Three-Dimensional Imaging of Organoids to Study Primary Ciliogenesis During ex vivo Organogenesis

Summary

Stem cell-derived organoids facilitate the analysis of molecular and cellular processes that regulate stem cell self-renewal and differentiation during organogenesis in mammalian tissues. Here we present a protocol for the analysis of the biology of the primary cilium in mouse mammary organoids.

Abstract

Organoids are stem cell-derived three-dimensional structures that reproduce ex vivo the complex architecture and physiology of organs. Thus, organoids represent useful models to study the mechanisms that control stem cell self-renewal and differentiation in mammals, including primary ciliogenesis and ciliary signaling. Primary ciliogenesis is the dynamic process of assembling the primary cilium, a key cell signaling center that controls stem cell self-renewal and/or differentiation in various tissues. Here we present a comprehensive protocol for the immunofluorescence staining of cell lineage and primary cilia markers, in whole-mount mouse mammary organoids, for light sheet microscopy. We describe the microscopy imaging method and an image processing technique for the quantitative analysis of primary cilium assembly and length in organoids. This protocol enables a precise analysis of primary cilia in complex three-dimensional structures at the single cell level. This method is applicable for immunofluorescence staining and imaging of primary cilia and ciliary signaling in mammary organoids derived from normal and genetically modified stem cells, from healthy and pathological tissues, to study the biology of the primary cilium in health and disease.

Introduction

Development of multicellular organisms and the maintenance of homeostasis in their adult tissues reside in a fine-tuned regulation between self-renewal and differentiation of stem cells, which orchestrate in time and space normal tissue development and regeneration¹. Subversion of this regulation causes developmental anomalies and cancers². Thus, understanding the molecular and cellular mechanisms that orchestrate stem cell self-renewal and differentiation is of key interest in developmental and cancer biology.

Recent development of ex vivo organogenesis methods, in which tissue stem cells generate three-dimensional organoids have transformed our capabilities to study the dynamics of stem cells during mammalian organogenesis and maintenance of tissue homeostasis in a dish³. Organoids represent a good alternative to cumbersome genetically modified animal models to study these processes. Protocols for the development of organoids from tissue stem cells of many organs have now been developed³, including small intestine and colon, stomach, liver, pancreas, prostate, and mammary gland³. Additionally, the development of somatic genome-editing techniques in organoid-forming stem cells now enables to quickly interrogate the molecular and cellular mechanisms that control their biology^4,5.

The primary cilium is a microtubule-based structure that is assembled at the surface of stem and/or differentiated cells of various tissues⁶. It is generally non-motile and is assembled as a single structure per cell⁷. Primary ciliogenesis is the dynamic process of assembling the primary cilium⁷. At the cell surface, the cilium acts as a cell signaling platform⁸. Thus, the primary cilium is thought to act as a key regulator of stem cell self-renewal and/or differentiation in many tissues, including the brain^9,10, the mammary gland^4,11, the adipose tissue¹², and the olfactory epithelium¹³, among others. Primary ciliogenesis and/or ciliary signaling are dynamically regulated in distinct cell lineages and at different developmental stages^4,13,14, but the underlying mechanisms remain to be largely determined.

Ex vivo organogenesis shows promise for the development of basic knowledge on the molecular and cellular mechanisms that control stem cell biology, including primary ciliogenesis and ciliary signaling. However, it relies on the ability to properly image whole mount organoids at the single cell level and at sub-cellular scales. We recently used a mouse mammary stem cell-derived organoid model to show that primary cilia positively control mouse mammary stem cell organoid-forming capacity⁴. Here we present a comprehensive protocol for the immunofluorescence staining of whole mount mouse mammary organoids (Figure 1A,B), which enables the analysis of primary cilia through light sheet microscopy during ex vivo organogenesis in three-dimension. Alternative methods were recently published for the immunofluorescence staining and imaging of organoids through confocal microscopy^15,16. This protocol focuses instead on the preparation and imaging of organoids through light sheet microscopy.

Protocol

NOTE: The protocol below is recommended for the staining of organoids that were grown in 5 wells of a 96 well plate and pooled together (> 100 organoids). Organoids were derived from mouse mammary stem cells. Donor mice were housed and handled in accordance with protocols approved by the Animal Care Committee of the University of Rennes (France).

1. Reagents

1. To prepare the fixative solution, dilute 125 µL of 16% paraformaldehyde (PFA) aqueous commercial solution in 375 µL of phosphate-buffered saline (PBS) to generate a 4% PFA solution.
   CAUTION: Manipulate PFA that is a toxic substance under a chemical hood.
2. To prepare the permeabilization buffer, dilute 1.5 µL of Triton X100 in 500 µL of PBS, to produce a 0.3% Triton X100 solution.
3. To prepare the blocking buffer, dilute 1.5 µL of Tween-20 and 75 µL of normal goat serum in PBS, to generate a 5% goat serum-0.1% Tween 20 solution.
4. To prepare the light sheet mounting medium, dissolve 1 g of ultrapure low melting point agarose in 100 mL of dH20 or PBS at 65 °C. Prepare 1 mL aliquots and store at room temperature.

2. Organoids recovery

1. Transfer organoids from the culture wells to a low binding polymer 1.7 mL tube, after pipetting them up and down 3 times in the wells, with an FBS-coated tip for which the end was cut (minimal diameter of the extremity 1.5 mm).
   NOTE: The coating prevents organoids from sticking to the tip. The low-binding polymer material enables to reduce the attachment of organoids to the side of the tube during the entire staining procedure.
2. Fill the tube with PBS, and spin down at 350 x g for 3 min. Remove the supernatant.

3. Fixation, permeabilization and blocking

1. Resuspend the organoids in 500 µL of 4% PFA. Incubate for 30 min at room temperature. Spin down at 350 x g for 30 s. Remove the supernatant.
   NOTE: From this step onwards, resuspend organoids in the different buffers by simply adding the buffers on the pellet of organoids without touching them. Perform all washing steps at room temperature.
2. Wash the organoids with 1 mL of PBS for 3 min. Spin down at 350 x g for 30 s. Remove the supernatant.
   PAUSE: Fixed organoids can be kept in PBS at 4 °C for at least a week.
3. Permeabilize the organoids by resuspending them in 500 µL of PBS-Triton X100 0.3% and incubate for 30 min at room temperature. Spin down the organoids at 350 g for 30 s.
4. Wash the organoids by resuspending them with 1 mL of PBS and incubate for 3 min. Spin down at 350 x g for 30 s. Remove the supernatant. Repeat once.
5. Optional: Resuspend the organoids in 500 µL of ice-cold methanol. Incubate at - 20 °C for 10 min. This step may be required for the staining of specific centrosomal markers.
6. Optional (if step 5 was performed): Wash the organoids with 1 mL of PBS for 3 min. Spin down at 350 x g for 30 s. Remove the supernatant. Repeat once.
7. Block non-specific antibody binding sites in organoids by resuspending them with 500 µL of blocking buffer (PBS, 5% goat serum, 0.1% Tween 20). Incubate 1 h and 30 min at room temperature. Spin down at 350 g for 30 s. Remove the supernatant.
8. Wash the organoids by resuspending them with 1 mL of PBS and incubate for 3 min. Spin down at 350 x g for 30 s. Remove the supernatant.

4. Labelling

1. Resuspend the organoids with 200 µL of blocking buffer with diluted primary antibodies and incubate overnight at 4 °C with mild shaking (60 rpm on a horizontal shaker). Place the tubes with a 45° angle with the horizontal plan of the shaker. It will maintain the organoids at the bottom of the tubes in the staining buffer.
2. Wash the organoids by resuspending them with 1 mL of PBS and incubate for 5 min. Spin down at 350 x g for 30 s. Remove the supernatant. Repeat twice.
   NOTE: Some organoids in the last step may stick to the side of the tube, resulting in organoid loss during aspiration of the supernatant after centrifugation, adding 0.2% (w/v) bovine serum albumin (BSA) to the PBS in the washing steps may reduce organoid loss.
3. Resuspend the organoids with 200 µL of blocking buffer with secondary antibodies and incubate for 1 h and 30 min with mild shaking (60 rpm on a horizontal shaker). Place the tubes with a 45° angle with the horizontal plan of the shaker. It will maintain the organoids at the bottom of the tubes in the staining buffer.
   NOTE: Hoechst (or other nuclear dyes, such as DAPI or DRAQ5) can be added to the buffer with secondary antibodies.
4. Wash the organoids by resuspending them with 1 mL of PBS and incubate for 5 min. Spin down at 350 x g for 30 s. Remove the supernatant. Repeat twice.
   ​NOTE: Some organoids in the last step may stick to the side of the tube, resulting in organoid loss during aspiration of the supernatant after centrifugation, adding 0.2% (w/v) BSA to the PBS in the washing steps may reduce organoid loss.

5. Preparation of the agarose sample for imaging

1. Melt light sheet mounting medium by incubating it at 65 °C. Once the medium has melted, incubate it at 37 °C for 5 min.
2. Resuspend the organoids in 100 µL of mounting medium using a 200 µL tip, with the extremity of the tip cut (minimal size of the extremity: 1.5 mm), by pipetting up and down twice.
3. Suck the mounting medium with the organoids in a glass capillary (green capillary, inner diameter: 1.5 mm) using a plunger. Incubate the capillary at room temperature for 5 min for the mounting medium to solidify.
   PAUSE: Capillary can be stored in PBS at 4 °C for a week before imaging.

6. Imaging

1. Using a light sheet microscope (e.g., ZEISS Lightsheet Z.1), image the organoids with 20x or 40x water immersion objectives.
   1. Place the glass capillary in the observation chamber. Locate the capillary with the front camera and place the tip of the glass capillary at the upper limit of the detection objective.
   2. Select the sample and set the optimal focus using the tip of the glass capillary in brightfield illumination. Push out the agarose sample slowly from the capillary and locate organoids within the solidified agarose.
   3. Keep the organoids to be imaged close to the tip of the capillary, to reduce movements of the agarose sample in the PBS. Rotate the capillary to set the optimal sample orientation and define the desired zoom.
   4. Select acquisition mode. Define the channels to image, set proper orientations of the light sheet, set the laser power for each channel (to reduce photobleaching, use low laser power), set the size of the image and the desired illumination side(s). Example of imaging parameters: Image size 1920 x 1920, illumination: dual side.
   5. Define the Z-stack to image by setting the Z-extremities of the stack using the Z-stack module and set the Z-step size to optimal.
2. Process the output file using the image processing module of the microscope software to navigate in the sample, obtain a Z-projection and 3D-representation.
3. Turn the sample and acquire a new image from a different angle or image other organoids.
4. Analyze images with an interactive microscopy image analysis software (e.g., Imaris) enabling (i) the visualization of the sample in 3D (ii) the segmentation of objects (iii) identification and quantitative analysis of objects.

Results

Ex vivo organogenesis methods are transforming our capabilities to study mammalian tissue development and maintenance of tissue homeostasis in a dish. The analysis of molecular and cellular mechanisms that regulate these processes, including primary ciliogenesis and ciliary signaling, relies on the ability to image organoids in three-dimension.

The protocol described above enables the staining of whole-mount mammary organoids. They arise from mammary stem cell-enriched basal cells tha...

Discussion

The detailed protocol presented here enables the staining and imaging of mouse mammary organoids that grow in semi-solid medium. This protocol is presumably applicable to the staining of organoids mimicking the architecture of various tissues that grow in semi-solid and solid media. For organoids that grow in 100% Matrigel with medium on top, the recovery and fixation steps slightly differ. The culture medium must be removed from the culture well. After a quick PBS wash, the fixative solution (4% PFA) may be directly add...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Anti-mouse IgG1 647	Thermo-Fisher	A21240
Anti-mouse IgG2A 488	Thermo-Fisher	A21131
Anti-rabbit 546	Thermo-Fisher	A11035
Arl13b	NeuroMab	73-287
EMS 16% Paraformaldehyde Aqueous Solution, EM Grade	Electron Microscopy Sciences	15710
FBS	Thermo-Fisher	10270106
gtubulin	Sigma-Aldrich	T5326
Hoechst 33342	Thermo-Fisher	62249
Integrin a6	Biolegend	313616
Light Sheet Capillary	Zeiss	701908
Light Sheet plunger	Zeiss	701998
Low binding Microcentrifuge tubes	BioScience	27210
Normal Goat Serum Blocking Solution	Vector labs	S-1000
PBS	Sigma-Aldrich	p3587
Slug	Cell Signaling Technology	9585
Triton-X100	Sigma-Aldrich	T9284
Tween-20	Euromedex	9005-64-5
UltraPure Low Melting Point Agarose	Thermo-Fisher	16520050