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.
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.
In contrast with other microscopic imaging methods1,2,3,4,5,6,7,8, 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 applications6,7,10,11,12, 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) mice14,15 (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.
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
2. Head-Cap installation
3. Imaging window craniotomy surgery
4. Post-surgery
5. Dye injection and EEG recording
6. In vivo fiber-optic-bundle-coupled pre-clinical confocal laser-scanning endomicroscopy (pCLE)
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 focus9,13.
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 awake mice during both ictal and interictal periods. Capture of blood flow events related to seizures requires recording for extended periods of time, so that aperiodic blood flow events (such as vasospasms) can be captured in response to both induced and naturally occurring epileptic seizures. The restraint system permitted stable blood flow recordings of deep brain microvessels and their proximal mural cells over long hours (Figure 6A-R). We found that, in entire microvessels, blood flow stoppages occurred at the positions of labelled mural cells.
We reliably located vessels in the hippocampus using a robotic stereotaxic device targeted with internal coordinates from a standard stereotaxic atlas. We verified the quality of the fiber recordings by comparing them to high-resolution two-photon imaging recordings of blood flow and equivalent vasospasms from cortical tissue, at depths that two-photon imaging can reach (Figure 6S-AA). We further verified the pCLE results using immunohistochemistry at recording sites, to show with traditional confocal microscopy that hippocampal mural cells were targeted correctly and that they were not only constricting but also spatially associated with strictures in microvessels far from arterioles15 (Figure 7).
The published findings with these methods indeed show abnormal hippocampal capillary vasospasms driven by seizures, as well as correlated cell death in Kv1.1 epileptic mutant mice and kainate model epileptic mice9,13. These results indicate a role for local ictal ischemia/hypoxia in cell death during seizures, due to abnormal vasospasms, and contribute to a growing body of work that expands on the potential mechanisms of ictal cell death.
Figure 1: Schematic representation of the experimental design. We recorded EEG while conducting confocal microscopy in hippocampal capillaries of both awake epileptic mice and WT littermates. After injecting the tail vein with fluorescein, we used a novel preclinical Confocal Laser-scanning Endomicroscopy (pCLE), with a fiber-objective with a tip diameter of 0.3 mm to record capillary blood flow dynamics deep in the brain. Yellow indicates increased blood flow, and red indicates steady or reduced flow. Please click here to view a larger version of this figure.
Figure 2: Protocol for head-cap implantation surgery. A) The anesthetized animal was placed on a heating pad and positioned within the stereotaxic frame (see Figure 3B, item j) and secured with mouse ear-bars (Figure 3A, item i) and jaw holder cuffs (Figure 3A, items g,h), over a heating pad. B) Exposure of the cranial sutures. C-D) Measurements of Bregma and Lambda. E) Visualization of drilling locations over the animal’s skull. The black points and square indicate the window over the hippocampus that will permit the pCLE insertion. The two red circles mark the insertion positions for the EEG recording wires. The white dot marks the insertion point of the EEG ground wires. F) The four black dots on the skull mark the future corners of the imaging window craniotomy over the hippocampus. The custom alignment piece (Figure 3A, item c), with the headpost implants (Figure 3A, item a) attached to the stereotactic microdrive positioner (Figure 3A, item d). G-H) Two additional dots, one posterior (G) and one anterior (H) mark the future location of head-cap anchor screws (Figure 3A, item b). I-J) Two small screws anchor the head-cap to the skull (Figure 3A, item b). K) Cyanoacrylate and dental cement are applied to the anchor screws from panels I & J. L) The imaging window craniotomy over the hippocampus, and the three holes for the EEG electrodes, have been drilled. M) The imaging chamber is built (Figure 3B, items i, m) by sculpting a berm with additional dental cement to encircle the three EEG holes and the imaging window. N) Overhead view of the finished head-cap. O) The recovered mouse back in its cage after the implantation surgery. Please click here to view a larger version of this figure.
Figure 3: Materials used in the Protocol (specific items enumerated with lower-case letters). A) Materials used to build the head-cap. (a) Two M1.6 x 12 mm machine screws (the headpost implants). (b)Two small bolts to secure the head-cap to the skull. (c) A custom L-shaped stainless-steel alignment piece, with two holes 4 mm apart, which secures the two M1.6 screws during the surgery. (d) A standard microdrive mounted on a stereotaxic device holds item c. (e) A custom mounting bar with an (f) alignment piece that matches the two holes from item c. (g) A bite bar and (h) nose clamp to stabilize the position of the animal’s head. B) Setup during the recordings. (i) The head-cap is secured to item f during the anesthetized implantation surgery. (j) A stereotaxic frame positions the microdrive (item d). (l) A 21G needle is attached to the microdrive (item d) and used to determine the positions of the Bregma and Lambda cranial sutures. (m) A simulated head-cap is pictured here, attached to item f, to demonstrate the positioning of the mouse in (n) the foam-lined restraining-tube (mouse not present). Item n molds itself to the shape of the mouse's body. C) Instruments and supplies needed to perform the surgery. Please click here to view a larger version of this figure.
Figure 4: Recording session setup. A) The animal’s head is fixed. B) Location of the EEG recording wires. White EEG wires are placed separately, in each side of the skull. Red EEG wires are placed together in the middle-bottom (ground) hole. C) The fiber-optic bundle is inserted into the target window. Please click here to view a larger version of this figure.
Figure 5: EEG recordings. A) Epidural EEG recordings are conducted with Physiotel F20-EET transmitters, simultaneously to the fiber-optic pCLE recordings in the awake animal. B) Location of the drilled holes for insertion of the leads for EEG recordings. The two red leads go to the two holes indicated by red dots. The two white leads are inserted together in the hole indicated by the white dot. The red square designates the target window over the animal’s hippocampus. The letter ‘B’ on the mouse’s skull indicates the bregma. C) The EEG recording wires are bent in an L-shape to enhance electrical contact between the skull and the dura matter. D) Examples of EEG recordings in awake mice. Please click here to view a larger version of this figure.
Figure 6: Recordings of mural cell vasoconstrictions in mice. A-R) In vivo fiber-optic dual-band pCLE image of capillary vasospasm (vessels in green, labeled with 2MD fluorescein-conjugated dextran) colocalized to mural cells (red, labeled via intravenous vein tail injection of Alexa Fluor 647) in awake knockout mice during seizure (arrow indicates mural cell and associated vasospasm). See Video 1 and Video 2. S-W) In vivo two-photon scanning laser microscopy (TPLSM) stack of capillary vessels (green) and mural cells (red) of a wild type mouse. See Video 3. X-Z) In vivo TPLSM of a mural-cell-localized (red) capillary constriction of kainic mice. See Video 4. AA) Quantification of vessel constriction from panels X-Z measured at white line. Scales for panels A-R= 5 µm; S-W= 25 µm; X-Z= 5 µm. Videos were recorded at 11.7 Hz. From Leal-Campanario et al.8. Please click here to view a larger version of this figure.
Figure 7: Immunohistochemistry of the recorded brain. A-B) Vessels at different magnifications injected with fluorescein dextran (green). DAPI-stained cellular nuclei in blue (white arrows). Scale bar for A= 200 μm; scale bar for B = 100 μm. C) Different section of the brain injected with fluorescein where a red blood cell (red arrow) is clearly seen. Scale bar = 50 μm. Please click here to view a larger version of this figure.
Video 1: Vasospasm and mural cell blood flow recorded (11.7 Hz) in a KO hippocampal capillary. See frames from the same recording in Figure 6A-H. Please click here to download this video.
Video 2: Vasospasm, leukocyte blockages, blood flow and mural cells recorded in a WT hippocampal capillary (11.7 Hz). See frames from the same recording in Figure 6I-R. Please click here to download this video.
Video 3: Non-uniform blood flow in in vivo WT mouse parietal cortex recorded with TPLSM. See frames from the same recording in Figure 6S-W. Please click here to download this video.
Video 4: Mural cell-driven (red) capillary (green) constriction from parietal cortex of seizing kainic animal recorded with TPLSM. See frames from the same recording Figure 6X-Z. Please click here to download this video.
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 implement the protocol we describe here. The surgery must be done under aseptic conditions and always under an operating microscope, while taking care to leave the dura mater intact when drilling the window over the hippocampus. Familiarity with the underlying vasculature is essential, as cutting above a major blood vessel creates the risk of undesirable bleeding.
Placing the animal correctly in stereotaxic alignment ensures that the anterior-posterior plane is level before attaching the head-cap. This safeguard facilitates locating brain loci with the help of a brain atlas.
Mouse motion can result in damage to vessels or erroneous recordings of vasospasms (that is, fiber motion rather than true vasomotion), so it is important to reduce animal motion during the recordings. Foam padding within the restraint tube helps extend the duration of the recording sessions while minimizing stress and movement from the mouse. Mice prefer to fit snugly within the foam, so proper fitting reduces excessive motion. Without this precaution, mice may struggle. If the mouse does show signs of struggle, the recording session should be terminated. Recordings may resume the next day, once physiological stress reactions have abated. In our recordings, the animals were evidently calm most of the time.
We note that the motion that most negatively impacts the pCLE imaging method occurs when the probe moves differentially to the brain tissue. Breathing, seizures, and motion of the animal are irrelevant unless they displace the fiber-probe with respect to the tissue. That is, when the imaging probe moves differentially to the tissue the entire imaging field moves as a unit, making motion artifacts evident. Thus, it is critical to always record from two or more vessels at a time: if all vessels move at once, it may be due to fiber instability, whereas if at least one vessel does not change, the changes seen in other vessels are likely due to actual variations in blood flow, including vasospasms.
One limitation of the pCLE imaging method compared to TPLSM is that the fiber-probe punctures the pia and is thus invasive to the brain. As the probe descends, it causes some damage to the neural tissue it passes through. Thus, recording from the same vessel on two consecutive days can be exceedingly difficult, even when employing the same exact coordinates in both sessions (and often only possible if the initial recording session has a short duration). It is therefore critical, when conducting chronic recording studies, to record at increasingly greater depths in subsequent recording sessions.
The results of the blood flow experiments suggest that excitotoxicity is not the only mechanistic pathway for ictal cell death and hippocampal sclerosis13. The finding that capillary blood flow restriction plays a role in ictal neurodegeneration paves the way for extending the methods described here to the identification of microscopic blood flow changes at any depth of the body, and in any tissue, for clinical diagnostic purposes.
The authors have nothing to disclose.
The project was funded by a Research Initiative Award from the American Epilepsy Society, and an award from the Arizona Biomedical Research Commission to S.L.M., as well as a challenge grant from Research to Prevent Blindness Inc. to the Department of Ophthalmology at SUNY Downstate Health Sciences University, the New York State Empire Innovation Program, and further grants from the National Science Foundation (0726113, 0852636, & 1523614), the Barrow Neurological Foundation, Mrs. Marian Rochelle, Mrs. Grace Welton, and Dignity Health SEED awards, and by federal grants from the National Science Foundation (0726113, 0852636, & 1523614) and by the National Institute of Health (Awards R01EY031971 AND R01CA258021), to S.L.M and S.M.C. This work was also supported by the Office of the Assistant Secretary of Defense for Health Affairs under Award No. W81XWH-15-1-0138, to S.L.M.. L.-C. was supported by a José Castillejo fellowship from the Spanish Ministry of Education. We thank O. Caballero and M. Ledo for their technical advice and assistance.
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, M1.6×0.35 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 |