Stereotaxic Infusion of Oligomeric Amyloid-beta into the Mouse Hippocampus

Summary

Here, we present a protocol for direct stereotaxic brain infusion of amyloid-beta. This methodology provides an alternative in vivo mouse model to address the short-term effects of amyloid-beta on brain neurons.

Abstract

Alzheimer’s disease is a neurodegenerative disease affecting the aging population. A key neuropathological feature of the disease is the over-production of amyloid-beta and the deposition of amyloid-beta plaques in brain regions of the afflicted individuals. Throughout the years scientists have generated numerous Alzheimer’s disease mouse models that attempt to replicate the amyloid-beta pathology. Unfortunately, the mouse models only selectively mimic the disease features. Neuronal death, a prominent effect in the brains of Alzheimer’s disease patients, is noticeably lacking in these mice. Hence, we and others have employed a method of directly infusing soluble oligomeric species of amyloid-beta - forms of amyloid-beta that have been proven to be most toxic to neurons - stereotaxically into the brain. In this report we utilize male C57BL/6J mice to document this surgical technique of increasing amyloid-beta levels in a select brain region. The infusion target is the dentate gyrus of the hippocampus because this brain structure, along with the basal forebrain that is connected by the cholinergic circuit, represents one of the areas of degeneration in the disease. The results of elevating amyloid-beta in the dentate gyrus via stereotaxic infusion reveal increases in neuron loss in the dentate gyrus within 1 week, while there is a concomitant increase in cell death and cholinergic neuron loss in the vertical limb of the diagonal band of Broca of the basal forebrain. These effects are observed up to 2 weeks. Our data suggests that the current amyloid-beta infusion model provides an alternative mouse model to address region specific neuron death in a short-term basis. The advantage of this model is that amyloid-beta can be elevated in a spatial and temporal manner.

Introduction

Amyloid plaque deposits, which are composed of amyloid-beta (Aβ_1-42), are a key feature of the pathology of Alzheimer’s disease (AD). Numerous studies have shown that high or toxic levels of recombinant oligomeric Aβ_1-42 elicit neuronal death, synaptic dystrophy, loss and dysfunction; as well as learning and memory deficits^1-4. Brain regions affected include the hippocampus, the cortex, and subcortical structures such as the basal forebrain and the amygdala^5,6. To date, there are multiple transgenic mouse models that attempt to simulate the Aβ_1-42 pathology of AD. Depending on the strain these animals prove to be useful in examining select pathological features of AD. Unfortunately, with the exception of 2 transgenic lines, APP23 and 5XFAD, these mice never fully replicate neuronal loss, a key aspect of AD. Even with the neuronal loss observed in APP23 and 5XFAD, the neuronal death observed was subtle, age dependent, and isolated to a few select regions^7,8.

The direct infusion of oligomeric Aβ_1-42 into the wild-type mouse brain provides an excellent in vivo model which replicates the neuronal death aspect of amyloidopathy^1,9,10. Unlike the commonly utilized transgenic mouse models the oligomeric Aβ_1-42 infusion model is ideal for acutely elevating Aβ_1-42 levels in a spatial and temporal manner. The advantage of using wild-type mice for this model obviates potential compensation or side effects from the mutations introduced in transgenic mouse lines. Past studies have shown that infusing toxic levels of Aβ_1-42 into the hippocampus elicits neuron death in the vicinity of the injection site within 1 week¹. Moreover, consistent with the observation that Aβ_1-42 is toxic for cholinergic neurons¹¹ the basal forebrain cholinergic neuron (BFCN) population which projects to the hippocampus is decreased 20-50% within 7-14 days following beta-amyloid infusion^1,10 in mice, effectively allowing for the examinations of isolated neuronal circuitry in the brain. Since BFCN project ipsilaterally to the dentate gyrus of the hippocampus¹², for the most part control/vehicle and oligomeric Aβ_1-42 solutions can be injected on either side of the brain allowing comparisons to be made between the left and right hemispheres¹.

In this report we will provide a detailed surgical and injection methodology for adult wild-type C57BL/6J mice. This mouse strain is chosen because of its wide use in research. Technically, any brain region can be targeted for infusion, however here we will use the dentate gyrus of the hippocampus as the target to illustrate the technique.

Protocol

Note: For all animal experimentation, Institutional and National guidelines for the care and use of laboratory animals were followed.

1. Prepare Surgical Instruments and Solutions for Surgery

1. Autoclave all stainless steel surgical instruments.
2. Prepare 70% ethanol by diluting 200 proof absolute ethanol with sterile molecular grade deionized distilled water.
3. Attach the 29 G needle to the Hamilton syringe. Clean the interior of Hamilton syringe and needle by drawing up and ejecting nanopure water repeatedly for 1 min. Repeat this procedure with 70% ethanol.
   1. Remove the plunger and needle from the Hamilton syringe. Air dry the parts in a laminar flow hood O/N.
   2. Irradiate the needle and syringe with ultraviolet light for 30 min before use.
4. Prepare saline solution by dissolving NaCl in molecular grade water to a final weight to volume concentration of 0.9%. Sterilize the solution by filtering it through 0.2 μm pore filter.

2. Prepare Oligomeric Aβ_1-42

1. Monomerize and resuspend recombinant human Aβ_1-42 in DMSO to 5 mM exactly as described in the publication of Fa' and others¹³ .
2. Dilute 5 mM Aβ_1-42 with sterile (1x) PBS to 100 µM on the day before surgery.
3. Mix the solution well by trituration.
4. Incubate Aβ_1-42 solution for 12 hr at 4˚C.
   Note: Control solutions such as diluting DMSO in PBS or scrambled/reverse Aβ_1-42 peptide should be prepared the same way as Aβ_1-42.

3. Determine Injection Coordinates

1. Use the adult mouse brain atlas to determine the exact anterior-posterior (AP), medial-lateral (ML), and dorsal-ventral (DV) coordinates for the brain region of interest¹⁴.
   ​Note: To illustrate the technique we picked the dentate gyrus of the hippocampus as the target with the following coordinates from the bregma: AP, -2.00 mm; ML, ±1.3 mm; DV, -2.2 mm. The negative sign preceding the AP value indicates that it is 2.00 mm posterior of the bregma. The ± sign preceding the ML value indicates left and right direction from the center. Lastly, the negative sign preceding the DV indicates it is moving ventrally from the surface of the brain.

4. Stereotaxic Frame Setup

1. Wear a surgical face mask, clean lab coat, and sterile surgical gloves.
2. Wipe down stereotaxic instrument and mouse adaptor with 70% ethanol.
3. Lay down sterile surgical drape on the counter.
4. Place the stereotaxic instrument with mouse adaptor on top of the sterile surgical drape.
5. Wipe the mouse heating pad with 70% ethanol. Position the heating pad over the stereotaxic frame bed. Use general purpose laboratory labeling tape to tape down the heating pad over the mouse adaptor bed.
   1. Attach the heating pad to the warm water recirculator pump according to manufacturer’s directions.
   2. Turn on the warm water recirculator pump and set the temperature to 37 °C, and then wait until it reaches temperature.
6. Affix the 50 µl Hamilton syringe with a 29 G needle onto the stereotaxic frame by screwing it onto the motorized vertical injecting shaft according to manufacturer’s directions.
7. Turn on the bead sterilizer and wait until it reaches the manufacturer’s preset temperature.
   Note: This is used for sterilizing stainless steel instruments between animals. It takes instruments 20 sec of contact with the beads for sterilization.
8. Turn on hot plate and set it to 42 °C and place a clean empty cage atop.
9. While the Hamilton syringe is fixed on the stereotaxic frame use the motorized stereotaxic injector to draw up the Aβ_1-42 solution.
   ​Note: Draw up more than 4 μl of the solution into the syringe and then use the stereotaxic injector to set the injecting volume. Operate the injector according to manufacturer’s instructions.

5. Animal Preparation

1. Determine the weight of the mouse using the weigh scale.
2. Anesthetize mouse with ketamine/xylazine cocktail at 100 mg/kg ketamine and 10 mg/kg xylazine by injecting intraperitoneally (IP). Monitor the depth of anesthesia by the loss of toe pinch reflex.
3. After the mouse is sedated take the hair clipper and shave its head to expose the skin over the skull.
4. Place the mouse on top of the heating pad on top of the stereotaxic bed.
5. Use the spatula to open the mouth and place the incisor teeth inside the teeth guard. Secure the nosepiece over the face of the mouse. Clamp it down gently and not too tight.
6. Position the bilateral ear crossbars into auditory meatus to secure the head. Move each crossbar in until it hits the skull, and then turn the screw to lock it.
   1. To make sure the head is secured use your index finger to gently push down on the head. If the head is properly secured it will not give out or move when pressed.
7. Apply a drop of eye cream or eye drop to the eyes to keep them moist. Make sure the eyes are moist throughout the surgical procedure.
8. To disinfect the surgical site use sterile cotton swabs to apply betadine solution to the skin over the skull, followed by 70% ethanol. Perform the alternating betadine and 70% ethanol cleaning 2 more times.

6. Surgery and Infusion

1. Use a scalpel to make a 2-3 mm incision in the midline of the scalp, and then use a straight fine scissors to extend the incision line to 1.0-1.5 cm to expose the sagittal suture, bregma, and lambda of the skull, landmarks which the stereotaxic coordinates are based on. Use micro clamps to keep the skin apart.
   Note: The bregma and lambda positions are explained in the mouse brain atlas “The Mouse Brain in Stereotaxic Coordinates”¹⁴.
2. Take a cotton swab, soak it in sterile saline, and use it to clean the skull. Then, use a clean dry cotton swab to dry the skull.
3. Use a sterile fine point pen to mark a dot on the bregma. Use the stereotaxic micromanipulator to position the Hamilton syringe so that the tip of the needle just touches the dot on the bregma.
   1. Record the DV coordinate.
   2. To check if the AP axis of the skull is level move the Hamilton syringe posteriorly so that the tip of the needle touches the lambda point.
   3. Record the DV position. Make sure the DV coordinates for bregma and lambda are within 0.5 mm.
      Note: The goal is to minimize the DV difference between bregma and lambda.
4. Use the micromanipulator to return the Hamilton syringe needle tip back to the bregma dot.
5. Record the starting AP position.
6. Calculate the ending AP position by subtracting 2.00 mm from the starting AP coordinate.
7. Use the micromanipulator to reposition the Hamilton syringe needle to the final AP coordinate.
8. Record the current ML coordinate.
9. Calculate the left and right ending ML coordinates by adding or subtracting 1.3 mm from the starting ML coordinate.
10. Use the micromanipulator to move the Hamilton syringe needle to either the ending ML coordinate.
    1. Touch the needle tip to the surface of the skull on both ending ML coordinates and record the DV coordinates and make sure the values are within 0.5 mm.
       Note: The goal is to minimize the DV difference between the two ending ML coordinates.
    2. While at an ending coordinate pull the needle up 0.5-1 cm over the skull. Take a sterile fine point pen to mark a dot on the surface of the skull. Repeat this step for the contralateral side of the skull.
11. Swing the Hamilton syringe clear out of the way.
12. Attach 0.8 mm drill head to the drill. Take the drill with both hands, set elbows on the surface of the table for stability, and position the drill head slightly above the sharpie dot on either the left or right ML coordinate. Activate the drill, lower the drill tip onto the skull surface to introduce a hole in the skull. Repeat this step for the contralateral side.
    Note: Some bleeding may occur from the newly introduced holes. If there is bleeding then use a clean dry cotton swab to dab the blood.
13. Move the Hamilton syringe back into position over either of the newly introduced ML holes. Lower the Hamilton needle just past the skull. At this point be careful not to puncture the brain. To make sure the needle has passed the skull take your index finger and gently push the needle against the skull to make sure the needle doesn’t exit the hole.
14. Record the starting DV coordinate.
15. Calculate the ending DV coordinate by subtracting 2.2 mm from the starting DV coordinate.
16. Lower the needle down to the ending DV coordinate.
17. Activate the stereotaxic injector pump to pump 4 µl of Aβ_1-42 into the dentate gyrus at a rate of 0.5 µl/min.
18. When the infusion is complete let the needle remain in place for an additional 1 min to minimize backflow of solution out of the injection site.
19. Move the needle to the other side of the brain and repeat steps 6.13-6.18.

7. Animal Removal and Postoperative Care

1. Unscrew the ear bars and face/nose guard. Remove the mouse from the apparatus.
2. Use a student standard pattern forceps to pull close the scalp and then seal the wound with suture.
3. Inject 1 ml of sterile saline into the mouse via IP for hydration.
4. Inject buprenorphine (0.1 mg/kg) subcutaneously to relieve pain.
5. Maintain the mouse in the clean empty cage set on top of 42 °C hot plate until it wakes, then return it to its housing cage with ample food and water.
6. Administer buprenorphine via subcutaneous injection every 12 hr for 3 days for pain relief.

8. Suture Removal

1. Anesthetize mouse with ketamine/xylazine cocktail at 100 mg/kg ketamine and 10 mg/kg xlyazine by IP. Note: This is done 7-10 days after surgery.
2. Use a student standard pattern forceps and straight fine scissors to remove the suture.
3. Inject 1 ml of sterile saline into the mouse via IP for hydration.
4. Maintain the mouse in a clean empty cage set on top of 42 °C hot plate until it wakes, and then return it to its housing cage with ample food and water.

9. Animal Sacrifice and Tissue Processing

1. Anesthetize the mouse and perfuse it with 4% paraformaldehyde¹⁵, remove the brain, and process it for cryosectioning¹⁶.
   Note: The timing of animal sacrifice depends on the experimenter. However, for our studies we chose 7 and 14 days post surgery.

Results

The present method of preparing human recombinant oligomeric Aβ_1-42 yields soluble oligomeric species consisting of monomers, dimers, trimers, and tetramers (Figure 1A). These low molecular weight Aβ_1-42 species, but not the fibrils and plaques, have been shown in numerous settings to be most toxic to neurons^1,4,9,17-19. To determine whether or not oligomeric Aβ_1-42 induces neuron death in the mouse brain Aβ_1-42 (4 µl of 100 µM stock solution) was in...

Discussion

To achieve a successful Aβ_1-42 injection the experimenter or surgeon must: 1) use aseptic technique; 2) correctly identify the brain region of interest with accurate coordinates; 3) be able to properly secure the mouse in the stereotaxic frame with the brain leveled in the AP and ML axis; 4) have the ability to operate the micromanipulator with precision; 5) ensure proper post-operative care. If these key steps are followed the mouse should survive the surgery with no observable infection.

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
Ketamine HCl (100 mg/ml)	Henry Schein Medical	1049007	100 mg ketamine per 1 kg animal
Xylazine (20 mg/ml)	Henry Schein Medical	not available	10 mg xylazine per 1 kg animal
Buprenex (0.3 mg/ml)	Henry Schein Medical	1217793	0.1 mg buprenex per 1 kg animal
Aβ1-42	David Teplow/UCLA	not available	100 μM; This amyloid was used in the paper
Aβ1-42	Bachem	H-1368	Can be used in place of amyloid from the Teplow lab
Aβ1-42	American Peptide	62-0-80B	Can be used in place of amyloid from the Teplow lab
Scrambled Aβ1-42	American Peptide	62-0-46B	Can be used as control peptide for comparing Aβ1-42
NU4 Antibody (Oligomeric Amyloid Antibody)	Gift from William Klein/Northwestern U.	not available	1:2,000 dilution
Anti-Amyloid Oligomeric Antibody  (Polyclonal Rabbit)	EMD Millipore	AB9234	May be used in place of Nu4; needs to  be tested by the end user
6E10 Antibody (Monoclonal Mouse) (Amyloid Antibody)	Biolegend	sig-39320	1:1,000 dilution
ChAT Antibody (Polyclonal Goat)	Millipore	AB144P	1:100 dilution
DeadEnd Fluorometric TUNEL system	Promega	G3250	Follow manufacturer's directions for use
Prolong Gold Antifade Reagent with DAPI	Invitrogen	P36935	Use when coverslipping slides
Fluorogold	Fluorochrome, LLC	not available	2% solution
Absolute Ethanol (200 proof)	Fisher Scientific	BP2818-4	For making 70% ethanol for sanitizing and disinfecting
Novex 10-20% Tricine gel	Life Technologies	EC6625BOX	For separating Aβ1-42
Novex Tricine SDS Running Buffer (10X)	Life Technologies	LC1675	For running 10-20% Tricine gels
Novex Tris-Glycine Transfer Buffer (25X)	Life Technologies	LC3675	For transferring 10-20% Tricine gels
SuperSignal Western Blot Enhancer	Thermo Scientific	46640	For enhancing Aβ1-42 signal; follow manufacturer's protocol
Protran BA79 Nitrocellulose Blotting Membrane, 0.1 μm	GE Healthcare Life Sciences	10402088	For transferring 10-20% Tricine gels
Xcell SureLock Mini-Cell	Life Technologies	EI0001	Electrophoresis aparatus for running 10-20% Tricine gels
GenTeal Lubricant Eye Gel	Novartis	not available	For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores
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Refresh Optive Lubricant Eye Drops	Allergan	not available	For keeping the mouse eyes moist during surgery; can be found in local pharmacy stores; Can be used in place of GenTeal
Betadine	Stoelting	50998	For sanitizing and disinfecting
Round/Tapered Spatula	VWR	82027-490	For opening animal mouth
Bulldog Serrefines Clamps (Jaw Dims. 9X1.6 mm; Length 28 mm)	Fine Science Tools	18050-28	Optional; For keeping scalp skin apart during injection
Straight Fine Scissors (Cutting edge 25 mm; Length 11.5 cm)	Fine Science Tools	14060-11	For cutting scalp
#3 Scalpel Handle	Fine Science Tools	10003-12
#11 Surgical Blade	Fine Science Tools	10011-00	For making scalp incision
Student Standard Pattern Forcep (Tip Dims. 2.5x1.5 mm; Length 11.5 cm)	Fine Science Tools	91100-12	For holding scalp closed during suturing
Trimmer Combo Kit	Kent Scientific	CL9990-1201	For shaving hair
T/Pump Warm Water Recirculator	Kent Scientific	TP-700	For warming animal during surgery
Resusable Warmining Pad (5" x 10")	Kent Scientific	TPZ-0510FEA	For attaching it to the T/Pump warm water recirculator to warm the animal during surgery
Cordless Micro Drill	Stoelting	58610	Use 0.8 mm steel burrs to drill holes in the skull
Lab Standard Stereotaxic Instrument with Mouse & Neonatal Rat Adaptor	Stoelting	51615
Just for Mouse Stereotaxic Instrument	Stoelting	51730	Can use this in place of Stoelting Cat. #51615
Quintessential Stereotaxic Injector	Stoelting	53311
Dry Glass Bead Sterilizer	Stoelting	50287	For sterilizing stainless steel instruments
Sterile Surgical Drape (18" x 26")	Stoelting	50981
Hamilton Syringe 50 ml, Model 705 RN SYR, NDL	Hamilton Company	7637-01	For brain injection; use different syringes for different solutions
29 G Needle, Small Hub RN NDL	Hamilton Company	7803-06	For attaching to the Hamilton syringe for brain injection
1 ml BD Tuberculin Syringes	VWR	BD309659	For administering anesthesia and saline
30 G Needle (0.5")	VWR	BD305106	For administering anesthesia and saline
Portable Electronic CS Series Scale (Ohaus)	VWR	65500-202	For weighing animals to determine anesthesia dose
Hot plate (Top Plate Dims. 7.25x7.25 in)	VWR	47751-148	For warming animals post-surgery
Sofsilk Silk Suture C-1 Cutting 3/8, 12 mm	Covidien	S1173	For closing wound
Vetbond Tissue Adhesive (3M)	Santa Cruz Biotechnology	sc-361931	Optional: for aiding in wound closure; Use with suture.
Cotton-Tipped Wooden-Shaft Sterile Applicators	Fisher scientific	22-029-488	For cleaning and drying surgical wound
Fisherbrand Superfrost Plus Microscope Slides	Fisher Scientific	12-550-15	For collecting brain sections
VWR Micro Cover Glass 24 X 50 mm	VWR	48393241	For mounting microscope slides
Thermo Scientific Nalgene Syringe Filter 0.2 μm	Fisher Scientific	194-2520	For sterilizing saline solution
Sterile dual tip skin markers by Viscot Medical	Medline	VIS1422SRL91	For marking coordinates on the skull