In Vivo Fiber-Coupled Pre-Clinical Confocal Laser-scanning Endomicroscopy (pCLE) of Hippocampal Capillaries in Awake Mice

Summary

Whereas multiphoton imaging is only effective at limited depths from the tissue’s surface, it is possible to achieve 3 µm resolution imaging at any depth via pCLE. Here, we present a protocol to conduct pCLE imaging to measure microvascular dynamics in the hippocampus of ictal and wild-type mice.

Abstract

The goal of this protocol is to describe fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE) in its specific application to elucidate capillary blood flow effects during seizures, driven by mural cells. In vitro and in vivo cortical imaging have shown that capillary constrictions driven by pericytes can result from functional local neural activity, as well as from drug application, in healthy animals. Here, a protocol is presented on how to use pCLE to determine the role of microvascular dynamics in neural degeneration in epilepsy, at any tissue depth (specifically in the hippocampus). We describe a head restraint technique that has been adapted to record pCLE in awake animals, to address potential side-effects of anesthetics on neural activity. Using these methods, electrophysiological and imaging recordings can be conducted over several hours in deep neural structures of the brain.

Introduction

In contrast with other microscopic imaging methods^{1,2,3,4,5,6,7,8}, in vivo fiber-optic-based confocal microscopy allows the measurement of blood flow dynamics in any brain region, at any depth, at high speed (up to 240 Hz depending on field-of-view size⁹). A fiber-optic probe enables in vivo confocal laser scanning imaging at 3 µm resolution because the tip of the probe (a lens-less objective made up of a bundle of 5000-6000 3 µm diameter individual fibers) can be positioned with a microelectrode’s accuracy, within 15 µm of the fluorescent target of interest. As with in vivo two-photon imaging, fluorophores must be previously introduced into the imaging target. For example, fluorescein dextran (or quantum dots) may be injected into the vasculature, or genetically-encoded fluorescent proteins can be transfected into cells, or fluorescent dyes such as Oregon Green BAPTA-1 can be bulk-loaded into cells, prior to imaging.

Recent research using these techniques has found that mural cell motor activity leading to ictal capillary vasospasms—sudden constrictions that occur at the position of the mural cells during seizures⁹—can contribute to neurodegeneration in the ictal hippocampus⁹. Whereas previous imaging studies showed in vitro and in vivo pericyte constrictions connected to drug applications^6,7,10,11,12, Leal-Campanario et al. found the first evidence of in vivo spontaneous capillary constrictions in the murine brain. To establish relevance to human temporal lobe epilepsy, they studied male (P30-40 old) knockout (KO) Kv1.1 (kcna1-null) mice^14,15 (JAX stock #003532), a genetic model of human episodic ataxia type 1¹⁵. Pericytes drove both pathological and physiological hippocampal mural vasoconstrictions⁹ in the spontaneously epileptic animals and their wild-type (WT) littermates. These observations were replicated in WT animals rendered epileptic with kainic-acid, thereby indicating their generalization to other forms of epilepsy. Leal-Campanario et al moreover determined, using novel stereological microscopy approaches, that apoptotic—but not healthy—neurons in epileptic animals were spatially coupled to the hippocampal microvasculature. Because excitotoxicity has no known spatial association to the vasculature, this result indicated that abnormal capillary vasospasmic ischemia-induced hypoxia contributes to neurodegeneration in epilepsy. Figure 1 shows a schematic of the general setup.

Protocol

The protocol follows the NIH Guidelines for the Care and Use of Laboratory Animals. All procedures were approved by the Barrow Neurological Institute’s Institutional Animal Care and Use Committee.

1. Stereotaxic positioning for craniotomy

1. Weigh and then anesthetize the mouse with a ketamine-xylazine (100 mg/kg–10 mg/kg i.p.) cocktail. Ensure that the animal is fully anesthetized by observing its lack of reaction to tail and/or toe pinch.
2. Place a heating pad under the mouse and use a rectal thermometer to continuously monitor its body temperature during the initial anesthetized implantation surgery (Figure 2A).
3. Apply ophthalmic ointment to prevent eye dryness.
4. Stabilize the head of the animal in a rodent stereotaxic frame, outfitted with a mouse adapter. To properly orient the mouse in the stereotaxic frame, place the animal’s teeth inside the hole of the bite-bar (Figure 2A and Figure 3A, item g). Tighten the nose-clamp until it is snug (Figure 2A,B and Figure 3A, item h). The mouse’s body positioning should be as straight as possible (as in Figure 2A).
   1. Further secure the animal’s head using mouse ear-bars with jaw holder cuffs (Figure 2A and Figure 3A, item i), to firmly clamp the zygomatic processes of the skull.
      NOTE: Once secured, the mouse’s head should have no range of horizontal motion.
5. Once the mouse is adjusted to the stereotaxic frame (Figure 3B, item j), clean and sterilize the scalp by wiping it with chlor-hexedine scrub 2-3 times, each time followed by an alcohol rinse. Then, apply topical lidocaine on top of the head.
6. Make an incision along the midline from posterior to anterior (start incision at the base of skull and complete it between the eyes; 12-15 mm) along the sagittal midline of the skull, using either fine scissors or a blade. Retract the borders of the scalp with four 28 mm bulldog serrefine clamps to expose the skull and maximize the working area (Figure 2B and Figure 3C).
7. Use a scalpel or microspatula (Figure 3C) to scrape the periosteum to the incision’s edges. Use tissue-separating scissor or forceps (Figure 3C) to carefully retract the muscles at the back of the neck (Figure 2B).
8. Dry the top of the skull with a cotton-tipped applicator, moistened with alcohol or 30% hydrogen peroxide, to optimize visualization of the cranial sutures. Apply saline to the skull once the connective tissue has been removed.
9. Once the mouse’s head is stabilized in the stereotaxic apparatus, place a syringe needle tip (21 G) in the electrode holder (Figure 3B, item l), and attach the electrode holder to the stereotaxic micromanipulator, lowering the needle tip to the level of the skull (Figure 2D).
10. Adjust the medial-lateral direction of the skull by measuring the height of the skull at any distance posterior to bregma (i.e., -2.0 mm), and two equal distances lateral from the midline on either side of the skull (i.e., ± 1.5 mm) (Figure 2D). Aim for a height difference across locations that is less than 0.05 mm in the dorsal-ventral direction.
    1. If greater than 0.05 mm, rotate the mouse’s skull using the bite bar, or reposition the mouse’s head by loosening the ear-bars carefully, laterally shifting the head and ear-bars as needed, and then retightening the ear-bars.
11. Once the skull is positioned in the stereotaxic device, determine and record the stereotactic coordinates of the bregma and lambda skull sutures along the anteroposterior (AP), medial lateral (ML), and dorsal ventral (DV) axes.
12. Identify the target point (right hippocampus) with the aid of a stereotaxic atlas. Then, find on the skull the target point coordinates relative to bregma (AP: -2.7 mm, ML: -2.7 mm) (Figure 2E).
13. Using a surgical marker, mark four corners of a 1.4 mm x 2 mm imaging window on the skull with dots, centered around the target point directly above the hippocampus (Figure 2F), for subsequent craniotomy.
14. Use the surgical marker to draw two additional dots: one over the top-left parietal bone (Figure 2G) and another over the right frontal bone (Figure 2H), to indicate the future placement of two bone screws (Figure 3A, item b) that will anchor the head-cap (Figure 3B, item m) to the skull.
15. Use the surgical marker to draw three more dots indicating where the epidural EEG recording electrodes will be inserted: one dot positioned 1 mm to either side of the midline, and one additional dot on the midline positioned 1 mm behind Lambda (Figure 2E, Figure 4A and Figure 5B).

2. Head-Cap installation

1. Using a 0.7 mm diameter burr (Figure 3C), carefully drill two small holes in the skull, at the dot positions indicating the bone screws placement (see step 1.15 above). Then, take two bone screws (stainless steel-slotted, length: 4 mm; diameter: 0.85 mm) previously disinfected with ethanol 70-80%, and screw them to the skull for additional head-cap stability (Figure 2I-J and Figure 3A, item b).
   1. Ensure that the screw tips do not protrude beyond the bottom of the bone into the skull cavity, risking damage to the brain. Note that one of the two bone screws will be anterior to the two head-cap screws, and the other posterior to them (Figure 2G-J).
2. Squirt saline over the skull to cool it down and clean it. Then dry the skull completely and apply a thin layer of cyanoacrylate around the screws.
3. Use a custom alignment piece (Figure 3A, item c) clamped to the microdrive mounted to the stereotaxic device (Figure 2F-J and Figure 3A, item d), so that the position of the head-cap is aligned along the midline, with the anterior ridge overlying bregma. This custom stainless-steel piece is L-shaped (angled at 90°), with two holes separated by 4 mm (Figure 2F-J and Figure 3A, item c).
   1. Use the stereotaxic manipulator to position the L-shaped alignment piece, with two Fillister Head Slotted drive screws attached (M1.6 x 0.35 Metric Coarse, 12 mm Length; Figure 3A, item a) over the bregma, until the base of the screws lies flat on the skull, being careful to not exert pressure on the skull (Figure 2I-J).
4. Apply cyanoacrylate followed by dental cement to attach the base of the screws to the skull. Be careful to not obstruct the imaging window reference marks over the hippocampus (Figure 2K).

3. Imaging window craniotomy surgery

1. At each of the positions of the craniotomy reference marks that indicate the corners of the imaging window (see step 1.14 above), drill a 0.7 mm diameter burr hole, followed by gently delineating the periphery of the 1.4 mm x 2 mm craniotomy window with the drill, with iteratively deeper cuts (Figure 2L).
2. Take care to not to drill too deeply or with too much force on the skull, which is thin and fragile, having a thickness of approximately 300 µm. Once the bone flap is complete, pry the loose flap up with the tip of a scalpel, being mindful to not damage the underlying dura mater.
3. Cover the open window with bone wax (Figure 2M).
4. Using a 0.9 mm diameter burr, drill three holes for future placement of the epidural EEG electrodes (see step 1.15 above) taking care to not to cut through the dura mater (Figure 2E,L).
5. Cover the three holes with bone wax.
6. Gently apply cyanoacrylate around the edges of the bone-wax covered imaging window and EEG holes, being careful to not seal the craniotomies (Figure 2L).
7. Build a wall of dental cement that encompasses the imaging window and EEG holes and binds it to the previously cemented head-cap (Figure 2M and Figure 3B, item m).
8. Let the cement dry for at least 10 minutes.
9. Close any loose wound margins with simple interrupted sutures, using 4-0 or 5-0 black braided silk (Figure 2N and Figure 3C).

4. Post-surgery

1. Use a cotton-tipped applicator to apply lidocaine ointment around the exposed skin to deaden sensation around the wound’s edge.
2. Use a sterile cotton-tipped applicator to apply Neosporin to the exposed skin.
3. Remove the animal from the stereotaxic frame (Figure 2N).
4. Inject ketoprofen (5 mg/kg) IP or IM for post-surgical analgesia.
5. Observe the animal until it is alert and moving (this will usually occur within minutes).
6. Place the animal in a warm cage and continue to monitor its recovery until it drinks or eats and is ready to transfer to animal housing.
7. Check on the animal at least once daily for approximately one week after the surgery, watching for any signs of listless behavior and/or inadequate eating/drinking (Figure 2O).

5. Dye injection and EEG recording

1. Wait a minimum of one day after the head-cap implantation surgery to initiate the recordings.
2. Place the mouse in a foam-restraining device molded to its body, within the stereotaxic frame (Figure 3B, items n & j).
3. Secure the head-cap (Figure 3B-I, item m) to a custom mounting bar which is attached to the stereotaxic frame (Figure 3A,B, items e & j, and Figure 4A). The long section of item e (Figure 3A,B) should have a length between 9.4 – 13 mm. Fix item f (a mounting plate with dimensions 1.5 cm x 0.5 cm, with two holes separated 4 mm from center to center) to item e (Figure 3A,B, items e, f and k). This alignment system will position the awake mouse securely to the stereotaxic frame throughout the imaging sessions (Figure 4).
4. Insert the epidural EEG recording electrodes (Figure 5A), placing the ground electrodes (white wires) within the posterior central hole, and the recording electrodes (red wires) in the anterior left and right holes (Figure 2E, Figure 4E and Figure 5E). Position each of the wires at an L-shape between the skull and the dura mater (Figure 5C) to optimize electrical contact.
5. Inject the tail vein with green fluorescein lysine-fixable dextran (0.2 mL/25 g body wt. of Fluorescein 5% w/w), to visualize blood flow within the hippocampus.
   1. To enhance dilation of the central vein of the mouse’s tail previous to injection, use one or more of the following strategies: a) Apply 70% ethanol to the central vein using a cotton-tipped applicator; b) Dip the tail into warm water (35-40 °C) for 1-3 minutes; c) Expose the tail to a heating lamp for a few minutes.
   2. Secure the mouse’s tail with two fingers and locate the central vein, which lies immediately below the skin. Insert the needle (ultrafine insulin syringe with integrated needle 30G), with the bevel up, into the vein, approximately halfway to two thirds from the base of the tail. Keep the needle parallel to the tail during the injection.
   3. Inject the dye slowly and steadily, taking care to avoid any changes in the position of the needle or tail.
      NOTE: There should be no resistance upon depressing the syringe plunger.
   4. After completing the injection, withdraw the needle and apply gentle pressure to the puncture site to seal the vessel.
   5. Allow clotting to occur.

6. In vivo fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE)

1. Directly following the tail vein injection (step 5.5.3 above), clamp the 300 µm beveled fiber-optic bundle (containing 5000–7000 3 µm fibers in each bundle) in the downward orientation to the mobile arm of the robotic stereotaxic drive.
2. Remove the bone wax from the craniotomy.
3. Remove the dura, using the bent tip of a syringe needle.
4. Using the robotic positioning system, first move the fiber objective to the appropriate anterior-posterior and left-right stereotaxic location, with the tip zeroed in the z-axis on the surface of the cortex.
5. Initiate the EEG recording (Figure 5D).
6. Initiate confocal laser-scanning of the back plane of the fiber-objective, followed directly by descending the objective slowly in the z-axis using the robotic microdrive’s z-controls, until it reaches the target depth within the brain (Figure 4C). View the imaging results in the microscopic software while descending.
   1. If the fiber tip is within the target X-Y-Z recording region, and two or more vessels enter the imaging window’s FOV, stop the descent and initiate video recordings. Obtain live full-field time-lapse movies of the blood flow at 11.7 Hz using Cellvizio Lab’s imaging software. (Higher speeds may be obtained with a smaller FOV). Recordings may take place for 4-5 hours at a time, in consecutive days if needed.
7. Using a 10 mL syringe with beveled silicon tubing fixed to the tip of the needle, feed the animal during the recordings by periodically providing paste made with mice pellets ground with water.
8. At the end of the imaging session, terminate the recordings.
9. Gently remove the EEG leads and clean them with Tergazyme.
10. Slowly remove the fiber-optic bundle from the animal’s brain by reversing the fiber along the z-axis of entry.
11. Release the animal from the head post and body restraints.
12. If appropriate, follow the necessary euthanasia and tissue processing protocols to visualize blood vessels and to perform immunohistochemistry.
13. If planning to record from the same animal in a future session, cover the imaging window with bone wax or silastic.

Results

We developed these methods to assess whether abnormal pericyte-driven capillary vasospasms in the hippocampus—occurring as the result of seizures—could cause frank hypoxia that contributes to cell death in the ictal focus^9,13.

The development of the head-cap and its proper installation afforded high-stability in the recordings, allowing simultaneous recording of EEG and blood flow deep in the hippocampus of wild-type and epileptic awak...

Discussion

We developed a head-cap restraint system for simultaneous electrophysiological and fiber-optic pCLE experiments in awake mice, reducing potential response contamination due to anesthetic drugs. The head-cap and mounting apparatus are straightforward to construct and are reusable for chronic awake-behaving imaging experiments. We checked the quality of the recordings against the gold-standard for in vivo microscopic blood flow imaging, TPLSM.

Proficient surgical skills are necessary to implemen...

Materials

List of materials used in this article

Name	Company	Catalog Number	Comments
0.7 mm diameter burr	Fine Science Tools	19007-07	For Screws No. 19010-00
0.9 mm diameter burr	Fine Science Tools	19007-09
ASEPTICO AEU-12C TORQUE PLUS	from Handpiece solution	AEU12C
Bull dog serrifine clump	Fine Science Tools	18050-28
CellVizio dual band	Mauna Kea Technologies
CellVizio single band	Mauna Kea Technologies
Confocal Microprobe 300 microns (Serie S)	Mauna Kea Technologies
Custom-made alignment piece	L-shaped (angled at 90 deg) and made of stainless steel with two holes drilled on it, with a 4 mm separation from center to center
Custom-made mounting bar	The long section piece of the mounting bar should be between 9.4 - 13mm. Fixed to this piece of the mounting bar, position a stainless-steel plate 1.5 cm long and 0.5 cm wide that has two holes drilled separated 4 mm from center to center, the same distance that the L-shaped alignment piece.
Cyanoacrylate adhesive-Super Glue
Dumont forceps #5	Fine Science Tools	11252-20
DuraLay Inlay Resin – Standard Package	Reliance Dental Mfg Co.	602-7395 (from patterson dental)
Fillister Head, Slotted Drive, M1.6x0.35 Metric Coarse, 12mm Length Under Head, Machine Screw	MSC industrial direct co.	2834117
Fine Point scissor	Fine Science Tools	14090-09
Fluorescein 5% w/w lysine-fixable dextran (2MD)	Invitrogen, USA	D7137
Halsey smooth needle holder	Fine Science Tools	12001-13
Kalt suture needle 3/8 curved	Fine Science Tools	12050-03
lab standard stereotaxic, rat and mouse	Stoelting Co. 51704	51670
Methocel 2%	Omnivision GmbH	PZN: 04682367	Eye ointment to prevent dryness.
Mouse Temperature controller, probe (YSI-451), small heating pad-TC-1000 Mouse	CWE Inc.	08-13000
PhysioTel F20-EET transmitters	DSI	270-0124-001
Robot Stereotaxic, Manipulator Arm, ADD-ON, 3 Axis, LEFT	Stoelting Co.C13	51704
Sel-Tapping bone screws	Fine Science Tools	19010-10
Standard Ear Bars and Rubber Tips for Mouse Stereotaxic	Stoelting Co	51648
Suture Thread - Braided Silk/Size 4/0	Fine Science Tools	18020-40
Tissue separating microspatula	Fine Science Tools	10091-121