Isolation and Micromass Culturing of Primary Chicken Chondroprogenitor Cells for Cartilage Regeneration

Much of the skeletal system develops by endochondral ossification, a process that takes place in early fetal life. This makes the early stages of chondrogenesis, i.e., when chondroprogenitor mesenchymal cells differentiate to chondroblasts, challenging to study in vivo. In vitro methods for the study of chondrogenic differentiation have been available for some time. There is currently high interest in developing fine‐tuned methodology that would allow chondrogenic cells to rebuild articular cartilage and restore joint functionality. The micromass culture system that relies on embryonic limb bud‐derived chondroprogenitor cells is a popular method for the study of the signaling pathways that control the formation and maturation of cartilage. In this protocol, we describe a technique fine‐tuned in our laboratory for culturing limb bud‐derived mesenchymal cells from early‐stage chick embryos in high density (Basic Protocol 1). We also provide a fine‐tuned method for high‐efficiency transient transfection of cells before plating using electroporation (Basic Protocol 2). In addition, protocols for histochemical detection of cartilage extracellular matrix using dimethyl methylene blue, Alcian blue, and safranin O are also provided (Basic Protocol 3 and Alternate Protocols 1 and 2, respectively). Finally, a step‐by‐step guide on a cell viability/proliferation assay using MTT reagent is also described (Basic Protocol 4). © 2023 The Authors. Current Protocols published by Wiley Periodicals LLC.


INTRODUCTION
In synovial joints, the articulating bone surfaces are covered by a layer of hyaline cartilage offering smooth and frictionless movement during locomotion. The dominant cell type in articular cartilage is the chondrocyte, which secretes an abundant extracellular matrix (ECM) with a unique architecture consisting of a high amount of water and a dense mixture of macromolecules, such as type II collagen, aggrecan, other matrix constituents (e.g., hyaluronan). Unlike the other connective tissue types, cartilage is avascular and receives oxygen and nutrients by diffusion. For articular cartilage, its main source is synovial fluid. Mature articular chondrocytes are therefore characterized by a low metabolic rate and anaerobic energy metabolism (Goldring, 2006).
The loss of articular cartilage due to trauma, degeneration or inflammation is one of the most prevalent challenges of orthopedics, as this can develop into osteoarthritis (OA). The clinical symptoms of OA include pain, stiffness, and restricted motion of joints, impairing the physical activity and wellbeing of the patient. Given that mature articular cartilage has very limited capacity for repair, the chondrocyte is one of the main targets of regenerative medicine to enhance pathways potentially restoring the cartilage ECM (Solanki et al., 2021). Articular cartilage develops early during ontogenesis, making it challenging to study the process in detail in a laboratory setting (Cancedda et al., 2000). A better understanding of the molecular mechanisms which lead to the differentiation of hyaline cartilage in the limb mesenchyme may help us develop more efficient regenerative approaches and enhance the healing response of articular cartilage.
A well-accepted in vitro model to recapitulate embryonic chondrogenesis is the high density micromass cell culture established from embryonic limb buds (Ahrens et al., 1977;Takacs et al., 2023). Limb bud-derived chondrogenic progenitor cells aggregate and then spontaneously differentiate into chondroblasts and chondrocytes, which produce hyaline cartilage ECM in micromass cell cultures in vitro ( Fig. 1. steps of chondrogenesis). A high initial in vitro seeding density (1.5-2 × 10 7 cells/ml) of the chondroprogenitor cells is indispensable to mimic the condensation phase of early skeletogenesis that happens in vivo. The high seeding density creates a microenvironment which supports chondrogenesis in the micromass cultures and prevents differentiation of the other progenitor cells, such as epithelial or muscle progenitors. In chicken embryo-derived micromass cultures, overt chondrogenesis is completed in 6 days in culture (Takacs et al., 2023).
This method provides an opportunity to apply various chemical, physical, or biological interventions at different stages of primary chondrogenic differentiation. Staining of the differentiating or mature micromass cell cultures using dyes (such as dimethyl methylene blue, Alcian blue, and safranin O) that visualize the cartilage ECM, which is particularly rich in highly negatively charged glycosaminoglycans by metachromasia, is a cost-effective method for estimating the efficacy of chondrogenesis and ECM production. Generating samples for mRNA and protein-based expression analyses from the micromass cultures is relatively straightforward (Koff et al., 1988).
Here, we report a refined protocol on setting up chondrifying micromass cultures of embryonic limb bud-derived chondrogenic progenitor cells obtained from earlystage (Hamburger-Hamilton developmental stage 23-24;Hamburger & Hamilton, 1951) chicken embryos used in our laboratory (Basic Protocol 1). In addition, we provide a refined approach on the transient transfection of cells with siRNA constructs using electroporation prior to micromass culturing (Basic Protocol 2). Confirming cartilage ECM production by simple metachromatic staining procedures is necessary for evaluating the efficacy of chondrogenesis; we therefore supply a protocol for the qualitative and quantitative assessment of chondrogenic differentiation and cartilage matrix production using dimethyl methylene blue (DMMB) staining and image analysis (Basic Protocol 3). If DMMB staining is not the preferred choice, we provide protocols for the qualitative assessment of cartilage matrix production using Alcian blue 8GX (Alternate Protocol 1) and safranin O staining (Alternate Protocol 2). Finally, a step-by-step guide on how to determine mitochondrial activity in cells of chondrifying micromass cultures with the MTT assay is also included (Basic Protocol 4).

STRATEGIC PLANNING
Here we provide a list of considerations in order to carefully prepare for the execution of the protocols.
Consider the full timeline of the experiment (Fig. 2). Keep in mind that a single experiment takes 6 days to complete. Plan the time of setting up the micromass cultures in a way that none of the critical days, in terms of workload (e.g., day 6), fall on a weekend. Obtain fertilized eggs no more than 7 days ahead of the experiment. Keep them in a cool place (∼18°C), away from direct sunlight. Choose a reliable poultry farm to obtain fertilized chicken eggs. They should use a single breed, e.g., the Ross hybrid breed. Given the key importance of embryos to be in the right developmental stage , a very precise timing of incubation in the egg hatcher is required. Although this depends on the egg hatcher model, it may be necessary to start the incubation late at night. For this, obtaining a plug-in timer or a programmable egg hatcher is recommended. During the incubation in the egg hatcher (∼4.5 days), maintaining appropriate humidity level (∼90%) is important. Make sure to refill water into the egg hatcher to supply humidity for extended periods (e.g., over the weekend). Alternatively, manually top up water every second day (depending on the egg hatcher model). Designate a dark room (or a dark corner of a room) for candling the eggs. Depending on the experimental plan (and the number of embryos to be processed), at least two staff members are required for setting up the micromass cultures.

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Current Protocols

Figure 2
Timeline of a typical micromass experiment.
One person is tasked with cracking the eggs and fishing out the embryos while the other person is dissecting the limb buds under a microscope. In case a high number of embryos need to be processed (e.g., over ∼50), involvement of three staff members is recommended. Egg cracking and limb bud dissection should not take more than 1 hour to avoid unnecessary stress to the chondroprogenitor cells. Check local ethical guidelines regarding work of early-stage chicken embryos. Before cracking, disinfect eggs by wiping them generously with 70% ethanol.
Once embryos are removed from the eggs, they should be kept in a sterile environment (in a laminar flow cabinet). Note that eggs are usually sourced from poultry farms. Please check whether any decontamination procedures are in place (e.g., treatment with ozone) to reduce the risk of infections (e.g., contagious poultry diseases that may be transmitted through eggs) of cell cultures in vitro.

NOTE:
All protocols involving animals must be reviewed and approved by the appropriate Animal Care and Use Committee and must follow regulations for the care and use of laboratory animals.

MICROMASS CULTURE OF CHICK EMBRYONIC LIMB BUD-DERIVED CELLS
Embryonic limb bud-derived chondrifying micromass cell cultures are used as an in vitro experimental model for hyaline cartilage formation. The chondroprogenitor cells in micromass cultures first proliferate and then differentiate into matrix-producing chondroblasts during the first 3 days of culture, and a well detectable hyaline cartilage-specific ECM forms by the sixth day of culture. To set up the chondrifying cultures, the distal parts of the fore and hind limbs of early-stage (Hamburger-Hamilton stages 23-24; ∼4.5 dayold; Hamburger & Hamilton, 1951) chicken embryos are dissected and pooled, the limb Takács et al.

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Current Protocols buds are dissociated using trypsin, the cell density is adjusted to 1.5 × 10 7 , and micromass cultures of required size are established.  ATTENTION: planning is advisable. Start the egg hatcher on a day so that the critical days of the experiment do not fall on a weekend. In our experience, the egg hatcher starts on a Friday at 2:00 am, and the rest of the procedure is carried out on the following Tuesday morning.

Materials
2. After the 104 hr incubation time, switch off the hatcher and remove the eggs. Use 70% (v/v) ethanol to wipe clean the surface of the eggs. Immediately afterwards, in a dimly lit room, use the egg candling device to assess the presence and position of the embryo within the egg ( Fig. 3A; Video file 1).
A proportion (usually ∼10%) of eggs will not contain an embryo, and these should be discarded. A graphite pencil is suitable for marking the position of the well vascularized chorioallantois area around the embryo on the shell (Fig. 3A). Some eggs will contain embryos that are not attached to the eggshell, this is typically the sign of reduced vitality, and we recommend against using these embryos.
We recommend putting the stereomicroscopes and dissecting tools (forceps, etc.) into the laminar hood which are required for further steps of the protocol.
3. Use a pair of curved micro forceps to crack the eggshell immediately next to the circular marking that indicates the position of the embryo. Remove enough of the shell to access the embryo with a filter spoon (Fig. 3B) and use it to remove the embryo ( Fig. 3C; Video file 2).
ATTENTION: Try to avoid removing the egg yolk together with the embryo as the lipid content of the egg yolk may interfere with limb bud removal by making the solution turbid.

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Current Protocols Video 1 Egg candling.
Video 2 Egg cracking and embryo retrieval.
This is a wash step, which helps rid the embryos of unwanted egg components, such as egg white, egg yolk, and any membranes.
5. Transfer the desired number of embryos (between 5 and 10) into separate Petri dishes containing 10 ml prewarmed (37°C) gentamycin-supplemented CMF-PBS and place the Petri dish under the stereomicroscope (  7. Once all limb buds from all embryos have been gathered, remove unwanted tissue debris (e.g., epithelial membranes, inappropriate parts of the embryo) from the Petri dish with the aid of a stereomicroscope. Remove the CMF-PBS by careful pipetting and transfer the limb buds to 15 ml trypsin-EDTA solution preincubated at 37°C in a 30-ml weighing bottle with a glass lid (Video file 5). Incubate 55 min at 37°C in a CO 2 incubator.
It is a good idea to gently mix the dissociating limb buds in the trypsin-EDTA solution by careful pipetting periodically or at least once approximately halfway through the incubation period (Video file 6).
Do not allow the digestion to proceed longer than 1 hr, even if the limb buds look intact. Excessive digestion will compromise the integrity of the cells and they will die.
8. Carefully dissociate the limb buds using a 5-ml pipette by careful pipetting several times. Terminate the enzymatic dissociation of limb buds by adding an equal volume of fetal bovine serum (Video file 6). Mix well and transfer the solution to a 50-ml centrifuge tube, and centrifuge 5 min at 150 × g, room temperature, to pellet the cells.
9. Carefully remove supernatant with a vacuum aspirator and resuspend pellet in 30 ml Ham's F12 culture medium supplemented with 10% FBS.
10. Centrifuge cell solution 10 min at ∼150 × g, room temperature. Resuspend cells in an adequate volume of Ham's F12 culture medium supplemented with 10% FBS.
11. Filter the cell suspension through a 20-μm nylon net filter (Video file 7).
This step is recommended to rid the cell suspension of larger clumps of tissue that failed to dissociate, and to generate a single-cell suspension for cell counting.

Setting a low volume of medium for resuspension is a safer option but it might make cell counting unnecessarily difficult (this, depending on the method, also increases the likelihood of mistakes). On the other hand, a too high resuspended volume may result in
Takács et al.

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Current Protocols Video 6 Dissociation of limb buds.
12. Determine the cell concentration of the suspension, preferably using an automated cell counter (Video file 8) and adjust the final volume with the same medium to achieve the desired concentration of 1.5 × 10 7 cells/ml.
It is ideal to set a 10× dilution for faster and more precise counting. Using trypan blue to assess cell vitality is optional (but recommended).
13. Inoculate droplets (of various size ranging from 15 μl to 100 μl) into 35-mm cell culturing dishes or 24-well plates that fit the purpose of the experiment (Video file 9). Allow cells to attach to the surface for 2 hr in a CO 2 incubator (37°C, 5% CO 2 and 90% humidity). 14. Flood cultures with medium (Ham's F12 supplemented with 10% FBS, 0.5 mM Lglutamine, and 1% penicillin/streptomycin solution) after the 2 hr attachment period. Make sure to add L-ascorbic acid (vitamin C) at 50 mg/L final concentration to the culture medium at this stage. Change the culture medium every other day for the duration of the experiment.

TRANSFECTION OF CELLS WITH SIRNA CONSTRUCTS USING ELECTROPORATION PRIOR TO MICROMASS CULTURING
High efficiency transfection of foreign nucleic acids into this model is particularly challenging. Transfections reagents that are gentle on limb bud cells (such as Lipofectamine 2000) usually result in low (∼25%) transfection efficacy, whereas other methods (such as Amaxa nucleofector technology) can achieve much higher efficiency (up to 97%) but at the same time can induce high rates of apoptosis (Bobick et al., 2014). Due to the necessity of plating droplets at a high initial cell density, the best window of opportunity for this intervention is while cells are in suspension (following step 11 of Basic Protocol 1). Below, we describe a modified version of the method described by Bobick et al. (2014). This method consists of square wave pulse electroporation of cells resuspended in a protective sucrose buffer containing siRNA. It results in a comparatively high transfection efficiency without significant harm on cell viability, and chondrogenic differentiation potential. Following electroporation, cells are pooled, resuspended in Ham's F12 culture medium, counted, and inoculated into cell culture dishes or plates that fit the purpose of the experiment.

Additional Materials (also see Basic Protocol 1)
Modified sucrose electroporation buffer (see recipe) Pre-and/or custom-designed siRNAs, including controls (Silencer Select, Thermo Fisher Scientific) 2.0-mm gap sterile electroporation cuvette (VWR, cat. no. 732-1136) Square wave electroporation system (BTX model ECM 830) Ice 1.5-ml Eppendorf tubes 1. Generate a single cell suspension of chondrogenic progenitor cells and determine cell density according to steps 1-12 of Basic Protocol 1.
2. Take the required amount of cell suspension, centrifuge 5 min at 150 × g, room temperature, and remove supernatant with a vacuum aspirator.
The sucrose content of the medium provides a protective environment that increases the viability of cells during and after electroporation (Bobick et al., 2014). 4. Add 2 μg siRNA to every 250 μl of cell suspension, mix well by pipetting, and transfer the entire volume to a 2.0-mm gap sterile electroporation cuvette.
Refrigerate the sterile electroporation cuvettes at 4°C overnight prior to use. Dilute and aliquot siRNA in advance according to the resuspension protocol recommended by the manufacturer, resulting in 100 μM final concentration (e.g., resuspension volume is 50 μl for 5 nmole siRNA, which consists of 10 μl 5 × siRNA buffer, and 40 μl nuclease-free H 2 O).

5.
Incubate cuvettes containing the solution at 4°C for 5 min then administer pulses with the electroporation system according to the following settings: three 400 V pulses, 150 s in length, at 100 ms intervals ( Fig. 4A-C).
ATTENTION: placing the electroporation cuvettes on ice immediately after pulsing is pivotal to the survival rate of transfected cells.
6. Incubate cells following electroporation at 4°C for 10 min, then at room temperature for 5 min, and at 37°C for 5 min to allow cells to sink to the bottom of the cuvette.
7. Pool the lower four-fifths of the suspension from each cuvette that belongs to the same experimental group (Fig. 4D) in 1.5-ml Eppendorf tubes, and pellet cells by centrifugation 5 min at 150 × g, room temperature.
Independent experimental groups are centrifuged and handled in separate centrifuge tubes.
8. Continue with step 12 of Basic Protocol 1.

QUALITATIVE AND QUANTITATIVE ASSESSMENT OF CARTILAGE MATRIX PRODUCTION USING DIMETHYL METHYLENE BLUE STAINING AND IMAGE ANALYSIS
1,9-dimethyl methylene blue is a cationic dye which preferentially binds to negatively charged sulfated glycosaminoglycans (sGAG). This staining technique is widely used in cartilage research (Templeton, 1988). The dye shows purple metachromasia in the presence of polyanionic sGAG. The pH must be kept at pH 1.5 to avoid erroneous estimation of sGAG content in newly produced cartilage ECM due to interference caused by polyanions such as hyaluronan, DNA, or RNA (Zheng & Levenston, 2015).

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Current Protocols initial exposure. Subsequent exposure may cause severe allergic reactions of the skin, eyes, and respiratory tract. Long-term or repeated exposure to low levels in the air or on the skin can cause asthma-like respiratory problems and skin irritation. Acute exposure can be highly irritating to the eyes, nose, and throat. Because of the serious potential hazards, precautions must be taken to eliminate or reduce the potential risk for exposure, e.g., by wearing appropriate PPE, and adhering to safe use practices. We have introduced three different indices for quantifying metachromasia based on the color of digital microscopic images. Since previous quantitative measures of metachromasia are based on comparison of optical densities at two different wavelengths (Hattori et al., 1996) or at two different polarization angles (Heuft & Bohm, 1978;Romhanyi, 1963), they are not easily applied under conditions available in ordinary cell biological laboratories. Since the novel indices are based on quantifying color, a brief introduction to the RGB color model is required. An arbitrary color can be defined by a number triplet with the first, second and third values specifying the intensity of the red, green, and blue colors, respectively. Color intensities are usually specified on a scale between 0-255. This color model, named RGB, can be represented by a cube in which the three spatial coordinates correspond to the intensity of either the red, green, or blue colors. Consequently, every different spatial position in the cube corresponds to a certain color (Fig. 5A).
The first metachromasia index is described by the following equation: where d is the distance between the points representing the color of the most purple, most metachromatic area in an experiment and the pixel under investigation (Fig.  5B). 255Ý3 is the main diagonal in the RGB cube, i.e., the largest possible distance in the cube. The second metachromasia index is defined according to the following equation: where d max is the distance between the most purple, most metachromatic color and the color of the gray background in the RGB cube. In the third index, d 1 is the distance between the projection of the color of the pixel under investigation on the line connecting the most purple and the gray colors in the RGB cube: . Follow these steps to evaluate the images (Fig. 5C) Figure 5D. c. Identify the most purple (most metachromatic) color, and the gray color (the background) by clicking on 'Pick purple' and 'Pick gray,' respectively. You can also manually specify these colors by entering values (

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Current Protocols e. The mean values in the blue fields in the red box can be copied to the clipboard by clicking on 'Copy to clipboard.' f. The results can be saved by clicking on 'Save results.' The pixelwise values of the three indices can be saved in three images, either in TIFF or JPG format. The user must specify a base name that is appended by '_1,' '_2' and '_3.' The three histograms are also saved in a single comma-delimited file.

QUALITATIVE ASSESSMENT OF CARTILAGE MATRIX PRODUCTION USING ALCIAN BLUE STAINING
Alcian blue staining technique is an alternative method to detect the sulfated GAGs in the cartilage ECM. The stained tissue parts are cyan or greenish-blue, and this staining effect is called "Alcianophilia." Alcian blue staining is pH sensitive; at pH 1.0 it stains sulfated GAGs only, but at pH 2.5 it also stains hyaluronan and sialomucins (Green & Pastewka, 1974).

QUALITATIVE ASSESSMENT OF CARTILAGE MATRIX PRODUCTION USING SAFRANIN O STAINING
Safranin O is a metachromatic dye which binds to highly polyanionic structures. This dye is widely applied to visualize GAGs and proteoglycans with an orange color (Kiraly et al., 1996;Rosenberg, 1971).

0.1% (w/v) safranin O solution (see recipe)
NOTE: Each step is performed at room temperature in a 24-well plate. Use ∼500 μl of solution per well at each step. Handle liquids using plastic Pasteur pipettes. To aspirate solvents, a vacuum aspirator may be used. 11. Let the specimens air-dry for 24 hr before examining under a microscope.

MEASUREMENT OF MITOCHONDRIAL ACTIVITY WITH THE MTT ASSAY
The MTT assay is a colorimetric method suitable for the analysis of cell viability, cell proliferation, and cytotoxicity. Living cells can reduce the yellow tetrazolium salt in the MTT reagent by mitochondrial oxidoreductase enzymes. This metabolic activity will lead to the formation of purple formazan crystals, which can be dissolved using a solubilization solution. At the end of the reaction, the absorbance of the purple-colored solution at a specific wavelength is measured by a spectrophotometer (van Meerloo et al., 2011).

Additional Materials (also see Basic Protocol 1-3)
MTT ( 6. At the end of the 2-hour-long incubation, remove the 24-well plate from the CO 2 incubator and aspirate the medium from the wells.
Make sure to completely remove the medium from the wells, as this would detrimentally affect the solubilizing ability of the solubilization solution.
7. Add 500 μl of the solubilization solution to each well.

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Current Protocols 10. Insert the 96-well plate into the microplate reader. Measure the optical density of the samples at 570 nm.
11. Analyze the optical density readings with the help of Microsoft Excel or other data analysis software.

Calcium and magnesium free phosphate buffered saline (CMF-PBS) stock solution (pH 7.2), 10×
80 g NaCl 3 g KCl 1.84 g Na 2 HPO 4 12H 2 O 0.2 g KH 2 PO 4 20 g D-glucose Bring volume to 1000 ml with deionized H 2 O Adjust to pH 7.2 with HCl or NaOH Sterilize using a 0.2-micron filter in a laminar flow hood Store up to 3 months at 4°C

Modified sucrose electroporation buffer
Add 272

Background Information
The original article that described the methodology of establishing chondrogenic micromass cell cultures from the limb buds of early-stage chick embryos was published 50 years ago (Ahrens et al., 1977). The underlying principle is that limb bud mesoderm cells, when retrieved from a specific developmental stage, are cultured in vitro in a sufficiently high seeding density in a microdrop. The cells form aggregates or precartilage nodules during the first 1-2 days and differentiate into chondrocytic cells, which produce an extracellular matrix from culture day 3. A critical cell mass is necessary for the chondroprogenitor cells to commence chondrogenic differentiation and extracellular matrix formation; when the mesenchymal cells are seeded in low-density, they acquire a fibroblastic phenotype and are incapable of chondrogenesis (Handschel et al., 2007).
In vitro cartilage regeneration methods rely on the re-aggregation approach, during which cells are first dissociated and then the dispersed cells are re-aggregated either by selforganization or into cellular spheres in a scaffold-free manner (Handschel et al., 2007). The capacity to spontaneously form regular, spatially organized patterns of cartilage nodules is preserved in the limb bud-derived progenitor cells.
A major advantage of this system lies in its simplicity. It allows for monitoring the temporal pattern of chondrogenesis in vitro (Takacs et al., 2023). It is also a highly adaptable and well-characterized model (Takacs et al., 2023) that has been used for assessing a variety of conditions including the application of soluble factors or mechanical stimuli (Rolfe et al., 2022). However, there are some major disadvantages of this method that are worth noting. In chondrogenic micromass cultures, hyaline cartilage is present in the form of cartilage nodules, interrupted by internodular areas where a mixture of mesenchymal cells, fibroblasts and chondroprogenitors is present, and the peripheral area of the round cell cultures does not contain any cartilage.
Thus, micromass cultures cannot be regarded as 'pure' cartilage, although the predominant tissue is hyaline cartilage. Another unfavorable feature of this method is that the hyaline cartilage which forms in the cell cultures is primarily transient cartilage, which undergoes chondrocyte hypertrophy; essentially recapitulating the main steps of endochondral ossification in the developing limbs. Gene expression profiling supports rapid chondrogenesis and transition to hypertrophy (Rolfe et al., 2022;Takacs et al., 2023). Cartilage formed in micromass cultures does not exhibit the unique organization of articular hyaline cartilage matrix. Another inherent feature of this model is that there are known differences between chicken and mammalian cartilage (Eyre et al., 1978), which should be considered when applying data generated using this model to mammalian or human chondrogenesis.
In summary, if the inherent limitations of this in vitro chondrogenesis model are considered, the primary micromass culture system still can provide an answer to questions which are unexplored in cartilage biology.

Critical Parameters
This model relies on freshly isolated progenitor cells from the developing limb buds of early-stage chicken embryos. Therefore, there are a couple of critical factors that influence the protocol and to which special attention should be paid. These are listed below. Consider the number of fertilized eggs to be procured for the experiment. A proportion (usually ∼10%) of eggs will not contain an embryo (or embryogenesis fails in an early period). Given that the eggs are sourced from a poultry farm, they can be covered in mud and farm detritus, and they must undergo a general rinse to clean the eggshell, followed by a generous wipe with 70% ethanol. The decontamination procedures are very important to reduce the risk of infections of the cell cultures in vitro. Adding gentamycin to CMF-PBS and Takács et al.

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Current Protocols trypsin is a good idea to reduce the risk of infections. The correct developmental stage of the embryo is of key importance (Ahrens et al., 1977). Embryos should be at Hamburger-Hamilton developmental stage 23-24 (Hamburger & Hamilton, 1951). Limb buds obtained from embryos earlier or later than this stage are unsuitable for micromass culturing. Therefore, we provide readers with a detailed morphological description of the external features of chicken embryos (mainly focusing on the maturation of limb buds) between Hamburger-Hamilton developmental stages 19 and 25 (see Supporting Information). The time window for harvesting the limb buds from embryos is ∼60 min, as the cells in the isolated limb buds start losing vitality. Unskilled personnel should only remove and collect limb buds under supervision. It is very important to remove and collect the distal parts of the developing limb buds only, as removing parts of the body wall and embryonic membranes together with the limb bud mesenchyme will detrimentally affect trypsin dissociation and may introduce inappropriate cell populations into the micromass cultures. Double-checking the pooled limb buds for any remaining membranes and inappropriate contaminants (e.g., parts of the tail, heart, or unidentifiable body parts) is a good idea. Do not allow the trypsin digestion of the limb buds to proceed longer than 60 min, even if the limb buds look intact. Excessive digestion will compromise the integrity of the cells and they will die. A critical cell mass is necessary to proceed with chondrogenic differentiation and extracellular matrix formation. Therefore, adjusting the appropriate seeding density (1.5 × 10 7 cells/ml) is critically important. When plating the cells, avoid air bubbles or irregularly shaped drops, as this will interfere with micromass culture morphology and chondrogenesis. Let the cells attach to the surface of plasticware/glassware for 2 hr. When feeding the cells, add medium to freshly established micromass cultures carefully. Let the culture medium run down the sides of the wells or Petri dishes so as not to disturb the cells. The cell attachments are very tenuous at this point and if the medium is introduced too quickly, the cells will be washed off the plate and die, adversely affecting chondrogenesis. Check the surface of the plasticware before culture. Surface (cell culture) treated plasticware is necessary for proper adhesion of micromasses. In our experience, Nunc, Eppendorf, Thermo, and Greiner brand tissue culture plasticware (6-, 24-well tissue culture plates, as well as 35-mm Petri dishes) appear to work the best. Adding 272 mM sucrose to the electroporation medium provides a cytoprotective environment, which appears critical in achieving high transfection efficiency/low cytotoxicity (Bobick et al., 2014). In our experience, transferring the electroporation cuvettes to a cold environment (i.e., on ice) immediately after pulsing proved to be of cardinal importance for the high survival rate of transfected cells. Providing living cells with enough incubation time following electroporation to sink to the bottom of the cuvette is also of high importance. Table 1 summarizes common problems with the protocols, their causes, and potential solutions.

Understanding Results
If the protocol is conducted properly, chondrogenesis proceeds spontaneously in the micromass cultures. Chondrogenesis progresses through the following stages: day of seeding (day 0), proliferation (culture days 1 and 2), chondrogenic differentiation (culture days 3 and 4), matrix production (starting from culture day 4). Around culture day 10, hypertrophic chondrocytes appear within the cultures.
Cell culture morphology nicely displays the dynamic development of micromass cultures (Fig. 6). Chondrogenesis commences in the center of micromass cultures (C) and gradually proceeds towards the periphery (P) as shown on the low-power magnification phase contrast photomicrographs ( Fig. 6B  and 6C). Proliferating chondroprogenitor cells are shiny, circular cells in Figure 6B. Their number gradually decreases in more mature cultures as they differentiate to chondroblasts. Chondrogenic cells form clusters Takács et al.

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Current Protocols   6B). Over time, the nodular regions tend to increase in size and number and (especially towards the center of culture) they coalesce, leaving fewer internodular regions (Fig. 6). In mature micromass cultures, an abundant, hyaline cartilage-like extracellular matrix is produced by culture day 6 as revealed by staining with the metachromatic dyes DMMB, Alcian blue, and safranin O (Fig. 7A). The method described in Basic Protocol 1 relies on pooling cells derived from forelimbs and hindlimbs. However, it is possible to micromass culture forelimbs and hindlimbs separately, although the forelimb and hindlimb mi-cromass cultures display differences in patterning and size of chondrogenic nodules (Butterfield et al., 2017).
The cells show active proliferation during the first culture days, which then gradually declines, as determined by MTT assay (Fig. 7B).
It is nearly impossible to achieve a high transfection efficiency of primary chondrogenic cells in micromass cultures. However, based on our experience using this protocol (Basic Protocol 2), it is possible to achieve a ∼50% downregulation at 48 hr post electroporation, as determined by quantitative RT-PCR. Please note that this figure is heavily dependent on the basal expression levels of the target gene (Fig. 7C).
It should also be noted that in contrast to conventional monolayer cell cultures, where cells grow only in two dimensions on the flat surface of a cell culture dish, micromass cultures are 3 dimensional "organoids", in which the cells form aggregates in multiple Takács et al.

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Current Protocols Figure 6 Microscopic cell culture morphology. (A) Morphology between culture days 1 and 15 (as indicated by numbers in right lower corner of each photomicrograph). As cultures mature, the thin layer of transparent culture becomes thicker, multilayered and glassy in appearance, making the term 'hyaline,' meaning transparent or glassy (hyalos in Greek) applicable. Photomicrographs were taken using a Leica stereomicroscope; 1× magnification. Scale bar, 500 μm. (B) Detailed morphology of a micromass culture on day 3 of culture. The phase contrast images clearly show the differences between the center (C) and periphery (P) of the culture. In the center, cells are polygonal and enter the process of chondrogenic differentiation. At the periphery, cells are more elongated with a fibroblastic morphology. In vitro chondrogenesis commences in the center and progresses towards the periphery. Day 3 cultures still display proliferating (spherical, shiny cells; black arrows). Nodule formation (N) takes place around culture day 3. Nodules are surrounded by internodular areas containing elongated cells. (C) Detailed morphology of a micromass culture on day 3 of culture. The nodules (N) reach the periphery (P) of the cultures and coalesce in the center. Due to the increased height of the culture, it is challenging to provide sharp image from each structure present in the view field. This is especially evident in the picture taken from the center (C) with a 40 × magnification. For panels B and C, photomicrographs were taken by phase contrast microscopy (Leica). Original magnification was 0.8×, 10×, and 40×. Abbreviations: C, center; P, periphery; N, nodule. layers, with a considerable amount of cartilage ECM also present in the cultures. The multilayered nature of the micromass cultures, and the abundant matrix in mature cultures may interfere with downstream applications such as single cell imaging in situ or live-cell calcium imaging, patch clamp experiments, and immunocytochemistry (since antibodies may bind to the matrix). When such applications are considered, the fact that older micromass cultures contain more cells and a higher amount of matrix should be taken into account.

Time Considerations
Given the long timeline of the experiment, including preparation time and incubation of eggs, advance planning is highly advisable.
Procure eggs a day in advance of incubation. Incubation of eggs in egg hatcher: 104 hr. Preparation time (prepare solutions such as CMF-PBS, trypsin, cell culture medium; disinfect laboratory surfaces, glassware, and dissection tools): ∼2 hr. Egg candling, cracking, embryo retrieval: ∼1 hr. Dissection of limb buds (can start parallel with egg cracking and embryo retrieval): showing an initially intense proliferation, which reaches a plateau in more mature cultures. The chart shows the time course of mitochondrial activity (cell viability) of micromass cultures; data points are average values of 6 biological replicates ± standard deviation (SD). (C) Transfection efficiency in chicken limb bud-derived chondroprogenitor cells using Basic Protocol 2. Normalized average expression data for two genes (EBF1 and ATOH8) are shown (left axis; EBF1 is in the 55 th percentile, ATOH8 is in the 74 th percentile ranked according to normalized expression data across all annotated genes in the dataset (BioProject IDs: PRJNA817177 and PRJNA938813; EMBL-EBI accession number: E-MTAB-12770)). Data points are average values of 3 biological replicates ± SD. Relative expression levels following transient silencing are also shown (53% and 85% of non-targeting control, respectively; right axis). Please note that the achieved silencing [as determined by RT-qPCR on culture day 2 (D2)] is dependent on the basal expression level of the target gene. ∼1 hr maximum. Dissociation of limb buds in trypsin-EDTA: 50 min. Washing & centrifuging cells, single cell suspension, plating of micromass cultures: ∼1 hr. Transient transfection of the cells, together with preparation time (Basic Protocol 2) takes ∼1 hr. Allow ∼2 hr for the cells to adhere. Feeding of plated micromass cultures: ∼30 min (depending on experimental setting). Micromass culturing takes 6 days for overt chondrogenesis to take place (cultures can be maintained for longer periods of time; see Fig. 6). Fixing and staining the cultures (Basic Protocol 3 and Alternate Protocols 1 and 2), including preparation time, takes ∼2 hr. An MTT assay, including preparation and 2-hour incubation time, solubilization of formazan crystals, and photometric quantification (Basic Protocol 4) takes ∼4 hr.
For more details and a visual representation of the timeline, please see Figure 2.