Confocal Imaging of Mitochondrial Movement within Oligodendrocytes in Organotypic Brain Slice Cultures

Abstract

Source: Kennedy, L. H., et. al. Visualization and Live Imaging of Oligodendrocyte Organelles in Organotypic Brain Slices Using Adeno-associated Virus and Confocal Microscopy. J. Vis. Exp. (2017)

This video demonstrates the imaging of oligodendrocyte mitochondria in organotypic mouse brain slices. The slice is placed inside a bath chamber with continuous fluid circulation and controlled temperature. After setting up various parameters of the confocal microscope, time-lapse images of mitochondria with red fluorescence are captured, followed by Z-stack images of the oligodendrocyte with green fluorescence.

Protocol

1. Preparation of Organotypic Slices

NOTE: This recipe uses two mouse pups at postnatal day 7-9 (p7-9), yielding 24 organotypic slices divided into two six-well culture dishes. Unless stated otherwise, all procedures should be done in a sterile hood, and nitrile or latex gloves should be used. Only cell culture-grade ingredients should be used.

1. Autoclave bottles, dissection tools (1 large scissor, 1 small scissor, 1 large flat spatula, 1 small rounded and sharpened spatula, and 1 forceps; see Materials list for details), glass pipettes, pipette tips, and tissue wipes were used for the slice preparation.
2. As half of the glass pipettes should be used with the large opening, break off the narrow end.
   1. Score with a diamond scriber pen (if available) around the pipette just below the point where the glass starts narrowing. Then, place the pointy end of the pipette into a nitrile glove or similar. Carefully break off the tip by bending the pipette while pushing it against a workbench.
   2. Then, blunt the broken end by rubbing it on a piece of sandpaper or melt it slightly on a Bunsen burner. The broken pipettes are used to transfer cut brain slices, with the broken end turned into the rubber bulb and the large open end touching the solution/slices.
      NOTE: This is not done in a sterile hood and can be done several days in advance.
3. Prepare culture dishes.
   NOTE: This can be done the day before slicing or at least 2 hours before slicing.
   1. Sterilize Polytetrafluoroethylene (PTFE) membranes ("confetti"). Use two sterile forceps to remove a single confetti from the surrounding blue plastic pieces. Then, sterilize by dipping twice into a Petri dish containing 96% ethanol (EtOH). Then, lay the confetti flat in a sterile large Petri dish to dry.
      NOTE: The confetti are hydrophilic PTFE membranes similar to the membranes in the culture inserts. The use of confetti aids live imaging because single slices can be easily transferred from the insert to the microscope setup by lifting the confetti with forceps (see 4.8. for details). Alternatively, the brain slices can be cultured without the confetti (in which the slices will attach directly to the membrane of the culture insert). In this case, the membrane insert must be cut with a scalpel to transfer the slice to the microscope bath.
   2. Add 1 mL of culture medium to each well of the six-well culture dishes.
   3. Place one culture insert in each well using sterile forceps. Avoid air bubbles between the insert and the culture medium.
      NOTE: To save money and the environment, it is possible to reuse the culture inserts. This can be done by thorough rinsing in distilled water (dH₂O) followed by incubation in 70% EtOH for a minimum of 24 hours until reuse. When preparing for a new culture, dry the inserts in a cell culture plate under ultraviolet (UV) light for 15-30 min.
   4. Place two (sterilized and dry) confetti on each of the culture inserts. Place the confetti towards the edges of the inserts to get minimal overlap.
   5. Put the plates in the cell culture incubator at 37 °C with 5% CO₂.
4. Bubble cold dissection medium with bioxide gas (95% O₂/5%CO₂). To keep the solution sterile, connect the tube from the gas cylinder to a sterile syringe filter (0.22 µm pore size).
   1. Couple the other end of the syringe filter to a sterile tube connected to a sterile glass pipette. Place the glass pipette in the solution, cover it with parafilm, and turn on the gas. Keep the dissection medium on ice.
5. Prepare the dissection/slicing area.
   NOTE: Ideally, this procedure should be done in a sterile hood. If this is not possible, the vibratome can be left on a workbench. In this case, it is recommended to use a hair net and face mask.
   1. Using a single-edge razor blade, cut out a rectangular piece (approximately 2 cm x 0.5 cm) of agarose gel and glue it onto the vibratome stage so that the long side faces the vibratome blade.
   2. Clean all surfaces and the vibratome parts with 70% EtOH and wipe with autoclaved tissue wipes.
      NOTE: It is important that all the pieces that will be in contact with the brain tissue are completely dry since EtOH can damage the cells.
   3. Attach a sterile razor blade to the vibratome and perform a vibration check to see if the vibratome has this function.
   4. Place autoclaved dissection tools, autoclaved tissue wipes, four small Petri dishes, a round-edge scalpel, two glass pipettes with rubber bulbs, two open-end glass pipettes (see pt. 1.1), and super glue (sprayed with EtOH) in the hood.
   5. Pour the bubbled dissection medium into the vibratome boat and into the four small Petri dishes.
      NOTE: This step should only be done immediately before dissection since the medium from this point on is not bubbled or kept on ice.
6. Dissect and mount brains.
   NOTE: To obtain healthy slices, this step must be done quickly and precisely. Some practice is needed.
   1. Euthanize animal #1 by dislocation of the neck or according to local guidelines and then decapitate with the large scissors.
   2. Using small, sharp scissors, quickly remove the skin by making a sagittal incision from the neck to the nose and pull it aside to reveal the skull.
      1. Cut along the sagittal suture, followed by lateral incisions over the interior part of the frontal bones and the lamboid suture (the border between cerebrum and cerebellum).
      2. Use forceps to bend open the skull on each side. The cranial nerves are cut off from the brain by inserting a rounded, sharp spatula under the ventral side of the brain and gently moving the spatula laterally.
   3. Use a single-edge razor blade to make a coronal cut rostral to the cerebellum.
      NOTE: This cut will determine the angle at which the slices are cut. It should, therefore, be made at a straight angle.
   4. Flip the brain out of the skull and into a small Petri dish containing a dissection medium.
   5. Repeat steps 1.6.1. - 1.6.4. for animal #2 and place this brain in a second Petri dish.
      NOTE: The animals are not sterile. Perform the dissection on one side of the hood and then move to the other, clean side once the brains have been removed from the skull.
   6. Change gloves.
   7. Attach brain to vibratome stage: add a thin layer of super glue to the stage just in front of the mounted agarose gel. Hold brain #1 with a large flat spatula with the freshly cut (caudal) side touching the spatula and the ventral side facing (and being as close as possible to) the agarose gel.
      1. Then, gently place the brain onto the glue by pushing it off the spatula with a set of forceps. Make sure to leave enough space for brain #2.
         NOTE: It is important that the brains are well attached to the stage but without excessive glue, as this can obstruct the cutting.
   8. Immediately after mounting brain #1, use the large spatula to add some drops of dissection medium on top of the brain. This will keep the brain moist, harden the glue, and prevent it from sticking to the sides of the brain.
   9. Repeat steps 1.6.7 and 1.6.8. for brain #2.
   10. Attach the stage onto the vibratome boat.
7. Cut 230 µm thick coronal slices with an amplitude of 1 - 1.5 mm, a frequency of 85 Hz, and a velocity of 0.2 mm/s. Collect slices approximately between 1.4 mm rostral and 2.1 mm caudal to Bregma.
   1. Collect slices after each slice has been cut by using open-end glass pipettes (pt. 1.1). Transfer slices to a Petri dish containing a dissection medium. Use a rounded scalpel to cut the slices into two pieces, dividing the two hemispheres.
8. When all the slices are cut, place slices into the culture dishes.
   1. Take the culture dish (one at a time) out of the incubator.
   2. Using the open-end glass pipettes, carefully transfer one slice to each of the confetti.
      NOTE: The slices should lay flat in the middle of the confetti. This requires some skill. However, if some slices are not perfect, it is better to leave them than to spend time trying to move them, as it is important to get all the slices into the incubator as fast as possible.
   3. Use a glass pipette (pointy end) to gently remove excess medium without touching the slice.
      NOTE: Slices should not be immersed in liquid during the incubation. Thus, the excess medium must be aspirated such that the slices are exposed to the air (a thin film of medium will still cover the slices). Slices that remain immersed in the liquid will usually be unhealthy or die (4 and our observations).
   4. Move the dish back into the incubator once all the confetti has one slice.
   5. Repeat steps 1.7.8. - 1.8.4. for dish #2.
9. Change the culture medium.
   NOTE: This should be done the day after slicing (day 1 in vitro, DIV1) and then every 2-3 days.
   1. Preheat the culture medium in a water bath or the incubator.
   2. In a sterile hood, use vacuum suction or a regular pipette with a sterile tip to aspirate culture medium from each well. This is done by gently tipping the dish (by about 30 degrees) and placing the pipette tip at the edge of the culture insert.
   3. Remove approximately 800 µL from each well (leaving just enough medium to keep the bottom of the insert covered in medium). Then, using a clean pipette, add 800 µL of the pre-heated medium to each well. Then, place the dishes back in the cell culture incubator.

2. Viral Transduction

1. Purchase or make Adeno-Associated Viruses (AAVs) with a plasmid of interest.
   NOTE: This protocol describes the use of AAV serotype 2/8 (AAV 2/8) carrying mitochondrial targeted dsred or GFP as a reporter gene under the transcriptional control of the myelin basic protein promoter12 (AAV2/8 MBP_mito_dsred or AAV2/8 MBP_mito_GFP) to visualize oligodendrocyte mitochondria. AAV 2/8 carrying GFP or tdtomato as reporter gene driven by the general CAG promoter (AAV 2/8 CAG_GFP or AAV 2/8 CAG_tdtomato) is used to visualize oligodendrocyte cytoplasm. Thus, the combination of AAV2/8 MBP_mito_dsred with AAV 2/8 CAG_GFP gives red mitochondria and green cytoplasm in oligodendrocytes, whereas combining AAV2/8 MBP_mito_GFP and AAV 2/8 CAG_tdtomato gives green mitochondria and red cytoplasm. The constructs are described in more detail elsewhere13.
2. On day 7 in vitro (DIV7), oligodendrocytes with the AAVs are transduced.
   CAUTION: Follow safety regulations when working with AAVs. Make sure to have the proper training and the Biosafety Level approval required. All the following points should be carried out in a Class II biosafety cabinet using gloves and lab coat. Here, application of AAV to the cortex area of the slices is described, as this is the area that shows the best recovery.
   1. Dilute the AAV in a pre-heated (37 °C) culture medium. Dilutions around 2 × 10⁹ genome copies/mL (GC/mL) are recommended to get a sparse transduction of oligodendrocytes which enables imaging of single cells within the slice. However, a pilot using different titers should be done to ensure a density of transduced oligodendrocytes that is suitable for the specific experiments. If two different AAVs are used, they should be mixed in the same solution and added to the slice simultaneously.
   2. Add around 1.3 µL diluted AAV to each slice. Make sure the outside of the pipette tip is dry. Pipette 1.3 µL diluted AAV and hold the pipette above the cortex as close as possible without touching the slice with the pipette tip (about 1 mm away).
   3. Then slowly push solution out of the pipette. When the solution touches the slice, move the pipette tip over the cortex to spread the solution over the whole cortex area.
      NOTE: This procedure should be done as fast as possible to avoid keeping the slices out of the hood for longer than necessary. Do this on one dish at the time and put the first dish back in the incubator before starting on dish #2. To get a satisfactory distribution of AAV without damaging the tissue requires some practice and a steady hand.
3. If an upright fluorescence microscope is available in the cell lab, protein expression can be checked during the time in culture by placing the dish under the microscope.
   NOTE: The dish should not be kept out of the incubator for more than 2 - 5 minutes.

3. Time-lapse Imaging

NOTE: The imaging can be performed whenever the expression levels of the fluorescent markers are sufficient, and the slices look healthy (usually DIV11-14).

1. Ensure that the microscope, perfusion, and heating are set up correctly so that the bubbled imaging solution is heated and can enter and exit the bath.
   NOTE: Various solutions exist for bath chambers. A simple version is presented here.
   1. Make in- and outlets of the bath by blunted syringe needles (bent at ~45°) that are attached to the sides of the bath by blu-tack/fun tack.
   2. Ensure that tubing goes from a bottle of continuously bubbled imaging solution via the peristaltic pump, through the heating tube of the heater unit, and then to the inlet syringe needle in the bath.
   3. Connect another tube to the syringe needle outlet. This tube then goes via the peristaltic pump and back to the bottle of imaging solution (or to a waste bottle).
2. Bubble imaging solution with bioxide gas.
   NOTE: This should be started at least 15 minutes before imaging and continue throughout the experiment.
3. Turn on peristaltic pump and heater and run solution through the bath to obtain optimal bath temperature (37 ± 0.5 °C). Ensure that the thermometer sensor connected to the temperature unit is submerged in the bath solution.
4. Turn on the microscope, fluorescence lamp, and appropriate lasers.
5. Set up a suitable light path for the sample.
   NOTE: This will vary depending onthe microscope setup and probe. General knowledge of how to use the confocal microscope is required and will not be dealt with here.
6. Use a water immersion objective with the appropriate magnification and numerical aperture (n.a.). We use a 40x water immersion plan-apochromat objective (n.a. 1.0).
7. Open pinhole to achieve an optical section of approximately 2 - 2.5 µm for the time-lapse video. This reduces the occurrence of mitochondria moving out of focus during time-lapse.
8. Transfer organotypic slice to the bath:
   1. Add a drop of imaging solution or culture medium to a small plastic Petri dish.
   2. In a sterile hood, use sterile forceps to transfer one organotypic slice from the membrane insert to the drop. The forceps should only touch the confetti and not the slice.
   3. Put the lid on the Petri dish.
   4. Immediately put the culture dish containing the rest of the slices back in the incubator.
   5. Bring a Petri dish containing the slice to the microscope.
   6. Turn off peristaltic pump while transferring slice to microscope bath. First, use forceps to lift confetti with slice from Petri dish to the top of the bath (the confetti will float). Then place the two tips of the forceps on each side of the confetti so that they touch the confetti without touching the slice, and push the confetti through the solution to the bottom of the bath. Center the confetti in the middle of the bath.
   7. Use forceps to place the hold-down anchor (harp) on top of the confetti without touching the slice.
      NOTE: A harp is a horseshoe-shaped platinum anchor that is used to prevent the slice from floating or drifting in the bath. For acute slices, thin strings run from one side of the harp to the other (resembling a music harp). For organotypic slices, the harp should be stringless and fit the size of the confetti in such a way that it can lay on top of the confetti without touching the slice.
   8. Turn the peristaltic pump back on.
9. Lower the objective down to the sample.
10. Select cell and region to image.
    NOTE: Avoid excessive exposure of the cells to laser light as this can cause cell toxicity (see tips in the points below).
    1. Use the eyepieces and the fluorescent lamp to find a healthy-looking oligodendrocyte with the correct expression. Typically, oligodendrocytes that have a strong overexpression of fluorescent protein appear unhealthy. Unhealthy oligodendrocytes will have processes with a blobby appearance, and the mitochondria will appear fragmented. In healthy oligodendrocytes, the soma and processes appear smooth, and mitochondria have various lengths (although they are typically much shorter than in neurons and astrocytes).
    2. Place the cell of interest in the middle of the field of view.
    3. Use the "live" function in the software (for Leica and Zeiss microscopes) to identify the cell on the screen and choose an area of interest, e.g., a part of the cell that contains several primary processes or myelin sheaths or a single process or myelin sheath.
       NOTE: To be able to image as much of the processes/sheaths as possible, choose areas containing processes that lay relatively flat in the x-y direction.
    4. Zoom in on the field of interest (typically producing an image with a pixel size of 0.14 - 0.30 µm).
    5. Using the "regions" tool, draw a rectangle around the area and choose the option to scan only the selected region. Scanning time, and thus laser exposure, is reduced by only scanning the selected region.
       NOTE: Horizontal rectangular regions will be scanned faster than vertical regions due to the shorter number of scanned lines needed. If a vertical rectangle region is scanned, the scanning time can be reduced by rotating the scan field by 90 degrees (using the rotation tool).
11. Adjust laser power and gain to get a good view of mitochondria.
    NOTE: Use the minimal laser power needed. If possible, turn up gain rather than increasing the laser power. In addition to causing cell toxicity, too high laser power will bleach the imaged objects over time, thus requiring a further increase of laser power or gain during scanning. The required laser power and gain will depend on the expression level and the microscope. With an LSM700 confocal microscope, we use a 555 nm laser at a power of 20-40 µW to image dsred or tdtomato, and a 488 nm laser is used at 20 - 30 µW to image GFP (laser power measured at the back aperture of the objective). Gain is set high for live imaging, usually in the range of 700 - 800 for GFP and 850-980 for dsred and tdtomato. The high gain may cause some more background noise, which is removed by increasing the digital offset (offset values typically lay between 0 and 15). In our experience, using MBP_mito_GFP gives a better signal-to-noise than MBP_mito_dsred.
12. Select scan speed.
    NOTE: Minimize scanning time by using a fast scanning speed and by drawing a small scanning region (see point 4.10.5 below). It is also possible to save scanning time by performing a bidirectional scan. However, this is not recommended due to poorer image quality.
13. Set frequency and duration of time-lapse recording, e.g., capture images every 2 seconds for a total of 20 minutes for mitochondrial imaging in oligodendrocytes.
    NOTE: The frequency and duration should be set according to the mobility of the objects of interest. A higher frequency will expose the specimen to more laser light, but faster-moving objects also require a shorter total duration of time-lapse videos (e.g. mitochondria in neurons, which move more frequently and at a much higher speed than mitochondria in oligodendrocytes, are typically imaged every second for a total of 2 minutes16).
14. Start time-lapse recording.
    NOTE: The mitochondria may move out of focus and/or drift out of the imaged region during the recording. To minimize vibrations and movement, adjust in- and outlets of the bath and reduce pump speed if needed. Small changes in focus usually still occur. Thus, continuous monitoring of the screen during imaging is required, with small adjustments of the focus during the recording.

Capture a z-stack of the whole cell for future identification of the cell and the imaged process.
NOTE: For better resolution, the pinhole should be reset to 1 airy unit for the z-stack.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Agarose	Sigma	A9539
BD Microlance 19G	BD	301500	Needles used for in- and outlet of bath
Bioxide gas	AGA	105701
Brand pipette bulbs	Sigma-Aldrich	Z615927	Pipette bulbs
Bunsen burner (Liquid propane burner)	VWR	89038-530
Cable assembly for heater controllers	Warner Instruments	64-0106	Temperature controller - thermometer part
Calcium chloride	Fluka	21100
Carbon dioxide	AGA	100309	CO2 for incubator
Cover glass, square Corning	Thermo Fischer Scientific	13206778	To attach under bath for live imaging. Seal with glue or petrolium jelly.
D-(+)-Glucose	Sigma	G7021
Delicate forceps	Finescience	11063-07	For dissection
Diamond scriber pen	Ted Pella Inc.	54463
Disposable Glass Pasteur Pipettes 230 mm	VWR	612-1702	Glass pipettes
Double edge stainless steel razor blade	Electron Microscopy Sciences	#7200	Razor blade for vibratome
Earle's Balanced Salt Solution (EBSS)	Gibco-Invitrogen	24010-043
Filter paper circles	Schleicher & Schuell	3,00,220	Filter paper used for filtration of PFA
Fun tack	Loctite	1270884	Use to connect/adjust position of in- and outlets in bath
Hand towel C-Fold 2	Katrin	344388
Harp, Flat for RC-41 Chamber,	Warner Instruments	64-1418	Harp to hold down confetti in bath. Cut off strings before use with organotypic slices. 1.5 mm, 13mm, SHD-41/15
HEPES, FW: 260.3	Sigma	H-7006
Holten LaminAir, Model 1.2	Heto-Holten	96004000M	Laminar flow hood
Horse serum, heat inactivated	Gibco-Invitrogen	26050-088
KCl	Sigma	P9541
LCR Membrane, PTFE,	Millipore	FHLC0130	Confetti
Leica VT1200	Leica	14048142065	Vibratome
MEM-Glutamax with HEPES	Thermo Fischer Scientific	42360024
MgCl2	R.P. Normapur	25 108.295
Micro Spoon Heyman Type B	Electron Microscopy Sciences	62411-B	Small, rounded spatula with sharpened end for dissection
Millex-GP filter unit	Millipore	SLGPM33RA	Syringe filter unit
Millicell cell culture insert, 30 mm	Millipore	PICM03050	Cell culture inserts
Minipuls 3 Speed Control Module	GILSON	F155001	Peristaltic pump for live imaging - Control module part (connect to two-cannel head)
Na2HPO4	Sigma-Aldrich	S7907
NaCl	Sigma-Aldrich	S9888
NaH2PO4	Sigma-Aldrich	S8282
NaHCO3	Fluka	71628
Nunclon Delta Surface	Thermo Fischer Scientific	140675	Culture plate
Nystatin Suspension	Sigma-Aldrich	N1638
Objective W "Plan-Apochromat" 40x/1.0 DIC	Zeiss	441452-9900-000	Water immersion objective used for live imaging. (WD=2.5mm), VIS-IR
Parafilm	VWR	291-1211
Paraformaldehyde, granular	Electron Microscopy Sciences	#19208
PC-R perfusion chamber	SiSkiYou	15280000E	Bath for live imaging
Penicillin-Streptomycin, liquid	Invitrogen	15070-063
Petri dish 140 mm	Heger	1075	Large Petri dish
Petri dish 92x16 mm	Sarstedt	82.1473	Medium Petri dish
Petridish 55x14,2 mm	VWR	391-0868	Small Petri dish
Phosphate buffered saline (PBS)	Sigma	P4417	PBS tablets
R2 Two Channel Head	GILSON	F117800	Peristaltic pump for live imaging - Two channel head part (requires control module)
Round/Flat Spatulas, Stainless Steel	VWR	82027-528	Large spatula for dissection
Sand paper	VWR	MMMA63119	Optional, for smoothing broken glass pipettes
Scissors, 17,5 cm	Finescience	14130-17	Large scissors for dissection
Scissors, 8,5	Finescience	14084-08	Small, sharp scissors for dissection
Single edge, gem blade	Electron Microscopy Sciences	#71972	Single edge razor blade
Single inline solution heater SH-27B	Warner Instruments	64-0102	Temperature controller - heater part
Steritop-GP Filter unit, 500 ml , 45mm	Millipore	SCGPT05RE	Filter to sterilize solutions
Super glue precision	Loctite	1577386
Surgical scalpel blade no. 22	Swann Morton Ltd.	209	Rounded scalpel blade
Temperature controller TC324B	Warner Instruments	64-0100	Temperature controller for live imaging (requires solution heater and cable assembly)
Trizma base	Sigma	T1503
Trizma HCl	Sigma	T3253
Water jacketed incubator series II	Forma Scientific	78653-2882	Incubator