Hippocampal Unicellular Recordings and Hippocampal-dependent Innate Behaviors in an Adolescent Mouse Model of Alzheimer’s disease

Materials and Reagents

Materials

1. Behavioral tasks
   Nesting
   1. Cotton square (5 cm x 5 cm, any commercial brand)
   Burrowing
   2. Handmade plastic black tube 20 cm long and 7.5 cm in diameter (The open end of the tube was elevated 2.5 cm from the floor (using two screws), to prevent accidental displacement of the tube’s content. The opposite end of the tube was blocked and touched the floor, as shown in Figure 1)
      
      
      Figure 1. Homemade plastic black tube used as a burrow
      
      
      
2. Electrophysiology
   1. Plastic tubing for perfusion system (Tubing Master Flex, catalog number: 065009-16; Tygon E-3603 and Tygon tubing, Cole Parmer, catalog number: 06460-34)
   2. Recording chamber and platform (Homemade, see Figure 8)
   3. Capillary glass tubing (Borosilicate standard wall with filament, length = 10 cm, Warner Instruments, catalog number: G150F-4)
   4. Camera with monitor (DAGE-MTI, catalog number: CCD-100)
   5. Double edge razor blades (Any commercial brand)
      
      
      

1. Na₂ATP (Sigma-Aldrich, catalog number: 34369078)
2. LiGTP (Sigma-Aldrich, catalog number: 85737048)
3. Potassium gluconic acid (Sigma-Aldrich, catalog number: 299274)
4. EGTA (Sigma-Aldrich, catalog number: 67425)
5. HEPES (Sigma-Aldrich, catalog number: 7365459)
6. Sucrose (Sigma-Aldrich, catalog number: S9378)
7. Potassium chloride (KCl) (Sigma-Aldrich, catalog number: P4504)
8. Sodium chloride (NaCl) (Sigma-Aldrich, catalog number: S9625)
9. Magnesium chloride (MgCl₂) (Sigma-Aldrich, catalog number: M8266)
10. Sodium bicarbonate (NaHCO₃) (Sigma-Aldrich, catalog number: S8875)
11. Calcium chloride (CaCl₂) (Sigma-Aldrich, catalog number: C3881)
12. D-glucose (Sigma-Aldrich, catalog number: D8270)
13. Cyanoacrylate (Any commercial brand)
14. Pentobarbital
15. Intracellular solution for patch-clamp recordings (see Recipes)
16. External solution (artificial cerebrospinal fluid (aCSF)) (see Recipes)
17. Internal solution (see Recipes)

Equipment

1. Behavioral tasks
   Nesting
   1. Individual cage (26 cm x 48 cm x 20 cm) with wood chip bedding
   Burrowing
   2. Individual cage (26 cm x 48 cm x 20 cm) with wood chip bedding
      
      
2. Electrophysiology
   1. Peristaltic pump (DYNAMAX, model: Rp-1)
   2. In line solution heater (Automatic Temperature Control, Warner Instruments, model: TC-324C)
   3. Temperature controller (Harvard Apparatus, model: TC-202A)
   4. Amplifier (Axon Instruments, model: Axo-clamp 2B amplifier)
   5. Connector block (National Instruments, model: BNC-2110)
   6. Digitizer (National Instruments, Digidata BNC-2110)
   7. Micromanipulator (Sutter Instrument Co, model: MP-225)
   8. Air table serial #119564, TMC and Faraday Cage (Homemade)
   9. Microscope (NIKON, model: E600-FN)
   10. Flaming/Brown Micropipette (Sutter Instrument Co, model: P-97)
   11. Vibratome (Thermo Scientific, model: Microm HM-650V)
       

Software

1. Any available software that controls amplifier and digitizer, for example, pCLAMP^TM Software. In our experiments we used software developed in LabView, National Instruments
   

Procedure

1. Behavioral tasks (both protocols are performed in the same behavioral room and all animals perform both tasks)
   Nesting
   1. Start the nesting protocol at 08:00 a.m., one hour before the dark phase (12-h light/12-h dark cycle). Place the animal into an individual cage filled only with wood-chip bedding (around 2 cm). After 60 min, place a cotton square (5 cm x 5 cm) on the bedding (Figure 2, see corner of the cage). Innately, animals use the cotton to build a nest. The following morning (08:00 a.m.), perform evaluation of nest.
      
      
      Figure 2. Place an animal in an individual cage with wood-chip bedding following by a cotton square 60 min later (scale bar = 5 cm)
      
      
      
   Burrowing
   1. Place each animal in an individual cage (26 cm x 48 cm x 20 cm) with the burrow tube filled up with 200 g of food pellets (these pellets are standard mouse food pellets normally supplied as their diet). The animals have access to water, but no access to food other than the one placed in the tube.
   2. Start the test at 06:00 a.m., three hours before the dark phase. Record the amount of food pellets displaced after the first 2 h of the test.
   3. The weight of the pellets burrowed for 2 h is then quantify by subtracting it from the original 200 g. The burrow tube is refill with pellets, and the test continue. Perform the final quantification of the displaced pellet weight after 12 h, as shown in figure 3.
      
      
      Figure 3. Plastic black tube with pellets displaced at time 0, 8 a.m. and 9 p.m.
      
      
      
2. Electrophysiology
   Patch-clamp (ethics approval should be obtained)
   1. Deeply anesthetize the animals with pentobarbital (63 mg/kg, alternatives ketamine and xylazin) to perform an intravascular perfusion (30 ml) of modified aCSF bubbled with carbogen (95% O₂ and 5% CO₂). At the end of the infusion, decapitate the animal and remove the brain in ice-cold aCSF. To obtain para-sagittal slices (Figures 4 and 5), remove the cerebellum and brainstem, separate the brain hemispheres and glue them by their medial portion at a 30° angle (Figures 4 and 5). First, glue the brain to a piece of filter paper and then, glue the filter paper (with the brain) to the agar block (Figures 4 and 5, agar size tends to vary according to brain size). Throughout this process, the brain tissue should be immersed in normal aCSF with continuous bubbled with carbogen.
      
      
      Figure 4. To obtain para-sagittal slices (see sagittal cut, red dotted line), separate the brain hemispheres and glue them by their medial portion at a 30° angle. First, glue the brain to a piece of filter paper and then, glue the filter paper (with the brain) to the agar block (see sagittal slice sequence). The agar block is glue to the adaptor tray (R = rostral, D = dorsal, C = caudal, V = ventral).
      
      
   2. Once the brain hemisphere is attached to the agar block, place the tissue in an adaptor tray submerged in ice-cold aCSF bubbled with carbogen. To obtain the parasagittal slices of the hippocampal formation (350-400 µm thick), couple the tray to a vibratome. Cut slices (dorsoventrally), and ventral slices containing hippocampus, 4-6 slices from each hemisphere (Figure 5). Immediately, transfer them into a nest beaker and incubate them for 1 h, at room temperature, with bubbled aCSF (Figure 6).
      
      
      Figure 5. Sagittal slice sequence. With the razor blade, separate the brain hemispheres (A), glue the brain to a piece of filter paper (B) and then, glue the filter paper (with the brain) to the agar block (C to E). Finally, transfer the plate into the slicing chamber of the vibratome and lower the blade holder (F). Once the slice is freed, transfer it to the nest beaker with recovery aCSF.
      
      
      Figure 6. Incubate sagittal slices in aCSF bubbled with carbogen at room temperature for at least one hour for recovery. The nest beaker could be prepared according to Papouin et al., 2018.
      
      
   3. After recovery, transfer the slices to the recording chamber located in a microscope equipped with optical components to perform Differential Interference Contrast (DIC) (Eclipse E600FN; Nikon, Melville, NY, Figure 7).
      
      
      Figure 7. Electrophysiology whole-cell patch-clamp set up and microscope
      
      
   4. In the recording chamber (Figures 8 and 9), place the slice in a well, which could contain up to 3 ml circulating aCSF bubbled with carbogen, at a temperature of 32 ± 0.5 °C. A total of 30 ml of aCSF continuously recirculate through the recording chamber at a flow rate of 17-20 ml/min (Figures 8 and 9).
      
      
      Figure 8. Slice holder and recording chamber. The slice holder is made of a bent and flattened stainless-steel wire crossed by horizontal nylon fibers (around 3 mm apart). The recording chamber is made of acrylic (measuring length in centimeters, cm).
      
      
      Figure 9. Mounting and holding the brain slice containing the hippocampus onto the recording chamber. The brain slice containing the hippocampal formation is placed onto the recording chamber and immobilized with the holder, leaving the recording area free for electrode location but fixed to maintain stable recording while the slice is perfused at high speed (17-20 ml/min).
      
      
   5. Perform recordings with glass pipettes (Warner Instruments, G150F-4; Handem, CT) made in a P-97 puller (Sutter Instruments, Novato, CA). The electrode resistance is 2-5 MΩ. Fill pipettes with an internal solution.
   6. The visual patch-clamp technique is on the whole-cell configuration in the current clamp mode (Figure 10). The biological signals are measured through an Axoclamp 2B amplifier (Axon Instruments, Foster City, CA). Always focus recordings on neurons located in the CA1 pyramidal area (CA1 has the least heterogeneous neuronal population and the pyramidal cells are arranged into well-defined layers, see Mondragón-Rodríguez et al., 2018a). Once the whole-cell configuration is achieved, hyperpolarizing and depolarizing current pulses are applied to evaluate the intrinsic properties of the recorded neurons (Figures 10 and 11). During this current injection protocol, the neurons remained at the original resting potential (variable between neurons). However, the same protocol was repeated while the cells had a -70 mV membrane potential, which was achieved mostly by applying hyperpolarizing current (for whole-cell patch-clamp video see Sunstrum and Inoue, 2019).
      
      
      Figure 10. Images of the CA1 pyramidal cell layer visualized through a differential interference contrast (DIC) microscope. The visual patch-clamp technique, in the whole-cell configuration, is achieved with pipette electrodes (highlighted with the dotted line) located on the cell’s soma (10x and 40x magnification on the left and right, respectively).
      
      
   Electrophysiological variables
   1. Resting membrane potential (trans-membrane voltage, Vm) is measured immediately after obtaining the whole-cell configuration.
   2. Fire frequency is measured as the number of action potentials (APs) generated by a neuron during a supra-threshold pulse of a one-second depolarizing current (Figure 11). To compare all neurons, the minimal depolarizing current injection required to evoke at least one AP (rheobase) is multiplied by two to quantify the frequency of the AP evoked by current injection.
   3. Fire frequency adaptation is calculated by dividing T2/T1, where T2 is the time between the last two APs of a train and T1 is the time elapsed between the first two APs of the same train. A value larger than one indicates adaptation of AP firing, whereas a value close to or less than one indicates no adaptation of AP firing or acceleration.
   4. Membrane resistance is measured using Ohm's law by dividing the change in voltage induced by the injected current. This measurement is performed for five hyperpolarizing current pulses.
      
      
      Figure 11. Characterization of the evoked firing pattern of a recorded whole-cell neuron. Injecting a one-second, 30 pA depolarizing current pulse (lower trace) induced a train of action potentials (upper trace), which exhibit the adaptation phenomenon (reduction in firing frequency during steady stimulation).
      
      
      
   Subthreshold oscillations
   To characterize subthreshold oscillations, bring the membrane potential to the level just below the threshold to trigger APs (although sporadic potential can be induced). Keep the membrane potential at the value at which maximal amplitude subthreshold oscillations could be observed while triggering the least amount of APs as possible. Such voltage is represented in Figure 12.
   
   
   Figure 12. Bring the membrane potential to the level just below the level where action potentials are triggered (see arrow)

Data analysis

1. Behavioral task
   Nesting
   Take images and evaluate the construction of the nest. Three categories are defined based on those proposed by Deacon (2012), exemplified next:
   Assign category 1 when the nestlet is not noticeably made (Figure 13A, >90% intact cotton). Assign category 2 when the nestlet is partially torn up (50 to 90 % of the cotton was shredded), with or without identifiable nesting (Figure 13B). Assign category 3 when a (near) perfect, crater-shaped nest is formed with walls higher than the mouse’s body weight, allowing the animal to curl up in the nest with more than 50% of its circumference (Figure 13C, >90% shredded cotton). In sum, the quantification will show the nest rating from 1 to 3, depending on the category (Figure 13).
   Note: In a blind experiment, genotypes of all animals are withheld (masked) until the experiments are complete.
   
   
   Figure 13. Nestlet is not noticeably made (A, see cotton square), nestlet partially torn up (B, see cotton square) and a near perfect nest (C, see cotton square)
   
   Burrowing
   Quantify the weight of displaced pellets (g) after 2 h and then, after 12 h (see Figure 3).
   
   
2. Electrophysiology
   Patch-clamp
   Record with a sampling frequency of 9 KHz. An HS-2 head stage, with a gain of 0.01 MU is used (Molecular Devices). Sample the signals by an analog-digital card coupled to a BNC-2110 connector (National Instruments) to convert the biological (analog) signal into a digital signal. Monitor the recorded signals online with an acquisition software instrument (LabView environment, National Instruments, Austin, TX) and store them in a computer for later analysis.
     Analyze the recordings off-line. It is important to mention that we always recorded one slice per animal or one neuron per slice. For classical power spectrum analysis of the membrane potential, segments of five seconds are analyzed using a Fast Fourier Transform algorithm with a Hamming window in Clampfit (Molecular Devices, Figure 14). The power spectrum, from 1 to 60 Hz, is integrated and normalized to control, meaning that the control power spectrum is integrated, and this value is arbitrarily set as 100%.
   
   
   Figure 14. For power spectrum analysis of the membrane potential, analyze segments using a Fast Fourier Transform algorithm with a Hamming window in Clampfit
   
   Expected results
   For expected results see Orta-Salazar et al., 2013 and Mondragón-Rodríguez et al., 2018a.
   

Recipes

1. Intracellular solution for patch-clamp recordings
   140 mM potassium gluconic acid
   10 mM EGTA
   2 mM MgCl₂
   10 mM HEPES
   2 mM Na₂ATP
   2 mM LiGTP
   pH 7.4
2. External solution (artificial cerebrospinal fluid (aCSF))
   119 mM NaCl
   3 mM KCl
   1.5 mM CaCl₂
   1 mM MgCl₂
   25 mM NaHCO₃
   30 mM D-glucose
   pH 7.4
3. Internal solution
   140 mM potassium gluconic acid
   1 mM CaCl₂
   10 mM EGTA
   2 mM MgCl₂
   4 mM Na₂ATP
   4 mM LiGTP
   10 mM HEPES
   pH 7.4
   

Acknowledgments

We thank Jessica Gonzalez Norris for proofreading and Ignacio Martínez García for technical support. This research was supported by the Dirección General de Asuntos del Personal Académico (DGAPA; Grant IN202018 and IN204519) and by CONACYT (Grants; 269021 and A1-S-7540). National Institute of Minority Health and Health Disparities Grant G12MD007591 from the National Institutes of Health and by the Semmes Foundation. S M-R was awarded a Cátedra position from CONACYT, México. This protocol was adapted or modified from Peña et al. (2010) and Deacon et al. (2012).

Competing interests

The authors declare no competing financial interests.

Ethics

Experimental procedures were approved by the Bioethics Committee of the Institute of Neurobiology, UNAM, Mexico. Approval ID-091/2015-2018.

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