Precise Brain Mapping to Perform Repetitive In Vivo Imaging of Neuro-Immune Dynamics in Mice

Summary

This protocol describes a chronic cranial window implantation technique that can be used for longitudinal imaging of neuro-glio-vascular structures, interactions, and function in both healthy and diseased conditions. It serves as a complementary alternative to the transcranial imaging approach that, while often preferred, possesses some critical limitations.

Abstract

The central nervous system (CNS) is regulated by a complex interplay of neuronal, glial, stromal, and vascular cells that facilitate its proper function. Although studying these cells in isolation in vitro or together ex vivo provides useful physiological information; salient features of neural cell physiology will be missed in such contexts. Therefore, there is a need for studying neural cells in their native in vivo environment. The protocol detailed here describes repetitive in vivo two-photon imaging of neural cells in the rodent cortex as a tool to visualize and study specific cells over extended periods of time from hours to months. We describe in detail the use of the grossly stable brain vasculature as a coarse map or fluorescently labeled dendrites as a fine map of select brain regions of interest. Using these maps as a visual key, we show how neural cells can be precisely relocated for subsequent repetitive in vivo imaging. Using examples of in vivo imaging of fluorescently-labeled microglia, neurons, and NG2⁺ cells, this protocol demonstrates the ability of this technique to allow repetitive visualization of cellular dynamics in the same brain location over extended time periods, that can further aid in understanding the structural and functional responses of these cells in normal physiology or following pathological insults. Where necessary, this approach can be coupled to functional imaging of neural cells, e.g., with calcium imaging. This approach is especially a powerful technique to visualize the physical interaction between different cell types of the CNS in vivo when genetic mouse models or specific dyes with distinct fluorescent tags to label the cells of interest are available.

Introduction

The central nervous system (CNS) is governed by a complex interplay of interactions between various resident cell types including neurons, glia and vessel-associated cells. Traditionally, neural cells were studied in isolated, co-cultured^1,2,3,4,5 (in vitro) or excised brain tissue (ex vivo)^6,7,8,9,10 contexts. However, there is need to further understand neural cell behavior and interactions in the native environment of the intact brain in vivo. In this protocol, we describe a method to map in vivo regions of interest and precisely re-image those regions in future imaging sessions to track the complex interactions between the various CNS cell types over extended periods of time.

The development of in vivo imaging approaches has provided significant gains for the proper understanding of neural function^{11,12,13,14,15}. Specifically, these approaches provide several advantages over traditional in vitro and ex vivo approaches. First, in vivo imaging systems have physiologically relevant cell and tissue components such as the vasculature with the full repertoire of cellular interactions to provide a complete understanding of neural network physiology. Second, recent findings suggest that when removed from their native environment, certain neural cells (such as microglia) lose important features of their identity and thus physiology^16,17 which can be preserved in the in vivo setting. Third, in vivo imaging systems provide the opportunity for stable longitudinal investigations of weeks to months to study CNS cellular interactions. Finally, given the growing evidence for contributions from the peripheral immune system^18,19 and the microbiome^20,21 in CNS physiology, in vivo systems provide a platform to interrogate such contributions and effects on CNS cells. Thus, approaches that employ longitudinal in vivo imaging to study neuro-immune physiology and interactions in healthy, injured, and diseased states are a great complementary addition to traditional approaches.

In this protocol, we describe a reliable approach to image different cell types in the brain including microglia, neurons and NG2⁺ cells as examples. Two approaches to visualize neural cells in vivo have been developed: the thinned skull approach and the open skull with a cranial window approach. Although thinned skull approaches are in use and are preferred because they overcome some of the disadvantages of the open skull approach such as glial cell activation, higher-than-physiological spine dynamics and the use of anti-inflammatory agents^22,23,24,25, thinned skull approaches also show a few critical drawbacks. First, the thinning procedure is a very delicate procedure that many researchers find difficult to perfect especially when re-thinning is necessary. This is the case because it is often difficult for experimenters to ascertain that they have thinned the skull to a ~20 µm depth. Second, for adequate comparisons between mice, thinning would need to be identical and a variety of thinning success between imaging sessions or mice could complicate visualization of neural structures. Third, when employed for longitudinal imaging, animals with thinned skulls can only be used for a limited number of sessions when re-thinning of the skull is employed. Forth, since some of the bone tissue still remains, clarity in depth of imaging could be compromised from the thinned skull approach allowing for great visualization of more superficial but not as much with deeper regions. In the light of this, deeper brain structures such as the hippocampus, cannot be successfully imaged with the thinned skull approach. These considerations raise the need for alternative and complementary approaches that could overcome these concerns.

Alternative to the thinned skull approach, the open skull window implantation approach uses a procedure in which the skull is replaced with an optically clear glass coverslip. This allows for a near-unlimited number of imaging sessions. Moreover, given the replacement of the skull with the glass coverslip, this method allows for a clear viewing window of fluorescently tagged brain cells for extensive periods of times from hours to months and, therefore, can be employed to study cell activity and interactions that are relevant for physiology, aging and pathology.

Overall, we detail steps that can be followed to do implant chronic cranial windows through a stereotaxic craniotomy that enables in vivo imaging of brain regions of interest. We also describe how the grossly stable brain vasculature or the fluorescently labeled dendrites could be used to generate a coarse or a fine map, respectively of the brain regions of interest. This approach can then be used for repeated imaging over several sessions. The importance of this technique, therefore, lies in its ability to image the long-term changes or stasis in brain elements including the arrangement, morphology, and interactions of the different cellular types.

Protocol

All steps are in accordance with the guidelines set and approved by the Institutional Animal Care and Use Committee of the University of Virginia.

1. Mouse preparation for cranial window implantation

NOTE: Various transgenic mouse lines with florescent tags are suitable for imaging.

1. Use CX3CR1^GFP/+ mice²⁶ to visualize microglia in vivo. Typically, juvenile to young adult 4 to 10-week-old mice that weigh 17-25 g are used.
   NOTE: Although, this approach is even apt for pre-weaned mice, the need to return the mice to their cage with their mothers for feeding, may complicate recovery if the mother does not take adequate care of pups post-surgery. Therefore, the use of mice post-weaning is recommended.
2. Anesthetize the mouse using isoflurane (5% flow in oxygen for induction for 1 min) in an anesthetic chamber. Check that the mouse doesn’t show any movement or twitching responses to toe and/or tail pinches. Take the mouse out of the chamber and in open air thoroughly shave the hair on the head between the ears from about the eye level to the top of the neck region using a hair trimmer.
   NOTE: The concentration of isoflurane used would depend on the size of the induction chamber. Therefore, for smaller chambers, 3-4% isoflurane can be used to effectively induce anesthesia while larger chambers will require up to 5%.
3. Move the mouse to the stereotactic surgery station nose cone for anesthesia (1.5-2% for maintenance for the surgery), stabilize its head using ear bars, and maintain the mouse on a heating pad to keep the body temperature warm.
4. Lubricate both eyes with eye ointment. Inject 100 µL of 0.25% bupivicaine (to provide local analgesia to the mouse that will last 8-12 h) and 100 µL of 4 mg/mL dexamethasone (to reduce the inflammation that may result from the surgery procedure) subcutaneously at the incision site. Allow the mouse to sit for at least 5 min before moving to the next step.
5. Clean the shaved head with three alternating swabs of betadine and 70% alcohol. Make a midline scalp incision using surgical blade or scissors extending from the back of the skull region between the ears to the frontal area between the eyes. The remaining skin is cut to expose the skull. 
6. Clean the connective tissue located between the scalp and the underlying skull with 3% hydrogen peroxide (H₂O₂) and localize the brain area to be imaged with stereotactic coordinates.
   NOTE: There is often some bleeding (step 1.5) from the incision on the skull surface. This bleeding usually resolves by itself within 3-5 min. Cleaning with the peroxide helps. Prior bupivacaine treatment (step 1.4) is also noted to limit the amount of bleeding during this time.

2. Mouse cranial window implantation surgery

1. Drill a circular opening ~4 mm into the skull using a dental drill bit (0.7 mm tip diameter) and carefully remove this portion of the skull using pointed forceps. For imaging the somatosensory cortex of 6-8-week-old mice, locate the center of the craniotomy at -2.5 posterior and ± 2.0 lateral to bregma. During drilling, regularly moisten the skull with sterile saline and cotton swabs to cool the brain, clean off bone debris and soften the skull bone for eventual removal.
   NOTE: The coordinates for the craniotomy would vary depending on the region of interest and the age of the mice.
2. After the skull is removed, carefully place a small coverglass (size #0 at 0.1 ± 0.02 mm thickness) moistened with saline in the craniotomy. Dry off excess saline using a sterile wipe.
3. Using a pointed applicator (such as a pipette tip or the pointed end of a broken wooden cotton swab stick), apply cyanoacrylate glue around the window and allow it to attach to the brain and skull. Apply the primer glue to the rest of the skull and cure it with a curing light for 20-40 s. Prepare a well around the window with the final glue and cure with a curing light for 20-40 s.
4. Glue a small head plate on to the skull on the contralateral hemisphere of the craniotomy first with the primer glue as a primer and then with the final glue. Cure both with the curing light for 20-40 s each.
   NOTE: Sutures are not needed if the skull is totally covered with the glue during this procedure.

3. Post-surgery care

1. Allow the mouse to wake up in the absence of anesthesia (recovery done on a heating pad shortens the recovery time) and return it to its home cage once fully awake. Inject one subcutaneous dose of buprenorphine SR (0.5 mg/kg) as post-operative analgesia that is sufficient for 72 h.
2. To facilitate a healthy recovery from the surgery, provide the mouse an extra soft food, which can be in the form of regular solid chow in water to soften the chow or food in the form of a gel.
   NOTE: A one-time provision of the soft food immediately after the surgery is sufficient.
3. Monitor the mouse daily for health and proper recovery for the first 72 h of the surgery procedure. Afterwards, perform imaging from as early as 2 weeks from the window implantation surgery.
   NOTE: If done well, mice recover well showing normal ambulatory behaviors, sufficient cage exploration, good hydration, stable weight gain and extensive interactions with other mice in the cage and other items in the cage. Mice showing lethargy, dehydration and greater than 10% weight loss following the surgery are euthanized and removed from the study.

4. Two-photon brain mapping for initial imaging

1. Anesthetize the mouse (Isoflurane, 5 % induction and 1.5 % maintenance). Stabilize the head using screws to mount the headplate on the two-photon microscope stage, being maintained on a heating plate at 35 ˚C. Inject intraperitoneally 100 µL of blood vessel dye such as Rhodamine B (2 mg/mL).
   NOTE: Imaging could also be done in awake mice without anesthesia. However, recent studies indicate that anesthesia affects microglial surveillance dynamics^27,28,29 and that head fixation for two photon imaging in awake mice increases stress even during chronic imaging for at least 25 days (see Juczewski et al., 2020)³⁰.
2. Clean the surface of the cranial window gently using a cotton swab dabbed in 70% ethanol. Put a few drops of water or saline on the cranial window and lower the objective lens into the solution since the objective is an immersion lens.
3. Hand-draw a coarse map to denote the major blood vessel landmarks in a lab notebook while looking through the eyepiece by epiflorescence. Use this drawing to identify the specific regions during two photon imaging. Alternatively, take pictures of the blood vessels either through a camera fitted to the microscope or through a hand-held camera or phone.
   NOTE: These hand-drawn images and pictures are to facilitate revisiting the same broad regions under the microscope before two photon imaging. These are not precise image mapping.
4. Under two photon imaging, collect images of florescent cells and vessels as needed. Take careful notes with appropriate coordinates to ensure that that the precise regions can be revisited for subsequent imaging. Collect several fields of view in this initial imaging session e.g. acquire z-stack images every 1-2 µm through a volume of tissue.
   NOTE: While collecting images by two photon, the blood vessel landmarks are used for coarse mapping. If fine mapping is needed, YFP-labeled dendrites from Thy1-YFP³¹ mice are used.
   1. Use these recommended parameters for imaging: a wavelength of 880-900 nm is optimal; for GFP and/or dsRed / Rhodamine excitation, a 565 nm dichroic mirror with 525/50 nm (green channel) and 620/60 nm (red channel) emission filters are used; for GFP and YFP separation, a 509 nm dichroic mirror with 500/15 and 537/26 nm emission filters are used; the power at the brain is maintained at 25 mW or below; image resolution is 1024 x 1024 pixels, the field of view taken with a 25X 0.9 NA objective at a 1.5X zoom factor is 295.24 x 295.24 µm.
5. At the end of the imaging, take the mouse off the stage, allow it to wake up from anesthesia and return to its home cage until a future imaging session.

5. Two-photon imaging and re-imaging

1. For future subsequent imaging sessions, which could be anywhere from a few hours to months after the initial imaging session, anesthetize the mouse (Isoflurane, 5% induction and 1.5% maintenance), mount on the two-photon microscope, maintain on a heating plate and re-inject 100 µl a blood vessel dye such as Rhodamine B (2 mg/mL).
2. Open the previously obtained images in ImageJ and, using these images as well as the notes from the previous session, identify the previously imaged areas and carefully re-image them. 
3. Repeat this for as long as the imaging window is clear or as essential for the extent of the study.

Results

To visualize microglial dynamics in vivo, double transgenic CX3CR1^GFP/+:Thy1^YFP mice were used. The Thy1-YFP H line is used as opposed to the Thy1-GFP M line to avoid florescence overlap of microglia (GFP) and neurons (YFP). Alternative approaches could use a reporter line in which microglia are labeled with e.g., tdTomato and then the Thy1-GFP M line can be used. A drawback of the H line is that YFP labels a lot of neurons and the label increases with increasing age (personal observation). The M li...

Discussion

The advent of in vivo two-photon imaging has opened opportunities to explore the plethora of cellular interactions and dynamics that occur in the healthy brain. Initial studies focused on using the open skull craniotomy approach to image neuronal dendrites by both acute and chronic imaging^37,38. This can also be used to elucidate neuroimmune interactions in the brain. This protocol describes a method for the reliable imaging of fluorescently tagged cells (especia...

Materials

Name	Company	Catalog Number	Comments
Coverglass (3mm)	Warner Instruments	64-0726
Cyanoacrylate glue (Krazy Glue)	Amazon	https://www.amazon.com/Krazy-Glue-Original-Purpose-Instant/dp/B07GSF31WZ/ref=sr_1_2?keywords=krazy+glue&qid=1583856837&s=pet-supplies&sr=8-2
Demi Ultra LED Curing Light System	Dental Health Products, Inc	910860-1
Dental Drill	Osada: www.osadausa.edu	EXL-M40
Drill Bit	Fine Science Tools	#19008-07
Eye Ointment	Henry Schien	1338333
iBond Total Etch (Primer glue)	Chase Dental Supply (Heraeus Kulzer)	66040094
Rhodamine B	Millipore Sigma	81-88-9 (R6626)
Tetris Evoflow glue (Final glue)	Top Dent (Ivoclar Vivadent)	#595956
Wahl Brav Mini+	Amazon	https://www.amazon.com/Wahl-Professional-Animal-BravMini-41590-0438/dp/B00IN24ILE/ref=asc_df_B00IN24ILE/?tag=hyprod-20&linkCode=df0&hvadid=167141013968&hvpos=&hvnetw=g&hvrand=12368793083893626704&hvpone=&hvptwo=&hvqmt=&hvdev=c&hvdvcmdl=&hvlocint=&hvlocphy=9008337&hvtargid=pla-332197544154&psc=1