Whatman filter paper 42 equivalent ratios

Preparation of acute sections of the hippocampus of rats and transgenic mice for the study of synaptic changes during aging and amyloid pathology


This article describes methods of making sections of the hippocampus of rats and transgenic mice for the study of synaptic changes with brain aging and age-related neurodegenerative diseases such as Alzheimer's disease.


The rodent hippocampal disc preparation is perhaps the most widely used tool for studying mammalian synaptic function and plasticity. The hippocampus can be quickly and easily obtained from rats and mice, and slices remain viable for hours in oxygenated artificial cerebrospinal fluid. In addition, basic electrophysisologic techniques are easily applied to the study of synaptic function in the hippocampal slices and have yielded some of the best biomarkers for cognitive impairment. The hippocampal disc is particularly popular for studying the synaptic plasticity mechanisms involved in learning and memory. Changes in the induction of long-term potentiation and depression (LTP and LTD) of synaptic effectiveness in sections of the hippocampus (or lack thereof) are often used to evaluate the neurological phenotype of cognitively impaired animals and / or to describe the mode of action of nootropic compounds . This article describes the procedure we use for preparing sections of the hippocampus of rats and transgenic mice for studying synaptic changes with brain aging and Alzheimer's disease (AD) 1-3 connected. Using aged rats and AD model mice present a unique set of challenges for researchers accustomed to using younger rats and / or mice in their research. Aged rats have thicker skulls and harder connective tissue than younger rats and mice, which can delay brain extraction and / or dissection and thereby negate or exaggerate real age differences in synaptic function and plasticity. Aging and amyloid pathology can also aggravate hippocampal damage sustained during the dissection procedure, again complicating any conclusions drawn from physiological assessment. Here we discuss the steps during the dissection procedure to minimize these problems. Examples of synaptic responses in "healthy" and "unhealthy" slices obtained from rats and mice are provided, as well as representatives of synaptic plasticity experiments. The possible effects of the other methodological factors on synaptic function in these animal models (e.g. recording solution components, stimulation parameters) are also discussed. While the focus of this article is on the use of ancient rats and transgenic mice, novices to the Physiology Slice should find enough detail here to begin their own studies with a variety of animal models.


1. Preparation of Ice Cold Oxygenated Artificial Liquor (ACSF)

  1. Prepare 2 L of "Approx 2+ -free " ACSF. In a 2L Erlenmeyer flask, add approximately 1.5L of sterile or double distilled H. 2 O, and begin stirring vigorously on a stir plate. Add the following ACSF components (in mM): 124 NaCl, 2 KCl, 1.25 KH 2 PO 4, 2 MgSO 4, 26 NaHCO 3 and 10 dextrose (see Table 1). Bring to a volume of 2 L with distilled H 2 O.
  2. With an aquarium bubbler and hose on a 95% O 2 / 5% CO 2 Air reservoir, oxygenate ACSF vigorously approx. 20-30 min.Check the pH value and, if necessary, adjust to 7.4 with NaOH or HCl.
  3. Pour 750 ml of oxygen approx 2+ free ACSF in a separate Erlenmeyer flask, cover the opening with parafilm and transfer to an ultrafreezer (-80 ° C) for 20-30 min. This media will be used for dissection of the brain and acute sections of the hippocampus *. Add 2 mM CaCl 2, to add the remaining 1.25 L volume of ACSF #, stir well and re-oxygenate with 95% O 2 / 5% CO 2. These media will be used for storing brain slices after dissection and for perfusion slices during electrophysiological recordings.
    * Ice cold approx 2 + free media is used to slow your metabolism and minimize Ca 2+ dependent Excitotoxicity when dissecting.
    # Changes in approx 2 + Dysregulation during aging and AD can have a big impact on the induction of Ca 2+ dependent synaptic plasticity 4-9. Conflicting reports of age differences in LTD may, in part, be due to the use of different ACSF Ca. 2 +: Mg2 + ratios during disc recordings (see 2,4,10). The importance of the Ca 2+ regulation and the ACSF Ca 2+ level addressed in greater detail in the discussion in aging studies.
  4. While the approx 2+ free ACSF freezes, prepare an acquisition chamber for maintenance of brain slices before and during electrophysiological recordings *. We use a custom macro-receiving chamber that contains four individual micro-chambers (see Figure 2). The macrochamber is filled with sterile, oxygen H 2 O and the micro-chambers are essentially islands that protrude above the surface of the water. Inside each microchamber, resting slices on a netted insert into a shallow basin of the oxygenated ACSF. Panes are not completely submerged, but sit at an interface with the humidified air. An isolated heating element within the macrochamber enables temperature adjustment.
    * Several varieties of holding chambers are also commercially available (e.g. Warner Instruments makes an immersion-style "Pre Chamber # BSC-PC) and should be used in aging and transgenic mouse slice studies. In a pinch, slices can be used for Be maintained for several hours in a small petri dish filled with ACSF. Make a small hole in the Petri lid for an oxygen supply tube. Be careful that oxygen bubbles do not physically move the slices. Slices will get adequate oxygen when the delivery tube It is only raised above the ACSF surface (gas dispersion should result in a depression in the surface of the liquid).

2. Brain Removal and Hippocampus Dissection in Aged (> 20-month-old rats)

  1. Prepare the dissection area * next to a large pelvis (see Figure 1 and Table 2). Place a small animal guillotine in the sink and lay out surgical instruments and materials for brain and hippocampal dissection including a folded paper towel, a No. 11 scalpel, beebee scissors, bone rongeurs, a "hippocampal tool" (a special one Dual spatulas from Fine Surgical Tools), small surgical scissors, a thin double-ended spatula, plastic Pasteur pipette, 110 mm diameter Whatman filter paper, a 100 mm lid glass petri dish filled with ice, and a plastic spoon. Also keep a plastic bag nearby for disposal of the carcass.
    * Brain extraction should be completed as soon as possible so it is a good idea to mentally "go through" the procedure and place your instruments in the order of use.
  2. Remove approx 2+ free ACSF from the freezer. The media should be partially, not entirely, frozen. Pour about 50 ml of ACSF into a beaker, cover with parafilm and place next to the dissection area. These media are used to briefly store the brain when it is removed. The remaining approx 2+ -free Media are used to fill the reservoir in the vibratome for piece preparation. This media can be reoxygenated, or simply covered with parafilm and refrigerated at 4 ° C.
  3. Euthanize the rat using methods approved by the Institutional Animal Care and Use Committee (IACUC). Our animals are in a small plexiglass chamber, which is gradually filled with 100% CO 2 is filled out. Loss of consciousness usually occurs within five minutes and is confirmed by the lack of reflex activity after pinching a toe.
  4. Decapitate the rat immediately rostrally from the first cervical vertebra and place the head on the folded paper towel. With the No. 11 scalpel, quickly make an incision in the center of the scalp starting near the nasal bone and running caudal to the occipital bone. Be sure to go completely through the skin muscle, fully exposing the sutures on the dorsal surface of the skull.
  5. Cut through the skull plates with beebee scissors. In young rats and mice, the skull can be removed quickly with the use of bone rongeurs alone. However, aged rats have thicker skulls than young adult rats and mice, which this procedure can be difficult. We found that cutting through the skull facilitates bone removal with rongeurs, reduces brain damage, and saves precious seconds. With the rat's head firmly on the tabletop, place the edge of the lower sheer of the beebee scissors in the superior area of ​​the foramen magnum at the back of the skull. Keeping the lower sheer tight against the skull's inner surface (and removal from the brain) *, cut through the back of the head plate and then suture the parietal plates along the midline. Go rostrally until you cut through the frontal skull plates.
    * It is very important that the pressure is directed away from the brain to prevent accidental measurement.
  6. Separate the occipital, parietal, and temporal skull plates from the brain. Keep your head firmly on the countertop for stability and leverage and slide the lower jaw of the rongeure under the left parietal plate maintaining pressure against the inside of the skull. Next, squeeze the rongeurs' jaws together and roll your wrist up and toward you to pull the parietal and occipital plates away from the brain. This should also cover most of the dorsal surface of the left hemisphere. If necessary, use the rongeurs to remove the left faceplate as well. Repeat this process for the other hemisphere. Once the plates are moved, quickly examine for each dura roll that the temporal plates can be attached and stretched across the surface of the brain. Pull this away with the rongeur or cut away with scissors *. Now slide the upper jaw of the rongeure between the brain and the right temple plate, again holding the pressure on the skull and away from the brain. Squeeze and rotate the temporal plate away from the brain. You will hear / feel "crunch" as you do this. Repeat for the left side.
    * The dura can be difficult to see, especially if the animal has been perfused transcardially with ACSF. However, if dura are not removed, they will be sliced ​​through the brain (and likely the hippocampus) like a razor when temporal plates are removed. Snipping the dura away near the temporal plates minimizes the likelihood of stabbing the brain.
  7. Extract the brain *. Quick removal of the parafilm from the approx 2+ free ACSF "mushy". Now slide the wide spatula head of the hippocampal tool between the ventral surface of the brain and the lower skull plates until it is completely under the brain. Move the spatula sideways from side to side and then back and forth a few times to sever intact cranial nerves. Now scoop up the brain with the hippocampus tool and plunge into approx 2+ free ACSF and cover with parafilm. Let the brain chill for about a minute. This is an opportune time to clean off the guillotine, discard the carcass, and reorganize the dissection area.
    * Hurry up for steps 2,3-2,7. We try to complete this procedure (from decapitating the brain by submerging it in ACSF) in 30-35 sec. In our experience, taking extractions longer than a minute appears to affect hippocampal disc health, especially for older rats. With these procedures, we did not observe any differences in extraction times between old and young rats for our studies.
  8. Unzip the hippocampi. Place the Whatman filter paper on the lid of the ice cold Petri dish and dampen the paper with ACSF using the plastic transfer pipette. Retrieve the brain from the ACSF with a spoon and place on the moistened Whatman paper. Use a scalpel to remove the cerebellum and around a quarter of the rostral frontal lobe. Run the scalpel through the intrahemispheric tear to completely separate the two hemispheres. Place one hemisphere back in the mushy ACSF and "stand" the other on the dissecting stage so that the coronal plane of the frontal lobe is facing down. You should clearly be able to distinguish the brain stem and midbrain (the white ones) from the overlying cortex (the pink / gray ones). Locate the upper and lower colliculi on the midbrain; these will look like two white "buttons" and will be at the "top" of the brain in this orientation. With the surgical scissors, gently hold the midbrain, and slide the scales spatula into the gap between the colliculi and the neocortex. Very gently, continue to slide the spatula down and pull the brain stem / midbrain / thalamus away revealing the inside of the lateral ventricle and the medial surface of the hippocampus. Use the sharp edge of the spatula to smoothly sever the fornix, a white bundle of fibers located at the front / back of the hippocampus. Use the scissors to gently pull further towards the brain stem / midbrain / thalamus without completely severing it from the rest of the brain. The cortex with hippocampus should now lay freely back from the brain stem *. Next, rotate the dissecting stage so that you are looking at the medial hippocampus and overlying cortex as if it were a sagittal incision (minus the thalamus). You should now see the white fimbria fibers that form a flat hyperbola at the bottom of the hippocampus in that plane. With the plastic transfer pipette, carefully inject some ACSF into the gap under the fimbria to help separate the hippocampus from the cortex. Slide the spatula into this gap so that the long side of the spatula is parallel with the long axis of the hippocampus. After the spatula is completely under the hippocampus, hold the brain stem / midbrain / thalamus firmly with the scissors and roll the spatula away from your body physically separating the hippocampus from the rest of the brain. Once the hippocampus is free, lightly cut away any remaining cortex, blood vessels, and white matter. Scoop a small amount of ACSF squishy on the other side of the dissection stage. Carefully position the hippocampus next to the slush and douse a few milliliters of ACSF with the plastic transfer pipette. Then remove the other hemisphere and repeat the dissection.
    * You must use the scissors to snip away any extra connective tissue, white matter, or blood vessels that prevent the brain stem from separating the cerebral cortex. The points of the scissors will be in close proximity to the hippocampus, so be sure to deliver very precise cuts (sharp scissors are a must).

3. Brain Removal and Hippocampus Dissection in Aged Transgenic Mice

  1. Euthanize and behead the mouse and make a midline incision on the scalp as described in Section 2.3. Mice have much thinner skulls than rats, which greatly simplifies brain extraction. Cutting through the skull with scissors is therefore not necessary. Use bone rongeurs with smaller jaws (Table 2), pull away, occipital, parietal, and temporal skull plates. As with the rat, remember to use controlled motions and pull the lower jaw of the rongeure into the inner surface of the skull and remove it from the brain as you remove the plates. Once the plates are removed, use the narrow end of the hippocampus tool to sever the remaining cranial nerves and scoop the brain in ice cold approx 2+ free ACSF.
  2. Prepare brain slices. Because of the smaller size of the mouse brain, dissecting the hippocampi can be a bit more difficult (though certainly doable) than when using rats. So, to make things easier, we remove the cerebellum and rostral tips of the frontal lobes, but do not dissect the hippocampi. Instead, halves of the brain are physically severed with a scalpel and left intact for vibratome incisions (see Section 4 below).

4. § Sliced ​​brain tissue with a vibrating microtome (vibratome) and transfer to the receiving chamber *

  1. Fill the vibratome reservoir with ice-cold approx 2+ free ACSF so that the cutting stage and blade are completely submerged. To make rat slices, cut the rostral and caudal tips of the hippocampus with a scalpel. These cuts allow you to vertically position the hippocampus close together like two pillars.This works best when the dentate gyrus of the hippocampus is facing each other and CA3 regions are aligned in the same direction. To make mouse slices, position each hemisphere vertically closely with the frontal lobe down. Glue brain tissue onto a mounting block and transfer it to the cutting phase of the vibratome. We usually prepare ~ 400 µM sections for synaptic physiology experiments. Thinner sections should be prepared for when fluorescence imaging is performed (e.g. examinations of the approx 2+ mirror and transients). Collect slices with a wide-mouth pipette or a brush # and transfer to a small petri dish with ice-cold approx 2+ free ACSF.
    * Using a vibratome minimizes damage to the top and bottom of the disc and is definitely used for studies requiring analysis of cells near the disc surface (e.g. voltage-clamp and approx 2+ imaging studies) recommended. However, cheaper alternatives are available and suitable for generating slices for extracellular physiology experiments. We have a small center of gravity controlled chopper 2,11 and a McIlwain tissue chopper 12 used with good success. For this procedure hippocampi are placed on a staged and cut with a vertical chopper. A brush is used to brush slices (one at a time) around a parcel bowl of ice-cold approx 2+ free ACSF filled to be transmitted. One problem with this approach is that the hippocampus can move between chops resulting in unequal parts. Also, be sure to remove as much of the white matter, and especially as much of the vascular system, as possible before chopping. This material will stick to the brush, the razor blade, or both, making slice transfer very difficult and increasing the likelihood of stretching or damaging the tissue.
    # If using a brush, try to rest the slice lengthwise over the bristles. Wrapping the disc around the brush tip can stretch unnecessary tissue.
  2. Transfer brain slices to the receiving chamber, where they are oxygenated to approx 2+ containing ACSF to be bathed. Gradually bring the chamber temperature to 27 ° to 32 ° C (+1 ° every five minutes). Let slices incubate for 1-1.5 h before electrophysiological experiments.

5. Elicit and Record CA3-CA1 Synaptic Responses

  1. For basic extracellular recordings in acute sections, your electrophysiology ward will need * include: a recording chamber, a perfusion system, a microscope with> 4x magnification capability, recording, stimulating, and ground electrodes; Macro and micromanipulators, a rigid vibration-proof table and Faraday cage, a stimulator, amplifier and analog-to-digital (A / D converter); Oscilloscope (preferably) and personal PC with appropriate software for acquisition.
    * Kerr Scientific Instruments offers a fantastic and inexpensive electrophysiology system (ie the Kerr Tissue Recording System) for performing a variety of basic disk electrophysiology experiments. This system has a small footprint, portable stimulators and amplifiers, and can be used on a standard laboratory bench without the need for a bulky Faraday cage.
    Of course, brain slices can also be used to perform numerous complex electrophysiological and imaging procedures that require additional equipment and materials. For example, our electrophysiology station contains an amplifier with fast voltage and current clamp functions, a fluorescent lighting system, and a digital camera. This station is for extracellular field recordings in brain slices as well as patch clamp recordings and fluorescence imaging in slices and cell cultures 3,12,13 used. See Table 3 for a complete list of devices and components.
  2. Using a wide-mouth pipette or small brush, transfer one or more segments to the recording chamber * allowing them to acclimate for 10-15 min prior to stimulation / recording. For our investigations, discs in ACSF and rest are inserted into an RC-22 chamber from Warner Instruments (see Figure 3) for offsetting. ACSF is gravity-fed by a regulator for flow adjustment, and preheated to 32 ° C by an in-line micro-heating element before reaching the intake chamber. A central vacuum line is used to remove ASCF.
    * Many different underwater and interface style intake chambers are available commercially. We observed that slices exhibit more robust synaptic responses in an interface chamber (i.e. slice sits at an interface with moist air and ACSF) 2,11. However, sensitivity is usually more stable when slices are submerged in ACSF. Drug perfusion is also more efficient in immersion chambers.
  3. Position stimulating and recording electrodes. With the stimulator or stimulus isolator switched on, but output selected to 0, position a stimulating electrode over the disc in the stratum radiatum region CA2 near the CA3 border (see Figure 4). We use a twisted platinum iridium wire to deliver 50-100 uS diphasic pulses to CA3 Schaffer Collateral (SC) fibers. The use of platinum-iridium wire and two-phase pulses can help minimize electrode polarization. Lower a recording electrode in the CA1 stratum radiatum, only breaking the surface of the disc. Our recording electrode is an Ag / AgCl wire in an ACSF filled glass micropipette (tip resistance of ~ 7 MOhm). Turn the output on the stimulator down to a moderate level (we're setting our Stimulus Isolator to ~ 150 uA) and start managing stimulus pulses and acquiring activity, slowly lowering with acquisition software, such as CLAMPEX from Axon Instruments, Inc. Apply the stimulating electrode at small intervals until a stimulus artificact is recorded in CA1. Next, continue to slowly lower both the exciting and recording electrodes at intervals while acquiring CA1 responses until the fiber volley (FV) and the excitatory postsynaptic potential (EPSP) amplitudes reach maximum values ​​(see Figure 4B).
  4. Make a synaptic strength curve and examine synaptic plasticity. To generate a synaptic strength curve, impulses deliver impulses SC in ever higher intensities and record the corresponding activity in CA1. The range and number of stimulus levels can vary, but should be sufficient to produce a sigmoid curve when plotted with either FV or EPSP values ​​(Fig. 5). The FV amplitude provides a reliable estimate of the proportion of presynaptic fibers activated, while the EPSP slope provides an unaffected measure of monosynaptic CA3-CA1 currents. Carrying the EPSP slope against the FV via stimulus levels therefore reflects the size of the EPSP per number of activated afferents and provides an excellent estimate of the CA3-CA1 synaptic strength. Typically, synaptic strengths are significantly reduced in the CA1 in aged rats and APP / PS1 mice, based on young rats and wild-type mice of the same age, see eg 4.14.
  5. Slices that show signs of ill health (i.e. a maximum EPSP amplitude <1 mV) or hyperexcitability (i.e. the occurrence of 2 or more population spikes) during the synaptic strength curve excluded from statistical analysis (see Figure 4C). We found that these discs rarely show stable responses over a 60 min period and / or show very variable responses after induction of synaptic plasticity. It deserves to be pointed out that "unhealthy slices" make up only about 10-20% of all slices transferred to the intake chamber. Also, in our experience, the frequency of identifying an unhealthy slice is very similar across age, species, and genotype.
  6. Induce long-term potentiation (LTP) or long-term depression (LTD) cut into slices. LTP and LTD are permanent increases (LTP) and decreases (LTD) of synaptic function in response to different patterns of synaptic activation. Both processes are generally believed to reflect critical mechanisms for learning and memory15 and provide useful results measures for studying cellular mechanisms of neural dysfunction and / or for assessing pharmacological strategies for amelioration of memory disorders and neurodegeneration 16.
    For synaptic plasticity experiments, set the stimulus strength for all slices to 1 mV * and begin baseline stimulation with a frequency of 0.033 Hz. EPSP slope values ​​should be stable for at least 20 min before the induction of LTP or LTD. Closely monitor EPSPs during this time and reset the stimulus intensity, if the slope fluctuates more than 10% and start a new baseline. Induce LTP with a second puff of 100 Hz stimulation or multiple short bursts (~ 10 pulses) of 100 Hz stimulation given every 200 ms. For LTD induction, deliver 900 stimulus pulses at a rate of 1 Hz. After induction of LTP or LTD, collect synaptic responses for an additional 60 min or more. Determine EPSP slope values ​​before and 60 min after high / low frequency stimulation are present, compared to the presence of LTP or LTD.
    Aged rats and APP / PS1 mice tended to show deficient LTP and improved LTD (see Figure 5), and these changes have been suggested to lead to impaired cognition in these animal models 6,16 contribute. However, unlike APP / PS1 mice, changes in LTP / LTD in old rats are variable across laboratories. Aged rats usually exhibit similar LTP levels compared to adults in response to "intense" (ie 100 Hz) stimulation, but show deficits when milder stimulation parameters are used (ie lower stimulus frequencies or fewer stimulus pulses) (see for an overview 4,6, 16). Also, some laboratories, including ours, have observed increased susceptibility to LTD induction in older rats 2,17-19, while other groups found no difference or reduced sensitivity in their old animals. As briefly described above (see discussion), subtle but crucial differences in the experimental protocol may account for these discrepancies.
    * The intensity of the stimulation and EPSP amplitude can influence LTP induction and can be an important cause of divergent results in the literature. This will be taken into account in the discussion.

6. Representative results

Our work, and the work of other groups, suggests that changes in astrocyte-based inflammatory signaling can trigger and / or accelerate neurological dysfunction during aging and AD 13,20,21. Recently we used Synaptic Strength, LTP and LTD as endpoint measures to assess the efficacy and mechanisms of action of several novel anti-inflammatory reagents in middle aged APP / PS1 mice (see 22 for the description of this model) and investigate Fischer aged 344 rats. The results provided below were made using the protocols described in this article.

One of the new anti-inflammatory adeno-associated virus (AAV) reagents that has been developed by our laboratory has been shown in pilot studies to significantly increase synaptic strength (p <0.05) and prevent LTP deficits (p <0.05) in middle aged (16 months old) APP / PS1 mice (n = 4-6 slices per treatment condition). Representative synaptic strength curves and LTP experiments from two different slices, collected from the same 16-month old APP / PS1 mouse, are shown in Figure 5A-C. One slice was extracted from the hemisphere treated with our novel AAV (Reagent A), while the other slice was treated with a control AAV reagent (Control). LTP was induced in both discs with two 1 sec trains of 100 Hz stimulation (10 sec intertrain interval). Note that the synaptic strength curve for the reagent A-treated slices is shifted to the left of the control slice, indicating higher synaptic strength. Note also that typical of middle aged APP / PS1 mice, LTP rapidly disintegrate to baseline in the control slice (e.g. 23). Conversely, LTPs disintegrate little in the disc treated with our novel reagent.

In a second study, we observed significant LTD in vehicle-treated rats in old age (85% of the pre-LTD baseline, p <0.05). In contrast, no LTD was observed in old rats treated with novel anti-inflammatory "Drug A" (97% of the pre-LTD baseline, not significant). No drug effects on synaptic strength were observed. Representative LTD experiments from this dataset (n = 8-10 rats per group) are shown in Figure 5D-F.

Figure 1. Tools and materials used for brain dissection. A, paper towel. B, scalpel. C, Beebee scissors. D, bone rongeurs (for rats). E, bone rongeurs (for mice / rats). F, plastic spoon. G, plastic transfer pipette. H, hippocampus tool. I, spatula. J, surgical scissors. K, glass petri dish.

Figure 2. Custom brain slice receiving chamber. A, macrochamber. B, lid. C, H 2 O-reservoir with perforated silicone tube. D, microchamber. E, ACSF feed tube (polyethylene). F, O 2 Supply pipe. G, port for temperature control. H, to insert the netted microchamber.

Figure 3. RC22 submerged chamber. A, recording chamber. B, ground electrode. C, aspiration needle.

Figure 4. Hippocampal disc illustration and extracellular signals. A, cartoon of a transverse hippocampal section used in electrophysiology experiments. CA = Ammon's horn. DG = dentate gyrus. SC = Schaffer collaterals. S radiatum = stratum radiatum. B, electrical stimulation of the SC (a CA3 axonic tract) eliciting a stimulus artifact, followed almost immediately by a presynaptic population spike or fiber volley (FV). The amplitude of the FV is directly proportional to the number of SC fibers activated. The slope of the negative ongoing phase of the excitatory postsynaptic potential field (EPSP) is directly related to the activation of depolarizing synaptic currents in CA1 pyramidal neurons in response to glutamate discharge from the SC terminals. C, Overlapping representative extracellular signals in CA1 stratum radiatum in response to nine different stimulus levels (3-50 uA) recorded in a "healthy" (left picture), "unhealthy" and "hyper-excitable" slice. Five curves were averaged per level. Healthy discs respond dynamically to this stimulus area and exhibit a single positive ongoing population spike (reflective CA1 neural discharge) at the higher stimulus levels. In the RC22 sinking chamber, maximum EPSPs typically in the range of 1.5 to 3 mV amplitude. Unhealthy slices (middle) often show a large FV, but a small maximum EPSP (<1 mV) and usually poor plasticity. Hyper-excitable discs (right) show two or more regenerative population spikes in the ascending portion of the EPSP. Responses in hyper-excitable discs are often unstable and are variably influenced by LTD / LTP stimulation.

Figure 5. Representative electrophysiology experiments performed on acute sections from middle aged (16 mos) APP / PS1 mice and aged (22 mos) Fisher 344 rats. Panels AB show data collected from APP / PS1 mice treated with a control group adeno-associated (AAV) virally constructed construct (Control) or a novel AAV (reagent A) that was developed by our laboratory group. In relation to the control slice, the slice treated with reagent A shows a clear left shift in the EPSP: FV curve (A), which indicates higher synaptic strength. The reagent-A-treated slice also shows robust and stable LTP (B) after delivery of two 1 sec, 100 Hz stimulus puffs, while the control slice exhibits deficient LTP, which is typical of this animal model. Panels DF show data collected from two single old rats that received chronic (4 weeks) intrahippocampal perfusions of a vehicle or a novel anti-inflammatory drug (Drug A). Basal synaptic strength was relatively unaffected by drug treatment (D). However, Drug A was very effective in preventing the induction of LTD (E). Panels C and F show representative EPSP waveforms from individual slices recorded before (pre) and 60 min after (post) the delivery of LTP / LTD stimulation. Note that stimulus artifacts are not shown.


The steps outlined in this protocol will help ensure that brain dissections are performed at least as quickly and efficiently in years as they are in young adult rats. We also provide sufficient detail for the beginner to set up their own slice studies LTP and LTD. If further research into aging and AD changes in synaptic function and plasticity is one of your goals, there are at least two other methodological questions, indicated above, that deserve further consideration. Initially in various laboratories have shown that the Ca 2 +: Mg2 + ratio in the ACSF uptake has a marked effect on the induction of synaptic plasticity in the hippocampal discs 2,10,24,25 can. In mammalian cells CSF, the approx 2 +: Mg2 + ratio of around one (see e.g. 26). However, ACSF approx 2 +: Mg2 + ratios closer to 2 are typically used in slice studies of synaptic function and plasticity. In early studies, this practice probably was adjusted to optimize the induction of LTP, then became routine for all plasticity studies. However, this practice can be problematic in aging and AD studies because of the well-characterized differences in neural Ca 2+ regulation. More precisely, approx 2+ influx and / or approx 2+ -induced Approx 2+ version is in old rats and / or AD model mice during neuronal activation 3,27-31 elevated. Induction of LTD is particularly sensitive to subtle changes in ACSF Ca 2+ mirror. Our protocol, which is 2 mM Ca 2 + and 2 mM Mg2 +, often leads to LTD for older but not young animals 2, while studies with an approx 2 + uses: Mg2 + ratio closer to two, robust LTD have been observed in adults in the absence of an age difference 2,10 or in conjunction with reduced LTD in old rats 32. These observations underscore the need to carefully consider ACSF Ca 2 + and Mg2 + concentrations when comparing Ca 2+ dependent Plasticity in old and young adult animals.

The second methodological problem concerns the strong dependence of LTP on postsynaptic depolarization 33 and the aging / genotype differences in synaptic strength. In a typical LTP experiment, baseline, and LTP stimulation intensity is usually set to produce half the maximum (or three quarters maximum) EPSP amplitude. The potential problem is that aged rats and mice show APP / PS1 mostly reduced synaptic strength relative to their younger and / or wild-type counterparts, so that baseline EPSP values ​​are also smaller in old rats and APP / PS1 mice. Smaller EPSPs can translate to less depolarization during LTP stimulation, resulting in a decreased likelihood of LTP induction 33. Because of this potential mix-up, it is difficult to determine whether these animals have a throughput deficit, a plasticity deficit, or both. That is, LTP induction mechanisms in age and / or APP / PS1 mice functionally intact (no plasticity deficit), but not sufficiently stimulated (throughput deficit) under these conditions. This distinction is important because the mechanisms for throughput and mechanisms for plasticity can react very differently to a specific pharmacological treatment. We try to minimize the effects of lower throughput on LTP induction by normalizing the EPSP amplitude to the same level (e.g. 1 mV) across all slices prior to LTP stimulation. Other strategies may be effective as well (e.g., use of a voltage or current clamp to balance the membrane potential between groups during LTP stimulation), and should be considered when studying LTP in these animal models.


The production of this video article was sponsored by Leica Microsystems.


Work supported by NIH AG027297, an award from the Kentucky Spinal Cord and Head Injury Research Trust, and a gift from the Kleberg Foundation.


SurnameCompanyCatalog NumberComments
NaClFisher ScientificBP358-1
KClFisher ScientificBP366-500
KH2PO4 (monobasic)Sigma-AldrichP5379-100G
CaCl2 (dihydrates)Sigma-AldrichC3306-250G
NaHCO3Fisher ScientificS233-500
C.6H12O6 (dextrose)Fisher ScientificBP350-1
Table 1. Reagents required
Erlenmeyer FlasksFisher ScientificFB-500-2000 FB-500-1000
Aquarium bubblerUsed for oxygenating media. Available at most pet stores
50 mL glass beakerFisher Scientific02-540GFor brain storage in ACSF
ParafilmFisher Scientific13-374-10
Small animal guillotineWorld Precision Instruments, Inc.DCAP-M
Flat paper towel
# 11 Feather surgical bladeFisher Scientific08-916-5B
Beebee bone scissorsFine science tools16044-10
Lempert RongeursRoboz Surgical Instruments Co.RS-8321Use for rats
Friedman-Pearson RongeursFine science tools16020-14Use for mice or rats
Hippocampus toolFine science tools10099-15
SpoonA plastic teaspoon will do
SpatulaFisher Scientific21-401-25ASpatula
Surgical iris scissorsFine science tools14058-09
plastic transfer pipetsFisher Scientific13-711-43
110mm Whatman filter paperFisher Scientific09-805EWhatman cat. 1001-110
Glass petri dishFisher Scientific
Leica VT1000P Manual Vibrating MicrotomeVibratomeVT1000P
0.1mm FA-10 Feather S bladeTed Pella, Inc.121-90.1mm FA-10 Feather S blade
Borosilicate Glass Pasteur Pipet (with rubber bulb)Fisher Scientific13-678-20AFor transferring slices: Tip is broken off and heat-polished for larger opening
35 mm Polysterine Culture dishCorning430588Used for collecting slices after dissection
Table 2. Tools and materials for dissection
Holding chamberCustom made
P-97 Horizontal Pipette PullerSutter Instrument Co.
Vibration isolation tableTechnical Manufacturing Corp.
Faraday cageCustom made
Pyrex Aspirator Bottle with Bottom SidearmCorning1220-1L
Gravity-controlled IV set with regulatorBaxter International Inc.2C8891
Central Vacuum LineAvailable in most modern labs
95% O2 / 5% CO2 Gas mix Scott-Gross Co.
TygonTM Lab tubing For O2/ CO2 deliveryFisher ScientificNon-toxic, non-oxidizing, comes in a variety of sizes.
Eclipse E600FN MicroscopeNikon Instrumentswith 10x and 40x objectives, near infared filter, and GFP, DS-Red2 filters
Cool Snap ES Digital CameraPhotometricsCool Snap ES Digital Camera
X-Cite Fluorescent IlluminatorEXFOX-Cite Fluorescent Illuminator
Microscope PlatformSiskiyou, Inc.Custom assembled
RC-22 Submersible recording chamberWarner Instruments64-0228Requires P-1 platform and stage adapter (Product # 64-0277 From Warner)
TC2BIP 2 / 3Ch Temperature controllerCell microcontrols
4 Axis Manual Miniature manipulatorSiskiyou, Inc.
Platinum Iridium Wire (0.002 in)World Precision Instruments, Inc.PTT0203
A365 stimulus isolatorWorld Precision Instruments, Inc.A365 stimulus isolator
Multiclamp 700b amplifierAxon Instruments
Digidata 1322A A / D converterAxon Instruments
PClamp softwareAxon Instruments
Personal Computer (Pentium 4)Dell
Table 3. Electrophysiology equipment and materials



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