. 3j).

8. Locate the specimen and set acquisition parameters according to your experimental design.

Organoid mounting for inverted LSM

For an inverted LSM setup, special FEP foil wells are designed to complement the geometry of the microscope objectives. The specimen is placed at the bottom of the well, where the double-sided light sheet illumination excites the fluorophores, while an inverted camera (placed on the bottom side of the well) detects emission. In the example Luxendo TruLive3D LSM, a maximum of 6 wells can be used simultaneously. Here we describe two approaches to mounting samples in V-shaped wells, depending on the type of the sample. The first (Sect. 3.9.1) is designed for cases where specific morphological features need to be preserved and therefore the ECM cannot be removed. The second approach (Sect. 3.9.2) is particularly useful when a larger number of smaller, morphologically uniform samples need to be imaged [22], and a temporary removal of ECM would not affect cellular behavior and organoid morphology. Prior to imaging, set the microscope environmental control to 37 °C and 5% CO₂ and allow the temperature and CO₂ levels to stabilize.

Mounting of large organoids with specific morphological features

1. Prepare the V-shaped wells: Open the sterile container and use tweezers to lift the sterile V-shaped wells from the attaching adhesive surface (see Note 37), place them in the manipulator holder, and mark the top edge (see Note 38) (Fig. 4a).

Fig. 4.

Equipment and crucial mounting steps for an inverted LSM. a Labelled V-shaped sample holders in a manipulation stand. b V-shaped imaging holders in an imaging chamber. c-e Crucial steps in sample mounting. Creation of Matrigel drop (c), spreading the drop on the bottom of the holder (d) and addition of organoids (e)

• 2.To facilitate the transport and maintain aseptic conditions, place the holder with wells on a 100 mm cell culture dish (Fig. 4a).
• 3.Using a stereomicroscope, locate and observe the inner bottom of the well.
• 4.Using the ECM dispenser (prepared in Sect. 3.6), add a small drop (up to 5 µL) of ECM gel to the bottom of the well (see Note 39) (Fig. 4c). After use, cap the needle and return the dispenser on ice.
• 5.Take a 10 µL pipette tip in your hand and using it spread the ECM drop lengthwise across the bottom of the well (Fig. 4d).
• 6.Use a pipette with a sterile cut 10 µL pipette tip to collect the organoids from the collection chamber drop. Observe the drops under a stereomicroscope and use no more than 3 µL of medium to collect the organoids.
• 7.Locate the bottom of the V-shaped well with the line of ECM gel and transfer the collected organoids to the center of the gel line (see Note 40) (Fig. 4d).
• 8.Once you have transferred all the desired organoids to the well (see Note 41), observe the volume of the medium in the well. If there is more than 3–5 µL of medium in the well, use sterile 10 µL pipette to remove the excess. Observe the process under the stereomicroscope and avoid picking up the organoids or ECM gel.
• 9.Use the ECM dispenser to add small drops of ECM to the top of the organoids (see Note 42). This will trap them in the ECM gel, provide the necessary growth conditions, and prevent unwanted movement during medium addition and image acquisition.
• 10.Repeat steps 3 through 9 to add organoids to the remaining wells.
• 11.When all the organoids are mounted, place the holder with the wells in the cell culture incubator for 5–10 min.
• 12.Fill the wells with preheated phenol red free organoid culture medium (such as basal organoid medium or MEGM) with growth factors and/or inhibitors according to your experiment, using a maximum of 400 µL per well.
• 13.Place the holder with the wells in the cell culture incubator for a minimum of 2 h.
• 14.Transfer the holder with the wells to the inverted LSM and place the wells in the stage holder of the microscope (Fig. 4b). Continue with your standard workflow to locate the sample and set up acquisition parameters according to your experimental design.

Mounting of small, single-cell-derived organoids with uniform morphological features

1. Digest the ECM: Add 4 µL of collagenase stock solution (equivalent to 300 U of collagenase) to each well (which already contains a 100 µL gel with organoids and 1 mL of MEGM medium). Resuspend the gels using the mechanical force of a 1000 µL pipette; pipette vigorously up and down 6–8 times (see Note 43). Incubate for 50 min at 37 °C, 5% CO₂ (see Note 44).

2. Resuspend the digesting suspension again, then collect and transfer the contents of the wells to a 15 mL centrifuge tube. Typically, a maximum of 6 initial 100 µL gels are transferred to one 15 mL centrifuge tube (see Note 45). Fill the tube(s) up to 14 mL with PBS.

3. Centrifuge the samples at 190 × g for 5 min (see Note 46).

4. Aspirate the supernatant gently, leaving approximately 600–1000 µL of culture medium to avoid disturbing the pellet. Resuspend the pellet vigorously in the remaining supernatant using a 1000 µL pipette, rinsing the walls of the tube (see Note 47).

5. Fill up the tube(s) with STOP medium. Add 5–8 µL of DNase I stock solution and incubate for 3–5 min at RT.

6. Centrifuge the samples at 190 × g for 5 min.

7. Aspirate the supernatant gently, leaving around 600–1000 µL of medium to avoid disturbing the pellet. Fill up the tube(s) with PBS and resuspend the pellet by pipetting with a 1000 µL pipette while rinsing the tube walls.

8. Centrifuge the samples at 190 × g for 5 min.

9. Aspirate as much supernatant as possible, without disturbing the cell pellet (see Note 48). Resuspend the pellet vigorously in the remaining supernatant.

10. Prepare a seeding 3D ECM master mix consisting of ice-cold PBS, collagen I and Matrigel in a volume ratio of 1:1:4 on ice by pipetting in this order (see Note 49).

11. Prepare imaging wells according to steps 1 and 2 in Sect. 3.9.1.

12. Take 100–200 µL of the ECM master mix and add it to the organoid pellet/suspension. Mix gently to avoid bubble formation.

13. Apply 10–20 µL of the ECM-organoid suspension in a line at the bottom of the V-shaped imaging well (see Note 50).

14. Allow the gels to solidify in the wells on a flat surface for 30 min at 37 C, 5% CO₂.

15. Add 400 µL of phenol red free organoid culture medium to each gel and place the wells in the cell culture incubator (at 37 °C, 5% CO₂) for at least 2 h to set.

16. Transfer the holder with the wells to the inverted LSM and place the wells to the microscope stage holder (Fig. 4b). Continue with your standard workflow to locate the sample and set up acquisition parameters according to your experimental design.

Image acquisition

Image acquisition protocols will vary depending on your experimental question and the fluorophores chosen. Prior to each acquisition, adjustments should be made to the light sheet characteristics and laser power, but this is beyond the scope of this protocol. In addition, each microscope and software are different, so be sure to familiarize yourself with your specific conditions. Here, we provide a quick guide to general procedures to follow and considerations to make during a time-lapse acquisition, which applies to both multiview and inverted LSM. Examples of data generated through this protocol can be found in the Fig. 5.

Fig. 5.

Examples of successful 5D experimental setups performed on inverted and multiview LSMs. a Maximum intensity projection (MIP) images of single cell-derived organoids carrying nuclear H2B-mCherry marker, stained with actin SPY650-FastAct dye. Full volumes in two channels were acquired from day 3 in culture every 30 min over the period of 72 h. Scale bars, 20 μm. b MIP images of epithelial fragment-derived organoids carrying EKAREV-NLS biosensor with nuclear CFP and YFP signals. Full volumes in two channels were acquired from day 4 in culture every 10 min over the period of 18 h. Scale bars, 40 μm. c MIP images of epithelial fragment-derived organoids carrying EKAREV-NLS biosensor construct with nuclear CFP and YFP signal. Full volumes in two channels were acquired from day 7 in culture every 3 min over the period of 4 h. Scale bars, 100 μm

1. Once you have positioned your holder in the field of view, use a bright field camera to locate your organoids.

2. Once you have located the organoid, mark the position in the software. Move the stage to locate another organoid and mark the position again. Repeat until you have located all available organoids (see Note 51).

3. Adjust the acquisition settings. Adjust the laser power and exposure time to obtain adequate signal intensity (see Note 52).

4. Set up the Z-volume. Use the brightest fluorescence channel and lower laser power to temporarily visualize the organoids. Find the start and end points of your organoids and mark them in the software. If you expect your organoids to grow or move, leave some space to make sure they will stay within the imaged area.

5. Repeat for all organoid locations.

6. Set up the region of interest (ROI) according to the largest organoid. To avoid excessive data generation, limit the x-y camera area by setting the ROI. If you expect your organoids to grow or move, leave enough space to ensure they stay within the imaged area.

7. Enter your time-lapse parameters to the software. In our experimental setup, we acquire the full z-volume of selected organoid positions every 10 min for at least 24 h.

8. Run the acquisition.

9. Check the positions the next day (after about 12–14 h). If organoids have not moved substantially and remain within the ROI, continue with acquisition. If the organoids have moved, stop the acquisition and manually correct the positions. Continue the acquisition for the remaining time.

10. After acquisition, remove the specimens and clean the microscope’s imaging chamber as usual. Samples can be fixed and stained for further analysis.

11. Keep in mind that the time-lapse acquisition combined with light sheet setup typically produces large amounts of data (several TBs). Consider data storage and analysis options when designing the experiments.

Notes

1. When selecting a fluorescent marker, be aware of its spectral characteristics. When combining multiple markers, ensure that the excitation and emission spectra do not overlap. If possible, avoid fluorochromes with excitation wavelength close to the UV spectral region, as UV light is detrimental to cell viability.

2. Phenol red free media should be used for acquisition. Specimens may be cultured in phenol red-containing media prior to acquisition, but in this case the medium should be changed to phenol red-free media at least 24 h prior to acquisition. Phenol red can quench the signal of certain fluorescent proteins and reduce the quality of acquired images. If the fluorescent protein signal is bright, the presence of phenol red should not be a problem. However, for signals of lower intensity, phenol red can significantly affect the signal-to-noise ratio.

3. Any phenol red free ECM gel can be used. Depending on the research question, growth factor-reduced or non-reduced ECM can be used.

4. FBS may contain various growth factors and other components that might affect organoid behavior in culture. It has been shown that FBS can negatively affect organoid formation and growth.

5. Use growth factor reduced and phenol red free BME, such as Matrigel growth factor reduced, phenol red free (356231, Corning), or Cultrex UltiMatrix (BME001-10, BioTechne GmbH). Store at −20 °C, thaw overnight at 4 °C before use. For collagen I, use rat tail collagen I from (e.g. 3447-020-01, BioTechne GmbH). Store at 4 °C.

6. Vials of BME gel (e.g. Matrigel or Cultrex) are normally stored at − 80 °C. To prepare working aliquots, thaw a vial overnight on ice at 4 °C. When working with BME gel, always keep it on ice and use pre-cooled tubes and pipette tips. Store aliquots (1 mL) at −20 °C.

7. The kit contains MEBM^™ Basal Medium (CC-3151, Lonza) and MEGM^™ SingleQuots^™ Supplements (CC-4136, Lonza). Mix the components according to the manufacturer’s instructions and store the medium at 4 °C for no longer than one month.

8. If an incubated shaker is not available, it can be replaced by a small shaker placed in an incubator (at 37 °C).

9. Any FEP tubing with the RI ranging 1.341–1.347 and wall thickness ≤ 0.20 mm can be used. The thinner the wall, the better image.

10. Alternatively, self-build sample holder tools (single well V-shaped trays) are also available from Bruker/Luxendo. FEP foil 0.025 mm (302463, RCT Reichets Chemietechnik GmbH + Co) is recommended for this option.

11. If necessary, use small sterile scissors to separate the mammary glands #4 and #5. This will prevent tearing and loss of tissue.

12. Place the tubes on ice to reduce loss of tissue viability. When working with larger tissue fragments or dissecting more mice, a small Petri dish containing a few mL of sterile PBS can be used for tissue collection. The collected tissue can be stored on ice in PBS for up to 24 h without much negative effect on organoid yield or viability.

13. Coating or pre-wetting the tissue culture plastic with BSA will reduce the cell/organoid loss by preventing them from adhering to the plastic. To coat a pipette, aspirate the solution to the maximum pipette volume, then return the solution to its stock tube. To coat a tube, pipette 1–2 mL of solution into the tube, close the cap, and shake gently by hand until all the tube walls are covered. Place the tube on the holder and aspirate the excess solution back to its stock tube. As an alternative to the 2.5% BSA solution, PBS or basic cell culture media (e.g. DMEM/F12) can be used.

14. It is recommended to first resuspend the pellet in 1 mL of the solution using an automated pipette with a 1000 µL pipette tip first to speed up the resuspension process, then add 3 mL of the solution to the total volume of 4 mL.

15. Separate the organoids from the stromal cell fraction by differential centrifugation. Stromal cells and other light suspension components will remain in the supernatant, while organoids (which are heavier because they are epithelial fragments with hundreds of cells) will stay in the pellet. To achieve 10 s centrifugation at 450 × g, set your centrifuge to 1 min, and observe when the speed reaches the 450 × g. From that moment count 10 s and manually stop the centrifugation.

16. Organoids can be kept on ice for several hours (up to 12 h).

17. The organoids in the counting drops are discarded after counting, so any plastic surface or even a microscope slide can be used. Sterility is not essential for this, so the surface can be washed and reused after use.

18. Freezing/thawing steps will slightly affect organoid viability and total number. When considering the amount to freeze per vial, calculate 10-20% more organoids than needed for the experimental setup.

19. This process is relatively quick and may take 2 to 3 min. Be careful not to over incubate the vial as this will affect organoid viability.

20. Since the freezing and thawing process may result in a slight loss of organoids, we recommend that you do not rely solely on the number of organoids listed on the cryovial. Count the organoids again to ensure the correct number of organoids for your experimental setup.

21. Primary organoids are highly variable in behavior and morphological response. Small numbers of organoids may limit observation of the desired phenotype. High numbers of organoids may result in unwanted organoid interactions (e.g., fusions) and ECM gel changes. We recommend seeding up to 200 organoids in 50 µL of ECM for optimal setup.

22. 1.5 mL centrifuge tubes have a V-shaped bottom which allows for easy manipulation of the organoid pellet, and to prevent bubble formation.

23. Use a 1000 µL automated pipette to slowly remove the supernatant. Do not to disturb the pellet. As the supernatant volume decreases, switch to using a 200 µL automated pipette for easier manipulation. Do not remove all of the supernatant, leave approximately 3 µL to facilitate resuspension of the organoids.

24. The size of the ECM discs should correspond to the volume of ECM you will use for the 3D culture. For smaller volumes, the disc size may be smaller than for larger volumes of the ECM. The disc provides a base for the ECM dome in which the organoids will grow. If the disc is too small, the ECM drop with organoids will overflow it, with the risk of losing the 3D dome shape and organoids attaching to the bottom of the well, which would ruin organoid morphogenesis.

25. This step can be performed at the RT, but if available, we recommend placing the well plate on a heated plate at 40°C. This will speed up the ECM polymerization process and prevent drop spillover.

26. Avoid creating bubbles and ensure sure that the needle and gel are not out of the ice for more than half a minute.

27. Cut a square piece of the parafilm corresponding to the size of the culture dish. Place the parafilm side on the bottom of the dish and press it firmly to the edges, avoiding wrinkles or bubbles. Remove the cover layer of the parafilm. To additionally sterilize the surface of the dish with parafilm, the dish can be left uncovered in the tissue culture hood under UV light for 20 min.

28. It is important to avoid touching the organoids with needles as this may damage the cell layers, cause additional stress to the organoids, alter cell behavior and cause lumen collapse in cystic organoids. However, it is important not to leave too much ECM gel around the organoid as this will affect the imaging quality and make mounting more difficult.

29. Use cut tips to minimize mechanical stress on the organoids during the transfer. Also, use small volumes for the transfer (up to 50 µL) to minimize the amount of phenol red co-transferred.

30. Cut the tube with a gentle cutting motion to avoid shape changes.

31. Hold the widest part of the pipette tip with your fingers for greater precision. Position the FEP fragment by rotating it around the needle.

32. The convex shape of the ECM gel may disappear at this step due to polymerization of the gel at higher temperatures and subsequent volume reduction.

33. It may be difficult to mount the organoids on the other side of the FEP tube. Therefore, the holder can be rotated to facilitate precise manipulation. Do not rotate the FEP fragment around the needle; instead, rotate the entire 100 mm cell culture dish. This will ensure optimal positioning for mounting but will risk movement of the organoids already mounted on the other side.

34. Typically, ECM gels are incubated at 37°C. Do not do this as the following steps will require the sample and gel to be out of the medium for a period of time and incubating the gel at 37 °C may result in overdrying of the gel.

35. This can be any type of a closed container or a box that will protect your sample during the transport. To prevent contamination or loss of sample, ensure that the FEP fragment with organoids does not touch the walls of the container or any other surface.

36. This is a critical moment where the loss of sample can occur. Keep your hands steady and insert the holder straight into the microscope stage. Do not touch the sides.

37. When manipulating the wells, avoid touching the FEP film as it is very thin and can be easily damaged, and any particles or shape changes on the FEP film may result in reduced image quality.

38. The area for marking the wells is quite small, so we recommend using a thin permanent marker and marking the wells with numbers corresponding to your experimental conditions or different samples.

39. During this step there is a risk of damaging the FEP sheet of the well as the ECM dispenser is actually a needle. This step can be performed with a low-volume automated pipette that can produce a 5 µL drop of ECM. However, this may be more challenging as the manipulation space under the stereomicroscope is limited and the drop should be placed as low as possible in the well.

40. It is best to collect all the organoids at once, but if this is not possible, repeat the collection until you have transferred all the organoids to the well.

41. The number of collected and mounted organoids will depend on your experimental settings, mainly the desired temporal resolution combined with the developmental stage and size of organoids in specific condition. We have successfully imaged up to 12 organoids in 10 min intervals, dispersed in four different wells (3 organoids per well). Still, within one well we mounted up to 6 organoids, of which 3 were imaged and 3 were used as non-illuminated controls.

42. Use the syringe to slowly add ECM gel onto the organoids. Do not apply larger drops or multiple drops at the same time as this will cause the organoids to float to the surface. It is important that the organoids remain as low as possible in the well due to the detection limitations of the microscope hardware. Depending on the placing and the number of the organoids, use around 5 drops to trap the organoids in place.

43. Mechanical disruption of the 3D ECM prior to incubation with the enzyme speeds up and ensures a complete digestion of the ECM. The gel must be completely dissolved in the medium before proceeding with the protocol.

44. Shorter incubation times or reduced enzyme concentration will result in an incomplete digestion of the 3D ECM. Efficient digestion of the ECM can be assessed by observing the pellets obtained in the next step of the protocol - after centrifugation. A typical defined organoid/cell pellet indicates complete EMC digestion. A larger, fuzzy, gradient-like pellet is found when the ECM is not sufficiently degraded and may result in loss of organoids (trapped in the haze) when the supernatant is removed.

45. Starting with 6 initial 100 μL gels in a 15 mL centrifuge tube is the most efficient in our experience. A larger number of initial gels results in more ECM and proportionally more digestive enzymes, which are not easily washed out in the subsequent steps. This increases the risk of residual digestive enzymes in the pellet, that would degrade the newly seeded 3D ECM and disrupt the newly cultured gels. Conversely, a smaller number of initial gels results in smaller, more unstable pellets, which may result in a low number of organoids after the washing/centrifugation steps.

46. Alternatively, centrifuge at 230 × g for 4 min.

47. We have found that organoids tend to adhere to the walls of the 15 mL tubes. By carefully rinsing the tubes every time, we recover higher yields of organoids.

48. To avoid disturbing the pellet, first aspirate as much supernatant as possible, leaving an excess volume of supernatant. Then remove the excess with a 200 μL pipette. Depending on the degree of digestion of the ECM, the pellet will have a different volume (larger, fuzzier when the digestion is incomplete). Under optimal conditions, the cell pellet will be small, and around 20–30 μL of supernatant can be left. If larger pellets are obtained, around approximately 50–80 μL of supernatant must be to be left to avoid reducing the yield/numbers of recovered organoids.

49. We are working with a 3D ECM composed of Matrigel and collagen I. Matrigel is routinely used to enable mammary organoid growth and development; the addition of collagen I facilitates other relevant processes such as cell migration, invasion, or branch elongation.

50. This forces the organoids to grow at the bottom (tip) of the V-shaped dish and gives the light sheet has the best access to the sample.

51. Due to the characteristics of the holders, some organoids may not be accessible. These cannot be imaged but can be used as non-illuminated controls.

52. Compromises must be made at this step depending on the phototoxicity of your fluorophores, the desired time resolution, and duration of the time-lapse acquisition.

Author contributions

M.B., M.J. and Z.S.K. conceptualized the study. M.B. performed most of the wet lab and imaging experiments and prepared data from them for presentation in this manuscript. A.A.K. performed imaging experiment presented in Fig. 5a and contributed data from it for presentation in this manuscript. M.B., M.J. and Z.S.K. wrote the main manuscript text, M.B. prepared all Figures. All authors reviewed the final manuscript.

Funding

This work was supported by grants from the Grant Agency of Masaryk University (MUNI/G/1775/2020 to Z.S.K. and MUNI/A/1398/2021), and Ministry of Education, Youth and Sports of the Czech Republic (MEYS CR; grant no. ERC CZ LL2323 FIBROFORCE to Z.S.K.) and the Baden Württemberg Foundation (MITI-SECC; Microbiom call to M.J.). M.B. is a holder of the Brno PhD Talent Scholarship, funded by the Brno City Municipality, and Christian Boulin Fellowship, funded by the European Molecular Biology Laboratory (EMBL). A.A.K. is supported by the MOLIT Institute for the postdoctoral fellowship at EMBL.

Data availability

No datasets were generated or analysed during the current study.

Declarations

Competing interests

Z.S.K. is the Editor-in-Chief of the Journal of Mammary Gland Biology and Neoplasia.