Protocol for electrophysiological measurements of circadian changes in excitability in dentate granule cells from adult mice

Summary

Many types of neurons exhibit a daily rhythm of intrinsic excitability. Here, we present a protocol for assessing circadian regulation of dentate granule cell excitability using a mouse model for conditional knockout of the molecular clock protein BMAL1. We describe steps for obtaining healthy oblique horizontal slices that contain the hippocampus and measuring intrinsic excitability and synaptic potentials by combining whole-cell patch-clamp recordings and perforant-path electric stimulation. We then detail procedures for validating single-cell genetic deletion of Bmal1 by immunohistochemistry.

For complete details on the use and execution of this protocol, please refer to Gonzalez et al.¹

Graphical abstract

Highlights

• •Strategy to obtain a model to study circadian physiology of the dentate gyrus
• •Steps for obtaining oblique horizontal brain slices containing the hippocampus
• •Steps for immunohistochemical validation of molecular clock conditional knockout
• •Adaptable for other brain areas and cell types

Publisher’s note: Undertaking any experimental protocol requires adherence to local institutional guidelines for laboratory safety and ethics.

Many types of neurons exhibit a daily rhythm of intrinsic excitability. Here, we present a protocol for assessing circadian regulation of dentate granule cell excitability using a mouse model for conditional knockout of the molecular clock protein BMAL1. We describe steps for obtaining healthy oblique horizontal slices that contain the hippocampus and measuring intrinsic excitability and synaptic potentials by combining whole-cell patch-clamp recordings and perforant-path electric stimulation. We then detail procedures for validating single-cell genetic deletion of Bmal1 by immunohistochemistry.

Before you begin

The dentate gyrus is the main entry point for neural activity into the hippocampus, integrating sensory and spatial information from the cortex in a manner that generates a neural representation of a context. A hallmark of the dentate gyrus is sparse neural activity wherein only a small fraction of the principal granule cells is active at any given time and active granule cells have a generally low firing rate. Sparse activity allows the dentate gyrus to serve as a “gate” that filters incoming information from the entorhinal cortex with a circadian regulation.² The dentate gate is controlled by a strong synaptic inhibition from local GABAergic interneurons^3,4 and intrinsic properties of the granule cells.⁵

This protocol describes a horizontal slice preparation for mouse dentate gyrus to assess daily oscillations in granule cell excitability. We detail step-by-step procedures for electrophysiological recordings of synaptic or current-elicited action potentials, or G-protein inward rectifying potassium (GIRK) and sodium leak (NaLCN) channels currents. Our protocol also provides information for immunohistochemical detection of BMAL1 to validate a cell-type specific approach for conditional knockout (cKO) of Bmal1 to disrupt the molecular clock in granule cells. This strategy allows us to record from granule cells that are wild type or cKO for Bmal1 within the same slice, using tdTomato expression as a reporter of Cre activity.

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All procedures were approved by the University of Alabama at Birmingham Institutional Care and use Committee (IACUC) in accordance with the US National Institute of Health Guide for the Care and Use of Laboratory Animals.

Breed adult Pomc-Cre:Bmal1^lox/lox:Ai14^H/H mice to disrupt the molecular clock in dentate granule cells

Timing: 12–15 weeks, for breeding and 8–12 weeks of development

The objective of this breeding strategy is to generate a mouse model to study the role of the molecular clock in the physiology of the dentate gyrus. The strategy involves conditional deletion of Bmal1, a core clock gene, in the principal dentate neurons, the granule cells (GCs).

Cross Pomc-Cre mice (Jackson #:010714; on a C57Bl/6J background)⁶ with Bmal1^lox mice (Jackson # 007668)⁷ to generate Pomc-Cre⁺:Bmal1^lox/lox mice. This pro-opiomelanocortin-alpha (Pomc) mouse line drives expression of Cre recombinase primarily in granule cells of the dentate gyrus, with efficiency of Cre targeting reported as 60%. Bmal1^lox have loxP sites flanking the Bmal1 gene, allowing for its conditional knockout in the presence of Cre recombinase. This mouse is a tool to study the role of the local circadian clock in physiological and behavioral regulation. Cross the resultant mice with Bmal1^lox/lox; Ai14^H/H reporter mice (Jackson #007914)⁸ to obtain Pomc-Cre⁺:Bmal1^lox/lox:Ai14^H/H (Bmal1 cKO) and Pomc-Cre⁻:Bmal1^lox/lox:Ai14^H/H (control) mice to use in experiments. The Ai14 reporter mouse carries a loxP-flanked STOP cassette preventing transcription of a red fluorescence protein (tdTomato). This strategy allows visualization of cell-autonomous molecular clock disruption since both Bmal1 wild type cells (tdTomato negative) and Bmal1 conditional knockout cells (tdTomato positive) coexist within the same slice. To control for potential effects of Cre expression, we also crossed Pomc-Cre mice with Ai14^H/H reporter mice (Pomc-Cre⁺:Bmal1^+/+:Ai14^H/H).

House the mice in standard cages with ad libitum access to food and water and maintain them in 12:12 light/dark (LD) cycle. Zeitgeber Time (ZT) 0 refers to lights on.

In order to confirm that differences in results obtained at different times during the light cycle are due to the molecular clock and not the light cycle, it is necessary to repeat experiments using mice housed in constant darkness (DD). For that purpose, house mice individually in cages equipped with running wheels in LD for at least 15 days before being released into DD for at least 21 days. Use circadian cabinets to control light regimen. Record wheel-running activity using ClockLab software and generate actograms using 6-min bins of activity and double plot for ease of examination.⁹ During DD, circadian time (CT) 12 equals activity onset and the beginning of subjective night.

Preparation of stock solutions for electrophysiology and immunohistochemistry

Timing: 2 h, one or few days prior to experiment

The amounts prepared are enough for several days of electrophysiological experiments.

For 2L (10× Artificial Cerebrospinal Fluid (ACSF) solution)

Salt	Final 1× [mM]	g/2 L (10×)
NaCl	125	146.1
KCl	2.5	3.7
NaH2PO4	1.25	3.4
CaCl2	2	5.9
MgCl2	1	4.1
ddH2O	-	Fill up to 2 L

Note: Store at 22 ± 2°C.

For 1L (10× cutting solution)

Salt	Final 1× [mM]	g/1 L (10×)
Choline·Cl	110	153.6
KCl	2.5	1.9
NaH2PO4	1.25	1.7
CaCl2	0.5	0.7
MgCl2	7	14.2
ddH2O	-	Fill up to 1 L

Note: Store at 4°C.

For 1 L (10× Phosphate Buffered Saline)

• 1.For 1 L add to 800 mL of ddH₂O.

Salt	G
NaCl	80
KCl	2
Na2HPO4	14.2
KH2PO4	2.7

• 2.Adjust pH to 7.4 with 10 M NaOH.

Caution: Follow laboratory safety guidelines. Avoid inhalation and contact. Wear appropriate personal protective equipment.

• 3.Adjust volume to 1000 mL with additional ddH₂O.

Note: Store at 22 ± 2°C.

For 1 L (1× Phosphate Buffered Saline)

• 4.Add 100 mL of 10× PBS to 850 mL of ddH₂O.
• 5.Adjust pH to 7.4 with 10 M NaOH.
• 6.Adjust volume to 1 L.

Note: Store at 4°C for up to 1 month.

For 250 mL (4% PFA)

• 7.Under a hood, add 10 g of paraformaldehyde to 200 mL of ddH₂O in a beaker.

Caution: Follow laboratory safety guidelines. Avoid inhalation and contact. Wear appropriate personal protective equipment.

• 8.Heat to about 55°C while stirring.

CRITICAL: Do not heat over 65°C.

• 9.When solution is 55°C, add drops of 10 M NaOH till it clears.
• 10.Add 25 mL of 10× PBS and let it cool down.
• 11.Adjust pH to 7.2 with 10 M HCl.
• 12.Add ddH₂O to 250 mL and filter using grade 4 Whatman filter paper.
• 13.Store at 4°C for up to two weeks.

CRITICAL: Wrap in foil to protect from light.

Tribromoethanol Stock

• 14.Add 25 g of 2,2,2,tribromoethanol (Avertin) in 15.5 mL of tert-amyl alcohol.
• 15.Stir using a magnetic stirrer for 12 h at 22 ± 2°C, protected from light using foil.
• 16.Store in foil covered scintillation vial in desiccator.
• 17.Stock can be stored indefinitely.

Note: Any alternative small glass bottles with cap can be used.

For 500 mL anti-freeze solution

• 18.
  Prepare Phosphate Buffer 0.05 M.

  • a.
    Prepare Solution A (0.2 M Sodium Phosphate Monobasic).

    • i.
      Add 13.79 g of NaH₂PO₄·H₂O in 500 mL of ddH₂O.
  • b.
    Prepare Solution B (0.2 Sodium Phosphate dibasic).

    • i.
      Add 14.19 g of Na₂HPO₄ in 500 mL of ddH₂O.
  • c.
    Add 46 mL of Solution A + 154 mL of Solution B in 600 mL of ddH₂O to a total of 800 mL.
  • d.
    Adjust pH to 7.4.
• 19.Add 250 mL of Phosphate Buffer 0.05 M + 150 mL of ethylene glycol + 100 mL of glycerol.
• 20.Stir for a few minutes until the solution is homogeneous.

Note: Store at −20°C for up to one year.

Blocking buffer for PROX1 immunohistochemistry (IHC)

Reagent	Final concentration	Amount for 10 mL
Triton X-100	0.4%	40 μL
Bovine serum albumin	3%	0.3 g
Normal Donkey Serum	10%	1 mL
Glycine	1%	0.1 mg
1× PBS	-	8.96 mL
Total	-	10 mL

Note: Dissolve BSA and glycine in PBS before adding Triton X-100. Blocking buffer without normal donkey serum can be made ahead and stored for up to 1 week. Prepare appropriate amount so that each brain slice will be able to receive at least 0.5 mL of blocking buffer.

Blocking buffer for BMAL1 IHC

Reagent	Final concentration	Amount for 10 mL
Triton X-100	0.3%	30 μL
Bovine serum albumin	5%	0.5 g
1× PBS	-	9.97 mL
Total	-	10 mL

Note: Prepare appropriate amount so that each brain slice will be able to receive at least 0.5 mL of blocking buffer.

Antibody buffer for BMAL1 IHC

Reagent	Final concentration	Amount for 10 mL
Triton X-100	0.3%	30 μL
Bovine serum albumin	1%	0.1 g
1× PBS	-	9.97 mL
Total	-	10 mL

Note: Prepare appropriate amount so that each brain slice will be able to receive at least 0.5 mL of blocking buffer.

Chloride a silver wire electrode and check the ground electrode

Timing: 30 min, one day before recording

The recording electrode is constructed using a silver wire that requires regular chloriding particularly if the chloride coating is diminishing. This becomes noticeable when the uniformity of the gray coating is disrupted, resulting in shiny silver areas appearing on the surface of the electrode.

• 21.Solder a wire finished in two crocodile clips to the positive pole and a wire finished in a resistor to the negative pole of a 9 V battery (Figure 1A).
• 22.Connect the crocodile clips to a recording electrode and to a silver wire spring of about 10 turns over 1 cm (ground electrode).
• 23.Submerge both silver wires and the resistor in a 5% NaCl solution for about 5 min.

CRITICAL: Hydrogen bubbles must be produced due to the reduction of H⁺ in the solution.

Figure 1.

Experimental setups

(A) Chloride station. (1) 9 V battery. (2) Crocodile clip. (3) Resistor. (4) 5% NaCl solution.

(B) (1) Staining jar and (2) lid after rubber coating. (3) In-house designed multi-well platform with nylon mesh.

Prepare the chamber to store slices with fluorophores (GFP, tdTomato)

• 24.Cover the outer surface of the staining jar and the glass lid with a multi-purpose rubber coating to prevent fluorescence bleaching (Figure 1B).
• 25.Recoat as many times as needed to make it dark.

CRITICAL: Preventing photo-bleaching is crucial.

Alternatives: Cover it with foil; however, rubber coating is permanent.

Key resources table

REAGENT or RESOURCE	SOURCE	IDENTIFIER
Antibodies
Rabbit anti-BMAL1 1:2,000	Novus Biologicals	Novus Cat# NB100-2288; RRID: AB_10000794
Goat anti-rabbit Alexa Fluor 647 1:1,000	Invitrogen	Thermo Fisher Cat#A21244; RRID: AB_143165
Donkey anti-goat Alexa Fluor 647 1:500	Invitrogen	Thermo Fisher Cat#A21447; RRID: AB_2535864
Goat anti h-PROX1 1:500	R&D Systems	R&D Systems Cat# AF2727; RRID: AB_2170716
Chemicals, peptides, and recombinant proteins
NaCl	Thermo Fisher Scientific	CAS 7647-14-5, S271-3
KCl	Sigma-Aldrich	CAS 7447-40-7, Cat#60128-250G-F
NaH2PO4	Thermo Fisher Scientific	CAS 10049-21-5, S369-1
CaCl2	Thermo Fisher Scientific	CAS 10035-04-8, C70-500
MgCl2	Thermo Fisher Scientific	CAS 7791-18-6, H35-500
D-(+)-glucose	Sigma-Aldrich	CAS 50-99-7, G7021-1KG
NaHCO3	Thermo Fisher Scientific	CAS 144-55-8, S233-500
Na-ascorbate	Sigma-Aldrich	CAS 134-03-2, A4034-100G
Na-pyruvate	Sigma-Aldrich	CAS 113-24-6, P2256-100G
Choline·Cl	Thermo Fisher Scientific	CAS 67-48-1, Cat#110290010
K-gluconate	Sigma-Aldrich	CAS 299-27-4, G4500-100G
HEPES	Sigma-Aldrich	CAS 75277-39-3, H7006-100G
EGTA	Sigma-Aldrich	CAS 67-42-5, O3777-10G
Phosphocreatine	Sigma-Aldrich	CAS 19333-65-4, P7936-1G
Mg-ATP	Sigma-Aldrich	CAS 74804-12-9, A9187-1G
Na-GTP	Sigma-Aldrich	CAS 36051-31-7, G8877-250MG
CsMeSO3	Sigma-Aldrich	CAS 2550-61-0, C1426 5G
Na2HPO4	Thermo Fisher Scientific	CAS 7558-79-4, BP332-500
NaH2PO4	Thermo Fisher Scientific	CAS 10049-21-5, S369-1
KH2PO4	Thermo Fisher Scientific	CAS 7778-77-0, BP362-500
BaCl2	Thermo Fisher Scientific	CAS 10361-37-2, B31-100C
CsCl2	Sigma-Aldrich	CAS 7647-17-8, 289329-25G
CsOH	Sigma-Aldrich	CAS 21351-79-1, 232041-10G
N-methyl-D-glutamine (NMDG)	Sigma-Aldrich	CAS 6284-40-8, M2004-1KG
Paraformaldehyde (PFA)	Thermo Fisher Scientific	CAS 30525-89-4, O4042-500
2,2,2, tribromoethanol (Avertin)	Sigma-Aldrich	CAS 75-80-9, T48402
Agar	Thermo Fisher Scientific	CAS 9002-18-0, BP1423-500
2-Methyl-2-butanol (tert-amyl alcohol)	Sigma-Aldrich	CAS 75-85-4, Cat#240486-100ML
Ethylene glycol	Sigma-Aldrich	CAS 107-21-1, Cat#102466-1L
Glycerol	Thermo Fisher Scientific	CAS 56-81-5, G31-1L
Normal donkey serum	Sigma-Aldrich	NACRES: NA.46, D9663
Bovine serum albumin (BSA)	Thermo Fisher Scientific	CAS 9048-46-8, BP1600-100
Triton X-100	Thermo Fisher Scientific	CAS 9002-93-1, BP151-100
Prolong Gold	Thermo Fisher Scientific	P36930
Isoflurane	VetOne	NDC 13985-528-60, V1 502017-250ML
SR95531 (Gabazine)	Abcam	CAS 104104-50-9, ab120042
CGP55845	Abcam	CAS 149184-22-5, ab120337
ML297	Sigma-Aldrich	CAS 1443246-62-5, SML0836
L-703,606	Sigma-Aldrich	CAS 351351-06-9, L119
NBQX	Abcam	CAS 118676-58-7, ab120045
(R)-CPP	Abcam	CAS 126453-07-4, ab120159
Tetrodotoxin citrate	Abcam	CAS 4368-28-9, ab120054
Picrotoxin	Abcam	CAS 124-87-8, ab120315
Apamin	Tocris	CAS 24345-16-2, Cat#178270
Experimental models: Organisms/strains
Mouse/C57BL/6J	JAX	Jackson#000664; RRID: IMSR_JAX:000664
Mouse/Ai14	JAX	Jackson#007914; RRID: IMSR_JAX:007914
Mouse/Pomc-Cre	JAX	Jackson#010714; RRID: IMSR_JAX:010714
Mouse/Bmal1 lox	JAX	Jackson#007668; RRID: IMSR_JAX:007668
Software and algorithms
PClamp	Molecular Devices	v.10.7.0.3
Clampfit	Molecular Devices	v.10.7.0.3
AxoGraph X	AxoGraph Scientific	v.1.7.6
Prism	GraphPad Software	v.9.2.0
SPSS	IBM	v.25
Fiji	imagej.net/software/fiji/	2.15.0
ClockLab software	Actimetrics	v.6
Others
MultiClamp 700A/B	Axon Instruments	N/A
Digidata digitizer	Axon Instruments	Model 1440A
Epifluorescence upright microscope	Olympus	BX51WI
Confocal microscope	Olympus	FV1200 + BX61WI
Fluoview software	Olympus	Ver.04.02
Camera controller	Hamamatsu	C2741
CCD camera	Sony	XC-ST50
Monitor	Pro Video	WM-1201B
Halogen power supply	Olympus	TH4-100
Fluorescence illumination system	Prior Scientific	Lumen 200
Filter set	Chroma	T540spuv-UF1
5× objective	Olympus	UPlanFI
60× objective	Olympus	LUMPlanFI
Constant current isolated stimulator	Digitimer Ltd.	DS3
Micromanipulator	Sutter Instrument	ROE-200
Filter cube	Semrock	TXRED-4040B-ZERO
Peristaltic flow pump	Rainin Dinamax	RP-1
Temperature controller	Warner Instruments	TC-324B
Pipette puller	Sutter Instrument	P-97
Borosilicate glass	Sutter Instrument	BF150-86-10
Syringe filter	Whatman	6789-0402
Pipette tips	Eppendorf	2028-09
Aquarium coarse air stones	Kordon	N/A
Vibratome (electrophysiology)	Leica Biosystems	VT1200S
Razor blades	Ted Pella	1221-4
Vibratome (IHC)	Pelco	3000
Water bath	Fisher Scientific	ISOTEMP 110
Spring scissors	Fine Science Tools	15000-00
Hardened scissors	Fine Science Tools	14090-09
Fine scissors	Fine Science Tools	14058-09
Surgical scissors	Fine Science Tools	14001-12
Forceps	Fine Science Tools	11252-00
Micro spatula	Ted Pella	13508
Scalpel handle	Fine Science Tools	10004-13
Scalpel blade	Fine Science Tools	10020-00
Winged infusion set	Terumo	SV∗25EL
Cotton swab	Puritan	867-WC
Needles	BD	305176
Insulin syringe	BD	329412
Superglue	Krazy Glue	-
Dissecting pan	Carolina	629004
Filter paper	Fisherbrand	09-801B
Glass staining jar with lid	Sigma-Aldrich	BR472200
Petri dish	Sigma-Aldrich	BR455751
Osmometer	Wescor	VAPRO 5520
Transfer pipettes	Fisherbrand	13-711-7M
Multi-purpose rubber coating	Performix	Plastic Dip
Kimwipes	Kimtech	-
Fume hood	Kewaunee Scientific	Air Supreme
Rotator	Barnstead	2309
Microscope slide	Fisherbrand	12-550-15
Microscope cover	Fisherbrand	12545M
Magnetic stirrer	Corning	PC 410-D
20 mL scintillation vial	Fisher Scientific	03-341-25L
Grade 4 filter paper	Whatman	1004-240
Carbogen cylinder (5% CO2/95% O2)	Airgas	UN3156
Circadian cabinets	Actimetrics	-
Running wheels	Coulbourn Instruments	CL300

Note: Use two-to-five-month old male and female mice for conducting experiments.

Alternatives: This protocol involves a Multiclamp 700A/B amplifier, pCLAMP and Clampex (Molecular Devices, CA, USA) software to record voltage- and ligand-gated currents as well as membrane potential, and a BX51WI Olympus microscope. Other amplifiers and programs can also be used for the recordings such as the EPC-10 patch-clamp amplifier and PATCHMASTER (HEKA Elektronik). Other upright microscopes are also suitable (Nikon, Leica, Zeiss, etc.).

Materials and equipment

For 50 mL standard physiological internal

Salt	Final [mM]	g/50 mL
K-gluconate	135	1.579
HEPES	10	0.119
EGTA	0.1	0.0019
KCl	3	0.0111
MgCl2	2	0.02
Phosphocreatine	10	0.127
Mg-ATP	2	0.05
Na-GTP	0.5	0.013
ddH2O	-	Fill up to 50 mL

Note: Stir solution with a magnetic stirrer. Adjust pH to 7.3 with 1 M KOH. Aliquot solution into small volumes (0.5–1 mL). Store at −20°C for up to one year.

CRITICAL: Limit freeze-thaw cycles to avoid reduction of stability of reagents.

Keep the solution over ice or in the refrigerator during patch clamp window.

Alternatives: Do not add Na-GTP for experiments without GTP (GTP^-).

For 50 mL Cs based internal

Salt	Final [mM]	g/50 mL
CsMeSO3	122	1.390
HEPES	9	0.1072
EGTA	0.45	0.0085
CaCl2	0.09	0.00066
MgCl2	1.8	0.0183
Phosphocreatine	14	0.1784
Mg-ATP	4	0.1014
Na-GTP	0.5	0.0131
ddH2O	-	Fill up to 50 mL

Note: Stir solution with a magnetic stirrer. Adjust pH to 7.3 with 1 M CsOH. Aliquot solution into small volumes (0.5–1 mL). Store at −20°C for up to one year.

CRITICAL: Limit freeze-thaw cycles to avoid reduction of stability of reagents.

Keep the solution over ice or in the refrigerator during recording.

For 1 L 1× cutting solution

Salt	Final [mM]	g/1 L
D-(+)-Glucose	25	4.5
NaHCO3	25	2.1
Na-ascorbate	1.3	0.2
Na-pyruvate	3	0.3
10× cutting solution	-	100 mL
ddH2O	-	Fill up to 1 L

Note: Stir solution with a magnetic stirrer. Store at 4°C for up to 3 days.

For 2 L 1× ACSF

Salt	Final [mM]	g/1 L
D-(+)-Glucose	25	9
NaHCO3	25	4.2
10× ACSF	-	200 mL
ddH2O	-	Fill up to 2 L

Note: Stir solution with a magnetic stirrer and measure the osmotic concentration using osmometer (follow manufacturer’s instructions). ACSF 1× should be discarded at the end of the recording day.

CRITICAL: The resulting osmolarity should be 290–300 mOsm, about 5 mOsm lower than any internal used to minimize osmotic perturbations.

For Na⁺ substitution experiments 1 L 1× NMDG solution

Salt	Final [mM]	g/1 L
NMDG	125	24.4
NaHCO3	25	2.1
NaH2PO4	1.25	0.2
ddH2O	-	Fill up to ∼800 mL

Note: Stir solution with a magnetic stirrer. Adjust pH to 7.3 with 12 N HCl.

Salt	Final [mM]	g/1 L
Glucose	25	4.5

• •Bubble with carbogen.

Salt	Final [mM]	g/1 L
CaCl2	2	0.3
MgCl2	1	0.2
ddH2O	-	Fill up to 1 L

For 1 L 1× PBS

• •Add 100 mL of 10× PBS to 850 mL of ddH₂O.
• •Adjust pH to 7.4.
• •Adjust volume to 1 L.

Note: Store at 4°C for up to 1 month.

For tribromoethanol (Avertin) working solution (2% in PBS)

• •Mix 0.5 mL of Avertin stock with 40 mL of 1× PBS.
• •Heat while stirring under hood to prevent precipitation until clear.
• •Cover with foil.

Note: Store at 4°C for up to 2 weeks.

For 4% agar

• •Add 8 g of Agar to 200 mL of ddH₂O.
• •Heat in microwave until all the agar melts.
• •Pour into petri dish and wrap with Parafilm.

Note: Store at 4°C for up to 1 month.

Step-by-step method details

Electrophysiological recordings

Acute brain slice preparation

Timing: ∼2 h

This step of the experimental protocol involves anaesthetizing the mouse for trans-cardial perfusion with ice-cold carbogenated cutting solution (5% CO₂ / 95% O₂) to obtain horizontal slices containing the ventral hippocampus (troubleshooting problem 1). In our experience trans-cardial perfusion significantly improves the viability of the slices and the survival of interneurons. The process of obtaining the slices typically takes between 15 to 20 min from the time of anaesthetizing the mouse. Anaesthetize animals at ZT 5.5 or ZT 11.5. This schedule avoids potential photic interference resulting from light exposure outside of the 12:12 cycle. In the original paper¹ slices were also prepared at ZT 0 and ZT 18. When cutting slices at ZT18 or from DD mice take extra precautions described in “optional” notes.

• 1.
  In a glass bottle take 250 mL of 1× cutting solution (stored at 4°C) and saturate with carbogen for 10 min.

  • a.
    Cool the solution by keeping it in −20°C for ∼50 min.

    • i.
      After 30 min shake the solution every 5 min until flash freezing occurs.
• 2.While cutting solution is bubbling, prepare 2 L 1× ACSF (see materials and equipment).
• 3.
  While the cutting solution is freezing, prepare bench material and dissection equipment.

  • a.
    Ensure all tools necessary for slice preparation are ready and organized for quick access (see Figure 2C).
  • b.
    Pre-warm at 37°C the storage chamber filled with 1× ACSF and bubble it with carbogen.

    Note: slices will be placed on a nylon mesh mounted over an in-house designed multi-well platform. The platform will be placed elevated over 4 small cups to facilitate ACSF circulation. Typically, slices are submerged 15 mm (Figure 2E).

  • c.
    Vibratome set up (Timing: 10 min). Example for Leica VT1200 S (Figures 2A and 2B).

    • i.
      Turn the vibratome on.
    • ii.
      Using a cotton swab gently wipe the razor blade with acetone to remove oil or other adhesives.

      CRITICAL: For safety wear gloves, do not touch the sharp side of the blade.

    • iii.
      Insert the razor blade into the blade holder, fasten securely with hex-key Allen screwdriver and adjust the angle to the right line. This clearance angle mark represents −21°.
    • iv.
      Check the position of the blade using the VibroCheck (follow the operating instructions in VT1200S manual, pages 37–40).
    • v.
      Set the thickness to 300 μm with 1 mm of amplitude and the sectioning speed to 0.1 mm/s.
    • vi.
      Install the ice bath and buffer tray.
    • vii.
      Fill the ice bath with crushed ice.
    • viii.
      Over a bucket with crushed ice, place a glass petri-dish with a Whatman round filter paper.
    • ix.
      Connect a source of carbogen with the buffer tray and the petri-dish using an aquarium air curtain (recommended) or any other connector (a small micro-filter-candle, a needle, a yellow pipette tip).
  • d.
    Fill the buffer tray and petri-dish with ice-cold cutting solution and start bubbling with carbogen.
  • e.
    Load a 1 mL insulin syringe with 0.7–0.8 mL of Avertin.
  • f.
    Load a 30 mL syringe with 20 mL of ice-cold carbogenated cutting solution.

    • i.
      Attach a winged infusion set.
    • ii.
      Purge the air by pressing the plunger.
    • iii.
      Place the loaded syringe in a dissecting pan with vinyl dissecting pad.
• 4.
  Trans-cardial perfusion with ice-cold cutting solution.

  • a.
    Anesthetize 10–14 weeks-old mouse.

    • i.
      Place the mouse into a isoflurane sealed plexiglass container connected with tubing to a second container for isofluorane that is also connected to a source of air, all placed in a fume hood.
    • ii.
      Volatilize isoflurane to anesthetize the mouse.
    • iii.
      When breathing slows down (below one breath per second), deeply anesthetize the mouse with Avertin by intraperitoneal injection.

      Note: Confirm deep anesthesia by loss of paw pinch reflex.

      Optional: For experiments at ZT 18 or DD handle the mouse under dim light (< 10 lux / red light), and once deeply anaesthetized cover the eyes of the mouse with the thumb of a black nitrile glove to avoid light interference. To facilitate breathing, cut the tip of the glove with a scissor to stick out the nose.

  • b.
    Place the mouse on its back on a dissecting pan and fix the forelegs to the pad using pins or adhesive tapes.
  • c.
    Lift the abdominal skin using forceps and trim from abdomen to the rib cage using fine scissors.
  • d.
    Incise the abdominal muscle to expose the liver and the chest cavity.
  • e.
    Incise the diaphragm carefully and cut the ribs to fully expose the heart.

    Note: Fix the rib cage to the pad using a needle to orientate the heart and facilitate perfusion.

  • f.
    Trim connective tissue around the heart if needed.

    CRITICAL: Avoid damage to any organ.

  • g.
    Pierce the left ventricle with the winged infusion set and cut the right atrium with spring scissors to expel blood.
  • h.
    Press the plunger steadily to perfuse ice-cold carbogenated cutting solution.

    CRITICAL: Keep the needle in the left ventricle. Liver will turn pale yellow after a successful perfusion.

• 5.Decapitate the mouse with surgical scissors rostral to the cervicothoracic junction.
• 6.
  Harvest the brain (Timing: 1 min).

  • a.
    Remove the skin from the scalp.
  • b.
    Open the skull with a single sagittal cut from caudal to rostral from the foramen magnum along the middle line using hardened fine scissors.
  • c.
    Remove the parietal bone in the lateral direction using a forceps or a rongeur.

    Note: Additional cuts in the base of the parietal bone may facilitate this process.

    CRITICAL: Meningeal connections between brain and skull following bone removal can damage cortical structures. Avoid any compressive force.

  • d.
    Remove the brain using a long-round tip spatula, in the rostro-caudal direction cutting the cranial nerves, to scoop out the brain into the petri-dish prepared in step 3.

    CRITICAL: Step 6 should be carried out carefully within 1 min to improve quality of slices.

    Pause point: rest the brain in cold ACSF for 1 min to cool down.

• 7.
  “Dentate gyrus cut”¹⁰ (Figure 3).

  • a.
    Orientate the brain with the ventral plane down and the dorsal plane up.
  • b.
    Cut and remove the cerebellum using a scalpel.
  • c.
    Make a sagittal cut through the corpus callosum to separate both hemispheres.
  • d.
    Proceed with the “Dentate gyrus cut”.

    • i.
      Lay the hemisphere down on the sagittal plane (i.e., the midline cut surface) with the temporal surface up (1^st flip; arrow in Figure 3B, also shown in Figure 3C after the flip).
    • ii.
      Cut ∼ 10% off from the dorsal surface using a scalpel held at two angles: α angle 10° (Figure 3D); β angle 20° (Figure 3E). Note that the angles are exaggerated in Figures 3D and 3E.
    • iii.
      Place the hemisphere on the cut surface with the ventral surface up (2^nd flip). This will leave the hemisphere it the proper orientation for mounting on the specimen plate.

Note: Repeat step d with the other hemisphere.

CRITICAL: Keep in mind the orientation of the brain hemispheres. The angle of the cut can be varied to optimize the slice for each hippocampal subfield.

• 8.
  Mount the brain in the specimen plate (Figure 2D).

  • a.
    With the screw hole of the specimen plate close to the researcher, coat the left side of the specimen plate in an angle of 45° from right to left with a thin layer of cyanoacrylate glue (∼ 1 cm²) using the brush of the glue applicator.
  • b.
    Rotate the specimen plate 180°. Now the screw hole will be far from the researcher.

    • i.
      Using a fold spatula in your right hand rest the cutting surface of the left hemisphere on the base of the spatula and dry it by touching it on a Kimtech tissue.
    • ii.
      Hover the left hemisphere over the glued surface and with the help of a spatula in the left hand (placed on the interhemispheric sagittal cut), gently slide the hemisphere onto the glue by moving the right hand (Figure 3F).

      Note: Left-handed researchers might need a mirrored adjustment to get same results

      CRITICAL: Do not move your left hand or the tissue will flip onto the glue surface in an incorrect position (Figure 3F).

  • c.
    Turn 180° the specimen plate. Now the screw hole will be close to the researcher again.
  • d.
    Coat the right side of the specimen plate in an angle of 45° from left to right with a thin layer of cyanoacrylate glue (∼ 1 cm²).

    • i.
      Repeat step 8.b.i. and 8.b.ii.
  • e.
    With the help of a screwdriver place the specimen plate into the buffer tray keeping the V-shape of the tissue with the cortical surface facing the blade (Figure 2D).
• 9.Raise the platform of the vibratome until the cutting solution reaches the blade.
• 10.Set the cutting window, pressing 1^st and 2^nd cutting window edge buttons, covering the whole hemispheres.
• 11.Raise the platform until the blade is at the brain surface. Usually, the first two slices are discarded or raise the platform (∼ 600 μm).
• 12.Start slicing, from rostral-lateral to caudal-medial.
• 13.Help cutting with a paintbrush n°2 and transfer the slices using an open-ended plastic Pasteur pipette into the storage chamber at 37°C.
• 14.After 30 min at 37°C remove storage chamber from the bath and keep it at 22 ± 2°C.

Note: Slices will be ready to use after 30 min at 22 ± 2°C, however for this protocol we wait an extra hour to use them.

Figure 2.

Preparations to obtain horizontal brain slices

(A) Picture of the vibratome set.

(B) Amplification of the cutting head stage to show correct angle (1).

(C) Tools. (1) surgical scissors, (2) fine scissors, (3) hardened scissors, (4) spring scissors, (5) forceps, (6) scalpel, (7) scalpel blade, (8) spatula, (9) folded spatula, (10) paintbrush, (11) cotton swab, (12) insulin syringe, (13) Petri dish, (14) winged infusion set.

(D) Specimen platform with the brain hemispheres orientated in a “V” shape and glued on the “dentate gyrus cut”. Note that each hemisphere is glued on the dorsal cut surface so that the orientation of the slices are horizontal, with slight modifications determined by the α and β angles (see Figure 3).

(E) Example of the horizontal slices stored in a multi-well plate with nylon mesh bottom that is placed inside a darkened glass slide box with ACSF that is continuously bubbled with carbogen.

Figure 3.

Schematic depicting the steps for the "dentate gyrus cut"

(A) View of the brain from the top. The right top corner represents the spatial coordinates of the entire brain. The right bottom corner in green shows the hippocampal formation. Dashed lines represent trimming areas.

(B) Frontal view of the right hemisphere. Arrow represents the first flip to lay the hemisphere on the midline cut.

(C) Frontal view of the right hemisphere after the first flip. Red line indicates the angle of 10° (α) for the "dentate gyrus cut" and dark line represents the angle of 20° (β). This cut surface will be glued to the specimen platform after a second flip. Dashed lines in the right bottom corner represent the new “horizontal” plane after “dentate gyrus cut”.

(D) Picture showing the α angle.

(E) Picture showing the β angle.

(F) Cartoon showing maneuver detailed in step 8.b.ii. 3D representation of the projections in A-C with the 3D Brain Explorer (https://mouse.brain-map.org/static/brainexplorer).^14.

Patch-clamp electrophysiology

Timing: 3 h

The main goal of the experiment is to compare excitability and characterize the intrinsic properties of the dentate gyrus granule cells at different times during the circadian cycle, under different pharmacological conditions and with cKO of Bmal1 (troubleshooting problem 2). For this purpose, the recording window will be limited to 3 h during the light (ZT 8–11) and dark (ZT 14–17) cycle. However, slices prepared under this protocol would be healthy for recordings up to 6 h.

Note: Perform experiments in the current-clamp mode, evoking action potentials by electric stimulation of the perforant path or eliciting action potentials by current injection. If needed, conduct experiments in the voltage-clamp mode to measure currents generated by specific G-protein regulated ion channels.¹¹

• 15.
  Preparation before recording.

  • a.
    Perfuse the chamber with carbogenated ACSF at a rate of 2 mL/min.

    Note: We use a preheater station to set the temperature to 30°C. The chamber solution is constantly exchanged by vacuum suction and outlet into a prebleached Erlenmeyer flask (5 L).

  • b.
    Prepare extra-beakers containing pharmacological agents, as required for the planned experiments (for example, gabazine 10 μM; CGP55845 10 μM; ML297 10 μM).
  • c.
    Thaw an aliquot of internal solution and load a 1 mL insulin injection syringe.
  • d.
    Connect the syringe to a pipette filter unit connected to a patch pipette filler.

    CRITICAL: Keep it on ice to minimize ATP and GTP autohydrolysis.

    Use clean filters to avoid clogged pipette tips.
  • e.
    Load another 1 mL insulin syringe with ACSF and connect it with a pipette filler.
  • f.
    Pull borosilicate glass electrodes using a two-step horizontal puller having adjusted the settings so that the resistance is 2–4 MΩ for patch pipettes and 1–2 MΩ for stimulation pipettes.

    CRITICAL: Place the pipettes in a closed container to protect them from airborne dust.

    Note: Parameters can be modified to reach a good balance between seal formation and a suitable series resistance after breaking membrane patch.

    Alternatives: A microforge can be used to heat the pipette tip after pulling; this will increase resistance and typically enhance seal formation.

  • g.
    Turn on the rig and configure the acquisition software.
• 16.Transfer a slice using an open-ended plastic transfer pipette (5 mL) into the recording chamber.

CRITICAL: Avoid air bubbles from touching the tissue to prevent surface damage.

• 17.
  Place a nylon thread-containing platinum harp on the slice.

  • a.
    Orient the slice to make sure nylon threads are not over the dentate gyrus and there is access to maneuver two pipettes.
• 18.
  Fill a 1–2 mΩ pipette with ACSF (stimulation pipette).

  • a.
    Insert it in the stimulator holder
  • b.
    Place it adjacent to the hippocampal fissure near the crest (See Figure 4A), between the suprapyramidal blade and apex of the granule cell layer (defined as the midpoint between the suprapyramidal and infrapyramidal blades).

Note: That location allows to stimulate both lateral and medial perforant path and minimize direct stimulation of local interneurons.

• 19.Using a 60× objective, visualize both dentate gyrus blades to find the area with packed, rounded, healthy granule cells. See Figure 4C.

Note: We visualize granule cells expressing tdTomato using epifluorescence illumination and a Texas Red filter set. In the absence of fluorescent labels, we target granule cells in the middle of the granule cell layer to avoid immature adult-born granule cells and semilunar granule cells.

CRITICAL: Adult-born granule cells and semilunar granule cells can be identified by distinct intrinsic excitable properties and dendritic morphology.¹² Semilunar granule cells present a low IR (bellow 200 MΩ) while adult-born granule cells exhibit a high IR (over 500 MΩ) compared to mature granule (200–500 MΩ).

• 20.
  Fill a 2–4 mΩ pipette with internal solution.

  • a.
    Insert it into the electrode holder.
  • b.
    Fasten the pipette tightly with the O-ring gasket.

Note: The chloride electrode must be submerged in the internal solution. This electrode is connected through a gold pin to the head stage of the amplifier.

• 21.
  Submerge the tip of the recording pipette into the recording solution.

  • a.
    Apply positive pressure with the aid of an insulin syringe connected to the head stage.
  • b.
    Adjust the current offset to zero.

Note: Headstage, amplifier and digitizer settings should be optimized for each specific experiment. In our case, for voltage clamp experiments recording conditions are: Offset potential 10 V/V; scale membrane current 5 V/nA; output gain 10; Rf 500 MΩ, Bessel filter 2 kHz and sample at 10 kHz. For current clamp experiments: Command current (0.5 V/nA); scale membrane potential (50 V/V); output gain 50 Rf 500 MΩ and Bessel filter 2 kHz. The test pulse parameters are set to 5 mV for 20 ms at 50 Hz.

• 22.With the aid of a micromanipulator move the pipette tip into the field of view and lower it until the target cell and the pipette are in the same plane.
• 23.Adjust the current offset to zero again if needed.
• 24.
  Bring the patch pipette close to the granule cell soma until a small invagination (dimple) of the cell membrane is visualized (Figure 4D).

  • a.
    Release positive pressure and apply slow, continuous negative pressure using buccal suction to bring the outer leaflet of the cell membrane to the pipette tip (Figure 4E).

    CRITICAL: the appropriate amount of pressure is critical to make good seal. The membrane test should show near 0 pA response and over 1 GΩ pipette resistance, indicating successful “cell-attached” configuration. For high quality whole cell recordings, a general rule is that the seal resistance must be >10× the input resistance of the cell. For mature granule cells, the membrane seal resistance should be >4GΩ.

  • b.
    Set the holding voltage to −70 mV to prevent cell depolarization when the membrane patch is broken.
  • c.
    Cancel the pipette capacitive transients that can be revealed during giga-seal formation, which result from current charging and discharging the pipette’s capacitor.
• 25.
  Apply brief, intense negative suction pulse to rupture the membrane patch enclosed in the pipette (Figure 4F).

  • a.
    Carefully, apply additional brief suctions if the initial break is incomplete.

    CRITICAL: Excess suction can disrupt the giga-seal and the cell can be lost.

  • b.
    Electrical access is achieved once you break the patch reaching the “whole cell” configuration.

    Note: This is manifested by the appearance of large capacitive transients and a reduction in the membrane resistance.

    CRITICAL: Do not attempt the same cell or reuse patch pipette if the patching maneuver fails.

• 26.Allow several minutes (3–5 min) to complete the cell dialysis while monitoring the seal resistance.

Note: Values of electrochemical parameters such as holding current, membrane capacitance, membrane resistance, and access resistance depend on the cell type, internal solution used, experimental conditions, and experimenter technique.

CRITICAL: It is essential that these parameters are stable for the duration of all experiments. Stability of recordings must be established by trial-and-error testing prior to experiments, and throughout each experiment.

• 27.
  For voltage clamp experiments measuring sodium channel leak current.

  • a.
    Record 5 min of basal holding current in presence of synaptic and potassium blockers.
  • b.
    Replace ACSF with NMDG solution.

Figure 4.

Electric stimulation and patch clamp

(A) DIC image of a ventral hippocampal slice with stimulation pipette placed at the crest of the dentate gyrus (Objective 5×).

(B) DIC image of the granule cell layer of the dentate gyrus showing healthy well-packed cells suitable to be patched (Objective 60×).

(C) TdTomato epifluorescence of image showed in panel B, with the bright fluorescence dimmed to facilitate identification of individual cells.

(D–F) Schematic depicting the steps for whole cell configuration. (D) Pipette approaching with positive pressure generating a modest invagination of the cell membrane. (E) Cell attached configuration after removing positive pressure and applying negative suction (buccal suction or syringe aspiration) to achieve GΩ seal. (F) Whole cell configuration after applying brief pulses of negative pressure. Cartoons created with Biorender.com.

Immunohistochemistry

In this major step, we use immunohistochemistry to validate our genetic model. We use BMAL1 IHC to quantify Bmal1 deletion after measuring BMAL1 immunoreactivity in linear regions of interest in both, Pomc-Cre:Ai14 and Bmal1 cKO mice. We also assess the efficiency of Cre targeting quantifying co-localization between the GC marker Prospero homeobox protein 1 (PROX1) and the Ai14 reporter (TdTomato).

Intracardiac perfusion and harvesting the brain

Timing: 30 min per mouse

In this step, mice are perfused by intracardiac administration of 4% Paraformaldehyde (PFA). The harvested brain is left over 12 h in 4% PFA (troubleshooting problem 3).

• 28.
  Prepare for perfusion.

  • a.
    Obtain a 3-way stopcock with extension tubing.
  • b.
    Fill a 30 mL syringe with 1× PBS (25 mL/mouse).
  • c.
    Fill a separate 30 mL syringe with 4% PFA solution (20 mL/mouse).
  • d.
    Connect a butterfly needle to the male luer lock port of the 3-way stopcock and the syringes to the female luer port of the 3-way stopcock.
  • e.
    Pump 5 mL of the 1× PBS into the main line.
• 29.
  Anesthetize the mouse.

  • a.
    Administer 0.2 mL of isoflurane in an anesthetic drop jar.
  • b.
    Administer 0.8 mL of Avertin working solution intraperitoneally using an insulin syringe with a 27G needle.
  • c.
    Lay the mouse on the dissecting tray.
  • d.
    Perform the toe pinch test to ensure that the mouse is completely unresponsive to painful stimuli.
  • e.
    Pin the mouse to the dissecting tray using BD 20G 1^1/2 needles.
• 30.
  Expose the heart.

  • a.
    Incise the skin over the abdomen with a forceps and a fine scissors.
  • b.
    Incise the diaphragm taking care to not to puncture the beating heart.
  • c.
    Cut the ribcage on both sides of the sternum up to the level of the collar bones using a fine scissors.
  • d.
    Lift the sternum with a forcep to expose the heart.
  • e.
    Pin the overturned sternum to the dissection board using a needle.
• 31.
  Intracardiac perfusion.

  • a.
    Remove the pericardial sac using a forceps.
  • b.
    Cut into the right auricle using a micro spring scissor.
  • c.
    Insert the butterfly needle into the left ventricle of the heart.
  • d.
    Perfuse in 20 mL of 1× PBS.
  • e.
    Switch the stopcock to perfuse in 20 mL of 4% PFA.

Note: Indications of a good perfusion include stiffening of the tail and body and gradual paling of the liver. Additionally, fluid should not inflate the lungs or exit through the nose or mouth. Gentle consistent pressure will help achieve a good perfusion.

• 32.
  Collect the brain.

  • a.
    Remove the butterfly needle and the pins holding the mouse to the dissection board.
  • b.
    Cut off the head at the base of the skull using surgical scissors.
  • c.
    Cut through the midline of the scalp using hardened fine scissors to expose the skull.
  • d.
    Cut transversely along the width of the interparietal bone using hardened fine scissors.
  • e.
    Cut transversely between the eye sockets.
  • f.
    Cut anteriorly trough the midline of the skull along the sagittal suture using a micro spring scissors.
  • g.
    Remove the left and right flaps of the skull to expose the brain using forceps. Use care to avoid pressure or punctures to the brain.
  • h.
    Remove the brain using a spatula.
  • i.
    Place the brain into a glass jar with 4% PFA store 12–18 h at 4°C.

Sectioning

Timing: 1 h per brain

In this step, the brain is sectioned into 50 μm thick slices (troubleshooting problem 4).

• 33.
  Prepare.

  • a.
    Fill about half of each well in a 48-well plate with anti-freeze solution.
  • b.
    Obtain an injector blade and clean the blade with a disposable cotton swab that has been dipped in acetone.
  • c.
    Attach the blade to the vibratome and adjust the blade angle to 21°.
  • d.
    Remove the brain from its container filled with 4% PFA and place on filter paper.
  • e.
    Cut off the olfactory bulb and cerebellum using a single edge razor blade.
• 34.
  Mount the brain to the specimen holding platform.

  • a.
    Cut a small block of agar that is roughly equal in size to the brain and superglue it to the mounting block, just behind where the brain will be placed, to serve as a support.
  • b.
    Apply superglue on the location on the mounting block where the brain will be placed.
  • c.
    For horizontal sections, place the brain on the glue in front of and flush against the agar with the ventral side down and anterior (nose) facing the blade.
  • d.
    Allow glue to harden for about 2 min.
• 35.
  Collect 50 μm sections of the brain.

  • a.
    Attach the mounting block to the platform.
  • b.
    Turn on the vibratome.
  • c.
    Set vibratome to manual mode.
  • d.
    Fill the buffer tray with 1× PBS.
  • e.
    To collect sections containing the hippocampus, discard initial sections (∼500 μm thick) until hippocampus is evident.
  • f.
    Collect the slices with a paintbrush.
  • g.
    Transfer tissue to the well plate that has been filled with anti-freeze solution.

Immunostaining BMAL1

Timing: approximately 32 h

• 36.
  Rinse anti-freeze solution from brain slices.

  • a.
    Obtain desired tissue samples and place into a 48 multi-well plate (1 slice/well).
  • b.
    Wash 3 times with at least 0.5 mL of 1× PBS for 10 min on shaker at 22 ± 2°C while protecting from light.
• 37.
  Permeabilization and blocking of brain slices.

  • a.
    Obtain blocking buffer solution for BMAL1 IHC.
  • b.
    Add 0.5 mL of blocking buffer per well.
  • c.
    Incubate for 1 h at 22 ± 2°C on shaker while protecting from light.
  • d.
    Remove the blocking buffer solution.
• 38.
  Incubate in primary antibody.

  • a.
    Dilute in Anti-BMAL1 polyclonal rabbit primary antibody in 1:2000 ratio in antibody buffer solution.
  • b.
    Add 0.5 mL of primary antibody buffer solution to each well.
  • c.
    Incubate 12–18 h in cold room (4°C) on shaker while protecting from light.

Alternatives: Use a refrigerator if a cold room is not available.

• 39.
  Incubate in secondary antibody.

  • a.
    Wash 3 times with at least 0.5 mL of 1× PBS per well for 5 min on shaker at 22 ± 2°C while protecting from light.
  • b.
    Obtain antibody buffer solution and add in goat anti-rabbit Alexa Fluor 647 secondary antibody in 1:1000 ratio.
  • c.
    Add 0.5 mL of secondary antibody buffer solution to each well.
  • d.
    Incubate for 2 h at 22 ± 2°C on shaker while protecting from light.
  • e.
    Remove secondary antibody buffer solution.
• 40.Wash slices 3 times with at least 0.5 mL of 1× PBS for 10 min on shaker at 22 ± 2°C while protecting from light.
• 41.
  Mount with Prolong Gold.

  • a.
    Fill a petri dish with 1× PBS.
  • b.
    Place microscope-slide inside the petri dish.
  • c.
    Use paintbrush to transfer slices to petri dish filled with 1× PBS.
  • d.
    Use paintbrush to gently guide slices onto the appropriate location on the microscope-slide.
  • e.
    Remove slide from 1× PBS bath.
  • f.
    Slightly tilt the microscope slide.
  • g.
    Remove the excess 1× PBS on slices by gently touching a Kimtech wipe to the beaded solution, taking care not to touch the tissue.
  • h.
    Dip a plastic pipette into Prolong Gold mounting solution and touch the pipet tip on the slide.
  • i.
    Align a coverslip to the edge of the microscope slide and gently drop the coverslip to place the coverslip on the slide.
  • j.
    Store the microscope slide in a dark place at 22 ± 2°C for at least 24 h to allow medium to solidify before imaging.
• 42.Acquire images using a confocal microscope (Figure 5).

Figure 5.

Representative examples of successful immunohistochemistry

(A) Confocal image (Objective 20×) of PROX1 IHC staining.

(B) Confocal image (Objective 60×) of BMAL1 IHC staining.

(C) Confocal high-magnification image from a Bmal1 cKO. Dash line shows region of interest (ROI).

Immunostaining PROX1 (Prospero homeobox protein 1 used as a marker for granule cells)

Timing: approximately 32 h

• 43.
  Rinse anti-freeze solution from brain slices.

  • a.
    Obtain desired tissue samples and place into a 48 multi-well plate (1 slice/well).
  • b.
    Wash 3 times with at least 0.5 mL of 1× PBS for 10 min on shaker at 22 ± 2°C while protecting from light.
• 44.
  Permeabilization and blocking of brain slices.

  • a.
    Obtain blocking buffer solution for PROX1 IHC.
  • b.
    Add 0.5 mL of blocking buffer per well.
  • c.
    Incubate for 1.5 h at 22 ± 2°C on shaker while protecting from light.
  • d.
    Remove the blocking buffer solution.
• 45.
  Incubate in primary antibody.

  • a.
    Obtain antibody buffer solution for PROX1 IHC and dilute in Goat Anti-PROX1 primary antibody in 1:500 ratio.
  • b.
    Add 0.5 mL of primary antibody buffer solution to each well.
  • c.
    Incubate 12–18 h in cold room (4°C) on shaker while protecting from light.

Alternatives: Use a refrigerator if a cold room is not available.

• 46.
  Incubate in secondary antibody.

  • a.
    Wash 3 times with at least 0.5 mL of 1× PBS per well for 5 min on shaker at 22 ± 2°C while protecting from light.
  • b.
    Obtain antibody buffer solution and add in donkey anti-goat Alexa Fluor 647 secondary antibody in 1:500 ratio.
  • c.
    Add 0.5 mL of secondary antibody buffer solution to each well.
  • d.
    Incubate for 2–3 h at 22 ± 2°C on shaker while protecting from light.
  • e.
    Remove secondary antibody buffer solution.
• 47.Wash slices 3 times with at least 0.5 mL of 1× PBS for 10 min on shaker at 22 ± 2°C while protecting from light.
• 48.
  Mount with Prolong Gold.

  • a.
    Fill a petri dish with 1× PBS.
  • b.
    Place microscope-slide inside the petri dish.
  • c.
    Use paintbrush to pick up slices.
  • d.
    Place slices in the petri dish filled with 1× PBS.
  • e.
    Use paintbrush to gently guide slices onto the appropriate location on the microscope-slide.
  • f.
    Remove slide from 1× PBS bath.
  • g.
    Slightly tilt the microscope slide.
  • h.
    Remove the excess 1× PBS on slices by gently touching a Kimtech wipe to the beaded solution, taking care not to touch the tissue.
  • i.
    Dip a plastic pipette into Prolong Gold mounting solution and touch the pipet tip on the slide.
  • j.
    Align a coverslip to the edge of the microscope slide and gently drop the coverslip to place the coverslip on the slide.
  • k.
    Store the microscope slide in a dark place at 22 ± 2°C for at least 24 h to allow medium to solidify before imaging.
• 49.Acquire images using a confocal microscope.

Expected outcomes

Using this slice preparation method will yield high-quality slices from 2- to 3-month-old adult mice with viable neurons. Thanks to the "dentate gyrus cut," dendritic processes of granule cells will remain intact, as they project into deeper layers of the slice. Granule cells recorded in whole-cell configuration typically exhibit action potentials with amplitudes of +100 mV above threshold, uncorrected resting membrane potentials more negative than −70 mV, and input resistances generally ranging between 200 and 400 MΩ. Electrical stimulation of the perforant path at the apex under these conditions reliably generates excitatory postsynaptic potentials (EPSPs) and can recruit action potentials depending on the time of day. Although whole-cell patch-clamp recordings from single neurons in acute brain slices have certain limitations¹³ (such as series resistance, space clamp errors or junction potentials), it remains a valuable approach for detailed analysis of synaptic and intrinsic properties. Employing combined electrophysiological and anatomical techniques, along with the utilization of cell-type-specific Cre mice, represents an advantageous strategy for labeling and manipulating specific circuits or projection pathways.

Quantification and statistical analysis

To calculate rhythmic activity of intrinsic properties use Cosinor analysis with the following equation:

$$\mathrm{f}\left(\mathrm{t}\right)=\text{Mesor}+\mathrm{A}\times \mathrm{Cos}[(2\pi \mathrm{t}/\mathrm{T})+\text{Acrophase}]$$

where:

Mesor is the mean of the oscillation.

A is the amplitude of the oscillation.

T is the period (24 h).

Acrophase is the timing of the cosine maximum.

t is a time point.

Images from both Bmal1 cKO and control samples were captured using identical microscope settings to ensure consistency. Subsequent analysis was conducted without adjustment using FIJI software. A linear region of interest (ROI) was carefully drawn across the soma of 3–5 neighboring tdT⁺ and tdT⁻ cells in each image (see Figure 5C). The "Plot Profile" option in FIJI was then utilized to obtain an arbitrary gray value along each point of the ROI in both the red and green channels. The correlation coefficient (R value) at each point was calculated using Prism software, allowing for quantitative comparison between the two channels.¹

Limitations

To achieve successful patch clamp recordings, several critical steps must be meticulously executed. Firstly, the preparation of viable slices demands rigorous maintenance of equipment, storage containers, and tools. Following dissection, thorough rinsing and cleaning of all equipment is essential. The storage chamber should undergo a similar process, using ethanol and then be rinsed under deionized water to ensure proper sanitation. Additionally, the preparation of oblique horizontal slices may require optimization, considering variations in brain size and shape among different animal strains, age and sex. Furthermore, incubation in a well-perfused storage chamber is vital for observing network activity in slices. Fluorescence in slices obtained from transgenic mouse lines offers a valuable method for targeting specific neurons. However, it is important to note that fluorescence loss due to prolonged exposure and over time is inevitable. To mitigate this, limiting the exposure of slices to white light during cutting (switching off vibratome light), storage (using appropriately the lid of the chamber), slice transfer (reducing the time of light exposure) or recording (keeping the recording room in darkness) can help reduce the loss of fluorescence. Moreover, optimizing camera settings is essential to obtain the best fluorescent and contrast images with minimal exposure.

Troubleshooting

Problem 1

Inadequate slice viability resulting in failure of high-quality electrophysiology recordings (steps 1–14).

Potential solution

To address the persistent poor slice health, several potential solutions can be considered.

• •Properly calibrate instruments used to prepare solutions to maintain accurate composition.
• •Keep electrophysiology tools separate from other dissection tools (such as IHC set).
• •Verify that the storage chamber has reached the appropriate temperatures before slicing.
• •Minimize oxygen deprivation during brain dissection and tissue block preparation by improving technical skills and completing procedures quickly.
• •Reduce mechanical stress and avoid excessive disturbance during slice transfers.
• •Slightly vary the angle of slice preparation to optimize it, noting that slices at different locations along the septal-temporal axis will exhibit different consequences.¹⁰

Problem 2

Recurring issues with electrophysiological recordings (steps 15–27).

Potential solution

• •
  Offset unstable.

  • ○
    Re-chloride the electrodes.
  • ○
    Rule out a possible leak in the recording chamber.
• •
  Difficulty to seal or to break in.

  • ○
    Check the osmolarity of the internal solution and external solution.
  • ○
    Prepare new stocks for internal and external.
  • ○
    Pull smaller pipettes to improve seals or bigger pipettes to facilitate patch rupture.
  • ○
    Adjust positive pressure while approaching the soma.
• •
  Electrical stimulation fails to reliably evoke synaptic EPSPs.

  • ○
    Replace stimulating pipette (if clogged).
  • ○
    Verify intact stimulating circuit with ground wire in bath.
  • ○
    Check batteries of the Isolated Pulse Stimulator and verify output using an oscilloscope.
• •
  Noisy recordings.

  • ○
    Chapter 11 of The Axon Guide contains many useful suggestions regarding equipment set-up and grounding

Problem 3

Low quality intracardiac perfusion.

Potential solution

• •
  Liver does not turn pale, and tail is relaxed during perfusion.

  • ○
    Reposition the butterfly needle in the right ventricle.
  • ○
    Increase technical skills.
• •
  Lungs inflate and solution leaks through nose during perfusion.

  • ○
    Reposition the butterfly needle in the right ventricle.
  • ○
    Improve technical skills.

Problem 4

Problems with IHC.

Potential solution

• •
  Antibody do not penetrate or strong staining at tissue edges.

  • ○
    Prolong antibody incubation time and/or increase Triton X-100 concentration.
• •
  High background.

  • ○
    Improve intracardiac perfusion.
  • ○
    Extend the time of blocking and/or washing.

Resource availability

Lead contact

Further information and request for resources and reagents should be directed to and will be fulfilled by the lead contact, Linda Overstreet-Wadiche (lwadiche@uab.edu).

Technical contact

Questions about the technical specifics of performing the protocol should be directed to and will be answered by the technical contact, Jose Carlos Gonzalez (jcgonza@uab.edu).

Materials availability

This study did not generate new unique reagents.

Data and code availability

This study did not generate unique datasets.