Lineage Tracing of Inducible Fluorescently-Labeled Stem Cells in the Adult Mouse Brain

All methods described here have been approved by the Institutional Animal Care and Use Committee (IACUC) of The Ohio State University.

1. Creation of a lineage tracing Tet-On mouse line, breeding strategies, and genotyping for experimental cohort mice

NOTE: Here, we outline creation of a Tet-On mouse system with Rosa-mTmG as the fluorescent reporter gene that undergoes Cre recombination, however various other Cre-recombination-driven reporter lines can be utilized with the Tet-On system as well.

1. Set up a mating cage of one oTet-Cre (+/-) male with one or more R26 (mTmG) (+/+) females. See Figure 1 for an overview of the experimental mouse mating setup.
2. Resulting pups (F1) will be oTet-Cre (+/-) or oTet-Cre (-/-) and R26 (mTmG) (+/-). Perform genotyping according to the primers in Supplementary Table 1 and protocols in Supplementary Table 2.
3. At weaning, perform germline testing by taking an ear punch from each weanling. Examine the ear punch under an epifluorescent microscope in the GFP/FITC spectrum to determine whether the animal had undergone germline recombination in development. If so, the animal will express GFP in every cell, and the ear punch will fluoresce with mGFP signal. These animals cannot be used for matings or experiments.
   NOTE: Ear clips from mice that are not germline show only endogenous fluorescence under the microscope (Figure 1C), while germline animals show a bright GFP signal (Figure 1D). Control ears can be from mice that do not contain fluorescent transgenes. As a note, ear hairs have high endogenous fluorescence and must not be used as a determining factor (Figure 1C). Additionally, differences in endogenous fluorescence are observed between mice of white vs. black coat color, and controls from mice of the same coat color as mice being tested will need to be used in this analysis.
4. Mate one male oTet-Cre (+/-) x R26 (mTmG) (+/-) with one or more female oTet-Cre (-/-) x R26 (mTmG) mice.
5. The resulting generation (F2) will be ¼ R26 (mTmG) (+/+) and ½ Cre (+/-). Cross these animals with mTert-rtTA (+/+) animals by mating one oTet-Cre (+/-) x R26 (mTmG) (+/+) with one or more mTert-rtTA (+/+) animals.
   NOTE: We used mTert-rtTA (+/+) animals, but this process can be performed with any rtTA mouse line³⁰.
6. For experimental animals, set up breedings to generate mTert-rtTA (+/+) or (+/-) x oTet-Cre (+/-) x R26 (mTmG) (+/+) or (+/-).

2. Doxycycline induction of Cre and pulse-chase design

1. When animals are at the correct age for the study to begin, replace their drinking water with dox water: drinking water containing 5% sucrose and 2 mg/mL doxycycline hyclate. Use Cre-negative animals that receive the same doxycycline pulse-chase as experimental animals as a control group for the resulting fluorescent expression patterns.
   1. After preparation, store dox water at 4 °C for up to 1 month. See Table 1 for more information. Dox water may be administered to animals in drinking bottles for up to 5 days before being changed out, as fungal growth can occur if dox water is left out at room temperature for longer periods of time.
      NOTE: A doxycycline pulse of at least 2 days is required for induction of mGFP signal when utilizing the mTmG transgene (initial mTmG paper).
      NOTE: We have not tested doxycycline concentrations other than 2 mg/mL. Previous literature has identified that this concentration has the highest effects when administered through drinking water²⁹. The amount of doxycycline delivered may differ depending on the amount of drinking done by each animal. Doxycycline injection or gavage may also be done in order to be consistent between animals⁴⁰.
2. After the pulse period has ended, remove dox water from the cages and replace with normal drinking water. The animals will now be in the 'chase' period until death.

3. Transcardial perfusion and brain dissection

1. Remove a mouse from its cage and restrain the animal using the left thumb and forefinger. Inject the animal via intraperitoneal (i.p.) injection with a solution of 100 mg/mL ketamine and 20 mg/mL xylazine in 0.9% sterile saline for a final concentration of 200 mg/kg ketamine and 20 mg/kg xylazine. See Table 1 for more information.
   CAUTION: The protocols for anesthesia may differ depending on the country. Additionally, there are effects of anesthetics on the brain including changes in microglial morphology and action and corticosterone levels that must be understood when performing perfusions for the purpose of brain collection and analysis^41,42.
2. After approximately 1-2 min the animal will lose its righting reflex. To test this, simply roll the animal on its back and if it cannot roll back onto its feet, it has lost the righting reflex.
3. After approximately 5-10 min the animal will lose its pedal reflex. To check the pedal reflex, use the thumb and forefinger to pinch each of the animal's feet. If there is no response, in the form of a jump or muscle spasm, use the nails of the thumb and forefinger to pinch each of the animal's feet. If there is no response, the animal has lost the pedal reflex.
   NOTE: If the animal responds to the pedal reflex for greater than 15 min after injection, an additional injection of ½ of the initial ketamine/xylazine injection may be required. Age, sex, genotype, phenotype, and treatment may affect the response of the animal to this drug cocktail.
4. After the righting and pedal reflexes are lost, test the ocular reflex by lightly contacting the mouse eyeball with a gloved finger. A lack of response indicates that the ocular reflex has been lost.
5. Secure the mouse in a supine position with paws pinned to a pinnable work surface.
6. Make an incision through the skin with surgical scissors along the thoracic midline approximately 1-2 cm below the xiphoid process. Cut superior until at the xiphoid process.
7. Grasp the cartilage of the xiphoid process with forceps. Insert scissors and cut through the musculature and ribcage diagonally along the mouse's right side to the level of the clavicles.
   1. Raise the thoracic musculature with forceps and slice through the diaphragm until the heart can be visualized. Then make an identical diagonal cut along the mouse's left side, making sure to prevent any contact with the heart. Use a hemostat to keep the thoracic musculature away from the cavity.
      NOTE: At this point, the mouse will not be able to breathe any longer, so work quickly to prevent death prior to the next steps.
8. Secure the beating heart with blunt forceps and insert a dispensing needle into the left ventricle through the base of the aortic arch.
9. Clamp the needle base to the left ventricle just above the insertion site.
10. Make an incision in the right atrium, and at the first sign of blood flow, begin perfusing the animal with 20 mL of 1x PBS at 8.11 mL/minute.
11. After the 1x PBS has perfused the body, switch to the appropriate fixative for the downstream application and perfuse the mouse with 20 mL of fixative.
    1. For frozen sectioning and immunostaining, perfuse animal with the glyoxal fixative for brain clearing, perfuse animal with 4% PFA in 1x PBS (pH 7.4).
       NOTE: Signs of a successful perfusion include animal stiffness (slight stiffness with glyoxal, intense stiffness with PFA) and a lack of visible blood vessels throughout the tissues.
12. Decapitate the mouse and remove the brain by inserting scissors into the brainstem and cutting along the squamosal suture through to the frontomaxillary suture on each side. Then cut across the fontonasal suture to remove the skull cap, being careful not to damage the olfactory bulb. From there, turn the skull upside down, and use curved forceps to remove the brain, making sure to sever the optic nerve.
13. Place the perfused brain in a tissue cartridge and immerse in the fixative used to perfuse the animal overnight at 4 °C.

4. Brain treatment, slicing, and immunostaining

1. Brain treatment and slicing
   1. Remove brain cartridges from the glyoxal fixative and incubate in 15% sucrose in 1x PBS for 2 days at 4 °C or until brains sink.
   2. Remove brain cartridges from 15% sucrose and incubate in 30% sucrose in 1x PBS for 2 days at 4 °C or until brains sink.
      NOTE: See Table 1 for more information.
   3. Remove brains from 30% sucrose in 1x PBS and separate brain into various 'blocks' using a coronal brain block or sagittal brain block.
   4. Embed coronal or sagittal brain sections in OCT compound by placing brain sections on a mixture of dry ice and ethanol (EtOH). After OCT has turned white and become solid, remove from dry ice-EtOH mixture and store wrapped in parafilm, inside an air-tight bag at -20 °C.
   5. Remove frozen blocks from the freezer and place inside a cryostat with an internal temperature set to -20 °C. Use OCT to attach the block to a metal chuck so that the tissue is solidly attached. Allow ~5 min for the tissue to freeze to the chuck. Slice the tissue at 7 µm, placing each tissue slice on a charged glass slide.
   6. Allow for slides to bake overnight at 37 °C, and then place at -20 °C until used for immunostaining.
2. Immunostaining
   1. Warm slides to room temperature (RT) and post-fix with ice-cold acetone for 15 min.
   2. Wash slides for 5 min in 1x rinse buffer (e.g., IHC Select TBS Rinse Buffer), shaking at 60 rpm at RT between each step.
   3. Permeabilize for staining of nuclear antigens with 0.3% Triton X-100 for 10 min at RT. Permeabilize for staining of cytoplasmic antigens with 0.3% Tween-20 for 10 min at RT.
   4. Perform antigen retrieval by microwaving slides in 50-100 mL of 1x DAKO Antigen Retrieval Solution on low for 10 min, twice. Then rinse with rinse buffer for 5 min at RT.
   5. Incubate slides in 0.3% Typogen Black in 70% EtOH for 20 min at RT and then wash with rinse buffer.
   6. Draw a hydrophobic barrier around each tissue without allowing tissues to dry. Then block for 20 minutes at 37 °C with 1 drop of Millipore Blocking Reagent per tissue. Then add 100 μL primary antibody diluted in antibody diluent to each tissue and incubate overnight at 4 °C.
   7. The next day, incubate sections in Alexa Fluor secondary antibodies for 10 min at RT. Then wash slides in rinse buffer.
   8. Cover brain sections in 100 μL of 1:500 anti-GFP antibody conjugated to AlexaFluor 488  to boost the endogenous GFP signal marking traced cells and incubate overnight at 4 °C.
   9. The next day, wash with rinse buffer 2x, and then wash in running DI water for 5 min.
      NOTE: Be sure to gently wash with DI water, taking care to avoid any running water on slides themselves that could remove brain sections from slides.
   10. Counterstain with 100 ng/mL 4′,6-diamidino-2-phenylindole (DAPI) for 5 min. Then wash in running DI water for 5 min.
   11. Add a drop of mounting medium on top of each tissue section and seal with a 22 mm x 22 mm coverslip. Imaging is recommended on the day of mounting to ensure optimal signal.

5. iDISCO brain clearing (adapted from³⁹)

1. Fixation and sectioning
   1. Post-fix brains in 4% PFA overnight at 4 °C and then again for 1 h at RT.
   2. Wash brains in 1x PBS for an hour, twice, at RT. Then cut into 1 mm thick sections using the necessary brain block and place into 5 mL microcentrifuge tubes containing 1x PBS.
2. Methanol pre-treatment (with methanol)
   NOTE: Due to the possibility of methanol impacting immunostaining of certain proteins, the authors would like to point the reader to Renier et al. 2014 for a protocol to a methanol free iDISCO protocol alternative³⁹. See Table 1 for more information regarding the following solutions in this section.
   1. Wash with 1x PBS for 1 h twice (shaking) at RT.
   2. Wash in 50% methanol (in PBS) for 1 h (shaking) at RT.
   3. Wash in 80% methanol for 1 h (shaking) at RT.
   4. Wash in 100% methanol for 1 h twice (shaking) at RT.
   5. Bleach samples with 5% hydrogen peroxide (H₂O₂) in 20% dimethyl sulfoxide (DMSO)/methanol (1 volume 30% H₂O₂/1 volume DMSO/4 volume methanol, ice cold) at 4 °C overnight (shaking).
   6. After bleaching, wash samples in methanol for 1 h twice (shaking) at RT.
   7. Wash in 20% DMSO/methanol for 1 h twice (shaking) at RT.
   8. Wash in 80% methanol for 1 h (shaking) at RT.
   9. Wash in 50% methanol for 1 h (shaking) at RT.
   10. Wash in PBS for 1 h twice (shaking) at RT.
   11. Wash in PBS/0.2% Triton X-100 for 1 h twice (shaking) at RT.
3. Immunostaining of clearing brains
   1. Incubate samples in 1x PBS/0.2% Triton X-100/20% DMSO/0.3 M glycine at 37 °C overnight on an orbital shaker.
   2. Block in 1x PBS/0.2% Triton X-100/10% DMSO/6% goat serum at 37 °C for 3 days on an orbital shaker.
   3. Wash samples in 1x PBS/0.2% Tween-20/10 µg/mL heparin sodium salt from porcine mucosa for 1 h twice at 37 °C, and then incubate in 1x PBS/0.2% Tween-20/10 µg/mL heparin/5% DMSO/3% goat serum containing 1:500 concentration of rabbit anti-GFP AlexaFluor 488 at 37 °C on an orbital shaker for 2 days.
   4. Wash samples in 1x PBS/0.2% Tween-20 with 10 µg/mL heparin for 1 h at 37 °C on orbital shaker, three times, and then once a day for 2 days.
   5. Incubate samples overnight in 10 mL of 50% v/v Tetrahydrofuran (THF)/H₂O in a 15 mL conical tube.
   6. Incubate samples for 1 h in 10 mL of 80% THF/H₂O.
   7. Incubate samples twice for 1 h in 100% THF.
   8. Dry samples with a sterile wipe and incubate in dichloromethane until they sink to the bottom of the vial (approximately 40 min). Do not incubate >60 min.
      NOTE: Be sure to handle dichloromethane under a fume hood.
   9. Incubate samples in 18 mL of dibenzyl ether (DBE) until clear (>2 h).
   10. Store samples in DBE at RT until ready to image.
       ​NOTE: For best results, it is important to image samples as soon as possible after clearing is complete.

6. Imaging of stained slides or cleared brains

1. To image immunostained brain sections, analyze slides via confocal microscopy, where co-expression of multiple antibodies can be confirmed. We use a Leica Stellaris 5 microscope with HyD S hybrid detectors, but any point scanning confocal microscope will work. The objectives include HC PL APO 10x/0.40 CS2, HC PL APO 40x/1.30 OIL CS2, and HC PL APO 63x/1.40.
   1. Configure the correct laser lines and emission filters. The required lasers and corresponding laser intensities are subject to the microscope and fluorophore combinations being used.
      NOTE: We utilize a combination of diode 405 nm and white light lasers to fine tune excitation and emission spectra for each fluorophore, or groups of fluorophores, being used to reduce and eliminate crosstalk. Similar outcomes can be achieved using traditional laser/filter combinations as well but pay close attention to channel bleed through. Bleed through can be identified by turning on a single laser line in conjunction with each of the desired emission filter combinations to see what channels are impacted by each specific laser. For example, if the diode 405 nm laser is producing visible fluorescence in the GFP emission filter, there is bleed through occurring. Ensure that multiple channels be imaged sequentially frame by frame to greatly reduce any bleed through from occurring.
   2. For co-stained Tet-On brains with an mTmG reporter, select DAPI (ex. 405 nm, em. 450 nm), GFP (ex. 488 nm, em. 509 nm), tdTomato (ex. 555 nm, em. 582 nm), and the corresponding far-red fluorophore to match the fluorophore used on the secondary antibody. We recommend Alexa Fluor 647 (ex. 651 nm, em. 667 nm).
   3. Establish settings first with a Cre- control brain stained with both GFP and the fluorescent secondary. These brain slices will control for background fluorescence from the fluorescent antibodies and will show no real GFP signal.
   4. Analyze each brain section from top to bottom, left to right, to verify that no signal has been missed in the analysis. If imaging the same brain section at various magnifications, it is recommended that the lower magnifications are utilized first, as the higher magnification will more quickly bleach the mTomato signal.
2. To image cleared brains, use an inverted confocal microscope with an automated stage and automated tiling function. We use the Leica Stellaris 5 microscope. Because the samples are so thick (>500 µm), larger working distances are required which limits the magnification that can be effectively achieved. We use the HC PL APO 10x/0.40 CS2 objective for all cleared brain imaging.
   1. Start by laying a cleared brain section flat against a glass bottom dish and submerge in dibenzyl ether.
      NOTE: Dibenzyl ether is corrosive to most objective lenses and most plastics. For this reason, any interior plastic in the glass bottom dish needs to be coated with quick drying silicone elastomer.
   2. Configure the correct laser lines and emission filters. For co-stained Tet-On brains with an mTmG reporter, select DAPI, GFP, tdTomato, and the corresponding far-red fluorophore to match the fluorophore used on the secondary antibody. Set the desired resolution, we recommend at least 1024 x 1024. Set pinhole to 1 AU.
   3. Establish the laser intensity and detector gain first with a Cre- control brain stained with both GFP and the fluorescent secondary. These brain slices will control for background fluorescence from the fluorescent antibodies and will show no real GFP signal.
   4. Once laser intensities and detector gain have been optimized, map out the brain for imaging. Start by locating the entire perimeter of the brain section to set the X and Y axis of the image acquisition. Next, identify the Z axis by setting the focal point to approximately the tissue's center depth, and use Z-positioning controls to locate the "highest" point of the tissue (most positive Z-value). Scan various regions of the tissue increasing the maximum Z-axis height as necessary.
      1. Repeat for the "lowest" (most negative Z-value) point required to obtain the entire thickness of the tissue. The Stellaris 5 has a limit of ± 250 um of the focal point. This limits optical sections to 500 µm thick in the Z-axis. The 3D area of the brain section is now known.
   5. Set Z-step size to 6 µm and start acquiring the image. Acquisition time is dependent on several factors and can range from hours to days depending on the desired parameters. To decrease image acquisition time try decreasing resolutions, increasing laser scan speed, or increasing step size.
   6. Once the tiled image of the entire cleared brain section has been captured, analyze as desired.