A Static Self-Directed Method for Generating Brain Organoids from Human Embryonic Stem Cells

Summary

This protocol was generated as a means to produce brain organoids in a simplified, low cost manner without exogenous growth factors or basement membrane matrix while still maintaining the diversity of brain cell types and many features of cellular organization.

Abstract

Human brain organoids differentiated from embryonic stem cells offer the unique opportunity to study complicated interactions of multiple cell types in a three-dimensional system. Here we present a relatively straightforward and inexpensive method that yields brain organoids. In this protocol human pluripotent stem cells are broken into small clusters instead of single cells and grown in basic media without a heterologous basement membrane matrix or exogenous growth factors, allowing the intrinsic developmental cues to shape the organoid's growth. This simple system produces a diversity of brain cell types including glial and microglial cells, stem cells, and neurons of the forebrain, midbrain, and hindbrain. Organoids generated from this protocol also display hallmarks of appropriate temporal and spatial organization demonstrated by brightfield images, histology, immunofluorescence and real-time quantitative reverse transcription polymerase chain reaction (qRT-PCR). Because these organoids contain cell types from various parts of the brain, they can be utilized for studying a multitude of diseases. For example, in a recent paper we demonstrated the use of organoids generated from this protocol for studying the effects of hypoxia on the human brain. This approach can be used to investigate an array of otherwise difficult to study conditions such as neurodevelopmental handicaps, genetic disorders, and neurologic diseases.

Introduction

Due to myriad practical and ethical limitations, there has been a great deal of difficulty in studying the human brain. While studies utilizing rodents have been critical to our understanding of the human brain, the mouse brain has many dissimilarities^1,2. Interestingly, mice have a neuronal density that is at least 7 times less than the primate brain^3,4. Although primates are closer to humans than rodents from an evolutionary standpoint, it is not practical for most researchers to work with them. The purpose of this protocol was to recapitulate many important features of the human brain using a simplified and less expensive method without the need for a heterologous basement membrane matrix or exogenous growth factors while maintaining brain cell diversity and cellular organization.

Formative work from the Sasai lab used the serum free culture of embryoid bodies (SFEBq) method to generate two- and three- dimensional neuronal cell types from signalized embryonic stem cells (ESCs)^5,6. Many human brain organoid methods have followed a relatively similar path from signalized ESCs^7,8. In contrast, this protocol starts with clusters of detached human ESCs (hESCs), similar to the initial steps of seminal work of the Thomson and Zhang laboratories prior to the plating steps^9,10 as well as the initial step of the brain organoid protocol of the Pasca laboratory before the addition of exogenous growth factors¹¹. Basement membrane matrices (e.g., matrigel) have been utilized in many brain organoid protocols and it has been shown to be an effective scaffold⁸. However, most commonly used basement membrane matrices do not come without complications as they co-purify with unknown quantities of growth factors with batch to batch variability during production¹². In addition, these matrices can complicate imaging, and increase the risk of contamination and cost.

While human brain organoids can be used to answer many questions, there are certain limitations to bear in mind. For one, starting from embryonic stem cells, organoids more closely resemble immature brains than aged brains and as such may not be ideal models for diseases that occur in old age, like Alzheimer's disease. Second, while our protocol found markers of forebrain, midbrain and hindbrain development which are useful to study the effect of a treatment or disease on cells from multiple brain regions in concert, other protocols can be followed to concentrate on specific brain regions^13,14. Finally, another limitation of organoid models is that of size, while the average length of a human brain approximately 167 mm, brain organoids made with the use of agitation grow up to 4 mm⁸ and the organoids formed by this protocol grow to 1-2 mm by 10 weeks. Nonetheless, this protocol provides an important tool to study human brain tissue and the interaction of multiple cell types.

Protocol

1. Stem Cell Maintenance

1. Maintain H9 hESCs on a layer of growth factor reduced basement membrane matrix (see the Table of Materials, henceforth simply referred to as matrix) according to the manufacturer's instructions.
   1. To coat one 6-well plate or one 10 cm dish, combine 100 µL of matrix with 5.9 mL of ice-cold Dulbecco's modified Eagle medium (DMEM)/F12 media. Wrap plates in paraffin film and store overnight at 4 °C. Use them on the next day for passaging cells after the excess matrix/media is aspirated.
2. Culture the cells week to week at approximately a 1:12 split ratio every 7 days. Maintain the cells using mTESR-1 media in a 37 °C, low oxygen incubator (5% O₂, 5% CO₂). Refresh media daily. Weed out differentiating cells from the culture between passages using glass tools.
3. H9 cells should be passaged four to six days prior to utilizing them to produce organoids. The cells should be passaged at approximately a 1:8 ratio of cell clusters. To do this, start by rinsing the cells with DMEM F12 media and dissociate the cells with a neutral protease (e.g., dispase, henceforth referred to simply as protease), rinse with DMEM/F-12, and plate as 30-60 cell clusters across 4 plates (6-well or 10 cm) at ~20% confluency. Two days prior to harvest, transition them to a regular incubator (21% O₂, 5% CO₂). Plates should reach ~80% confluency when starting organoid formation.

2. Dissociation of the hESCs for Organoid Culture

1. Aliquot the protease stock solution (5 U/mL).
   NOTE: We typically freeze down 1 mL aliquots at -20 °C for use over several months.
2. Dilute the protease stock solution to the working concentration by adding 1 mL of the stock solution plus 5 mL of DMEM/F12 for each 6-well or 10 cm plate of hESCs.
3. Aspirate and remove cell culture media, then cover the hESCs with the protease solution. Place plates in the incubator for 10-15 min or until the edges of the colonies round up and begin to separate from the matrix.
4. Tilt the plate, aspirate the protease solution, and gently wash the cells with DMEM/F12 three times. Use 2 mL/well for each wash when using a 6-well plate and 6 mL when using a 10 cm plate. Make sure colonies stay attached to the matrix when performing this step.
5. Add back about 1.5 mL of fresh mTESR media to each well (or 5 mL for a 10 cm plate) and flush the cells off the plate using gentle pipetting.
6. Using a 10 mL pipette, gently aspirate and dispense hESC within the plate until they reach approximately 1/30^th of their original size. Colony clusters should resemble ~250-350 µm sized squares at the completion of these steps.

3. Generation of Organoids

1. Transfer cells into a single ultra-low attachment T75 flask containing 30 mL of mTESR media without basic fibroblast growth factor (bFGF).
2. The next day, tilt the flask(s) such that the live cells pool in the corner (this may take 5-10 min on the first day, but will get quicker as the clusters get larger).
   NOTE: If there are a large number of cells that have adhered to the bottom of the flask at this step or any subsequent steps, transfer the cells to a new flask. It is normal to have a high population of dead cells for the first two days. When performing media changes, be sure to remove as much of the cell debris as possible.
3. Once the cells settle, aspirate off the media and dead cells leaving about 10 mL of media containing the live cells.
4. Add ~20 mL of low bFGF media (DMEM/F12 supplemented with 1x N2, 1x B27, 1x L-glutamine, 1x NEAA, 0.05% bovine serum albumin (BSA), and 0.1 mM monothioglycerol (MTG) supplemented with 30 ng/mL bFGF).
5. Check the cells on day 2. If most of the cells look healthy and bright, there is no need to do anything. However, if more than a third of the cells appear dark, replace the media (using the same tilting technique as in step 3.2) with ~20 mL of low bFGF media supplemented with 20 ng/mL bFGF.
6. On day 3, replace half of the media (using the tilting technique in step 3.2) with 20 mL of low bFGF media supplemented with 10 ng/mL bFGF.
7. On day 5, replace half of the medium (using the tilting technique in step 3.2) with 20 mL of neural induction media (NIM: DMEM/F12, 1x N2 supplement, 0.1 mM MEM NEAA, 2 µg/mL heparin).
   NOTE: If there are any large clusters of cells or organoids that are much larger than the others, they should be removed from the culture. Size is estimated by appearance under the microscope; for example, using an eyepiece with reticle. The majority of organoids are similarly sized (roughly 100 ± 20 µm). We removed organoids that were approximately 2x smaller or larger than the others.
8. Replace half of the medium (~15 mL) (using the tilting technique) with NIM every other day.
9. After 3 weeks in culture, add 100x penicillin/streptomycin to the media (NIM: DMEM/F12, 1x N2 supplement, 0.1 mM MEM NEAA, 2 µg/mL heparin) at a final concentration of 1x if desired. Refresh the media every other day.
   NOTE: In this fashion, we maintained the organoids for up to 6 months in culture.

4. RNA Extraction and Preparation

1. Gently extract approximately 15 organoids (depending upon size) from the flask using a 10 mL pipette and place into a 1.5 mL tube.
   1. Gently pellet the organoids in the centrifuge (200 x g for 1 min), and rinse with 1x Dulbecco's PBS (DPBS) three times.
2. Extract RNA using validated system or protocol (e.g., RNeasy kit).
3. Measure the optical density value of each sample at 260 and 280 nm.
4. Prepare cDNA using a validated system or protocol (e.g., iScript cDNA synthesis kit).
5. Perform qRT-PCR using pre-validated primers (Table 1) including at least one housekeeping gene.

5. Immunohistochemistry

1. Fixation
   1. Prepare a 4% paraformaldehyde (PFA) solution and place it at 4 °C.
   2. Using a sterile razor, cut the tip off of a sterile transfer pipette.
   3. Gently extract organoids using the cut transfer pipette, as they can be easily broken apart, especially when they grow large, and place them into a 6-well plate with additional media or DPBS.
   4. Tilt the plate, aspirate the media, and replace with 1x DPBS. Rinse the cells with 1x DPBS two additional times.
   5. Replace the DPBS with 4% PFA solution and place on a shaker at 4 °C.
      NOTE: While we fixed for 2 days (for small organoids) to 7 days (for organoids >3 months), shorter times (e.g., 16-24 h) may also be possible.
   6. Prepare 30%, 20% and 10% sucrose solutions in DPBS.
   7. After fixation in PFA, replace with the 10% sucrose solution and place on a shaker at 4 °C for 24 h.
   8. Replace the 10% sucrose with 20% sucrose and place on a shaker at 4 °C for 24 h.
   9. Replace the 20% sucrose with 30% sucrose and place on a shaker at 4 °C for 24 h.
2. Frozen sections
   1. Prepare a flat layer of dry ice and place a labeled plastic mold on top of it.
   2. Pour a thin layer of optimal cutting temperature medium (OCT) into the mold and let it start to harden (within a few seconds).
   3. Place a few organoids on the top of the OCT in the mold using a transfer pipette with the tip cut off and pay close attention to the location of the organoids.
   4. Slowly add in OCT until the mold is full and the organoids are covered. Let it harden completely for an additional 10 min.
      NOTE: While freezing over 10 min helps ensure ideal relative placement of multiple organoids for sectioning, it is possible to use an ethanol-dry ice mix or liquid nitrogen to freeze more quickly.
   5. Mark the relative location of the organoids with a marker to make it easier to find them when cutting.
   6. Place the molds in a bag or box and store at -80 °C until ready to cut the sections.
   7. Using a cryostat, slice 10 µm sections and place the tissue onto labeled, positively charged slides.
3. Staining
   1. Prepare blocking solution (0.3% Triton X-100, 4% normal donkey serum in PBS).
   2. Use a hydrophobic pap pen to draw around the perimeter of the tissue.
   3. Rinse the slides with PBS 3 times for 5 min each.
   4. Replace PBS with blocking solution for 1 h at room temperature.
   5. Replace the blocking solution with antibody solution (antibody at appropriate concentration, 0.1% Triton X-100, 4% normal donkey serum in PBS) at 4 °C overnight.
   6. On the following day, wash the slide 3 times with PBS for 10 min each.
   7. Replace PBS with the appropriate secondary antibody (at the appropriate concentration) diluted in antibody solution for 1 h at room temperature.
   8. Rinse 3 times for 10 min each time with 1x PBS.
   9. Apply the 4',6-diamidino-2-phenylindole (DAPI) stain and rinse three times for 10 min each with 1x PBS.
   10. Affix coverslips to the front of the slides with mounting solution, let dry at room temperature in the dark, and store in the dark at 4 °C.

Results

Figure 1 shows representative brightfield images of several time points to demonstrate what the cells/organoids look like throughout the different stages of the protocol. The hESCs were removed from the tissue culture plate, broken into small pieces, and placed in a T75 ultra-low attachment flask where they formed spheres. It is important to note that the cells look bright and similar in size, without dark, dying cells in the centers of these clusters. The cells were gradually weaned off bFG...

Discussion

Similar to other organoid models, this is an artificial system that comes with several caveats. Although there was little batch to batch variation in terms of overall expression levels, individual organoids did exhibit differences. For example, the location of Sox-2 positive areas were not identical in every organoid (Figure 3). While qPCR is suitable to look for overall changes in batches of cells, additional techniques such as single cell RNAseq will be utilized in future studies to gather more informa...

Materials

Name	Company	Catalog Number	Comments
Alexa Fluor 488 goat anti-mouse	Thermo Fisher Scientific, Waltham, MA, USA	A11029
Alexa Fluor 546 goat anti-rabbit	Thermo Fisher Scientific, Waltham, MA, USA	A11035
B27 Supplement	Gibco, Waltham, MA, USA	17504-044
bFGF	Life Technologies, Carlsbad, CA, USA	PHG0263
BSA	Sigma-Aldrich, St. Louis, MO, USA	A9647
BX43 microscope	Olympus, Shinjuku, Tokyo, Japan
DAPI stain	Thermo Fisher Scientific, Waltham, MA, USA	D1306
Dispase	STEMCELL Technologies, Vancouver, Canada	07913
DMEM/F12	Thermo Fisher Scientific, Waltham, MA, USA	11330-032
DPBS	Gibco, Waltham, MA, USA	10010023
FluroSave	MilliporeSigma, Burlington, MA	345789
GFAP antibody	NeuroMab, Davis, CA	N206A/8
Growth Factor Reduced Matrigel (Matrix)	Corning, Corning, NY, USA	356231
H9 hESCs	WiCell, Madison, WI, USA	WA09
Heparin	Sigma-Aldrich, St. Louis, MO, USA	9041-08-1
iQ SYBR Green Supermix	Bio-Rad, Hercules, CA, USA	1708880
iScript cDNA Synthesis Kit	Bio-Rad, Hercules, CA, USA	1708891
L-glutamine	Gibco, Waltham, MA, USA	25030-081
Monothioglycerol	Sigma-Aldrich, St. Louis, MO, USA	M6145
mTESR media	STEMCELL Technologies, Vancouver, Canada	85850
N2 NeuroPlex	Gemini Bio Products, West Sacramento, CA, USA	400-163
Nanodrop	Thermo Fisher Scientific, Waltham, MA, USA	ND-2000
NEAA	Gibco, Waltham, MA, USA	11140-050
Normal Donkey Serum (NDS)	ImmunoResearch Laboratories Inc., West Grove, PA, USA	017-000-121
OCT	Sakura Finetek, Torrance, CA, USA	25608-930
PFA	Electron Microscopy Sciences, Hatfield, PA	RT15710
qPCR machine	Bio-Rad, CFX96, Hercules, CA, USA	1855196
RNeasy kit	Qiagen, Hilden, Germany	74104
Sox2	MilliporeSigma, Burlington, MA	AB5603
TMS-F microscope	Nikon, Melville, NY, USA
Triton X-100	Sigma-Aldrich, St. Louis, MO, USA	T8787-100ML
Ultra-low attachment T75 flasks	Corning, Corning, NY, USA	3814