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Method Article

In Vivo Calcium Imaging of Neuronal Ensembles in Networks of Primary Sensory Neurons in Intact Trigeminal Ganglia

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DOI:

10.3791/68284

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August 1st, 2025

 ,  ,  ,  ,  ,  ,  , 
1Department of Oral & Maxillofacial Surgery, School of Dentistry, University of Texas Health Science Center at San Antonio, 2Department of Endodontics, School of Dentistry, University of Alabama at Birmingham, 3Division of Life Sciences, College of Life Sciences and Bioengineering, Incheon National University, 4Programs in Integrated Biomedical Sciences, Translational Sciences, Biomedical Engineering, Radiological Sciences, University of Texas Health Science Center at San Antonio

In This Article

Summary

This protocol describes in vivo GCaMP calcium imaging of intact trigeminal ganglion (TG) neurons. This description includes TG exposure surgery, in vivo confocal microscopy imaging of TG neurons, application of somatic stimuli during TG neurons microscopic imaging, and analysis of in vivo GCaMP calcium imaging data.

Abstract

Calcium imaging is an important tool for monitoring signaling mechanisms that either influx Ca2+ into the cytosol or release Ca2+ from intracellular Ca2+ stores. The use of Pirt-GCaMP3 mice, which have the Ca2+sensor GCaMP3 inserted downstream of the Pirt promoter region and expressed in nearly all (>95%) primary sensory neurons, allows for simultaneous monitoring of the activity of >3,000 neurons via Ca2+ imaging of trigeminal ganglion (TG) primary sensory neurons. This allows investigators to study neural network responses to noxious stimuli and somatosensory and hormonal responses in vivo, which would otherwise be very difficult to detect using other methods.

Specifically, the ability to monitor the TG area innervating the face and other areas of the head separately allows for monitoring primary sensory neurons activated by stimuli of the face area before later layers of neuron processing. The number of neurons generating Ca2+ transients and the amplitude of Ca2+ transients, which indicate their sensitivity to different sensory modalities, can also be observed. Additionally, investigators can measure the diameter of neurons, which is an indicator of the type of fiber activated (non-noxious mechano-fibers vs. noxious nociceptive fibers, Aβ, Aδ, and C fibers). Therefore, Pirt-GCaMP3 calcium imaging of TG would be a powerful tool to dissect specific sensory modalities and neuronal cell types to study trigeminal origin pain (orofacial and craniofacial, dental or/tooth pain, headache, migraine, temporomandibular joint pain, trigeminal neuralgia), itch and touch and any other somatosensations, as well as hormonal responses.

Introduction

Primary sensory neurons directly innervate tissues, including the skin, and relay sensory signals to the central nervous system (spinal cord or brainstem). Trigeminal ganglion (TG) neurons innervate the head and respond to sensory stimuli, including pain from the teeth, periorbital region, orofacial and craniofacial regions, temporomandibular joint1,2, and meninges3,4,5. TG neurons vary in size, level of myelination, and gene expression profiles6,7,8,9. Smaller neurons are nociceptive, whereas larger neurons generally respond to nonpainful mechanical stimuli10,11. Dysfunction of trigeminal ganglion neurons can sensitize both the neurons themselves and central nervous system (CNS) sensory neurons11,12.

In a normal physiological context in non-pathological conditions, firing neurons in vivo are necessary for pain, but until the last two decades, tools for studying intact sensory ganglia in vivo have been limited to monitoring relatively few neurons at a time13. Due to normally low cytoplasmic calcium concentrations, calcium influx during action potentials, and the use of calcium as a cytoplasmic signal, calcium dynamics are powerful surrogates to monitor action potentials or cellular activity in vivo using calcium-sensitive fluorescent indicators. The rationale for this method is that fluorescence microscopy can simultaneously monitor these indicators in hundreds to thousands of neurons simultaneously in peripheral sensory ganglia of mice13,14,15,16,17,18,19,20,21,22,23,24 and rats25,26. This is a powerful advantage over methods that can monitor a more limited number of neurons in vivo.

The goal of this method is direct observation of cytoplasmic Ca2+ dynamics in vivo in the TG in response to mechanical, temperature, chemical, and hormonal stimuli in headache pain models19,23,24. This method can be used as a complement to pain or other somatosensory tests in the orofacial area. In vivo trigeminal Ca2+ imaging has been used to characterize thermal and sensory coding19,22, study classes of mechanoreceptors27, discover a new class of mechanosensitive neuron28, monitor trigeminal responses to normal mechanical stresses on the meninges29, study the effects of hormones on pain sensitivity24, characterize orofacial pain models30, and study contributors to alcohol withdrawal-induced pain23.

Pirt-GCaMP3 mice allow monitoring of neural activity in vivo because they have the Ca2+ sensor GCaMP3 inserted into the promoter region of the Pirt gene, which is highly expressed in all (>95%) primary sensory neurons12,18. This paper demonstrates a technique for performing in vivo TG surgical exposure and the collection and analysis of calcium indicator microscopy data for the right-side TG of Pirt-GCaMP3 mice using confocal laser scanning microscopy.

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Protocol

All procedures described herein are performed under protocols approved by the Institutional Animal Care and Use Committee. The surgery is non-survival. Sections 1 and 2 of this procedure must be performed immediately upon initiation. Section 3 may be performed at a later date.

1. Securing and surgery of the mouse for right side TG imaging

NOTE: Pirt-GCaMP3 C57BL/6J mice18 need to have adult body size (usually 8 weeks or older), regardless of sex. Bilateral TG imaging is possible. Times listed here are estimates by an experienced technician. If bleeding is more than normal, it may take several minutes longer for each given time.

  1. Prepare a sterile saline solution with 40 mg/mL ketamine and 6 mg/mL xylazine. Prepare enough volume for surgery and euthanasia after imaging (minimum 9 µL per gram of mouse body weight).
    NOTE: Many kinds of anesthesia have been used during in vivo imaging. Caution: Ketamine is harmful if injected, swallowed, or in contact with eyes. Handle with care.
  2. Sterilize all surgical instruments in a bead sterilizer. Wash instruments with 70% ethanol.
    NOTE: Alternatively, autoclave the instruments beforehand.
  3. Inject 2.25 µL of ketamine/xylazine per gram of body weight (90 mg/kg and 13.5 mg/kg ketamine and xylazine, respectively) i.p. into Pirt-GCaMP3 mice 15-25 min prior to surgery. Do not exceed 120 mg/kg ketamine.
  4. After 15-25 min of anesthetic injection (step 1.3), pinch the hindpaw with forceps to ensure that the mouse is on a surgical anesthetic plane.
    NOTE: The pinch should be intense enough to cause vocalization in a conscious mouse, but not intense enough to cause injury. It is important to not leave the mouse at this step longer than necessary to avoid a drop in body temperature. Keeping the cage warm is helpful, if possible.
  5. Place the mouse on a heating pad to ensure the body temperature is maintained at 37 °C and place the mouse's head in a stereotactic mask (Figure 1A,B), with the head tilted approximately 15° to the left (Figure 1B).
    NOTE: A stereotactic mask that both stabilizes the head and provides a route to administer isoflurane is described here. If a procedure does not use isoflurane or a stereotactic mask is not used, another method of stabilizing the head should be used to prevent movement.
  6. Apply ophthalmic ointment to the eyes to prevent dryness and irritation, since irritation induces trigeminal nerve activity.
  7. Shave the right side of the mouse's head.
    NOTE: This step takes 90 s. The investigator may briefly remove the animal from the heating pad for shaving.
  8. Make a 9 mm x 5 mm rectangular incision between the right ear and the right eye (Figure 2A). Stop bleeding with a hemostatic swab or laboratory wipe.
    NOTE: This step takes 2 min. The investigator may use hemostatic forceps or a retractor to keep the incision open. This is a non-survival procedure, meaning additional cleaning is unnecessary. However, the investigator may swab the surgical site with povidone iodine. The largest allowable incision size is given. If the surgeon is sufficiently skilled, a smaller incision is preferable to a larger incision to minimize tissue damage. The TG innervates this tissue and could be damaged.
  9. Cut the tissue (periosteum) on the skull surface just ventral to the eye (Figure 2B).
  10. Using a dental drill and taper fissure TNT bur 31p, drill a hole approximately 9 mm x 5 mm in the dorsal skull centered on the incision site (Figure 2C). Apply the bur as lightly as possible to the skull surface.
    NOTE: This step takes approximately 2 min. If the surgeon is sufficiently skilled, a smaller hole is preferable to a larger hole.
  11. Tilt the head until the TG is clearly visible (Figure 2C). If there is bleeding onto the TG, aspirate the blood, or remove the blood by wicking with a hemostatic swab or laboratory wipe. Ensure that the TG is clearly visible without removing any cortical tissue (Figure 2C).
    NOTE: We routinely image TG for 3-6 h or more without signs of distress in neurons, but a researcher can choose to place a coverslip or other covering to prevent the TG from drying.
  12. Move the animal and heating pad to the microscope stage and place the mouse's nose into the stereotaxic nose cone. Hook up the oxygen/isoflurane lines so that the mouse receives continuous isoflurane anesthesia (Figure 3A).
    NOTE: This step takes 3 min. Continuous isoflurane requires oxygen tanks, gas delivery tubes, and an anesthesia vaporizer. It is also possible to maintain anesthesia throughout the procedure by periodic injections of anesthetic.
  13. Position the stereotaxic frame so the objective of the microscope is 9 mm directly above the cranial opening (Figure 3B).
    NOTE: Exact distance depends on the objective, microscope, and mouse.
  14. Insert the rectal thermometer. Connect the power cord to the rectal thermometer and heating pad. Connect the anesthesia mask to the isoflurane in the oxygen line with 1% isoflurane.

2. TG imaging

NOTE: Use green (FITC) setting 495 nm, emission 519 nm, detection wavelength 500-580 nm, GaAsP-Pmt1 imaging device, GaAsP-PMT detector, or use the recommended settings for the microscope. See Table 1 for a summary of software settings.

  1. Use a 5x/0.25 M27 objective, microscope manufacturer's software, and a vertical confocal laser scanning microscope. Locate the TG surface with the microscope.
    NOTE: Objective choice depends on the microscope and investigator preference.
  2. Adjust the objective and nose cone to make the TG surface as flat as possible and have the largest possible surface area in the focal plane.
    NOTE: A flatter TG provides a sharper image, causes software to compose sharper movies, images more neurons, and simplifies and improves the accuracy of analysis compared to a more tilted TG.
  3. Maintain the mouse under anesthesia by applying 1-1.5% isoflurane to the oxygen flow through the anesthesia mask.
    NOTE: The animal must be monitored closely throughout the procedure by pinching the hindpaw every 20-30 min to ensure that the animal remains under anesthesia. The percentage of isoflurane required to maintain anesthesia will depend on the individual mouse. If the animal moves during a test pinch, the isoflurane must be increased. If breathing becomes shallow, the isoflurane must be decreased. CAUTION: Isoflurane is harmful if inhaled and may cause dizziness or drowsiness. Avoid inhalation and make sure the work area has sufficient ventilation.
  4. Load the microscope high-speed scanning protocol: voxel size 4.160 µm x 4.160 µm x 14 µm, 512 x 512 pixels, 10 optical slices Z-stack, 1.02 airy units (AU)/33 µm, 15% 488 nm laser power/75 mW, pixel time 1.52 µs, line time 0.91 ms, frame time 465 ms, LSM scan speed 8, bidirectional scanning, GaAsP-PMT detector gain 550 V, digital gain 1.
    NOTE: Optimal settings may vary between microscopes and animals.
  5. Scan the TG in short bursts of 6-8 cycles. Create orthogonal projections of the scans (one scan makes one movie frame) to create a movie and check the image clarity and consistency between frames. If necessary, adjust the stereoscopic frame position and optical section thickness and repeat this step until a clear, consistent movie is created.
    NOTE: This step must be repeated if the mouse's head moves or the investigator repositions the mouse or adjusts the focal plane.
  6. Load the microscope high resolution scanning protocol: voxel size 0.520 µm x 0.520 µm x 14 µm, 4096 x 4096 pixels, 6 optical slice Z-stack, 1.02 airy unit (AU)/33 µm, 20% 488 nm laser power/100 mW, pixel time 0.52 µs, line time 4.95 ms, frame time 20.26 s, LSM scan speed 6, bidirectional scanning, GaAsP-PMT detector gain 550 V, digital gain 1.
    NOTE: Optimal settings depend on the microscope and animal. If cells are labeled with td-Tomato, set the microscope to scan the red channel 592 nm excitation/614 nm emission, detection wavelength 600-700 nm in addition to the green channel.
  7. Create a high-resolution image of the TG.
    NOTE: This image can be useful for counting neurons, measuring the diameter of dim neurons, or providing a detailed image to distinguish if the same neuron responded to two different stimuli or two different neurons responded.
  8. Load the microscope high-speed scanning protocol (see step 2.4).
  9. Record spontaneous activity for 80 cycles (approximately 10 min). Create an orthogonal projection movie and check that the images are clear and consistent enough to analyze.
    NOTE: This produces enough frames to analyze spontaneous activity. The number can be decreased or increased depending on how important spontaneous activity is to a particular experiment.
  10. To apply stimulation, set the microscope to scan 15-20 cycles.
    NOTE: Be cautious when applying stimuli to not move the mouse's head. Moving the TG by even microns during the stimulus can disrupt imaging. The scanning allows cycles to establish a baseline, a stimulus, and to observe the neurons after the stimulus is removed.
  11. Wait until cycles 1-5 are completed to generate a baseline. Apply stimulation during scans 6-10. Wait at least 5 min after each stimulus to avoid desensitizing neurons.
    NOTE: Each stimulus requires a separate 15-20 cycles scan. Apply mechanical stimulation first, followed by cold, heat, and chemical stimulation. Apply low-frequency stimulation (e.g., low mechanical force, temperature close to room temperature) before high-frequency stimulation (high mechanical force, temperature far from room temperature). Be careful not to move the TG when applying stimulation. If scans can be clearly heard, stimulation can be applied immediately after scan 5. However, any method that provides consistent timing of stimulation will work. See the discussion about stimulus order.
  12. For von Frey in the area innervated by V2 part of the TG, hold the filament and apply it repeatedly to the ipsilateral area below the eye and above the mouth from immediately after scan 5 to immediately after scan 10.
  13. For von Frey in the area innervated by V3 part of the TG, hold the filament and apply repeatedly to the ipsilateral area just below the ear from immediately after the end of scan 5 until immediately after the end of scan 10.
  14. For cold and heat stimuli, cool or heat the water beaker to slightly below (for cold) or above (for heat) the desired temperature and then, start scanning using a plastic transfer pipette. Apply the stimulus with the transfer pipette immediately after scan 5 to the ipsilateral area just below the eye, above the mouth (for V2), or just below the ear (for V3).
    NOTE: Ensure the temperature is within 1 °C of the desired temperature for scan 5. To avoid desensitizing the neurons, do not apply the stimulus if the beaker temperature is not correct. Instead, cool or warm the water again and try again.
  15. At the end of the experiments, inject 50 mM potassium chloride subcutaneously in the V2 or V3 areas starting on cycle 6 to activate and identify all responsive neurons.
  16. After all stimuli have been administered and recorded, euthanize the animal by overdose of ketamine/xylazine (300 mg per kg of ketamine, 30 mg per kg of xylazine, or 5 µL per g of the solution prepared in step 1.1), and then decapitate.

3. Protocol for analysis

  1. Open the orthogonal projection file by using the drag and drop option. Select the image type from Image | Type | RGB Color. Optional: Use the StackReg31 plugin under Plugins | jars | StackReg to correct motion artifacts and aligning.
    NOTE: The investigator can choose the image type based on preference. RGB simplifies the creation of color images for publication.
  2. Open the ROI Manager using Analyze | Tools | ROI Manager.
  3. Select active neurons by drawing an ROI using the Ellipse or Rectangle tool in the toolbar and then place it in the ROI file by clicking the Add button in the ROI Manager window or by pressing the "t" key.
    NOTE: Save ROI files frequently. ROIs can be restored by dragging and dropping the ROI file into ImageJ and clicking the Show All box at the bottom of the ROI Manager window. Saving an ROI .zip file simplifies analysis compared to saving ROIs as an overlay. A neuron that fluctuates during the baseline period should be excluded as spontaneously active, rather than as responding to the stimulus.
  4. In Analyze | Measurement Settings, check Mean Gray Value (optional, but recommended: uncheck all other boxes under Measurement Settings).
  5. Use the ROI window in More | Multi-Measure to measure intensity.
    NOTE: Sometimes, adjacent neurons are too close to draw a separate ROI. These neurons can be included in the active neurons count but cannot be included in transient intensity measurements.
  6. Multi-measure generates a CSV file in a new window labeled "Results." Save the CSV file by clicking on File | Save As. Open the .csv file with spreadsheet software and save the file as a spreadsheet.
    NOTE: See Supplemental File 1 for an analysis template.
  7. Calculate the Ca2+ transient intensity as ΔF/F0 = (Ft- F0)/F0, where Ft is the pixel intensity in the ROI at the time point of interest and F0 is the baseline intensity determined by the mean intensity of the 2-4 frames preceding the Ca2+ transient for spontaneous activity or the first 1-5 frames for stimulus-induced Ca2+ transients. Exclude all neurons that generate Ca2+ peaks before stimulation and do not analyze activity after stimulation.
  8. For Ca2+ transient intensity analysis, randomly sample approximately the same number of neurons from each ganglion to prevent the ganglia that generates the largest number of responding neurons from dominating the analysis. Exclude peaks with ΔF/F0 < 0.15.
    NOTE: There are alternative published methods for analyzing and including and excluding neurons15,16,19,22,32,33,34,35.
  9. Measure neuron diameter by calculating the average length of a line drawn in ImageJ along the longest and shortest diameters using the Line tool in the toolbar.
    NOTE: An investigator can also measure the area and elliptical parameters of ROIs to estimate the diameter.

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Results

Imaging of large numbers of neurons and specific trigeminal branches by confocal scanning of the Pirt-GCaMP trigeminal ganglion
After surgical TG exposure in Pirt-GCaMP3 mice, confocal microscopy allows imaging of over >3,000 neurons simultaneously (Figure 4). This provides the powerful advantage of observing thousands of TG neurons simultaneously in their normal physiological context. Spontaneous Ca2+ transients can be monitored in the absence of stimulati...

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Discussion

Pain disorders of the temporomandibular joint and apical periodontitis affect approximately 5%36 and 30-70%37,38 of the general population, respectively. Primary sensory neurons detect noxious stimuli in the skin, oral mucosa, teeth, and periorbital regions, and their plasticity contributes to persistent pain. Cell culture and explants can be used to study neurons but are removed from their normal physiological context. Pirt-GCaMP3 calcium...

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Disclosures

The authors declare no competing financial interests.

Acknowledgements

This research was supported by National Institutes of Health Grants R01NS0128574 and R01DE031477 (to Y.S.K.), and a Rising STAR Award from the University of Texas system (Y.S.K.).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
Anased Injection (Xylazine)Covetrus, Akorn33197
Flurar 5x/0.25 M27 obejctiveCal Zeiss420130-9900-000
Cotton Tipped ApplicatorsMcKesson24-106-1S
Curved HemostatFine Science Tools13007-12
DC Temperature ControllerFHC40-90-8D
DC Temperature Controller Heating PadFHC40-90-2-05
Dumont Ceramic Coated ForcepsFine Science Tools11252-50
FHC DC Temperature ControllerFHC40-90-8D
Fluriso (Isoflurane)MWI Animal Health, Piramal Group501017
Friedman-Pearson RongeursFine Science Tools16221-14
GelFoamPfizer09-0353-01
ImageJNational Institutes of Healthij153
Ketaset (Ketamine)ZoetisKET-00002R2
Model 923-B Mouse Gas Anesthesia Head Holder (Adult mouse)Kopf Instruments923-B
Luminescent Green Stage TapeJSITON/ AmazonB803YW8ZWL
M35 Compact Electric Lab Handpiece SystemBuffalo Dental Manufacturing Co., Inc.M35
Matrx VIP 3000 Isoflurane VaporizerMidmark91305430
Micro dissecting scissorsRobozRS-5882
Micro dissecting spring scissorsFine Science Tools15023-10
Micro dissecting spring scissorsRobozRS-5677
Mini Rectal Thermistor ProbeFHC40-90-5D-02
Operating scissorsRobozRS-6812
Pirt-GCaMP3 C57/BL/6J miceJohns Hopkins UniversityN/AEither sex can be imaged equally well. Mice should be at least 8 weeks old due to weak or intermittent Pirt promoter expression in younger mice.
Potassium chlorideFisher ScientificBP366-500
Stereotaxic frameKopf Model 923-B923-B
Sterotactic mask (active) for mice <30g with tubesRWD Life Sciences68663
Taper Fissure TNT Bur (31P)Buffalo Dental Manufacturing Co., Inc.31P-GOLD
td-Tomato C57BL/6J miceJackson Laboratory7909
Top Plate, 6in x 10inNewport290-TP
Transfer pipette, 3mLCole ParmerUX-06226-23
TrpV1-Cre C57BL/6J miceJackson Laboratory17769
Zeiss LSM 800 confocal microscopeCal ZeissLSM800
Zeiss Zen 2.6 Blue Edition SoftwareCal ZeissZen (Blue Edition) 2.6

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