Long-Lived Organotypic Slice Culture Model Of Rat Basolateral Amygdala.

Abstract

Organotypic slice cultures (OTCs) have been employed in the laboratory since the early 1980s and proved to be useful for the study of a number of neural systems (Gähwiler, 1981, Gähwiler et al., 1997). Our recent work focuses on the development of a behavioral stress resilience induced by repeated daily injections of Neuropeptide Y (NPY) into the basolateral amygdala (BLA). Resilience develops over weeks, persisting to 8 weeks. To unravel the cellular mechanisms underlying NPY-induced stress resilience we developed in vitro OTC cultures of BLA. Herein, we provide an optimized protocol that consistently yields viable and healthy OTCs containing the BLA and surrounding tissue using the “interface method” prepared with slices taken from postnatal (P) day 14 rats. We explain key points to optimizing tissue viability, discuss mitigating or avoiding pitfalls that can arise, to aid in the successful implementation of this technique. We show that principal neurons (PNs) in BLA OTCs (8 weeks in vitro = EP [equivalent postnatal] 70), develop into networks that are electrophysiologically very similar to those from acute slices obtained from older rats (P70), and respond to pharmacological treatments in a comparable way. Furthermore, we highlight how these cultures be used to further understand the molecular, cellular and circuit-level neuropathophysiological changes underlying stress disorders. BLA OTCs provide long-term physiological and pharmacological results whose predictions were borne out in vivo, supporting the validity of BLA OTC as a model to unravel BLA neurocircuitry. Recent preliminary results also support the successful application of this approach to preparing long-lived OTCs of BLA and neocortex from mice.

INTRODUCTION

A stressor initiates a complex response that promotes an organism’s survival. This stress response requires the concerted effort of many brain areas to elicit the appropriate defensive actions. However, when an individual is exposed to repeated or severe stress, this normally adaptive response can result in pathology, such as posttraumatic stress disorder. The basolateral amygdala (BLA), located deep in the medial temporal lobes (Krettek and Price, 1978), is the chief regulator of emotional responses to stress, integrating sensory input and experience to assess and choreograph behavioral output through an intricate network of downstream projections (LeDoux, 1992).

To study the complex neurocircuitry underlying the development of chronic stress-related pathology, in vitro experiments performed on acute ex vivo slices from rodent brains (e.g, Rainnie, 1999, Ehrlich et al., 2012, Ryan et al., 2014) or in vivo preparations are used. However, when examining the mechanisms that underlie the acquisition or mitigation of stress-related pathology, both acute ex vivo brain slices (Silveira Villarroel et al., 2018, Humpel, 2015a), and in vivo recordings have limitations. To study molecular and cellular mechanisms governing these emotional responses, a reduced model system is needed that best recapitulates the long-term plasticity responses of neural circuitry experiencing prolonged or repeated stress.

Organotypic slice culture (OTC) preparations have been used as an alternative to the classical in vitro preparations for many years (Gähwiler et al., 1997). “Organotypic” thin tissue explant cultures resemble their in vivo counterparts in organizational structure and cellular phenotypes and thus represent an intermediate model between dispersed cell cultures and in vivo models (Gähwiler, 1981a,b; Gähwiler et al., 1997; De Simoni et al., 2003; Humpel 2015a,b; Stoppini, 1991). As OTCs maintain much of the structural, cytoarchitectural and synaptic organization of the original tissue, they also conserve much of their network activities, permitting flexible, controlled manipulations of specific neural circuits (Pena, 2010). OTCs can be successfully cultured for prolonged periods, even months, allowing examination of the long-term effects of acute and sustained treatments. OTCs have been employed to study mechanisms underlying widely different research question s lncluding network development and plasticity, alterations in neuronal networks through gene transfection, synchronous neuronal network activity and morphological changes due to neurodegeneration, and have served as neural models to study Alzheimer’s and Parkinson’s diseases (Gahwiler et al., 1997, Pena, 2010, Humpel, 2015a). In addition, their use in preclinical drug screening can provide proof of principal for new drug candidates on models of the target brain system, and facilitate drug discovery and safety testing (Sundstrom et al., 2005. Magelhäes et al., 2018).

OTCs can be cultured in two ways: the original “roller-tube” method (Gahwiler, 1981b) or the “interface method”, in which they are grown at the air-medium interface using semipermeable membranes, avoiding the need for rotation (Stoppini et al., 1991). Consideration of the age of the animal (tissue) is important for OTC viability, as tissue from embryonic animals typically survives better; yet postnatal (<P12) donors are often preferred due to their relative structural maturity (Humpel, 2015a). Recently, a number of groups have succeeded in culturing OTCs from adult brain tissues, however most of these studies require special conditions and are largely the exception to the rule (Lossi et al., 2009, Kim et al., 2013, Humpel, 2015b). We recently published work using BLA OTCs using the interface method, established from Sprague-Dawley rats at postnatal day 14 (P14) that can be maintained for up to 12 weeks (Michaelson et al., 2020). Here we describe in detail the methods developed to establish and validate OTCs of BLA for long-term pharmacological experiments to parallel in vivo studies. Significant developmental changes occur within the rat BLA during the first postnatal month (Ehrlich et al., 2012), so we sought long-lived OTCs from the latest possible time postnatally, such that the OTCs could reach the equivalent postnatal age of the in vivo experiments (Silveira-Villarroel et al, 2018) (OTCs prepared from rats older than P18 were not viable for more than a few days, while those prepared from P14 rats developed successfully and could be readily maintained up to equivalent postnatal (EP) P70 (i.e., 8 weeks in vitro, Fig. 1). Comparison of electrophysiological and morphological properties of BLA principal neurons from acutely prepared slices (P14 and P70 aged rats) to those obtained from OTCs suggest BLA OTCs develop and behave in a fashion similar to slices acutely prepared from P70 animals. In these OTCs, repeated treatment of OTCs with either NPY or CRF to mimic an in vivo protocol resulting in prolonged behavioral stress resilience or stress vulnerability, respectively, yielded consistent and prolonged structural changes in BLA OTCs principal neurons, which predicted identical changes in BLA principal neurons in vivo (Michaelson et al., 2020). The BLA OTC model proves to be a valuable and innovative tool to generate mechanistic insights into pathways regulating physiological and pathophysiological emotional states and promises to aid in rational drug design for the therapeutic treatment of patients with anxiety-related disorders.

Figure 1:

Timeline comparison for in vivo and OTC experiments. Upper band represents timeline for OTC preparation, while lower band represents the in vivo treatment of naïve rats. OTCs are prepared and placed in culture at P14 – P16, while rats receive cannula implants into BLA at 6 – 8 weeks postnatal (Silveira Villarroel et al. 2018). At 4 weeks in culture (equivalent postnatal 42 days = EP42), OTCs received experimental manipulation treatments (e.g. 5 x daily media changes with vehicle or with vehicle + NPY), while after acclimatization, rats received e.g. 5x daily bilateral infusions of vehicle alone or vehicle + NPY. Following 2 weeks control media changes, OTCs could be studied for changes in electrophysiological properties over the next 2 weeks. After 2-4 weeks, rats could be used to prepare acute ex vivo brain slices for electrophysiological experiments.

Strategic Planning

Animals

Two different age groups of male Sprague-Dawley rats are used for experimentation. P14 pups are housed with the dam prior to preparation of OTCs and for a subset of ex vivo electrophysiological and morphological studies. The second set of rats, aged five-weeks, are grouped housed (2-3 animals per cage) with 12:12 hours light: dark schedule and food and water are supplied ad libitum. All animal procedures were approved by the University of Alberta Animal Care and Use Committee: Health Sciences, in accordance with the guidelines of the Canadian Council on Animal Care.

Experimental setup for OTCs

Traditionally, OTCs of brain tissue are obtained from young (P0-P10) animals, using the “interface-method” (Stoppini et al., 1991) format to take advantage of their ease of preparation and high viability. To study biological mechanisms that more closely resemble those from the adult brain, we sought to prepare OTCs from rats as old as possible that consistently produced viable, cultures that thrived over an extended period (Fig. 1). However, this protocol can be adapted for shorter culturing timepoints and for producing healthy OTCs prepared from younger animals. OTCs were prepared from P14 animals, and cultured to the equivalent postnatal (EP) age of 6 weeks when culture conditions were experimentally varied, and at EP 9-10 weeks, further experiments, including electrophysiological recordings and intracellular labeling were performed.

We prepared OTCs from four P14 rats per week (completed in a single day), but this can be adjusted to meet specific research needs. Each animal produces roughly four 350 μm thick, BLA-containing slices per hemisphere, totaling eight slices per animal. The procedure can be completed in ~ 1 – 1.5 hours/animal by an experienced individual. Time is vital during the procedure, as a slow dissection will negatively affect the outcome, but meticulous care must also be taken when handling the tissue for optimal health. Note that, while we describe the slicing using our homemade slicing chamber, it can be readily adapted for use with other commercially available vibrating slicers.

Basic Protocol 1: ORGANOTYPIC SLICE CULTURE

Here we provide a technical procedure and materials required for the preparation of OTCs from P14 rats for pharmacological manipulation of biological activity, electrophysiological recordings, and immunohistochemistry. This procedure can be used and adapted to investigate other brain regions, for example, neocortex and hippocampus, and has recently been successfully adapted for use in the mouse.

CAUTION: Wear disposable gloves, N95 respirator, and a clean, calf length lab coat with cuffed sleeves during all procedures that require strict sanitation. Otherwise, follow your lab’s requirements for PPE.

Materials

70% ethanol

100% acetone

Corning Pyrex 100 mL media storage bottles

Paper towel

Nitrile gloves

Calf-length lab coat with cuffs

N95 respirator

Laminar flow biosafety cabinet with UV light

Cell culture incubator set to 37°C, 5% CO₂ (bone dry)

Glass bead sterilizer

Aquaguard-1

Water bath

Aquaguard-2

15 mL and 50 mL sterile centrifuge tubes

Complete medium (see Reagents and Solutions)

Complete slicing solution ( see Reagents and Solutions)

Anti-mitotic solution (see Reagents and Solutions)

10 mL and 25 mL sterile serological pipettes with controller

1μL, 10μL, 100μL and 1000μL single channel micropipettes with sterile tips

1.5 mL and 5 mL sterile microcentrifuge tubes

0.22μm vacuum filter

0.22μm sterile syringe filters

1 L vacuum flask

Bleach

500 mL Stericup® with Steritop® vacuum filter unit (or equivalent)

5% CO₂, 95% O₂

Vibratome or equivalent vibrating slicer

Stainless steel double-edge razor blades for tissue sectioning with vibrating microtome

Low melting point agarose

Single edge stainless steel “industrial” razor blades

60x10mm and 100x10mm disposable petri dishes

Sterile 24-well plates

Sterile Millicell Cell Culture Inserts 12mm, 0.4μm

Sterile 1mL syringes

25G 5/8” needles

Ice bucket

Ice

Small animal guillotine, sharpened

Large surgical scissors (14-cm length)

Small straight dissecting scissors (10.5cm length)

Straight fine forceps (2) (e.g. e.g. Dumont AA-style tweezers )

Micro-tapered stainless steel spatula (e.g. Fisher Handi-hold microspatula) – polished with crocus cloth)(2)

100 mm filter paper

Cyanoacrylate glue (“superglue” – conventional thin viscosity)

Fine sable hair, watercolor-type paintbrush

Dissection stereomicroscope

Peptides and drugs relevant to your experiments

Preparation (one week prior)

1. Using sturdy, sharp, non-surgical scissors, prepare thin, double-edged stainless steel razor blades by halving them (lengthwise, at the flanges joining the two sides) for the vibratome. Store in a screw cap-sealable glass bottle (e.g Corning Pyrex 100 mL media storage bottles) in 100% acetone (kept anhydrous over Molecular Sieves - Fisher Grade 514 Type 4A) to remove oil and grease from blades.

2. Prepare 3L of stock 70% ethanol and one 500 mL bottle for immediate use.

3. Decontaminate and start incubator, if necessary. Add 10mL of Aquaguard-1 to each liter of water in the water tray.

4. Decontaminate and prepare water bath by adding 10mL Aquaguard-2 for each liter of water in the bath.

5. Prepare anti-mitotic solution and pharmacological reagents and store at −20°C.

Preparation (two working days prior)

6. Prepare complete media and slicing solution and store at 4°C.

Preparation (one working day prior)

7. Test agarose incubation. Turn on water bath and dissolve 400 mg of low melting point agarose in 20 mL slicing solution (2%wt/vol) in a 50mL Falcon tube with the lid screwed on to the point that it can't be lifted off, but not so tight that the lid doesn't wiggle (not airtight!). Heat in a microwave, in a water bath or in water on a hotplate until completely dissolved. Tighten lid completely, then store in water bath at 42°C.

Agarose must remain in liquid phase for four hours. Adjust temperature of water bath as needed.

8. Sterilize all surgical instruments (except sharps – which can be sterilized with alcohol) by autoclaving prior to use.

9. Sterilize biosafety cabinet according to institutionally-mandated environmental health and safety protocols.

10. Place sterilized equipment, vibratome, micropipettes, and glass bead sterilizer in sterilized biosafety cabinet.

11. Test vibratome settings with an agarose block prepared from the agarose test. Add a sterile vibratome blade to the vibratome, cut a 2cm l x 1cm h x 1cm w piece of agarose and glue to the slicing chamber. Cut slices while adjusting vibratome settings so that agarose slices are uniform and intact with no cracks and minimal chatter.

Estimated time to complete this protocol – One week prior ~ 30 minutes plus time to make solutions (this could take several hours), 2 working days prior – 1 to 1.5 hours to make solutions. Day before ~ 3 hours

Preparation (day of)

12. Collect vibratome blades from storage, wash acetone off immediately with water and air dry. Alcohol sterilize and air dry shortly before use.

13. Cut off the bottom of a screw-top 50 mL Falcon tube at the 35 mL mark to utilize as a chamber for the liquid agarose and brain block.

14. Dissolve 400 mg of low melting point agarose in 20 mL slicing solution (2%wt/vol) in a 50mL Falcon tube and store in water bath at 42°C as in Step 7 above. This will be enough for two animals and will remain liquid in the tube until needed for the second animal.

15. Turn on laminar flow cabinet and clean surface of the hood using 70 % ethanol. Any material entering the laminar flow hood must be sprayed and wiped with 70% ethanol prior to being placed inside. Use UV irradiation to sterilize the hood for ~20 min prior to use. Clear a workspace on a bench near the hood to use for decapitation.

16. Sterilize and place 100 mm plastic Petri dish, cyanoacrylate, ice bucket with ice, syringe and needle, industrial razor blade, paintbrush, 24-well plates, paper towel, Falcon tube half (from step 2) and vibratome razor blade, and 50 mL Falcon tubes, pipettes and tips in sterilized biosafety cabinet. Test cyanoacrylate on a piece of filter paper prior to use to ensure it is liquid and emerges from the tube at a controlled rate – if not, discard and replace with new tube.

17. Add slicing solution to 24-well plates by filling wells halfway with plastic 25 mL serological pipet using a pipette controller. Fill an entire row (6 wells) for each hemisphere of BLA to be sliced. For four animals, add solution to two, 24-well plates.

18. Add 15 mL slicing solution to a 50 mL Falcon tube, 1 for each animal. Store 1 tube at −20°C until slushy for immediate use, and store the remainder at 4°C for later use.

19. Add ice to the Petri dish, close with cover and place in ice bucket.

20. Mount razor blade into vibratome blade holder.

21. Place tube of slushy slicing solution into ice bucket.

Dissection

22. At the clean workbench, spray neck of P14 rat with 70% ethanol, decapitate immediately using the guillotine and remove fur from scalp using surgical scissors.

23. Bring head into laminar flow hood for remainder of dissection and slicing.

24. Using small straight dissecting scissors with the tips pointed upward (to avoid damaging the brain), cut the skull along the midline from the foramen magnum vertically to the top of the cerebellum then along the suture to the front. Make 2 additional cuts to the left and right from the midline at the lambdoid suture and anterior to the coronal suture. This will facilitate the gentle removal of the skull.

25. Peel away the skull with care using forceps. Avoid contact with the cortical surface if possible. Once brain is fully exposed, use a polished microspatula to slide gently underneath brain (from caudal to rostral) and lift the brain into the slushy slicing solution. Screw on the cap and store tube in ice bucket.

We use crocus cloth (1500 – 2000 grit) – to polish all surfaces of the microspatula smooth.

26. Let brain cool in tube on ice for six minutes. While waiting, remove ice-filled Petri dish from ice bucket, dry water from top surface, then place filter paper on top and wet with slicing solution.

27. Remove warm liquid agarose from water bath and pour 10 mL into the cut off top of the 50 mL Falcon tube (cap down). Return unused agarose to water bath when finished.

28. Remove brain from slushy slicing solution and place on wet filter paper covered petri dish (Fig. 2A).

Figure 2.

Preparation of P 14 rat brain for OTC. A-C ventral brain surface as blocked (Circle of Willis indicated in A), D Block rostral face up (dorsal neocortex has been trimmed), E. Brain in agarose in cut Falcon tube. F. Tube removed from brain block. G. Agarose block removed from tube cap. H. Close-up of blocked brain (rostral face). Arrows indicate orientation of brain in block. All images were taken with a handheld camera

29. Gently remove meninges and major surface blood vessels from the ventral brain surface with a pair of straight forceps. This will prevent the vessels from interfering with the slicing. Particular care should be taken to carefully remove the vessels at the circle of Willis that invest the medial aspects of the nearby cortex.

30. Use single-edge (industrial) razor blade to make 2 coronal cuts (starting at ventral surface) and 1 transverse cut to isolate the region anterior and posterior to the amygdala and remove the dorsal neocortex as shown in Fig. 2B-D.

Agarose and Brain Block

31. Guide brain block into Falcon tube half containing liquid agarose and position the brain block to one end of the tube while maintaining correct axis orientation (Fig. 2E; ventral part facing away). Place the tube on ice. Note: agarose needs to be just warm enough to stay liquid but not too hot to damage the tissue.

32. Once agarose has solidified, remove block by removing cap then loosening block with micro-tapered spatula. Place on filter paper on top of petri dish. Using a single-edged industrial blade, cut a block of excess agarose for a backstop and trim the block containing the brain by making cuts parallel to the edges of the brain block (Fig. 2F-H).

33. Prepare the agarose “backstop” with cyanoacrylate on the side that will face down and adhere to the bottom of the slicing chamber. Glue the backstop to the bottom of the slicing chamber right up against the plastic block to cushion the blade from touching the plastic during slicing.

34. Prepare the brain block with cyanoacrylate on the caudal (now the bottom) side. The ventral portion of the brain must face the blade, with the rostral brain surface facing up. Place brain block in slicing chamber up against the backstop agarose block. Wait for block to completely adhere to the bottom of the chamber (Fig. 3A). Curing of cyanoacrylate is accelerated by contact with aqueous solutions like saline, so gently applying a few drops of slicing solution to the top and sides of the block with a pipette before the next step can set the glue and help prevent excess glue from forming a skin when filling the chamber with saline. This skin can interfere with slicing.

Figure 3.

Sectioning chamber and slice preparation for BLA OTC. A. Brain in agarose block glued into slicing chamber, ventral face out, rostral up. Note larger agarose “backstop” block behind brain block (arrow). B. 1 mL syringe with bent needle used to trim brain slices. C. P14 brain slice in slicing chamber. Lines indicate cuts to isolate slice containing BLA (⟡). D. Darkfield image of P14 brain slice. Outline indicates approximate boundaries of OTC slice placed on insert membrane. All images were taken with a handheld camera except D, which was taken with a compound microscope.

Slicing

35. Remove slicing solution from 4°C storage and place in slicing chamber. Pour slicing solution gently into the deepest parts of the chamber and allow it to fill to near the top of the chamber, submerging the brain block completely. Position slicing chamber in appropriate holder of vibratome.

36. Position razor blade of vibratome approximately 700μm rostral to where the amygdala begins and make first cut, using a slow advance and relatively wide blade excursion (e.g. 0.05mm/s and 1 mm). After completing first test slice, raise the blade 100μm before reversing the blade. (NB – depending on the vibratome, e.g Leica VT1000S, this might be an automated feature). Continue to cut test slices at 300 – 350 μm thickness until the beginning of the BLA is visible, always raising the blade before reversing.

37. Continue to cut 350 μm thick coronal slices containing BLA. Move each slice to the glass slide bottom of the chamber, then remove excess agarose and trim slice as shown in Figures 3B and C with a bent 26 ga 5/8” needle affixed to 1 mL syringe (used as a handle). Holding the bent part of the needle parallel to the bottom of the dish, gently press down with the needle (the tubing, not the tip) to cut the desired section free of the remainder of the slice (Fig. 3D)

38. Use an industrial razor blade to remove the tapered tip section of a disposable 1000μL pipette tip and attach to the 1000 μL Gilson (or equivalent) pipettor. Collect the slice from the slicing chamber with utmost care using the modified 1 mL pipette tip and gently pipette into 24-well plate containing cold slicing solution (Fig. 4A). It is essential to avoid bending or warping the slice. Repeat until BLA is no longer visible in brain block.

Figure 4. OTC placement on inserts and quality check.

A. placement of slices and inserts in 24 well plate during plating. B. Healthy OTC of EP 21. Note preserved ventral curvature of clearly layered neocortex and entorhinal fissure (upper left side of slice). outlined teardrop shape is BLA. C, D, Damaged slices with elevated white tissue (red outlines - discard any such slices). E. Healthy EP70 OTC on insert membrane. BLA enclosed is within square area. As in Figure 2, images all taken with handheld camera except E, which was taken with a compound microscope.

39. If needed, use a dissecting microscope to determine which slices contain BLA. There should be four slices containing BLA per hemisphere for a total of eight slices per animal. Label slices (on plate cover) and store at 4°C until next animal is finished.

40. Repeat necessary steps from preparation prior to dissecting the second animal. Store slices from the second animal in the other 24 well plate, separate from those from the first animal. Turn the glass bead sterilizer on immediately after slicing the second brain.

41. Allow slices from the second animal to rest for at least 30 minutes at 4°C.

42. While above slices are resting, add 350 μL of fresh cold culture media to the middle 4 wells of each of the 2 upper rows of a new 24-well plate (Fig. 4A). Bead-sterilize forceps not used during dissection; once cooled, use them to place culture inserts into the lower 8 wells. The membrane will be below the surface of the solution.

43. Withdraw the plate containing slices from the first animal and place on top of ice in the ice bucket in the biosafety cabinet.

44. Sterilize the sable brush by dipping in ethanol and immediately rinsing in sterile slicing solution in one of the wells not containing a brain slice, or a well containing a slice that does not have BLA. Pat the brush dry on fresh filter paper. Prolonged contact with the ethanol can damage the expensive brush.

45. Prepare a 1 mL pipette tip as per Slicing Step 4. Set the micropipette to 200 μL and gently withdraw the first slice of animal 1. Carefully pipette the slice onto the membrane of the culture insert. Allow the slice to float down and prevent contact between the pipette tip and the insert if at all possible.

46. Remove excess slice solution using a micropipette set to 100μL. Place the pipette tip away from the slice and lightly withdraw slicing solution, while not drawing the slice away from the center of the insert membrane. Ensure the slice is not in contact with the side of the membrane insert when finished.

47. Gently position slice in the middle of the insert membrane using the broad edge of the fine sable brush (Fig. 4B). Avoid using the tip of the brush.

48. Using the recently-sterilized forceps, lift the insert and place into the corresponding upper well of the 24 well plate containing media. Immediately add one drop of fresh media gently next to the slice to prevent slice from drying.

49. Continue to transfer remaining slices to inserts for that animal (Fig.4A).

50. Check that the slices have not changed position since being plated. If they have, guide them once again using the sable brush and remove any visible excess solution using a new 100μL pipette tip. There should be a slight dampness around the slice, but no solution collecting in the edges of the insert where it is attached to the membrane.

51. Once all slices are positioned, label plate and place in incubator and repeat plating procedure for second animal. Repeat necessary preparation and following steps for remaining animals.

Estimated time to complete this protocol – can be up to 6 hours initially, but with practice two animals can be prepared in 4 or so hours. With assistance in preparing solutions, etc. (this is very helpful in this step!) this can be even shorter.

ANTI-MITOTICS

An anti-mitotic solution is prepared to limit glial proliferation in the slices. Use the anti-mitotic solution only once, 48 hours after slices were first incubated. Return slices to normal media after 24 hours exposure to the anti-mitotic solution.

Preparation

52. Dilute antimitotic solution stock (10⁻⁴M stock– see solutions, below) 1:100, i.e. 1 aliquot of 1 mL anti-mitiotic stock in 100 mL media, resulting in a 10⁻⁶M final solution.

Procedure

53. After 48 h in culture, incubate cultures with 300 μL of anti-mitotic solution for 24 h to reduce glial proliferation (prepare solution using warmed culture media with stock anti-mitotic solution to a final concentration of 10⁻⁶ M).

54. Following 24 h anti-mitotic incubation, change media 3x/week with 300 μL of fresh warmed media.

Estimated time to complete this protocol – The task itself is 90 minutes, but it is best if the prepared anti mitiotic solution is then left to equilibrate to the correct pH in the incubator for at least 4 or more hours prior to changing the media .

Basic Protocol 2: EXCISION OF OTCs FROM INSERTS

The OTC slices bond inextricably with the membranes of the inserts, so for electrophysiological recordings, we had to excise the membrane with the OTC from the inserts before placing them into the fixed stage recording chamber of an upright infrared DIC videomicroscope. Moreover, we routinely used a bicarbonate-CO₂ buffer system in the aCSF, so we equilibrate it with 5%CO₂, 95% O₂ (carbogen) for the slices. This would be unnecessary when using aCSF with a non-CO₂ –dependent buffer.

Materials

Cultured OTC slices

Prepared aCSF

Water bath set to 37°C

5 mL plastic pipettes

#11 scalpel blades

Clamp-style forceps or (preferably) blade breaker (e.g. Fine Science tools Item No. 10053-09

Breakable razor blades (e.g. breakable scalpel blades Fine Science Tools 10050-00)

Vannas-Tübingen spring scissors, 3-5 mm blade, angled to side, or equivalent

Fine forceps ( e.g. Dumont AA-style tweezers)

Angled forceps (e.g. FST 11049-10)

60 x10mm petri dish

Two platinum (or other inert, relatively dense metal) rings from partly flattened, 1mm wire, the larger diameter ring with fine fibers* glued on (“harp”), the smaller one just large enough to weigh down the membrane

Micro-tapered spatula, with approximately 8-10 mm at the tip of the wider end bent at a 30° angle to transfer membrane from petri dish to recording chamber

*Excellent, non-autofluorescent fibers can be unspun from unwaxed polyester dental floss, then wound at the appropriate spacing around the ring while it is stuck to a small plastic weighing dish with double-sided tape, and glued in place with small drops of 5-minute epoxy.

Preparation

1. Pipette 25 mL of Carbogen-bubbled aCSF into a tube and incubate in water bath to 37°C. This must be done in less than 30 minutes or the aCSF will begin to precipitate. Turn on glass bead sterilizer at the same time.

2. Prepare microscope area with #11 scalpel blades, blade breaker or clamp style forceps, fine tip forceps, angled forceps, and plastic pipette tips.

3. Wear PPE. Pipette ~15 mL aCSF from the warmed tube into a 35mm petri dish.

4. Spray gloves and forearms with 70% ethanol. Remove well plate from incubator and place 35mm petri dish and well plate inside biosafety cabinet. Bead-sterilize fine forceps and wait for them to cool.

5. Using fine forceps, remove the insert with slice to be used and place gently in heated aCSF. Cover immediately with petri dish lid. Keep records in multiple places regarding which slice has been removed at that particular time and date, e.g. on cover of well plate and in lab book.

6. Spray 70% ethanol on well plate, gloves, and forearms. Return well plate to incubator and take slice in petri dish and remaining amount of aCSF in tube to dissecting stereomicroscope to be excised.

Procedure

7. Place petri dish on stage of stereomicroscope. Gently pipette remaining aCSF along the inside of the insert until the insert is full. Pipette remaining aCSF into petri dish until the insert is floating.

8. Using fine Graefe forceps or other fine tweezers, gently invert the insert so the bottom of the membrane is facing upwards. Do this without collecting air bubbles in the chamber of the insert as this will damage the slice.

9. Using #11 scalpel blade, or a razor blade fragment held in a razor blade breaker, or equivalent thin blade, begin cutting the membrane away from the sides of the insert. It is best to do this in as few strokes as possible.

10. Once the larger cuts have been complete, use angled spring scissors or similar to make the final cuts to leave the slice floating freely in the aCSF. It is imperative that the membrane and slice be flipped upwards as soon as possible to preserve the correct orientation (slice facing up). A fiber optic dissecting light can be used to illuminate the slice from the side to distinguish the desired orientation, moving the light to optimize visualization using light and shadow.

11. Immediately drain most of the aCSF from the petri dish back into the tube and carry the slice inside the petri dish to the recording chamber of the microscope.

12. Turn the aCSF inflow off and stop suction from the recording chamber.

13. Using bent micro-tapered spatula, scoop the slice out of the petri dish and place the membrane slice side up in the chamber as close to the center of the chamber as possible. Using a pipette, place drops of aCSF on the surface of the membrane to encourage the slice to sink. Do this in a way that minimizes forces on the slice.

14. Once the slice is on the bottom of the recording chamber, quickly focus on the slice using the low magnification (e.g. 5x) objective. Use the field diaphragm to concentrate the light in the approximate location of the BLA.

15. Place the larger harp with strings aligned parallel with the orientation of the recording micropipette, and bracketing, but not touching the BLA. Place the second ring with its edges to surround the slice, ensuring that there is access to the BLA for the micropipette.

16. Restore flow of aCSF and suction. It is important that the suction is placed in the same spot, as the amount of aCSF in the chamber will affect the stability of the temperature between each experiment.

Estimated time to complete this protocol – will take some, practice but should be done in 30 min or less for maximum viability.

Support Protocol 1: CHANGING MEDIA

Media must be changed three times per week to maintain culture health. It is best to maintain a regular date and time schedule to preserve integrity of the experiment. Carrying out the media changes at the same time each day, and doing them spaced apart relatively equally, e.g. Monday, Wednesday, and Friday will also ensure that media changes are performed regularly. To change media, the trays must be withdrawn from the incubator in full PPE and using appropriate aseptic practices.

Materials List

25 mL sterile serological pipettes with controller

Nitrile gloves

Calf-length lab coat with cuffs

Face mask

Complete medium

25 mL sterile serological pipettes with controller

1 L vacuum flask

Fine forceps

10% bleach

70% ethanol

Preparation

1. Don PPE: full length lab coat with cuffed sleeves, nitrile gloves, and a mask.

2. Collect materials required to change media, including the appropriate number of tubes to heat media, serological pipettes of relevant size (25 mL), and retrieve media from refrigerator.

3. Using aseptic technique, prepare quantities of media required to change media in all cultures. Each well must be filled to 300μm. It is best to prepare a little bit of extra media in case of spills or misfiring of the pipette. Therefore, calculating 350μL per culture slice is adequate. Media must be incubated in amounts of no more than 25 mL per tube.

4. When screwing the top of the tube to seal the media, leave the lid loose enough that it is not completely sealed but that the lid does not come off when pulling it upwards. This permits the carbon dioxide in the incubator to begin adjusting the pH. Place the media in the incubator to warm, but be aware that the pH equilibrates more slowly than does the temperature. Viability is enhanced if this is considered. It is best to have a sterilized rack within the incubator to stand the media tube up. It is difficult to maintain a sanitized environment if the media spills or leaks.

5. Depending on the size of the incubator and volume of media in each tube, the media will take a varying amount of time to heat to the required 37°C. Therefore, a sample should be prepared prior to changing the media to determine how long it takes to reach the correct temperature.

6. Discard PPE and return media to the refrigerator. Dispose of all contaminated materials in autoclavable bag.

7. Twenty minutes prior to changing the media, turn on UV light in laminar hood and glass bead sterilizer.

8. After media has reached the appropriate temperature, don PPE again.

Procedure

9. Prepare vacuum flask inside the biosafety cabinet. Vacuum flask should contain 25 mL of bleach to start. Turn on vacuum flask and attach glass pipette to the tube.

10. Spray gloves and midway up the forearm with 70% ethanol. Open incubator and stand next to the incubator to remove well plate. Do not stand inside the glass door of the incubator. Well plates should only be removed one at a time. Remove one tube of media from incubator.

11. Sterilize fine forceps in glass bead sterilizer and allow them to cool before using.

12. Remove the lid from the well plate. Pipet 300μL of media into the wells corresponding with the ones already filled with media and an insert, e.g. if the upper 8 wells are filled, fill the lower 8 with fresh media.

13. Using sterilized fine forceps, move each insert from old media into new media. It is best to hold the insert on an angle to place the insert gently into the media. It is imperative that no air bubbles are beneath the membrane attached to the insert.

14. Once all inserts are in new media, place forceps in sterilizer again.

15. Prepare the glass pipette with the vacuum flask suction turned on. Attach a 10μL or 100μL plastic pipette tip to the glass pipette using suction. Use this to remove the media from the used wells. It is helpful to slightly tip the well plate to ensure complete removal of the media.

16. Once finished, discard pipette tip and cover well plate. Remove forceps from bead sterilizer to cool. Spray gloves, forearms, and well plate with 70% ethanol and return well plate to incubator. Repeat for all well plates that are due to be changed.

Estimated time to complete this protocol – once the setup is complete and solutions equilibrated, the changing of media for 8 plates should take no more than about 15–20 minutes. Care is important, so emphasize that over speed, especially initially.

Support Protocol 2: DRUG INCUBATIONS

Drug incubations are carried out much the same as media changes, so only the test compounds (e.g., peptides or drugs) must be prepared in advance to incubating the media. Serial dilution technique must be used to prepare the final solutions. Keep a vortex mixer near or within the biosafety cabinet to ensure proper mixing. Be aware that mixing the media produces bubbles and it is ideal for the media to settle without bubbles prior to distributing. Alternatively, use of the pipettor and sterile pipette to gently draw up and expel the solution a few times can mix the solution with less agitation.

Preparation

1. Culture OTCs for 4 weeks prior to any experimental manipulations/procedures (6 weeks equivalent postnatal, EP). For drug incubations starting on culture week 5, change media daily according to the length of the treatment, each time adding fresh drug diluted in fresh warmed media.

2. Thaw solutions of the required reagents (peptides, drugs, etc.) and place in biosafety cabinet.

3. Prepare all materials indicated in treatment protocol and calculate serial dilutions required to reach final concentration.

4. Place sterile 5 mL microcentrifuge tubes in biosafety cabinet.

5. Place cold media in the biosafety cabinet with required serological pipets and tubes from ‘Changing Media’ preparation.

6. Prepare in advance all tubes of drug solution with required volume of media for each final concentration of drugs to be tested that day..

7. Pipette drug or solution into intermediate solutions according to serial dilution calculations, and use a new pipette each time until final concentration for each solution is reached.

8. Cap the microcentrifuge tubes and the media and place in the incubator until the media is changed.

Procedure

NB. In our experiments, we mimicked the in vivo protocol we used to study NPY-induced stress resilience, which entailed 5 daily intra-BLA infusions of NPY. The following reflects this, but could readily be altered to reflect other in vivo protocols.

Carry out ‘changing media’ procedure with the following modifications.

9. Drug incubations will be done daily from Monday to Friday. Media will be changed in control slices according to the same schedule using plain media or a diluted control solution such as saline or an inactivated peptide.

10. Drug incubations will be left to incubate over the weekend on the Friday of each treatment week and changed back to regular media three times a week subsequently.

11. Following drug incubations, return to using drug-free normal media, and continue to culture with normal media changes for the remainder of the time course.

Estimated time to complete this protocol – Preparation of the solutions will depend on the complexity of the experiment, but once they are completed, figure about 50% more time than for conventional media changes.

Support Protocol 3: FIXATION OF SLICES

When all electrophysiological recordings have been completed for a given slice, it is then fixed for staining and subsequent imaging. This was an essential step to correlate the changes in whole cell capacitance with changes in morphology caused by repeated treatment with NPY or CRF (Michaelson et al., 2020). While we initially followed a fixation protocol that used 10% formalin, we subsequently determined that use of the 4% paraformaladehyde protocol significantly improved results.

Materials

4% paraformaldehyde solution (freshly prepared, or stored at 4°C)

5 mL Disposable serological pipettes

Fine forceps (like Dumont AA). Do not use any tweezers that touch fixative to handle live tissue!

Glass vial with screw top lid (e.g. 10 mL liquid scintillation vial)

PBS with 0.2% sodium azide

Procedure

1. Prepare glass vial with 1 to 2 mL of bubbled aCSF using a pipette. Label it appropriately (e.g., with the date, name of experimenter, slice ID, and experimental conditions).

2. Stop both suction and aCSF inflow to the recording chamber.

3. Using fine forceps, gently remove the rings holding down the OTC (smaller one first) and attached membrane. Pick up the slice with the forceps (by the membrane, carefully avoiding the relevant tissue!) and place it gently in the vial. Submerge it in aCSF, if necessary using a couple of gently pipetted drops of aCSF to submerge slice. An alternative is to use a fine flat spatula with a bend in the blunt side blade to slide under the slice in the chamber (holding the edge of the slice with fine forceps to keep it on the spatula if needed), then lift slice with spatula from chamber and gently introduce into aCSF in vial.

4. Wear the appropriate PPE to handle 4% paraformaldehyde. Gently pipette the aCSF from the vial and replace it with 1 – 2 mL of fixative, ensuring the surface of the slice is completely submerged.

5. Seal the (labeled) lid of the vial and refrigerate (4°C) the slice for 36-60 hours (no longer – the quality of slices deteriorates rapidly if left in fixative too long). Immediately thereafter, gently remove PFA (dispose of appropriately) and replace with PBS containing 0.2% sodium azide. Store slices at 4°C for up to 28 days, although it is highly recommended to process and mount the slices within one week.

Estimated time to complete this protocol –This takes about 5 minutes of active intervention overall, excluding the waiting times as indicated.

REAGENTS AND SOLUTIONS

Note: All solutions below must be filter-sterilized before making aliquots for storage or use. Prepare all solutions using HPLC grade, ultrapure water or equivalent and analytical grade reagents.

50% D-glucose solution:

Prepare by dissolving 50 g of D-glucose into 100 mL of HPLC grade ultrapure water, sterilize via syringe filter and store at −20°C as 6 and 8 mL aliquots.

Kynurenic acid (3 mM):

Prepare the stock solution by dissolving 61.15 mg of kynurenic acid into 100 mL of HBSS, sterile filter and store at as 5 mL aliquots at −20°C. Sonication facilitates dissolving the kynurenic acid.

L-ascorbic acid (50 mM):

Prepare the solution by dissolving 352.24 mg of L-ascorbic acid into 40mL of HBSS, sterile filter and store at −20°C as 4 mL aliquots.

Penicillin/streptomycin:

From 5000 U/mL stock solution, store at −20°C as 1 mL aliquots.

Slicing solution:

To a full 500 mL bottle of HBSS, add one 5 mL aliquot of 3 mM kynurenic acid and one 6 mL aliquot of 50 % D-glucose solution, store at 4°C.

Culture medium (for ~ 400 mL):

To a large beaker, add 100 mL heat-inactivated horse serum, 200 mL MEM, 84 mL HBSS, 2 mL Glutamax, 8 mL 50% D-glucose, 2 mL penicillin/streptomycin and 4 mL L-ascorbic acid. Stir and adjust pH to 7.2 using NaOH. If necessary, adjust osmolarity to 320mOsm using HPLC grade ultrapure water. Bubble the final solution with 5% CO₂ for 60 seconds prior to filtering with Stericup® in biosafety cabinet. Store at 4°C and discard any unused media after 28 days.

Anti-mitotic stock solution (10-4 M):

Add 3 mg of each cytosine-β-D-arabino-furanoside, uridine and 5-Fluro-2’ deoxyuridine to 100 mL HBSS, sterile filter and store at −20°C as 1 mL aliquots.

aCSF:

124 NaCl, 3 KCl, 1.3 MgSO₄, 1.4 NaH₂PO₄, 10 glucose, 26 NaHCO₃, and 2.5 CaCl₂. Osmolarity 320 mOsm/L, adjust if needed with NaCl or HPLC grade water. Bubble with 5% CO_2, 95% O₂ for 10 minutes prior to storing at 4°C. Do not use stock solution and do not prepare more than in a day in advance. This may vary based on experimental conditions, but osmolality has proven to be critical for this and the internal solution.

4% paraformaldehyde fixative solution:

Using appropriate PPE and working in a fume hood throughout, add paraformaldehyde to prepared PBS (4 gramsPFA/100 mL PBS) in a beaker (with stir bar), heat beaker gently on stirring hotplate in fume hood to 55°C, add 2 pellets NaOH/100 mL, then adjust to pH 7.5. Filter and store at 4°C . Use within 1 week.

Phosphate-buffered saline (PBS):

(0.02 M sodium phosphate with 0.15 M sodium chloride). To prepare 1L PBS, dissolve 2.84g Na₂HPO₄ and 8.77g NaCl in HPLC-quality water to make 1.00L

COMMENTARY

Background Information:

When we developed this long-lived OTC preparation, we initially sought a tool to identify pharmacology and mechanisms underlying the long-term changes in behavior (social interaction) seen with repeated administration of NPY into rat BLA. The preparation that resulted was not only novel in its longevity, but also unexpectedly pointed the way to a surprising type of reversible remodeling of dendritic structures. The results from the OTC experiments not only predicted the changes we would subsequently observe in vivo but also allowed us clear mechanistic experiments into the underlying signaling of the bivalent changes in BLA PN structure (Silveira Villarroel et al., 2018. Michaelson et al., 2020).

Here we provide a detailed protocol which should facilitate the adoption of the BLA organotypic slice preparation, and which moreover appears to be effective for OTC preparations of amygdala and neocortex in mice (Michaelson et al. unpublished results). In our hands, the EP70 OTC PNs retain most key properties of their counterparts in P70 rat, appear to be synaptically wired in similar ways and certainly retain much of the neuropeptide pharmacology in the system. Nonetheless, OTCs clearly undergo some remodeling after being placed in vitro, including loss of coventional anatomical markers such as the internal capsule, and thinning of the slices over time. This thinning or flattening, commonly described common in OTC preparations, serves as a macroscopic criterion of healthy slices, is consistent with the increases in cell density seen in OTCs from other brain regions (Guy et al., 2011, Humpel, 2015a), and which can be seen here as well(eg. Fig.5). Moreover, the OTC preparation is of necessity an axotomized system (as are acute ex vivo slices), so neurons lose both afferent and efferent connections with distant brain regions. In other OTC models, the degree of connectivity between neurons appears to depart from that of acute slices and approaches the higher levels observed in vivo (Holopainen and Lauren, 2003, Pena, 2010, Okamoto et al., 2014). This increase in connectivity may well be attributed to synaptogenesis and neuronal sprouting, which occurs within days to weeks in hippocampal OTCs (Stoppini et al., 1993, Robain et al., 1994). This implies that OTCs may develop altered local synaptic circuits and can be a limitation of the technique.

Figure 5: Overview and higher magnification images of BLA and principal neurons.

Infrared DIC optics. A,B P70 acute ex vivo slice. C,D P14 acute ex vivo slice. E,F EP70 OTC slice. E.C. – External Capsule. Inset panel E: Higher magnification of external capsule remnant (arrows). Scale bars: A,C,E – 200 μm, B,D,F, Inset – 20 μm.

Nevertheless, our comparison of electrophysiological properties of principal cells at different stages of development in acute slices are in agreement with previously reported findings of changes in BLA in vivo, consistent with normal development (Ehrlich et al., 2012). Moreover, the majority of electrical features, RMP, rheobase, input resistance, I_h-amplitude and density, and sIPSC and sEPSC frequencies, of BLA OTC principal neurons were closer to neurons in age-matched acute slices than to those in P14 rats. Furthermore, the immunohistochemical analysis of OTCs is consistent with most of the major features in the corresponding adult brain. Together, this suggests that BLA OTCs mature in a manner comparable, though not identical, to that seen in vivo.

Despite their ability to model, mirror and indeed predicted the changes seen in vivo, an important caveat for potential users is the constraints on expansion of the dendritic trees seen in the OTCs. The smaller dendritic architecture of BLA OTCs likely explains the differences in amplitudes of spontaneous postsynaptic currents between OTCs and acute slices. Synaptic currents in non-spherical cells are particularly sensitive to the lack of space-clamp, a distortion of the measured current with increased distance from the current source (Spruston and Johnston, 2008). This results in poorer voltage clamp at more distal dendritic membranes and is exacerbated by the extent and complexity of the dendritic tree. Synaptic events recorded at the soma of larger PNs are slower and smaller in amplitude than the local synaptic currents injected at distal dendritic regions (Bar-Yehuda and Korngreen, 2008), consistent with the differences observed between synaptic events recorded in EP70 PNs relative to those in P70 acute slices. This might be an important consideration for those interested in biophysical aspects of synaptic transmission.

We never observed compound postsynaptic bursting activity in acutely prepared slices, while this was almost ubiquitous in OTCs, to a greater or lesser degree. While rarely seen in acute BLA slices, phase-locked oscillations in the high delta/low theta frequency band occur frequently in the BLA in vivo, and have been shown to contribute to learning processes (Lesting et al., 2011, Ryan et al., 2012). In acute BLA slices, these bursts result from synchronized, rhythmic network activity and can be induced either by elevations in bath K⁺ concentration or by electrical stimulation of the slice (Rainnie, 1999, Chung and Moore, 2009). Studies in OTCs from neocortex demonstrated that spontaneous activity increases with development in vitro, culminating in bursts, or “up and down” states, with a decrease in intrinsic excitability (Johnson and Buonomano, 2007). While it seems plausible for bursting activity observed in our OTCs to be a component of normal physiological behavior, the loss of the normal afferent and efferent projections typically results in the up-regulation in growth of more localized synapses, unnaturally enhancing local neural circuits to increase overall network activity in the OTCs. In the only other report we were able to find on BLA OTCs (Bender et al., 2017), early postnatal (P3) corticoamygdalar slices were described that survived ~2 weeks. While reporting biochemical responses and some morphological data (including electron microscopy of synapses), this study unfortunately did not report electrophysiological responses in these neurons.

Together the results here and in our previous publication (Michaelson et al., 2020) clearly validate numerous critical similarities of long-term BLA OTCs with their in vivo counterparts, and certainly directly predicted unanticipated and suprising findings in vivo. This makes OTCs of the BLA a potentially very attractive tool with which to study stress related circuitry and related neuroplasticity. The OTC model of the BLA (or of other constellations of amygdalar nuclei) should prove useful as tool to generate mechanistic insights into the physiology governing stress, stress resilience and the signal transduction processes underlying plastic changes caused by various stress mediators. Moreover, the apparently transferability of this approach to the mouse opens the door to taking advantage of the myriad genetic models available (particularly those that are conditionally inducible) for detailed mechanistic and developmental studies in vitro.

Critical Parameters

Prior to beginning, it is essential to consider the time requirements and constraints of the organotypic slice protocol. To acquire the skill and sensitivity to produce healthy cultures has taken past members of our lab up to a year of consistent work, though some achieved this considerably more rapidly. Best results will be achieved if the procedure is repeated many times early on, e.g., producing cultures every day for a week to start and then once every two weeks until the desired viability is reached. One of the appeals of slice cultures is the reduction in the number of animals necessary to complete experiments, but this reduction will not be achieved until the protocol has been perfected and mastered.

The viability of the cultures relies on strict adherence to the initial protocol at first, and afterwards making small, systematic changes to suit your individual lab environment and the speed of the individual performing the preparation. Common issues are further described in the troubleshooting section.

The protocol for the organotypic slice culture appears simple, but there are many fine details that will contribute to a successful outcome. All small details must be initially adhered to, including temperature and time specifications. Based on our own experience, it will generally take the experimenter numerous attempts to become proficient at the dissection. Systematically keeping logs of procedures and any variations or errors at all stages of the procedure is highly recommended, as is regular microscopic inspection of the slices to monitor their health. Mistakes in the procedure will produce a family of patterns of ill health in the slices, and will become characteristic over time.

Animal Handling and Stress

While euthanasia of rat pups is less challenging than with their mature counterparts, we have found nonetheless that care must be taken to avoid stress to the animals. Pups that will be euthanized that day should be kept in a temperature-controlled room, with the accustomed light/dark cycle, and kept together until removed one at a time for euthanasia to reduce stress. Any pup briefly left alone should be kept in a cage or a secure box with bedding for warmth and comfort and access to adequate water and food.

P14 rats often have not opened their eyes yet, and gentle handling will ensure they remain relaxed prior to euthanasia. Interaction with the pup can stress it, and contribute to excess bleeding with decapitation and increased likelihood of a failed slice culture due to contamination.

Euthanasia and Dissection

Inducing anaesthesia is itself stressful to animals, so here we prefer decapitation without prior anaesthesia for the euthanasia (Holson, 1992), which also helps prevent contamination of the brain tissue. Use of a well-sharpened guillotine rather than scissors is strongly recommended for ease and speed of handling. Euthanasia must take place as close to the biosafety cabinet as possible to prevent contamination of the tissue and extending the time of the dissection. The guillotine can be set up on a cart near the biosafety cabinet, and moved away when not required. Guillotine and area must be well cleaned between dissections to prevent stress to the next animal.

After decapitation, it is best to hold the head gently from the front by the muzzle to secure it while removing the brain. The skull is fragile, so care must be taken to avoid pressure or other mishandling of the brain. Notably, dropping the brain suddenly into the aCSF or taking too long to submerge it, can negatively affect viability. Dissection instruments are best sterilized via autoclave or bead sterilizer, then allowed to cool to room temperature prior to dissection.

Agarose and Brain Block

The agarose must be heated until boiling and shaken to ensure uniformity, before allowing it to cool to just above its melting point. It is important to ensure the agarose remains liquid from preparation until embedding, and it must remain so for 4 hours during OTC preparation. In our lab, agarose remained liquid at 42°C, but should be tested intermittently as it can vary with the age of the agarose and other environmental factors. Tubes must be submerged to slightly below the level of the liquid agarose to prevent the formation of film on top, which will weaken the brain block.

If the agarose brain block cracks while preparing it for the slicing stage, there is a chance the slices will become uneven. It must sit on ice long enough to harden the agarose, but not too long so as to unduly lengthen the time of the experiment.

Slicing

It is essential to practice slicing the kind of blocks and thicknesses with your instrument prior to using live animals for the procedure. Using fixed brain tissue for this is discouraged because the fixed material does not mimic the properties of fresh tissue, and the fixative can contaminate your slicing chamber. Practicing on plain agarose blocks can help you determine the appropriate speed and frequency to contribute to a smooth surface on each side of the slice, without distorting or pressing on the brain block or slices. In preparing slices for culture, it is important to trim the block as closely to the correct brain area as possible, to speed up sectioning and prevent mechanical stress to, or distortion of, the brain block in the slicing chamber, especially when cutting thicker sections while approaching the region of interest. Ideally, one will begin to section only 1 mm or so above the first slice to be used for culture – this does take some practice. It is best to take two practice slices, with at least the last one being 350 microns, and always raising the vibratome blade to 100 microns above the cut surface prior to reversing the blade (this avoids dragging the blade edge at high speed on the fresh surface), then lowering the blade the extra 100 microns to ensure the desired slice thickness. While not essential during the approach slices, this does make a big difference with the slices that will be placed into culture. As mentioned elsewhere, a vibratome that automatically raises the blade during retraction is an advantage here, but doing this manually rapidly becomes routine.

It is especially important to make sure that the block is trimmed small enough so slices will fit into a glass pipette or cut-off pipettor tip without the tissue being bent or folded. The vibration frequency for cutting should be set low enough that the liquid in the chamber remains sufficiently calm so slices remain relatively still for manipulation and cutting. Having to pipette the slice out more than once will contribute to tissue damage.

Incubation

The ideal incubation time is between 30 and 60 minutes. Do not incubate slices from consecutive animals in the same dish, as maintaining the tissue at 4 °C during this time is critical for OTC viability.

Plating

Pipette the slices onto individual, dry, sterile inserts. It is most efficient to do this procedure with two slices at a time. Remove the slicing solution and place the inserts into the media so they can begin to moisten the bottom of the slices. If necessary, add a drop of media NEXT TO (not on top of!) a slice placed on the insert, using a 10 μL pipette to moisten the edges and prevent it drying in the middle. If the slice is not plated correctly, if it dries out, or if it is plated without lying flat on the insert membrane, it will raise in the middle and create a dry spot right where the BLA is. It is of utmost importance to position slices correctly as soon as possible to prevent damaging the tissue through repeated movement. A moistened brush with a fine, flexible tip (see materials) is the best way to gently guide the tissue to the middle of the insert membrane; tissue must not remain in contact with the outer plastic edge of the insert, as the slices will eventually be cut out of the inserts. Once all 8 slices have been mounted on their inserts, observe the slices and make sure none of them are floating or have moved from where they have been plated, and that the inserts appear slightly moist around the slices, and the surface of the slices doesn’t look dry. Apply 1 μL drops of media, one at a time, to the edge of the slices until they no longer appear dry, but not so much that they start to float. This will take some practice. If there is too much liquid in the inserts, carefully remove some with the 10 μL pipette without disturbing (or touching!) the slice itself.

Post-dissection

After 24 hours in the incubator, it is important to check the slices to ensure they remain moist and haven’t dried out. If slices are drying out after 24 hours, it is likely they won’t be viable so make sure to record this in your journal and review the protocol notes to determine see if this was a mistake in following the protocol correctly or arose from a systematic error in the protocol. Either conclusion should be clearly annotated, or procedures revised if necessary. At this timepoint, you can add a couple of drops of media to the edge of the inserts, but it may already be too late. Sometimes the slices will swell, which correlates with the amount of time they spent in slicing solution prior to plating. Continue to monitor the slices at each media change.

Once the slices adhere to the insert membrane, they will thin to about 80-100 microns over roughly two weeks. Healthy slices appear uniform in thickness, and are translucent after a couple of weeks (Fig. 4B). If slices appear white, thick, or puffy (Figs. 4C, D), they are likely to be non-viable and should be removed from the well plate and the incubator.

Until the slices are routinely satisfactory, it can be difficult to tell the location of the BLA after the slices have thinned out (Fig. 4E). We have found it ideal to take photos of the slices under a dissecting microscope (we use a handheld small digital camera or even a cell phone aimed through the eyepiece of a stereomicroscope, see Figs. 4B-D) to determine the location of the BLA while it is still visible. With experience, this may eventually no longer be required.

Troubleshooting

Non-viable cultures

While strict adherence to the procedure and notes in this protocol will eventually produce the best results, it takes repeated experience and dedicated attention to consistently reproduce a high volume of viable cultures. It is reasonable to expect non-viable cultures from the initial attempts at this protocol, but the number of viable cultures will increase with practice.

Healthy cultures will always be thin and translucent, and as such this is the number one indicator for the health of cultures. Any slices that remain thick and white or orange-brown are non-viable and may be discarded to conserve time and materials (Figs. 4C, D). Still, there will be cultures that appear healthy, yet prove unhealthy when prepared for electrophysiology. Again, depending on resources it may be best to discard these slices, especially if there are healthy cultures incubating and waiting to be used.

Once examined with a compound microscope (we use infrared differential interference contrast – ir-DIC), healthy cultures will appear uniform, with cells appearing quite obvious and easy to identify (Fig 5). If the area of interest demonstrates lots of bubbles and debris, this part of the culture is non-viable.

Other factors contributing to unhealthy cultures in the incubation phase:

The pH of media is unstable – thus media must always be freshly prepared every 28 days or less. The media should always be buffered with CO₂ post-filtration, as vacuum filtration removes dissolved gases in the media and will raise the pH slightly. The media should also be exposed to CO₂ in the incubator for at least one hour prior to media changes.

Old/expired reagents – reagents and solutions must always be made fresh when specified, and stored at the required conditions without exception. Any expired reagents must be properly disposed of. If in doubt, throw it out.

Agarose temperature – if the agarose is too hot, it will contribute to non-viability of the cultures. Measuring the temperature of the agarose while preparing the brain block is impractical, so keep the agarose at the lowest possible temperature that maintains it in the liquid phase prior to use.

Aggressive or hasty dissection and preparation technique – those with prior dissection experience and delicate fine motor skills will have an advantage here. It takes time to acquire the sensitivity and delicate touch required to produce great cultures. This is simply a matter of time and practice. It is recommended to manipulate the slices as little as possible. Each time the slice is manipulated there is a risk of damaging it.

Slices too dry or too moist – it is imperative when mounting slices on the inserts to ensure they do not dry out, but also to be sure that, in keeping them moist, not too much media is left in the insert. Additionally, it is important to use a timer to track the duration of slices exposure to 4°C incubation; if left to incubate too long, they will swell too much and become non-viable.

Factors contributing to slice death during electrophysiological experiment phase:

Aggressive or hasty handling of OTCs can negatively affect outcomes. Care in handling the OTC while removing from the insert, placing in the chamber and positioning correctly is essential. Sanitation – all hardware and utensils that come into contact with the slice should be as clean as possible. Saline delivery lines and the recording chamber should be flushed and rinsed, respectively, with distilled water daily after experiments, and flushed with several mL 1N HCl weekly to prevent biological contamination, especially fungus.

Temperature – temperature of water bath should be calibrated on a regular basis and the recording chamber temperature should be checked. Flow rates are key if the saline is heated outside the chamber, as is the case in our lab. aCSF – should always be made fresh, we have found it best to not use concentrated stock solutions of salts to dilute into aCSF. Likewise, we do not reuse aCSF that has been heated and carbogenated one day, then refrigerated overnight.

Viability of BLA OTCs

Once the protocols are routinely implemented, BLA OTCs obtained from P14 animals can be maintained up to EP70 and beyond. BLA slices will typically thin over time in vitro from 350 μm to ~ 80-100 μm, and the external and internal capsules will greatly diminish, making determination of the boundaries of the BLA, such as the external capsule, using DIC optics at 4-5x magnification difficult in OTCs. However, under higher magnification (60x), the majority of cultures will retain traces of white fibrous tracts which can be used to identify BLA borders (arrows, inset Fig.5E).

UNDERSTANDING RESULTS

The neuronal architecture of the amygdala in juvenile does not reach maturity until after the first postnatal month (Berdel et al., 1997, Morys et al., 1998, Berdel and Morys, 2000, Bouwmeester et al., 2002, Davila et al., 2008, Ehrlich et al., 2012, 2013). Therefore, for OTCs prepared from P14 rats to be considered a useful model of this brain region in development and in adulthood, it was critical to compare electrophysiological properties between principal neurons in acutely prepared juvenile (P14) and mature (P70) BLA principal neurons, and those from OTCs. The section below describes the comparisons we made, and indicates what to expect when the OTCs are implemented successfully.

Comparison of PN membrane properties between EP70 OTCs and ex vivo P14 and P70 BLA

These data were recently reported in some detail elsewhere (Michaelson et al., 2020), so comparisons will be summarized here; interested readers should consult the cited paper for more details. In brief, PNs in BLA OTCs mature in culture to develop electrophysiological attributes distinct from those of PNs in acute ex vivo BLA slices both from P14 and from P70 rats, which in turn differ from one another.

The resting membrane potential of PNs in EP70 OTCs is similar to that of P14 neurons, while the P70 BLA PNs rest at a significantly more negative potential than either. The resting membrane potential (RMP) and especially the membrane input resistance (R_m) of BLA OTC principal cells more closely resemble those of acute P70 than of acute P14 PNs (Figs. 6A, B), even though they still differ significantly from one another. The rheobase (here defined as the current required to depolarize a neuron from rest to action potential threshold) of the OTCs is far closer to that of the P70 than to that of the P14 ex vivo neurons (Figs. 6C,D), consistent with the higher R_m of the P14 neurons relative to the others. The action potential properties, in particular accommodation (reduction in firing rate during a sustained depolarization) and the afterhyperpolarization of OTC PNs are also closer in appearance to, though not identical with, those of the P70 rats than those of the P14 rats (Figs. 6D, E).

Figure 6: Passive and active membrane properties of PNs from BLA EP70 OTCs are closer to those in acute slices from P70 than P14 rats.

A. Comparison of resting membrane potential of PNs from acute ex vivo BLA slices from P14 and P70 rats and from EP70 OTC slices. P70 neurons RMPs were significantly more negative than either P14 or OTC cells, which did not differ significantly from each other. B. Input resistance (R_m) was significantly higher in P14 neurons than either P70 or OTC neurons, while R_m in P70 and OTC cells were still different but closer in value. C. Consistent with the input resistance comparisons in B, the rheobase for P70 and OTC neurons was considerably greater than that of the P14 neurons, while the P70 and OTC neurons were still different but much closer. D. Responses to current ramps of P14, P70 and OTC neurons. Action potentials in response to a ramp from 0 to 300 mV over 800 ms in P14, P70 and EP70 OTC neurons. The P14 neurons fire far more action potentials than either of the other neurons, and the overshoot decreases significantly, consistent with increasing inactivation of the Na⁺ channel. The P70 neuron fires only a few Aps, while the OTC cell fires 4. Note that the P70 and OTC cells have greater afterhyperpolarizations following each action potential, and that there is significant hyperpolarization after the end of the ramp with both P70 and OTC cells. E. Action potentials in response to current steps. There is a relatively high firing rate in the P14 and P70 neurons, with a more modest firing rate and pronounced accommodation is seen in the OTC cell. For all figures: *P< 0.05; **P<0.01, ***P< 0.001, ****P<0.0001.

Morphological comparison of EP70 BLA PNs with P14 and P70 BLA PNs

The dendritic tree of BLA PNs undergoes extensive remodeling in the first postnatal month (Ryan et al., 2014). Most of the critical changes in structure and function occur by P28 (Ehrlich et al., 2012, 2013, King et al., 2013, Ryan et al., 2014). Somewhat surprisingly, despite their relatively low membrane input resistance (R_m), the OTC PNs have the smallest whole cell capacitance (Fig. 7A). In healthy OTCs (EP 70), PN properties will differ relative to acute ex vivo P14 and P70 cells in the three preparations. Neurons from the three populations were filled with neurobiotin via the pipette after electrophysiological characterization, then slices were fixed, stained, cleared and optically sectioned by confocal microscopy (Silveira-Villarroel et al., 2018, Michaelson et al., 2020). Reagents used for immunocytochemistry illustrated below are provided in the legends to Figure 11. As depicted in Figure 7B, the shape of dendritic arbors of principal neurons from all three groups were broadly similar, (some are more pyramidal and some more stellate in form) but differed markedly in their overall extent. While PN size increases with age ex vivo (Fig. 7B), but cells from EP70 OTCs are clearly smaller than acute neurons from either age.

Figure 7. BLA OTCs PNs are physically more compact than either acute P14 or P70 PNs.

A. Whole cell membrane capacitance is lowest in OTC cells. B. Confocal reconstruction of neurons recorded, neurobiotin-filled, fixed and stained. Note the compact dendritic extent of OTC neurons relative to the acute neurons. C. Sholl analysis reveals significant differences in dendritic extent between OTCs and both P14 and P70 acute neurons. D. Total dendritic length is very different between all 3 types of neurons, with OTCs having the smallest. E. Total branch points (an index of dendritic tree complexity) are also significantly different between the 3 neuron types, with OTCs being the least branched and P70s the most.

Figure 11. Immunohistochemical characterization of BLA OTC explants and rat brain BLA using markers for excitatory and inhibitory neurons.

Panels A-H demonstrate single labeling for specific neuronal markers in OTC (A-D) and perfused rat BLA (E-H). Characteristic patterns for CaMKII and PV are noted in the OTC (I-K) which parallel that in rat BLA in situ (L-N). SOM and NPY are co-expressed within cell bodies and fibers in the OTCs (O-Q) and rat BLA (R-T). Arrows indicate double-labeled cells; larger arrowheads indicate single labeled SOM cells; smaller arrowheads note co-expression of SOM and NPY in fibers. Scale bars (H, K, N, T) = 30 μm. Free floating tissues were treated as described previously (Rostkowski et al., 2009) and processed for double labeling using the following antigens: HCN1 (1μg/mL; Neuromab #75-110; RRID:AB_2115181); CaMKII (pyramidal cell marker; 1:2000; MilliPore #MAB8699; RRID:AB_2067919); and markers for interneuronal populations: GAD (glutamic acid decarboxylase; 1:500; MilliPore); PV (parvalbumin; 1:5000, Swant; RRID:AB_10000344), SOM (somatostatin; 1:500; Millipore #AB345; RRID:AB_518503) and NPY (Neuropeptide Y; 1:2000; Peninsula T-4069.0400; RRID:AB_518503). After 48 hour incubations with primary antibodies, the appropriate signals were visualized using secondary antibodies (CY3-streptavidin; 1:250, Jackson ImmunoResearch) and AlexaFluor 488 (1:500, Molecular Probes). Immunoreactivity was visualized using confocal microscopy (FV10i, Olympus or Leica TCS SP8 with a 63x objective), and images were captured at 10x or 60x magnification (with additional 2x or 3x digital zoom respectively) and analyzed using Fluoview1000 (Olympus) software. Z-series confocal stacks (0.5 μm steps, 5 μm thickness) were generated for each fluorophore at every randomly sampled site from BLA in rat brain or OTC. Images were imported into Adobe Photoshop and brightness and contrast was adjusted

Quantification of total dendritic length and numbers of branch points (an index of dendritic complexity) differed between the populations studied (Michaelson et al., 2020, and Methods above). BLA OTC PNs at EP70 are considerably smaller than neurons either from P14 or P70 ex vivo slices (we did not analyze soma size due potential errors incurred during detaching the pipette from patched neurons). Overall, total dendritic lengths in OTC PNs were only about 42% of that of the P70 PNs and only about 65% that of the P14 PNs (Fig. 7C), consistent with the low capacitance values observed (Fig. 7A). The number of branch points increase significantly with age in ex vivo BLA while OTC PNs had fewer branches compared to both acute preparations (Fig. 7E). The distribution of intersections in the Sholl analysis (Fig. 7E) clearly reflects the differences in dendritic extent and complexity of the three populations. This substantial difference in size, especially between the OTCs and the P14 neurons coupled with the low R_m observed, indicates a considerably higher ion channel density in OTC PN plasma membranes, a potential consideration if planning cell-attached or excised-patch recordings from OTC neurons

H-current (I_h)

I_h is prominent in BLA PNs (Womble and Moises, 1993, Park et al., 2007), and increases steadily with age in rat BLA PNs during the entire first postnatal month (Ehrlich et al., 2012). I_h is modulated in opposing directions by NPY and CRF, both acutely ex vivo and with repeated treatments in vivo (Giesbrecht et al., 2010, Silveira Villarroel et al., 2018) and in vitro (Michaelson et al., 2020). Ex vivo P14 neurons already exhibit a robust I_h, which considerably greater in ex vivo P70 PNs (Fig. 8A). We found that peak I_h amplitudes in OTC PNs are intermediate between the P14 and P70 PNs (Fig. 8B). However, because of their much smaller size, I_h density (pA/pF) was significantly higher in OTC neurons compared with acute P14 cells, but were indistinguishable between EP70 OTC neurons and ex vivo P70 neurons (Fig. 8C). This suggests that normal developmental upregulation in I_h expression occurs in OTC neurons.

Figure 8: I_h in BLA OTCs develops similarly to I_h in vivo.

Family of superimposed current traces from V_h −55 mV to −135 in 10 mV increments in an EP70 OTC PN. A₂ Blue trace (to −135 mV) from A₁ shown superimposed with recordings of I_h in response to the same step in BLA PNs from acute ex vivo slices from P14 (grey trace) and P70 (black trace) rats. Note similar kinetics and current magnitude between the OTC and the P70 acute neuron, while I_h in less strongly expressed at P14. B. Current-voltage (I-V) relationships for I_h in P14, P70 and EP70 neurons. Numbers of cells as indicated in legend. Blue asterisks indicate difference between P14 and OTC, # indicate difference between P70 and P14, and black asterisks between OTC and P70. C. Comparison of I_h density (pA/pF) in these neurons. Note I_h density does not differ between EP70 and P70 neurons

Development of spontaneous inhibitory and excitatory postsynaptic currents

Parvalbumin-expressing GABAergic interneurons of the BLA do not reach mature levels until P25-30, while many excitatory inputs to BLA from thalamic and cortical regions are continually being refined until P26 (Berdel and Morys, 2000, Bouwmeester et al., 2002). Consistent with this, the numbers of synaptic contacts nearly triple in BLA during this period (Berdel et al., 1997, Morys et al., 1998). We therefore next studied the synaptic connections formed with PNs in our cultures in relation to sIPSCs and sEPSCs recorded in the two ex vivo preparations.

Spontaneous postsynaptic currents (sPSCs) were examined by holding neurons at −55 mV in voltage clamp with a conventional K⁺ gluconate internal pipette solution (Silviera Villarroel et al., 2018, Michaelson et al., 2020). At that potential, GABA_A receptor-mediated, inhibitory (sIPSCs) are outward, and manifest as upward deflections in the current trace, while AMPA and NMDA receptor-mediated components of excitatory (sEPSCs) are seen as downward (inward current) deflections (Fig. 9A), allowing for the simultaneous analysis of sEPSC and sIPSC activity in a given neuron. In such recordings, the mean sIPSC amplitude for EP70 OTC principal neurons was significantly greater than that seen in neurons from either P14 or P70 PNs (Fig. 9B), consistent with the greater electrical compactness of the OTC neurons. However, the sIPSC frequencies (expressed as inter-event intervals – IEI) did not significantly differ between OTC principal neurons and those from the P70 PNs, while sIPSC frequencies in P14 cells were lower than in either P70 or OTC neurons (Fig. 9C). Similarly, sEPSCs recorded in EP70 OTC PNs had sharply larger amplitudes than seen in P14 and P70 neurons (Fig. 9D), also consistent with the compact electrotonic structure of the OTC cells. As with sIPSCs, the mean frequency of sEPSCs was similar between EP70 and P70 PNs, while it was considerably lower in P14 neurons (Fig. 9E). Overall, this suggests that the EP70 PNs receive generally similar levels of synaptic input to P70 neurons.

Figure 9: Synaptic event amplitudes are larger in EP70 than in P70 PNs, but frequencies are similar.

A. Representative traces of spontaneous synaptic events in PNs from acute P14 and P70 slices and EP70 OTC. Vh = −55 mV so spontaneous IPSCs (sIPSC - upward deflections) and spontaneous EPSC’s (sEPSC - downward deflections) can be observed simultaneously (Michaelson et al, 2020, Giesbrecht et al, 2010, Silveira Villarroel et al, 2018). B sIPSC amplitudes are similar in P14 and P70 PNs, but are less than half that seen in OTC neurons. C. sIPSC intervals are considerably longer in the P14 neurons than in either P70 or OTC PNs. D. Similarly, sEPSC amplitudes in OTCs are double those in P14 and P70 PNs, while E. sEPSC frequencies are considerably greater both in P70 and OTC PNs than in P14 neurons. These suggest that the synaptic inputs of OTCs are equivalent in activity with the P70 BLA but the smaller OTC neurons are electrically far more compact.

A somewhat unexpected finding was the occurrence of compound postsynaptic bursting activity within the BLA of OTC slices, which was completely absent from acutely prepared slices (Figs. 10A-C). OTC slices generated bursts at a frequency of 0.23 ± 0.02 Hz (Fig. 10C), while none were observed in either group of ex vivo slices. Compound bursts (Fig. 10D) are consistent with synchronous activity in the neural network, requiring both GABA_A receptor activation in projection cells and glutamatergic input to GABAergic interneurons and has been postulated to facilitate emotional states within the BLA (Rainnie, 1999, Chung and Moore, 2009). As an aside, this activity can sometimes interfere with some measurements in particularly active slices, but can be readily blocked with antagonists of ionotropic glutamate and GABA receptors if needed.

Figure 10: Compound postsynaptic bursts occur in principal neurons from BLA OTCs. A-C.

Continuous traces of activity (120 s duration) in representative neurons from P14, P70 and EP70 OTC neurons held in voltage clamp at −55 mV. OTC PNs are noticably more active, and undergo periodic episodes of spontaneous bursting activity. Trace in C truncated at the bottom for clarity. D. Burst highlighted in red in trace C is shown at full amplitude (same scale as in C) and at an expanded timescale (indicated in bar above). Note the barrages of sIPSCs which precede the more prolonged barrage of sEPSCs, which occurs at each burst event.

Immunohistochemical characterization of OTCs

In addition to the electrophysiological data demonstrating appropriate properties of acute and OTC PNs, we used immunohistochemistry (Rostkowski et al., 2009) to assess whether the OTCs exhibited qualitative profiles of interneuronal markers consistent with the appropriate development of BLA circuitry (McDonald, 2020). Due to the overall increases in cell densities relative to normal brain tissue caused by the inherent thinning of the OTCs, quantitative assessment of neuronal populations exhibiting the different interneuronal markers as done previously (Rostkowski et al., 2009) is problematic. CaMKII immunoreactivity (CaMKII-ir), a marker for BLA glutamatergic neurons, is robustly evident within OTCs, labeling cells that are typical for glutamatergic principal neurons (Fig. 11 A, E). The diameter of these cell bodies (15-25 μm) is typical of that for BLA PNs in vivo. Additionally, CaMKII-ir is distributed within the neuropil which is associated with staining of dendrites and spines. Consistent with the expression of a robust I_h, OTC PNs express HCN1 channel immunoreactivity just as can be seen in adult rat brain, though the staining pattern in cells appears lighter (Figs. 11 D,H).

GABAergic interneurons provide high levels of inhibitory tone to BLA PNs (Ehrlich et al., 2012, Mackay et al., 2019). Immunoreactive glutamic acid dehydrogenase (GAD-ir) a marker of GABA synthesis, is present throughout the OTCs, and displays a diffuse staining pattern in fibers both in the adult rat BLA and the OTCs. While no GAD-ir positive cell bodies are noted in OTCs, this is similar to GAD-ir seen in freshly preserved BLA tissue. Using other neuropeptide and calcium binding protein markers provides a more detailed description of the anatomy of BLA interneurons. Parvalbumin (PV) interneurons represent 40% of the interneuronal population of the BLA and provide strong innervation to glutamatergic neurons as well as forming synapses with other PV neurons (Muller et al., 2005; 2006; Rostkowski et al., 2009). Numerous PV-ir cells are scattered throughout the OTCs, and there are numerous discrete PV-ir punctae associated with the formation of peri-cellular baskets around CaMKII-ir somata (Figure 11 I-N). Of interest to our research is the presence of NPY and SOM populations of interneurons. Within the BLA, NPY and SOM provide significant innervation to PNs which is inhibitory in nature. Immunoreactivity for both of these peptides is present in cells and noted fibers (‘string of beads’ appearance) within the OTCs. We also note that there is extensive co-expression of NPY with SOM, yet single-labeled SOM neurons are also detected which recapitulates the anatomical findings from perfused rat brain (Rostkowski et al., 2009, McDonald, 2020).

These results are consistent with the idea that the BLA OTCs prepared, using the protocols we describe, closely adhere to the properties of their postnatal age equivalents, with a few notable exceptions, including the increased cellular density and relatively compact size, and the significantly increased ion channel density.

Slicing chamber design

To help reduce the volumes of solutions often required in vibrating microtomes, we adopted a homemade design from the Saggau laboratory (Qian et al., 1997). This has the advantages of being inexpensive and versatile, and is light and small enough to permit relatively simple mounting of brain blocks in the desired orientation (Fig 12). In a vibrating slicer with a large specimen chamber such as the TPI Vibratome, the specimen chamber can be filled with ice to help maintain a stable cold temperature in the small slicing chamber, adding to the viability of slices.

Figure 12. Slicing chamber for OTC preparation.

Compare with photograph in Figure 3A. Dimensions of the chamber used are given in mm; elevations from the rear (top profile) and left (left profile) surround overhead view (center), perspective drawing from above front shown below. All components are Plexiglas acrylic of dimensions shown, except for a well bottom (right) formed by gluing a conventional glass microscope slide beneath the wide slot at the bottom of the chamber which has been milled out completely. The slide permits a clearer inspection of slice quality. This version is designed to be clamped in the specimen vise of a standard TPI 1000 or 1500 vibratome. This allows crushed ice to be placed the large specimen bath of the vibratome to help keep the saline cold, while slicing solution is restricted to the slicing chamber itself, saving on reagents and keeping slices close to the operator for ready inspection and manipulation. Additional modifications to the dimensions of the chamber to accommodate different types of vibrating microtomes can readily be undertaken.