Combining Imaging and Electrophysiology to Visualize and Record Spreading Depolarizations in Mice

Summary

This protocol demonstrates the combination of imaging and electrophysiology to reliably detect spreading depolarizations in adult mice following a mild traumatic brain injury.

Abstract

Spreading Depolarizations (SDs) are massive events in the brain that often go undetected due to their slow propagation through gray matter. Because SD detection can be elusive, it is optimally confirmed using multiple methods. This protocol describes methods for combining imaging and electrophysiology to detect SDs in a manner that most laboratories can reliably and easily adopt. SDs occur following traumatic brain injuries, stroke, subarachnoid hemorrhages, ischemia, and migraine aura. Historically, SDs have been recorded using DC amplifiers, which can resolve the slow extracellular shift and the depression in high-frequency activity. However, DC amplifiers are nearly impossible to use for chronic in vivo recordings. This protocol employs a common AC amplifier for in vivo electrophysiology recordings to confirm high-frequency depression, along with non-invasive imaging necessary to detect the propagating wave of SD. These methods can be reliably adopted and/or modified for most experimental approaches to confirm the presence or absence of SDs following brain injury.

Introduction

Spreading depolarizations (SDs) are associated with various neurological conditions, including traumatic brain injury (TBI), stroke, subarachnoid hemorrhages, ischemia, and migraine aura¹. SDs have been implicated in the progression of tissue damage in strokes and TBIs². The exact mechanism underlying tissue loss progression remains unclear^3,4. However, SDs occur when the metabolic demands of neurons are not met with sufficient oxygen and glucose, as the function of the ATP-dependent sodium-potassium pump becomes inadequate to maintain ion gradients. To understand the role of SDs in tissue loss, precise techniques are needed to measure and study them, which is crucial for advancing scientific knowledge. SDs are significant neurological events that can often go undetected due to experimental design and/or instrumentation limitations.

The role of spreading depolarizations (SDs) in injury progression is currently being elucidated, but their role in mild traumatic brain injuries (mTBIs) remains unclear. Recent studies using the techniques described in this manuscript have shown that SDs are initiated in closed-skull mTBI mouse models, even when there is no overt tissue damage or bleeding^5,6. These studies suggest that SDs may contribute to acute neurological impairments and neuroinflammation. Additionally, the presence of mTBI-induced SDs has been closely associated with acute behavioral deficits^6,7. Therefore, it is likely that SDs are initiated in all TBI models, but their role remains difficult to ascertain without confirmation. Confirming the presence of SDs is crucial for accurately assessing the outcomes of mTBIs.

Spreading depolarizations (SDs) are identified by a significant drop in extracellular potential, known as the "DC shift," and by a depression of high-frequency activity⁸. The "DC shift" typically takes seconds to resolve (~10 s)⁹. To capture the "DC shift," a DC amplifier and silver wire electrodes are required. However, in vivo electrophysiology is usually performed with AC amplifiers, which are designed to filter out low-frequency events to reduce motion artifacts. Unfortunately, this low-frequency filtering also removes the "DC shift." Additionally, silver is toxic and unsuitable for chronic recordings. Commonly used AC amplifiers can detect the depression of high-frequency activity, but without the accompanying "DC shift," confirming the presence of an SD is challenging.

Spreading depolarizations (SDs) can also be detected using various imaging techniques. Most commonly, SDs are imaged with laser speckle contrast imaging, which measures cerebral blood flow through the intact skull¹⁰. SDs significantly influence cerebral vasculature and blood flow. In most species other than mice, SDs induce a propagating wave of hyperperfusion that travels with the leading edge of the SD¹¹. SDs can also be imaged using calcium or glutamate fluorescent indicators¹², though this requires loading tissue with dyes or genetically encoded fluorescent indicators^12,13. Interestingly, SDs can be observed by the naked eye due to intrinsic optical signals (iOS) associated with the SD¹⁴. iOS are complex signals composed of autofluorescent molecules like NAD(P)H and FADH^15,16, tissue swelling¹⁷, and potential changes in cerebral blood flow. Despite their complexity, iOS signals are reliable and can be captured in wild-type mice¹⁸. This protocol describes the method for imaging iOS because this technique can be implemented in any lab without the need for specialized viruses or transgenic animals.

The overall goal of this method is to experimentally confirm the presence or absence of a spreading depolarization (SD) following a mild traumatic brain injury (mTBI) using simultaneous imaging and electrophysiology. In vivo electrophysiology, facilitated by advanced amplifiers, is a commonly used technique. However, these amplifiers are unable to detect the characteristic "DC shift." Therefore, imaging techniques, as described in this protocol, are employed to confidently confirm the presence or absence of SDs. The advantage of combining imaging and electrophysiology is that imaging can detect the propagation of the SD, while electrophysiology can identify the associated high-frequency depression.

Overall, this method is suitable for investigating the presence of spreading depolarizations (SDs) following a traumatic brain injury (TBI). Its application could enhance understanding of cortical network function after TBI and SDs. This protocol can be adapted from a closed skull impact model to other models, such as controlled cortical impact, fluid percussion, or weight drop. Implementing this protocol across multiple laboratories could elucidate the role of SDs across varying TBI severities. Any established TBI model can be used, followed by the iOS imaging and electrophysiology procedures described in this protocol.

Protocol

All procedures were performed in accordance with the Institute's Animal Care and Use Committee (IACUC) at the University of New Mexico. C57BL/6J mice were obtained from a commercial source. Equal numbers of male and female mice were used within the age range of 8-14 weeks. All mice weighed approximately 20 g. Details of the animals, reagents, and equipment used in this study are listed in the Table of Materials. Supplementary File 1 contains the list of abbreviations.

1. Preparing the skull for imaging and electrophysiology

1. Anesthetize the animal in an isoflurane induction box with 2%-3% isoflurane in 100% oxygen, ensuring proper anesthesia depth with no flinching from a paw pinch.
2. Administer preoperative analgesics (Slow-release Buprenorphine, 3.25 mg/kg, SQ) before placing the animal into a stereotaxic frame and maintain anesthesia (2% isoflurane).
3. Add ophthalmic ointment and maintain the animal's body temperature with a heating pad that has body temperature feedback.
4. Shave the hair above the skull where the incision will be, down the midline from the ocular fissure to lambda.
5. Remove the clipped hair before sterilizing the skin with an antiseptic and disinfectant scrub and solution. Finish with 70% ethanol, and repeat three times to ensure a sterile surgical site.
6. Using a #10 scalpel blade, make an incision down midline from rostral to caudal, immediately followed by the application of medical grade cyanoacrylate glue to the incision. Spread the skin laterally to expose the skull.
7. Add another drop or two of cyanoacrylate glue over the skull to add additional protection to the skull.
   NOTE: Glue application is time-sensitive as the skull will begin to go opaque when exposed to the air for longer than 10 s. This will obscure the visualization of the SD through imaging methods.
8. While the glue is drying, use sterile surgical scissors to increase the incision size over the cerebellum. Once the glue is set, begin the implantation of skull screw electrodes.
   1. Sterilize skull screws by a bead sterilizer for 20-30 s prior to implantation.
   2. Place the skull screws in a region distant from the SD initiation site to maintain an open visual window.
      NOTE: For chronic recordings in the primary visual cortex, three skull screws are used. The first screw is placed approximately at y = -6.00 mm from the bregma, in the cerebellum, serving as a reference electrode. The recording electrodes are positioned at y = -2.79 mm from bregma and x = ±2.5 mm from the midline, within the left and right hemispheres.
   3. With a 0.8 mm size drill bit, drill pilot holes through the skull. To ensure no damage to the underlying tissue, -0.5 mm in z is ideal. Use #00-90 screws that are pre-soldered to gold pins with stainless steel wire.
   4. Turn the screws down into the skull.1.5-2 full turns will get the screw through the skull and in contact with the cortical surface.
   5. Wind excess wire around the screw, allowing some slack in the wire for headstage placement later.
   6. Using the same procedures, place the other skull screws.
   7. Once all skull screws are in place, plug all the gold pins into the desired headstage connector. Hold the connector in place using the stereotaxic arm and hold the connector pins close above the screw heads while ensuring the wires are not touching.
   8. Seal the entire area with cyanoacrylate glue, making sure not to disrupt the imaging area in front of the skull screws.
   9. Build up dental cement around the base of the gold pins, ensuring that the gold pins from the skull screws are not connected to the gold pins in the head stage.
   10. Allow the dental cement to cure before removing the headstage connector.
   11. Remove the animal from the stereotaxic frame and place it back into the home cage to recover. Follow standard postoperative care and keep animals on a heating pad while recovering from anesthesia.
       NOTE: After surgery, the animals were single-housed and monitored for pain daily. Follow the local institutional guidelines for analgesia. If using a slow-release buprenorphine monitor for pain and potentially give an additional dose at 72 h.
9. Remove any objects that may catch on the electrodes, including housing and enrichment. Use standard bedding with ad libium access to food and water.
   NOTE: Allow for a two-day recovery period between surgery and recording.

2. Detecting spreading depolarizations with intrinsic optical signal (iOS) imaging and AC- electrophysiology

1. Anesthetize the animals with 2% isoflurane in 100% oxygen.
   NOTE: The imaging part of this procedure should take ~5 min, and the amount of isoflurane exposure should be consistent.
2. Place the animal into a stereotaxic frame. Through the nose cone, maintain 1%-2% isoflurane in oxygen.
   NOTE: It is critical to maintain proper depth of anesthesia, as isoflurane is known to reduce SDs. Maintaining 60-70 breaths/min is recommended throughout the experiment.
3. Place the animal beneath the camera and ensure the camera is focused on the cortical surface by adjusting the focus ring.
4. Apply mineral oil to the imaging area to increase transparency.
5. Turn on the computers, LED panel board, and electrophysiology equipment.
   NOTE: LED panel board works best when positioned in front of the animal to illuminate the imaging window while overhead lights are off. The light was set to ~2,400 lux (95% power) with a cool temperature setting of 4000K.
6. Load setting for iOS using Micro-Manager^19,20,21,22.
   1. Press Live to view the mouse under the camera and center the animal's skull under the camera. Close the "Live" window.
   2. Press Multi-D Acq.
   3. In the Multi-Dimensional Acquisition window, select the desired time points.
      NOTE: A count of 750 images at an interval of 250 ms is recommended.
   4. Save images to the desired file path and name prefix.
7. Connect the gold pins from the animal to the headstage connector and then to the headstage.
8. Load setting for Intan software.
   1. Click on Choose file format icon, and in pop-up window, select One File Per Signal Type Format. This will save the file as a ".dat" for further use in MATLAB.
   2. In the BW tab under Hardware Bandwidth, select amplifier bandwidth of 0.13-500 Hz.
   3. Under Filter Display Selector, select WIDE to cover the widest range of signals.
   4. To select the desired file path to save, click on the Select Filename icon.
9. Perform impact using an electromagnetic impactor.
   NOTE: Impactor settings are 4 m/s impact speed and 5 mm deflection.
   1. Turn the impactor on and ensure that the tip retracts and extends appropriately by flicking the 3-way toggle to retract the tip and extend the tip. The tip should be extended before placing the animal underneath the tip.
   2. Moving quickly, place the animal on a foam bed and slide the bed beneath the tip.
   3. Align the impactor tip at midline and 1 mm rostral of bregma.
   4. Lower the impactor tip until it is firmly touching the top of the animal's head.
      NOTE: Do not push the tip down into the head.
   5. Push the lever to retract the impactor tip and then manually lower it 5 mm before activating the impact lever.
   6. Immediately remove the animal from the foam bed and place the animal beneath the camera for imaging within 5 s, as the SD is initiated from the impact.
10. Start iOS imaging by clicking on Acquire! in the Multi-Dimensional Acquisition window. Image for 4 min to capture the entire SD.
11. Start the electrophysiology recording by clicking on the Record icon on the Intan software.
12. Remove the animal from the foam pad and allow the animal to recover in an open arena or home cage.
    NOTE: It is recommended that electrophysiology be recorded for at least 1 h to allow the cortical network function to recover.
13. Upon the completion of the recording, unplug the animal from the headstage and connector and place the animal back into its cage.
    NOTE: It is recommended to allow 24 h between each impact and recording session. The animal can remain in its home cage for the entire time outside of recording sessions.

3. iOS image visualization and quantification

1. In ImageJ^19,20,21,22, open the image stack by clicking on the File > Import > Image sequence.
2. Manually scroll through images to ensure the lack of movement or any other artifacts.
3. To generate the plot of the SD, create an ROI by using the cursor to draw a square.
   NOTE: It is recommended to use a 30 x 30-pixel square. One may use any shape or size, but ensure ROI size does not change across animals.
4. Create a plot by clicking on Image > Stacks > Plot Z-axis Profile. On the newly created graph, click on the live button. Go back to the image stack of the SD and move the ROI around until the graph shows the SD curve. The plot should look like a Sin wave with an initial rise followed by a drop below the baseline and a slow return to baseline within a few minutes.
   NOTE: The plots can be expressed as arbitrary units (AU) or as percent baseline using the first few images as the baseline average.
5. On the generated plot, click on List to generate a list of values that can be plotted in other graph-generating software or Matlab.
6. For visualization, turn the grayscale image into a lookup table in ImageJ by Image > Lookup Tables > Fire.

4. Electrophysiology analysis

NOTE: It is recommended that the expertise of the Buzsaki lab and their MATLAB-generated code be heavily utilized for visualizing and analyzing electrophysiology data. The analysis will require downloading MATLAB with an institutional activation code and the Buzsaki lab "Buzcode" zip file via GitHub (https://github.com/buzsakilab/buzcode).

1. In the Intan electrophysiology software, ensure that the exported raw data is saved in ".dat" format.
   NOTE: Matlab can be installed from the MathWorks website. An institutional or individual activation code is necessary for the purchase and use of Matlab. Some prior experience with running code in Matlab is necessary for performing this analysis.
2. Use the Buzaki code bz_LoadBinary function to load the data into Matlab.
   NOTE: Example "'data' = bz_LoadBinary ('full data file path','frequency', sampling frequency, nChannels, number of channels);
3. Filter the AC noise using a notch filter centered at 60 Hz.
   NOTE: Example notch_freq = 60; [b,a] = iirnotch(notch_freq/(sampling_frequency/2), notch_bandwidth/(sampling_frequency/2)); filtered_data = filtfilt(b, a, data);
4. Isolate the high-frequency activity using a bandpass filter with a range of 0.5-100 Hz, in accordance with the recommendations of the COSBID group for analyzing and visualizing spreading depolarizations (SDs).
5. Calculate the power spectrogram using the pspectrum command in MatLab.
6. To visualize, plot the high-frequency and the power spectrogram on the same time scale and in one column.
   NOTE: From the plots, there will be an extension of the high-frequency depression and loss of power.

Results

Figure 1 illustrates a graphical representation of the experimental setup. The recording skull screws are positioned in the primary visual cortex, with one skull screw electrode placed in the cerebellar vermis as a ground electrode. The skull screws and gold pins are secured to the skull using cyanoacrylate glue and dental cement. It is crucial to limit the dental cement to the caudal part of the skull to leave space for impact and imaging. The impact location, centered at midline and a...

Discussion

Spreading depolarizations are crucial in various disease models, including traumatic brain injury (TBI), stroke, and migraine¹. Accurate detection of SDs is essential for understanding their role in acute behavioral deficits and/or the activation of the neuroimmune system following mild traumatic brain injuries (mTBIs)^23,24. To mitigate the risk of missing SDs when using a single method, a multi-method approach is necessary for effective v...

Materials

Name	Company	Catalog Number	Comments
#00-90 x .118 Machine Screw	US Micro Screw	00.90-118-M-SS-P, Amazon
0.8 mm diameter drill bit	PCB Drill Bit	WYTP21, Amazon
16-A Headstage	Intan	C3335
2.1 mm Burrs micro drill bit	Fine Science Tools	19007-21
AC amplifier	Intan	RHD recording system
C57BL/6J mice	Jackson Laboratory	Jax#000664
CCD camera	Mightex	TCE-1304-U
Extension lens	Zeikos	ZE-ETN
Ethiqa XR (slow release Buprenorphine)	Midwest Veterinary Supply	Product Number 267.10000.3
Gold connector pin (female)	A-M Systems	520100
Gold connector pin (male)	A-M Systems	520200
ImageJ2/Fiji	Image J	NA
Impactor One	Leica	39463930
LED panel board	Supon	L122t
MATLAB	MathWorks	Institutional or individual activation code
Medical grade cyanoacrylate glue	MasterBond	MB287
Micro-Manager Camera Software	Image J	NA
Nail drill/polisher	Makartt Nail Drill Electric Nail File	JD700
Omnetic connectors	Omnetics	A79014-001
RHX Data Acquisition Software	Intan	NA
Stainless Steel uncoated wire (0.005")	A-M Systems	792800
Stereotaxic Frame	Kopf Intruments	1923-B
Teets 'cold cure' dental cement	A-M systems	525000, 526000
XLR camera lens	Nikon	NIKKOR 50 mm f/1.8-16