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Method Article

Head Implants for the Neuroimaging of Awake, Head-Fixed Rats

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DOI:

10.3791/64324

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September 7th, 2022

 ,  ,  , 
1Department of Neurobiology and Behavior, School of Biological Sciences, University of California Irvine, 2Department of Biomedical Engineering, School of Engineering, University of California, Irvine, 33Center for Neurobiology of Learning and Memory, University of California, Irvine

In This Article

Summary

A detailed new procedure for functional imaging of awake, head-fixed rats is described.

Abstract

Anesthetics, commonly used in preclinical and fundamental scientific research, have a depressive influence on the metabolic, neuronal, and vascular functions of the brain and can adversely influence neurophysiological results. The use of awake animals for research studies is advantageous but poses the major challenge of keeping the animals calm and stationary to minimize motion artifacts throughout data acquisition. Awake imaging in smaller-sized rodents (e.g., mice) is very common but remains scant in rats as rats are bigger, stronger, and have a greater tendency to oppose movement restraints and head fixation over the long durations required for imaging. A new model of neuroimaging of awake, head-fixed rats using customized hand-sewn slings, 3D-printed head implants, head caps, and a headframe is described. The results acquired following a single trial of single-whisker stimulation suggest an increase in the intensity of the evoked functional response. The acquisition of the evoked functional response from awake, head-fixed rats is faster than that from anesthetized rats, reliable, reproducible, and can be used for repeated longitudinal studies.

Introduction

Most of the basic, preclinical, and translational scientific neuroimaging investigations are acquired from anesthetized animals1,2. Anesthetics ease experimentation but continuously influence the brain's and body's metabolism, blood pressure, and heart rate3. The type of anesthetic and the duration and route of administration add confounding variables to data interpretation that could contribute to reproducibility and translational failures4. A major bottleneck of awake, head-fixed rat neuroimaging studies is the requirement to keep the rat stationary and calm throughout the preparation and data acquisition processes. Small movements produce unwarranted motion artifacts, which can adversely affect data analysis and interpretations.

A new model of neuroimaging from awake, head-fixed rats using customized slings, three-dimensional (3D)-printed head implants, head caps, and a headframe has been devised that offers several advantages for easy experimentation. The 3D head implant is light and covers a small portion of the skull needed for transfixing. The 3D-printed head implants and caps are designed using computer-aided design (CAD) software. The protocols of whisker stimulation, data acquisition, data analysis, and results from anesthetized rats have been described in detail in previous work5,6,7.

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Protocol

All procedures were compliant with National Institute of Health guidelines and approved by the University of California, Irvine Animal Care and Use Committee. Seven males and one female rat (Sprague-Dawley, weight: 185-350 g) were used in this study. After study completion, the rats were sacrificed using carbon dioxide overdose.

1. Design of different components

  1. Design of the head implant:
    1. Make the head implant using CAD software (Figure 1C) and design it to image the area posterior to the bregma and adjacent to the midline centered on the somatosensory cortex. Ensure that the head implant covers an area of 0.9 mm to 1.9 mm on the skull away from the imaging area.
    2. Use only three screws to anchor the head implant on the rat's skull. Design all the screw holes so that they remain on the opposite side of the midline in the contralateral hemisphere of the imaged hemisphere.
    3. Place a bar, hollowed from the inside, in the upper part of the head implant to allow wires to fix the head cap to the head implant as shown in Figure 1D.
  2. Design of the head cap:
    1. Ensure that the head cap covers the imaging area completely and protects it from any sort of trauma as shown in Figure 1A, B. Add a curvature to the head cap so that it aligns to the shape of the head without causing difficulty to the animal's daily activities in the standard enriched cages.
    2. Cut the inner side of the head cap in a wider rectangular shape so that the upper part of the head implant can fit into it as shown in Figure 1E. Perpendicular to this rectangle, cut two other rectangular regions to anchor the head cap to the head implant.
    3. Pass one wire through the upper hollowed bar of the head implant for fixation of the head cap on the rat head as shown in Figure 1E-G. Pass the second wire in the same way.
      NOTE: These wires can be easily removed using pliers or forceps. The 3D printing files are provided (file format: STL) as Supplemental File 1 and Supplemental File 2.
  3. Design of the head frame:
    1. Design the head frame in a way that one cut part can move through the upper bar of the head implant and is fixed using a clamp.
    2. Angle the other cut part to provide extra strength for keeping the rat head-fixed to make the contralateral side completely accessible for imaging. For the purpose of this study, cut the steel plate with tin snips to produce the head frame (Figure 1H, I).
      NOTE: This part can be 3D printed as well.

2. Initial rat training

  1. Allow rats to acclimate to the vivarium environment in their cages for 2-3 days.
  2. Start handling the rat in a quiet room. Open the cage and have the experimenter put their hand inside the cage near the rat for 15-20 min to let the rat get habituated.
  3. Once the rat displays calmness by not getting startled or running away from the experimenter's hands, gently pick the rat up for handling. Handle the rat for 30-45 min each day before sling training.

3. Sling training

  1. Train the rats for at least 2-3 days in the slings before the surgical implantation of the head implant and head cap.
  2. Arrange the sling setup as shown in Figure 2A. Clean the sling setup using ethanol wipes.
    NOTE: All the slings are hand-sewn and made of a netting material either on the bottom or on both sides as shown in Figure 2A, B.
  3. For sling training, anesthetize the rats using 4% isoflurane for induction and 1% for maintenance until there is no hind paw pinch reflex.
  4. Under isoflurane anesthesia, place the rats on a flexible plastic sheet measuring 20 cm x 8 cm (length x width), where 10 cm x 8 cm of the plastic sheet is fully covered with the softer part of the Velcro.
    NOTE: Anesthetizing the rats for sling training is an optional step, primarily used to reduce stress and anxiety.
  5. For the first 2 days of the training, put the rat snuggly into a baby sock (size 0-3 months) with the head out through a small hole incised at the end of the sock.
  6. Wrap a small piece of absorbent pad around the lower body part to keep the rat dry and collect excrements.
  7. Wrap the rat in a breathable cotton cloth (size: 25 cm x 25 cm). Place the rat on a plastic sheet that has Velcro strips glued to it.
  8. Further secure the rat to the plastic sheet using 0.5 cm wide Velcro strips at a distance of 3-6 mm from each other.
  9. Secure the rat in the sling. Remove the gas anesthesia. Allow the rat to recover from gas anesthesia in the sling.
  10. When the rat starts whisking, offer a few drops of 10% sucrose solution as a reward every 10-15 min.
  11. Randomly present the rat with the sensory stimuli that will be used during imaging (here whisker stimulation, every 15-25 min) to make it accustomed to sensory stimuli. Manually stimulate the whiskers at random intervals.
  12. Train the rat in the sling for 1 h on day 1, 2 h on day 2, and 3 h on day 3 as shown in Figure 2C.

4. Presurgical preparation

  1. Print the head implant and head cap using the 3D printer (Figure 1).
  2. Sterilize all the surgical instruments and headpieces (implants and caps) by immersing the equipment in the Metricide28 germicide for 10 hours. Rinse tools thoroughly with sterile water just before surgery.
  3. Expose the rat to 4% isoflurane and then maintain at 1%-2% isoflurane until there is no hind paw pinch reflex. This surgery can be performed under many types of anesthesia, such as isoflurane, sodium pentobarbital, and ketamine-xylazine.
  4. Inject atropine (0.05 mg/kg) intramuscularly to reduce mucous secretions to help in breathing.
  5. Shave the head of the rat 5 mm centered around the midline using a hair trimmer starting from between the eyes to the back of the ears.
  6. Monitor the partial oxygen saturation and heart rate through a pulse oximeter and heart rate monitor probe secured to the hind leg of the rat.
  7. Wipe the rat’s head and the surrounding area three times with alternating rounds of betadine and 70% alcohol wipes.
  8. Fix the rat in a stereotaxic system.
  9. Insert a petroleum jelly-lubricated rectal probe to measure the rat's body temperature and maintain it through the heating blanket's feedback system to avoid hypothermia after anesthetic administration.
  10. Administer local anesthetic lidocaine hydrochloride at a concentration of 20 mg/ml, 0.07 mg/kg +/-0.2 body weight subcutaneously at the surgical site.
  11. Apply ophthalmic ointment to both eyes to prevent drying.
  12. Administer 2% local anesthetic subcutaneously over the surgical site.
  13. Inject 3 mL of lactated ringer's solution at room temperature subcutaneously to prevent dehydration and provide nourishment during surgery.

5. Surgery

  1. Remove the part of the skin over the surgical site (4 mm diameter centered around the midline and center of the head) using sharp surgical scissors. Dissect and remove part of the skin (~2 mm diameter, over the left somatosensory cortex) between the ear and eye on the temporal part of the head.
  2. Remove, using a scalpel, the underlying skin (pericranium) tissue to expose the skull. Clean the skull using sterilized cotton gauze.
  3. Retract/resect temporal muscle to expose desired size for imaging area [7.5 mm by 7.5 mm for this study]. 
  4. Expose the skull on the contralateral hemisphere for the head implant. Place the head implant on the skull to ascertain the location of anchoring screws for the implant as shown in Figure 2D-F.
  5. Mark the skull for drilling the screws using India Ink with drill bit 1. Drill the burr holes for the screws using dental drill bit 3. Screw the head implant in place.
  6. Dry the skull using sterile gauze. Apply a thin layer of tissue adhesive around and beneath the head implant to glue it to the skull. Apply a layer of dental cement to further support the head implant in place and let the cement dry for 2-3 min.
    NOTE: The use of tissue adhesive in addition to dental cement ensures a strong hold8.
  7. Using dental drill bit 3, thin a 7.5 mm x 7.5 mm area on the left side of skull just posterior to the bregma and lateral to the midline. Thin the skull to ~50 µm as shown in Figure 3A.
  8. Apply topical antibiotic ointment over the surgical site and then cover it with a thin layer of silicone rubber to protect the thinned skull as shown in Figure 3B. Cover the surgical site using the head cap as shown in Figure 3C. Fix it in place with the two small pieces of wires going through both the head implant and head cap as shown in Figure 3D, E. Apply silicone rubber to cover the head cap and skull to stabilize the head cap further on the rat's head as shown in Figure 3F.
    NOTE: Silicon rubber provides additional protection to thinned skull.
  9. Inject the rat with flunixin meglumine (2.5 mg/kg) subcutaneously for pain and inflammation management. To prevent infection, inject Enrosite antibiotic enrofloxacin (22.7mg/ml, 10mg/kg +/-.01), intraperitoneally.
  10. Move the rat to the recovery chamber to help maintain its body temperature with a warming blanket and a heat lamp. Monitor the rat continuously until it regains consciousness and can maintain sternal recumbency.
  11. Return the rat to its separate cage once it fully recovers.
  12. For the next 3 days, administer flunixin and buprenorphine to alleviate inflammation and pain and enrosite to prevent infection twice daily.

6. Awake imaging

  1. Anesthetize the rat with 4% isoflurane for induction and 1% for maintenance when there is no hind paw pinch reflex. Inject acepromazine (0.3-0.5 mg/kg) subcutaneously.
    NOTE: This concentration of acepromazine is below mild sedation levels and only helps keep the rats calm throughout the imaging process.
  2. Using customized strips of Velcro, fix the rat on the plastic sheet used during the training procedures. Wrap the lower body part using an absorption pad and place the rat snugly in the sling.
  3. Remove the silicon rubber. Remove the head cap by removing the fixation wires. Fix the headframe in the head implant as shown in Figure 2G.
  4. Lock the headframe in clamps as shown in Figure 2H, I.
  5. Remove gas anesthesia. Flush the imaging area with saline 3x and clean with wet gauze. Dry the imaging area and make a well, using petroleum jelly, around the imaging area. Fill the well with sterilized saline and cover with a glass slide (Figure 2E).
  6. Refer to the acquisition procedures for intrinsic signal optical imaging, the whisker stimulation protocol, and data analysis and presentation, which have been discussed in detail previously6,7.
  7. Throughout the experiment, monitor the rats for signs of agitation and restlessness, which can be further reduced by covering the eyes of the rats with a soft cloth or gauze (optional).

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Results

The representative optical imaging signals from a single trial of an anesthetized rat and the summed response (of 40 collected trials) of an awake rat are shown (Figure 4). The signal intensity for single-whisker stimulation of an awake rat can be visualized at a higher threshold than for the anesthetized rat, showing a stronger signal from the awake animal. The C2 whiskers of rats are stimulated at 5 Hz for 1 s, and the functional response is displayed as a fractional change compared to the...

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Discussion

The use of awake, head-fixed rat imaging offers many advantages in terms of ease and customization. The custom-designed slings allow the rats to be wrapped through breathable netting material, eliminating the need to enclose animals in closed, plastic restraining chambers for extended periods of time10,11. Rats are kept calm and stress-free throughout the long durations of successive imaging sessions using a very low dose of acepromazine below the levels of mild ...

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Disclosures

The authors have no conflicts of interest to disclose.

Acknowledgements

We acknowledge Clara Jones, James Stirwalt, Linh Hoang, Young Joon Ha, and Amirsoheil Zareh for their help during training of the rats and preparation of the slings. Funding was provided by the National Institutes of Health (NIH, Grant Number: NS119852) and Leducq Foundation (Grant Number:15CVD02).

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Materials

List of materials used in this article
NameCompanyCatalog NumberComments
RatsCharles RiverSprague Dawley
IsofluranePivetal21295098General anesthetic
Lidocaine HCl 2% injectionPhoenixL-2000-04Local anesthetic
Atropine sulfate injectionVedco5098907512Help in respiration
Lactated Ringer's injection solutionVedco50989088317
Flunixin injectionVedco6064408670Pain management
Enrosite injection (Enrofloxacin 2.27%)VetOne501084Avoid infection
PromAce injection (Acepromazine maleate)Beohringer Ingelheim136059
Animax ointmentDechra Veterinary Products122-75active ingredients of nystatin 1000units per gram, neomycin sulfate 2.5mg per gram, thiostrepton 2500 units per gram, and triamcinolone acetonide 1mg per gram
Puralube ophthalmic ointmentDechra Veterinary Products211-38
Povidone-iodine PVP prep padsMedlineMDS093917Betadine generic
Isopropyl alcohol swabsBD326895
Vetbond tissue adhesive3M1469SB
Bur (drill bit), standard operatory carbideSS White Burs14829#3 bur
Screws, 00-90 x 1/8 flat head stainless steelJ.I. MorrisF0090CE125Anchor screws
Stereotaxic systemKopf Instruments1430
Homeothermic heating blanketHarvard Apparatus50-7220-F
Pulse oximeter & heart rate monitorKent ScientificMouseStat Jr.
PetrolatumFisher ScientificP66-1LBVaseline generic
Wire, bare copperFisher Scientific15-545-2C20 gauge
Teets Cold Cure powderPearson DentalC73-0054 active ingredient: Methyl Methacrylate
Teets Cold Cure liquidPearson DentalC73-0078 active ingredient: Methyl Methacrylate
Silicone mold rubberSmooth-OnBody Double Fastsilicon polymer
Metricide 28 (Germicide)MetrexOct-05
India ink, blackPelikan301051
Dental drillNSK DentalUltimate XL-F
3D printerPrusa Researchi3 MK3S+
Sew on fastenersVelcro90030
Pet screening utility fabricJoann10173334Netting material
Bur (drill bit), standard operatory carbideSS White Burs14829#1 bur

References

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Awake ImagingNeuroimaging RatsFunctional ResponseThin Skull Imaging3D Printed ImplantsStereotaxic SystemCalcium ImagingFunctional Ultrasound
Head Implants for the Neuroimaging of Awake, Head-Fixed Rats
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